Colloids and Surfaces B: Biointerfaces 123 (2014) 191–198
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Osteoblast activity of MG-63 cells is enhanced by growth on a lactoferrin-immobilized titanium substrate Sung Eun Kim a,1 , Young-Pil Yun a,1 , Jae Yong Lee b , Kyeongsoon Park c , Dong Hun Suh d,∗ a Department of Orthopedic Surgery and Rare Diseases Institute, Korea University Medical College, Guro Hospital, #80, Guro-dong, Guro-gu, Seoul 152-703, Republic of Korea b Department of Biomedical Science, College of Medicine, Korea University, Anam-dong, Seongbuk-gu 136-701, Republic of Korea c Division of Bio-imaging, Chuncheon Center, Korea Basic Science Institute, 192-1 Hyoja 2-dong, Chuncheon, Gangwon-do 200-701, Republic of Korea d Department of Orthopedic Surgery, Korea University Medical College, Ansan Hospital, Gojan 1-dong, Danwon-gu, Gyeonggi-do 425-707, Republic of Korea
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Article history: Received 25 March 2014 Received in revised form 25 August 2014 Accepted 5 September 2014 Available online 23 September 2014 Keywords: Titanium Lactoferrin Heparin–dopamine Osteoblast Osteogenic differentiation
a b s t r a c t The aim of this study was to develop a lactoferrin (LF)-immobilized titanium (Ti) substrate to enhance the osteoblast activity of MG-63 cells. Ti substrates were first modified through heparin–dopamine (Hep–DOPA) anchorage. Then, LF was immobilized on the Hep–Ti substrates via electrostatic interactions. Hep–Ti substrates, with or without LF, were evaluated by scanning electron microscopy (SEM), X-ray photoelectron spectroscopy (XPS), and contact angle measurements. Sustained release of LF on the Ti substrates was observed over a 28-day period. In vitro studies of osteoblast activity showed increased alkaline phosphatase activity and calcium deposition by MG-63 cells cultured on LF-immobilized Ti substrates as compared to those cultured on pristine Ti substrates, indicating that LF-immobilized Ti substrates were effective at enhancing osteoblast activity. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Titanium (Ti) and its alloys are widely used in the orthopedic and dental fields due to their low toxicity, excellent mechanical properties, and low corrosive properties [1–3]. Despite these excellent properties, Ti alone does not promote osseo-integration with the host bone at the implant surface. Poor bonding of Ti implants with the host bone can bring about unpredictable bone loosening over time, resulting in implant failure and patient discomfort [4,5]. To promote osseo-integration between the Ti surface and host bone, attempts have been made to modify and functionalize the Ti surface though calcium phosphate/hydroxyapatite (HAp) coating, biomolecule immobilization, and surface topography modification [4,6–9]. Bone morphogenic protein-2 (BMP-2) has been shown to promote osteo-inductive activity, bone regeneration, and cartilage regeneration. Additionally, BMP-2 has been used in combination with various materials such as collagen gels, sponges, hyaluronic acid, dextran, chitosan, fibrin scaffolds, and Ti to promote bone formation and osseo-integration [10–16]. More recently, BMP-2 mixed with components such as collagen sponges (INFUSE Bone
∗ Corresponding author. Tel.: +82 31 412 4945; fax: +82 31 487 9502. E-mail address: [email protected] (D.H. Suh). 1 These authors contributed equally to this paper. http://dx.doi.org/10.1016/j.colsurfb.2014.09.014 0927-7765/© 2014 Elsevier B.V. All rights reserved.
Graft; Medtronic, Minneapolis, MN, USA) and biphasic calcium phosphate (BioC® ; 70% tri-calcium phosphate (TCP): 30% HAp) has been approved by bodies such as the Food and Drug Administration (FDA) and the Korea Food and Drug Administration (KFDA) for operations such as posterior-lateral spine fusions, tibial fractures, and alveolar ridge augmentations in the fields of orthopedic and dental surgery. Although clinical treatment with BMP-2 has been successful, BMP-2 has well-known drawbacks, such as a short in vivo half-life, high cost, and induction of excessive bone formation [17,18]. Lactoferrin (LF) is an 80-kDa iron-binding glycoprotein in the transferrin family of proteins [19]. It is produced by many different exocrine glands, and is widely distributed among body fluids such as milk, saliva, tears, bile, and pancreatic fluid [20]. Previous studies have demonstrated that LF has inhibitory biological effects against tumors, inflammatory processes, bacteria, and fungi [21,22]. In addition, LF has been reported to induce osteoblast and osteoblast cell-line proliferation, as well as osteoblast differentiation [23–25]. LF has also been reported to inhibit osteoblastic cells from apoptosis induced by serum withdrawal [26]. Takayama et al. [25] reported that LF incorporated in collagen membranes promoted alkaline phosphatase (ALP) activity and osteocalcin (OCN) production by human osteoblast-like cells (MG-63 cells). Previous studies have also demonstrated that LF incorporated in gelatin hydrogels significantly induced 3T3E1 (mouse-derived
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Fig. 1. Schematic diagram of the immobilization of lactoferrin (LF) to heparin–dopamine (Hep–DOPA)-coated Ti substrate.
osteoblast) cell proliferation in vitro, and, through sustained release from the hydrogels, enhanced bone regeneration at defect sites in vivo [27]. Furthermore, mesenchymal stem cells treated with biomimetic hydroxyapatite nanocrystals functionalized with LF showed enhanced osteogenic differentiation [28]. More recently, Ying et al. reported that LF promotes proliferation and osteogenic differentiation of human adipose-derived stem cells (hADSCs) [29]. Although the above studies suggest that LF may be a promising therapeutic agent to treat bone diseases or regenerate bone defects, no prior study has investigated the effect of LF-immobilized titanium (Ti) substrates on osteogenesis induction of MG-63 cells. We therefore set out to evaluate the effects of LF-immobilized Ti substrates on MG-63 osteoblast activity, which we did by evaluating F-actin staining, cell proliferation, alkaline phosphatase (ALP) activity, and calcium content. 2. Materials and methods 2.1. Materials Titanium (Ti) discs (diameter × height: 1.2 cm × 0.3 cm) were kindly supplied by Neobiotech Co., Ltd (Seoul, Korea). Heparin (average MW ca. 12,000 Da) was purchased from Pharmacia Hepar Co. (Franklin, OH, USA). Dulbecco’s modified Eagle’s medium (DMEM), fetal bovine serum (FBS), phosphatebuffered saline (PBS), and penicillin-streptomycin were purchased from Gibco BRL (Rockville, MD, USA). Dopamine hydrochloride, lactoferrin (LF, origin: human milk, Cat. No: L4894), 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDAC), and Nhydroxysuccinimide (NHS) were obtained from Sigma Chemical Co. (St. Louis, MO, USA). PierceTM BCA protein assay kit was purchased from Thermo Fisher Scientific (Rockford, IL, USA). MG 63 cells (human osteosarcoma cell line) were obtained from the Korea Cell Line Bank (KCLB No. 21427, Seoul, Korea). All other chemicals were of the purest analytical grade available. 2.2. Immobilization of heparin–dopamine (Hep–DOPA) and lactoferrin (LF) on the surface of titanium (Ti) Prior to preparing functionalized Ti substrates, we modified pristine Ti substrates using Hep–DOPA conjugated materials. In
brief, pristine Ti substrates were immersed in Hep–DOPA (2 mg) and dissolved in 10 mM Tris·HCl (pH 8.0) buffer in the dark, overnight. Heparin-coated Ti (Hep–Ti) substrates were rinsed with distilled water (DW) and dried under N2 . Lactoferrin (LF) was immobilized on the surface of Hep–Ti through electrostatic interaction between the amine groups of LF and the carboxylic/sulfate groups of heparin. Briefly, different amounts of LF (10, 50, or 100 g/mL) and Hep–Ti substrate were immersed in 0.1 M MES buffer (pH 5.6) and gently shaken for 6 hr. LF (10, 50, or 100 g)/Hep–Ti was washed and dried at room temperature (RT) (Fig. 1).
2.3. Characterization of pristine Ti and surface-modified Ti Surface morphologies of pristine Ti and surface-modified Ti were investigated by scanning electron microscopy (SEM, S4800, Hitachi, Japan). Each sample was coated with gold using a sputtercoater (Eiko IB, Japan), and SEM observations were performed at an operating voltage of 3 kV. To confirm the surface chemical compositions of all samples, X-ray photoelectron spectroscopy (XPS) was carried out using a K-alpha spectrometer XPS system (ESCALAB250) and a Theta Probe AR-XPS System (Thermo Fisher Scientific, UK) with an Al K␣ X-ray source (1486.6 eV photons) at the Korea Basic Science Institute Busan Center. The C1s hydrocarbon peak at 284.84 eV was used as the reference for all binding energies. The area of each peak was normalized to the total peak area (for all atomic elements) to calculate surface atom percentage. Hydrophilic properties of pristine Ti and surface-modified Ti were measured by contact angle (SEO Phoenix 300, Suwon, Korea). Toluidine blue was used to quantify the amount of heparin immobilized on the surface of pristine Ti. Briefly, heparin-modified Ti (Hep–Ti) was immersed in 1 mL of 0.2% (v/v) NaCl solution containing 1 mL of 0.005% (w/v) toluidine blue solution. After gentle shaking for 30 min, 2 mL of hexane was added. Hep–Ti was removed from solution, and aqueous-phase absorbance was measured using a micro-plate reader (Bio-Rad, Hercules, CA, USA) at a wavelength of 620 nm. To analyze the amount of LF immobilized on Hep–Ti, a PierceTM BCA protein assay kit was used. In brief, after a 6-hr reaction, LF immobilized on Hep–Ti was removed from the MES buffer and then rinsed with PBS. After the reaction, the amount of
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LF remaining in the MES buffer was measured to indirectly evaluate LF immobilization on the Ti substrate surface. 2.4. Lactoferrin (LF) release study To evaluate the amount of LF released from LF/Hep–Ti in the 10/50/100-g LF conditions, each sample was immersed
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in 1 mL PBS buffer (pH 7.4) and gently shaken at 100 rpm at 37 ◦ C. PBS was collected and replaced with fresh PBS at predesigned time intervals of 1, 3, 5, and 10 h and 1, 3, 5, 7, 14, 21, and 28 days. Concentration of released LF was analyzed with a PierceTM BCA protein assay kit in accordance with the manufacturer’s instructions using a micro-plate reader at 562 nm.
