Characterization of Genetically Modified Hucpvcs As an Osteogenic Cell Source

Characterization of Genetically Modified HUCPVCs as an Osteogenic Cell Source.

Catalina Estrada Vallejo

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Institute of Biomaterials & Biomedical Engineering University of Toronto

Characterization of Genetically Modified HUCPVCs as an Osteogenic Cell Source.

Catalina Estrada Vallejo

Doctor of Philosophy, 2013

Institute of Biomaterials & Biomedical Engineering University of Toronto

Abstract

Tissue engineering and ex vivo gene therapy can be used synergically as tool to regenerate bone, which overcome the problems of currently available bone replacements. Recently, a new source of mesenchymal stromal cells (MSCs) has been found in the umbilical cord; human umbilical cord perivascular cells

(HUCPVCs) provide an alternative to bone marrow derived MSCs and due to their easy harvest, fast expansion, and non-immunogeneic and immunomodulatory phenotype we hypothesized that HUCPVCs are a putative candidate cell source for osteogenic ex vivo gene therapy. This work proposes the generation of cocktails of genetically modified HUCPVCs and their cryopreservation as an “off the shelf” therapeutic. This approach involves the engineering of osteogenic cell populations, by genetically modifying HUCPVCs using recombinant adenoviruses to deliver four fundamental genes for bone formation: bone morphogenetic protein 2 (BMP-2), runt-related transcription factor 2 (Runx2), Osterix (OSX/SP7) transcription factor and vascular endothelial growth factor (VEGF). Our results show that HUCPVCs can be efficiently modified by adenoviruses and can be cryopreserved without affecting the production efficiency and bioactivity of proteins of interest produced by the cells. Moreover, overexpression of BMP2, Runx2 and SP7 enhances ALP activity levels in HUCPVCs and upregulates ALP, OPN, COL1A1 and OCN gene expression; data that provides the first evidence of the effects of combinational expression of BMP2, Runx2 and SP7. Furthermore, we report for the first time the genetic modification of human BMSCs to express SP7 and Runx2, which enhances their ALP activity and matrix mineralization capacity. ii

Acknowledgments

I would like to extend my sincerest gratitude to my supervisor John E. Davies for his support, guidance, encouragement and for giving me the opportunity to work in such a great environment.

Thank you to Bernhard Ganss for opening the door of your lab to me and for the continuous help and support during this process. I also wish to thank the members of the Ganss lab especially James Holcroft for all his technical support.

Thank you to my committee Drs. Armand Keating and Craig Simmons for their time, assistance and insight into this work.

To ALL the Boneheads, you have made this journey so much easier and fun. Thanks for your moral support and advice on different fields. I was really lucky to be part of such an amazing group, it was a pleasure to work with all of you!

Finally, I thank my parents Martha and Luis, my brother Esteban and the rest of my family for their unconditional support and love. To Dave and my friends thanks for the love and fun times. I am pretty lucky to have such a great team by my side.

iii

Table of Contents

Abstract ...... ii Aknowledments ...... Error! Bookmark not defined. Table of Contents ...... iv List of Tables ...... x List of Figures ...... xi List of Abbreviations ...... xiii

1. Introduction ...... 1 1.1. Bone and Osteogenesis ...... 1 1.2. Need for Bone Substitutes ...... 2 1.3. Bone Tissue Engineering ...... 4 1.3.1. Cells for Bone Tissue Engineering ...... 4 1.3.1.1. Mesenchymal Stromal Cell ...... 5 1.3.1.1.1. Bone Marrow Stromal Cells ...... 6 1.3.1.1.2. Human Umbilical Cord Perivascular Cells ...... 6 1.3.2. Osteoinductive Factors in Bone Tissue Engineering ...... 7 1.4. Orthopedic Gene therapy ...... 8 1.4.1. Gene delivery vectors ...... 9 1.4.1.1. Non- viral Vectors ...... 9 1.4.1.2. Viral Vectors ...... 10 1.4.2. Target Genes for Orthopedic Gene therapy ...... 12 1.5. Bone Morphogenetic Protein -2 ...... 12 1.5.1. BMP-2 Synthesis and Posttranslational Modification ...... 13 1.5.2. Furin in BMP-2 Cleavage ...... 13 1.5.3. BMP-2 Signaling Cascade ...... 14 1.5.4. BMP-2 Regulators and Inhibitors ...... 16 1.5.5. BMP-2 ex vivo Gene Therapy...... 16 1.6. Vascular Endothelial Growth Factor A ...... 20 1.6.1. VEGF ex vivo Gene Therapy ...... 22 iv

1.7. Runt-related Transcription Factor 2 ...... 25 1.7.1. Runx2 ex vivo Gene Therapy ...... 27 1.7.1.1. Runx2 and BMP2 Combinational ex vivo Gene Therapy ...... 30 1.8. Osterix Transcription Factor ...... 32 1.8.1. Osterix ex vivo Gene Therapy ...... 34 1.9. Rationale ...... 37 1.10. Hypothesis ...... 38

2. Materials and Methods ...... 39 2.1. HUCPVCs Isolation, Thawing and Culture...... 39 2.2. Genetic Modification of HUCPVCs ...... 39 2.2.1. FuGENE 6 ...... 40 2.2.1.1. Flow Cytometry ...... 40 2.2.2. Lipofectamine LTX ...... 41 2.2.3. Amaxa Nucleofector II ...... 41 2.2.3.1. Longevity of GFP Expression After HUCPVCs Nucleofection ...... 42 2.2.4. Retroviral Transduction of HUCPVCs ...... 42 2.2.4.1. Production of Runx2- and Mock-retrovirus ...... 42 2.2.4.2. Viral Infection of HUCPVCs ...... 43 2.2.4.3. Immunofluorescence ...... 44 2.2.4.4. Osteogenic Cultures of Runx2-HUCPVCs and Mock-HUCPVCs ...... 45 2.2.4.5. ALP Staining ...... 45 2.2.4.6. Von Kossa Staining ...... 45 2.2.4.7. RT-PCR ...... 46 2.2.4.7.1. Primer Design ...... 46 2.3. Recombinant Adenoviral Transfection of HUCPVCs ...... 47 2.3.1. Construction of Recombinant Adenoviruses ...... 47 2.3.1.1. Cloning the Genes of Interest ...... 47 2.3.1.2. Homologous Recombination in vivo in Bacteria ...... 53 2.3.1.3. Recombinant Adenoviral Plasmids Amplification ...... 54 2.3.1.4. Generating Recombinant Adenoviruses in AD-293 Packaging Cells ...... 54

2.3.1.5. Amplification of Recombinant Adenoviruses ...... 55 2.3.2. HUCPVCs Infection with Recombinant Adenoviruses ...... 55 2.3.2.1. Viral titer and Multiplicity of Infection ...... 56 2.3.3. Confirmation of mRNA Production of the Genes of Interest in Genetically modified HUCPVCs ...... 57 2.3.3.1. RT-PCR ...... 57 2.3.3.1.1. Primer Design ...... 57 2.3.4. Verification of the Production of the Proteins of Interest by Modified HUCPVCs.58 2.3.4.1. Western Blots ...... 58 2.3.4.2. ELISA ...... 59 2.3.5. Longevity of GFP Expression by HUCPVCs in Different Growing Conditions ...... 60 2.3.6. Bioactivity Test of BMP2 Produced by BMP2- HUCPVCs ...... 60 2.3.6.1. C2C12 Cell Culture and Assay ...... 61 2.3.6.2. Alkaline Phosphatase Assay ...... 61 2.3.6.3. Protein Assay ...... 62 2.3.7. Effect of Liquid Nitrogen Storage on Genetically Modified HUCPVCs ...... 62 2.3.7.1. Cryopreservation of HUCPVCs...... 63 2.3.7.2. Bioactivity of BMP2 Produced by Post-Liquid Nitrogen BMP2-HUCPVCs vs. Pre-Liquid Nitrogen BMP2-HUCPVCs...... 63 2.3.7.3. Comparison of Efficiency of Protein Production Between Pre- and Post-Liquid Nitrogen VEGF-HUCPVCs ...... 63

2.4. In vitro Evaluation of Cocktails of Genetically Modified HUCPVCS ...... 64 2.4.1. Cell Culture of Different Groups and Cocktails of Genetically Modified HUCPVCs64 2.4.2. ALP Staining and Quantification ...... 65 2.4.3. Von Kossa Staining ...... 66 2.4.4. RNA Extraction ...... 66 2.4.5. Quantitative RT-PCR ...... 66 2.5. In Vivo Osteoinductive and Osteogenic Potential of Intramuscular Injected Genetically Modified HUCPVCs ...... 67 2.5.1. Animal Subjects ...... 67 2.5.2. Intramuscular Cell Delivery ...... 68

2.5.3. X-rays ...... 69 2.5.4. Micro-computed Tomography ...... 69 2.6. rhFurin Cleavage of Pro-BMP2 secreted by BMP2-HUCPVCs...... 69 2.6.1. HUCPVCs, rBMSC and hBMSCs BMP2-adenoviral Infection ...... 69 2.6.2. ELISA ...... 70 2.6.3. Treatment of BMP2-HUCPVCs Conditioned Media with rhFurin ...... 70 2.6.3.1. ELISA ...... 70 2.6.3.2. Western Blot ...... 70 2.6.3.3. Test of bioactivity of Cleaved BMP2...... 71 2.7. Osteogenic Cultures of Genetically Modified Human Bone Marrow Stromal Cells (HBMSCs) vs. HUCPVCs ...... 71 2.7.1. HBMSCs and HUCPVCs Culture ...... 71 2.7.2. HBMSCs and HUCPVC infection ...... 72 2.7.3. Osteogenic cultures of genetically modified HBMSCs and HUCPVCs ...... 72 2.7.4. ALP staining and Quantification ...... 72 2.7.5. Alizarin Red Staining ...... 72 2.7.6. Calcium Content Quantification ...... 73 2.7.7. Osteogenic Cultures in Naive and Engineered HBMSCs ...... 73 2.8. Statistical Analysis ...... 74

3. Results ...... 75 3.1. Genetic modification of HUCPVCs by non-viral methods...... 75 3.1.1. FuGENE 6 ...... 75 3.1.2. Lipofectamine LTX ...... 76 3.1.3. Nucleofection ...... 77 3.1.3.1. Longevity of GFP expression after HUCPVCs nucleofection ...... 78 3.2. HUCPVCs retroviral transduction ...... 79 3.2.1. Effects of Runx2 over-expression in HUCPVCs ...... 80 3.3. Recombinant adenovirus fabrication ...... 82 3.4. Recombinant Adenoviral Infection of HUCPVCs ...... 84

vii

3.4.1. Production of the transcripts of the genes of interest by genetically modified HUCPVCs ...... 85 3.4.2. Production of the Proteins of Interest by Genetically modified HUCPVCs ...... 86 3.4.3. Longevity of GFP expression in HUCPVCs after gene transfection with recombinant adenoviruses...... 88 3.4.4. Bioactivity of the BMP2 produced by BMP2-HUCPVCs ...... 89 3.4.5. Effects of cryopreservation on protein production and bioactivity ...... 90 3.4.5.1. Bioactivity of BMP2 produced by pre- and post-liquid nitrogen BMP2- HUCPVCs. 90 3.4.5.2. Production of VEGF by pre- and post-liquid nitrogen VEGF-HUCPVCs ...... 91 3.4.6. Effects of over-expression of BMP2, Runx2 and SP7 in HUCPVCs...... 92 3.4.7. Effects of the intramuscular delivery of genetically modified HUCPVCs...... 99 3.5. Furin Cleavage of Pro-BMP2 produce by BMP2-HUCPVCs ...... 100 3.5.1. Production of mature BMP2 by different cell types after modification with Ad- BMP2 101 3.5.2. rhFurin cleavage of Pro-BMP2 produced by BMP2-HUCPVCs ...... 101 3.5.3. Bioactivity test of BMP2-HUCPVCs conditioned media after rhFurin treatment 102 3.6. Osteogenic Cultures of Genetically Modified Human Bone Marrow Stromal Cells (HBMSCs) vs. Genetically Modified HUCPVCs ...... 103 3.6.1. Recombinant adenoviral infection of HBMSCs and HUCPVCs ...... 103 3.6.1. Effects of over-expression of Runx2 and SP7 in HBMSCs vs. HUCPVCs...... 104 3.6.2. Osteogenic cultures in naive and engineered HBMSCs ...... 110

4. Discussion ...... 112 4.1. HUCPVCs are genetically modifiable by non- viral and viral methods...... 112 4.2. Cryopreservation does not have an effect on production efficiency and bioactivity of proteins of interest produced by genetically modified HUCPVCs...... 115 4.3. HA tagged BMP-2 produced by BMP2-HUCPVCs is not properly processed into mature-BMP2...... 117 4.4. Overexpression of Runx2 and SP7 causes upregulation of different osteogenic markers but does not have an effect on matrix mineralization in HUCPVCs in vitro or in vivo. 121

viii

4.5. Overexpression of Runx2 or SP7 enhances ALP activity and matrix mineralization in HBMSCs...... 125 4.6. Conclusions ...... 128 4.7. Future Directions ...... 128 References ...... 131 Appendix 1 ...... 156 Appendix 2 ...... 159

List of Tables

Table 2.1 – PCR Primers for Osteoblastic Genes ...... 47 Table 2.2 – Primers Used to Amplify Human cDNA and to Introduce the BglII and XhoI Restriction Sites at the 5’ and 3’ ends of the Amplicons Respectively...... 52 Table 2.3 – PCR temperature conditions for the amplification of the genes of interest...... 52 Table 2.4 – Primers for the Specific Detection of Transcripts Produced as a Result of The Genetic Modification with Recombinant Adenoviruses ...... 58 Table 2.5 – Quantitative RT-PCR Primers for Osteogenic Genes ...... 67

Table 3.1 – Conditions and Efficiencies of HUCPVCs Transfection with FuGENE 6...... 75 Table 3.2 – Conditions and Efficiencies of HUCPVCs Transfection with Lipofectamine LTX...... 76 Table 3.3 – Programs, Transfection Efficiencies and Cell Viability of HUCPVCs Nucleofection. . 77 Table 3.4 – Recombinant Adenoviruses Constructed Using the AdEasy XL Adenoviral Vector System...... 83

List of Figures

Figure 1.1 - Human Umbilical Cord Perivascular Cells (HUCPVCs)...... 7 Figure 1.2 - BMP-2 Processing...... 14 Figure 1.3 - Bone Morphogenetic Protein Receptor Binding and Smads Signal Transduction. .. 15 Figure 1.4 - Vascular Endothelial Growth Factor (VEGF) Receptor Binding and Signal Transduction...... 22

Figure 2.1 – Production of Recombinant Adenovirus Using the AdEasy XL Adenoviral Vector System ...... 49 Figure 2.2 – (A) pShuttle-IRES-hrGFP-2 map. (B) Multiple Cloning Site Region (MCS)...... 50 Figure 2.3 – Electrophoresis (A) pShuttle-IRES-hrGFP-2 (B) PCR Amplified Runx2, SP7, VEGF and (C) BMP2...... 50 Figure 2.4 – Intramuscular Delivery of Genetically Modified HUCPVCs ...... 68

Figure 3.1– Fluorescence Microscopy of HUCPVCs Transfected with FuGENE 6 ...... 75 Figure 3.2 – HUCPVCs Transfection with Lipofectamine LTX...... 76 Figure 3.3 – Microscopy of HUCPVCs Transfected with Lipofectamine LTX ...... 77 Figure 3.4 – Longevity of GFP Expression after HUCPVCs Nucleofection...... 78 Figure 3.5 – HUCPVCs Retroviral Transduction...... 79 Figure 3.6 – Runx2 Immunofluorescence...... 80 Figure 3.7 – Alkaline Phosphatase Staining after Retroviral Transduction of HUCPVCs...... 81 Figure 3.8 – Gene Expression Detected by RT-PCR after Retroviral Transduction with Runx2.. 82 Figure 3.9 – Screening of Seven Different Clones of pBMP2-rAV After the Recombination Process...... 84 Figure 3.10 – Genetically Modified HUCPVCs Expressing GFP after Recombinant Adenoviral Infection...... 85 Figure 3.11– Flow Cytometry of HUCPVCs after Recombinant Adenoviral Infection...... 85 Figure 3.12 – RT-PCR of the Genes of Interest after Recombinant Adenoviral infection...... 86 Figure 3.13 – Western Blots of the Protein of Interest...... 87 Figure 3.14 – ELISA Result of the Secreted Factors of Interest...... 88 Figure 3.15 – Longevity of GFP Expression under Different Growing Conditions after Recombinant Adenoviral Infection of HUCPVCs...... 89 Figure 3.16 – Bioactivity Test of BMP2 Secreted by BMP2-HUCPVCs...... 90 Figure 3.17 – Bioactivity Test of BMP2 Produced by pre- vs. post-Liquid Nitrogen BMP2- HUCPVCs...... 91

Figure 3.18 – VEGF Production Over Time by pre- and post-Liquid Nitrogen VEGF-HUCPVCs. .. 92 Figure 3.19 – ALP Histochemical Staining of Genetically Modified HUCPVCs...... 93 Figure 3.20 – Measurement of Alkaline Phosphatase Activity in BMP2-, Runx2- and SP7- HUCPVCs...... 94 Figure 3.21 – Quantification of Alkaline Phosphatase Activity in Cocktails of Genetically Modified HUCPVCs...... 95 Figure 3.22 – Quantification of Alkaline Phosphatase Activity of Genetically Modified HUCPVCs...... 95 Figure 3.23 – ALP mRNA Expression in Genetically Modified HUCPVCs...... 96 Figure 3.24 – Collagen 1A1 mRNA Expression in Genetically Modified HUCPVCs...... 97 Figure 3.25 – Osteocalcin mRNA Expression in Genetically Modified HUCPVCs...... 98 Figure 3.26 – Osteopontin mRNA expression in genetically modified HUCPVCs...... 99 Figure 3.27– X-rays of the Region of Interest after Intramuscular Delivery of HUCPVCs...... 100 Figure 3.28 – Treatment of BMP2-HUCPVCs Conditioned Media with rhFurin ...... 102 Figure 3.29 – Bioactivity Test of BMP2 Cleaved by rhFurin...... 103 Figure 3.30 – Genetically Modified HBMSCs and HUCPVCs Expressing GFP after Recombinant Adenoviral Infection...... 104 Figure 3.31 – Flow Cytometry of HBMSCs and HUCPVCs after Recombinant Adenoviral Infection...... 105 Figure 3.32 – Alkaline Phosphatase Staining in Modified HBMSCs and HUCPVCs...... 106 Figure 3.33 – Alkaline Phosphatase Activity Measurement in Genetically Modified HBMSCs and HUCPVCs...... 107 Figure 3.34 – Comparison of Alkaline Phosphatase Activity between HBMSCs and HUCPVCs. 108 Figure 3.35 – Alizarin Red Staining of Engineered HBMSCs and HUCPVCs...... 109 Figure 3.36 – Calcium Content Quantification in Modified HBMSCs Osteogenic Cultures at Day 21...... 110 Figure 3.37 – Alizarin Red Staining of Engineered and Naive HBMSCs...... 111

Figure 4.1– Western Blots from BMP2-HUCPVCs Conditioned Medium and Cells Lysate...... 118

xii

List of Abbreviations

Ad-BMP2 BMP-2 adenovirus Ad-Mock Mock adenovirus Ad-Runx2 Runx2 adenovirus Ad-SP7 SP7 adenovirus Ad-VEGF VEGF adenovirus ADSC Adipose derived stomal cell ALP Alkaline phosphatase β2M Beta-2-microglobulin BMP-2 Bone morphogenetic protein 2 BMP2-HUCPVC HUCPVC modified with Ad-BMP2 BMPs Bone morphogenetic proteins BSA Bovine serum albumin BSP Bone sialoprotein CFU-F Colony-forming unit-fibroblast COL1A1 Collagen type I alpha I ECM Extracellular matrix ELISA Enzyme-Linked ImmunoSorbent Assay FBS Fetal bovine serum FGF Fibroblast growth factor GFP Green fluorescent protein HBMSC Human bone marrow stromal cells HUCPVCs Human umbilical cord perivascular cell IGF Insulin growth factor MDSC Muscle derived stromal cell Mock-HUCPVC HUCPVC modified with Ad-Mock MSC Mesenchymal Stromal Cell LN Liquid nitrogen OCN Osteocalcin OPN Osteopontin Osx Osterix PBS Phosphate buffered saline PDGF Platelet-derived growth factor qRT-PCR Quantitative PCR rhBMPs Recombinant human bone morphogenetic proteins rhfurin Recombinant human furin RT-PCR Reverse transcription polymerase chain reaction Runx2 Runt-related transcription factor 2 Runx2-HUCPVC HUCPVC modified with Ad-Runx2 SP7 SP7 transcription factor (Osterix) SP7-HUCPVC HUCPVC modified with Ad-SP7 TBS Tris Buffered Saline TGF-β transforming growth factor–β VEGF-A Vascular endothelial growth factor-A VEGF-HUCPVC HUCPVC modified with Ad-VEGF

xiii

1. Introduction

1.1. Bone and Osteogenesis

Bone is a specialized connective tissue with several important functions in the human body. The skeleton provides structural and mechanical support for locomotion while bones such as the ribs, skull, and vertebrae also provide protection to vital organs including the heart, lungs, brain, and spinal cord. Bone also acts as a reservoir of calcium, phosphate ions and growth factors, and its marrow cavity serves as housing and factory for haematopoiesis (1–3).

Bone tissue consists of both inorganic (70%) and organic components (30%) (4). The organically synthesized matrix is primarily made of type I collagen (95%) and multiple noncollagenous calcium binding and signaling proteins including osteopontin, bone sialoprotein, osteocalcin, and osteonectin. This organic matrix is embedded with crystalline salts, primarily calcium and phosphate, in an hydroxyapatite lattice structure for which the stoichiometric formula is

(Ca10(PO4)6(OH)2) (5), although biological apatite may contain as many as 25 atomic substitutions within this stable phosphate lattice.

Bone is a dynamic tissue that is constantly being remodeled throughout life (6). Bone remodeling is accomplished by the processes of resorption and osteogenesis (7,8). Osteoclasts, multinucleated cells of hematopoietic origin, are responsible for bone resorption through mineral dissolution and enzymatic digestion of the organic matrix (9). On the other hand, osteoblasts, cells of mesenchymal origin, are responsible for generating bone tissue by secreting matrix and mineral deposition (10). Bone formation or osteoblastogenesis is marked by four major phases: lineage commitment, proliferation, synthesis of the extracellular matrix (ECM), and mineralization (11–13). During the first stage MSCs commitment and differentiation to the osteogenic lineage is regulated by master transcription factors Runx2 and Osterix (14–18). Furthermore, different paracrine, autocrine, and endocrine factors such as bone morphogenetic proteins (BMPs), transforming growth factor–β (TGF-β), fibroblast growth factor (FGF), insulin growth factor (IGF), vascular endothelial growth factor (VEGF), vitamin D3, 1

glucocorticoids, parathyroid hormone, and estrogen modulate osteoblast differentiation (8,11,14). During the proliferative phase preosteoblastic cells express genes related with growth (histones, c-myc and c-fos) (11,13) and genes related to the formation of the ECM principally type I collagen, fibronectin and TGF-β (12,13). The third stage of osteoblastogenesis is characterized by a dramatic increment in the production of ALP, a membrane bound glycosylphosphatidylinositol protein generally recognized as one of the earliest markers of osteoblast differentiation (13). During this stage cells start to cluster and form multilayers in culture followed by a decline in DNA synthesis and histone production (11). Moreover, while cells continue differentiating they start producing non-collagenous proteins of the ECM such as osteopontin, osteonectin, bone sialoprotein, and osteocalcin (19). The culmination of the maturation process towards becoming an osteoblast is marked by the mineralization of the collagenous extracellular matrix (11,20).

Bone is one of the few organs that retains the ability to regenerate through adult life. The process of bone repair is highly complex and involves the coordination of many different cells, transcription factors and growth and differentiation factors to produce the desired response. Following bone damage, the general reparative process involves hematoma formation and inflammation, leading to development of a soft tissue callus surrounding the defect site, which is eventually mineralized and remodeled into mature lamellar bone (21). This process occurs naturally after injury to bone tissue; however, this process may be hindered by infection, lack of blood supply, fracture instability, substantial bone loss, malnutrition, smoking, diabetes and metabolic bone disorders among others (22). When the normal endogenous mechanisms are not able to restore the lost bone such as in non-union fractures, removal of bone tumors, or large-scale traumatic bone injury, clinical therapy for healing is necessary.

1.2. Need for Bone Substitutes

Bone substitutes are required for the healing of large bone defects, including orthopedic, craniofacial, and dental reconstructions. After blood, bone is the most commonly transplanted tissue (23). Per year approximately 6.5 million fractures occur in the United States and 10-15% 2

are difficult to heal (24,25). Annually about 500,000 bone grafts are performed in the U.S. (26) and 2.2 million worldwide (23). Usually bone tissue loss is treated using autografts, allografts, implantation of substitute materials and recently, using recombinant bone morphogenetic proteins (rBMPs).

In autografts bone is harvested from one site of the patient, typically from the iliac crest, and implanted into an osseous defect of the same patient. Autografts remain the gold standard for bone substitutes avoiding immunological issues and providing both an osteoconductive matrix for bone cell adhesion and the cells and bioactive molecules to induce and sustain the regenerative process (27). Although successful in many cases, this procedure is limited by the availability of healthy tissue, donor site morbidity, and pain associated with the harvest (28).

Allografts represent approximately 1/3 of all bone grafts in North America, where cadaveric bone is taken from an unrelated donor and frozen until use (27). Allografts are widely used to address the issue of donor tissue availability for autografts; however, they suffer from reduced biological and mechanical properties after processing and possible disease transmission and immunological host response (29), leading to graft failure rates as high as 50% (30,31).

Synthetic materials which represent approximately 10% of bone substitutes (32) including titanium, ceramics, and synthetic hydroxyapatite, are subject to various disadvantages such as obvious lack of osteogenic or osteoinductive component, toxicity, wear, biodegradability, elicitation of inflammatory responses, adverse host tissue necrosis or resorption due to relatively poor biomechanical compatibility and fail to adequately integrate and remodel with the surrounding native tissue (2,32,33).

Molecular therapy using rBMPs (rhBMP-2 and rhBMP7) has been approved by the FDA for human use in long bone non-unions where autograft treatment is not feasible or has failed, for lumbar spinal fusion (34,35), oromaxillofacial procedures and a variety of orthopedic disorders (36). Although the use of rBMPs in clinical repair of bone defects has been successful several negative factors contribute to the continued search for alternatives. For example short

systemic half-life, uncontrolled release of rBMPs from the carrier, diffusion of BMP away from the delivery site, undesired ectopic bone formation and the use of supra-physiological doses (35,37) that can lead to post therapy problems and that causes this procedure to be very costly.

Due to the limitations that accompanied all the available options of bone substitutes there is an obvious need for new alternatives that can ideally be osteoinductive, osteogenic, osteoconductive, angioinductive and allow blood vessel invasion, degrade within an appropriate time frame, and be biomechanically stable.

1.3. Bone Tissue Engineering

As an alternative approach to satisfy the need for bone substitutes tissue engineering "applies the principles of engineering and life sciences towards the development of biological substitutes that can restore, maintain or improve tissue function" (38). There are many approaches to bone tissue engineering, but all involve one or more of the following fundamental components: harvested cells, scaffolding matrix and regulatory/inductive/stimulatory signals to direct osteoblastic differentiation (39–41).

1.3.1. Cells for Bone Tissue Engineering

The ideal cell for cell-based tissue engineering should be easy to obtain and expand, be non- immunogenic, be genetically modifiable, ethically non-controversial and be capable to differentiate to the osteogenic lineage. Most of the cells currently being investigated in the bone engineering field are multipotent mesenchymal stromal cells (MSCs).

1.3.1.1. Mesenchymal Stromal Cell

Multipotent mesenchymal stromal cells (MSCs) were initially described by Friedenstein and co-workers as a subpopulation of bone marrow cells capable to differentiate to bone (42,43). Later they developed a method to isolate fibroblast-like cells from the marrow based on their ability to adhere to tissue culture plastic (44), these adherent cells capable of forming colonies when plated at low density were termed colony-forming unit-fibroblast (CFU-F) (45). Afterwards the same group showed for the first time the multipotential of these cells to produce bone and cartilage when implanted intraperitoneally in diffusion chambers (46). Years later, Caplan (47) gave these cells the name, Mesenchymal Stem Cells and described them as cells capable of differentiating to all cells of the mesodermal lineage (48).

Over the past years new sources of MSCs have been reported including umbilical cord blood (49), dental pulp (50), synovial membrane (51), adipose tissue (52), placenta (53), Wharton’s jelly (54), scalp tissue (55), umbilical cord perivascular cells (56), skeletal muscle perivascular cells (57), amniotic fluid (58), and breastmilk (59). Due to different sources of MSCs, terminology and diverse methods of isolation and characterization of the cells it has been difficult to compare the experimental outcomes among studies having a negative impact in the MSC field. To help standardized the characterization of MSC and facilitate the exchange of data the International Society for Cellular Therapy (ISCT) has proposed the minimal criteria to define human multipotent mesenchymal stromal cell: “First, MSC must be plastic-adherent when maintained in standard culture conditions. Second, MSC must express CD105, CD73 and CD90, and lack expression of CD45, CD34, CD14 or CD11b, CD79a or CD19 and HLA-DR surface molecules. Third, MSC must differentiate to osteoblasts, adipocytes and chondroblasts in vitro” (60).

In addition to this minimal set of standards it is important to highlight that MSCs have been reported to be immunoprivileged and immunosuppressive (61–64) reason why they can be used as an allogeneic source of cells for different therapies. In fact, last year Osiris Therapeutics (http://www.osiris.com) a US-based MSC company received authorization from

Health Canada to market MSCs as a stem cell therapy for the treatment of acute graft-versus host disease in children with their product "Prochymal" based in human bone marrow mesenchymal stromal cell.

1.3.1.1.1. Bone Marrow Stromal Cells

Bone marrow stromal cells (BMSCs) are the most common source of MSCs and are the main cells used in bone tissue engineering. Although BMSCs have been used successfully in the healing of bone defects (65–68), the harvest of bone marrow requires an invasive procedure and the yield of CFU-F is low, approximately 1:10,000 at birth diminishing to 1:250,000 in adult marrow (69). Moreover it has been demonstrated that the osteogenic capacity of the cells decreases with the age of the donor (70) and extensive in vitro culture to obtain sufficient cell numbers for therapeutic applications causes morphology changes, reduction in proliferation rate, and loss of osteoblastic differentiation capacity (71–73) which sometimes limit their usefulness. Therefore there is a need for alternative MSCs sources that can overcome these problems by providing higher yield of MSCs and easier acquisition and faster expansion.

1.3.1.1.2. Human Umbilical Cord Perivascular Cells

Human Umbilical Cord Perivascular Cells (HUCPVCs) are obtained by the dissection of the vessels from the umbilical cord, followed by the digestion of the perivascular tissue (Figure 1.1) (56). HUCPVCs lack the expression of CD45, CD34, MHC-II and express CD73, CD105, CD90, CD44, plus the pericyte markers 3G5, CD146 and NG2 (56,74). They are capable of osteogenic, chondrogenic, adipogenic, and myogenic differentiation in vitro (74,75) and they have been shown to contribute to the healing of bone and cartilage (75), skin (76) and tendon (77). To date, HUCPVCs have shown many similarities with human BMSCs in terms of differentiation capacity, surface markers (74) and non-immunogeneic and immunomodulatory phenotype (78–80) while exhibiting a higher proliferation rate (74) and a greater CFU-F frequency (1:333)

(56) when compared to BMSCs (1:10,000 in newborns) (69). Moreover, HUCPVCs are easy and safe to obtain since they are harvest from a tissue that is normally discarded after birth and are ethically non-controversial. Altogether the previous mentioned characteristics indicate that HUCPVCs could be a suitable allogeneic source of cells for cell-based tissue engineering.

Figure 1.1 - Human Umbilical Cord Perivascular Cells (HUCPVCs). (A) Scanning electron microscopy of the umbilical artery excise from the umbilical cord. The white dotted line delineates the perivascular tissue from which the HUCPVCs are harvested. (B) HUCPVCs morphology in vitro is fibroblastic.

1.3.2. Osteoinductive Factors in Bone Tissue Engineering

Several approaches have been used to induce cell differentiation to the osteogenic lineage, the most commonly utilized is the exposure of MSCs to dexamethasone and other factors in vitro; however it has been suggested that these ex vivo manipulations will likely not be sustained after in vivo implantation (81). The delivery of soluble molecules that initiate signaling cascades to promote osteogenesis have primarily focused on BMPs (82) especially BMP-2 and BMP-7 since they have been approved by the FDA for different clinical applications (34–36) but other growth factors have also been examined in the healing of bone defects, including platelet-derived growth factor (PDGF) (83,84), fibroblast growth factor (FGF) (85) and insulinlike growth factor (IGF) (86). Although these approaches have been successful, high doses of bioactive factors are often necessary because of their short biological half-lives (33)

and the lack of optimal matrices for controlled and sustained delivery limiting the benefits of this approach (81).

