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Bio-inspired calcium phosphate/biopolymer nanocomposite fibrous scaffolds for hard tissue regeneration Chae, Taesik 2015

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  BIO-INSPIRED CALCIUM PHOSPHATE/BIOPOLYMER NANOCOMPOSITE FIBROUS SCAFFOLDS  FOR HARD TISSUE REGENERATION  by  Taesik Chae  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in  THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES  (Materials Engineering)  THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)    March 2015   Taesik Chae, 2015                                                                                                                  ii  Abstract This study discloses an original process for making calcium phosphate (CaP)/biopolymer nanocomposite fibrous scaffolds by biomimetic in-situ synthesis and electrospinning. Electrospinning (ES) produces non-woven nanofibrous mesh structure with 3D interconnected pores and a high surface area by applying electrostatic force to polymer-based solution. The resulting topography of the scaffolds mimics the natural extracellular matrix of human tissues, with the potential application in tissue engineering, drug delivery, and wound dressing. We have demonstrated that the possibility of inclusion of CaP into biopolymer nanofibers, inspired by mineralized collagen fibrils in bone tissue, makes ES an attractive processing route for preparation of the nanocomposites for bone tissue regeneration. Two different nanocomposite fibers were explored; i) poly(lactic acid) (PLA) with dicalcium phosphate anhydrate (DCPA) and ii) alginate with hydroxyapatite (HAp). In-situ synthesized DCPA in non-aqueous PLA solution were electrospun into self-fused and intra-nano porous networks. Homogeneous dispersion of DCPA nanocrystallites in the PLA nanofibers was induced by controlling the interaction of Ca2+ ions and the carbonyl groups in PLA, providing nucleation sites for DCPA during the in-situ synthesis. It is shown that the nucleation and growth of HAp on electrospun alginate nanofibers was generated at the [–COO–]–Ca2+–[–COO–] linkage sites on electrospun alginate nanofibers impregnated with PO43– ions during cross-linking treatment of alginate. This novel in-situ synthesis developed in this work resulted in the uniform distribution of the CaP nanophases and avoided agglomeration of the inorganic nanoparticles fabricated by the conventional  mechanical blending method. Rat calvarial osteoblasts were stably attached and proliferated faster on the CaP/biopolymer nanocomposites fibrous scaffolds than the pure polymer scaffolds, respectively. Mineralized bone-like nodules deposited after 6 weeks of seeding on DCPA/PLA scaffolds. The unique nanofibrous architectures combined with the CaP nanophases were engineered using ES and the novel biomimetic in-situ synthesis. It is anticipated that the nanocomposite systems mimicking the mineralized collagen fibrils in bone tissue could be advantageous in bone tissue regenerative medicine applications.   iii  Preface  Parts of Chapter 5 have been published in the Journal of Biomedical Materials Research Part A (T. Chae, H. Yang, F. Ko, T. Troczynski. Bio-inspired dicalcium phosphate anhydrate/poly(lactic acid) nanocomposite fibrous scaffolds: In-situ synthesis and electrospinning. J Biomed Mater Res Part: A 2014;102(2):514‒522). I conducted all of the experiments and data analysis, and wrote the manuscript as a first author. Prof. Tom Troczynski, Prof. Frank Ko, and Dr. Heejae Yang advised me with suggestions to improve the paper.   Parts of Chapter 6 and Chapter 7 have been published in the Journal of Materials Science: Materials in Medicine  (T. Chae, H. Yang, V. Leung, F. Ko, T. Troczynski. Novel biomimetic hydroxyapatite/alginate nanocomposite fibrous scaffolds for bone tissue engineering. J Mater Sci Mater Med 2013;24(8):1885‒1894). I conducted all of the experiments and data analysis, and wrote the manuscript as a first author. Prof. Tom Troczynski, Prof. Frank Ko, and Dr. Heejae Yang advised me with suggestions to improve the paper. Dr. Victor Leung conducted the optimization of the alginate electrospinning process.             iv  Table of Contents  Abstract ···································································································· ii Preface····································································································· iii Table of Contents ························································································ iv List of Tables ···························································································· viii List of Figures ····························································································· x List of Symbols and Abbreviations ································································ xviii Acknowledgements ······················································································ xx Dedication ······························································································· xxi Chapter 1 Introduction and Rationale·································································· 1 Chapter 2 Literature Review ············································································ 5 2.1 Bone tissue ························································································· 5 2.1.1 Structure and composition of bone ························································ 5 2.1.2 Healing and repair of bone ·································································· 8 2.2 Biomaterials for bone tissue ···································································· 10 2.2.1 Bioceramic calcium phosphates ··························································· 11 2.2.1.1 Hydroxyapatite ·········································································· 13 2.2.1.2 Dicalcium phosphate anyhydrate ····················································· 15 2.2.2 Biopolymers ·················································································· 16 2.2.2.1 Poly(lactic acid) ········································································· 16 2.2.2.2 Alginate ··················································································· 19 2.2.3 Calcium phosphate/biopolymer composite scaffolds ·································· 21 2.3 Electrospinninng ·················································································· 24 2.3.1 Tissue engineering scaffolds ······························································· 28 2.3.2. Drug delivery devices ······································································ 30 Chapter 3 Scope and Objectives ······································································· 32 3.1 Scope of the investigation ······································································· 32 3.2 Objectives ························································································· 34   v  Chapter 4 Materials and Methodology ······························································· 36 4.1 Fabrication of DCPA/PLA nanocomposite fibrous scaffolds ······························ 36 4.1.1 Raw materials ················································································ 36 4.1.2 In-situ synthesis of DCPA/PLA nanocomposite suspension ·························· 36 4.1.3 Electrospinning of DCPA/PLA nanocomposite suspension ·························· 38 4.2 Fabrication of HAp/alginate nanocomposite fibrous scaffolds ···························· 39 4.2.1 Raw materials ················································································ 39 4.2.2 Preparation of alginate/PEO solution ····················································· 39 4.2.3 Electrospininng of alginate/PEO solution ··············································· 40 4.2.4 In-situ synthesis for HAp/alginate nanocomposite fibrous scaffolds ················ 41 4.3 Characterization of CaP/biopolymer nanocomposite fibrous scaffolds ·················· 42 4.3.1 X-ray diffraction ············································································· 42 4.3.2 Scanning electron microscopy ····························································· 42 4.3.3 Scanning and transmission electron microscopy ······································· 42 4.3.4 Fourier transform infrared spectroscopy ················································· 43 4.3.5 Thermogravimetric analysis ······························································· 43 4.3.6 Tensile properties of scaffolds ···························································· 43 4.3.7 In-vitro bioactivity analysis ································································ 44 4.4 Rat calvarial osteoblast culture on scaffolds ················································· 45 4.4.1 Scanning electron microscopy ····························································· 45 4.4.2 Immunocytochemistry staining ··························································· 46 4.4.3 MTS assay ···················································································· 47 4.4.4 DAPI staining ················································································ 47 4.4.5 Tetracycline labeling ········································································ 48 4.4.6 Multiphoton microscopy ··································································· 48     vi  Chapter 5 Dicalcium Phosphate Anhydrate/Poly(lactic acid) Nanocomposite Fibrous Scaffolds ·································································································· 50 5.1 Fabrication of DCPA/PLA scaffolds ·························································· 50 5.1.1 Biomimetic in-situ synthesis of DCPA in PLA solution ······························ 50 5.1.2 Electrospinning DCPA/PLA scaffolds ··················································· 52 5.1.3 Effects of PEG on DCPA/PLA/PEG scaffolds ········································· 57 5.1.4 Uniform DCPA/PLA nanofibers after re-electrospinning process ··················· 59 5.2 Characterization of DCPA/PLA scaffolds ···················································· 61 5.2.1 Chemical interactions between DCPA and PLA during in-situ synthesis ·········· 61 5.2.2 Quantification of DCPA in DCPA/PLA scaffolds ····································· 64 5.2.3 Tensile properties of scaffolds ···························································· 67 5.2.4 In-vitro bioactivity of DCPA/PLA scaffolds ············································ 80 5.3 Conclusions ······················································································· 85 Chapter 6 Hydroxyapatite/Alginate Nanocomposite Fibrous Scaffolds ························· 86 6.1 Fabrication of HAp/alginate scaffolds ························································ 86 6.1.1 Electrospinning and cross-linking of alginate scaffolds ······························· 86 6.1.2 Fabrication of HAp/alginate scaffold via in-situ synthesis ···························· 89 6.2 Characterization of HAp/alginate scaffolds ·················································· 95 6.2.1 Chemical interaction between HAp and alginate during in-situ synthesis ·········· 95 6.2.2 Quantification of HAp in HAp/alginate scaffolds ······································ 99 6.2.3 Tensile properties of HAp/alginate scaffolds ········································· 100 6.2.4 In-vitro bioactivity of HAp/alginate scaffolds ········································ 110 6.3 Conclusions ····················································································· 114 Chapter 7 In-vitro Osteoblasts Response on Scaffolds ··········································· 115 7.1 Dicalcium phosphate anhydrate/poly(lactic acid) scaffolds ······························ 115 7.1.1 Attatchment of osteoblasts on scaffolds ··············································· 115 7.1.2 Proliferation of osteoblasts on scaffolds ··············································· 118 7.1.3 Bone-like nodule formation on scaffolds ·············································· 124   vii  7.2 Hydroxyapatite/Alginate scaffolds ·························································· 131 7.2.1 Attatchement of osteoblasts on scaffolds ·············································· 131 7.2.2 Proliferation of osteoblasts on scaffolds ··············································· 135 7.3 Conclusions ····················································································· 141 Chapter 8 Conclusions and Future Work ··························································· 143 8.1 DCPA/PLA nanocomposite fibrous scaffolds ············································· 143 8.2 HAp/alginate nanocomposite fibrous scaffolds ············································ 146 8.3 Future work ····················································································· 148 References ······························································································ 151 Appendix ································································································ 168 A.1 Details of statistical analysis of MTS assay ··············································· 168 A.2 Details of fiber diameter measurement ····················································· 177               viii  List of Tables  Table 2.1 Acronyms, formulas, Ca/P ratios, and solubility products of representative calcium phosphates. ····················································································· 12  Table 2.2 Fabrication techniques for 3D porous composite scaffolds and their advantages and disadvantages. ······················································································· 23  Table 4.1 Compositions of  in-situ synthesized DCPA/PLA and DCPA/PLA/PEG nanocomposite suspensions and reference solution. ················································ 37  Table 4.2 Compositions of H-Alginate solutions and reference solution for electrospinning.40  Table 4.3 Raw materials and ion concentration of original Kokubo's SBF solution [140]. ··· 44  Table 5.1 Calcium phosphate content in DCPA/PLA scaffolds measured by TGA (n = 3). ·· 64 Table 5.2 Mechanical properties (in tension) of PLA and DCPA/PLA nanofibrous scaffolds (n = 6). ····································································································· 68  Table 6.1 HAp content in cross-linked/in-situ synthesized H-Alginate scaffolds measured by TGA (n = 3). ·························································································· 99  Table 6.2 Mechanical properties (in tension) of cross-linked Ref-Alginate and cross-linked/in-situ synthesized H-Alginate scaffolds (n = 6). ········································· 101  Table 8.1 Comparison of CaP/biopolymer nanocomposite fibrous scaffolds properties studied in this study. ··················································································· 147  Table A.1 Measured optical density of MTS assay of PLA-based scaffolds at day 1. ······· 167  Table A.2 Measured optical density of MTS assay of PLA-based scaffolds at day 3. ······· 168  Table A.3 Measured optical density of MTS assay of PLA-based scaffolds at day 7. ······· 170  Table A.4 Measured optical density of MTS assay of alginate-based scaffolds at day 1. ··· 171  ix  Table A.5 Measured optical density of MTS assay of alginate-based scaffolds at day 3. ··· 173  Table A.6 Measured optical density of MTS assay of alginate-based scaffolds at day 7. ··· 174                      x  List of Figures  Figure 2. 1 The 7 hierarchical levels of bone tissue from nano to macro-scales [4]. ............... 6  Figure 2.2 Hierarchical structural organization of bone: (a) cortical and cancellation bone; osteons with Haversian systems; (c) lamellae; (d) fiber assemblies of collagen fibrils; (e) bone mineral crystals, collagen molecules, and non-collagenous proteins [39]. ...................... 7  Figure 2.3 The healing stages of a bone fracture: (a) fracture hematoma formation and inflammation, (b) soft callus formation, (c) hard callus formation, and (d) remodeling [47]. . 9  Figure 2.4 Solubility isotherms of calcium phosphate phases at 37 °C [58]. TCa and Tp are the total molar concentrations of calcium and phosphate ions, respectively. ......................... 13  Figure 2.5 Synthesis of poly(lactic acid) [17]. ....................................................................... 18  Figure 2.6 Structure of alginic acid extracted from Lessonia trabeculata marine algae; (a) D-mannuronic residue: M, (b) L-guluronic residue: G, and (c) polysaccharide chains of MM, GG and MG blocks [95]. ............................................................................................... 19  Figure 2.7 The egg-box model for alginate gellation with calcium ions. Guluronic acid (G) blocks of alginate are held together by a number of calcium ions [94]. ................................. 20  Figure 2.8 Typical morphologies of 3D porous scaffolds produced by different techniques and structure of cancelleous bone. (a) thermally induced phase separation [104], (b) solvent casting and particles leaching [107], (c) solid free-form [109], (d) microsphere sintering [110], and (e) cancellous bone [111]....................................................................................... 22  Figure 2.9 Elastic modulus vs. compressive strength of biodegradable polymers, bioactive ceramics, and composites [6]. ................................................................................................. 24  Figure 2.10 Schematic illustration of electrospinning system and process. ........................... 25  Figure 2.11 SEM micrograph of electrospun poly(vinyl alcohol) nanofibrous scaffold. ....... 26    xi  Figure 4.1 Electrospinning system for the fabrication of DCPA/PLA nanocomposite fibrous scaffolds (NANON, MECC Co., Ltd., Japan). ....................................................................... 38  Figure 4.2 Electrospinning system for the fabrication of HAp/alginate nanocomposite fibrous scaffolds (NEU-1, Kato Tech. Co. Ltd., Japan). ......................................................... 41  Figure 5.1 XRD patterns of (a) Ref-PLA, (b) DCPA/PLA-3, (c) DCPA/PLA-4, and (d) DCPA/PLA-8. The sample (e) was synthesized in the absence of PLA with the same CNT concentration as (b) showing all characteristic crystalline DCPA peaks, matched with PDF # 01-070-1425. ........................................................................................................................ 51  Figure 5.2 TEM micrographs of (a) DCPA/PLA-4 dense nanofiber, (b) DCPA/PLA-4 porous nanofiber, (c) Ref-PLA nanofiber, and (d) Ref-DCPA/PLA-Mech nanofiber. The insets are the selected area electron diffraction (SAED) patterns on the spots with the white arrows. .................................................................................................................................... 52  Figure 5.3 (a) STEM micrograph and (b–d) EDS mapping of DCPA/PLA-4 nanocomposite fiber. ........................................................................................................................................ 53  Figure 5.4 SEM micrographs of (a) Ref-PLA nanofibrous scaffold and (b) Ref-DCPA/PLA-Mech nanocomposite fibrous scaffold. ................................................................................... 54  Figure 5.5 SEM micrographs of (a) DCPA/PLA-2, (b) DCPA/PLA-3 and (c) DCPA/PLA-4 nanocomposite fibrous scaffolds. The white arrows and circles indicate intra-nanopores and self-fused nanofibers, respectively. ......................................................................................... 55  Figure 5.6 Fiber diameter distributions of (a) Ref-PLA nanofibrous scaffold, (b) DCPA/PLA-2, (c) DCPA/PLA-3, and (d) DCPA/PLA-4 nanocomposite fibrous scaffolds. .. 56  Figure 5.7 SEM micrographs of (a) DCPA/PLA/PEG-2, (b) DCPA/PLA/PEG-4, (c) DCPA/PLA/PEG-6, and (d) DCPA/PLA/PEG-8 nanocomposite fibrous scaffolds. .............. 58 Figure 5.8 Fiber diameter distributions of (a) DCPA/PLA/PEG-2, (b) DCPA/PLA/PEG-4, (c) DCPA/PLA/PEG-6, and (d) DCPA/PLA/PEG-8 nanocomposite fibrous scaffolds.......... 59  Figure 5.9 SEM and TEM (inset) micrographs of (a) Ref-PLA and (b) Re-DCPA/PLA/PEG-4 nanofibrous scaffolds. ............................................................................. 60    xii  Figure 5.10 Fiber diameter distributions of (a) Ref-PLA and (b) Re-DCPA/PLA/PEG-4 nanofibrous scaffolds. ............................................................................................................. 61  Figure 5.11 FT-IR spectra of (a) DCPA/PLA-4 nanocomposite fibrous scaffold and (b) Ref-PLA nanofibrous scaffold. ...................................................................................................... 62  Figure 5.12 Chemical reaction scheme of in-situ synthesized dicalcium phosphate anhydrate in poly(lactic acid) solution. .................................................................................. 63  Figure 5.13 Representative TGA graphs of (a) Ref-PLA nanofibrous scaffold, (b) DCPA/PLA-2, (c) DCPA/PLA-3, and (d) DCPA/PLA-4 nanocomposite fibrous scaffolds. .. 65  Figure 5.14 Representative TGA graphs of (a) Ref-PLA nanofibrous scaffold, (b) DCPA/PLA/PEG-2, and (c) DCPA/PLA/PEG-4 nanocomposite fibrous scaffolds. .............. 66  Figure 5.15 Representative TGA graphs of (a) Ref-PLA nanofibrous scaffold, (b) Re-DCPA/PLA/PEG-2, and (c) Re-DCPA/PLA/PEG-4 nanocomposite fibrous scaffolds. ........ 67  Figure 5.16 Tensile stress-strain curves of Ref-PLA scaffold. ............................................... 69  Figure 5.17 Tensile stress-strain curves of DCPA/PLA/PEG-2 scaffold. ............................... 70  Figure 5.18 Tensile stress-strain curves of DCPA/PLA/PEG-4 scaffold. ............................... 71  Figure 5.19 Tensile stress vs. strain curves of Re-DCPA/PLA/PEG-2 scaffold. .................... 72  Figure 5.20 Tensile stress vs. strain curves of Re-DCPA/PLA/PEG-4 scaffold. .................... 73  Figure 5.21 SEM micrographs of Micro-Mech-DCPA/PLA-4 nanocomposite fibrous scaffold. .................................................................................................................................. 75  Figure 5.22 Tensile stress vs. strain curves of Micro-Mech-DCPA/PLA-2 scaffold. ............. 76  Figure 5.23 Tensile stress vs. strain curves of Micro-Mech-DCPA/PLA-4 scaffold. ............. 77  Figure 5.24 Representative stress-strain curves of PLA and DCPA/PLA scaffolds. .............. 79  xiii  Figure 5.25 SEM micrographs of (a, d) Ref-PLA, (b, e) Re-DCPA/PLA/PEG-2, and (c, f)  Re-DCPA/PLA/PEG-4 scaffolds after immersed in simulated body fluid solution for 36 hours (a, b, c) and 10 days (d, e, f). ........................................................................................ 81  Figure 5.26 XRD patterns of (a) Ref-PLA, (b) Re-DCPA/PLA/PEG-2, and (c) Re-DCPA/PLA/PEG-4 scaffolds after immersed in simulated body fluid solution for 10 days. HAp peaks were identified and assigned from PDF # 01-072-1243. ..................................... 82  Figure 5.27 Energy-dispersive X-ray spectroscopy spectrum on the surface of Re-DCPA/PLA/PEG-4 scaffold after soaking in simulated body fluid solution for 10 days. ...... 83  Figure 6.1 SEM micrographs of (a) as-electrospun sodium Ref-Alginate and (b) cross-linked calcium Ref-Alginate scaffolds, and (c) fiber diameter distribution measurement of a as-electrospun sodium Ref-Alginate and b cross-linked calcium Ref-Alginate scaffolds. . 87  Figure 6.2 FT-IR spectra of (a) PEO, (b) sodium alginate, (c) as-electrospun sodium Ref-Alginate scaffold, and (d) cross-linked calcium Ref-Alginate scaffold. ................................ 88  Figure 6.3 SEM micrographs of (a) as-electrospun H-Alginate-5 and (b) cross-linked/in-situ synthesized H-Alginate-5 scaffolds. ................................................................................ 90  Figure 6.4 EDS spectroscopy of as-electrospun H-Alginate-5 scaffold with P element area mapping. ................................................................................................................................. 91  Figure 6.5 Fiber diameter distributions of (a) as-electrospun H-Alginate-5 and (b) cross-linked/in-situ synthesized H-Alginate-5 scaffolds. ................................................................. 92  Figure 6.6 SEM micrographs of  cross-linked/in-situ synthesized (a) H-Alginate-2 and (b) H-Alginate-8 scaffolds. ........................................................................................................... 93  Figure 6.7 XRD patterns of (a) cross-linked Ref-Alginate, cross-linked/in-situ synthesized (b) H-Alginate-2, (c) H-Alginate-5, and (d) H-Alginate-8 scaffolds. HAp peaks were identified and assigned from PDF # 01-072-1243. ................................................................. 94  Figure 6.8 FT-IR spectra of (a) cross-linked Ref-Alginate, cross-linked/in-situ synthesized (b) H-Alginate-2, (c) H-Alginate-5, and (d) H-Alginate-8 scaffolds. .................................... 96   xiv  Figure 6.9 Chemical structure of (a) “egg box” model of calcium alginate, (b) “egg box” model of calcium alginate with precursor ions for HAp nucleation, and (c) mineralized “egg-box” structure with HAp, and (d) illustration of cross-linked/in-situ synthesized HAp/alginate nanocomposite fibrous scaffold. ...................................................................... 97  Figure 6.10 SEM micrograph of cross-linked Mech-H-Alginate-2 scaffold fabricated using a mechanical blending/electrospinning method. The white arrows indicate the agglomerated HAp particles at micro-meter levels. ............................................................... 98  Figure 6.11 TGA patterns of (a) crossed-linked Ref-Alginate, cross-linked/in-situ synthesized (b) H-Alginate-2, (c) H-Alginate-5, and (d) H-Alginate-8 scaffolds. ............... 100  Figure 6.12 Tensile stress-strain curves of cross-linked Ref-Alginate nanofibrous scaffold.102  Figure 6.13 Tensile stress-strain curves of cross-linked/in-situ synthesized H-Alginate-2 nanocomposite fibrous scaffold. ........................................................................................... 103  Figure 6.14 Tensile stress-strain curves of cross-linked/in-situ synthesized H-Alginate-5 nanocomposite fibrous scaffold. ........................................................................................... 104  Figure 6.15 Tensile stress-strain curves of cross-linked Mech-H-Alginate-2 nanocomposite fibrous scaffold. .................................................................................................................... 106  Figure 6.16 Tensile stress-strain curves of cross-linked Mech-H-Alginate-5 nanocomposite fibrous scaffold. .................................................................................................................... 107  Figure 6.17 Representative tensile stress-strain curves of alginate and HAp/alginate nanofibrous scaffolds. ............................................................................................................110  Figure 6.18 SEM micrographs of (a) cross-linked Ref-Alginate nanofibrous scaffold  and (b) cross-linked/in-situ synthesized H-Alginate-5 nanocomposite fibrous scaffold after immersed in simulated body fluid solution for 36 hours. ...................................................... 