Biomimetic surface modification of dental implant for enhanced osseointegration

BIOMIMETIC SURFACE MODIFICATION OF DENTAL IMPLANT FOR ENHANCED OSSEOINTEGRATION RAJESWARI RAVICHANDRAN (B.Tech, ANNA University) A THESIS SUBMITTED FOR THE DEGREE OF MASTER OF ENGINEERING DIVISION OF BIOENGINEERING NATIONAL UNIVERISTY OF SINGAPORE 2009 1 ACKNOWLEDGEMENT I would like to express my sincere appreciation to those who have helped and contributed to this thesis. I would like to thank Professor Michael Raghunath who has shown faith in me and given me tremendous encouragement throughout my tenure. I would like to express my sincere thanks to Professor Seeram Ramakrishna for his excellent supervision and guidance throughout this project. I would like to express my heartfelt gratitude to Dr Clarisse Ng and Dr Susan Liao, who have provided unmatched guidance and support, throughout this project. I would also like to thank Professor Casey Chan, Dr. Damian Pliza and Dr. Venugopal for giving me invaluable advice, discussion, and suggestions. I would also like to thank all Prof Seeram’s lab members for their assistance in the completion of this project. I would like to thank the Division of Bioengineering and the Faculty of Dentistry for their constant support. Last but not the least I would like to thank my parents for their profound love and support. 2 TABLE OF CONTENTS ACKNOWLEDGEMENTS TABLE OF CONTENTS LIST OF FIGURES LIST OF TABLES LIST OF APPENDICES LIST OF ABBREVIATIONS Chapter 1: Introduction 1.1 Background 1 1.2 Clinical problems associated with osseointegration 2 1.3 Hypothesis and objectives 4 Chapter 2: Literature Review 2.1 Introduction 5 2.2 Surface modification techniques 6 2.2.1 Modification of scaffolds using surface adhesive molecules 2.2.2 Cell – substrate interaction 9 13 3 2.3 Tissue Engineering 2.3.1 Introduction 14 2.3.2 Nanofiber fabrication by electrospinning 14 2.3.3 Modifications of the electrospun nanofibers 19 2.3.4 Potential application of Mesenchymal stem cells for osseointegration 20 Chapter 3: Biomimetic surface modification of dental implant by advanced electrospinning 3.1 Introduction 21 3.2 Materials and Methods 22 3.2.1 Mechanical Polishing/ etching 22 3.2.2 Pretreatment of Ti 22 3.2.3 Electrospinning of PLGA and PLGA/Collagen nanofibers on the Ti discs 23 3.2.4 Biomineralization using Calcium-Phosphate dipping method 25 3.2.5 Cell adhesion study 26 3.2.6 Surface characterization analysis 27 3.2.7 Surface roughness analysis 28 4 3.2.8 Fourier transform infrared spectroscopy (FT-IR) and X-ray photoelectron spectroscopy (XPS) 28 3.2.9 Water contact angle measurement 29 3.2.10 Statistical analysis 29 3.3 Results and Discussion 29 3.3.1 Surface characterization analysis 29 3.3.2 Surface Roughness analysis 34 3.3.3 Fourier transform infrared spectroscopy (FT-IR) 35 3.3.4 Water contact angle measurement 38 3.3.5 X-ray photoelectron spectroscopy (XPS) 39 3.3.6 Cell culture analysis 40 3.4 Conclusion 46 Chapter 4: Mesenchymal stem cells proliferation and differentiation studies on the modified implant surfaces 4.1 Introduction 47 4.2 Material and methods 48 4.2.1 Mesenchymal stem cells culture 48 4.2.2 Cell Morphology study 49 5 4.2.3 Cell Proliferation study 49 4.2.4 Alkaline phosphatase activity 50 4.2.5 Cell mineralization study 50 4.2.6 Statistical analysis 51 4.3 Results and Discussion 51 4.3.1 Cell Morphology study 51 4.3.2 Cell Proliferation study 56 4.3.3 Alkaline phosphatase activity 58 4.3.4 Cell mineralization study 61 4.4 Conclusion 70 Chapter 5: Conclusions and Recommendations 5.1 Conclusions 72 5.2 Recommendations 73 6 LIST OF FIGURES Figure 1.1 A model Ti dental implant 3 Figure 2.1 Scaffold architecture affects cell binding and spreading 6 Figure 2.2 Schematic diagram of electrospinning set-up 18 Figure 3.1 Electrospinning set up 24 Figure 3.2 Electric field pattern a) 18kV at the needle tip and 10kV at the ring electrode, b) 18kV at the needle tip and 14kV at the ring electrode 24 Figure 3.3 Biomineralization procedure 26 Figure 3.4 SEM images of a) untreated Ti, b) Ti after surface modification c) Ti 34 coated with PLGA nanofibers at 1000X magnification d) Ti coated with PLGA/Collagen nanofibers at 1000X magnification e) Ti coated with PLGA nanofibers at 5000X magnification f) Ti coated with PLGA/Collagen nanofibers at 5000X magnification g) Ti coated with functionalized PLGA/Collagen nanofibers h) Ti coated with functionalized PLGA/Collagen nanofibers Figure 3.5 AFM image of pretreated Ti showing the surface roughness 35 Figure 3.6 FTIR results for a) cpTi treated and untreated, b) Ti6Al4V alloy treated and untreated, c) PLGA and PLGA/Collagen nanofibers coated over the Ti surface. 37 Figure 3.7 XPS results showing the Ti2p peaks in the treated samples 39 7 Figure 3.8A Adhesion of hMSCs on the a) untreated cpTi implants, b) cpTi 42 implant coated with PLGA nanofibers, c) cpTi implant coated with PLGA/Collagen nanofibers, d) cpTi implant coated with PLGA/HA, e) cpTi implant coated with PLGA/Collagen/HA nanofibers at 500x Figure 3.8B Adhesion of hMSCs on the a) untreated Ti6Al4V implants, 43 b) Ti6Al4V implant coated with PLGA nanofibers, c) Ti6Al4V implant coated with PLGA/Collagen nanofibers, d) Ti6Al4V implant coated with PLGA/HA, e) Ti6Al4V implant coated with PLGA/Collagen/HA nanofibers at 500x Figure 3.9 Percentage attachment efficiency of hMSCs on cpTi and Ti6Al4V alloy 45 Figure 4.1 SEM images of the hMSC morphology on day 7 on a) untreated Ti, b) 53 Treated Ti coated with PLGA nanofibers, c) Treated Ti coated with PLGA/Collagen nanofibers, d) Treated Ti coated with functionalized PLGA nanofibers, e) Treated Ti coated with functionalized PLGA/Collagen nanofibers. Figure 4.2 SEM images of the hMSC morphology on day 14 on a) untreated Ti, b) 54 Treated Ti coated with PLGA nanofibers, c) Treated Ti coated with PLGA/Collagen nanofibers, d) Treated Ti coated with functionalized PLGA nanofibers, e) Treated Ti coated with functionalized PLGA/Collagen nanofibers. Figure 4.3 SEM images of the hMSC morphology on day 21 on a) untreated Ti, b) 55 Treated Ti coated with PLGA nanofibers, c) Treated Ti coated with PLGA/Collagen nanofibers, d) Treated Ti coated with functionalized PLGA nanofibers, e) Treated Ti coated with functionalized PLGA/Collagen nanofibers. Figure 4.4 MTS assay for hMSC cells proliferation on a) cpTi based 57 scaffolds untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers b) Ti-6Al-4V based scaffolds - untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers for day 7, 14 and 21. * represents p≤ 0.05 statistical difference. Control refers to the Tissue Culture Plate (TCP); TiK refers to Ti6Al4V alloy. Figure 4.5 ALP activity for hMSC cells on a) cpTi based 60 scaffolds - untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, 8 coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers b) Ti-6Al-4V based scaffolds - untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers for day 7, 14 and 21. * represents p≤ 0.05 statistical difference. Control refers to the Tissue Culture Plate (TCP); TiK refers to Ti6Al4V alloy. Figure 4.6 Quantitative data for Alizarin red staining on hMSC cells on a) cp Ti 63 scaffolds b) Ti-6Al-4V scaffolds for days 7, 14 and 21. * represents p≤ 0.05 statistical difference Figure 4.7A Optical image of the ARS stained hMSCs on the cp Ti scaffolds 64 on day 7 a) untreated Ti, b) Treated Ti coated with PLGA nanofibers, c) Treated Ti coated with PLGA/Collagen nanofibers, d) Treated Ti coated with functionalized PLGA nanofibers, e) Treated Ti coated with functionalized PLGA/Collagen nanofibers. Figure 4.7B Optical image of the ARS stained hMSCs on the cpTi 65 scaffolds on day 14 a) untreated cpTi, b) Treated cpTi coated with PLGA nanofibers, c) Treated cpTi coated with PLGA/Collagen nanofibers, d) Treated cpTi coated with functionalized PLGA nanofibers, e) Treated cpTi coated with functionalized PLGA/Collagen nanofibers. Figure 4.7C Optical image of the ARS stained hMSCs on the cpTi scaffolds 66 on day 21 a) untreated cpTi, b) Treated cpTi coated with PLGA nanofibers, c) Treated cpTi coated with PLGA/Collagen nanofibers, d) Treated cpTi coated with functionalized PLGA nanofibers, e) Treated cpTi coated with functionalized PLGA/Collagen nanofibers. Figure 4.8A Optical image of the ARS stained hMSCs on the Ti6Al4V 67 scaffolds on day 7 a) untreated Ti6Al4V, b) Treated Ti6Al4V coated with PLGA nanofibers, c) Treated Ti6Al4V coated with PLGA/Collagen nanofibers, d) Treated Ti6Al4V coated with functionalized PLGA nanofibers, e) Treated Ti6Al4V coated with functionalized PLGA/Collagen nanofibers. Figure 4.8B Optical image of the ARS stained hMSCs on the Ti6Al4V 68 scaffolds on day 14 a) untreated Ti6Al4V, b) Treated Ti6Al4V coated with PLGA nanofibers, c) Treated Ti6Al4V coated with PLGA/Collagen nanofibers, d) Treated Ti6Al4V coated with functionalized PLGA nanofibers, e) Treated Ti6Al4V coated with functionalized PLGA/Collagen nanofibers. Figure 4.8C Optical image of the ARS stained hMSCs on the Ti6Al4V 69 scaffolds on day 21 a) untreated Ti6Al4V, b) Treated Ti6Al4V coated with PLGA nanofibers, c) Treated Ti6Al4V coated with PLGA/Collagen nanofibers, d) Treated Ti6Al4V coated with 9 functionalized PLGA nanofibers, e) Treated Ti6Al4V coated with functionalized PLGA/Collagen nanofibers. 10 LIST OF TABLES Table 2.1 Different types of implant surface modifications and their surface 7 roughness and contact angle. Table 2.2: Various fabrication techniques along with their advantages and 12 disadvantages Table 2.3 commonly used polymers and their properties Table 2.4. Factors that affect the electrospinning process and fiber morphology Table 3.1 Optimization of electrospinning parameters by varying the time and 15 16 32 concentration for PLGA nanofibers Table 3.2 Optimization of electrospinning parameters by varying the time and 32 concentration for PLGA/Collagen nanofibers Table 3.3 Water contact angle measurements for treated and untreated cpTi and 38 Ti6Al4V alloy Table 3.4 Water contact angle measurements for PLGA and PLGA/Collagen 39 nanofibers Table 3.5: Average number of cells adhered to the Ti samples 44 11 LIST OF APPENDICES Appendix A: Optical image of hMSC morphology cultured on TCP Appendix B: FESEM EDX results showing cell mineralization. 12 LIST OF ABBREVIATIONS AFM atomic force microscopy ECM extracellular matrix HFP 1,1,1,3,3,3-hexafluoro-2-propanol kDa unit of 1000 Dalton PBS phosphate buffered saline PLGA poly(lactic acid)-co-poly(glycolic acid) SEM scanning electron microscopy w weight v volume XPS x-ray photoelectron spectrometry avg average 13 SUMMARY The introduction of dental implants has changed the way dentists approach the replacement of missing teeth. The clinical success of dental implants is related to their osseointegration, which is a property virtually unique to titanium and has enhanced the science of joint replacement techniques. Generally, the time between implant placement and implant loading ranged from 3 months in the mandible to 6 months in the maxilla, for machined surfaces. However, the trend towards a shorter healing time is largely driven by consumer demands as many patients are unhappy waiting long periods of time for their prosthesis. In order to achieve rapid osseointegration, it is necessary that the implant surface has an improved capture ratio which will provide a critical number of mesenchymal stem cells (MSCs) necessary for successful bone integration [63]. We have proven that the fabrication of a nanofibrous scaffold offers the possibility to optimize stem cell capture as well as cell adhesion and proliferation, as the nanofibers mimic the ECM matrix. It is our hypothesis that this improved capture ratio will provide a critical number of MSCs necessary for successful bone integration. Thus the healing time can be reduced, leading to enhanced initial osseointegration. In this study, we have proven the feasibility of creating a nanotextured surface on titanium by using a simple acid/alkali treatment. The surface roughness can be tailored by modifying the etching/ polishing procedures. Besides we have demonstrated that the cell 14 adhesion can be increased by coating the titanium surface with nanofibers. This is because the nanofibers mimic the natural ECM and hence improve cell attachment. Through our advanced electrospinning set-up we have achieved more fiber deposition at a shorter interval of time than conventional electrospinning. Moreover we have shown that the adhesion efficiency of the human bone marrow derived MSCs was the maximum on the biomineralized PLGA/Collagen nanofibers coated Ti compared to the other samples. Furthermore, incorporation of biomolecular cue like collagen and nano-HA have enhanced the cell proliferation, osteogenic differentiation and cell mineralization. To our knowledge, dental implant using functionalized nanofibers as a surface modification is a novel idea to enhance osseointegration using the bone regeneration concept. 15 Chapter 1 Introduction 1.1 Background In the past 20 years, the number of dental implant procedures has increased steadily worldwide, reaching about one million dental implantations per year [1]. Dental implants are useful for restoration of oral function, including mastication and speech, as well as for aesthetic improvement in patients with tooth loss. The clinical success of dental implants is related to their early osseointegration. The other implant related to the early osseointegration is total joint replacement, which is an effective treatment for relieving pain and restoring range of motion. Osseointegration may be defined as the direct structural and functional connection between living bone and the surface of a loadbearing artificial implant, typically made of titanium. It is a property virtually unique to titanium and hydroxylapatite, and has enhanced the science of medical bone, and joint replacement techniques. As long as implants are positioned correctly and infection is avoided, they will generally last for many years. Geometry and surface topography are crucial for the short- and long-term success of the implants. These parameters are associated with delicate surgical techniques, a prerequisite for a successful early clinical outcome. High success rates for dental implants are reported in healthy patients with good bone quality. In the future, with an aging population, more patients may be considered for dental implants; osseointegration of dental implants under less than optimal circumstances and reduced bone healing quality may then be encountered. In such cases, enhanced bone formation around the implant would be an important criterion. 16 This may be achieved by implant coatings that are able to interact actively with the surrounding tissues. 1.2 Clinical problems associated with osseointegration: There are two types of responses exhibited by the body after implantation. The first type involves the formation of a soft fibrous tissue around the implant. This fibrous tissue does not ensure proper osseointegration and leads to the clinical failure of the dental implant. The second type of bone response is related to direct bone–implant contact without an intervening connective tissue layer. This is the desired response after implantation. From the clinical point of view, during osseointegration, two factors play an important role: primary stability (mechanical stability) and secondary stability (biological stability after bone remodelling). Primary stability is the mechanical stability of the implant as soon as the implant is placed into the bone. It gradually decreases in the bone remodelling process. Secondary stability involves the formation of new bone with the implant after bone remodelling. Primary stability is fully replaced by secondary stability when the healing process is completed. However, at one point, the implant stability decreases during the stability conversion, a process also called the ―dip‖. Many implant failures occur during this period, and this period seems to be critical to the successful integration of the implant [2]. 