As alternative strategy gene therapy can be implemented to deliver therapeutic proteins via appropriate target cells in a more physiological and consistent manner when compare to the previous mentioned approaches (25) and can also be applied to improve the osteogenic potential of the chosen cells for bone tissue engineering .

1.4. Orthopedic Gene therapy

Originally, gene therapy was developed as a strategy to correct genetic diseases caused by single gene mutations (87,88), however, its potential is recently being investigated in the orthopedic field to promote the healing of bone defects by inducing the local expression of genes that are involved in the regenerative process (89–94). Gene therapy provides an alternative to protein therapy (e.g rhBMPs, rhPDGF) and also allows the expression of transcription factors, receptors and other intracellular proteins as therapeutic agents that otherwise could not be delivered to the defect site.

Gene therapy requires key steps to occur correctly in order to produce successfully a functional protein: transduction/transfection, known as the process by which the gene of interest is introduced via a vector into the target cell; transcription, step in where the DNA template is copied into messenger RNA (mRNA); translation, process in where the protein is synthesized from the mRNA template (91,95,96); and finally, in some cases the produced protein requires post-translational modifications (e.g glycosylation, cleavage) to insure its full biological activity (97).

Orthopedic gene therapy can be performed in vivo or ex vivo. In the in vivo approach the vector is directly delivered to the bone defect while in the ex vivo approach, strategy chosen for this thesis work, autologous or allogeneic cells are expanded in tissue culture and genetically manipulated in vitro to subsequently be implanted into the bone defect (90,92,98). 8

Although in vivo gene therapy is technically more simple and less expensive due to the cell-free approach, ex vivo gene therapy allows higher gene transfer efficiency and target specificity and obviates many of the safety concerns related to direct exposure to some gene vectors (89– 91,96).

1.4.1. Gene delivery vectors

Vectors are the tools used to deliver the therapeutic gene to the target cell. Ideally, gene delivery method should be non-toxic, non-immunogenic, cost effective, easy to produce, have higher transfection/transduction efficiency, have the ability to allow external control of protein expression, mediate the desired level of gene expression during an appropriate length of time, transduce/transfect dividing and non-dividing cells and integrate the gene into the target cell genome without disruption of normal cell function (91,93,99). Although some of the current vectors partially fulfill the above criteria, the ideal vector has yet to be developed. The existing variety of vectors that are currently use for gene transfer have their own advantages and limitations. They can be divided in non-viral and viral methods.

1.4.1.1. Non- viral Vectors

Non-viral vectors use physical or chemical methods to insert the therapeutic gene into the target cells (94). These methods offer several advantages like easy manufacturing and use, low production cost, no or low immunogenicity, no risk of transmission of infectious diseases and flexibility towards the molecular size of loaded DNA. But they also present limitation such as transient gene expression and lower transfection efficiency compared to viral vectors (93,94). Most studies of gene therapy for bone regeneration using non-viral methods have been conducted with liposomal vectors and Nucleofection.

Liposomes are formed by self-assembly of cationic lipids that have polar heads and non-polar tails (93). The positively charged liposomes interact with the negatively charge molecules of

DNA protecting them against degradation, creating positively charged particles, and facilitating internalisation, intracellular trafficking, and processing (91). The positively charged DNA-liposome complex fuses with the negatively charged cell membrane entering the target cell by endocytosis, then the DNA is released into the cytoplasm as a result of the destabilization of the endosomal membrane (100). Nowadays different lipid-based transfection agents are commercially available, like FuGENE™ 6 and Lipofectamine™ used in this thesis work. All specific formulations of such lipid-based transfections reagents are proprietary to the distributing suppliers.

Nucleofection is a trademark that combines cell-type specific reagents with the use of pulsed electrical fields to form holes in the plasma membrane in order to increase DNA diffusion and direct DNA delivery into the nucleus (101). This technique has been successfully applied for the transfection of hard-to-transfect cell lines and primary cells, including human MSCs as applied in this thesis work. Transfection efficiencies achieved by this technique can be higher than the ones obtained by liposomal transfections but with lower cell viability (102).

1.4.1.2. Viral Vectors

Viral vectors are considered to be the most efficient vectors for gene transfer but they are more complicated to manufacture and they raise greater safety concern. Viruses used as gene therapy vectors are genetically modified to disrupt the native propagation machinery of the virus, making them replication incompetent (90). The viral vectors that are used the most in ex vivo gene therapy are adenoviruses, retroviruses and lentiviruses, each of them with different advantages and limitations.

Adenoviruses contain a linear, double-stranded DNA packaged within a nonenveloped icosahedral capsid (98). Adenoviruses enter the cell by receptor mediated endocytosis by binding to the coxsackie/adenovirus receptor (CAR) which is broadly distributed among most host cells (98,103). After infection, adenoviruses do not normally integrate into the host genome; instead, they remain in the nucleus as an episome which is not replicated during cell 10

division resulting in the expression of the gene only for a limited period of time (104). Adenoviral vectors derived from human serotype 5 are the most popular vectors for ex vivo gene therapy aimed at bone regeneration (91,99) and the vector of choice for the major part of this thesis work. They are attractive delivery vehicles due to their capacity to infect a broad range of dividing and non-dividing cell types with high efficiency, they can be produced in high titer and as they do not integrate into the host genome they do not carry the risk of disrupting endogenous gene function (98,105). The primary disadvantage of adenoviral vectors is a potential host immune response, which targets the cells transfected by the virus (94).

Retroviruses are enveloped viruses that enter the cell through receptor-ligand and membrane fusion events. After internalization, viral RNA is reverse transcribed by the viral reverse transcriptase into double stranded DNA that is randomly integrated into the genome and then replicated as the target cell divides (106–108). Retroviruses can infect many cell types at a high efficiency but do not infect non-dividing cells and production of high titers is challenging (94,98). Also the random integration in the host genome may disrupt a host gene at the site of incorporation or cause abnormal expression of nearby host genes affecting cell function by insertional mutagenesis limiting its application (109,110).

Lentiviruses are a specialized family of retroviruses that are capable of infecting non-dividing cells (111). Even though lentiviruses integrate into the host cell genome, there is evidence that the integration sites are significantly more limited than other retroviral family classes (112). The biggest limitation of using this vector is the safety concerns due to the lethal nature of many of the original viruses used to develop lentiviral vectors being human immunodeficiency virus 1 (HIV-1) the most common example (94,108).

1.4.2. Target Genes for Orthopedic Gene therapy

Many genes that are involved in the formation and regeneration of bone can be considered for gene therapy applications. During the past years different genes have been used to enhance bone formation in vivo with varying results, some of these genes encode for soluble factors like BMP-2 (113,114) , BMP-4 (115,116), BMP-6 (117), BMP-7 (118,119), BMP-9 (120), FGF-2 (121) , VEGF-A (122), TGF-β (123), IGF-1 (124), PDGF (125) and RANK-L (126); transcription factors like Runx2 (127) and Osterix (128); LIM mineralization protein-1 (LMP-1) (129) and LMP-3 (130); parathyroid hormone (PTH) (131); and the constitutively active form of the BMP receptor Alk-2 (132).

For the experimental work of this thesis two soluble factors (BMP-2 and VEGF-A) and two transcription factors (Runx2 and osterix) were chosen. Further information is provided in the next sections.

1.5. Bone Morphogenetic Protein -2

Bone morphogenetic proteins (BMPs) are a group of phylogenetically conserved signaling molecules — members of the transforming growth factor β (TGF-β) superfamily of secreted growth and differentiation factors (133,134). The discovery and characterization of BMPs began when Marshal Urist reported the induction of bone formation when implanting demineralized bone matrices into ectopic sites in rabbits and rodents (135). Of the 20 members of the BMP family, BMP-1 through BMP-7 are expressed in skeletal tissue, and BMP- 2, -4 and -6 are the most detectable in osteoblast cultures (133). BMP-2 is the most thoroughly evaluated BMP and its clinical use was approved by the FDA in 2002. Thus, although other BMP members have osteogenic potential, BMP-2 is the growth and differentiation factor of choice for this thesis work, and a detailed description of its mode of action is given below.

BMP-2 has been shown to recruit osteoblasts progenitors, induce osteogenesis, and contribute to the maintenance of the osteoblastic phenotype (136). BMP-2 induced osteoblastic

differentiation is associated with high levels of ALP activity, increased production of extracellular matrix proteins including type I collagen, bone sialoprotein and osteocalcin (137), upregulation of transcription factors such as Osterix and Runx2 (138), and increased responsiveness to calcitropic hormones such as parathyroid hormone (PTH) and lα,25- dihydroxyvitamin D3 (139) .

1.5.1. BMP-2 Synthesis and Posttranslational Modification

BMP-2 is synthesized as a pre-pro-protein which consists of an amino terminal signal peptide, a pro-domain and a carboxyl terminal mature protein (140,141). For BMP-2 synthesis, the DNA encoding BMP-2 is first transcribed into mRNA in the nucleus and then the pre-sequence mediates translocation into the lumen of the endoplasmic reticulum (ER) where BMP-2 is translated into amino acids to form the BMP-2 precursor (141). Following cleavage of the signal peptide, pro-BMP-2 undergoes glycosylation and dimerization in the Golgi apparatus (140,142). For the final step to become a mature bioactive protein known to have 144 amino acids and a molecular mass of 18kDa (143), pro-BMP-2 is cleaved on the C-terminal side of the proprotein convertase recognition sequence (R-X-X-R) by proteases that belong to the proprotein convertase family such as the serine endoprotease furin (134,144) (Figure 1.2). Once secreted from the cell, the active BMP-2 dimer binds to BMP-2 receptors and activates its signaling cascade.

1.5.2. Furin in BMP-2 Cleavage

Proprotein convertases (PC) are secretory mammalian serine proteinases responsible for proteolytic cleavage of BMPs. The family of PCs comprises nine members, PC1/3, PC2, furin, PC4, PC5/6,PACE4, PC7, SKI-1/S1P, and PCSK9 (145). One of the most studied and characterized member of the PC family is furin, a calcium-dependent serine protease that acts on proteins in the trans-Golgi network (TGN), secretory granules, cell surface, or endosomes (145,146). Furin is ubiquitously expressed at various levels in all tissues, having a widespread

role in the processing of multiple precursors including growth factors and their receptors, proteases and adhesion molecules (145,146). Furin processes pro-proteins at the dibasic sequence motif R-X-X-R, which is usually found at the C-terminal of the pro-domain (147). When the third AA is either an arginine (R) or a lysine (K), the sequence R-X-K/R-R provides an optimum processing site (147). It is well known that furin can cleave pro-BMP2 at the optimal site R-E-K-R (134).

Figure 1.2 - BMP-2 Processing. BMP-2 is synthesized as a pre-pro-protein that consists of an amino terminal signal peptide, a pro-domain and a mature-domain. In the first step the signaling peptide is cleaved to form pro-BMP2. Then, two pro-BMP2 monomers dimerize to form the pro-BMP2 dimer. Finally pro-domains are cleaved to form the mature BMP2 dimer which is then secreted into the extracellular space. Image modified from Zhou et al. (148).

1.5.3. BMP-2 Signaling Cascade

BMP-2 forms a disulfide linked homodimer that binds to its cell surface receptor and can activate its signaling cascade via two independent pathways (149). In the first pathway, Smad- dependent, BMP-2 binds a heterodimeric complex of type I and type II transmembrane serine/threonine kinase receptor proteins (BMP-2RI and BMP-2RII), inducing autophosphorylation of the intracellular domain, which, in turn, phosphorylates one of the receptor-regulated Smad proteins (R-Smads : Smads 1, 5 and 8) (133). Once phosphorylated by the type I receptor, the R-Smads associate with Co-Smad (Smad4) forming the complex R- 14

Smads/Co-Smads that then translocates to the nucleus and regulates transcription of target genes by interacting with various transcription factors such as Runx2 and transcriptional coactivators or co-repressors (133,150) (Figure 1.2). In the second pathway, Smad independent, BMP-2 binds to BMP-2RII which then recruits BMP-2RI, initiating a phosphorylation cascade via the p38 mitogen activated kinase (MAPK) pathway possibly mediated by Tak1/Tab1 (134,149,151).

Figure 1.3 - Bone Morphogenetic Protein Receptor Binding and Smad Signal Transduction. BMP-2 binds a heterodimeric complex of type I and type II transmembrane serine/threonine kinase receptor proteins (BMP-2RI and BMP-2RII), inducing autophosphorylation of the intracellular domain, which, in turn, phosphorylates one of the receptor-regulated Smad proteins (R-Smads) (133). Once phosphorylated by the type I receptor, the R-Smads associate with Co-Smad forming the complex R- Smads/Co-Smads that then translocates to the nucleus and regulates transcription of target genes by interacting with various transcription factors and transcriptional coactivators or co-repressors (133,150). Extracelular interaction of BMP-2 can be blocked by noggin, chordin, gremlin and follistatin. (152,153). Smurf1 can block R-Smad signaling by targeting their degradation (154,155). Also, the inhibitory Smads (I-Smads) physically preventents the interaction between BMP-2RI and R-Smads inhibiting subsequent activation (156).

1.5.4. BMP-2 Regulators and Inhibitors

BMP-2-induced osteogenesis has several known points of regulation. At the extracellular level the proteins noggin, chordin, gremlin and follistatin can block the extracellular interaction of BMPs with their receptors by competitive dimerization (152,153). At the intracellular level, the protein Smurf1 can block Smad1 and Smad5 signaling by targeting their degradation (154,155). Also, the inhibitory Smads (I-Smads: Smad6 and Smad7) block BMP-2 signaling by physically preventing the interaction between BMP-2RI receptor and R-Smads inhibiting subsequent activation (156).

1.5.5. BMP-2 ex vivo Gene Therapy

Therapy using recombinant bone morphogenetic proteins (rhBMPs) is FDA approved. Nevertheless, the short biological half-life of rhBMPs and lack of optimal matrices for controlled and sustained delivery result in high and expensive doses to achieve healing improvement, which limits the usefulness of these proteins. In the search to overcome these limitations new strategies are being explored. Over the past years BMP-2 ex vivo gene therapy has been widely studied and encouraging results have been reported. The success of strategies using BMP-2 engineered cells in vivo is based in their capacity to perform as potent inducers of bone formation, as constant secreted BMP-2 stimulates the differentiation of native osteoprogenitors that can help in the bone formation and repair process (157). BMP-2 ex vivo gene therapy has been extensively used to induce bone formation in different anatomical sites and although diverse gene delivery strategies such as Nucleofection (158), liposomes (159) and lentiviral infection (160), have been explored successfully, the approach that has shown the most promise for BMP2 ex vivo therapy is the implantation of MSCs genetically modified using adenoviral vectors (Ad-BMP2) (161). For example, Lieberman et al., (113) were one of the firsts to report the use of BMP2-producing cells to heal bone defects and since then have extensively investigated the usefulness of this method. In one of their first in vivo studies, rat bone marrow stromal cells were genetically modified with Ad-BMP2 and 5 × 106 cells were delivered in demineralized bone matrix into a 8mm segmental femoral defect in rats. After 12 16

weeks, radiological and histological analysis revealed that BMP2-BMSCs were able to heal the defect while control groups (LacZ-BMSCs, naive-BMSCs and carrier alone) showed little to no bone formation. Moreover, they compared bone formation in defects treated with BMP2- BMSCs and defects treated with a clinical dose of rhBMP-2 and found that defects treated with BMP2-BMSCs were filled with denser trabecuar bone. Similar results were reported in a later study by the same group (114) in where human adipose tissue derived mesenchymal stromal cells (ADSCs) were modified with Ad-BMP2 and delivered in a collagen–ceramic carrier into a 6mm critical size femoral defect. After 8 weeks, they found that BMP2-ADSCs were able to heal the defect and when compared to defects treated with rhBMP2 no statistically significant difference was detected between the extent of healing in both groups. After successfully showing the effectiveness of BMP2 ex vivo gene therapy to heal critical size segmental defects in rats, this group also investigated the effects of this approach in spinal fusion in a rat model (162). After infection with Ad-BMP2, 5 × 106 human-derived bone marrow cells (BMSCs) were delivered in a collagen sponge into the decorticated transverse processes of L4–L5. After 12 weeks, specimens were assessed by manual palpation and radiological and histological analysis revealing solid fusion between the transverse processes of L4 and L5 in the group that was treated with BMP2-BMSCs while, no evidence of fusion was found in the control groups (LacZ- BMSCs, naive- BMSCs and collagen sponge alone) showing an evident osteoinductive effect of the BMP2 produced by the engineered cells. Years later this groups use the same spinal fusion model to study the difference of delivering rat bone marrow stromal cells producing BMP2 for short term (modified with Ad-BMP2) and producing BMP2 for long-term (modified with BMP2- Lentivirus) (160). After 8 weeks of treatment specimens were analysed by manual palpation, MicroCT and histology revealing that the group that was treated with long term BMP2- expressing cells presented solid fusions between L4 and L5 with abundant bone mass that extended to adjacent levels while the groups that were treated with short term BMP2- expressing cells or rhBMP2 presented spinal fusions but made of thin trabecular bone. Although the results obtained in this study suggest that long-term BMP2 expression might be more beneficial when compared to short-term there is a concern regarding the side effects since prolong expression of BMP2 could lead to undesired bone formation. To understand this, new studies have to be design to evaluate the long term effect of BMP2 expression in different

defect models. Altogether this group has produced robust and consistent evidence that shows that BMP2 ex vivo gene therapy using different MSCs, murine and human, efficiently induce bone healing in murine models.

In the search for understanding the effects of utilizing cells modified with different types of gene delivery vectors Park et al (159) compared the outcome of delivering rat BMSCs modified by Ad-BMP2 or Liposomes (Metafectene) in the healing of a mandible defect in rats. 1 x106 cells were seeded in a collagen sponge and delivered into a 6mm critical size defect. At different time points specimens were collected and analyzed by x-rays and histology. Data indicated that the defects treated with ad-BMP2 modified-cells were filled with new bone after 4 weeks while the defects treated with liposome-BMP2 modified-cells were filled with bone after 6 weeks. In addition, it was reported that control groups (ad-LacZ modified-cells, naive cells and collagen alone) only presented minor regeneration at the margin of the defects indicating that the defect healing was only possible due to the effects of the BMP2 produced by the engineered cells. Interestingly, the results obtained in this study show that although the BMP2 produced by liposome-BMP2 modified-cells was two-fold lower than the amount produced by the ad-BMP2 modified-cells it was enough to induce the healing of the defect at a later time point. This might indicate that for certain defects the transient production of small doses of BMP2 by the engineered cells is enough to accomplish bone healing.

Other groups have investigated the use of cells different than MSCs for BMP2 ex vivo gene therapy. For example Gugala et al (163) investigated if the osteoinductive effect of BMP2 overexpressing cells was independent of cell type by comparing the amount of heterotopic bone formation among different type of cells. For this study human BMSCs, primary human skin fibroblasts, and the human diploid fetal lung cell line (MRC-5) were genetically modified with an adenovirus encoding the BMP2 gene; 5x106 cells were injected in 100 µl of PBS into the quadriceps muscle of NOD/SCID mice. After 14 days radiographic analysis showed that considerable amounts of heterotopic bone were formed in the muscle of the limbs injected with the different BMP2 expressing cells while no bone formation was detected in the muscles in where the LacZ-cells were injected. After quantification of bone formation no statistically

significant difference was found between the amount of formed bone by the different BMP-2 expressing cells. Interestingly, immunohistochemical staining for human mitochondrial protein was performed in the area in where BMP2-BMSCs were injected only detecting human cells at days 2, 3 and 5 after injection while on days 6 and 7, no human cells could be found in the sections, indicating that mainly host cells were responsible for the new bone formation. This results suggest that since the cells were no delivered seeded in a scaffold they were not retained in the anatomical site of interest for too long, however the BMP2 secreted by this different cells for the initial days of the study was enough to elicit ectopic bone formation.

Based in the success of ex vivo gene therapy in small animal models it only seems logical to start exploring its effects in larger animals. In view of this, Chang et al (164), investigated the effect of BMP2 expressing cells in skull defects in miniature swine. Autologous BMSCs were genetically modified with Ad-BMP2 and 3 X108 cells were delivered into a 4 cm defect in two layers of Pluronic F127 separated by a gelatin/tricalcium/phosphate/ceramic/glutaraldehyde scaffold, after 6 months the animals were euthanized and the specimens were analyzed by MicroCT, biomechanical test and histology. The group that was treated with BMP2-BMSCs was found to achieved complete full thickness repair while incomplete bone formation was found in the centre of the defects of animals that were treated with naive cells; also statistically significant difference was found in the stiffness of the new bone formed in the group treated with BMP2-BMSCs when compared with the one treated with naive cells. In a different animal model Dai et al (165) studied the potential of autologous BMP2-BMSCs to heal segmental tibia defects in goats. BMSCs were genetically modified with Ad-BMP2 and seeded in biphasic calcined bone coated in collagen. After 5 days of in vitro incubation constructs carrying about 2×108 cells were delivered into a 2.6 cm critical size defect. 26 weeks after implantation radiographic examinations and biomechanic measurements revealed that the defects in the group treated with BMP2-BMSCs healed completely while defects in controls groups (LacZ- BMSCs, naive BMSCs and scaffold alone) failed to healed. These two studies are an indication that autologous MSCs genetically modified to expressed BMP2 transiently are sufficient to achieve the healing of critical size defects in large animals which reflects in a better way than murine models, the challenge encountered when healing bone defects in humans.

Taken together, the results obtained by BMP2 ex vivo gene therapy indicate that it could be use as a powerful therapeutic tool to induce bone formation. However, despite of these positive results BMP2 ex vivo gene therapy has not reached the clinical trial phase therefore, future research should focus in further experimentation in larger animals for longer time points to understand the long term effect of this therapy. Also, it is important to develop cost- effective strategies to culture large quantities of cells and their genetic manipulation as it would be needed for human applications.

1.6. Vascular Endothelial Growth Factor A

Vascular endothelial growth factor A (VEGF-A), normally referred to as VEGF, is a member of the VEGF family that includes VEGF-B, VEGF-C, VEGF-D and placental growth factor (PLGF) (166). VEGF is a critical growth factor for vascular development and has been proven to promote proliferation, migration, differentiation and survival of vascular endothelial cells (167). VEGF was first identified as an endothelial-specific growth factor found in the media conditioned by bovine pituitary follicular cells (168). Now it is known that is expressed by many cells including human bone marrow derived MSCs (166), macrophages, endothelial cells, smooth muscle cells, fibroblasts, hypertrophic chondrocytes (169), and osteoblasts (170,171). VEGF is inducible by hypoxia and it has been reported that different growth factors including TGF-α, TGF-β, FGF, platelet-derived growth factor, insulin-like growth factor-1, epidermal growth factor and keratinocyte growth factor, and the transcription factor Runx2 (172) play a role in the upregulation of VEGF mRNA expression (173).

Structurally, VEGF is a disulfide-bonded homodimeric glycoprotein with a molecular mass of 34–45 kDa that has receptor-binding sites at each pole of the dimer (167). The human VEGF gene undergoes alternative splicing to produce six different isoforms of VEGF encoding

121,145, 165, 189, and 206 amino acids termed VEGF-A121, VEGF-A145, VEGF-A165, VEGF-A189, 20

and VEGF-A206, respectively (174). In all the various isoforms of VEGF the amino acids that are encoded by exons 1-5 are conserved (175). The first four exons encode the signal peptide, the dimerization and glycosylation sites and the sequences recognized by VEGF receptors. Exon 5 encodes a ten amino acid sequence that contains the main site of cleavage by plasmin and matrix metalloproteinases (176,177) while exons 6-8 encode the heparin binding site and neuropilin binding site (178,179). The VEGF isoforms have distinct biological activities, such as differential interaction with heparan sulfate proteoglycans (HSPGs), neuropilin (NRP) and the extracellular matrix (ECM) (178). The two isoforms of VEGF that have received the most attention are VEGF-A121 and VEGF-A165. The latter binds to HSPGs and NRPs whereas VEGF-A121 is more freely diffusible because it does not bind to HSPGs or NRPs decreasing its retention to the cell surface and ECM (180). For these reasons VEGF-A121 is the angiogenic factor of choice for this thesis work.

Similar to most peptide growth factors, VEGF exerts its biologic effect by binding to receptors present on the cell surface (181). VEGF-Receptor 1 (VEGF-R1; flt-1) and VEGF-Receptor 2 (VEGF-R2;KDR/flk-1) are transmembrane tyrosine kinase receptors that have seven immunoglobulin-like domains in the extracellular domain (173). Upon binding of VEGF to the extracellular domain of its receptor, dimerization and autophosphorylation of the intracellular receptor occurs and a cascade of downstream proteins are activated leading to biologic effects such as proliferation, migration, and survival of endothelial cells (Figure 1.4) (181). Today VEGF-R2 is recognized as the major mediator of the mitogenic, angiogenic and permeability- enhancing effects of VEGF (173) and the binding affinity of VEGF to this receptor can be enhanced by neuropilin-1 (NRP1), a non-kinase co-receptor (182). In contrast, it has been observed that VEGFR-1 is not effective at mediating the mitogenic signal and is thought to perform an inhibitory role during early embryonic development by sequestering VEGF and preventing its interaction with VEGFR-2 (173). In addition, a splice variant of VEGFR-1, soluble Flt-1 (sFlt-1) acts as a potent antiangiogenic molecule by binding circulating VEGF (183).

Figure 1.4 - Vascular Endothelial Growth Factor (VEGF) Receptor Binding and Signal Transduction. Upon binding of VEGF to the extracellular domain of its receptor, dimerization and autophosphorylation of the intracellular receptor tyrosine kinases occurs and a cascade of downstream proteins are activated leading to biologic effects such as proliferation, migration, and survival of endothelial cells. Only the major proteins in each pathway are depicted. Phosphoinositide 3-kinase (PI3K); protein kinase B (Akt/PKB); p38 mitogen-activated protein kinase (p38MAPK); mitogen and extracellular kinase (MEK); extracellular regulated kinase (Erk). (Modified from Rini, B et al.)(181)

1.6.1. VEGF ex vivo Gene Therapy

VEGF mediated angiogenesis is crucial in bone formation (184), fracture repair (170) and endochondral ossification (185). VEGF has also been shown to promote mesenchymal stem cell (MSC) chemotaxis (186) and play an important role in chondrocyte differentiation, osteoblast differentiation, and osteoclast recruitment (187). Due to these important roles of VEGF in bone formation VEGF ex vivo gene therapy is starting to be explored to enhance bone healing in animal models. As an example, Li et al. (122) transfected rabbit fibroblast with the plasmid pcDNA-VEGF encoding the VEGF gene using SuperFect reagent. After transfection, even though the authors did not report the confirmation or quantification of VEGF protein production, 5x106 cells were delivered in a Gelfoam to a 10 mm bone defect created in rabbit tibiae. After 12 weeks, bone healing was evaluated by x-rays, MicroCT and histology, finding

that rabbits that were treated with the VEGF-cells had new bone bridging the defect while the defects treated with naive cells or saline were filled by fibrotic or sparsely ossified tissue. In addition, immunohistochemistry for CD31, a marker for endothelial cells, revealed more positively stained vessels in the group treated with the VEGF-cells than in the group treated with naive cells indicating the angiogenic effect caused by the VEGF produced by the engineered cells. The results obtained by this group provide strong evidence of the enhancing effect of VEGF over-expressing non-osteogenic cells in bone healing. Furthermore, results indicate that although VEGF expression was transient, it was enough to initiate and enhance bone healing in the rabbit defect model chose by the authors.

Recent studies have shown that VEGF plays a key role in regulating the interaction between osteogenesis and angiogenesis (188). Therefore some groups are exploring the relationship of VEGF and some members of the BMP family. For example, Samee et al (189) investigated the effect of overexpression of BMP2 and VEGF in human periosteal cells in vitro and in vivo. Cells were transfected using Lipofectamine 2000 with the plasmids pEGFP-N1-BMP2 encoding the human BMP2 gene and/or pcDNA3.1-VEGF encoding the human VEGF gene. After transfection cells were cultured under osteogenic conditions and at day 14 and 21 it was found that ALP activity was significantly higher in the BMP2/VEGF group when compared to BMP2, VEGF and naive cells. Also, matrix mineralization was analyzed, although bone nodules were formed in all the groups it was found that at day 21 and 28 the combinational group presented more bone nodules than the other groups. To test the effects of the modified cells in vivo porous β-TCP scaffolds were seeded with 1.25 x 10 6 cells and placed into a muscle pouch in nude mice. Histological analysis 4 weeks after implantation showed that the combinational group had formed the highest amount of ectopic bone while at 8 weeks no difference was found between the BMP2/VEGF and the BMP2 group. Furthermore, CD31 immune staining showed a high density of blood vessels in the scaffolds that were seeded with VEGF and the combinational group confirming the angioinductive effect of VEGF secreted by the cells.

Peng et al., (25) investigated the interaction between VEGF and BMP4 in ectopic bone formation and healing of critical size defects in mice by delivering mouse muscle-derived stem

cells (MDSCs) transduced via retrovirus with either the BMP4 or VEGF transgene. In terms of ectopic bone formation it was found that the group that received BMP4-MDSCs combined with VEGF-MDSCs (ratio of 5:1 BMP4-/VEGF-expressing cells) significantly formed more bone than the BMP4 group. Moreover, CD31 immune staining showed more active capillaries in the BMP4+VEGF group than in the BMP4 group at 4 days demonstrating the angiogenic effect of VEGF-cells. With respect to the healing of critical size calvarial defects it was reported that supplying VEGF-MDSCs alone in Gelfoam was not sufficient to initiate the cascade of bone regeneration; however, when combined with BMP4-MDSCs, they acted synergistically to increase bone healing by enhanced host MSC recruitment and survival at the defect site which accelerated endochondral bone formation. In addition, it was shown that the defects that received BMP4-MDSCs combined with VEGF-MDSCs had statistically significantly higher bone density than the group containing BMP4-cells alone. Results obtained by the same group using BMP2- MDSCs instead of BMP4-MDSCs showed very similar outcomes, where the mix of BMP2- and VEGF-cells enhanced bone healing by improving angiogenesis, which in turn led to accelerated cartilage resorption and enhanced mineralized bone formation. Interestingly, the results of the two studies were compared and it was reported that ex vivo gene therapy with VEGF plus BMP4 is more effective than ex vivo gene therapy with VEGF plus BMP2 (190). Taken together the results of these studies provide robust evidence of the synergistic interaction between VEGF and BMPs in bone healing and formation.

In conjunction the results obtained so far by ex vivo gene therapy suggest that while VEGF alone may not be capable of inducing bone regeneration, its combined application with BMPs produces synergistic effects to enhance bone formation yielding better result than when BMPs are used alone. Although these studies are the first steps they indicate that the combinational gene therapy could be an efficient strategy to induce bone formation and bone healing. To solidify these findings more studies in human cells and bigger animal models are needed. Also, it is imperative to answer important questions related to the ratio of BMPs- to VEGF-cells that should be used, or if it is better to co-express the two genes in the same cells.

1.7. Runt-related Transcription Factor 2

Runt-related Transcription Factor 2 (Runx2, Cbfa1, Osf2, AML3 and PEBP2_A) (191) is a member of the Runt domain family of transcription factors essential for osteoblast differentiation (192,193), endochondral and intramembranous bone formation and turnover (194), vascular invasion of developing skeleton (195) and chondrocyte maturation and hypertrophy (196,197). Originally, Runx2 was characterized as an osteoblast-specific transcriptional activator (192) that binds to the OSE2, a cis-acting element in the promoter of the osteocalcin (OCN) gene (198). Runx2 has been identified and localised on human chromosome 6p21 (197) and its expression is initiated from two different promoters, P1 and P2, which give rise to the two major Runx2 isoforms, type I and type II. Runx2 type II is initiated at the distal promoter P1 while Runx2 type I transcription is initiated at the proximal promoter P2 (199). Runx2 type I begins with the MRIPVD amino terminus (200) and was originally cloned as a T-cell specific factor (201). However, this isoform is also expressed in other non-osseous tissues in addition to osteoblasts (202,203). Runx2 type II begins with the MASNS amino terminus and it was originally cloned as a bone specific factor (192). Different studies indicate that only protein levels, and DNA binding activity of Runx2 type II are physiologically regulated in response to osteoinductive stimulation and osteoblastic maturation (204). Thus, it is generally accepted that Runx2 type II is the isoform highly regulated and expressed in mature osteoblasts and hypertrophic chondrocytes (197).

Consistent with its role as a master organizer of bone formation, alterations in Runx2 expression levels are associated with skeletal diseases. Runx2-deficient mice have normally patterned cartilaginous skeletons but display a complete lack of endochondral and intramembranous bone formation due to the absence of osteoblasts and hypertrophic chondrocytes (194,205). Transgenic mice studies have shown that overexpression of Runx2 using type I collagen promoter inhibits late stage osteoblast differentiation by blocking maturation of osteoblasts, causing osteopenia with multiple fractures in mice (206). Moreover, Runx2 haplo-insufficiency causes the pathogenic skeletal phenotype cleidocranial

dysplasia (CCD) in mice and humans, characterized by short stature, hypoplastic clavicles and dental abnormalities (200,207).