111  Figure 6.19 XRD patterns of (a) cross-linked Ref-Alginate nanofibrous scaffold  and (b) cross-linked/in-situ synthesized  H-Alginate-5 nanocomposite fibrous scaffold after immersed in simulated body fluid solution for 36 hours. HAp peaks were identified and assigned from PDF # 01-072-1243. .......................................................................................112   xv  Figure 7.1 SEM micrographs of RCO cells cultured on (a) Ref-PLA and (b) Re-DCPA/PLA/PEG-4 scaffolds for 7 days. The magnified inset SEM micrographs of the DCPA/PLA scaffold shows closely and stably attached and spread RCO cells on the electrospun nanofibrous structure at day 3 of post cell-seeding. The multiple filopodial connections on the surface were made along the nanofibers. ··································· 116  Figure 7.2 Immunofluorescence images of RCO cells cultured on (a) Ref-PLA scaffold for 1 day, (b) Ref-PLA for 7 days, (c) Re-DCPA/PLA/PEG-4 scaffold for 1 day, and (d) Re-DCPA/PLA/PEG-4 scaffold for 7 days. Actin filaments were labeled red with phalloidin (left), nuclei were stained blue with DAPI (middle), and the two fluorescent images were merged (right). ························································································· 117  Figure 7.3 MTS assays of cultured RCO cells on (a) PLA film, (b) Ref-PLA, (c) Re-DCPA/PLA/PEG-2, (d)  Re-DCPA/PLA/PEG-4, and (e) Micro-Mech-4 scaffolds for 7 days. Triplicate samples were tested for each scaffold. Statistical significance was selected at p < 0.05, using one-way ANOVA for the specimens at each day. ··································· 119  Figure 7.4 Average number of RCO cells cultured on (a) PLA film, (b) Ref-PLA, (c) Re-DCPA/PLA/PEG-2, (d)  Re-DCPA/PLA/PEG-4, and (e) Micro-Mech-4 scaffolds for 7 days. Triplicate samples were tested for each scaffold. Nuclei stained RCO cells with DAPI were counted using ImageJ software. Six images per scaffold were taken, and triplicate samples were prepared for each scaffold. ····································································· 121  Figure 7.5 Fluorescent images of DAPI stained nuclei of RCO cells cultured on (a) PLA film, (b) Ref-PLA, (c) Re-DCPA/PLA/PEG-2, (d)  Re-DCPA/PLA/PEG-4, and (e) Micro-Mech-4 scaffolds for 7 days. ········································································· 122  Figure 7.6 (a, b) SEM micrographs and (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Ref-PLA scaffold for 2 weeks. ······································· 125  Figure 7.7 (a, b) SEM micrographs and (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Ref-PLA scaffold for 4 weeks. ······································· 126  Figure 7.8 (a, b) SEM micrographs and (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Re-DCPA/PLA/PEG-4 scaffold for 4 weeks. ······················ 127  Figure 7.9 (a, b) SEM micrographs and  (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Ref-PLA scaffold for 6 weeks. ······································· 127   xvi  Figure 7.10 Epifluorescence images of bone-like nodules produced by RCO cells cultured on Re-DCPA/PLA/PEG-4 scaffold for (a) 2, (b) 4, and (c) 6 weeks. ·························· 128  Figure 7.11 (a−f) MPM micrographs of RCO cells cultured on Re-DCPA/PLA/PEG-4 scaffold for 6 weeks and 3D images of (g) the scaffold and (h) cell layers. The 2D micrographs were scanned with every 1 µm stack. The numbers indicate each stack of the scanned images. Amira® software was used to produce 3D images from 2D images. The mineralized deposits (white arrows) were mostly found between the cell layers and the nanofiber surface. The osteoblasts migrated through the nanofibrous structure, as shown in the rectangles. ·························································································· 129  Figure 7.12 SEM micrographs of RCO cells cultured on (a) cross-linked Ref-Alginate and (b) cross-linked/in-situ synthesized H-Alginate 5 scaffolds for 7 days. The white arrows indicate more closely and stably attached RCO cells on the HAp/alginate than the pure alginate scaffold at day 1 of post cell-seeding. The magnified inset SEM micrograph of the HAp/alginate scaffold shows the visible development of multiple filopodial connections to the surface at day 7 of post cell-seeding. ··························································· 131  Figure 7.13 Immunofluorescence images of RCO cells cultured on (a) cross-linked Ref-Alginate scaffold for 1 day, (b) cross-linked Ref-Alginate scaffold for 7 days, (c) cross-linked/in-situ synthesized H-Alginate-5 scaffold for 1 day, and (d) cross-linked/in-situ synthesized H-Alginate-5 scaffold for 7 days. Actin filaments were labeled red with phalloidin (left), nuclei were stained blue with DAPI (middle), and the two fluorescent images were merged (right). ········································································· 133  Figure 7.14 MTS assays of cultured RCO cells on (a) alginate film, (b) cross-linked Ref- Alginate, (c) cross-linked/in-situ synthesized H-Alginate-2, (d) cross-linked/in-situ synthesized H-Alginate-5, and (e) cross-linked Mech-H-Alginate-5 scaffolds for 7 days. Triplicate samples were tested for each scaffold. Statistical significance was selected at p < 0.05, using one-way ANOVA for the specimens at each day. ··································· 135  Figure 7.15 Average number of RCO cells cultured on (a) alginate film, (b) cross-linked Ref-Alginate, (c) cross-linked/in-situ synthesized H-Alginate-2, (d) cross-linked/in-situ synthesized H-Alginate-5, and (e) cross-linked Mech-H-Alginate-5 scaffolds for 7 days. Nuclei stained RCO cells with DAPI were counted using ImageJ software. Six images per scaffold were taken, and triplicate samples were prepared for each scaffold. ················ 137      xvii  Figure 7.16 Fluorescent images of DAPI stained nuclei of RCO cells cultured on (a) alginate film, (b) cross-linked Ref-Alginate, (c) cross-linked/in-situ synthesized H-Alginate-2, (d) cross-linked/in-situ synthesized H-Alginate-5, and (e) cross-linked Mech-H-Alginate-5 scaffolds. ··············································································· 138                       xviii  List of Symbols and Abbreviations  C concentration of solution d fiber diameter   H distance between spinneret capillary and collector ht fiber diameter I electric current L length of capillary  Q solution feeding rate R radius of capillary Vc critical voltage for electrospinning jet formation x displacement of centerline of electrospinning jet γ surface tension of solution ε dielectric permittivity η solution viscosity in poise or intrinsic viscosity of solution   A/W   apatite/wallastonite     ACP   amorphous calcium phosphate    BMP-2  bone morphogenetic protein-2     CaP   calcium phosphates    CNT   calcium nitrate tetrahydrate    DCPA   dicalcium phosphate anhydrate    DCPD   dicalcium phosphate dihydrate    DMF   dimethyl formamid    DMSO   dimethyl sulfoxide    ECM   extra-cellualr matrix  EDS       energy dispersive spectroscopy   ES   electrospinning     xix  FDA   Food and Drug Administration FT-IR    Fourier transform infrared spectroscopy    GAG   glycosaminoglycan    HAp   hydroxyapatite    HMSc   human bone marrow-derived mesenchymal stem cells  MPM     multiphoton microscopy    MTS   3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-               tetrazolium    OCP   octacalcium phosphate PBS       phosphate-buffered saline    PCL   poly(ε-carprotacton)  PEG   poly(ethylene glycol)   PGA   poly(glycolic acid)    PLA   polyl(actic acid)    PLGA   poly(lactic-co-glycolic acid)  RCO      rat calvarial osteoblast   SBF   simulated body fluid  SEM      scanning electron microscopy    SFF   solid free form  SHG      second harmonic generation    SPM   sodium  phosphate monobasic dihydrate    TCP   tricalcium phosphate  TEM      transmission electron microscopy TGA      thermogravimetric analysis   TIPS   thermally induced phase separation  TPEF     two-photon excited fluorescence   TTCP   tetracalcium phosphate     xx  Acknowledgements  First of all, I would like to acknowledge and express my sincere gratitude to my supervisor, Prof. Tom Troczynski, for his comprehensive scientific guidance and advice and for his patience in training me. I would also like to thank my supervisory committee members, Prof. Frank Ko and Prof. Rizhi Wang, for supporting and guiding my research. I wish to extend my appreciation to Dr. Heejae Yang for his valuable suggestions for this project. I could not complete this dissertation without the support and help from Prof. Donald Brunette and Ms. Hai-sle Moon in the Department of Oral Biological and Medical Science. Mr. Clifford Kim also gave me full support to make me approach the final steps towards graduation.    I also owe thanks to the other graduate students whom I spent years together in the laboratories, Dr. Shuxin Zhou, Dr. Azadeh Goudarzi, Dr. Hamidreza Zargar, Dr. Ali Shafeie and Dr. Victor Leung, and to the staffs in the Department of Materials Engineering and the Department of Oral Biological and Medical Science for all their help. The last but not least, I would like to acknowledge the financial support for this study by the Natural Science and Engineering Council of Canada (Discovery Projects “Process engineering for smart bioceramics” NSERC RGPIN 121269-09).         xxi  Dedication  I do not believe I could have completed this research without the love and support of my family. This dissertation is dedicated to my wife, Sunghee Kwak, and my children, Kyoungjoo and Soomin. You were always there for me, and it has made all the difference. Last but not least, this dissertation could not exist without the sacrifice of my parents, Sangmuk Chae and Soonrae Jeong, and parents-in law, Pyongchi Kwak and Soonjung Park.                                                                                                                                                                          1  Chapter 1 Introduction and Rationale  The acute need for hard tissue substitute materials due to limited supplies of autografts and possibility of disease transmission and anti-immunization response from allografts and xenografts is major motive for research in hard tissue reconstruction/regeneration. During the last several decades, biomaterials (synthetic or natural materials suitable for use in constructing and/or replacing defected tissues) have been successfully used, but not induced complete regeneration of the tissues [1]. Tissue regenerative engineering has received a lot of attention in recent years as a new methodology to search for solutions to restore damaged tissue functions. It is a multidisciplinary filed of science and engineering incorporating materials science, biomedical engineering, cell biology, and advanced stem cell science [2, 3]. Successful tissue regeneration necessitates the design of an advanced smart biomaterial as a temporary extra-cellular matrix that is capable of inducing host cells to assume highly specialized functions [3].   Lessons learned from the structure and compositions of bone tissue lead to novel designs of calcium phosphate/biopolymer nanocomposites with sufficient biological and mechanical properties, mimicking natural bone tissue [1‒3]. A hierarchically structured nanocomposite of bone tissue is basically composed of mineralized collagen fibrils. The biologically "engineered" biomineralization process stimulates amorphous calcium phosphate and carbonated hydroxyapatite (HAp) nanocrystals to nucleate in the spaces between collagen fibrils and to grow over parallel to the fibrils [4]. Bioceramic calcium phosphates (CaP) have been used as a main component for bone reconstruction and/or regeneration [5]. HAp, especially in poorly crystalline form, is the closest inorganic phase to bone mineral [4]. As HAp has high biocompatibility, bioactivity and osteoconductivity, it has been successfully used in the forms of granules, dense or porous blocks, cements, and coatings [5, 6]. HAp is rather stable in biological environment, and its resorption characteristics depend on the level of crystalline disorder [6, 7]. Recently, dicalcium phosphate anhydrate (DCPA) and  2  dicalcium phosphate dihydrate (DCPD) also have received attention due to their favorable osteoinductive and bioresorbable characteristics [8]. Dissolution (i.e. resorption) of hard tissue implants is required for allowing simultaneous replacement with new bone tissues, and finally obtaining complete repair without remnants of the artificial substitutes. Osteoinductivity is another key characteristic of such materials as it plays a critical role in curing large size bone defects [8‒10].    Nanometer scale fibrous materials inspired by the fundamental building blocks of human tissues have been designed for various biomedical and medicine applications [11]. Electrospinning (ES) is one of the simplest and cost-effective methodologies used to fabricate nanofibrous structures by applying an electrostatic field to polymeric solutions. It produces non-woven nanofibers with 3D interconnected pores and a high surface area that mimic the natural extra-cellular matrix (ECM) of human tissues, and it is of great interest in tissue regenerative medicines [11‒13]. Various types of biopolymers have been electrospun into multi-functional scaffolds from natural polymers of alginate [14], chitosan [15], silk [16] to synthetic polymers of poly(lactic acid) (PLA) [17], poly(glycolic acid) (PGA) [18] , poly(lactic-co-glycolic acid) (PLGA) [12], and poly(ε-carprotacton) (PCL) [19] . It has been demonstrated that they can act as tissue engineering scaffolds where cells adhere and proliferate for the regeneration of new functional host tissues [2] . They can also be loaded with pharmaceutical agents including antibiotics, anticancer drugs, proteins, and DNA for subsequent controlled targeted release and delivery into the surgery sites for more effective curing process [11]. At the same time, these types of novel fibrous biomaterials can provide bacterial protection, thermal insulation, and mechanical barrier as a wound healing dresser [11, 13, 17].   There are initial indications that the incorporation of CaP into the biopolymer nanofibers could make electrospinning more attractive for use in the preparation of organic/inorganic nanocomposites in hard tissue engineering applications. Electrospinning of mechanically  3  blended polymer solution with pre-processed CaP nanoparticles has been used in composite fabrication [20−22, 246, 247]. Li et al. [20] produced silk fibroin fibrous scaffolds containing bone morphogenetic protein-2 (BMP-2) and HAp nanoparticles (Berkley Advanced Biomaterials Inc., USA), which generated in vitro bone formation from human bone marrow-derived mesenchymal stem cells (hMSC). Cotton wool-like PLGA fibers, including amorphous tricalcium phosphate nanoparticles (prepared by flame spray pyrolysis), demonstrated bioactivity and osteogenic differentiation from hMSC [21]. Nie and Wang [22] studied the release of BMP-2 plasmid DNA from an electrospun PLGA blend with HAp nanoparticles (Berkley Advanced Biomaterials Inc., USA). Although these previously studied CaP/biopolymer composite fibrous scaffolds demonstrated better biological performance than the pure biopolymers, the mechanical blending caused severe agglomeration of the CaP nanoparticles and, consequently, irregular distribution of the aggregates within/on the fibers. The nanocomposite systems need to be improved with controlled and uniform distribution of the CaP nanophases for homogeneous nanocomposite fibers, such as the mineralized collagen fibrils of bone tissue. Sol-gel technique and biomineralization in the simulated body fluid (SBF) solution were also applied with ES for processing of CaP/biopolymer composite fibrous scaffolds. CaP-based sol, prepared in water-soluble polymer solutions, was electrospun into fibrous mats, but the firing step necessary for the synthesis of the CaP inorganic phase gave rise to losing mechanical stability of the fibrous matrix [23, 24].   In this study, we propose a novel process for the development of calcium phospahte/biopolymer nanocomposite fibrous scaffolds via biomimetic in-situ synthesis and electrospinning, including i) DCPA/PLA and ii) HAp/aginate systems. Electrospinning of in-situ synthesized CaP within a synthetic biopolymer solution has been selected for the first time in the DCPA/PLA system. HAp/alginate nanocomposite scaffolds through biomimetic nucleation and growth of HAp nanocrystals on electrospun alginate nanofibers are also original systems disclosed only in this work. It was postulated that the in-situ  4  synthesis of CaP within the biopolymeric matrix can induce improved dispersion and distribution of the inorganic phase by controlling the chemical interactions with the organic matrix, than a simple blending method. The inorganic phases are expected to increase biocompatibility, bioactivity and bioconductivity, and enhance structural stability of the nanocomposite fibrous scaffolds for bone tissue regenerative engineering. PLA has been selected as it is one of the Food and Drug Administration (FDA) approved synthetic polyesters. It is biocompatible, biodegradable, and thermoplastic. Its fibrous scaffolds have shown promising results for variety of medical applications, such as culturing mouse neutral stem cells [25] and human fibroblast cells [26], delivering antibacterial silver nanoparticles [27] and tetracycline hydrochloride [28] and biosensing E.coli [29]. Electrospun PLA nanofibers can make the degradation products of lactic acid and oligomers quickly escape from the surface to neighboring pores, and eventually out of the scaffold, so that the possibility of adverse tissue response from local acidicity is minimized. Drug release periods can also be extended due to the slower degradation rate resulting from minimized autocatalytic hydrolysis [30, 31]. Alginate, a natural polysaccharide derived from brown algae, has been previously studied for drug delivery [33], tissue engineering scaffolds for skin [34], cartilage [35] and bone [36], and wound dressing [37]. Such interest in alginate is attributed to its chemical structure, which is composed of (1-4) linked β-D-mannuronic acid (M unit) and α-L-guluronic acid (G unit). These units are sequentially distributed (MM, GG, and MG blocks) along the molecular chains [14, 35]. This structure resembles glycosaminoglycan (GAG), one of the major components of the natural ECM in human tissue [35]. It also has favorable properties of natural biopolymers over synthesized biodegradable polymers such as non-toxicity, biocompatibility, hydrophilicity and relatively low cost for production [32, 33]. Alginate forms a stable hydrogel in the presence of low concentrations of divalent cations, such as Ca2+, Sr2+, and Ba2+, through ionic interactions with the carboxylic functional groups on G units in the alginate molecular chains [38].   5  Chapter 2 Literature Review  2.1 Bone tissue  Bone is one of the specialized connective tissues, providing the five main functions: mechanically supporting the body, protecting internal organs, aiding body movements, producing blood cells, and storing fat and minerals[41]. It is a hierarchically structured nanocomposite with relatively high strength and toughness. Typical bone has tensile strength of 50 − 150 MPa, compressive strength of 130 ‒ 180MPa, and elastic modulus of 12 ‒ 18 GPa [6, 39, 40]. The crystallinity, porosity, and composition of bone adjust to the dynamic biological and biomechanical environments. This makes the properties of bone vary from location to location and with time. Bone is the tissue that can undergo spontaneous regeneration and remodel its structure. It is accomplished through a delicate balance between the osteogenic (bone forming) and osteoclastic (bone removing) processes. Bone tissue is constantly being created and destroyed to insure optimal bone density in any particular location within the body. It can adapt to static and dynamic stresses with self-sensing ability. If excessive stress (i.e. more than the normal physiological condition) is applied, more osteogenic activity happens. Conversely, if less stress is applied, the balance tilts towards osteoclastic activity. Bone loss is observed in people unable to use their limbs e.g. when immobilized due to accident or disease. Fortunately, bone can also self-heal and self-recycle [4, 41, 42].  2.1.1 Structure and composition of bone Bones are classified as long (femur and humerus), short (ankle and wrist bones), flat (sternum andskull), or irregular (vertebral, pelvis, and fitting none of the other categories). Figure 2.1 shows the 7 hierarchical structure of bone. Macroscopically, bone tissue is divided into spongy (cancellous) and compact (cortical) bone. Porous spongy bone is located at the ends of long bones and the core of all bones. Compact bone forms the shaft and outer   6   Figure 2. 1 The 7 hierarchical levels of bone tissue from nano to macro-scales [4].   covering of bones. Figure 2.2 shows the microscopic view of bone tissue. Cylindrical columns oriented parallel to the long axis of bone and surrounding a central cavity are called osteons. The cavity forms pathways for blood vessels, nerves, and nutrients. The hole is called Haversian canal surrounded by concentric rings termed lamellae. The lamellae are filled with numerous dark spots, called lacunae, and fine dark lines, termed canaliculi. The   7    Figure 2.2 Hierarchical structural organization of bone: (a) cortical and cancellation bone; osteons with Haversian systems; (c) lamellae; (d) fiber assemblies of collagen fibrils; (e) bone mineral crystals, collagen molecules, and non-collagenous proteins [39].   interior of bone is filled with areolar connective tissue, blood  vessels, and  bone marrow producing blood cells. Lamellae are composed of collagen fibers which are bundles of mineralized collagen fibril in nano-scales. The mineralized collagen fibril is the basic building block of bone tissue. Type I collagen of two α (I) and one α (II) molecule chains make up a triple-helix to form a cylinder of 300 nm x 1.5 nm collagen fibril. The collagen fibril is mineralized predominantly with a carbonated hydroxyapatite (Ca10(PO4, CO3)6(OH)2), called dahllite, with dimensions of ~ 50 nm x 25 nm x 2.5 nm. The dahllite crystals form in the holes between collagen fibrils and grow over parallel to the fibrils [41, 42].     8  Depending on age, location, and, function of bone tissue, bone minerals are a mixture of carbonated hydroxyapatite and amorphous calcium phosphate with different ratios. From macroscopic bone to the mineralized collagen fibril, bone tissue is the hierarchically structured nanocomposite, composed of ~ 70 wt % of calcium phosphates, ~ 20 wt % of collagen proteins, and ~ 10 wt % of water and non-collagen proteins like proteoglycans and glycoproteins [4]. Collagen fibrous network provides structural support to cells. Bone cells, mainly osteoblasts, osteocytes, and osteoclasts, are present in the lacunae. Osteoblasts are bone forming cells derived from either mesenchymal stem cells or bone marrow stromal cells. Osteocytes are matured osteoblasts for maintaining the mineralized tissue with limited bone forming and resorption and for providing nutrition to bone. Osteoclasts are multi-nucleated giant cells derived from hematopoietic stem cells found in circulating blood. They are the cells responsible for bone resorption by synthesizing hydrochloric acid, collagenase, and enzymes that dissolves the bone minerals and the collagen matrix [43].  2.1.2 Healing and repair of bone Bone defects result from many different reasons, such as trauma, birth defects, cancer, osteoporosis, and various orthopaedic surgeries. If the rate of osteoclastic activity that dissolve bone tissue exceeds the rate at which osteoblasts create bone tissue, bone density will decrease and result in weaker, brittle, and vulnerable to bone fracture. This is known as osteoporosis [44]. Bone fracture becomes a common injury due to the modern high-speed transportation, active recreation, and due to multiple processes related to general aging of the population. If bone is only slightly fractured, it is typically immobilized superficially by a cast or splint and allowed to self-heal. Figure 2.3 illustrates the healing stages of bone fracture. Soon after a fracture occurs, the blood vessels within the bone rupture, creating a blood clot (fracture hematoma) in the fracture site within 6 to 8 hours. Localized swelling and inflammation follows. Macrophages and inflammatory leukocytes move into the injury sites and produce the pro-inflammatory agents that initiate healings (Figure 2.3 (a)). Inflammation triggers cell division and the growth of new blood vessels. Chondrocytes   9    Figure 2.3 The healing stages of a bone fracture: (a) fracture hematoma formation and inflammation, (b) soft callus formation, (c) hard callus formation, and (d) remodeling [47].   secrete collagen and proteoglycans, creating fibro-cartilage fills in the gaps between the fractured bones and forming bony soft callus (Figure 2.3 (b)). Over the several weeks, osteoclasts resorb the dead bone, and osteogenic cells become active and differentiate into osteoblasts. The soft bony callus is then converted into hard  bony callus by the action of endochondral ossification (Figure 2.3 (c)).  Remodeling of the fracture site begins gradually in order to restore the original internal structure and shape (Figure 2.3 (d)). The last stage is much longer than the previous ones [42, 47].  If the fracture is very severe, it must be treated surgically by repositioning the bones and fixing them in place with the use of plates, screws, and pins. These procedures require hospitalization and sometimes follow-up surgery to remove the hardware, accompanied with additional pain and cost. Most such procedures are successful, but there are bacterial or fungal infections reported in 10 to 15 % of all surgeries performed [45]. These must be treated with further surgery to remove the dead bone and treatment with high amounts of antibiotics to treat the infecions [46].    10  2.2 Biomaterials for bone tissue The demand for bone tissue treatments to cure trauma or diseases has increased with the extended life span of humans. Autograft harvested from a patient is an ideal bone graft because it possesses all the characteristic necessary for new bone growth, such as osteoconductivity (supporting the attachment of osteoblasts), osteogenicity (producing bone tissue by cells), and osteoinductivity (inducing non-differentiated stem cells or osteoprogenitor cells to differentiate into osteoblasts) [1, 3, 7]. However, harvesting the autograft requires an additional surgery which causes extra pain and cost of a patient. Furthermore, there is the limitation of tissue supply. Allograft transferred from donors is another option to reconstruct bone defects. Although it can avoid donor site morbidity and limited supply of autograft, there is a risk of disease transmission from a donor to a recipient [7, 50]. To minimize the risk, allograft tissue is treated through tissue freezing and drying, sterilization via gamma radiation, electron beam radiation, and ethylene oxide, but the risk cannot be completely eliminated. It has been reported that Hepatitis B and C transmissions occurred in 1990s, and septic arthritis and Clostridium Sordelli infections in early 2000s [51]. The sterilization methods can also alter mechanical and biological properties of the allograft tissue.   Due to the limitation and disadvantages of autograft and allograft, intensive research has been undertaken to find suitable biomaterials with sufficient biological and mechanical properties comparable to natural bone tissues. Biomaterials are synthetic or natural materials designed to replace partially or fully, permanently or temporarily the living tissues. From the materials science point of view, the biomaterials are generally categorized into metallic, polymeric, ceramic, and composite biomaterials [52].      11  2.2.1 Bioceramic calcium phosphates The word “ceramics” comes from the Greek word “Keramos” meaning “Pottery,” or “Potter’s Clay.” Ceramics can be defined as “inorganic non-metallic” materials [53]. Ceramics have relatively high elastic modulus, melting temperature, and chemical resistance compared to metals and polymers, due to the prevalent ionic and covalent bonding. On the other hand, low fracture toughness is the unavoidable disadvantage of ceramics [54]. The imperfections such as micro-pores or cracks create stress concentration during loading, leading to breakdown of ceramic body rather than more desirable plastic deformation [54]. Bioceramics are generally defined as ceramic materials used for the repair, reconstruction, and replacement of diseased or damaged hard tissues [54]. Regarding the degree of an implant-tissue interaction, bioceramics are broadly categorized into bioinert, bioactive, and bioresorbable [55]. Nearly bioinert ceramics undergo little or no chemical change during long-term exposure to the physiological environment and develop a few µm thick fibrous layers at the interface. Alumina and Zirconia used in load-bearing hip and knee joint replacement and dental implant are the representative examples of the bioinert ceramics [55]. Bioactive glass and glass-ceramics, such as Bioglass®, Cervital® glass-ceramics, Apatite/Wallastonite (A/W) glass-ceramics, are the representative examples of bioactive ceramics. Glass-ceramics have amorphous glass and crystalline ceramic properties, processed by "controlled crystalllization" [240]. Bioactive ceramics stimulate a specific biological reaction at the interface, which results in the formation of bond between the implant and the soft or hard tissues. This interface has been referred to as the ‘bonding zone’ consisting of mineralized organic mesh network [54]. Calcium phosphates (CaP) are not only bioactive but also bioresorbable. After implantation into a body, bioresorbable ceramics degrade biochemically by the combined action of body fluids and macrophages. The resorbed area is replaced by surrounding endogenous tissues. These characteristics make the bioresorbable ceramics a prime candidate for temporary bone fillers, bone tissue engineering scaffolds hybridized with biodegradable polymers, and drug delivery applications [54, 56, 57]. Table 2.1 shows acronyms, formulas, Ca/P ratio, and solubility products for various    12  bioceramic calcium phosphates. While a number of variables have an effect on the biodegradation of calcium phosphates, the general order of solubility at pH of around 7 is as follows:    ACP > DCPD ≈ DCPA > TTCP > OCP > TCP > HAp   Table 2.1 Acronyms, formulas, Ca/P ratios, and solubility products of representative calcium phosphates.   Acronym Formula Ca/P Solubility product  Ksp Tetracalcium phosphate   TTCP  Ca4P2O9 2.00             ‒ Hydroxyapatite   HAp  Ca10(PO4)6(OH)2 1.67  5.5 × 10‒118 M [62] Amorphous calcium phosphate   CaxHy(PO4)z·nH2O  n=3‒4.5; 15‒20% H2O ‒             ‒ Tricalcium phosphate (α)   α-TCP   α-Ca3(PO4)2 1.50  2.8 × 10‒29 M [60] Tricalcium phosphate (β)   β-TCP   β-Ca3(PO4)2 1.50  2.5 × 10‒30 M [60] Octacalcium phosphate   OCP  Ca8H2(PO4)6·5H2O 1.33  2.5 × 10‒99 M [61] Dicalcium phosphate anhydrate   DCPA  CaHPO4 1.00  9.2 × 10‒7 M [59] Dicalcium phosphate dihydrate   DCPD CaHPO4·2H2O 1.00  1.9 × 10‒7 M [59]   Stability of various CaP is illustrated as a function of pH in Figure 2.2-1 [58]. HAp is relatively insoluble and the most stable phase at pH above 4.0. For pH lower than 4.0, DCPD is more stable than any other. Through dissolution and re-precipitation process, unstable phases of CaP transform into stable phases at a given pH.  Normal physiological environment keep pH 7.2, but it decreases around the tissue injury sites or inflammation [58]. Such changes of pH may affect stability of calcium phosphates. For example, accelerated dissolution of HAp coatings on dental implants has been observed for cases where improper implantation procedure triggered local inflammation and pH decrease.   13    Figure 2.4 Solubility isotherms of calcium phosphate phases at 37 °C [58]. TCa and Tp are the total molar concentrations of calcium and phosphate ions, respectively.    2.2.1.1 Hydroxyapatite Human bone and teeth contain calcium and phosphorus mostly in the form of partially or fully crystalline carbonated hydroxyapatite and fluorapatite [63]. Hydroxyapatite (HAp) has the chemical formula of Ca10(PO4)6(OH)2. It belongs to the hexagonal group with lattice constants a = 0.942 nm and c = 0.688 nm. The atomic ratio of Ca/P is 1.67, and the density is 3.163 g/cm3. After Levitt et al. [64] suggested a preparation method of HAp from mineral fluorapatite in 1969, there have been many efforts to synthesize and commercialize  14  HAp. Nowadays, HAp powder is usually synthesized in one of the following four processes: (1) precipitation, (2) hydrolysis, (3) solid-state reaction, and (4) hydrothermal reaction [63]. Due to its composition similar to bone mineral, synthetic HAp is more biocompatible and osteoconductive than any metal, polymer and even other ceramics. With such biochemical properties, it has been the most feasible bone substitute biomaterial. HAp allows ingrowth of new bone and promotes adhesion of matrix-producing cells and organic molecules, forming a 3 – 5 μm thick amorphous zone with high concentration of phosphate and calcium ions. This is the area where bone mineral crystals form, and as this region matures, the ‘bonding zone’ between the HAp implant and natural bone tissue shrinks to 50 – 200 nm thickness providing the high interfacial strength and thus load transfer capabilities [65].   The mechanical properties of synthetic HAp ceramics depend on crystallinity, grain and defects size, additives, porosity, etc. In general, the mechanical properties decrease significantly with increasing amount of mircoporosity and grain size. High crystallinity, low porosity and small grain size lead to higher elastic modulus, strength, and toughness of HAp. It has been reported that the flextural strength and fracture toughness of dense HAp are much lower in dry conditions than wet [66]. Comparing the mechanical properties of synthetic HAp ceramics with those of human bone shows that dense HAp has a reasonably good compressive strength but significantly lower fracture toughness than cortical bone, and the overall properties of porous HAp are clearly poorer than those of cancellous bone [39, 43, 133]. Hence, HAp ceramics cannot be used as a stand-alone biomaterial for load bearing applications. HAp ceramics have therefore been used in orthopedics and dentistry over the last two decades in the forms of powders, coatings, dense or porous solid blocks, and injectable pastes. Examples include tooth root replacement, augmentation of alveolar ridge, maxillofacial reconstructions, coating for dental and orthopedic metal implants, and pulp capping [54].    15  2.2.1.2 Dicalcium phosphate anyhydrate Among calcium phosphates, dicalcium phosphate anhydrate (DCPA; CaHPO4), also known as monetite, is also bioactive, bioresorbable, and above all osteoinductive. DCPA is a stable phase under acidic conditions (pH < 4.8), and dissolves and/or hydrolyzes to HAp depending on implanted sites and physiological conditions [67]. It is also found in small amounts in urinary and dental stones [68]. Triclinic crystallographic structure of DCPA has the following lattice parameters: a = 6.910, b = 6.627, c = 6.998 Å, α =96.34°, β = 103.82° and γ = 88.33° with density of 2.92 g/cm3 [69]. DCPA nanoparticles with various morphologies such as spheres, nanofibers, and bundles of nanowires were obtained in cetyltrimethylammonium bromide (CTAB) reverse micelle solution by precipitation from CaCl2 and (NH4)2HPO4 at pH 3.0 [68]. Monetite whiskers have been synthesized by hydrothermal treatment of monocalcium phosphate monohydrate suspension at 160 ºC [70].   