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Parathyroid hormone effects on marrow stromal cells for potential bone regeneration applications : delivery… Yang, Chiming 2009

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PARATHYROID HORMONE EFFECTS ON MARROW STROMAL CELLS FOR POTENTIAL BONE REGENERATION APPLICATIONS: DELIVERY SYSTEMS DEVELOPMENT AND BIOLOGICAL CHARACTERIZATION by  Chiming Yang  B.A.Sc. (Engineering Sciences  —  Biomedical Engineering), University of Toronto, 2001  M.A.Sc. (Chemical Engineering & Applied Chemistry and The Institute of Biomaterials and Biomedical Engineering), University of Toronto, 2003  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENT FOR THE DEGREE OF DOCTOR OF PHILOSOPHY  in The Faculty of Graduate Studies (Pharmaceutical Sciences)  •THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)  June 2009  © Chiming Yang 2009  ABSTRACT Despite the inherent ability of bone tissue to regenerate upon damage, there are incidences such as ‘critical’ defects where the damaged and lost bone will not repair or regenerate itself. Thus, the natural bone regeneration process must be augmented with the application of therapeutic agents (growth factors, hormones, cells). In addition, in every possible scenario where surgical interventions are performed, there exists the risk of infections that must be minimized and managed with the administration of antibiotics.  The overall goals of this thesis were to engineer, develop, and characterize biodegradable and bioresorbable polymeric microsphere and porous scaffold delivery systems for parathyroid hormone (PTH) and marrow stromal cells (MSC5), respectively, for enhancing the innate regenerative capacity of bone, and to investigate the effects of continuous and pulsatile PTH treatments on MSCs to better understand its regulatory actions on MSC differentiation, proliferation and clonogenicity. In addition, the development and characterization of biodegradable and bioresorbable polymeric microsphere delivery systems for the antibiotic, fusidic acid (FA) for potential localized application in bone infection were also undertaken.  PTH-loaded  poly(lactic-co-glycolic  acid)  (PLGA)  and  poly(hydroxybutyrate-co  hydroxyvalerate) (PHBV) microspheres were developed. However, these initial formulations did not achieve the precise level of controlled release of PTH required for MSCs. Osteogenic differentiation of MSCs was found to increase with continuous PTH treatment, and decrease with pulsatile PTH exposure. The observed effects of PTH were strongly dependent on the presence of dexamethasone. PTH treatments did not influence MSC proliferation but was  11  found to increase the colony forming unit-fibroblast (CFU-F) content within MSC cultures. Biocompatible, biodegradable and bioresorbable porous gelatin-alginate scaffolds produced by microwave vacuum drying were found to support MSC attachment, proliferation and differentiation. However, MSC differentiation (osteogenic, chondrogenic, adipogenic) were suppressed in vivo compared to in vitro when seeded on these scaffolds.  In the process of formulating FA-loaded PLGA and PHBV microspheres, an interesting phase separation phenomenon of FA in PLGA but not in PHBV polymer was observed. Phase separated FA formed distinct, large, completely amorphous, spherical FA-rich solid microdomains throughout the PLGA microsphere, and on the microsphere surface. FA release kinetics from the microsphere formulations were controlled by selective formulation factors determined from factorial design experiments.  Thus, the data presented in this thesis contribute to our understanding of PTH effects on MSCs, the responses of MSCs on porous gelatin-alginate scaffolds as well as the solid-state characteristics and release of FA loaded in PLGA and PHBV microspheres.  111  TABLE OF CONTENTS  ABSTRACT . ii iv  TABLE OF CONTENTS  Viii  LIST OF TABLES LIST OF FIGURES  ix  LIST OF ABBREVIATIONS  xv xvii  ACKNOWLEDGEMENTS CO-AUTHORSHIP STATEMENT CHAPTER  1.1  xx  Introduction  1  Project overview  1  1:  1.2 Marrow stromal cells 1.2.1 Biological characteristics of marrow stromal cells 1.2.2 Effects of growth factors on marrow stromal cell proliferation and differentiation 1.3 Parathyroid hormone in bone regeneration 1.3.1 Biological functions and activities of parathyroid hormone 1.3.2 Anabolic and catabolic effects of parathyroid hormone on bone  6 6  9 12 12 17  1.4 Delivery systems for growth factors and cells in bone regeneration applications.. 22 24 1.4.1 Controlled release of growth factors 1.4.2 25 Delivery of growth factors using microspheres 26 1.4.2.1 Protein loading methods 1.4.3 28 Porous scaffold delivery systems for cells 1.4.3.1 Porous scaffoldfabrication technologies 30 1.4.4 Current delivery strategies for growth factors and cells in bone regeneration 32 applications 1.4.5 Biomaterials selected for porous scaffold fabrication 35 1.4.5.1 Gelatin 35 38 1.4.5.2 Alginate Localized delivery of antibiotics for orthopaedic infection applications 1.5 1.5.1 Antibiotics in bone cement 1.5.2 Antibiotic loaded microspheres 1.5.3 Mechanisms of drug release 1.5.4 Factors influencing drug release 1.6  Development of fusidic acid loaded microspheres for bone infections  iv  41 42 43 44 47 49  1.6.1 Fusidic acid 1.6.1.1 Chemistry 1.6.1.2 Pharmacology 1.6.2 Biodegradable polymers selected for fusidic acid loaded microspheres 1.6.2.1 Poly(hydroxybutyrate-co-hydroxyvalerate) 1.6.2.2 Poly(lactic-co-glycolic acid) 1.6.3 Formulation approaches for localized delivery of fusidic acid  49 50 51 52 53 55 57  1.7 Research rationale, goals and objectives 1.7.1 Research rationale 1.7.2 Research goals 1.7.3 Research objectives  58 58 60 60  1.8  62  References  CHAPTER 2:  Effects of continuous and pulsatile parathyroid hormone on bone marrow stromal cells: Possible implications for bone regeneration 94 2.1  Introduction  94  2.2 Materials and methods 2.2.1 Chemicals 2.2.2 Isolation and cell culture of rat marrow stromal cells (MSC5) 2.2.3 Flow cytometry 2.2.4 Rat MSC differentiation assays 2.2.5 PTH treatment of rat MSCs 2.2.6 Alkaline phosphatase (ALP) activity 2.2.7 Histochemistry staining 2.2.8 Quantitative real time-polymerase chain reaction (qRT-PCR) 2.2.9 Cell count and MTT cell proliferation assay 2.2.10 Colony-forming unit fibroblast (CFU-F) assay 2.2.11 Dataanalysis  98 98 98 99 99 100 101 102 103 103 104 105  2.3 Results 106 2.3.1 Rat MSC characterization 106 2.3.2 Effects of PTH on rat MSC differentiation and proliferation in the presence of DEX 109 2.3.3 Effects of PTH on rat MSC differentiation and proliferation in the absence of DEX 115 2.3.4 Effects of PTH on rat MSC colony forming unit-fibroblast (CFU-F) 119 2.4  Discussion  122  2.5  Conclusion  127  2.6  References  128  CHAPTER 3: The differential in vitro and in vivo responses of bone marrow stromal cells on novel porous gelatin-alginate scaffolds 132  3.1  Introduction  132  v  3.2 Materials and methods 3.2.1 Materials 3.2.2 Fabrication of porous gelatin-alginate scaffolds using microwave vacuum drying 3.2.3 Mercury intrusion porosimetry and scanning electron microscope (SEM) analyses 3.2.4 Equilibrium water uptake measurements 3.2.5 Biodegradation and bioresorption 3.2.6 Isolation and cell culture of rat marrow stromal cells (MSCs) 3.2.7 In vitro culturing of MSCs on scaffolds 3.2.8 Implantation of MSC seeded scaffolds into NOD/SCID mice 3.2.9 Recovery of GFP MSCs from scaffolds 3.2.10 MSC proliferation assessment using BrdU incorporation 3.2.11 Immunohistochemistry 3.2.12 Quantitative real time-polymerase chain reaction (qRT-PCR) 3.2.13 Colony-forming unit fibroblast (CFU-F) assay 3.2.14 Dataanalysis  137 137  3.3 Results 3.3.1 Scaffold physical characterization 3.3.2 Scaffold bioresorption 3.3.3 Multi-potentiality of rat MSCs 3.3.4 In vitro and in vivo proliferation of MSC seeded on scaffolds 3.3.5 In vitro and in vivo differentiation of MSCs seeded on scaffolds MSCs CFU-F potential after in vitro culture and in vivo implantation 3.3.6  146 146 149 152 152 155 160  3.4  Discussion  161  3.5  Conclusion  166  3.6  References  167  .  137 138 139 139 140 141 141 142 142 143 144 144 145  CHAPTER 4: Fusidic acid loaded PLGA and PHBV microspheres for bone infections: Formulations development and solid-state characterization 172  4.1  172  Introduction  174 4.2 Materials and methods 174 4.2.1 Chemicals 4.2.2 Factorial design for formulation of FA-loaded PLGA and PHBV microspheres 174 4.2.3 Fabrication of FA-loaded PLGA and PHBV microspheres 175 4.2.4 176 Casting of FA and PLGA films 4.2.5 Preparation of amorphous FA drug 176 176 4.2.6 Microsphere particle size determination 4.2.7 177 FA encapsulation efficiency 4.2.8 177 In vitro FA release from PLGA and PHBV microspheres 4.2.9 Scanning electron microscope (SEM), backscattering SEM (BSEM), and laser confocal microscope analyses 178 178 4.2.10 Raman spectroscopy  vi  4.2.11 4.2.12 4.2.13 4.2.14  X-ray powder diffraction (XRPD) Differential scanning calorimetry (DSC) Micromanipulation and video imaging of microsphere formation Data analysis  179 179 180 181  4.3 Results 4.3.1 Factorial design of FA-loaded PLGA and PHBV microspheres 4.3.2 Surface characterization studies 4.3.3 Phase separation of FA in PLGA microspheres 4.3.4 Miscibility and phase separation of FA and PLGA in solvent-cast films 4.3.5 XRPD characterization of FA-loaded PLGA and PHBV microspheres 4.3.6 Thermal analysis of FA-loaded PLGA and PHBV microspheres 4.3.7 Drug release profiles of FA-loaded PLGA and PHBV microspheres  182 182 187 195 201 203 206 212  4.4  Discussion  217  4.5  Conclusion  224  4.6  References  225  CHAPTERS:  Summarizing discussion, conclusions and suggestions for future work... 230  5.1  Summarizing discussion and conclusions  230  5.2  Suggestions for future work  243  5.3  References  244  Appendix A: Parathyroid hormone loaded PLGA and PHBV microspheres: Formulations development and release kinetics  250  A.1 Materials and Methods A.1.1 Chemicals A.1.2 Fabrication of PTH-loaded PLGA and PHBV microspheres A. 1.3 Microsphere particle size determination A. 1.4 Scanning electron microscope (SEM) analyses A. 1.5 In vitro PTH release from PLGA and PHBV microspheres  250 250 251 254 254 254  A.2  Results  255  A.3  References  259  Appendix B: Additional figures for Chapter 4: Fusidic acid loaded PLGA and PHBV microspheres for bone infections: Formulations development and solid-state 260 characterization B.1  260  Additional Figures  Appendix C: ANOVA Result Tables for Factorial Design Experiments  265  Appendix D: UBC Research Ethics Board Certificates of Approval  269  vii  LIST OF TABLES  Table 3.1: Summary of the physical and bioresorption properties of the porous gelatin alginate scaffolds  148  Table 4.1: 2 full factorial design matrix and measured responses of FA-loaded PLGA micro spheres  183  Table 4.2: 2 full factorial design matrix and measured responses of FA-loaded PHBV microspheres  184  Table 4.3: Summary of the thermal properties of FA and FA-loaded PLGA (85/15) microspheres obtained using DSC  210  Table 4.4: Summary of the thermal properties of FA and FA-loaded PHBV (12% HV) microspheres obtained using DSC  211  Table A-i: Summary of formulation conditions for PTH loaded PLGA micro spheres. Polymer concentration used was 10% (w/v)  253  Table A-2: Summary of formulation conditions for PTH-loaded PHBV microspheres. Span 80 concentration used was 0.5% (v/v), where Span 80 = Sorbitan monooleate 253 Table C-i: ANOVA table for effect of polymer and drug concentration on the mean diameter of FA-loaded PLGA microspheres 266 Table C-2: ANOVA table for effect of (1) drug loading and (2) polymer and drug concentration on the encapsulation efficiency of FA-loaded PLGA micro spheres  267  Table C-3: ANOVA table for effect of polymer and drug concentration on the mean diameter of FA-loaded PHBV microspheres 268  viii  LIST OF FIGURES  Figure 1.1: Schematic representation of the amino acid sequence of parathyroid hormone (PTH)  14  Figure 1.2: Schematic representation of PTH regulatory effects on osteoblasts and osteoclast to maintain calcium homeostasis and bone turnover. In response to reduced ionized calcium concentration in the blood, PTH secreted by parathyroid cells exit the blood stream and binds to the PTH receptor 1 found in osteoblasts. PTH binding stimulate osteoblasts to increase their expression of RANKE, which can then bind to osteoclast precursors exiting the circulation. The binding of RANKL stimulates these precursors to fuse, forming new osteoclasts, which ultimately enhance the resorption of bone, releasing calcium in the process 16 Figure 1.3: Schematic representation of the proposed cellular mechanisms involved in the anabolic effect of pulsatile PTH. Pulsatile PTH has been proposed to increase osteoblast number by (A) increasing the development of osteoblasts, (B) inhibiting osteoblast apoptosis, (C) reactivating lining cells to resume their matrix synthesizing function, (D) inducing osteoblast progenitors to exit from the cell cycle for differentiation to osteoblasts in response to locally produced autocrine/paracrine growth factors, and (E) favoring MSCs differentiation towards osteoprogenitors at the expense of adipogenic differentiation 20 Figure 1.4: Schematic representation of a typical amino acid sequence of gelatin  36  Figure 1.5: Chemical structures of alginate. (A) f3-D-mannuronic acid (M) and cx-L glucuronic acid (G) monomer units of alginate, (B) typical copolymer structure of alginate.39 Figure 1.6: Schematic representation of the mechanisms of polymer degradation. (A) Bulk degradation and (B) surface degradation 46 Figure 1.7: Chemical structure of fusidic acid (FA)  50  Figure 1.8: Chemical structure of poly(hydroxybutyrate-co-hydroxyvalerate) (PHBV) copolymer  53  Figure 1.9: Chemical structure of poly(lactic-co-glycolic acid) (PLGA) copolymer  55  Figure 2.1: Representative FACS dot plot illustrating the population distribution of rat MSCs after 3 5 passages. Hematopoietic cell population (< 4%) gated for both GFP and CD45 is shown in the upper right quadrant 107 —  Figure 2.2: Multi-potentiality of rat MSCs. Histochemical staining results of the differentiation of (A) rat MSCs (GFP) along three lineages after 3 weeks. (B) Osteogenic (mineralized bone nodule, arrowhead), (C) adipogenic (intracellular fat-containing vacuoles, arrows), and (D) chondrogenic (cartilage) 108  ix  Figure 2.3: Effects of continuous and pulsatile PTH treatments on ALP activity and staining in the presence of DEX. MSC were cultured for up to seven 48-h cycle resulting in (A) 4 days, (B) 8 days, and (C) 14 days exposure to PTH. PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5. *]J < 0.05 as compared to control 110 Figure 2.4: Gene expression profiles of MSC after continuous and pulsatile PTH treatments in the presence of DEX. MSC were cultured for up to seven 48-h cycle illustrating the changes in (A) PTHR1 and (B) ALP gene expressions after 8 and 14 days exposure to PTH. PTH-C = continuous PTH and PTH-P pulsatile PTH as described in Section 2.2.5. *] < < 0.05 as compared to 0.05 as compared to control. #p < 0.05 as compared to PTH-P. PTH-C 113 Figure 2.5: Proliferation kinetics of MSC after continuous and pulsatile PTH treatments in the presence of DEX. MSC were cultured for up to seven 48-h cycle illustrating the changes in (A) total number of cells and (B) the proliferation kinetics as measured by the MTT assay, monitoring absorbance at 570 mm PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5 114 Figure 2.6: Effects of continuous and pulsatile PTH treatments on gene expressions in the absence of DEX. MSC were cultured for seven 48-h cycle (i.e. 14 days). (A) PTHR1 and (B) ALP gene expression. PTH-C continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5 116 Figure 2.7: Effects of continuous and pulsatile PTH treatments on ALP activity in the absence of DEX. MSC were cultured for seven 48-h cycle (i.e. 14 days). PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5  117  Figure 2.8: Effects of continuous and pulsatile PTH treatments on cell proliferation kinetics in the absence of DEX as measured by MTT assay, monitoring absorbance at 570 nm. MSC were cultured for seven 48-h cycle (i.e. 14 days). PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5 118 Figure 2.9: Effects of continuous and pulsatile PTH treatments on CFU-F by bulk and limiting dilution assays. MSC were exposed to continuous and pulsatile PTH (10 or 100 nM) for three to four 48-h cycles (i.e. 6 8 days), (A) in expansion medium without DEX, (B) in osteogenic medium without DEX, and (C) in osteogenic medium with DEX before cells were re-plated and cultured for another 6 8 days in expansion medium only to allow for the formation of CFU-F. PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5. < 0.05 as compared to control 121 —  —  Figure 3.1: Schematic diagram of the microwave vacuum drying process to fabricate porous scaffolds. (A) Formulation of hydrogel with or without additional agents such as cross linkers and drugs. (B) Hydrogel cast into molds. (C) Hydrogel is placed into microwave vacuum drying chamber, where electromagnetic microwave energy penetrates into the hydrogel and is converted to thermal energy for the subsequent in situ vaporization of water which generates an expansive force to create porous structures in the matrix. (D) Dry porous scaffolds 134  x  Figure 3.2: Digital photographs and SEM micrographs of high cross-linked porous gelatin alginate scaffolds. (A) and (C) are top views, (B) side view, and (D) cross-sectional view of the scaffold 147 Figure 3.3: Representative images of the biodegradation and bioresorption of high and low cross-linked porous gelatin-alginate scaffolds before and after subcutaneous implantation in NOD/SCID mice for 7 and 21 days. [Note: Scaffolds were rehydrated in culture medium before implantation which resulted in the pink colouration.] 150 Figure 3.4: Representative image of the neovascularization (arrows) surrounding high crosslinked porous gelatin-alginate scaffolds after subcutaneous implantation in NOD/SCID mice for 21 days. [Note: Also notice on the right-hand side the absence of the low cross-linked scaffold which was completely bioresorbed after 21 days, with only a dashed-circle representing the missing scaffold.] 151 Figure 3.5: Representative fluorescence microscope images of MSCs on high cross-linked porous gelatin-alginate scaffold after 1 week (A) in vitro culture (Hoechst stained, arrows), and (B) in vivo subcutaneous implantation (GPF, arrows) 153 Figure 3.6: MSC proliferation after 7 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin-alginate scaffolds measured using BrdU incorporation in proliferating cells and analyzed using FACS. Mean ± SD,n=3. 154 Figure 3.7: MSC osteogenic gene expressions after 7 and 21 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin alginate scaffolds. Mean ± SD, n 3, *p <0.05 compared to in vivo at the same time point. N/D = not detected 157 Figure 3.8: MSC chondrogenic gene expressions after 7 and 21 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin alginate scaffolds. Mean ± SD, n = 3, *p <0.05 compared to in vivo at the same time point. N/D = not detected 158 Figure 3.9: MSC adipogenic gene expression after 7 and 21 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin alginate scaffolds. Mean ± SD, n = 3, *p <0.05 compared to in vivo at the same time point. 159 Figure 3.10: MSC CFU-F forming ability after 8 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin-alginate scaffolds. Mean ± SD, n =4 for CFU-F 160 Figure 4.1: Formulation factors that influence FA-loaded PLGA microspheres. (A) Effects of polymer and drug concentration on mean diameter,p <0.0001, (B) effects of polymer and drug concentration on the encapsulation efficiency of FA, p <0.01, and (C) effects of initial drug loading on encapsulation efficiency of FA, p < 0.01. Results are expressed as mean ± 95% CI 185 xi  Figure 4.2: The effect of polymer and drug concentration on the mean diameter of FA-loaded PHBV microspheres, p < 0.0001. Results are expressed as mean ± 95% CI 186 Figure 4.3: SEM images illustrating the surface morphologies of the eight FA-loaded PLGA microsphere formulations described in Table 4.1 189 Figure 4.4: SEM images illustrating the effects of different drug loading on FA-loaded PLGA (85/15) microsphere surface morphologies. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer(w/w) 190 Figure 4.5: SEM images illustrating the surface morphologies of the eight FA-loaded PHBV microsphere formulations described in Table 4.2 191 Figure 4.6: SEM images illustrating the effects of different drug loading on FA-loaded PHBV (12% HV) microsphere surface morphologies. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w) 192 Figure 4.7: Detailed surface and interior morphologies of 30% (w/w) FA loaded PLGA (85/15) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) Laser confocal microscope images of PLGA microsphere, (B) BSEM images of PLGA microsphere, and (C) BSEM images of sectioned PLGA microsphere 193 Figure 4.8: Detailed surface morphologies of 30% (w/w) FA loaded PHBV (12% HV) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A C) BSEM images of PHBV microsphere at different magnifications 194 —  Figure 4.9: Raman spectroscopy images of FA distribution in 30% (w/w) FA-loaded PLGA (85/15) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light microsphere montage, 50x magnification, (B) distribution of PLGA-rich regions (green) across microsphere, (C) distribution of FA-rich regions (red) across microsphere, and (D) combined distribution of the two regions across microsphere 197 Figure 4.10: SEM images illustrating the effects of different drug loading on FA-loaded PLGA (85/15) microsphere surface morphologies (A D) and the changes after 7 days of drug release (E H). All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w) 198 —  —  Figure 4.11: Time lapsed video images illustrating the formation of a single 30% (w/v) FA loaded PLGA (85/15) microsphere with an initial polymer and drug concentration of 10% (w/v). The initial FA!PLGA/DCM droplet was blown from a micropipette into 0.O1M SDS aqueous solution at room temperature. Arrows illustrates the phase separated FA-rich microdomains, while star indicate the micropipette. The full video showing the FA phase separation phenomenon and microsphere solidification process is available upon request to the author of this thesis. [Note: for scale, screen height is 25 Elm] 199  xii  Figure 4.12: Raman spectroscopy images of FA distribution in 30% (w/w) FA loaded PHBV (12% HV) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light microsphere montage, 20x magnification, (B) distribution of PHBV (green) across microsphere, and (C) distribution of FA (red) across microsphere 200 Figure 4.13: SEM images of different weight % FA in PLGA (85/15) films solvent-cast from DCM illustrating the miscibility characteristics of FA and PLGA. All films were cast at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w) 202 Figure 4.14: XRPD patterns of solid-state (A) FA as received, (B) 30% (w/w) FA loaded PLGA (85/15) microspheres, and (C) control (no drug) PLGA (85/15) microspheres. All microspheres were manufactured at a polymer and drug concentration of 10% (w/v) 204 Figure 4.15: XRPD patterns of solid-state (A) FA as received, (B) PHBV (12% HV) polymer as received, (C) control (no drug) PHBV (85/15) microspheres, and (D) 30% (w/w) FA loaded PHBV (12% HV) microspheres. All microspheres were manufactured at a polymer and drug concentration of 10% (w/v) 205 Figure 4.16: DSC thermograms of(A) first heating cycle of FA loaded PLGA (85/15) microspheres illustrating the enthalpy relaxation temperature, T of the PLGA polymer and the glass transition temperature, Tg of FA, (B) second heating cycle (after quench cooled) of FA loaded PLGA (85/15) microspheres showing the Tg of the PLGA polymer. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w) 207 Figure 4.17: DSC thermogram of pure amorphous FA drug illustrating the glass transition temperature, Tg 208 Figure 4.18: DSC thermograms of FA loaded PHBV (12% HV) microspheres illustrating the glass transition temperature, Tg and double melting temperature, Tmi and Tm2 of the PHBV polymer. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w) 209 Figure 4.19: FA release profiles from 10% FA loaded PLGA microsphere formulations. See Table 4.1 for the details of each formulation (Run) 213 Figure 4.20: FA release profiles from 30% FA loaded PLGA microsphere formulations. See Table 4.1 for the details of each formulation (Run) 214 Figure 4.21: FA release profiles from 10% FA loaded PHBV microsphere formulations. See Table 4.2 for the details of each formulation (Run) 215 Figure 4.22: FA release profiles from 30% FA loaded PHBV microsphere formulations. See Table 4.2 for the details of each formulation (Run) 216  xlii  Figure A-i: Schematic diagram of the water-in-oil-in-water (W/O/W) double emulsion technique to encapsulate growth factors (hydrophilic drugs). (I) Growth factor dissolved in an aqueous solvent is emulsified in a non-miscible organic polymer solution to form the first water-in-oil (W/O) emulsion. (II) The primary W/O emulsion is transferred to an excess secondary aqueous medium containing a stabilizer, at which point homogenization again or intensive stirring is applied to form the water-in-oil-in-water (W/O/W) double emulsion. (III) Stabilization of the double emulsion is achieved by constant mechanical agitation (stirring) and subsequent removal (evaporation) of the organic solvent. (IV) Hardening of the polymer surrounding the growth factor to produce growth factor-loaded micro spheres 252 Figure A-2: SEM images of parathyroid hormone (PTH) loaded PLGA microspheres. See Table A-i for details of each run 256 Figure A-3: SEM images of parathyroid hormone (PTH) loaded PHBV microspheres. See 256 Table A-2 for details of each run Figure A-4: Cumulative in vitro release profiles of PTH loaded PLGA microspheres in PBS (pH 7.4) @ 37 °C, n = 3 See Table A-i for details of each run 257 Figure A-5: Cumulative in vitro release profiles of PTH loaded PHBV microspheres in PBS (pH 7.4) @ 37 °C, n = 3. See Table A-2 for details of each run 258 Figure B-i: SEM images illustrating the phase separation of FA from different PLGA composition matrices. PLGA (50/50) = poly(lactic-co-glycolic acid) with 50/50 lactic/glycolic acid molar ratio. PLGA (85/15) = poly(lactic-co-glycolic acid) with 85/iS lactic /glycolic acid molar ratio. PLLA = poly(L-lactic acid) polymer, where it can be consider to be PLGA (100/0) with iOO% lactic acid. All microsphere formulations were manufactured at a polymer and drug concentration of iO% (w/v) and 30 % FA loadings (w/w) 261 Figure B-2: Raman spectroscopy images of FA distribution in 30% (w/w) FA-loaded PLGA (50/50) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light micro sphere montage, 1 OOx magnification, (B) distribution of PLGA-rich regions (green) across microsphere, and (C) distribution of FA-rich regions (red) across microsphere. 262 Figure B-3: Raman spectroscopy images of FA distribution in 30% (w/w) FA-loaded PLLA microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light microsphere montage, lOOx magnification, (B) distribution of PLLA-rich regions (green) 263 across microsphere, and (C) distribution of FA-rich regions (red) across microsphere Figure B-4: SEM images illustrating the effects of emulsif’ing agents on the phase separation of FA from PLGA (85/15) microspheres. PVA = poly(vinyl alcohol). All micro sphere formulations were manufactured at a polymer and drug concentration of i 0% (w/v) and 30% FA loadings (w/w) 264  xiv  LIST OF ABBREVIATIONS aFGF  Acid fibroblast growth factor  ALP  Alkaline phosphatase  aP2  Acid-binding protein 2  bFGF  Basic fibroblast growth factor  BMP  Bone morphogenetic protein  BSA  Bovine serum albumin  BSP  Bone sialoprotein  CFU-F  Colony forming unit-fibroblast  ECM  Extracellular matrix  FA  Fusidic acid  FGF  Fibroblast growth factor  LPL  Lipoprotein lipase  MIC  Minimum inhibitory concentration  MRSA  Methicillin-resistant staphylococcus aureus  MSC  Marrow stromal cell  MSCs  Marrow stromal cells  OC  Osteocalcin  PDGF  Platelet-derived growth factor  PHA  Polyhydroxyalkanoate  PHB  Poly(hydroxybutyrate)  PHBV  Poly(hydroxybutyrate-co-hydroxyvalerate)  p1  Isoelectric point  PLGA  Poly(lactic-co-glycolic acid)  PMMA  Poly(methylmethacrylate)  PPARy  Peroxisome proliferator-activated receptors-y  PTH  Parathyroid hormone  PTHR1  PTH receptor 1  PVA  Poly(vinyl alcohol)  xv  RANK  Receptor activator of nuclear factor-id3 ligand  RANKL  Receptor activator of nuclear factor-id3  S. aureus  Staphylococcus aureus  S. epidermidis  Staphylococcus epidermidis  Sox9  SRY (sex determining region Y)-box 9  Tg  Glass transition temperature  TGF-b  Transforming growth factor-b  Tm  Melting temperature  Tr  Enthalpy relaxation temperature  VEGF  Vascular endothelial growth factor  W/O  water-in-oil  W/O/W  water-in-oil-in-water  xvi  ACKNOWLEDGEMENTS I would like to express my sincerest gratitude to my supervisor Dr. Helen Burt. The completion of this project would not have been possible without her research expertise, continuous guidance and encouragement. Dr. Burt was not only an excellent supervisor that gave me all the necessary resources and critical evaluations to complete my research, she was also a very good friend that provided many insightful advices into my professional and personal development.  I would also like to give special thanks to my research and PhD defense committee members: Drs. Kathleen MacLeod, Urs Hafeli, Fabio Rossi, Thomas Oxiand, Goran Ferniund, Michael Underhill, and Rizhi Wang for their effort, invaluable suggestions and recommendations. Particularly, I want to thank Dr. Fabio Rossi for accepting me as an honorary member of his lab and provided me with facilities and resources for all cell culture and animal experiments, in addition to his scientific expertise and thoughtful discussion.  Moreover, I wish to thank Dr. Hanspeter Frei for his training and participation in cell culture and animal experiments, for inspiring discussion and motivation, for his value points of view on research and life, and for his wonderful friendship. Thank you to John Jackson for his technical expertise, informative scientific discussion and invaluable perspectives. I am also grateful for the technical expertise of Lin Yi for help with cell culture experiments, histological staining, and for organizing all necessary supplies.  xvii  In addition, I would like to acknowledge Dr. Timothy Durance for the shared microwave vacuum drying technology and thank Parastoo Yaghmaee for her informative discussion and helpful advice on the scaffold fabrication and characterization. Furthermore, I would like to thank Dr. John McNeill and his graduate student Linda Tran, who graciously allowed me to use their radioactive facilities, and provided assistance with the initial setup and training. Thanks you to Anita Lam in UBC Chemistry for her expertise on x-ray powder diffraction analysis and data interpretation, Dr. David Plackett in Risø National Laboratory for Sustainable Energy, Technical University of Denmark and Dr. Tim Smith in Renishaw plc, Wotton-under-Edge, UK for their expertise and the access of their confocal Raman microscopes, and Dr. David Needham in Department of Mechanical Engineering and Material Science, Duke University for his micropipette experimental setup, data collection and expertise on single microsphere formation.  Thank you to Julia Lin and Ben Wasserman for their hard work through the summers. Thanks to all my past and present colleagues in Drs. Burt and Rossi laboratories for making the labs fun and memorable. Kevin Letchford, Sam Gilchrist (and the SeaBass Production), Leon Wan, Clement Mugabe, Antonia Tsallas, Michelle Chakraborti, Lucy Ye, Christ Springate, Tobi Higo, Karen Long, John Lu, Rita Zhao, Praveen Elamanchili, Melanie ter Borg, Katherine Haxton, Wes Wong, Aaron Joe, Yasmine Even, Leslie So, Stephane Corbel and Andy Johnson, thanks you all for your help, suggestions, support, friendship and laughter.  xviii  Finally, I am truly grateful to my family for their timely encouragements, unwavering support and unconditional love over the years. Specially, I am extremely thankful to Carol Chen for her enthusiastic support, enduring patience, and devoted love.  For financial support, I would like to thank the National Sciences and Engineering Research Council of Canada (NSERC) and University of British Columbia for scholarships. Project funding from the Canadian Institutes of Health Research (CIHR) New Emerging Team (NET) Grant is also gratefully acknowledged.  xix  Co-AUTHORSHIP STATEMENT This thesis is comprised of three manuscripts of which I am the principal author. In all instances, I was the primary individual responsible for the identification and design of the research program, conduction of all practical aspects of the research experiments, analysis and interpretation of the findings, and preparation of the manuscripts. The contribution of co authors was through the provision of intellectual discussion and editorial assistance.  xx  Chapter 1: Introduction 1.1 Project overview In humans, the skeletal system is composed of over 200 bones that are extremely dynamic and well-organized [1]. Bone tissue formation occurs throughout the lifespan of an individual beginning from embryonic development, and continuing with growth and remodeling [2]. Despite the inherent ability of bone tissue to regenerate upon damage, there are incidences such as ‘critical’ defects found in trauma, tumors, lesions, and congenital abnormalities as well as in revision total hip replacements where the lost and damaged bone will not regenerate or repair itself.  There is a great cost, both economic and human, associated with the clinical need to find effective treatments for bone regeneration, estimated to be in the billions of dollars [3]. Moreover, with increased life expectancy and the striking age demographic changes occurring in Canada, it is clear that the incidence of bone injuries and joint arthroplasties will undoubtably increase dramatically. The number of people age 65 and over was 13.7% of the population in 2008 [4], and will increase to 23.4% by 2031, according to Statistics Canada projections [5].  Current clinical treatments for the restoration of the normal structure and function of the bony deficits that occur in orthopaedic injuries and joint arthroplasties rely on the use of surgical bone graft transplantations from various natural sources [6, 7]. Although autograft transplantation (from the patient) is the gold standard for orthopaedic treatments, it is  1  restricted by its limited supply, requirement for a secondary surgery, and the possible morbidity and increased post-operative pain to the patient [8, 9]. Other treatment alternatives including allografts (from human donors) and xenograft transplantations (from non-human sources) might resolve the issue of limited supply but introduce the risk of disease transmission and immunogenic responses [10-13]. Where surgical interventions are required for the enhancement and promotion of bone regeneration, the risks of infection are real and must be minimized and managed using antibiotics. Bacterial infection remains a major concern when there is the use of orthopaedic implants, despite modern day sterilization and aseptic procedures. The reported infection rates are in the range of 0.5 arthroplasties [14-16], but much higher rates, 2  —  —  5% for total joint  20% for revision procedures have been  documented [17]. In addition, infections often result in the removal of the orthopaedic implants and the need for follow-up operations [14].  This thesis is focused on two main areas of study; investigation of the delivery of stem cells (marrow stromal cells) and a hormone (parathyroid hormone) for bone regeneration, and localized delivery of an antibiotic (fusidic acid) for bone infection.  Marrow stromal cells (MSC5) also known as mesenchymal stem cells and multipotent stromal cells [18-23] are non-hematopoietic stromal cells that can be harvested relatively easily, isolated from adult bone marrow and various other tissues, and have high proliferative capacity [24]. In addition, the ability of ex vivo expanded MSCs to generate multiple cell types including osteoblasts, to facilitate the processes of bone repair and regeneration is well established [23], making these cells an attractive source for tissue regeneration and tissue  2  engineering [24-301. The successful regeneration of tissues in numerous animal models has led to preclinical and clinical trials using MSCs for bone repair and regeneration [24, 31, 321.  Moreover, studies have demonstrated that bone repair and regeneration were significantly enhanced when stimulatory factors, along with MSC, were delivered concurrently [33-3 51. Parathyroid hormone (PTH), the only clinically approved anabolic therapy for osteoporosis [361 has stimulated tremendous research interest due to its dual anabolic and catabolic actions on bone formation. PTH has been shown to enhance bone regeneration, in for example, fracture healing  [371  and may increase the relative osteogenic progenitor content of  MSC cultures to better promote bone tissue regeneration. However, the effects of different regimens of PTH treatment on the differentiation and proliferation of MSCs have not been investigated.  