Runx2 controls osteoblast development by binding to specific DNA sequences to activate or repress the transcription of different genes. Runx2 target genes include regulators of cell growth, components of the bone extracellular matrix, angiogenesis, and signaling proteins for development of the osteoblast phenotype and bone turnover (208). By binding to the osteoblast-specific cis-acting element OSE2 which is present in the promoter of several osteogenic genes (191), Runx2 controls the expression of osteocalcin (OCN) (198), osteopontin (OPN) (209), BSP (210), and collagen type I alpha I (Col1A1) (211). In addition, other genes related to the bone development and turnover such as vascular endothelial growth factor (VEGF) (172), collagenase III (212), and osteoprotegerin (213) are upregulated by Runx2 expression.

Runx2 expression and transcriptional activity is highly regulated by multiple transcription factors, signal transduction pathways and cofactors. Runx2 is transcriptionally upregulated by the BMP/Smads pathway (204), Wnt/LRP5/b-catenin pathway (208), fibroblast growth factors (FGFs) (214), and retinoic acid (212), and is downregulated by the 1,25-(OH) 2-vitamin D3/VDR/VDRE pathway (199,215) and tumor necrosis factor-a (TNF-a) (216). Runx2 is also regulated by the glucocorticoid family of steroid hormones (217), the parathyroid hormone (PTH) and parathyroid-related peptides (PTHrPs) and integrins (195). Factors upstream of Runx2 such as Msx2 (199), Dlx5 (218), STAB2 (219) and Dlx3 (220) activate the expression of Runx2 while Twist is known to act as an antagonist by directly binding to the Runt DNA-binding domain of Runx2 decreasing its binding to DNA (221). Cofactors are known to interact with Runx2, some to co-activate, while others to co-suppress by affecting Runx2 DNA binding activity and/or transactivation potential. STAT1 (222), YAP (223) and histone deacetylases HDACs (195) are all known to co-suppress Runx2, while core-binding factor-beta (CBF-β) (224), TAZ (225), MOZ and MORF (226) are known to co-activate Runx2. In addition, Runx2 has been shown to regulate the activity of its own promoter both positively (227) and negatively (228).

1.7.1. Runx2 ex vivo Gene Therapy

Due to the fundamental role of Runx2 in bone formation, some groups have been exploring Runx2 ex vivo gene therapy to improve bone regeneration. For example, Zhao et at. (81) studied the in vitro and in vivo effects of Runx2 overexpression in mouse bone marrow stromal cells (BMSCs) after being genetically modified using Runx2 adenovirus (Ad-Runx2). Their in vitro studies revealed that Runx2 protein levels in Runx2-BMSCs remained relatively constant for 6 days and then gradually declined. In addition, they found that overexpression of Runx2 caused an evident increase of ALP activity, and production of BSP and OCN mRNA, that peaked at day 6 and declined thereafter. In addition, matrix mineralization was detected in Runx2- BMSCs at day 6, a week earlier than in their control LacZ-BMSCs. To test the potential of Runx2-BMSCs to form bone ectopically, in vivo, they seeded 2x106 cells in either a type I collagen hydrogel or a gelatin sponge (Gelfoam) and implanted them subcutaneously in mice. After 3 weeks, radiological and histological analysis revealed that both Runx2-BMSCs and LacZ- BMSCs formed bone, but the mineralized area of the Runx2-BMSCs group was 2.7-fold greater than in the LacZ-BMSCs group when seeded in collagen, and 8.3-fold higher in Gelfoam — demonstrating the increased osteogenic activity in the Runx2-cells. In a later study from the same group (127) the potential of Runx2-BMSCs to heal critical size defects was tested by delivering a gelatin sponge seeded with 2x106 cells into 5mm calvarial defects in mice. After 7 weeks, radiographic, MicroCT and histological analyses revealed that only limited bone was formed at the margins of defects in the LacZ-BMSCs group while, defects were completely closed in the Runx2-BMSCs group. Similar results were reported by Zheng et al. (229) where the overexpression of Runx2 in mouse BMSCs was achieved using adenoviral infection. In vitro, ALP activity at day 7 was markedly higher in Runx2-BMSCs when compared to LacZ-BMSCs and naive cells; and mineralized nodules and OCN and BSP mRNA expression were only detected in the Runx2-BMSCs. In vivo, cells were delivered in collagen type I into a 5mm skull defect in mice; after 4 weeks samples were analyzed by histology. Specimens from groups that were treated by collagen alone or not treated showed no evidence of osseous healing, defects that were treated with naive cells showed minimal newly formed bone, while specimens that were treated with Runx2-cells showed nearly complete osseous closure of the created defects. 27

Collectively, these three studies provide robust evidence of the enhancing effect of Runx2 over-expressing mouse BMSCs in bone healing.

In another study, rat adipose derived stromal cells (ADSCs) genetically modified with Ad-Runx2 were found to express upregulated mRNA levels of OPN, OCN, BSP and Col1A1 when compared to the control EGFP-ADSCs in vitro. It was also found that Runx2-ADSCs expressed high levels of ALP activity with a peak at day 10 post-infection and mineralized matrix by day 10 while EGFP-ADSCs had very low levels of ALP activity and no detected matrix mineralization when cultured under non-osteogenic conditions. To test the osteogenic potential of the Runx2- ADSCs in vivo 2 × 106 cells were seeded in PLA scaffolds and implanted in the back of nude mice. After 4 weeks, histological analysis showed that a considerable amount of cartilage and bone was formed by the Runx2-ADSCs while little, or no, mineralized tissue was observed in the control groups (230). Altogether these results corroborate the previous independent studies, and indicate that although Runx2 genetic modification via adenoviral infection is transient it seems to be enough to initiate and enhance bone formation when murine mesenchymal stromal cells are used.

Runx2 overexpression induced by infection with Runx2-retrovirus encoding the murine Runx2 gene has been extensively studied in different cells by the Garcia group at the Georgia Institute of Technology. In one of their earliest publications Byers et al (231) investigated the effects of Runx2 overexpression in different types of cells in vitro by analyzing RNA from cell cultures by real time PCR. Results revealed that Runx2 overexpression caused the upregulation of OCN and COLI in MC3T3-E1 immature osteoblast-like cells, NIH3T3 cells, and C3H10T1/2 fibroblasts and also caused the upregulation of BSP in MC3T3-E1 and, to a lower extent, in C3H10T1/2 cells. In addition, it was found that Runx2 enhanced ALP activity in all the cells employed, but to a lower level in NIH3T3. Interestingly, it was observed that overexpression of Runx2 in MC3T3-E1 cells resulted in enhanced matrix mineralization compared with controls while cultures from NIH3T3 and IMR-90 fibroblasts overexpressing Runx2 did not produce mineralized matrix and only 25% of Runx2-expressing C3H10T1/2 cell cultures produced mineralized nodules indicating that the effects of Runx2 overexpression are cell type dependent. Work by the same

authors in a later publication reported the effects of Runx2 overexpression in rat bone marrow stromal cells after cultured under osteogenic conditions. Real time PCR was performed in samples from cultures at day 1, 3 and 7 and showed that levels of OCN and COLI were upregulated at all time points, in contrast, BSP and OPN levels were found to be only upregulated at day 3 while at the other time points they were similar to control cultures. Moreover, ALP activity at day 7 was found to be around 2-fold higher in Runx2-BMSCs when compared to controls and culture mineralization was detected from day 7 in Runx2-BMSCs while at day 14 in the control group indicating Runx2 enhanced BMSC osteogenic differentiation (232).

Further studies to understand the effect of Runx2 overexpression mediated by retroviral infection were performed in mouse myoblasts. In vitro work by Gersbach et al (233) revealed that Runx2-myoblasts expressed upregulated levels of OCN, BSP, OSX, and Dlx5 mRNA and ALP activity was at least 10-fold higher in Runx2-myoblasts when compared to controls. Furthermore, matrix mineralization was detected in Runx2-myoblasts by day 14 while it was undetected in the controls indicating that overexpression of Runx2 stimulated the transdifferentiation of mouse myoblasts into a mineralizing cell. In a later publication by the same group (234) mouse myoblasts were genetically modified using a retroviral system encoding the Runx2 gene under the control of the tetracycline-inducible (tet-off) promoter. The system was tested in vitro by adding anhydrotetracyline (aTc) to cell culture medium at different concentrations. Detection of Runx2 at the protein level showed that Runx2 expression was controlled by aTc in a dose-dependent manner; the same was noticed with osteoblastic gene expression, ALP activity and matrix mineralization. To test the system in vivo, 5 x 10 5 cells were seeded in collagen disks and implanted into the hind limbs of immunocompetent mice for 28 days, during the experimental period some of the mice were provided drinking water with aTc. After robust histology and MicroCt analyses of the specimens it was found that implants in mice that did not receive aTc formed mineralized tissue; while mineralization was not detected in implants of mice that received aTc or implants that were seeded with unmodified myoblasts. While the tetracycline-inducible (tet-off) approach of this study is of great value to the field of ex vivo gene therapy, especially when

using vectors that allow for long-term expression of the gene of interest like retroviruses, it would be more relevant to show similar results with a tet-on system since it makes more sense to provide the drug only when the expression of the gene is needed, allowing more control of the outcome of the therapy.

After successfully showing that Runx2 overexpression caused the transdifferentiation of mouse myoblasts into mineralizing cells the Garcia group studied the effects of Runx2 expression in rat dermal fibroblasts (235). To test the in vitro osteogenic potential of Runx2-fibroblasts 5x105 cells were seeded in collagen disks and cultured them for up to 42 days in osteogenic conditions. Osteogenic gene expression analyzed by real time RT-PCR showed upregulation of ALP, OCN and BSP mRNA expression compared to controls. Furthermore, MicroCT analysis detected an approximately 2-fold increase in mineral volume in scaffolds seeded with Runx2- fibroblasts when compared with scaffolds seeded with control-fibroblasts. To test their in vivo potential, scaffolds were subcutaneously implanted into the back of syngeneic rats. After 4 weeks, MicroCT and histology showed that Runx2-fibroblasts formed mineralized tissue in vivo, whereas minimal mineralization was found in control group. These results prove that Runx2 overexpression via retroviral infection can be use as a strategy for the conversion of dermal fibroblasts into a mineralizing cell source.

1.7.1.1. Runx2 and BMP2 Combinational ex vivo Gene Therapy

Based on the reported studies, Runx2 ex vivo gene therapy seems promising as a route to enhance bone healing in clinical applications. However, it has been reported that Runx2 therapy is not as effective as BMP2 ex vivo gene therapy (236,237). Therefore, some groups are starting to test the combined effect of overexpressing Runx2 and BMP-2 in bone formation. As an example, Yang et al. (237) genetically modified the pluripotent C3H10T1/2 murine cell line with a mix of Ad-Runx2 and Ad-BMP2 and compared the in vitro and in vivo results with cells only infected by Ad-Runx2 or Ad-BMP2. The in vitro work revealed that BMP2/Runx2-cells exhibited higher levels of ALP activity and mineralization than BMP2- or Runx2-cells alone, but BMP2-cells showed higher levels of mineralization compared to Runx2- 30

cells. Also, OCN mRNA levels were found to be higher in the BMP2/Runx2-cells followed by the Runx2-cells. In addition, similar results were obtained when 5x106 engineered cells were seeded in gelatin sponges and implanted subcutaneously in immunodeficient mice. After 4 weeks, histological analysis revealed that LacZ-cells only generated fibrous tissue while Runx2- cells produced a small amount of cartilage and bone. BMP2-cells and BMP2/Runx2-cells generated large ossicles containing bone, cartilage and a marrow cavity, nevertheless, it was found that BMP2/Runx2-cells produced significantly (P<0.05) more mineral content and bone area than the BMP2-cells indicating a synergistic interaction between Runx2 and BMP2.

In a recent study and in the first report found using overexpression of Runx2 in human cells, Lee et al., (238) investigated the interaction between BMP2 and Runx2 in adipose-derived stromal cells (ASCs). Cells were transfected using microporation to deliver a BMP2/Runx2 bicistronic vector encoding the human BMP2 gene tagged with the HA epitope at the C- terminal region and the human Runx2 gene tagged with the Myc epitope. In vitro studies in non-osteogenic conditions revealed that BMP2/Runx2-ASCs expressed higher mRNA levels of OPN, OCN and COL1A1 than BMP2-ASCs. In addition, it was reported that ALP activity and calcium content was higher in the BMP2/Runx2-ASCs when compared to BMP2-ASCs. To test the osteogenic and osteoinductive capacity of the engineered cells, PLGA scaffolds were seeded with 1X 106 cells and implanted subcutaneously in nude mice. After 6 weeks, samples were analyzed by histology and revealed, according to the authors, a “significant increase” in bone formation in the BMP2/Runx2-ASCs compared to the BMP2- ASCs. However, the only histological images they provided failed to present strong evidence of bone formation. Also RNA was extracted from the specimens and analyzed by RT-PCR finding that expression of OPN, OCN and COL I was higher in BMP2/Runx2-ASCs when compared to BMP2-ASCs while in the mock-ASCs expression of these genes was undetected. Although this is the first study published where Runx2 is overexpressed in human MSCs the authors only investigated the effects of its overexpression combined with BMP2 but did not study the effects of Runx2 overexpression alone; moreover, the images provided from the in vitro and in vivo work do not strongly support what its claimed by the authors therefore, there is no robust evidence yet of the osteoinductive effects of Runx2 overexpression in human cells.

Even though Runx2 was discovered more than a decade ago the field of Runx2 gene therapy is still undeveloped. Despite of the various studies that have investigated the effects of Runx2 overexpression in different types of non-human cells with promising results there is a need for new research that focuses on the effects of Runx2 overexpression in human cells. Furthermore, combinational therapy is a strategy that is starting to be explored and that needs to be further studied to understand the interaction between Runx2 and other important proteins related to bone formation such as BMP2 and VEGF. Moreover, the effectiveness of Runx2 ex vivo therapy have to be tested in more challenging animal models since the murine models used to date might not reflect all the issues that can be encountered in human bone defects.

1.8. Osterix Transcription Factor

Osterix (Osx, Sp7) is a member of the Sp/KLF (Krüppel Like Factor) family of transcription

factors that contains three C2H2-type zinc-finger motifs (239). Over a decade ago Osterix was identified as a BMP-2-induced gene in the premyoblastic murine cell line C2C12 by Nakashima et al. (240). The murine Osterix is a 428 amino acid polypeptide with a molecular mass of 46 kDa that contains a C-terminal zinc finger motif of 85 amino acids which serves as the DNA binding domain, that recognizes GC boxes (CGCCC) and GT boxes (CACCC) (241). Recently, two isoforms of human osterix have been identified, Sp7-α and -β, sharing 95% sequence homology with murine Osx (242,243). Osterix is expressed specifically in osteoblasts and transiently in differentiating chondrocytes (244). Osterix deficient mice exhibit impairment of osteoblastogenesis and expression of osteoblast markers such as osteonectin, osteopontin, osteocalcin and bone sialoprotein, absence of mineralized bone matrix, and perinatal lethality (240), indicating that Osterix, like Runx2, is an essential transcription factor for bone development. Despite the similarities between the phenotype of Osterix and Runx2 deficient mice, important differences were reported such as the inhibition of chondrocyte hypertrophy found in the Runx2 mutants but not in the Osterix knockouts (245), suggesting that Osterix, unlike Runx2, is not required for chondrocyte hypertrophy (18). In addition Osterix expression 32

was not found in Runx2 deficient mice whereas Runx2 was normally expressed in Osterix deficient embryos (240). Thus, Osterix acts downstream of Runx2 to induce differentiation of preosteoblasts into fully functional osteoblasts (138).

Functional roles for Osterix in bone and cartilage are starting to be elucidated. Forced expression of Osterix in vitro has been reported to induce expression of osteopontin (246), ALP (247), osteocalcin and collagen type 1A1 (240). Also, Osterix is known to inhibit chondrocyte differentiation while promoting the commitment of progenitor cells to the osteogenic phenotype and their differentiation into mature osteoblasts (248), and is thought to inhibit osteoblast proliferation through the inhibition of the Wnt signaling pathway by directly activating the transcription of the Wnt antagonist Dkk1 (249).

The regulation of Osterix expression during osteoblastogenesis is complex and involves various growth factors in osteoblasts and chondrocytes (250,251). BMP2 induces Osterix expression by inducing the transcription of Runx2 which upregulates Osterix by binding directly to a responsive element in the promoter of the Osterix gene (138). This BMP-2 induction and subsequent upregulation of osteoblast-specific genes involves Dlx5, Smad transducers, and the mitogen activated protein kinase (MAPK) pathway (11). Like BMP-2, insulin-like growth factor- 1 (IGF-1) can induce Osterix expression in undifferentiated mesenchymal stem cells through Runx2-dependent (252) and Runx2-independent (251) pathways. Ascorbic acid (253) , 1,25 (OH)2 vitamin D3 (254) and the homebox family member Msx2 (255) are also implicated in upregulation of Osterix expression. Negative regulators of osteoblastogenesis, such as tumour necrosis factor-α (TNF-α) (256) , tumour suppressor p53 (257) and epidermal-growth factor receptor (EGFR) (258) can inhibit Osterix expression. Interestingly, Osterix can regulate its own expression through a feedback mechanism by interacting with its own promoter (259).

1.8.1. Osterix ex vivo Gene Therapy

Osterix ex vivo gene therapy for bone regeneration has only recently been explored and the few studies reported to date present encouraging results. For example, Tu et al (128,260) reported the genetic modification of mouse bone marrow derived MSCs using retroviral infection to overexpress the mouse Osterix gene. In a first in vitro study (260), it was reported that overexpression of Osterix increased Osterix-BMSCs proliferation and osteogenic potential by upregulating the expression of osteoblastic markers, including alkaline phosphatase, bone sialoprotein, osteocalcin, and osteopontin. Furthermore, it was shown by RT-PCR that Runx2 was not upregulated as a result of Osterix overexpression, finding similar levels between the Osterix-BMSCs and the cells transduced with the empty vector. Although the expression of Osterix mRNA by the Osterix-BMSCs was reported the authors failed to provide evidence of Osterix production at the protein level, which is important since the presence of the mRNA is not a complete indication that the protein is being produced, and in studies that involve genetic modification the confirmation of the production of the protein of interest by the modified cells should be one of the first steps. In a later in vivo study from the same group (128), 5x106 of BMSCs modified via retroviral infection were delivered to a 4mm mouse critical- sized calvarial defect using type I collagen sponges as carriers. Five weeks later, after radiology and histological examination it was found that Osterix-BMSCs accelerated bone healing and formed five times more bone than the BMSCs transduced with the empty vector and the naive BMSCs; and 27 times more bone when compared to defects treated with the collagen carrier alone. In addition, immunohistochemistry for BSP resulted in intense staining in areas of new bone formation in the group treated with the Osterix-BMSCs while in moderate staining in the group treated with BMSCs transduced with the empty vector.

In another study, Kim et al. (246) reported the insufficiency of Osterix to induce osteogenic differentiation in NIH-3T3 fibroblasts after transfection using lipofectamine PLUS with the plasmid pcDNA3.1–HA–Osterix encoding the mouse Osterix gene. After culturing the cells in osteogenic media for a period of 25 days they detected upregulation of the osteopontin gene when compared to the cells transfected with the empty vector, but did not detect upregulation

of other bone-related markers, including collagen type I, alkaline phosphatase, osteocalcin, or osteonectin. Moreover, Osterix overexpressing cells failed to produce a mineralized matrix. Even though, it would be relevant and more conclusive to see the effect of stable long-term Osterix overexpression in NIH-3T3 these results indicate that the forced expression of Osterix is not sufficient to induce in vitro mineralization in this non-osteogenic cell line. Interestingly, similar results were reported by Byers et al (231) after stable long-term expression of Runx2 in NIH-3T3 failed to direct in vitro mineralization.

In a recent study, Lai et al. (261) investigated the effects of Osterix ex vivo gene therapy in the osteodistracted zone of a mandibular lengthening model in rabbits. Bone marrow derived MSCs (BMSCs) were transfected using Lipofectamine 2000 with the plasmid pEGFP-OSX encoding the Osterix gene with a transfection efficiency of about 48%. After genetic modification, a group of nine rabbits was treated by injecting 1 × 107 of OSX-BMSCs suspended in 0.2 mL of physiological saline into the osteodistraction zone, while in two more groups rabbits received the same dosage of naive BMSCs or injection of saline alone. After 6 weeks, osteogenesis was achieved between the 2 bone fragments in all the specimens of the 3 groups but results obtained by radiographic and histological analysis showed significant differences among the groups. According to the authors, even though they failed to show a representative x-ray for each of the 3 groups, the gray density analysis of the x-rays at both 2 and 6 weeks confirmed statistically significant differences among groups, in which the radiodensity of the callus in the group that received the Osterix-BMSCs was greater when compared with the other two groups indicating greater bone mineral density. In addition, histology slides prepared from the distraction gaps of all the samples showed that the group that received the Osterix-BMSCs presented thicker, homogeneous and denser trabeculae when compared to the other groups. Also, it was found by immunohistochemical analysis that BSP expression at two weeks was evidently increased within the gaps of the Osterix-BMSCs group when compared to the other groups, a finding that correlates with the results reported previoulsy by Tu et al (128). Although the authors did not report the confirmation of expression of Osterix at the protein level by the BMSCs after the genetic modification, the immunohistochemical and

histological evidence shows that the delivery of BMSCs transiently overexpressing Osterix is enough to improve bone formation in an ostedistracted site.

Finally, in the only study published so far using human cells, Wang et al., (247) transfected umbilical cord–derived mesenchymal stromal cells (UC-MSCs) using Lipofectamine 2000 to deliver the plasmid pEGFP-Osx containing the Osterix gene. Although the authors failed to provide evidence of the production of Osterix at the protein level by the Osx-UC-MSCs, the in vitro results obtained by RT-PCR at days 7 and 14 showed that overexpression of Osterix in UC- MSCs enhanced expression of bone matrix proteins including collagen type I, alkaline phosphatase, osteocalcin and bone sialoprotein. Moreover, to investigate the potential of Osx- UC-MSCs to form bone in ectopic sites, an unreported number of cells were seeded in PLGA scaffolds and implanted subcutaneously in nude mice. Four weeks after implantation histological and histomorphometric analyses were performed and although the only histological images they provided failed to present strong evidence of bone formation the authors claimed that a small amount of bone was formed in the control groups (scaffolds implanted alone, scaffolds seeded with naive UC-MSCs and scaffolds seeded with Mock-UC- MSCs) while approximately four times more bone was formed in the Osx-UC-MSCs group. In addition, quantitative real-time PCR was performed in RNA extracted from the different implanted scaffolds revealing that ALP, OCN, OPN and COL I gene expression was upregulated in the scaffolds seeded with Osx-UC-MSCs when compared with the control groups. While this is the first report attempting to show the usefulness of Osterix ex vivo gene therapy using human cells it lacks power since the results reported were obtained with cells from only two different donors, also no information was provided regarding the capacity of Osx-UC-MSCs to mineralize matrix in vitro and the in vivo histology data does not clearly show bone formation.

Despite the recognition of Osterix as a fundamental gene for bone formation since 2002 there are only few published studies that have investigated the potential of Osterix overexpression to enhance bone formation in a clinical setting. Therefore, there is a need for more studies that can provide robust evidence of the effects of Osterix overexpression particularly in human

cells. Also there should be no hesitation in starting to investigate the combination of Osterix with other genes related to bone formation as it might be a way to enhance the osteogenic effects of Osterix ex vivo gene therapy (261).

1.9. Rationale

Bone marrow stromal cells (BMSCs) are currently the "gold standard" cell source for mesenchymal cell-based therapies and last year for the first time allogeneic human BMSCs received authorization from Health Canada to be marketed as a stem cell therapy for acute graft-versus host disease in children (262). This could represent the beginning of the commercial approval of other allogeneic MSC-based products for the treatment of different conditions. Recently, human umbilical cord perivascular cells (HUCPVCs) have been reported to have many similarities with human BMSCs in terms of differentiation capacity, surface markers (74–76), transfectability (74), and non-immunogeneic and immunomodulatory phenotype (78– 80), while exhibiting a higher proliferation rate (74) and the advantage of being easier and safer to obtain since they are harvested from a tissue that is normally discarded after birth.

During the past years the majority of MSCs that have been used in the orthopedic ex vivo gene therapy field have been BMSCs. Genetic modification of these cells with different exogenous genes such as BMP-2 (113,159,160,162), Runx2 (81,263,264), osterix (128,260,261) have successfully improved their bone formation capacity. Furthermore, it has been shown that the overexpression of a combination of two genes VEGF/BMP2 (190), VEGF/BMP4 (25), BMP2/Runx2 (238) or Runx2/SP7 (265) leads to improved results in terms of bone formation when compared to single gene overexpression. Due to the similarities between BMSCs and HUCPVCs, and significantly higher HUCPVCs proliferation rate that allows the production of vast numbers of cells the main purpose of this work is to determine the potential of HUCPVCs as a putative candidate cell source for osteogenic ex vivo gene therapy by combining for the

first time the overexpression of four genes related to bone formation: BMP-2, Runx2, SP7 and VEGF.

1.10. Hypothesis

HUCPVCs are a putative cell source for osteogenic ex vivo gene therapy that when genetically modified and employed in cocktails overexpressing different osteogenic related genes would be more biologically effective than populations overexpressing a single gene, and which biological activity could be maintained upon thaw after being cryopreserved.

2. Materials and Methods

Materials used in this thesis work are listed in the Appendix 1 with their respective supplier and catalog number.

2.1. HUCPVCs Isolation, Thawing and Culture.

Frozen aliquots of human umbilical cord perivascular cells (HUCPVCs) were kindly provided by Tissue Regeneration Therapeutics, Inc (TRT, Toronto, ON). HUCPVCs were harvested according to the protocol described by Sarugaser et al (56). The cells were thawed by quickly submerging and swirling the sample vial in a 42°C water bath for 1-2min - until cells were completely thawed but still cold - followed by the addition of 1 mL of cold alpha-MEM (Invitrogen) to the sample vial. After 2min, the solution containing the cells was transferred to a 15mL tube and 2 mL of alpha-MEM were added, 2 min later, 8 mL more of alpha-MEM for a total volume of 13ml. Cell suspension was gently mixed and 500µl were used to count viable cells using the Vi- CELL XR Cell Viability Analyzer (Beckman Coulter) which employs the trypan blue dye exclusion method, the remaining in the 15 ml tube was centrifuged at 285xg for 5 min at 4°C. After centrifugation cell pellet was resuspended in alpha-MEM supplemented with 5% fetal bovine serum (FBS, HyClone) and 10% antibiotic stock solution (0.1% Penicillin (Sigma-Aldrich); 1% Gentamicin (Sigma-Aldrich); and 0.3 mg/mL amphotericin B (Sigma-Aldrich)) and 1x106 cells 2 were plated per 150 cm flasks. During cell culture flasks were kept at 37°C and 5% CO2, and medium was changed 2-3 times a week; when confluence was around 80-90% cells were passaged at a ratio of 1:3 using 0.05% trypsin/0.02% EDTA solution (Invitrogen).

2.2. Genetic Modification of HUCPVCs

Most of the work in this thesis is based in the genetic modification of HUCPVCs, therefore different non-viral and viral strategies were assessed to assure that the most efficient method

of genetically modifying HUCPVCs was chosen. Different transfection/transduction methods are detailed in the following sections.

2.2.1. FuGENE 6

FuGENE 6 (Roche Diagnostics) transfection reagent is a proprietary, nonliposomal formulation made of a blend of lipids and other components that form a complex with DNA, when added to the culture medium the complex is transported into the cells. To test this transfection method HUCPVCs from 5 different cords were mixed and seeded at a concentration of 2x105 cells/well in 6-well plates and allowed to attach overnight. The day after, when confluence reached 70- 80%, cells were transfected with the plasmid pmaxGFP (3.4kbp) (Amaxa) using FuGENE 6. Following the manufacturer's protocol for adherent cells three different ratios of plasmid DNA (µg) to volume of FuGENE 6 (µl) were used, 2:3, 1:3 and 1:6. Briefly, FuGENE 6 reagent was diluted with serum-free medium and then mixed with plasmid DNA. After 20 minutes of incubation at room temperature the obtained solution ~100µl was added to the HUCPVCs culture medium (2ml/well). Three days post transfection gene expression was assessed by both fluorescence microscopy (Olympus IX81 with a Photometrics CoolSnap HQ2 camera) and flow cytometry (Beckman Coulter, CYTOMICS FC 500).

2.2.1.1. Flow Cytometry

For the detection of GFP expression 1x105 modified cells were resuspended in 500µl of alpha- MEM and flow cytometry (Beckman Coulter, CYTOMICS FC 500) was performed using a blue Argon-ion laser at 488nm and detecting the emitted signal by the FL1 detector. For each sample the number of events was set to 5000. As a negative control non-modified cells were used and data was analyzed using the CXP Cytometry List Mode Data Acquisition and Analysis Software (Beckman Coulter, 2006).

2.2.2. Lipofectamine LTX

Lipofectamine LTX (invitrogen) is a cationic lipid formulation that interacts with the plasmid DNA to form positively charged micelles or liposomes. These lipid-nucleic acid complexes, when added to the culture medium fuse with the cell membrane, become internalized, and subsequently expressed. To transfect HUCPVCs using Lipofectamine LTX four different mixtures were prepared: 500ng of DNA/1.25µl of Lipofectamine, 750ng of DNA/1.25µl of Lipofectamine, 500ng of DNA/2µl of Lipofectamine and 750ng of DNA/2µl of lipofectamine following the protocol for adherent mammalian cells as suggested by the supplier. Briefly, HUCPVCs were seeded at a concentration of 3 x104 cells/well in 24-well plates and left to attach overnight. The day after, when confluence reached 70-80% cells were transfected by diluting the plasmid pmaxGFP (Amaxa) in Opti-MEM Medium (Invitrogen, 31985-062) and combining it with Lipofectamine LTX. After 30 minutes of incubation the obtained solution (~100µl) was added to the HUCPVC culture media (500µl/well) and the medium was changed after 6 hrs. Three days post transfection gene expression was assessed by both fluorescence microscopy and flow cytometry.

2.2.3. Amaxa Nucleofector II

The nucleofection technique employs the delivery of an electric pulse of defined length and magnitude to cells that are suspended in optimized cell-type-specific solutions containing the plasmids to be transfected, delivering the plasmid DNA straight to the cell nucleus. The nucleofector has different electrical settings that are displayed as distinct programs. In order to choose the best protocol for HUCPVC transfection, 7 programs were tested with the Cell Line Optimization Nucleofector kit (Amaxa) in accordance with the manufacturer’s instructions. Briefly, 1x106 HUCPVCs were resuspended in 100µl of Nucleofector Solution V and mixed with 2µg of pmaxGFP and then transferred to a nucleofection cuvette. After delivery of the electric pulse cells were mixed with 500µl of pre-warmed medium and transferred to 3 wells of 6-well plates each containing 2ml culture medium. In addition, a negative control for later measurement of cell survival and transfection efficiency was set by exposing the same number 41

of cells to all the steps but the electric pulse. After 2 days of incubation transfection efficiency was measured by flow cytometry and cell survival of the nucleofected cells was assessed by comparing the number of viable cell obtained in the negative control with the viable cells counted in each of the 7 different tested programs using the Vi-CELL XR Cell Viability Analyzer (Beckman Coulter) which employs the trypan blue dye exclusion method.

2.2.3.1. Longevity of GFP Expression After HUCPVCs Nucleofection

To assess the level of GFP expression over time HUCPVCs were transfected by nucleofection using the program X-001 (which was found to be the best of the seven tested) and then cultured in alpha-MEM supplemented with 5% FBS. After 3, 7, 12 and 15 days cells were trypsinized and GFP expression was quantified by flow cytometry.

2.2.4. Retroviral Transduction of HUCPVCs

2.2.4.1. Production of Runx2- and Mock-retrovirus pTJ66-Runx2 retroviral expression vector carrying Runx2 murine cDNA and pTJ66 (empty vector), were both kindly provided by Dr. Andres Garcia (Georgia Institute of Technology, USA) (231). To obtain more plasmids copies, DH5α competent E. coli cells (Invitrogen) were transformed by heat shock at 42°C followed by incubation for 1 hour in LB Broth culture media (20g/L of LB broth (BioShop),dH2O) in a shaking incubator at 37°C and 300rpm. Cells were then pellet at 4000g in a table top centrifuge at room temperature and resuspended in 200µl of LB Broth to be plated onto LB-Ampicillin Agar (20g/L of LB broth, 16g/L of Agar (BioShop), 10mg/ml of ampicillin (BioShop), dH2O). Plates were incubated for 16 hours at 37°C and 3 well isolated colonies were picked with a sterilized loop to inoculate 3mL individual cultures of LB- Ampicillin broth (20g/L of LB broth, 10mg/ml of ampicillin, dH2O) that were incubated in a shaking incubator for 16 hours at 37°C. Plasmids were purified from the pelleted D5Hα cells using the Qiagen Miniprep Kit (Qiagen). Eluted DNA was then diagnosed by restriction

digestion with HindIII restriction enzyme (New England Biolabs) in a total reaction volume of

20µl containing 5µl DNA, 1µl of HindIII, 2µl of NEBuffer 2 and 12µl of ddH2O for 2 hours at 37°C; and ran on a 1% agarose (BioShop) gel resulting in three bands for the pTJ66-Runx2 (673, 1537 and 6336 bases in length) while in pTJ66 resulted in one band (6818 bases in length) meeting the expect pattern. To isolate enough plasmid DNA for the generation of retroviruses 100 ml of LB-Ampicillin broth were inoculated with 100µl of the leftover bacterial cultures previously tested, and incubated for 16 hours in a 37°C shaking incubator, set at 300 rpm. The plasmids were purified using EndoFree Plasmid Maxi Kit (Qiagen) and the concentration and quality of the DNA were verified using spectrophotometric quantitation (NanoDrop1000, Thermo Fisher Scientific) (266).