Dicalcium phosphate dihydrate (DCPD) transforms (by losing its crystal water) into DCPA on heating at above 110 ºC [71]. In the in-vivo animal study by Habibovic et al., [8] 3D printed DCPA and DCPD cements implanted at intramuscular sites in goats led to ectopic immature and woven bone formation induced by the supersaturated local environments of Ca2+ and PO43– ions  formed by dissolution of the implants surface after 6 – 9 weeks, indicating the osteoinductive properties of the DCPA and DCPD cements. It has been demonstrated that osteoinductive biomaterials stimulate more bone formation when used as tissue engineered constructs in goat muscles [72] and perform better in clinically critical size orthopedic defects in goats [9]. Although details of osteoinduction mechanisms remain unclear, it has led to the recognition of several biomaterial parameters that can be tuned in order to guide the biological response. DCPA and DCPD have been successfully used as starting materials in the preparation of the powder components of self-hardening calcium phosphate cements [73‒75]. DCPA or DCPD cements recently receive more attention due to their superior in-vivo bioresorbability than the conventional apatite cements [76, 77]. Biodegradation of hard tissue implants should also be considered to allow simultaneous replacement with new bone tissues and obtain complete repair without remnants of the  16  artificial substitutes. Dicalcium phosphates are one of the fastest resorbable CaP materials with the resorption rate of DCPD at a pH of 5.5 being about three times and three hundred times faster than that of tricalcium phosphate and carbonated HAp, respectively [10].   2.2.2 Biopolymers Biopolymers are generally categorized as natural and synthetic, linear and cross-linked, and degradable and non-degradable [78]. Collagen, gelatin, chitosan, and alginate are the examples of natural biopolymers. Nylon, polyethylene, polypropylene, and polyacrylate are non-degradable synthetic polymers. Poly(lactic acid) (PLA), poly(glycolic acid) (PGA), and poly(lactic-co-glycolic acid) (PLGA) are representative linear degradable synthetic polymers [78]. The main advantages of the biopolymers, compared to metallic and ceramic biomaterials, are ease of manufacturing to produce desired shapes (latex, film, sheet, fibers, etc.), ease of secondary processing, reasonably low cost, and availability with specified (desired) mechanical and physical properties [7, 78]. The synthetic biodegradable polymers have widely been used in medical disposable supplies, prosthetic materials, implants, wound dressing, drug delivery, and tissue engineering. A wide range of degradation rates can be achieved through varying the monomer ratio and crystallinity of the polymers. Advantageous attributes of natural polymers include their hydrophilicity, non-toxicity, less immune reactions, as well as enhanced cell adhesion and proliferation [78].   2.2.2.1 Poly(lactic acid) Poly(lactic acid) (PLA) has generated great interest as one of the most innovative materials for a wide range of applications from packaging, agricultural products, disposable materials, and sport wear. It is a biocompatible, biodegradable, and thermoplastic polymer, which makes it highly attractive for biological and medical applications [17]. PLA breaks down into monomeric components in physiological environments by enzyme reactions, hydrolytic cleavage of ester bonds, and also by exposure to aerobic or anaerobic microorganisms. The degradation product is lactic acid, which is metabolically innocuous, but local acidity due to  17  abrupt release of the acidic degradation products may cause adverse tissue response [79, 80]. Thicker samples can lead to heterogeneous degradation which may be ascribed to the following two phenomena; (1)  easier diffusion of soluble oligomers from the surface into the external medium than from inside (2)  neutralization of carboxylic end groups located at the surface by the external solution.  These reactions reduce acidity at the surface, but degradation rate in the bulk is enhanced by autocatalysis [6]. The glass transition temperature of PLA is 55 – 65 ºC and melting temperature 170 – 180 ºC. Owing to its thermoplastic nature, it is possible to easily shape the polymer for implants and other surgical applications. PLA has been approved by the Federal Drug Administration (FDA) for the use as a suture material [93]. PLA has compressive strength of 35 – 150 MPa, tensile strength of 25 – 50 MPa, and elastic modulus of 1.2 – 3.0 GPa, similar range to those of cancellous bone tissue [234]. It has already shown favorable results in the fixation of bone fractures and osteotomies [81‒83]. Tissue engineering is the most recent application area where PLA was found to promote the regeneration and reconstruction of human organs. PLA also shows excellent biocompatibility and good osteoconductivity, applicable for bone tissue reconstruction and regeneration [6, 86]. Drugs such as growth factors, antibiotics, or thrombin inhibitors combined with the polymer can be released locally during its hydrolytic degradation and/or morphological changes [84, 85].   PLA was first synthesized by Carothers in 1932 as a low molecular weight product by heating lactic acid in vacuum [17]. Dupont was awarded a patent for a process to produce high molecular weight PLA. Lactic acid was produced prevalently by petrochemical route until 1990. Today, corn starch is converted into lactic acid by bacterial fermentation using an optimized strain of Lactobacillus [17]. Polymerization of lactic acid to high molecular weight can be achieved in two ways as illustrated in Figure 2.5; (1) direct condensation and  18  (2) formation of a cyclic dimmer intermediate. In direct condensation, solvent is used under high vacuum and temperature for the removal of water produced in the condensation, which lead to achieve low to intermediate molecular weight PLA. In the solvent free process, a cyclic intermediate dimer, referred to as lactide, is produced and purified by distillation. Catalytic ring-opening polymerization of the lactide results in PLA with controlled molecular weight [17, 87].       Figure 2.5 Synthesis of poly(lactic acid) [17].      19  2.2.2.2 Alginate Alginate is an anionic natural polysaccharides, prevalently derived from marine brown algae. Sodium alginate is the sodium salt of alginic acid (NaC6H7O6) extracted from the cell walls of the algae. Potasium alginate is the potasium salt of alginic acid (KC6H7O6) from seaweed.  Alginate quickly absorbs water and forms viscous substance when binding with water [118]. It is also produced by two bacterial genera, Pseudomonas and Azotobacter. Bacterial, which are useful for the production of micro or nanostructures [119]. Alginate has been extensively  studied for biomedical engineering applications such as a tissue engineering scaffold for skin [88], cartilage [14, 35], bone [36], liver [89] and cardiac tissue [90], wound dressing [91], and drug delivery carriers [96]. Such interest in alginate is attributed to its unique chemical structure, which is composed of (1-4) linked β-D-mannuronic acid (M unit) and α-L- guluronic acid (G unit). These units are sequentially distributed (MM, GG, and MG blocks)      Figure 2.6 Structure of alginic acid extracted from Lessonia trabeculata marine algae; (a) D-mannuronic residue: M, (b) L-guluronic residue: G, and (c) polysaccharide chains of MM, GG and MG blocks [95].  20  along the molecular chains (Figure 2.6). This structure resembles glycosaminoglycan (GAG),  one of the major components of the natural ECM in human tissue [14]. Alginate forms stable hydrogels in the presence of divalent cations, Ca2+, Sr2+, and Ba2+, at low concentrations through the ionic interaction between the cation and carboxylic functional groups on G units located on the alginate molecular chain [82]. According to the model presented by Grant et al. [92], the divalent cations bridge the negatively charged GG blocks and form an egg-box structure, which encloses the cations and creates "junction zones" for cross-linking of alginate, as shown in Figure 2.7.      Figure 2.7 The egg-box model for alginate gellation with calcium ions. Guluronic acid (G) blocks of alginate are held together by a number of calcium ions [94].         21  2.2.3 Calcium phosphate/biopolymer composite scaffolds Advantageous properties of two or more types of materials can be combined to best address mechanical and physiological demands of the natural bone tissue. By taking advantages of compliant biopolymers and stiff bioceramic calcium phosphates (CaP), mechanically reinforced composites can be achieved with enhanced biological capability. Poor bioactivity and osteoconductivity of biopolymers can be counteracted by increasing volume fraction and higher surface area to volume ratio of the CaP inclusions [241, 242]. The morphology and size of the inorganic particles dispersed in polymers has strong influence on the final properties of the reinforced composite with strong emphasis on uniform dispersion of nano particles. For example, 3.0 wt % of exfoliated clay nanoparticles have been demonstrated to increase the flextural modulus of PLA by 20 % through in-situ synthesis technique [134]. Addition of the bioactive inorganic phases to biodegradable polymers can also alter the degradation behavior by allowing rapid exchange of protons in water for alkali, providing a pH buffering effect at the polymer surface and modifying the acidic polymer degradation [98–100]. The inclusion of bioceramic CaP has been shown to modify surface and bulk properties of composite scaffolds by increasing the hydrophilicity and water absorption of hydrophobic polymer matrix, thus altering the scaffold degradation kinetics. It has been reported that PLA composites filled with HAp particles hydrolyzed homogeneously due to water penetrating the interfacial regions [101].  There are numerous forming techniques to obtain 3D highly porous structures. Figure 2.8 shows typical morphologies of 3D composite scaffolds, as compared to cancellous bone structure, and Table 2.2-2 compares advantages and disadvantages of the fabrication techniques leading to the composite scaffolds. Thermally induced phase separation (TIPS) quenches and freezes polymer solution with inorganic phase powder. The organic solvent is then sublimated until the scaffold reaches a constant weight [102–104]. Solvent casting/particle leaching involves the dissolution of the polymer in organic solvents, mixing with inorganic second phase powder and leaching the particles, and casting the solution into a mould. Subsequently, the solvent is evaporated and the particles are washed-out with   22    Figure 2.8 Typical morphologies of 3D porous scaffolds produced by different techniques and structure of cancelleous bone. (a) thermally induced phase separation [104], (b) solvent casting and particles leaching [107], (c) solid free-form [109], (d) microsphere sintering [110], and (e) cancellous bone [111].      23  Table 2.2 Fabrication techniques for 3D porous composite scaffolds and their advantages and disadvantages.  Fabrication technique Advantages Disadvantages Ref. Thermally induced  phase separation  (TIPS)   High porosities (~95%) Highly interconnected pore structures Anisotropic and tubular pores possible Control of structure and pore size  by varying preparation conditions  Long time to sublime solvent  Shrinkage issues Small scale production Use of organic solvents [101–104]     Solvent casting/ particles leaching  Controlled porosity Controlled interconnectivity (if  particles are sintered) Generally isotropic structure Use of organic solvents [105–107]     Solid free-form  Porous structure can be tailored to  host tissue Protein and cell encapsulation possible Good interface with medical imaging Resolution needs to be improved  to the micro-scale Some methods use organic solvents [108, 109]     Microsphere sintering  Graded porosity structures possible Controlled porosity Can be fabricated into complex shapes Interconnectivity is an issue Use of organic solvents [110]   proper solvents [105–107]. Solid free form (SFF) process, also known as rapid prototyping, uses computer tomography and magnetic resonance imaging, for the accurate design of scaffold structure. Most recently, construction of the scaffold is often done using 3D printing technology. SFF fabrication is used to develop scaffolds with controlled micro and macroporous structures [108, 109]. In microsphere sintering, microspheres of a ceramic/polymer composite are synthesized first by using emulsion/solvent evaporation technique, and sintering the composite microspheres yields a 3D porous scaffold [110].The mechanical properties of currently available porous scaffolds are generally inferior to the relevant properties of bone, as shown in Figure 2.9. It is seen, however, that some dense polymers may match cancellous bone properties and approach cortical bone properties. The dense bioactive ceramics are close to the properties of cortical bone as well. Porous scaffolds are at least one order of magnitude weaker than cancellous bone and orders of magnitude weaker than the cortical bone.     24    Figure 2.9 Elastic modulus vs. compressive strength of biodegradable polymers, bioactive ceramics, and composites [6].    2.3 Electrospinninng  Electrospinning (ES) is a simple and efficient process that can produce polymer fibers ranging from nanometers to micrometers using an electrostatic forces. In 1934, the first patent regarding electrospinning was disclosed by Formhals [126], but a variety of its applications from biomedical, textile to electrical/electronic engineering have emerged since 1990s [120‒122]. Figure 2.10 schematically illustrates an eletrospinning unit. The electrospinning system basically consists of a spinneret (practically syringe and needle) containing solution, a conductive ground collector placed a distance from the spinneret, and a power supply which is used to apply a electrical potential between the spinneret and collector. A drop of polymer solution forms at the tip of the spinneret due to gavity and is held in place by surface tension. As the applied voltage increases a droplet of solution at  25  the tip of the spinneret will charge, and the repulsion of charges accumulating on the droplet surface overcomes the surface tension of the droplet forming a conical shape. When the electrical potential reaches a critical value, a charged fluid jet is emitted from the droplet. Geoffrey Taylor analyzed the formation of cone and jet emission under an electrical field, and proposed the mathematical model, known as "Taylor cone" [229, 243];                                                                                              2         H2        2L                                    VC =  4         ln         − 1.5  0.117πγR                                                   (Eq.2.3-1)                                   L2         R  where Vc: critical voltage for electrospinning jet formation, H: distance between spinneret capillary and collector, L: length of capillary, R: radius of capillary, and γ: surface tension of solution.       Figure 2.10 Schematic illustration of electrospinning system and process.   26  The ES jet follows a straight path along the electric field direction at the early stage, then undergoes a rapid bending instability, and follows a spiraling path during passage towards the collector, which results in extensive stretching of the jet [11]. The stretching process is accompanied by a rapid evaporation of the solvent, which also leads to a reduction in the diameter of the jet and increase in surface area of  the jet, resulting in the formation of fibers in a nanometer scale. The dried fibers are typically deposited with random orientation as a non-woven mat on the collector, as shown in Figure 2.11, but can be aligned with accessories of electrospinning unit. In general, low molecular weight polymers have a higher tendency to form droplets, while higher molecular weight polymer with entanglements of chains tend to form fibers. The ultra-fine diameter of electrospun fibers results from enormous elongation. The high rate of stretching also produces a high degree of molecular orientation, which makes electrospun nanofibers birefringent [122‒125].     Figure 2.11 SEM micrograph of electrospun poly(vinyl alcohol) nanofibrous scaffold.     27  During electrohydrodynamic jet-emission in ES process, bending and stretching instabilities can cause the jet to either break into droplets (electrospraying) or promote the formation of fibers. The different resulting morphology is determined by the balance between surface tension and axial stress generated by capillary, viscoelastic, and electrostatic forces. The morphology of nanofibers depends on the complex interaction between the electric field (stretching the jet axially), surface tension (producing droplets to minimize surface area), and the rheology of the solution. In 1971, Baumgarten highlighted the effects of solution viscosity, surrounding gas, flow rate, etc on the fiber diameter and jet length [132]. It was found that as the solution viscosity increases the fiber diameter increases. The relationship between fiber diameter and solution viscosity was empirically established by the equation as following;                          d = η·0.5                                                                                          (Eq. 2.3-2) where d: fiber diameter and η: solution viscosity in poise.  In 2003, Fridrikh et al. [127] developed a model to predict the diameter of electrospun fibers as a function of solution properties,  such as conductivity, dielectric properties, surface tension, and processing parameters, such as solution feeding rate and electric current;                                                                     1                                     Q2           2              3                      d =    γε                                                                                               (Eq.2.3-3)                                      I2    π(2ln x − 3)  where d: fiber diameter, γ: surface tension of solution, ε: dielectric permittivity, Q: solution feeding rate, I: electric current, and x: displacement of centerline of electrospinning jet.    Ko et al. [128, 129] also suggested a simple method to predict the fiber diameter by applying theory from Berry. Berry stated that the degree of entanglement of polymer chains    28  in solution could be described by a dimensionless number called the Berry number, Be  [130]:                         Be = [η]C                                                                                          (Eq. 2.3-4)                        d = a Bec                                                                                           (Eq. 2.3-5)                                                                       where η: intrinsic viscosity of solution, C: concentration of solution, d: fiber diameter, and a and c: constant.  When polymer is dissolved in a solvent with dilute concentration, the polymer molecules are so far apart in the solvent that polymer chains rarely entangled. When the polymer concentration increases, the polymer molecules interact and become entangled. The polymer solution at Be = 1 found to be electrospinable solution generating nanofibers. As Be increases, the fiber diameter increases until Be approaches up to 4 [128, 131]. The ability of electrospinning to generate relatively large-size of non-woven nanofibrous structures makes it an ideal process for biomedical scaffold fabrication. In addition to variety of materials, from biodegradable, non-degradable, natural to synthetic biopolymers formable through electrospinning, such unique fibrous architecture with interconnected 3D pores and a high surface area  mimicking natural extra-cellular matrix (ECM) make it particularly suitable for biomedical applications. It has been demonstrated that they can act as tissue engineering scaffolds, drug delivery devices, and wound dressing [11, 13, 17].  2.3.1 Tissue engineering scaffolds Tissue engineering is an interdisciplinary field that applies the principles of life science and engineering to reconstruct or regenerate malfunctioned tissues and organs. It is well known that biological tissues consist of well-organized hierarchical fibrous structures (ECM), where cells are embedded [41,112, 244]. Hence, it has long been hypothesized that a tissue-engineered scaffold should mimic an ECM with specific requirements in order to  29  introduce it in the body as a part of not a foreign body but a native tissue. The purpose of tissue engineering scaffold is to act as an ECM where cells could adhere and grow, and thus to guide the development of new and fully functional tissues. The scaffold must be biocompatible without any adverse tissue reaction and should not interrupt physiological processes. Its 3D interconnected porous structure must sustain mechanical stability until a damaged tissue/organ is fully regenerated. For clinical and commercial success, scaffold production must be simple and versatile enough to allow a wide array of configurations to accommodate the size, shape, and strength of the target tissue. Once tissue regeneration is completed, the scaffold must be removed via degradation or absorption, ideally without leaving any residues [11, 13, 17].   Electrospinning has drawn recent interest in producing tissue engineering scaffolds due to its relative ease of use, adaptability, and ability to fabricate fibers with diameters in nanometer scales closely mimicking the size of collagen fibers found in the ECM. The inherently high surface area to volume ratio of electrospun nanofibrous scaffolds can enhance cell attachment, and high level of interconnected 3D porosity can accommodate large number of cells, vascularization, and diffusion of nutrients and oxygen. Some research has encountered limitations with regards to cell infiltration into nanofibrous membrane due to their relatively small pore sizes [2–4]. However, electrospun scaffolds have shown to have great potential in a number of tissues engineering applications, including: vasculature [113], bone [20–22], neural [16], and tendon/ligament [100].   He et al. [113] examined the effects of electrospun collagen-blended PLA-co-PCL nanofibrous scaffold on human coronary artery endothelial cell (HCAEC) viability and attachment. In an in vitro study, they demonstrated that incorporation of collagen was able to enhance HCAEC viability, spreading, and attachment. Nie and Wang [22] examined the release of DNA from electrospun scaffolds consisting of a blend of PLGA and HAp with various HAp contents (0%, 5%, and 10%) for bone tissue engineering applications. The  30  authors noted that encapsulated DNA/chitosan nanoparticles with HAp enhanced transfection efficiency leading to higher cell attachment and viability. For neural tissue engineering applications, Yang et al. [112] examined the performance of both aligned and random PLA electrospun scaffolds. The authors observed that neural stem cells (NSCs) oriented parallel to the aligned fibers, giving directed neurite outgrowth. However, the NSCs cultured on random PLA scaffolds did not show a directed orientation. Sahoo et al. [114] electrospun PLGA nanofibers onto a knitted PLGA scaffold in order to provide a large surface area for cell attachment. Knitted scaffolds were fabricated from PLGA microfibers (10:90 PLA:PGA, 3 yarns, 20 filaments/yarn, 25 µm diameter of filament; Shanghai Tianqing Biomaterial, Shanghai, China). PLGA nanofibers were electrospun onto the surface of the knitted PLGA to make hybrid nano-microfibrous scaffolds. The authors then examined porcine bone marrow stromal cell attachment, proliferation, and ECM synthesis on the electrospun/knitted composite scaffold as compared to a knit PLGA scaffold in which cells were immobilized using a fibrin gel. They found that cell proliferation and cellular activity were both increased in the electrospun/knit composite scaffold.   2.3.2. Drug delivery devices Drug delivery is a rapidly developing biomedical research field, which requires interdisciplinary expertise in biotechnology, pharmacology, microbiology, biochemistry, polymer chemistry, and materials engineering. Controlled drug delivery has attracted the attention of many researchers as it allows release of drugs with precision at the specific sites. This eliminates adverse effects like over dosage and toxicity of the drug. Advantages of using such system include maintenance of drug levels within desired range, fewer administrations, optimal use of the drug, and better patient compliance. The material used for drug delivery should be biocompatible, chemically inert, easily processable, and physically and mechanically stable [84].    31  Electrospun scaffolds are of great interest in drug delivery as electrospinning affords great flexibility in selecting materials, their morphology, and other physical-biological-chemical characteristics for drug delivery applications. Either biodegradable or non-degradable materials can be used to control drug release via diffusion and scaffold degradation or diffusion alone [11, 13]. Additionally, due to the flexibility in material selection a number of drugs can be delivered including: antibiotics, anticancer drugs, proteins, and DNA. Using the various electrospininng techniques, a number of different drug loading methods can also be utilized: coatings, embedding, and encapsulation [13].                        32  Chapter 3 Scope and Objectives  3.1 Scope of the investigation The principal motivation of the present work is search for novel processing methods to achieve homogenous CaP/biopolymer nanocomposite fibrous scaffolds. As the mineralized collagen fibrils with carbonated hydroxyapatite nanocrystals are a foundation of bone tissue, it is considered that CaP/biopolymer composites would be the most feasible scaffolds for bone tissue engineering. The addition of bone-mineral like CaP is expected to improve structural stability and provide bioactivity, osteoconductivity, and osteoinductivity. Electrospinning produces a non-woven nanofibrous mesh structure with 3D interconnected pores. This unique fibrous nanostructure, mimicking natural ECM, has shown  in the past promising performance in multi-functional scaffolds for biomedical applications. The nanofibers with a high surface area to volume ratio allow cellular migration and proliferation in tissue engineering scaffolds and load/release of larger quantities of drugs for drug delivery. Simplicity and efficiency are additional advantages of electrospinning for scaffold fabrication. It was also hypothesized that the incorporation of CaP into biopolymer fibers could make electrospinning more attractive for the preparation of organic/inorganic scaffolds. However, previous studies mostly used mechanically blended CaP/biopolymer solutions for electrospinning, which caused severe agglomeration of the CaP nanoparticles and consequently irregular distribution of CaP aggregates within/on fibers.   In this work, novel strategic methods for in-situ biomimetic formation of CaP have been researched and developed. The nucleation and growth of CaP nanocrystals within biopolymer matrices, mimicking the biomineralization process of the mineralized collagen fibrils, have been studied to overcome the drawbacks of the traditionally processed CaP/biopolymer composite fibrous scaffolds. Electrospun CaP/biopolymer nanocomposite fibrous scaffolds were processed via two different routes; i) DCPA/PLA and ii)  33  HAp/alginate. It was postulated that the in-situ synthesis of CaP within the biopolymer environment could induce better distribution of the inorganic phase within the organic matrix than a simple blending method, resulting in homogenous CaP/biopolymer nanocomposite scaffolds for better mechanical stability and cellular response. The nanocomposites were characterized as follows:  X-ray diffraction for characterization of the CaP in the biopolymer matrices,  Fourier transform infrared spectroscopy for verification of the presence and chemical interaction of the CaP with the biopolymers,  Scanning and transmission electron microscopy and energy dispersive spectroscopy for observation of the distributed CaP in the biopolymer nanofibers,   Scanning electron microscopy for observation of the morphology of the nanocomposite fibrous scaffolds,  Thermogravimetric analysis for quantitative analysis of the CaP amount in the scaffolds.  Tensile strength properties of the scaffolds using a micro-tensometer.  Comparative characterization of osteoblast response to the scaffolds using MTS assay for proliferation, DAPI staining for cell number counting, and scanning electron microscopy and confocal microscopy for observation of cell morphology.  PLA and PLGA have been used for tissue engineering and drug delivery scaffolds. Biodegradation rate of PLGA can be controlled by changing the ratio of lactic and glycolic monomers, which makes it more favorable for drug delivery applicatons than PLA [232].  In the present research, PLA was chosen as a representative synthetic biodegradable polymer over PLGA due to the following reasons. PLA has higher mechanical and structural  stability in physiological conditions than PLGA [230−232]. The primary function of tissue regeneration engineering scaffold is to provide a stable physical construct  34  until seeded cells attach, proliferate, and differentiate into mature cells for the process of targeted tissue regeneration. PLA breaks down inside the body within 6 months to 2 years with gradual degradation which is enough period of time for bone tissue regeneration. On the other hand, PLGA degrades within from a few months to 6 months, depending on the ratio of the monomers. PLGA has been used in a number of clinical applications but  few applications are reported  in the orthopedic area [232]. PLGA cannot be utilized in load bearing applications due to its low mechanical strength,  low stiffness, and lack of osteogenic bioactivity [234]. Alginate was chosen as another biopolymer matrix for calcium phosphate in-situ synthesis. It is a natural polysaccharide which is biocompatible, non-toxic, non-immunogenic, and biodegradable. With these features, alginate has been widely used as a biomaterial in many fields such as cell immobilization, tissue engineering, drug deliery, and wound healing dressing [14, 35, 88−91, 235]. The unique hydrogel properties, when cross-linked with divalent cations, and the possession of the hydroxyl and carboxyl functional groups give it chances of the surface modification for targeted applications.     We believe that the electrospinning of in-situ synthesized DCPA into PLA nanofibrous scaffolds and processing of HAp/alginate nanocomposite fibrous scaffolds through biomimetic nucleation and growth of HAp on electrospun alginate nanofibers are originally demonstrated for the first time in this work,     3.2 Objectives In view of the above scope of the investigation, the broad objective is to study and achieve a well-controlled and reproducible process to obtain homogeneous CaP/biopolymer nanocomposite fibrous scaffolds, and to characterize the scaffolds. The specific objectives of the proposed research are as follows:  35   1. Study, characterization, and optimization of in-situ formation of calcium phosphate in the presence of PLA molecular chains. 2. Process development and optimization for fibrous scaffolds by electrospinning of the in-situ synthesized DCPA/PLA composite solutions. 