The primary method of delivery of MSCs for bone tissue regeneration has been the use of three-dimensional porous scaffolds engineered from biocompatible and biodegradable biomaterials such as gelatin [38,  391  and alginate [40]. In an attempt to mimic the natural  extracellular matrix (ECM), scaffolds provide the initial framework for MSCs to attach, survive, proliferate, and differentiate to form new ECM [411. The scaffold not only retains the delivered MSCs within the defect site, it can also function as a substrate for tissue ingrowth and vascularization.There have been many in vitro and in vivo studies that have demonstrated tissue regeneration with the delivery of MSCs on porous scaffolds, but few have directly compared in vitro and in vivo responses including proliferation and  3  differentiation of MSCs on scaffolds [42, 43] and have not explicitly examined the fate and changes in the MSC population after in vivo transplantation.  Clinically, treatment [44-47] and prevention [48-51] of orthopaedic infections are accomplished primarily with prolonged systemic antibiotic therapy. However, local delivery of antibiotics can offer significant advantages in the management and prevention of orthopaedic infections [52]. The antibiotic, fusidic acid (FA) has been available since the 1960s [53] and is most active against Staphylococcus aureus, Staphylococcus epidermidis, and coagulase-negative staphylococci, including strains that are methicillin-resistant that commonly cause prosthetic joint infections [54]. Nevertheless, systemic delivery of FA can lead to many undesired side effects such as hyperbilirubinemia and thrombophlebitis [55-57]. We have proposed the localized and controlled delivery of FA to orthopaedic surgical sites using polymeric microspheres technology for the management and prevention of bone infections. There are no documented studies using this approach.  The goals of this project were to develop and evaluate localized delivery systems for PTH and MSCs for bone regeneration applications, and FA for bone infection applications. Although some early formulation work was undertaken to develop controlled release polymeric microspheres for PTH, it was evident in the early phase of the work (drug release and cell culture studies), that very precise control of exposure of MSC to PTH (continuous and pulsatile) was required and was not achievable using PTH-loaded microspheres. Accordingly, the first part of the thesis examines the effects of continuous and pulsatile (intermittent) PTH treatment on the osteoprogenitor content of rat MSCs in vitro, to better  4  understand the regulatory effects of PTH on MSC differentiation, proliferation and clonogenicity. In the second part of the thesis, studies were focused around understanding the in vitro and in vivo responses of MSCs seeded on porous gelatin-alginate scaffolds in the absence of any additional growth factors or hormones, such as PTH. The final part of the thesis outlines the formulation development and characterization of FA-loaded biodegradable polymeric microspheres, as the first step in developing formulation approaches for antibiotics with potential for localized treatment of bone infections.  5  1.2 Marrow stromal cells Osteogenesis takes place via a multi-step differentiation cascade thought to originate from multipotent mesenchymal cells and proceeding through osteoprogenitors and osteoblasts to generate mature bone osteocytes [58]. Marrow stromal cells (MSCs) also referred to as mesenchymal stem cells, multipotent stromal cells and mesenchymal stromal cells [18-23] are non-hematopoietic stromal cells first isolated from bone marrow by Friedenstein and colleagues [59, 60]. The extent to which MSCs participate in the regeneration of tissues such as bone, cartilage, and fat during the adult lifespan is the subject of tremendous research activity and has been extensively reviewed [20, 23, 24, 27, 61-63]. Moreover, the relative ease of isolation and the ability of in vitro expanded MSCs to generate multiple cell types, including osteoblasts, chondrocytes, and adipocytes is well established [19, 64-66] and makes these cells an attractive source for tissue regeneration and tissue engineering studies [24-301. The successful regeneration of tissues in animal models has led to preclinical and clinical trials using MSCs for bone repair and regeneration [24, 31, 32].  1.2.1  Biological characteristics of marrow stromal cells  As first demonstrated by Friedenstein et al., MSCs preferentially adhere to plastic and display a fibroblast-like morphology [60]. When a single-cell suspension is plated at low density, MSCs form colonies, with each colony derived from a single progenitor cell defined as a Colony Forming Unit-Fibroblast (CFU-F) [67, 68]. Phenotypically, MSCs express numerous surface markers, but unfortunately none are specific to MSCs. Nonetheless, it is generally agreed that MSCs do not express hematopoietic markers such as CD45, CD34,  6  CD 14, and CD1 lb. Furthermore, MSCs are devoid of the costimulatory molecules CD8O, CD86, CD4O, and the adhesion molecules CD3 1 (platelet/endothelial cell adhesion molecule [PECAM]-l), CD18 (leukocyte function-associated antigen-i [LFA-1]), and CD56 (neuronal cell adhesion molecule-i) [64, 65, 69]. On the other hand, MSCs express a number of surface receptors including CD 105 (SH2 or endoglin), CD73 (SH3/4), CD44, CD90 (Thy-i), CD7 1, CD1 17 (c-kit), and STRO-1 as well as the adhesion molecules CD49e, CD62, CD 106 (vascular cell adhesion molecule [VCAM- 1]), CD 166 (activated leukocyte cell adhesion molecule [ALCAM]), intercellular adhesion molecule (ICAM)-1, and CD29 [64, 65, 69]. However, the expression of these markers is variable due to the differences in tissue source, species of origin, isolation method, and culture conditions. Expression is also influenced by secreted factors from accessory cells in the initial passages of the MSCs [64, 65].  A hallmark characteristic of MSCs is their capacity to be induced to differentiate along multiple mesenchymal lineages including osteogenic (bone), chondrogenic (cartilage) and adipogenic (fat) in vitro [65, 69]. Osteogenic differentiation of MSCs to osteoblasts involves culturing of a confluent monolayer of MSCs with ascorbic acid, -glycerophosphate, and dexamethasone for 2  —  3 weeks. Osteoblasts are identified by the formation of proteoglycan  rich bone nodules that stain positive with alizarin red and Von Kossa techniques, and the increased expression of alkaline phosphatase (ALP), bone sialoprotein (BSP), and osteocalcin (OC), as well as the accumulation of calcium over time [70]. For chondrogenic differentiation, MSCs are cultured in a pelleted micromass in the presence of transforming growth factor-13 (TGF-J3) to differentiate into mature chondrocytes that produce an abundance of glycosaminoglycans within the extracellular matrix, which stain intensely with toluidine  7  blue and alcian blue. The differentiated chondrocytes also express aggrecan and SRY (sex determining region Y)-box 9 (Sox9) [70]. To promote adipogenic differentiation, MSCs are cultured with dexamethasone and insulin, leading to adipocyte differentiation and accumulation of lipid-rich vacuoles within the cell cytoplasm that can be stained with oil red 0. Increased expression of peroxisome proliferator-activated receptors (PPARy), lipoprotein lipase (LPL), and acid-binding protein 2 (aP2) is also observed in the differentiated adipocytes [70]. In addition to the mesenchymal lineages, MSCs under appropriate in vitro conditions have been shown to differentiate along other pathways [71], to form tenocytes [72], skeletal myocytes [73, 74], neurons [19, 75], endothelial cells [76, 77], and epithelial cell types [19].  While bone marrow was the first source of MSCs to be described, MSCs and MSC-like cells with similar biological characteristics have been isolated from various other tissue sites including skeletal muscle [78, 79], adipose tissue [80], amniotic fluid [81], synovial membrane [82], fetal tissues [83, 84], umbilical [85] and peripheral blood [86, 87]. Even in bone marrow, one of the most abundant sources, MSC frequency is quite low [88], 10,000 nucleated cells of newborn [21], and declines with age to  -  1 in  1 in 2,000,000 nucleated  cells in an 80 year old [21, 89-91]. MSC frequency also varies with species [92]. Nonetheless, under optimum culture conditions such as low-cell density plating or addition of growth factors, MSCs have a tremendous self-renewal capacity and can readily expand more than a thousand-fold in a relatively short period, generating billions of cells from a limited amount of starting materials and still maintaining their differentiation potentials [9396].  8  It is important to note that the MSC population described in the literature to date, is heterogeneous in nature due to the lack of specific and unique phenotypic markers for the identification of MSCs [97]. Current isolation techniques are based on MSC preferential adherence to plastic surfaces, together with enrichment by expansion and passaging in relatively deprivational, serum-only containing medium [95, 96] and isolation with monoclonal antibodies such as STRO-l [98, 99] or CD49a molecule [100]. However, the resulting cultures may still contain cell types displaying a range of developmental potentials and at different stages of commitment [19, 20, 64, 65].  1.2.2  Effects of growth factors on marrow stromal cell proliferation and differentiation  Whether MSCs self-renew and proliferate as multipotent stem cells capable of differentiating in multiple mesenchymal lineages, or differentiate into committed progenitors of a particular lineage, is likely regulated by a myriad of both systemic and local stimulatory factors including growth factors and hormones [63, 88, 101]. Among the numerous growth factors that have been shown to affect MSC proliferation and differentiation, bone morphogenetic proteins (BMPs) are the most studied and have been clinically approved for bone regeneration [102, 103]. BMPs are a 15-member subfamily of the transforming growth factor-f3 (TGF-13) super family of growth factors [103]. In bone morphogenesis, BMP-2, -4,  -  6, -7 and -9 are known to play critical roles, with recombinant BMP-2 and -7 being used clinically for spinal fusion and long bone non-unions, respectively [104, 105]. BMP-2, -4, and -7 have been shown to stimulate MSC differentiation to osteoblasts to improve bone  9  healing [106]. Studies have demonstrated that the addition of BMP-2 to MSCs cultures in vitro induces osteoblast differentiation, increasing bone nodule formation and calcium content [107, 108], as well as enhancing the bone-forming capacity of implanted MSCs in vivo [109].  Another key growth factor that modulates the fate of MSCs is fibroblast growth factor (FGF) [110-112]. FGFs are a family of 11 growth factors characterized by their affinity for the glycosaminoglycan heparin-binding sites of cells [113]. The most abundant FGFs are acidic FGF (aFGF or FGF-1) and basic FGF (bFGF or FGF-2), both promoting the proliferation and differentiation of a variety of cells including osteoblasts. Specifically, bFGF is a potent modulator of MSCs, capable of increasing MSC proliferation rate and maintaining MSC differentiation potential during initial in vitro culture. bFGF likely selects for a more primitive subpopulation of osteoprogenitor cells in the heterogeneous MSC cultures and favors its self-renewal and proliferation, eventually enhancing bone formation in vivo [114, 115]. In addition, bFGF was shown to increase the proliferation rate and prolong the life-span of MSCs without affecting their differential potentials [94, 116]. In other studies, bFGF was found to increase both CFU-F number and size, and maintain the fibroblast-like morphology of MSC [115, 117]. Furthermore, Hanada et al. demonstrated that bFGF can have synergistic effects on the stimulation of MSCs proliferation and differentiation with BMP-2 [108].  Relative to growth factors, the effects of hormones and, in particular, parathyroid hormone (PTH), on the proliferation and differentiation of MSCs have not been clearly delineated [118]. PTH is a polypeptide of 84 amino acids that regulates calcium homeostasis and bone  10  turnover. Only a few in vitro studies have examined PTH interactions with MSCs, and they were performed with the addition of other growth factors and stimulatory agents. Simmons et , and found enhanced osteogenic 3 al. cultured MSCs with PTH, BMP-6 and vitamin D differentiation [119]. When cultured under adipogenic conditions containing insulin, isobutyl-methylxanthine and troglitazone, the addition of PTH in a pulsatile (intermittent) manner,  suppressed  adipogenic  differentiation  and  possibly promoted  osteogenic  differentiation in MSCs [120]. In contrast to in vitro experiments, published in vivo studies have highlighted the complex and sometimes conflicting effects of PTH on MSC. Whereas daily administration of PTH was not shown to affect the number of MSCs extracted from femoral marrow isolates from mice [121-123], other studies demonstrated that daily injection of PTH in rats increased MSC proliferation [124, 125]. Using a tissue engineered bone growth model in vivo, where ectopic ossicles were generated from subcutaneous implanted MSCs, Pettway et al. demonstrated that PTH injections increased MSC proliferation in mice [126]. The differences observed with in vivo MSC proliferation after PTH administration might be due to the effects of different animal models. In various in vivo animal models, administration of PTH has been shown to promote osteogenic differentiation of MSCs, albeit indirectly through PTH-induced paracrine factors [125, 127, 128].  Thus, an understanding of how growth factors and hormones ultimately control MSC self renewal and proliferation versus differentiation is of great importance for any tissue regeneration applications of MSCs.  11  1.3 Parathyroid hormone in bone regeneration All musculoskeletal tissues secrete and respond to growth factors, cytokines and hormones that are involved in injury, disease, and repair processes as well as normal growth and development [129, 1301. One hormone in particular, parathyroid hormone (PTH), is a critical regulator of calcium concentration and bone metabolism and has drawn a lot of research interest due to its dual anabolic and catabolic actions on bone formation. Although its mechanism of action is complex and not fully understood [131], PTH is the only clinically approved anabolic therapy for osteoporosis [36, 132]. Furthermore, PTH has also demonstrated beneficial effects in bone regeneration during fracture repair and implant fixation [37]. Thus, an enhanced understanding of the functions of PTH in bone regeneration will likely lead to more widespread use.  1.3.1  Biological functions and activities of parathyroid hormone  Parathyroid hormone (PTH) (isoelectric point, p1 9), amino acid composition illustrated in Figure 1.1, is an endogenous polypeptide of 84 amino acids, synthesized and secreted by cells of the parathyroid glands, and is a key endocrine regulator of bone metabolism. The main function of PTH is to regulate calcium homeostasis by maintaining the calcium-ion ) concentration of the extra- and intracellular fluids within physiological limits, with the 2 (Ca overall effect of conserving calcium [133]. PTH acts directly on the skeleton to promote calcium and phosphate release from bone, while it acts on the kidney to enhance calcium reabsorption and inhibit phosphate reabsorption [134]. Furthermore, PTH acts indirectly on  12  the intestine to increase calcium absorption, which is mediated by the increase in 1,25dihydroxyvitamin D 3 formation in the kidney [135].  13  LLspAGIy.  ProALLGuAspAVaILVaIGIuSerHisGIuLSerLeGIyGIuAIaAsp  Asp  Vaif  Vai  Thr  L  Al  Ser  Gin  H  Figure 1.1: Schematic representation of the amino acid sequence of parathyroid hormone (PTH)  The liberation of calcium from the skeleton is through a process of enhanced bone resorption by osteoclasts mediated by osteoblast signaling [1361 and is illustrated schematically in Figure 1.2. In response to reduced ionized calcium concentration in the blood, parathyroid cells secrete intact PTH, which may undergo further cleavage and still remain active, providing the N-terminus is preserved and at least 31 amino acids remain [137]. Intact and active PTH fragments then bind to the PTH receptor 1 (PTHR1) found on osteoblasts [137]. PTH binding stimulates osteoblasts to increase their expression of receptor activator of nuclear factor-KB ligand (RANKL), which can then bind to osteoclast precursors containing receptor activator of nuclear factor-icB (RANK), a receptor for RANKL. The binding of RANKL to RANK stimulates these precursors to fuse, forming new osteoclasts which ultimately enhance the resorption of bone, releasing calcium in the process [137]. Similarly, an increase in extracellular calcium concentrations is also detected by parathyroid cells, inhibiting the secretion of PTH to achieve overall calcium homeo stasis [137].  15  .  •.•.• Parathyroid Hormone  Red Blood Cells  New Lining Cells  Bone  Osteoclasts  i  Medullary Cavity Osteoblasts  Cortical Compact Bone Cancellous Spongy Bone  I Bone Marrow  Figure 1.2: Schematic representation of PTH regulatory effects on osteoblasts and osteoclast to maintain calcium homeostasis and bone turnover. In response to reduced ionized calcium concentration in the blood, PTH secreted by parathyroid cells exit the blood stream and binds to the PTH receptor 1 found in osteoblasts. PTH binding stimulate osteoblasts to increase their expression of RANKL, which can then bind to osteoclast precursors exiting the circulation. The binding of RANKL stimulates these precursors to fuse, forming new osteoclasts, which ultimately enhance the resorption of bone, releasing calcium in the process.  As stated above, PTH-fragments based on differing numbers of amino acid, with a preserved N-terminus, PTH (1-3 1), PTH (1-34), and PTH (1-38) have all shown similar potency and pharmacological profile as the full length hormone [136]. The recombinant human PTH (134) N-terminus fragment [1381 and the full length recombinant human PTH (1-84) [135] are both clinically approved anabolic therapies for osteoporosis.  1.3.2  Anabolic and catabolic effects of parathyroid hormone on bone  PTH is a multifunctional and complex hormone that can affect bone differentially. It exerts both anabolic and catabolic effects on bone, depending on the nature and duration of the exposure, but the mechanisms underlying these actions are not well understood and are the subject of numerous studies [134, 139-142]. Pulsatile (intermittent) delivery of PTH has been shown to lead to increased bone mass [143], mineral density [144], mechanical strength [145, 1461, as well as improved bone microarchitecture [147], fracture healing [148, 149], and implant fixations [150], whereas continuous administration of PTH results in bone resorption and hypercalcemia [151, 152].  The dual anabolic and catabolic effects of PTH were first demonstrated by Selye, where once daily injections of parathyroid gland extract resulted in denser bone formation in rats, but increased doses led to a bone resorption response [153]. Subsequent to these data and the eventual purification of PTH [154], extensive research has been conducted on the anabolic effects of PTH in animals and humans (see reviews [139-141, 155]). The anabolic effects of PTH were achieved by repeated transient (intermittent or pulsatile) exposure to the skeleton because PTH was cleared from the circulation within 2  17  —  3 hours after injections [156, 157].  As a result of understanding this unique exposure-related mechanism of action, PTH was clinically approved for anabolic therapy in the treatment of osteoporosis [138]. Pulsatile (daily) subcutaneous injections of PTH (1-34) and PTH (1-84) have been demonstrated to increase bone mineral density, reduce vertebral and non-vertebral fracture risk in postmenopausal women, elderly men, and women with glucocorticoid-induced osteoporosis [135, 158-1641.  Beyond its promising effects in treating osteoporosis, the anabolic actions of pulsatile PTH can also have beneficial effects in fracture healing by increasing callus formation and mechanical strength of the repairing bone [145, 149, 165-167]. Furthermore, in response to the need for bone repair and bone formation to facilitate fixation of orthopaedic implants, Skripitz et al. showed that pulsatile administration of PTH increased the density of regenerating bone and reduced the areas of soft tissue at the implant-tissue interface to enhance the fixation of stainless steel implants in a dose- and time-dependent manner [168, 169]. Skripitz et al. also demonstrated that pulsatile administration of PTH increased the bonding tensile strength of bone to poly(methylmethacrylate) (PMMA) bone cement [1701. The use of pulsatile PTH treatment to accelerate the consolidation of newly generated bone after distraction osteogenesis, a technique for bone lengthening, was investigated by Seebach et al. Their results suggested PTH might be useful as a stimulator of bone formation in order to improve regenerated bone stability while consolidation occurs [1711.  The cellular mechanisms that control the anabolic effects of pulsatile PTH treatment are not well understood. Pulsatile PTH treatment first stimulates bone formation and subsequently  18  stimulates both resorption and formation, but the overall balance remains positive with augmented bone formation [172, 173]. Even so, the degree of bone anabolism caused by pulsatile PTH can be limited by its ability to also stimulate bone resorption, establishing a feedback ioop that contributes to the gradual loss of the anabolic effect with continued administration of the hormone [160]. The proposed cellular mechanisms involved in the anabolic effect of pulsatile PTH are shown in Figure 1.3. Published evidence suggests that there is a positive correlation between the anabolic effects of pulsatile PTH to stimulate bone formation and an increase in osteoblast numbers and activity (action (A) in Figure 1.3) [139]. The increased number of matrix-synthesizing osteoblasts has been shown to result from numerous possible actions by PTH. Predominately, pulsatile PTH has a direct action in delaying osteoblast apoptosis (action (B) in Figure 1.3) resulting in a corresponding increase in osteoblast numbers and this effect has been demonstrated to account for the anabolic effects [123, 174, 175]. Activation of resting bone lining cells by pulsatile PTH to become osteoblasts has also been proposed (action (C) in Figure 1.3) [176, 177]. The inmiediate actions of pulsatile PTH on osteoprogenitors to increase osteoblast numbers are still unclear [118, 139], but it has been shown that short term exposure to PTH can induce osteoblast progenitors to exit from the cell cycle (action (D) in Figure 1.3), at which point, paracrine regulator molecules may then promote osteoblast differentiation [178, 179]. It has also been proposed that pulsatile PTH may favour the differentiation of MSCs toward osteogenic progenitors at the expense of other lineages, such as adipogenesis (action (E) in Figure 1.3), to enhance osteogenesis [120].  19  >  Parathyroid Hormone  CeUs  Bone  Osteoblasts  Apoptotic Osteoblasts  e.  e  Osteoprogeniters Expansion of Osteoprogeniters  Adipocytes  Bone  :  Figure 1.3: Schematic representation of the proposed cellular mechanisms involved in the anabolic effect of pulsatile PTH. Pulsatile PTH has been proposed to increase osteoblast number by (A) increasing the development of osteoblasts, (B) inhibiting osteoblast apoptosis, (C) reactivating lining cells to resume their matrix synthesizing function, (D) inducing osteoblast progenitors to exit from the cell cycle for differentiation to osteoblasts in response to locally produced autocrine/paracrine growth factors, and (E) favoring MSCs differentiation towards osteoprogenitors at the expense of adipogenic differentiation.  20  The mechanism whereby continuous PTH exerts the catabolic effect on bone is even less well understood. Even though bone deposition occurs with continuous PTH treatment, bone resorption still offsets bone deposition leading to an overall bone loss [177]. Continuous PTH treatment does not affect the life span nor the number of osteoblasts [175]. However, it has been suggested that continuous PTH treatment increases the expression of RANKL by osteoblasts to increase osteoclast numbers and enhance osteoclastogenesis [175, 180-182]. Bellido et al. suggested that the reduced expression of sclerostin by osteocytes, mediated by continuous treatment PTH was another possible mechanism for the hormonal control of osteoblastogenesis by PTH [180].  Therefore, the dual anabolic and catabolic effects of PTH on bone is complex, with the duration of PTH exposure (pulsatile vs. continuous) rather than the peak or total concentration determining the final anabolic or catabolic response [156].  21  1.4 Delivery systems for growth factors and cells in bone regeneration applications Bone is a dynamic tissue that is continuously remodeled due to the balance between the activities of two different cell populations, the osteoclasts and the osteoblasts that are responsible for bone resorption and deposition, respectively [183]. When bone is damaged, it has the capacity to repair itself, provided the damage does not result in defects of a ‘critical size’. However, in cases of bone defects, lesions, trauma, and congenital abnormalities where there are severe bone tissue losses resulting in critical-sized bony deficits, the damaged bone will not regenerate spontaneously [184], resulting in severe pain and disability for millions of people worldwide. Moreover, with increased life expectancy and aging of the population, the incidence of bone injuries will undoubtedly increase, with a greater clinical need for effective treatments [1851.  Restoration of the normal structure and function requires replacement of the missing bone and this may be accomplished by surgical bone graft transplant of natural tissue from various sources [7, 186, 187]. Autograft transplantation has long been considered the gold standard procedure because of its immunocompatibility and has demonstrated its importance in enhanced bone-healing, spinal fusion, and fracture repair [188]. This is due to the fact that autografts contain a host of osteogenic bone cells including MSCs, osteoprogenitors and osteoblasts, an osteoconductive collagen matrix (natural scaffold) suitable for new and existing cell attachment and migration, as well as osteoinductive proteins and growth factors endogenous to bone [8, 189]. Yet, autografts are still constrained by several factors such as limited quantity of donor tissue, and the requirement for secondary surgery at the site of  22  tissue harvest that might possibly lead to morbidity and increased post-operative pain [8, 9]. Other than autografts, alternative approaches include allograft and xenograft transplantations that are obtained from human donors and other species, respectively [6, 1901. Although the issue of limited supply is resolved, there exists a minimal but real risk of disease transmission and immunogenic responses with allografts and xenografts [10-131. For these reasons, a potential solution to the clinical problem could be bone tissue engineering, wherein the selfrepairing capacity of bone is augmented to elicit the regeneration that would not proceed otherwise [20, 24, 32, 188, 189, 191, 1921.  The bone repair and regeneration process is a complex cascade of biological events controlled by numerous growth factors, cytokines and honnones that provide local signals to regulate and direct the mechanisms of tissue regeneration [193, 1941. The regeneration process involves an increase in the number of matrix-forming cells to ultimately reconstruct the extracellular matrix (ECM) that will remodel to bone. Locally secreted growth factors first mediate the migration of MSCs to the injured site, regulate their differentiation to osteoprogenitors  and  subsequently  direct  the  proliferation and  differentiation  of  osteoprogenitors towards osteoblasts to initiate the regeneration sequence that includes revascularization, ECM production and remodeling [28, 62, 1181. Consequently, bone tissue engineering attempts to replicate this natural regeneration process by using the combination of growth factors, cells, and biodegradable scaffolds [195, 196]. Delivery of growth factors stimulates endogenous and exogenous cell migration, proliferation and differentiation and delivery of exogenous cells compensates for the deficiency in the number or function of endogenous cells. Biodegradable scaffolds function as three-dimensional delivery matrices  23  for growth factors and cells, and allows for cellular and vascular infiltration to regenerate the tissue.  1.4.1  Controlled release of growth factors  For endogenous and exogenous (transplanted) cells to regenerate tissues, they would need to perform a multitude of functions such as migration, proliferation, and differentiation, all of which are driven and orchestrated by growth factors, cytokines and hormones [1971. Growth factors have been infused into the systemic circulation or given via a bolus injection at the defect site. However, due to their short half-life, potential non-target tissue toxicity, and rapid diffusion from the application site [1981, these delivery methods have not met the physiological requirements of the tissue repair processes and may have contributed to the lack of success of clinical trials [199-2011.  Local delivery of growth factors has been shown to be clinically successful but there is still substantial room for improvement. The local delivery of BMP-2 (Infuse®; Medtronic Sofamor Danek, Memphis, Tennessee) and BMP-7 (OP-i Device®; Stryker Biotech, Hopkinton, Massachusetts) for improving bone-healing [102, 103], has been achieved by simply dispersing the growth factors onto collagen scaffolds. However, this has provided relatively little control over the rate of delivery, conformation, presentation, clearance, or degradation of the delivered growth factors [202]. Furthermore, the inefficiency of these delivery strategies is highlighted by the very large supraphysiological doses of BMPs that are required (e.g. 3500 ig of BMP-7), since only a small fraction actually comes in contact with target cells to stimulate new bone formation [202]. As a consequence, it has been suggested  24  that controlled release delivery systems are needed to mimic the in vivo release profiles of growth factors produced during natural bone tissue morphogenesis or repair.  In designing controlled release strategies, growth factor immobilization via association or tethering of growth factor directly to the scaffold through covalent linkages [203-205] or physical adsorption (complexation) [206] have been evaluated for tissue regeneration. The growth factor is placed directly in the cell microenvironment to avoid mass transfer limitations and reduces potentially undesirable diffusion to distant sites resulting in systemic toxicity. However, immobilization can result in growth factor denaturation and improper presentation of the active region that leads to reduced bioactivity. There are numerous other strategies for the controlled delivery of growth factor for tissue engineering and they have been extensively reviewed [195, 207-222]. One of the most common and straightforward methods by which growth factors have been incorporated into polymers for controlled release is through the formation of microspheres [223, 224] as described below.  1.4.2  Delivery of growth factors using microspheres  Encapsulation of growth factors in polymeric microsphere delivery systems can achieve localized and controlled delivery to improve bone regeneration [195, 196, 209, 225]. The use of biodegradable polymeric microspheres for the local delivery of growth factors not only provides controlled release that reduce dosing frequency and the total amount of growth factor administered, it also protects the growth factors from a proteolytic environment which can cause denaturation, unfolding and cleavage [198, 209]. In addition, the release profile of the growth factor can be modulated by changing the physicochemical properties of the  25  encapsulating polymer, such as molecular weight [226] and polymer composition, to better match that of the regeneration process. Methods of loading growth factors for delivery using biodegradable polymers are described below.  1.4.2.1 Protein loading methods The most commonly used methods for loading growth factors into biodegradable polymers are spray drying, phase separation (coacervation), and double emulsion, with each approach having their own advantages and disadvantages [227]. In the spray drying method, biodegradable polymers such as poly(lactic-co-glycolic acid) (PLGA) are dissolved in a volatile organic solvent (dichioromethane or acetone) and lyophilized or precipitated dehydrated growth factor particles are suspended and dispersed in the polymer solution by high speed homogenization or sonication [228]. The dispersion is then atomized using a stream of heated air, forming microdroplets where the solvent evaporates instantaneously yielding growth factor-loaded micro spheres in sizes ranging from 1  —  100 pm depending on  the atomizing conditions [227]. The resulting microspheres are collected from the air-stream by a cyclone separator and residual organic solvents are then removed by vacuum drying. The advantages of spray drying include a well-defined control of particle size, control of drug release profiles, and an ability to tolerate small changes in the polymer specifications. Conversely, the disadvantages of spray drying include the use of high temperatures and lyophilized protein with the possibility of protein aggregation and denaturation [227].  The use of phase separation (coacervation) to encapsulate growth factors in microspheres is accomplished by first dispersing lyophilized or dehydrated solid protein particles in a  26  polymer solution in an organic phase (dichioromethane) similar to the initial steps of spray drying. However, silicon oil is then added to the dispersion at a defined rate, reducing the polymer solubility in its own solvent (dichioromethane) to create a polymer-rich liquid phase (coacervate) that encapsulates the dispersed protein particles. Initially formed microspheres are then subjected to hardening and washing using a secondary organic solvent such as heptane. Although phase separation avoids the use of high temperature, it still requires the use of lyophilized protein that might be aggregated or denatured prior to encapsulation. Moreover, the extensive use of organic solvents increases the likelihood of residual solvent in the final microspheres and the phase separation process itself is quite sensitive to changes in the polymer properties [2271.  The water-in-oil-in-water (W/O/W) double emulsion technique [2291 has been used extensively to encapsulate growth factors in microspheres for controlled release [230-232]. In this process, growth factor dissolved in an aqueous solvent is emulsified in a non-miscible organic polymer solution to form the primary water-in-oil (W/O) emulsion [233]. Typically, volatile dichloromethane is used as the organic solvent and the homogenization step to form the emulsion is carried out using either a high speed homogenizer or a sonicator. The primary W/O emulsion is then rapidly transferred to an aqueous medium containing a stabilizer, usually poly(vinyl alcohol) (PVA), at which point homogenization or intensive stifling is applied to form the water-in-oil-in-water (W/O/W) double emulsion. Subsequent removal (evaporation) of the organic solvent by heat, vacuum or both results in phase separation and hardening of the polymer to produce growth factor-loaded microspheres. Alternatively,  27  instead of solvent evaporation, solvent extraction can be used to produce microspheres loaded with growth factors [234].  The major advantages of the double emulsion method include high yields and encapsulation efficiencies and it is an easily scalable process [227]. However, the denaturation of growth factors at the organic-aqueous interface in the primary W/O emulsion has been the principal obstacle to maintaining protein integrity and bioactivity during encapsulation [235, 236]. Several methods have been used to stabilize proteins during the emulsification process. One method involves the addition of surfactants or proteins such as albumin to minimize the fraction of the desired protein at the interface [237-240]. For example, in the encapsulation of recombinant human growth hormone in PLGA microspheres (Nutropin®, Genentech, California, USA) polysorbate 20 was added to the formulation to stabilize the protein [241], and bovine serum albumin, added to the aqueous phase of insulin-growth factor, was found to significantly reduce its degradation in PLGA microspheres [242]. Another method to stabilize protein in the emulsion process is the addition of small molecule osmolytes such as trehalose or mannitol in the protein phase, which stabilizes the proteins by preferential hydration of its native conformation during the removal of water from the encapsulated protein solution [243].  1.4.3  Porous scaffold delivery systems for cells  The engineering of biomaterials into three-dimensional (3-D) structures, referred to as scaffolds, attempts to mimic key features of native ECM, and is essential for the success of local cell delivery in tissue regeneration [221, 244, 245]. Scaffolds perform a multitude of  28  functions in tissue regeneration ranging from physical support for the development of new functional tissues to vehicles for cell delivery, and the release of therapeutic biomolecules [20, 32, 189, 192, 195, 221].  