To produce retroviral stocks 70% confluent Phoenix-amphotropic helper-free retrovirus producer cells growing in D-MEM supplemented with 10% FBS at 37°C and 5% CO2 were transfected in T75-flasks by calcium phosphate co-precipitation by mixing in one tube 450μl of dH20, 50μl of 2.5M CaCl2 (Sigma-Aldrich) and 15μg of plasmid. Then the DNA mixture was added dropwise into 2X HBS buffer (0.28M NaCl (Sigma-Aldrich), 0.05M HEPES (Sigma-Aldrich),

1.5 mM Na2HPO4 (Sigma-Aldrich) in ddH2O pH 7), vortexed and incubated at room temperature for 20 minutes. Before adding the resulting CaCl2/HBS/DNA solution chloroquine (Sigma-Aldrich) was added to the Phoenix cultures to a final concentration of 25μM. After 24 hours the medium was changed and the temperature of the incubator was changed to 32°C due to more stability of the virus at this temperature. 48, 60, and 72 hours after transfection supernatants containing retrovirus were collected, centrifuged at 1500g for 5 minutes to pellet cell debris, filtered through a 0.45 μm filter and stored at -80°C until use. From this step two different viruses were obtained: Runx2- and Mock-retrovirus.

2.2.4.2. Viral Infection of HUCPVCs

Following the suggested protocol by the Garcia Lab (231), prior to retroviral infection HUCPVCs were plated in 6-well plates at a density of 1X105 cells/well and incubated in growth media at

37°C. At 50-80% confluence cells were transduced with Runx2- or Mock-retrovirus by adding

0.2ml/cm2 of retroviral supernatant, supplemented with 4μg/ml of Polybrene (Millipore) to increase the efficiency of infection, followed by the centrifugation of the whole 6-well plate at 1200g for 30 minutes at 32° C. After infection, the medium was replaced with regular growth medium (alpha-MEM supplemented with 5% FBS and 10% antibiotic stock solution), and the cells were returned to incubation at 37°C. To increase the transduction efficiency, a second identical infection was performed 10-12 hours later. Three days post-transduction cells were evaluated for the presence of GFP by fluorescence microscopy and flow cytometry. Two types of virally infected HUCPVCs were obtained in this procedure: Runx2-HUCPVC and Mock- HUCPVC.

2.2.4.3. Immunofluorescence

Runx2 expression by Runx2-HUCPVCs was assessed by immunostaining of cultured cells in 12- well plates. All steps were carried out at room temperature. After fixing the cultures using 10% neutral buffer formalin (Sigma, HT501128) for 10 minutes the samples were washed 3 times in PBS for 5 minutes then the cells were permeabilized by adding 800µl of PBS (Gibco) with 0.5% Triton-X (Sigma-Aldrich) for 10 minutes, and blocked using 800µl of PBS with 1% BSA and 0.05% Tween for 1 hour. Following blocking, the samples were incubated for 1 hour with 1:400 Runx2 Rabbit Polyclonal Antibody (Santa Cruz Biotechnology) in blocking solution, washed 3 times with PBS for 5 minutes each, and incubated for 1 hour with 1:1000 Alexa Fluor 555 Goat Anti-Rabbit IgG (Invitrogen), and 1:3000 Hoechst 33342 (Invitrogen) diluted in blocking solution for 1 hour. After 3 final washes with PBS, the samples were photographed using an inverted fluorescent microscope (Olympus IX81with a Photometrics CoolSnap HQ2). To optimize antibody dilutions, HeLa cells (ATCC, CMR2-CCL2) were used as a negative control and HepG2 cells (ATCC) as a positive control for Runx2 expression. The antibody ratios were chosen based in the dilution that gave the best staining in HepG2 cells with no background staining in HeLa cells.

2.2.4.4. Osteogenic Cultures of Runx2-HUCPVCs and Mock-HUCPVCs

After viral infection Runx2-HUCPVCs and Mock-HUCPVCs were seeded in 6-well plates at a concentration of 1x105 cells/well and cultured with osteogenic inductive media (alpha-MEM -8 supplemented with 5% FBS, 10% antibiotics stock, 10 M Dexamethasone (Sigma-Aldrich), 50μg/ml ascorbic acid (Sigma-Aldrich) and 5mM β-glycerophosphate (Sigma-Aldrich)) for 3, 7, 14, 21 and 30 days with media changes every 2-3 days.

2.2.4.5. ALP Staining

Histochemical staining of cultures was performed to identify cells that expressed the alkaline phosphatase (ALP) enzyme. Cultures were rinsed in 1x Tris Buffered Saline (TBS) (Bio-Rad) three times and fixed in 10% ice-cold neutral formalin buffer (Sigma-Aldrich) for 15 minutes. Cultures were then rinsed with distilled water twice and incubated with 1ml of a solution containing 0.005g Naphthol AS MX-P04, (Sigma-Aldrich), 200µl N,N-Dimethylformamide (Sigma, D4254), 25ml TrisHCL (0.2M, pH 8.3), 25ml distilled water and 0.03g Fast Red Violet LB salt (Sigma-Aldrich) for 45 minutes at room temperature. Following the incubation period, cultures were rinsed in distilled water 3 times and pictures were taken with a light microscope (Nikon, DIAPHOT).

2.2.4.6. Von Kossa Staining

Cultures at day 21 were rinsed with 1x TBS three times and fixed in 10% ice-cold neutral formalin buffer for 15 minutes. Cultures were then rinsed with distilled water twice and incubated in the dark for 30 minutes with 1ml of 2.5% Silver Nitrate (AgNO3 (BioShop) 2.5g and distilled water 100 mL). Following the incubation period plates were rinse with distilled water 3 times.

2.2.4.7. RT-PCR

To detect the expression of osteogenic genes, total RNA from monolayer cultures in 6-well plates of Runx2-HUCPVCs and Mock- HUCPVCs at day 3, 7, 14 and 21 was isolated using 1ml/well of Tri Reagent (Sigma-Aldrich) following the manufacturer’s recommended procedure. RNA content and quality was measured using the NanoDrop1000 (Thermo Fisher Scientific) and genomic DNA elimination and reverse transcription was performed according to the manufacturer’s instructions using 0.2-1µg of total RNA in the presence of gDNA Wipeout Buffer 7X, RNAse-free water, reverse transcriptase, RT Buffer 5X, and RT Primer mix, all from the QuantiTect Reverse Transcription Kit (Qiagen). PCR was carried out on the resulting cDNA following manufacturer’s recommendations in a 10.5µl reaction containing 1µl of 10X AccuPrime™ PCR Buffer I, 1µl primers mix, 0.5µl of cDNA and 0.05µl of AccuPrime™ Taq High Fidelity (Invitrogen) at an annealing temperature of 60°C for 35 cycles. The PCR products were then analyzed using 1% agarose gel electrophoresis with ethidium bromide staining. Gels were photographed using a gel photodocumentation system (Gel Doc 1000, Bio-Rad). Primer sequences are provided in Table 2.1.

2.2.4.7.1. Primer Design

All the primers were designed across exon junctions, in such a way that any product coming from the genomic DNA would be significantly larger than the expected PCR product coming from the transcript. For each specific gene, primers were designed with the online program Primer 3 (v. 0.4.0) (http://frodo.wi.mit.edu/primer3/input.htm) by providing the transcript sequence of the genes found in http://www.ensembl.org. In addition, primers were virtually tested with AmplifX software (http://ifrjr.nord.univ-mrs.fr/AmplifX).

Table 2.1 – PCR Primers for Osteoblastic Genes

Gene Forward primer Reverse primer Product Size GAPDH 5'-ACCACAGTCCATGCCATCAC-3’ 5'-TCCACCACCCTGTTGCTGTA-3’ 452 bp ALP 5'-CCTCCTCGGAAGACACTCTG-3’ 5'-AGACTGCGCCTGGTAGTTGT-3’ 239 bp Colα1 5'-AAATGGAGCTCCTGGTCAGA-3’ 5'-AGCTTCACCCTTAGCACCAA-3’ 184 bp OCN 5'-AGAGTCCAGCAAAGGTGCAG-3’ 5'-TCAGCCAACTCGTCACAGTC-3’ 171 bp OPN 5'-CCCACAGACCCTTCCAAGTA-3’ 5'-GGGGACAACTGGAGTGAAAA-3’ 244 bp Runx2-H 5'-CCTCCTACCTGAGCCAGATG-3’ 5'-ATGAAATGCTTGGGAACTGC-3’ 231 bp Runx2-M 5'-GCCGGGAATGATGAGAACTA-3’ 5'-GGACCGTCCACTGTCACTTT-3’ 232 bp H: human specific M: murine specific

2.3. Recombinant Adenoviral Transfection of HUCPVCs

2.3.1. Construction of Recombinant Adenoviruses

Four different recombinant replication defective adenoviruses carrying human cDNA fragments of BMP2, Runx2, SP7 or VEGF-A121 were constructed using the AdEasy™ XL Adenoviral Vector System (Stratagene) (Figure 2.1). The system employs the high efficiency of homologous recombination in E. coli coupled with selectable antibiotic resistance markers, to produce a recombinant adenovirus by a double recombination event between the adenoviral backbone plasmid vector, pAdEasy-1, and a shuttle vector carrying the gene of interest.

2.3.1.1. Cloning the Genes of Interest pShuttle-IRES-hrGFP-2 (Stratagene) was chosen as the shuttle vector since its design allows the monitoring of the expression of the gene of interest at the single-cell level due to the simultaneous expression of hrGFP from the same transcript through and internal ribosome entry site (IRES). The gene of interest is also cloned in frame with a C-terminal 3X HA tag to facilitate purification, detection (e.g. all the genes of interest can be detected at the protein

level using an anti-HA antibody; mRNA produced by the modified cells as a result of the genetic modification can be specifically distinguished by detecting mRNA containing the HA tag sequence) (Figure 2.2)(267).

Plasmid containing the cDNA of human Runx2 was purchased from Origene (SC302270, Accession # NM_001024630); human VEGFA (Clone ID# 6006890, Accession # BC_065522) and SP7 cDNA (Clone ID# 8069055, Accession # BC_101549) were purchased from Open Biosystems; a plasmid containing the human BMP2 cDNA (Accession #NM_001200) was obtained from Dr. Bernhard Ganss (University of Toronto, Faculty of Dentistry).

To insert the genes of interest into the multiple cloning site (MCS) region of the shuttle vector the restriction sites BglII and XhoI (Figure 2.2-B) were chosen. To do so, pShuttle-IRES-hrGFP-2 was digested with BglII and XhoI restriction enzymes in a total reaction volume of 60µl containing 50µl of pShuttle-IRES-hrGFP-2, 2µl of BglII (New England Biolabs), 2µl XhoI (New England Biolabs), 6µl of NEBuffer 3 (New England Biolabs) and 0.1mg/mL of BSA. This digestion was conducted for 2 hours at 37°C and fractioned in a 1% agarose gel (Figure 2.3-A) then gel- purified using the QIAquick Gel Extraction Kit (Quiagen) following the manufacturer’s recommended procedure.

In order to prepare the human cDNAs for ligation into the shuttle vector they were amplified from the respective plasmids by PCR using forward primers containing a BglII restriction site and reverse primers containing the XhoI site. In addition, a G nucleotide had to be added upstream of the start codon in all the forward primers to keep the sequences in frame with the 3X HA tag. PCR was performed in a total reaction volume of 25µl containing 2µl of 1/10 diluted template plasmid containing the gene of interest, 1µl of 0.1 μg/μL forward primer, 1µl of 0.1 μg/μL reverse primer, 2.5µl of 10x pfx Buffer, 2.5µl of 10mM dNTP mix, 1µl 50mM MgSO4, 1µl Platinum pfx DNA Polymerase (Invitrogen) and 14µl of DNase/RNase-Free Distilled Water (Invitrogen). Primers and PCR conditions are indicated in Table 3.2 and Table 3.3 respectively.

Figure 2.1 – Production of Recombinant Adenovirus Using the AdEasy XL Adenoviral Vector System (267).

Figure 2.2 – (A) pShuttle-IRES-hrGFP-2 map. (B) Multiple Cloning Site Region (MCS) (267).

Figure 2.3 – Electrophoresis (A) pShuttle-IRES-hrGFP-2 (B) PCR Amplified Runx2, SP7, VEGF and (C) BMP2 – all BglII/XhoI digested and run in a 1% agarose gel ready to be gel purified.

After PCR amplification, the products were digested with BglII and XhoI restriction enzymes in a total reaction volume of 60µl containing 50µl of PCR product of the gene of interest, 2µl of BglII (New England Biolabs), 2µl XhoI (New England Biolabs), 6µl of NEBuffer 3 (New England Biolabs) and 0.1mg/mL of BSA. This digestion was conducted for 2 hours at 37°C and fractioned in a 1% agarose gel, bands of the expected size were excised and purified using the QIAquick Gel Extraction Kit (Figure 2.3-B, 2.3-C). The final DNA fragments (BglII- gene of interest-XhoI) (full sequences can be found in the Appendix 2) were ligated in a total reaction volume of 20µl containing 1µl of BglII/XhoI linearized pShuttle-IRES-hrGFP-2, 6µl of BglII/XhoI digested gene of interest, 2µl of T4 DNA Ligase buffer, 1µl of T4 DNA Ligase (New England Biolabs) and 10µl of dH2O. The ligation was conducted at 16°C overnight.

Finally, 10µl aliquots of the ligation reaction containing the new shuttle vectors (pBMP2- Shuttle, pRunx2-Shuttle, pSP7-Shuttle and pVEGF-Shuttle) were transformed into 50µl of DH5α competent E.coli cells (Invitrogen) by heat shock at 42°C, followed by incubation for 1 hour in LB Broth culture media in a shaking incubator at 37°C and 300rpm. Cells were then pellet at 4000g in a table top centrifuge at room temperature and resuspended in 200µl of LB Broth to be plated onto LB-Kanamycin Agar (20g/L of LB broth (BioShop), 16g/L of Agar (BioShop), 10mg/ml of kanamycin (BioShop), dH2O). Plates were incubated for 16 hours at 37°C and 4 isolated colonies were selected for screening. Stab cultures of the colonies were incubated for 16 hours at 37°C in 3mL of LB-Kanamycin Broth (20g/L of LB broth, 10mg/ml of kanamycin, dH2O). Plasmids were isolated from bacteria cultures by QIAprep Spin Miniprep Kit (Qiagen). Eluted DNA was then diagnosed by restriction digestion with BglII/XhoI to confirm the presence of appropriately sized inserts in the shuttle vector. In addition, plasmid constructs were sequenced at the TCAG Sequencing Server (Hospital for Sick Children, Toronto, ON) to verify the accuracy of the insert sequence. Primers used for the sequencing reactions were: forward 5´-CTCACGGGGATTTCCAAGTC-3´ and reverse 5´-ATGCAGTCGTCGAGGAATTG-3´).

Table 2.2 – Primers Used to Amplify Human cDNA and to Introduce the BglII and XhoI Restriction Sites at the 5’ and 3’ ends of the Amplicons Respectively.

BMP2 SP7 Bgl II (forward) Bgl II (for) 5’-GTTC-AGATCTGATGGTGGCCGGGA -3’ 5’-GTTC-AGATCTGATGGCGTCCTCCCT-3’

Xho I (reverse) Xho I (rev) 5’-GTTC-CTCGAGGCGACACCCACAACC-3’ 5’-GTTC-CTCGAGCTCCAGCAAGTTGC-3’ Product size: 1201 bp Product size: 1303 bp

Runx2 VEGF 121 Bgl II (for) Bgl II (for) 5’-GTTC-AGATCTGATGGCATCAAAC-3’ 5’-GTTC-AGATCTGATGGCAGAAGGAG-3’

Xho I (rev) Xho I (rev) 5’-GTTC-CTCGAGATATGGTCGCCAAA-3’ 5’-GTTC-CTCGAGCCGCCTCGGCTT-3’ Product size: 1576 bp Product size: 370 bp

Color code: Restriction site Start codon Target cDNA

Table 2.3 – PCR temperature conditions for the amplification of the genes of interest.

Step Temperature (°C) Duration (h:m:s) 1 94.0 0:04:00 2 94.0 0:00:30 3 60.0 0:00:30 4 72.0 0:00:30 5 94.0 0:00:30 6 59.0 0:00:30 7 72.0 0:00:30 8 94.0 0:00:30 9 58.0 0:00:30 10 72.0 0:00:30 11 94.0 0:00:30 12 57.0 0:00:30 13 72.0 0:00:30 14 94.0 0:00:30 15 56.0 0:00:30 16 72.0 0:00:30 17 94.0 0:00:30 18 55.0 0:00:30 19 72.0 0:00:30 20 Back to step 17 Repeat 35 times 21 72.0 0:10:00 22 4.0 Hold 52

2.3.1.2. Homologous Recombination in vivo in Bacteria

This stage employs the efficient homologous recombination machinery in BJ5183-AD-1 cells, to produce a recombinant adenovirus plasmid by a double recombination event between the pAdEasy-1 (adenoviral backbone plasmid vector contained in BJ5183-AD-1 cells), and the pShuttle vector carrying the gene of interest (Figure 2.1). From this stage five different shuttle vectors were used, the four as described in the previous step, pBMP2-Shuttle, pRunx2-Shuttle, pSP7-Shuttle and pVEGF-Shuttle, and pMock-Shuttle which is the pShuttle-IRES-hrGFP-2 (Figure 2.2-A) without any gene cloned in the MCS, as negative control.

In order to get the shuttle vectors ready for the recombination step the restriction enzyme PmeI (New England Biolabs) was used to linearize the shuttle plasmids in a total reaction volume of 80µl containing 25µl of Shuttle vector, 8µl of NEBuffer 4, 0.8µl of BSA, 2µl of PmeI and 44.2µl of dH2O at 37°C for 16 hours. After the digestion the enzyme was removed using QIAquick PCR Purification Kit (Qiagen) and the linearized vectors were treated with alkaline phosphatase (New England Biolabs) in a total reaction volume of 60µl containing 50µl of

Linearized Shuttle vector, 2µl of alkaline phosphatase, 6µl of NEBuffer 3 and 2µl of dH2O at 37°C for 30 minutes and the gel purified using the QIAquick Gel Extraction Kit.

To produce the final recombinant adenoviral plasmids BJ5183-AD-1 (Stratagene) cells containing the pAdEasy-1 plasmid (the adenoviral backbone plasmid vector) were transformed by electroporation (GenePulser, Bio-Rad) with the linearized shuttle vectors and plated on LB- kanamycin agar plates overnight. The following morning 5-10 small well isolated colonies were picked with a sterilized loop from each recombination reaction and cultured in LB-kanamycin broth. Plasmids were purified using the Qiagen Miniprep Kit, digested with PacI restriction enzyme (New England Biolabs) in a total reaction volume of 20µl containing 10µl of miniprep recombinants, 1µl of PacI, 2µl of NEBuffer 1, 2µl of 1/10 BSA and 5µl of dH2O and ran on a

0.8% agarose gel to screen for proper recombination.

2.3.1.3. Recombinant Adenoviral Plasmids Amplification

In this part of the process individual positive recombinant adenovirus plasmids, identified by restriction digestion with PacI in the previous step, were used to transform XL10-Gold ultracompetent E. coli cells (Stratagene) to create a continuous source of the recombinant adenovirus plasmid DNA. Following the AdEasy™ XL Adenoviral Vector System manual (267) colonies obtained after the XL10-Gold transformation were cultured in 500 ml LB-kanamycin broth for ~16 hours in a 37°C shaking incubator, set at 300 rpm. Plasmid DNA was isolated using EndoFree Plasmid Maxi Kit (Qiagen, 12362) for 10 different recombinant adenoviral constructs, two for each gene: pBMP2-rAV(1), pBMP2-rAV(2), pRunx2-rAV(1), pRunx2-rAV(2), pSP7-rAV(2), pSP7-rAV(3), pVEGF-rAV(1), pVEGF-rAV(2), pMock-rAV(7) and pMock-rAV(8).

Before the virus generation in AD-293 cells recombinant adenoviral plasmids were linearized with PacI restriction enzyme (New England Biolabs) in a total reaction volume of 60µl containing 50µl of recombinant adenovirus plasmid, 2µl of PacI, 6µl of NEBuffer 1, 0.6µl of BSA and 1.4µl of dH2O at 37°C for 16 hours. To remove buffer and enzyme from the restriction reactions ethanol precipitation was carried out by adding 150µl of 95% Ethanol and 6µl of NaOAc (3M pH:4.8) followed by centrifugation at 12000g for 30 minutes at 4°C. Supernatant was removed and 500µl of 70% ethanol were added. After a brief centrifugation DNA pellet was air dried and resuspended in 50µl of H2O. DNA content and quality was measured using NanoDrop1000 Spectrophotometer.

2.3.1.4. Generating Recombinant Adenoviruses in AD-293 Packaging Cells

AD-293 cells (Stratagene) an adenovirus packaging cell line derived from HEK293 were grown in DMEM (Gibco) supplemented with 10% FBS at 37°C and 5% CO2 as per supplier’s recommendations. For transfection, cells were seeded at a concentration of 4X106 cells per T75 flask in 10 ml of growth media. At a confluence of ~70% cells were transfected by calcium phosphate co-precipitation as described in section 3.2.4.1 Transfection efficiency and virus

production was monitored by GFP expression using fluorescence microscopy. Transfected cells were maintained in a 37°C, 5% CO2 incubator for 15-20 days without passaging and changing the medium, 1–2 ml of fresh growth medium were added every 5–7 days. Cells were scraped off the flasks with a cell scraper and transfered to a 50ml tube and centrifuged for 10 minutes at 500g at 4°C. All but 3ml of medium was removed and the pellet was resupended by vortexing. Adenoviruses were released from the cells by performing four freeze–thaw–vortex cycles, centrifuged again to pellet cell debris and stored as a primary virus stock at -80°C.

2.3.1.5. Amplification of Recombinant Adenoviruses

To achieve higher titer, viruses were amplified as follows (268): AD-293 cells were seeded at a concentration of 6X106 cells per T75 flask in 10 ml of growth medium. At 80-90% confluency 1ml of primary virus stock was added to the flask. After 3 to 5 days of infection when cells started to detach cells and medium were collected in a 50ml tube and centrifuged for 10 minutes at 500g at 4°C. All but 6ml of medium was removed and the pellet was resupended by vortexing. Four freeze–thaw–vortex cycles were performed and adenoviruses were centrifuged again to pellet cell debris and stored as a secondary virus stock at -80°C. The final amplification round was done by seeding 1.4X107 cells per T150 Flask in 25ml of growth medium. At 90% confluency 1.5ml of secondary virus stock was added to the flask. After 3 to 5 days adenoviruses were collected as described before and passed through a 0.45μm filter and stored at -80°C in 1ml aliquots for future use. After this stage 5 different virus preparations were obtained Ad-BMP2, Ad-Runx2, Ad-SP7, Ad-VEGF and Ad-Mock.

2.3.2. HUCPVCs Infection with Recombinant Adenoviruses

HUCPVCs were plated in 6-well plates at a concentration of 3 X 105/well in alpha-MEM supplemented with 5% FBS, approximately 18 hours later, when 90% confluency was reached, cells were exposed to the virus overnight diluted in 1000µl of growth medium per well. In the morning cells were washed three times with PBS and fresh growth medium was added. After 55

48 hours, cell viability was measured with the Vi-cell and transfection efficiency was assessed by florescence microscopy and flow cytometry.

2.3.2.1. Viral titer and Multiplicity of Infection

Adenoviruses were tested by infecting HUCPVCs with different volumes of virus until finding the optimal Multiplicity of infection (MOI) that resulted in maximum efficiency of transfection and cell viability. The titer (infectious units/ml) of each virus was calculated using the following formula (269,270):

The MOI was calculated using the following formula:

For all subsequent experiments HUCPVCs were infected at a MOI of 10 which achieved transfection efficiencies higher than 90% and cell viability higher than 85%.

Since HUCPVCs were infected only with one of the viruses, five different groups of genetically modified HUCPVCs were generated, BMP2-, Runx2-, SP7-, VEGF- and Mock-HUCPVCs.

2.3.3. Confirmation of mRNA Production of the Genes of Interest in Genetically modified HUCPVCs

2.3.3.1. RT-PCR

Total RNA from monolayer cultures in 6-well plates of BMP2-HUCPVCs, Runx2-HUCPVCs, SP7- HUCPVCs, VEGF-HUCPVCs and Mock HUCPVCs was isolated using 1ml/well of Tri Reagent (Sigma-Aldrich) following the manufacturer’s recommended procedure. RNA content and quality was measured using the NanoDrop1000 (Thermo Fisher Scientific) and genomic DNA elimination and reverse transcription was performed as described in section 2.2.4.2 PCR was carried out on the resulting cDNA following manufacturer’s recommendations in a 10.5µl reaction containing 1µl of 10X PCR Buffer, 1µl primers mix, 0.3µl of 50M MgCl2, 0.25µl of 10 mM dNTP, 0.5µl of cDNA and 0.04µl of Platinum® Taq DNA Polymerase (Invitrogen) at an annealing temperature of 60°C for 35 cycles. The PCR products were then analyzed using 1% agarose gel electrophoresis with ethidium bromide staining. Gels were photographed using a gel photodocumentation system (Gel Doc 1000, Bio-Rad). Presence of mRNA transcripts for the genes of interest was confirmed by using specific primers (Table 2.4) to detect only transcripts that have the XhoI restriction site and the HA tag sequence (part of the recombinant adenoviral vector) (Figure 2.2-A). This strategy allows only the detection of transcripts being produced as a result of the genetic modification.

2.3.3.1.1. Primer Design

To ensure detection of part of the HA tag and the XhoI restriction site reverse primers were design by hand writing the inverse complementary sequence of the last 10 nucleotides of the genes of interest sequence, the 6 nucleotides that formed the XhoI restriction site and the first 5 nucleotides of the HA tag (this is part of the sequence in where the gene of interest is fused with the recombinant vector - full sequence of all the genes of interest are provided in the

Appendix 2). Forward primers were designed using the online tool Primer-BLAST (http://www.ncbi.nlm.nih.gov/tools/primer-blast/) by entering the full sequence for each gene (provided in the Appendix 2) and by entering the hand designed reverse primer for the corresponding gene (Table 2.4). In addition, primers were virtually tested with AmplifX software (http://ifrjr.nord.univ-mrs.fr/AmplifX).

Table 2.4 – Primers for the Specific Detection of Transcripts Produced as a Result of The Genetic Modification with Recombinant Adenoviruses

Gene Forward primer Reverse primer Product size Β-actin 5’-ACCATCACGCCCTGGTGCCT-3’ 5’-ACCCATGCCCACCATCACGC-3’ 199 bp BMP2 5’-GGACTGCACAGGGACACGCC-3’ 5’-GGGTACTCGAGGCGACACCCA-3’ 553 bp Runx2 5’-TTCCGCCATGCACCACCACC-3’ 5’-TGGGTACTCGAGATATGGTCGCC-3’ 167 bp SP7 5’-GCACCCACGGAGAACCAGGC-3’ 5’-TGGGTACTCGAGCTCCAGCAAGT-3’ 182 bp VEGF 5’-TGTGTGCCCCTGATGCGATGC-3’ 5’-GGGTACTCGAGCCGCCTCGGC-3’ 224 bp Color code: HA tag XhoI Restriction site Target cDNA

2.3.4. Verification of the Production of the Proteins of Interest by Modified HUCPVCs.

2.3.4.1. Western Blots

To detect the expression of the genes of interest at a protein level BMP2-, Runx2-, SP7- and VEGF-HUCPVCs were washed with PBS and lysed by adding Cellytic M Cell Lysis Reagent (Sigma-Aldrich) with 1X Protease Inhibitor Cocktail (Sigma-Aldrich) and centrifuged at 500g at 4°C for 5 minutes to remove cell debris. Samples were boiled in protein loading buffer (60 mM

Tris, 10% glycerol, 2% SDS, 0.1% bromophenol blue and 400mM DTT). SDS-PAGE was carried out on a 12% Minigel (Mini-PROTEAN II Tube Cell, Bio-Rad), then blotted onto an Immobilon-P

membrane (Millipore) using a semi-dry transfer apparatus at 100mA per gel for 1 hour. Once transferred, the membranes were incubated in blocking solution (5% skim milk in TBS with 0.1% Tween 20 (Bio-rad)) overnight. Membranes were then incubated for 1 hour at room temperature with gentle rocking with 1:2000 anti-HA (Covance, MMS-101R) or 1ug/ml anti-SP7 (Abcam) or 1ug/ml anti-Runx2 antibody (Abcam) in blocking solution. Unbound primary antibody was removed from the membranes by washing five times with TBS with 0.1% Tween (TBS-T) for 5 minutes each time. The membranes were then incubated with the corresponding secondary antibody 1:5000 Goat pAb to Ms IgG (HRP) (Abcam) or 1:10000 Goat Anti-Rabbit IgG (H+L)-HRP Conjugate (Bio-Rad) diluted in blocking solution at room temperature under gentle rocking. After 1 hour membranes were washed five times with TBS-T for 5 minutes each time. Immunoreactive signals were detected using the Immun-Star Western C chemiluminescent kit (Bio-Rad) according to the manufacturer’s instructions, capturing the signal on a 8” x 10” X-ray film (Clonex). The X-ray films were developed and photographed using a gel photodocumentation system (Gel Doc 1000, Bio-Rad Laboratories).

2.3.4.2. ELISA

To measure the amount of BMP2 and VEGF-A121, BMP2-HUCPVCs, VEGF-HUCPVCs, Mock- HUCPVCs and naive HUCPVCs from three different cords (n=3) were individually seeded in 6- well plates (105 cells/ml), 24 hrs after, conditioned media were collected and centrifuged at 500g at 4°C for 5 minutes to remove cell debris. The quantity of BMP2 and VEGF-A was measured by Enzyme-Linked ImmunoSorbent Assay (ELISA) using Human BMP2 DuoSet (R&D Systems, DY355) and Human VEGF DuoSet (R&D Systems, DY293B) as per manufacturer’s instructions. Briefly, MaxiSorp plates (NUNC, 439454) were coated overnight with the capture antibody. In the morning wells were washed with wash buffer (0.05% Tween 20 in PBS) three times and blocked with 1% BSA in PBS, followed by incubation for 1 hour with 100µl of conditioned media per microwell. After washing, the plates were incubated for 1 hour with detection antibody. Following another wash step Streptavidin-HRP was added to each well for 20 minutes and washed. Substrate Solution (R&D Systems) was added to the plate for color

development and stopped after 20 minutes with Stop Solution (R&D Systems). Finally, sample absorbance at 450 nm was recorded using a Microplate Reader (Synergy HT Multi-Mode, BioTek). Using the standards, a standard curve was constructed and the amount of BMP2 or VEGF-A in the samples was interpolated.

2.3.5. Longevity of GFP Expression by HUCPVCs in Different Growing Conditions

To evaluate the effect of proliferation on the longevity of HUCPVC genetic modification with recombinant adenoviruses cells were grown in different conditions and studied by measuring the number of GFP positive cells by flow cytometry at different time points. HUCPVCs from 4 different cords were mixed and transfected with Ad-Mock as explained in section 2.3.2 . Three days after infection cells were trypsinized, GFP expression was quantified by flow cytometry (day 0) and cells were seeded in 12-well plates with alpha-MEM under different conditions: 1% FBS, 2% FBS, 5% FBS and Mitomycin C treated HUCPVCs in 5% FBS. Before transfection with Ad-Mock a portion of HUCPVCs were treated with Mitomycin C which inhibits DNA replication by forming crosslinks between the complementary strands of DNA arresting cell growth (271). Briefly, HUCPVCs were plated in 6-well plates at a concentration of 3X105/well after attachment overnight cell were incubated in culture medium supplemented with 15 μg/ml of Mitomycin C (Sigma-Aldrich) for 2 hours. Following the incubation, the mitomycin C supplemented medium was removed and the cells were washed 3 times with PBS before adding fresh medium. After 6 hours HUCPVCs were transfected with Ad-Mock. The percentage of GFP positive cells was measured at day 3, 7, 14 and 21 by flow cytometry for each growth condition.