3. Investigation of the effects of secondary polymeric source, poly(ethylene glycol), on the electrospinning process of DCPA/PLA nanocomposite solutions. 4. Process development and characteriza  tion of HAp/alginate nanocomposite fibrous scaffolds via biomimetic nucleation/growth of HAp on electrospun alginate nanofibers. 5. Systematic investigation of the effects of the process parameters on the morphology of the HAp/alginate scaffolds, including concentration of the precursor ion. 6. Characterization of the physico-chemical properties and biological in-vitro properties of the CaP/biopolymer scaffolds.                  36  Chapter 4 Materials and Methodology  4.1 Fabrication of DCPA/PLA nanocomposite fibrous scaffolds  4.1.1 Raw materials For dicalcium phosphate anhydrate synthesis, calcium nitrate tetrahydrate (CNT; Ca(NO3)2·4H2O, Cat #:13477-34-4) as a Ca2+ precursor, and sodium phosphate monobasic dihydrate (SPM; NaH2(PO4)·2H2O, Cat #: 13472-35-0) as a PO43–  precursor were purchased from Fisher Scientific, Canada. Polyl(actic acid) (PLA; Catalogue #: 2002D,  Natureworks, USA) and poly(ethylene glycol) (PEG; MW:20,000, Cat #: 25322-68-3, Alfa Aesar, USA) were used as the polymeric sources for electrospinning. Organic solvents of tetrahydrofuran (THF, Cat #: 109-99-9), dimethyl formamide (DMF, Cat #: 68-12-2), and dimethyl sulfoxide (DMSO, Cat #: 67-68-5) from Fisher Scientific, Canada were used to dissolve CNT, SPM, PLA, and PEG for preparation of the electrospinning solution.  4.1.2 In-situ synthesis of DCPA/PLA nanocomposite suspension A 10 wt% PLA solution was prepared in THF/DMF (70/30 by wt %) on a magnetic stirring hot plate at 35 ºC for 48 hrs. The CNT was completely dissolved in the PLA solution, providing Ca2+ precursor ions. The SPM dissolved in the DMSO at 60 ºC on the same plate for 24 hrs was drop-wise added into the CNT/PLA solution to supply PO43– precursor ions for the DCPA in-situ synthesis. It was hypothesized that the Ca2+ ions interact with the carbonyl groups of the PLA, making PLA/Ca2+ complexes and subsequently functioning as nucleation sites for the DCPA, supplemented by PO43– ions. The composite suspension was stirred for 24 hrs on the plate. The PEG dissolved in the DMF was ad-mixed into the composite suspension for 6 hrs as a secondary polymeric source. The PEG was used as a co-polymeric source to aid electrospinning of PLA/DCPA nanocomposite suspension. A mass of 1.0 g of 20 wt% Triton X-100 (Cat #:807423, Sigma Aldrich, USA) solution in the DMSO was added as a surfactant into the composite suspension. The suspension was ultra- 37  sonicated before electrospinning. The in-situ synthesized nanocomposite suspension was referred to as DCPA/PLA and DCPA/PLA/PEG depending on the presence of PEG in  the suspension. Two reference samples were prepared for comparison. A pure 10 wt% PLA solution dissolved in THF/DMF (70/3 by wt%) was referred to as Ref-PLA. Mechanically  blended PLA solution with pre- synthesized DCPA was referred to as Ref-DCPA/PLA-Mech. The CNT was dissolved in THF/DMF (70/30 by wt%), and the SPM dissolved in the DMSO was drop-wise added into the PLA-free CNT solution to synthesize DCPA nanocrystals. The compositions of the all samples are compiled in Table 4.1. The organic solvents of THF, DMF, and DMSO were chosen among the others, such as alcohols, chloroform, dichloromethane, and so on,  since they mix homogeneously and become clear solution, not making emulsion and/or precipitation.  The concentration of PLA solution was decided, based on the electrospun nanofiber morphologies depending on the concentration from  6 wt % to 12 wt %. The 10 wt % PLA solution produced uniformly deposited nanofibers without the foramtion of beads  and dried solution at the tip of syringe while electrospinning.      Table 4.1 Compositions of  in-situ synthesized DCPA/PLA and DCPA/PLA/PEG nanocomposite suspensions and reference solution.    CNT (g) SPM (g) THF (g) DMF (g) DMSO (g) PLA (g) PEG (g) DCPA/PLA-1 0.25 0.10 24.00 10.20 10.00 3.80 − DCPA/PLA-2 0.50 0.20 24.00 10.20 10.00 3.80 − DCPA/PLA-3 0.75 0.30 24.00 10.20 10.00 3.80 − DCPA/PLA-4 1.00 0.40 24.00 10.20 10.00 3.80 −         DCPA/PLA/PEG-1 0.25 0.10 24.00 15.20 10.00 3.80 3.80 DCPA/PLA/PEG-2 0.50 0.20 24.00 15.20 10.00 3.80 3.80 DCPA/PLA/PEG-3 0.75 0.30 24.00 15.20 10.00 3.80 3.80 DCPA/PLA/PEG-4 1.00 0.40 24.00 15.20 10.00 3.80 3.80 DCPA/PLA/PEG-5 1.25 0.50 24.00 15.20 10.00 3.80 3.80 DCPA/PLA/PEG-6 1.50 0.60 24.00 15.20 10.00 3.80 3.80 DCPA/PLA/PEG-7 1.75 0.70 24.00 15.20 10.00 3.80 3.80 DCPA/PLA/PEG-8 2.00 0.80 24.00 15.20 10.00 3.80 3.80         Ref-PLA − − 24.00 10.20 − 3.80 − Ref-DCPA/PLA-Mech 1.00 0.40 24.00 10.20 10.00 3.80 −  38  4.1.3 Electrospinning of DCPA/PLA nanocomposite suspension Electrospinning was performed using the electrospinning system (NANON, MECC Co., Ltd., Japan) shown in Figure 4.1. The solution was placed in a 10 ml disposable syringe (BD, USA) with a needle (PrecisionGlideTM 18G 1½, BD, USA) mounted horizontally in a syringe holder. A syringe pump ejected the solution at a rate of 1.0 ml/hr, and a voltage of 28 kV was applied at the needle tip. The spinning distance between the needle tip and the collection target was 20 cm. The tip moved horizontally over a distance of 20 mm at a rate of 20 mm/sec. The cylinder grounded target covered with Al foil rotated at 300 rpm to collect the electrospun fibers. The fibrous scaffold was washed-out with de-ionized water twice by vacuum filtration and freeze-dried in a lyophilizer (FreeZone 2.5. Labconco Co.,     Figure 4.1 Electrospinning system for the fabrication of DCPA/PLA nanocomposite fibrous scaffolds (NANON, MECC Co., Ltd., Japan).  39  USA) for 24 hrs. The scaffolds were named as DCPA/PLA and DCPA/PLG/PEG depending on the addition of PEG before electrospinning, respectively. NANON electrospinning unit enabled to electrospin DCPA/PLA and DCPA/PLA/PEG nanocomposite suspensions and collect the nanocomposite fibers, using the high ratation speed of drum. Furthermore, the DCPA/PLA/PEG scaffolds were re-dissolved with the THF/DMF solution and electrospun with the same parameters. These samples were referred to as Re-DCPA/PLA/PEG scaffolds.     4.2 Fabrication of HAp/alginate nanocomposite fibrous scaffolds 4.2.1 Raw materials Sodium alginate (medium viscosity, Cat #: 9005-38-3) and poly(ethylene oxide) (PEO; Mw=900,000, Cat #: 25322-68-3) were purchased from Sigma-Aldrich, Canada. Triton X-100 (Cat #: 807423, Fisher Scientific, Canada) was used as a surfactant. De-ionized (D·I) water was produced in the lab, and dimethyl sulfoxide (DMSO; Cat #: 67-68-5) was purchased from Fisher Scientific, USA. For HAp synthesis, calcium nitrate tetrahydrate (CNT; Ca(NO3)2·4H2O, Cat #:13477-34-4) as a Ca2+ precursor and sodium phosphate monobasic dihydrate (SPM; NaH2(PO4)·2H2O, Cat #: 13472-35-0) as a PO43–  precursor were purchased from Fisher Scientific, Canada.     4.2.2 Preparation of alginate/PEO solution   40 g of 4.0 wt% aqueous sodium alginate solution  and 40 g of 5.0 wt% aqueous PEO solution were prepared with Vortex mixing, ultra-sonication, and magnetic stirring for 3 days. PEO was used to make the sodium alginate solution electrospinnable. The function of PEO for electrospinning alginate/PEO solution will be explained in the section of 6.1.1. 4.67 g of 9.0 wt % Triton X-100 solution in the DMSO was produced with magnetic  40  stirring. The three solutions were mixed together into the final compositions of 1.9 wt% sodium alginate, 2.4 wt% PEO, 0.5 wt% Triton X-100, 5.0 wt% DMSO, and 90.2 wt% water. The SPM dissolved in 0.5 mL D·I water was vigorously mixed into the alginate/PEO solution using a vortex mixer to provide precursor ions for HAp in-situ synthesis, namely PO43–. The alginate solutions mixed with SPM concentrations of 0.04 to 0.32 g are referred to as H-Alginate 1 to H-Alginate 8, respectively. The reference solution without SPM is referred to as Ref-Alginate. All the components of the solutions are shown in Table 4.2.   Table 4.2 Compositions of H-Alginate solutions and reference solution for electrospinning.   SPM  (g) Na-alginate (g) PEO  (g) Triton X-100 (g) DMSO  (g) D·I water (g) H-Alginate-1 0.04 0.756 0.945 0.198 2.007 36.09 H-Alginate-2 0.08 0.756 0.945 0.198 2.007 36.09 H-Alginate-3 0.12 0.756 0.945 0.198 2.007 36.09 H-Alginate-4 0.16 0.756 0.945 0.198 2.007 36.09 H-Alginate-5 0.20 0.756 0.945 0.198 2.007 36.09 H-Alginate-6 0.24 0.756 0.945 0.198 2.007 36.09 H-Alginate-7 0.28 0.756 0.945 0.198 2.007 36.09 H-Alginate-8 0.32 0.756 0.945 0.198 2.007 36.09 Ref-Alginate ‒ 0.756 0.945 0.198 2.007 36.09   4.2.3 Electrospininng of alginate/PEO solution 40 mL of alginate solution was electrospun to fabricate nanofibrous scaffold by using the electrospinning system (NEU-1, Kato Tech. Co. Ltd., Japan) shown in Figure 4.2. The solution was placed in a 10 mL disposable syringe (BD, USA) with a needle (PrecisionGlideTM 18G 1½, BD, USA). The syringe pump ejected the solution at a rate of 0.08 mm/min, and a voltage of 20 kV  applied at the needle tip. The grounded drum rotated at a rate of 2.50 m/min to collect electrospun nanofibers. The spinning distance between the tip and the drum was 20 cm.   41    Figure 4.2 Electrospinning system for the fabrication of HAp/alginate nanocomposite fibrous scaffolds (NEU-1, Kato Tech. Co. Ltd., Japan).   4.2.4 In-situ synthesis for HAp/alginate nanocomposite fibrous scaffolds To induce biomimetic in-situ formation of HAp on the alginate nanofibers, the electrospun H-Alginate scaffolds were stirred in aqueous 1.0 M CNT solution for 24 hrs, to supply Ca2+ which act as precursor ions for both cross-linking of the alginate and HAp nucleation/growth. 1 N NaOH (Cat #: SS266, Fisher Scientific, Canada) solution was titrated into the solution for pH 10. The Ref-Alginate scaffold was dipped in the CNT solution without titration for 3 hrs. The in-situ synthesized H-Alginate scaffolds were washed with D·I water and vacuum-filtration twice, frozen, and then freeze-dried in a lyophilizer (FreeZone 2.5. Labconco Co., USA) for 24 hrs. Another reference scaffold was fabricated by a mechanical blending/electrospinning method to compare morphology and distribution of HAp nanoparticles with the in-situ synthesized scaffolds. HAp nanoparticles were pre-synthesized using a precipitation method in aqueous solution at pH 10. The SPM concentration was the same as the H-Alginate-5, and Ca2+ ions were provided from CNT with a Ca/P ratio of 1.67. 20 mL of the SPM solution was drop-wise added into the 20 mL CNT solution. This solution was titrated with 1 N NaOH and stirred for 24 hrs. The HAp precipitates were washed with D·I water three times using a centrifuge, and vigorously  42  mixed with the alginate/PEO solution using a vortex mixer and ultra-sonication before electrospinning.    4.3 Characterization of CaP/biopolymer nanocomposite fibrous scaffolds 4.3.1 X-ray diffraction  The DCPA/PLA nanocomposite suspension was dried on a glass slide at room temperature and washed-out with de-ionized water to remove by-products from the synthesis. Completely dried glass-slide was inserted on the sample holder. In case of the HAp/alginate samples, the lyophilized HAp/alginate fibrous scaffold was stacked on the sample holder. The nucleation and phase transformations of the inorganic phases were  monitored by X-ray diffractometer (XRD; Rigaku Multiflex, Japan) with CuKα radiation at 40 kV and 20 mA. Diffraction data was collected from 2θ = 10 – 70º with a step size of 0.02º.   4.3.2 Scanning electron microscopy The morphology of the electrospun fibrous scaffolds was examined using scanning electron microscopy (SEM, Hitach S-3000, Japan) at an operating voltage of 5.0 kV. The surface of the scaffolds was coated by gold sputtering. The fiber diameter distribution was measured at 50 different locations on the SEM micrograph and analyzed by ImageJ software.  4.3.3 Scanning and transmission electron microscopy Transmission electron microscopy (TEM; Hitachi H-800, Japan) was used to verify formation and dispersion of the DCPA  in the PLA nanofibers and to perform an analysis  an analysis of selected area electron diffraction (SAED) patterns. The electrospun nanofibers were directly deposited on a carbon-coated Cu grid and sandwiched with a plain  43  Cu grid to firmly hold the nanofibers while an accelerating voltage of 100 kV. The TEM was changed into SEM mode for energy dispersive spectroscopy (EDS) mapping to analyze the element distribution of the DCPA/PLA nanocomposite fibers.   4.3.4 Fourier transform infrared spectroscopy The presence and interaction of the inorganic phases in the biopolymer polymer nanofibers were investigated by Fourier transform infrared spectroscopy (FT-IR; Perkin Elmer 2000, USA) over the range of 400 to 4000 cm-1 at a resolution of 0.5 cm-1. The fibrous scaffold mixed with KBr powder in a mass ratio of 1:20 was pressed into a pellet by a hydraulic press for sample preparation.   4.3.5 Thermogravimetric analysis Thermogravimetric analysis (TGA, Linseis L81/1750, USA) was used to quantify the inorganic phases in the fibrous scaffolds by heating up to 600 ºC and 1200 ºC at a heating rate of 5 ºC min–1 for the DCPA/PLA and HAp/alginate scaffolds, respectively.  4.3.6 Tensile properties of scaffolds Tensile properties of nanofibrous scaffolds were measured using a micro-tensile tester (KEG-G1, Kawabata, Japan). The scaffold were cut to strip of 0.3 cm in width by 3.0 cm in length, and then glued on a paper sample frame using double sided tape. The gauge length for tensile tests were 2.4 cm. After mounting the sample, the paper frame was cut in the middle, and the strip was elongated at an  extension rate of 0.02 cm/s. The load in grams was measured as a function of time with a load sensitivity of 100g/V. The time was converted to displacement by multiplying by the extension rate. The displacement was converted to strain by dividing by the gauge length. The measured load in grams was converted to specific stress (g/tex) using the following equation [139].                                                               Force (g) / specimen width (mm)                Specific stress (g/tex) =                                                                     Areal density (g/m2)                             (Eq. 4.3-1)  44  The areal density is simply the weight (g) of the strip divided by the area (m2) of the strip. The specific stress (g/tex) was then converted to Engineering stress (MPa) using following equation.          Engineering Stress (MPa) = 9.8 × Specific stress (g/tex) × density of sample (g/cm3)                                                                                                                                  (Eq. 4.3-2)  The average tensile properties of each scaffolds were calculated from ten different strip samples.  4.3.7 In-vitro bioactivity analysis  In-vitro bioactivity of the nanofibrous scaffolds was tested using simulated body fluid (SBF) solution. SBF solution has nearly equal ion concentration to those of human blood plasma, which has been used for in-vitro bioactivity simulation since two decades ago. 1L of original Kokubo's SBF solution was prepared by following the literature (Table 4.3) [140]. The scaffold was filled with the SBF solution in a 20 mL of capped vial and stored in a warm water bath at 37 °C for 36 hours and 10 days, respectively. At the point of 24 hours, it was changed with the fresh SBF solution. The scaffolds were washed with D·I water twice slightly and freeze-dried using the lyophilizer for 24 hours. The formation of bone-like apatite was confirmed using XRD, SEM, and EDS analysis.      Table 4.3 Raw materials and ion concentration of original Kokubo's SBF solution [140].  Raw materials  Ion concentrations    NaCl 8.035 g/L    Na+ 142.00    NaHCO3 0.355 g/L    K+    5.00    KCl 0.225 g/L    Mg+    1.50    Na2HPO4·2H2O 0.231 g/L    Ca2+    2.50    K2HPO4·3H2O 0.311 g/L    HCO3‒    4.20    CaCl2 0.292 g/L    HPO42‒    1.00    Na2SO4 0.072 g/L    SO42‒    0.50   1M HCl 0.160 mL    Cl‒ 147.96  45  4.4 Rat calvarial osteoblast culture on scaffolds Osteoblasts cells obtained from newborn rat calvaria were cultured as previously described [135]. Briefly, rat calvarial osteoblast (RCO) cells were cultured on tissue culture plastic (Falcon, Becton Dickinson Labware, USA) in α–minimal essential medium (α–MEM; Cat #: 36453 Stem Cell Technologies, Canada) supplemented with 100 mg/mL penicillin G (Cat #: P 3032, Sigma-Aldrich, Canada), 50 mg/mL gentamicin (Cat #: G1264 Sigma-Aldrich, Canada), 3 mg/mL amphotericin B (Cat #: A2411, Sigma-Aldrich, USA), and 15 % newborn bovine serum (Cansera, Canada) at 37 °C in a humidified atmosphere of 95 % air and 5 % CO2. Before cell-seeding, scaffolds were cut into 1 cm x 1 cm pieces, sterilized by argon-gas glow discharge treatment, glued with autoclaved silicone grease (Dow Corning, USA) to attach them to the bottom of a 24-well cell culture plate (Falcon, Becton Dickinson Labware, USA), and pre-soaked in the basic α–MEM overnight. The RCO cells were trypsinized (0.25 % trypsin, 0.1 % glucose, and citrate–saline buffer pH 7.8) prior to confluent growth. The RCO cells were counted electronically (Cell Counter, Coulter Electronics Limited, England) and plated onto the surface of the Alginate-based  scaffolds at 2 × 104 cells/mL. The medium was changed 24 hrs after post cell-seeding and then every other day. The RCO cells were cultured up to 7 days for the observation of cell morphology using SEM and immunocytochemistry staining, and the analysis of cell proliferation using MTS assay and nucleus counting. The RCO cells were cultured on the PLA-based scaffolds to detect formation of bio-mineralized hard tissue, bone-like nodule,  up to 6 weeks.  4.4.1 Scanning electron microscopy The RCO cells were seeded on the scaffolds at 2 × 104 cells/mL, and cultured for 1, 3, 5, and 7 days for SEM sample preparation to observe the morphology of the cells. The RCO cells were washed with phosphate-buffered saline (PBS) twice for 5 min. each fixed with 3.7 % formaldehyde solution for 4 hrs, washed with PBS twice for 5 min. each, and dehydrated using a series of increasing ethanol concentrations (50, 70, 80, 90, 95, and  46  100 %), each concentration for 1 hr. In the case of the alginate-based scaffolds, the basic α–MEM medium  was used instead of PBS for the washing procedure. The scaffolds were then critical point-dried and coated by gold sputtering to investigate the morphology of the RCO cells using scanning electron microscopy (SEM, Hitach S-3000, Japan) at an operating voltage of 5.0 ‒ 10.0 kV.  4.4.2 Immunocytochemistry staining The RCO cells were seeded on the scaffolds at 2 × 104 cells/mL, and cultured for 1, 3, 5, and 7 days for immunocytochemistry staining to observe cytoskeletal organization of the cells. The samples were washed with PBS twice for 5 minutes each, and fixed with 3.7 % formaldehyde for 20 minutes., and washed with PBS three times for 1 minute each. Permeabilization was performed in 0.05 % Triton X-100 (Sigma-Aldrich, Canada) for 5 minutes, and washed with PBS three times for 5 minutes each. The primary antibody used in this study was a mouse lgG1 isotype monoclonal Anti-Vinculin FITC (Vinculin; Clone hVIN-1, Sigma-Aldrich, Canada). 200 µL of the diluted Vinculin solution in PBS at a concentration of 1:50 was added into each well of the cell culture plate and incubated for 2 hours. Phalloidin‒Tetramethylrhodamine B isothiocyanate (Phalloidin- TRITC; catalogue #: P1951, Sigma-Aldrich, Canada) was used for F-actin staining. 200 µL of the diluted Phalloidin solution in PBS at a concentration of 1:500 was added into the sample well of the cell culture plate for 40 minutes. The samples were washed with PBS three times quickly. For nucleus staining, 600 µL of  the diluted DAPI (Catalogue #: D8417, Sigma-Aldrich, Canada) solution in PBS at a concentration of 0.5 µg/mL was added into the sample for 20 minutes at room temperature. The samples were washed with PBS twice for 5 minutes each and then mounted on a slide in Fluoromount-GTM slide mounting medium (Catalogue #: OB100-01, Fisher Scientific, Canada).  In the case of the alginate-based scaffolds, the basic α–MEM medium  was used instead of PBS for the washing procedure. Labeling was visualized on a Nikon Eclipse C1 confocal laser scanning microscope, and images were captured using Nikon Eclipse software.  47  4.4.3 MTS assay  The RCO cells were seeded on the scaffolds at 2 × 104 cells/mL, and cultured for 1, 3, 5, and 7 days. The number of viable cells on the scaffolds was quantified using the metabolic MTS assay (Promega™ CellTiter 96™ AQueous Nonradioactive Cell Proliferation Assay, catalogue #: PR-G 5421, Fisher Scientific, Canada). 200 µL of the MTS reagent was added into each well of the cell culture plate. After 4 hours of incubation, 200 µL of the supernatants were transferred in triplicates to a 96 well plates (Nunc MicroWell 96-Well Microplate, catalogue #: 1256566, Fisher Scientific, Canada), and the absorbance of the formazan produced by cellular reduction of the MTS was measured at 490 nm on a microplate reader (Model 3550, Bio-Rad, USA).  4.4.4 DAPI staining  To assess cell number on the surfaces of the scaffolds,  the RCO cells were seeded at 2 × 104 cells/mL, and cultured for 1, 3, 5, and 7 days. To calculate cell number, the cells  were stained with DAPI (Catalogue #: D8417, Sigma-Aldrich, Canada). The scaffolds were washed with PBS twice for 5 minutes each, and the cells were fixed using 70 % ethanol for 20 minutes. The scaffolds were washed again with PBS for 5 minutes, and 600 µL of  the diluted DAPI solution in PBS at a concentration of 0.5 µg/mL was added into each well of the cell culture plate for 10 minutes. The samples were washed with PBS twice for 5 minutes each. Glass coverslips were mounted onto the samples with Fluoromount-GTM slide mounting medium (Catalogue #: OB100-01, Fisher Scientific, Canada). In the case of the alginate-based scaffolds, the basic α–MEM medium  was used instead of PBS for the washing procedure. The labelled nuclei of the cells were visualized on a Zeiss Axioplan 2 microscope, and images were captured using Nikon Eclipse software. The number of stained nuclei was counted using ImageJ software.     48  4.4.5 Tetracycline labeling  The RCO cells of 2 × 105 cells/mL were cultured on the PLA-based nanofibrous scaffolds  for from 2 to 6 weeks. Cell culture procedures are similar to as described  above, but after 1 week of post-seeding, 31.5 mg/mL β-Glycerophosphate disodium salt hydrate (Cat #: G9442, Sigma-Aldrich, Canada) and 0.58 mg/mL L-ascorbic acid phosphate magnesium salt n-hydrate (Cat #: 013-19641Wako Pure Chemical Industries, Richmond, VA) were added to the α-MEM supplemented with the antibiotics and 15% fetal bovine serum to promote mineralization process of the cells as well as tetracycline  (Cat #: 3258, Sigma-Aldrich, Canada) to label mineralized deposit. The medium was changed three times a week. The scaffolds were washed with PBS twice for 5 minutes each, fixed with 3.7 % formaldehyde for  2 hours, and rinsed with PBS three times quickly. Glass coverslips were mounted onto the samples with Fluoromount-GTM slide mounting medium (Catalogue #: OB100-01, Fisher Scientific, Canada). Bone nodules labeled by tetracycline were visualized on a Zeiss Axioplan 2 microscope using light (λmax = 488 nm), and images were captured using Nikon Eclipse software.  4.4.6 Multiphoton microscopy   Multiphoton microscopy (MPM) was utilized for identification of the RCO cell layers grown on the PLA-based scaffolds and as well as infiltration of the cells into the scaffolds. MPM is a minimally invasive and high resolution optical imaging technique to be able to image cells and tissues  [136]. MPM excites and detects nonlinear signals of two-photon excited fluorescence (TPEF) and second harmonic generation (SHG). TPEF signals can be observed from endogenous fluorescent biochemical species such as reduced nicotinamide adenine dinucleotide and flavin adenine dinucleotide, two major fluorophores in cell cytoplasm and elastin fibers. On the other hand, SHG signals come from non-centrosymmetric structures such as type I collagen, an abundant connective tissue found in biological tissues. Therefore, MPM can image cellular and extracellular matrix structures by TPEF and SHG. contrasts. Images in MPM are formed by scanning a focused laser  49  beam and mapping the generation of the signals from the biological samples. MPM provides depth resolved imaging with sub-micrometer resolution. It typically has a field-of-view of a few hundred micrometers in depth and width and a relatively slow speed of 1 frame per second  [136‒138].  The details of the MPM system used in this study  can be found  in Ref [136]. The cell culture procedures are the same as for the tetracycline labeling. The samples were washed with PBS twice for 5 minutes each. The samples were transferred into a Nunc® 4-well cell culture plate (Cat #:Z688746, Sigma-Aldrich, Canada), were attached onto the bottom with the silicon grease, and filled with PBS for MPM observation.                     50  Chapter 5 Dicalcium Phosphate Anhydrate/Poly(lactic acid) Nanocomposite Fibrous Scaffolds  5.1 Fabrication of DCPA/PLA scaffolds 5.1.1 Biomimetic in-situ synthesis of DCPA in PLA solution Dicalcium phosphate anhydrate (DCPA), known as monetite, was successfully in-situ synthesized within the non-aqueous PLA solution at a pH = 4.0 ± 0.5. The XRD patterns in Figure 5.1 (c and d) show that low crystalline DCPA peaks at around 2θ = 26º and 30º were observed for DCPA/PLA-4 and DCPA/PLA-8. In the case of the DCPA/PLA-3 in Figure 5.1 (b), no crystallographic peaks for any calcium phosphates (CaP) were detected, indicating the formation of amorphous calcium phosphate (ACP). Conversely, Figure 5.1 (e) shows that in the absence of PLA, highly crystalline DCPA was synthesized with the same CNT and SPM concentrations as the DCPA/PLA-3. Investigations of the CaP synthesis in organic environments have demonstrated that the organic molecular chains suppress and/or delay the transformation of ACP to crystalline CaP, possibly by hindering the transportation of precursor ions during nucleation and crystal growth [141, 142]. In this system, it appears that the PLA molecular chains also obstruct the diffusion of Ca2+ and PO43– ions during DCPA nucleation and crystallization, resulting in the formation of ACP or low crystalline DCPA depending on the concentration of the Ca2+ and PO34– ions. A comparison of Figure 5.1 (a) with Figure 5.1 (b–d) indicates that the crystallinity of the PLA was not affected by the presence of ACP and DCPA in the nanocomposite systems.        51    Figure 5.1 XRD patterns of (a) Ref-PLA, (b) DCPA/PLA-3, (c) DCPA/PLA-4, and (d) DCPA/PLA-8. The sample (e) was synthesized in the absence of PLA with the same CNT concentration as (b) showing all characteristic crystalline DCPA peaks, matched with PDF # 01-070-1425.        52  5.1.2 Electrospinning DCPA/PLA scaffolds The in-situ synthesized DCPA/PLA suspensions were electrospun into nanocomposite fibers. The presence and dispersion of the DCPA in the PLA nanofibers was compared with  Ref-PLA and Ref-DCPA/PLA-Mech by examining the TEM micrographs (refer to Table 4.1 for the nomenclature of the materials processed in this work). Figure 5.2 (a) shows that the in-situ precipitated DCPA nanocrystallites, approximately 5 to 50 nm diameter and confirmed by the SAED pattern in the inset, were dispersed uniformly over the PLA nanofiber, unlike the smooth and plain Ref-PLA nanofibers shown in Figure 5.2 (c). Interestingly, intra-nanopores ranging from 20 to 200 nm were produced mostly in fibers of several hundred nanometers in diameter, as shown in Figure 5.2 (b). In contrast, the mechanically blended/electrospun Ref-DCPA/PLA-Mech fiber shown in Figure 5.2 (d)      Figure 5.2 TEM micrographs of (a) DCPA/PLA-4 dense nanofiber, (b) DCPA/PLA-4 porous nanofiber, (c) Ref-PLA nanofiber, and (d) Ref-DCPA/PLA-Mech nanofiber. The insets are the selected area electron diffraction (SAED) patterns on the spots with the white arrows.     53  exhibited a cluster of severely agglomerated DCPA nanocrystals approximately 3.0 μm large. Such an agglomeration of the inorganic phases was previously reported when electrospinning mechanically blended CaP/biopolymer composite solution [20‒22]. The SAED pattern on the cluster shows higher crystallinity of the pre-synthesized DCPA with more distinctive dots and rings than the patterns in the in-situ synthesized DCPA/PLA-4 nanocomposite fibers (Figure 5.2 (a and b)), which is in accordance with the XRD patterns in Figure 5.1 (c) and (e). In Figure 5.3, the STEM micrograph and the EDS mapping of the DCPA/PLA-4 nanocomposite fiber support the improved dispersion of the DCPA nanocrystallites with the uniform distribution of Ca, O, and P elements over the nanofiber.        Figure 5.3 (a) STEM micrograph and (b–d) EDS mapping of DCPA/PLA-4 nanocomposite fiber.  54  The nanocomposite suspensions from DCPA/PLA-1 to DCPA/PLA-4 were electrospun into non-woven fibrous scaffolds. Electrospinning the DCPA/PLA-5 and the suspensions containing greater amounts of DCPA did not produce continuous fibers, but rather short fibers mixed with beads and sprayed droplets on a macro scale. This finding is most likely due to the presence of a hypercritical amount of the DCPA nanocrystallites and by-products from the synthesis reaction, which cannot be carried by the PLA solution during electrospinning. The morphology of the DCPA/PLA scaffolds was studied using the SEM.  Compared with the Ref-PLA scaffold in Figure 5.4 (a), branched nanofibers were      Figure 5.4 SEM micrographs of (a) Ref-PLA nanofibrous scaffold and (b) Ref-DCPA/PLA-Mech nanocomposite fibrous scaffold.    55  easily detectable in the DCPA/PLA nanocomposite fibrous scaffolds, shown as white circles in Figure 5.5. Intra-nanopores in the fibers (indicated by white arrows) were also observed, as seen in the TEM micrograph in Figure 5.5 (b). The porous nanofibers were generated during the electrospinning process without any post-treatment, such as selective dissolution and photo- or thermal-degradation of one of the polymeric sources from the co-electrospun fibers as reported previously [143‒146]. The intra-nanopores were possibly generated by the evaporation of the non-volatile DMSO (TB.P=189 ºC) remaining in the fibers after electrospinning. The presence of such a remnant solvent may also cause a self-fusing phenomenon of the nanofibers. In the case of the Ref-DCPA/PLA-Mech scaffold in Figure 5.4 (b), a cluster of the agglomerated DCPA nanocrystals with an approximate      Figure 5.5 SEM micrographs of (a) DCPA/PLA-2, (b) DCPA/PLA-3 and (c) DCPA/PLA-4 nanocomposite fibrous scaffolds. The white arrows and circles indicate intra-nanopores and self-fused nanofibers, respectively.  56  size of 2.0 μm was observed, which resembles the TEM image of Figure 5.2 (d). The analysis of the fiber diameter distribution by the ImageJ software, Figure 5.6, shows that the nanocomposite fiber diameter ranged broadly from about 100 nm to about 3.0 μm,  due to the self-fusing effects, whereas the Ref-PLA nanofibers ranged from about 80 to about 500 nm. The fibers with the diameter larger than 3.0 μm were considered as beads and not counted for fiber diameter distribution analysis. In case of the DCPA/PLA-4 scaffolds, the sum of the frequency is not 100 % due to the presence of the beads. The fiber diameter of the DCPA/PLA scaffolds increased with the amount of the inorganic phases.     Figure 5.6 Fiber diameter distributions of (a) Ref-PLA nanofibrous scaffold, (b) DCPA/PLA-2, (c) DCPA/PLA-3, and (d) DCPA/PLA-4 nanocomposite fibrous scaffolds.      