Fig. 2. (A) SEM images and (B) wide-scanned XPS spectra of pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti.
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2.5. Cytotoxicity and F-actin staining Cytotoxicities of all Ti substrates were tested using a cell counting kit-8 (CCK-8) (Dojindo, Japan). This kit is based on a highly water-soluble tetrazolium salt, WST-8 [2-(2-methoxy4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2Htetrazolium, monosodium salt], that produces a water-soluble formazan dye upon reduction in the presence of an electron mediator. Cytotoxicity of both pristine Ti and surface-modified Ti was evaluated via an indirect method. Specifically, 1 × 104 MG-63 cells were seeded on 96-well plates and incubated at 37 ◦ C for 24 h in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 50 g/mL ascorbic acid, 10 nM dexamethasone, and 10 mM -glycerophosphate in the presence of 100 U/mL penicillin and 100 g/mL streptomycin. Pristine and surface-modified Ti samples were incubated at 37 ◦ C for 24 h in DMEM medium to obtain extraction medium. DMEM medium was aspirated over the (pristine Ti or modified Ti) surface containing MG-63 cells before cells were washed with PBS, and fresh DMEM medium added. MG-63 cells were incubated for 1 or 2 days. At predetermined time intervals, the extraction medium was aspirated, CCK-8 solution added, and MG-63 cells were incubated for 1 hour at 37 ◦ C. Live-cell optical density (OD) was measured using a micro-plate reader at 450 nm. Relative cell viability (%) was calculated as [OD]test/[OD]control × 100. To observe the morphology of MG-63 cells adhered on pristine Ti and surface-modified Ti, surfaces with 1 × 105 seeded MG-63 cells were transferred to a 24-well tissue-culture plates. After 24 h of culture, substrates were placed in a cytoskeleton buffer (5 × 10−3 M NaCl, 150 × 10−3 M MgCl2 , 5 × 10−4 M Tris-base and 0.5% triton X100) for 5 min at 4 ◦ C, followed by addition of blocking buffer (5% FBS, 0.1% Tween-20 and 0.02% sodium azide in PBS) for 30 min at 37 ◦ C. Cells/substrates were incubated with rhodamine-phalloidin (1:200) and DAPI (1:10,000) and observed by confocal laser scanning microscopy (LSM700, Zeiss, Germany). 2.6. Cell proliferation CCK-8 kit for cell proliferation assay was also used, because cell proliferation measured using CCK-8 correlates well with results obtained using the [3 H]-thymidine incorporation assay, and thus, the CCK-8 assay can be substituted for the [3 H]-thymidine incorporation assay. MG-63 cells were seeded at a concentration of 1 × 105 cells/mL on the surface of pristine or surface-modified Ti in a 24-well tissue-culture plate. At days 3 and 7 of incubation, substrates were rinsed with PBS, and then CCK-8 solution was added to the substrates followed by a 1-h incubation. Reagents were transferred to 96-well plates. Cell proliferation results are presented as optical density measured at 450 nm using a micro-plate reader.
1 × 105 MG-63 cells were seeded on the surface of pristine Ti, Hep–Ti, or Hep–Ti modified with 10, 50, or 100 g LF in a 24well tissue-culture plate. At day 21 of culture, cells/substrates were rinsed with PBS, and 0.5 N HCl was added. Cells were centrifuged at 13,500 rpm for 1 min. The supernatant was used for calcium content measurements with a QuantiChromTM Calcium Assay Kit (DICA-500, BioAssay Systems, USA) in accordance with the manufacturer’s instructions. Concentration of calcium was measured by a micro-plate reader at 612 nm. 2.8. Statistical analysis Data are presented as mean ± standard deviations (n = 5). Statistical comparisons were carried out by one-way ANOVA using Systat software (Systat Software, Inc., Chicago, IL, USA). Differences were considered statistically significant at * P < 0.05 and ** P < 0.01. 3. Results 3.1. Characterization of pristine Ti and surface-modified Ti Ti morphology, both in the presence of and absence of surface modification, was observed by SEM (Fig. 2A). Adding Hep–Ti or LF (at any concentration, from 10 to 100 g) did not have an observable effect on Ti morphology. XPS measurement was carried out to investigate the surface chemical composition of pristine Ti and surface-modified Ti substrates. As shown in Fig. 2B and Table 1, wide-scanned XPS spectra of pristine Ti, Hep–Ti, and LF/Hep–Ti (at 10, 50 and 100 g LF) were evaluated. Successful heparin anchorage on the surface of pristine Ti was confirmed by an increase in the C1s content from 61.13% to 67.74%, an increase in the S2p content from 0% to 2.12%, and an increase in the N1s content from 2.15% to 3.65% as compared to pristine Ti (Table 1). Successful immobilization of increasing doses of LF on Hep–Ti was confirmed by increases in N1s
Table 1 Percentages of C1s, O1s, N1s, Ti2p, and S2p expressed at the surface of pristine and surface-modified Ti, as analyzed by XPS. Substrate
C1s
O1s
N1s
Ti2p
S2p
Pristine Ti Hep–Ti LF3.6 /Hep–Ti LF17.1 /Hep–Ti LF38.9 /Hep–Ti
61.13% 67.74% 72.19% 72.32% 71.36%
30.07% 21.76% 18.31% 17.18% 17.04%
2.15% 3.63% 5.74% 6.41% 6.97%
6.65% 4.75% 0.82% 0.92% 1.12%
– 2.12% 2.94% 3.17% 3.51%
2.7. Alkaline phosphatase (ALP) activity and calcium deposition ALP activity was measured at days 3, 7, and 10 of culture. Briefly, MG-63 cells were seeded at a density of 1 × 105 cells/mL in a 24well culture plate on the surface of pristine Ti, Hep–Ti, or Hep–Ti modified with 10, 50, or 100 g LF. Cells/substrates were rinsed with PBS at pre-determined times, and 1× RIPA buffer was added to lyse cells. Lysed cells were centrifuged at 13,500 rpm for 1 min. A 100 L p-nitrophenyl phosphate solution was added to the supernatant, and the reaction was allowed to proceed for 30 minute at 37 ◦ C. 1 N NaOH was added to stop the reaction. Optical density was determined at 405 nm using a micro-plate reader. Enzymatic activity was expressed as micromolecules of p-nitrophenol produced per min per g of protein. Protein content was determined using a modified Bradford assay using BSA as the standard. Calcium content was investigated after 21 days of incubation. In brief,
Fig. 3. Release kinetics of lactoferrin (LF) from LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti. Error bars represent mean ± SDs (n = 5).
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Fig. 4. (A) Cytotoxicity of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 1 and 2 days of culture. (B) F-actin staining images of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 24 hr of incubation.
and S2p contents. Mean ± standard deviations of contact angles of pristine Ti, Hep–Ti, and 10, 50, 100 g LF/Hep–Ti were 75.1 ± 1.8◦ , 54.6 ± 2.7◦ , 30.1 ± 2.7◦ , 29.1 ± 3.3◦ , and 26.1 ± 2.9◦ , respectively. 5.47 ± 0.64 g/sample of heparin was grafted on Ti (loading
efficiency (LE): 0.27%). Amounts of LF in the 10, 50 and 100 g LF/Hep–Ti cases were 3.6 g (LE: 36.0%), 17.1 g (LE: 34.2%), and 38.9 g (LE: 38.9%); these samples were designated as LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti, respectively.
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Fig. 5. Cell proliferation of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 3 and 7 days of incubation. Error bars represent mean ± SDs (n = 5). * P < 0.05 and ** P < 0.01.
3.2. In vitro LF release kinetics Fig. 3 shows the in vitro release kinetics of LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti as measured by the PierceTM BCA protein assay. LF exhibited sustained release kinetics for up to 28 days. On the first day, 1.65 ± 0.61 g, 4.53 ± 0.75 g, and 9.18 ± 0.40 g of LF was released from LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti, respectively. At 28 days, cumulative amount of LF released from LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti was 3.07 ± 1.16 g, 11.87 ± 0.98 g, and 22.98 ± 1.49 g, respectively, indicating that the amount of LF remaining was 14.8%, 30.6%, and 40.9%, respectively. 3.3. Cytotoxicity test and F-actin staining MG-63 cell viability on pristine Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti is shown in Fig. 4A. Cell viability for all groups was over 95% for incubation periods of 24–48 h, indicating that the Ti substrates were not cytotoxic to MG-63 cells. F-actin staining after 24 h of incubation revealed typical long and straight actin stress fibers on all Ti substrates (Fig. 4B). Long and straight actin stress fiber structures on the three LF/Hep–Ti substrates were more developed than those on pristine Ti and Hep–Ti substrates. Approximately two-fold more MG-63 cells adhered to the LF38.9/Hep–Ti substrate than the pristine Ti and Hep–Ti substrates. 3.4. Cell proliferation MG-63 cell proliferation on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti was evaluated at days 3 and 7 of incubation (Fig. 5). For each substrate, MG-63 cell proliferation increased gradually in a time-dependent manner. There were no significant differences in cell proliferation between MG-63 cells grown on pristine Ti and those grown on Hep–Ti at days 3 and 7. However, there were significant differences in MG-63 cell proliferation on LF38.9 /Hep–Ti as compared with pristine Ti at day 3 and day 7 (* P < 0.05 or ** P < 0.01). At day 7 of incubation, cell proliferation on the Ti substrates containing LF was significantly different from that on pristine Ti (* P < 0.05 and ** P < 0.01). 3.5. Alkaline phosphatase (ALP) activity and calcium deposition Fig. 6A shows ALP activity in MG-63 cells cultured on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti, for
incubation periods of up to 10 days. After 3 days, no significant differences in ALP activity between pristine Ti and surface-modified Ti were evident. However, there was a significant difference in the ALP activity of MG-63 cells between LF38.9 /Hep–Ti and pristine Ti at 7 and 10 days (* P < 0.05 and ** P < 0.01). ALP activity was 1.05 ± 0.05 for pristine Ti, 1.10 ± 0.08 for Hep–Ti, 1.24 ± 0.07 for LF3.6 /Hep–Ti, 1.36 ± 0.09 for LF17.1 /Hep–Ti, and 1.53 ± 0.06 M/min/g protein for LF38.9 /Hep–Ti, respectively. At 10 days, significant differences in ALP activity were observed between MG-63 cells cultured on Ti substrates containing LF and those cultured on pristine Ti (* P < 0.05 and ** P < 0.01). The amount of calcium deposited by MG-63 cells cultured on all Ti substrates was assessed after 21 days of culture (Fig. 6B). Amount of deposited calcium was 10.32 ± 1.0 for pristine Ti, 11.26 ± 1.0 for Hep–Ti, 14.52 ± 0.8 for LF3.6 /Hep–Ti, 16.30 ± 1.6 for LF17.1 /Hep–Ti, and 19.2 ± 1.6 g for LF38.9 /Hep–Ti. Calcium deposition by MG-63 cells cultured on Ti substrates containing LF was significantly higher than for MG-63 cells grown on pristine Ti (* P < 0.05 and ** P < 0.01). Moreover, a significant difference in the amount of calcium deposited by MG-63 cells grown on LF38.9 /Hep–Ti versus LF3.6 /Hep–Ti was observed (* P < 0.05). However, the amount of calcium deposited by MG-63 cells cultured on LF38.9 /Hep–Ti and LF17.1 /Hep–Ti was not significantly different after 21 days of culture.