Scaffolds provide the initial framework for cells to attach, survive, proliferate, and differentiate to form new ECM [41]. Scaffolds must also be both macroscopically stable and microscopically dynamic; two opposing yet complimentary characteristics possessed by natural ECM. For macroscopic stability, scaffolds must create a space and serve as a spaceholder to prevent encroachment of undesired surrounding tissues into the regenerating site until new tissue can form, and provide a substrate for cell adhesion and migration [202]. The microscopic dynamics of tissue formation, however, require that scaffolds must also be degradable to enable cellular remodeling of the microenvironment. Finally, scaffolds must allow for vascular ingrowth that provides both nutrient delivery and waste removal [246]. Therefore, the ideal scaffold system for the delivery of cells should have the following properties [244]: •  biocompatible, non-toxic, non-inflammatory and non-immunogenic,  •  biodegradable in synchrony with tissue regeneration or capable of being remodeled,  •  bioresorbable, where degraded by-products do not elicit any adverse reactions and are metabolized by host cells via normal physiological pathways,  •  architectures and porosity to allow host blood vessel colonization and vascularization of the entire structure,  •  very porous and highly permeable to allow for proper transport of nutrients and metabolites,  •  appropriate pore sizes and surface properties for the candidate cells infiltration, adhesion, survival, migration, and proliferation,  29  •  adequate mechanical properties to provide the correct microstress environment for cells or mechanical support to the implantation site,  •  reproducible fabrication and sterilizable.  1.4.3.1 Porous scaffoldfabrication technologies Porous scaffolds can have porosities ranging from as low as 30% to more than 95%, and pore sizes ranging from ten to hundreds of microns [247, 248]. High porosity and interconnected pore structure not only ensures that migrating cells and infiltrating vascular networks can penetrate throughout the entire scaffold but also provides void space for ECM formation, and the transport of nutrients, metabolic waste and biodegradation by-products [249]. A number of fabrication technologies have been applied to process biomaterials into solid 3-D scaffolds of high porosity and surface area for cells delivery [189, 250-252]. Conventional and innovative techniques for scaffold fabrication include, solvent casting combined with particulate leaching [253], emulsion freeze-drying [254], rapid prototyping methods (laser stereolithography, 3-D inkjet printing, selective laser and heat sintering) [250, 25 5-259], and the more recently developed microwave vacuum drying technique [260, 261], among others.  Solvent casting, in combination with particulate leaching is a very simple and common technique to fabricate porous scaffolds. First described by Mikos and associates [262, 263], this procedure involves dispersing water-soluble particles of defined size, such as salt, sugars, or polymer spheres, in a matrix consisting of the scaffold material dissolved in an organic solvent. The solvent is evaporated under vacuum, resulting in the polymer solidifying around the particles. The entrapped water-soluble particle can then be leached out of the scaffold by numerous rinses in water, thus creating a defined pore structure. However, this  30  process works only for thin membranes or 3-D specimens with very thin wall sections (few millimeters), otherwise, it is not possible to remove the soluble particles from within the polymer matrix [225]. Other disadvantages include the existence of a “skin layer” of nonporous polymer at the surface, extensive use of toxic organic solvents, time required for solvent evaporation and particle leaching (days-to-weeks), residual particles in the matrix, and insufficient interconnectivity [225].  The emulsion freeze-drying technique involves creating an emulsion by homogenizing a polymer solvent solution and water. The mixture is then rapidly quenched in liquid nitrogen, and the solvent and water are removed by freeze-drying. Advantages of this technique include the ability to control pore sizes from 15 to 200 im and the ability to obtain porosity> 90% [225]. Conversely, the disadvantages of the emulsion freeze-drying method are the closed pore structure in the resulting matrix and length of time required for drying (days).  Rapid prototyping systems are some of the most promising scaffold fabrication techniques that can process a wide range of biomaterials into complex and custom-made shapes [257, 264-267]. For example, in laser stereolithography, intricate 3-D computer models of the tissue to be regenerated are first interpreted by a rapid-prototyping machine and then fabricated into the corresponding scaffold by combining a photopolymerizable polymer in a liquid, layer by layer, with each subsequent layer being built on top of the previous one to form the entire 3 -D structure, at which point, a final curing step is performed to complete the fabrication [268]. Although rapid prototyping systems possess the ability to fine tune the scaffold porosity, pore size and shape, and have a completely interconnected pore network,  31  these techniques still have drawbacks that include the use of high temperatures and solvents during processing that could affect or compromise any added growth factors or drugs [257].  Recently, Durance et al. [269, 270] have developed a novel, versatile, rapid and reproducible method, which also avoids the use of toxic organic solvents and high temperatures that can be used to fabricate highly porous scaffolds from a number of gel-forming biomaterials such as gelatin and alginate. This technique is known as microwave vacuum drying and is described in Chapter 3 of this thesis. Briefly, a hydrogel matrix is subjected to microwave radiation under vacuum. The water boils at a low temperature and vaporization creates porous structures in the matrix.  1.4.4  Current delivery strategies for growth factors and cells in bone regeneration applications  As noted earlier, bone regeneration is a dynamic process involving a cascade of signaling growth factors, cytokines, and hormones released at various concentrations, times and combinations. Thus, controlled release at a constant rate of a single stimulatory factor will not mimic the physiological cues needed for normal regenerative processes. Researchers are now beginning to design more sophisticated delivery systems that can deliver growth factors with variable release profiles such as pulsatile (on-off) release, and multiple growth factors with different release kinetics to better match the signals that regulate the mechanisms and pathways governing bone regeneration [221]. For example, given that PTH was known to elicit an anabolic effect on bone when delivered intermittently, but a catabolic effect when delivered continuously, Liu et al. designed and fabricated an implantable device that  32  delivered PTH in a pulsatile manner [2711. The delivery system, fabricated using alternating PTH-loaded alginate and surface-erodable polyanhydride isolation film layers, demonstrated multi-pulse PTH release with the lag time between two adjacent pulses modulated by the composition and the thickness of the polyanhydride film layer [2711. Using a similar approach, Jeon et al. produced pulsatile PTH releasing devices that were made from alternating PTH-loaded and blank microsphere layers pressure-sintered into cylinders [272]. Sequential and alternating delivery of two bioactive molecules, both with pulsatile kinetics were demonstrated by Lui et al. [271]. Alternating pulses of PTH and simvastatin hydroxyacid were achieved by Jeon et al. by pressure-sintering alternating PTH- and simvastatin hydroxyacid-loaded microsphere layers [273].  Additional temporal and spatial control of release via the combination of different delivery systems, such as the incorporation of protein-loaded microspheres into protein-containing scaffolds, has provided distinct release kinetics for multiple growth factors and has shown enhanced tissue formation [274, 275]. Richardson et al. designed a single delivery system to release vascular endothelial growth factor (VEGF) and platelet-derived growth factor (PDGF) with differential kinetics for promoting angiogenesis [275]. The PLGA scaffold containing both growth factors was fabricated by mixing PDGF-loaded PLGA microspheres together with lyophilized VEGF and PLGA particles, then pressure-sintering, gas-forming and particulate-leaching to form porous scaffolds. VEGF was largely associated with the surface of the scaffold and was rapidly released first, whereas, the PDGF was encapsulated into microspheres, distributed throughout the matrix, and released much slower with its release kinetics regulated by the degradation of the different PLGA polymers used to form  33  the microspheres [275]. Dual delivery of VEGF and PDGF led to a high density of blood vessels and the formation of thicker, larger and more mature vessels relative to the delivery of either growth factor alone [275]. Using similar PDGF-loaded microspheres embedded into VEGF-containing scaffold system, Chen et al. developed an anisotropic system based on a porous bi-layered PLGA scaffold that exposed VEGF in one spatial region, and delivered VEGF and PDGF in another adjacent region, leading to significantly enhanced blood vessel size and maturation [274].  In order to better reproduce the natural bone tissue morphogenesis and development process, Mooney’s group have simultaneously delivered multiple growth factors and cells, to mimic the complexity of the regeneration processes [40, 276]. Alginate scaffolds containing a combination of BMP-2 and TGF-f3, along with MSCs were mixed together, cross-linked and subcutaneously implanted into SCID mice to investigate bone formation [40]. They showed that the delivery of dual growth factors, along with MSCs, produced more effective and efficient bone growth than either individually delivered BMP-2 or TGF-133 [40]. In other work, VEGF, condensed plasmid DNA encoding for BMP-4, and MSCs were delivered via a scaffold to enhance bone regeneration [276]. VEGF and plasmid DNA were mixed with PLGA particles, and scaffolds were fabricated using a high pressure gas foaming process to produce porous scaffolds for subsequent MSC seeding. The combined local angiogenic (VEGF), and sustained osteogenic (BMP-4) factors with MSCs were shown to produce the greatest quantity and quality of bone formation compared to individual factors or MSCs alone, indicating that the cells and growth factors may have acted in an additive or synergistic manner to regenerate bone tissue [276].  34  1.4.5  Biomaterials selected for porous scaffold fabrication  The design and selection of a biomaterial is a critical step in the development of scaffolds for tissue regeneration applications [277]. The ideal properties of a scaffold biomaterial have been discussed in Section 1.4.3. Both natural and synthetic polymers have been proposed to facilitate the regeneration of damaged or dysfunctional bone tissues. However, since natural biopolymers such as gelatin and alginate possess properties that most closely resemble those of ECM, they hold considerable advantages as scaffold materials. In addition, the biodegradation by-products are biocompatible, they generally do not elicit any adverse reactions, and are readily metabolized by cells [278, 279].  1.4.5.1  Gelatin  Gelatin is a naturally occurring polymer that is derived from collagen. Its biocompatibility and tolerability is well documented, and it is commonly used for food, pharmaceutical, and medical applications [206, 280-282]. Gelatin is an amphoteric protein that is non-toxic, nonimmunogenic, non-inflammatory, biodegradable, bioresorbable, and inexpensive to produce. It is a heterogeneous mixture of single or multi-stranded polypeptides, each with extended left-handed proline helix conformations and containing between 300  —  4000 amino acids.  Each strand is composed of a unique sequence of amino acids and contains a large number of glycine (almost 1 in 3 residues, arranged every third residue), proline and 4-hydroxyproline residues. A typical amino acid sequence is -alanine-glycine-proline-arginine-glycine glutamic acid-4-hydroxyproline-glycine-proline-, shown in Figure 1.4.  35  Figure 1.4: Schematic representation of a typical amino acid sequence of gelatin  Gelatin is obtained from the partial hydrolysis of Type I collagen isolated from animal skin, tendons, ligaments, and bone, as well as fish skin extracted with very dilute acid. There are two types of gelatin, depending on whether or not the preparation involves an alkaline pre treatment, which converts the amide groups of the asparagine and glutamine residues to their respective carboxylic acids, aspartic acid and glutamic acid, and results in higher viscosity. Alkaline pre-treatment makes use of bovine hides and bones to produce acidic (Type B) gelatin with an isoelectric point (p1) between 4.8 to 5.2. Acid pre-treatment does little to affect the amide groups present, and generates basic (Type A) gelatin, with a p1 ranging from 7 to 9, commonly derived from porcine and fish skins [283].  Gelatin is susceptible to most proteases, but they do not break gelatin down into peptides containing much less than 20 amino acids. Aqueous gelatin solutions undergo a progressive hydrolytic degradation, depending on temperature and pH. Hydrolytic degradation of gelatin proceeds most slowly at neutral pH. The acid pre-treated (Type A) gelatin is more susceptible to alkaline degradation than acid degradation, while alkaline pre-treated (Type B) gelatin is the reverse [283]. Gelatin forms a thermoreversible gel with water. Above a concentration of --1% (w/v), at temperatures above 32 °C, gelatin forms a homogeneous  36  solution in aqueous or any hydrogen bond forming solvent. However, it forms a physical gel when cooled to room temperature. With further increase in the temperature above the gelation temperature, 32°C, gelatin is soluble again [2841. Thus, gelatin-based delivery systems for long-term biomedical applications must be cross-linked to improve the stability of the polymer [206, 280, 282]. Porous gelatin scaffolds have been used as a delivery vehicle for MSCs in bone [38, 39], cartilage [285, 286], and adipose [287] tissue regeneration studies. Furthermore, gelatin microspheres encapsulating either growth factors [288-293] or cells [294-296] and incorporated into a secondary scaffold or matrix for tissue engineering applications have been investigated [278].  There are several commercially available gelatin-based biomedical products that are being used in tissue engineering applications [2781. These include Gelfoam® (Pfizer, USA) and Surgifoam® (Ethicon Inc., USA), which are highly porous, absorbable gelatin sponges derived from porcine skin used as a hemostatic agent, CultiSpher-G and S® (Percell Biolytica AB, Sweden), which are macroporous gelatin beads used as microcarrier cell culture systems, and Gelfilm® (Pfizer, USA) that is an absorbable gelatin film designed for neuro, thoracic and ocular surgeries. In a study by Ponticiello et al., Gelfoam® was used as a delivery vehicle for MSCs in cartilage regeneration therapy. Adult human MSC were seeded throughout the porous gelatin scaffold after a 2-h incubation period. When cultured for 21 days in vitro the MSC-loaded gelatin construct produced a cartilage-like ECM containing sulfated  glycosaminoglycans  and  type-IT  collagen.  Following  implantation in  an  osteochondral defect in the rabbit femoral condyle, the porous gelatin scaffolds were shown  37  to be biocompatible, with no evidence of immune response or lymphocytic infiltration at the site [285].  1.4.5.2 Alginate Alginate is a naturally derived anionic polysaccharide copolymer consisting of individual blocks of f3-D-mannuronic acid (M units) and cL-L-glucuronic acid (G units), interspersed with random copolymers of M and G units, shown in Figure 1.5. The length of the M- and G blocks and their random distribution along the polymer chain varies depending on the source of the alginate [297, 298]. Commercially, alginate is mainly extracted from the structural components of three different species of marine brown algae, although the capsular polysaccharides in some soil bacteria are another source [299]. Similar to gelatin, alginate is biocompatible,  non-toxic,  non-immunogenic,  non-inflammatory,  biodegradable,  bioresorbable, and inexpensive to produce, making alginate another widely used natural polymer for pharmaceutical and biomedical applications [297, 299, 300]. Alginate also shows good hemocompatibility [301].  38  A OHQ  13.-D-mannuronate (M)  a-L.guluronate (G)  B  M  M  M  -OOC M  C  M  Figure 1.5: Chemical structures of alginate. (A) 13-D-mannuronic acid (M) and a-L glucuronic acid (G) monomer units of alginate, (B) typical copolymer structure of alginate.  Alginate can undergo a reversible gelation in aqueous solution where the G blocks of adjacent polymer chains cooperatively bind with divalent cations such as Ca , or Ba 2 , 2 , Sr 2 but not Mg 2 and monovalent cations, resulting in the formation of ionic inter-chain bridges which leads to gelling, almost independent of temperature. The overall gel stiffness depends on the polymer composition (i.e., the M/G ratio), and the stoichiometry of the alginate with the chelating cation [297]. In general, alginates with higher G content develop stiffer and more porous gels that maintain their integrity longer, whereas alginates rich in M residues result in softer and less porous gels that tend to dissolve more readily [302]. Alginate gels gradually and uncontrollably dissolve, losing their mechanical integrity, due to the movement of cross-linking divalent cations from the alginate hydrogel to the surrounding medium. This process is prevented by introducing covalent cross-links, similar to that of gelatin hydrogel,  39  to immobilize the polymer chains via covalent bonds [277]. Alginate degradation can be controlled by manipulating polymer molecular weight and composition, or even by partial oxidation, to create reactive side groups that spontaneously form covalent linkages with the closest hydroxyl groups in the chains [303].  Alginate hydrogels and microspheres are currently being evaluated for several orthopaedic applications including growth factor delivery and MSC delivery [40, 286], cell encapsulation [304, 305], and tissue regeneration [306, 307]. In terms of commercially available products, alginate has been mainly used for wound dressings such as NuDenn® (Johnson & Johnson, USA), Curasorb® (Kendall, USA), and AlgiSite® (Smith & Nephew, USA). Some tissue engineering driven products are now beginning to emerge, such as the alginate-recovered chondrocyte (ARCTM) process (Articular Engineering LCC, USA) which uses alginate beads for cell delivery [278].  40  1.5 Localized delivery of antibiotics for orthopaedic infection applications Orthopaedic surgeries are routinely performed to restore structure and function to millions of people disabled by accidents and diseases. However, in every possible surgical scenario, the risks of infection are real, even with today’s advancements in surgical techniques, ultra-clean operation rooms and development of new antibiotics [308]. For example, prosthetic joint infections [17] such as those seen in hip replacements are difficult to treat because they are often associated with necrosis of bone, resorption of bone matrix and poor vascular perfusion accompanied by infection of the surrounding soft tissues [309, 310]. Clinically, the first line of defense for the treatment [44-47] and prevention [48-51] of orthopaedic infections has been prolonged systemic antibiotic therapy. However, serious problems can arise, including a failure to produce therapeutic tissue concentrations of the antibiotics because of relatively low vascularity within necrotic bone and implant sites in prosthetic joint infections, as well as the potential for the development of bacterial resistance and deep-seated mycoses [46, 311].  Local delivery of antibiotics offers significant advantages in the management and prevention of orthopaedic infections [312, 313]. By direct application of antibiotics to the site of infection or potential infection, it is possible to achieve higher tissue levels and for a longer period of time, while simultaneously avoiding systemic side effects. These high local levels of directly applied antibiotics facilitate delivery by diffusion to avascular areas of the wound that are inaccessible by systemic antibiotics, which often can only be delivered in concentrations that may result in resistance [314].  41  1.5.1  Antibiotics in bone cement  Poly(methylmethacrylate) (PMMA) bone cement and PMMA beads have long been investigated and used for local antibiotic delivery in orthopaedic surgeries [313-315]. The decline in antibacterial activity of fusidic acid (FA) released in vitro from PMMA bone cement over the course of 3 to 9 days was illustrated by the groups of Medcraft et a!. [316] and Hill et a!. [317]. Using in vitro assays, Neut et a!. showed that the combination FA and gentamicin loaded in PMMA bone cement was more effective against clinical bacteria isolates than the combination of gentamicin with clindamycin, or gentamicin alone [318, 319]. A prospective study by Coombs et al. to evaluate the prophylactic use of both FA and gentamicin-loaded PMMA bone cement used in joint replacement surgery found that no patients receiving antibiotic-loaded cement developed chronic infection [320]. The sodium salt of FA was incorporated into PMMA bone cement by Quinlan and Mehigan, and was released in rabbits for a period of 45  —  187 days but at relatively low concentrations (0.014  —  0.530 ig/ml) [321].  Although PMMA bone cement and beads have been widely used clinically for localized delivery of antibiotics, PMMA has several disadvantages. PMMA is nondegradable and must be surgically removed from the implantation site following drug release for possible bone regeneration. Furthermore, the major drawback associated with the use of PMMA is the very poor release characteristics of this matrix. Typically, there is a sub-optimal antibiotic elution profile since the vast majority of the loaded antibiotic is retained in the matrix and not released [322-325].  42  1.5.2  Antibiotic loaded microspheres  Polymeric microspheres are another delivery system that has been investigated for localized and  controlled  antibiotics  delivery.  Tetracycline  loaded  poly(hydroxybutyrate-co  hydroxyvalerate) (PHBV) micro spheres of various hydroxyvalerate content were prepared by the double emulsion method for the treatment of periodontal disease by Sendil et al. [326]. They found the concentration of the emulsifiers used influenced the encapsulation efficiency appreciably, whereas in vitro drug release rates were influenced by polymer composition [326]. Composite microspheres prepared from blending PHBV with either wollastonite (bioceramic) [327] or poly(lactic-co-glycolic acid) (PLGA) (50/50) [328] for the controlled release of gentamicin or vancomycin, respectively, have also been investigated for the treatment of bone infections. In both studies, the additional bioceramic or polymer in the antibiotic-loaded PHBV microspheres affected the drug release kinetics [327, 328].  The use of PLGA beads [329-332] and microspheres [333-337] to delivery a variety of antibiotics for the treatment of bone infections has been under active investigation. For instance, the group led by Liu et al. manufactured vancomyin PLGA (50/50) beads by compression and sintering, and achieved high local concentrations of the antibiotic for more than 55 days in the bony cavities of rabbits, with minimal systemic exposure [330, 331]. PLGA beads of various polymer composition, individually loaded with clindamycin, tobramycin, or vancomyin were also shown to release high concentration of all the antibiotics in vitro for 4 to 8 weeks by Mader et al. [332]. Jacob et al. encapsulated cefazolin and ampicillin in PLGA microspheres for the prevention and treatment of bone infection caused by S. aureus and found that in all cases, the rabbits treated with the antibiotic loaded PLGA  43  microspheres either did not develop any infection, or infection was eliminated as compared to control groups with systemic antibiotic delivery or no antibiotic delivery [333-335]. In vitro release kinetics and in vivo efficacy of teicoplanin-loaded PLGA microspheres in a bone infection model in rabbits has been investigated [337].  1.5.3  Mechanisms of drug release  Polymeric controlled release drug delivery systems [14, 221] can be generally divided into two fundamental types: reservoir and matrix systems [338]. In the reservoir system, a core of drug is surrounded by a rate-limiting polymer barrier, whereas in the matrix system, the drug is uniformly distributed throughout a polymer matrix [338]. Matrix type systems were used in this work and mechanisms of drug release are described only for these systems. Controlled drug release from matrix systems generally occurs by one of three mechanisms: (i) diffusion, (ii) chemical reaction, and (iii) solvent activation [338], with the first two mechanisms dominating and described below.  Drug release from matrix systems occurs via diffusion down the concentration gradient. The homogenously distributed drug can be either dissolved in the polymeric matrix, termed a monolithic solution system, or the drug can be dispersed as solid particles if the drug has limited solubility in the polymer, and is termed a monolithic dispersion system [339]. For monolithic solution systems, initial concentration gradient is absent within the polymer matrix prior to the onset of drug release. However, as drug begins to release from the surface of the matrix, a concentration gradient is established to provide the driving force for drug diffusion from the interior of the matrix towards the surface for subsequent release [339].  44  In monolithic dispersion systems, the amount of initial drug loading can be used to further classify the systems. At low drug loadings (< 5% by volume), systems are referred to as simple monolithic dispersion [339]. Drug release involves the initial dissolution of the drug into the polymer followed by diffusion to the surface of the device. At slightly higher drug loadings (5  —  10% by volume), monolithic dispersion systems are referred to as complex  monolithic dispersion, since release mechanism becomes more complex [339]. Fluid-filled cavities on the surfaces of the device, formed from the loss of drug near the surfaces enhance drug release. Even though the cavities are not connected to form continuous pathways to the surface, they increase the overall permeability of the drug in the polymer matrix. When drug loadings are greater than about 20% by volume, the cavities become interconnected to form continuous fluid-filled channels to the surfaces of the device through which the majority of the drug is released. These devices are known as monolithic matrix systems, and the solubility and diffusivity of the drug in the fluid-filling channels determine its rate of release [339].  In the case of chemical controlled polymeric drug release systems, the rate of drug release is dictated by the rate of polymer degradation. There are two mechanisms by which polymers can degrade, bulk (homogenous) degradation and surface (heterogeneous) degradation [340, 341], which are schematically illustrated in Figure 1.6.  45  A  B  Jr  Drug dissolved or dispersed in the polymer Jr  .  Jr .  .  ••  :  •  Figure 1.6: Schematic representation of the mechanisms of polymer degradation. (A) Bulk degradation and (B) surface degradation.  For bulk degradation (Figure 1 .6A), the influx of water into the polymer matrix is faster than the degradation of the individual chains into water-soluble fragments. Thus, degradation occurs throughout the entire polymer matrix and erosion kinetics of the degraded fragments are non-linear, and usually characterized by a discontinuity [341]. The size of the delivery devices generally remains constant for a considerable portion of time, until a critical point,  46  where bulk degradation leads to the disintegration of the whole matrix into smaller pieces [340]. Examples of degradable polymers that undergo bulk degradation include polyesters such as PLGA and PHBV.  In the case of surface degradation (Figure 1 .6B), polymer degrades much faster than the penetration of water into the matrix. Thus, degradation and erosion are limited to the surface of the polymer only. The structural integrity and the original geometric shape of the device are maintained, but its size is steadily reduced as eroding polymers are lost on the surface during the degradation process [342]. Typically, the mass loss kinetics are linear and predictable, therefore the rate of drug release can be directly related to the rate of polymer erosion [340]. Poly(anhydrides) [342, 343] and poly(ortho esters) [344] are degradable polymers that undergo surface degradation.  1.5.4  Factors influencing drug release  A number of factors can influence the kinetics of drug release from a polymeric matrix. For example, the geometric shape and size of the device that increases surface area will result in a higher drug release rate [339]. Moreover, the initial drug loading in the device also affects the rate of drug release. As drug loading is increased, not only is a higher concentration gradient established, but the formation of cavities and channels for diffusion also increases, to increase drug release [339].  The degree of crystallinity of the polymer can affect the diffusion of the drug as well as the degradation and erosion of the polymer matrix. Crystalline regions within the polymer are  47  less accessible to water and drug molecules than amorphous regions, where the free volume is higher [345]. Therefore, lowering the crystallinity of a polymer will increase the rate of drug release due to the increase in both drug diffusion and polymer degradation [346, 347]. The molecular weight of a polymer also affects drug diffusion and polymer degradation. As the molecular weight of a polymer decreases, the free volume of the polymer increases due to a greater number of polymer chain ends [348]. As a result, permeability of the polymer matrix increases leading to higher drug diffusion and polymer degradation to increase drug release rate. In copolymers such PLGA and PHBV, polymer composition affects the hydrophobicity of polymer matrix to influence drug release rates. For example, PLGA (50/50) is more hydrophilic than PLGA (85/15), thus water penetration into the matrix is greater for the 50/50 PLGA, leading to more rapid polymer degradation that increases the rate of drug release [231].  Other critical factors affecting drug release rates include the fundamental properties of the drug such as diffusion coefficient, solubility, molecular weight as well as its partition coefficient, and have been reviewed [349, 350].  48  1.6 Development of fusidic acid loaded microspheres for bone infections The increase in antimicrobial resistance to clinically used antibiotics in recent years has necessitated a search for alternatives and one promising approach has been the re-evaluation of older generation antibiotics. Due to their limited use, many antibiotics such as fusidic acid have remained active against a large number of currently prevalent bacterial isolates, and are re-emerging as valuable alternatives for the treatment of difficult-to-treat infections [351]. Furthermore, even though antibiotic-loaded PMMA bone cement has been the primary method of local antibiotic therapy in orthopaedic surgeries over the past three decades [313315], the fundamental problems associated with PMMA described in Section 1.5.1, warranted the investigation of alternative localized delivery systems such as polymeric microspheres.  1.6.1  Fusidic acid  The appearance of methicillin-resistant Staphylococcus aureus (MRSA) as causal pathogens in orthopaedic infections has challenged clinicians and microbiologists to find new antibiotics or combinations that are effective in deep infection, but are also tolerable with prolonged treatments [352, 353]. Even though introduced in the 1960s, like many older drugs, fusidic acid (FA) has never been intensively studied or widely applied to orthopaedic infections. Interestingly, FA may be considered a valuable antibiotic for the management of staphylococcal and other Gram-positive infections [354, 355]. FA has retained activity against MRSA and other multi-drug resistant Gram-positive bacteria [355].  49  1.6.1.1  Chemistry  FA, shown in Figure 1.7, is a tetracyclic triterpenoid, derived from the fermentation broth of the fungus Fusidium coccineum and is chemically related to the antibiotics helvoic acid and cephalosporin P 1 [356]. FA and the sodium salt of FA, termed Fucidin® were first introduced into clinical practice in the 1960’s [53]. FA is a white crystalline powder with a molecular weight of 516.7 g/mol (anhydrous). It is a weak acid with a pKa of 5.35, and is mostly ionized in plasma and tissue at the physiological pH of 7.4. FA is almost insoluble in water and has a high octanol/water partition coefficient [54], while the sodium salt of FA is readily water soluble [357]. Due to its structural similarity to bile salts, FA forms micelles at concentrations in the range of 1.44  —  4.56 mM [358]. Above 11 mM FA also tends to self-  aggregate in aqueous solution [357].  2H Me OAc  Me HO H Me  Figure 1.7: Chemical structure of fusidic acid (FA)  50  1.6.1.2 Pharmacology FA is an antimicrobial agent that acts as an inhibitor of protein synthesis in microorganisms [359]. It interferes with the translocation of the elongation factor G (EF-G) from the ribosome by stabilizing the ribosome-guanosine diphophate-EF-G complex. This prevents binding of aminoacyl tRNA to the ribosome and thereafter stops transfer of additional amino acids to the growing polypeptide [360]. FA is mainly bacteriostatic, but at higher concentrations, may be bactericidal [361]. FA is primarily active against Gram-positive organisms, both aerobes and anaerobes, and is most active against Staphylococcus aureus (S. aureus), S. epidermidis, and coagulase-negative staphylococci including strains that are methicillin-resistant (i.e. MRSA) that commonly cause prosthetic joint infections [45, 57, 354-356, 362-364]. FA is also effective against oxacillin-resistant and quinolone-resistant staphylococci [365].  The activity of FA against infections is dependent upon the drug reaching concentrations in the tissues above the minimum inhibitory concentration (MIC) for the particular organism. Concentrations of FA between 0.03 to 0.12 mg/L inhibit virtually all strains of Staphylococcus [54, 57, 361, 364], with MICs for MRSA being a little higher, ranging from 0.03 to 0.8 mg/L. MICs are markedly increased in the presence of serum due to high protein binding of FA [366], and antimicrobial activity is also reduced in alkaline media [54]. More detailed analyses of the antimicrobial activity of FA have been provided by Verbist [366], and Mandell [54].  51  The side effects of FA include hyperbilirubinemia, thrombophiebitis, venospasm, and reversible jaundice for intravenous dosing [54]. For oral dosing, side effects including epigastric pain, anorexia, vomiting and diarrhea, hyperbilirubinemia and reversible jaundice have been reported [55]. A drug-induced reversible immune-mediated thrombocytopenia has also been described in which FA-dependent platelet-reactive antibodies bind platelets only in the presence of FA [367]. Furthermore, FA has been shown to affect bile acid uptake and secretion of biliary lipids because it competes with the same transport system as bile acid [368]. FA undergoes extensive metabolism in the liver and is primarily excreted in the bile as metabolites with weak or no biological activity. The major metabolites are glucuronic acid conjugate, dicarboxylic acid metabolite, and hydroxyl metabolite. Approximately 2% of the drug can be recovered unchanged in the feces with less than 1% of the administered dose excreted in the urine  [54]. A comprehensive review of the pharmacology and  pharmacokinetics of FA is provided by Turnidge [57].  1.6.2  Biodegradable polymers selected for fusidic acid loaded microspheres  Various biodegradable and bioresorbable carriers of antibiotics for the treatment and prevention of bone infections have been studied. Natural biopolymers such as poly(hydroxybutyrate-co-hydroxyvalerate) (PHBV) are ideal biomaterials [369-371] for antibiotic delivery [326, 327, 372-375] due to their biocompatibility, minimal toxicity and lack of inflammatory response, as well as their complete biodegradation into biocompatible by-products. In addition, the use of synthetic biodegradable and bioresorbable polymers such as poly(lactic-co-glycolic acid) (PLGA) [376, 377] for antibiotic delivery has also been widely studied [329, 333-337, 378].  52  1.6.2.1 Poly(hydroxybutyrate-co-hydroxyvalerate) Poly(hydroxybutyrate-co-hydroxyvalerate) (PHBV), shown in Figure 1.8, is a natural aliphatic polyester that can be produced from a variety of renewable resources [379]. PHBV is a member of the polyhydroxyalkanoate (PHA) family of biopolymers. It is biosynthesized by various microorganisms  such as Rhodospirillum rubdum, Raistonia eutropha,  Pseudomonas oleovorans, among others, under conditions of nutrient limitation as an intracellular carbon and energy storage compound [380, 381]. Poly(hydroxybutyrate) (PHB) homopolymer and PHBV copolymer with different composition of hydroxybutyrate and hydroxyvalerate content are biosynthesized using different mixtures and concentrations of the carbon sources such as glucose, sucrose and propionic acid [382]. PHBV can be biosynthetically produced in vitro via PHA-polymerase catalyzed polymerization or in vivo with batch, fed-batch, and continuous (chemo stat) cultures [383].  