57  5.1.3 Effects of PEG on DCPA/PLA/PEG scaffolds The solution properties, such as concentration, viscosity, conductivity and surface tension, and electrospinning parameters, such as applied voltage, solution feeding rate and electrospinning distance, are the main factors influencing process efficiency and the size and morphology of the electrospun fibers. In the polymeric solution with fixed electrospinning parameters, fiber diameters generally increase with solution viscosity and solution concentration, but decrease with solution electrical conductivity. The incorporation of an inorganic phase causes the interactions in the solution to become complex, and the electrospinning outcome does not always follow the general rules. Indeed, electrospinning the DCPA/PLA nanocomposite suspensions was not as efficient as the Ref-PLA solution, likely due to the presence of the DCPA nanocrystallites, chemical reaction by-products, and non-compatibility of solvent DMSO to PLA. One of the ways to increase the electrospinability of such complex solutions is to add a second polymeric additive, co-solvent and/or surfactant that modifies the solution properties to stabilize the electrospun jet flows [14].Poly(ethylene oxide) (PEO) has been used widely  to control sol-gel transition of silk fibroin [20], alginate [14], and collagen [147] solutions. In this study, poly(ethylene glycol) (PEG) was ad-mixed into the DCPA/PLA suspensions as a temporary co-polymeric carrier to aid the PLA incorporating the DCPA nanocrystallites. Figure 5.7 shows the DCPA/PLA/PEG nanocomposite fibrous scaffolds after being washed with de-ionized water to remove the temporary PEG additive. The structure of the nanocomposite fibrous scaffolds remained stable after the PEG leaching-out process. The electrospun jets from the DCPA/PLA/PEG nanocomposite suspensions became more stable and continuous than those from the DCPA/PLA suspensions, resulting in the ability to fabricate the DCPA/PLA/PEG-8 scaffolds. Noticeable enhancement is shown in the DCPA/PLA/PEG-4 scaffold in Figure 5.7 (b) when compared to the DCPA/PLA-4 scaffold in Figure 5.5 (c). The higher frequency occurrence in the sub-micron fiber diameters of the DCPA/PLA/PEG scaffolds in Figure 5.8 indicates a decrease in the fiber diameters compared to the DCPA/PLA fibers in Figure 5.6. The fibers with the diameter larger than 3.0 μm were considered as beads and not counted for fiber diameter distribution analysis. In case of the  58  DCPA/PLA/PEG-8 scaffolds, the sum of the frequency is not 100 % due to the presence of the beads.    Figure 5.7 SEM micrographs of (a) DCPA/PLA/PEG-2, (b) DCPA/PLA/PEG-4, (c) DCPA/PLA/PEG-6, and (d) DCPA/PLA/PEG-8 nanocomposite fibrous scaffolds.  59    Figure 5.8 Fiber diameter distributions of (a) DCPA/PLA/PEG-2, (b) DCPA/PLA/PEG-4, (c) DCPA/PLA/PEG-6, and (d) DCPA/PLA/PEG-8 nanocomposite fibrous scaffolds.   5.1.4 Uniform DCPA/PLA nanofibers after re-electrospinning process The addition of poly(ethylene glycol) (PEG) into the in-situ synthesized DCPA/PLA suspensions gave rise to the formation of more stable electrospinning jet and the decrease of fiber diameters, as shown in the previous section. However, a uniform nanofibrous structure like the Ref-PLA scaffold was not achieved. The DCPA/PLA/PEG scaffolds after washing and freeze-drying procedures were re-dissolved and electrospun with the same electrospinning parameters.  The SEM micrographs in Figure 5.9 demonstrate that the nanofibrous structure of Re-DCPA/PLA/PEG-4 scaffold is as uniform and consistent as the neat PLA scaffold. The presence of DCPA nanocrystals was confirmed using TEM as shown in the inset of Figure 5.9 (b). It was found that the fiber diameters of the Re- 60  DCPA/PLA/PEG-4 scaffold was 354 ± 85.6 nm, slightly larger than 338 ± 92.1 nm of the Ref-PLA scaffold (Figure 5.10). It is considered that the uniform nanofibrous structure of the Re-DCPA/PLA/PEG-4 scaffold was achieved because the chemical reaction by-products produced by the DCPA in-situ synthesis were removed by the washing procedure, and only PLA soluble solvents of THF and DMF were used for preparation of the Re-DCPA/PLA/PEG-4 suspension for electrospinning.     Figure 5.9 SEM and TEM (inset) micrographs of (a) Ref-PLA and (b) Re-DCPA/PLA/PEG-4 nanofibrous scaffolds.  61    Figure 5.10 Fiber diameter distributions of (a) Ref-PLA and (b) Re-DCPA/PLA/PEG-4 nanofibrous scaffolds.   5.2 Characterization of DCPA/PLA scaffolds 5.2.1 Chemical interactions between DCPA and PLA during in-situ synthesis The FT-IR spectrum in Figure 5.11 (a) indicates the presence of DCPA in the DCPA/PLA-4 nanocomposite fibers with the phosphate bands at 470 and 956.5 cm-1 (PO4(v1)), 1046 cm-1 (PO4(v3)), and 577.5 cm-1 (PO4(v4)), compared to the Ref-PLA nanofibers in Figure 5.11 (b). It was also found that the C=O bond bands at 1760.5 and 711.5 cm-1 in the Ref-PLA nanofibers shifted to the lower wavenumbers of 1758 and 707.5 cm-1, respectively, in the DCPA/PLA-4 nanocomposite fibers. Such a shifting phenomenon in the carbonyl group regions was previously reported during the mineralization of poly(propylene fumarate) [142] and collagen [150] with HAp by the interaction of Ca2+ ions and the carbonyl groups in the polymers. It is well known that ultra-fine particles tend to flocculate in solution   62    Figure 5.11 FT-IR spectra of (a) DCPA/PLA-4 nanocomposite fibrous scaffold and (b) Ref-PLA nanofibrous scaffold.    unless they have relatively large zeta potential [148] or have organic molecules covering their surface forming steric obstacles against agglomeration [51]. During the in-situ synthesis, it is highly conceivable that the Ca2+ ions from the CNT form chelate bonding with unpaired electrons from the oxygen in the carbonyl groups of PLA, PLA/Ca2+ complexes function as nucleation/crystal growth sites for the DCPA, as illustrated in Figure 5.12. Correspondingly, neighboring PLA molecular chains surround the DCPA nanocrystallites. This synthetic mechanism that mimics the biomineralization of the  63  mineralized collagen fibrils leads to preventing of agglomeration of the DCPA nanocrystallites and induces their uniform distribution in the PLA nanofibers through electrospinning.      Figure 5.12 Chemical reaction scheme of in-situ synthesized dicalcium phosphate anhydrate in poly(lactic acid) solution.       64  5.2.2 Quantification of DCPA in DCPA/PLA scaffolds  The quantification of the inorganic phases in the DCPA containing scaffolds was verified using thermogravimetric analysis (TGA). Figures 5.13, 5.14, and 5.15 show the representative TGA graphs of the DCPA/PLA, DCPA/PLA/PEG, and Re-DCPA/PLA/PEG scaffolds, respectively. Mass loss starting at about 160 ºC caused by the evaporation of physically absorbed water molecules on the nanofibers increased with the inorganic phase contents in the scaffolds. The thermal degradation of the PLA occurred at about 300 ºC in the case of the Ref-PLA scaffold, but began at lower temperatures for the nanocomposites, correlated with the contents of the inorganic phases within the scaffolds. The nanocrystallites of the inorganic phases interfering with the arrangement of the PLA molecular chains seem to facilitate the thermal decomposition of the polymer. The thermal degradation of PEG is visible at around 350 ºC [245]. The DCPA/PLA/PEG scaffolds in Figure 5.14 (b and c) did not show weight loss at around 350 ºC, confirming that the PEG added into the DCPA/PLA nanocomposite suspension before electrospinning was removed after the washing-out process. The TGA tests were repeated three times on each scaffold for the calculation of the inorganic phases amount. The average weight percentages of amorphous calcium phosphate and dicalcium phosphate anhydrate within the nanocomposite scaffolds were summarized in Table 5.1. It was found that the addition of PEG (DCPA/PLA/PEG scaffolds) and the re-dissolving/electrospinning process (Re-DCPA/PLA/PEG scaffolds) did not significantly affect the amount of the reinforced inorganic phases within/on the PLA nanofibers. These values of the calcium phosphate    Table 5.1 Calcium phosphate content in DCPA/PLA scaffolds measured by TGA (n = 3).   Calcium phosphate content in scaffolds (wt %)      DCPA/PLA-2                               4.11 ± 0.25      DCPA/PLA-4                             10.29 ± 0.80      DCPA/PLA/PEG-2                               4.25 ± 0.36      DCPA/PLA/PEG-4                               9.99 ± 1.19        Re-DCPA/PLA/PEG-2                               5.41 ± 0.87      Re-DCPA/PLA/PEG-4                               9.53 ± 0.44  65  content were used to calculate areal density of the DCPA/PLA nanocomposite fibrous scaffolds for the measurement of tensile properties.       Figure 5.13 Representative TGA graphs of (a) Ref-PLA nanofibrous scaffold, (b) DCPA/PLA-2, (c) DCPA/PLA-3, and (d) DCPA/PLA-4 nanocomposite fibrous scaffolds.    66    Figure 5.14 Representative TGA graphs of (a) Ref-PLA nanofibrous scaffold, (b) DCPA/PLA/PEG-2, and (c) DCPA/PLA/PEG-4 nanocomposite fibrous scaffolds.     67    Figure 5.15 Representative TGA graphs of (a) Ref-PLA nanofibrous scaffold, (b) Re-DCPA/PLA/PEG-2, and (c) Re-DCPA/PLA/PEG-4 nanocomposite fibrous scaffolds.    5.2.3 Tensile properties of scaffolds Distribution of nanomaterials in polymer matrix and interfacial adhesion between nanomaterials and the polymer are the two major factors that determine the reinforcement effects of the nanomaterials. In this study, the effects of the reinforced DCPA nanophase on the mechanical properties under tension of the scaffolds were observed by measuring tensile strength, elastic modulus, and strain at fracture using a micro-tensile tester. Six samples of each scaffold were tested for a statistical analysis. Table 5.2 summarizes the mechanical properties of the all scaffolds tested. Figure 5.16 shows the tensile stress-strain   68  Table 5.2 Mechanical properties (in tension) of PLA and DCPA/PLA nanofibrous scaffolds (n = 6).     curves of the Ref-PLA scaffolds. The Ref-PLA sample had the average tensile strength of 24.78 ± 2.03 MPa, elastic modulus of 362 ± 21 MPa, and failure strain of 0.92 ± 0.09. The PLA only scaffold possessed non-comparable ductility to the other DCPA containing PLA nanocomposite fibrous scaffolds, as shown in the failure strain data in Table 5.2. They continued to elongate even further after the initial break and were completely fractures between 80 and 100 % strain. Figure 5.17 shows the tensile stress-strain curves of the in-situ synthesized and electrospun DCPA/PLA/PEG-2 scaffolds. While the scaffolds showed the similar tensile strength to the Ref-PLA scaffolds, they had an increased elastic modulus of 412 ± 11 MPa, but dramatically dropped to zero stress with the failure strain of 0.22 ± 0.05. Figure 5.18 shows the tensile stress-strain curves of the in-situ synthesized and electrospun DCPA/PLA/PEG-4 scaffold. The tensile strength and elastic modulus of the DCPA/PLA/PEG-4 scaffold were lower than those of the DCPA/PLA/PEG-2 scaffold. This result may be attributed to the increase of fiber diameters and broader range of fiber diameters of the DCPA/PLA/PEG-4 scaffold than the DCPA/PLA/PEG-2 scaffold. The brittleness of the DCPA/PLA/PEG scaffolds increased with the content of DCPA nanocrystallites, because the inorganic phases were located within and on the nanofibers and broke the continuous molecular PLA chains, resulting in surface defects around the DCPA nanocrystallites. Figure 5.19 and Figure 5.20 are the stress-strain curves of the Re-DCPA/PLA/PEG-2 and Re- DCPA/PLA/PEG-4 scaffolds, respectively. After re-dissolving and electrospinning processes of the DCPA/PLA/PEG scaffolds, the Re-DCPA/PLA/PEG   Tensile Strength (MPa) Elastic Modulus (MPa) Failure  Strain Ref-PLA 24.78 ± 2.03 362 ± 21 0.92 ± 0.09 DCPA/PLA/PEG-2 24.63 ± 1.36 412 ± 11 0.22 ± 0.05 DCPA/PLA/PEG-4 17.78 ± 1.11 374 ± 17 0.18 ± 0.03 Re-DCPA/PLA/PEG-2 25.76 ± 1.77 454 ± 20 0.35 ± 0.09 Re-DCPA/PLA/PEG-4 26.91 ± 1.34 534 ± 14 0.25 ± 0.03 Micro-Mech-DCPA/PLA-2 22.08 ± 1.71 398 ± 12 0.77 ± 0.10 Micro-Mech-DCPA/PLA-4 19.76 ± 1.21 416 ± 21 0.52 ± 0.04  69    Figure 5.16 Tensile stress-strain curves of Ref-PLA scaffold.  70    Figure 5.17 Tensile stress-strain curves of DCPA/PLA/PEG-2 scaffold.  71    Figure 5.18 Tensile stress-strain curves of DCPA/PLA/PEG-4 scaffold.  72    Figure 5.19 Tensile stress vs. strain curves of Re-DCPA/PLA/PEG-2 scaffold.  73    Figure 5.20 Tensile stress vs. strain curves of Re-DCPA/PLA/PEG-4 scaffold.  74  scaffolds had similar tensile strengths to the value of the Ref-PLA scaffold, but noticeable increase of the elastic moduli was observed. The graph patterns show that in the plastic deformation regions, these scaffolds were stiffer than the Ref-PLA scaffold. The Re-DCPA/PLA/PEG-4 scaffold showed the highest tensile strength of 26.91 ± 1.34 MPa and the highest elastic modulus of 534 ± 14 MPa compared to the other DCPA containing nanocomposite scaffolds and the neat PLA scaffold. The elastic modulus of the Re-DCPA/PLA/PEG-4 increased 47.5 % over the Ref-PLA. This result can be explained by the homogeneously dispersed and distributed DCPA nanocrystallites within the uniform PLA nanofiber matrix, as described in the section of 5.1.4. When matrix is under stress, the reinforced inorganic nanophases can take more mechanical loading from the matrix [157, 158]. When the load is applied to the Re-DCPA/PLA/PEG scaffolds, the DCPA nanocrystallites are believed to induce a stress transfer effect, which can enhance strength of the matrix. It has been known that the stress transfer from polymer matrix to nanophase fillers is carried by three main mechanisms; i) micromechanical interlocking, ii) chemical bonding between the nanophase and the matrix, and iii) weak van der Waals bonding between the nanophase and the matrix [159]. Even though the Re-DCPA/PLA/PEG scaffolds showed the much more brittle nature with the sharp breakage than the Ref-PLA scaffold, they were slightly further elongated, compared to the DCPA/PLA/PEG-2 and the DCPA/PLA/PEG-4 scaffolds, possibly due to the more uniform PLA nanofibrous structure.   Using commercial DCPA sub-micron particles, effects of the size and dispersion of DCPA phases on the tensile properties were compared with those of the Re-DCPA/PLA/PEG nanocomposite scaffolds. The commercial particles were mechanically mixed with the PLA solution and electrospun into scaffolds, referred to as Micro-Mech DCPA/PLA-2 and Micro-Mech DCPA/PLA-4, which contain similar amount of the DCPA phase to the other nanocomposite fibrous scaffolds. Figure 5.21 shows the presence of the DCPA nanoparticles (a) and the highly agglomerated chunk of the DCPA micron particles (b and c) on the surface of the PLA nanofibers. Figure 5.22 and Figure 5.23 show the tensile   75    Figure 5.21 SEM micrographs of Micro-Mech-DCPA/PLA-4 nanocomposite fibrous scaffold.    stress-strain curves of the Micro-Mech DCPA/PLA-2 and Micro-Mech DCPA/PLA-4 scaffolds, respectively. In comparison with the Re-DCPA/PLA/PEG scaffolds, the tensile strength and elastic moduli of the Micro-Mech-DCPA/PLA scaffolds were deteriorated. In case of the Micro-Mech DCPA/PLA-4, approximately 20 % of the values were reduced compared to those of the Re-DCPA/PLA/PEG-4. While the failure strain of the Micro-Mech-DCPA/PLA scaffolds decreased with the DCPA amount, they showed more ductile curve patterns than those of the Re-DCPA/PLA/PEG scaffolds. The overall curve patterns of the Micro-Mech-DCPA/PLA scaffolds were similar to those of the Ref-PLA scaffold, rather than those of the Re-DCPA/PLA/PEG scaffolds. The addition of the submicron  76    Figure 5.22 Tensile stress vs. strain curves of Micro-Mech-DCPA/PLA-2 scaffold.  77    Figure 5.23 Tensile stress vs. strain curves of Micro-Mech-DCPA/PLA-4 scaffold.  78  DCPA phase within/on the PLA nanofibers slightly decreased the tensile strength, but slightly increased the elastic modulus, as compared to the Ref-PLA scaffold. Previous studies by others [160,  161] reported that the calcium phosphate/biopolymer nanocomposite fibrous scaffolds, processed by mechanical mixing/electrospinning, produced deteriorated tensile strength and elastic modulus compared to the neat biopolymer scaffolds. Such opposite outcomes of the Re-DCPA/PLA/PEG and the Micro-Mech-DCPA/PLA scaffolds over the Ref-PLA scaffold demonstrate how important to control particle size and distribution of the reinforced nanomaterials within the polymer matrix for maximizing its effects for the mechanical properties. The chemical interactions between the reinforced DCPA nanocrystallites and the PLA nanofiber matrix, resulted from the in-situ synthesis route, is also believed to play a key role to enhance the mechanical properties by increasing interfacial shear stress at the interface. Xu et al. [161] fabricated uniform distribution of HAp nanoparticles within electrospun PLA nanofibers and induced better interaction/adhesion of the particles with the PLA matrix through the surface modification of the HAp with PLA. The PLA-grafted-HAp/PLA scaffold produced increased tensile properties over the neat PLA nanofibers and the HAp/PLA nanofibers without the surface modification. Figure 5.24 shows the representative tensile stress-strain curves obtained from the all PLA and DCPA/PLA scaffolds.    79    Figure 5.24 Representative stress-strain curves of PLA and DCPA/PLA scaffolds.  80  5.2.4 In-vitro bioactivity of DCPA/PLA scaffolds  Kokubo et al. [171] introduced a simulated body fluid (SBF) with the composition close to that of human blood plasma. The SBF solution has been used to evaluate in-vitro bioactivity of biomaterials over several decades [164‒171]. Bone-like apatite is formed on the surfaces of various bioactive materials in the body, such as bioactive glasses [167, 168], hydroxyapatite (HAp) [169], and bioactive polymers [170]. The bone-like mineral makes the bioactive materials favorable for bonding to bone tissue. Such biomineralization process is one of the key properties of bone tissue engineering scaffolds for successful bone regeneration [1, 3]. In this study, the Ref-PLA and Re-DCPA/PLA/PEG scaffolds were immersed in the SBF solution for 36 hrs and for 10 days, respectively, to assess in-vitro bioactivity of the scaffolds. Figure 5.25 (a‒c) shows SEM micrographs of the surface of the Ref-PLA, Re-DCPA/PLA/PEG-2, and Re-DCPA/PLA/PEG-4 scaffolds after 36 hrs of immersion. Cauliflower-like spherical crystals smaller than 1 µm formed on the surface of the Ref-PLA nanofibers. The presence of the crystals noticeably increased  on the nanocomposite scaffolds containing the DCPA nanocrystallites. After immersed for 10 days, the surface of the PLA nanofibers was almost covered with the larger crystals that were connected to each other (Figure 5.25 (d)). On the Re-DCPA/PLA/PEG scaffolds, the cauliflower-like precipitates lined the entire nanofiber surface, and even covered  all the porous spaces with overlapping of the spherical crystals. Figure 5.25 (f) shows that the crystals on the Re-DCPA/PLA/PEG-4 scaffold has sharper facets and are distributed in a more uniform size in comparison with those on the Ref-PLA scaffold. The phase of the precipitated crystals was analyzed using X-ray diffractometer (XRD). Figure 5.26 shows the diffraction patterns of the Ref-PLA (a), Re-DCPA/PLA/PEG-2 (b), and Re-DCPA/PLA/PEG-4 scaffolds, immersed in the SBF solution for 10 days. While broad diffraction signals at around 2θ = 32° were observed from the Ref-PLA scaffold, the intensity of the signals increased with the nanocomposite scaffolds containing the DCPA nanocrystallites. In case of the Re-DCPA/PLA/PEG-4 scaffold, weak characteristic peaks for HAp were observed as indicated in Figure 5.26 (c). Figure 5.27 shows energy-   81   Figure 5.25 SEM micrographs of (a, d) Ref-PLA, (b, e) Re-DCPA/PLA/PEG-2, and (c, f) Re-DCPA/PLA/PEG-4 scaffolds after immersed in simulated body fluid solution for 36 hours (a, b, c) and 10 days (d, e, f).      82    Figure 5.26 XRD patterns of (a) Ref-PLA, (b) Re-DCPA/PLA/PEG-2, and (c) Re-DCPA/PLA/PEG-4 scaffolds after immersed in simulated body fluid solution for 10 days. HAp peaks were identified and assigned from PDF # 01-072-1243.   dispersive X-ray spectroscopy (EDS) spectrum on the surface of the Re-DCPA/PLA/PEG-4 scaffold. The elemental analysis indicates that the surface is composed mostly of carbon, oxygen, calcium, and phosphorus. Carbon and oxygen are the main components of PLA, and hence calcium and phosphorus are believed to be detected from the crystals on the PLA nanofibers. The EDS analysis of the Ref-PLA scaffold, after immersed in SBF solution for 10 days, also showed similar spectrum result to that of the Re-DCPA/PLA/PEG-4 scaffold. Based on the SEM, EDS, and XRD analysis, the crystals formed on the nanofibers after incubation in the SBF solution are found to be similar to the   83            Figure 5.27 Energy-dispersive X-ray spectroscopy spectrum on the surface of Re-DCPA/PLA/PEG-4 scaffold after soaking in simulated body fluid solution for 10 days.  bone-like apatite. The cauliflower-like morphology is typical structure of the in-vitro mineralized apatite phase in a SBF solution [164‒171]. The crystal deposits with broad/weak X-ray diffraction patterns resemble those of poorly crystalline, nano-sized apatite present in bone tissue [162]. The mineralized layer of the crystals on the scaffolds after 10 days of the incubation indicates that steady deposition of the bone-like apatite from nucleation and crystallization sites proceeded. This result confirms that both the pure PLA scaffold and the DCPA containing scaffolds have bioactive potentials.  Elements Concentration        Carbon 71.71 at% 57.54 wt%        Oxygen 22.29 at% 23.82 wt%        Sodium   0.33 at%   0.50 wt%        Phosphorus   1.78 at%   3.67 wt%        Chlorine   0.47 at%   1.12 wt%        Calcium   3.03 at%   8.12 wt%        Gold   0.40 at%   5.22 wt%  84  Interestingly, the higher degree and faster rate of the mineralization was observed for the Re-DCPA/PLA/PEG scaffolds than for the Ref-PLA scaffolds. This reflects the abundant sites of bone-like apatite nucleation on the nanocomposite scaffolds. It is highly conceivable that the DCPA nanocrytallites on the PLA nanofibers function as nucleation sites and stimulate the bone-like mineral formation in the SBF solution. It has been demonstrated that on bioactive surface amorphous calcium phosphate forms and transforms into bone-like apatite in a SBF solution [163]. In the system of the Re-DCPA/PLA/PEG, it can be assumed that at pH of 7.4, the highly active DCPA dissolves and increases the ion concentrations of calcium and phosphorus near the surface of the PLA nanfibers, which would trigger the deposition of amorphous calcium phosphate and its transformation into the apatite phase. The increased apatite phase formation has been also reported by increasing local calcium and phosphate ion concentrations using the other reinforced calcium phosphates and bioactive glasses on biopolymers [164‒166]. Therefore, two possible mechanisms of the bone-like apatite formation on the DCPA/PLA nanocomposite scaffolds can be suggested as follows: i) formation of the amorphous calcium phosphate and its transformation into bone-like apatite in the supersaturated SBF solution. ii) dissolution and re-precipitation of DCPA into bone-like apatite in the supersaturated SBF solution.  In other studies, no or negligible amount of the mineralized crystals was formed on the electrospun PLA [164] and PLGA [21, 165, 166] nanofibers. Conversely, this study showed that the neat PLA scaffold also possesses comparable bioactivity to the DCPA/PLA nanocomposite scaffolds. This difference could be attributed to the samples preparation for the in-vitro bioactivity test. Unlike the others, the Ref-PLA and the DCPA/PLA/PEG scaffolds were soaked in, washed with de-ionized water twice using vacuum filtration, stored in a refrigerator for 24 hours, and lyophilized in order to remove the residual organic solvents. This process is believed to influence on the hydrolysis activity of the PLA  85  nanofibers. Zhang et al. [172] tested the effect of PLA hydrolysis in distilled water on in-vitro biomineralization in a SBF solution. It was reported that after immersion in distilled water at 37°C for 15 days before incubation in the SBF, the number of apatite particles formed on the PLA samples significantly increased, compared to the PLA samples without the hydrolysis treatment. The partially hydrolyzed PLA after the hydrolysis treatment is believed to generate negatively charged COOH and OH groups on the surface in the SBF. Through electrostatic force and hydrogen bonding, the calcium and phosphate ions in the SBF solution could be accumulated by COO−, COOH, and OH groups, which makes the nucleation of apatite phase possible on the hydrolyzed PLA surface.   5.3 Conclusions For the first time, DCPA was in-situ synthesized in this work in a non-aqueous PLA solution environment, which can be electrospun into nanocomposite fibers with intra-nanopores. The chemical interaction sites of the PLA/Ca2+ complexes served as the nucleation substrates for DCPA during the bio-mimetic synthesis, and induced improved dispersion of the DCPA nanocrystallites in the PLA nanofibers. This process avoided the severe agglomeration of DCPA nanoparticles processed by electrospinning the mechanically blended composite suspension. The incorporation of PEG, as a temporary co-polymeric additive, enabled greater stability in the formation of the electrospun jets, decreased the fiber diameter, and permitted more uniform fiber morphology, while increasing the amount of DCPA within the nanofibers. After re-dissolving and electrospinning process, the uniformly controlled PLA nanofibrous structure with the homogeneous distribution of the DCPA nanocrystallites was achieved, referred to as the Re-DCPA/PLA/PEG scaffolds. The tensile strength and elastic modulus of the Re-DCPA/PLA/PEG scaffolds were enhanced by the load transfer effect, compared to those of the neat PLA scaffold. The in-vitro bioactivity of the Re-DCPA/PLA/PEG scaffolds was found to greater than that of the neat PLA scaffolds.     86  Chapter 6 Hydroxyapatite/Alginate Nanocomposite Fibrous Scaffolds  6.1 Fabrication of HAp/alginate scaffolds  6.1.1 Electrospinning and cross-linking of alginate scaffolds Electrospinning sodium alginate solution is challenging as gelation starts at relatively low concentrations. At low concentrations, the solution does not contain a sufficient amount of alginate to generate continuous fibers during electrospinning. It was reported that sodium alginate forms a polyelectrolyte in aqueous solutions, and a repulsive force among the polyanions along the alginate molecular chains prevents entanglement of the chains during fiber formation [149]. Previous studies [32, 149] demonstrated that non-ionic poly(ethylene oxide) (PEO) and poly(vinyl alcohol) (PVA) are effective additives to control sol-gel transition and to reduce the solution’s viscosity, making electrospinning sodium alginate solution possible. In this work, sodium alginate was successfully electrospun into nanofibers with the addition of PEO and surfactant Triton X-100 (Refer to the sections of 4.2.2  and 4.2.3). As-electrospun Ref-Alginate nanofibers were randomly deposited into a non-woven nanofibrous scaffold with 3D interconnected pores, as shown in Figure 6.1 (a). The fiber diameter distribution analysis in Figure 6.1 (c) shows that fiber diameters were distributed between 50 nm to 250 nm with an average of 141 ± 45 nm. Figure 6.1 (b) shows that the nanofibers in the calcium Ref-Alginate scaffold were fused during the cross-linking treatment, which leads to a broader fiber diameter distribution, between 100 and 350 nm, and increased average fiber diameter, 207 ± 66 nm, as shown in Figure 6.1 (c). The sodium Ref-Alginate scaffold was cross-linked to calcium alginate using Ca2+ cations from the calcium nitrate tetrahydrate (CNT) aqueous solution, making the scaffold hydro-resistant in aqueous environments. It has been shown that divalent cations, such as Ca2+, Sr2+, and Ba2+, interact with the carboxyl functional groups of the GG blocks (sequential distribution of α-L-guluronic acid (G unit)) of sodium alginate [38]. According to the   87    Figure 6.1 SEM micrographs of (a) as-electrospun sodium Ref-Alginate and (b) cross-linked calcium Ref-Alginate scaffolds, and (c) fiber diameter distribution measurement of a as-electrospun sodium Ref-Alginate and b cross-linked calcium Ref-Alginate scaffolds.    88  model presented by Grant et al. [92], the divalent cations bridge the negatively charged GG blocks and form an “egg-box” structure, which encloses the cations and creates “junction zones” for the cross-linking of alginate.  Figure 6.2 (a) shows the FT-IR spectrum of PEO with the characteristic bands of OH at 3450 cm–1, CH at 2890 cm–1, and COC at 1100 cm–1. FT-IR spectra in Figure 6.2 (b–d) reveal the molecular interactions of sodium alginate and PEO before and after the cross-linking treatment. The broad hydroxyl band of sodium alginate at 3435 cm–1 in Figure 6.2 (b) shifted to a higher wavenumber, 3445 cm–1, with a sharper band both before and after      Figure 6.2 FT-IR spectra of (a) PEO, (b) sodium alginate, (c) as-electrospun sodium Ref-Alginate scaffold, and (d) cross-linked calcium Ref-Alginate scaffold.  89  cross-linking of the Ref-Alginate scaffold, as shown in Figure 6.2 (c and d). This shift is attributed to the interactions of sodium alginate and PEO through hydrogen bonding between the etheric oxygen of PEO and the hydroxyl groups of sodium alginate [32, 149]. Such molecular interactions were previously found to reduce the degree of ionization of sodium alginate in aqueous solutions with PEO, which helped increase entanglement of the sodium alginate chains and allowed successful electrospinning of the alginate/PEO solution. The characteristic peaks of asymmetric carboxyl (1625 cm–1) and symmetric carboxyl (1420 cm–1) bands of sodium alginate in Figure 6.2 (b) also shifted to higher wavenumbers, 1630 cm–1 and 1445 cm–1, for the cross-linked calcium Ref-Alginate scaffold in Figure 6.2 (d). This is possibly due to the interaction between the Ca2+ ions and carboxyl groups during the cross-linking treatment. The removal of PEO in the cross-linked calcium Ref-Alginate scaffold was confirmed by the disappearance of the finger print regions of PEO between 1400 cm–1 and 950 cm–1.  6.1.2 Fabrication of HAp/alginate scaffold via in-situ synthesis As-electrospun H-Alginate-5 and cross-linked/in-situ synthesized H-Alginate-5 nanofibrous scaffolds are shown in Figure 6.3 (a) and (b), respectively. The as-electrospun H-Alginate-5 was composed of random nanofibers similar to the as-electrospun Ref-Alginate (Figure 6.3 (a)), but contained homogeneously distributed PO43– ions as confirmed by EDS mapping of the P element  shown in Figure 6.4. Conversely, the cross-linked/in-situ  synthesized H-Alginate-5 clearly showed homogeneously deposited nanocrystals along the nanofibers. Due to the nanocrystals on the fused nanofibers, the average fiber diameter increased to 192 ± 50 nm from the 115 ± 28 nm average diameter of the as-electrospun fibers as indicated by the analysis of the fiber diameter distribution in Figure 6.5. The degree of nanocrystal deposition increased with the amount of SPM incorporated into the alginate nanofibers. Figure 6. 6 shows that the cross-linked/in-situ synthesized H-Alginate-2 scaffold possessed locally deposited nanocrystals, but in the H-Alginate-8 scaffold, the nanocrystals covered almost all plenary dimensions of the fibrous   90    Figure 6.3 SEM micrographs of (a) as-electrospun H-Alginate-5 and (b) cross-linked/in-situ synthesized H-Alginate-5 scaffolds.    