17 Figure 1: A typical Ti dental implant 18 1.3 Hypothesis and Objectives: Hypothesis This project is to develop a surface modification system for dental implant using electrospun nanofiber and biomineralization to fabricate a biomimetic substrate. We hypothesized that both substrate topographical and biochemical cues promote mesenchymal stem cells (MSCs) adhesive behaviors, followed by proliferation and differentiation, which are crucial for enhanced osseointegration. Objectives:  Modify the implant surface to produce nanotextured topography.  Develop a nanofibrous coating from biodegradable synthetic polymers and/or natural polymers to mimic extracellular matrix.  Functionalization of the nanofiber by biomineralization.  Evaluate adhesion, proliferation of MSCs on the modified implant surface.  Investigate osteogenic differentiation and mineralization of MSCs on the modified implant surface. 19 Chapter 2 Literature Review 2.1 Introduction The native Extracellular Matrix (ECM) consists of nano- to micro- structured fibers (proteins and proteoglycans). This hierarchical organization presents a defined environment with nano-scale intermolecular binding interactions that will affect the morphological and functional development of the cells. Recent studies have shown the importance of nano-textured implant surface for tissue engineering applications [3]. Cells that were cultured on micro-size fibrous scaffolds were flattened and the cells spread as if they were cultured on flat surfaces (Figure 2.1) [4]. Scaffolds with nano-scale architectures have larger surface area to adsorb proteins and present many more binding sites to cell membrane receptors would be more biomimetic to support better cell-matrix interactions [4]. Thus the presentation of suitable topographical cues is an important aspect to consider when designing tissue engineered scaffolds. 20 Figure 2.1 Scaffold architecture affects cell binding and spreading [4] 2.2. Surface modification techniques To generate topographical cues on the implant surface, in order to enhance osseointegration process, several surface modification techniques have been tried as shown in the Table 2.1. The nanostructured surfaces of nanometallic and nanoceramic materials have several advantages compared to the conventional surfaces. These include, (i) they possess greater surface roughness resulting from both decreased grain size and possibly decreased diameter of surface pores, (ii) enhanced surface wettability due to greater surface roughness and (iii) greater numbers of grain boundaries. There are a number of physical and chemical techniques that can be used for the surface modification or activation of an implant surface. Among these methods, chemical modifications seem to be relatively simple and inexpensive. Hence it is widely used. There have been various techniques tried out in the past to improve the surface roughness of the implant like plasma treatment, acid-etching and heat treatment. For example, the TPS (titanium 21 plasma sprayed) surfaces used by Straumann recommended a healing period of 12 weeks [5] and this was reduced to 6 to 8 weeks with the introduction of the SLA (sand blasted, acid etched) surface [6]. The differences in the contact angle and the surface roughness of the implant surface owing to the various surface modification techniques were shown in Table 2.1 Table 2.1 Different types of implant surface modifications and their surface roughness and contact angle. Type of implant Surface roughness Contact (μm) cpTi (Commercially pure Ra = 0.22 ± 0.01 angle References (°) 55.4 ± 4.1 [7], [8] Ti) Ti6Al4V Ra = 0.23 ± 0.01 56.3 ± 2.7 [7], [8] TPS Ra = 7.01 ± 2.09 n.d. [7] SLA Sa = 1.15 ± 0.05 138.3 ± 4.2 [9] Modified SLA Sa = 1.16 ± 0.04 0 [9] Plasma-sprayed HA coating Ra = 1.06 ± 0.21 57.4 ± 3.2 [10], [11] Mitsuru Takemoto et al., compared HCl–Alkali-heat treatment, alkali and heat treatment and water–acid–alkali treatments [12]. He demonstrated that dilute HCl treatment 22 effectively removed sodium from the sodium titanate layer of alkali-treated porous titanium and contributed to the formation of the titania layer on the surface of porous bioactive titanium. Furthermore, the HCl–Alkali-heat treated implants possessed a more complex surface when compared to other treatments, which may have been caused by an etching effect of the dilute HCl treatment. The results of this study indicated that chemistry and topography were related to material-induced osteoinduction as the dilute HCl treatment was considered to give both chemical (titania formation and sodium removal) and topographic (etching) effects on the titanium surface [12]. Timothy et al., adopted porous bone metal implant strategy to improve implant fixation, as it allows for the ingrowths of bone and also reduces the Young’s modulus of the implant material to better match that of bone. Besides bone ingrowths it also reduces the risks associated with the bone resorption due to stiffness mismatch [13]. It was demonstrated that the treatment of Ti with a NaOH solution followed by heat treatment at 873 K forms a crystalline phase of sodium titanate layer on the Ti surface resulting in improved adhesion of apatite coating prepared by incubation in simulated body fluid (SBF). The authors concluded that the released sodium ions from the sodium titanate layer caused the formation of Ti–OH groups that react with the calcium ions from the SBF and form calcium titanate, which then could act as nucleation sites for apatite crystal formation [14, 15]. Lewandowska et al., characterized the chemical composition and morphology of titanium surfaces exposed to acidic, alkaline or polymer solutions. It was found that there were large differences in the morphology of Ti pretreated with different procedures whereas 23 only minor differences in the chemistry of the surfaces. In all the cases TiO2 being the principle chemical component [16]. The Ti metal spontaneously forms a protective TiO2 layer in the atmosphere. When the Ti implant is inserted into the human body, the surrounding tissues directly contact the TiO2 layer on the implant surface. The surface characteristics of the TiO2 layer determine the biocompatibility of Ti implant. Therefore, it is important to use appropriate surface modifications to increase the biocompatibility of the Ti implant for long-term clinical applications. Several chemical etching agents like sodium hydroxide, hydrogen peroxide and hydrofluoric acid have been used to improve the TiO2 layer, which is responsible for the excellent corrosion resistance of the implant. In the body, however, mechanical friction and chemical influences might lead to rupture or weakening of the TiO2 layer, leading to a corrosion processes and the formation of wear debris in such regions [17]. Meanwhile, Nishiguchi et al., compared the bone-bonding ability of alkali- and heattreated titanium with that treated in NaOH without subsequent heat treatment. It was concluded that the NaOH-treated titanium without heat treatment had no bone-bonding ability due to its unstable reactive surface layer. He also demonstrated that soaking the implant in NaOH solution stimulated the bone ingrowths onto the surface of the implant [18]. 2.2.1 Modification of the implant surface using surface adhesive molecules In native tissues, ECM presents their adhesion proteins such as laminin, collagen, fibronectin, and vitronectin to effect cell attachment through the binding between integrin 24 receptors on the cell surfaces. Therefore much work is done to enhance the biocompatibility of polymeric tissue engineered scaffolds to create a biochemical-like environment on the biomaterial surfaces [3]. Biomolecules such as adhesive proteins like collagen, RGD peptides, fibronectin and growth factors like basic fibroblast growth factor and epidermal growth factor that can be easily recognized by the cells can be coupled onto the biomaterials to induce biorecognition mechanisms of the interaction of cells and polymeric biomaterial scaffolds. These modifications can preserve the mechanical integrity of polymeric scaffolds while creating an ECM-like environment to the scaffolds. The surface chemistry of the implant also plays an important role in deciding the cell characteristics. For example it was reported that arginine-glycine-aspartic acid (RGD)-coated Ti disks greatly promoted attachment and decreased apoptosis of MC3T3-E1 osteoprogenitor cells. Coating the nanofibers with RGD or another positively charged molecule, such as calcium ion or poly-lysine, may promote the attachment of cells. [19] Currently, the most popular surface treatment for commercial artificial joints and dental implants is plasma-spray coating with hydroxyapatite (HA). Plasma-sprayed hydroxyapatite on titanium has been reported to show beneficial effects such as osteoconductivity and direct-bone bonding ability [20]. However, the process has disadvantages attributed to the high temperatures used during the process, such as the possibility of fracture at the interface between the titanium and the HA due to the residual stress at the interface, and changes in the composition, porosity, crystallinity, and structure of the plasma-sprayed hydroxyapatite [21]. Therefore, new HA coating methods 25 have attracted great interests in recent years for replacing the high temperature techniques like plasma spraying. Besides, clinical trials were done by Wang et al., on canine trabecullar bone. He studied the osseointegration of uncoated, Plasma- sprayed -HA-coated and electrodeposition – HA -coated Ti–6Al–4V in a canine trabecular bone at 6 h, 7 days and 14 days postimplantation. The Plasma sprayed -HA was found to provide higher bone apposition ratio than those exhibited by the bare alloy and electrodeposited-HA, owing to their earliest mineralization (6 h—7 days) in the form of nano-ribbon cluster mineral deposits with a Ca/P atomic ratio lower than that of hydroxyapatite [22]. In another study, pure titanium was subjected to various surface modifications and examined in terms of morphology, chemical characteristics and wettability. The results showed that etching in alkaline or acid solutions resulted in significant changes in surface morphology; a characteristic feature for the presence of sub-microporosity [23]. An earlier work done by Nicula et al., compared cp Ti, Ti–Al–V, Ti–Al–V–Cr and Ti– Al–Mn–V–Cr prepared by high-energy ball-milling method, to achieve a microtextured suface. Optimal cell adhesion was observed for the Ti–Mn–V–Cr–Al alloy, which might be due to the surface morphology of this specimen (high-roughness, porosity in the micron range). Thus the results showed that the surface properties are important for implant materials, since the surface topography influences the mechanisms of cell adhesion and growth [24]. 26 The biomimetic scaffolds for tissue engineering can be manufactured by various processes like electrospinning, phase separation, self–assembly and lithography. Comparisons between the various techniques are shown in Table 2.2 Table 2.2: Various fabrication techniques along with their advantages and disadvantages Fabrication Advantages Drawbacks technique Laser Uniform distribution of pore size, simple Reduced resolution and deposition and fast. Self assembly Can generate fibrous networks capable Lack poor surface finish mechanical of supporting cells in three dimensions. strength, Limited Cell-seeding problems associated with amphiphilic using prefabricated scaffolds eliminated materials, nanofibrous random and very short owing to nanofibers. spontaneous assembly. Lithography Relatively good resolution. Time consuming and expensive. Electrospinning The properties of electrospun nanofibers, Electrospinning yields a such as fiber diameter, can be controlled flat mat that has limited readily via manipulation of spinning three dimensionality parameters. Capable of mimicking the and suffers from cell stem cell niche. infiltration problems because of the small pore size of the mats Phase A nanofibrous 3D scaffold can be Nanofiber distribution is separation constructed. Has controllable high subject to the the 27 porosity and surface-to- volume ratios. processing. 2.2.2 Cell – substrate interaction The implant's surface properties, surface chemistry, surface energy, topography and roughness influence the initial cell response at the cell - material interface, ultimately affecting the neo-tissue formation. Recent studies have shown higher osteoblasts adhesion and enhanced alkaline phosphatase activity on rough Ti and Ti-6Al-4V [25, 26]. It is well known that cell response is affected by the physicochemical parameters of the biomaterial surface, such as surface energy, surface charges or chemical composition. Topography is one of the most crucial physical cues for stem cells and recently it has been proven that nanotopography plays the main influencing factor, rather than microtopography [27]. Though the surface modification techniques like grit-blasting, plasma treatment, sand blasting, have been successful, the time required for osseointegration ranges from 3 to 6 months. Osteoblasts adhesion on nanostructured surfaces was first reported in 1999 by Webster et al., [28]. He demonstrated that osteoblasts adhesion was improved when they were cultured on nanostructured surfaces, compared to the conventional micro surfaces. Specifically, alumina with grain sizes between 49 and 67 nm and titania with grain sizes 28 between 32 and 56 nm enhanced osteoblast adhesion compared to their respective micrograined materials. It has been proved that the contact of cells to the surface of the biomaterials results in changes to the cell shape and bioactivity depending on the topography of the surface [29]. For instance, cells cultured on pure Ti and Ti alloy exhibit differences in cell response even though both are covered with TiO2 oxide layer. These differences may be attributed to the surface morphology and chemistry differences between the two. 2.3 Tissue Engineering 2.3.1 Introduction The current medical need is to address bone graft problems such as implant failure owing to lack of tissue regeneration around the implant surface, resulting in poor bone remodelling and loosening of the implants. In recent years, tissue engineering has revolutionized the direction of research for orthopaedic applications because of the success of nanotechnological advancements in creating new fabrication techniques for nano-scale materials such as nanofibers and nanofibrous scaffolds. Previous studies conducted by Ngiam et al., proved that n-HA on PLGA and PLGA/Collagen had a positive modulation on early capture of osteoblasts compared to the non-functionalized nanofibers. However no studies have been reported on the influence of hMSCs on the functionalized nanofibers. The main advantage of using hMSCs for tissue engineering applications is because of its direct clinical applications [30]. 