0  V  Hydroxybutyrate  HydroxyvaLerate  Figure 1.8: Chemical structure of poly(hydroxybutyrate-co-hydroxyvalerate) (PHBV) copolymer  53  The physical and thermal properties of PHBV copolymers can be regulated by varying their molecular weight and copolymer compositions [384]. PHBV copolymers are less crystalline, more flexible and more readily processable than PHB homopolymers [278]. Nevertheless, PHBV displays high degrees of crystallinity (50  —  70%) throughout a wide range of  copolymer compositions [385] and behaves as a thermoplastic with melting temperatures, Tm ranging from 50 to 180 °C [385, 386]. The glass transition temperature, Tg decreases from 4 to -18 °C as the hydroxyvalerate fraction increases from 0 to 100 mol% in the copolymer [384]. PHBV possesses piezoelectric characteristics, where it has the ability to generate an electric potential in response to applied mechanical stress and vice versa [387-390]. The piezoelectricity of PHBV can lead to interesting applications in orthopaedic devices since bone also demonstrates piezoelectric characteristics and electrical stimulation is thought to promote bone healing and repair [385].  The biodegradability of PHBV is reported to be much lower than that of PLGA, but not lower than poly(L-lactic acid) [391]. The degradation rate can be controlled by varying the copolymer composition, and increases with increasing hydroxyvalerate content of the PHBV copolymers [392-394]. PHBV degrades mainly by hydrolysis through surface erosion but can also be degraded by PHA depolymerases secreted by many types of microorganism in the biological environment [385, 395, 396]. In vivo, PHBV degrades to 3-hydroxybutyric acid, which is a normal constituent of human blood and 3-hydroxyvaleric acid [381]. The low toxicity of PHBV copolymers may be at least partly due to their biocompatible degradation  54  by-products. The synthesis, physical properties and degradation of PHBV in biomedical applications has been recently reviewed [384, 385, 396, 397]. 1.6.2.2 Poly(lactic-co-glycolic acid) Poly(lactic-co-glycolic acid) (PLGA) belongs to the poly(cL—hydroxy acid) polymer family and is by far the most extensively studied synthetic biomaterial used to develop controlled release drug delivery systems [377, 378, 398-403]. PLGA, shown in Figure 1.9, is an aliphatic polyester and a copolymer of lactic acid and glycolic acid. Ring opening polymerization of the cyclic dimers, lactide and glycolide is used to synthesize high molecular weight (> 10,000 g/mol) PLGA [404].  0  0 Lactic Acid  Glycolic Acid  Figure 1.9: Chemical structure of poly(lactic-co-glycolic acid) (PLGA) copolymer  As two enantiomeric isomers of lactide exist (D- and L-), the physical and thermal properties of PLGA are dependent on the particular isomer of lactide and the molar ratio of lactic acid to glycolic acid repeat units in the copolymer. Both poly(L-lactic acid) (PLLA) and  55  poly(glycolic acid) (PGA) homopolymers are semi-crystalline, and PLGA copolymers synthesized with L-lactide exhibit some degree of crystallinity when either lactic acid or glycolic acid is present over 70 mol% [405]. PLGA synthesized from D,L-lactide is amorphous in nature [377, 406]. The Tg of PLGA is above 37 °C and decreases correspondingly with the decrease in lactic acid content and molecular weight of the copolymer [398, 407]. PLGA with molar ratio of lactic acid to glycolic acid of 85 to 15% (85/15) possesses a Tg in the range of 50 range of 45  —  —  55 °C, while the Tg of PLGA (50/50) is in the  50 °C [404].  PLGA degrades by bulk hydrolysis of the ester linkages in the backbone of the copolymer with the degradation occurring at a uniform rate throughout the PLGA matrix [407]. During the degradation process, the number of carboxylic acid end groups increases as the individual polymer chains are cleaved, which further catalyzes the degradation process (autocatalysis) [402, 407]. The molar ratio of lactic acid to glycolic acid in the copolymer, polymer molecular weight, the degree of crystallinity, and the Tg of the PLGA all influence the rate of polymer degradation [407, 408]. Degradation is more rapid for lower crystalline and lower molecular weight polymers as well as PLGA with a lower Tg [404, 407]. However, the relationship between the copolymer composition and the degradation rate is not linear as demonstrated by the faster degradation rate of PLGA (50/50) than either PLA or PGA homopolymer [404]. PLGA degrades into its constituent monomers: lactic acid and glycolic acid. Lactic acid is eliminated from the body as carbon dioxide and water through the Kreb’s cycle. Glycolic acid is either excreted unchanged in the kidney or it also enters the Kreb’s  56  cycle and is eventually eliminated as carbon dioxide and water [409]. The biodegradability of PLGA has been further reviewed in bone tissue applications [410, 4111.  1.6.3  Formulation approaches for localized delivery of fusidic acid  FA has been used primarily for the treatment of staphylococcal infections [45, 320, 354, 412, 413]. Detailed reviews of virtually every study and use of FA systemically for the treatment of orthopaedic infections for the last four decades have been provided by Atkins et al. [354], Coombs [320] and Anderson [53]. Even though localized and controlled delivery of FA has been suggested to be suitable for treating and preventing bone infection in orthopaedic medicine, there are very few such studies. Bouillet et a!. described the successful treatment of 16 out of 19 patients with S. aureus bone infection in various locations by curettage of the lesions and packing the bone cavities with biodegradable plaster of Paris (calcium sulphate hemihydrate) beads loaded with FA and amoxicillin [414]. Using the W/O/W double emulsion technique, Cevher et a!. loaded the sodium salt of FA into PLGA microspheres and investigated both the in vitro drug release and in vivo efficacy against MRSA osteomyelitis [415]. Sodium FA-loaded PLGA microspheres were implanted in the proximal tibia of rats with methicillin-resistant MRSA osteomyelitis and showed significantly lower colony forming units of MRSA than controls [415].  57  1.7 Research rationale, goals and objectives  1.7.1  Research rationale  Orthopaedic surgeries are always required when damaged bone tissues are ‘critical’ in size and unable to repair or regenerate themselves, such as those found in revision total hip replacements. Thus, the natural bone regeneration process must be augmented with the application of exogenous therapeutic agents such as growth factors, hormones and/or cells. However, there exists the risk of infections that must be minimized and managed with the application of antibiotics. Our laboratory has developed strong collaborations with other research groups interested in the development and characterization of delivery systems for both bone regeneration and bone infection in revision total hip replacements, and other orthopaedic applications.  MSCs are ideal cells for bone regeneration applications [416] since they are relatively easy to harvest, and can be isolated from adult bone marrow and various other tissues. Additionally, they have high proliferative capacity, and can be easily expanded in vitro whilst still retaining their differentiation potential to generate multiple cell types [18, 20], including osteoblasts to augment bone regeneration [417]. As a result of the successful use of PTH to enhance bone regeneration [37], and studies demonstrating improved bone regeneration when growth factors were delivered along with MSCs [33-35], we hypothesized that PTH may increase the relative osteogenic progenitor content of MSC cultures to better promote bone tissue regeneration. However, as a first step, it was important to understand the effects of different regimens of PTH treatment on MSCs differentiation, proliferation and  58  clonogenicity in order to fully maximize the potential of using the combination of MSCs with PTH for bone regeneration.  FA is a suitable antibiotic for the management and prevention of orthopaedic infections because of its effectiveness against a wide spectrum of pathogens that commonly cause prosthetic joint infections including MRSA and other multi-drug resistant Gram-positive bacteria [54]. However, systemic delivery of FA can lead to many serious toxicities, thus necessitating its localized delivery. By direct application of antibiotics to the site of infection or potential infection, it is possible to deliver a lower dose but still achieve higher tissue levels and for a longer period of time, as well as decreasing the potential for systemic side effects [14].  The ideal biomaterials for PTH, MSCs, and FA delivery should be biocompatible (non-toxic, non-inflammatory, non-immunogenic), biodegradable and bioresorbable. In addition, they should be fabricated reproducibly and be sterilizable and cost effective. Since the delivery of MSCs attempts to replace the missing bone tissue, natural biomaterials such as gelatin and alginate used to fabricate porous scaffolds meet many the above criteria in addition to possessing physicochemical properties that mimic the natural ECM [278, 279]. In order to achieve controlled and predictable release of PTH and FA over prolonged periods of time, encapsulation of these agents in biodegradable polymeric microspheres was felt to be an appropriate choice of delivery technology. The well established biocompatibility and biodegradability of PLGA and its extensive use in orthopaedic devices and clinically approved drug delivery systems has led us to select PLGA as a biomaterial for PTH and FA  59  delivery. PHBV also demonstrates biocompatibility, biodegradability and a piezoelectric characteristic that is thought to promote bone healing and repair. Hence PHBV was also selected as a potential suitable biomaterial.  1.7.2  Research goals  The overall goals of this project were to engineer, develop, and characterize biodegradable and bioresorbable polymeric microsphere and porous scaffold delivery systems for PTH and MSCs, respectively, for enhancing the innate regenerative capacity of bone, and to investigate the effects of continuous and pulsatile PTH treatments on MSC to better understand its regulatory actions on MSC differentiation, proliferation and clonogenicity. In addition, the development and characterization of biodegradable and bioresorbable polymeric microsphere delivery systems for the antibiotic, FA for the potential localized application in bone infection were also undertaken in this project.  1.7.3  Research objectives  (1) To detennine the effects of continuous and pulsatile PTH treatments on rat MSCs differentiation and proliferation.  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The ease of isolation and the ability of in vitro expanded marrow stromal cells (MSCs, also known as mesenchymal stem cells [2]) to generate multiple cell types, including osteoblasts, chondrocytes, and adipocytes are well established and make these cells an attractive source for tissue engineering and regenerative medicine (see reviews [3-5]). Whether MSCs proliferate as multipotent stem cells capable of differentiating into multiple mesenchymal lineages, or as committed progenitors of a particular lineage, is likely regulated by a plethora of growth factors, cytokines and hormones [6].  Parathyroid hormone (PTH) exerts both anabolic and catabolic effects on bone depending on the nature and duration of exposure. Pulsatile (intermittent) delivery of PTH has been shown to lead to increased bone mass, strength, mineral density, and improved bone microarchitecture and fracture healing [7], whereas continuous administration of PTH results in bone resorption and hypercalcemia [8]. PTH is the only clinically approved anabolic treatment for osteoporosis where intermittent subcutaneous injections of PTH have been 1  A version of this chapter has been published. Yang C et al., (2009) Effects of continuous and pulsatile PTH treatments on rat bone marrow stromal cells. Biochem Biophys Res Commun 380(4):791—796  94  demonstrated to reduce vertebral and non-vertebral fracture risk in postmenopausal women, elderly men, and women with glucocorticoid-induced osteoporosis [9].  Based on the anabolic responses obtained in vivo with pulsatile PTH administration [10-131, it is tempting to hypothesize that a similar pulsatile treatment may be beneficial in increasing the relative osteogenic progenitor content of MSC cultures and might enhance bone regeneration. However, the effects of different regimens of PTH treatment on osteogenic gene expression in MSC cultures other than in an adipogenic medium [14] have not been investigated. Additionally, the osteogenic potential of MSC cultures is known to be affected by glucocorticoids such as dexamethasone (DEX). For example, studies have shown that treatment with DEX greatly increases the differentiation of rat MSC into osteoblasts triggered by bone morphogenetic proteins (BMP5) [15]. However, despite the great interest in PTH as a therapeutic and the fact that in vitro studies frequently include DEX, the combined effects of PTH and DEX on the osteogenic potential of MSC cultures have not been examined.  In preliminary work described in Appendix A, we developed PTH-loaded polymeric microspheres in an attempt to deliver PTH in a controlled manner. PTH was encapsulated in poly(lactic-co-glycolic  acid)  (PLGA)  and  poly(hydroxybutyrate-co-hydroxyvalerate)  (PHBV) microspheres using the double emulsion solvent evaporation method. The release data showed consistent biphasic release profiles with a pronounced burst phase over 24 hours followed by a prolonged period of slower, controlled release (Figures A-4 and A-5). It was evident that these formulations were not suitable for providing the precise and reproducible  95  delivery of PTH to MSCs in either continuous or pulsatile regimens. Therefore, the studies described in this chapter used PTH (1-34) peptide solutions as follows. For continuous administration, MSCs were exposed to PTH throughout the culture experiments with addition of PTH every 24 h. For pulsatile administration, MSCs were exposed to PTH for the first 6 h only of each 48 h incubation cycle.  Several groups have explored the development of localized and controlled release systems to deliver PTH in either a continuous [16] or pulsatile [17-19] manner to osteoblastic cell lines [16-19] but not primary cultures like MSCs. And although these research groups have shown that the continuous released PTH was biologically active [16, 19] and PTI-I delivered in a pulsatile fashion elicited greater osteoblastic activity [17, 18], none of these studies actually delivered PTH directly to the osteoblastic cell lines they were investigating [16-19]. Rather, in all the studies, PTH eluted from in vitro release experiments was collected and then used in separate experiments to expose the cells to either continuous [16] or pulsatile [17-19] PTH regimens, in order to assess the bioactivity of the released PTH.  The objectives of this work were to investigate the effects of continuous and pulsatile PTH in the presence and absence of DEX on the osteoprogenitor content and expansion of rat bone marrow-derived MSCs in vitro. Continuous and pulsatile PTH (10 nM) with and without DEX (10 nM) were used. Osteoprogenitor content was quantified by assessing alkaline phosphatase (ALP) enzyme activity, ALP histochemical staining, as well as PTH receptor 1 (PTHR1), ALP, and osteocalcin (OC) mRNA expression using qRT-PCR. The effects of PTH on MSC culture expansion was assessed by total cell count and cell proliferation  96  kinetics. The ability of PTH treatments to promote the maintenance of clonogenic progenitors under osteogenic conditions was investigated with CFU-F assays.  97  2.2  2.2.1  Materials and methods  Chemicals  Recombinant human parathyroid hormone (PTH) peptide 1-34 purchased from Bachem (Bachem California Inc., Torrance, CA) was dissolved in 4 mM HC1 containing 0.1% bovine serum albumin (BSA) and stored at -30°C before dilution to the appropriate concentration immediately prior to use. Mesencult® Basal medium (cat # 05401), fetal bovine serum (FBS, cat # 06471), osteogenic FBS (cat # 06473), 13-glycerophosphate, ascorbic acid, dexamethasone (DEX) were purchased from StemCell Technologies (Vancouver, BC Canada). Insulin, BSA, paraformaldehyde (PFA), lOx trypsinlEDTA solution, p-nitrophenyl phosphate (pNPP), naphthol AS-MX phosphate, diazonium salt (fast red violet LB salt), Alizarin  Red  sodium  mono  sulfonate,  MTT  (3 -(4,5 -Dimethylthiazol-2-yl)-2,5-  diphenyltetrazolium bromide) and DMSO were all purchased from Sigma-Aldrich (Mississauga, ON, Canada) at tissue culture or spectrophotometric grade. Reversetranscription reagents using the Superscript III RT system were acquired from Invitrogen TM (Burlington, ON, Canada).  2.2.2  Isolation and cell culture of rat marrow stromal cells (MSCs)  MSCs were isolated from 6-week Sprague Dawley rats expressing transgenic green fluorescent protein (GFP) (NBRP, Japan). All procedures were approved by the University of British Columbia, Animal Care Committee. Briefly, femora and tibiae harvested from euthanized rats were crushed to release total bone marrow cells. MSCs were subsequently isolated by virtue of their adherence to tissue culture plastic cultured in Mesencult® medium 98  supplemented with 15% FBS and 100 tg/mL penicillinlstreptomycin at 37 °C, and 5% C0 , 2 expanded to passage two and cryopreserved until they were used. Cells used for all PTH treatment experiments were further expanded to passage 3  —  5. MSCs were cultured in  Mesencult® Basal medium supplemented with 15% FBS, 5 mM f3-glycerophosphate, 50 .ig/ml ascorbic acid, and penicillin!streptomycin at 37 °C, and 5% C0 , unless otherwise 2 noted.  2.2.3  Flow cytometry  Fluorescence Activated Cell Sorting (FACS) analysis was used to determine the extent of hematopoietic contamination of the isolated and passaged MSCs. Briefly, trypsinized MSCs were centrifuged then resuspended in primary mouse anti-rat CD45 biotin antibody (BD Pharmigen) and incubated at room temperature for 30 mi  After incubation, cells were  washed, incubated with streptavidin APC-cy7 conjugated secondary antibody (CALTAG) for 30 mm at room temperature, and then washed with PBS. Cells were immediately filtered through a 0.22 iim mesh to remove any cell aggregates and FACS analyzed. Samples were run on a BD FACSCalibur SE flow cytometry system and samples gated for GFP and CD45 staining.  2.2.4  Rat MSC differentiation assays  The differentiation potential of the MSC preparations used for the PTH treatment experiments was determined along three lineages: osteogenic (bone), adipogenic (fat) and chondrogenic (cartilage) [2]. Differentiation was induced for 3 weeks, and the conditions  99  used were as follows. (1) For osteogenic differentiation, cells were plated at a density of 15 x 1  2 and cultured in Mesencult® medium supplemented with 15% osteogenic FBS cells/cm  (StemCell Technologies), 5 mM 3-g1ycerophosphate, 50 jig/ml ascorbic acid, and 10 nM of dexamethasone; (2) For adipogenic differentiation, cells were plated at a density of 60 x 1 0 2 and cultured in Mesencult® medium supplemented with 15% FBS, 100 nM cells/cm dexamethasone, and 6 ng/ml insulin; (3) For chondrogenic differentiation, 5 x 1  cells were  spun down into a pellet and cultured in Mesencult® medium supplemented with 10% FBS, 50 jig/ml ascorbic acid, 10 nM dexamethasone, and 10 ng/ml TGF-13 . Media for all 1 differentiation assays contained 100 j.ig/mL penicillinlstreptomycin and were changed every 48 hr.  2.2.5  PTH treatment of rat MSCs  MSCs were thawed and seeded at a density of 5 x 1  2 in 12-well tissue culture cells/cm  plates and cultured for 24 h in Mesencult® medium supplemented with 15% FBS without  13-  glycerophosphate, ascorbic acid, and PTH (1-34). After 24 h of culture, medium was changed to fresh medium supplemented with f3-glycerophosphate and ascorbic acid (i.e. osteogenic culture conditions). From this time point (Day 0) onward, MSCs were first treated with 10 nM PTH and 10 nM DEX for up to 14 days. For all experiments, MSCs were divided into three groups according to the mode of PTH treatment: control (no PTH treatment) group, pulsatile PTH group (PTH-P), and continuous PTH group (PTH-C). For the pulsatile treatment group, the cells were exposed to PTH for the first 6 h of each 48-h incubation cycle, and then cultured in the absence of PTH for the remainder of the cycle [201. In the continuous treatment group, the cells were exposed to PTH from Day 0 to the end of the 100  culture experiment, with addition of PTH every 24 h to maintain constant concentration because it has been shown in the presence of cells, PTH concentration decreases  50% after  24 h [20]. For the entire duration of the experiment, the culture media for all three groups were changed every 48 h. In separate experiments to investigate functional interactions between DEX and the effects of continuous and pulsatile PTH treatments on MSC differentiation and proliferation, MSCs were cultured with PTH (10 nM) in the absence of DEX for seven 48-h cycles (i.e. 14 days).  2.2.6  Alkaline phosphatase (ALP) activity  ALP enzyme activity was determined from cell extracts. Briefly, after 3 weeks of culture, the MSCs were washed twice with PBS and then lysed with a probe sonicator (Ultrasonic Cell Disruptor, Misonix Incorporated) for 20 sec in 0.5 mL of 0.1 M Tris buffer (pH 7.2) containing 0.1% Triton X-100 at 4 °C. Sonicated cell lysates were then centrifuged for 10 minutes at 1500 g and 4 °C, and the supematants were used for the assays. ALP activity was determined spectrophotometrically from conversion of p-nitrophenyl phosphate (pNPP, Sigma, St. Louis, MO) to p-nitrophenol by adding 50 iL of cell extract to 150 pL of buffersubstrate solution at 37 °C. The buffer-substrate solution contained 50 mM pNPP, 2 mM 2 and 100 mM glycine buffer at pH 10.4. The colour change of pNPP to p-nitrophenol MgCl showed a linear relationship with time and was monitored spectrophotometrically at 405 nm (SpectraMax 190, Molecular Devices) every minute for 30 mm and quantified by comparison with a standard curve. ALP activity was normalized to the total protein (cell) content of the sample, which was measured spectrophotometrically using a Micro BCA protein assay kit (Pierce Chemical Co., Rockford, IL).  101  2.2.7  Histochemistry staining  To visualize ALP activity in MSC cultures, the cells in each tissue culture well were fixed with 2% PFA in PBS for 10  —  30 mm and incubated with substrate solution for 30  —  60 mm  at 37 °C, and then rinsed three times with PBS to remove excess staining. The substrate solution used for staining contained 0.1 mg/mi naphthol AS-MX phosphate and 0.6 mg/ml diazonium salt (fast red violet LB salt) in Tris buffer, pH 8.74 [201. ALP activity was then imaged with a light microscopy.  In MSC differentiation assays, bone nodules (mineralized bone matrix) were visualized after fixation with 2% PFA in PBS for 10—30 mm and stained with freshly dissolved Alizarin red S solution (20 mg/mL, pH 4.2). The staining solution was removed after 1 mm and each well was washed three times with PBS to remove excess staining. Adipogenesis was assessed by staining intracellular fat-containing vacuoles with oil Red 0. After 4% PFA fixation for 30  —  60 mm at 37 °C and three washes with PBS, adipogenic differentiated MSC were stained with 1 part 0.55% of oil Red 0 in isopropanol and 0.75 part water at 37 °C for 3h, washed five times with PBS and then imaged. Chondrogenic differentiation was determined from cryo-sections of pellet cultures stained with 10 mg/mL alcian blue in 3% acetic acid solution (pH 2.5) at room temperature overnight. Excess staining was removed with 3% acetic acid, followed by distilled water washes.  102  2.2.8  Quantitative real time-polymerase chain reaction (qRT-PCR)  To quantify gene expressions after PTH treatments, mRNA from cultured MSCs was extracted, reverse-transcribed and subjected to quantitative real time PCR. Briefly, cultured and PTH treated cells were trypsinized from plates using PBS containing 0.5 mg/mi trypsin and 0.2 mg/mi EDTA, and mRNA was extracted using QIAGEN RNeasy® Mini Kit according to manufacturer’s instructions. Subsequently,  1 ig of total RNA was reverse-  transcribed using the Superscript III RT system to generate complementary DNA (cDNA). The TaqMan® probes-based detection system (Applied Biosystems) was used and qRT-PCR reactions were performed in a Roche LightCycler® 480 real-time PCR system (Laval, Quebec, Canada). TaqMan® primers assay for glyceraldehyde 3-phosphate dehydrogenase (GAPDH), PTH receptor 1 (PTHR1), alkaline phosphatase (ALP), and osteocalcin (OC) genes were from Applied Biosystems and their manufacturer’s assay numbers were as follows: GAPDH, Hs02758991_gl (cross-reative for rat); PTHR1, Rn00571596_ml, ALP, Rn0056493 1_mi, and OC, Rn00566386_gl.  2.2.9  Cell count and MTT cell proliferation assay  Cell numbers were assessed by counting trypsinized cells using a hemocytometer. Trypan blue staining was used to exclude non-viable cells. Cell proliferation kinetics were assessed using a MTT (3 -(4,5 -Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) colorimetric assay. Briefly, cultured and PTH treated cells were washed with PBS, then 50 il of MTT solution (5 mg/mI dissolved in PBS) were added to each well and the samples were incubated at 37 °C for 4 h. Subsequently, MTT solution was removed and the cells were washed with PBS. The purple formazan salt product resulting from the reduction of the yellow MTT was 103  solublized with the addition of 100 tl of DMSO and quantified spectrophotometrically at 570 nm (SpectraMax 190, Molecular Devices).  2.2.10 Colony-forming unit fibroblast (CFU-F) assay The presence of CFU-F in MSC cultures after PTH treatments was assessed by bulk assay as follows. MSCs were seeded at a density of 5 x  2 in 12-well tissue culture plates cells/cm  and cultured for 24 h in expansion medium (Mesencult® Basal medium supplemented with 15% FBS and antibiotics only). After 24 h of culture, medium was either changed to fresh expansion medium or to osteogenic medium (expansion medium supplemented with f3glycerophosphate and ascorbic acid). MSC were divided into three groups according to the mode of PTH treatment: control (no PTH treatment) group, continuous Pill group (PTH-C), and pulsatile PTH group (PTH-P), as above, and treated for three to four 48-h cycles (i.e. 6  —  8 days). All groups were treated with 100 nM PTH. Following these treatments, cells were trypsinized and re-plated at low density (100 2 cells/cm in 60 mm diameter dishes and ) cultured in expansion medium for 6 to 8 days, with medium change after 3 or 4 days. Subsequently, cells were washed with PBS, fixed with methanol for 5  —  10 mm, and stained  with Giemsa staining solution. Colonies containing> 20 cells per foci were then counted under a microscope. The frequency of CFU-F was calculated as following:  Frequency of CFU  —  F  =  #ofCFU-F # of cells re plated  Equation (1.1)  —  For limiting dilution experiment, MSCs were seeded in 96-well culture plates at different cell densities in osteogenic medium supplemented with 10 nM DEX, and the treatment groups 104  were subjected to either continuous or pulsatile PTH (10 nM) for 8 days as above. The frequency of CFU-F was determined at the lowest cell density where CFU-Fs were present.  2.2.11 Data analysis For statistical analysis, a one-way ANOVA followed by a Tukey-Kramer’s post-hoc test or two-way (factorial) ANOVA followed by a Bonferroni’s post-hoc test in GraphPad Prism® version 5.00 for Windows (GraphPad Software, San Diego, CA, USA) were used to determine level of significance. For all experiments, results are expressed as mean ± SD (n 3  —  =  5), and data presented are representative of the results obtained in three independent  experiments.  105  2.3 Results  2.3.1  Rat MSC characterization  MSC cultures were derived from rat bone marrow and used at early passages. Under such conditions, these cultures often still contain a variable number of hematopoietic cells. To assess the extent of hematopoietic contamination in our experiments, we quantified the frequency of cells expressing the pan-hematopoietic marker CD45 within MSC preparations using flow cytometry. We found that following 3 to 5 passages, all MSC cultures used in this study contained less than 4% CD45 hematopoietic cells as demonstrated by a representative FACS analysis gated for both GFP and CD45 shown in the upper right quadrant of Figure 2.1. A defining characteristic of marrow stromal cultures is their ability to differentiate along multiple mesenchymal lineages when exposed to the appropriate inducing signals. To confirm that MSC preparations used in this study satisfy this criterion, we exposed them to osteogenic, adipogenic and chondrogenic conditions for 3 weeks in vitro. Isolated MSCs were found capable of differentiating along the osteogenic, adipogenic and chondrogenic lineages to produce bone nodules, fat vacuoles, and cartilage, respectively after 3 weeks of in vitro differentiation induction (Figure 2.2).  106  1 0’  1  +  LC) cj  o  C-) 101  10 100  101  102  1  GFP Figure 2.1: Representative FACS dot plot illustrating the population distribution of rat MSCs after 3 5 passages. Hematopoietic cell population (<4%) gated for both GFP and CD45 is shown in the upper right quadrant. —  107  A  MSC B C  QO  C b’  Cartilage Figure 2.2: Multi-potentiality of rat MSCs. Histochemical staining results of the differentiation of (A) rat MSCs (GFP) along three lineages after 3 weeks. (B) Osteogenic (mineralized bone nodule, arrowhead), (C) adipogenic (intracellular fat-containing vacuoles, arrows), and (D) chondrogenic (cartilage).  2.3.2  Effects of PTH on rat MSC differentiation and proliferation in the presence of DEX  To investigate the effects of continuous and pulsatile PTH on the differentiation of MSCs along the osteogenic lineage in the presence of DEX, early passage cultures were exposed to PTH (10 nM) for 14 days, as described in the Section 2.2.5, and the emergence of alkaline phosphatase positive cells was quantified using enzymatic activity assays and histochemical staining (Figure 2.3). After 4 days of PTH treatment, a decrease in ALP activity was observed in the group treated with pulsatile PTH compared with control, although this difference was not statistically significant (Figure 2.3A). There was no difference between continuous PTH and control groups at this time point (Day 4). At later time points (Day 8 and 14) however, MSC cultures treated with continuous PTH contained significantly higher levels of ALP activity as well as an increased number of ALP expressing cells compared to control. In contrast, pulsatile PTH treatment significantly decreased both ALP activity and numbers of ALP expressing cells (Figures 2.3B and 2.3C). In all samples, we observed a steady decrease in the measured ALP activity from Day 4 to 14. This may be due to the fact that undifferentiated cells continued to expand, while differentiated cells withdrew from the cell cycle, leading to an effective dilution of ALP expressing cells.  109  Control  PTH-C (lOnM)  PTH-P (lOnM)  Day4 0.008 0.0061  0.005-I 0.  0.003 < .  0.002 0.001  0.000 Control  B  PTH-P (lOniti)  DayS *  0.006  f  PTH-C (lOnM)  0.004 0.003 0002 0.001  I—  0.00c  -  Control  C  PTH-C (lOnM) PTH-P (lOnM)  Day 14  0.0051  *  0.004-f  *  p0.0031 .  I  0.002.j  o.ooi-t 0.00 Control  PTH-C (lOnM) PTH-P (lOnM)  Figure 2.3: Effects of continuous and pulsatile PTH treatments on ALP activity and staining in the presence of DEX. MSC were cultured for up to seven 48-h cycle resulting in (A) 4 days, (B) 8 days, and (C) 14 days exposure to PTH. PTH-C = continuous PTH 0 < 0.05 as compared to control. and PTH-P = pulsatile PTH as described in Section 2.2.5. f  Next, we analyzed the effect of different regimens on changes in the transcription of osteogenic differentiation markers by qRT-PCR. Previous studies have identified PTHR1 and ALP as two osteogenic lineage markers that appear to be ubiquitously expressed by all osteogenic precursors. In contrast, the expression of most other markers including osteopontin and bone sialoprotein is limited to specific substages of the osteogenic differentiation hierarchy, and it appears to depend on a variety of other conditions such as for example, their anatomical source [21, 221. Thus, we focused our analysis on the ubiquitous PTHR1 and ALP transcripts, and a mature osteogenic differentiation marker, OC. Gene expression profiles for all three markers, PTHR1, ALP, and OC over the time of PTH treatment are illustrated in Figure 2.4. Pulsatile PTH treatment significantly down-regulated the expression of PTHR1 compared with the control cultures throughout the duration of this study, supporting the notion that it may inhibit the production of osteogenic cells. In contrast, continuous PTH appeared to up-regulate the expression of PTHR1 but was not significantly different when compared to control. On the other hand, continuous PTH treatment significantly up-regulated the expression of PTHR1 in comparison with pulsatile PTH (Figure 2.4A). Continuous PTH treatment was found to up-regulate the expression of ALP mRNA, confirming the results of enzyme activity assays and strongly suggesting that PTH regulates ALP expression at the transcriptional level. Pulsatile PTH delivery did not down regulate ALP expression compared to controls, although a significant difference was observed in comparison to continuous PTH treatment (Figure 2.4B). Moreover, the observed comparative expression of PTHR1 and ALP mRNA for all three groups correlated with the expression of a more mature osteogenic lineage marker, OC (Figure 2.4C).  111  To assess whether the observed effects of pulsatile vs. continuous PTH treatments may be explained at least partially through differential effects on cell proliferation, we measured mitogenic activity in the treated cultures using cell counts and MTT assays. No significant differences between in total cell count or cell proliferation kinetics were observed between continuous and pulsatile PTH treated samples (Figure 2.5). Thus, changes in cell proliferation are unlikely to play a major role in the observed effects of PTH treatments.  112  PTHRI  ALP  B  A I 0  I 0 0  C, 0  I  0 1  C 0  I  (n C) 1  x  Lii  I 0 a C,  0.0015  oc  C  Day 8 Day 14  .2 o.ooio  g  I  0.0005  Control  PTH-C (lOnM)  PTH-P (lOnM)  Figure 2.4: Gene expression profiles of MSC after continuous and pulsatile PTH treatments in the presence of DEX. MSC were cultured for up to seven 48-h cycle illustrating the changes in (A) PTHR1 and (B) ALP gene expressions after 8 and 14 days exposure to PTH. PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5. *13 < 0.05 as compared to control. #p.< 0.05 as compared to PTH-P. @13 < 0.05 as compared to PTH-C.  A  B  Day4 Day8  0.5  Day 14  0.4•  -%  C.,  0  0.3•  x 0)  (0 (0  Control  PTH-C (lOnM)  PTH-P (lOnM)  E  • -A -4-.  DayO  Day8  Control PTh-C (lOnM) Pm-P (lOnM)  Dayl4  Figure 2.5: Proliferation kinetics of MSC after continuous and pulsatile PTH treatments in the presence of DEX. MSC were cultured for up to seven 48-h cycle illustrating the changes in (A) total number of cells and (B) the proliferation kinetics as measured by the MTT assay, monitoring absorbance at 570 nm. PTH-C continuous PTH and PTH-P pulsatile PTH as described in Section 2.2.5. =  =  2.3.3  Effects of PTH on rat MSC differentiation and proliferation in the absence of DEX  The steroid hormone dexamethasone has well-documented effects on the ability of cells of the osteogenic lineage to differentiate [15, 23-25]. To detect functional interactions between the activation of the steroid signaling pathway and the effects of continuous and pulsatile PTH treatments on MSC differentiation and proliferation, MSCs were cultured with PTH in the absence of DEX for seven 48-h cycles (i.e. 14 days). In these conditions, neither continuous nor pulsatile treatments had any detectable effect on MSC differentiation or proliferation. Specifically, no difference was observed between continuous PTH, pulsatile PTH and untreated control cultures in PTHR1 or ALP mRNA expression, (Figure 2.6), or ALP activity (Figure 2.7). Moreover, cells proliferated at the same rate independent of the presence or absence of PTH (Figure 2.8).  115  —  I 0  ON  CD o 4-’ 4)  . 4-’ a) C  .2 U)  A  PTHRI  B  I 0 0..  ALP  (3 0  4-’  a)  .? 4-’  (U 4) 1..  C  ° U)  I Figure 2.6: Effects of continuous and pulsatile PTI-1 treatments on gene expressions in the absence of DEX. MSC were cultured for  seven 48-h cycle (i.e. 14 days). (A) PTHR1 and (B) ALP gene expression. PTH-C described in Section 2.2.5.  =  continuous PTH and PTH-P  =  pulsatile PTH as  0. 00  0.01 C.)  PTH-C (lOnM) PTH-P (lOnM) Figure 2.7: Effects of continuous and pulsatile PTH treatments on ALP activity in the absence of DEX. MSC were cultured for seven 48-h cycle (i.e. 14 days). PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5.  117  0.5  E  0.4  C 1%. L()  0.3 C.) I  0  --  -A-  < 0.1 --  0.0  I  I  DayO  Day8  Control PTH-C(lOnM) PTH-P (lOnM)  Dayl4  Figure 2.8: Effects of continuous and pulsatile PTH treatments on cell proliferation kinetics in the absence of DEX as measured by MTT assay, monitoring absorbance at 570 nm. MSC were cultured for seven 48-h cycle (i.e. 14 days). PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5.  