2.3.6. Bioactivity Test of BMP2 Produced by BMP2- HUCPVCs

BMP2-HUCPVCs and MOCK-HUCPVCs from different cords (n=5) were cultured in alpha-MEM supplemented with 2% FBS from cells grown at a concentration of 1x105 cells/ml in 6-well plates. The conditioned media were collected after 48 hours. To test the bioactivity of the 60

BMP2 produced by the BMP2-HUCPVCs, C2C12 cells were grown in these conditioned media. C2C12 is a mouse muscle derived cell line that when exposed to BMP2 are driven from a default myogenic to an osteogenic pathway, characterized by induced levels of ALP activity (137). The ALP activity of the cells increases proportionately to the amount of BMP present in the medium (272).

2.3.6.1. C2C12 Cell Culture and Assay

Frozen aliquots of C2C12 cells (ATCC) were kindly provided by Dr. Sean Peel (Faculty of Dentistry, University of Toronto). The cells were thawed and expanded at 37°C and 5% CO2 in αMEM supplemented with 10% FBS and 10% antibiotics. Cells were passaged at 60-80% confluence to prevent cell differentiation. C2C12 cells were seeded in 24 well plates at a concentration of 50x103 cells/well in 500µl of growth media and were allowed to attach overnight. In the morning the medium was changed to test medium which consisted of centrifuged BMP2-HUCPVCs or MOCK-HUCPVCs conditioned media supplemented with 15% FBS. Each conditioned medium sample was tested in 3 wells. Additionally, as a negative control C2C12 cells were grown in αMEM supplemented with 15% FBS and, as a positive control, with αMEM supplemented with 15% FBS plus 40ng/mL rhBMP2 kindly provided by Dr. Sean Peel. After 3 days of incubation, cultures were terminated by rinsing three times with 1X TBS (Bio- Rad) and adding 500µl of Cellytic M Cell Lysis reagent per well, sealing the 24-well plate with parafilm, and freezing at –20°C for future use.

2.3.6.2. Alkaline Phosphatase Assay

The standard method to measure ALP activity is to incubate the samples with a substrate buffer consisting of p-nitrophenol phosphate (pNPP) in an alkaline buffer. In the presence of ALP the colourless pNPP is cleaved to release p-nitrophenol (pNP) which is yellow. The amount of pNP released is then determined colorimetrically and interpolated on a strandard curve

generated using different concentrations of pNP. ALP activity is expressed as nmol pNP generated/min.

To prepare the samples, frozen culture dishes were thawed and cell layers were scraped into Eppendorf tubes and vortexed. Cell lysates were centrifuged at 12000g for 10 min at 4°C to pellet cell debris and supernatants were transferred to new tubes. To measure ALP activity 20µl of cell lysates were pipetted into the wells of a 96-well microplate in triplicate and incubated with 200µl substrate buffer (25ml of p-nitrophenol phosphate (4mg/ml in H2O) (Sigma-Aldrich), 25ml alkaline buffer solution (Sigma-Aldrich)) at 37°C. After 30 minutes the reactions were stopped by adding 20µl of 1N NaOH and the absorbance was measured at 405 nm using a Synergy HT Multi-Mode Microplate Reader (BioTek). The standard curves were generated using p-nitrophenol (pNP) solution (Sigma-Aldrich), which is the end product of the ALP assay, diluted in 0.02 N NaOH.

2.3.6.3. Protein Assay

The Pierce 660nm Protein Assay (ThermoScientific) was used to quantify the total protein content by incubating 10µl of the cell lysates with 150µl of the Protein Assay Reagent for 5 minutes as per manufacture’s recommendations. The absorbance was measured at 660nm using a Synergy HT Multi-Mode Microplate Reader (BioTek). The standard curve was generated using various concentrations of BSA diluted in Cellytic M Cell Lysis reagent.

2.3.7. Effect of Liquid Nitrogen Storage on Genetically Modified HUCPVCs

To determine if the process of freeze-store-thaw has an effect on the production and bioactivity of proteins expressed in genetically modified HUCPVCs different assays were performed comparing HUCPVCs before and after storage in liquid nitrogen.

2.3.7.1. Cryopreservation of HUCPVCs

Three days after HUCPVC infection, cells were trypsinized and centrifuged at 285g for 5 minutes at 4°C. Cells were resuspended in freezing medium (700µL of alpha-MEM, 200µL of FBS and 100µL of DMSO (Sigma-Aldrich)) at a concentration of 1x106 cells/ml and transferred to a cryovial (VWR, 66008-754). The vials were placed in the fridge for 5 minutes to allow the DMSO to penetrate and then transferred to a Mr. Frosty container (Sigma-Aldrich) and placed in -70°C freezer overnight. The next day, the vials were transferred to liquid nitrogen storage and were kept there for at least 2 days. These cryopreserved cells are referred to as post-liquid nitrogen (post-LN) cells. Before using the cells for the following experiments LN-stored-cells were thawed and cell viability (recovery) was measured as explained in section 2.1.

2.3.7.2. Bioactivity of BMP2 Produced by Post-Liquid Nitrogen BMP2- HUCPVCs vs. Pre-Liquid Nitrogen BMP2-HUCPVCs

To test if the cryopreservation of BMP2-HUCPVCs has an effect on the bioactivity of the BMP2 produced pre- and post-LN BMP2-HUCPVCs and pre- and post-LN MOCK-HUCPVCs from five different cords (n=5) were cultured in alpha-MEM supplemented with 2% FBS, after 48 hours the conditioned media were collected. To assay the bioactivity of the BMP2 produced by the BMP2-HUCPVCs, C2C12 cells were exposed to the conditioned media for 3 days. The activity of secreted BMP2 from pre- and post-LN BMP2-HUCPVCs was compared using the alkaline phosphatase activity assay on C2C12 cells that were grown in conditioned medium as described under section 2.3.6.

2.3.7.3. Comparison of Efficiency of Protein Production Between Pre- and Post-Liquid Nitrogen VEGF-HUCPVCs

To evaluate if the cryopreservation of VEGF-HUCPVCs has an effect on the efficiency of protein production, pre- and post-LN VEGF-HUCPVCs and pre- and post-LN MOCK-HUCPVCs from three different cords (n=3) were cultured in alpha-MEM supplemented with 2% FBS, after 63

conditioning the medium for 24 hours it was collected at 3, 7, 11, 14, 17 and 21 days for the determination and comparison of VEGFA levels by ELISA using Human VEGF DuoSet (R&D Systems) as per manufacturer’s instructions. The ELISA procedure is further explained in section 2.3.4.2.

2.4. In vitro Evaluation of Cocktails of Genetically Modified HUCPVCS

To determine how the configuration of “cocktails” affects the levels of expression of multiple osteogenic markers, different genetically modified HUCPVCs were mixed, cultured and analyzed for the expression of osteogenic markers through different methods.

2.4.1. Cell Culture of Different Groups and Cocktails of Genetically Modified HUCPVCs

HUCPVCs derived from different umbilical cords (n value is determined in each section) were genetically modified (section 2.3.2) with Ad-BMP2, Ad-Runx2, Ad-SP7 or Ad-Mock (For this in vitro study VEGF-HUCPVCs were not utilized since they were only intended to be used to induce angiogenesis in vivo.) Three days later cells were trypsinized, cell viability was measured by the Vi-cell and transfection efficiency was assessed by flow cytometry. Different groups and cocktails of genetically modified HUCPVCs were cultured in triplicates in 24-well plates at a concentration of 3 x 104 cells/well in osteogenic media (alpha-MEM supplemented with 2% FBS, 10% antibiotic stock solution, 50µg/ml ascorbic acid (Sigma, A4544), 10-8M dexamethasone (Sigma-Aldrich) and 3.5mM -glycerophosphate (Sigma-Aldrich)).

The different groups and cocktails of cells that were cultured were:

1. BMP2-HUCPVCs

2. Runx2-HUCPVCs

3. SP7-HUCPVCs

4. BMP2-HUCPVCs + Runx2-HUCPVCs (50% of each)

5. BMP2-HUCPVCs + SP7-HUCPVCs (50% of each)

6. Runx2-HUCPVCs + SP7-HUCPVCs (50% of each)

7. BMP2-HUCPVCs + Runx2-HUCPVCs+ SP7-HUCPVCs (33.3% of each)

8. Mock-HUCPVCs

These proportion of cells chosen for each cocktail (mix of different types of genetically modified HUCPVCs) were originally decided to study the equal roll of each type of cell to later based in the results obtained start modifying the ratios depending on the outcomes.

2.4.2. ALP Staining and Quantification

At day 7 histochemical staining of cultures was performed to identify cells that exhibited the ALP enzyme. Cultures were rinsed in TBS three times and fixed in 10% ice-cold neutral formalin buffer (Sigma, HT501128) for 15 minutes. Cultures were then rinsed with distilled water and incubated with 500µl of ALP substrate solution (0.005g of Naphthol AS MX-P04, (Sigma- Aldrich), 200µl of N,N-Dimethylformamide (Sigma-Aldrich) 25ml TrisHCL (O.2M, pH 8.3), 25ml distilled water and 0.03g Fast Red Violet LB salt (Sigma-Aldrich)) for 45 minutes at room temperature. Following the incubation period, cultures were rinsed in distilled water 3 times and photographed using an inverted light microscope.

ALP was also quantified at day 3, 7 and 14 cultures from six different cords (n=6) were terminated by rinsing with TBS three times, then 500µl of Cellytic M Cell Lysis reagent were added and 24-well plates were sealed with parafilm and stored at –20°C for future use. ALP activity was measured and normalized to protein content following the processes in sections 2.3.6.1 - 2.3.6.3.

2.4.3. Von Kossa Staining

At day 21, 28 and 35 cultures of the different groups and cocktails of HUCPVCs were stained with von Kossa for the detection of calcium deposition as described in section 2.2.4.1.

2.4.4. RNA Extraction

At day 2, 6 and 9 RNA from monolayer cultures from three different cords (n=3) was isolated using 500µl of Tri Reagent (Sigma-Aldrich) following the company’s recommended procedure. RNA was then purified by ethanol precipitation described in section 2.3.1.3. RNA content and quality was measured using NanoDrop1000.

2.4.5. Quantitative RT-PCR

Gene specific primers for ALP, COL1A1, OCN, OPN and Beta-2-microglobulin (β2M) (Table 3.5) were designed across exon junctions when possible, using the NCBI online tool Primer-BLAST (http://www.ncbi.nlm.nih.gov/tools/primer-blast/) by providing the accession number of the osteogenic gene of interest found in the NCBI nucleotide search (http://www.ncbi.nlm.nih.gov/nuccore). In addition, primers were virtually tested with AmplifX software (http://ifrjr.nord.univ-mrs.fr/AmplifX). qRT-PCR was carried out in triplicates using iScript One-Step RT-qPCR with SYBR Green (Bio- Rad, 170-8893) according to the manufacturer’s recommendations in a total reaction volume of 10µl containing 6µl of 2X SYBR Green RT-PCR Reaction Mix, 0.2µl of iScript Reverse

Transcriptase for One-Step RT-PCR, 18ng of RNA, 1µl of primer mix and Nuclease-free H2O at an annealing temperature of 60°C for fluorescence measurement. Quantification was done utilizing the Pfaffl method (273), in which a mathematical model determines the relative quantification of a target gene (e.g. ALP, COL1A, OCN, OPN) in comparison to a reference gene (e.g. β2M). The relative fold expression (R) of a target gene is calculated based on the real-time

PCR efficiency (E) and the Ct deviation of an unknown sample (e.g. BMP2-HUCPVC, SP7- HUCPVCs, Runx2-HUCPVCs + SP7-HUCPVCs) versus the control (Mock-HUCPVCs) and expressed normalized to β2M using the following formula:

Table 2.5 – Quantitative RT-PCR Primers for Osteogenic Genes

Gene Forward primer Reverse primer Efficiency Product Size β2M CTCCGTGGCCTTAGCTGTG TTT GGAGTACGCTGGATAGCCT 96% (1.96) 69bp ALP GTATGAGAGTGACGAGAAA GTTCCAGATGAAGTGGGAGTG 92% (1.92) 109bp COL1A1 CCCCTGGAAAGAATGGAGA TCCAAACCACTGAAACCTCTG 93% (1.93) 148bp OCN CTGGCCAGGCAGGTGCGAA CGGGTAGGGGACTGG GGCTC 99% (1.99) 140bp OPN CCCACAGACCCTTCCAAGTA GGGGACAAC TGGAGTGAAAA 93% (1.93) 244bp

2.5. In Vivo Osteoinductive and Osteogenic Potential of Intramuscular Injected Genetically Modified HUCPVCs

2.5.1. Animal Subjects

All procedures carried out on animals were approved by the University of Toronto Animal Care Committee (Protocol # 20009531). Six, 6 week old, male NOD/SCID mice (Charles River, Strain code 394) were kept in a sterile facility and given free access to food and water throughout the study period.

2.5.2. Intramuscular Cell Delivery

NOD/SCID mice were anesthetized using cone inhalation of isofluorane in O2 (flow rate 400 ml/min) and N2O (flow rate 800 ml/min), 3-5% induction, 1.5-2.5% maintenance. After the animals were anesthetized the skin over the area to be injected was shaved (Figure 2.4-A). 5x106 genetically modified HUCPVCs (mixed at equal proportions from 5 different cords) suspended in 100µl of sterile PBS were delivered through intramuscular injection into the hind limb quadriceps muscle using a 1 ml syringe and a 25 G needle (Figure 2.4-B). After 6 weeks mice were euthanized using carbon dioxide inhalation. For this study four different experimental groups were used:

1. Mock-HUCPVCs

2. VEGF-HUCPVCs

3. BMP2-HUCPVCs + Runx2-HUCPVs + Sp7-HUCPVCs (33% each)

4. BMP2-HUCPVCs + Runx2-HUCPVs + Sp7-HUCPVCs + VEGF-HUCPVCs (25% each)

A. B.

Figure 2.4 – Intramuscular Delivery of Genetically Modified HUCPVCs (A) Mice were anesthetized using cone inhalation; skin over the area to be injected was shaved and disinfected. (B) Intramuscular injection into the quadriceps muscle.

2.5.3. X-rays

Ectopic bone formation was monitored at 2, 4 and 6 weeks post intramuscular injection. Animals were anesthetized as described in section 2.5.2 and ventrodorsal view x-rays were taken for 0.40 sec at 50kvp on a digital x-ray machine (CR 30-X, AGFA HealthCare).

2.5.4. Micro-computed Tomography

To detect any calcified tissue formed in the area where the cells were injected hind limbs were harvested at week 6 and fixed in 10% neutral formalin buffer. After 24 hours samples were placed in a poly-methyl-metacrylate (PMMA) holder with formalin and scanned using a microcomputed tomography system (MicroCT40, Scanco Medical, Basserdorf, Switzerland) at 70 kVp and 114 μA in high-resolution mode with a X, Y, and Z resolution of 6 μm.

2.6. rhFurin Cleavage of Pro-BMP2 secreted by BMP2-HUCPVCs

In order for BMP2 to become a mature bioactive protein Pro-BMP2 has to be cleaved by Furin (148) at the dibasic sequence motif R-X-X-R (147). In the following experiments rhfurin was used to investigate if it was possible to increase the yield of mature bioactive BMP-2 secreted by BMP2-HUCPVCs.

2.6.1. HUCPVCs, rBMSC and hBMSCs BMP2-adenoviral Infection

HUCPVCs, hBMSCs and rBMSCs were genetically modified using the Ad-BMP2 and the Ad- Mock. Adenoviral infections in hBMSCs and rBMSCs were optimized by trying different concentrations of virus until finding the optimal Multiplicity of infection (MOI) that resulted in maximum efficiency of transfection and cell viability. Cells were infected as described in

section 3.3.2. at a MOI of 10 for HUCPVC, 50 for hBMSCs and 80 for rBMSCs, MOIs that yielded transfection efficiency >85% and cell viability >80% for each specific type of cell.

2.6.2. ELISA

Following infection cells were seeded in 6-well plates at a concentration of 1 X 105 cells/ml in alpha-MEM supplemented with 2% FBS for HUCPVCs and 5% FBS for hBMSCs and rBMSCs After 24 hours conditioned media were collected and centrifuged at 500g at 4°C for 5 minutes to remove cell debris. The amount of BMP2 was measured by ELISA using Human BMP2 DuoSet (explained in detail section 2.3.4.2.)

2.6.3. Treatment of BMP2-HUCPVCs Conditioned Media with rhFurin

BMP2-HUCPVCs were cultured in 6-well plates at a concentration of 1 X 105 cells/ml in alpha- MEM alone, after 48 hours conditioned media were collected and incubated with different concentrations (1000, 100, 10 and 0 ng/ml) of rhfurin (R&D Systems) in 2X cleavage buffer (50mM Tris, 2mM CaCl2, 1% Triton-X, pH 9.0) at 1:1 ration for 16 hours at 30°C.

2.6.3.1. ELISA

After furin treatment the BMP2 in the cleavage reactions (1 part conditioned medium: 1 part 2x cleavage buffer) was measured by ELISA using Human BMP2 DuoSet (R&D Systems) (section 2.3.4.2.) to determine if higher levels of mature BMP2 were obtained after furin treatment.

2.6.3.2. Western Blot

Following furin treatment 20µl of cleavage reaction were boiled in protein loading buffer followed by SDS-PAGE on a 12% gel. Once the proteins were transferred to the membrane it

was probe with 1:2000 anti-HA antibody and 1:5000 Goat pAb to Ms IgG (HRP) (western blot procedure explained in detail in section 2.3.4.1.)

2.6.3.3. Test of bioactivity of Cleaved BMP2

After treatment of BMP2-HUCPVCs conditioned media with 1000ng/ml or 0ng/ml of rhfurin the cleavage reaction solution (1 part conditioned medium: 1 part 2X cleavage buffer) was treated to remove the containing Triton-X by using DetergentOUT (G-Bioscienses) as per manufacturer’s instructions. To test bioactivity of cleaved BMP2 C2C12 cells were plated in 24- well plates at a concentration of 5X104 cells/well in 500µl and allowed to attach overnight. In the morning media were changed to the cleavage reaction solution supplemented with 15% FBS and 10% antibiotics stock. After 3 days cultures were terminated by rinsing with 1X TBS (Bio-Rad) three times and adding 500µl of Cellytic M Cell Lysis reagent. Plates were sealed with parafilm and frozen at –20°C for future use. ALP activity assay was performed as explained in sections 2.3.6.1 - 2.3.6.3.

2.7. Osteogenic Cultures of Genetically Modified Human Bone Marrow Stromal Cells (HBMSCs) vs. HUCPVCs

2.7.1. HBMSCs and HUCPVCs Culture

HBMSCs were kindly provided by Dr. Armand Keating (Princes Margaret Hospital). The cells were expanded at 37°C and 5% CO2 in low Glucose D-MEM supplemented with 10% FBS and 10% antibiotic stock solution (1% Penicillin, 1% Gentamicin and 0.3 mg/mL amphotericin B). Medium was changed 2-3 times a week; when confluence was around 80-90% cells were passaged at a ratio of 1:3 using 0.05% trypsin/0.02% EDTA solution. HUCPVCs were cultured as described in section 2.1.

2.7.2. HBMSCs and HUCPVC infection

HBMSCs and HUCPVCs from three different donors each (n=3) were infected as described in seccion 2.6.1. by exposing the cells to Ad-Runx2, Ad-SP7 or Ad-Mock. After 48 hours cell viability was measured with the Vi-cell and transfection efficiency was assessed by flow cytometry.

2.7.3. Osteogenic cultures of genetically modified HBMSCs and HUCPVCs

To test and compare the osteogenic capacity of genetically modified HBMSCs and HUCPVCs cultures were kept in parallel and cells were cultured in triplicates in 24-well plates at a concentration of 3 x 104 cells/well in osteogenic medium (DMED low glucose, 10% FBS, 50µg/ml ascorbic acid, 10-8M dexamethasone and 3.5mM -glycerophosphate added when cells reached confluence around day 12). Four different groups of modified cells were cultured for the two different types of cells: Runx2, SP7, Runx2/SP7 (50% of each) and Mock.

2.7.4. ALP staining and Quantification

At day 7 and 14 histochemical staining of cultures was performed as described in section 2.4.2 to identify cells that exhibited ALP.

ALP was also quantified at day 7 and 14 by a colorimetric assay as explained in section 2.3.6.1 - 2.3.6.2. and normalized to protein content as described in section 2.3.6.3.

2.7.5. Alizarin Red Staining

To detect mineralized areas in the culture dishes at day 21 cultures were rinsed in TBS three times and fixed in 10% ice-cold neutral formalin buffer for 15 minutes. Cultures were then

rinsed with distilled water and incubated with 400µl of 2% Alizarin Red solution (Alizarin Red (Sigma-Aldrich), 2.0 g, 100ml distilled water, ammonium hydroxide, pH 4.2) for 5 minutes at room. Following the incubation period, cultures were rinsed in distilled water 3 times and photographed.

2.7.6. Calcium Content Quantification

At day 21 genetically modified HBMSCs cultures were rinsed with TBS two times and decalcified with 300µl/well of 0.6M HCl for 24 hours at 4°C in a moving platform. After removing the HCl supernant, the remaining cell layers were solubilized with 300µl/well of a solution of NaOH 0.1M and SDS 0.1% for protein concentration analysis as described in section 2.3.6.3. The calcium content in the HCl supernatant was determined by the QuantiChrome Calcium Assay Kit following the manufacturer's protocol and normalized by protein concentration.

2.7.7. Osteogenic Cultures in Naive and Engineered HBMSCs

To investigate if the genetic modification was having an effect in the osteogenic differentiation of HBMSCs, cultures of Mock-HMBSCs were cultured in parallel and compared with Naive- HBMSCs (non-engineered) while cultured in osteogenic medium (DMED low glucose, 10% FBS, 50µg/ml ascorbic acid, 10-8M dexamethasone and -glycerophosphate added when cells reached confluence around day 12). For this experiment two different concentrations of - glycerophosphate were tested 3.5mM and 5mM; and as a negative control Mock-HBMSCs and Naive-HBMSCs were cultured in growth medium (D-MEM Low Glucose, 10% FBS).

2.8. Statistical Analysis

Statistical analysis was performed by means of a two-sample Student’s T-test after logarithmic transformation in Excel 2007 (Microsoft, Redmond, WA) or by analysis of variance (ANOVA) and Tukey's test for pairwise comparisons using SigmaPlot 12 (Systat Software). Each figure indicates which statistical analysis was used, number of samples and the p value.

3. Results

3.1. Genetic modification of HUCPVCs by non-viral methods.

3.1.1. FuGENE 6

To assess transfection capacity HUCPVCs were transfected under different conditions (Table 3.1) at a confluence of 70-80% in 6-well plates with a GFP reporter plasmid (pmaxGFP) using the nonliposomal reagent FuGene 6. Three days post transfection it was determined by flow cytometry that a ratio of 1:3 of plasmid DNA to volume of FuGENE yielded the highest transfection efficiency, achieving 8.82% of GFP positive cells, followed by the 2:3 ratio reaching 6.28% of transfection efficiency (Table 3.1).

Table 3.1 – Conditions and Efficiencies of HUCPVCs Transfection with FuGENE 6.

DNA:FuGENE 6 Transfection Efficiency 1:3 8.82% 2:3 6.28% 1:6 Not enough cell survival to perform flow cytometry n=1 (HUCPVCs from 5 different cords were pooled)

Figure 3.1– Fluorescence Microscopy of HUCPVCs Transfected with FuGENE 6 (Original magnification 9.37X).

3.1.2. Lipofectamine LTX

HUCPVCs were transfected under different settings with a GFP reporter plasmid using Lipofectamine LTX a liposomal-based transfection method at a confluence of 70-80% in 24-well plates (Table 3.2). After three days GFP expression in the cells was analysed by flow cytometry and results suggested that by maintaining the levels of DNA but increasing the concentration of Lipofectamine it was possible to double the transfection efficiency (Figure 3.2). The maximum transfection efficiency achieved was ~20% when using 500ng of DNA and 2µl of Lipofectamine LTX.

Table 3.2 – Conditions and Efficiencies of HUCPVCs Transfection with Lipofectamine LTX.

µl of Lipofectamine LTX ng of Plasmid DNA Transfection Efficiency 1.25 500 5.19% 1.25 750 10.77% 2 500 20.93% 2 750 18.23%

Figure 3.2 – HUCPVCs Transfection with Lipofectamine LTX. After transfecting HUCPVCs with Lipofectamine LTX flow cytometry was used to measure the percentage of GFP positive cells. Transfection efficiency is augmented by increasing Lipofectamine concentration. n=1 (HUCPVCs from 5 different cords were pooled) 76

A. B.

Figure 3.3 – Microscopy of HUCPVCs Transfected with Lipofectamine LTX . (A) Phase-contrast microscopy of HUCPVCs. (B) Fluorescence microscopy of the same field in A. (Original magnification 9.37X).

3.1.3. Nucleofection

HUCPVCs nucleofection with a GFP reporter plasmid was tested with different programs (Table 3.3), in most of the programs transfection efficiency determined by flow cytometry was >50%, but in some cases high rates of cell death were found using the Vi-CELL XR Cell Viability Analyzer. The program that best fitted HUCPVCs was X-001 achieving up to 79% transfection efficiency and >70% of cell survival when compared to cells that were not nucleofected as described in section 2.2.3. (Table 3.3).

Table 3.3 – Programs, Transfection Efficiencies and Cell Viability of HUCPVCs Nucleofection.

Program Efficiency of Transfection Cell Survival X-001 79% 70% A-020 50% 65.7% X-005 69% 47.3% T-020 62% 41.6% D-023 68% 41% L-029 67% 35% T-030 52% 23.3%

3.1.3.1. Longevity of GFP expression after HUCPVCs nucleofection

After transfecting HUCPVCs with the X-001 nucleofector program the longevity of GFP expression was measured by flow cytometry at days 3, 7, 12 and 15 post-transfection. HUCPVCs growing in media supplemented with 5% FBS lost ~16% of GFP expression within 7 days and a dramatic decrease was observed from day 7 to 12, losing ~ 70% of GFP expression (Figure 3.4).

Figure 3.4 – Longevity of GFP Expression after HUCPVCs Nucleofection. HUCPVCs post-transfection efficiency at days 3, 7, 12 and 15. HUCPVCs lost a significant amount of GFP expression during the first 12 days post-transfection. n=1 (HUCPVCs from 5 different cords were pooled)

Even though non viral methods were capable of genetically modifying HUCPVCs the transfection efficiencies achieved with FuGENE 6 and Lipofectamine LTX were very low, less than 25%, which was not enough for our future goal of creating 70-100% positive populations of HUCPVCs expressing osteogenic genes. Although, HUCPVC nucleofection achieved high efficiency levels (~79%) the stability of gene expression diminished dramatically over time. Therefore, it was decided to investigate the genetic modification of HUCPVCs by different viral methods.

3.2. HUCPVCs retroviral transduction

HUCPVCs were transduced in 6-well plates at a 70% confluence with a viral construct expressing murine Runx2- and Mock-retrovirus as described in section 2.2.4.2. Three days post-transduction cells were evaluated for the presence of GFP by fluorescence microscopy and flow cytometry, indicating transduction efficiencies of ~90% for the Mock virus and ~77% for the Runx2 virus (Figure 3.5). Immunofluorescence was used to detect the production of Runx2 at the protein level, Runx2 was detected in the nucleus of Runx2-HUCPVCs but not in Mock-HUCPVCs (Figure 3.6).

Figure 3.5 – HUCPVCs Retroviral Transduction. (A) Runx2-HUCPVCs and (B) Mock-HUCPVCS fluorescence microscopy together with phase contrast (Original magnification 9.37X) and transfection efficiency measured by flow cytometry.

A. B.

Figure 3.6 – Runx2 Immunofluorescence. Fluorescence microscopy of (A) Mock-HUCPVCs (B) Runx2- HUCPVCs (Green: GFP, Blue: Hoechst, Red: Runx2). Runx2 expression is localized in the nucleus of Runx2-HUCPVCs but not in Mock-HUCPVCs. (Original magnification 19.75X). After genetic modification of HUCPVCs with Runx2 or the Mock retrovirus, Runx2 expression was assessed by Immunofluorescence using 1:400 Runx2 Rabbit Polyclonal Antibody, 1:1000 Alexa Fluor 555 Goat Anti- Rabbit IgG, and 1:3000 Hoechst 33342.

3.2.1. Effects of Runx2 over-expression in HUCPVCs

As an early marker of osteoblastic differentiation ALP activity was detected by histochemical staining. In Runx2-HUCPVCs ALP activity was noticeable from day 7 with an evident increase at day 21. Despite being cultured in osteogenic medium Mock-HUCPVCs expressed very low ALP (Figure 3.7). ALP expression was also detected by RT-PCR in Runx2-HUCPVC and Mock- HUCPVCs from day 3. Although, both expressed ALP the bands in Runx2-HUCPVCs are more intense than in the mock indicating an upregulation of the gene (Figure 3.8). Moreover using species specific primers murine Runx2 gene expression was detected in Runx2-HUCPVCs at all time points but not in Mock-HUCPVCs (Figure 3.8), confirming the expression of Runx2 caused by the retroviral infection. While the gene expression of human Runx2 was detected in both Runx2- and Mock-HUCPVCs at all time points (Figure 3.8) only Runx2 at the protein level was detected in Runx2-HUCPVCs by immunofluorescence (Figure 3.6). In addition, Runx2- and Mock-HUCPVCs were examined for matrix mineralization by Von Kossa staining at day 21 but all the cultures failed to mineralize (not shown).

Figure 3.7 – Alkaline Phosphatase Staining after Retroviral Transduction of HUCPVCs. (A) Images of entire wells and (B) light microscopy of selected areas of wells (original magnification 12.8X). After genetic modification HUCPVCs were cultured in osteogenic medium for different time points. In Runx2- HUCPVCs ALP expression was detected from day 7 with an evident increase at day 21. The morphology of the positive cells (red) was found to be less fibroblastic and more enlarged when compared to the Mock-HUCPVCs. Mock-HUCPVCs expressed very low ALP despite of being under osteogenic conditions. 81

Figure 3.8 – Gene Expression Detected by RT-PCR after Retroviral Transduction with Runx2. Only Runx2-HUCPVCs (+) expressed murine Runx2 (Runx2-M). ALP is expressed by Runx2- and Mock- HUCPVCs (-) but at significantly higher levels in Runx2-HUCPVCs than in Mock-HUCPVCs. Relatively low levels of human Runx2 (Runx2-H) and Collagen 1 Alpha 1 were detected in both Runx2- and Mock- HUCPVCs at all time points.

Although retroviral infection causes long lasting expression of the gene of interest its random integration in the host genome may disrupt endogenous cell function by insertional mutagenesis (109). After the results achieved by retroviral-mediated transfer of murine-Runx2 into HUCPVCs it was decided to explore the genetic modification of HUCPVCs with recombinant adenoviruses carrying human genes based on that fact that adenoviruses do not normally integrate into the host genome resulting in the expression of the gene only for a limited period of time (104) which is probably more suitable for Bone engineering approaches.

3.3. Recombinant adenovirus fabrication

Five different recombinant replication defective adenoviruses were constructed using the AdEasy XL Adenoviral Vector System (267) as described in section 3.3.1.; four carrying the human cDNA of BMP2, Runx2, SP7 or VEGF-A121 and one empty as the mock (Table 3.4). First, genes of interest were cloned into the shuttle vector pShuttle-IRES-hrGFP-2 (Figure 2.2) which design allows the monitoring of the expression of the gene of interest at the single-cell level due to the simultaneous expression of hrGFP from the same transcript through an internal ribosome entry site (IRES). The gene of interest was also cloned in frame with a C-terminal 3X

HA tag to facilitate purification, detection (e.g. all the genes of interest can be detected at the protein level using an anti-HA antibody; mRNA produced by the modified cells as a result of the genetic modification can be specifically distinguished by detecting mRNA containing the HA tag sequence). After obtaining 5 different shuttle vectors pBMP2-Shuttle, pRunx2-Shuttle, pSP7- Shuttle, pVEGF-Shuttle and pMock-Shuttle, homologous recombination in vivo in bacteria was used to produce the recombinant adenovirus plasmids by a double recombination event between the pAdEasy-1 (adenoviral backbone plasmid vector) and the pShuttle vector carrying the gene of interest. The different resulting recombinant adenoviral constructs for each gene of interest were digested with PacI restriction enzyme and ran on a 0.8% agarose gel to screen for proper recombination (Figure 3.9). After verifying that the constructs matched the pattern expected five different recombinant adenoviral plasmids pBMP2-rAV, pRunx2-rAV, pSP7-rAV, pVEGF-rAV and pMock-rAV were linearized and used to transfect AD-293 adenovirus packaging cells to obtain five different recombinant adenoviruses: Ad-BMP2, Ad-Runx2, Ad- SP7, Ad-VEGF and Ad-Mock.

Table 3.4 – Recombinant Adenoviruses Constructed Using the AdEasy XL Adenoviral Vector System.