29 2.3.2 Nanofiber fabrication by electrospinning Electrospinning is a simple and versatile technique that can produce non-woven ECMlike nanofiber scaffolds with nano-topographical cues to interact with the cells. Synthetic polymeric nanofibers such as poly(ε-caprolactone) (PCL) [31], poly(L-lactic acid) (PLLA) [32], poly(glycolic acid) (PGA) [33] and poly(lactic-co-glycolic acid) (PLGA) [34], and natural-occurring polymeric nanofibers such as collagen [35] and gelatin [36] have been widely explored for applications in the different areas of tissue engineering such as skin, cartilage, bone, blood vessel, heart, and nerve [31 - 40]. The properties of the commonly used polymers are discussed in Table 2.3. Table 2.3 Commonly used polymers and their properties Polymer Properties Degradation rate Reference PGA Aliphatic polyester, 6-12 months [41] > 24 months [42 – 44] Crystalline, semi permeable PLLA Aliphatic polyester, crystalline, porous; Roughlooking due to the open-pore structure PLGA Semi pemeable 6 – 12 months [40] PCL Semi permeable, amorphous < 12 months [45] Collagen Semi pemeable 1 – 9 months [46] 30 Electrospinning process utilizes an electric field generated by an applied voltage that subsequently introduces surface charges to the polymer solution. This results in the formation of a Taylor cone polymeric droplet at the tip of the spinneret. Once the electric potential that is created at the droplet surface exceeds a critical value, the electrostatic forces will overcome the solution surface tension to initiate a polymer jet stream. The charged jet is accelerated towards the grounded collector and undergoes bending instability, elongation, and solvent evaporation or jet solidification which leads to rapid thinning of the jet and deposition of dry fibers in a random manner onto the collector [33, 41, 42]. The experimental set up for electrospinning is shown in Figure 2.2. Several factors can affect the electrospinning process and fiber morphology (Table 2.4). Table 2.4. Factors that affect the electrospinning process and fiber morphology [47]. Process Parameter Viscosity/concentration Effect on fiber morphology  Low concentrations/viscosities yielded defects in the form of beads and unction; increasing concentration/viscosity reduced the defects;  Fiber diameters increased with increasing concentration/viscosity. Conductivity  Increasing the conductivity aided in the production of uniform bead-free fibers; 31  Higher conductivities yielded smaller fibers in general (except PAA and polyamide-6). Polymer molecular  weight Dipole Increasing molecular weight reduced the number of beads and droplets. moment and  dielectric constant Flow rate Successful spinning occurred in solvents with a high dielectric constant.  Lower flow rates yielded fibers with smaller diameters;  High flow rates produced fibers that were not dry upon reaching the collector. Field strength/voltage  At too high voltage, beading was observed;  Correlation between voltage and fiber diameter was ambiguous. Distance between tip  and collector A minimum distance was required to obtain dried fibers;  At distance either too close or too far, beading was observed. Fiber morphology  Smooth fibers resulted from metal collectors;  Aligned fibers were obtained using a conductive 32 frame, rotating drum, or a wheel-like bobbin collector; Ambient parameters  Yarns and braided fibers were also obtained.  Increased temperature caused a decrease in solution viscosity, resulting in smaller fibers;  Increasing humidity resulted in the appearance of circular pores on the fibers. Figure 2.2. Schematic diagram of electrospinning set-up In a work done by Ma et al., three different materials, silicon (Si), silicon oxide (SiO2), and titanium oxide (TiO2), were used to construct nanofibers for surface coating of Ti alloy Ti-6Al-4V. The results demonstrated that TiO2 nanofibers coated over the Ti alloy facilitated a higher adhesion potential and higher cellular differentiation capacity than Ti alloy and tissue culture–treated polystyrene surfaces (TCP). Thus, surface modification 33 using nanofibers of various materials was proved to alter the attachment, proliferation, and differentiation of osteoprogenitor cells in vitro [48]. It was also reported that nanofibrous poly (L-lactide) (PLLA) scaffold fabricated by phase separation and particle-leaching method showed biological function similar to those of the collagen fibers of bone [49]. These results might implicate the possibility that a nanofibrous surface can improve the osseointegration of implants. However, these nanofibrous materials such as carbon and organic polymer are difficult to be immobilized on titanium surface because of their low reactivity with titanium [50]. 2.3.3 Modifications of the electrospun nanofibers At present HA has been widely used as bioceramics in orthopaedics and dentistry due its osteoconductive properties [29]. In the native bone tissue, HA nanocrystals grow in intimate contact within collagen fibers, building up a nano-structured composite. However, HA has a disadvantage that is attributed to low mechanical strength. Hence the combination of a load bearing biomaterial like titanium with the osteoconductive properties of HA is very attractive. HA related bone formation is believed to begin with surface dissolution of the HA, which releases calcium and phosphate ions into the vicinity around the implant. Reprecipitation of carbonated apatite then occurs on the coating surface, thereby enhancing osteoblasts adhesion onto the surface. 34 Immobilization reaction of TiO2 nanofibers on the titanium plate was done by treating the Titanium plates firstly in alkali and then in acid solutions. When immersed in NaOH, the passive oxide layer of titanium dissolves to form amorphous titanate layer containing Na+ ions. Immediately after immersion in simulated body fluid (SBF), Na+ ions from the amorphous layer will be exchanged by H3O+ ions from the surrounding fluid resulting in the formation of Ti–OH layer. And then hydroxyapatite was formed on titanium surface by ionic bonding between Ti–O anions and Ca2+ cations in SBF [51]. Thus, biomineralization originated from native process may provide some effective way for osseointegration. In another study collagen fibrils/carbonate-hydroxyapatite coating has been electrodeposited on Ti plates using Ca (NO3)2 and NH4H2PO4 solutions in a type I collagen molecule suspension [52]. 2.3.4 Potential application of mesenchymal stem cells for osseointegration Stem cells are unspecialized cells that can self renew indefinitely and differentiate into several somatic cells with proper environmental cues. In stem cell niche, the stem cell– ECM interactions are very crucial for different cellular functions like adhesion, proliferation and differentiation. Most recently, the importance of nanometric scale surface topography, and roughness of biomaterials is, besides chemical surface modifications, increasingly becoming recognized as a crucial factor as synthetic ECM for cell survival and host tissue acceptance. 35 Recent work by Muschler [53, 54] demonstrated that it is possible to capture MSCs on substrate such as allograft bone. He has developed a system where it is able to capture MSCs on allograft bone with an enrichment-factor of 3-4x at best. A much higher theoretical capture ratio is possible. The fabrication of a nanofibrous scaffold offers the possibility to optimize cell capture as well as cell adhesion and proliferation. Furthermore, MSCs derived from the bone marrow of neonatal rats, were used for seeding on electrospun PCL scaffolds by Yoshimoto et al., [31]. MSCs not only attached favourably and grew well on the surface of these scaffolds, but the MSCs were also able to migrate inside the scaffold up to 114 µm within 1 week of culture. Gelatin/PCL shows better biocompatibility than PCL nanofibrous material. The enhanced adhesion and proliferation of MSCs on nanofibers matrix also showed up on PLA and silk electrospun nanofibers [50, 51]. Hosseinkhani et al., investigated mesenchymal stem cell (MSC) behavior on self-assembled peptide-amphiphile (PA) nanofiber scaffolds [55] Significantly enhanced osteogenic differentiation of MSC occurred in the 3D PA scaffold compared to 2D static tissue culture. 36 Chapter 3 Biomimetic surface modification of dental implant materials by advanced electrospinning 3.1 Introduction Implants can be divided into smooth (machined or turned) and rough on the basis of surface roughness. The techniques used for preparing the surface roughness maybe either additive or subtractive in nature. Additive techniques involve coating the implants with titanium or HA using plasma spraying technique or sintering. Subtraction techniques involve the use of sandblasting or acid/alkali etching treatments. Titanium (Ti) has been widely used as implant materials in the dentistry and orthopaedics owing to their excellent mechanical properties and biocompatibility [1]. Some of these properties, in particular the biological response of titanium, are strongly determined by the surface characteristics— its morphology, chemistry and physical properties. Ti and Ti alloy facilitate new bone formation and provide long-lasting bone-implant stability. In addition to being bio-inert and nontoxic, requirements for the next generation of biomaterials include enhanced cell attachment and differentiation to accelerate osseointegration of implants. Modified or coated Ti and its alloys have become candidates for next-generation implants. Surface properties may be changed by applying various surface modifications while the crucial bulk properties such as tensile strength and fatigue resistance remain unchanged. However implant failures do occur owing to 37 loosening of the implants. One of the main strategies to enhance osteoconduction is the use of a nanofiber-coated surface [56]. A nanofiber coating on Ti constructs a rough surface, which may stimulate bone formation by triggering specific cell responses. Our strategy is to design and fabricate biomimetic and bioactive implant surfaces that resemble the native extracellular matrix (ECM) as closely as possible so as to create conducive living milieu that will induce cells to function naturally. In this context, our current endeavor is to use the natural polymer collagen along with PLGA as a matrix and to deposit n-HA (nano – HA) by Calcium-phosphate (Ca-P) dipping method so as to develop biomimetic n-HA containing nanocomposite nanofibers. 3.2 Materials and Methods 3.2.1 Mechanical Polishing/ etching: Pure Titanium (15mm diameter) and Titanium alloy (Ti- 6Al- 4V) discs (25mm diameter), purchased from Northwest Institute for Non – Ferrous Metal Research (Xian, Shanxi, P.R. China) were mechanically polished using 320 grit and 400 grit SiC papers, till a mirror finish was achieved. The discs were further polished using alumina (1M) cloth for a smoother finish. The discs were then cleaned with ethanol using an ultrasonicator for 15min. This ensures the removal of the impurities arisen due to the mechanical polishing. The mechanical treatment was followed by chemical etching using 4% HNO3 in ethanol for 1min. The discs were then allowed to dry at room temperature. 38 3.2.2 Pretreatment of Ti The polished/etched Ti plates were immersed in 10N (Normality) NaOH solution at 600C for 24 hrs. The samples were then allowed to cool to the room temperature, followed by treatment with 10N HCl solution for 1hr. The samples were then dried. Titanium implants after the alkali treatment retained sodium and the sodium titanate layer with limited formation of titania layers. To overcome these problems, in addition to water treatment, a dilute hydrochloric acid (HCl) treatment was done, which almost completely removes sodium, even from deep pores [12]. 3.2.3 Electrospinning of PLGA and PLGA/Collagen nanofibers on the Ti discs The materials used for electrospinning were Type I collagen (Koken Co. Tokyo, Japan), PLGA (100,000 Da, Aldrich Chemical Company, Inc., St. Louis, U.S.) and 1,1,1,3,3hexafluoro-2-propanol (HFP, Aldrich Chemical Company, Inc., St. Louis, U.S.). PLGA (75:25) pellets were dissolved in HFP at a w/v ratio of 15%. The electrospinning parameters, the w/v ratio of PLGA in HFP and the fiber deposition time were optimized till uniform nanofibers without bead formation was obtained, as shown in Table 3.1. Electrospinning of blended PLGA/Collagen (50:50 w/w ratio) was also done following the same procedure (Table 3.2). The polymer solution was then loaded into a syringe (Becton Dickinson, BD, N.J, U.S.) and a high voltage electric field (DC high voltage power supply from Gamma High 39 Voltage Research, Florida, U.S.) was applied to draw the fibers from the spinneret (27G1/2 needle, Becton Dickinson, BD, N.J, U.S.) onto the collector plate, over which the Ti disc was placed. The experimental setup was shown in figure 3.1. The spinneret was first grounded to give a flat tip in order to produce continuous and uniform nanofibers. A constant feed rate of 1 mL/h was applied using a syringe pump (KD Scientific Inc., M.A., U.S.). Polymer solution spinneret Ring electrode Collector Figure 3.1 Electrospinning set up 40 Needle tip a Ring electrode b Z Z X X Figure 3.2 Electric field pattern a) 18kV at the needle tip and 10kV at the ring electrode 18kV at the needle tip and 14kV at the ring electrode b) In order to achieve focussed electrospinning, an additional ring electrode was used. A high voltage electric field supply was connected to the ring electrode as shown in Figure 3.1. The voltages across the two power supplies were optimized by analysing the electric field pattern developed from the two voltages, as shown in Figure 3.2. It was found that when the voltage at the needle was 18kV and the voltage at the ring electrode was 14kV, the electrospinning process was limited. This was because, at 14kV there were starting to appear irregularities in the electric field, as shown in the Figure 3.2. However when the voltage at the ring electrode was reduced to 10kV the electric field was more uniform without any features that could disturb electrospinning process. As the voltage was increased, the fiber deposition spot decreased. 41 The electrospun nanofibers were subsequently vacuum dried so that any residual solvent present could be removed. 3.2.4 Biomineralization using Calcium-Phosphate dipping method Biomineralization of nano-hydroxyapatite (n-HA) was achieved using Ca-P dip method. The Ti plates coated with electrospun PLGA and PLGA/Collagen nanofibers were initially immersed in 0.5M CaCl2 solution (Aldrich Chemical Company, Inc., St. Louis, U.S.), for 10 min. The samples were then rinsed in deionised water for 1min. The samples were then immersed in 0.3M of Na2HPO4 (Merck & Co. Inc., N.J, U.S.), for 10min and rinsed for 1min in DI water. This entire procedure was considered as 1 cycle. The scaffolds were subjected to 3 cycles of the above treatment. The first cycle was for 10min and the subsequent cycles were for 5min in each solution. After that, the scaffolds were removed and freeze dried overnight. The above process is schematically illustrated in Figure 3.3. 42 Biomineralization Procedure: Ti plate xxxxxxxxxxx x Nanofibers Freeze Dry (overnight) Figure 3.3 Biomineralization procedure 3.2.5 Cell adhesion study All the titanium samples were sterilized under UV light for 2 hrs. The discs were then washed with phosphate buffered saline (PBS) thrice for 15min each. Human bone marrow derived mesenchymal stem cells (hMSCs) (PT-2501, Lonza, USA) was cultured in DMEM low glucose medium (DMEM, Invitrogen, CA, U.S.) with 10% FBS (Invitrogen, CA, U.S.) and 1X Antibiotics (Sigma-Aldrich Chemical Company Inc., St. Louis, U.S) until 3 passages. The cells were then trypsinized and seeded onto all the samples (untreated Ti, Ti + PLGA nanofibers, Ti + PLGA/Collagen nanofibers, Ti + 43 PLGA nanofibers + n-HA, Ti + PLGA/Collagen nanofibers + n-HA) at the concentration of 10,000 cells per well. The well plates were incubated for different time points – 10, 20, 30 and 60 minutes at room temperature. After the incubation time, the media was removed and the plates were washed thrice with PBS to remove the unbound cells. The attachment efficiency of each sample was then evaluated using field emission scanning electron microscopy (FESEM). The samples were fixed with 3% glutaraldehyde for 3 hours. The scaffolds were then rinsed with distilled water for 15min and then dehydrated with a series of ethanol gradients starting from 30% to 50%, 75%, 90% and 100% (v/v). Subsequently the samples were treated with HMDS (Hexamethyldisilazane) solution and allowed to air – dry at room temperature in the fume hood. The samples were then gold coated (JEOL JFC-1200 fine coater, Japan) and the cells were counted using FESEM. Five locations were chosen on each scaffold – four corners and centre. The average number of cells was then counted from the chosen locations. The distance of the location was measured using Image J software. The total number of cells was then calculated for the entire dimension of the scaffold. 