4. Discussion Ti implant materials should ideally provide not only a good environment for osteoblast adhesion, proliferation, and differentiation, but also an environment that promotes interaction and osseo-integration between implants and host bone. However, failed Ti implants have been shown to lack implant/host bone interactions and have poor healing capabilities. To address these problems, Ti implants have been modified by surface modification and immobilization with biomolecules and bioactive proteins to improve osteoblast activity and osseo-integration [16,30–33]. The main objective of this study was to develop a lactoferrin (LF)immobilized Ti substrate to enhance osteoblast activity. To prepare LF-immobilized Ti substrate, the surface of the Ti substrate was first functionalized with Hep–DOPA in a slightly basic environment by chemical immobilization between oxidized catechol groups and the Ti surface, mimicking a mussel adhesion mechanism [34–36]. Surface functionalization of Ti with heparin has several advantages. Because of their highly negative charge and high binding affinities for growth factors, heparin molecules can facilitate the immobilization of growth factors on the surface of Ti substrate through electrostatic interactions between the growth factors and heparin, and can minimize rapid diffusion and regulate the sustained release of growth factors or drugs from substrates to improve their efficacies [16,32,37–40]. Indeed, we confirmed that LF was successfully immobilized on the surface of Hep–Ti substrates and that the amount of LF immobilized increased with increasing feed amounts of LF. As reported in previous in vitro release studies, Hep–Ti substrates effectively prevented high initial burst effects and also showed sustained and extended release of LF for 28 days [16,41,42]. Furthermore, all Ti substrates examined in this study were non-toxic to MG-63 cells, indicating that they are safe to use. Initial adhesion of osteoblast cells to the substrate is important for long-term stability and cell differentiation [43]. Furthermore, promotion of cell attachment by the substrate affects the proliferation capacity of cells [43]. Compared to osteoblast cells cultured on pristine Ti and Hep–Ti substrates, osteoblast cells grown on LF/Hep–Ti substrates were more spread out with numerous filopodia in all directions. In particular, cells cultured on LF38.9 /Hep–Ti, which was the substrate with the highest concentration of LF,
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Fig. 6. (A) Alkaline phosphatase (ALP) activity of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 3, 7, and 10 days of incubation. (B) Amount of calcium deposited by MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 21 days of incubation. Error bars represent mean ± SDs (n = 5). * P < 0.05 and ** P < 0.01.
had a flattened, mature, spreading morphology with scattered focal adhesion that increased contact between the cells and the LF38.9 /Hep–Ti surface, indicating that LF/Hep–Ti substrate promoted osteoblast cell attachment. Consistently, osteoblast cell proliferation increased gradually with increasing amount of LF immobilized on the Hep–Ti substrates. Our cell adhesion and proliferation results indicate that the LF-immobilized Ti substrates promoted cell adhesion and proliferation. It is possible that the presence of heparin on Ti substrate also affected cell attachment and proliferation. However, as we reported in our previous studies [16,32], cell adhesion and proliferation of osteoblasts on Hep–Ti substrate were not significantly different compared to those on pristine Ti. This may be because the amount of heparin immobilized on the Ti substrate in this study is too low to affect cell adhesion and proliferation. Three principal osteoblast differentiation processes can be recognized: proliferation, extracellular maturation, and mineralization [44]. Each stage is characterized by the expression of distinctive osteogenic markers. ALP activity is an early marker of osteoblastic differentiation and commitment of stem cells to the osteoblastic phenotype [3,43,45–47]. After osteogenic differentiation, cells begin to secrete a mineral matrix, leading to calcium deposition, which is a marker of mature osteoblasts [16,32,33]. In the present study, we compared the osteogenic activity of osteoblast-like cells (MG-63) cultured on LF-immobilized Ti substrates to those cultured on pristine Ti and Hep–Ti substrates. ALP activity and calcium content of osteoblasts cultured on pristine Ti and Hep–Ti without LF were not significantly different during the experiment periods. However, ALP activity and calcium content of osteoblasts cultured on LF/Hep–Ti substrates increased gradually in time- or dose-dependent manners. It has been reported that LF may stimulate osteogenic differentiation of osteoblast cells via receptor-mediated mechanisms. Among various receptors, lowdensity lipoprotein receptor-related protein 1 (LRP1) is the only receptor that can transduce the LF signal in osteoblasts. LF stimulates osteoblast differentiation mainly through LRP-1 independent protein kinase A (PKA) and p38 signaling pathways [48]. In addition, osteoblast proliferation and differentiation induced by LF appear to be mutually independent intracellular events that are regulated by the ERK1/2 and p38 pathways, respectively [49]. We demonstrated in this study that LF-immobilized Hep–Ti substrates significantly enhanced MG-63 cell proliferation, osteoblast ALP activity, and calcium deposition. These results suggest that LF-immobilized Hep–Ti
substrates are promising alternative Ti implants for regeneration of bone defects. 5. Conclusions We successfully developed LF-immobilized Ti substrates following Hep–DOPA anchorage. LF immobilized on Ti substrates showed sustained release over a 28-day period. LF-immobilized Ti substrates enhanced osteoblastic activity of MG-63 cells as compared to pristine Ti substrates, as shown by a significant increase in ALP activity and calcium deposition. These results suggest that LFimmobilized Ti substrates could effectively promote regeneration of bone defects. Further in vivo studies are needed to investigate the effects of dose-dependently-increasing the amount of LF immobilized to the Ti substrate. Acknowledgement This study was supported by a grant from Korea University (K1300081). References [1] D. Buser, U.C. Belser, N.P. Lang, The original one-stage dental implant system and its clinical application, Periodontology 2000 17 (1998) 106–118. [2] S.G. Steinemann, Titanium—the material of choice? Periodontology 2000 17 (1998) 7–21. [3] L. Zhu, X. Ye, G. Tang, N. Zhao, Y. Gong, Y. Zhao, J. Zhao, X. Zhang, Biomimetic coating of compound titania and hydroxyapatite on titanium, J. Biomed. Mater. Res. Part A 83 (2007) 1165–1175. [4] Z. Shi, K.G. Neoh, E.T. Kang, C.K. Poh, W. Wang, Surface functionalization of titanium with carboxymethyl chitosan and immobilized bone morphogenetic protein-2 for enhanced osseointegration, Biomacromolecules 10 (2009) 1603–1611. [5] D.A. Puleo, A. Nanci, Understanding and controlling the bone-implant interface, Biomaterials 20 (1999) 2311–2321. [6] F. Rupp, L. Scheideler, N. Olshanska, M. de Wild, M. Wieland, J. Geis-Gerstorfer, Enhancing surface free energy and hydrophilicity through chemical modification of microstructured titanium implant surfaces, J. Biomed. Mater. Res. Part A 76 (2006) 323–334. [7] S. Piskounova, J. Forsgren, U. Brohede, H. Engqvist, M. Stromme, In vitro characterization of bioactive titanium dioxide/hydroxyapatite surfaces functionalized with BMP-2, J. Biomed. Mater. Res. Part B: Appl. Biomater. 91 (2009) 780–787, Applied biomaterials. [8] K. Oya, Y. Tanaka, H. Saito, K. Kurashima, K. Nogi, H. Tsutsumi, Y. Tsutsumi, H. Doi, N. Nomura, T. Hanawa, Calcification by MC3T3-E1 cells on RGD peptide immobilized on titanium through electrodeposited PEG, Biomaterials 30 (2009) 1281–1286.