Recombinant Adenovirus cDNA Insert Size Ad-BMP2 1183 bp Ad-Runx2 1552 bp Ad-SP7 1284 bp Ad-VEGF-A 366 bp Ad-Mock No insert

The inserted cDNA corresponds to the BglII-gene of interest-XhoI obtained after PCR amplification (detailed sequences are reported in the Appendix 2). bp; Base Pairs

Figure 3.9 – Screening of Seven Different Clones of pBMP2-rAV After the Recombination Process. Supercoiled (uncut) plasmids (even lanes 2-14), pBMP2-shuttle uncut (lane 15) and PacI digested potential recombinant adenovirus plasmids (uneven lanes 1-13). All but two clones (5 and 7) matched the pattern expected after a proper recombination. Restriction of recombinant adenoviral plasmids with PacI should yield a large fragment observed to migrate next to the 23kb marker and a smaller fragment of either 3.0 kb or 4.5 kb. Uncut recombinants give a large smear at the top of the gel very close to the wells and a smaller band that runs just below 23 kb (267).

3.4. Recombinant Adenoviral Infection of HUCPVCs

HUCPVCs were genetically modified with five different recombinant adenoviruses at a MOI of 10 as described in section 3.3.2. Five different groups of genetically modified HUCPVCs were obtained: BMP2-HUCPVCs, Runx2-HUCPVCs, SP7-HUCPVCs, VEGF-HUCPVCs and Mock- HUCPVCs. In all the groups of modified cells the transfection efficiency measured by flow cytometry was >90% (Figure 3.11) and cell viability assessed by the Vi-Cell was >85%.

A. B.

Figure 3.10 – Genetically Modified HUCPVCs Expressing GFP after Recombinant Adenoviral Infection. (A) Phase contrast, (B) fluorescence microscopy of the same field.

Figure 3.11– Flow Cytometry of HUCPVCs after Recombinant Adenoviral Infection. Results for the five different groups of genetically modified HUCPVCs. In all of the groups transfections were >90%. The red signal in all the graphs represent naive cells (non-modified) use as a negative control.

3.4.1. Production of the transcripts of the genes of interest by genetically modified HUCPVCs

mRNA expression of the genes of interest was assessed by RT-PCR detecting only the transcripts being produced as a result of the genetic modification by the recognition of the genes tagged with the HA sequence (part of the recombinant adenoviral vector) as described

in section 3.3.3. Specific transcripts were only found in the cells modified with the specific genes of interest (Figure 3.12). For example, HA tagged BMP2 was only found in BMP2- HUCPVCs but not in the other genetically modified cells.

Figure 3.12 – RT-PCR of the Genes of Interest after Recombinant Adenoviral infection. Presence of mRNA transcripts for the genes of interest was confirmed by using specific primers to detect only transcripts that have the XhoI restriction site and the HA tag sequence (part of the recombinant adenoviral vector) (Figure 2-A). This strategy allows only the detection of transcripts being produced as a result of the genetic modification. HA tagged specific transcripts were only found in the cells modified with the specific genes of interest but not in the other genetically modified cells. bp (Base pairs)

3.4.2. Production of the Proteins of Interest by Genetically modified HUCPVCs

To detect the production of the genes of interest at a protein level western blots were use to detect the proteins from cell lysates by using anti-HA antibody and specific antibodies in the case of Runx2 and SP7. Antibodies against BMP2 and VEGF-A were used in ELISAs to detect the amount of protein secreted by the genetically modified cells. Since Runx2 and SP7 are transcription factors they are not secreted by the cell therefore ELISAs were not performed. BMP2 protein was detected at approximately 23kDa by anti-HA antibody, likewise, VEGFA-121 was detected at ~20kDa. SP7 was found at ~48kDa using the anti-HA and anti-SP7 antibody. Runx2 was detected by anti-HA and anti-Runx2 antibody at ~60 kDa (Figure 3.13).

Figure 3.13 – Western Blots of the Protein of Interest. Western blots with specific and/or anti-HA antibody of protein isolated from genetically modified HUCPVCs (cell lysates). The production of BMP2 and VEGF-A was shown with specific antibodies by ELISAs against BMP2 and VEGF-A (Figure 3.14).

The amount of the secreted proteins was quantified by ELISA. On average it was found that BMP2-HUCPVCs produced 87.7 ± 12.78 pg/ml/105cells/24hr, BMP2 secretion was not detected in Mock-HUCPVCs or naive HUCPVCs. In the same way it was found that VEGF-HUCPVCs produce 1469.4 ± 296.9 pg/ml/105cells/24hr and VEGF-A secretion was not detected by this method in Mock-HUCPVCs or naive HUCPVCs (Figure 3.14).

Figure 3.14 – ELISA Result of the Secreted Factors of Interest. (A) BMP2 production by BMP2-HUCPVCs (87.7 ± 12.78 pg/ml/105cells/24hr) and (B) VEGF-A production by VEGF-HUCPVCs (1469.4 ± 296.9 pg/ml/105cells/24hr) measured in conditioned medium. BMP2 and VEFG-A production was not detected in Mock-HUCPVCs conditioned medium. Error bars indicate the standard deviation of 3 independent biological repeats.

3.4.3. Longevity of GFP expression in HUCPVCs after gene transfection with recombinant adenoviruses.

The longevity of GFP expression over time after recombinant adenoviral gene transfection was tested under different growing conditions in Mock-HUCPVCs and measured by flow cytometry as described in section 3.3.5. During the first 3 days GFP expression was very similar between genetically modified HUCPVCs from all the different growing conditions (Figure 3.15). From day 7 onwards differences in the amount of GFP positive cells were noticeable between groups. At day 7 Mock-HUCPVCs growing in media supplemented with 5% FBS lost ~15% of GFP expression while expression in cells treated with Mitomycin C stayed stable. By day 21 cells growing in 5% FBS lost ~85% of GFP expression, cells in 2% FBS and 1% FBS lost ~55% and ~25% respectively whereas cells in the Mitomycin C group showed a stable number of GFP positive cells (Figure 3.15). These findings suggest that proliferation has an effect on gene expression stability since the group of cells that were growth arrested (Mitomycin C treated) kept a constant percentage of GFP positive cells while the cells proliferating lost GPF expression at different rates depending on the concentration of FBS in the media.

Figure 3.15 – Longevity of GFP Expression under Different Growing Conditions after Recombinant Adenoviral Infection of HUCPVCs. HUCPVCs from 4 different cords were mixed and transfected with Ad-Mock; GFP expression was quantified by flow cytometry at different time points and under different growing conditions: 1% FBS, 2% FBS, 5% FBS and Mitomycin C treated. GFP expression was found to be stable in cells treated with Mitomycin C while it decreases overtime in proliferating cells. Error bars indicate standard deviation of the triplicate flow cytometry measurements.

3.4.4. Bioactivity of the BMP2 produced by BMP2-HUCPVCs

Bioactivity of the BMP2 secreted by BMP2-HUCPVCs was tested by exposing C2C12 cells to the conditioned media as described in section 2.3.6. Three days later the alkaline phosphatase activity in C2C12 cells was measured indicating higher ALP expression in C2C12 cells exposed to conditioned media of BMP2-HUCPVCs when compared to C2C12 cells exposed to Mock- HUCPVC conditioned medium suggesting that the BMP2 secreted by the modified cells is causing an osteoinductive effect (Figure 3.16).

Figure 3.16 – Bioactivity Test of BMP2 Secreted by BMP2-HUCPVCs. C2C12 cells exposed to conditioned medium (CM) from BMP2-HUCPVCs expressed higher amounts of ALP when compared to C2C12 cells exposed to CM of Mock-HUCPVCs. As a positive control rhBMP2 was added to the medium of C2C12 cells triggering the production of high amounts of ALP. Error bars indicate the standard deviation of 5 independent biological repeats. T-test after logarithmic transformation.

3.4.5. Effects of cryopreservation on protein production and bioactivity

3.4.5.1. Bioactivity of BMP2 produced by pre- and post-liquid nitrogen BMP2-HUCPVCs.

Bioactivity of the BMP2 secreted by pre- and post-liquid nitrogen (LN) BMP2-HUCPVCs was tested by exposing C2C12 cells to the conditioned medium as described in section 2.3.7.2. Three days later alkaline phosphatase activity in C2C12 cells was measured finding consistent results between pre- and post-LN cells; higher ALP activity was seen in C2C12 cells exposed to pre- and post-LN BMP2-HUCPVCs conditioned medium when compared to C2C12 cells exposed to pre- and post-LN Mock-HUCPVCs conditioned medium (Figure 3.17).

Figure 3.17 – Bioactivity Test of BMP2 Produced by pre- vs. post-Liquid Nitrogen BMP2-HUCPVCs. Conditioned medium by pre- and post-liquid nitrogen (LN) BMP2-HUCPVCs has a similar osteoinductive effect in C2C12 cells by increasing the production of ALP when compared to C2C12 cells that were exposed to the conditioned medium of pre- and post-LN Mock-HUCPVCs. Error bars indicate the standard deviation of 5 independent biological repeats. T-test after logarithmic transformation.

3.4.5.2. Production of VEGF by pre- and post-liquid nitrogen VEGF- HUCPVCs

After thawing, recovery of engineered HUCPVCs was determined to be 85.37 ± 5.07% (n=6) as explained in section 3.1. VEGF secretion in pre- and post-LN VEGF-HUCPVCs was measured by ELISA at different time points as described in section 2.3.7.3. VEGF-A secretion in pre- and post-LN stored cells was found to behave similarly over time (Figure 3.18) and no statistical differences were found between the secreted VEGF by pre- and post-LN VEGF-HUCPVCs at the same time point. VEGF production by the cells was found to diminish approximately 60% from day 3 to 7 and continued to fall progressively until day 21 when the last measurement was performed.

Figure 3.18 – VEGF Production Over Time by pre- and post-Liquid Nitrogen VEGF-HUCPVCs. VEGF secretion measured by ELISA in pre- and post-liquid nitrogen (LN) VEGF-HUCPVCs behaves very similarly over time; no statistical differences were found between the secreted VEGF by pre- and post- LN VEGF-HUCPVCs at the same time point. VEGF production by the cells was found to diminish approximately 60% from day 3 to day 7 and continued to diminish progressively. Error bars indicate the standard deviation of 3 independent biological repeats. T-test after logarithmic transformation.

3.4.6. Effects of over-expression of BMP2, Runx2 and SP7 in HUCPVCs.

Different groups of genetically modified HUCPVCs were cultured in osteogenic media. The different groups were: BMP2-HUCPVCs; Runx2-HUCPVCs; SP7-HUCPVCs; BMP2-HUCPVCs + Runx2-HUCPVCs; BMP2-HUCPVCs + SP7-HUCPVCs; Runx2-HUCPVCs + SP7-HUCPVCs; BMP2- HUCPVCs + Runx2-HUCPVCs+ SP7-HUCPVCs; and Mock-HUCPVCs. Details described in section 2.4.1.

Production of ALP as an early osteogenic marker was studied in genetically modified HUCPVCs by histochemical staining at day 7 (Figure 3.19). BMP2-HUCPVCs were found to express low amounts of ALP while Runx2-HUCPVCs and SP7-HUCPVCs expressed the most ALP followed by the cocktail of Runx2/SP7. The cocktails BMP2/Runx2, BMP2/SP7 and BMP2/Runx2/SP7

expressed more ALP than BMP2-HUCPVCs whereas no positive staining was found in Mock- HUCPVCs.

Figure 3.19 – ALP Histochemical Staining of Genetically Modified HUCPVCs. After 7 days of culture BMP2-HUCPVCs (A) express low amounts of ALP. Runx2-HUCPVCs (B) and SP7-HUCPVCs (C) expressed the most ALP followed by the cocktails of Runx2/SP7 (F). No positive staining (red) was found in Mock- HUCPVCs (H). These images are representative of 4 independent biological repeats.

To quantified ALP activity at days 3, 7 and 14 a colorimetric assay was employed as described in section 2.4.2. At all time points ALP activity was found to be significantly higher (P<0.01) in Runx2- and SP7-HUCPVCs when compared to Mock-HUCPVCs. Moreover, ALP activity in SP7- and Runx2-HUCPVCs was higher when compared to BMP2-HUCPVCs (P<0.01) being at least 3- fold higher by day 7 (Figure 3.20). When comparing all the cocktails (mix of different groups of genetically modified HUCPVCs) with the Mock, ALP activity was found to be significantly higher at all time points (P<0.01) (Figure 3.21); at day 7 ALP activity in BMP2/Runx2 and BMP2/SP7 93

was around 2-fold higher, while in Runx2/SP7 and BMP2/Runx2/SP7 was more than 3-fold higher. Comparing the cocktails of modified HUCPVCs with the cultures over-expressing only one gene ALP activity in BMP2-HUCPVCs was significantly lower than the rest of the cocktails (P<0.05) followed by the cocktails BMP2/Runx2 and BMP2/SP7 (Figure 3.22). At all time points Runx2- , SP7-HUCPVCs and the Runx2/SP7 cocktail expressed the highest amounts of ALP activity among the samples followed by the BMP2/Runx2/SP7 cocktail.

Figure 3.20 – Measurement of Alkaline Phosphatase Activity in BMP2-, Runx2- and SP7-HUCPVCs. ALP activity is significantly higher in Runx2- and SP7-HUCPVCs when compared to Mock-HUCPVCs. Moreover, ALP activity in SP7- and Runx2-HUCPVCs is higher when compared to BMP2-HUCPVCs at all time points (P<0.01) being at least 3-fold higher by day 7. Error bars indicate the standard error of 6 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons.

Figure 3.21 – Quantification of Alkaline Phosphatase Activity in Cocktails of Genetically Modified HUCPVCs. All the different cocktails of genetically modified HUCPVCs were found to have a significantly higher (P<0.01) ALP activity than the Mock. At day 7 ALP activity in BMP2/Runx2 and BMP2/SP7 was around 2-fold higher, while in Runx2/SP7 and BMP2/Runx2/SP7 was more than 3-fold higher. Error bars indicate the standard error of 6 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons.

Figure 3.22 – Quantification of Alkaline Phosphatase Activity of Genetically Modified HUCPVCs. Comparing the cocktails of modified HUCPVCs with the cultures over-expressing only one gene ALP activity in BMP2-HUCPVCs was significantly lower than the rest of the cocktails (P<0.05) followed by the cocktails BMP2/Runx2 and BMP2/SP7. At all time points Runx2- , SP7-HUCPVCs and the Runx2/SP7 cocktail had the highest amounts of ALP activity among the samples followed by the BMP2/Runx2/SP7 cocktail. Error bars indicate the standard error of 6 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons. 95

To assess the behaviour of other osteogenic markers gene quantification by RT-qPCR was done utilizing the Pfaffl method (273), in where a mathematical model determines the fold change of mRNA expression of an unknown sample versus the mock and normalize it to the house keeping gene (section 2.4.4.).

Gene expression was analyzed at days 2, 6 and 9 finding that ALP was upregulated in all the samples at all time points when compared to the Mock-HUCPVCs and normalized to Beta-2 microglobulin (β2M) leves (Figure 3.23). ALP mRNA levels in BMP2-HUCPVCs were found to be lower when compared to the rest of the samples while levels in Runx2-HUCPVCs and Runx2/SP7 were the highest being at least 10-fold higher than the Mock at day 6. By day 9 ALP mRNA expression decreased in all the samples when compared to days 2 and 6; despite of this decrement, ALP levels in all the groups but BMP2-HCUPVCs were found to be at least 4-fold higher than in the mock.

Figure 3.23 – ALP mRNA Expression in Genetically Modified HUCPVCs. ALP mRNA is upregulated in all the samples when compared to the Mock-HUCPVCs and normalized to Beta-2 microglobulin (β2M) levels. BMP2-HUCPVCs expressed lower amounts of ALP when compared to the rest of the samples while levels in Runx2-HUCPVCs and Runx2/SP7 were the highest being at least 10-fold higher than the Mock at day 6. By day 9 a decrease of ALP levels can be noticed in all the groups. The dashed line depicts the mean value of Mock-HUCPVCs. Error bars indicate standard deviation of 3 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons. 96

Figure 3.24 – Collagen 1A1 mRNA Expression in Genetically Modified HUCPVCs. mRNA levels of COL1A1 are upregulated in all the samples when compared to the Mock-HUCPVCs but by day 9 levels are lower than at day 2 and 6. The dashed line depicts the mean value of Mock-HUCPVCs. Error bars indicate standard deviation of 3 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons.

In all the groups of genetically modified HUCPVCs and cocktails, COL1A1 (Figure 3.24) was found to be upregulated when compared to Mock-HUCPVCs and normalized to β2M levels but by day 9 levels started to diminish when compared to days 2 and 6. OCN mRNA levels (Figure 3.25) were found upregulated in all the samples when compared to the Mock-HUCPVCs, at day 2 OCN mRNA expression was at least 1.5-fold higher for all the samples. In general OCN expression was found to behave similarly among the samples at all time points. OPN mRNA levels (Figure 3.26) were also upregulated in all the samples when compared to the Mock- HUCPVCs. At day 2 OPN mRNA levels of all the samples were more than 5-fold higher than in Mock-HUCPVCs. At day 6 the levels of OPN were significantly higher in BMP2-HUCPVCs when compared to Runx2-, SP7-HUCPVCs and Runx2/SP7 cocktail (P>0.05). The levels of these last samples diminished to 3-fold higher while for BMP2-HUCPVCs OPN levels were 10-fold higher

than the mock-HUCPVCs. Also, it seems that cocktails that contained BMP2-HUCPVCs like in the case of BMP2/Runx2, BMP2/SP7 and BMP2/Runx2/SP7 kept the levels at least 5-fold higher than the Mock. By day 9 the levels of OPN decreased in all the groups, although BMP2- HUCPVCs were still expressing significantly (P<0.01) higher levels than the rest of the groups minus the BMP2/Runx2/SP7 cocktail.

Furthermore, the overexpression effect of BMP2, Runx2 and SP7 in matrix mineralization was examined by von Kossa staining of cultures at day 21, 28 and 35 but all the cultures failed to mineralize (not shown).

Figure 3.25 – Osteocalcin mRNA Expression in Genetically Modified HUCPVCs. mRNA levels of OCN are upregulated in all the samples when compared to the Mock-HUCPVCs. By day 9 a decrease in OCN levels can be noticed in all the groups. The dashed line depicts the mean value of Mock-HUCPVCs. Error bars indicate standard deviation of 3 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons.

Figure 3.26 – Osteopontin mRNA expression in genetically modified HUCPVCs. mRNA levels of OPN are upregulated in all the samples when compared to the Mock-HUCPVCs. At day 6 the levels of OPN are significantly higher in BMP2-HUCPVCs when compared to Runx2-, SP7-HUCPVCs and Runx2/SP7 cocktail (P>0.05). By day 9 the levels of OPN are decreasing in all the groups although, BMP2-HUCPVCs are still expressing higher levels than the rest of the samples. The dashed line depicts the mean value of Mock-HUCPVCs. Error bars indicate standard deviation of 3 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons.

3.4.7. Effects of the intramuscular delivery of genetically modified HUCPVCs.

Three days post infection of HUCPVCs four experimental groups (Mock-HUCPVCs, VEGF- HUCPVCs, BMP2/Runx2/SP7 and BMP2/Runx2/SP7/VEGF) were tested for osteoinductive and osteogenic capacity by delivering the cells through intramuscular injection into the hind limb quadriceps muscle of NOD/SCID mice. After 2, 4 and 6 weeks x-rays were taken to monitor the formation of calcified tissue. At week 6 it was not possible to detect the formation of evident mineralized tissue in the x-rays (Figure 3.27). In addition, after euthanasia micro-computed 99

tomography of the hind limbs was performed but no obvious calcified formation was found in the area of injection (not shown).

Figure 3.27– X-rays of the Region of Interest after Intramuscular Delivery of HUCPVCs. 6 weeks after intramuscular delivery of genetically modified HUCPVCs it was not possible to detect the formation of calcified tissue in the area of injection.

3.5. Furin Cleavage of Pro-BMP2 produce by BMP2-HUCPVCs

After finding that BMP2-HUCPVCs produced low amounts of mature BMP2, 87.7 ± 12.78 pg/ml/105cells/24hr (Figure 3.13) it was hypothesized that the pro-BMP2 produced by BMP2- HUCPVCs was not being efficiently processed. In order to address this hypothesis additional experiments were performed which showed the following.

100

3.5.1. Production of mature BMP2 by different cell types after modification with Ad-BMP2

HUCPVCs, hBMSCs and rBMSCs were genetically modified with Ad-BMP2. BMP2 quantities in the conditioned media were measured by ELISA. Low amounts, 60-90 pg/ml/105cells/24hrs of mature BMP2 were found in all the samples, indicating that independent of the cell type or species the yield of mature BMP2 is low when cells are genetically modified with our Ad-BMP2.

3.5.2. rhFurin cleavage of Pro-BMP2 produced by BMP2-HUCPVCs

To assess whether the low production of mature BMP2 could be caused by low efficiency cleavage of pro-BMP2, HUCPVCs were genetically modified with Ad-BMP2. BMP2-HUCPVC conditioned medium was then treated with different concentrations (1000, 100, 10 and 0 ng/ml) of recombinant human furin (rhfurin). After cleavage reactions it was found by ELISA (Figure 3.28-A) that rhfurin increases the yield of mature BMP2 secreted by BMP2-HUCPVCs in a dose dependent manner. At all the different rhfurin concentrations the yield of mature BMP2 was significantly (P<0.005) higher than in the untreated samples. At a concentration of 1000ng/ml a greater than 5-fold increase of mature BMP2 was observed. In addition, western blot results (Figure 3.28-B) showed that in the reactions in which high levels (1000 ng/ml) of rhfurin were used it was possible to detect a band at around 23kDa, which is the expected size of mature BMP2, confirming the ELISA findings. Also, a band at ~45kDa which is the expected size of Pro-BMP2 was detected in all the reactions indicating that pro-BMP2 is being produced by BMP2-HUCPVCs but not being efficiently cleaved and turned into mature BMP2.

101

A. B.

Figure 3.28 – Treatment of BMP2-HUCPVCs Conditioned Media with rhFurin (A) ELISA. rhfurin increases the yield of mature BMP2 secreted by BMP2-HUCPVCs in a dose dependent manner. At all the different rhfurin concentrations the yield of mature BMP2 was significantly (P<0.005) higher than in the untreated samples. Error bars indicate standard deviation of 5 independent biological repeats. T-test after logarithmic transformation. (B) Western Blots. A band of ~ 23 kDa (expected size of mature BMP2) was only detected after rhfurin treatment (1000ng/ml), the ~45kDa band find in the treated and untreated samples corresponds to pro-BMP2.

3.5.3. Bioactivity test of BMP2-HUCPVCs conditioned media after rhFurin treatment

Bioactivity of the BMP2 cleaved by rhfurin was tested by exposing C2C12 cells to the cleavage reaction solution as described in section 2.6.3.3. Three days later the alkaline phosphatase activity in C2C12 cells was measured indicating significantly (P<0.02) higher levels of ALP activity in C2C12 cells exposed to the cleavage reaction solution containing 1000ng/ml of rhfurin when compared to C2C12 cells exposed to the cleavage reaction solution containing 0ng/ml of rhfurin (Figure 3.29). These results suggest that the mature BMP2 contained in the cleaved reaction solution treated with 1000ng/ml of rhfurin is bioactive since it triggers the upregulation of ALP in C2C12 cells.

102

Figure 3.29 – Bioactivity Test of BMP2 Cleaved by rhFurin. C2C12 cells exposed to the cleavage reaction solution containing 1000ng/ml of rhfurin expressed significant (P<0.02) higher levels of ALP activity when compared to C2C12 cells exposed to the cleavage reaction solution containing 0ng/ml Error bars indicate standard deviation of 3 independent biological repeats. T-test after logarithmic transformation.

3.6. Osteogenic Cultures of Genetically Modified Human Bone Marrow Stromal Cells (HBMSCs) vs. Genetically Modified HUCPVCs

3.6.1. Recombinant adenoviral infection of HBMSCs and HUCPVCs

HBMSCs and HUCPVCs were genetically modified with Ad-Runx2, Ad-SP7, or Ad-Mock as described in section 3.7.2., and after 48hrs, transfection efficiency was assessed by florescence microscopy (Figure 3.30) and flow cytometry (Figure 3.31). Three different groups of genetically modified HBMSCs and HUCPVCs were obtained: Runx2, SP7 and Mock. In all the groups of modified cells the transfection efficiency was >85% and cell viability assessed by the Vi-cell was >85%.

103

A. B.

C. D.

Figure 3.30 – Genetically Modified HBMSCs and HUCPVCs Expressing GFP after Recombinant Adenoviral Infection. HBMSCs (A) brightfield microscopy and (B) fluorescence microscopy of the same field. HUCPVCs (C) brightfield microscopy and (D) fluorescence microscopy of the same field.

3.6.1. Effects of over-expression of Runx2 and SP7 in HBMSCs vs. HUCPVCs.

To test and compare the osteogenic capacity of genetically modified HBMSCs and HUCPVCs cultures were kept in parallel in osteogenic media. Four different groups of modified cells were cultured for the two different types of cells: Runx2, SP7, Runx2/SP7 (50% of each) and Mock.

Production of ALP as an early osteogenic marker was studied by histochemical staining at day 7 and 14 revealing that HBMSCs expressed more ALP than HUCPVCs indicated by the abundant and intense pink staining found in the different samples of HBMSCs (Figure 3.32). In HBMSCs and HUCPVCs Runx2, SP7 and the cocktail Runx2/SP7 expressed higher amounts of ALP when compared to the Mock. Despite of being cultured in osteogenic media Mock-HUCPVCs expressed very low ALP while in Mock-HBMSCs the ALP positive stained area was comparable to the ALP positive area in Runx2-HUCPVCs, SP7-HUCPVCs and Runx2/SP7 HUCPVCs cocktail. 104

Figure 3.31 – Flow Cytometry of HBMSCs and HUCPVCs after Recombinant Adenoviral Infection. HBMSCs (A) and HUCPVCs (B). Transfection efficiency was >85% in all the samples. The red signal in all the graphs represent the naive cells (non-modified) use as a control.

To measure the ALP activity levels at days 7 and 14 a colorimetric assay was employed as described in section 2.4.2. At all time points ALP activity was significantly higher in Runx2- HBMSCs, SP7-HBMSCs and the cocktail Runx2/SP7 when compared to Mock-HBMSCs (P<0.05) and Mock-HBMSCs cultured in normal grow media (without osteogenic supplements) (P<0.05) (Figure 3.33-A). At day 7 and 14 ALP activity was at least 4-fold and 2.5-fold higher respectively in Runx2-HBMSCs, SP7-HBMSCs and Runx2/SP7 when compared to Mock-HBMSCs. Although ALP activity levels in genetically modified HUCPVCs were found to be significantly lower than in HBMSCs (Figure 3.33) genetic modifications seem to have a similar effect but at a different scale; Runx2-HUCPVCs, SP7-HUCPVCs and the cocktail Runx2/SP7 had higher ALP activity when compared to Mock-HUCPVCs (P<0.05) (Figure 3.33-B) and Mock-HUCPVCs cultured in normal growth media (P<0.05). In addition, a decreased in ALP activity can be noticed between day 7

105

and 14 in genetically modified HUCPVCs while in HBMSCs ALP activity seems to be more stable (Figure 3.33).

Figure 3.32 – Alkaline Phosphatase Staining in Modified HBMSCs and HUCPVCs. After 7 and 14 days of culture under osteogenic conditions HBMSCs expressed the most ALP when compared to HUCPVCs. In HBMSCs and HUCPVCs Runx2, SP7 and the cocktail Runx2/SP7 expressed higher amounts of ALP when compared to the Mock. Despite of being cultured in osteogenic media Mock-HUCPVCs expressed very low ALP while in Mock-HBMSCs the ALP positive stained area was comparable to the ALP positive area in Runx2-HUCPVCs, SP7-HUCPVCs and Runx2/SP7 HUCPVCs cocktail.

106

A. B.

Figure 3.33 – Alkaline Phosphatase Activity Measurement in Genetically Modified HBMSCs and HUCPVCs. (A) ALP activity is significantly higher in Runx2-HBMSCs, SP7-HBMSCs and the cocktail Runx2/SP7 when compared to Mock-HBMSCs (P<0.05) and Mock-HBMSCs cultured in normal grow media (without osteogenic supplements) (P<0.05). (B) ALP activity in genetically modified HUCPVCs was significantly lower than in HBMSCs but samples were found to followed the same trend than HBMSCs; Runx2-HUCPVCs, SP7-HUCPVCs and the cocktail Runx2/SP7 had higher ALP activity when compared to Mock-HUCPVCs (P<0.05) and Mock-HUCPVCs cultured in normal growth media (P<0.05). In addition, a decreased in ALP activity can be noticed between day 7 and 14 in genetically modified HUCPVCs (B) while in HBMSCs (A) ALP activity seems to be more stable. Error bars indicate standard deviation of 3 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons.

When ALP activity is directly compared between HBMSCs and HUCPVCs the difference in the levels is very evident (Figure 3.34). In HBMSCs ALP activity levels are at least 7-fold higher than in all the samples of HUCPVCs all time points. ALP activity levels in Runx2-HUCPVCs, SP7- HUCPVCs and Runx2/SP7 cocktail compared with the levels found in Mock-HBMSCs at day 7 and by day 14 ALP activity diminish being similar to the ALP activity found in Mock-HBMSCs cultured in non-osteogenic conditions.

107

A. B.

Figure 3.34 – Comparison of Alkaline Phosphatase Activity between HBMSCs and HUCPVCs. (A) day 7 and (B) day 14. ALP activity levels are at least 7-fold higher in HBMSCs when compared to HUCPVCs. ALP activity levels of Runx2-HUCPVCs, SP7-HUCPVCs and Runx2/SP7 are comparable with the levels found in Mock-HBMSCs at day 7 and by day 14 ALP levels are comparable with the ones found in Mock- HBMSCs cultured in non-osteogenic conditions. Error bars indicate standard deviation of 3 independent biological repeats.

To assessed mineralization capacity of the engineered cells cultures were fixed and stained with alizarin red at days 21 and 28 (Figure 3.35). Runx2-HBMSCs, SP7-HBMSCs and the cocktail Runx2/SP7 were found highly positive when compared to Mock-HBMSCs at days 21 and 28 (Figure 3.35-A). Evident matrix mineralization was only noticed in Mock-HBMSCs by day 28. Under the growing conditions implemented in this study (10% FBS) HUCPVCs proliferated faster than HBMSCs and by day 16 started to roll off the culture dishes which caused the loss of most of the samples. Only some wells containing Runx2-HUCPVCs, SP7-HUCPVCs and Runx2/SP7 cocktail were stained at the same time with the same alizarin red solution and only a light purple staining was detected revealing a big difference in calcium deposition capability between the HBMSCs and HUCPVCs (Figure 3.35-B). No mock-HUCPVCs cultures made it to day 21, therefore, there is not alizarin red staining for the HUCPVCs negative control.

108

Day 21

Day 28

Day 21

Figure 3.35 – Alizarin Red Staining of Engineered HBMSCs and HUCPVCs. (A) Runx2-HBMSCs, SP7- HBMSCs and the cocktail Runx2/SP7 are highly positive (red) when compared to the Mock-HBMSCs at day 21 and 28. Evident matrix mineralization is only noticed in the Mock-HBMSCs by day 28. (B) Only a light purple staining was detected in Runx2-HUCPVCs, SP7-HUCPVCs and the cocktail Runx2/SP7 revealing a big difference in calcium deposition capability between the HBMSCs and HUCPVCs. Staining was not performed in Mock-HUCPVCs cultures at day 21 and in all the HUCPVCs cultures at day 28 due to fast proliferation of the cells causing the loss of the samples.

Calcium content measured by a colorimetric assay (described in section 2.7.6) at day 21 in Runx2-HBMSCs, SP7-HBMSCs and Runx2/SP7 cultures was in average 246.33±100.1,

109

398.11±54.04 and 350.03±16.41 µg of Ca/mg of protein respectively, values that are significantly higher (P<0.01) than the calcium quantified in the Mock-HBMSCs cultures, 26.31±5.59 µg of Ca/mg of protein (Figure 3.36). Calcium content could not be measured in genetically modified HUCPVCs due to fast proliferation causing the cultures to roll off the dishes before the time point.

Figure 3.36 – Calcium Content Quantification in Modified HBMSCs Osteogenic Cultures at Day 21. Calcium content in Runx2-HBMSCs, SP7-HBMSCs and the cocktail of Runx2/SP7 HBMSCs was significantly higher (P<0.01) than in Mock-HBMSCs. Calcium content could not be measured in genetically modified HUCPVCs due to fast proliferation causing the cultures to roll off the dishes. Error bars indicate standard deviation of 3 independent biological repeats. ANOVA and Tukey's test for pairwise comparisons.

3.6.2. Osteogenic cultures in naive and engineered HBMSCs

Evident mineralization was not detected by day 21 in Mock-HBMSCs (Figure 3.35-A) that were cultured in osteogenic media supplemented with 3.5mM -glycerophosphate therefore it was tested if the genetic modification was having a negative effect in the osteogenic capacity of the cells by comparing osteogenic cultures of Mock-HBMSCs and Naive-HBMSCs (non-engineered). Moreover, cells cultured with osteogenic media supplemented with 3.5mM or 5mM - glycerophosphate were compared to cells cultured in normal growing media.