3.2.6 Surface characterization analysis The effect of the pretreatment on titanium was characterised using field-emission scanning electron microscopy (FESEM) (Quanta 200F, FEI, Oregon, U.S.). Prior to which the samples were gold coated (JEOL JFC-1200 fine coater, Japan). The biomineralized and the non-biomineralized electrospun nanofibers were also characterised using FESEM, to analyse the distribution of HA formed owing to the in 44 vitro biomineralization procedure. The diameter of the nanofibers was measured using image analysis software (Image J, National Institutes of Health, Bethesda, U.S.). 3.2.7 Surface roughness analysis The surface roughness of the Titanium discs were analysed before and after the pretreatment procedure using Profilometer (Surftest SV-400, Mitutoyo, U.S.), using a PC50 filter and at the rate of 0.5mm/s. Five discs were measured from each of the different substrates and the average roughness value Ra was calculated. The discs were further characterized using the Atomic Force Microscope (AFM) in tapping mode (Dimension 3100 AFM, Veeco Instruments Inc., CA, U.S.). The roughness data was then analysed using the Nanoscope software (Digital Systems, US). Similar to the profilometer analysis, five discs were chosen in random for each of the procedures and the average roughness value was analysed. 3.2.8 Fourier transform infrared spectroscopy (FTIR) and X-ray photoelectron spectroscopy (XPS) The TiO2 oxide layer on the titanium samples before and after the pretreatment was analysed using ATR mode FTIR (Bio-Rad FTIR FTS 3500). A universal sampling aperture at a grazing angle of 670 with respect to the surface was used. A spectral resolution of 4 cm-1 in the 400 – 4000cm-1 range was employed to analyze the TiO2 oxide 45 layer. Besides the electrospun PLGA and PLGA/Collagen nanofibers were also analyzed using FTIR to determine the functional groups present in them. The chemical composition of the untreated and the pretreated Ti discs were analyzed using XPS (Kratos AXIS HSi, X-ray Source: Mono Al K alpha, 15 kV, 10 mA (150 w)). The following were the parameters used: Photoelectron accept angle: 90 degree. Base pressure: 1.0X10 -9 Torr, working pressure: 1.0X10-8 Torr. 3.2.9 Water contact angle measurement The contact angle of the Ti discs was measured before and after the pretreatment procedure to study the influence of the treatment procedure on the wettability of the substrate. The contact angle measurements were done using VCA Optima Surface Analysis system (AST products, Billerica, MA). Distilled water was used for drop formation. 3.2.10 Statistical analysis Values (at least triplicate) were averaged and expressed as mean ± standard deviation (SD). Each experiment was repeated twice for cell adhesion. Diameter of nanofibers was calculated from 5 SEM images by randomly selecting 10 fibers from each SEM image. Statistical differences were determined by Student two-sample t test. Differences were considered statistically significant at p ≤ 0.05. 46 3.3 Results and Discussion 3.3.1 Surface characterization analysis Electrospinning is a very simple and versatile process by which polymer nanofibers with diameters ranging from a few nanometers to several micrometers can be produced using an electrostatically driven jet of polymer solution. Figure 3.1 shows the conventional electrospinning set-up. In a typical electrospinning process, an electrical potential is applied between a droplet of a polymer solution, held at the end of a capillary tube and a grounded target. When the applied electric field overcomes the surface tension of the droplet, a charged jet of polymer solution is ejected. The route of the charged jet is controlled by the electric field. This inherent property of the electrospinning process, favors the control of deposition of polymer fibers onto any target substrate [57]. In the work by Theron et al., an electrostatic field-assisted assembly technique was described, which in combination with an electrospinning process was used to position and align nanofibers on a tapered and grounded wheel-like bobbin [58]. Our experiment suggested the targeted deposition of the nanofibers onto the cpTi (commercially pure Ti) and Ti alloy (Ti6Al4V) discs. Results were also reported by Hohman, Shin, Rutlege and Brenner, who studied electrospinning with regard to electrically forced jet and instabilities, and proposed a stability theory for electrified fluid jets [59 - 62]. It was demonstrated that at increasing field strengths, the electrical instabilities are enhanced [59]. This was in correlation with the results obtained in our study using advanced electrospinning technique; as shown in Figure 3.2. Wherein, on the left diagram it is 18kV at the needle and 10kV at the ring; on the right it is 18kV at the needle and 14kV at the ring. As shown, for 14kV there are starting to appear irregularities in the electric field. 47 A bit higher voltage and this irregularity was bigger, also indicating a decrease in the fiber deposition diameter. In fact it is repelling the electric field from the ring, but still too small to stop the electrospinning action. On the contrary electric field for 10kV at the ring is much more uniform without any features that could disturb electrospinning process. Green lines indicated in the Figure 3.2 are only for representation of the decrease of the fiber deposition spot. These are protons moving in given electric field and cannot be referred to as nanofiber. Additionally in correlation with the earlier studies it is seen that with higher voltage, fiber deposition spot should be smaller as proven in the experiment. The deposition of n-HA was achieved by a feasible Ca-P dipping method as shown in Figure 3.3. The SEM images (Figure 3.4a) of the untreated Ti discs revealed no distinctive surface topography. However after the pretreatment, the SEM images show topographical distribution of α and β grains (Figure 3.4b). This shows that the acid treatment has led to conversion of the initial microtextured surface to a nanotextured surface. The morphology of the PLGA and the PLGA/Collagen nanofibers were analyzed using SEM images (Figure 3.4c – 3.4f). It was not possible to measure the tensile strength of the PLGA and PLGA/Collagen coated implant surface owing to the hard nature of the implant material and the thin layer of nanofiber coating on the implant surface. The tensile properties of PLGA and PLGA/Collagen nanofibers have been reported by Kun Ma et al., where Young’s modulus (MPa) of PLGA and PLGA/Collagen nanofibers was reported to be 190.42 ± 9.97 and 40.43 ± 3.53 respectively. Ultimate tensile stress (MPa) was 4.82 ± 0.33 and 1.22 ± 0.12 for PLGA and PLGA/Collagen nanofibers respectively [63]. The deposition time and the concentration parameters were varied till an optimum 48 fiber diameter was achieved as shown in Table 3.1 and Table 3.2. From the SEM micrographs it was seen that as the fiber deposition time increased beyond 15 seconds, the amount of fiber deposited also increased. In order to ensure osseointegration, it is therefore desirable that the cells contact both the nanofibers and the nanotopography of the Ti substrate. For the deposition time of 10 seconds and 5 seconds, the fiber deposition was not uniform as can be seen from the SEM images. Hence 15 seconds was chosen as the optimum deposition time for further studies. From the Tables 3.1 and 3.2, it was seen that as the polymer concentration was increased, the fiber diameter also increased. For polymer concentrations less than 10% and 15% in the case of PLGA/Collagen and PLGA respectively, even though the fiber diameter was smaller, bead formation and nonuniformity of fibers was seen. Hence for further studies PLGA nanofibers were spun at a polymer concentration of 15% and deposition time of 15 seconds and PLGA/Collagen at a concentration of 10% and deposition time of 15 seconds. The deposition of n-HA after the biomineralization treatment using Ca-P dipping method was also shown using FESEM (Figure 3.4 g and 3.4 h). HA nanocrystals grow in intimate contact within collagen fibers, building up a nanostructured composite. However, HA has disadvantage attributed to low mechanical strength for implant applications. Hence the combination of a load bearing biomaterial like titanium with the osteoconductive properties of HA is very attractive. It was found that the n-HA deposition was uniform and more predominant on the PLGA/Collagen nanofibers than on the PLGA nanofibers. The attachment of nano-HA was more on PLGA/Collagen compared to PLGA nanofibers because collagen is more hydrophilic and mimics the natural bone, thereby favouring nano-HA deposition. 49 Table 3.1 Optimization of electrospinning parameters by varying the time and concentration for PLGA nanofibers Electrospinning PLGA nanofibers at various concentrations (15%, 18% and 20% w/v) and time periods (5 seconds to 2 minutes) Nanofiber diameter ± SD (micro meter) Time 15% 18% 20% 2 min 0.68 ± 0.282 1.761 ± 0.371 1.800 ± 0.213 1.5 min 0.774 ± 0.227 1.212 ± 0.39 0.914 ± 0.151 1min 0.543 ± 0.153 1.114 ± 0.357 1.089 ± 0.267 30sec 0.768 ± 0.314 1.326 ± 0.479 1.067 ± 0.194 15 sec 0.957 ± 0.357 1.615 ± 0.472 1.731 ± 0.386 10sec 0.996 ± 0.344 1.721 ± 0.413 1.264 ± 0.269 5sec 0.759 ± 0.415 1.535 ± 0.594 1.381 ± 0.449 Table 3.2 Optimization of electrospinning parameters by varying the time and concentration for PLGA/Collagen nanofibers Electrospinning PLGA/Collagen nanofibers at various concentrations (10% and 15% w/v) and time periods (5 seconds to 2 minutes) Nanofiber diameter ± SD (micro meter) Time 10% 15% 2 min 0.549 ± 0.213 0.827 ± 0.116 1.5 min 0.279 ± 0.085 0.898 ± 0.176 1min 0.368 ± 0.089 0.801 ± 0.147 30sec 0.251 ± 0.093 0.776 ± 0.136 15 sec 0.378 ± 0.068 0.817 ± 0.151 10sec 0.410 ± 0.093 0.828 ± 0.185 50 5sec a) c) 0.310 ± 0.089 0.783 ± 0.454 b) d) 51 e) f) g) h) Figure 3.4 SEM images of a) untreated cpTi, b) cpTi after surface modification c) cpTi coated with PLGA nanofibers at 1000X magnification d) cpTi coated with PLGA/Collagen nanofibers at 1000X magnification e) cpTi coated with PLGA nanofibers at 5000X magnification f) cpTi coated with PLGA/Collagen nanofibers at 5000X magnification g) cpTi coated with functionalized PLGA nanofibers h) cpTi coated with functionalized PLGA/Collagen nanofibers 3.3.2 Surface Roughness analysis It is generally accepted that rough, textured and porous surfaces are able to stimulate cell attachment, differentiation and formation of the extracellular matrix [64]. Moreover, an 52 appropriate surface roughness can produce beneficial mechanical interlocking at the initial adhesion stage and aid in further cell adhesion [65]. Profilometer analysis revealed that the Ra value of the untreated cp Ti and Ti-6Al-4V discs were 0.306 μm and 1.529 μm respectively. However after the pretreatment procedure, the Ra values reduced to 0.022 μm and 0.042 μm respectively. This proves that the microtextured samples, after the pretreatment have attained a nanotextured surface. Thus the chemical pretreatment procedure using NaOH and HCl was a very feasible procedure to achieve nanotopography. Nanotopography plays a very important role in stem cell adhesion, proliferation and differentiation [63, 66]. Moreover, an appropriate surface roughness can produce beneficial mechanical interlocking at the initial adhesion stage and aid in further cell adhesion [64, 65]. Cell adhesion and proliferation were reported to be more on a nanotextured surface than on a microsurface. Hence it was essential that an implant surface has a nanotexture, in order to accelerate the osseointegration process in vivo. It was not possible to measure the surface roughness of the untreated Ti, using AFM, as it was highly rough. But the Ra results from AFM (Figure 3.5) for the treated samples complemented the profilometer results. The Ra values for the pretreated Ti and Ti alloy were calculated using the Nanoscope software and were found to be 15.2nm and 18.9 nm respectively. 53 Figure 3.5 AFM image of pretreated Ti showing the surface roughness 3.3.3 FTIR The FTIR results (Figure 3.6) showed an increase in the TiO2 oxide layer thickness in the pretreated samples. The TiO2 peak in the case of Ti was located at 667 cm-1. Though the untreated samples also showed the TiO2 peak, it was more significant in the case of the pretreated samples. The increase in the TiO2 oxide layer improves the biocompatibility of the implant [67]. Besides the new peaks in the 400-800 cm-1 wavenumber range, in the pretreated samples corresponds to the other oxides of Ti like TiO and Ti2O3. 54 In pure PLGA nanofibers the C=O stretch and the C–O stretch hovered around 1761 cm−1 and 1088 cm−1 respectively. Amide I and amide II of collagen were detected at 1658 cm−1 and 1544 cm−1 in PLGA/Collagen nanofibers. a) 55 b) c) Figure 3.6 FTIR results for a) pure Ti treated and untreated, b) Ti-6Al-4V alloy treated and untreated, c) PLGA and PLGA/Collagen nanofibers coated over the Ti surface. 56 3.3.4 Water contact angle measurement The contact angle results showed that the pretreatment procedure has improved the hydrophilicity of the scaffold. The contact angle of the control samples were 106.10 ±1.70 for cp Ti and 71.10 ± 5.40 for Ti alloy. The contact angle was reduced to 16.02 ± 0.80 and 11.04 ± 1.10 for cp Ti and Ti alloy respectively after the pretreatment as shown in Table 3.3. A decrease in the water contact angle indicates that the substrate has become more hydrophilic. An increase in the wettability of the scaffold was said to improve cell adhesion [66]. The functional groups present in collagen, i.e. carboxyl groups and carbonyl groups [68] and [69], served as nucleation sites for apatite formation and consequently, uniform distribution of n-HA was apparent on the outer and inner surfaces of the PLGA/Collagen nanofibers compared to the PLGA nanofibers. Besides being a favorable site for nucleation, the –COOH functional groups of collagen increased the hydrophilicity of the nanofibers. The contact angle measurements were taken after depositing the PLGA and PLGA/Collagen nanofibers over the scaffold by electrospinning. The PLGA scaffolds had a contact angle of 9.9 + 0.3° and the PLGA/Collagen scaffolds had a value of 0 as shown in Table 3.4. This was because collagen is very hydrophilic in nature. The hydroxyl groups present in the collagen forms hydrogen bonds with water molecules thus imparting the relevant hydrophilicity. Hence after incorporating collagen to PLGA, the water contact angle decreases. 