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[9] R.J. Miron, C.J. Oates, A. Molenberg, M. Dard, D.W. Hamilton, The effect of enamel matrix proteins on the spreading, proliferation and differentiation of osteoblasts cultured on titanium surfaces, Biomaterials 31 (2010) 449–460. [10] G.N. King, N. King, A.T. Cruchley, J.M. Wozney, F.J. Hughes, Recombinant human bone morphogenetic protein-2 promotes wound healing in rat periodontal fenestration defects, J. Dental Res. 76 (1997) 1460–1470. [11] G. Zellin, A. Linde, Importance of delivery systems for growth-stimulatory factors in combination with osteopromotive membranes. An experimental study using rhBMP-2 in rat mandibular defects, J. Biomed. Mater. Res. 35 (1997) 181–190. [12] J. Kim, I.S. Kim, T.H. Cho, K.B. Lee, S.J. Hwang, G. Tae, I. Noh, S.H. Lee, Y. Park, K. Sun, Bone regeneration using hyaluronic acid-based hydrogel with bone morphogenic protein-2 and human mesenchymal stem cells, Biomaterials 28 (2007) 1830–1837. [13] N. Okafuji, T. Shimizu, T. Watanabe, A. Kimura, S. Kurihara, Y. Arai, K. Furusawa, H. Hasegawa, T. Kawakami, Three-dimensional observation of reconstruction course of rabbit experimental mandibular defect with rhBMP-2 and atelocollagen gel, Eur. J. Med. Res. 11 (2006) 351–354. [14] F. Chen, Z. Wu, Q. Wang, H. Wu, Y. Zhang, X. Nie, Y. Jin, Preparation and biological characteristics of recombinant human bone morphogenetic protein-2-loaded dextran-co-gelatin hydrogel microspheres, in vitro and in vivo studies, Pharmacology 75 (2005) 133–144. [15] Y.I. Chung, K.M. Ahn, S.H. Jeon, S.Y. Lee, J.H. Lee, G. Tae, Enhanced bone regeneration with BMP-2 loaded functional nanoparticle-hydrogel complex, J. Control. Release 121 (2007) 91–99. [16] S.E. Kim, S.H. Song, Y.P. Yun, B.J. Choi, I.K. Kwon, M.S. Bae, H.J. Moon, Y.D. Kwon, The effect of immobilization of heparin and bone morphogenic protein-2 (BMP2) to titanium surfaces on inflammation and osteoblast function, Biomaterials 32 (2011) 366–373. [17] T.J. Cho, L.C. Gerstenfeld, T.A. Einhorn, Differential temporal expression of members of the transforming growth factor beta superfamily during murine fracture healing, J. Bone Miner. Res. 17 (2002) 513–520. [18] R.S. Sellers, R. Zhang, S.S. Glasson, H.D. Kim, D. Peluso, D.A. D’Augusta, K. Beckwith, E.A. Morris, Repair of articular cartilage defects one year after treatment with recombinant human bone morphogenetic protein-2 (rhBMP-2), J. Bone Joint Surg. Am. 82 (2000) 151–160. [19] M.H. Metz-Boutigue, J. Jolles, J. Mazurier, F. Schoentgen, D. Legrand, G. Spik, J. Montreuil, P. Jolles, Human lactotransferrin: amino acid sequence and structural comparisons with other transferrins, Eur. J. Biochem./FEBS 145 (1984) 659–676. [20] B. Lonnerdal, S. Iyer, Lactoferrin: molecular structure and biological function, Annu. Rev. Nutr. 15 (1995) 93–110. [21] H. Onishi, Lactoferrin delivery systems: approaches for its more effective use, Expert Opin. Drug Deliv. 8 (2011) 1469–1479. [22] S.A. Gonzalez-Chavez, S. Arevalo-Gallegos, Q. Rascon-Cruz, Lactoferrin: structure, function and applications, Int. J. Antimicrob. Agents 33 (301) (2009) e301–e308. [23] J. Cornish, K. Palmano, K.E. Callon, M. Watson, J.M. Lin, P. Valenti, D. Naot, A.B. Grey, I.R. Reid, Lactoferrin and bone; structure–activity relationships, Biochem. Cell Biol. 84 (2006) 297–302. [24] Y. Takayama, K. Mizumachi, Effect of bovine lactoferrin on extracellular matrix calcification by human osteoblast-like cells, Biosci. Biotechnol. Biochem. 72 (2008) 226–230. [25] Y. Takayama, K. Mizumachi, Effect of lactoferrin-embedded collagen membrane on osteogenic differentiation of human osteoblast-like cells, J. Biosci. Bioeng. 107 (2009) 191–195. [26] A. Grey, T. Banovic, Q. Zhu, M. Watson, K. Callon, K. Palmano, J. Ross, D. Naot, I.R. Reid, J. Cornish, The low-density lipoprotein receptor-related protein 1 is a mitogenic receptor for lactoferrin in osteoblastic cells, Mol. Endocrinol. 18 (2004) 2268–2278. [27] R. Takaoka, Y. Hikasa, K. Hayashi, Y. Tabata, Bone regeneration by lactoferrin released from a gelatin hydrogel, J. Biomater. Sci. Polym. Ed. 22 (2011) 1581–1589. [28] M. Montesi, S. Panseri, M. lafisco, A. Adamiano, A. Tampieri, Effect of hydroxyapatite nanocrystals functionalized with lactoferrin in osteogenic differentiation of mesenchymal stem cells, J. Biomed. Mater. Res. A (March) (2014), http://dx.doi.org/10.1002/jbm.a.35170.
[29] X. Ying, S. Cheng, W. Wang, Z. Lin, Q. Chen, W. Zhang, D. Kou, Y. Shen, X. Cheng, L. Peng, H. Zi Xu, C. Zhu Lu, Effect of lactoferrin on osteogenic differentiation of human adipose stem cells, Int. Orthop. 36 (2012) 647–653. [30] K. Das, S. Bose, A. Bandyopadhyay, Surface modifications and cell-materials interactions with anodized Ti, Acta Biomater. 3 (2007) 573–585. [31] D. Lin, Q. Li, W. Li, M. Swain, Bone remodeling induced by dental implants of functionally graded materials, J. Biomed. Mater. Res. Part B: Appl. Biomater. 92 (2010) 430–438. [32] D.W. Lee, Y.P. Yun, K. Park, S.E. Kim, Gentamicin and bone morphogenic protein-2 (BMP-2)-delivering heparinized-titanium implant with enhanced antibacterial activity and osteointegration, Bone 50 (2012) 974–982. [33] S.Y. Lee, Y.P. Yun, H.R. Song, H.J. Chun, D.H. Yang, K. Park, S.E. Kim, The effect of titanium with heparin/BMP-2 complex for improving osteoblast activity, Carbohydr. Polym. 98 (2013) 546–554. [34] X. Fan, L. Lin, J.L. Dalsin, P.B. Messersmith, Biomimetic anchor for surfaceinitiated polymerization from metal substrates, J. Am. Chem. Soc. 127 (2005) 15843–15847. [35] H. Lee, N.F. Scherer, P.B. Messersmith, Single-molecule mechanics of mussel adhesion, Proc. Natl. Acad. Sci. U.S.A. 103 (2006) 12999–13003. [36] H. Lee, S.M. Dellatore, W.M. Miller, P.B. Messersmith, Mussel-inspired surface chemistry for multifunctional coatings, Science 318 (2007) 426–430. [37] P.H. Chua, K.G. Neoh, E.T. Kang, W. Wang, Surface functionalization of titanium with hyaluronic acid/chitosan polyelectrolyte multilayers and RGD for promoting osteoblast functions and inhibiting bacterial adhesion, Biomaterials 29 (2008) 1412–1421. [38] T. Ishibe, T. Goto, T. Kodama, T. Miyazaki, S. Kobayashi, T. Takahashi, Bone formation on apatite-coated titanium with incorporated BMP-2/heparin in vivo, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod. 108 (2009) 867–875. [39] J.S. Park, K. Park, D.G. Woo, H.N. Yang, H.M. Chung, K.H. Park, Triple constructs consisting of nanoparticles and microspheres for bone-marrow-derived stromal-cell-delivery microscaffolds, Small 4 (2008) 1950–1955. [40] H.J. Moon, Y.P. Yun, C.W. Han, M.S. Kim, S.E. Kim, M.S. Bae, G.T. Kim, Y.S. Choi, E.H. Hwang, J.W. Lee, J.M. Lee, C.H. Lee, D.S. Kim, I.K. Kwon, Effect of heparin and alendronate coating on titanium surfaces on inhibition of osteoclast and enhancement of osteoblast function, Biochem. Biophys. Res. Commun. 413 (2011) 194–200. [41] J.B. Huh, S.E. Kim, H.E. Kim, S.S. Kang, K.H. Choi, C.M. Jeong, J.Y. Lee, S.W. Shin, Effects of the immobilization of heparin and rhPDGF-BB to titanium surfaces for the enhancement of osteoblastic functions and anti-inflammation, J. Adv. Prosthodont. 3 (2011) 152–160. [42] J.S. Park, K. Park, D.G. Woo, H.N. Yang, H.M. Chung, K.H. Park, PLGA microsphere construct coated with TGF-beta 3 loaded nanoparticles for neocartilage formation, Biomacromolecules 9 (2008) 2162–2169. [43] R. Ravichandran, J.R. Venugopal, S. Sundarrajan, S. Mukherjee, S. Ramakrishna, Precipitation of nanohydroxyapatite on PLLA/PBLG/Collagen nanofibrous structures for the differentiation of adipose derived stem cells to osteogenic lineage, Biomaterials 33 (2012) 846–855. [44] T.A. Owen, M. Aronow, V. Shalhoub, L.M. Barone, L. Wilming, M.S. Tassinari, M.B. Kennedy, S. Pockwinse, J.B. Lian, G.S. Stein, Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix, J. Cell Physiol. 143 (1990) 420–430. [45] K. Serigano, D. Sakai, A. Hiyama, F. Tamura, M. Tanaka, J. Mochida, Effect of cell number on mesenchymal stem cell transplantation in a canine disc degeneration model, J. Orthop. Res. 28 (2010) 1267–1275. [46] U. Stucki, J. Schmid, C.F. Hammerle, N.P. Lang, Temporal and local appearance of alkaline phosphatase activity in early stages of guided bone regeneration. A descriptive histochemical study in humans, Clin. Oral Implants Res. 12 (2001) 121–127. [47] C. Heinemann, S. Heinemann, A. Bernhardt, H. Worch, T. Hanke, Novel textile chitosan scaffolds promote spreading, proliferation, and differentiation of osteoblasts, Biomacromolecules 9 (2008) 2913–2920. [48] D. Naot, A. Chhana, B.G. Matthews, K.E. Callon, P.C. Tong, J.-M. Lin, J.L. Costa, M. Watson, A.B. Grey, J. Cornish, Molecular mechanisms involved in the mitogenic effect of lactoferrin in osteoblasts, Bone 49 (2011) 217–224. [49] W. Zhang, H. Guo, H. Jing, Y. Li, X. Wang, H. Zhang, L. Jiang, F. Ren, Lactoferrin stimulates osteoblast differentiation through PKA and p38 pathways independent of lactoferrin’s receptor LRP1, J. Bone Mineral Res. 29 (2014) 1232–1243.