91    Figure 6.4 EDS spectroscopy of as-electrospun H-Alginate-5 scaffold with P element area mapping.   scaffold like a coated film. The phase of the nanocrystals on the alginate nanofibers was confirmed by XRD analysis. No characteristic crystallographic peaks were observed in the XRD pattern of the cross-linked Ref-Alginate  (Figure 6.7 (a)). However, the XRD patterns in Figure 6.7 (b–d) show that the characteristic HAp peak at 2θ ≈ 32º was detected from the cross-linked/in-situ synthesized H-Alginate nanocomposite fibrous scaffolds. This poorly crystalline HAp formation is attributed to the alginate molecular chains suppressing and/or delaying the transformation of amorphous calcium phosphate to crystalline HAp by hindering the transportation of precursor ions during nucleation and crystal growth [141, 142].   92    Figure 6.5 Fiber diameter distributions of (a) as-electrospun H-Alginate-5 and (b) cross-linked/in-situ synthesized H-Alginate-5 scaffolds.         93    Figure 6.6 SEM micrographs of  cross-linked/in-situ synthesized (a) H-Alginate-2 and (b) H-Alginate-8 scaffolds.      94    Figure 6.7 XRD patterns of (a) cross-linked Ref-Alginate, cross-linked/in-situ synthesized (b) H-Alginate-2, (c) H-Alginate-5, and (d) H-Alginate-8 scaffolds. HAp peaks were identified and assigned from PDF # 01-072-1243.    95  6.2 Characterization of HAp/alginate scaffolds 6.2.1 Chemical interaction between HAp and alginate during in-situ synthesis The investigation of FT-IR spectra in Figure 6.8 also indicated the precipitation of HAp nanocrystals on the alginate nanofibers and suggests molecular level interactions between the HAp and alginate during the cross-linking and in-situ synthesis treatments. Figure 6.8 (b–d) show that the precipitated nanocrystals on the nanofibers are HAp with phosphate bands at approximately 1030 cm–1 (PO4(v3)), 950 cm–1 (PO4(v1)), 600 cm–1 (PO4(v4)), and 565 cm–1 (PO4(v4)). As displayed by the dotted circles, the double split of the PO4(v4) bands is typical of crystalline HAp [24, 25]. The asymmetric carboxyl (1630 cm–1) and symmetric carboxyl (1445 cm–1) bands of the cross-linked calcium Ref-Alginate in Figure 6.8 (a) shifted to lower wavenumbers, 1600 cm–1 and 1417 cm–1, 1592 cm–1 and 1413 cm–1, and 1590 cm–1 and 1410 cm–1, for the cross-linked/in-situ synthesized H-Alginate-2, H-Alginate-5, and H-Alginate-8 nanocomposite scaffolds, respectively. It appears that the formation of [–COO–]–Ca2+–[–COO–] linkage sites in the alginate “egg box” structures serve as nucleation substrates for HAp, resulting in the shift of the carboxyl bands. These shifts were also presented in previous studies on the chemical interactions between HAp and the functional groups of biopolymers [50, 142].  Figure 6.9 illustrates the mechanism of the in-situ synthesized HAp formation on the alginate nanofibers. Based on the results of SEM, XRD, and FT-IR in this study, it is conceivable that the as-electrospun H-Alginate nanofibers containing PO43– ions cross-link into calcium alginate in the aqueous Ca2+ solution at pH 10 (Figure 6.9 (a)); PO43– ions (from the alginate fibers), Ca2+, and OH– ions (from the aqueous environment) diffuse to the [–COO–]–Ca2+–[–COO–] linkages for HAp nucleation (Figure 6.9 (b)). Thus, the HAp nanocrystals precipitate and grow on the specific nucleation sites along the alginate nanofibers that function as a template for in-vitro biomineralization (Figure 6.9 (c and d)), mimicking the process, structure, and chemical composition of the mineralized collagen fibrils in bone tissue. The novelty of the present work relates to the in-situ HAp formation,  96    Figure 6.8 FT-IR spectra of (a) cross-linked Ref-Alginate, cross-linked/in-situ synthesized (b) H-Alginate-2, (c) H-Alginate-5, and (d) H-Alginate-8 scaffolds.    97   Figure 6.9 Chemical structure of (a) “egg box” model of calcium alginate, (b) “egg box” model of calcium alginate with precursor ions for HAp nucleation, and (c) mineralized “egg-box” structure with HAp, and (d) illustration of cross-linked/in-situ synthesized HAp/alginate nanocomposite fibrous scaffold.   98  in contrast to the other reported methods [150−155]. Figure 6.3 and Figure 6.6 show that the amount of the precipitated HAp nanocrystals on the alginate nanofibers increased with  the SPM concentrations in the electrospinning solutions. This phenomenon results from the increase of super-saturated degree of PO43− ions around the [–COO–]–Ca2+–[–COO–] linkage sites on the alginate nanofibers. The electrospinning of a mechanically blended biopolymer solution with pre-processed CaP nanoparticles has been commonly used to fabricate nanocomposite fibers. However, nanoparticles tend to flocculate in solution due to surface charge effects, and agglomeration of the nanoparticles in the biopolymer solution before and during electrospinning causes irregular distribution of the aggregates within/on the fibers [21, 22, 156]. It is evident that the biomimetic approach of the current work for fabricating HAp/alginate nanocomposite fibrous scaffolds induced a more homogenous and uniform distribution of the HAp nanocrystals on the alginate nanofibers (Figure 6.3 (b)) than the conventional mechanical mixing/electrospinning method which resulted in the severely agglomerated HAp nanoparticles at micrometer levels (Figure 6.10).     Figure 6.10 SEM micrograph of cross-linked Mech-H-Alginate-2 scaffold fabricated using a mechanical blending/electrospinning method. The white arrows indicate the agglomerated HAp particles at micro-meter levels.  99  6.2.2 Quantification of HAp in HAp/alginate scaffolds The qunatification of the HAp phase in the cross-linked/in-situ H-Alginate nanocomposite fibrous scaffolds was verified using thermogravimetric analysis (TGA). Figure 6.11 shows representative TGA graphs of the cross-linked Ref-Alginate and the cross-linked/in-situ synthesized H-Alginate-2, H-Alginate-5, and H-Alginate-8 scaffolds. Mass loss starting at about 100 ºC was caused by the evaporation of physically absorbed  moisture on the nanofibers. The rapid weight loss occurred at between 200 ºC and 300 ºC due to the preliminary degradation of the alginate [173]. The mass loss kept slowly at the higher temperatures due to the further decomposition of the alginate. The mass change began to be stabilized at about 800 ºC. It was reported that metal alginate decomposes and produces the residue mainly containing corresponding metal carbonate and also metal oxide at 800 ºC [173, 174]. In this study, it was hypothesized that the residue of the cross-linked/in-situ synthesized H-Alginate scaffolds contains mainly calcium carbonate and calcium oxide with hydroxyapatite at 900 ºC. Therefore, the calculation of the HAp phase amount was carried out by subtracting the Ref-Alginate residue wt% from the H-Alginate residue wt %. The temperature, 900 ºC, was chosen because the mass loss of a few samples was not completely stabilized at 800 ºC. The TGA tests were repeated three times on each scaffold. The average weight percentages of the HAp phase within the nanocomposite scaffolds were summarized in Table 6.1.  The nanocomposites were found to contain 4.25 ± 2.40, 12.49 ± 2.94, 19.04 ± 1.17 wt % of the HAp within the H-Alginate-2, H-Alginate-5, and H-Alginate-8 scaffolds, respectively. These values were used to calculate areal density of the HAp/alginate nanocomposite fibrous scaffolds for the measurement of tensile properties.  Table 6.1 HAp content in cross-linked/in-situ synthesized H-Alginate scaffolds measured by TGA (n = 3).       HAp content in scaffolds (wt %)      H-Alginate-2                               4.25 ± 2.40      H-Alginate-5                             12.49 ± 2.94      H-Alginate-8                             19.04 ± 1.17  100    Figure 6.11 TGA patterns of (a) crossed-linked Ref-Alginate, cross-linked/in-situ synthesized (b) H-Alginate-2, (c) H-Alginate-5, and (d) H-Alginate-8 scaffolds.    6.2.3 Tensile properties of HAp/alginate scaffolds Mechanical properties under tension of the alginate and HAp/alginate scaffolds were measured using the same testing methodology as that used for the DCPA/PLA scaffolds. This was done to observe the effects of the HAp nanocrystals on the alginate nanofibers. Table 6.2 summarizes the tensile strength, elastic modulus, and failure strain of the all scaffolds tested. The Mech-H-Alginate-2 and Mech-H-Alginate-5 scaffolds were also evaluated in order to compare the effects of HAp size and distribution on the tensile properties with the cross-linked/in-situ synthesized H-Alginate scaffolds. Figure 6.12 shows the tensile stress-strain curves of the cross-linked Ref-Alginate scaffold. The Ref-   101  Table 6. 2 Mechanical properties (in tension) of cross-linked Ref-Alginate and cross-linked/in-situ synthesized H-Alginate scaffolds (n = 6).     Alginate scaffold had average tensile strength of 27.45 ± 2.73 MPa, elastic modulus of 0.654 ± 0.075 GPa, and failure strain of 0.0347 ± 0.0026. The alginate only scaffold possessed the most ductile properties over the other HAp containing alginate scaffolds, as shown in the failure strain data in Table 6.2. The tensile strength constantly increased and vertically dropped to zero at failure. Figure 6.13 shows the tensile stress-strain curves of the cross-linked/in-situ synthesized H-Alginate-2 scaffold. While the tensile strength of the scaffold slightly increased, the elastic modulus increased to 1.114 ± 0.171 GPa, 70.3 % increase compared to that of the Ref-Alginate. However, the scaffold dropped to zero stress with the failure strain of 0.0233 ± 0.0022, 32.9 % decrease compared to that of the Ref-Alginate. Figure 6.14 shows the tensile stress-strain curves of the cross-linked/in-situ synthesized H-Alginate-5 scaffold. This HAp/alginate scaffold had the highest tensile strength and elastic modulus among the scaffolds tested. The tensile strength increased 79.3 %, and the elastic modulus increased 158.4 %, compared to those of the Ref-Alginate scaffold. Interestingly, the H-Alginate-5 scaffold broke at the higher failure strain than that of the H-Alginate-2 scaffold, even though the initial breakage occurred at the failure strain between 0.001 and 0.002. The tensile stress kept increasing, while overcoming further several breakages as shown in Figure 6.14. Such noticeable enhancement of the mechanical properties can be attributed to the homogeneously dispersed and distributed HAp nanocrytals on the alginate nanofibers, as described in the section of 6.1.3. When matrix is under stress, the reinforced inorganic nanophases can take more mechanical loading from the matrix [157, 158]. When the load is applied to the H-Alginate scaffolds,  the HAp  Tensile Stress (MPa) Elastic Modulus (GPa) Failure  Strain   Ref-Alginate 27.45 ± 2.73 0.654 ± 0.075 0.0347 ± 0.0026   H-Alginate-2 32.09 ± 1.81 1.114 ± 0.171  0.0233 ± 0.0022   H-Alginate-5 49.23 ± 2.09 1.690 ± 0.208 0.0292 ± 0.0031   Mech-H-Alginate-2 18.54 ± 1.74 1.023 ± 0.092 0.0217 ± 0.0026   Mech-H-Alginate-5   8.77 ± 1.97 0.934 ± 0.114 0.0087 ± 0.0010  102    Figure 6.12 Tensile stress-strain curves of cross-linked Ref-Alginate nanofibrous scaffold.  103    Figure 6.13 Tensile stress-strain curves of cross-linked/in-situ synthesized H-Alginate-2 nanocomposite fibrous scaffold.  104    Figure 6.14 Tensile stress-strain curves of cross-linked/in-situ synthesized H-Alginate-5 nanocomposite fibrous scaffold.  105  nanocrystals are believed to induce a stress transfer effect, which can enhance strength of the matrix. It has been known that the stress transfer from polymer matrix to nanophase fillers is carried by three main mechanisms; i) micromechanical interlocking, ii) chemical bonding between the nanophase and the matrix, and iii) weak van der Waals bonding between the nanophase and the matrix [159]. More detailed studies are required to unveil mechanisms of the HAp nanocrystals for increasing the mechanical properties of the HAp/alginate scaffolds.    Effects of HAp size and dispersion on the mechanical properties in tension were observed by comparing two distinctive HAp/alginate composites; i) the Mech-H-Alginate scaffolds and  ii) the cross-linked/in-situ synthesized H-Alginate scaffolds. The Mech-H-Alginate scaffolds, processed by mechanical mixing and electrospinning of the pre-synthesized HAp nanocrystals with the alginate solution, contains the locally distributed HAp particles with the broad range of sizes from nano to micro levels, as shown in Figure 6.10. Figure 6.15 and 6.16 show the tensile stress-strain curves of the Mech-H-Alginate-2 and the Mech-H-Alginate-5 scaffolds, respectively. The tensile strength of the Mech-H-Alginate-2 was 18.54  ± 1.74 MPa, almost half value of the cross-linked/in-situ synthesized H-Alginate-2 scaffold. Unlike the H-Alginate nanocomposite scaffolds, the mechanical properties of the Mech-H-Alginate scaffolds were deteriorated with increasing the HAp amount. The Mech-H-Alginate-5 scaffold had the lowest tensile strength and failure strain among the all scaffold tested. The elastic modulus was higher than that of the Ref-Alginate, but lower than any other HAp containing scaffolds. Furthermore, it was the most brittle with the failure strain of 0.0087 ± 0.0010, more than 50 % decrease compared to the other scaffolds. The non-uniformly distributed and agglomerated HAp particles, processed by the mechanical mixing/electrospinning, produced deteriorated the tensile strength and elastic modulus of the Mech-H-Alginate scaffolds compared to the homogeneously distributed HAp nanocrystal containing alginate scaffolds. This result also demonstrates how important to control particle size and distribution of the reinforced nanomaterials within the   106    Figure 6.15 Tensile stress-strain curves of cross-linked Mech-H-Alginate-2 nanocomposite fibrous scaffold.  107    Figure 6.16 Tensile stress-strain curves of cross-linked Mech-H-Alginate-5 nanocomposite fibrous scaffold.  108  polymer matrix for maximizing its effects for the mechanical properties. It is also highly considered that the chemically interacted deposition of the HAp nanocrystals on the alginate nanofibers, through the biomimetic in-situ synthesis, would play an important role to enhance the tensile properties of the H-Alginate nanocomposite fibrous scaffolds. Further characterization studies are required to investigate what toughening mechanism play a major role for the increased strength and modulus of the H-Alginate scaffolds.   Figure 6.17 shows the representative tensile stress-strain curves obtained from all alginate and HAp/alginate fibrous scaffolds. There are two types of bone; cortical and cancellous bone. Cortical bone has tensile strength in the range of 50 − 151 MPa, compressive strength in the range of 130 ‒ 180MPa, and elastic modulus of 12 ‒ 18 GPa [6, 39, 40]. On the other hand, cancellous bone has compressive strength in the range of 5 − 10 MPa and elastic modulus of 50 − 100 MPa [236, 237]. The tensile strengths and elastic moluli of the DCPA/PLA and the HAp/alginate nanocomposite fibrous scaffolds are in the range between cancellous bone and cortical bone tissues. Compared to the values of the PLA-based scaffolds (Table 5.2), the in-situ synthesized HAp/algiante scaffolds had much higher tensile strength and elastic modulus, but lower failure strain (Table 6.2).  Representatively, the H-Alginate-5 scaffold had 82.9 % higher strength, 216 % higher elastic modulus, but 88.3 % lower failure stain than those of the Re-DCPA/PLA/PEG-4 scaffold.     109     110  Figure 6.17 Representative tensile stress-strain curves of alginate and HAp/alginate nanofibrous scaffolds. 6.2.4 In-vitro bioactivity of HAp/alginate scaffolds  Bioactivity and biodegradability of scaffolds are the key necessary requirements for bone tissue regeneration to make them bonded with natural bone tissues. Calcium phosphates, representatively hydroxyapatite, have been added into a biodegradable polymer matrix to meet these requirements [175]. However, conventional composites of calcium phosphate particles and biopolymers resulted in limited functionality of the bioactive phase, because most of the particles were embedded in the polymer matrix. Hydroxyapatite coating technology on organic  and metal substrates has received much attention, since it increases exposed surface area of the phase combined with the matrix. Kokubo and his co-workers [176−178]  developed a biomimetic process to coat hydroxyapatite layers on the substrates using a reaction between bioactive glass and a simulated body fluid (SBF). The bone-like hydroxyapatite layers on the surface make the bioactive materials favorable for bonding to bone tissue [167−170]. In this study, hydroxyapatite/alginate nanocomposite system was developed using the novel biomimetic in-situ synthesis route  without a SBF solution with thecomposition close to that of human blood plasma. The bioactivity of the pure alginate and the hydroxyapatite/alginate scaffolds was compared using the same SBF solution utilized in the bioactivity test of the DCPA/PLA nanocomposite fibrous scaffold in section 5.2.4. The cross-linked Ref-Alginate and the across-linked/in-situ synthesized H-Alginate scaffolds were immersed in the SBF solution for 36 hrs and 10 days, respectively. Figure 6.18 shows SEM micrographs of the surface of the Ref-Alginate (a) and the H-Alginate 5 (b) scaffolds after 36 hours of immersion. In case of the pure alginate scaffold, the nanofibrous structure on the surface collapsed and fused into a film, as shown in the magnified image. On the other hand, the non-woven naonfibrous structure with the coated HAp nanoparticles, in case of the hydroxyapatite/alginate scaffold, partly collapsed and changed into uneven and bumpy surface with cauliflower-like spherical crystals in around 500 nm sizes. The phase of the precipitated crystals was analyzed using X-ray diffractometer (XRD). Figure 6.19 shows the diffraction patterns of the Ref-Alginate (a)  111  and the H-Alginate-5 scaffolds (b), immersed in the SBF solution for 10 days. One    Figure 6.18 SEM micrographs of (a) cross-linked Ref-Alginate nanofibrous scaffold  and (b) cross-linked/in-situ synthesized H-Alginate-5 nanocomposite fibrous scaffold after immersed in simulated body fluid solution for 36 hours.    112     Figure 6.19 XRD patterns of (a) cross-linked Ref-Alginate nanofibrous scaffold  and (b) cross-linked/in-situ synthesized  H-Alginate-5 nanocomposite fibrous scaffold after immersed in simulated body fluid solution for 36 hours. HAp peaks were identified and assigned from PDF # 01-072-1243.   diffraction signal around  at 2θ = 31.5° was observed from the H-Alginate scaffold, while no characteristic peaks were found from the Ref-Alginate scaffold. The characteristic cauliflower-like HAp crystals in the H-Alginate scaffold were much smaller and less precipitated than those of the DCPA/PLA nanocomposite scaffolds in Figure 5.25. This might cause the less visible peak on the XRD diffraction pattern.          113   The cross-linking of alginate with bivalant metal cations, such as Ca2+, Ba2+, and Sr2+ is attributed to the interaction of two carboxyl groups on the alginate chains with the cations at the "egg-box" junction zones [38, 92]. The cross-linked alginate structure is stable in water, but in a SBF solution with sufficient Na+ ions the dissolution of the cross-lined alginate structure occurs due to the exchange of Na+ ions in the solution with Ca2+ ions bound to the carboxyl groups in the alginate chains [179]. This study also showed that the pure alginate nanofibers cross-linked with Ca2+ ions dissolved and lost its non-woven mat structures. However, it is worth mentioning that the surface modified alginate nanofibers with the HAp nanocrystals in the cross-linked/in-situ synthesized H-Alginate-5 scaffold survived in the SBF solution for 36 hrs. It is believed that the HAp nanocrystals formed on the alginate nanofibers obstruct the ion exchange process between Na+ ions and Ca2+ ions, resulting in retaining the structural integrity of the nanofibrous structure in the SBF solution. Other investigators used a double cross-linking method, cationic cross-linking and covalent cross-linking, to increase stability of the alginate structure in aqueous environments. The covalent cross-linking at hydroxyl and carboxyl functional groups in alginate chains  was made possible using epichlorohydrin, glutaraldehyde, methylene bis(4-cyclohexylisocyanate) (HMDI), and acetal diacrylate (ADA) cross-linkers [180−184]. However, toxicity of the cross-linkers are in question for their use in biomaterials. The poorly crystalline HAp precipitates were only observed only from the cross-linked/in-situ synthesized H-Alginate-5 scaffold. The HAp nanocrystals coating on the alginate nanofibers through the novel in-situ synthesis strategy are believed to function as substrates for heterogeneous nucleation and crystal growth of the HAp precipitates. The in-vitro HAp precipitate formation in the SBF solution showed visible difference betweeen the PLA based scaffold and the alginate-based scaffolds. The size and crystallinity of the cauliflower-like HAp precipitates on the HAp/alginate scaffolds were smaller and lower than thouse of the PLA-based scaffolds, respectively. The unstable structural construct of the alginate in the sodium ions containing solution (SBF solution) is believed to prevent  114  crytal growth of the HAp precipiates with time. Even though the pure alginate nanofibers have the functional hydroxy and carboxyl groups on the surface, HAp precipitation was not induced probably due to the stably cross-linked "egg-box" structures at the carboxyl sites at the early stages of the immersion and the unstable surface of the alginate nanofibers with time. Hosoya et al. [185] also observed no formation of HAp precipiates on pure alginate gel, but silane modified alginate gel provided nucleation sites for HAp.            6.3 Conclusions HAp/alginate nanocomposite fibrous scaffolds were successfully fabricated by the biomimetic in-situ synthesis of HAp on electrospun alginate nanofibers. The alginate nanofibers were cross-linked by Ca2+ ions at GG blocks (according to “Egg box” model), which is believed to serve as nucleation sites for HAp. While the scaffolds were agitated in the solution at a pH = 10, impregnated PO43– (from the fibers), Ca2+, and OH– (from the solution) ions diffused towards the [–COO–]–Ca2+–[–COO–] linkage sites for the nucleation and growth of the HAp nanocrystals along the alginate nanofibers. This novel processing method induced homogenous deposition of HAp nanocrystals on the nanofibers, unlike the severely agglomerated, micrometer-large HAp particles processed by the conventional mechanical blending/ electrospinning method. The tensile strength and elastic modulus of the HAp/alginate nanocomposite fibrous scaffold increased with the HAp content. This is believed to be due to the stress transfer effect. The cauliflower-like HAp crystals precipitated on the surface of the HAp/alginate scaffold, demonstrating in-vitro bioactivity of the nanocomposite unlike the pure alginate scaffold. It is also worth mentioning that the surface modified alginate nanofibers with the HAp nanocrystals in the cross-linked/in-situ synthesized H-Alginate-5 scaffold survived in the SBF solution for 36 hrs. The pure alginate nanofibers cross-linked with Ca2+ ions dissolved and lost its non-woven structures.  115   Chapter 7 In-vitro Osteoblasts Response on Scaffolds  7.1 Dicalcium phosphate anhydrate/poly(lactic acid) scaffolds 7.1.1 Attatchment of osteoblasts on scaffolds  Rat calvarial osteoblast (RCO) cell morphology on the PLA nanofibers with the different surface chemistry (PLA vs. DCPA/PLA) was studied in vitro for 7 days (Refer to the section and its subsections of 4.4.). The morphological response and interaction between the cells and the nanofibers were observed using scanning electron microscopy (SEM). Figure 7.1 shows representative morphology of the RCO cells cultured on the Ref-PLA scaffold and the Re-DCPA/PLA/PEG-4 nanocomposite fibrous scaffold for 7 days. There was no significant difference of the cell morphology between the two samples. The surface modification of the PLA nanofibers with the DCPA nanocrystallites did not affect the morphological response of the RCO cells. The osteoblasts attached and spread on the surface without any preferred orientation on the nanofibrous surface at day 1. The cells migrated aside on the surface, fused through the nanofibers, and even migrated under the layers of the nanofiber network at day 3. It was also observed that the cells interacted and integrated well with the surrounding nanofibers. The inset SEM micrographs of Figure 7.1 (b) show that the cell growth was guided by the nanofiber topography, and the multiple filopodial connections to the surface were made along the nanofibers, resulting in stable and close adhesion of the RCO cells. At day 7 of post cell-seeding, the cells reached confluence on the surface, forming multi-cellular network. The morphology of the RCO cells on the scaffolds was also visualized using an immunostaining method. Figure 7.2 shows representative immunofluorescent images of the RCO cells cultured on the Ref-PLA scaffold and the Re-DCPA/PLA/PEG-4 scaffold for 1 day and 7 days. Actin filaments of the osteoblast cells, one of the primary cytoskeletal components, were labeled red with phalloidin. Nuclei of the cells were labeled blue with DAPI. These immunostaining images  116  support the SEM results of the RCO cell adhesion morphology on the scaffolds. The cells    Figure 7.1 SEM micrographs of RCO cells cultured on (a) Ref-PLA and (b) Re-DCPA/PLA/PEG-4 scaffolds for 7 days. The magnified inset SEM micrographs of the DCPA/PLA scaffold shows closely and stably attached and spread RCO cells on the electrospun nanofibrous structure at day 3 of post cell-seeding. The multiple filopodial connections on the surface were made along the nanofibers.  stably attached with no preferred orientation at day 1. The confluent growth of the multi layered RCO cells on the scaffolds was also observed, as shown in Figure 7.2 (b) and (d). The bio-inspired nanofibrous architecture, produced by electrospinning, has been reported to enhance attachment, adhesion, and proliferation of cells [11, 12]. Cell adhesion and morphology on the surface of scaffold play important roles for cellular activities. More  117  detailed explanation will be discussed in the next section of 7.2.    Figure 7.2 Immunofluorescence images of RCO cells cultured on (a) Ref-PLA scaffold for 1 day, (b) Ref-PLA for 7 days, (c) Re-DCPA/PLA/PEG-4 scaffold for 1 day, and (d) Re-DCPA/PLA/PEG-4 scaffold for 7 days. Actin filaments were labeled red with phalloidin  118  (left), nuclei were stained blue with DAPI (middle), and the two fluorescent images were merged (right).    7.1.2 Proliferation of osteoblasts on scaffolds An ideal tissue engineering scaffold should be mechanically stable and capable of functioning biologically, in order to induce host cells for successful tissue regeneration. General properties required for design of tissue engineering scaffolds are biocompatibility, biodegradability with nontoxic by-products, and adequate porosity to allow for cell infiltration and vascularization and to make diffusion of nutrients and wastes easier. 75 to 150 µm pore diameters are required for new bone tissue ingrowth into scaffold. For vascular graft, the effective pore diameter for cell ingrowth are between 20 and 60 µm [238]. Specific properties required for bone tissue engineering scaffolds are bioactivity (ability of a material to integrate with the natural bone tissue), osteoconductivity (ability of a material for bone-forming cells to attach, migrate, grow, and form new bone tissue), and osteoinductivity (ability of a material to induce bone tissue regeneration) [1, 6, 204]. Biocompatibility can be classified into surface and structural biocompatibility [12]. Surface biocompatibility is associated with the surface chemistry of the material. The surface chemistry is closely related to the adsorption of biological molecules and the cellular activities such as adhesion and migration [205, 206]. Structural biocompatibility is associated with the architecture and dimensions of the material. It was reported that cell population is influenced by the scale of architectural structure, and cell adhesion by the topography of the material  [207−209]. An MTS cell proliferation assay was used to assess cultured RCO cells viability on the PLA and PLA/DCPA nanocomposite scaffolds. Effects of the electrospun nanofibrous topography on the proliferation of the RCO cells were also observed by comparing the Ref-PLA scaffold with the cast PLA film.   The working principle of an MTS assay is as follows. Living cells produce oxidoreductase enzymes, which reduce the yellow tetrazolium dye of MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3 carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium) to a purple formazan product in the presence of phenazine methosulfate. The absorbance of the colored solution  119  is quantified by measuring at 490 to 500 nm wavelength by a spectrophotometer. The  degree of the light absorption is proportional to the number of living cells [200]. The MTS assay results of the RCO cells seeded on the scaffolds at 2 × 104 cells/mL and cultured for 7 days are shown in Figure 7.3. The average values of optical density were produced from triplicate samples of each scaffold. Statistical significance was selected at p < 0.05, using one-way ANOVA for the specimens at each day. The PLA film, fabricated by solvent casting and drying, was used as another control sample to compare cell attachment and proliferation with the electrospun nanofibrous structure of PLA. Overall, the viable cells on each scaffold increased with time for 7 days of culture periods. The RCO cells proliferated much faster on the electrospun PLA nanofibrous structure over the film structure, as shown in Figure 7.3 (a and b). At day 1 of the post cell-seeding, it was found that the optical density of the PLA film was the lowest among all the scaffolds tested. After 7 days of culture, the optical density of the Ref-PLA scaffold was about twice higher than that of the      Figure 7.3 MTS assays of cultured RCO cells on (a) PLA film, (b) Ref-PLA, (c) Re-DCPA/PLA/PEG-2, (d)  Re-DCPA/PLA/PEG-4, and (e) Micro-Mech-4 scaffolds for 7  120  days. Triplicate samples were tested for each scaffold. Statistical significance was selected at p < 0.05, using one-way ANOVA for the specimens at each day.  