110

After Alizarin Red staining at day 21 it was found that cultures of Mock- and Naive-HBMSCs behave very similar since evident matrix mineralization was found in both groups cultured in osteogenic media supplemented with 5mM -glycerophosphate but not in the ones supplemented with 3.5mM -glycerophosphate (Figure 3.37-A). At day 28 some small mineralized areas were detected in cultures supplemented with 3.5mM -glycerophosphate while cultures supplemented with 5mM -glycerophosphate showed intense staining in most of the plate area (Figure 3.37-B). No evident difference in mineralization was observed between engineered cells and naive cells indicating that genetic modification does not affect mineralization capacity.

A. B.

Figure 3.37 – Alizarin Red Staining of Engineered and Naive HBMSCs. (A) At day 21 it was found that cultures of Mock- and Naive-HBMSCs behave very similar. Evident matrix mineralization (red) was found in cells cultured in osteogenic media supplemented with 5mM -glycerophosphate but not in the ones supplemented with 3.5mM -glycerophosphate. (B) At day 28 some small mineralized areas were detected in cultures supplemented with 3.5mM -glycerophosphate while cultures supplemented with 5mM -glycerophosphate showed intense staining in most of the plate area. No evident difference was observed in mineralization between engineered cells and naive cells indicating that genetic modification does not affect mineralization capacity.

111

4. Discussion

Bone marrow stromal cells (BMSCs) are currently the "gold standard" cell source for mesenchymal cell-based therapies and last year for the first time allogeneic human BMSCs received authorization from Health Canada to be marketed as a stem cell therapy for acute graft-versus host disease in children (262). This could represent the beginning of the commercial approval of other allogeneic MSC-based products for the treatment of different conditions. During the recent years human umbilical cord perivascular cells (HUCPVCs) have been reported to have many similarities with human BMSCs in terms of differentiation capacity, surface markers (74–76) and non-immunogeneic and immunomodulatory phenotype (78–80), while exhibiting a higher proliferation rate (74), a greater CFU-F frequency (56) and the advantage of being easier and safer to obtain since they are harvested from a tissue that is normally discarded after birth. Based on this, we hypothesised that HUCPVCs could potentially be a putative candidate cell source for osteogenic/osteoinductive ex vivo gene therapy, and this thesis studied this potential.

4.1. HUCPVCs are genetically modifiable by non- viral and viral methods.

To explore the potential use of HUCPVCs as an allogeneic cell source for osteogenic/osteoinductive ex vivo gene therapy strategies, part of this thesis work aimed to assess the feasibility of genetically modifying HUCPVCs with different techniques. Initially conventional non-viral methods such as FuGene, Lipofectamine and Nucleofection were utilized to transfect HUCPVCs with the plasmid pmaxGFP. Transfection of HUCPVCs with the chemical methods, FuGene and Lipofectamine, resulted in ~8% and ~20% transfection efficiency respectively. These efficiencies are found within the ranges reported by others when transfecting BMSCs with FuGene (3.6% - 22%) (102,274,275) and Lipofectamine (11% - 48%) (261,275–277) and are comparable to the efficiencies previously reported by Baksh et al. (278) after transfecting HUCPVCs with FuGene. Having obtained low transfection efficiencies in

112

HUCPVCs with the previous chemical methods and based in the success by Aluigi et al. (102) and Aslan et al. (158) at nucleofecting human BMSCs at higher efficiencies, ~73% and ~68% respectively, Nucleofection was performed in HUCPVCs achieving an efficiency of ~79% with a cell survival of 70%, data that is comparable with the results reported by Baksh et al. (278). However, despite the high nucleofection efficiency, HUCPVCs lost ~70% of GFP expression after 12 days of cell culture. This behaviour reflected that reported by Baksh et al. (278) who nucleofected HUCPVCs with a DsRed plasmid construct and found that HUCPVCs lost >80% of DsRed expression from day 4 to day 21. In the same study they compared the gene expression behaviour of nucleofected HUCPVCs and human BMSCs, reporting that BMSCs lost only 45% expression of DsRed from day 4 to day 21 compared to >80% in HUCPVCs, a difference that could be explained by the higher proliferation rate of HUCPVCs causing a faster dilution of the transgene expression. Additionally, Kim et al. (279) reported high HUCPVC nucleofection efficiency when using the plasmid pmaxGFP (~3.5 Kilobases) while very low efficiency (~6%) when nucleofecting HUCPVCs with a plasmid of 18.5 Kilobases demonstrating that this transfection method could be affected by the size of the plasmid, limiting its use when the plasmid containing the therapeutic gene is of a larger size.

To explore the genetic modification of HUCPVCs by viral methods retroviruses carrying the GFP gene or the murine form of Runx2 were tested. Three days after infection the efficiencies of transduction with both viruses were > 75%, similar to those reported by Byers et al. (232) when modifying rat BMSCs with the same retroviruses (>50%) and comparable to those reported by Gysin et al. (280) (>60%). Although this gene delivery method proved to be efficient to genetically modify HUCPVCs it required additional cell manipulation due to the two rounds of infection needed to compensate for the incapacity of the retrovirus to infect cells that were not dividing (94,98). In addition, the risk of disrupting endogenous cell function by retroviral-mediated insertional mutagenesis of a proto-oncogene or tumor suppressor (109), and the belief that for osteogenic/osteoinductive strategies the expression of the gene of interest should be transient to avoid undesired bone formation, it was decided to assess different gene delivery methods.

113

Adenoviruses are the most popular gene delivery method for ex vivo gene therapy aimed at bone regeneration (91,99) due to their high level of transfection efficiency, their capacity to infect a broad range of dividing and non-dividing cell types, their feasibility to be produced at a high titer and as they do not integrate into the host genome they do not carry the risk of disrupting endogenous gene function (94,98). To genetically modify HUCPVCs, recombinant adenoviruses carrying the GFP reporter gene were constructed using the Adeasy XL Adenoviral Vector System (Section 2.3.1). After exposing HUCPVCs to the recombinant adenoviruses flow cytometry was performed and transfection efficiency was determined to be >85% with cell viability >80%. It is known that adenoviruses remain in the nucleus as an episome which is not replicated during cell division resulting in the transient expression of the gene of interest (104). Therefore, longevity of gene expression after HUCPVC adenoviral infection was assessed at different time points and under different concentrations of FBS, and compared to growth arrested HUCPVCs (by Mitomycin C treatment). During the first 3 days GFP expression was found to be very similar between genetically modified HUCPVCs from all the different growing conditions (Figure 3.15). At day 7 modified HUCPVCs growing in medium supplemented with 5% FBS lost ~15% of GFP expression while expression in cells treated with Mitomycin C stayed stable. By day 21, cells growing in 5% FBS lost ~85% of GFP expression, cells in 2% FBS and 1% FBS lost ~55% and ~25% respectively whereas cells in the Mitomycin C group showed a stable number of GFP positive cells. Based on the data acquired, it was decided to carry out subsequent in vitro experiments with media supplemented with 2% FBS to maximize longevity of gene expression while guaranting proper cell growth and differentiation as previously demonstrated by Sarugaser et al. (75) when differentiating naive HUCPVCs into the osteogenic lineage and culturing them long-term (more than 10 passages) utilizing an FBS concentration of 2%.

During this thesis work different in vitro tests were performed to compare the performance of HUCPVCs and HBMSCs after genetic modification via adenoviral infection. In order to find the ideal infection conditions that would yield similar efficiency of transfection and cell viability in the two cell types, different virus concentrations were tested until finding the optimal multiplicity of infection (MOI) for each type of cell. Interestingly, it was observed that in order

114

to genetically modify more than 85% of the HUCPVC population an MOI of 10 was required while HBMSCs required an MOI of 50, indicating that HUCPVCs need less adenovirus concentration than HBMSCs to be infected with the same efficiency. It is well known that adenoviruses enter the cell by receptor mediated endocytosis by binding to the coxsackie/adenovirus receptor (CAR) (98,103) and therefore the efficiency of infection depends on the level of expression of the CAR in the target cell (281). Indeed, after performing microarray analysis on HUCPVCs and HBMSCs Tissue Regeneration Therapeutics - suppliers of HUCPVCs - found that CXADR, the gene that encodes the CAR was expressed at a higher level in HUCPVCs by 5.41-fold (p<0.05), which makes HUCPVCs more suitable for applications that require genetic modification via adenoviral infections.

4.2. Cryopreservation does not have an effect on production efficiency and bioactivity of proteins of interest produced by genetically modified HUCPVCs.

It was important to evaluate the effects of cryopreservation of modified HUCPVCs since one of the proposals of this thesis work was to genetically modify HUCPVCs with different genes fundamental for bone formation and regeneration and store them in liquid nitrogen so a bank of cryopreserved engineered human MSCs could be established ready to be used in osteogenic/osteinductive ex vivo gene therapy.

It is well documented that cryopreservation of human MSCs has no effects in their proliferation capacity, osteogenic differentiation (71,282,283), and cell surface antigen expression (CD34-, CD45-, CD14-, HLA-DR-, CD29+, CD44+, CD73+, CD90+, CD 166+ and CD105+) (284,285). The effects of the cryopreservation process in engineered MSCs have not been extensively assessed but in a recent study McGinley et al. (286) genetically modified rat BMSCs with a lentivirus carrying the GFP gene and examined the effects of cryopreservation detecting no negative impact on transgene expression levels after thawing. Also, Mumaw et al. (287) modified pig and sheep BMSCs with Ad-BMP2, microencapsulated the cells in Poly(ethylene glycol) (PEG), and cryopreserved them in liquid nitrogen in a solution containing 10% DMSO.

115

After comparing pre- and post-cryopreserved microencapsulated BMP2-MSCs it was concluded that the cryopreservation process did not have an effect on cell viability and biological activity of the microencapsulated BMP2-MSCs since both groups were capable of generating similar quantities of heterotopic bone in vivo.

To determine if the cryopreservation of genetically modified HUCPVCs had an effect on the production efficiency and bioactivity of the proteins of interest expressed by the cells different assays were performed comparing HUCPVCs before and after storage in liquid nitrogen. Although, for these experiments HUCPVCs were only stored for less than a week in liquid nitrogen, it is known that the critical parts of the cryopreservation process are the freezing and thawing steps, since is during these phases when cellular damage is at higher risk (288,289). Following thawing of engineered HUCPVCs recovery was determined to be 85.37 ± 5.07%. VEGF secretion in pre- and post-LN VEGF-HUCPVCs was measured by ELISA at days 3, 7, 11, 14, 17 and 21 finding that the cryopreservation process did not have an effect in VEGF expression as VEGF secretion in pre- and post-LN stored HUCPVCs was found to behave similarly over time (Figure 3.18). VEGF production by both groups diminished by approximately 60% from day 3 to 7 and continued to fall progressively over time. As discussed in the previous section (Section 5.1) this rapid decrease in VEGF expression, which was normalized to cell number, could be the result of HUCPVCs fast proliferation and the transient gene expression due to the gene delivery vector used to modified the cells.

When evaluating if the cryopreservation process had an effect on the biological activity of genetically modified HUCPVCs, BMP2 bioactivity was assessed by exposing C2C12 cells to the medium conditioned by pre- and post-LN BMP2-HUCPVCs or pre- and post-LN Mock-HUCPVCs. C2C12 cells exposed to the pre- and post-LN BMP2-HUCPVCs conditioned medium expressed higher levels of ALP activity when compared to C2C12 cells exposed to pre- and post-LN Mock- HUCPVCs conditioned medium (Figure 3.17) indicating the presence of bioactive BMP2 in the conditioned medium of pre- and post-LN BMP2-HUCPVCs. Furthermore, ALP activity in C2C12 cells was found to behave very similar after treatment with media conditioned by pre- and

116

post-LN cells indicating that cryopreservation of the cells did not affect the biological activity of the protein of interest produced by the engineered HUCPVCs.

4.3. HA tagged BMP-2 produced by BMP2-HUCPVCs is not properly processed into mature-BMP2.

BMP-2 Recombinant adenovirus (Ad-BMP2) was constructed using the AdEasy XL Adenoviral Vector System (267) by cloning the full length of the human BMP-2 cDNA (sequence can be found in the Appendix 2) into the pShuttle-IRES-hrGFP-2 vector (Figure 3.2). This shuttle vector was chosen to facilitate the monitoring of the expression of the gene of interest at the single- cell level due to the simultaneous expression of hrGFP from the same transcript through and internal ribosome entry site (IRES). BMP-2 was also cloned in frame with a C-terminal 3X HA tag to facilitate purification and detection. As a result of the tagging BMP-2 can be detected at the protein level using an anti-HA antibody and BMP-2 mRNA produced as a result of the genetic modification can be specifically distinguished by detecting the mRNA containing the HA tag sequence.

After HUCPVC infection with the Ad-BMP2, GFP expression was detected by flow cytometry in >90% of the BMP2-HUCPVCs. In addition, the presence of HA tagged BMP-2 transcripts was detected by RT-PCR (Figure 3.12) and BMP-2 at the protein level was detected at ~23kDa in western blots (Figure 3.13) but when measuring the concentration of BMP-2 in the conditioned medium by ELISA low production of BMP-2 was found (87.7 ± 12.78 pg/ml/105cells/24hr). To test if the inefficient production of BMP-2 was being caused by HUCPVCs, human BMSCs and rat BMSCs were genetically modified with Ad-BMP2. These two different cell types were chosen due to their common use in ex vivo gene therapy research and their reported capability of producing high levels of BMP-2 after genetic modification with vectors carrying the BMP-2 gene (not fused to a HA tag) (159,290,291). After achieving >85% transfection efficiency BMP-2 quantities in the conditioned media were measured and low amounts (60-90 pg/ml/105cells/24hrs) of BMP-2 were also found in modified human BMSCs and rat BMSCs,

117

indicating that independent of the cell type or species the yield of BMP-2 is low when cells are genetically modified with the Ad-BMP2 constructed during this thesis work.

BMP-2 is synthesized as an inactive pre-pro-protein which consists of an amino terminal signal peptide, a pro-domain and a carboxyl terminal mature protein (140,141) (Figure 1.2 and Figure 5.1). In order for BMP-2 to become a mature protein pro-BMP-2 has to be cleaved on the C- terminal side of the proprotein convertase recognition sequence (R-X-X-R) by proteases that belong to the proprotein convertase family such as the serine endoprotease furin (134,144) which mainly acts on proteins in the trans-Golgi network (TGN) (145,146,292). When western blots from BMP2-HUCPVCs conditioned medium and cells lysate were performed with anti-HA antibodies two bands were detected in the conditioned medium, one at ~45kDa which corresponds to the predicted size of 3X HA tagged Pro-BMP2 and one at ~23kDa that corresponds to the expected size of 3X HA tagged mature BMP-2. In the cell lysate three bands were detected, two of the same size found in the conditioned medium plus a ~55kDa band that could correspond to pre-pro-BMP2 (Figure 4.1).

Figure 4.1– Western Blots from BMP2-HUCPVCs Conditioned Medium and Cells Lysate. Anti-HA antibodies were used to detect the presence of HA tagged BMP-2. In the conditioned medium two bands were detected, one at ~45kDa that represents Pro-BMP2 and one at ~23kDa that corresponds to the expected size of 3x HA tagged mature BMP-2. In the cells lysate three bands were detected, two of the same sizes found in the conditioned medium plus a ~55kDa band that could correspond to pre-pro- BMP2.

118

To confirm if the ~45kDa band found in the western blots corresponds to pro-BMP2 and to assess if low production of mature BMP-2 is being caused by inefficient intracellular cleavage of pro-BMP-2, BMP2-HUCPVCs conditioned medium was treated with different concentrations of rhfurin. After cleavage reactions it was found by ELISA (Figure 3.28-A) that rhfurin increased the measurements of mature BMP2 contained in the BMP2-HUCPVCs conditioned medium in a dose dependent manner. In all the conditioned media treated with rhfurin the concentration of mature BMP-2 was significantly (P<0.005) higher than in the untreated samples. When furin was used at a concentration of 1000ng/ml an increase greater than 5-fold was observed in levels of mature BMP-2. In addition, western blots (Figure 3.28-B) showed that in the reactions in which high levels of rhfurin (1000 ng/ml) were used it was possible to detect a band that was not detected in untreated samples around 23kDa, which is the expected size of mature 3x HA tagged BMP-2. These results confirmed that BMP2-HUCPVCs are producing pro-BMP2, but is not being efficiently cleaved intracellularly.

Bioactivity of the BMP-2 cleaved by rhfurin was also tested by exposing C2C12 cells to the cleavage reaction solution. When C2C12 cells are exposed to BMP-2 their ALP activity increases proportionately to the amount of BMP-2 present in the medium (272). When alkaline phosphatase activity in C2C12 cells was measured, significantly higher levels (P<0.02) were found in C2C12 cells exposed to the cleavage reaction solution containing 1000ng/ml of rhfurin when compared to C2C12 cells exposed to the cleavage reaction solution containing 0ng/ml of rhfurin (Figure 3.29). These results suggest that there is more mature bioactive HA tagged BMP-2 contained in the cleaved reaction solution treated with 1000ng/ml of rhfurin which triggered the upregulation of ALP activity in C2C12 cells.

Knowing that HUCPVCs were producing pro-BMP2 as detected in western blots (Figure 4.1), and confirmed by the furin cleavage experiment (Figure 3.28), and having shown that cells that normally are capable of efficiently produce mature BMP-2 were not able after being modified with our Ad-BMP2 we believe that possibly what is causing low efficiency in the cleavage of pro-BMP2 is the design of the adenoviral vector itself. It was hypothesized that the C-terminal 3X HA might be impeding the efficient cleavage of pro-BMP2 by covering the proprotein

119

convertase recognition sequence when passing through the TGN where BMP-2 is processed into a mature protein by furin. Previously it has been reported that C-terminal or N-terminal tags might influence the cleavage, expression, solubility, and bioactivity of proteins. Although, information about tags altering the cleavage and activity of BMP-2 was not found, there are existing reports about the effects of tags in other proteins. For example Byun et al. (293) reported that tagging of Rem protein on the N-terminal allowed protein accumulation, cleavage and activity, whereas tagging on the C-terminus had the opposite effect. Pulkki et al (294) demonstrated that the presence of a C-terminal FLAG tag in BMP-15 inhibited its bioactivity. Moreover, Mason et al (295) reported that a C-terminal His tag had an effect on the iron release from recombinant human serum transferrins (hTF), while the presence of a His tag at the N-terminal of hTF had no effect on the rate of iron release. Also, Christensen et al. (296) showed that when OPN is tagged at the C-terminal it reduces its capability to promote cell adhesion via the αVβ3-integrin, whereas modification of the N-terminal does not influence the binding.

Although, some of the data acquired regarding the issue of low production of BMP-2 by BMP2- HUCPVCs could indicate that the 3X HA tag may be affecting the cleavage it is not conclusive to prove or deny our hypothesis since the only way is to redesign the adenoviral vector and go back to the cloning stage and removed the 3X HA tag. However, since this was not the main aim of this thesis work and the problem was detected during the last stage no further work was performed regarding this issue. The important lesson to learn is that BMP-2 is a complex protein that has to go through different post-translational modification in order to become a bioactive protein and therefore a powerful tool for different gene therapy applications. In the end what was shown is that when HUCPVCs are genetically modified with our Ad-BMP2 they produce low amounts of mature BMP-2 and produce pro-BMP2 that is not properly cleaved intracellularly, but when the HA tagged pro-BMP2 contained in the conditioned media is treated with rhfurin it is cleaved and the resulting mature BMP-2 is bioactive.

120

4.4. Overexpression of Runx2 and SP7 causes upregulation of different osteogenic markers but does not have an effect on matrix mineralization in HUCPVCs in vitro or in vivo.

One of the novel proposals of this thesis work was to create a cocktail of genetically modified HUCPVCs expressing different genes fundamental for bone formation. The motivation behind overexpressing more than one gene was based on different studies in which engineered MSCs expressed the combination of two genes VEGF/BMP2 (190), VEGF/BMP4 (25), BMP2/Runx2 (238) or Runx2/SP7 (265) yielding improved results in terms of bone formation when compared to single gene overexpression. Indeed, to our knowledge, only two genes have been previously combined and, therefore, we are the first to propose combining 4 genes: BMP-2, Runx2, SP7 and VEGF, to maximize such an effect. Since each gene has a different function in bone formation, it was hypothesized that they might complement each other and act synergically to enhance bone formation.

To initially test the in vitro behaviour of different groups of modified HUCPVCs (BMP2- HUCPVCs, Runx2-HUCPVCs and SP7-HUCPVCs) and different cocktails (BMP2-HUCPVCs + Runx2-HUCPVCs, BMP2-HUCPVCs + SP7-HUCPVCs, Runx2-HUCPVCs + SP7-HUCPVCs, BMP2- HUCPVCs + Runx2-HUCPVCs+ SP7-HUCPVCs) cultures were examined at different time points by different techniques (VEGF-HUCPVCs were not used in vitro since originally they were only intended to induce angiogenesis in vivo). Alkaline phosphatase activity, commonly accepted as a surrogate osteogenic marker, was quantified at days 3, 7 and 14 and exhibited significantly higher (P<0.01) levels in Runx2- and SP7-HUCPVCs, when compared to Mock-HUCPVCs, which complements the ALP mRNA expression and the histochemical findings; and corroborates other reports of overexpressing Runx2 (81,229,230) or SP7 (247,260) in MSCs sourced from different tissues/species. No significant differences were found between Runx2- and SP7- HUCPVCs ALP activity or mRNA expression at any time point indicating that overexpression of Runx2 and SP7 seems to have a similar effect on ALP expression in HUCPVCs: a result that differs from the report of Lee et al. (265) who found higher levels of ALP at day 7 but not at day 14 in Runx2-ADSCs when compared to SP7-ADSCs.

121

When comparing all the cocktails with the Mock, ALP activity was significantly higher at all time points (P<0.01) (Figure 3.21): a result that correlated with the ALP mRNA expression and staining. The cocktails with lowest ALP were BMP2/Runx2 and BMP2/SP7, which were affected by the low ALP expression in BMP2-HUCPVCs but had higher ALP expression than the latter. On the other hand, Runx2/SP7 and BMP2/Runx2/SP7 cocktails had the highest ALP with activity at least 3-fold higher, and mRNA expression of at least 7-fold higher, when compared to the Mock cells. Overall Runx2-, SP7-HUCPVCs and the Runx2/SP7 cocktail had the best performance in terms of ALP expression among the samples followed by the BMP2/Runx2/SP7 cocktail. Based on these results it seems that the mix of different groups of cells in cocktails does not have an additive effect in the ALP expression since none of the cocktails expressed higher levels than Runx2- and SP7-HUCPVCs; but it has a complementary effect as seen when BMP2-HUCPVCs were mixed with Runx2- and/or SP7-HUCPVCs, significantly increasing (P<0.05) the level of ALP activity (Figure 3.22); an effect that can also be seen in the ALP staining (Figure 3.19). Moreover, it is important to highlight that ALP activity and mRNA expression peaked at day 6-7 and started to diminish after that, a behaviour that has been previously reported after modifying mouse BMSCs (81) and rat BMSCs (230) with Ad-Runx2.

Similarly, it was found that although expression of other osteogenic genes such as COL1A1, OCN and OPN was upregulated when compared to Mock-HUCPVCs by day 9 levels had decreased almost to levels comparable with the mock indicating that the effect of the BMP2, Runx2 and SP7 genetic modification via adenoviral infection was not lasting enough to cause a robust effect in the modified cells; a finding that could be explained by the low longevity of transgene expression due to fast proliferation of HUCPVCs (discussed in Sections 4.1 and 4.2).

When testing the capacity of the different groups and cocktails of genetically modified HUCPVCs to mineralize matrix, all cultures failed to mineralize as indicated by negative von Kossa staining at days 21, 28 and 35, despite being cultured in osteogenic media. These unexpected results differ with previous reports by Sarugaser et al. (56,75) and Baksh et al (278) who showed matrix mineralization in naive HUCPVCs cultures after being maintained in osteogenic medium for 7-14 days and 35 days respectively. Moreover, Wen et al. (297)

122

reported positive von Kossa staining in osteogenic cultures of human umbilical cord-MSCs (hUC-MSCs) — cells that were harvested in a similar manner to HUCPVCs. Furthermore, others have successfully showed matrix mineralization in osteogenic cultures of MSCs isolated by enzymatic digestion of small pieces (1-2 mm3) of the umbilical cord (UC) (298–301) and in osteogenic cultures of cells harvested from the UC after removing the vessels (54,302,303).

Although we cannot explain the differences between these results and ours, several factors may have had a role in the different outcome. For example, harvesting procedures especially the ones that include the vessels during the enzymatic digestion, could lead to an increased harvest of osteoprogenitor cells by digesting the UC vessel walls. Previously, it has been shown that in the subendothelial layer of the human umbilical cord vein there is a "MSC-like cell" population capable of expressing ALP after being exposed to osteogenic medium (304). Also, it is well known that blood vessels contain calcifying vascular cells (305) capable of forming spontaneous calcified nodules in vitro without the addition of exogenous factors (306,307). The latter might explain the spontaneous bone nodule formation observed by Sarugaser et al. (56) in HUCPVCs cultures in non-osteogenic conditions; an occurrence that could have been caused by overdigestion of the perivascular area of the UC vessels followed by the digestion of the actual vessel walls. Also, culture conditions such as FBS batch and concentration could lead to significant differences in proliferation and differentiation of the cells (308) due to variations in levels of hormones, growth and differentiation factors and other important proteins (309). The majority of reports referenced in this section (54,278,297–303) utilized osteogenic media supplemented with 10% FBS while we used 2% FBS; a concentration chosen to maximize longevity of gene expression (Section 4.1) while guarantying proper cell growth and differentiation as previously demonstrated by Sarugaser et al. (75). Furthermore, osteogenic medium components (ascorbic acid, dexamethasone and -glycerophosphate) and their concentrations play an important role in the success of osteogenic cultures; small changes in the concentration of -GP can have a dramatic impact on the outcome of matrix mineralization as shown in this thesis work (Figure 3.37). In eight (54,297–303) studies referenced in this section osteogenic medium was supplemented with 10mM -glycerophosphate (-GP) while Sarugaser et al. (56,75) and Baksh et al. (278) used 5mM -GP. On the other hand, we added 123

3.5 mM -GP to the cultures only after detecting cell multilayering by phase microscopy to avoid dystrophic mineralization (310).

To test the osteoinductive and osteogenic capacity of engineered HUCPVCs four different experimental groups (Mock-HUCPVCs, VEGF-HUCPVCs, BMP2/Runx2/SP7 and BMP2/Runx2/SP7/VEGF) were injected intramuscularly in NOD/SCID mice but after 6 weeks we failed to detect the formation of calcified tissue in the area of injection by either x-ray or Micro-CT. Previously, heterotopic bone formation has been shown by others when delivering cells engineered to secrete BMP2 (163,311,312) using the method chosen for this in vivo work which was the intramuscular injection of 5x106 cells suspended in 100 μl PBS. We believe that the main reason for our negative result was that the small amount of mature BMP2 produced by BMP2-HUCPVCs after the inefficient cleavage of proBMP2 (see Section 4.3) was not enough to induce bone formation and although Hillger et al. (141) reported that 0.25µmole of proBMP2 induced ectopic bone formation we never attempted to quantify the amount of secreted proBMP2 by BMP2-HUCPVCs therefore it is unknown if BMP2-HUCPVCs were producing non-bioactive proBMP2 or if the quantity produced was not enough to induce bone formation in an heterotopic site. For our proposed cocktail strategy BMP2 is fundamental to effective bone formation by inducing the osteogenic differentiation of naïve cells. Since the transcription factors Runx2 and SP7 do not have a paracrine effect in the naive cells it would explain why it has been reported that Runx2 therapy is not as effective as BMP2 ex vivo gene therapy (236,237). Although heterotopic bone formation has been shown when delivering cells overexpressing Runx2 (81,230,237,265) and SP7 (265) the reported work always involved the delivery of the cells in scaffolds. In our case the cells were delivered without a scaffold which might have had an impact in the final results as the injected cells might have dispersed due to the absence of a scaffold to contain them in the area of delivery.

BMP2 ex vivo gene therapy using different sources of animal and human cells has been shown to be a powerful therapeutic tool to induce bone formation in several in vivo studies (114,162– 165) whereas research in the Runx2 and SP7 ex vivo gene therapy field is not abundant and most of the work has been done in non-human cells. Only three papers were found using the

124

genetic modification of human cells, two in which ADSCs were microporated with plasmids carrying Runx2 and/or SP7 (265) and BMP2 or Runx2/BMP2 (238) and one using lipofectamine to deliver the SP7 gene into umbilical cord-derived mesenchymal stromal cells (247). Out of these three reports only Lee et al. (265) provided convincing evidence of matrix mineralization in vitro and in vivo after modifying ADSCs with Runx2 and/or SP7. The low number of reports using human cells engineered to express the transcription factors Runx2 and SP7 could be an indication of research hindered by challenges, of the type discussed in detail herein, but not normally addressed in the published literature.

4.5. Overexpression of Runx2 or SP7 enhances ALP activity and matrix mineralization in HBMSCs.

Matrix mineralization was not detected in cultures of genetically modified HUCPVCs (see section 5.4). Thus, as an alternative, HBMSCs were genetically modified to demonstrate, as shown by others in rat BMSCs (81,230,232,265), that the adenoviral Runx2, or SP7, constructs could enhance matrix mineralization in vitro. For this experiment HBMSCs were cultured in parallel with HUCPVCs and under the same conditions; their osteogenic potential was assessed by evaluating ALP activity and matrix mineralization.

When comparing the effect of genetic modification of both HUCPVCs and HBMSCs with Runx2 and SP7 (Figure 3.33) it was observed that overexpression of these genes had a higher effect in HUCPVCs by increasing their ALP activity levels at least 10-fold while approximately 4-fold in HBMSCs. Despite this greater effect, in general ALP activity levels in HBMSCs were found to be much higher (~7-fold) than in HUCPVCs (Figure 3.34), for example ALP levels of Runx2- HUCPVCs, SP7-HUCPVCs and Runx2/SP7 HUCPVCs were comparable with the levels found in Mock-HBMSCs (Figure 4.34) which is a strong indication of the low intrinsic ALP levels found in HUCPVCs; a result that differed from a previous report by Baksh et al. (278) but was comparable with a report from Wen et al. (297) who found a significant difference (P<0.05) in ALP activity levels in osteogenic cultures of naive HBMSCs and human umbilical cord-MSCs

125

(hUC-MSCs) — cells that were harvested in a similar manner to HUCPVCs. Higher inherent levels of ALP in HBMCSc could be explained by the reported existence of at least two classes of osteoprogenitors in this cell source, those that need to be induced and those that are committed (313). This is also an explanation for the formation of heterotopic bone after in vivo implantation of naive human BMSCs in ectopic sites (314–317).

When evaluating matrix mineralization it was observed that Runx2-HBMSCs, SP7-HBMSCs and the cocktail Runx2/SP7 were highly positive for alizarin red staining when compared to the Mock-HBMSCs (Figure 4.35). These results correlated with the quantification of calcium content of the cultures (Figure 4.36) where significant differences were found between the Mock-HBMSCs (P<0.01) and Runx2-HBMSCs, SP7-HBMSCs and the cocktail Runx2/SP7, although no significant differences were found among the latter. These results could indicate that overexpression of Runx2 and SP7 have a similar effect in calcium deposition in HBMSCs and that the mix of Runx2- and SP7-HBMSCs in cocktails does not have an additive effect in matrix mineralization since the amount of calcium deposition quantified in the cocktail was not higher than in Runx2- and SP7-HBMSCs. In a similar study Lee et al. (265) transiently overexpressed Runx2 and/or SP7 in human ADSCs; although calcium content of the cultures was not quantified, alizarin red staining showed enhanced calcium deposition in Runx2-, SP7- ADSCs and cells co-expressing both genes when compared to the control. Our results corroborate their findings, although they showed that Runx2 overexpression had a greater effect in calcium deposition in ADSCs when compared to SP7, while we detected no significant difference in calcium deposition among Runx2- and SP7-HUCPVCs cultures. Furthermore, we detected earlier matrix mineralization (at least a week) in Runx2-HBMSCs, SP7-HBMSCs and the cocktail Runx2/SP7 than in the mock cultures (Figure 3.35-A). This was also reported by both Zhao et al. (81) after transient overexpression of Runx2 in mouse BMSCs, and by Byers et al. (232) after modifying rat BMSCs with a retrovirus carrying the Runx2 gene.

Since this was meant to be a comparative study both cell types needed to be seeded at the same cell density and have the same FBS conditions. Although, HUCPVCs osteogenic cultures during this thesis work were supplemented with 2% FBS to maximize longevity of gene

126

expression (Section 4.1), for this study 10% FBS concentration was chosen since unlike HUCPVCs, HBMSCs do not grow well in medium supplemented with 5-2% FBS (75). Thus, perhaps the conditions employed during these experiments are not optimal for the osteogenic differentiation HUCPVCs. Indeed, due to their higher proliferation rate, most of the HUCPVCs samples rolled-up and off the culture dishes under these conditions, as has been reported previously when comparing the proliferative potential of HUCPVCs with HBMSCs (278,297). Nevertheless, some HUCPVC samples did stain a light purple with alizarin red (Figure 4.35-B), although this differed significantly with what was found in HBMSC cultures (Figure 4.35-A). It is possible that, under the conditions employed during these experiments, the longevity of expression of Runx2 and SP7 in HUCPVCs is insufficient to have an enhancing effect in matrix mineralization although it is enough to significantly enhance matrix mineralization in HBMSCs cultures.