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Colloids and Surfaces B: Biointerfaces journal homepage: www.elsevier.com/locate/colsurfb
Osteoblast activity of MG-63 cells is enhanced by growth on a lactoferrin-immobilized titanium substrate Sung Eun Kim a,1 , Young-Pil Yun a,1 , Jae Yong Lee b , Kyeongsoon Park c , Dong Hun Suh d,∗ a Department of Orthopedic Surgery and Rare Diseases Institute, Korea University Medical College, Guro Hospital, #80, Guro-dong, Guro-gu, Seoul 152-703, Republic of Korea b Department of Biomedical Science, College of Medicine, Korea University, Anam-dong, Seongbuk-gu 136-701, Republic of Korea c Division of Bio-imaging, Chuncheon Center, Korea Basic Science Institute, 192-1 Hyoja 2-dong, Chuncheon, Gangwon-do 200-701, Republic of Korea d Department of Orthopedic Surgery, Korea University Medical College, Ansan Hospital, Gojan 1-dong, Danwon-gu, Gyeonggi-do 425-707, Republic of Korea
a r t i c l e
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Article history: Received 25 March 2014 Received in revised form 25 August 2014 Accepted 5 September 2014 Available online 23 September 2014 Keywords: Titanium Lactoferrin Heparin–dopamine Osteoblast Osteogenic differentiation
a b s t r a c t The aim of this study was to develop a lactoferrin (LF)-immobilized titanium (Ti) substrate to enhance the osteoblast activity of MG-63 cells. Ti substrates were first modified through heparin–dopamine (Hep–DOPA) anchorage. Then, LF was immobilized on the Hep–Ti substrates via electrostatic interactions. Hep–Ti substrates, with or without LF, were evaluated by scanning electron microscopy (SEM), X-ray photoelectron spectroscopy (XPS), and contact angle measurements. Sustained release of LF on the Ti substrates was observed over a 28-day period. In vitro studies of osteoblast activity showed increased alkaline phosphatase activity and calcium deposition by MG-63 cells cultured on LF-immobilized Ti substrates as compared to those cultured on pristine Ti substrates, indicating that LF-immobilized Ti substrates were effective at enhancing osteoblast activity. © 2014 Elsevier B.V. All rights reserved.
1. Introduction Titanium (Ti) and its alloys are widely used in the orthopedic and dental fields due to their low toxicity, excellent mechanical properties, and low corrosive properties [1–3]. Despite these excellent properties, Ti alone does not promote osseo-integration with the host bone at the implant surface. Poor bonding of Ti implants with the host bone can bring about unpredictable bone loosening over time, resulting in implant failure and patient discomfort [4,5]. To promote osseo-integration between the Ti surface and host bone, attempts have been made to modify and functionalize the Ti surface though calcium phosphate/hydroxyapatite (HAp) coating, biomolecule immobilization, and surface topography modification [4,6–9]. Bone morphogenic protein-2 (BMP-2) has been shown to promote osteo-inductive activity, bone regeneration, and cartilage regeneration. Additionally, BMP-2 has been used in combination with various materials such as collagen gels, sponges, hyaluronic acid, dextran, chitosan, fibrin scaffolds, and Ti to promote bone formation and osseo-integration [10–16]. More recently, BMP-2 mixed with components such as collagen sponges (INFUSE Bone
∗ Corresponding author. Tel.: +82 31 412 4945; fax: +82 31 487 9502. E-mail address: [email protected] (D.H. Suh). 1 These authors contributed equally to this paper. http://dx.doi.org/10.1016/j.colsurfb.2014.09.014 0927-7765/© 2014 Elsevier B.V. All rights reserved.
Graft; Medtronic, Minneapolis, MN, USA) and biphasic calcium phosphate (BioC® ; 70% tri-calcium phosphate (TCP): 30% HAp) has been approved by bodies such as the Food and Drug Administration (FDA) and the Korea Food and Drug Administration (KFDA) for operations such as posterior-lateral spine fusions, tibial fractures, and alveolar ridge augmentations in the fields of orthopedic and dental surgery. Although clinical treatment with BMP-2 has been successful, BMP-2 has well-known drawbacks, such as a short in vivo half-life, high cost, and induction of excessive bone formation [17,18]. Lactoferrin (LF) is an 80-kDa iron-binding glycoprotein in the transferrin family of proteins [19]. It is produced by many different exocrine glands, and is widely distributed among body fluids such as milk, saliva, tears, bile, and pancreatic fluid [20]. Previous studies have demonstrated that LF has inhibitory biological effects against tumors, inflammatory processes, bacteria, and fungi [21,22]. In addition, LF has been reported to induce osteoblast and osteoblast cell-line proliferation, as well as osteoblast differentiation [23–25]. LF has also been reported to inhibit osteoblastic cells from apoptosis induced by serum withdrawal [26]. Takayama et al. [25] reported that LF incorporated in collagen membranes promoted alkaline phosphatase (ALP) activity and osteocalcin (OCN) production by human osteoblast-like cells (MG-63 cells). Previous studies have also demonstrated that LF incorporated in gelatin hydrogels significantly induced 3T3E1 (mouse-derived
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Fig. 1. Schematic diagram of the immobilization of lactoferrin (LF) to heparin–dopamine (Hep–DOPA)-coated Ti substrate.
osteoblast) cell proliferation in vitro, and, through sustained release from the hydrogels, enhanced bone regeneration at defect sites in vivo [27]. Furthermore, mesenchymal stem cells treated with biomimetic hydroxyapatite nanocrystals functionalized with LF showed enhanced osteogenic differentiation [28]. More recently, Ying et al. reported that LF promotes proliferation and osteogenic differentiation of human adipose-derived stem cells (hADSCs) [29]. Although the above studies suggest that LF may be a promising therapeutic agent to treat bone diseases or regenerate bone defects, no prior study has investigated the effect of LF-immobilized titanium (Ti) substrates on osteogenesis induction of MG-63 cells. We therefore set out to evaluate the effects of LF-immobilized Ti substrates on MG-63 osteoblast activity, which we did by evaluating F-actin staining, cell proliferation, alkaline phosphatase (ALP) activity, and calcium content. 2. Materials and methods 2.1. Materials Titanium (Ti) discs (diameter × height: 1.2 cm × 0.3 cm) were kindly supplied by Neobiotech Co., Ltd (Seoul, Korea). Heparin (average MW ca. 12,000 Da) was purchased from Pharmacia Hepar Co. (Franklin, OH, USA). Dulbecco’s modified Eagle’s medium (DMEM), fetal bovine serum (FBS), phosphatebuffered saline (PBS), and penicillin-streptomycin were purchased from Gibco BRL (Rockville, MD, USA). Dopamine hydrochloride, lactoferrin (LF, origin: human milk, Cat. No: L4894), 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDAC), and Nhydroxysuccinimide (NHS) were obtained from Sigma Chemical Co. (St. Louis, MO, USA). PierceTM BCA protein assay kit was purchased from Thermo Fisher Scientific (Rockford, IL, USA). MG 63 cells (human osteosarcoma cell line) were obtained from the Korea Cell Line Bank (KCLB No. 21427, Seoul, Korea). All other chemicals were of the purest analytical grade available. 2.2. Immobilization of heparin–dopamine (Hep–DOPA) and lactoferrin (LF) on the surface of titanium (Ti) Prior to preparing functionalized Ti substrates, we modified pristine Ti substrates using Hep–DOPA conjugated materials. In
brief, pristine Ti substrates were immersed in Hep–DOPA (2 mg) and dissolved in 10 mM Tris·HCl (pH 8.0) buffer in the dark, overnight. Heparin-coated Ti (Hep–Ti) substrates were rinsed with distilled water (DW) and dried under N2 . Lactoferrin (LF) was immobilized on the surface of Hep–Ti through electrostatic interaction between the amine groups of LF and the carboxylic/sulfate groups of heparin. Briefly, different amounts of LF (10, 50, or 100 g/mL) and Hep–Ti substrate were immersed in 0.1 M MES buffer (pH 5.6) and gently shaken for 6 hr. LF (10, 50, or 100 g)/Hep–Ti was washed and dried at room temperature (RT) (Fig. 1).
2.3. Characterization of pristine Ti and surface-modified Ti Surface morphologies of pristine Ti and surface-modified Ti were investigated by scanning electron microscopy (SEM, S4800, Hitachi, Japan). Each sample was coated with gold using a sputtercoater (Eiko IB, Japan), and SEM observations were performed at an operating voltage of 3 kV. To confirm the surface chemical compositions of all samples, X-ray photoelectron spectroscopy (XPS) was carried out using a K-alpha spectrometer XPS system (ESCALAB250) and a Theta Probe AR-XPS System (Thermo Fisher Scientific, UK) with an Al K␣ X-ray source (1486.6 eV photons) at the Korea Basic Science Institute Busan Center. The C1s hydrocarbon peak at 284.84 eV was used as the reference for all binding energies. The area of each peak was normalized to the total peak area (for all atomic elements) to calculate surface atom percentage. Hydrophilic properties of pristine Ti and surface-modified Ti were measured by contact angle (SEO Phoenix 300, Suwon, Korea). Toluidine blue was used to quantify the amount of heparin immobilized on the surface of pristine Ti. Briefly, heparin-modified Ti (Hep–Ti) was immersed in 1 mL of 0.2% (v/v) NaCl solution containing 1 mL of 0.005% (w/v) toluidine blue solution. After gentle shaking for 30 min, 2 mL of hexane was added. Hep–Ti was removed from solution, and aqueous-phase absorbance was measured using a micro-plate reader (Bio-Rad, Hercules, CA, USA) at a wavelength of 620 nm. To analyze the amount of LF immobilized on Hep–Ti, a PierceTM BCA protein assay kit was used. In brief, after a 6-hr reaction, LF immobilized on Hep–Ti was removed from the MES buffer and then rinsed with PBS. After the reaction, the amount of
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LF remaining in the MES buffer was measured to indirectly evaluate LF immobilization on the Ti substrate surface. 2.4. Lactoferrin (LF) release study To evaluate the amount of LF released from LF/Hep–Ti in the 10/50/100-g LF conditions, each sample was immersed
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in 1 mL PBS buffer (pH 7.4) and gently shaken at 100 rpm at 37 ◦ C. PBS was collected and replaced with fresh PBS at predesigned time intervals of 1, 3, 5, and 10 h and 1, 3, 5, 7, 14, 21, and 28 days. Concentration of released LF was analyzed with a PierceTM BCA protein assay kit in accordance with the manufacturer’s instructions using a micro-plate reader at 562 nm.