57 Table 3.3 Water contact angle measurements for treated and untreated cp Ti and Ti6Al4V SAMPLE WATER CONTACT ANGLE(°) Before pretreatment Ti 106.10 ± 1.7 After pretreatment Ti 16.02 ± 0.8 Before pretreatment Ti alloy 71.10 ± 5.4 After pretreatment Ti alloy 11.04 ± 1.1 Table 3.4 Water contact angle measurements for PLGA and PLGA/Collagen nanofibers SAMPLE WATER CONTACT ANGLE (°) PLGA nanofibers 9.9 ± 0.3 PLGA/Collagen nanofibers 0 3.3.5 XPS Figure 3.7 shows distinct peaks obtained in the range 450 – 470eV in the treated Ti sample, corresponding to Ti 2p. We find that in the treated samples new peaks arise in this range corresponding to the Ti oxides like Ti2O3, TiO2 and TiO. Besides the Oxygen 1s peak also increased indicating an increase in the oxide layer formed over Ti. The XPS results proved that owing to the pretreatment of Ti, the oxide layers have increased, which in turn improves the biocompatibility of the implant surface [67]. Improving the oxide layer also favours enhanced initial osseointegration. The peaks for Na 1s and Cl 2p 58 increased in the pretreated samples owing to the NaOH/HCl pretreatment procedure followed. Figure 3.7 XPS results showing the Ti2p peaks in the treated samples 3.3.6 Cell culture analysis As depicted in the Figures 3.8A and 3.8B, the biomineralized nanofibers show enhanced cell adhesion when compared to the non- biomineralized nanofibers and the untreated titanium samples. In the untreated samples no significant adhesion of cells occurs even after 60min. This suggests that the untreated cp Ti and Ti alloy samples are not suitable for early cell adhesion. However at 10min, no statistical difference (p ≤ 0.05) was found between the Ti coated with PLGA/Collagen and the Ti coated with PLGA/HA. Also in the case of Ti alloy at 10min there was no significant difference between Ti alloy coated with PLGA and PLGA/HA. No statistical difference was also observed between Ti alloys coated with PLGA/Collagen and PLGA/Collagen/HA. This maybe because 10min 59 duration was too short for HA to cause a significant cell adhesion. At 30min and 60min the adhesion onto the Ti/PLGA/Collagen/HA and Ti alloy/PLGA/Collagen/HA substrate was statistically significant from all the other Ti and Ti alloy samples respectively, indicating that maximum adhesion of hMSCs occurred on the mineralized PLGA/Collagen scaffolds compared to the other scaffolds. Figure 3.8A and 3.8B shows one of the SEM images taken for the cp Ti and Ti alloy samples respectively at the time intervals – 10, 20, 30 and 60 min. Studies have reported lower cell adhesion and proliferation on less organized surfaces (i.e. sandblasted ones) [70]. Hence for this reason, in the present study, regular and uniform surface roughness on the surface of all samples was produced, resulting in homogeneous surface texture on all the cpTi and Ti alloy disks. This study thus proves that the nano-hydroxyapatite coated on the nanotextured titanium surface improves the initial cell attachment, which is very crucial for enhanced osseointegration. It was found that the cell adhesion was more on the biomineralized scaffolds compared to the nonbiomineralized scaffolds. This is because collagen along with n-HA synergistically enhances early cell capture. Besides, calcium ions have also been suggested to be advantageous to cell growth [71]. However the exact mechanisms by which calcium phosphate ceramics improve bone bonding are not clearly understood, although it is known that the bioactivity of ceramics is related to the dissolution rate and that the early cellular response is of primary importance [72]. Similarly adhesion study has also been reported earlier by Kun Ma et al., using bone marrow derived haematopoitic stem cells on PLGA and PLGA/Collagen nanofibers coated with a surface adhesion molecule, E selectin. The study revealed that the haematopoitic stem cells capture efficiency on the 60 PLGA/Collagen nanofiber scaffold after coated with E-selectin, significantly increased cell capture percentage from 23.40% to 67.41% within 30 min and from 29.44% to 70.19% within 60 min of incubation at room temperature [63]. Nevertheless our results have indicated nearly 75% cell adhesion on to the bioceramic coated surface on both cpTi and Ti alloy samples as shown in Figure 3.9. Table 3.5 indicates the statistics for the number of cells adhered to the cpTi and Ti alloy implant surfaces at various time intervals. The rationale for conducting a short-term cell adhesion study on the nanofibrous scaffolds was to assess the viability of avoiding extended culture periods of cell seeding on the substrates, thereby reducing the down-time from material preparation to the material implantation in the patient, preferably in-situ during surgery. 10 min 20 min 30 min 60 min a)Untreated cpTi b)cpTi + PLGA 61 c)cpTi+PLGA/Col d)cpTi+PLGA/HA e)cpTi+PLGA/Col/HA Figure 3.8A: Adhesion of hMSCs on the a) untreated cpTi implants, b) cpTi implant coated with PLGA nanofibers, c) cpTi implant coated with PLGA/Collagen nanofibers, d) cpTi implant coated with PLGA/HA, e) cpTi implant coated with PLGA/Collagen/HA nanofibers at 500x 10 min 20 min 30 min 60 min a) untreated Ti6Al4V 62 b) Ti6Al4V+PLGA c)Ti6Al4V + PLGA/Col d)Ti6Al4V + PLGA/HA e)Ti6Al4V+PLGA/Col/HA Figure 3.8B: Adhesion of hMSCs on the a) untreated Ti6Al4V implants, b) Ti6Al4V implant coated with PLGA nanofibers, c) Ti6Al4V implant coated with PLGA/Collagen nanofibers, d) Ti6Al4V implant coated with PLGA/HA, e) Ti6Al4V implant coated with PLGA/Collagen/HA nanofibers at 500x 63 Table 3.5: Average number of cells adhered to the Ti samples Sample 10 min 20 min 30 min 60 min cpTi 44 108 132 188 cpTi coated with PLGA nanofibers 683 1230 2188 3420 2482 3120 4601 biomineralized 746 1668 3842 5880 biomineralized 1080 3260 5460 7512 cpTi coated with PLGA/Collagen 1025 nanofibers cpTi coated with PLGA nanofibers cpTi coated with PLGA/Collagen nanofibers Ti alloy 32 48 85 105 Ti alloy coated with PLGA nanofibers 592 1185 2018 3220 2380 3175 4500 1775 3724 5800 3120 5516 7475 Ti alloy coated with PLGA/Collagen 1105 nanofibers Ti alloy coated with biomineralized 820 PLGA nanofibers Ti alloy coated with biomineralized 1185 PLGA/Collagen nanofibers 64 Ti P Ti P er s er s /H A LG A/ HA lag en ol Ti P ib fib no f na no na LG A/ C lag en LG A nt re at ed 100 ol LG A/ C Ti P Ti u MSC capture percentage (%) Ti P Ti P lag en ol er s er s /H A LG A/ HA lag en Ti P ib no f fib nt re at ed na no na LG A LG A/ C LG A/ Co l Ti P Ti u MSC capture percentage (%) 100 Adhesion study on pure Ti 80 60 40 10 min 20 20 min 0 30 min 40 60 min Adhesion study on Ti6Al4V 80 60 10 min 20 20 min 0 30 min 60 min Figure 3.9: Percentage attachment efficiency of hMSCs on cpTi and Ti6Al4V alloy 65 3.4 Conclusion Our work has proved the feasibility of creating a nanotextured surface on titanium by simple acid/alkali treatment. The surface roughness can be tailored by modifying the etching/ polishing procedures. Besides we have demonstrated that the cell adhesion can be increased by coating the titanium surface with nanofibers. This is because the nanofibers mimic the natural ECM and hence improve cell attachment. Through our electrospinning set up we were able to achieve precise fiber deposition at a shorter interval of time. We have increased the fiber deposition efficiency by our set up compared to the conventional electrospinning. Moreover we have shown that the adhesion efficiency of the hMSCs was the maximum on the cpTi and Ti alloy samples coated with biomineralized PLGA/Collagen nanofibers compared to the other samples, owing to the synergistic effect of collagen and n-HA. 66 Chapter 4 Mesenchymal stem cells proliferation and differentiation studies on the modified implant surfaces 4.1 Introduction When a biomaterial is implanted into the human body, it is unavoidable that blood will contact the implant surface. Upon contact, the implant surface could be covered with a layer of plasma proteins that mediate the next cellular responses. Therefore, the surface characteristics of an implant should not only enhance the osteogenic cell–material interactions but also optimize the initial blood–material interactions. The success of a bone implant depends on how early the osseointegration is achieved [73]. Hence the surface of the implants ought to be modified to improve early osseointegration. Albrektsson et al., proposed six factors as especially important for successful osseointegration. These include the implant material, implant design, surface conditions, and status of the bone, the surgical technique and the implant loading conditions [74]. Nanofibers have demonstrated excellent cell adhesion and differentiation. Ultimately, one of the main goals is to attract and induce the osteoprogenitor cells to differentiate into osteoblasts. We hypothesize that coating surfaces with nanofibers and the presence of biomolecules like n-HA would affect the proliferation and differentiation of osteoprogenitors to osteoblasts. Therefore, nanofibrous modification of dental and bone implants might enhance osseointegration. There have been various techniques tried out in the past to improve the surface roughness 67 of the implant like plasma treatment, acid-etching and heat treatment. It is our hypothesis that biomimetic bone like composite-coated metallic implants with loading capability from the metal core and having a bioactive surface like nanofibers with nano-HA will accelerate bone formation and implant fixation. 4.2 Material and methods 4.2.1 Mesenchymal stem cells culture All the titanium samples were sterilized under UV light for 2 hrs. The discs were then washed with phosphate buffered saline (PBS) thrice for 15min and were eventually incubated with DMEM overnight before cell seeding. Human Mesenchymal stem cells (PT-2501, Lonza, USA) was cultured in DMEM low glucose medium (DMEM, Invitrogen, CA, U.S.) with 10% FBS (Invitrogen, CA, U.S.) and 1X Antibiotics (SigmaAldrich Chemical Company Inc., St. Louis, U.S) until 3 passages. The cells were then trypsinized from the 75 cm2 cell culture flasks by adding 1 ml of 0.25% trypsin containing 0.1% EDTA, purchased from GIBCO Invitrogen, USA. Detached cells were centrifuged, counted by tryphan blue assay using a hemocytometer and seeded on the scaffolds at a cell density of 1.0 × 104 cells/well for 24 well plates (pure Ti based samples) and 2.0 × 104 cells/well for the 6 well plates (Ti alloy based samples) was added and left in incubator for facilitating cell growth. The well plates were incubated for 60 minutes at room temperature to favor cell adhesion as described in chapter 3. After the incubation time, the media was removed and the plates were washed thrice with PBS to remove the unbound cells. Fresh media was then added to the wells and the plates were then transferred to the incubator. The well plates were cultured for days 7, 14 and 21 to 68 carry out further cell culture analysis like proliferation, differentiation and mineralization. The pure Ti samples – untreated Ti, treated Ti coated with PLGA nanofibers, treated Ti coated with PLGA/Collagen nanofibers, treated Ti coated with PLGA nanofibers+nanoHA and treated Ti coated with PLGA/Collagen nanofibers+nano-HA, were cultured onto the 24 well plates. The Ti-6Al-4V samples – untreated Ti-6Al-4V, treated Ti-6Al-4V coated with PLGA nanofibers, treated Ti-6A-4V coated with PLGA/Collagen nanofibers, treated Ti-6Al-4V coated with PLGA nanofibers+nano-HA and treated Ti-6Al-4V coated with PLGA/Collagen nanofibers+nano-HA, were cultured onto 6 well plates. The cells were cultured and analyzed for their proliferation and differentiation on days 7, 14 and 21. The optical images of hMSCs cultured on Tissue Culture Plate (TCP) at 24, 48, 72 and 96 hrs were shown in Appendix A 4.2.2 Cell Morphology Study The cell morphology was analyzed using FESEM. After 6 days of seeding the hMSCs, the media was removed from the wells and the samples were fixed with 3% glutaraldehyde for 3 hours. The scaffolds were then rinsed with distilled water for 15min and then dehydrated with a series of ethanol gradients starting from 30% to 50%, 75%, 90% and 100% (v/v). Subsequently the samples were treated with HMDS (Hexamethyldisilazane) solution and allowed to air – dry at room temperature in the fume hood. The samples were then gold coated and the cells morphology was analyzed using FESEM. The mineral secreted by the cells was analyzed using FESEM equipped with 69 EDX. The same procedure was repeated for day 14 and day 21. The results are shown in Appendix B. 4.2.3 Cell Proliferation Study The cell proliferation on different scaffolds was analyzed using MTS assay (CellTiter 96 AQueous One solution reagent, purchased from Promega, Madison, WI). The principle behind the MTS assay involves the reduction of yellow tetrazolium salt [3-(4, 5- dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2(4-sulfophenyl)-2H-tetrazolium] in MTS to form purple formazan crystals by the dehydrogenase enzymes secreted by mitochondria of metabolically active cells forms the basis of this assay. The formazan dye shows absorbance at 492 nm and the amount of formazan crystals formed is directly proportional to the number of cells. After 6 days of seeding, the media was removed from the well plates and the scaffolds were washed in PBS. The scaffolds were then incubated in a 1:5 ratio mixture of MTS assay and serum free DMEM medium for 3 – 5 hrs at 37°C in a 5% CO2 incubator. After the incubation period, the samples were pipetted out into 96 well plates. The absorbance was then calibrated at 490 nm using a spectrophotometric plate reader (Fluostar Optima, BMG Lab Technologies, Germany). The same procedure was repeated for day 14 and day 21 samples. 70 4.2.4 Alkaline phosphatase activity The osteogenic differentiation of hMSCs was analyzed by measuring the alkaline phosphatase (ALP) activity. ALP activity was measured using Alkaline Phosphate Yellow Liquid substrate system for ELISA (Sigma Life Sciences, USA). In this reaction, ALP catalyzes the hydrolysis of colorless organic phosphate ester substrate, pnitrophenylphosphate (pNPP) to a yellow product, p-nitrophenol, and phosphate. After 6 days of seeding the scaffolds with the hMSCs were washed thrice with PBS. 400 μl of pNPP liquid was added to the scaffolds and incubated for 30 min till the colour of solution becomes yellow. The reaction was then arrested by the addition of 100 μl of 2 N NaOH solution following which the yellow color product was aliquoted in 96-well plate and read in spectrophotometric plate reader at 405 nm. 4.2.5 Cell mineralization Study The amount of minerals secreted by the hMSCs can be both qualitatively and quantitatively analysed by using Alizarin red staining. Alizarin Red-S (ARS) is a dye that binds selectively to the calcium salts and hence can be used for mineral staining. The Ti and Ti alloy scaffolds with hMSC cells was washed thrice with PBS and fixed in ice-cold 70% ethanol for 1 h. These constructs were then washed twice with distilled water and stained with ARS (40 mM) for 20 min at room temperature. After several washes with distilled water, the scaffolds were observed under inverted optical microscope and images were taken using image software (Leica FW4000, version v 1.0.2). The stain was eluted by incubating the scaffold with 10% cetylpyridinium chloride for 1 h. The absorbance of 71 the collected dye was then read at 540 nm in spectrophotometer (Thermo Spectronics, Waltham, MA, USA). 4.2.6 Statistical analysis Values (at least triplicate) were averaged and expressed as means ± standard deviation (SD). Statistical differences were determined by Student two-sample t test. Differences were considered statistically significant at p ≤ 0.05. 4.3 Results and Discussion 4.3.1 Cell Morphology Study The hMSC morphology was analyzed on the day 7, 14 and 21 using FESEM. The Figures 4.1 – 4.3 show the SEM micrographs of cell interaction with the nanofibers as well as the Ti scaffold. Since the duration of electrospinning was short, only a thin layer of the electrospun fibers have been deposited onto the Ti plates. Hence the cells begin to migrate further beyond the fibers and interact with the Ti discs. Nanotopography favors cell adhesion, proliferation and differentiation [1, 4, 63, 66, 86]. Since both cp Ti and the Ti6Al4V alloy have a nanotextured surface it is believed to enhance the cell – scaffold interactions. From the cell morphology as shown in Figure 4.1, it is seen that by day 7 the hMSCs cultured on the treated Ti coated with functionalized PLGA/Collagen nanofibers with nano-HA have extended their filopodia and contacted the adjoining cells and proliferated. The morphology of the cells remains rounded in the case of the untreated 72 scaffolds, indicating that the surface is not suitable for cell culture. Since the duration of electrospinning was short, only a thin layer of the electrospun fibers have been deposited onto the Ti plates. Hence the cells begin to migrate further beyond the fibers and interact with the Ti discs. The nanotopography of a scaffold surface favors cell adhesion, proliferation and differentiation. The cell spreading, with spindle-like and polygonal like cell shapes, was also observed on HA-based composites on days 14 and 21 (Figure 4.2 and 4.3) of culture and physical contact between cells were maintained via filopodia or lamellipodia [75]. By virtue of these observations, n-HA or n-HA in combination with collagen would result in greater cell motility due to better-developed filopodia and lamellipodia, as reported earlier [76]. Vanessa et al. [77] concluded that the treatment of titanium and titanium alloy implant surfaces with discrete crystalline deposits like HA renders them bone bonding, and it is the increase in complexity of the resultant surface which is the driving force for the bonding mechanism at the bone – implant interface. Theoretically, the osteoconductive properties of HA would provide reproducible attachment of implants to the skeleton by osseointegration and bone ingrowth. The morphology of the cells cultured on cp Ti and Ti-6Al-4V alloy was similar as the scaffolds were subjected to the same surface treatment procedures. 