118  2.3.4  Effects of PTH on rat MSC colony forming unit-fibroblast (CFU-F)  The number of cells capable of initiating a colony (colony forming units, CFUs) contained in primary culture is routinely used as a measure of the presence of immature, proliferative progenitors. In particular the enumeration of CFUs generating fibroblastic colonies (CFU-Fs) as a measure of self-renewal potential of MSC cultures is well supported in the literature [26]. To determine the effects of the different regimens of PTH treatment on the abundance of CFU-Fs, MSC cultures were treated with continuous or pulsatile PTH (100 nM) in expansion medium and osteogenic medium in the absence of DEX for four 48-h cycles (8 days). The data shown in Figure 2.9 demonstrate that continuous PTH treatment significantly increased the frequency of CFU-F in both expansion and osteogenic conditions (Figure 2.9A and 2.9B). A trend toward a similar increase was also observed with pulsatile PTH although in this case the differences were not statistically significant (Figure 2.9A and 2.9B). Strikingly, the frequency of CFU-F was dramatically decreased under osteogenic conditions compared to expansion conditions, independently of the treatment regimen used, supporting the use of this assay to detect immature cells (Figure 2.9A vs. 2.9B).  In another experiment, limiting dilutions of cells were plated in osteogenic conditions with the addition of DEX, and the cells subjected to either continuous or pulsatile PTH (10 nM) treatments. Both treatments were found to increase the frequency of CFU-Fs compared to control (Figure 2.9C), consistent with the results obtained in the absence of DEX in bulk assays (Figure 2.9B). The observed CFU-F frequency difference between the bulk (Figure 2.9B) and limiting dilution (Figure 2.9C) experiments is most likely attributed to the biological variation of the MSCs in isolation processes from two different rats and the  119  addition of DEX. The above findings suggest that PTH is capable of increasing the frequency of immature cells in MSCs cultures, independent of whether it is delivered in a continuous or pulsed fashion.  120  Bulk Assay: Expansion Medium 0.03  Bulk Assay: Osteogenic Medium  B  A  L  0002  0 u.Q  u0  0.00  -.  Control  PTH-C (lOOnM)  F.rl-P(1  Limiting Dilution Assay: Osteogenic Medium with DEX  L’J 0.03 LI.  C  —  0002 wt. ::.::  L  0.01 u.O  Control  PTH-C(lOnM)  PTH-P(lOnM)  Figure 2.9: Effects of continuous and pulsatile PTH treatments on CFU-F by bulk and limiting dilution assays. MSC were exposed to continuous and pulsatile PTH (10 or 100 nM) for three to four 48-h cycles (i.e. 6 8 days), (A) in expansion medium without DEX, (B) in osteogenic medium without DEX, and (C) in osteogenic medium with DEX before cells were re-plated and cultured for another 6—8 days in expansion medium only to allow for the formation of CFU-F. PTH-C = continuous PTH and PTH-P = pulsatile PTH as described in Section 2.2.5. *P < 0.05 as compared to control. —  2.4 Discussion Mesenchymal stromal cells have been proposed as a convenient source of cells for use in bone tissue engineering and regeneration therapies due to their accessibility, high content of osteoprogenitors and chondroprogenitors and especially of cells displaying self-renewal ability and thus potentially capable of continuing to replenish the osteoprogenitor compartment long term [27-29]. The effects of DEX [24, 25] and various other extrinsic factors on these cells have been studied [15, 23]. In contrast, and surprisingly given their clinical importance, the effects of continuous and pulsatile PTH have been thoroughly investigated on osteogenic cells [10-13, 20, 30-33] (also see reviews [7, 34]), but not on these complex MSC cultures. The study in this chapter provides a first assessment of the effects of different modalities of PTH treatment on the ability of MSCs to generate osteogenic cells, and to promote the maintenance of clonogenic progenitors under osteogenic conditions.  MSC cultures are heterogeneous in nature, with one of the key contaminants represented by hematopoietic cells expressing the CD45 marker [3 5-37]. In this study, the rat MSC populations were examined for CD45 positive cell content prior to PTH and DEX treatments, and found to contain less than 4% CD45 hematopoietic cells (Figure 2.1). This level of contamination is substantially lower than what has been reported in previous studies [24, 25], and it is unlikely that it would significantly affect the interpretation of the findings presented in this chapter. In addition and in agreement with previous literature [2] it was confirmed that the MSC cultures used in this study contained cells capable of differentiating into osteoblasts, adipocytes and chondrocytes (Figure 2.2).  122  This study used ALP activity assays and ALP expressing cells (Figure 2.2), as well as analysis of PTHR1, ALP and OC gene expressions (Figure 2.3), to show that continuous treatment with PTH promotes osteogenic differentiation from mesenchymal stromal cells, while pulsatile PTH treatment results in inhibition. Previously, Simmons et al. [38] investigated PTH-mediated cyclic adenosine monophosphate (cAMP) production in rat MSC and found no effect after 15 mm  of PTH stimulation. However, this study did observe a  differentiation effect with PTH, and thus suggests there might be an ‘optimal’ PTH treatment concentration and duration to influence and even control rat MSC differentiation, similar to that observed for rat calvariae osteoblast differentiation [20, 39]. In support of this notion, the results presented in this chapter showed that continuous (daily) treatment with PTH enhanced MSC differentiation along the osteogenic lineage. In contrast, pulsatile PTH treatment, where MSCs were exposed to only the first six hours in a 48-h cycle, inhibited MSC differentiation along the osteogenic lineage.  These results are noticeably different from the findings of studies on rat calvariae osteoblasts and osteoblastic cell lines, where it has been consistently shown that continuous PTH inhibits osteoblast differentiation, while pulsatile PTH stimulates osteoblast differentiation [10-13, 20, 30-33]. Such disparity might be attributed to the fact that in this study, the effects of PTH take place on the generation of osteoprogenitors from immature cells, rather than on downstream maturation events evident when more differentiated populations are used. This explanation is in agreement with our understanding of the anabolic effects of pulsatile PTH in vivo, where the increase in bone formation is due to the direct action of PTH on mature  123  matrix-synthesizing osteoblasts, delaying their apoptosis to ultimately increase their numbers [101. Another potential reason for the apparent discrepancies between the results presented in this chapter and published reports could be ascribed to differences in the culture conditions. For example, previous studies using calvariae osteoblast and osteoblastic cell lines were performed in the absence of DEX which, in this study, is critical to observe differences in the effects of the two treatments. Moreover, the importance of differences in culture conditions is also evidenced by the apparently contrasting results found in our work compared to that of Rickard et a!., who showed that under adipogenic conditions pulsatile PTH inhibited adipogenic and possibly promoted osteogenic differentiation [14]. Under such adipogenic conditions, it is possible that the enhanced osteogenic differentiation observed may be a result of the stochastic differentiation of immature cells in the cultures, whose frequency would be increased by PTH treatment according to the findings presented here, rather than the promotion of osteoprogenitor differentiation. Indeed Rickard et al. reported that intermittent PTH treatment did not affect ALP, OC, bone sialoprotein and Runx2 mRNA expressions, but in fact, decreased osteopontin and PTHR1 mRNA expressions [14], as seen in the current study. The effects of PTH treatments on the expression of adipogenic differentiation markers such as peroxisome proliferator-activated receptor gamma (PPARy) and lipid protein lipase (LPL) were also examined in this study (data not shown), but under the examined osteogenic culture conditions, highly variable expression levels were found suggesting that an effect on this lineage may only be observed when cells are primed toward an adipogenic fate [14]. Thus, while we have described the promotion and suppression of osteogenic differentiation by continuous and pulsatile PTH respectively, conclusions  124  concerning PTH effects on other lineages such as adipogenesis cannot be drawn without further experimentation.  DEX has been shown to alter the differentiation of rat MSC when administered alone [15, 23-26]. We showed the presence of DEX to be critical to reveal the effects exerted by PTH on MSC differentiation along the osteogenic lineage. In the absence of DEX, neither continuous nor pulsatile PTH treatment had any detectable effect on ALP activity, or expression of PTHR1 and ALP mRNA levels. The notion that DEX alone can alter the composition of rat MSC cultures may help to shed light on the mechanism of action of PTH in the experiments described here. As previous studies have suggested, there exist at least two osteoprogenitor populations in rat bone marrow, a more abundant, less differentiated population, and a less represented, more mature population [40]. The less differentiated population requires glucocorticoids such as DEX to undergo differentiation, whereas the more differentiated population is capable of differentiation without the addition of glucocorticoids. Since research has shown DEX increases rat MSC differentiation [15, 2326], the differentiation effects exerted by continuous and pulsatile PTH in the presence of DEX suggest that PTH mainly affects the more differentiated osteoprogenitor population. This subpopulation is increased in the presence of DEX, which stimulates its generation by pushing earlier osteogenic cells to differentiate. Thus, in the absence of DEX, there may not be a sufficient number of mature osteoprogenitor present in MSC cultures to detect the effect of PTH treatments. Similarly, Kostenuik et al. noted that PTH treatments had no effect on MSCs with the absence of DEX in the culture media [41].  125  It has been suggested that osteoprogenitor differentiation and proliferative (self-renewal) capacity are inversely correlated [1]. Nonetheless, in this study, continuous and pulsatile PTH treatments did increase and decrease rat MSC osteogenic differentiation, respectively, in the absence of any detectable change in overall MSC proliferation. An unequivocal interpretation of these findings is difficult in light of the known heterogeneity of MSC cultures. However, these findings suggest that PTH action may be carried out through a mechanism that is significantly different from that underlying the effects of other regulatory stimuli such as BMP-2 [15] and calcitriol [23], which have been shown to affect both the differentiation and proliferation of rat MSCs. Potentially, the effects of PTH may be more specific, affecting differentiation potential and/or kinetics directly rather than indirectly through changes in proliferation efficiency.  Another interesting finding stemming from the current study is that treatment with PTH, either continuous or pulsatile, increased the CFU-F content of rat MSCs in all media conditions investigated. These results suggest PTH may play a role in the self renewal of clonogenic progenitors found in MSC cultures. CFU-Fs are themselves a heterogeneous cell population, with only a minor portion of clonal colonies initiated by multipotential stem cells [42, 43]. Thus it cannot be concluded that PTH enhances the self renewal of multipotent cells without further experimentation, although this possibility must be kept in consideration. Nevertheless, the effect of PTH on CFU-F frequency appears to be distinct from that on osteogenic differentiation, as it is not influenced by treatment regimen.  126  2.5 Conclusion The results of this chapter demonstrated the effects of continuous and pulsatile PTH treatments on rat MSC differentiation and proliferation in vitro. Osteogenic differentiation of rat MSC was found to increase with continuous PTH treatment, and decrease with intermittent PTH exposure. The observed effects of PTH were strongly dependent on the presence of DEX. Conversely, MSC proliferation was not influenced by PTH, independent of treatment regimen and presence or absence of DEX. Furthermore, this work raises the possibility that PTH treatment may modulate stem/progenitor cell activity within MSC cultures.  127  2.6 References 1. 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Onyia JE, Helvering LM, Gelbert L, Wei T, Huang 5, Chen P, et al. Molecular profile of catabolic versus anabolic treatment regimens of parathyroid hormone (PTH) in rat bone: an analysis by DNA microarray. J Cell Biochem 2005 May 15;95(2):403-418. 32. Robinson JA, Susulic V, Liu YB, Taylor C, Hardenburg J, Gironda V, et al. Identification of a PTH regulated gene selectively induced in vivo during PTH-mediated bone formation. J Cell Biochem 2006 Aug l;98(5):1203-1220. 33. Wang YH, Liu Y, Rowe DW. Effects of transient PTH on early proliferation, apoptosis, and subsequent differentiation of osteoblast in primary osteoblast cultures. Am J Physiol Endocrinol Metab 2007 Feb;292(2) :E594-603. 34. Compston JE. Skeletal actions of intermittent parathyroid hormone: effects on bone remodelling and structure. Bone 2007 Jun;40(6):1447-1452. 35. Colter DC, Class R, DiGirolamo CM, Prockop DJ. Rapid expansion of recycling stem cells in cultures of plastic-adherent cells from human bone marrow. 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Skeletal unloading causes resistance of osteoprogenitor cells to parathyroid hormone and to insulin-like growth factor-I. J Bone Miner Res 1999 Jan;14(1):21-31. 42. 910.  Aubin JE. Advances in the osteoblast lineage. Biochem Cell Biol 1998;76(6):899-  43.  Aubin JE. Bone stem cells. J Cell Biochem Suppi 1998;30-31:73-82.  131  Chapter 3: The differential in vitro and in vivo responses of bone marrow stromal cells on novel porous gelatin-alginate scaffolds 2  3.1 Introduction Loss or deterioration of musculoskeletal tissue due to disease, trauma or surgical intervention often leads to premature disability and a reduced quality of life for patients. Restoring the normal structure and function of these defects requires replacement or repair of damaged tissue for which tissue engineering and stem cell therapy hold tremendous potential [1]. The extent to which marrow stromal cells (MSC5, mesenchymal stem cells [2]) participate in the regeneration of tissues such as bone, cartilage, and fat is under active investigation [3-15]. The successful regeneration of tissues in animal models has led to preclinical and clinical trials using MSCs for bone repair and regeneration [10, 16, 17].  The delivery of MSCs for tissue regeneration primarily uses 3-D porous scaffolds to support MSC seeding, proliferation, differentiation and implantation. The use of a scaffold, will not only retain the cells within the defect site, but can also function as a substrate for tissue ingrowth and vascularization. In addition to having the appropriate porosity and surface properties for the candidate cells to adhere, infiltrate, and migrate, the ideal scaffold should also  be  biocompatible,  non-inflammatory,  non-immunogenic,  biodegradable,  and  bioresorbable. It should be readily fabricated in a reproducible manner and sterilizable. A large number of fabrication technologies have been applied to engineer biomaterials into porous scaffolds of high porosity and surface area for cell delivery [18-22]. These techniques 2  A version of this chapter has been submitted. Yang C et at., (2009) The differential in vitro and in vivo responses of bone marrow stromal cells on novel porous gelatin-alginate scaffolds.  132  include solvent casting combined with particulate leaching [23], emulsion freeze-drying [24], electrospinning [25, 26] and rapid prototyping [27-29], among others. There are disadvantages associated with these methods that vary, depending on the particular fabrication process, and these have been reviewed [22, 30-32].  In this work, the microwave vacuum drying process [33, 34], depicted in Figure 3.1 was employed to engineer porous scaffolds. In this approach, a solid or semi-solid hydrogel is formed by dissolving suitable biomaterials (e.g. gelatin and alginate) in an aqueous solvent. The gel solution is cast and allowed to set with or without the addition of cross-linker, and if necessary cut into desired shapes and sizes. An optional step might be freezing the gel to allow the formation of ice crystals that act as “porogens”. Ice crystal size will ultimately affect the size and number of pores in the final scaffold. Size is controlled by adjusting the freezing rate and freezing temperature of the gel; low temperatures and fast freezing will result in small crystals, while slow freezing at higher temperature yields larger crystals [3336].  133  Hydrogel Formulation  —  Hydrogel Moulds  A B Water Vapour  ‘1 4 D Under Vacuum  Porous Scaffolds  Figure 3.1: Schematic diagram of the microwave vacuum drying process to fabricate porous scaffolds. (A) Formulation of hydrogel with or without additional agents such as crosslinkers and drugs. (B) Hydrogel cast into molds. (C) Hydrogel is placed into microwave vacuum drying chamber, where electromagnetic microwave energy penetrates into the hydrogel and is converted to thermal energy for the subsequent in situ vaporization of water which generates an expansive force to create porous structures in the matrix. (D) Dry porous scaffolds.  134  The hydrogel matrix is then placed into a microwave chamber that is under vacuum. Applied electromagnetic microwave energy penetrates into the interior of the hydrogel matrix and is converted to thermal energy for the subsequent in situ vaporization of water which generates an expansive force to create porous structures in the matrix. The absolute pressure in the chamber is maintained between 30 and 760 mmHg, and the applied microwave energy is typically between 150 and 5000 Watts per kilogram of initial mass. Since the microwave energy is applied under vacuum, solvents will boil at a much lower temperature than at atmosphere pressure. For example, at a pressure of 24 mmHg, water will boil at 25 °C without any addition of energy (heat). After drying, porous scaffolds can be further postprocessed if needed or desired. For example, cross-linking for additional stability or reshaping, if it was not done prior to microwave vacuum drying. The advantage of the microwave vacuum drying technique is that the process is under low vacuum where a pressure gradient is generated to enhance the drying rate (in the timeframe of minutes), and reduce the boiling point of water. Since the scaffold is always kept at relatively low temperature (< 37 °C) and the drying process occurs rapidly at low oxygen pressure, the biomaterial itself and any incorporated biological factors and drugs that are sensitive to thermal degradation and/or oxidation are not affected or compromised [33, 35, 36].  Despite the many in vitro and in vivo studies that have demonstrated tissue regeneration with the delivery of MSCs on porous scaffolds, only a limited number of studies have compared in vitro and in vivo responses of MSCs on scaffolds [37, 38]. Using different degrees of mineralized bovine bone-derived matrix as scaffolds, Mauney et al. [37] examined the ability of seeded MSCs to differentiate along the osteogenic lineage in vitro for 14 days and its  135  bone-forming capacity in vivo when implanted subcutaneously in nude mice for 8 weeks. Under in vitro conditions, mineralized bone-derived scaffolds showed higher osteogenic differentiation capacity compared to fully demineralized scaffolds, a trend not found in vivo, where the extent and frequency of bone formation were similar for all the different mineralized bone-derived scaffolds [37]. In another study, Mendes and colleagues demonstrated that the in vitro and in vivo osteogenic potential of MSCs was not always correlated [38]. When MSCs were cultured on tissue culture plates with the addition of osteogenic differentiation factor dexamethasone, Mendes et al. found a consistent increase in the relative amount of cells expressing alkaline phosphatase (ALP). However, when MSCs were seeded on porous hydroxylapatite scaffolds and implanted subcutaneously in nude mice, nearly half of the samples (6 out of 14) did not form bone in vivo [38]. Even though these studies [37, 38] examined the in vitro and in vivo osteogenic potential of MSCs, they did not explicitly examine the fate and changes in the MSC population after in vivo transplantation.  Therefore, in this chapter, the physical characterization and bioresorption, as well as the differential in vitro and in vivo responses of MSCs seeded on porous gelatin-alginate scaffolds, prepared with different cross-link density using the novel microwave vacuum drying process, were investigated. Specifically, MSCs were retrieved from explanted scaffolds after in vivo implantation and compared directly to MSCs cultured in vitro. MSC proliferation and self-renewing potential were quantified using BrdU incorporation and CFU F assay, respectively. MSC differentiation potential along three lineages: osteogenic, chondrogenic and adipogenic were examined through gene expressions with qRT-PCR.  136  3.2  3.2.1  Materials and methods  Materials  Gelatin (type A, from porcine skin, bloom  300), alginate (alginic acid sodium salt from  brown algae), N-hydroxysuccinimide (NHS, 98%), N-ethyl-N ‘-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC,  98%), and paraformaldehyde (PFA) were all obtained  from Sigma-Aldrich (Oakville, ON, Canada). Mesencult® Basal medium (cat # 05401), fetal bovine serum (FBS, cat # 06471), osteogenic FBS (cat # 06473), 13-glycerophosphate, ascorbic acid, and dexamethasone were purchased from StemCell Technologies (Vancouver, BC, Canada). Allophycocyanin (APC, Alexa Fluor 647) fluorescent protein conjugated anti BrdU antibody (APC-anti-BrdU) and reverse-transcription reagents using the Superscript III RT system were acquired from Invitrogen TM (Burlington, ON, Canada). Bromodeoxyuridine (BrdU, 10 mg/mL) solution was purchased from BD Biosciences (Mississauga, ON, Canada). Non-Obese Diabetic/Severe Combined Immunodeficient (NOD/SCID) mice, 3  —  6  weeks old were purchased from The Jackson Laboratory (Bar Harbor, Maine USA).  3.2.2  Fabrication of porous gelatin-alginate scaffolds using microwave vacuum drying  To engineer porous gelatin-alginate scaffolds, 50% (w/v) gelatin and 5% (w/v) alginate were first dissolved in distilled water at 60°C and 25°C, respectively. In a Teflon® container, 6 g of gelatin, 2 g of alginate, and 8 mL of cross-linking solution were mixed thoroughly and crosslinked at 4°C for 24 h to produce a hydrogel. For high cross-linked hydrogel, the crosslinking solution contained 34 mM EDC and 17 mM NHS dissolved in distilled water, while for low cross-linked hydrogel, the cross-linking solution contained 17 mM EDC and 8.5 mM 137  NHS. Hydrogel disks of the desired shape and size (10 mm diameter x 6 mm height) were subsequently stamped out using a cork borer, soaked in distilled water at room temperature for 12 h and then washed 3 times with distilled water to remove excess cross-linking agents. The hydrogel was placed into a microwave vacuum dryer (EnWave Corporation, Vancouver, Canada) for drying. 300 watts of microwave power under an absolute pressure of 30 mmHg was applied for 30 mm to vaporize the water in the hydrogel and produce the dried scaffold. To expose the rough and porous structures of the scaffold, the dried outer ‘skin’ layer produced after drying was removed using a sharp razor blade, and scaffolds were stored under vacuum until use or analysis. For all in vitro and in vivo studies, the scaffolds used were approximately 10 mm in diameter and 6 mm in height before they were rehydrated.  3.2.3  Mercury intrusion porosimetry and scanning electron microscope (SEM) analyses  The total porosity, median pore size, and pore size distribution of the scaffolds were measured using a mercury intrusion porosimeter (Micromeritics TM AutoPore III, Folio Instruments Inc., ON, Canada). Mercury injection capillary pressures used for measurements ranged from 0.5 to 33,000 psia. Penetrometer (solid, 3cc bulb, 1.190cc stem volume) with approximately 26  —  38% of the stem volume filled was used for all the samples analyzed,  which were approximately 40  —  60 mg in weight. SEM analysis was performed on the  surface and cross-section of scaffolds sheared from the center using a sharp razor blade and mounted on an aluminum stud. All samples were sputter-coated with a layer of 60:40 alloy of gold:palladium using a Denton Vacuum Desk II sputter-coater (Moorestown, NJ) at 50  138  millitorr. SEM images were then captured using a Hitachi S-3000N system (Tokyo, Japan) scanning at 10  3.2.4  —  20 keV.  Equilibrium water uptake measurements  Scaffolds were accurately weighed, placed into 30 mL of distilled water in centrifuge tubes and shaken at 30 rpm and 37 °C for 24 h. Subsequently, ‘wet’ scaffolds were removed, gently patted dry with Kimwipe® and weighed. The amount of water uptake was calculated using the following equation:  % Water Uptake  =  Ww—WD  x  100%  Equation (2.1)  where, WD is the weight of the dry scaffold and Ww is the weight of the scaffold after 24 h of water uptake.  3.2.5  Biodegradation and bioresorption  All animal work was done in accordance with the University of British Columbia Animal Care Committee requirements and all scaffolds were sterilized with 0.25 Mrad of gamma irradiation produced from a Cobalt-60 source (GammaCell 200, Atomic Energy Canada Ltd.) before they were used in any in vitro cell culture or in vivo animal experiments. Briefly, high and low cross-linked scaffolds were first rehydrated in culture medium at 37 °C for 24 h and the dimensions (diameter and height) of the rehydrated scaffolds were recorded. After rehydration, high and low cross-linked scaffolds were approximately 14 mm diameter x 8 mm height and 16 mm diameter x 9 mm height, respectively, in dimensions. One high and  139  one low cross-linked scaffold each was subcutaneously (Sc) implanted into the backs of the same NOD/SCID mice for 7 and 21 days, with n  =  5 for each time point. Specifically,  following isoflurane anesthesia, two longitudinal skin incisions  (‘—‘  1 cm in length) were made  on the dorsal surface of each mouse. Blunt dissection scissors was then used to form sc pouches where the incisions were made and a single scaffold was implanted into each pouch. Following implantation, each incision was closed using surgical staples. After sacrifice, scaffolds were carefully extracted, fixed with 4% PFA for 30 mill and the final dimensions (diameter and height) of the scaffolds were measured for biodegradation and bioresorption evaluation. The extent of bioresorption of the scaffold was determined from volume loss, and was calculated using the following equation:  % Volume Loss  =  1 V  —  VF  x 100%  Equation (2.2)  where, V 1 is the initial rehydrated volume of the scaffold and VF is the fmal volume of the scaffold after either 7 or 21 days. The volume of the scaffold was calculated using the formula, V  3.2.6  itr h 2 , where r and h are the hydrated radius and height, respectively.  Isolation and cell culture of rat marrow stromal cells (MSCs)  MSCs were isolated from 6-week Sprague Dawley rats expressing transgenic green fluorescent protein (GFP) (NBRP, Japan). All procedures were approved by the University of British Columbia, Animal Care Committee. Briefly, femora and tibiae harvested from euthanized rats were crushed to release total bone marrow cells. MSCs were subsequently isolated by virtue of their adherence to tissue culture plastic cultured in Mesencult® medium supplemented with 15% FBS and 100 ig/mL penicillinlstreptomycin at 37 °C, and 5% CO . 2 140  expanded to passage two and cryopreserved until they were used. MSCs used for all in vitro and in vivo experiments were further expanded to passage 3  —  6 and cultured in Mesencult®  medium supplemented with 15% FBS and 100 tg/mL penicillinlstreptomycin, unless otherwise noted.  3.2.7  In vitro culturing of MSCs on scaffolds  To investigate the in vitro responses of MSCs on the porous gelatin-alginate scaffold, MSCs were seeded and cultured as follows: Firstly, the high cross-linked scaffold, rehydrated with culture medium at 37 °C for 24 h was drained of excess fluid by centrifugation at 1500 rpm for 2 mm in centrifuge tube containing a 70 pm mesh support. I x 106 MSCs resuspended in 200 tL of medium were then gently added drop-wise to the surfaces of individual scaffolds in 24-well non-tissue culture plates and incubated at 37 °C and 5% CO 2 for 20 mm to allow the MSCs to adhere to the scaffold. Subsequently, each culture well containing a scaffold seeded with MSCs was topped off with an additional 1 mL of medium and cultured at 37 °C and 5% CO 2 for various lengths of time, depending on the experiment. For all experiments, culture media were changed every 24 to 48 h, and the recovery of MSCs from the scaffold for subsequent analyses was performed according to the procedure in Section 9.  3.2.8  Implantation of MSC seeded scaffolds into NOD/SCID mice  The in vivo responses of MSCs on the porous gelatin-alginate scaffold were determined from MSCs extracted from scaffold after sc implantation into NOD/SCID mice. Briefly, 1 x 106 MSCs were seeded onto rehydrated high cross-linked scaffold as described above and  141  cultured for 24 h at 37 °C and 5% CO 2 to allow MSCs to attach to the scaffold. MSC seeded scaffolds were then implanted subcutaneously according to the surgical procedure in Section 5 for different time periods, depending on the experiment, with n  =  3  —  4. After sacrifice,  MSC seeded scaffolds were explanted and MSCs were recovered as described below.  3.2.9  Recovery of GFP MSCs from scaffolds  After in vitro and in vivo experiments, GFP MSCs were recovered from high cross-linked scaffolds by enzymatic digestion with 2 mE of collagenase D (1.5 U/mL)/dispase 11(2.4 U/mL) solution at 37 °C for 1 h. The digested sample was then filtered through a 70 urn cell strainer and the desired GFP MSCs population was sorted and collected using a Fluorescence Activated Cell Sorter (FACS, BD FACSVantage SE) flow cytometry system that gated for the GFP MSCs. Sorted and collected GFP MSCs were immediately used for analysis.  3.2.10 MSC proliferation assessment using BrdU incorporation In vitro and in vivo MSC proliferation on the high cross-linked porous gelatin-alginate scaffold was assessed for 7 days by the incorporation of BrdU into the DNA during DNA synthesis of proliferating cells. MSCs were either cultured in vitro or sc implanted into NOD/SCID mice as previously described. For in vitro labeling of proliferating cells, 10 jiL of 1 mM BrdU diluted in culture medium was added to each well containing MSCs seeded scaffold on day 3, 4, 5 and 6 of the seven day culture period. For in vivo labeling, mice were given drinking water containing 0.8 mg/mL BrdU starting on day 3 till the end of the seven  142  days. In addition, on days 3 and 5, 100 j.iL of 1 mg!mL BrdU solution were also injected directly into the scaffold seeded with MSCs that were sc implanted to ensure the total dose of BrdU was approximately equal among all the mice. After 7 days of the in vitro and in vivo studies, MSCs on scaffolds were digested using collagenase/dispase solution and filtered as described above. Cellular incorporation of BrdU was then detected with APC-anti-BrdU antibody following membrane permeabilization and MSC proliferation was quantified by Fluorescence Activated Cell Sorter system (FACS, BD FACSCalibur SE) gated for GFP and APC labeled cells, with n  =  3 for statistical analysis. Proliferation of GFP MSCs was  determined as a percentage, by taking the total number of double positive GFP and APC cells divided by the total number of GFP cells recovered and was calculated using the following equation:  % proliferation =  # of GFPand APC cells x 100% # ofGFPcells+(# of GFPandAPC cells)  Equation (2.3)  3.2.11 Immunohistochemistry To visualize and qualitatively assess the viability and attachment of MSCs on the porous gelatin-alginate scaffold after in vitro culturing and in vivo implantation, scaffolds seeded with MSCs were fixed with 2% PFA in PBS for 30 mm  and processed as follows. After  fixation, scaffolds were embedded in Tissue Tek® (Sakura), cryo-sectioned and stained with Hoechst to label the DNA of the MSCs. Stained sections were then imaged with a fluorescence microscope using both GFP and DAPI filters.  143  3.2.12 Quantitative real time-polymerase chain reaction (qRT-PCR) To quantify MSC gene expression after 7 and 21 days of in vitro culturing and in vivo implantation on high cross-linked porous gelatin-alginate scaffold, mRNA from retrieved MSCs was extracted, reverse-transcribed and subjected to quantitative real time PCR. mRNA was extracted from FACS sorted GFP MSCs using QIAGEN RNeasy® Mini Kit according to the manufacturer’s instructions. Subsequently,  1 ig of total RNA was reverse-  transcribed using the Superscript III RT system to generate complementary DNA (cDNA). The TaqMan® probes-based detection system (Applied Biosystems) was used and qRT-PCR reactions were performed in an Applied Biosystems 7900HT Fast Real-Time PCR system (Foster City, CA, USA). TaqMan® primers assay for all genes examined were from Applied Biosystems and their manufacturer’s assay numbers were as follows: glyceraldehyde 3phosphate dehydrogenase (GAPDH), Hs0275 8991 _g 1 (cross-reactive for rat); osterix, Rn01761789_ml; Runx2 (Cbfal), Mm03003491_ml (cross-reactive for rat); alkaline phosphatase (ALP), Rn0056493 1_mi; osteocalcin (OC), Rn00566386_gl; bone sialoprotein (BSP), Rn00561414 ml; aggrecan, Rn00573424_ml; SRY (sex determining region Y)-box 9 (Sox9), RnO 1751 069_mH; lipoprotein lipase (LPL), Rn005 61482_mi; and peroxisome proliferator-activated receptor-gamma (PPARy), Mm00440945_m 1 (cross-reactive for rat).  3.2.13 Colony-forming unit fibroblast (CFU-F) assay The presence of CFU-F in the MSC population after 8 days of in vitro culturing and in vivo sc implantation on the high cross-linked porous gelatin-alginate scaffold were assessed by bulk assay as follows. After recovery from scaffolds, FACS sorted MSCs were re-plated at low density (100 cells/cm ) in 12-well tissue culture plates and cultured in medium for 15 2  144  days, with medium change every 3 or 4 days. Subsequently, MSCs were washed with PBS, fixed with methanol for 5  —  10 mm, and stained with Giemsa staining solution. Colonies  containing> 20 cells per foci were then counted under a microscope, with n  =  4 for statistical  analysis.  3.2.14 Data analysis For statistical analysis, the Student’s t-test (unpaired, two-tailed) in GraphPad Prism® version 5.00 for Windows (GraphPad Software, San Diego, CA, USA) was used to determine level of significance. For all experiments, results are expressed as mean ± SD (n  =  3  —  5), and data  presented are representative of the results obtained in two independent experiments. Note for the purpose of statistical analysis of gene expression (qRT-PCR) data, in the event where no expression was detected or expression level was below the limit of quantiation, a zero value was assigned.  145  3.3 Results  3.3.1  Scaffold physical characterization  Porous gelatin-alginate scaffolds fabricated by a microwave vacuum drying process are shown in Figure 3.2 and their physical properties are summarized in Table 3.1. The effects of cross-link density were investigated with the production of two different cross-linked scaffolds: high and low. It was found that the total porosity of high cross-linked scaffold was approximately 82%, which was significantly lower than the low cross-linked scaffold that had a total porosity of approximately 88%. However, the median pore diameter and the pore size distribution of the high and low cross-linked scaffolds were found to be similar. The median diameter was between 1 greater than 90 Jim, and 70  —  —  2 tim, with only approximately 3  78% in the range of 1  —  —  5% of the pores being  90 !Im. SEM images illustrated in  Figure 3.2 confirmed the wide range of pore sizes in the scaffold and revealed that many of the larger pores were not interconnected (closed). There was no significant difference in the equilibrium water uptake of the high or low cross-linked scaffolds (Table 3.1).  146  !‘/1’/1’/1’Iuiij1i1i1lIi  Figure 3.2: Digital photographs and SEM micrographs of high cross-linked porous gelatin-alginate scaffolds. (A) and (C) are top views, (B) side view, and (D) cross-sectional view of the scaffold.  Table 3.1: Summary of the physical and bioresorption properties of the porous gelatin-alginate scaffolds  Physical Properties  Bioresorption  Total Porosity  Median Pore Diameter  >  Pore Size 90 pm  Pore Size 1 —90 jim  <  Pore Size I jim  Water Uptake  Volume Loss after 7 Days  Volume Loss after 21 Days  Sample  (%)a  (pm)a  (%)a  (%)a  (%)a  (% wt)b  (%)c  (%)c  Highx-linked  81.73±1.23*  1.14±0.31  3.31 ± 1.68  69.50±3.37  27.19±6.54  589 ±32  17±11  13±16#  Low x-linked  87.88 ± 2.21  1.91 ± 0.66  4.60 ± 1.70  77.63 ± 5.48  17.77 ± 5.53  627 ± 66  60 ± 32  100 ± 0  Mean ± SD, 4. *p < 0.005 compared to low x-linked scaffolds b % weight increase after 24h soaking in distilled deionized water @ 37°C. Mean ± SD, n = 5 C Bioresorption after subcutaneous implantation in NOD/SCID mice. Mean ± SD, n = 5. **p <0.05, linked scaffolds a  <  5 x 10-6 compared to low x  3.3.2  Scaffold bioresorption  The biodegradation and volume loss leading to bioresorption of the high and low crosslinked porous gelatin-alginate scaffolds were assessed by subcutaneous implantation in NOD/SCID mice for 7 and 21 days. As illustrated in Figure 3.3 and summarized in Table 3.1, the low cross-linked scaffolds bioresorbed much more rapidly than the high cross-linked scaffolds. After 7 days, high cross-linked scaffold volume was reduced by approximately 17%, but retained much of its original shape. Moreover, even after 21 days of implantation, high cross-linked scaffolds still maintained their shape with no further volume loss. After 7 days, low cross-linked scaffolds lost  60% of their volume and no longer retained the  original shape. After 21 days, low cross-linked scaffolds were completely bioresorbed. None of the scaffolds showed any evidence of fibrous capsule formation. Furthermore, the scaffolds were found to ‘bond or adhere’ to both the upper subcutaneous tissue layer and the lower muscle layer at the implant site, and there was evidence of neovascularization around the scaffold (Figure 3.4).  