Fig. 2. (A) SEM images and (B) wide-scanned XPS spectra of pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti.
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2.5. Cytotoxicity and F-actin staining Cytotoxicities of all Ti substrates were tested using a cell counting kit-8 (CCK-8) (Dojindo, Japan). This kit is based on a highly water-soluble tetrazolium salt, WST-8 [2-(2-methoxy4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2Htetrazolium, monosodium salt], that produces a water-soluble formazan dye upon reduction in the presence of an electron mediator. Cytotoxicity of both pristine Ti and surface-modified Ti was evaluated via an indirect method. Specifically, 1 × 104 MG-63 cells were seeded on 96-well plates and incubated at 37 ◦ C for 24 h in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 50 g/mL ascorbic acid, 10 nM dexamethasone, and 10 mM -glycerophosphate in the presence of 100 U/mL penicillin and 100 g/mL streptomycin. Pristine and surface-modified Ti samples were incubated at 37 ◦ C for 24 h in DMEM medium to obtain extraction medium. DMEM medium was aspirated over the (pristine Ti or modified Ti) surface containing MG-63 cells before cells were washed with PBS, and fresh DMEM medium added. MG-63 cells were incubated for 1 or 2 days. At predetermined time intervals, the extraction medium was aspirated, CCK-8 solution added, and MG-63 cells were incubated for 1 hour at 37 ◦ C. Live-cell optical density (OD) was measured using a micro-plate reader at 450 nm. Relative cell viability (%) was calculated as [OD]test/[OD]control × 100. To observe the morphology of MG-63 cells adhered on pristine Ti and surface-modified Ti, surfaces with 1 × 105 seeded MG-63 cells were transferred to a 24-well tissue-culture plates. After 24 h of culture, substrates were placed in a cytoskeleton buffer (5 × 10−3 M NaCl, 150 × 10−3 M MgCl2 , 5 × 10−4 M Tris-base and 0.5% triton X100) for 5 min at 4 ◦ C, followed by addition of blocking buffer (5% FBS, 0.1% Tween-20 and 0.02% sodium azide in PBS) for 30 min at 37 ◦ C. Cells/substrates were incubated with rhodamine-phalloidin (1:200) and DAPI (1:10,000) and observed by confocal laser scanning microscopy (LSM700, Zeiss, Germany). 2.6. Cell proliferation CCK-8 kit for cell proliferation assay was also used, because cell proliferation measured using CCK-8 correlates well with results obtained using the [3 H]-thymidine incorporation assay, and thus, the CCK-8 assay can be substituted for the [3 H]-thymidine incorporation assay. MG-63 cells were seeded at a concentration of 1 × 105 cells/mL on the surface of pristine or surface-modified Ti in a 24-well tissue-culture plate. At days 3 and 7 of incubation, substrates were rinsed with PBS, and then CCK-8 solution was added to the substrates followed by a 1-h incubation. Reagents were transferred to 96-well plates. Cell proliferation results are presented as optical density measured at 450 nm using a micro-plate reader.
1 × 105 MG-63 cells were seeded on the surface of pristine Ti, Hep–Ti, or Hep–Ti modified with 10, 50, or 100 g LF in a 24well tissue-culture plate. At day 21 of culture, cells/substrates were rinsed with PBS, and 0.5 N HCl was added. Cells were centrifuged at 13,500 rpm for 1 min. The supernatant was used for calcium content measurements with a QuantiChromTM Calcium Assay Kit (DICA-500, BioAssay Systems, USA) in accordance with the manufacturer’s instructions. Concentration of calcium was measured by a micro-plate reader at 612 nm. 2.8. Statistical analysis Data are presented as mean ± standard deviations (n = 5). Statistical comparisons were carried out by one-way ANOVA using Systat software (Systat Software, Inc., Chicago, IL, USA). Differences were considered statistically significant at * P < 0.05 and ** P < 0.01. 3. Results 3.1. Characterization of pristine Ti and surface-modified Ti Ti morphology, both in the presence of and absence of surface modification, was observed by SEM (Fig. 2A). Adding Hep–Ti or LF (at any concentration, from 10 to 100 g) did not have an observable effect on Ti morphology. XPS measurement was carried out to investigate the surface chemical composition of pristine Ti and surface-modified Ti substrates. As shown in Fig. 2B and Table 1, wide-scanned XPS spectra of pristine Ti, Hep–Ti, and LF/Hep–Ti (at 10, 50 and 100 g LF) were evaluated. Successful heparin anchorage on the surface of pristine Ti was confirmed by an increase in the C1s content from 61.13% to 67.74%, an increase in the S2p content from 0% to 2.12%, and an increase in the N1s content from 2.15% to 3.65% as compared to pristine Ti (Table 1). Successful immobilization of increasing doses of LF on Hep–Ti was confirmed by increases in N1s
Table 1 Percentages of C1s, O1s, N1s, Ti2p, and S2p expressed at the surface of pristine and surface-modified Ti, as analyzed by XPS. Substrate
C1s
O1s
N1s
Ti2p
S2p
Pristine Ti Hep–Ti LF3.6 /Hep–Ti LF17.1 /Hep–Ti LF38.9 /Hep–Ti
61.13% 67.74% 72.19% 72.32% 71.36%
30.07% 21.76% 18.31% 17.18% 17.04%
2.15% 3.63% 5.74% 6.41% 6.97%
6.65% 4.75% 0.82% 0.92% 1.12%
– 2.12% 2.94% 3.17% 3.51%
2.7. Alkaline phosphatase (ALP) activity and calcium deposition ALP activity was measured at days 3, 7, and 10 of culture. Briefly, MG-63 cells were seeded at a density of 1 × 105 cells/mL in a 24well culture plate on the surface of pristine Ti, Hep–Ti, or Hep–Ti modified with 10, 50, or 100 g LF. Cells/substrates were rinsed with PBS at pre-determined times, and 1× RIPA buffer was added to lyse cells. Lysed cells were centrifuged at 13,500 rpm for 1 min. A 100 L p-nitrophenyl phosphate solution was added to the supernatant, and the reaction was allowed to proceed for 30 minute at 37 ◦ C. 1 N NaOH was added to stop the reaction. Optical density was determined at 405 nm using a micro-plate reader. Enzymatic activity was expressed as micromolecules of p-nitrophenol produced per min per g of protein. Protein content was determined using a modified Bradford assay using BSA as the standard. Calcium content was investigated after 21 days of incubation. In brief,
Fig. 3. Release kinetics of lactoferrin (LF) from LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti. Error bars represent mean ± SDs (n = 5).
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Fig. 4. (A) Cytotoxicity of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 1 and 2 days of culture. (B) F-actin staining images of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 24 hr of incubation.
and S2p contents. Mean ± standard deviations of contact angles of pristine Ti, Hep–Ti, and 10, 50, 100 g LF/Hep–Ti were 75.1 ± 1.8◦ , 54.6 ± 2.7◦ , 30.1 ± 2.7◦ , 29.1 ± 3.3◦ , and 26.1 ± 2.9◦ , respectively. 5.47 ± 0.64 g/sample of heparin was grafted on Ti (loading
efficiency (LE): 0.27%). Amounts of LF in the 10, 50 and 100 g LF/Hep–Ti cases were 3.6 g (LE: 36.0%), 17.1 g (LE: 34.2%), and 38.9 g (LE: 38.9%); these samples were designated as LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti, respectively.
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Fig. 5. Cell proliferation of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 3 and 7 days of incubation. Error bars represent mean ± SDs (n = 5). * P < 0.05 and ** P < 0.01.
3.2. In vitro LF release kinetics Fig. 3 shows the in vitro release kinetics of LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti as measured by the PierceTM BCA protein assay. LF exhibited sustained release kinetics for up to 28 days. On the first day, 1.65 ± 0.61 g, 4.53 ± 0.75 g, and 9.18 ± 0.40 g of LF was released from LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti, respectively. At 28 days, cumulative amount of LF released from LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti was 3.07 ± 1.16 g, 11.87 ± 0.98 g, and 22.98 ± 1.49 g, respectively, indicating that the amount of LF remaining was 14.8%, 30.6%, and 40.9%, respectively. 3.3. Cytotoxicity test and F-actin staining MG-63 cell viability on pristine Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti is shown in Fig. 4A. Cell viability for all groups was over 95% for incubation periods of 24–48 h, indicating that the Ti substrates were not cytotoxic to MG-63 cells. F-actin staining after 24 h of incubation revealed typical long and straight actin stress fibers on all Ti substrates (Fig. 4B). Long and straight actin stress fiber structures on the three LF/Hep–Ti substrates were more developed than those on pristine Ti and Hep–Ti substrates. Approximately two-fold more MG-63 cells adhered to the LF38.9/Hep–Ti substrate than the pristine Ti and Hep–Ti substrates. 3.4. Cell proliferation MG-63 cell proliferation on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti was evaluated at days 3 and 7 of incubation (Fig. 5). For each substrate, MG-63 cell proliferation increased gradually in a time-dependent manner. There were no significant differences in cell proliferation between MG-63 cells grown on pristine Ti and those grown on Hep–Ti at days 3 and 7. However, there were significant differences in MG-63 cell proliferation on LF38.9 /Hep–Ti as compared with pristine Ti at day 3 and day 7 (* P < 0.05 or ** P < 0.01). At day 7 of incubation, cell proliferation on the Ti substrates containing LF was significantly different from that on pristine Ti (* P < 0.05 and ** P < 0.01). 3.5. Alkaline phosphatase (ALP) activity and calcium deposition Fig. 6A shows ALP activity in MG-63 cells cultured on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti, for
incubation periods of up to 10 days. After 3 days, no significant differences in ALP activity between pristine Ti and surface-modified Ti were evident. However, there was a significant difference in the ALP activity of MG-63 cells between LF38.9 /Hep–Ti and pristine Ti at 7 and 10 days (* P < 0.05 and ** P < 0.01). ALP activity was 1.05 ± 0.05 for pristine Ti, 1.10 ± 0.08 for Hep–Ti, 1.24 ± 0.07 for LF3.6 /Hep–Ti, 1.36 ± 0.09 for LF17.1 /Hep–Ti, and 1.53 ± 0.06 M/min/g protein for LF38.9 /Hep–Ti, respectively. At 10 days, significant differences in ALP activity were observed between MG-63 cells cultured on Ti substrates containing LF and those cultured on pristine Ti (* P < 0.05 and ** P < 0.01). The amount of calcium deposited by MG-63 cells cultured on all Ti substrates was assessed after 21 days of culture (Fig. 6B). Amount of deposited calcium was 10.32 ± 1.0 for pristine Ti, 11.26 ± 1.0 for Hep–Ti, 14.52 ± 0.8 for LF3.6 /Hep–Ti, 16.30 ± 1.6 for LF17.1 /Hep–Ti, and 19.2 ± 1.6 g for LF38.9 /Hep–Ti. Calcium deposition by MG-63 cells cultured on Ti substrates containing LF was significantly higher than for MG-63 cells grown on pristine Ti (* P < 0.05 and ** P < 0.01). Moreover, a significant difference in the amount of calcium deposited by MG-63 cells grown on LF38.9 /Hep–Ti versus LF3.6 /Hep–Ti was observed (* P < 0.05). However, the amount of calcium deposited by MG-63 cells cultured on LF38.9 /Hep–Ti and LF17.1 /Hep–Ti was not significantly different after 21 days of culture.