PLA film. The incorporated DCPA nanocrystallites within/on the PLA nanofibers were found to promote growth of the RCO cells. The Re-DCPA/PLA/PEG-4 nanocomposite fibrous scaffold showed the highest optical densities among the samples at day 3 and 7. The RCO cells proliferated faster on the Re-DCPA/PLA scaffolds with the DCPA content, as shown in Figure 7.3 (b−d). The cells proliferation on the DCPA/PLA scaffolds depending on the size and distribution of the DCPA crystals was compared using the Re-DCPA/PLA/PEG-4 and the Micro-Mech-4 scaffold. The two scaffolds contained similar amount of DCPA, but the mechanically mixed/electrospun Micro-Mech-4 scaffold possessed highly agglomerated DCPA crystals over the size of sub-micrometers irregularly distributed on the surface of the scaffold (Figure 5.21). It was found that from day 3 of the post cell-seeding, the RCO cells on the Re-DCPA/PLA/PEG-4 scaffold grew faster than the Micro-Mech-4 scaffold, while no significant difference was observed at day 1. The optical densities of the Micro-Mech-4 were not significantly different from the values of the Ref-PLA throughout the culture periods. The RCO cells proliferation on the different surface topographies (non-woven nanfibers vs. film) and the different surface chemistry (PLA vs. DCPA/PLA) was tested, with another method using a DAPI stained cell number counting. Figure 7.4 shows the average number of the RCO cells proliferated on the scaffolds for 7 days. Nuclei of the RCO cells were stained with DAPI and visualized with fluorescent microscope. Six images per scaffold were taken, and the nuclei of the cells were counted using ImageJ software. Triplicate samples were prepared for each scaffold. Figure 7.5 demonstrates representative fluorescent images of the stained nuclei of the RCO cells cultured on the scaffolds for 7 days. This cell number counting studies also demonstrated that the electrospun PLA nanofibrous topography promoted faster RCO cells growth than the PLA film surface, as indicated in Figure 7.4 (a) and (b). Electrospinning produces non-woven nanofibers with 3D interconnected pores and a high surface area that mimic the natural extra-cellular matrix (ECM) of human tissues, and it is of great interest  121  in tissue regenerative medicines [2, 12, 13]. Poly(lactic acid) (PLA) has been electrospun into scaffolds for tissue engineering. The unique nanofibrous topography is favorable for    Figure 7.4 Average number of RCO cells cultured on (a) PLA film, (b) Ref-PLA, (c) Re-DCPA/PLA/PEG-2, (d)  Re-DCPA/PLA/PEG-4, and (e) Micro-Mech-4 scaffolds for 7 days. Triplicate samples were tested for each scaffold. Nuclei stained RCO cells with DAPI were counted using ImageJ software. Six images per scaffold were taken, and triplicate samples were prepared for each scaffold.   cell attachment, diffusion of oxygen and nutrients, vascularization, and loading high quantities of pharmaceutical agents. It is anticipitated that the electrospun PLA nanofibers could make the degradation products of lactic acid and oligomers quickly escape from the  surface to neighboring pores, and eventually out of the scaffold, so that the possibility of adverse tissue response from local acidity would be limited [13, 17, 31]. Figure 7.5 (b)  122  shows that the entire surface of the Ref-PLA scaffold was covered by the populated RCO cells at day 7 of post cell-seeding. The cell number was the highest on the Re-   Figure 7.5 Fluorescent images of DAPI stained nuclei of RCO cells cultured on (a) PLA film, (b) Ref-PLA, (c) Re-DCPA/PLA/PEG-2, (d)  Re-DCPA/PLA/PEG-4, and (e) Micro-Mech-4 scaffolds for 7 days.     123    DCPA/PLA/PEG-4 scaffold after 7 days of culture, as shown in Figure 7.4 (d) and Figure 7.5 (d). The cell number on the PLA scaffolds was positively correlated with the DCPA concentration except the Micro-Mech 4 scaffold, which is consistent with the results of the MTS assay. It was reported by many other ressearchers that osteogenic cell activities were stimulated by adding calcium phosphate nanocrystals into polymeric nanofibrous scaffolds, representatively HAp. Li et al. [20] produced silk fibroin fibrous scaffolds containing bone morphogenetic protein-2 (BMP-2) and HAp nanoparticles which generated in vitro bone formation from human bone marrow-derived mesenchymal stem cells (hMSC). Cotton wool-like PLGA fibers, including amorphous tricalcium phosphate nanoparticles, demonstrated bioactivity and osteogenic differentiation from hMSC [21]. Nie and Wang  [22] studied the release of BMP-2 plasmid DNA from an electrospun PLGA blend with HAp nanoparticles. This can be explained by upregulated expression of genes by HAp [210], direct mitogenic effects of HAp crystals [211], and also by the effects of HAp reinforcement functioning as s stronger support for cell colonization [212, 213]. Therefore, the enhanced proliferation of the ROC cells on the Re-DCPA/PLA/PEG scaffolds observed in this study,  is believed to be attributed to the reinforced DCPA nanocrystallites within the PLA nanofibers.   Recently, dicalcium phosphate anhydrate (DCPA) and dicalcium phosphate dihydrate (DCPD) have drawn attention as a result of their osteoinductivity and bioresorbability. They possess stable phases under acidic conditions (pH < 4.8), and thus dissolve and/or hydrolyze to HAp depending on the implanted sites and physiological conditions [67]. In the in-vivo animal study by Habibovic et al. [8], 3D printed DCPA and DCPD cements implanted at intramuscular sites in goats led to ectopic immature and woven bone formation induced by the supersaturated local environment of Ca2+ and PO43– ions, indicating the osteoinductive properties of the DCPA and DCPD cements. Osteoinductive  124  biomaterials have been shown to perform better in orthopedic defects at clinically critical sizes and stimulate more new bone formation [9, 72]. Biodegradation of hard tissue implants should also be considered to allow simultaneous replacement with new bone tissues and obtain complete repair without remnants of the artificial substitutes. Dicalcium phosphates are one of the fastest resorbable CaP materials with the resorption rate of DCPD at a pH of 5.5 being about three times and three hundred times faster than that of tricalcium phosphate and carbonated HAp, respectively [10]. Conversely, there was no significantly difference on the proliferation rate between the Ref-PLA scaffold and the Micro-Mech-4 scaffold. The proliferation rate was much lower than that of the Re-DCPA/PLA/PEG-4 scaffold. It has shown that the behavior of osteogenic cells on calcium phosphate surfaces can be altered by the surface topography, roughness, crystal size and grain size of the particles which play important roles in particle-cell interactions [214−218]. It can be explained, therefore, that the DCPA effects on the cell proliferation between the Re-DCPA/PLA/PEG scaffolds and the Micro-Mech-4 scaffold differed by the DCPA size and distribution on the PLA nanofibers. The DCPA nanocrystallites reinforced PLA nanocomposite fibrous scaffolds promoted faster proliferation rate of the osteoblast cells than the mechanically mixed and electrospun PLA nanofibers containing the micro meter level DCPA crystals.   7.1.3 Bone-like nodule formation on scaffolds The investigation of mineral deposition and cell layers on the Ref-PLA scaffold and the Re-DCPA/PLA/PEG-4 scaffold was carried out using various tools of microscopy. 2.0 × 105 cells/mL were seeded on each scaffold and cultured for 2, 4 and 6 weeks. Cell culture procedures were similar to those of the cell attachment and proliferation studies (Refer to the section of 4.4.5.). However, after 1 week of post cell-seeding, 31.5 mg/mL β-glycerophosphate disodium salt hydrate and 0.58 mg/mL L-ascorbic acid phosphate magnesium salt n-hydrate were added to the α-MEM supplemented, in order to promote mineralization process of the cells, as well as tetracycline to label mineralized deposits.  125  SEM/EDS was used to visualize the cells and the mineralized deposits on the scaffold. Bone-like nodules labeled by tetracycline were visualized using epifluorescence microscopy ( λmax = 488 nm) and a low light-intensity CCD camera. Previous studies using the same rat calvarial osteoblasts demonstrated that tetracycline clearly labels mineralized deposits [220, 221]. Figure 7.6 is SEM micrographs and its EDS mapping of the RCO cells cultured on the Ref-PLA scaffold for 2 weeks. Figure 7.6 (a) shows the RCO cell layers on the surface of the Ref-PLA nanofibers. Around 5 µm size of extracelluar deposit was found on the top surface of the cell layers, as shown in 7.6 (b). Energy dispersive spectroscopy (EDS) mapping of the deposit indicates that it consisted with Ca, P, C, and O elements. C and O elements were distributed all over the area, but Ca and P elements, originated from the osteoblasts, were distributed mainly on the area of the deposit (Figure 7.6 (c and d)). The mineralized deposit and the proliferated growth of the cells confirm that the Ref-PLA scaffold is biocompatible and osteoconductive. The larger size of the mineral deposits around 20 µm were found on the Ref-PLA scaffold and the Re- DCPA/PLA/PEG-4 scaffold after culturing the RCO cells for 4 weeks. Figure 7.7 (a) shows multiple cell layers                   126    Figure 7.6 (a, b) SEM micrographs and (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Ref-PLA scaffold for 2 weeks.   Figure 7.7 (a, b) SEM micrographs and (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Ref-PLA scaffold for 4 weeks.   127   covering the entire surface of the Ref-PLA scaffold. Visible mineralized deposits (red arrows) were discretely distributed under the cell layers. Unlike the others, globular deposit was visualized as shown in Figure 7.7 (b). The Re-DCPA/PLA/PEG-4 scaffold in Figure 7.8 demonstrates the mineralized deposit with uneven and bumpy crystals covered by the osteoblasts and the nanoporous extracellular substance (red arrows). The structure of mineralized deposit should be at least 50 µm in diameter to be considered as bone-like nodule [221]. After 6 weeks of post cell-seeding, bone-like nodule with the size over 50 µm was found under the RCO cell layers cultured on the Re-DCPA/PLA/PEG-4 scaffold, as shown in Figure 7.9. The size increase of the mineralized deposits on the nanocomposite scaffold with the cell culture periods was also observed from the epifluorescence microscopy images in Figure 7.10. In case of the Ref-PLA scaffold, the mineralized deposits also grew with time (not shown here). In accordance with the SEM micrograph  in Figure 7.9, around 50 um size of the bone-like nodule was observed on the Re-   Figure 7.8 (a, b) SEM micrographs and (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Re-DCPA/PLA/PEG-4 scaffold for 4 weeks.   128    Figure 7.9 (a, b) SEM micrographs and  (c, d) EDS mapping of bone-like nodule produced by RCO cells cultured on Ref-PLA scaffold for 6 weeks. DCPA/PLA/PEG-4 scaffold after 6 weeks of post cell-seeding. These mineralized deposits result from mature osteoblast cells that are able to produce minerals [218, 219]. The bone-like nodule observation, in this research, suggests that the DCPA/PLA nanocomposite fibrous scaffold possesses great potential for bone tissue regeneration.       Figure 7.10 Epifluorescence images of bone-like nodules produced by RCO cells cultured on Re-DCPA/PLA/PEG-4 scaffold for (a) 2, (b) 4, and (c) 6 weeks.    129   The RCO cells cultured on the Ref-PLA scaffold and the Re-DCPA/PLA/PEG-4 scaffold were characterized using multiphoton microscopy (MPM), in order to observe cross-section of both the cell layers and the scaffolds. MPM is a minimally invasive and high-resolution optical imaging tool that can be used for structural and functional imaging of biological tissues [136]. MPM excites and detects nonlinear signals such as two-photon excitation fluorescence (TPEF) and  second harmonic generation (SHG). TPEF signals originate from autofluorescence of tissue and/or exogenous fluorophores, SHG signals come mainly from collagen fibers [136−138]. Figure 7.11 (a−f) show 2D cross-section images of the RCO cell layers and the DCPA/PLA nanocomposite fibrous structure, after 6 weeks of post cell-seeding. The density of the seeded cells was the same as that of the bone-like nodule formation. The 2D micrographs were scanned with every 1 µm stack. The numbers on the micrographs indicate each stack of the scanned images. Figure 7.11 (a) shows the top layer of the cells covering entire surface of the scaffold. Figure 7.11 (b)  130    Figure 7.11 (a−f) MPM micrographs of RCO cells cultured on Re-DCPA/PLA/PEG-4 scaffold for 6 weeks and 3D images of (g) the scaffold and (h) cell layers. The 2D micrographs were scanned with every 1 µm stack. The numbers indicate each stack of the scanned images. Amira® software was used to produce 3D images from 2D images. The mineralized deposits (white arrows) were mostly found between the cell layers and the nanofiber surface. The osteoblasts migrated through the nanofibrous structure, as shown in the rectangles.    131  shows the sub-layer of the cells with the globular mineral deposits (white arrows) produced by the top layer osteoblasts. In the region of the red rectangle on the micrographs of Figure 7.11 (b) shows no cells but the nanofibers, but the RCO cell was found further down into the inner cross-section as shown in Figure 7.11 (c). The white rectangles in the micrographs of Figure 7.11 (c−f) also clearly indicate that the RCO cell migrated through the nanofibrous structure, called cell infiltration. When nanofibers are created during electrospinning, they are deposited randomly on the target layer by layer, and various diameter of pores from a few to hundred micrometers are formed. It has been hypothesized that cells use amoeboid movement to migrate through the pores in an electrospun nanofibrous structure (dynamic nanofibrous architecture) by pushing the loosely lying fibers aside to expand the pores [12]. Many others demonstrated the observation of cell infiltration into the electrospun nanofibrous scaffolds, using cell staining and histology [224−228]. All of the 2D scanned images were compiled and processed with Amira® software to produce 3D images of the scaffold and the cell layers, as shown in Figure 7.11 (g and h).   7.2 Hydroxyapatite/Alginate scaffolds 7.2.1 Attatchement of osteoblasts on scaffolds  The morphological response of the rat calvarial osteoblast (RCO) cells to different surface chemistry of the alginate and HAp/alginate nanofibrous scaffolds was observed using scanning electron microscopy (SEM). Figure 7.12 (a) shows representative morphology of the RCO cells cultured on the cross-linked Ref-Alginate scaffold for 7 days. In general, the RCO cells were spherically shaped and anchored to the surface of the Ref-Alginate scaffold at day 1. At day 3, the osteoblasts spread more extensively without any preferred orientation and were round-flattened with closer adhesion to the surface at day 7. Figure 7.12 (b) demonstrates representative morphology of the RCO cells cultured on the cross- linked/in-situ synthesized H-Alginate 5 scaffold for 7 days. At day 1, the osteoblasts were  132    Figure 7.12 SEM micrographs of RCO cells cultured on (a) cross-linked Ref-Alginate and (b) cross-linked/in-situ synthesized H-Alginate-5 scaffolds for 7 days. The white arrows indicate more closely and stably attached RCO cells on the HAp/alginate than the pure alginate scaffold at day 1 of post cell-seeding. The magnified inset SEM micrograph of the HAp/alginate scaffold shows the visible development of multiple filopodial connections to the surface at day 7 of post cell-seeding.     133  attached more closely and stably to the surface of the HAp/alginate scaffold than the cells on the pure alginate scaffold, as indicated by the white arrows. The most common morphology of the RCO cells at day 3 was an elongated shape in a unidirectional manner. The osteoblasts were further flattened, stretched, and elongated into a spindle-shape with well-developed leading edges and multiple filopodial connections (the magnified inset SEM micrograph) to the surface of the HAp/alginate scaffold at day 7. The RCO cells on the cross-linked/in-situ synthesized H-Alginate-2 scaffold also displayed more stable attachment and elongated spindle shape than those on the Ref-Alginate scaffold (not shown here). The cross-linked/in-situ synthesized H-Alginate 8 scaffold was excluded from the cellular response study due to its loss of the nanofiber structure. Figure 7.13 shows representative immunostaining images of the RCO cells cultured on the cross-linked Ref-Alginate scaffold and the cross-linked/in-situ synthesized H-Alginate-5 scaffold for 7 days. Actin filaments of the osteoblast cells, one of the primary cytoskeletal components, were labeled red with phalloidin. Nuclei of the cells were labeled blue with DAPI. These immunofluorescent images support the SEM results of the RCO cells adhesion morphology on the pure alginate and HAp/alginate scaffolds. At day 1, the RCO cells attached more stably and spread on the cross-linked/in-situ synthesized H-Alginate 5 nanocomposte fibrous scaffold than the cross-linked Ref-Alginate scaffold (Figure 7.13 (a and c)). The osteoblast cells were elongated further and proliferated faster with well-developed leading edges on the nanocomposite scaffold at day 7 of post cell-seeding (Figure 7.13 (d)). Figure 7.13 (a) and (b) show that most of the RCO cells on the pure alginate formed round shape during 7 days of culture, similar to the SEM micrograph of Figure 7.12 (a).    Cell adhesion is an important cellular process that directly influences cell growth, proliferation, differentiation, and migration on the surface of a scaffold [186, 187]. Cell shape is also known to regulate cell behaviors, such as cell growth and differentiation, ECM metabolism, gene expression, and cell life destiny [188–190]. The adhesion and shape of cells are mainly affected by the surface topography and surface chemistry of a   134    Figure 7.13 Immunofluorescence images of RCO cells cultured on (a) cross-linked Ref-Alginate scaffold for 1 day, (b) cross-linked Ref-Alginate scaffold for 7 days, (c) cross-linked/in-situ synthesized H-Alginate-5 scaffold for 1 day, and (d) cross-linked/in-situ synthesized H-Alginate-5 scaffold for 7 days. Actin filaments were labeled red with phalloidin (left), nuclei were stained blue with DAPI (middle), and the two fluorescent images were merged (right).     135  scaffold [187]. Initial osteoblast adhesion and spreading play a key role in early bone tissue healing processes such as osteoconduction and de novo bone formation [191]. Therefore, optimization of the surface chemistry and topographical geometry has become essential to the development of ideal scaffolds for successful hard tissue regeneration. In the present study, the nanofibrous structure with 3D interconnected pores and a high surface area resembling the natural ECM of human tissues was achieved using electrospinning technology. This biomimetic nanofibrous surface topography has been reported to enhance adhesion, proliferation, and differentiation of cells [11, 12]. Furthermore, the surface chemistry of the electrospun alginate nanofibers was modified with the deposition of the HAp nanocrystals via the novel in-situ synthesis route. The observation of the in-vitro RCO cells response clearly indicates that the osteoblasts were more stably adhered, attached, and spread on the surface of the HAp/alginate nanocomposite fibrous scaffolds than on the pure alginate scaffold. It is highly considered that the HAp surface modification played a major role for the improved cell adhesion on the alginate nanofibers. More details of the HAp nanocrystals functions will be explained in the next section.  7.2.2 Proliferation of osteoblasts on scaffolds  An MTS cell proliferation assay was used to assess cultured RCO cells viability on the alginate and HAp/alginate scaffolds. Effects of the electrospun nanofibrous topography and the surface modification of alginate with the HAp nanocrystals on the proliferation of the RCO cells were also observed. Figure 7.14 shows the MTS assays of the RCO cells seeded on the scaffolds at 2 × 104 cells/mL and cultured for 7 days. The average values of optical density were produced from triplicate samples of each scaffold. Statistical significance was selected at p < 0.05, using one-way ANOVA for the specimens at each day. Alginate film was used as another control sample to compare cell attachment and proliferation with the electrospun nanofibrous structure of Ref-Alginate. Overall, the viable cell numbers on each scaffold increased with time for 7 days of culturing periods. The nanofibrous structure was found to promote cell growth over the film structure, as shown in Figure 7.14 (a and b).  136    Figure 7.14 MTS assays of cultured RCO cells on (a) alginate film, (b) cross-linked Ref-Alginate, (c) cross-linked/in-situ synthesized H-Alginate-2, (d) cross-linked/in-situ synthesized H-Alginate-5, and (e) cross-linked Mech-H-Alginate-5 scaffolds for 7 days. Triplicate samples were tested for each scaffold. Statistical significance was selected at p < 0.05, using one-way ANOVA for the specimens at each day.   After 7 days of culture, the optical density of the cell culture medium on the Ref-Alginate scaffold was about 1.5 times higher than that of the alginate film. The cross-linked/in-situ synthesized H-Alginate-5 scaffold showed the highest optical density among the samples throughout the culture periods. The RCO cells proliferated faster on the H-Alginate scaffolds with the HAp nanocrystals content. The RCO cells proliferation on the HAp/alginate scaffolds depending on the size and distribution of the HAp crystals was compared using the cross-linked/in-situ synthesized  H-Alginate-5 scaffold and the cross-linked Mech-H-Alginate-5 scaffold. The two scaffolds contained similar amount of the HAp, but the mechanically  mixed/electrospun Mech-H-Alginate possessed highly agglomerated HAp crystals over the size of sub-micrometers irregularly distributed on the surface of the scaffold. It was found that at day 7 of the post cell-seeding, the RCO cells  137  grew faster on the cross-linked/in-situ synthesized H-Alginate-5 scaffold than the cross-linked Mech-H-Alginate-5 scaffold, while no significant difference was observed during 3 days of culture periods.  Tissue regenerative engineering searches for solutions to restore damaged tissue or organ functions. Successful tissue regeneration necessitates the design of an advanced smart biomaterial as a temporary extra-cellular matrix (ECM) scaffold that is capable of inducing host cells to assume highly specialized functions [201]. An ideal scaffold also must be mechanically stable, which is dependent on the selection of the material composition, the architectural design, and the cell-material interactions. The cell proliferation studies performed in this work showed that the electrospun nanofibrous scaffold can provide the RCO cells with more favorable architecture than the alginate film. Electrospinning produces non-woven nanofibrous topography with a high surface area/volume and 3D interconnected pores resembling the structure of natural ECM. The high surface area with its 3D features is advantageous features for cells to attach and proliferate. An electrospun nanofibrous scaffold has around 90 % of porosity, which provides more structural space for cell accommodation and makes the exchange of nutrient and metabolic waste between the scaffold and environment efficiently [12]. The RCO cells proliferation on the electrospun alginate scaffold for RCO cells was also compared with the alginate film, using a DAPI stained cell number counting method. Figure 7.15 demonstrates the average number of RCO cells cultured on the scaffolds for 7 days. Nuclei of the RCO cells were stained with DAPI and visualized with fluorescent microscope. Six images per scaffold were taken, and  the nuclei of the cells were counted using ImageJ software. Triplicate samples were prepared for each scaffold. Figure 7.16 shows representative fluorescent images of the stained nuclei of the RCO cells cultured on the scaffolds for 7 days. The cell growth curves in Figure 7.15 (a and b) show that from day 3 of the culture, the RCO cell numbers on the nanofibrous surface increased more rapidly than those of the film surface.   138     Figure 7.15 Average number of RCO cells cultured on (a) alginate film, (b) cross-linked Ref-Alginate, (c) cross-linked/in-situ synthesized H-Alginate-2, (d) cross-linked/in-situ synthesized H-Alginate-5, and (e) cross-linked Mech-H-Alginate-5 scaffolds for 7 days. Nuclei stained RCO cells with DAPI were counted using ImageJ software. Six images per scaffold were taken, and triplicate samples were prepared for each scaffold.    The MTS assay indicated that the RCO cells on the surface modified alginate nanofibers with the HAp nanocrystals proliferated faster than those on the surface of the pure alginate nanofibers. The increase of the cell growth rate with the HAp content on the scaffolds was also observed from the cell number counting studies, as shown in Figure 7.15 (b−d) and Figure 7.16 (b−d). The cell attachment studies in the section of 7.2-1 showed that the RCO cells spread extensively and elongated into a spindle shape forming well developed multi    139    Figure 7.16 Fluorescent images of DAPI stained nuclei of RCO cells cultured on (a) alginate film, (b) cross-linked Ref-Alginate, (c) cross-linked/in-situ synthesized H-Alginate-2, (d) cross-linked/in-situ synthesized H-Alginate-5, and (e) cross-linked Mech-H-Alginate-5 scaffolds.     140  filopodia on the HAp/alginate nanocomposite fibrous scaffolds. The enhanced cell-material adhesion is believed to improve RCO cellular  interaction on the surface of the HAp/alginate nanocomposite fibrous scaffolds, and hence, and hence resulted in the faster growth rate of the RCO cells on the in-situ synthesized/cross-linked H-Alginate scaffolds than the Ref-Alginate scaffold. Interestingly, the RCO cells grew faster on the surface modified with the homogeneously deposited HAp nanocrystals along the alginate nanofibers (H-Alginate-5) than that with the highly agglomerated HAp crystals on the alginate nanofibers (Mech-H-Alginate-5) (Figure 7.14 (d and e), Figure 7.15 (d and e)). This result also emphasizes how it is important to control size of the HAp nanocrystals and its distribution for the surface modification of biopolymer surfaces, in order to maximize its effectiveness for cellular activities.    The remarkable biocompatibility, osteoconductivity, and non-toxicity of HAp have been shown to promote attachment, proliferation, and differentiation of osteoblasts cultured on HAp incorporated biopolymer scaffolds [192, 197–199]. Peter et al. [192] reported that HAp nanoparticles added to a chitosan-gelatin porous scaffold improved the formation of focal adhesion and allowed for substantial cell spreading as a result of enhanced protein adsorption on the surface by increasing the surface area and binding sites with the HAp. An initial cell-material interaction is considered to be a process mediated by the ECM proteins [202]. Alginate is so hydrophilic that it has limited protein adsorption capacity, which induces less affinity of cell adhesion to the surface [203]. On the other hand, HAp surfaces are known to passively adsorb RGD peptides and increase the adhesion of cells [193, 194]. Other in-vitro biological studies have reported that HAp addition is an effective method to buffer the acidic environment which is caused by the degradation of biopolymer matrices, resulting in more favorable conditions for cellular activities. The ionic dissolution of HAp significantly enhanced in-vitro and in-vivo osteogenesis by forming super-saturated microenvironments with Ca2+ and PO43– precursor ions for new bone formation [195, 196].   141  The in-vitro cellular responses of RCO cells were different on the two distinctive CaP/biopolymer nanocomposite fibrous scaffolds. The RCO cells attached more closely, spread more extensively, and proliferated faster on the PLA-based scaffolds than the alginate-based scaffolds. The surface topography of the electrospun nanofibrous scaffolds was similar, but the surface chemistry of the synthetic polyester PLA and the natural polysaccharide alginate was different. Alginate has highly hydrophilic surface, which causes it to have limited protein adsorption capacity and lack of cellular interactions [239]. On the others hand, the RCO cells interacted actively with the PLA surface, resulting in the fusion of the filopodial of the osteoblasts with the nanofibers and the  migration of the cells under the layers of the PLA naofibers. Futhermore, the ionically cross-linked alginate-based scaffolds in the cell culture medium was unstable due to the counter-ion exchange between Ca2+ and Na2+. This may cause the RCO cells not only to attach stably but also to proliferate on the alginate-based scaffolds, compared to the PLA-based scaffolds.         7.3 Conclusions  There was no change of the osteoblasts morphology on the surface modified PLA nanofibers with the DCPA nanocrystallites, compared to the pure PLA nanofibers. However, the cell proliferation studies using the MTS assay and cell number counting with DAPI staining confirmed that noticeably positive increase of the cell growth was achieved from the DCPA/PLA nanocomposite fibrous scaffolds. After 6 weeks of cell culture, bone-like nodules, over 50 µm size of the mineralized deposits, were produced by the mature osteoblast cell layers on the nanocomposite fibrous scaffolds.  The initial attachment of the RCO cells on the HAp/alginate scaffolds was more stable than attachment on the pure alginate. The osteoblasts were stretched and elongated into a spindle-shape with well-developed leading edges and multiple filopodial formation on the HAp/alginate scaffolds compared to the flattened and round shaped cells on the pure alginate. The MTS assay and  142  the cell number counting study confirmed that the RCO cells grew faster on the electrospun alginate nanofibrous structure than the film surface, and the biomimetic surface modification with the HAp nanocrystals induced much faster proliferation rate of the cells on the scaffold.  The enhanced osteoblast cellular activities on the calcium phosphate/biopolymer nanocomposite fibrous scaffolds suggested their potential advantages to be used as bone tissue regeneration matrix over the pure biopolymer nanofibrous scaffolds. In this in-vitro studies, the effects of the calcium phosphates addition into the biopolymers noticeably increased by controlling its nano sizes and distribution within/on the nanofibers through the novel in-situ synthesis strategies.                        143  Chapter 8 Conclusions and Future Work   This study of the bio-inspired nanocomposite fibrous scaffolds for hard tissue regeneration has investigated materials synthesis, processing and fabrication, and  continued with materials characterization and in-vitro cellular response analysis. The fundamental building blocks of hierarchically-structured bone tissue, mineralized collagen fibrils with calcium phosphates, inspired us to engineer calcium phosphate/biopolymer nancomposites through biomimetic in-situ synthesis and electrospinning. All research Objectives, as listed in Section 3.2, were addressed. Two distinctive nanocomposite systems were explored in details: i) synthetic biodegradable polymer of poly(lactic acid) (PLA) with dicalcium phosphate anyhydrate (DCPA) and ii) natural polysaccharide of alginate with hydroxyapatite (HAp). The following  conclusions were drawn from this study.       8.1 DCPA/PLA nanocomposite fibrous scaffolds DCPA was, for the first time, in-situ synthesized in non-aqueous PLA solution environment composed of tetrahydrofuran (THF), dimethyl formamid (DMF), and dimethyl sulfoxide (DMSO) at a pH = 4.0 ± 0.5. The formation of crystalline DCPA was suppressed by the presence of PLA molecular chains which is believed to hinder the diffusion of Ca2+ and PO43– ions necessary for the crystallization of DCPA. In the presence of PLA, amorphous calcium phosphate or poorly crystalline DCPA was formed, depending on the calcium nitrate tetrahydrate (CNT) concentration used for the synthesis, while highly crystalline DCPA was produced in the absence of PLA.   The in-situ synthesized DCPA/PLA composite solutions were electrospun into nanocomposite fibrous scaffolds.  