73 a) c) b) d) e) Figure 4.1 SEM images of the hMSC morphology on day 7 on a) untreated Ti-6Al-4V, b) Treated Ti-6Al-4V coated with PLGA nanofibers, c) Treated Ti-6Al-4V coated with PLGA/Collagen nanofibers, d) Treated Ti-6Al-4V coated with functionalized PLGA nanofibers, e) Treated Ti6Al-4V coated with functionalized PLGA/Collagen nanofibers. 74 a) b) c) d) e) Figure 4.2 SEM images of the hMSC morphology on day 14 on a) untreated Ti-6Al-4V, b) Treated Ti-6Al-4V coated with PLGA nanofibers, c) Treated Ti-6Al-4V coated with PLGA/Collagen nanofibers, d) Treated Ti-6Al-4V coated with functionalized PLGA nanofibers, e) Treated Ti-6Al-4V coated with functionalized PLGA/Collagen nanofibers. 75 a) b a) ) c) d) e) Figure 4.3 SEM images of the hMSC morphology on day 21 on a) untreated Ti -6Al-4V, b) Treated Ti-6Al-4V coated with PLGA nanofibers, c) Treated Ti-6Al-4V coated with PLGA/Collagen nanofibers, d) Treated Ti-6Al-4V coated with functionalized PLGA nanofibers, e) Treated Ti-6Al-4V coated with functionalized PLGA/Collagen nanofibers. 76 4.3.2 Cell Proliferation Study The MTS assay was used to study the cell proliferation of hMSCs on cpTi and Ti alloy surfaces as shown in Figures 4.4a and 4.4b respectively. It was seen that even though the cell proliferation rate initially was high in the TCP (Tissue Culture Plate), by day 21 the cell proliferation was maximum in the functionalized nanofiber coated Ti. By day 21 the cells seeded onto the biomineralized PLGA/Collagen scaffolds had proliferated by 257% compared to day 7. In the case of cpTi samples on day 7, significant difference was observed between the mineralized PLGA/Collagen nanofibers and the untreated Ti samples. The samples coated with PLGA nanofibers were statistically different from the sample coated with PLGA/Collagen nanofibers and the mineralized scaffolds; indicating that PLGA alone was not sufficient to significantly enhance the cell proliferation rate. Additionally, for the HA coated PLGA/Collagen nanofibers, the cell proliferation was higher compared to the HA coated PLGA nanofibers. This was because of the presence of collagen, which is a principle component of the ECM. The rate of cell proliferation was rather slow in the untreated Ti samples. This was because the samples have a low cell capture ratio as seen in the cell adhesion study in chapter 3. The proliferation in the case of Ti coated with the PLGA and PLGA/Collagen nanofibers was also high as the nanofibers mimic the ECM and thereby enhance the cell proliferation rate. However owing to the presence of collagen the proliferation rate was higher in the PLGA/Collagen nanofibers compared to the PLGA nanofibers. Thus it was seen that functionalization of the nanofibers enhance the cell proliferation compared to the non-functionalized scaffolds. As shown in the Figures 4.4a and 4.4b, the results for both the cp Ti and Ti6Al4V alloy were similar. This maybe because the surface treatment and coating on 77 both the substrates were similar, indicating that both cpTi and Ti6Al4V alloy are imparting similar mechanical and biological cues. a) * * b) * Figure 4.4 MTS assay for hMSC cells proliferation on a) cpTi based scaffolds – untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers b) Ti-6Al-4V based scaffolds - untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers for day 7, 14 and 21. * represents p≤ 0.05 statistical difference. Control refers to the Tissue Culture Plate (TCP); TiK refers to Ti6Al4V alloy. 78 4.3.3 Alkaline phosphatase activity Alkaline phosphatase is a membrane bound enzyme and its activity is used as an osteoblastic differentiation marker [78], as it is produced only by cells showing mineralized ECM [79]. The ALP activity indicates the osteogenic differentiation capacity of the cells. The ALP activity for the cpTi and Ti alloy samples are depicted in Figures 4.5a and 4.5b. It was seen that even though initially at day 7 the ALP activity was similar on all the scaffolds, the activity started to increase from day 14. There was not much increase in the ALP activity for the untreated scaffolds, indicating that the cells cultured on the untreated surfaces have not undergone osteogenic differentiation. This maybe because the untreated scaffold have no biological and mechanical cues capable of inducing the osteogenic differentiation of hMSCs. However TCP showed more ALP activity compared to the untreated Ti surfaces, indicating that the untreated surfaces were not suitable for cell differentiation compared to TCP. Comparison of the nonfunctionalized scaffolds show the ALP activity was higher for Ti coated with PLGA/Collagen compared to the scaffolds coated with PLGA nanofibers (p ≤ 0.05). There was also statistically significant difference between the PLGA/HA scaffolds and the PLGA/Collagen/HA scaffolds indicating an enhanced ALP activity in the presence of both collagen and HA. By day 21, significant difference was observed between the mineralized scaffolds and the untreated and the non – mineralized scaffolds. As explained earlier this increase in the ALP activity upon the incorporation of collagen was due to its presence. Type I Collagen being the principle component of the organic part of the bone matrix, induces the hMSCs to differentiate into bone cells. Thus the functionalized nanofibers show the ability to induce osteogenic differentiation of mesenchymal stem 79 cells. However the hMSC undergoing osteogenic differentiation was more in the biomineralized PLGA/Collagen fibers rather than the biomineralized PLGA nanofibers. This difference is due to the presence of collagen in addition to HA which synergistically enhances the osteogenic differentiation capacity of the hMSCs. However in the case of cpTi, no significant difference was observed between the mineralized scaffolds and the non – mineralized scaffolds on the days 7 and 14. This maybe because cpTi surface, unlike the Ti alloy, is significantly slow in inducing the hMSCs differentiation. By day 21 however significant difference was observed between the mineralized and non – mineralized scaffolds, indicating that collagen and HA being the native constituent of the bone, synergistically induces hMSC differentiation. Ohgushi et al. [80] suggested that mesenchymal cells could be influenced to differentiate into osteoblasts in the presence of bioceramics. Our study has shown that the presence of bioceramics like HA has triggered the differentiation of hMSC into osteoblasts as proved by the enhanced ALP activity in the mineralized implants. As shown in the Figure 4.5a and 4.5b, the results for both cpTi and Ti-6Al-4V alloy were similar. This similarity is due to the surface treatment employed and the nanofibrous coating. Similar results have been found by others [81 84], who concluded that both rough Ti and Ti-6Al-4V surfaces enhance alkaline phosphatase activity and mineralization. But none of the above studies involved coating the rough Ti and Ti alloy discs with a mineralized nanofibrous coating. It has been shown in vivo, that there are many cells which are capable of differentiating into osteoblastic cells and hence contributing to the production of extracellular matrix, and that differentiation is encouraged most by the HA coating on the titanium [85]. 80 a) * * b) * * Figure 4.5 ALP activity for hMSC cells on a) cpTi based scaffolds - untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers b) Ti-6Al-4V based scaffolds - untreated, coated with PLGA nanofibers, coated with PLGA/Collagen nanofibers, coated with PLGA/HA and coated with PLGA/Collagen/HA nanofibers for day 7, 14 and 21. * represents p≤ 0.05 statistical difference. Control refers to the Tissue Culture Plate (TCP); TiK refers to Ti6Al4V alloy. 81 4.3.4 Cell mineralization study Mineralization refers to the cell-mediated deposition of extracellular calcium and phosphorus salts where anionic matrix molecules take up the calcium ions and the phosphate ions and serve as nucleation and growth sites leading to calcification [86]. The cell mineralization on the cpTi and Ti alloy samples have been represented quantitatively in Figures 4.6a and 4.6b respectively, and qualitatively in Figures 4.7(A-C) and 4.8(A-C) respectively. From Figures 4.6a and 4.6b, it was seen that the cells cultured on the functionalized nanofiber scaffolds start secreting their minerals by day 7. In the case of Ti alloy, even as early as day 7 significant difference (p ≤ 0.05) was observed between the mineralized scaffolds and the non – mineralized scaffolds. This suggests that the HA deposited on the fibers stimulates the mineralization of the cells. HA serves as biological cues for stimulating the osteogenic differentiation and mineralization of hMSCs. The untreated cpTi and Ti alloy samples (Figure 4.7A (a) and 4.8A (a)) showed similar limited mineralization profile. Since it has not taken up the Alizarin red stain, it appears green under the optical microscope. They did not favor cell mineralization owing to the absence of any biological cues. However, the Ti coated with PLGA/Collagen favored more cell mineralization than Ti coated with PLGA nanofibers. The presence of collagen stimulates the secretion of minerals by the hMSCs. The cell mineralization was the maximum for the functionalized PLGA/Collagen nanofibers. The presence of collagen along with HA induces a synergic effect for the mineral secretion. In accordance to the cell proliferation and the differentiation studies, the results were similar for both cp Ti and Ti6Al4V alloy. Harris et al., demonstrated an increase in the extracellular matrix production by osteoblastic cells when cultured on HA coatings [87]. Rough surface may 82 allow the osteoblastic cells to obtain more points of adhesion, as described by Niederauer et al., [88], and to produce more extracellular matrix [89]. Although similar results have not been reported with regard to hMSC, Tenenbaum et al., [90] have shown that, given the right environment, many cells are capable of behaving in an osteoblast-like way and it may be that the correct microenvironment is provided by the HA and collagen present in the nanofiber surface coating. From the qualitative representation of the ARS staining as shown in Figures 4.7(A-C) and 4.8(A-C), it was seen that the ARS staining was more preponderant on the functionalized PLGA/Collagen nanofibers coated scaffolds compared to the other scaffolds. This was because the cell mineralization was more on the functionalized nanofibers and hence they take up more alizarin red stain giving a bright red appearance. The untreated Ti scaffolds had not taken up any stain due to the absence of the cell mineralization. Hence it gives a green appearance on the optical microscope. Moreover, the ARS staining was increased by day 21 compared to the day 7 and day 14 on cpTi and Ti alloy samples coated with nanofibers indicating that the cell mineralization has increased. More alizarin red staining uptake can be noticed on the samples coated with mineralized PLGA/Collagen nanofibers on the cpTi and Ti alloy samples as indicated in Figure 4.7C and 4.8C compared to the day 7 (Figure 4.7A and 4.8A) and day 14 (Figure 4.7B and 4.8B). The mineralization was further analyzed using FESEM EDX (Supplementary data). The Ca/P ratio of mineral was 1.3 on day 7 on the functionalized PLGA/Collagen nanofibers with nano-HA. This ratio increased further as the cell mineralization increases. It was seen that the Ca/P ratio on day 14 and day 21 was 1.5 and 2.03 respectively. The EDX data were attached in the Appendix B. 83 a) * * b) * * Figure 4.6 Quantitative data for Alizarin red staining on hMSC cells on a) cp Ti scaffolds b) Ti6Al-4V scaffolds for days 7, 14 and 21. * represents p≤ 0.05 statistical difference 84 a) b) c) d) e) Figure 4.7A: Optical image of the ARS stained hMSCs on the cpTi scaffolds on day 7 a) untreated cpTi, b) Treated Ti coated with PLGA nanofibers, c) Treated cpTi coated with PLGA/Collagen nanofibers, d) Treated cpTi coated with functionalized PLGA nanofibers, e) Treated cpTi coated with functionalized PLGA/Collagen nanofibers. 85 a) b) c) d) e) Figure 4.7B Optical image of the ARS stained hMSCs on the cpTi scaffolds on day 14 a) untreated cpTi, b) Treated cpTi coated with PLGA nanofibers, c) Treated cpTi coated with PLGA/Collagen nanofibers, d) Treated cpTi coated with functionalized PLGA nanofibers, e) Treated cpTi coated with functionalized PLGA/Collagen nanofibers. 86 a) c) b) d) e) Figure 4.7C Optical image of the ARS stained hMSCs on the cpTi scaffolds on day 21 a) untreated cpTi, b) Treated cpTi coated with PLGA nanofibers, c) Treated cpTi coated with PLGA/Collagen nanofibers, d) Treated cpTi coated with functionalized PLGA nanofibers, e) Treated cpTi coated with functionalized PLGA/Collagen nanofibers. 87 a) b) c) d) e) Figure 4.8A Optical image of the ARS stained hMSCs on the Ti-6Al-4V scaffolds on day 7 a) untreated Ti-6Al-4V, b) Treated Ti-6Al-4V coated with PLGA nanofibers, c) Treated Ti-6Al-4V coated with PLGA/Collagen nanofibers, d) Treated Ti-6Al-4V coated with functionalized PLGA nanofibers, e) Treated Ti-6Al-4V coated with functionalized PLGA/Collagen nanofibers. 88 a) c) b) d) e) Figure 4.8B Optical image of the ARS stained hMSCs on the Ti-6Al-4V scaffolds on day 14 a) untreated Ti-6Al-4V, b) Treated Ti-6Al-4V coated with PLGA nanofibers, c) Treated Ti-6Al-4V coated with PLGA/Collagen nanofibers, d) Treated Ti-6Al-4V coated with functionalized PLGA nanofibers, e) Treated Ti-6Al-4V coated with functionalized PLGA/Collagen nanofibers. 89 a) b) c) d) e) Figure 4.8C Optical image of the ARS stained hMSCs on the Ti-6Al-4V scaffolds on day 21 a) untreated Ti-6Al-4V, b) Treated Ti-6Al-4V coated with PLGA nanofibers, c) Treated Ti-6Al-4V coated with PLGA/Collagen nanofibers, d) Treated Ti-6Al-4V coated with functionalized PLGA nanofibers, e) Treated Ti-6Al-4V coated with functionalized PLGA/Collagen nanofibers. 