4. Discussion Ti implant materials should ideally provide not only a good environment for osteoblast adhesion, proliferation, and differentiation, but also an environment that promotes interaction and osseo-integration between implants and host bone. However, failed Ti implants have been shown to lack implant/host bone interactions and have poor healing capabilities. To address these problems, Ti implants have been modified by surface modification and immobilization with biomolecules and bioactive proteins to improve osteoblast activity and osseo-integration [16,30–33]. The main objective of this study was to develop a lactoferrin (LF)immobilized Ti substrate to enhance osteoblast activity. To prepare LF-immobilized Ti substrate, the surface of the Ti substrate was first functionalized with Hep–DOPA in a slightly basic environment by chemical immobilization between oxidized catechol groups and the Ti surface, mimicking a mussel adhesion mechanism [34–36]. Surface functionalization of Ti with heparin has several advantages. Because of their highly negative charge and high binding affinities for growth factors, heparin molecules can facilitate the immobilization of growth factors on the surface of Ti substrate through electrostatic interactions between the growth factors and heparin, and can minimize rapid diffusion and regulate the sustained release of growth factors or drugs from substrates to improve their efficacies [16,32,37–40]. Indeed, we confirmed that LF was successfully immobilized on the surface of Hep–Ti substrates and that the amount of LF immobilized increased with increasing feed amounts of LF. As reported in previous in vitro release studies, Hep–Ti substrates effectively prevented high initial burst effects and also showed sustained and extended release of LF for 28 days [16,41,42]. Furthermore, all Ti substrates examined in this study were non-toxic to MG-63 cells, indicating that they are safe to use. Initial adhesion of osteoblast cells to the substrate is important for long-term stability and cell differentiation [43]. Furthermore, promotion of cell attachment by the substrate affects the proliferation capacity of cells [43]. Compared to osteoblast cells cultured on pristine Ti and Hep–Ti substrates, osteoblast cells grown on LF/Hep–Ti substrates were more spread out with numerous filopodia in all directions. In particular, cells cultured on LF38.9 /Hep–Ti, which was the substrate with the highest concentration of LF,
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Fig. 6. (A) Alkaline phosphatase (ALP) activity of MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 3, 7, and 10 days of incubation. (B) Amount of calcium deposited by MG-63 cells grown on pristine Ti, Hep–Ti, LF3.6 /Hep–Ti, LF17.1 /Hep–Ti, and LF38.9 /Hep–Ti after 21 days of incubation. Error bars represent mean ± SDs (n = 5). * P < 0.05 and ** P < 0.01.
had a flattened, mature, spreading morphology with scattered focal adhesion that increased contact between the cells and the LF38.9 /Hep–Ti surface, indicating that LF/Hep–Ti substrate promoted osteoblast cell attachment. Consistently, osteoblast cell proliferation increased gradually with increasing amount of LF immobilized on the Hep–Ti substrates. Our cell adhesion and proliferation results indicate that the LF-immobilized Ti substrates promoted cell adhesion and proliferation. It is possible that the presence of heparin on Ti substrate also affected cell attachment and proliferation. However, as we reported in our previous studies [16,32], cell adhesion and proliferation of osteoblasts on Hep–Ti substrate were not significantly different compared to those on pristine Ti. This may be because the amount of heparin immobilized on the Ti substrate in this study is too low to affect cell adhesion and proliferation. Three principal osteoblast differentiation processes can be recognized: proliferation, extracellular maturation, and mineralization [44]. Each stage is characterized by the expression of distinctive osteogenic markers. ALP activity is an early marker of osteoblastic differentiation and commitment of stem cells to the osteoblastic phenotype [3,43,45–47]. After osteogenic differentiation, cells begin to secrete a mineral matrix, leading to calcium deposition, which is a marker of mature osteoblasts [16,32,33]. In the present study, we compared the osteogenic activity of osteoblast-like cells (MG-63) cultured on LF-immobilized Ti substrates to those cultured on pristine Ti and Hep–Ti substrates. ALP activity and calcium content of osteoblasts cultured on pristine Ti and Hep–Ti without LF were not significantly different during the experiment periods. However, ALP activity and calcium content of osteoblasts cultured on LF/Hep–Ti substrates increased gradually in time- or dose-dependent manners. It has been reported that LF may stimulate osteogenic differentiation of osteoblast cells via receptor-mediated mechanisms. Among various receptors, lowdensity lipoprotein receptor-related protein 1 (LRP1) is the only receptor that can transduce the LF signal in osteoblasts. LF stimulates osteoblast differentiation mainly through LRP-1 independent protein kinase A (PKA) and p38 signaling pathways [48]. In addition, osteoblast proliferation and differentiation induced by LF appear to be mutually independent intracellular events that are regulated by the ERK1/2 and p38 pathways, respectively [49]. We demonstrated in this study that LF-immobilized Hep–Ti substrates significantly enhanced MG-63 cell proliferation, osteoblast ALP activity, and calcium deposition. These results suggest that LF-immobilized Hep–Ti
substrates are promising alternative Ti implants for regeneration of bone defects. 5. Conclusions We successfully developed LF-immobilized Ti substrates following Hep–DOPA anchorage. LF immobilized on Ti substrates showed sustained release over a 28-day period. LF-immobilized Ti substrates enhanced osteoblastic activity of MG-63 cells as compared to pristine Ti substrates, as shown by a significant increase in ALP activity and calcium deposition. These results suggest that LFimmobilized Ti substrates could effectively promote regeneration of bone defects. Further in vivo studies are needed to investigate the effects of dose-dependently-increasing the amount of LF immobilized to the Ti substrate. Acknowledgement This study was supported by a grant from Korea University (K1300081). References [1] D. Buser, U.C. Belser, N.P. Lang, The original one-stage dental implant system and its clinical application, Periodontology 2000 17 (1998) 106–118. [2] S.G. Steinemann, Titanium—the material of choice? Periodontology 2000 17 (1998) 7–21. [3] L. Zhu, X. Ye, G. Tang, N. Zhao, Y. Gong, Y. Zhao, J. Zhao, X. Zhang, Biomimetic coating of compound titania and hydroxyapatite on titanium, J. Biomed. Mater. Res. Part A 83 (2007) 1165–1175. [4] Z. Shi, K.G. Neoh, E.T. Kang, C.K. Poh, W. Wang, Surface functionalization of titanium with carboxymethyl chitosan and immobilized bone morphogenetic protein-2 for enhanced osseointegration, Biomacromolecules 10 (2009) 1603–1611. [5] D.A. Puleo, A. Nanci, Understanding and controlling the bone-implant interface, Biomaterials 20 (1999) 2311–2321. [6] F. Rupp, L. Scheideler, N. Olshanska, M. de Wild, M. Wieland, J. Geis-Gerstorfer, Enhancing surface free energy and hydrophilicity through chemical modification of microstructured titanium implant surfaces, J. Biomed. Mater. Res. Part A 76 (2006) 323–334. [7] S. Piskounova, J. Forsgren, U. Brohede, H. Engqvist, M. Stromme, In vitro characterization of bioactive titanium dioxide/hydroxyapatite surfaces functionalized with BMP-2, J. Biomed. Mater. Res. Part B: Appl. Biomater. 91 (2009) 780–787, Applied biomaterials. [8] K. Oya, Y. Tanaka, H. Saito, K. Kurashima, K. Nogi, H. Tsutsumi, Y. Tsutsumi, H. Doi, N. Nomura, T. Hanawa, Calcification by MC3T3-E1 cells on RGD peptide immobilized on titanium through electrodeposited PEG, Biomaterials 30 (2009) 1281–1286.