149  High cross-linked porous scaffolds  After 7 days  Low cross-linked porous scaffolds  Complete bioresorption of scaffold Before  After 7 days  Before  After 21 days  Figure 3.3: Representative images of the biodegradation and bioresorption of high and low cross-linked porous gelatin-alginate scaffolds before and after subcutaneous implantation in NOD/SCID mice for 7 and 21 days. [Note: Scaffolds were rehydrated in culture medium before implantation which resulted in the pink colouration.]  Figure 3.4: Representative image of the neovascularization (arrows) surrounding high cross linked porous gelatin-alginate scaffolds after subcutaneous implantation in NOD/SCID mice for 21 days. [Note: Also notice on the right-hand side the absence of the low cross-linked scaffold which was completely bioresorbed after 21 days, with only a dashed-circle representing the missing scaffold.]  151  3.3.3  Multi-potentiality of rat MSCs  MSC cultures were derived from rat bone marrow and used at relatively early passages (P3  —  P6). A defining characteristic of marrow stromal cultures is their ability to differentiate along multiple mesenchymal lineages when exposed to the appropriate inducing signals. To confirm that MSC preparations used in this study satisfy this criterion, we exposed them to osteogenic, adipogenic and chondrogenic conditions for 3 weeks in vitro.  Isolated and  passaged MSCs were found capable of differentiating along the osteogenic, adipogenic and chondrogenic lineages to produce bone nodules, intracellular fat-containing vacuoles, and cartilage, respectively after 3 weeks of in vitro differentiation induction, as shown in Figure 2.2.  3.3.4  In vitro and in vivo proliferation of MSC seeded on scaffolds  MSC viability and proliferation on the high cross-liuked porous gelatin-alginate scaffold were assessed for 7 days by the incorporation of BrdU into proliferating cells. MSCs were able to attach, survive and proliferate on the high cross-linked scaffolds both in vitro and in vivo. The attachment and viability of the MSCs are shown in Figure 3.5. Moreover, through FACS analysis of GFP cells with BrdU incorporation, it was found that there was no difference in the proliferation of the MSCs cultured in vitro and implanted in vivo, 29 and 32 ± 19%, respectively (Figure 3.6).  152  ±  12%  A  B  Figure 3.5: Representative fluorescence microscope images of MSCs on high cross-linked porous gelatin-alginate scaffold after 1 week (A) in vitro culture (Hoechst stained, arrows), and (B) in vivo subcutaneous implantation (GPF, arrows).  60’ C  0  = Cu  I  0)  40’  .5 I  .  .  ........ F____wF_%::q: I. I. .  0  0)  .....  ••••Wa••••t •••••N••  20’  C-)  .  .  .  .  .  . ..  .  iii I.  I.  .  . . .. . . . _ — — — • • • • • • • •• • • •••••••••••• • • • • • • • •• • • • • • • • • • •• • • •••••••••••• • • • • • • • •• • • I • — — — — — — —  F 0’  ..  I  in vitro  in vivo  Figure 3.6: MSC proliferation after 7 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin-alginate scaffolds measured using BrdU incorporation in proliferating cells and analyzed using FACS. Mean ± SD,n3.  154  3.3.5  In vitro and in vivo differentiation of MSCs seeded on scaffolds  Using qRT-PCR, expressions in the transcription of osteogenic, chondrogenic and adipogenic lineage differentiation markers were quantified after 7 and 21 days of in vitro culture and in vivo implantation of GFP MSCs seeded on high cross-linked scaffolds. With the use of FACS sorting, GFP MSCs transplanted in vivo were recovered and analyzed without any contamination from host cells. For osteogenic differentiation, early transcription factors, Runx2 and Osterix, and late stage osteogenic markers, ALP, OC and BSP were assessed and their mRNA expression profiles are illustrated in Figure 3.7. In general, for all five osteogenic markers (Runx2, Osterix, ALP, OC and BSP) it was found that the in vitro culturing conditions increased MSC osteogenic differentiation compared to the in vivo implantation environment, except for Runx2 which had a significantly lower in vitro expression after 7 days. However, after 21 days, the in vitro Runx2 expression was higher compared with in vivo, following the same trend as the other osteogenic markers.  MSC chondrogenic differentiation was examined by the mRNA expressions of early marker, Sox9 and later marker, aggrecan, and the results are shown in Figure 3.8. Overall, mRNA transcript levels of both Sox9 and aggrecan were expressed higher in vitro compared to in vivo, similar to all the osteogenic markers. One exception was at Day 7 where aggrecan was found to be expressed significantly lower in vitro than in vivo, but by Day 21, in vitro expression was significantly higher than in vivo. For adipogenic differentiation, PPARy and LPL mRNA expressions were measured and shown in Figure 3.9. It was found that the earlier differentiation marker, PPARy was expressed higher in vitro compared to in vivo only  155  at day 7, while the later stage marker, LPL was expressed higher in vitro than in vivo at both day 7 and 21.  156  Runx2 0.0025  Osterix *  0.0020  in vitro in vivo  T  T  0  2  -i 0 . 0 04 0.003-i  1  in vitro in viva  I  0.002 0.001  0.0015 0.0010 C  0  T 15 °°°° 0.000101  *  0.0005  0.00005.1  0 LU  0.0000  LU  Day 21  Day 7  a  BSP  0.005  *  Invitro  in vivo 0.00010  Day 21  oc  ALP 0.00015  “a y 7  in vitro in vivo  0 0.004 0  a  0.00015  in vitro invivo  *  (9 0.00010  0.003  ‘a 0.002  = 0.00005  C  0.00000  I.  0  LU  Ui  C  0  0.00005  ‘I’  0.00000 Dar 7  Day 21  NID N/D Day 7  NID Day 21  Figure 3.7: MSC osteogenic gene expressions after 7 and 21 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin-alginate scaffolds. Mean ± SD, n = 3, *p <0.05 compared to in vivo at the same time point. N/D = not detected.  Sox9 x  Aggrecan  0.  0.005  0 Go  ‘S a,  in vitro in vivo  0 0  0.  >  I  a, 0  :: 0.001  U, U, a,  1 *  • •••••  I  x  LU  Day7  ...  N/D  I.  Day 21  in vitro in vivo  *  (. 0.004  Day 7  Day 21  Figure 3.8: MSC chondrogenic gene expressions after 7 and 21 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin-alginate scaffolds. Mean ± SD, n 3, *p <0.05 compared to in vivo at the same time point. N/D = not detected.  PPARy  II  0.0025 0.0020 0.0015  LPL x  c:  in vitro in vivo  <  0.0010 0.0005  0.20  in vivo  0.15  c• *  0.05  1  in vitro  i t r  1T  I  Figure 3.9: MSC adipogenic gene expression after 7 and 21 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin-alginate scaffolds. Mean ± SD, n = 3, *p <0.05 compared to in vivo at the same time point.  3.3.6  MSCs CFU-F potential after in vitro culture and in vivo implantation  The number of cells capable of initiating a colony (colony forming units, CFUs) contained in primary culture is routinely used as a measure of the presence of immature, proliferative progenitors. In particular, the enumeration of CFUs generating fibroblastic colonies (CFU Fs) as a measure of self-renewal potential of MSC cultures is well supported in the literature [39]. There was no significant difference found in the ability of the MSC cultures to form CFU-F after in vitro culture or in vivo implantation on high cross-linked scaffolds, with in vitro and in vivo conditions resulting in 8  ±  3 and 4  ±  4 CFU-F per 1 x 1 o 3 cells re-plated,  respectively (Figure 3.10).  x1O  r  I  a) a  .1  ..............  ..  :•:•:•::•x•:  :—:•:—:—:—:•:•:  LL  5c Li.  0 ___.___._._._._._.___.  *  0’  _  LA... I  I  in vitro  in vivo  Figure 3.10: MSC CFU-F forming ability after 8 days of in vitro culture and in vivo subcutaneous implantation in NOD/SCID mice on high cross-linked porous gelatin-alginate scaffolds. Mean ± SD, n 4 for CFU-F.  160  3.4 Discussion Although an enormous range of different biomaterials have been fabricated into porous scaffolds using conventional and emerging technologies [18-22], gelatin and alginate are still among the most studied biomaterials for cell delivery due to their proven biocompatibility, biodegradability and broad biomedical applications, and their use has been comprehensively reviewed by Malafaya et al. [40]. The study in this chapter describes the engineering of porous gelatin-alginate scaffolds using a novel process developed by our collaborator, Dr. Tim Durance [33], termed microwave vacuum drying, and highlights the differential in vitro and in vivo responses of MSCs seeded on these porous scaffolds.  The processing time for porous gelatin-alginate scaffolds using microwave vacuum drying was very short, less than 30 minutes, and produced scaffolds with greater than 80% total porosity regardless of the extent of cross-linking investigated (Table 3.1). Increasing the concentration of the cross-linking agents (EDC/NHS) lowered the total porosity of the scaffolds, but produced similar median pore size and pore size distributions (Table 3.1). The reduction in total porosity with increased cross-link density is a function of increasing crosslinkages between the polymer chains and has been demonstrated by other groups [41, 42]. The irregular pore shape, small pore sizes and the limited interconnectivity with larger pores demonstrated in this study (Figure 3.2) and by Sundaram et al. [34] using the same technique were probably the biggest drawbacks of using microwave vacuum drying.  Cross-linking with EDC/NHS occurs via a reaction with carboxyl groups in gelatinlalginate to form an activated O-acylurea ester group. This intermediate may react with an available amino group in gelatin forming a stable amide cross-linkage or may hydrolyze, reforming the  161  carboxyl group and releasing a soluble urea product. NHS is added to improve cross-linking by reacting with either the carboxyl group or activated O-acylurea to form an activated ester which is much more stable in solution, and can then form cross-linkages [43]. Similar to this study, Choi et al. [44] have also reported changes in EDC concentrations that led to different degrees of cross-linkage but did not affect the pore size of the gelatin-alginate sponges produced. The main advantage of using EDC/NHS is its water solubility and ease of removal, thus enhancing biocompatibility [45].  Cross-link density in the scaffolds was shown to have a pronounced effect on biodegradation, mass/volume loss and bioresorption following implantation (Table 3.1 and Figure 3.3). Low cross-link density scaffolds were shown to be biodegraded and bioresorbed rapidly, being completely bioresorbed within 21 days. High cross-link density scaffolds were able to resist biodegradation and bioresorption much longer, retaining its original shape and losing only 13% of the original volume after 21 days. The porous gelatin-alginate scaffold was primarily composed of gelatin  (-  97% dry wt) and since gelatin biodegrades by proteolysis, the  increased cross-link density was likely to stereochemically reduce the access of proteases to gelatin chains, resulting in slower biodegradation and bioresorption [44, 46]. More interestingly, the porous gelatin-alginate scaffolds demonstrated excellent biocompatibility eliciting no fibrous encapsulation (Figure 3.4), unlike other studies where fibrous connective tissues were found to surround the subcutaneously implanted atelocollagen [47] and poly(ethylene glycol)-based [48] scaffolds in SCID mice. In addition, the observed angiogenesis and neovascularization (Figure 3.4) would likely assist the survival of the seeded MSCs, enhance the tissue integration process and favour de novo tissue regeneration.  162  The ability of the seeded MSCs to attach, survive and proliferate on the porous scaffold both in vitro and in vivo further supports the biocompatibility of the porous gelatin-alginate scaffolds (Figure 3.5). The comparable in vitro and in vivo proliferation of the MSCs suggests that adequate nutrients and metabolite transport were available to the seeded MSCs. In vivo, new blood vessels surrounded the scaffold and seeded MSCs, allowing implanted cells to proliferate at the same rate as in nutrient rich in vitro environments. However, MSCs were only found on the surfaces of the scaffolds either cultured in vitro or implanted in vivo, with no evidence of migration into the interior region of the scaffold (Figure 3.5). The lack of cell migration into the scaffold can be attributed to a few factors. MSCs were only seeded on the surfaces of the scaffold and without any chemoattractant in the interior of the scaffold, MSC migration would be limited, if at all. In addition, the small pore sizes and the lack of interconnectivity (Figure 3.2) in the scaffold likely served as a physical barrier preventing any possible cell migration.  In agreement with previous literature [2] we have confirmed that the MSC cultures used in this study contained cells capable of differentiating into osteoblasts, chondrocytes and adipocytes (Figure 2.2). By examining the transcription of the differentiation markers for osteogenesis (Runx2, osterix, ALP, OC and BSP, Figure 3.7), chondrogenesis (Sox9 and aggrecan, Figure 3.8), and adipogenesis (PPAR’y and LPL, Figure 3.9) it was determined that MSCs respond differently when cultured in vitro compared to being implanted in vivo. When implanted subcutaneously, MSCs generally expressed all three lineage differentiation markers at lower levels compared to in vitro culture conditions (Figure 3.7  —  3.9), with the  exception of Runx2 (Figure 3.7) and aggrecan (Figure 3.8) at Day 7, where both were found to be expressed lower in vitro than in vivo. Such anomalies might be attributed to the well  163  documented heterogeneity of the complex MSC culture [49-52]. Particularly, at the earlier time point (Day 7), the heterogeneous MSC culture used for the in vivo experiments might contain a small number of osteoprogenitors and chondroprogenitors which could produce the observed differences in the two more specific markers, Runx2 and aggrecan, for osteoblast and chondrocytes, respectively [49]. But by the end of 21 days, where the MSC population is expected to become more homogeneous due to their common microenvironment, and cause them to differentiate along the same lineage, both Runx2 and aggrecan were expressed significantly higher in vitro compared to in vivo, following the same trend as all the other differentiation markers investigated.  Nonetheless, it appears that the complex milieu of the subcutaneous environment where the MSCs were implanted, suppressed all lineage differentiation markers compared to in vitro expansion culture conditions. Being in the subcutaneous tissue, where neither osteogenesis nor chondrogenesis actually occurs in vivo, it is possible that the endogenous biological signals in this microenvironment only served to suppress such differentiation by MSCs. Moreover, since adipogenesis found in the subcutaneous tissue is primarily the result of more differentiated adipocyte precursors [53], the involvement of MSCs might be very minor, as reflected in their lack of differentiation found in vivo. The disparity between the in vitro and in vivo differentiation of MSCs found in this study has also been reported in the literature  [38]. Mendes et al. showed that although MSCs cultured on tissue culture plastic were consistently able to differentiate along the osteogenic lineage in vitro as measured by ALP, collagen-I and osteopontin expressions, MSCs seeded on porous hydroxylapatite scaffolds  and implanted subcutaneously in nude mice did not always correlate with in vivo bone  164  formation [381. Thus, the findings in this and other studies highlight the fact that signals that affect differentiation in vivo remain largely indeterminate and cannot be easily replicated in vitro.  Even though MSC differentiation along the osteogenic, chondrogenic and adipogenic lineages, were found to be suppressed under in vivo compared to the in vitro environment, the CFU-F content of the MSCs was found to be comparable in both conditions (Figure 3.10). Therefore, the self-renewal potential of MSC cultures was not significantly modulated by in vivo implantation.  165  3.5 Conclusion A versatile and simple scaffold fabrication process termed microwave vacuum drying was used to produce porous gelatin-alginate scaffolds for tissue engineering applications. The microwave vacuum drying technique demonstrated many advantages including rapid processing time, elimination of toxic organic solvent, and avoidance of high temperature, among others. The total porosity of the fabricated scaffolds was controlled by changing the cross-link density but the pore size and pore size distribution was not affected. Although highly porous, the scaffold had relatively small pores and limited interconnectivity that might have affected MSC migration into the interior region of the scaffold. The porous gelatin alginate  scaffold  demonstrated  excellent  biocompatibility  with  angiogenesis  and  neovascularization on the surfaces and was bioresorbed in vivo, the extent of bioresorption depending upon the cross-link density. MSCs were able to attach and proliferate at the same rate on the scaffolds, and the self-renewal potential of MSC cultures was similar during in vitro culture and in vivo implantation. However, the subcutaneous microenvironment was found to suppress MSC differentiation along all lineages tested compared to in vitro conditions. 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Cell 2008 Oct 17;135(2):240-249.  171  Chapter 4: Fusidic acid loaded PLGA and PHBV microspheres for bone infections: Formulations development and solid-state 3 characterization  4.1 Introduction Even with today’s advancements in surgical techniques, ultra-clean operation rooms and development of new antibiotics, orthopaedic surgeries can still be complicated by infections [1]. Thus, the treatment [2-5] and prevention [6-9] of orthopaedic infections have utilized prolonged systemic antibiotic therapy. However, serious problems can arise from this approach, including a failure to produce therapeutic tissue concentrations of the antibiotics because of relatively low vascularity within necrotic bone and implant in prosthetic joint infections and potential for development of bacterial resistance and deep-seated mycoses [3, 101. Local delivery of antibiotics offers significant advantages in the management and prevention of orthopaedic infections [11]. Fusidic acid (FA) has been available since the 1960s [12] and is most active against Staphylococcus aureus, Staphylococcus epidermidis, and coagulase-negative staphylococci, including strains that are methicillin-resistant that commonly cause prosthetic joint infections [13]. However, systemic delivery of FA can lead to many serious toxicities [14-16], thus necessitating its localized delivery.  By direct application of antibiotics to the site of infection or potential infection, it is possible to achieve higher tissue levels and for a longer period of time, while simultaneously avoiding systemic side effects. These high local doses of antibiotics facilitate delivery by diffusion to  A version of this chapter has been published. Yang C et al. (2009). PLGA and PHBV Microsphere Formulations and Solid-State Characterization: Possible Implications for Local Delivery of Fusidic Acid for the Treatment and Prevention of Orthopaedic Infections. Pharm Res 26(7): 1644-1656  172  avascular areas of the wound that are inaccessible by systemic antibiotics, which often can only be delivered in concentrations that may result in resistance [17]. Particularly, the use of PLGA [18-22] and PHBV [23, 24] microspheres to locally deliver a variety of antibiotics for the treatment of bone infections has been demonstrated.  Although localized and controlled delivery of FA has been proposed to treat and prevent bone infection in orthopaedic medicine, the literature is limited. There are only a few reports of the local administration of FA, which include the studies of FA released from nonbiodegradable poly(methylmethacrylate) (PMMA) bone cement [2, 25, 26], bioresorbable plaster of Paris (calcium sulphate hemihydrate) beads [27-29], and delivery of sodium fusidate (sodium salt of FA) from PLGA microspheres [30].  Therefore, in this chapter, we investigated the development and characterization of FA loaded PLGA and PHBV microspheres using a factorial design approach to identify critical and interacting formulation factors. We show that encapsulation of FA in PLGA (but not PHBV) microspheres results in an interesting phase separation phenomenon of FA-rich spherical domains throughout the microsphere matrix and surface. From real time recordings of single microdroplets produced and positioned using micromanipulation video microscopy, this was attributed to a phase separation and coalescence of liquid phase FA-rich microdroplets produced within the microsphere during solvent evaporation and their exclusion from the phase-separated PLGA matrix upon hardening.  173  4.2  4.2.1  Materials and methods  Chemicals  Fusidic acid (FA, M. W. 516.709 g/mol), poly(3-hydroxybutyric acid-co-3-hydroxyvaleric acid) (PHBV, 5 and 12 wt % hydroxyvaleric acid, HV), poly(vinyl alcohol) (PVA, 98% hydrolyzed, Mw 13  —  23 g/mol), dichioromethane (DCM), acetonitrile (ACN), chloroform,  methanol, phosphoric acid, and the different reagents needed to prepare phosphate buffered saline (PBS, pH 7.4) solution were all obtained from Sigma-Aldrich (Oakville, ON, CA). Poly(DL-lactic-co-glycolic acid) (PLGA, 85/15 LA/GA) with an intrinsic viscosity of 0.61 dL/g in chloroform (equivalent to Mw  -  86,000 g/mol) was obtained from Birmingham  Polymers Inc. (Birmingham, AL, USA) and poly(DL-lactic-co-glycolic acid) (PLGA, 50/50 LA/GA) with intrinsic viscosity of 0.58 dL/g in hexafluoroisopropanol (equivalent to Mw 84,000 g/mol) was obtained from Lactel® Absorbable Polymers (Peiham, AL, USA). Poly(L -lactic acid) (PLLA, Mw 100,000 g/mol) was obtained from Polysciences Inc (Warrington, PA, USA).  4.2.2  Factorial design for formulation of FA-loaded PLGA and PHBV microspheres  To develop FA-loaded PLGA microsphere formulations, a 2 full factorial design with three factors at two input levels was utilized to produce eight different formulations. The three factors chosen for investigation were initial drug loading (10 and 30% w/w), PLGA composition (50/50 and 85/15), and polymer and drug concentration (5 and 10% w/v). Similarly, FA-loaded PHBV microsphere formulations were developed using a 2 full  174  factorial design where the three factors chosen for investigation were initial drug loading (10 and 30% w/w), PHBV composition (5 and 12 wt % HV), and polymer and drug concentration (5 and 10% w/v). The two direct measured results of the FA-loaded PLGA and PHBV microsphere formulations were mean particle diameter and encapsulation efficiency of FA. The design matrices with all three factors and measured responses for FA-loaded PLGA and PHBV microsphere formulation are shown in Table 4.1 and 4.2, respectively.  4.2.3  Fabrication of FA-loaded PLGA and PHBV microspheres  FA encapsulated micro spheres were synthesized by the solvent evaporation method [31]. FA and PLGA were dissolved in DCM according to the factorial design parameters (Table 4.1), and added drop-wise into 100 mL of 2.5% (w/v) PVA solution with an overhead propeller stirring at 600 rpm to disperse the organic phase. The dispersion was stirred continuously for 2.5 h at room temperature under the fume hood to evaporate the organic solvent. The FA loaded PLGA microspheres were collected by centrifuging at 3000 rpm for 5 mm  and  subsequently washed four times with distilled water. The microspheres were then vacuum dried at room temperature and stored in a desiccator for further analysis. FA-loaded PHBV microspheres were fabricated using the same procedures described above according to the factorial design parameters listed in Table 4.2.  To examine the effects of initial drug loading on the microsphere formulation, FA-loaded PLGA (85/15) microspheres with initial FA loading between 0  —  50% (w/w), and at a  polymer and drug concentration of 10% (w/v) were fabricated in the same manner as described above. In addition, the effects of different polymers on the microsphere  175  formulation were also studied using 30% (w/v) FA-loaded PLLA microspheres prepared at a polymer and drug concentration of 10% (w/v) as above.  4.2.4  Casting of FA and PLGA films  Films containing varying weight % of FA and PLGA were solution cast at a concentration of 10% (w/v) on Teflon® templates applied to glass slides. FA and PLGA were dissolved in DCM and the weight % of FA to PLGA (w/w) solutions were: 0.1, 0.5, 1, 2, 10, 20, 30, 80, 90, 98, 99, 99.5 and 99.9% (w/w) FA.  4.2.5  Preparation of amorphous FA drug  Amorphous FA was prepared by dissolving the drug in DCM in a glass vial at a concentration of 100 mg/mL followed by rapid removal of the solvent at 100 °C in an oil bath under a stream of nitrogen gas. Subsequently, amorphous FA was dried for 6 days under a 25 inl-Ig vacuum at ambient temperature until the sample weight was constant.  4.2.6  Microsphere particle size determination  The mean particle size and size distributions of FA-loaded PLGA and PHBV microspheres were determined using a Malvern Mastersizer 2000 (Malvern Inc., Malvem, Worcestershire, UK), laser diffraction particle size analyzer. Briefly,  5  —  10 mg of micro spheres were  suspended in 5 mL of distilled water with two drops of 1% polysorbate 80 (Tween 80) and sonicated for 2 mm to prevent aggregation of microspheres.  176  4.2.7  FA encapsulation efficiency  To determine FA encapsulation efficiency in the microsphere formulations, 5 mg of microspheres were dissolved in 1 mL of acetonitrile (ACN) or chloroform, and then 5 mL of PBS (pH 7.4) was added to precipitate the polymer. Subsequently, the sample was centrifuged at 3000 rpm for 5 mm to spin down the precipitated polymer. The organic phase of the solution (ACN or chloroform) was further filtered with 0.45 jim PTFE syringe filter prior to HPLC (Waters® Millennium System) analysis that utilized a mobile phase of 50/30/20 (v/v/v) ACN/methanol/0.O1M phosphoric acid solution, flowing at 1 ml/min through a C 18 reverse phase Novapak column (Waters®), with a 20ji1 sample injection volume, and detection 2 at 235 nm. FA content was quantified against a standard curve prepared by dissolving FA in ACN over a range of 0.01 to 1.0 mg/mL.  4.2.8  In vitro FA release from PLGA and PHBV microspheres  In vitro FA release from PLGA and PHBV microsphere formulations was carried out in PBS (pH 7.4) at 37 °C. For release studies, 5 mg of FA-loaded microspheres were placed into 15 mL of PBS and the samples were tumbled end-over-end at 10 rpm in a thermostatically controlled oven at 37 °C. At specified time points, the sample tubes were centrifuged at 3000 rpm for 5 mm, 5 mL of samples were then withdrawn for HPLC analysis to determine the amount of drug released. The remaining medium was removed and replaced with fresh PBS (pH 7.4) to maintain sink conditions. Concentrations of FA in the release medium were measured directly using the above JEIPLC method.  177  4.2.9  Scanning electron microscope (SEM), backscattering SEM (BSEM), and laser confocal microscope analyses  The morphologies and structures of the FA-loaded microsphere formulations and cast films were characterized using a combination of scanning electron microscope (SEM), backscattering SEM (BSEM), and 3D laser confocal microscope. For SEM analysis, samples were sputter-coated with a layer of 60:40 alloy of gold:palladium using a Denton Vacuum Desk II sputter-coater (Moorestown, NJ) at 50 millitorr. SEM images were then captured using a Hitachi S-3000N system (Tokyo, Japan) scanning at 10 —20 keV. To avoid obscuring the fine detailed surface characteristics of the FA-loaded PLGA and PHBV microspheres, uncoated samples were analyzed using either a Hitachi S-4700 Field Emission Scanning Electron Microscope (FESEM, Tokyo, Japan) operated at 1 keV to produce BSEM images or a Keyence VK-9700 3D laser confocal microscope that employed two light sources: a short waveform laser light and a white light source.  4.2.10 Raman spectroscopy High spatial resolution Raman spectroscopy surface mapping analyses of FA-loaded PLGA, PHBV and PLLA microsphere formulations were kindly performed by Dr. Tim Smith of Renishaw, plc (Wotton-under-Edge, UK). Specifically, Raman spectra were obtained on a Renishaw RIvllOO confocal Raman Microscope (Renishaw, plc), recorded at a spatial resolution of 1  —  3 tm on a 62  —  157 tm x 69  —  163 im image area producing up to 3968  Raman mapped spectra as the laser scanned the analysis area. Images were subsequently created using the component method (using FA and polymer reference spectra) and coloured  178  images were generated from StreamLineTM images of Anadin Extra tablet as reference with argon ion laser excitation at X = 785 nm.  4.2.11 X-ray powder diffraction (XRPD) X-ray powder diffraction (XRPD) patterns of FA and FA-loaded PLGA and PHBV microsphere formulations were acquired using a Bruker D8 Advance (Madison, WI) diffractometer in Bragg-Brentano configuration with a Cu source at 25°C. Samples were scanned from 2  —  50° 29, using a step size of 0.020°, and a step time of 1 second per step.  Approximately 200  —  300 mg of sample was packed onto a standard Bruker sample holder  with sample spinning during data acquisition to avoid preferential orientation of sample.  4.2.12 Differential scanning calorimetry (DSC) DSC analyses of the FA-loaded PLGA and PHBV microspheres as well as amorphous FA drug were performed on a TA Instruments DSC Ql00 (New Castle, DE, USA) with liquid nitrogen cooling system. Accurately weighed samples  (--‘  2  —  5 mg) were hermetically sealed  in aluminum pans and heated from 25°C to 250°C at a rate of 10°C/mm under nitrogen flow. The initial heat scan was followed by a rapid quench cooling scan from 25 0°C to -80°C at a rate of 35°C/mm and then a second heating scan from -80 °C to 250°C at a rate of 10°C/mm. For FA-loaded PLGA microspheres, the peak temperature of the first endothermic transition in the first heating cycle was recorded as the temperature at which enthalpy relaxation occurred (Tr), while the glass transition temperature (Tg) was taken as the midpoint of the heat capacity change in the second heating cycle and was clearly distinguishable from the  179  enthalpy relaxation. Since PHBV does not possess an enthalpy relaxation, Tg was taken as the midpoint of the heat capacity change in the first heating cycle for FA-loaded PHBV microspheres. In addition, the double endothermic transitions (melt-recrystallization remelting) of PHBV [32, 33] were recorded in the first heating cycle. For amorphous FA drug, Tg was taken as the midpoint of the heat capacity change in the first heating cycle.  4.2.13 Micromanipulation and video imaging of microsphere formation Real-time recordings of the formation of single FA-loaded PLGA microspheres were captured for the 30% (w/w) FA-loaded PLGA (85/15) formulation at a polymer and drug concentration of 10% (w/v). Briefly, drug and polymer solution in DCM was formed at the tip of a 5 im diameter borosilicate glass micropipette, in a solution of 0.01 M SDS solution contained in a customized design glass chamber placed under a conventional inverted light microscope with 60x oil immersion objective, connected to a CCD camera, monitor and video recorder [34]. Once a single droplet of the drug and polymer solution was formed at the tip of the micropipette, it was held there by gentle suction pressure, allowing the DCM to “evaporate”, (i.e. to dissolve from the droplet into the aqueous phase), as it would in the normal bulk-suspension microsphere fabrication process. Control PLGA and FA-loaded PHBV microsphere formations and their solidification processes were similarly video imaged as above.  180  4.2.14 Data analysis For the factorial design studies, statistical analyses were performed using a factorial design software package, Design-Expert® (Version 7.1.1, Stat-Ease, Inc., Minneapolis, MN). This software performs an analysis of variance (ANOVA) along with post-ANOVA analyses which include examination of the normal probability plot of the studentized residuals for normality of residuals and the Box-Cox plot for power transformations. Level of significance was considered to be p  <  0.05. The results of the ANOVA table for the factorial design  experiments can be found in Appendix C. For all other statistical analyses, the Student’s t test (unpaired, two-tailed) in GraphPad Prism® version 5.00 for Windows (GraphPad Software, San Diego, CA, USA) was used to determine level of significance.  181  4.3  4.3.1  Results  Factorial design of FA-loaded PLGA and PHBV microspheres  To investigate FA-loaded PLGA and PHBV microsphere formulations, full factorial designs were used to determine the effects of three factors: initial drug loading, polymer composition, and polymer and drug concentration on the resulting mean diameter and encapsulation efficiency of the microsphere formulations. Tables 4.1 and 4.2 summarize the two 2 full factorial design matrices of the three factors examined along with the measured mean diameter and encapsulation efficiency for each PLGA and PHBV microsphere formulation (run), respectively.  Out of the three factors investigated for the FA-loaded PLGA micro spheres, only the increase in polymer and drug concentration from 5 to 10% (w/v) had an effect and resulted in a statistically significant increase in the mean diameter of the microspheres, Figure 4. 1A. The encapsulation efficiency of FA in PLGA microspheres was significantly affected by both the change in the polymer and drug concentration, and the change in initial drug loading. As the polymer and drug concentration increased from 5 to 10% (w/v), the encapsulation efficiency of FA decreased significantly in PLGA microspheres, Figure 4.1B. An increase in drug loading from 10 to 30% (w/w) in the PLGA microspheres produced a significant increase in FA encapsulation efficiency, Figure 4.1 C. The change in polymer composition of PLGA from 50/50 to 85/15 (LA/GA) had no effect on the mean diameter or encapsulation efficiency of the FA-loaded PLGA microsphere formulations.  182  Table 4.1: 2 full factorial design matrix and measured responses of FA-loaded PLGA microspheres  :  Run  Factor 1: Factor3: Drug Loading Factor 2: Polymer and Drug a Polymer Compositi Concentration (% wlv) (% wlw) b  1 10 PLGA 50/50 2 30 PLGA50/50’ PLGA85/15c 3 10 PLGA85/15c 4 30 10 5 PLGA5O/50” PLGA5O/50b 6 30 7 10 PLGA85/15’ PLGA85/15c 8 30 % wiw weight % of FA relative to polymer b PLGA with monomer molar ratio of 50/50 lactic to glycolic acid C PLGA with monomer molar ratio of 85/15 lactic to glycolic acid  5 5 5 5 10 10 10 10  Response 1: Mean Diameter (jm) 52 59 62 52 89 95 108 103  Response 2: Encapsulation Efficiency (%) 82 99 95 98 75 92 79 90  Table 4.2: 2 full factorial design matrix and measured responses of FA-loaded PHBV microspheres  Run 1  00  Factor I Factor 3 Drug Loading Factor 2: Polymer and Drug a Polymer Composition Concentration (% w!v) (% wlw)  PHBV5b 10 PHBV5b 2 30 PHBVI2C 3 10 PHBV12C 4 30 5 10 PHBV5’ PHBV5L 6 30 PHBV12C 7 10 PHBV12C 8 30 % w/w = weight % of FA relative to polymer b PHBV with 5% wt of hydroxyvalerate (HV) in the copolymer PHBV with 12% wt of hydroxyvalerate (HV) in the copolymer ‘  5 5 5 5 10 10 10 10  Response I Mean Diameter (rim) 70 76 95 63 144 141 148 138  Response 2 Encapsulation Effic,ency(%) 104 126 116 107 143 103 104 100  A ion  B  -  C  100.0—  100.0.  93.8-  93.0.  S .5  :s S  °-  Lii  t LII  07.5—  I  87.0.  .5  62.  47.  75.0— 5.00  6.2.5  I 7.50  I 0.75  Polymer and Drug Concentration (% wlv)  10.00  75.0. 5.00  6.20  7.00  I 0.75  Polymer and Drug Concentration (% w/v)  I 10.00  I 10.0  I 15.0  I 20.0  20.0  Drug Loading (% wlw)  Figure 4.1: Formulation factors that influence FA-loaded PLGA microspheres. (A) Effects of polymer and drug concentration on mean diameter, p <0.0001, (B) effects of polymer and drug concentration on the encapsulation efficiency of FA, p < 0.01, and (C) effects of initial drug loading on encapsulation efficiency of FA, p <0.01. Results are expressed as mean ± 95% CI.  30.0  For the FA-loaded PHBV microspheres, only the increased polymer and drug concentration from 5 to 10% (w/v) had an effect and significantly increased the mean diameter of the microspheres, Figure 4.2. Drug loading and polymer composition had no effects on the mean diameter. Moreover, FA encapsulation efficiency in the PHBV microspheres was not affected by any of the three factors investigated.  156—  133  —  2 ) ‘:1)  4-.  2 (U ci  110—  (U G) 86  63  —  —  I  I  I  I  I  5.00  6.25  7.50  8.75  10.00  Polymer and Drug Concentration (% wfv) Figure 4.2: The effect of polymer and drug concentration on the mean diameter of FA loaded PHBV microspheres, p < 0.0001. Results are expressed as mean ± 95% CI.  186  4.3.2  Surface characterization studies  SEM images revealed the detailed surface morphologies of the FA-loaded PLGA microsphere formulations (Figure 4.3). All FA-loaded PLGA microspheres showed spherical and relatively uniform protrusions (bumps) on their surfaces regardless of the changes in the three formulation factors. The only difference observed was the increase in size and height of the protrusion on the surfaces with an increase in initial FA loading. This is clearly demonstrated in Figure 4.4 with FA-loaded PLGA (85/15) microspheres and FA loading up to 50% (w/w). On the other hand, the surfaces of control (no drug) PLGA microspheres were found to be smooth (Figure 4.4A).  