90 4.4 Conclusion The studies reported so far [64, 91, 92] have used various cell culture models representing osteoblasts and various stages in their lineage. The present study used human mesenchymal stem cells, which are capable of differentiating into osteoprogenitor cells and osteoblasts. However the increase in proliferation rate in the biomineralized samples, which also showed an enhanced mineralization, is in contrary to the theory that proliferation is down-regulated when extracellular matrix maturation is induced and mineralization occurs [93]. This absence in down-regulation of the hMSCs cell number on the biomineralized scaffolds which also showed enhanced mineralization maybe because of the presence of surface cues like HA present on the implants which triggers proliferation as well as mineralization simultaneously, without inhibiting one of them. Similar results were obtained by Deepika et al., [66] who cultured osteoblasts on the HA sprayed and HA blended PCL/Gelatin nanofibers; both proliferation and mineralization increased continuously and was maximum on the HA sprayed nanofibrous scaffold. Cell behaviour such as adhesion, spreading and proliferation represent the initial phase of cell–scaffold interaction that subsequently effect differentiation and mineralization [86]. Animal studies done in the past have suggested that HA stimulates bone to bridge gaps, induces fibrous connective tissue (FCT) metaplasia to bone, and increases bone mineralization when compared to uncoated implants of equal size, material, and structure [94]. This is in correlation to the increased mineralization observed on the HA coated Ti samples compared to the untreated Ti samples, in our study. Although in the earlier studies the deposition of HA was by plasma spray technique, the process has disadvantages attributed to the high temperatures used during the process, such as the 91 possibility of fracture at the interface between the titanium and the HA due to the residual stress at the interface, and changes in the composition, porosity, crystallinity, and structure of the plasma sprayed HA [87]. Our study demonstrated the deposition of HA on to the nanofibers by a feasible Ca–P dipping method, thereby overcoming the disadvantages of the plasma-spraying technique, in addition to increasing the bonebonding ability. The difference in the bone reaction between HA coated and uncoated implants, as reported by Alzubaydi et al., [95] not only suggests a high osteoconductive potential of the coated HA material but also its osteoinductivity, which is very much essential for early osseointegration. However most techniques used to deposit inorganic Ca-P coatings involve either extremely high temperatures or other non-physiological conditions that impede the incorporation of biomolecules such as collagen [96- 99]. Hence our method by which collagen is electrospun along with PLGA followed by the dipping method to deposit Ca-P is advantageous. This in-situ method of producing n-HA on polymeric nanofiberous scaffolds coated on nanotextured implant surface, may be a probable option for future implant materials. 92 Chapter 5 Conclusions and Recommendations 5.1 Conclusions This work has proved the feasibility of creating a nanotextured surface on titanium by simple acid/alkali treatment. The surface roughness can be tailored by modifying the etching/ polishing procedures. Another strategy to enhance osteoconduction is the use of a nanofiber-coated surface. Besides we have demonstrated that the cell adhesion can be increased by coating the titanium surface with nanofibers. This is because the nanofibers mimic the natural ECM and hence improve cell attachment. The cell attachment can be increased further by depositing n-HA onto the nanofibers. The n-HA biomolecule improves the adhesion efficiency of the hMSCs compared to the non-biomineralized nanofibers. Through our electrospinning set up we were able to achieve fiber deposition at a shorter interval of time. We have increased the fiber deposition efficiency by our experimental set up compared to the conventional electrospinning. A combination of structural, mechanical and biological properties of an implant material play a critical role in cell seeding, proliferation and new tissue formation in orthopedic research. Nano-biomaterials should promote cell adhesion and be optimized for ECM production, mineralization and subsequent tissue regeneration. PLGA/Collagen/HA nanofibers coated implant surfaces fabricated by a modified advanced electrospinning technique, and hMSCs grown on them showed higher cell proliferation, and increased 93 ALP activity and mineralization, compared to the PLGA, PLGA/HA and PLGA/Collagen nanofiber coated implant surfaces. Hence, electrospun biomimetic PLGA/Collagen/HA nanofibers coated Ti surfaces hold great potential for adhesion, proliferation, differentiation and mineralization of hMSCs.Our results suggested that the nanotextured oxidised titanium surfaces, both cpTi and Ti alloy, coated with biomineralized PLGA/Collagen nanofibers enhanced the initial cell capture ratio. Incorporating biomolecular cue like collagen and n-HA have enhanced the cell proliferation, osteogenic differentiation and cell mineralization. Thus the healing time can be reduced, leading to enhanced initial osseointegration. To our knowledge, functionalized nanofiber treated dental implant is a novel idea for enhanced osseointegration using bone regeneration concept. The complete bone integration between dental implant and host bone will be enhanced by following three biomimetic aspects: 1. Natural ECM like nanofiber coated on the dental implant; 2. The biomineralization treatment; 3. Controllable MSCs incorporation with dental implant. 5.2 Recommendations From the above results it can be seen that osseointegration of the implant can be improved by coating the implant surface with biomineralized PLGA/Collagen nanofibers. This is very useful to reduce the duration required for osseointegration as the biomineralized scaffolds have shown tremendous promise for early cell capture. This enhances the bone bonding ability of the implant in vivo, resulting in early 94 osseointegration. For sure, animal study in a rabbit model will prove this concept ultimately. On the other hand, growth factors like bone morphogenic proteins (BMPs), especially BMP-7 can be incorporated into the nanofibers and the increase in the early osseointegration will be possible. 95 REFERENCES [1] Guehennec Le, Soueidan A, Layrolle P, Amouriq Y. Surface treatments of titanium dental implants for rapid osseointegration. Dental Materials. (2007); 23(7): 844-854 [2] Jimbo R, Sawase T, Shibata Y, Hirata K, Hishikawa Y, Tanaka Y, Bessho K, Ikeda T and Atsuta M. Enhanced osseointegration by the chemotactic activity of plasma fibronectin for cellular fibronectin positive cells. Biomaterials (2007); 28 (24): 34693477 [3] Ma Z.W, Kotaki M, Inai R. and Ramakrishna S. Potential of nanofiber matrix as tissue-engineering scaffolds. Tissue Engineering (2005); 11(1-2): 101-109. [4] Stevens M.M, and George J.H. Exploring and engineering the cell surface interface. Science, (2005); 310(5751): 1135-1138. [5] Palmer R, Smith B, Howe L, et al. Implants in Clinical Dentistry. London, UK: Martin Dunitz Ltd; (2002); 23: 273. [6] Cochran D, Oates T, Morton D, Jones A, Buser D, Peters F, et al. Clinical field trial examining an implant with a sand-blasted, acid-etched surface. J Periodontol (2007); 78(6): 974-982. [7] Bagno A and Di Bello C, Surface treatments and roughness properties of Ti -based biomaterials, J Mater Sci Mater Med (2004); 15: 935–949. [8] Mabboux F, Ponsonnet L, Morrier J.J, Jaffrezic N and Barsotti O, Surface free energy and bacterial retention to saliva-coated dental implant materials—an in vitro study, Colloids Surf B Biointerfaces (2004); 39: 199–205 96 [9] Giavaresi G, Fini M, Cigada A, Chiesa R, Rondelli G and Rimondini L et al., Mechanical and histomorphometric evaluations of titanium implants with different surface treatments inserted in sheep cortical bone, Biomaterials (2003); 24: 1583–1594. [10] Buser D, Broggini N, Wieland M, Schenk R.K, Denzer A.J and Cochran D.L et al., Enhanced bone apposition to a chemically modified SLA titanium surface, J Dent Res (2004); 83: 529–533 [11] Ferraz M.P, Monteiro F.J, Serro A.P, Saramago B, Gibson I.R and Santos J.D, Effect of chemical composition on hydrophobicity and zeta potential of plasma sprayed HA/CaO-P2O5 glass coatings, Biomaterials (2001); 22: 3105–3112. [12] Takemoto M, Fujibayashi S, Neo M, Suzuki J, Matsushita T, Kokubo T and Nakamura T. Osteoinductive porous titanium implants: Effect of sodium removal by dilute HCl treatment. Biomaterials (2006); 27 (13): 2682-2691 [13] Timothy D. Sargeant, Mustafa O. Guler, Scott M. Oppenheimer, Alvaro Mata, Robert L. Satcher, David C. Dunand, Samuel I. Stupp. Hybrid bone implants: Selfassembly of peptide amphiphile nanofibers within porous titanium. Biomaterials (2008); 29 (2): 161-171 [14] Wei M, Kim H.M, Kokubo T, Evans J.H. Effects of titanium surface modifications on bonding behavior of hydroxyapatite ceramics and titanium by hydrothermal hotpressing. Mater. Sci. Eng., C Bio S. (2002); 20: 125. [15] Kim H.M, Kokubo T, Fujibayashi S, Nishiguchi S, Nakamura T. Effect of nanotitanium hydride on formation of multi-nanoporous TiO2 film on Ti. J. Biomed. Mater. Res (2000); 52: 553. 97 [16] Lewandowska M, Roguska A, Pisarek M, Polak B, Janik-Czachor M, Kurzydłowski K.J. Morphology and chemical characterization of Ti surfaces modified for biomedical applications. Biomolecular Engineering (2007); 24 (5): 438-442 [17] Tsaryk R, Kalbacova M, Hempel U, Scharnweber D, Unger R.E, Dieter P, Kirkpatrick C.J, Peters K. Response of human endothelial cells to oxidative stress on Ti6Al4V alloy. Biomaterials (2007); 28(5): 806-813 [18] Nishiguchi S, Nakamura T, Kobayashi M, Kim H.M, Miyaji F and Kokubo T, The effect of heat treatment on bone-bonding ability of alkali-treated titanium, Biomaterials (1999); 20: 491–500. [19] Secchi A.G, Grigoriou V, Shapiro I.M, Cavalcanti-Adam E.A, Composto R.J, Ducheyne P, and Adams C.S. RGDS peptides immobilized on titanium alloy stimulate bone cell attachment, differentiation and confer resistance to apoptosis. J Biomed Mater Res A (2007); 83: 577. [20] Bornstein M.M, Valderrama P, Jones A.A, Wilson T.G, Seibl R, Cochran D.L, et al. Bone apposition around two different sandblasted and acid-etched titanium implant surfaces: a histomorphometric study in canine mandibles. Clin Oral Implants Res (2008); 19(3): 233-241. [21] Manara S, Paolucci F, Palazzo B, Marcaccio M, Foresti E, Tosi G, Sabbatini S , Sabatino P, Altankov G, Roveri N. Electrochemically-assisted deposition of biomimetic hydroxyapatite–collagen coatings on titanium plate. Inorganica Chimica Acta (2007); 361: 1634-1645. 98 [22] Wang H, Eliaz N, Xiang Z, Hsu H.P, Spector M and Hobbs L.W, Early bone apposition in vivo on plasma-sprayed and electrochemically deposited hydroxyapatite coatings on titanium alloy, Biomaterials 27 (2006): 4192–4203. [23] Lewandowska M, Roguska A, Pisarek M, Polak B, Janik-Czachor M, Kurzydłowski K.J. Morphology and chemical characterization of Ti surfaces modified for biomedical applications Biomolecular Engineering (2007); 24 (5): 438-442. [24] Nicula R, Lüthen F, Stir M, Nebe B, Burkel E. Spark plasma sintering synthesis of porous nanocrystalline titanium alloys for biomedical applications. Biomolecular Engineering (2007); 24 (5): 564-567 [25] Lincks J, Boyan B.D, Blanchard C.R, et al. Response of MG63 osteoblast-like cells to titanium and titanium alloy is dependent on surface roughness and composition. Biomaterials (1998); 19(23): 2219 - 32. [26] Ong J.L, Carnes D.L, Cardenas H.L, Cavin R. Surface roughness of titanium on bone morphogenetic protein-2 treated osteoblast cells in vitro. Implant Dent (1997); 6(1): 19-24. [27] Hoet P.H.M, Bruske-Hohlfeld I, Salata O.V. Nanoparticle Interaction with Biological Membranes J. Nanobiotechnol. (2004); 2: 2. [28] Webster T.J, Ergun C, Doremus R.H, Siegel R.W, Bizios R. Specific proteins mediate enhanced osteoblast adhesion on nanophase ceramics. J Biomed Mater Res (2000); 51(3): 475–83. 99 [29] SLActive - the third generation in implant surface technology [Internet 01 Oct 2008]. Availablefrom:http://www.straumann.com/com_index/ci_media_relations/ci_pr_presskit _bl_launch.htm/ci_presskit_backgrounder_slactive.pdf [30] Ngiam M, Liao S, Avinash Patil J, Cheng Z, Chan C.K, Ramakrishna S. The fabrication of nano-hydroxyapatite on PLGA and PLGA/Collagen nanofibrous composite scaffolds and their effects in osteoblastic behavior for bone tissue engineering. Bone (2009); 45(1): 4 – 16. [31] Yoshimoto H, Shin Y.M, Terai H. and Vacanti JP. A biodegradable nanofiber scaffold by electrospinning and its potential for bone tissue engineering. Biomaterials (2003); 24(12): 2077-2082. [32] Li W.J, Cooper J.A, Mauck R.L. and Tuan R.S. Fabrication and characterization of six electrospun poly (alpha-hydroxy ester)-based fibrous scaffolds for tissue engineering applications. Acta Biomaterialia, (2006); 2(4): 377-385. [33] Li W.J, Laurencin C.T, Caterson E.J, Tuan R.S. and Ko F.K. Electrospun nanofibrous structure: A novel scaffold for tissue engineering. Journal of Biomedical Materials Research, (2002); 60(4): 613-621. [34] Matthews J.A, Wnek G.E, Simpson D.G. and Bowlin G. L. Electrospinning of collagen nanofibers. Biomacromolecules (2002); 3(2): 232-238. 100 [35] Zhang Y.Z, Ouyang H.W, Lim C.T, Ramakrishna S. and Huang Z.M. Electrospinning of gelatin fibers and gelatin/PCL composite fibrous scaffolds. Journal of Biomedical Materials Research Part B-Applied Biomaterials (2005); 72B (1): 156-165. [36] Rho K.S. et al., Electrospinning of collagen nanofibers: effects on the behavior of normal human keratinocytes and early-stage wound healing. Biomaterials (2006); 27: 1452-1461. [37] Fertala A, Han W.B, and Ko F.K. Mapping critical sites in collagen II for rational design of gene-engineered proteins for cell-supporting materials. J. Biomed. Mater. Res., (2001); 57: 48-58. [38] Xu C.Y. et al., Aligned biodegradable nanofibrous structure: a potential scaffold for blood vessel engineering. Biomaterials (2004); 25: 877-886. [39] Zong X. et al., Electrospun fine-textured scaffolds for heart tissue constructs. Biomaterials (2005); 26: 5330-5338. [40] Bini T.B, Gao S.J, Tan T.C, Wang S, Lim A, Lim B.H. and Ramakrishna S. Electrospun poly(L-lactide-co-glycolide) biodegradable polymer nanofibre tubes for peripheral nerve regeneration. Nanotechnology (2004); 15(11): 1459-1464. [41] Schlosshauer B, Dreesmann L, Schaller H.E. and Sinis N. Synthetic nerve guide implants in humans: A comprehensive survey. Neurosurgery (2006); 59(4): 740-747. [42] Evans G.R.D, Brandt K, Niederbichler A.D, Chauvin P, Hermann S, Bogle M, Otta L, Wang B. and Patrick C.W. Clinical long-term in vivo evaluation of poly(L-lactic acid) 101 porous conduits for peripheral nerve regeneration. Journal of Biomaterials SciencePolymer Edition (2000); 11(8): 869-878. [43] Evans G.R.D, Brandt K, Widmer M.S, Lu L, Meszlenyi R.K, Gupta P.K, Mikos A.G, Hodges J, Williams J, Gurlek A, Nabawi A, Lohman R. and Patrick C.W. In vivo evaluation of poly (L-lactic acid) porous conduits for peripheral nerve regeneration. Biomaterials (1999); 20(12): 1109-1115. [44] Widmer M.S, Gupta P.K, Lu L.C, Meszlenyi R.K, Evans G.R.D, Brandt K, Savel T, Gurlek A, Patrick C.W. and Mikos A.G, Manufacture of porous biodegradable polymer conduits by an extrusion process for guided tissue regeneration. Biomaterials (1998); 19(21): 1945-1955. [45] Den Dunnen W.F.A, Meek M.F, Robinson P.H. and Schakernraad J.M. Peripheral nerve regeneration through P(DLLA-epsilon-CL) nerve guides. Journal of Materials Science-Materials in Medicine (1998); 9(12): 811-814. [46] Li S.T, Archibald S.J, Krarup C. and Madison R.D. Peripheral Nerve Repair with Collagen Conduits. Clinical Materials (1992); 9(3-4): 195-200. [47] Murugan R. and Ramakrishna S. Design strategies of tissue engineering scaffolds with controlled fiber orientation. Tissue Engineering (2007); 13(8): 1845-1866 [48] Ma T, Gutnick J, Salazar B, Larsen M.D, Suenaga E, Zilber S, Huang Z, Huddleston J, Smith R.L and Goodman S. Modulation of allograft incorporation by continuous infusion of growth factors over a prolonged duration in vivo. Bone (2007); 41 (3): 386392 102 [49] Liu X, Won Y.J, and Ma P.X. Porogen-induced surface modification of nano-fibrous poly (l-lactic acid) scaffolds for bone tissue engineering, Biomaterials, (2006); 27(21): 3980-3987 [50] Lewandowska M, Roguska A, Pisarek M, Polak B, Janik-Czachor M, Kurzydłowski K.J. Morphology and chemical characterization of Ti surfaces modified for biomedical applications. Biomolecular Engineering (2007); 24 (5): 438-442. [51] Lim J.I, Bin Yu, Woob K.M, Yong-Keun Lee. Immobilization of TiO2 nanofibers on titanium plates for implant applications. Applied Surface Science (2008); 255(1): 2456-2460. [52] Manara S, Paolucci F, Palazzo B,. Marcaccio M, Foresti E, Tosi G, Sabbatini S, Sabatino P, Altankov G, Roveri N. Electrochemically-assisted deposition of biomimetic hydroxyapatite–collagen coatings on titanium plate. Inorganica Chimica Acta (2007); 361:1634-1645. [53] Muschler GF. Apparatus and methods for repairing an implantable graft. 