198
S.E. Kim et al. / Colloids and Surfaces B: Biointerfaces 123 (2014) 191–198
[9] R.J. Miron, C.J. Oates, A. Molenberg, M. Dard, D.W. Hamilton, The effect of enamel matrix proteins on the spreading, proliferation and differentiation of osteoblasts cultured on titanium surfaces, Biomaterials 31 (2010) 449–460. [10] G.N. King, N. King, A.T. Cruchley, J.M. Wozney, F.J. Hughes, Recombinant human bone morphogenetic protein-2 promotes wound healing in rat periodontal fenestration defects, J. Dental Res. 76 (1997) 1460–1470. [11] G. Zellin, A. Linde, Importance of delivery systems for growth-stimulatory factors in combination with osteopromotive membranes. An experimental study using rhBMP-2 in rat mandibular defects, J. Biomed. Mater. Res. 35 (1997) 181–190. [12] J. Kim, I.S. Kim, T.H. Cho, K.B. Lee, S.J. Hwang, G. Tae, I. Noh, S.H. Lee, Y. Park, K. Sun, Bone regeneration using hyaluronic acid-based hydrogel with bone morphogenic protein-2 and human mesenchymal stem cells, Biomaterials 28 (2007) 1830–1837. [13] N. Okafuji, T. Shimizu, T. Watanabe, A. Kimura, S. Kurihara, Y. Arai, K. Furusawa, H. Hasegawa, T. Kawakami, Three-dimensional observation of reconstruction course of rabbit experimental mandibular defect with rhBMP-2 and atelocollagen gel, Eur. J. Med. Res. 11 (2006) 351–354. [14] F. Chen, Z. Wu, Q. Wang, H. Wu, Y. Zhang, X. Nie, Y. Jin, Preparation and biological characteristics of recombinant human bone morphogenetic protein-2-loaded dextran-co-gelatin hydrogel microspheres, in vitro and in vivo studies, Pharmacology 75 (2005) 133–144. [15] Y.I. Chung, K.M. Ahn, S.H. Jeon, S.Y. Lee, J.H. Lee, G. Tae, Enhanced bone regeneration with BMP-2 loaded functional nanoparticle-hydrogel complex, J. Control. Release 121 (2007) 91–99. [16] S.E. Kim, S.H. Song, Y.P. Yun, B.J. Choi, I.K. Kwon, M.S. Bae, H.J. Moon, Y.D. Kwon, The effect of immobilization of heparin and bone morphogenic protein-2 (BMP2) to titanium surfaces on inflammation and osteoblast function, Biomaterials 32 (2011) 366–373. [17] T.J. Cho, L.C. Gerstenfeld, T.A. Einhorn, Differential temporal expression of members of the transforming growth factor beta superfamily during murine fracture healing, J. Bone Miner. Res. 17 (2002) 513–520. [18] R.S. Sellers, R. Zhang, S.S. Glasson, H.D. Kim, D. Peluso, D.A. D’Augusta, K. Beckwith, E.A. Morris, Repair of articular cartilage defects one year after treatment with recombinant human bone morphogenetic protein-2 (rhBMP-2), J. Bone Joint Surg. Am. 82 (2000) 151–160. [19] M.H. Metz-Boutigue, J. Jolles, J. Mazurier, F. Schoentgen, D. Legrand, G. Spik, J. Montreuil, P. Jolles, Human lactotransferrin: amino acid sequence and structural comparisons with other transferrins, Eur. J. Biochem./FEBS 145 (1984) 659–676. [20] B. Lonnerdal, S. Iyer, Lactoferrin: molecular structure and biological function, Annu. Rev. Nutr. 15 (1995) 93–110. [21] H. Onishi, Lactoferrin delivery systems: approaches for its more effective use, Expert Opin. Drug Deliv. 8 (2011) 1469–1479. [22] S.A. Gonzalez-Chavez, S. Arevalo-Gallegos, Q. Rascon-Cruz, Lactoferrin: structure, function and applications, Int. J. Antimicrob. Agents 33 (301) (2009) e301–e308. [23] J. Cornish, K. Palmano, K.E. Callon, M. Watson, J.M. Lin, P. Valenti, D. Naot, A.B. Grey, I.R. Reid, Lactoferrin and bone; structure–activity relationships, Biochem. Cell Biol. 84 (2006) 297–302. [24] Y. Takayama, K. Mizumachi, Effect of bovine lactoferrin on extracellular matrix calcification by human osteoblast-like cells, Biosci. Biotechnol. Biochem. 72 (2008) 226–230. [25] Y. Takayama, K. Mizumachi, Effect of lactoferrin-embedded collagen membrane on osteogenic differentiation of human osteoblast-like cells, J. Biosci. Bioeng. 107 (2009) 191–195. [26] A. Grey, T. Banovic, Q. Zhu, M. Watson, K. Callon, K. Palmano, J. Ross, D. Naot, I.R. Reid, J. Cornish, The low-density lipoprotein receptor-related protein 1 is a mitogenic receptor for lactoferrin in osteoblastic cells, Mol. Endocrinol. 18 (2004) 2268–2278. [27] R. Takaoka, Y. Hikasa, K. Hayashi, Y. Tabata, Bone regeneration by lactoferrin released from a gelatin hydrogel, J. Biomater. Sci. Polym. Ed. 22 (2011) 1581–1589. [28] M. Montesi, S. Panseri, M. lafisco, A. Adamiano, A. Tampieri, Effect of hydroxyapatite nanocrystals functionalized with lactoferrin in osteogenic differentiation of mesenchymal stem cells, J. Biomed. Mater. Res. A (March) (2014), http://dx.doi.org/10.1002/jbm.a.35170.
[29] X. Ying, S. Cheng, W. Wang, Z. Lin, Q. Chen, W. Zhang, D. Kou, Y. Shen, X. Cheng, L. Peng, H. Zi Xu, C. Zhu Lu, Effect of lactoferrin on osteogenic differentiation of human adipose stem cells, Int. Orthop. 36 (2012) 647–653. [30] K. Das, S. Bose, A. Bandyopadhyay, Surface modifications and cell-materials interactions with anodized Ti, Acta Biomater. 3 (2007) 573–585. [31] D. Lin, Q. Li, W. Li, M. Swain, Bone remodeling induced by dental implants of functionally graded materials, J. Biomed. Mater. Res. Part B: Appl. Biomater. 92 (2010) 430–438. [32] D.W. Lee, Y.P. Yun, K. Park, S.E. Kim, Gentamicin and bone morphogenic protein-2 (BMP-2)-delivering heparinized-titanium implant with enhanced antibacterial activity and osteointegration, Bone 50 (2012) 974–982. [33] S.Y. Lee, Y.P. Yun, H.R. Song, H.J. Chun, D.H. Yang, K. Park, S.E. Kim, The effect of titanium with heparin/BMP-2 complex for improving osteoblast activity, Carbohydr. Polym. 98 (2013) 546–554. [34] X. Fan, L. Lin, J.L. Dalsin, P.B. Messersmith, Biomimetic anchor for surfaceinitiated polymerization from metal substrates, J. Am. Chem. Soc. 127 (2005) 15843–15847. [35] H. Lee, N.F. Scherer, P.B. Messersmith, Single-molecule mechanics of mussel adhesion, Proc. Natl. Acad. Sci. U.S.A. 103 (2006) 12999–13003. [36] H. Lee, S.M. Dellatore, W.M. Miller, P.B. Messersmith, Mussel-inspired surface chemistry for multifunctional coatings, Science 318 (2007) 426–430. [37] P.H. Chua, K.G. Neoh, E.T. Kang, W. Wang, Surface functionalization of titanium with hyaluronic acid/chitosan polyelectrolyte multilayers and RGD for promoting osteoblast functions and inhibiting bacterial adhesion, Biomaterials 29 (2008) 1412–1421. [38] T. Ishibe, T. Goto, T. Kodama, T. Miyazaki, S. Kobayashi, T. Takahashi, Bone formation on apatite-coated titanium with incorporated BMP-2/heparin in vivo, Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endod. 108 (2009) 867–875. [39] J.S. Park, K. Park, D.G. Woo, H.N. Yang, H.M. Chung, K.H. Park, Triple constructs consisting of nanoparticles and microspheres for bone-marrow-derived stromal-cell-delivery microscaffolds, Small 4 (2008) 1950–1955. [40] H.J. Moon, Y.P. Yun, C.W. Han, M.S. Kim, S.E. Kim, M.S. Bae, G.T. Kim, Y.S. Choi, E.H. Hwang, J.W. Lee, J.M. Lee, C.H. Lee, D.S. Kim, I.K. Kwon, Effect of heparin and alendronate coating on titanium surfaces on inhibition of osteoclast and enhancement of osteoblast function, Biochem. Biophys. Res. Commun. 413 (2011) 194–200. [41] J.B. Huh, S.E. Kim, H.E. Kim, S.S. Kang, K.H. Choi, C.M. Jeong, J.Y. Lee, S.W. Shin, Effects of the immobilization of heparin and rhPDGF-BB to titanium surfaces for the enhancement of osteoblastic functions and anti-inflammation, J. Adv. Prosthodont. 3 (2011) 152–160. [42] J.S. Park, K. Park, D.G. Woo, H.N. Yang, H.M. Chung, K.H. Park, PLGA microsphere construct coated with TGF-beta 3 loaded nanoparticles for neocartilage formation, Biomacromolecules 9 (2008) 2162–2169. [43] R. Ravichandran, J.R. Venugopal, S. Sundarrajan, S. Mukherjee, S. Ramakrishna, Precipitation of nanohydroxyapatite on PLLA/PBLG/Collagen nanofibrous structures for the differentiation of adipose derived stem cells to osteogenic lineage, Biomaterials 33 (2012) 846–855. [44] T.A. Owen, M. Aronow, V. Shalhoub, L.M. Barone, L. Wilming, M.S. Tassinari, M.B. Kennedy, S. Pockwinse, J.B. Lian, G.S. Stein, Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix, J. Cell Physiol. 143 (1990) 420–430. [45] K. Serigano, D. Sakai, A. Hiyama, F. Tamura, M. Tanaka, J. Mochida, Effect of cell number on mesenchymal stem cell transplantation in a canine disc degeneration model, J. Orthop. Res. 28 (2010) 1267–1275. [46] U. Stucki, J. Schmid, C.F. Hammerle, N.P. Lang, Temporal and local appearance of alkaline phosphatase activity in early stages of guided bone regeneration. A descriptive histochemical study in humans, Clin. Oral Implants Res. 12 (2001) 121–127. [47] C. Heinemann, S. Heinemann, A. Bernhardt, H. Worch, T. Hanke, Novel textile chitosan scaffolds promote spreading, proliferation, and differentiation of osteoblasts, Biomacromolecules 9 (2008) 2913–2920. [48] D. Naot, A. Chhana, B.G. Matthews, K.E. Callon, P.C. Tong, J.-M. Lin, J.L. Costa, M. Watson, A.B. Grey, J. Cornish, Molecular mechanisms involved in the mitogenic effect of lactoferrin in osteoblasts, Bone 49 (2011) 217–224. [49] W. Zhang, H. Guo, H. Jing, Y. Li, X. Wang, H. Zhang, L. Jiang, F. Ren, Lactoferrin stimulates osteoblast differentiation through PKA and p38 pathways independent of lactoferrin’s receptor LRP1, J. Bone Mineral Res. 29 (2014) 1232–1243.