FA-loaded PHBV microspheres surface morphologies were also affected by initial FA loading, but were quite different in appearance to FA-loaded PLGA microspheres, with recessed spherical dimples on the surfaces of all 30% (w/w) FA-loaded formulations (Figure 4.5). The surfaces of control (no drug) PHBV microspheres were relatively smooth, although not as smooth as control PLGA microspheres. As the FA loading increased beyond 20% (w/w) in the PHBV microspheres, the recessed spherical dimples appeared to increase in size (Figure 4.6).  Fine surface morphological details of the FA-loaded PLGA and PHBV microspheres were obtained with the use of laser confocal and backscattering SEM (BSEM) analyses where there are no requirements for a coating layer and are shown in Figures 4.7 and 4.8, respectively. The spherical protrusions on the PLGA microsphere surfaces appeared to possess distinct boundaries (Figure 4.7A and 4.7B). In addition, the spherical microdomains were found throughout the entire PLGA matrix as demonstrated in the microsphere cross-  187  section in Figure 4.7C, although these inclusions were not as large as the ones at the surface. BSEM images of FA-loaded PHBV microspheres (Figure 4.8B and 4.8C), revealed cracked, rough and pitted surfaces.  188  Polymer and Drug Concentration (5% wlv) PLGA (50150)  PLGA (85115)  Polymer and Drug Concentration (10% wlv) PLGA (50150)  PLGA (85115)  10% FA,  00  30% FA  Figure 4.3: SEM images illustrating the surface morphologies of the eight FA-loaded PLGA microsphere formulations described in Table 4.1.  O%FA  30% FA  10% FA  40% FA  20% FA  50% FA  Figure 4.4: SEM images illustrating the effects of different drug loading on FA-loaded PLGA (85/15) microsphere surface morphologies. All micro sphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w).  Concentration (5% wlv) PHBV (5% HV)  PHBV (12% HV)  Concentration (10% wlv) PHBV (5% HV)  PHBV (12% HV)  10% FA  (Run #8)  30% FA,  Figure 4.5: SEM images illustrating the surface morphologies of the eight FA-loaded PHBV microsphere formulations described in Table 4.2.  0% FA in PHBV  10% FA in PHBV  20% FA in PHBV  30% FA in PHBV  Figure 4.6: SEM images illustrating the effects of different drug loading on FA-loaded PHBV (12% HV) microsphere surface morphologies. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w).  192  —.-.-.-.-.-.-.-  04700 05kv 25  0 SE(L 4/2705  Figure 4.7: Detailed surface and interior morphologies of 30% (w/w) FA loaded PLGA (85/15) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) Laser confocal microscope images of PLGA microsphere, (B) BSEM images of PLGA microsphere, and (C) BSEM images of sectioned PLGA microsphere.  193  Figure 4.8: Detailed surface morphologies of 30% (w/w) FA loaded PHBV (12% HV) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A C) BSEM images of PHBV microsphere at different magnifications. —  194  4.3.3  Phase separation of FA in PLGA microspheres  The existence of phase separated regions in the FA-loaded PLGA microspheres was confirmed and identified chemically by Raman spectroscopy surface mapping analyses (Figure 4.9). The PLGA polymer-rich regions (Figure 4.9B, shown in green), and FA-rich regions (Figure 4.9C, shown in red) could be clearly identified, and upon merging of the two images, it was evident that the spherical protrusions were a FA-rich microdomain phase distributed throughout the PLGA-rich matrix of the microsphere (Figure 4.9D). SEM images of 10, 20 and 30% (w/w) FA-loaded PLGA microspheres before and after 7 days of drug release in PBS (pH 7.4) shown in Figure 4.10, also support the Raman findings that the spherical protrusions on the surfaces of the microspheres were primarily composed of FA, since the protrusions were eliminated following 7 days of drug release (i.e. the protruded FA microdomains have dissolved off and formed depressions on the surface).  Real-time video images of a single FA-loaded PLGA microsphere are illustrated in Figure 4.11 (shown as time separated screen captures, with the full length video available upon request to the author of this thesis), and demonstrate the phase separation process on the surface and within the interior of the forming microsphere (for scale, screen height is ‘25 tim). As DCM solvent evaporates from the initial 4.11A, time  =  40 !lm microsphere droplet (Figure  0 sec), the overall size of the microsphere droplet begins to decrease (Figure  4.1 1B, time = 3 see). Further DCM evaporation leads to numerous phase-separated FA liquid microdroplets being formed throughout the microsphere (Figure 4.11 C, time  =  5 see) and  shortly after, there was evidence of extensive coalescence of these FA-rich liquid phase microdroplets (Figure 4.11D, time  =  8 see) to form larger phase-separated FA-rich  195  microdomains (Figure 4.11 E, time  =  9 sec) that are clearly visible within the bulk and also on  the surface of the microsphere as spherical protrusions at the end of the solidification process (Figure 4.11F, time= 13 see).  Raman spectroseopy surface mapping of the FA-loaded PHBV microsphere (Figure 4.12A), showed that the PHBV polymer (Figure 4.12B, illustrated in green), and FA drug (Figure 4.1 2C, illustrated in red) were both distributed uniformly throughout the entire microsphere with no apparent phase separation of FA from the polymer matrix. Real-time video recordings were also made of single FA-loaded PHBV microspheres and showed no phase separation of FA consistent with the Raman spectroscopy mapping (data not shown).  196  B  A  -20  0  20  40  60  00  C  D  tO  20  30  40  50  60  Figure 4.9: Raman spectroscopy images of FA distribution in 30% (w/w) FA-loaded PLGA (85/15) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light microsphere montage, 50x magnification, (B) distribution of PLGA rich regions (green) across microsphere, (C) distribution of FA-rich regions (red) across microsphere, and (D) combined distribution of the two regions across microsphere.  0% FA in PLGA  10% FA in PLGA  20% FA in PLGA  30% FAin PLGA  After 7 days of drug release  Go  0% FA in PLGA  10% FA in PLGA  20% FA in PLGA  30% FA in PLGA  Figure 4.10: SEM images illustrating the effects of different drug loading on FA-loaded PLGA (85/15) microsphere surface morphologies (A D) and the changes after 7 days of drug release (E H). All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w). —  —  Time  Time  =  =  0 sec  8 sec  Time  =  3 sec  Time  =  9 sec  Time  Time  =  =  5 sec  13 sec  Figure 4.11: Time lapsed video images illustrating the formation of a single 30% (w/v) FA loaded PLGA (85/15) microsphere with an initial polymer and drug concentration of 10% (w/v). The initial FA/PLGA/DCM droplet was blown from a micropipette into 0.O1M SDS aqueous solution at room temperature. Arrows illustrates the phase separated FA-rich microdomains, while star indicate the micropipette. The full video showing the FA phase separation phenomenon and microsphere solidification process is available upon request to the author of this thesis. [Note: for scale, screen height is -25 urn].  B  A  C  C C  -  5Op  5flp  : 5op  I  Figure 4.12: Raman spectroscopy images of FA distribution in 30% (w/w) FA loaded PHBV (12% HV) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light microsphere montage, 20x magnification, (B) distribution of PHBV (green) across microsphere, and (C) distribution of FA (red) across microsphere.  4.3.4  Miscibility and phase separation of FA and PLGA in solvent-cast films  We evaluated FA and PLGA miscibility characteristics and the phase separation phenomenon in cast films where drug loading can be completely and accurately controlled. It was shown that the miscibility limit for FA in PLGA was approximately 1% (w/w) (Figure 4.1 3A). At FA loadings of 2% (w/w) and above, the films were phase-separated (Figure 4.1 3B). At 30% (w/w) FA loading, the distinctive spherical microdomains were formed within the PLGA matrix (Figure 4.13C). At the other extreme of the two component phase diagram, even at very low concentrations (0.1% w/w) PLGA was not miscible in FA and the two components were completely phase separated (data not shown).  201  1%FA  2%FA  30% FA  Figure 4.13: SEM images of different weight % FA in PLGA (85/15) films solvent-cast from DCM illustrating the miscibility characteristics of FA and PLGA. All films were cast at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w).  4.3.5  XRPD characterization of FA-loaded PLGA and PHBV microspheres  To determine whether phase separated, FA-rich microdomains were crystalline in nature, XRPD was carried out and results are shown in Figure 4.14. Even at the highest drug loading of 30% (w/w) FA in the PLGA microspheres, there was no evidence of crystallinity in the sample, illustrated by the characteristic amorphous halo in the XRPD diffraction pattern (Figure 4.14B). All other FA-loaded PLGA microspheres produced similar amorphous x-ray patterns. Only the crystalline diffraction pattern of the PHBV polymer was obtained for the FA-loaded PHBV microspheres, with no evidence of crystallinity of FA in the microspheres (Figure 4.15).  203  500  400  -300  200  100  0 10  20  30  2-Theta Scale Figure 4.14: XRPD patterns of solid-state (A) FA as received, (B) 30% (w/w) FA loaded PLGA (85/15) microspheres, and (C) control (no drug) PLGA (85/15) microspheres. All microspheres were manufactured at a polymer and drug concentration of 10% (w/v). -  800  D  700  600  C .500  400  B  300  200  A  100  0 2  10  20  30  40  2-Theta Scale -  Figure 4.15: XRPD patterns of solid-state (A) FA as received, (B) PHBV (12% HV) polymer as received, (C) control (no drug) PHBV (85/15) microspheres, and (D) 30% (w/w) FA loaded PHBV (12% HV) microspheres. All microspheres were manufactured at a polymer and drug concentration of 10% (w/v).  5(  4.3.6  Thermal analysis of FA-loaded PLGA and PHBV microspheres  DSC scans of FA-loaded PLGA microspheres showed an enthalpy relaxation endotherm (Figure 4.1 6A) and Tg (Figure 4.1 6B) for PLGA at  46  —  50 °C. A Tg for FA at  117  —  118°C (Figure 4.1 6A) was obtained for microspheres with greater than 20% (w/w) FA loading that correlated with the Tg of the pure FA drug in the amorphous state shown in Figure 4.17. DSC analyses of FA-loaded PHBV microspheres revealed a Tg for PHBV around 57  —  59°C and double melting endotherms in the range of 133  —  152°C (Figure 4.18)  corresponding to PHBV’ s melting-recrystallization-remelting process upon heating [32, 33]. All the thermal events for FA-loaded PLGA and PHBV microspheres are summarized in Tables 4.3 and 4.4, respectively. The presence of FA was found to increase the enthalpy relaxation temperature, Tr and the Tg of PLGA (Table 4.3), whereas in FA-loaded PHBV microspheres, the presence of FA had no effect on Tg, but decreased the melting, Tmi and remelting, Tm 2 of PHBV polymer (Table 4.4).  206  u_  u_  a’  a’  I  C  E,o Up  Temperature (°C)  Exo Up  Temperature (C)  Figure 4.16: DSC thermograms of(A) first heating cycle of FA loaded PLGA (85/15) microspheres illustrating the enthalpy relaxation temperature, T r of the PLGA polymer and the glass transition temperature, Tg of FA, (B) second heating cycle (after quench cooled) of FA loaded PLGA (85/15) microspheres showing the Tg of the PLGA polymer. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w).  U CU  a) I  -0.5  GO  Exo Up  Temperature (C)  Figure 4.17: DSC thermogram of pure amorphous FA drug illustrating the glass transition temperature, Tg.  0 U 4-.  1)  Exo Up  Temperature (CC)  Figure 4.18: DSC thermograms of FA loaded PHBV (12% HV) microspheres illustrating the glass transition temperature, Tg and double melting temperature, Tmi and Tm2 of the PHBV polymer. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and % FA loadings were relative to polymer (w/w).  Table 4.3: Summary of the thermal properties of FA and FA-loaded PLGA (85/15) microspheres obtained using DSC  Polymer Transitions  .  Formulations 0% FA loaded PLGA (85/1 5) 10% FA loaded PLGA (85/15) 20% FAloaded PLGA(85/15) 30% FAloaded PLGA(85/15) a  AHr (Jig) 8.8 ± 0.5 8.9 ± 0.4 8.2±0.4 7.8±0.1  b  T (°C) C AC Ji(g C) 45.9 ± 1.2 0.51 ± 0.06 49.9 ± 0.1# 0.53 ± 0.03 49.6± 1.1 0.52±0.03 48.1 ±0.7# 0.49±0.02  T (°C)  AC Ji(g °C)  --  --  --  118.4±0.2 116.8±0.7  --  0.02±0.01 0.03±0.01  Tr = enthalpy relaxation temperature. The peak temperature of the first endothermic transition in the first heating cycle = enthalpy relaxation. The first endothermic transition in the first heating cycle Tg = glass transition temperature. The midpoint of the heat capacity change in the second heating cycle for FA-loaded PLGA microspheres and first heating cycle for FA-loaded PHBV microspheres. * p < 0.005 compared to 0% FA loaded PLGA (85/15) <0.05 compared to 0% FA loaded PLGA (85/15) Results are expressed as mean ± SD of 3 separate DSC scans of different samples b  C  Tr (°C) a 46.6 ± 0.6 50.1 ± 0.3* 50.9±0.9* 51.0±0.3*  Fusidic Acid Transitions  Table 4.4: Summary of the thermal properties of FA and FA-loaded PHBV (12% HV) microspheres obtained using DSC •:.  Polymer Transitions  -  Formulations 0% FA loaded PHBV 10% FA loaded PHBV 20% FA loaded PHBV 30% FA loaded PHBV  a Tg  Tg (°C) a 59.0 ± 1.6 57.2 ± 0.8 57.1 ± 2.1 57.3 ± 2.0  JI(g C) 0.31 ± 0.09 0.49 ± 0.08 0.44 ± 0.05 0.45 ± 0.07  b  Tmi (°C) 139.8 ± 0.9 136.4 ± 0.9* 133.8 ± 0.8* 134.1 ± 0.2*  1 (Jig) AHm 10 ± 3 13 ± 2 5± 1 13 ± 3  C  d  Tm (°C) 2 152.8 ± 0.7 150.1 ± 0.9# 147.3 ± 0.5# 147.2 ± 0.3#  AHm (Jig) 2 8± 1 10 ± 1 2± 1 9±2  e  T (°C)  AC Ji(g °C)  --  --  --  --  --  --  --  --  glass transition temperature. The midpoint of the heat capacity change in the first heating cycle. first melting temperature. The peak temperature of the first endothermic transition in the first heating cycle 1 enthalpy of melting. The first endothermic transition in the first heating cycle ARm d Tm2 remelting temperature after recrystallization. The peak temperature of the second endothermic transition in the first heating cycle 2 = enthalpy of melting after recrystallization. The second endothermic transition in the first heating cycle AHm * p < 0.00 icompared to 0% FA loaded PHBV p < 0.05 compared to 0% FA loaded PHBV Results are expressed as mean ± SD of 3 separate DSC scans of different samples b  -‘  Fusidic Acid Transitions  .  -  .  =  =  4.3.7  Drug release profiles of FA-loaded PLGA and PHBV microspheres  FA-loaded PLGA and PHBV microsphere drug release profiles were determined in PBS (pH 7.4) and are shown separately in groups according to initial drug loading, 10 and 30% (w/w) FA loading in Figure 4.19 to Figure 4.22. Overall, cumulative release of FA from all PLGA microspheres demonstrated similar biphasic release profiles with a rapid burst phase followed by a phase of slow controlled release over 21 days (Figure 4.19 and 4.20). The formulation factors influencing drug release were the increase in FA loading (from 10 to 30%) and decreased PLGA hydrophobicity (LAJGA ratio decreased), both of which produced a larger burst phase of release, leading to higher overall FA released.  The cumulative release of FA from PHBV microsphere formulations also demonstrated biphasic release profiles with a rapid burst phase followed by a phase of slow controlled release over 21 days (Figure 4.21 and 4.22). Higher initial FA loading (10 to 30%) produced a larger initial burst release of FA leading to higher cumulative released. The change in polymer composition had no effect on the release profiles of FA from PHBV microspheres, but as the polymer and drug concentration increased from 5 to 10% (w/v), the initial burst release of FA was reduced regardless of the initial FA loading.  212  10% FA loaded PLGA microspheres  I  Run -*- Run -- Run -4- Run  0.0  2.5  5.0  7.5  #1 #3 #5 #7  PLGA (50/50) PLGA (85/15) PLGA (50/50) PLGA (85/15)  10.0 12.5 15.0 17.5 20.0 22.5  Time (Days) Figure 4.19: FA release profiles from 10% FA loaded PLGA microsphere formulations. See Table 4.1 for the details of each formulation (Run).  213  30% FA loaded PLGA microspheres 11  Run Run -‘- Run -4-- Run  00  25  5.0  7.5  #2 #4 #6 #8  PLGA (50/50) PLGA (85/15) PLGA (50/50) PLGA (85/15)  10.0 12.5 15.0 17.5 20.0 22.5  Time (Days) Figure 4.20: FA release profiles from 30% FA loaded PLGA microsphere formulations. See Table 4.1 for the details of each formulation (Run).  214  10% FA loaded PHBV microspheres  i  --  Run #1 PHBV(5%HV) Run #3 PHBV(12% HV)  Run#5PHBV(5%HV) -4- Run #7 PHBV(12%HV) -‘-  0.0  2.5  5.0  7.5  10.0 12.5 15.0 17.5 20.0 22.5  Time (Days) Figure 4.21: FA release profiles from 10% FA loaded PHBV microsphere formulations. See Table 4.2 for the details of each formulation (Run).  215  30% FA loaded PHBV microspheres I I  --  Run #2 PHBV(5%HV) Run #4 PHBV(12%HV) Run #6 PHBV(5% HV)  -4- Run#8PHBV(12%HV)  0.0  2.5  5.0  7.5  10.0 12.5 15.0 17.5 20.0 22.5 Time (Days)  Figure 4.22: FA release profiles from 30% FA loaded PHBV microsphere formulations. See Table 4.2 for the details of each formulation (Run).  216  4.4 Discussion Fabrication of microspheres using solvent evaporation is a variable and complex process with many adjustable parameters [31, 35, 36]. An enormous amount of time and effort would be required to test and optimize all factors through “one-factor-at-a-time” experiments. Therefore, in this study a full factorial design experiment [37] was performed to systematically and efficiently determine the important and possible interacting factors in the formulation of FA-loaded PLGA and PHBV microspheres. From the factorial design experiment, higher polymer and drug concentrations used in the microsphere preparation were found to not only increase the mean diameter, but also decrease the encapsulation efficiency of the FA-loaded PLGA microspheres (Figure 4.1A and 4.1B). Increased microsphere diameters can be explained by both higher polymer/drug concentration and increased viscosity of the internal organic phase leading to more rapid solidification of the polymer [38]. It is possible that an increased number and rate of formation of FA-rich microdroplets formed within the solidifying microspheres may have led to increasing partitioning of FA out of the microsphere and into the external phase, resulting in the decreased encapsulation efficiency.  Increasing the initial drug loading produced a corresponding increase in encapsulation efficiency for PLGA microspheres (Figure 4.1 C) because FA is very poorly water soluble (FA solubility in aqueous phase, pH 5.7 was determined to be  50 ig/mL) and at higher FA  loadings, less of the drug is lost in the aqueous phase, resulting in higher encapsulation efficiency. Similarly, when formulating allopurinol, a hydrophobic drug in solid liposheres, El-Gibaly et al. found that encapsulation efficiency increased with initial drug loading and  217  attributed it to the poor water solubility of the drug and its reduced overall loss as drug loading increased [39]. Furthermore, Jalil et al. found encapsulation of phenobarbitone in poly(D,L-lactic acid) microspheres also increased with initial drug loading [40].  The increase in mean diameter of FA-loaded PHBV microsphere with increased polymer and drug concentration (Figure 4.2) was similarly due to the increased viscosity of the organic phase as observed for the FA-loaded PLGA microspheres.  A combination of SEM, BSEM, laser confocal microscope, Raman spectroscopy analysis and miscibility determination with solvent-cast films all support the observation that FA phase separates from PLGA polymers. Phase separation of FA occurred in PLGA matrices under all conditions of PLGA composition investigated, including PLGA (50/50), (85/15), and PLLA, illustrated in SEM images for all three compositions in Appendix B (Figure B-i), and Raman spectroscopy mappings for PLGA (50/50, Figure B-2), PLGA (85/15, Figure 4.9) and PLLA (Figure B-3). Additionally, phase separation of FA occurred regardless of the changes in any other formulation factors such as initial drug loading or drug and polymer concentration (Figure 4.3). Gangrade et al. found that changing the emulsifying agents (PVA to gelatin) significantly altered the micro spheres properties [41]. However, in our study, changing the emulsifying agent PVA to gelatin in the external aqueous phase during the fabrication process did not affect the phase separation of FA from PLGA (see Appendix B, Figure B-4). Increasing FA loadings from 10 to 30% (w/w) greatly exceeded the miscibility of FA in PLGA  (- 1% w/w) and produced larger FA-rich phase  218  separated microdomains that  may also contain small amounts of PLGA chain segments and residual DCM and water solvent.  The separated FA-rich phase was present throughout the PLGA matrix illustrated by microsphere sections (Figure 4.7C) and seen by real-time video recordings of single microspheres during their formation (Figure 4.11). The representative video images of the FA-loaded PLGA microsphere showed that the formation of FA-rich microdomains was driven by the coalescence of highly concentrated FA microdroplets within the liquid phase PLGA microsphere prior to final hardening. As the DCM solvent “evaporated”, FA-rich microdroplets (resolution is about 1 jim) began to phase separate out and continued DCM evaporation led to decreases in the overall microsphere size. The video clearly showed individual FA-rich microdroplets coalescing together until larger and stable microdroplets were formed throughout the solidified microsphere. Interestingly, in the final stages of solidification when most of the solvent has “evaporated”, these phase-separated FA-rich domains are seen to appear and grow at the interface, and soon form rounded protrusions as seen in the video (and SEM) images.  It is as though they are indeed being physically  excluded from the solidifying PLGA rich matrix. Similarly, the FA phase separation and coalescence phenomena were seen in video recordings for the different FA loadings (10 and 20%) in PLGA. They were not however, observed with FA-loaded PHBV microspheres, and were not present in control (no drug) PLGA microspheres (data not shown). Panyam et al [42] have previously observed phase separation of hydrophobic drug from PLGA and PLLA polymers, and suggested the result was due to the different solid-state solubility of the drug in the polymers, while Vasanthavada et al. [43, 44] also investigated the mechanism and  219  kinetics of phase separation of small molecules in polymer as a result of solid-state solubility. However, in both the Panyam et al. and Vasanthavada et al. studies, no distinctive spherical microdomains of the phase separated drug were observed. The phase separated FA-rich spherical microdomains observed throughout the PLGA matrix in this study resemble the formation of microdomains in composite (blended) polymer microspheres, where the microdomains were the results of non-equilibrium of two phase separated polymers [45). In any event, the most striking observation from the real-time video recordings of single microspheres is that the FA-rich phase, that must still contain some DCM solvent, is liquid, and so is subject to the same effects of interfacial tension, forming minimum interfacial areas and subject to coalescence if such liquid domains touch in the shrinking liquid PLGA-DCM microsphere.  Interestingly, FA was not phase separated when formulated in PHBV, which is another polymer in the family of poly(x-hydroxy acid) polymers like PLGA and PLLA since FA was found to be distributed uniformly over the microsphere surface (Figure 4.12). The subtle differences in molecular structure of the polymer chain such as the extra methyl group found in the backbone of the repeat units of PHBV compared to PLGAIPLLA polymer could be a possible explanation for FA to not undergo phase separation from the PHBV matrix. The extra methyl group might contribute to PHBV being more hydrophobic than PLGA!PLLA, thus allowing FA, a hydrophobic drug, to have greater solid-state miscibility with PHBV.  The FA-rich solid microdomains phase separated out in the PLGA microsphere were found to be amorphous due to the lack of crystalline peaks in the XRPD patterns (Figure 4.14), and  220  absence of a melt endotherm and presence of a Tg in the DSC thermograms at high FA loadings (Table 4.3). The inhibition of FA recrystallization even at 30% (w/w) FA loading could be attributed to multiple factors as follows: the constant, mechanically agitated solvent evaporation process; the presence of residual solvents (both DCM and water); the presence of a very small amount of dispersed PLGA or PLGA chain segments, and even the kinetics required for recrystallization. The absence of crystalline drug in PLGA particle formulations has also been reported for other hydrophobic drugs that phase separated from the polymer matrix [42, 46].  An FA melting event (175 °C) was not detected for all drug loadings. At higher FA loadings (20 and 30%), DSC analyses showed a Tg of FA at  116  —  118 °C (Figure 4.16A) which was  the same as the Tg of the pure amorphous FA drug, 117 °C (Figure 4.17). Together with the XRPD analyses, it seems clear that the phase separated FA microdomains in PLGA microspheres were in the amorphous state. During the microsphere formation process, polymer chains are solidified in non-equilibrium conformations that gradually relax towards equilibrium with time [47]. This relaxation process is thermodynamically driven and is known as enthalpy relaxation [48]. Enthalpy relaxation events (Figure 4.16) were observed concurrently at the Tg of the PLGA microspheres. The miscible amount of FA within the PLGA matrix of the FA-loaded microspheres may as well have acted as an anti-plasticizing agent reducing the molecular mobility (or free volume) of the PLGA chains as shown by the increase in both the Tr and Tg [49, 50]. Since up to a maximum of 1% FA was miscible in the PLGA matrix, all the FA-loaded microspheres produced the same extent of increase in enthalpy relaxation, Tr and Tg compared to control (Table 4.3). An increase in T and Tg were similarly observed for other reported PLGA and PLLA microsphere formulations in which  221  the added PVA surfactant [51], and paclitaxel drug [52], respectively, acted as antiplasticizers.  On the contrary, for the FA-loaded PHBV microspheres, the uniformly distributed FA within the solid polymer matrix had no effect on the Tg (-57  —  59 °C), but lowered the melt  temperature, Tmi (—-140 °C), and re-melt temperature, Tm 2 (—152°C) of PHBV [32]. Taken together with the absence of any Tg for FA even at 30% drug loading (w/w) (Table 4.4) and Raman spectroscopy analysis showing the uniformly distributed FA molecules throughout the PHBV matrix (Figure 4.12C), we suggest that the microspheres were composed of a single phase solid solution of FA in PHBV.  The burst phase of the biphasic release profiles of all FA-loaded PLGA microsphere formulations was likely a result of the dissolution of the phase separated FA located on the surface (Figure 4.19 and 4.20). FA-loaded PLGA 30% (w/w) microspheres formulations produced a larger burst release compared to 10% (w/w) FA-loaded formulations probably due to the increased amount of FA phase separated on the surface as shown in Figures 4.4D vs. 4.4B as larger surface protrusions. The higher burst release for the PLGA (50/50) compared to the PLGA (85/15) microsphere formulations may be attributed to the increased hydrophilicity of the PLGA (50/50) polymer matrix, allowing more water penetration and promoting rapid dissolution and transport of FA. The subsequent release after the initial burst phase was very slow for all the PLGA microsphere formulations and this was controlled by the slow diffusional release of FA through the polymer matrix. After 14 days of release, microsphere formulations with PLGA (50/50) demonstrated an increased drug release when compared to PLGA (85/15) microspheres, most likely due to the contribution by polymer  222  degradation from the bulk PLGA (50/50) which begins at slow degradation of PLGA (85/15) that occurs at  -  2  —  4 weeks compared to the  15 —20 weeks [53, 54].  FA-loaded PHBV microsphere formulations also showed a significant burst release due to FA being distributed on the surfaces as well as the cracked, rough, and possibly porous micro sphere surfaces. The smaller burst release observed for the 10% (w/v) polymer and drug concentration microsphere formulations compared to their corresponding 5% (w/v) formulations was likely due to the fact the 10% (w/v) microspheres were larger in size, thus having a smaller surface area to volume and resulting in a slower transport rate of FA from the micro sphere.  223  4.5 Conclusion In the process of formulating FA in PLGA and PHBV microspheres, we observed an interesting phase separation phenomenon of FA in PLGA but not in PHBV polymer. It was found that FA was miscible in the PLGA polymer matrix up to a maximum of 1% (w/v) and interacted with the PLGA polymer chains, as demonstrated by the increase in Tr and Tg. Above 1% (w/w), liquid microdroplets of FA phase separated from the PLGA matrix during DCM solvent removal and, driven by a coalescence behavior of these liquid DCM-FA-rich domains,  formed  distinct,  large,  completely  amorphous,  spherical  FA-rich  solid  microdomains throughout the microsphere, and on the microsphere surface.  The use of a full factorial design experiment showed that the mean diameter of FA-loaded PLGA and PHBV microspheres was only dependent on the drug and polymer concentration, while the encapsulation efficiency of FA in PLGA microsphere was affected by both the drug and polymer concentration and the initial FA loading. The biphasic drug release profiles showed an initial burst release for all FA-loaded PLGA and PHBV microsphere formulations, depending upon FA loading and polymer. 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Degradation rates of oral resorbable implants (polylactates and polyglycolates): rate modification with changes in PLA/PGA copolymer ratios. Journal of Biomedical Materials Research 1977; 11:711-719.  229  Chapter 5: Summarizing discussion, conclusions and suggestions for future work  5.1 Summarizing discussion and conclusions Orthopaedic surgeries and the delivery of therapeutic agents such as growth factors, hormones and stem cells hold tremendous promise for the treatment of ‘critical’ size bone defects found in injuries and joint arthroplasties. In addition, infections can complicate these surgical procedures, especially when implants are used, thus necessitating the administration of antibiotics. Our laboratory has developed strong collaborations with other research groups interested in the development and characterization of delivery systems for both bone regeneration and bone infection designed for revision total hip replacements, and other orthopaedic applications. Therefore, the overall goals of this thesis were to engineer, develop, and characterize biodegradable and bioresorbable polymeric microsphere and porous scaffold delivery systems for PTH and MSCs, respectively, for potential application in enhancing the innate regenerative capacity of bone. The effects of continuous and pulsatile PTH treatments on MSCs were investigated to better understand its regulatory actions on MSC differentiation, proliferation and clonogenicity in order to fully maximize the potential of using the combination of MSCs with PTH for bone regeneration. In addition, the development and characterization of biodegradable and bioresorbable polymeric microsphere delivery systems for FA for localized application in bone infection were also undertaken in this project.  230  MSCs are a convenient source of cells for use in bone tissue regeneration due to their accessibility in adult tissue of various sources, and their high content of osteoprogenitors, especially of cells displaying self-renewal ability and thus, potentially capable of continuing to replenish the osteoprogenitor compartment long term [1-3]. As a result of the successful use of PTH to enhance bone regeneration [4, 5], and studies demonstrating improved bone regeneration when growth factors were delivered along with MSCs [6-8], we hypothesized that PTH may increase the relative osteogenic progenitor content of MSC cultures to better promote bone tissue regeneration.  Surprisingly, the effects of continuous and pulsatile PTH have only been thoroughly investigated on osteogenic cells and cell lines [9-17] (also see reviews [18, 19]), but not on these complex MSC cultures. In preliminary work, we developed PTH-loaded polymeric microspheres in an attempt to deliver PTH in a controlled manner. PTH was encapsulated in PLGA and PHBV microspheres using the double emulsion solvent evaporation method. The release data showed biphasic release profiles with a pronounced burst phase over 24 hours followed by a prolonged period of slower, controlled release. It was evident from initial cell culture experiments that these formulations were not suitable for providing the precise and reproducible delivery of PTH to MSCs in either continuous or pulsatile regimens. Other research groups have also explored the development of controlled release systems to deliver PTH in either a continuous [20] or pulsatile [2 1-23] manner. However, none of these studies used primary cultures like MSCs or actually delivered PTH directly to the osteoblastic cell lines they were investigating [20-23]. Rather, in all the studies, PTH eluted from in vitro release experiments was collected and then used in separate experiments to expose the cells  231  to either continuous [20] or pulsatile [21-23] PTH regimens, in order to assess the bioactivity of the released PTH.  Therefore, as a first step to understand the effects of different regimens of PTH treatment on MSCs differentiation, proliferation and clonogenicity, studies described in this work used PTH (1-34) peptide solutions rather than PTH (1-34) peptide released from any delivery systems. The selection of PTH (1-34) over the full length PTH (1-84) was based on the following reasons. Firstly, the 1-34 PTH-fragment with the preserved N-terminus still shows similar potency and pharmacological profile as the full length hormone [24]. Secondly, PTH (1-34) is a clinically approved anabolic therapy for osteoporosis [25], and extensive studies have used PTH (1-34) to investigate its effects on osteogenic cells and cell lines [9-17] (also see reviews [18, 19]).  The design of our experiments was adapted from similar published studies [9, 26, 27]. For pulsatile administration, MSCs were exposed to PTH (10 nM) for the first 6 hours only of each 48-hour incubation cycle, by replacing the culture media after 6 hours of exposure. For continuous administration, MSCs were exposed to PTH throughout the culture experiments with an initial dose of 10 nM and an additional 10 nM dose every 24 hours. The culture media for all treatment groups (control, continuous and pulsatile) were changed every 48 hours. Replenishing PTH every 24 hours for continuous treatment was a necessary step, since PTH concentrations have been shown to decrease significantly  (-  50% in 24 h) in the  presence of cells [9], most probably due to proteolytic degradation. Although the continuous PTH treatment group might have experienced some variation in concentration of PTH within  232  each 48-hour incubation cycle, we believe that this variation in PTH concentration had little effect on our findings, since it has been shown by Ishizuya et al. [9] that the dose-dependent effects of PTH in range of 2 to 13 nM on osteoblastic cells are comparable.  Previous work using rat calvariae osteoblasts and osteoblastic cell lines showed that continuous PTH inhibits osteoblast differentiation, while pulsatile PTH stimulates osteoblast differentiation [9-17]. We based our working hypothesis on these findings, in other words, that the effects of continuous vs. pulsatile PTH would show similar results with MSC cultures. However, using ALP activity assays and staining of ALP expressing cells, as well as analysis of PTHR1, ALP and OC gene expressions, it was shown that continuous treatment with PTH promotes osteogenic differentiation from MSCs, while pulsatile PTH treatment results in inhibition. Thus, our studies showed quite different effects. We believe this may be attributed to multiple reasons that include the use of uncommitted and immature cells (MSC5) as opposed to osteogenic committed osteoprogenitors (calvariae osteoblasts) and osteoblastic cell lines used in previous work. Furthermore, the use of different culture conditions such as the addition of DEX, in our studies, might be a contributing factor. DEX has been shown to affect MSC differentiation mediated by the production of cAMP and through the protein kinase A (PKA) signaling pathway. In addition, the anabolic effects of pulsatile PTH on mature matrix-synthesizing osteoblasts have also been shown to be a result of the stimulation of cAMP production, and subsequent activation of the PKA pathway. Thus, it is possible that the effects of continuous and pulsatile PTH treatments on MSCs observed in our studies might also be mediated by means of the cAMP/PKA pathway.  233  However, detailed studies are required to delineate the interactions of PTH and DEX, and the possible mechanism of PTH on MSCs.  