2000 US patent 6049026. [54] Muschler GF. Composite bone marrow graft material with method and kit. 2004 US patent 6723131 B2. [55] Hosseinkhani H, Hosseinkhani M, Tian F, Kobayashi H, Tabata Y. Osteogenic differentiation of mesenchymal stem cells in self-assembled peptide-amphiphile nanofibers. Biomaterials (2006); 27: 4079 103 [56] Dong W, Zhang T, McDonald M, Padilla C, Epstein J, Ryan Tian Z. Biocompatible nanofiber scaffolds on metal for controlled release and cell colonization. Nanomedicine: Nanotechnology, Biology and Medicine (2006); 2(4): 248-252. [57] Frenot A, Chronakis I.S., Polymer nanofibers assembled by electrospinning, Curr. Opin. in Colloid and Interface Sci. (2003); 8: 64–75. [58] Theron A, Zussman E, Yarin A.L. Electrostatic fields-assisted alignment of electrospun nanofibers, Nanotechnology. (2001); 12: 384 –90. [59] Shin M.Y, Hohman M.M, Brenner M, Ruteldge G.C. Experimental characterization of electrospinning: the electrically forced jet and instabilities. Polymer. (2001); 42: 9955 –67. [60] Hohman M.M, Shin M.Y, Ruteldge G.C, Brenner M. Electrospinning and electrically forced jets. I. Stability theory, Phys. Fluids. (2001); 13: 2201 –20. [61] Hohman M.M, Shin M.Y, Ruteldge G.C, Brenner M. Electrospinning and electrically forced jets. II. Applications, Phys. Fluids. (2001); 13: 2221 –36. [62] Shin M.Y, Hohman M.M, Ruteldge G.C, Brenner M. Electrospinning: a whipping fluid jet generates submicron polymer fibers, Appl. Phys. Lett. (2001); 78: 1149 –51. [63] Kun Ma, Chan C.K, Liao S, William Hwang Y.K, Qi Feng, Ramakrishna S. Electrospun nanofiber scaffolds for rapid and rich capture of bone marrow-derived hematopoietic stem cells. Biomaterials (2008); 29: 2096 – 2103. [64] Boyan B.D, Hummert T.W, Dean D.D, Schwartz Z. Role of material surfaces in regulating bone and cartilage cell response, Biomaterials. (1996); 17: 137-46. 104 [65] Yang Y, Bumgardner J.D, Cavin R, Carnes D.L, Ong J.L. Osteoblast Precursor Cell Attachment on Heat-treated Calcium Phosphate Coatings, Journal of Dental Research. (2003); 82: 449 - 453. [66] Gupta D, Venugopal J, Mitra S, Giri Dev V.R, Ramakrishna S. Nanostructured biocomposite substrates by electrospinning and electrospraying for the mineralization of osteoblasts. Biomaterials (2009); 30: 2085–2094. [67] Lausmaa J. Mattson L, Rolander U, Kasemo B. Chemical composition and morphology of titanium surface oxides, Mater. Res. Soc. Symp. Proc. (1986); 55: 351- 9. [68] Zhang W, Huang K.L, Liao S.S, Cui F.Z. Nucleation sites of calcium phosphate crystals during collagen mineralization, J. Am. Ceram. Soc. (2003); 86: 1052–4. [69] Zhang W, Liao S.S, Cui F.Z. Hierarchical self-assembly of nanofibrils in mineralized collagen, Chem. Mater. (2003); 15: 3221–6. [70] Anselme K, Bigerelle M, Noel B, Dufresne E, Judas D, Lost A. Qualitative and quantitative study of human osteoblast adhesion on materials with various surface roughnesses, J. Biomed. Mater. Res. (2000); 49: 156 - 66. [71] Cheung H.S, Story M.T, McCarty D.J. Mitogenic effects of HA and calcium pyrophosphate dihydrate crystals on cultured mammalian cells, Arthritis. Rheum. (1984); 27: 665 - 74. [72] Ducheyne P, Qiu Q. Bioactive ceramics: the effect of surface reactivity on bone formation and bone cell function, Biomaterials. (1999); 20: 2287-303. 105 [73] Adell R, Lekholm U, Rockler B, Brånemark P.I. A 15-year study of osseointegrated implants in the treatment of the edentulous jaw. Int J Oral Surg (1981); 10(6): 387-416. [74] Albrektsson T, Brånemark P.I, Hansson H.A. and Lindstrom L. Osseointegrated titanium implants. Requirements for ensuring a long-lasting, direct bone-to-implant anchorage in man. Acta Orthop. Scand. (1981); 52: 155 - 70. [75] Ogata K, Imazato S, Ehara A, Ebisu S, Kinomoto Y, Nakano T. Comparison of osteoblast responses to hydroxyapatite and hydroxyapatite/soluble calcium phosphate composites, J. Biomed. Mater. Res. (2005); 72: 127–35. [76] Zhu X, Eibl O, Scheideler L, Geis-Gerstorfer J. Characterization of nano hydroxyapatite/collagen surfaces and cellular behaviours, J. Biomed. Mater. Res. (2006); 79: 114–27. [77] Vanessa C.M, Rahim M, John E.D. The effect of discrete calcium phosphate nanocrystals on bone-bonding to titanium surfaces, Biomaterials, (2007); 28: 4748 – 4755. [78] Gotoh Y, Hiraiwa K, Narajama M. In vitro mineralization of osteoblastic cells derived from human bone, Bone Miner. (1990); 8: 239–50. [79] Stein G.S, Lian J.B, Owen T.A. Relationship of cell growth to the regulation of tissue-specific gene expression during osteoblast differentiation, FASEB J. (1990); 4: 3111–23. [80] Ohgushi H, Goldberg V.M, Caplan A.I. Heterotopic osteogenesis in porous ceramics induced by marrow cells, J. Orthop. Res. (1989); 7: 568 - 578. 106 [81] Groessner-Schreiber B, Tuan R.S. Enhanced extracellular matrix production and mineralization by osteoblasts cultured on titanium surfaces in vitro, J. Cell Sci. (1992); 101: 209-17. [82] Zreiqat H, Howlett C.R. Titanium substrata composition influences osteoblastic phenotype: in vitro study, J. Biomed. Mater. Res. (1999); 47: 360-6. [83] Lincks J, Boyan B.D, Blanchard C.R. Response of MG63 osteoblast-like cells to titanium and titanium alloy is dependent on surface roughness and composition, Biomaterials. (1998); 19: 2219-32. [84] Ong J.L, Carnes D.L, Cardenas H.L, Cavin R. Surface roughness of titanium on bone morphogenetic protein-2 treated osteoblast cells in vitro, Implant Dent. (1997); 6: 19-24. [85] Chehroudi B, Ratkay J, Brunette D. The role of implant surface geometry on mineralization in vivo and in vitro; a transmission and scanning electron microscopy study, Cells Mater. (1992); 2: 89-104. [86] Venugopal J.R, Low S, Choon A.T, Kumar A.B, Ramakrishna S. Nanobioengineered electrospun composite nanofibers and osteoblasts for bone regeneration, Artif. Organs. (2008); 32: 388–97. [87] Courteney-Harris R.G, Kayser M.V, and Downes S. Comparison of the early production of extracellular matrix on dense hydroxyapatite and hydroxyapatite coated titanium in cell and organ culture, Biomaterials. (1995); 16: 489-495. 107 [88] Niederauer G, McGee T, Keller J, Zaharias R. Attachment of epithelial cells and fibroblasts to ceramic materials, Biomaterials. (1994); 15: 342-352. [89] Gomi K, Davies J. Guided bone tissue elaboration by osteogenic cells in vitro, J. Biomed. Mater. Res. (1993); 27: 429-431. [90] Tenenbaum H, Heersche J. Differentiation of osteoblasts and formation of mineralised bone in vitro, Calcif. Tissue Int. (1982); 34: 76-79. [91] Martin J.Y, Schwartz Z, Hummert T.W, Schraub D.M, Simpson J, Lankford Jr J. Effect of titanium surface roughness on proliferation, differentiation, and protein synthesis of human osteoblast-like cells (MG63), J. Biomed. Mater. Res. (1995); 29: 389401. [92] Lincks J, Boyan B.D, Blanchard C.R. Response of MG63 osteoblast-like cells to titanium and titanium alloy is dependent on surface roughness and composition, Biomaterials. (1998); 19: 2219 - 2232. [93] Rammelt S, Illert T, Bierbaum S, Scharnweber D, Zwipp H, Schneiders W. Coating of titanium implants with collagen, RGD peptide and chondroitin sulphate, Biomaterials. (2006); 27: 5561–71. [94] Soballe K, Tansen E.S, Rasmussen H.B, Bunger C. Hydroxyapatite coating converts fibrous anchorage to bony fixation during continuous implant loading, Trans. Orthopaed. Res. Soc. (1992); 17: 270 - 8. 108 [95] Alzubaydi T.L, Alameer S.S, Ismaeel T, Aihijazi A.Y, Geetha M. In vivo studies of the ceramic coated titanium alloy for enhanced osseointegration in dental applications, J. Mater. Sci: Mater. Med. (2009); 20: 35–42. [96] DeGroot K, Geesink R, Klein C.P, Serekian P. Plasma sprayed coatings of hydroxylapatite, J. Biomed. Mater. Res. (1987); 21: 1375–81. [97] Jansen J.A. Wolke J.G, Swann S, Van der Waerden J.P, De Groot K. Application of magnetron sputtering for producing ceramic coatings on implant materials, Clin. Oral Implants Res. (1993); 4: 28–34. [98] Leeuwenburgh S, Wolke J, Schoonman J, Jansen J. Electrostatic spray deposition (ESD) of calcium phosphate coatings, J. Biomed. Mater. Res. A. (2003); 66: 330–4. [99] LeGeros R.Z, LeGeros J.P, Kim Y, Kijkowska R, Zheng R, Bautista C. Calcium phosphates in plasma-sprayed HA coatings, Ceram. Trans. (1995); 48: 173–89. 109 Appendix A Figure: MSC proliferation on TCP at 24, 48, 72 and 96 hrs. 110 Appendix B FESEM-EDX results on Ti alloy coated with PLGA/Collagen/HA on day 7, 14 and 21. 111 1/1 View000 --------------------------Title : IMG1 --------------------------Instrument : 6700F Volt : 15.00 kV Mag : x 3,000 Date : 2009/06/16 Pixel : 512 x 384 --------------------------- 10 µm 001 Acquisition Parameter Instrument : 6700F Acc. Voltage : 15.0 kV Probe Current: 1.00000 nA PHA mode : T3 Real Time : 60.87 sec Live Time : 50.00 sec Dead Time : 18 % Counting Rate: 3595 cps Energy Range : 0 - 20 keV 001 4400 4000 3600 3200 2400 CaKa PKa Counts 2800 2000 CaKb 1600 1200 800 400 0 0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 keV ZAF Method Standardless Quantitative Analysis Fitting Coefficient : 0.7151 Element (keV) mass% Error% At% Compound P K 2.013 32.94 1.62 38.86 Ca K 3.690 67.06 3.41 61.14 Total 100.00 100.00 AnalysisStation mass% Cation K 33.0606 69.2746 1/1 View001 --------------------------Title : IMG1 --------------------------Instrument : 6700F Volt : 15.00 kV Mag : x 1,600 Date : 2009/06/16 Pixel : 512 x 384 --------------------------- 20µm 001 Acquisition Parameter Instrument : 6700F Acc. Voltage : 15.0 kV Probe Current: 1.00000 nA PHA mode : T3 Real Time : 60.51 sec Live Time : 50.00 sec Dead Time : 17 % Counting Rate: 3487 cps Energy Range : 0 - 20 keV 0.00 1.00 2.00 CaKa CaKb PKa 001 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 keV ZAF Method Standardless Quantitative Analysis Fitting Coefficient : 0.9557 Element (keV) mass% Error% At% Compound P K* 2.013 40.19 12.62 46.51 Ca K* 3.690 59.81 27.19 53.49 Total 100.00 100.00 AnalysisStation mass% Cation K 41.1509 61.4710 1/1 View000 --------------------------Title : IMG1 --------------------------Instrument : 6700F Volt : 15.00 kV Mag : x 3,000 Date : 2009/06/16 Pixel : 512 x 384 --------------------------- 10 µm 001 Acquisition Parameter Instrument : 6700F Acc. Voltage : 15.0 kV Probe Current: 1.00000 nA PHA mode : T3 Real Time : 57.00 sec Live Time : 50.00 sec Dead Time : 12 % Counting Rate: 2651 cps Energy Range : 0 - 20 keV 001 6000 5500 5000 4500 Counts 4000 3500 3000 2500 1500 CaKa CaKb PKa 2000 1000 500 0 0.00 1.00 2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 keV ZAF Method Standardless Quantitative Analysis Fitting Coefficient : 0.9655 Element (keV) mass% Error% At% Compound P K* 2.013 43.16 16.72 49.56 Ca K* 3.690 56.84 36.39 50.44 Total 100.00 100.00 AnalysisStation mass% Cation K 44.5089 58.2319 [...]... knowledge, dental implant using functionalized nanofibers as a surface modification is a novel idea to enhance osseointegration using the bone regeneration concept 15 Chapter 1 Introduction 1.1 Background In the past 20 years, the number of dental implant procedures has increased steadily worldwide, reaching about one million dental implantations per year [1] Dental implants are useful for restoration of oral... period of 12 weeks [5] and this was reduced to 6 to 8 weeks with the introduction of the SLA (sand blasted, acid etched) surface [6] The differences in the contact angle and the surface roughness of the implant surface owing to the various surface modification techniques were shown in Table 2.1 Table 2.1 Different types of implant surface modifications and their surface roughness and contact angle Type of. .. short- and long-term success of the implants These parameters are associated with delicate surgical techniques, a prerequisite for a successful early clinical outcome High success rates for dental implants are reported in healthy patients with good bone quality In the future, with an aging population, more patients may be considered for dental implants; osseointegration of dental implants under less than... called the ―dip‖ Many implant failures occur during this period, and this period seems to be critical to the successful integration of the implant [2] 17 Figure 1: A typical Ti dental implant 18 1.3 Hypothesis and Objectives: Hypothesis This project is to develop a surface modification system for dental implant using electrospun nanofiber and biomineralization to fabricate a biomimetic substrate We... be used for the surface modification or activation of an implant surface Among these methods, chemical modifications seem to be relatively simple and inexpensive Hence it is widely used There have been various techniques tried out in the past to improve the surface roughness of the implant like plasma treatment, acid-etching and heat treatment For example, the TPS (titanium 21 plasma sprayed) surfaces... average 13 SUMMARY The introduction of dental implants has changed the way dentists approach the replacement of missing teeth The clinical success of dental implants is related to their osseointegration, which is a property virtually unique to titanium and has enhanced the science of joint replacement techniques Generally, the time between implant placement and implant loading ranged from 3 months... nanostructured surfaces of nanometallic and nanoceramic materials have several advantages compared to the conventional surfaces These include, (i) they possess greater surface roughness resulting from both decreased grain size and possibly decreased diameter of surface pores, (ii) enhanced surface wettability due to greater surface roughness and (iii) greater numbers of grain boundaries There are a number of physical... cases, enhanced bone formation around the implant would be an important criterion 16 This may be achieved by implant coatings that are able to interact actively with the surrounding tissues 1.2 Clinical problems associated with osseointegration: There are two types of responses exhibited by the body after implantation The first type involves the formation of a soft fibrous tissue around the implant. .. differentiation, which are crucial for enhanced osseointegration Objectives:  Modify the implant surface to produce nanotextured topography  Develop a nanofibrous coating from biodegradable synthetic polymers and/or natural polymers to mimic extracellular matrix  Functionalization of the nanofiber by biomineralization  Evaluate adhesion, proliferation of MSCs on the modified implant surface  Investigate osteogenic... chemistry of the surfaces In all the cases TiO2 being the principle chemical component [16] The Ti metal spontaneously forms a protective TiO2 layer in the atmosphere When the Ti implant is inserted into the human body, the surrounding tissues directly contact the TiO2 layer on the implant surface The surface characteristics of the TiO2 layer determine the biocompatibility of Ti implant Therefore, it ... strength of the PLGA and PLGA/Collagen coated implant surface owing to the hard nature of the implant material and the thin layer of nanofiber coating on the implant surface The tensile properties of. .. integration of the implant [2] 17 Figure 1: A typical Ti dental implant 18 1.3 Hypothesis and Objectives: Hypothesis This project is to develop a surface modification system for dental implant using... roughness of the implant surface owing to the various surface modification techniques were shown in Table 2.1 Table 2.1 Different types of implant surface modifications and their surface roughness