The interaction between Ca2+ ions and the carbonyl groups in the PLA was confirmed by FT-IR. We believe that these sites serve as the nucleation sites for DCPA during the synthesis and thus induce the homogenous dispersion  144  of the DPCA nanocrystallites in the electrospun PLA fiber matrix. This process avoided the severe agglomeration of calcium phosphate processed by electrospinning mechanically blended solution with pre-prepared nanoparticles. The intra-nanopores within the DCPA/PLA nanocomposite fibers were generated probably due to the evaporation of DMSO after fiber formation. The nanofibers containing residual DMSO during electrospinning were self-fused, resulting in broad distribution of the fiber diameters from ~100 nm to ~3.0 um. The fiber diameters increased with the CNT concentration used for the DCPA synthesis, i.e. DCPA amount in the fibers. The incorporation of poly(ethylene glycol) (PEG), as a temporary co-polymeric additive, enabled greater stability in the formation of the electrospun jets, decreased the fiber diameter, and permitted more uniform fiber morphology, while increasing the amount of DCPA within the nanofibers.   After re-dissolving the DCPA/PLA/PEG nanocomposites and electrospinning process, the uniformly controlled PLA nanofibrous structure with the homogeneous distribution of the DCPA nanocrystallites was achieved, referred to as the Re-DCPA/PLA/PEG scaffolds. A micro tensile testing studies showed that the Re-DCPA/PLA/PEG-4 scaffold possessed the highest tensile strength of 26.91 ± 1.34 MPa and the highest elastic modulus of 534 ± 14 MPa among all PLA-based scaffolds. There was no significant difference of the tensile strength between the Re-DCPA/PLA/PEG-4 scaffold and Ref-PLA scaffold. On the other hand, the elastic modulus of the Re-DCPA/PLA/PEG-4 increased 47.5 % over the Ref-PLA. This result can be explained by the presence of homogeneously dispersed and distributed DCPA nanocrystallites within the uniform PLA nanofiber matrix. When the load is applied to the Re-DCPA/PLA/PEG scaffolds, the DCPA nanocrystallites are believed to induce a stress transfer effect, which can enhance strength of the matrix. Even though the Re-DCPA/PLA/PEG scaffolds showed the more brittle phenomenon than the Ref-PLA scaffold, they were slightly further elongated, as compared to the DCPA/PLA/PEG-2 and the DCPA/PLA/PEG-4 scaffolds, possibly due to the more uniform PLA nanofibrous structure.   145  The in-vitro bioactivity of the Re-DCPA/PLA/PEG scaffolds was found to be greater than that of the neat PLA scaffold. On the surface of the Re-DCPA/PLA/PEG-4 scaffold, the cauliflower-like precipitates lined the entire nanofiber surface, and even covered all the porous spaces with overlapping of the spherical crystals after 10 days of soaking in the simulated body fluid (SBF) solution. The crystals on the Re-DCPA/PLA/PEG-4 scaffold had sharper facets and were distributed in a more uniform size in comparison with those on the Ref-PLA scaffold. The phase of the precipitated crystals was analyzed using X-ray diffractometer (XRD). While broad diffraction signals at around 2θ = 32° were observed from the Ref-PLA scaffold, the intensity of the signals increased with the nanocomposite scaffolds containing the DCPA nanocrystallites. In case of the Re-DCPA/PLA/PEG-4 scaffold, weak characteristic peaks for HAp were observed.  The In-vitro cellular responses on the nanocomposite scaffolds were compared with the neat PLA scaffold, using rat calvarial osteoblast (RCO) cells. There was no change of the osteoblasts morphology on the surface modified PLA nanofibers with the DCPA nanocrystallites, compared to the neat PLA nanofibers. The RCO cells attached and spread on the nanofibrous surface without preferred directions, the cell growth was guided by the nanofiber topography, and the multiple filopodial connections to the surface were made along the nanofibers, resulting in stable and close adhesion of the RCO cells at the cell-matrix interfaces. At day 7 of post cell-seeding, the cells grew confluence on the entire surface of the scaffolds, forming multi-cellular network. The cell proliferation studies using the MTS assay and cell number counting with DAPI staining confirmed that noticeably positive increase of the cell growth was achieved from the Re-DCPA/PLA/PEG nanocomposite fibrous scaffolds for 7 days of cell culture. The enhanced cell-matrix interaction on the nanocomposite resulted in the deposition of the mineralized crystals over 50 µm size, called bone-like nodules, which were produced by the mature osteoblast cell layers after 6 weeks of post cell-seeding. The osteoblast cell infiltration into the nanofibrous structure was also observed by multiphoton microscopy.   146  8.2 HAp/alginate nanocomposite fibrous scaffolds The HAp/alginate nanocomposite fibrous scaffolds were successfully processed by the  biomimetic in-situ synthesis of HAp on electrospun alginate nanofibers. The alginate nanofibers were cross-linked by Ca2+ ions at GG blocks (according to “Egg box” model), which is believed to serve as nucleation sites for HAp. While the scaffolds were agitated in the solution at a pH = 10, impregnated PO43– (from the fibers), Ca2+, and OH– (from the solution) ions diffused towards the [–COO–]–Ca2+–[–COO–] linkage sites for the nucleation and growth of the HAp nanocrystals along the alginate nanofibers. As PO43–  ion concentration increased in the alginate/PEO solution for electrospinning, more HAp nanocrystals precipitated on the alginate fibers. This novel processing method induced homogenous deposition of HAp nanocrystals on the nanofibers, unlike the severely agglomerated, micrometer-large HAp particles processed by the conventional mechanical blending/electrospinning method.   The tensile strength and elastic modulus of the HAp/alginate nanocomposite fibrous scaffold increased with the HAp content, as predictd by the stress transfer effect. The cross-linked/in-situ synthesized H-Alginate 5 scaffold had the highest tensile strength of 49.23 ± 2.09 MPa and the highest elastic modulus of  1.690 ± 0.208 GPa. The tensile strength increased 79.3 %, and the elastic modulus increased 158.4 %, compared to those of the Ref-Alginate scaffold. Such noticeable enhancement of the mechanical properties can be explained by the homogeneously dispersed and distributed HAp nanocrytals on the alginate nanofibers. The non-uniformly distributed and agglomerated HAp particles, processed by the mechanical mixing/electrospinning, produced the deteriorated tensile strength and elastic modulus of the Mech-H-Alginate scaffolds compared to the H-Alginate nanocomposite scaffolds. This result  demonstrates the importance of the control of particle size and distribution of the reinforced nanomaterials within the polymer matrix for maximizing its effects for the mechanical properties. It is also highly anticipated that the chemically induced deposition of the HAp nanocrystals on the alginate nanofibers, through  147  the biomimetic in-situ synthesis, would play an important role to enhance the tensile properties of the H-Alginate nanocomposite fibrous scaffolds.  The in-vitro bioactivity test using the SBF solution showed that the cauliflower-like HAp crystals were precipitated on the surface of the H-Alginate scaffolds, demonstrating in-vitro bioactivity of the nanocomposite unlike the pure alginate scaffold. The cauliflower-like HAp crystals in the H-Alginate scaffolds were much smaller than those of the Re-DCPA/PLA/PEG nanocomposite scaffolds. This might cause the less HAp visible peak on the XRD diffraction pattern.   This study also demonstrated that the pure alginate nanofibers cross-linked with Ca2+ ions dissolved and thus lost its non-woven structures. The cross-linked alginate structure is stable in water, but in the SBF solution with sufficient Na+ ions the dissolution of the cross-lined alginate structure occurs due to the exchange of Na+ ions in the solution with Ca2+ ions bound to the carboxyl groups in the alginate chains. However, the surface modified alginate nanofibers with the HAp nanocrystals in the cross-linked/in-situ synthesized H-Alginate 5 scaffold survived in the SBF solution for 36 hrs. It is believed that the HAp nanocrystals formed on the alginate nanofibers obstruct the ion exchange process between Na+ ions and Ca2+ ions, resulting in retaining the structural integrity of the nanofibrous structure in the SBF solution.           The initial attachment of RCO cells on the HAp/alginate scaffolds was more stable than on pure alginate. The osteoblasts were stretched and elongated into a spindle-shape with well-developed leading edges and multiple filopodial formation on the HAp/alginate scaffolds, compared to the flattened and round shaped cells on the pure alginate. The MTS assay and the cell number counting study confirmed that the biomimetic surface modification with the HAp nanocrystals on the H-Alginate nanocomposite scaffolds induced much faster proliferation rate of the cells on the nanocompoiste scaffold than the neat alginate scaffold  148  and the mechanically blended/electrospun HAp/alginate scaffold. The cell-material interactions were noticeably enhanced due to the results of controlling the nanocrystal precipitation and its homogenous distribution on the naonfibers through the novel in-situ synthesis strategies.    The two types of the CaP/biopolymer nanocomposite fibrous scaffolds possess the distinctive nature of the biopolymer nanofiber matrix and surface modification with the different calcium phosphates. These materials charateristics induced various outcomes of the scaffold properties, including in-vitro bioactivity, mechanical properties under tension, and in-vitro osteoblast cellular activities. Table 8.1 below summarizes the comparion of the scaffold properties between the PLA-based scaffolds and the alginate-based scaffolds.  Table 8.1 Comparison of CaP/biopolymer nanocomposite fibrous scaffolds properties studied in this study.  Order of Scaffold Properties  In-vitro Bioactivity            DCPA/PLA  >  PLA  >  HAp/alginate  >  alginate  Tensile Strength            HAp/alginate  ≈  alginate, DCPA/PLA, PLA   Elastic Modulus            HAp/alginate  >  alginate  >  DCPA/PLA  >  PLA  Failure Strain            PLA  >  DCPA/PLA  >  Alginate  >  HAp/alginate   Osteoblast Attachment            DCPA/PLA, PLA  >  DCPA/PLA  >  PLA   Osteoblast Proliferation            DCPA/PLA  >  PLA  > HAp/alginate  >  alginate  Osteoblast Migration            DCPA, PLA  >  HAp/alginate  >  alginate   8.3 Future work The homogeneous calcium phosphate/biopolymer nanocomposite fibrous scaffolds were engineered by mimicking the mineralized collagen fibrils of bone tissue. The biocompatible, bioactive, biodegradable, and osteoconductive inorganic/organic hybrid systems enhanced structural stability and biological performance  of the scaffolds, suggesting their potential advantages to be used as bone tissue regeneration matrix over the  149  pure biopolymer scaffolds. However, the goal of clinical application is still far from being realized. Thus, a broad research and development effort involving materials optimization studies (as suggested in detail below), then animal studies, and ultimately human trials and materials certification processes, should be undertaken with the collaboration with clinicians in the future.  A most significant advantage of a conventional electrospinning process is the capacity for massive production of uniform non-woven nanometer scale fibrous structure with about 90 % 3D interconnected porosity. However, the majority of pore diameters is limited to several tens micrometer levels which are not sufficient enough for osteoblast cells easily to infiltrate across the nanofibrous structures and eventually for new bone tissue ingrowth throughout the structures. Various new types of electrospinning apparatus have been developed to address this issue. The future research should include applying the novel in-situ synthesis strategies into one of the innovative electrospinning technologies, in order to tailor a 3D scaffold with a nanofibrous topography and interconnected macropores.   In case of the DCPA/PLA nanocomposites, the effectiveness of the DCPA nanophase to address cellular performances was lower than expected, unlike that of the HAp/alginate nanocomposites. This is believed to result from the fact that the majority of the DCPA nanocrystallites were not exposed to the surface of the nanofibers, but rather entrapped within the nanofibers. This limited surface modification could be resolved by controlled coating of the nanofibers with another layer of calcium phosphate. The nanoscale calcium phosphate coating technology, such as sol-gel, electro-spraying, and electro-deposition, could be combined with electrospinning.   The HAp/alginate nanocomposite fibrous scaffolds demonstrated brittle phenomenon of their fracture pattern. Alginate itself is not ductile, due to its linear copolymer molecular structure. The addition of HAp nanocrystals coated on the alginate nanofibers caused brittle  150  breakage of the nanocomposite scaffolds with smaller failure strain than the pure alginate scaffold. Another essential next study should therefore address and tailor the brittle structure of the HAp/alginate scaffold, e.g. by adding plasticizing nanofibers through co-electrospinning. 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PLA film Ref-PLA Re-DCPA/PLA/PEG-2 Re-DCPA/PLA/PEG-4 Micro-Mech-4 0.609 0.711 0.684 0.688 0.720 0.656 0.724 0.707 0.699 0.672 0.621 0.687 0.727 0.719 0.683  One Way Analysis of Variance  Normality Test: Passed (P = 0.448) Equal Variance Test: Passed (P = 0.938)      Group Name             N     Missing  Mean      Std Dev      SEM  PLA film                         3                   0             0.628      0.0241 0.0139  Ref-PLA             3         0             0.707      0.0185 0.0107  Re-DCPA/PLA/PEG-2         3         0             0.706      0.0215 0.0124  Re-DCPA/PLA/PEG-4         3         0             0.702      0.0159 0.0092  Micro-Mech-4            3         0             0.692      0.0253 0.0146   Source of Variation          DF        SS               MS          F     P   Between Groups           4    0.0133       0.00331     7.244         0.005  Residual                      10    0.00457     0.000457    Total                                  14    0.0178      The differences in the mean values among the treatment groups are greater than would be expected by chance; there is a statistically significant difference  (P = 0.005).  Power of performed test with alpha = 0.050: 0.906   169  All Pairwise Multiple Comparison Procedures (Tukey Test): Comparisons for factor:        Comparison          Diff of Means       p          q                 P         P<0.050 PLA film vs. Ref-PLA              0.0787 5.0000      6.3710    8.0000e-3 Yes PLA film vs. Re-2               0.0772 5.0000      6.2500    9.0000e-3 Yes PLA film vs. Re-4               0.0733 5.0000      5.9390    0.0120             Yes PLA film vs. Micro               0.0630 5.0000      5.1020    0.0310             Yes       Ref-PLA vs. Re-2          1.5000e-3 5.0000     0.1210     1.0000       Do Not Test Ref-PLA vs. Re-4          5.3300e-3 5.0000     0.4320     0.9980       Do Not Test Ref-PLA vs. Micro              0.0157 5.0000     1.2690     0.8920           No       Re-2 vs. Re-4                     3.8300e-3 5.0000     0.3100     0.9990       Do Not Test Re-2 vs. Micro              0.0142 5.0000     1.1470     0.9210       Do Not Test       Re-4 vs. Micro              0.0103 5.0000     0.8370     0.9730       Do Not Test  A result of "Do Not Test" occurs for a comparison when no significant difference is found between two means that enclose that comparison.     A.1.1.2 At day 3 of post cell-seeding   Table A.2 Measured optical density of MTS assay of PLA-based scaffolds at day 3.  PLA film Ref-PLA Re-DCPA/PLA/PEG-2 Re-DCPA/PLA/PEG-4 Micro-Mech-4 0.864 1.172 1.343 1.504 1.230 0.919 1.199 1.357 1.479 1.202 0.830 1.323 1.413 1.431 1.349  One Way Analysis of Variance  Normality Test: Passed (P = 0.383) Equal Variance Test: Passed (P = 0.811)     170       Group Name             N     Missing  Mean      Std Dev      SEM  PLA film                         3                   0             0.871       0.0446 0.0258  Ref-PLA             3         0             1.231       0.0806 0.0465  Re-DCPA/PLA/PEG-2         3         0             1.371       0.0373 0.0215   Re-DCPA/PLA/PEG-4         3         0             1.471       0.0369 0.0213  Micro-Mech-4            3         0             1.260       0.0778 0.0449   Source of Variation          DF        SS               MS          F     P   Between Groups           4    0.622 0.155     44.943     <0.001  Residual                      10    0.0346 0.00346    Total                                 14    0.656    The differences in the mean values among the treatment groups are greater than would be expected by chance; there is a statistically significant difference  (P = <0.001).  Power of performed test with alpha = 0.050: 1.000  All Pairwise Multiple Comparison Procedures (Tukey Test):  Comparisons for factor:        Comparison          Diff of Means       p          q                 P         P<0.050 PLA film vs. Ref-PLA              0.3600 5.0000     10.6120    <0.001             Yes PLA film vs. Re-2               0.5000 5.0000     14.7200    <0.001             Yes PLA film vs. Re-4               0.6000 5.0000     17.6800    <0.001             Yes PLA film vs. Micro               0.3890 5.0000     11.4660    <0.001     Yes       Ref-PLA vs. Re-2               0.1400 5.0000       4.1080     0.0910              No Ref-PLA vs. Re-4               0.2400 5.0000       7.0680     4.0000e-3   Yes Ref-PLA vs. Micro               0.0290 5.0000       0.8540     0.9710       Do Not Test       Re-2 vs. Re-4                           0.1010 5.0000       2.9600     0.2940              No Re-2 vs. Micro               0.1110 5.0000       3.2540     0.2210       Do Not Test       Re-4 vs. Micro               0.2110 5.0000       6.2140     9.0000e-3 Yes   171  A.1.1.3 At day 7 of post cell-seeding   Table A.3 Measured optical density of MTS assay of PLA-based scaffolds at day 7.  PLA film Ref-PLA Re-DCPA/PLA/PEG-2 Re-DCPA/PLA/PEG-4 Micro-Mech-4 1.448 2.302 2.704 2.817 2.575 1.217 2.273 2.713 2.910 2.377 1.445 2.486 2.764 2.782 2.358  One Way Analysis of Variance  Normality Test: Passed (P = 0.406) Equal Variance Test: Passed (P = 0.903)      Group Name             N     Missing  Mean      Std Dev      SEM  PLA film                         3                   0             1.370        0.133 0.0765 Ref-PLA             3         0             2.354        0.116 0.0668  Re-DCPA/PLA/PEG-2         3         0             2.727        0.0325 0.0188   Re-DCPA/PLA/PEG-4         3         0             2.836        0.0662 0.0382  Micro-Mech-4            3         0             2.437        0.120 0.0695   Source of Variation          DF        SS               MS          F     P   Between Groups           4    4.039 1.010     99.262      <0.001  Residual                      10    0.102 0.0102    Total                                 14    4.141   The differences in the mean values among the treatment groups are greater than would be expected by chance; there is a statistically significant difference  (P = <0.001).  Power of performed test with alpha = 0.050: 1.000  All Pairwise Multiple Comparison Procedures (Tukey Test):     172  Comparisons for factor:        Comparison          Diff of Means       p          q                 P         P<0.050 PLA film vs. Ref-PLA              0.9840 5.0000    16.8900      <0.001             Yes PLA film vs. Re-2               1.3570 5.0000    23.3010      <0.001             Yes PLA film vs. Re-4               1.4660 5.0000    25.1820      <0.001             Yes PLA film vs. Micro               1.0670 5.0000    18.3150      <0.001             Yes       Ref-PLA vs. Re-2               0.3730 5.0000      6.4110         8.0000e-3 Yes Ref-PLA vs. Re-4               0.4830 5.0000      8.2920  1.0000e-3 Yes Ref-PLA vs. Micro               0.0830 5.0000      1.4250  0.8460  No       Re-2 vs. Re-4                           0.1100 5.0000      1.8800  0.6810  No Re-2 vs. Micro               0.2900 5.0000      4.9860  0.0350 Yes       Re-4 vs. Micro               0.4000 5.0000      6.8660  5.0000e-3 Yes   A.1.2 Alginate and HAp/alginate scaffolds A.1.2.1 At day 1 of post cell-seeding  Table A.4 Measured optical density of MTS assay of alginate-based scaffolds at day 1.  Alginate film Ref-Alginate H-Alginate-2 H-Alginate-5 Mech-H-Alginate-5 0.395 0.432 0.458 0.472 0.427 0.425 0.450 0.487 0.462 0.474 0.419 0.442 0.469 0.484 0.449  One Way Analysis of Variance  Normality Test: Passed (P = 0.257) Equal Variance Test: Passed (P = 0.758)        173      Group Name                   N   Missing Mean       Std Dev       SEM  Alginate film                       3                   0             0.627         0.0720  0.0416 Ref-Alginate            3        0             0.815       0.0576  0.0332  H-Alginate-2                        3                    0             0.929       0.0441  0.0255  H-Alginate-5                        3                    0             1.189       0.0208  0.0120  Mech-H-Alginate-5           3        0             1.056       0.0210  0.0121   Source of Variation          DF        SS               MS          F     P   Between Groups           4    0.564 0.141     62.225      <0.001   Residual                      10    0.0226 0.00226    Total                                  14    0.586  The differences in the mean values among the treatment groups are greater than would be expected by chance; there is a statistically significant difference  (P = <0.001).  Power of performed test with alpha = 0.050: 1.000  All Pairwise Multiple Comparison Procedures (Tukey Test):  Comparisons for factor:           Comparison          Diff of Means      p           q               P            P<0.050 Algi film vs. Ref-Algi            0.0283             5.0000      3.1430       0.2470         Do Not Test Algi film vs. H-2             0.0583             5.0000      6.4700      7.0000e-3 Yes Algi film vs. H-5             0.0598             5.0000      6.6360      6.0000e-3 Yes Algi film vs. Mech             0.0373             5.0000      4.1410      0.0880              No       Ref-Algi vs. H-2             0.0300             5.0000      3.3270      0.2060        Do Not Test Ref-Algi vs. H-5             0.0315             5.0000      3.4940      0.1740              No Ref-PLA vs. Micro             9.0000e-3 5.0000      0.9980      0.9510        Do Not Test       H-2 vs. H-5                         1.5000e-3 5.0000      0.1660      1.0000        Do Not Test H-2 vs. Mech                          0.0210             5.0000      2.3290      0.5030        Do Not Test       H-5 vs. Mech                         0.0225             5.0000      2.4960      0.4420        Do Not Test    174  A.1.2.2 At day 3 of post cell-seeding  Table A.5 Measured optical density of MTS assay of alginate-based scaffolds at day 3.  Alginate film Ref-Alginate H-Alginate-2 H-Alginate-5 Mech-H-Alginate-5 0.571 0.702 0.773 0.884 0.845 0.532 0.740 0.798 0.905 0.871 0.510 0.726 0.775 0.901 0.868  One Way Analysis of Variance  Normality Test: Passed (P = 0.710) Equal Variance Test: Passed (P = 0.522)      Group Name                   N   Missing Mean       Std Dev       SEM  Alginate film                       3                   0             0.538        0.0309  0.0178 Ref-Alginate            3        0             0.723        0.0190  0.0110 H-Alginate-2                        3                    0             0.782        0.0137  0.00793  H-Alginate-5                        3                    0             0.896        0.0113  0.00650  Mech-H-Alginate-5           3        0             0.861        0.0142  0.00821   Source of Variation          DF        SS               MS          F     P   Between Groups           4    0.240 0.0600    163.830 <0.001   Residual                      10    0.00367 0.000367    Total                                  14    0.244  The differences in the mean values among the treatment groups are greater than would be expected by chance; there is a statistically significant difference  (P = <0.001).  Power of performed test with alpha = 0.050: 1.000  All Pairwise Multiple Comparison Procedures (Tukey Test):     175  Comparisons for factor:           Comparison          Diff of Means       p             q                 P            P<0.050 Algi film vs. Ref-Algi            0.1850             5.0000       16.7220      <0.001              Yes Algi film vs. H-2             0.2440             5.0000       22.0750      <0.001              Yes Algi film vs. H-5             0.3590             5.0000       32.4490      <0.001              Yes Algi film vs. Mech             0.3230             5.0000       29.2380      <0.001              Yes       Ref-Algi vs. H-2             0.0592             5.0000         5.3530        0.0230           Yes  Ref-Algi vs. H-5             0.1740             5.0000        15.7270  <0.001    Yes Ref-PLA vs. Micro             0.1380             5.0000        12.5150  <0.001    Yes       H-2 vs. H-5                         0.1150             5.0000       10.3740  <0.001        Yes H-2 vs. Mech                          0.0792             5.0000         7.1620    4.0000e-3       Yes       H-5 vs. Mech                         0.0355             5.0000          3.2120    0.2310    No   A.1.2.3 At day 7 of post cell-seeding  Table A.6 Measured optical density of MTS assay of alginate-based scaffolds at day 7.  Alginate film Ref-Alginate H-Alginate-2 H-Alginate-5 Mech-H-Alginate-5 0.6855 1.0495 1.2125 1.4970 1.3870 0.6810 1.0930 1.2245 1.5050 1.2760 0.7065 1.1085 1.3765 1.6070 1.4355  One Way Analysis of Variance  Normality Test: Passed (P = 0.475) Equal Variance Test: Passed (P = 0.704)      Group Name                   N   Missing Mean       Std Dev       SEM  Alginate film                       3                   0             0.691        0.0136 0.00786 Ref-Alginate            3        0             1.084        0.0306 0.0177  H-Alginate-2                        3                    0             1.271        0.0914 0.0528  H-Alginate-5                        3                    0             1.536        0.0613 0.0354  Mech-H-Alginate-5           3        0             1.366        0.0818 0.0472     176  Source of Variation          DF        SS               MS          F     P   Between Groups           4    1.254 0.313 78.646 <0.001   Residual                      10    0.0399 0.00399    Total                                  14    1.293  The differences in the mean values among the treatment groups are greater than would be expected by chance; there is a statistically significant difference  (P = <0.001).  Power of performed test with alpha = 0.050: 1.000  All Pairwise Multiple Comparison Procedures (Tukey Test):  Comparisons for factor:           Comparison          Diff of Means       p             q                 P            P<0.050 Algi film vs. Ref-Algi            0.3930   5.0000       10.7740   <0.001   Yes Algi film vs. H-2             0.5800             5.0000       15.9180   <0.001   Yes Algi film vs. H-5             0.8450             5.0000       23.1940   <0.001   Yes Algi film vs. Mech             0.6750   5.0000       18.5250   <0.001   Yes       Ref-Algi vs. H-2             0.1870   5.0000         5.1450     0.0290   Yes  Ref-Algi vs. H-5             0.4530             5.0000       12.4200   <0.001   Yes Ref-PLA vs. Micro             0.2820             5.0000         7.7510     2.0000e-3   Yes       H-2 vs. H-5                         0.2650             5.0000         7.2760     3.0000e-3   Yes H-2 vs. Mech                          0.0950             5.0000         2.6070     0.4030    No       H-5 vs. Mech                         0.1700             5.0000         4.6690     0.0490   Yes         177  A.2 Details of fiber diameter measurement  A.2.1 Ref-PLA and DCPA/PLA scaffodS   Ref-PLA (nm) DCPA/PLA-2 (nm) DCPA/PLA-3 (nm) DCPA/PLA-4 (nm)     1 398 1207 1431   778     2 315 1012 1270 1266     3 144   887   715   844     4 305 1403   842 5485     5 175   887   992 6160     6 303 1190 1331 3816     7 305 1403 1068 4985     8 172 1775   715 4151     9 247   887 1122 4356   10 85   397 1331 1016   11 247 1431 1068 2830   12 163   794   842 2334   13 336   992   887 2830   14 163   992 1190 1740   15 268   397 1190 3967   16 179   397   992 1100   17 271   992   992 1208   18 243   794   715 1361   19 216   887 1012 1361   20 127 1157   397 1100   21 335 1403   887   943   22 195   198   715   943   23 137   444 1068 1148   24 148   627 1012 1374   25 222   887 1122   597   26 153   715   887   943   27 362   627   887 1148   28 229   818 1122 1100   29 305   715   818 1623   30 246   715   842   844   31 156   715   715 3778   32 134 1157 1829 1193   33 121   842 1122   844   34 193   444   887   800   35 240 1270   627   755   36 175 1122   992   800   37 134   715 1122   377   38 254   715 1255   597   39 405   715 1157   844  178   Ref-PLA (nm) DCPA/PLA-2 (nm) DCPA/PLA-3 (nm) DCPA/PLA-4 (nm)   40 201   627   887   778   41 161 1012 1122   943   42 326   397   887   377   43 148   397   887   778   44 189   444   887   422   45 271 1270   887 1148   46 170   818   595 1208   47 148 1157 1270   755   48 214 1707 1122   800   49 228   397   992 1924   50 208   794   715   566   A.2.2 DCPA/PLA/PEG scaffodS   DCPA/PLA/PEG-2 (nm) DCPA/PLA/PEG-4 (nm) DCPA/PLA/PEG-6 (nm) DCPA/PLA/PEG-8 (nm)     1   661 1882 4900 4880     2   592   715 7597 7104     3   296 1444 3396 2397     4   771 1255 5982 4437     5   674   627 2587 3691     6   748   887 3968 5862     7 1138   627 2422 7817     8   265   818 3577 9730     9   712 2218   561 11001   10 1431   627 1190 4283   11 1381 1012 1964 4167   12 1220 1207 1068 4533   13    887 1829   595 9172   14   837   818   397 1882   15 1220   444   627 2699   16 1508   397   444 2889   17 1431   627 1207 3230   18   805 1207   992 1600   19 1007 2218 1122 1122   20   545 1012   198 1331   21   296   842   887 2023   22   397   561   444 3003   23   418   281   715 1255   24   418   444   887 1872   25   296   627   595 3414  179   DCPA/PLA/PEG-2 (nm) DCPA/PLA/PEG-4 (nm) DCPA/PLA/PEG-6 (nm) DCPA/PLA/PEG-8 (nm)   26   661   198 1012 2192   27   805 2389 2043   794   28 2023   794 1190   397   29   837   444   397   715   30   529   444   561   992   31   397   715   818   595   32   529   561   281   715   33   805   715   715   444   34 1389   595   397   627   35 1138 1511   887   887   36   847   627 1444 1012   37   805   794   715   595   38 1444   281   627   444   39 1954   627   715   992   40   477 1157   281 1122   41   374   715 1600   715   42   661   444   715   561   43 1362 2262   444   444   44   661   715   198   198   45   545   444   281   794   46   561   281   198   561   47   712 1012   198   561   48 2662   992   444   887   49 1600   561   198 1068   50   674 1403 1587   561   A.2.3 Ref-PLA and Re-DCPA/PLA/PEG-4 scaffodS   Ref-PLA (nm) Re-DCPA/PLA/PEG-4 (nm)     1 160 501     2 308 275     3 336 193     4 424 314     5 302 370     6 404 471     7 351 336     8 308 371     9 237 390   10 202 332   11 405 358  180   Ref-PLA (nm) Re-DCPA/PLA/PEG-4 (nm)   12 502 374   13 405 390   14 275 275   15 306 282   16 248 390   17 405 428   18 420 391   19 408 254   20 409 274   21 300 519   22 351 341   23 325 281   24 404 273   25 333 428   26 299 291   27 506 387   28 304 282   29 300 292   30 296 196   31 408 370   32 317 292   33 306 362   34 217 332   35 290 391   36 279 493   37 256 401   38 603 428   39 230 295   40 216 398   41 606 293   42 159 281   43 300 196   44 404 493   45 238 392   46 404 338   47 541 602   48 279 394   49 302 428   50 336 292   

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