Neither continuous nor pulsatile PTH treatment produced any detectable change in overall MSC proliferation regardless of the presence or absence of DEX. Interpretation of the findings was difficult given the known heterogeneity of MSC cultures [28], which may contain differentiated, uni-, bi- and tripotential progenitors and primitive, self-renewing stem cells. Furthermore, as discussed previously in Section 1.2.2, the effects of PTH on MSC proliferation have been shown to be variable and the conflicting results obtained from in vivo studies can be attributed to the effects of different animal models used [10, 29-32]. Our finding that treatment with PTH, either continuous or pulsatile, increased the CFU-F content of rat MSCs, suggested that PTH might enhance the self renewal of clonogenic progenitors found in MSC cultures. However, CFU-Fs are also known to be heterogeneous in nature [33, 34], and therefore additional experimentation would be required to confirm that all CFU-Fs are derived from primitive stem cells.  The primary method to deliver MSCs for bone tissue regeneration has been the use of 3-D porous scaffolds engineered from biocompatible and biodegradable biomaterials such as gelatin [35, 36] and alginate [37]. In an attempt to replicate the natural ECM, scaffolds provide the initial framework for MSCs to attach, survive, proliferate, and differentiate to form new ECM [38]. Moreover, the scaffold not only retains the delivered MSCs within the defect site, it can also function as a substrate for tissue ingrowth and vascularization. While numerous fabrication technologies have been used to engineer porous scaffolds for cell  234  delivery [39-43], each of the existing processes has their associated disadvantages that have been described [43-46]. Despite the many in vitro and in vivo studies that have demonstrated tissue regeneration with the delivery of MSCs on porous scaffolds [37, 47-5 1], only a limited number of studies have compared in vitro and in vivo responses of MSCs on scaffolds [52, 53]. However, these studies did not explicitly examine the fate and changes in the MSC population after in vivo transplantation. Hence, we have described the use of a novel fabrication technique, termed microwave vacuum drying [54, 55], to engineer biodegradable and bioresorbable porous gelatin-alginate scaffolds with different cross-linking, capable of supporting MSCs delivery for bone regeneration applications. In addition, self-renewal capacity, differentiation potential, and clonogenicity of MSCs in vivo were directly compared to in vitro, to assess the differential responses of the MSCs seeded on these porous gelatin alginate scaffolds.  The microwave vacuum drying process used was able to fabricate highly porous (> 80% porosity) gelatin-alginate scaffolds rapidly (< 30 minutes). However, the resulting irregular pore shape, small median pore sizes  (  2 pm) and the limited interconnectivity with larger  pores in the gelatin-alginate scaffolds found in our studies were probably the biggest drawbacks of using microwave vacuum drying. It has been suggested that pore size in the range of 100  —  500 im is necessary to accommodate the size of cells as well as support cell  migration, ingrowth of vasculature and penetration of new tissues [56, 57]. Thus, the lack of cell migration to the interior of the scaffolds found in our studies both in vitro and in vivo, might partially be attributed to the small pore size and limited interconnectivity that acted as physical  barriers  preventing  any  possible  235  migration.  Furthermore,  without  any  chemoattractants (growth factors) in the interior the scaffolds, cell migration would be limited, if at all. Nonetheless, favorable processing conditions such the absence of organic solvents and low temperatures (< 37 °C) are advantages of microwave vacuum drying that could be utilized in future work for the incorporation of growth factors into porous scaffolds. This would allow for the potential co-delivery of growth factors and MSCs. Our original intent had been to disperse PTH-loaded microspheres within the porous gelatin-alginate scaffold to deliver PTH to the MSCs. However, as noted earlier, this approach was not pursued further when it became clear that precise control of PTH exposure to MSCs could not be achieved.  Although collagen is a commonly used biomaterial for porous scaffold fabrication [8, 5 8-62], we selected gelatin and alginate due to their well established biocompatibility and biodegradability, and their clinically approved used as biomedical products. Gelatin has also been shown to be a substrate for effective adherence of MSCs [74]. From initial screening studies, it was determined that the use of a blend of gelatin and alginate produced porous scaffolds with more suitable pore size, porosity and biodegradation profile than the individual materials alone. Cross-linking the gelatin-alginate scaffolds was shown to have a pronounced effect on biodegradation and bioresorption following subcutaneous implantation. Low cross-link density scaffolds were shown to be biodegraded and bioresorbed rapidly, being completely bioresorbed after 21 days. Whereas, high cross-link density scaffolds were able to resist biodegradation and bioresorption much longer, retaining the original shape and losing less than one quarter of the volume after the same period of time. Biodegradation of gelatin by proteolysis was likely a major biodegradation route since gelatin is known to  236  biodegrade through the actions of proteases [75]. A number of different proteases would be expected to be present at the implantation site as part of the natural wound healing process [76].  The porous gelatin-alginate scaffolds used in this work were capable of absorbing large amount of fluids and would therefore be unlikely to possess any mechanical strength in vivo. However, these scaffolds were designed to serve as delivery systems for either growth factors and/or cells, and they were not intended to provide any mechanical support. Thus, the mechanical stiffness and strength of these scaffolds were not measured.  Angiogenesis is an integral part of bone development and repair, hence, the establishment of a functional vascular network around and within an implanted scaffold is an essential pre requisite for the survival of transplanted cells and its integration with existing host tissue [63]. The rate and extent of revascularization in bone regeneration is critical because rapid revascularization favors osteoblastic differentiation, whereas prolonged hypoxia favors formation of cartilage or fibrous tissue [57, 63]. The porous gelatin-alginate scaffolds demonstrated excellent biocompatibility. There was no evidence adverse reactions such of chronic inflammation or fibrous encapsulation around the scaffold, unlike other studies where fibrous connective tissues were found to surround the subcutaneously implanted atelocollagen [64] and poly(ethylene glycol)-based [65] scaffolds in SCID mice. There was visual evidence of angiogenesis and neovascularization surrounding our implanted scaffolds that would have been likely to assist the survival of the seeded MSCs, allowing the implanted cells to proliferate at the same rate as in nutrient rich in vitro environments. The mechanism  237  for the observed angiogenesis is not fully known and may be partially due to inflammation associated with the natural wound healing process after the implantation of the scaffold [76]. In addition, undifferentiated MSCs have been shown to produce a number of pro-angiogenic factors, in particular in hypoxic conditions, and to induce angiogenesis when transplanted in a variety of locations, most noticeably in infarcted myocardium [77]. Thus, the induction of host derived angiogenesis would be expected in our experimental conditions even in the absence of inflammation.  By examining the transcription of the differentiation markers for osteogenesis (Runx2, osterix, ALP, OC and BSP), chondrogenesis (Sox9 and aggrecan), and adipogenesis (PPARy and LPL) it was determined that MSCs differentiation along all three lineages were suppressed under the complex in vivo environment compared to in vitro culturing conditions. The inhibition of MSC differentiation in vivo was likely due to the subcutaneous environment where they were implanted. Given that in subcutaneous tissues, where neither osteogenesis nor chondrogenesis actually occurs, it is likely the endogenous biological signals in this microenvironment only served to suppress such differentiation by MSCs. Furthermore, since adipogenesis found in the subcutaneous tissue is primarily the result of more differentiated adipocyte precursors [66], the involvement of MSCs might be very minor, as reflected in their lack of differentiation found in vivo. Thus, the findings in this work highlight the fact that MSC differentiation in vivo is complex and cannot be easily replicated in vitro.  238  It should be acknowledged that one of the limitations of the in vitro and in vivo studies was the small sample size (n  <  5) used in each experiment. However, for all cell culture and  animal studies, separate and repeat experiments were conducted. Nevertheless, for data showing no significant difference, it is possible these studies may not have had enough statistical power to detect differences.  Even with modem day sterilization and aseptic procedures, every orthopaedic surgery performed to restore bone functionalities or promote bone regeneration, can still be complicated by infections. Particularly, bacterial infection remains a major concern when there is the use of orthopaedic implants [67]. Therefore, orthopaedic surgeries are always accompanied by the administration of antibiotics [68, 69]. With this knowledge, our laboratory is developing delivery systems for therapeutic agents (growth factors, hormones, stem cells) that promote bone regeneration and concurrently exploring the localized delivery of antibiotics from biomaterials that treat and prevent infections. A composite delivery system capable of co-delivering growth factors, cells and antibiotics would represent a major advance in orthopaedic medicine.  We become interested in using the antibiotic FA, given that it may still be effective against a wide spectrum of pathogens that commonly causes prosthetic joint infections including those MRSA and other multi-drug resistant Gram-positive bacteria [70]. Thus, as a first strategy to develop a localized delivery system, we chose to develop biodegradable polymeric microspheres loaded with FA. Although there was a previous report of sodium fusidate (sodium salt of FA) being loaded into PLGA microsphere [71], no other studies had  239  investigated  FA-loaded  microspheres.  The  well  established  biocompatibility  and  biodegradability of PLGA and PHBV led us to select these two biomaterials for study. A factorial design strategy [72, 73] to formulation permitted an efficient approach to determining the conditions under which a high encapsulation efficiency of FA (> 90%) could be achieved. However, what became most interesting and a phenomenon that to our knowledge, has not been previously demonstrated in any drug loaded polymeric microsphere formulations, was the observation of highly regular spherical protrusions on the surface of PLGA, but not PHBV microspheres. Furthermore, the size and height of these protrusions increased with drug loading.  Using a combination of SEM, BSEM, laser confocal microscope, Raman spectroscopy analysis and miscibility determination with solvent-cast films as well as real-time video recordings of single microspheres formation, it was determined that FA phase separates from all PLGA polymers investigated, regardless of any changes in other formulation factors, to form distinct, completely amorphous, spherical FA-rich solid microdomains. Increasing FA loadings only served to produce larger FA-rich phase separated microdomains that were present throughout the PLGA matrix, and on the microsphere surface. The phase separation of FA was only identified with PLGA polymers and was not observed for PHBV polymers, even though both belong to the family of poly(a-hydroxy acid) polymers. The subtle differences in molecular structure of the polymer chain such as the extra methyl group found in the backbone of the repeat units of PHBV compared to PLGA polymer could be a possible explanation for FA to not undergo phase separation from the PHBV matrix.  240  The burst phase of the biphasic release profiles of all FA-loaded PLGA microsphere formulations was likely a result of the dissolution of the phase separated FA located on the surface. Increasing the initial FA loading or changing the polymer composition from PLGA (85/15) to PLGA (50/50) produced a larger burst. These results were due to the increased amount of FA phase separated on the surface and the increased hydrophilicity of the PLGA (50/50) polymer matrix, respectively. FA-loaded PHBV microsphere formulations also showed biphasic release profiles with a significant burst release due to FA being distributed on the surfaces as well as the cracked, rough, and possibly porous microsphere surfaces.  In conclusion, the major findings in this thesis were that osteogenic differentiation of rat MSCs increased with continuous PTH treatment, and decreased with pulsatile PTH exposure, with these effects being strongly dependent on the presence of DEX. However, MSC proliferation was not influenced by PTH, independent of the presence or absence of DEX. Moreover, increases in the CFU-F content of rat MSCs suggested that PTH might enhance the self renewal of clonogenic progenitors found in MSC cultures. Biocompatible, biodegradable and bioresorbable porous gelatin-alginate scaffolds produced by microwave vacuum drying were found to support MSC attachment, proliferation and differentiation. However, MSC differentiation (osteogenic, chondrogenic, adipogenic) were found to be suppressed in vivo compared to in vitro when seeded on these porous scaffolds.  In the process of formulating biocompatible and biodegradable FA-loaded PLGA and PHBV microspheres, an interesting phase separation phenomenon of FA in PLGA but not in PHBV polymer was observed. Phase separated FA formed distinct, large, completely amorphous,  241  spherical FA-rich solid microdomains throughout the PLGA microsphere, and on the microsphere surface. This suggests to us that PLGA matrices are not an appropriate choice for delivery of FA, whereas PHBV, where the drug was homogenously distributed throughout the polymer, may be better.  Thus, the data presented in this thesis contribute to our understanding of PTH effects on MSCs, the responses of MSCs on porous gelatin-alginate scaffolds as well as the solid-state characteristics and release of FA loaded in PLGA and PHBV microspheres.  242  5.2 Suggestions for future work Understanding that continuous PTH treatment promotes osteogenic differentiation of MSCs and knowing the limitations in the initial PTH-loaded microsphere formulations, it should be possible to optimize a PTH delivery system by dispersing PTH-loaded microspheres within porous scaffolds and fabricating using the microwave vacuum drying technique. The scaffold may function as a secondary controlled release mechanism to provide the precision desired for the continuous delivery of PTH. In vitro and in vivo evaluation of the PTH released from the composite porous scaffold delivery system to affect MSC differentiation should be performed. Once the therapeutic effects of PTH have been established, a ‘critical’ bone defect animal should be developed to investigate the efficacy and effectiveness of the combined delivery of PTH and MSCs to regenerate bone.  Having the ability to process various biomaterials that are hydrogel-like such as gelatin, alginate, collagen, fibrin, chitosan, hyaluronan, among others, and possessing favorable processing conditions (avoid the use of organic solvents and low temperature), porous scaffold cell delivery systems should be further developed using the microwave vacuum drying technique. This may include the optimization of the blend of gelatin and alginate to improve the scaffolds’ pore size and interconnectivity, and examination of other biomaterials listed above, separately or in blends. 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Eur Cell Mater 2008; 15:100-114. 64. Hidetsugu T, Paola RA, Hitoshi N, Mehmet G, Jin LY, Borkosky SS, et al. Mechanism of bone induction by KUSA/Al cells using atelocollagen honeycomb scaffold. J Biomed Sci 2007 Mar;14(2):255-263.  248  65. Alhadlaq A, Mao JJ. Tissue-engineered neogenesis of human-shaped mandibular condyle from rat mesenchymal stem cells. J Dent Res 2003 Dec;82(12):951-956. Rodeheffer MS, Birsoy K, Friedman JM. Identification of white adipocyte progenitor 66. cells in vivo. Cell 2008 Oct 17;135(2):240-249. 67. Trampuz A, Widmer AF. Infections associated with orthopedic implants. Cuff Opin Infect Dis 2006 Aug;19(4):349-356. 68. Esposito S, Leone S. Prosthetic joint infections: microbiology, diagnosis, management and prevention. International Journal of Antimicrobial Agents 2008;32(4):287293. 69. Ratnaraja NV, Hawkey PM. Current challenges in treating MRSA: what are the options? Expert Rev Anti Infect Ther 2008 Oct;6(5):601-618. Mandell LA. Fusidic Acid. In: Mandell GL, Bennett JE, Dolin R, editors. Mandell, 70. Douglas and Bennett’s Principles and Practice of Infectious Diseases. 5 edition ed. Philadelphia: Churchill Livingstone, 2000. p. 306. 71. Cevher E, Orhan Z, Sensoy D, Ahiskali R, Kan PL, Sagirli 0, et a!. Sodium fusidate poly(D,L-lactide-co-glycolide) microspheres: preparation, characterisation and in vivo evaluation of their effectiveness in the treatment of chronic osteomyelitis. J Microencapsul 2007 Sep;24(6):577-595. 72. Ruhé PQ, Boerman OC, Russel FGM, Spauwen PHM, Mikos AG, Jansen JA. Controlled release of rhBMP-2 loaded poly(dl-lactic-co-glycolic acid)/calcium phosphate cement composites in vivo. Journal of Controlled Release 2005; 106(1-2): 162-171. El-Gibaly I, Abdel-Ghaffar SK. Effect of hexacosanol on the characteristics of novel 73. sustained-release allopurinol solid lipospheres (SLS): factorial design application and product evaluation. International Journal of Pharmaceutics 2005 ;294( 1-2) :33-51. 74. Yang Y, Rossi FM, Putnins EE. Ex vivo expansion of rat bone marrow mesenchymal stromal cells on microcarrier beads in spin culture. Biomaterials 2007 Jul;28(20):3 110-3120. 75.  Veis A. Molecular Biology. Vol. 5. The Macromolecular Chemistry of Gelatin, 1964.  76. Patel ZS, Mikos AG. Angiogenesis with biomaterial-based drug- and cell-delivery systems. J Biomater Sci Polym Ed 2004; 1 5(6):70 1-726. Imanishi Y, Saito A, Komoda H, Kitagawa-Sakakida S, Miyagawa S, Kondoh H, et 77. al. Allogenic mesenchymal stem cell transplantation has a therapeutic effect in acute myocardial infarction in rats. J Mol Cell Cardiol 2008 Apr;44(4):662-671.  249  Appendix A: Parathyroid hormone loaded PLGA and PHBV microspheres: Formulations development and release kinetics  A.1 Materials and Methods A.1.1 Chemicals Recombinant rat parathyroid hormone (PTH) peptide 1-34 was synthesized and provided by our collaborator Dr. Fabio M. Rossi (Biomedical Research Centre, University of British Columbia, Vancouver, Canada). Poly(3 -hydroxybutyric acid-co-3-hydroxyvaleric acid) (PHBV, 5 and 12 wt % hydroxyvaleric acid, HV), poly(vinyl alcohol) (PVA, 87  —  89%  hydrolyzed), dichloromethane (DCM), bovine serum albumin (BSA), sorbitan monooleate (Span 80) and the different reagents needed to prepare phosphate buffered saline (PBS, pH 7.4) solution were all obtained from Sigma-Aldrich (Oakville, ON, CA). Poly(DL-lactic-co glycolic acid) (PLGA, 85/15 LA/GA) with an intrinsic viscosity of 0.61 dL/g in chloroform (equivalent to Mw  86,000 g/mol) was obtained from Birmingham Polymers Inc.  (Birmingham, AL, USA) and poly(DL-lactic-co-glycolic acid) (PLGA, 50/50 LA/GA) with intrinsic viscosity of 0.58 dL/g in hexafluoroisopropanol (equivalent to Mw  84,000 g/mol)  was obtained from Lactel® Absorbable Polymers (Pelham, AL, USA). PTH (1-34) (Rat) radioimmunoassay (RIA) kit was purchased from Phoenix Pharmaceuticals, Inc. (Belmont, CA, USA).  250  A.1.2 Fabrication of PTH-loaded PLGA and PHBV microspheres PTH encapsulated microspheres were synthesized by the water-in-oil-in-water (W/0/W) double emulsion solvent evaporation method [1] illustrated in Figure A-i. The materials used for each PTH-loaded PLGA and PHBV micro sphere formulation are listed in Table A-i and A-2, respectively. For example (Run #1 in Table A.i), PTH (1 mg) and BSA (5 mg) PLGA were first dissolved in 100 iL of distilled water by vortexing and sonication. Separately, 100 mg of PLGA (85/15) was dissolved in 1 mL of DCM. PTH/BSA solution, along with 50 iL of Span 80 were then added to the PLGA solution and homogenized at high speed to form the first 01W emulsion. Immediately, the first 01W emulsion was drop-wise added into 100 mL of 2.5% (w/v) PVA solution with an overhead propeller stirring at 600 rpm to form the second W/OIW emulsion. The resulting double emulsion was stirred continuously for 2.5 h at room temperature under the fume hood to evaporate the organic solvent. The PTH-loaded PLGA microspheres were collected by centrifuging at 3000 rpm for 5 mm and subsequently washed four times with distilled water. The microspheres were then vacuum dried at room temperature and stored in a desiccator for further analysis. All other PTH-loaded PLGA and PHBV microspheres were fabricated using the same procedures described above. However, for the PTH-loaded PHBV microspheres, chloroform instead of DCM was used and stirring was done for 4 h to evaporate the solvent.  251  Mechanical Mixer  -  Water-in-Oil (W/O) Emulsion  —  /r  ‘7  Organic Polymer Solution  Collected Microspheres  Water-in-Oil ,V/O) Emulsion  W/O  W/O/W  EMULSION  EMUlSION  SOLVENT EVAPORATION  HARDENING & PRECIPITATION  Figure A-i: Schematic diagram of the water-in-oil-in-water (W/O/W) double emulsion technique to encapsulate growth factors (hydrophilic drugs). (I) Growth factor dissolved in an aqueous solvent is emulsified in a non-miscible organic polymer solution to form the first water-in-oil (W/O) emulsion. (II) The primary W/O emulsion is transferred to an excess secondary aqueous medium containing a stabilizer, at which point homogenization again or intensive stirring is applied to form the water-in-oil-in-water (W/O/W) double emulsion. (III) Stabilization of the double emulsion is achieved by constant mechanical agitation (stirring) and subsequent removal (evaporation) of the organic solvent. (IV) Hardening of the polymer surrounding the growth factor to produce growth factorloaded microspheres.  Table A-i: Summary of formulation conditions for PTH loaded PLGA microspheres. Polymer concentration used was 10% (w/v) F-  Formulation Conditions Polymer BSA Composition (lactic:glycolic)’ (% wlw)b 5 85:15 0 85:15 5 50:50 0 50:50  Results  Mean Diameter Span 80 Run PTH (% WIW)a (% vIv)’ (pm) 1 1 5 144 2 1 5 152 3 1 0.5 101 4 1 0.5 93 a w/w = PTH/polymer, b w/w = BSAlpolymer, where BSA = bovine serum albumin, C lactic:glycolic lactic acid/glycolic acid monomer molar ratio of the polymer, d %v/v = volume % of Span 80 relative to the solvent, DCM, where Span 80 = Sorbitan monooleate cJ  Table A-2: Summary of formulation conditions for PTH-loaded PHBV microspheres. Span 80 concentration used was 0.5% (v/v), where Span 80 = Sorbitan monooleate.  a  b C  d  Run 1  PTH (% wiw)a 1  2  1  Formulation Conditions Polymer Concentration BSA (% wlw)” (% wIvY 5 10  5  5  3 1 5 5 4 1 0 5 w/w = PTH/polymer, w/w = BSA/polymer, where BSA bovine serum albumin, w/v = polymer/solvent, wt HV = % weight of hydroxyvalerate in the copolymer PHBV  p  Results  Polymer Composition (% wt HVY’ 12  Mean Diameter (jim) 205  12  127  5 5  147 119  A.1.3 Microsphere particle size determination PTH-loaded PLGA and PHBV microspheres mean particle size and size distributions were determined using a Malvern Mastersizer 2000 (Malvem Inc., Malvern, Worcestershire, UK), laser diffraction particle size analyzer. Briefly,  5  —  10 mg of micro spheres were suspended  in 5 mL of distilled water with two drops of 1% polysorbate 80 (Tween 80) and sonicated for 2 mm to prevent aggregation of microspheres.  A.1.4 Scanning electron microscope (SEM) analyses The morphologies of the PTH-loaded microsphere formulations were imaged using SEM. Briefly, samples were sputter-coated with a layer of 60:40 alloy of gold:palladium using a Denton Vacuum Desk II sputter-coater (Moorestown, NJ) at 50 millitorr. SEM images were then captured using a Hitachi S-3000N system (Tokyo, Japan) scanning at 10 —20 keV.  A.1.5 In vitro PTH release from PLGA and PHBV microspheres In vitro PTH release from PLGA and PHBV inicrosphere formulations was carried out in PBS (pH 7.4) at 37 °C. For release studies,  1.5 mg of PTH-loaded microspheres were  placed into 2 mL of PBS and the samples were tumbled end-over-end at 10 rpm in a thermostatically controlled oven at 37 °C. At specified time points, the sample tubes were centrifuged at 3000 rpm for 5 mm, and all the sample volume was then withdrawn and frozen at  —  20 °C until it was analyszed to determine the amount of PTH released. Fresh PBS (pH  7.4) added back to each sample to maintain sink conditions. Concentrations of PTH in the  254  release medium were measured using a radioimmunoassay kit according to manufacture’s instructions.  A.2 Results The effects of different formulation factors on PTH-loaded PLGA and PHBV microsphere mean diameter are shown in Table A-i and A-2, respectively. As the polymer composition was changed from PLGA (50/50) to PLGA (85/15), the mean diameter of PTH-loaded microspheres increased. Similarly, as polymer concentration increased in the PHBV microspheres, the mean diameter increased. The SEM images of the different formulations described in Table A-i and A-2 for the PTH-loaded PLGA and PHBV microspheres are illustrated in Figure A-2 and A-3, respectively. The morphologies of all the microspheres were not affected by the different formulation factors. They all possessed relatively smooth surfaces with a few scattered pinholes visible on the surfaces. In addition, the in vitro PTH release profiles from PLGA and PHBV microsphere formulations are shown in Figure A-4 and A-5, respectively. All microsphere formulations demonstrated a biphasic release profile with an initial burst release followed by a slow controlled release.  255  Run#1  Run#2  Run#3  Run#4  Figure A-2: SEM images of parathyroid hormone (PTH) loaded PLGA microspheres. See Table A-i for details of each run.  L’J ON  Run #1  Run #2  Run #3  Run #4  Figure A-3: SEM images of parathyroid hormone (PTH) loaded PHBV microspheres. See Table A-2 for details of each run.  50’  ø.  40’ Run -*- Run —v- Run Run -‘  (uu) wE >w  #1 #2 #3 #4  .! , 20• E— (-‘I D C  o  5  15  10  20  25  Time (Days) Figure A-4: Cumulative in vitro release profiles of PTH loaded PLGA microspheres in PBS (pH  7.4)  37 °C, n = 3 See Table A-i for details of each run.  257  WE  -‘-  Run #1 Run #2 Run #3  >4_  -4--  Run#4  (fl.  -.•  -*-  .!D,  0  5  10  15  20  25  Time (Days) Figure A-5: Cumulative in vitro release profiles of PTH loaded PHBV microspheres in PBS (pH 7.4)  @ 37 °C, n  =  3. See Table A-2 for details of each run.  258  A.3 References 1. Sinha VR, Trehan A. Biodegradable microspheres for protein delivery. Journal of Controlled Release 2003 ;90(3) :261-280.  259  Appendix B: Additional figures for Chapter 4: Fusidic acid loaded PLGA and PHBV microspheres for bone infections: Formulations development and solid-state characterization B.1 Additional Figures  260  PLGA (50/50)  PLGA (85/1 5)  PLLA  Figure B-i: SEM images illustrating the phase separation of FA from different PLGA composition matrices. PLGA (50/50) = poly(lactic-co-glycolic acid) with 50/50 lactic/glycolic acid molar ratio. PLGA (85/15) = poly(lactic-co-glycolic acid) with 85/15 lactic /glycolic acid molar ratio. PLLA poly(L-lactic acid) polymer, where it can be consider to be PLGA (100/0) with 100% lactic acid. All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and 30 % FA loadings (w/w).  A  B  C  Figure B-2: Raman spectroscopy images of FA distribution in 30% (w/w) FA-loaded PLGA (50/50) microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light microsphere montage, 100x magnification, (B) distribution of PLGA rich regions (green) across microsphere, and (C) distribution of FA-rich regions (red) across microsphere.  A  B  C  Figure B-3: Raman spectroscopy images of FA distribution in 30% (w/w) FA-loaded PLLA microspheres fabricated at a polymer and drug concentration of 10% (w/v). (A) White light microsphere montage, 100x magnification, (B) distribution of PLLA-rich regions (green) across microsphere, and (C) distribution of FA-rich regions (red) across microsphere.  CN  PVA  Gelatin  Figure B-4: SEM images illustrating the effects of emulsifying agents on the phase separation of FA from PLGA (85/15) microspheres. PVA = poly(vinyl alcohol). All microsphere formulations were manufactured at a polymer and drug concentration of 10% (w/v) and 30% FA loadings (w/w).  Appendix C: ANOVA Result Tables for Factorial Design Experiments  265  Table C-i: ANOVA table for effect of polymer and drug concentration on the mean diameter of FA-loaded PLGA microspheres Factor: Polymer and Drug Concentration Response: Mean Diameter ANOVA for selected factorial model Analysis of variance table [Partial sum of squares Type Ill] -  Source Model C-Polymer & Drug Concentration Residual Cor Total  Sum of Squares 3612.5 3612.5 289.5 3902  Std. Dev. Mean C.V.% PRESS  6.946221995 77.5 8.96286709 514.6666667  Factor Intercept C-Polymer and Drug Concentration  Coefficient Estimate 77.5 21.25  df 1 1 6 7  df 1 1  Mean Square 3612.5 3612.5 48.25  F Value 74.87046632 74.87046632  R-Squared Adj R-Squared Pred R-Squared Adeq Precision  0.925807278 0.913441825 0.868101828 12.23686776  Standard Error 2.455860338 2.455860338  95% Cl Low 71.49072645 15.24072645  Final Equation in Terms of Coded Factors: Mean Diameter =  77  +  21.25C  Final Equation in Terms of Actual Factors: Mean Diameter  =  13.75  +  8.5*Polymer and Drug Concentration  0.0001  95% Cl High 83.50927355 27.25927355  VIF 1  Table C-2: ANOVA table for effect of(1) drug loading and (2) polymer and drug concentration on the encapsulation efficiency of  FA-loaded PLGA microspheres Factors: (1) Drug Loading; (2) Polymer and Drug Concentration Response: Encapsulation Efficiency ANOVA for selected factorial model Analysis of variance table [Partial sum of squares Type Ill] -  L’J ON  Source Model A-Drug Loading C-Polymer & Drug Concentration Residual Cor Total  Sum of Squares 440.79025 220.0802 220.71005 66.41995 507.2102  Std. Dev. Mean C.v. % PRESS  3.64472084 89.615 4.06708792 170.035072  Factor Intercept A-Drug Loading C-Polymer and Drug Concentration  df 2 1 1 5 7  Coefficient Estimate 89.615 5.245  df 1 1 1  -5.2525  Mean Square 220.395125 220.0802 220.71005 13.28399  F Value 16.591 03364 16.56732653 16.61474075  R-Sq uared Adj R-Squared Pred R-Squared Adeq Precision  0.869048473 0.816667863 0.664764092 9.406671441  Standard Error 1.288603411 1.288603411 1.288603411  95% Cl Low 86.30253967 1.932539672 -8.564960328  0.0062 0.0096 0.0096  95% Cl High 92.92746033 8.557460328 -1 .940039672  I  VIF I 1  Final Equation in Terms of Coded Factors: Mean Diameter  =  89.615  Mean Diameter  =  94.8825  +  5.245*A 5.2525*C -  Final Equation in Terms of Actual Factors: +  0.5245*Drug Loading 2.101*Polymer and Drug Concentration -  Table C-3: ANOVA table for effect of polymer and drug concentration on the mean diameter of FA-loaded PHBV microspheres Factor: Polymer and Drug Concentration Response: Mean Diameter ANOVA for selected factorial model Analysis of variance table [Partial sum of squares Type III] -  Source Model C-Polymer & Drug Concentration Residual Cor Total Std. Dev. Mean  Sum of Squares 8911. 125 8911.125 620.75 9531.875  PRESS  10.17144696 109.375 9.299608652 1103.555556  Factor Intercept C-Polymer and Drug Concentration  Coefficient Estimate 109.375 33.375  df 1 1 6 7  df 1 1  Mean Square 8911.125 8911.125 103.4583333  F Value 86.13250101 86.13250101  R-Squared Adj R-Squared Pred R-Squared Adeq Precision  0.934876402 0.924022468 0.884224714 13.12497627  Standard Error 3.596149561 3.596149561  95% CI Low 100.5755393 24.57553933  Final Equation in Terms of Coded Factors: Mean Diameter =  109.375  +  33.375*C  Final Equation in Terms of Actual Factors: Mean Diameter =  9.25  +  13.35*Polymer and Drug Concentration  <0.0001 <0.0001  95% CI High 118.1744607 42.17446067  VIF 1  Appendix D: UBC Research Ethics Board Certificates of Approval  !ac S  THE UNIVERSITY OF BRITISH COLUMBIA -  ANIMAL CARE CERTIFICATE Application Number: A05-035l Investigator or Course Director: Fabio Rossi Department: Medicine, Faculty of Animals:  Mice multiple nbred and transgenic strains 1177  Start Date:  March 22, 2005  April 30, 2008  Funding Sources: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title:  Stem Cell Network (SCN) Networks of Centres of Excellence (NCE) -  Cell therapy for muscular disease Jesse s Journey Foundation Targeted cell fusion to myofibers: a novel therapeutic approach Canadian Institutes of Health Research (CIHR) Role of CD34 in muscle stemlprogenitor cell function  NeVOÜCS of Centres of Excellence (NCE) Development of stable viral vectors for the expression of inducible genes in stem cells Canadian Institutes of Health Research (CIHR) Functional role of hernatogenous inflanunatory cells in amyotrophic lateral sclerosis  269  Funding Agency: Funding Title:  Funding Agency: Funding Title:  Canadian Institutes of I-Iealth Research (CIHR) Mesenehymal stem cells and biomaterials in bone regeneration: a team approach Networks of Centres of Excellence (NCE)  Funding Title:  Investigation of SET domain proteins with regard to their role in the developmental plasticity of adult stem cells  Funding Agency: Funding Title:  Natural Sciences and Engineering Research Council of Canada (NSERC) Bone marrow-derived stem cells for treatment of neurological disease Canadian Institutes of Health Research (CIHR) Mesenchymal stem cells and hiomaterials in bone regeneration: A team approach  Funding Agency:  Michael Smith Foundation for Health Research  Funding Title:  Characterization of mesenchymal stem cell differentiation and self-renewal in the context of an in vivo model of bone regeneration  Funding Agency:  Networks of Centres of Excellence (NCE)  Funding Title:  Identification of phosphoregulation pathways involved in hemotopoetic stem cell selfrenewal  Funding Agency:  Michael Smith Foundation for Health Research  Funding Title:  Identification of Phosphoregulation pathways involved in Hematopoetic Stem cell self-renewal  Funding Agency: Funding Title:  Funding  I Agency:  I Funding Title:  II  Mesenchymal stem cells and biomaterials in bone regeneration: a team approach  Funding Agency:  Funding Agency: Funding Title:  I  Canadian Institutes of Health Research (CIHR)  Funding Agency:  Muscular Dystrophy Association (US) Identification and engineering of circulating myogenic progenitors Canadian Institutes of Health Research (QHR) Microenviromnental control of stem cell fate Muscular Dystrophy Association of Canada  270  Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title:  Circulating myogenic progenitors: a lineage analysis Canadian Institutes of Health Research (CIHR) Mesenchymal stem cells and hiomaterials in bone regeneration Stem Cell Network (SCN)  -  Networks of Centres of Excellence (NCE)  Haematopoletic stem cell aging and CD34 expression Michael Smith Foundation for Health Research Stem cell aging  Funding Agency:  Canadian Institutes of Health Research (CIHR)  Funding Title  Regeneration of hard & soft periodontal connective tissues utilizing engineered mesenchymal stem cells  Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title:  Canadian Institutes of Health Research (CIl-IR) Mesenchymal stem cells and biomaterials in bone regeneration: A team approach Michael Smith Foundation for Health Research Identification of circulating cells with a myogenic potential Michael Smith Foundation for Health Research Engineered mesenchymal stem cells for osteoporotic fracture prevention  Funding Agency:  Canadian Institutes of Health Research (CIHR)  Funding Title:  Usin& mesenchymall stem cells and nonoengineered membranes to regenerate hard and soft pendontal connective tissues  Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Fimding Title:  Various Sources Adult Stem Cells: Maintenance and transdifferentiation Canadian Institutes of Health Research (CIHR) Mesenchymal stem cells and biomaterials in bone regeneration: A team approach Stem Cell Network (SCN)  -  Networks of Centres of Excellence (NCE)  Lentivector engineering for functional studies in stem cells  271  Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title: Funding Agency: Funding Title:  Unfunded title:  Canadian Institutes of Health Research (CIHR) Adult stem cells for fracture prevention Networks of Centres of Excellence (NCE) Stem cell plasticity Canadian Institutes of Health Research (CIHR) Microenvironmental control of stem cell fate Michael Smith Foundation for Health Research Identification of circulating cells with a myotenic potential Michael Smith Foundation for Health Research Identification of circulating cells with a myogenic potential Canadian Institutes of Health Research (CIHR) Mesenchymal stem cells and hiomaterials in bone regeneration: a team approach Canadian Institutes of Health Research (CIHR) Microenvironmental control of stem cell fate Canadian Institutes of Health Research (CIHR) Mesenchymal stem cells and biomaterials in bone regeneration: a team appmach  N/A  The Animal Care Committee has examined and approved the use of animals for the above experimental project This certificate is valid for one year from the above start or approval date (whichever is later) provided there is no change in the experimental procedures. Aimual review is required by the CCAC and some granting agencies.  A copy of this certificate must be displayed in your animal facility. Office of Research Services and Administration 102, 6190 Agronomy Road, Vancouver, BC V6T lZ3 Phone: 604-827-5111 Fax: 604-822-5093  272  

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