While others have shown improved bone formation in vivo and in vitro after genetically modifying BMSCs from mouse (81,229), rat (232) and rabbit (261) to overexpress Runx2 or SP7, to our knowledge this is the first report to show enhanced ALP activity and matrix mineralization in human BMSCs after genetic modification with Runx2 or SP7.

The hypothesis underlying the work undertaken herein was that HUCPVCs are a putative cell source for osteogenic ex vivo gene therapy that when genetically modified and employed in cocktails overexpressing different osteogenic related genes would be more biologically effective than populations overexpressing a single gene, and which biological activity could be maintained upon thaw after being cryopreserved. The work reported through the previous sections addressed the hypothesis and demonstrates that HUCPVCs can be cryopreserved without affecting the production efficiency and bioactivity of proteins of interest produced by the cells. Furthermore, under the experimental conditions employed during this work and considering the inefficient expression of BMP2 by the modified cells due to problems in the vector design it would be unreasonable to conclude that genetically modified HUCPVCs are not an ideal osteogenic cell source since more experimentation has to be performed with all the

127

osteogenic factors working properly and under the appropriate environment before any definitive conclusion can be reached.

4.6. Conclusions

From the work reported herein, it can be concluded that:

 HUCPVCs are genetically modifiable by non-viral and viral method; and can be cryopreserved without affecting the production efficiency and bioactivity of proteins of interest produced by the cells.  HUCPVCs are more effectively infected by adenoviruses than HBMSC but HBMSCs have higher osteogenic potential than HUCPVCs.  Transient overexpression of Runx2 and SP7 in HUCPVCs does not enhance matrix mineralization, but overexpression of Runx2 and SP7 enhances ALP activity and matrix mineralization in HBMSCs.  Overexpression of Runx2 and SP7 enhances ALP activity levels and upregulates ALP, OPN, COL1A1 and OCN gene expression in HUCPVCs.

4.7. Future Directions

It was concluded that 3x HA tagged BMP2 is not efficiently processed into mature BMP2 (Section 4.3). To prove that this was, indeed, the cause of the problems reported, it would be necessary to redesign the adenoviral vector and remove the C-terminal 3X HA tag fused to the BMP2 gene. If this resulted in the BMP2-HUCPVCs efficiently producing mature BMP2 this could be tested both by the in vitro and in vivo assays employed herein. It would also, of course, be important to retest the contribution of BMP2-HUCPVCs to the different cocktails tested.

For undetermined reasons all genetically modified HUCPVCs cultures failed to form mineralized matrix in vitro despite being grown in osteogenic medium. As discussed in Section 4.4 many 128

factors such as composition of the osteogenic media and the batch of FBS used may have played a role in this outcome. Therefore, since the effect of BMP2 in HUCPVCs could not be tested — due to the small amount produced by BMP2-HUCPVCs — it will be interesting to supplement the osteogenic medium with rhBMP2 to establish if this osteoinductive protein can act as the missing cue to accomplish HUCPVC differentiation.

Runx2 and SP7 overexpression enhanced matrix mineralization in HBMSCs but not in HUCPVCs. Even though, during this thesis work it was found that HBMSCs have a higher osteogenic potential than HUCPVCs it is not understood why the genetic modification of HUCPVCs with Runx2 and SP7 did not induce matrix mineralization. Thus, to better understand these results it will be important to quantify and compare the production of these transcription factors in Runx2-HUCPVCs and Runx2-HBMSCs and, SP7-HUCPVCs and SP7-HBMSCs by performing Runx2 and SP7 ELISAs from the cell lysates. Significant differences in these levels might indicate inadequate production of these transcriptions factors, which could help to explain the negative outcome obtained in HUCPVCs. However, if the levels of expression are found to be similar it could indicate that, due to the lower osteogenic capacity of HUCPVCs, they require the levels of overexpression to be higher in order to elicit matrix mineralization or that downstream genes are not being properly turned on.

The advantage of our proposed cocktail strategy is its flexibility to allow the modification of its configuration by simply changing the proportion of the different types of cells (BMP2-, Runx2-, SP7- and VEGF-HUCPVCs) within the cocktail. The tools and information acquired during this work are valuable for future in vivo testing in which it will be possible to assess the effect on bone healing of differently configured cocktails and thus find a "recipe" that can provide more effective and robust bone formation than others. The first tests should be done in simple animal models to demonstrate that the strategy can induce heterotopic bone formation. Only then could more challenging bone defect scenarios be addressed. As discussed previously (Section 4.4) we believe that the delivery of the cells without a scaffold might have had an impact on the negative results. Thus, it would be worthwhile to seed the cells in a scaffold to avoid the dispersion of the cells after delivery.

129

One of the objectives of this thesis work was to genetically modify HUCPVCs and store them in liquid nitrogen to assess the possibility of creating banks of cryopreserved engineered human MSCs to be used in osteogenic ex vivo gene therapy. Our in vitro results showed that genetically modified HUCPVCs can be cryopreserved and thawed without affecting the production efficiency and bioactivity of proteins of interest produced by the cells. However, in vivo tests were not performed on these cells due to the issues with the low BMP2 production. Thus, once the latter issue is resolved, parallel in vivo tests have to be carried out utilizing pre- and post-liquid nitrogen genetically modified HUCPVCs to find out if the cryopreservation process has an effect on the bone formation capacity of these cells in vivo.

Bone healing is a complex process that requires the upregulation and downregulation of several genes at different time points, hence it will be of value to modify the adenoviral constructs to incorporate inducible expression systems such as TET-On and TET-off that will allow the control of the level and time of expression of desired genes within the cocktail. By controlling their expression it may be possible to recapitulate the sequence in which these genes of interest are turned on and off during the normal process of bone formation and therefore provide a more efficient bone healing strategy than is currently available today.

130

References

1. Martin RB. The importance of mechanical loading in bone biology and medicine. J Musculoskelet Neuronal Interact. 2007 Mar;7(1):48–53.

2. Yaszemski MJ, Payne RG, Hayes WC, Langer R, Mikos AG. Evolution of bone transplantation: molecular, cellular and tissue strategies to engineer human bone. Biomaterials. 1996 Jan;17(2):175–85.

3. Jensen ED, Gopalakrishnan R, Westendorf JJ. Regulation of gene expression in osteoblasts. Biofactors. 2010 Jan;36(1):25–32.

4. Salter RB. Textbook of Disorders and Injuries of the Musculoskeletal System: An Introduction to Orthopaedics, Fractures, and Joint Injuries, Rheumatology, Metabolic Bone Disease, and Rehabilitation. Lippincott Williams & Wilkins; 1999.

5. Marks S, Hermey D. The Structure and Development of Bone. Principles of Bone Biology. San Diego: Academic Press; 1996. p. 3–14.

6. University RAP of BSWS. Biology of Aging : Observations and Principles: Observations and Principles. Oxford University Press; 2006.

7. Davies JE. Understanding peri-implant endosseous healing. J Dent Educ. 2003 Aug;67(8):932–49.

8. Ducy P, Schinke T, Karsenty G. The osteoblast: a sophisticated fibroblast under central surveillance. Science. 2000 Sep 1;289(5484):1501–4.

9. Teitelbaum SL. Bone resorption by osteoclasts. Science. 2000 Sep 1;289(5484):1504–8.

10. Aubin, JE, Liu, F. The Osteoblast Lineage. Principles of Bone Biology. San Diego: The Academic Press; 1996. p. 51–68.

11. Javed A, Chen H, Ghori FY. Genetic and Transcriptional Control of Bone Formation. Oral and Maxillofacial Surgery Clinics of North America. 2010 Aug;22(3):283–93.

12. Neve A, Corrado A, Cantatore FP. Osteoblast physiology in normal and pathological conditions. Cell Tissue Res. 2011 Feb;343(2):289–302.

13. Stein GS, Lian JB, Owen TA. Relationship of cell growth to the regulation of tissue-specific gene expression during osteoblast differentiation. FASEB J. 1990 Oct;4(13):3111–23.

14. Eriksen EF. Cellular mechanisms of bone remodeling. Rev Endocr Metab Disord. 2010 Dec;11(4):219–27.

15. Sinha KM, Zhou X. Genetic and molecular control of Osterix in skeletal formation. J Cell Biochem. 2012 Dec 5; 131

16. Komori T. Regulation of osteoblast differentiation by transcription factors. J Cell Biochem. 2006 Dec 1;99(5):1233–9.

17. Marie PJ. Transcription factors controlling osteoblastogenesis. Arch Biochem Biophys. 2008 May 15;473(2):98–105.

18. Karsenty G. Transcriptional control of skeletogenesis. Annu Rev Genomics Hum Genet. 2008;9:183–96.

19. Lian JB, Javed A, Zaidi SK, Lengner C, Montecino M, van Wijnen AJ, et al. Regulatory controls for osteoblast growth and differentiation: role of Runx/Cbfa/AML factors. Crit Rev Eukaryot Gene Expr. 2004;14(1-2):1–41.

20. Robey, PG, Boskey, A. The Biochemistry of Bone. Osteoporosis. San Diego: The Academic \press; 1996. p. 95–182.

21. Duvall CL, Taylor WR, Weiss D, Wojtowicz AM, Guldberg RE. Impaired angiogenesis, early callus formation, and late stage remodeling in fracture healing of osteopontin-deficient mice. J Bone Miner Res. 2007 Feb;22(2):286–97.

22. Perumal V, Roberts CS. (ii) Factors contributing to non-union of fractures. Current Orthopaedics. 2007 Aug;21(4):258–61.

23. Giannoudis PV, Dinopoulos H, Tsiridis E. Bone substitutes: an update. Injury. 2005 Nov;36 Suppl 3:S20–27.

24. Braddock M, Houston P, Campbell C, Ashcroft P. Born again bone: tissue engineering for bone repair. News Physiol Sci. 2001 Oct;16:208–13.

25. Peng H, Wright V, Usas A, Gearhart B, Shen H-C, Cummins J, et al. Synergistic enhancement of bone formation and healing by stem cell-expressed VEGF and bone morphogenetic protein-4. J Clin Invest. 2002 Sep;110(6):751–9.

26. Bostrom MP, Saleh KJ, Einhorn TA. Osteoinductive growth factors in preclinical fracture and long bone defects models. Orthop Clin North Am. 1999 Oct;30(4):647–58.

27. De Long WG Jr, Einhorn TA, Koval K, McKee M, Smith W, Sanders R, et al. Bone grafts and bone graft substitutes in orthopaedic trauma surgery. A critical analysis. J Bone Joint Surg Am. 2007 Mar;89(3):649–58.

28. Finkemeier CG. Bone-grafting and bone-graft substitutes. J Bone Joint Surg Am. 2002 Mar;84-A(3):454–64.

29. Vaccaro AR. The role of the osteoconductive scaffold in synthetic bone graft. Orthopedics. 2002 May;25(5 Suppl):s571–578.

132

30. Mankin HJ, Hornicek FJ, Raskin KA. Infection in massive bone allografts. Clin Orthop Relat Res. 2005 Mar;(432):210–6.

31. Sorger JI, Hornicek FJ, Zavatta M, Menzner JP, Gebhardt MC, Tomford WW, et al. Allograft fractures revisited. Clin Orthop Relat Res. 2001 Jan;(382):66–74.

32. Bucholz RW. Nonallograft osteoconductive bone graft substitutes. Clin Orthop Relat Res. 2002 Feb;(395):44–52.

33. Bueno EM, Glowacki J. Cell-free and cell-based approaches for bone regeneration. Nat Rev Rheumatol. 2009 Dec;5(12):685–97.

34. Boden SD. The ABCs of BMPs. Orthop Nurs. 2005 Feb;24(1):49–52; quiz 53–54.

35. Bishop GB, Einhorn TA. Current and future clinical applications of bone morphogenetic proteins in orthopaedic trauma surgery. Int Orthop. 2007 Dec;31(6):721–7.

36. Latzman JM, Kong L, Liu C, Samadani U. Administration of human recombinant bone morphogenetic protein-2 for spine fusion may be associated with transient postoperative renal insufficiency. Spine. 2010 Apr 1;35(7):E231–237.

37. Uludag H, D’Augusta D, Golden J, Li J, Timony G, Riedel R, et al. Implantation of recombinant human bone morphogenetic proteins with biomaterial carriers: A correlation between protein pharmacokinetics and osteoinduction in the rat ectopic model. J Biomed Mater Res. 2000 May;50(2):227–38.

38. Langer R, Vacanti JP. Tissue engineering. Science. 1993 May 14;260(5110):920–6.

39. Reddi AH. Symbiosis of biotechnology and biomaterials: applications in tissue engineering of bone and cartilage. J Cell Biochem. 1994 Oct;56(2):192–5.

40. Bruder SP, Fox BS. Tissue engineering of bone. Cell based strategies. Clin Orthop Relat Res. 1999 Oct;(367 Suppl):S68–83.

41. Lanza R, Langer R, Vacanti JP. Principles of tissue engineering. 2nd ed. San Diego, Calif; 2000.

42. Friedenstein AJ, Piatetzky-Shapiro II, Petrakova KV. Osteogenesis in transplants of bone marrow cells. J Embryol Exp Morphol. 1966 Dec;16(3):381–90.

43. Friedenstein AJ, Petrakova KV, Kurolesova AI, Frolova GP. Heterotopic of bone marrow. Analysis of precursor cells for osteogenic and hematopoietic tissues. Transplantation. 1968 Mar;6(2):230–47.

44. Friedenstein AJ, Chailakhjan RK, Lalykina KS. The development of fibroblast colonies in monolayer cultures of guinea-pig bone marrow and spleen cells. Cell Tissue Kinet. 1970 Oct;3(4):393–403.

133

45. Friedenstein AJ, Deriglasova UF, Kulagina NN, Panasuk AF, Rudakowa SF, Luriá EA, et al. Precursors for fibroblasts in different populations of hematopoietic cells as detected by the in vitro colony assay method. Exp Hematol. 1974;2(2):83–92.

46. Friedenstein AJ, Chailakhyan RK, Gerasimov UV. Bone marrow osteogenic stem cells: in vitro cultivation and transplantation in diffusion chambers. Cell Tissue Kinet. 1987 May;20(3):263–72.

47. Caplan AI. Mesenchymal stem cells. Journal of Orthopaedic Research. 1991 Sep;9(5):641– 50.

48. Caplan AI. The mesengenic process. Clin Plast Surg. 1994 Jul;21(3):429–35.

49. Erices A, Conget P, Minguell JJ. Mesenchymal progenitor cells in human umbilical cord blood. Br J Haematol. 2000 Apr;109(1):235–42.

50. Gronthos S, Mankani M, Brahim J, Robey PG, Shi S. Postnatal human dental pulp stem cells (DPSCs) in vitro and in vivo. Proc Natl Acad Sci USA. 2000 Dec 5;97(25):13625–30.

51. De Bari C, Dell’Accio F, Tylzanowski P, Luyten FP. Multipotent mesenchymal stem cells from adult human synovial membrane. Arthritis Rheum. 2001 Aug;44(8):1928–42.

52. Zuk PA, Zhu M, Ashjian P, De Ugarte DA, Huang JI, Mizuno H, et al. Human adipose tissue is a source of multipotent stem cells. Mol Biol Cell. 2002 Dec;13(12):4279–95.

53. In ’t Anker PS, Scherjon SA, Kleijburg-van der Keur C, de Groot-Swings GMJS, Claas FHJ, Fibbe WE, et al. Isolation of mesenchymal stem cells of fetal or maternal origin from human placenta. Stem Cells. 2004;22(7):1338–45.

54. Wang H-S, Hung S-C, Peng S-T, Huang C-C, Wei H-M, Guo Y-J, et al. Mesenchymal stem cells in the Wharton’s jelly of the human umbilical cord. Stem Cells. 2004;22(7):1330–7.

55. Shih DT, Lee D-C, Chen S-C, Tsai R-Y, Huang C-T, Tsai C-C, et al. Isolation and characterization of neurogenic mesenchymal stem cells in human scalp tissue. Stem Cells. 2005 Aug;23(7):1012–20.

56. Sarugaser R, Lickorish D, Baksh D, Hosseini MM, Davies JE. Human umbilical cord perivascular (HUCPV) cells: a source of mesenchymal progenitors. Stem Cells. 2005 Feb;23(2):220–9.

57. Crisan M, Yap S, Casteilla L, Chen C-W, Corselli M, Park TS, et al. A perivascular origin for mesenchymal stem cells in multiple human organs. Cell Stem Cell. 2008 Sep 11;3(3):301– 13.

58. Nadri S, Soleimani M. Comparative analysis of mesenchymal stromal cells from murine bone marrow and amniotic fluid. Cytotherapy. 2007;9(8):729–37.

134

59. Patki S, Kadam S, Chandra V, Bhonde R. Human breast milk is a rich source of multipotent mesenchymal stem cells. Hum Cell. 2010 May;23(2):35–40.

60. Dominici M, Le Blanc K, Mueller I, Slaper-Cortenbach I, Marini F, Krause D, et al. Minimal criteria for defining multipotent mesenchymal stromal cells. The International Society for Cellular Therapy position statement. Cytotherapy. 2006;8(4):315–7.

61. Barry FP, Murphy JM. Mesenchymal stem cells: clinical applications and biological characterization. Int J Biochem Cell Biol. 2004 Apr;36(4):568–84.

62. Singer NG, Caplan AI. Mesenchymal stem cells: mechanisms of inflammation. Annu Rev Pathol. 2011;6:457–78.

63. Le Blanc K. Immunomodulatory effects of fetal and adult mesenchymal stem cells. Cytotherapy. 2003;5(6):485–9.

64. Nauta AJ, Fibbe WE. Immunomodulatory properties of mesenchymal stromal cells. Blood. 2007 Nov 15;110(10):3499–506.

65. Bruder SP, Kurth AA, Shea M, Hayes WC, Jaiswal N, Kadiyala S. Bone regeneration by implantation of purified, culture-expanded human mesenchymal stem cells. J Orthop Res. 1998 Mar;16(2):155–62.

66. Bruder SP, Kraus KH, Goldberg VM, Kadiyala S. The effect of implants loaded with autologous mesenchymal stem cells on the healing of canine segmental bone defects. J Bone Joint Surg Am. 1998 Jul;80(7):985–96.

67. Quarto R, Mastrogiacomo M, Cancedda R, Kutepov SM, Mukhachev V, Lavroukov A, et al. Repair of large bone defects with the use of autologous bone marrow stromal cells. N Engl J Med. 2001 Feb 1;344(5):385–6.

68. Petite H, Viateau V, Bensaïd W, Meunier A, de Pollak C, Bourguignon M, et al. Tissue- engineered bone regeneration. Nat Biotechnol. 2000 Sep;18(9):959–63.

69. Caplan AI. Why are MSCs therapeutic? New data: new insight. J Pathol. 2009 Jan;217(2):318–24.

70. Mueller SM, Glowacki J. Age-related decline in the osteogenic potential of human bone marrow cells cultured in three-dimensional collagen sponges. J Cell Biochem. 2001;82(4):583–90.

71. Bruder SP, Jaiswal N, Haynesworth SE. Growth kinetics, self-renewal, and the osteogenic potential of purified human mesenchymal stem cells during extensive subcultivation and following cryopreservation. Journal of Cellular Biochemistry. 1997;64(2):278–94.

135

72. Banfi A, Muraglia A, Dozin B, Mastrogiacomo M, Cancedda R, Quarto R. Proliferation kinetics and differentiation potential of ex vivo expanded human bone marrow stromal cells: Implications for their use in cell therapy. Exp Hematol. 2000 Jun;28(6):707–15.

73. Derubeis AR, Cancedda R. Bone marrow stromal cells (BMSCs) in bone engineering: limitations and recent advances. Ann Biomed Eng. 2004 Jan;32(1):160–5.

74. Baksh D, Yao R, Tuan RS. Comparison of proliferative and multilineage differentiation potential of human mesenchymal stem cells derived from umbilical cord and bone marrow. Stem Cells. 2007 Jun;25(6):1384–92.

75. Sarugaser R, Hanoun L, Keating A, Stanford WL, Davies JE. Human mesenchymal stem cells self-renew and differentiate according to a deterministic hierarchy. PLoS ONE. 2009;4(8):e6498.

76. Zebardast N, Lickorish D, Davies JE. Human umbilical cord perivascular cells (HUCPVC): A mesenchymal cell source for dermal wound healing. Organogenesis. 2010 Dec;6(4):197– 203.

77. Emrani H, Davies JE. Umbilical Cord Perivascular Cells: A Mesenchymal Cell Source for Treatment of Tendon Injuries. The Open Tissue Engineering and Regenerative Medicine Journal ,. 2011;4:112 The Open Tissue Engineering and Regenerative Medicine Journal , 2011, 4, 112–9.

78. Ennis J, Götherström C, Le Blanc K, Davies JE. In vitro immunologic properties of human umbilical cord perivascular cells. Cytotherapy. 2008;10(2):174–81.

79. Ennis J, Sarugaser R, Gomez A, Baksh D, Davies JE. Isolation, characterization, and differentiation of human umbilical cord perivascular cells (HUCPVCs). Methods Cell Biol. 2008;86:121–36.

80. Gómez-Aristizábal A, Ng C, Ng J, Davies JE. Effects of two mesenchymal cell populations on hepatocytes and lymphocytes. Liver Transpl. 2012 Nov;18(11):1384–94.

81. Zhao Z, Zhao M, Xiao G, Franceschi RT. Gene transfer of the Runx2 transcription factor enhances osteogenic activity of bone marrow stromal cells in vitro and in vivo. Mol Ther. 2005 Aug;12(2):247–53.

82. Ebara S, Nakayama K. Mechanism for the action of bone morphogenetic proteins and regulation of their activity. Spine. 2002 Aug 15;27(16 Suppl 1):S10–15.

83. McGuire MK, Scheyer ET, Schupbach P. Growth factor-mediated treatment of recession defects: a randomized controlled trial and histologic and microcomputed tomography examination. J Periodontol. 2009 Apr;80(4):550–64.

136

84. Park YJ, Lee YM, Lee JY, Seol YJ, Chung CP, Lee SJ. Controlled release of platelet-derived growth factor-BB from chondroitin sulfate-chitosan sponge for guided bone regeneration. J Control Release. 2000 Jul 3;67(2-3):385–94.

85. Nakamura K, Kawaguchi H, Aoyama I, Hanada K, Hiyama Y, Awa T, et al. Stimulation of bone formation by intraosseous application of recombinant basic fibroblast growth factor in normal and ovariectomized rabbits. J Orthop Res. 1997 Mar;15(2):307–13.

86. Busch O, Solheim E, Bang G, Tornes K. Guided tissue regeneration and local delivery of insulinlike growth factor I by bioerodible polyorthoester membranes in rat calvarial defects. Int J Oral Maxillofac Implants. 1996 Aug;11(4):498–505.

87. Anderson WF. September 14, 1990: the beginning. Hum Gene Ther. 1990;1(4):371–2.

88. Cavazzana-Calvo M, Hacein-Bey S, de Saint Basile G, Gross F, Yvon E, Nusbaum P, et al. Gene therapy of human severe combined immunodeficiency (SCID)-X1 disease. Science. 2000 Apr 28;288(5466):669–72.

89. Evans CH. Gene therapy for bone healing. Expert Rev Mol Med. 2010;12:e18.

90. Evans C. Gene therapy for the regeneration of bone. Injury. 2011 Jun;42(6):599–604.

91. Fischer J, Kolk A, Wolfart S, Pautke C, Warnke PH, Plank C, et al. Future of local bone regeneration - Protein versus gene therapy. J Craniomaxillofac Surg. 2011 Jan;39(1):54–64.

92. Baltzer AWA, Lieberman JR. Regional gene therapy to enhance bone repair. Gene Ther. 2004 Feb;11(4):344–50.

93. Santos JL, Pandita D, Rodrigues J, Pêgo AP, Granja PL, Tomás H. Non-viral gene delivery to mesenchymal stem cells: methods, strategies and application in bone tissue engineering and regeneration. Curr Gene Ther. 2011 Feb;11(1):46–57.

94. Kofron MD, Laurencin CT. Bone tissue engineering by gene delivery. Adv Drug Deliv Rev. 2006 Jul 7;58(4):555–76.

95. Anderson WF. Human gene therapy. Nature. 1998 Apr 30;392(6679 Suppl):25–30.

96. Oakes DA, Lieberman JR. Osteoinductive applications of regional gene therapy: ex vivo gene transfer. Clin Orthop Relat Res. 2000 Oct;(379 Suppl):S101–112.

97. Castilho L dos R. Animal cell technology: from biopharmaceuticals to gene therapy. Routledge; 2008.

98. Phillips JE, Gersbach CA, García AJ. Virus-based gene therapy strategies for bone regeneration. Biomaterials. 2007 Jan;28(2):211–29.

137

99. Hong D, Chen H-X, Ge R, Li J-C. Genetically engineered mesenchymal stem cells: The ongoing research for bone tissue engineering. Anat Rec (Hoboken). 2010 Mar;293(3):531– 7.

100. Hoekstra D, Rejman J, Wasungu L, Shi F, Zuhorn I. Gene delivery by cationic lipids: in and out of an endosome. Biochem Soc Trans. 2007 Feb;35(Pt 1):68–71.

101. Gresch O, Engel FB, Nesic D, Tran TT, England HM, Hickman ES, et al. New non-viral method for gene transfer into primary cells. Methods. 2004 Jun;33(2):151–63.

102. Aluigi M, Fogli M, Curti A, Isidori A, Gruppioni E, Chiodoni C, et al. Nucleofection is an efficient nonviral transfection technique for human bone marrow-derived mesenchymal stem cells. Stem Cells. 2006 Feb;24(2):454–61.

103. Fechner H, Haack A, Wang H, Wang X, Eizema K, Pauschinger M, et al. Expression of coxsackie adenovirus receptor and alphav-integrin does not correlate with adenovector targeting in vivo indicating anatomical vector barriers. Gene Ther. 1999 Sep;6(9):1520–35.

104. Tepper OM, Mehrara BJ. Gene therapy in plastic surgery. Plast Reconstr Surg. 2002 Feb;109(2):716–34.

105. Partridge KA, Oreffo ROC. Gene delivery in bone tissue engineering: progress and prospects using viral and nonviral strategies. Tissue Eng. 2004 Feb;10(1-2):295–307.

106. Oligino TJ, Yao Q, Ghivizzani SC, Robbins P. Vector systems for gene transfer to joints. Clin Orthop Relat Res. 2000 Oct;(379 Suppl):S17–30.

107. Kurian KM, Watson CJ, Wyllie AH. Retroviral vectors. MP, Mol Pathol. 2000 Aug;53(4):173–6.

108. Franceschi RT. Biological approaches to bone regeneration by gene therapy. J Dent Res. 2005 Dec;84(12):1093–103.

109. Yi Y, Hahm SH, Lee KH. Retroviral gene therapy: safety issues and possible solutions. Curr Gene Ther. 2005 Feb;5(1):25–35.

110. Uren AG, Kool J, Berns A, van Lohuizen M. Retroviral insertional mutagenesis: past, present and future. Oncogene. 2005 Nov 21;24(52):7656–72.

111. Zhang X-Y, La Russa VF, Bao L, Kolls J, Schwarzenberger P, Reiser J. Lentiviral vectors for sustained transgene expression in human bone marrow-derived stromal cells. Mol Ther. 2002 May;5(5 Pt 1):555–65.

112. Bushman FD. Integration site selection by lentiviruses: biology and possible control. Curr Top Microbiol Immunol. 2002;261:165–77.

138

113. Lieberman JR, Daluiski A, Stevenson S, Wu L, McAllister P, Lee YP, et al. The effect of regional gene therapy with bone morphogenetic protein-2-producing bone-marrow cells on the repair of segmental femoral defects in rats. J Bone Joint Surg Am. 1999 Jul;81(7):905–17.

114. Peterson B, Zhang J, Iglesias R, Kabo M, Hedrick M, Benhaim P, et al. Healing of critically sized femoral defects, using genetically modified mesenchymal stem cells from human adipose tissue. Tissue Eng. 2005 Feb;11(1-2):120–9.

115. Rundle CH, Miyakoshi N, Kasukawa Y, Chen S-T, Sheng MH-C, Wergedal JE, et al. In vivo bone formation in fracture repair induced by direct retroviral-based gene therapy with bone morphogenetic protein-4. Bone. 2003 Jun;32(6):591–601.

116. Huang Y-C, Kaigler D, Rice KG, Krebsbach PH, Mooney DJ. Combined angiogenic and osteogenic factor delivery enhances bone marrow stromal cell-driven bone regeneration. J Bone Miner Res. 2005 May;20(5):848–57.

117. Ishihara A, Shields KM, Litsky AS, Mattoon JS, Weisbrode SE, Bartlett JS, et al. Osteogenic gene regulation and relative acceleration of healing by adenoviral-mediated transfer of human BMP-2 or -6 in equine osteotomy and ostectomy models. J Orthop Res. 2008 Jun;26(6):764–71.

118. Breitbart AS, Grande DA, Mason JM, Barcia M, James T, Grant RT. Gene-enhanced tissue engineering: applications for bone healing using cultured periosteal cells transduced retrovirally with the BMP-7 gene. Ann Plast Surg. 1999 May;42(5):488–95.

119. Krebsbach PH, Gu K, Franceschi RT, Rutherford RB. Gene therapy-directed osteogenesis: BMP-7-transduced human fibroblasts form bone in vivo. Hum Gene Ther. 2000 May 20;11(8):1201–10.

120. Helm GA, Alden TD, Beres EJ, Hudson SB, Das S, Engh JA, et al. Use of bone morphogenetic protein-9 gene therapy to induce spinal arthrodesis in the rodent. J Neurosurg. 2000 Apr;92(2 Suppl):191–6.

121. Guo X, Zheng Q, Kulbatski I, Yuan Q, Yang S, Shao Z, et al. Bone regeneration with active angiogenesis by basic fibroblast growth factor gene transfected mesenchymal stem cells seeded on porous beta-TCP ceramic scaffolds. Biomed Mater. 2006 Sep;1(3):93–9.

122. Li R, Stewart DJ, von Schroeder HP, Mackinnon ES, Schemitsch EH. Effect of cell-based VEGF gene therapy on healing of a segmental bone defect. J Orthop Res. 2009 Jan;27(1):8– 14.

123. Li BC, Zhang JJ, Xu C, Zhang LC, Kang JY, Zhao H. Treatment of rabbit femoral defect by firearm with BMP-4 gene combined with TGF-beta1. J Trauma. 2009 Feb;66(2):450–6.

139

124. Shen FH, Visger JM, Balian G, Hurwitz SR, Diduch DR. Systemically administered mesenchymal stromal cells transduced with insulin-like growth factor-I localize to a fracture site and potentiate healing. J Orthop Trauma. 2002 Oct;16(9):651–9.

125. Chang P-C, Cirelli JA, Jin Q, Seol Y-J, Sugai JV, D’Silva NJ, et al. Adenovirus encoding human platelet-derived growth factor-B delivered to alveolar bone defects exhibits safety and biodistribution profiles favorable for clinical use. Hum Gene Ther. 2009 May;20(5):486–96.

126. Ito H, Koefoed M, Tiyapatanaputi P, Gromov K, Goater JJ, Carmouche J, et al. Remodeling of cortical bone allografts mediated by adherent rAAV-RANKL and VEGF gene therapy. Nat Med. 2005 Mar;11(3):291–7.

127. Zhao Z, Wang Z, Ge C, Krebsbach P, Franceschi RT. Healing cranial defects with AdRunx2-transduced marrow stromal cells. J Dent Res. 2007 Dec;86(12):1207–11.

128. Tu Q, Valverde P, Li S, Zhang J, Yang P, Chen J. Osterix overexpression in mesenchymal stem cells stimulates healing of critical-sized defects in murine calvarial bone. Tissue Eng. 2007 Oct;13(10):2431–40.

129. Viggeswarapu M, Boden SD, Liu Y, Hair GA, Louis-Ugbo J, Murakami H, et al. Adenoviral delivery of LIM mineralization protein-1 induces new-bone formation in vitro and in vivo. J Bone Joint Surg Am. 2001 Mar;83-A(3):364–76.

130. Lattanzi W, Parrilla C, Fetoni A, Logroscino G, Straface G, Pecorini G, et al. Ex vivo- transduced autologous skin fibroblasts expressing human Lim mineralization protein-3 efficiently form new bone in animal models. Gene Ther. 2008 Oct;15(19):1330–43.

131. Fang J, Zhu YY, Smiley E, Bonadio J, Rouleau JP, Goldstein SA, et al. Stimulation of new bone formation by direct transfer of osteogenic plasmid genes. Proc Natl Acad Sci USA. 1996 Jun 11;93(12):5753–8.

132. Koefoed M, Ito H, Gromov K, Reynolds DG, Awad HA, Rubery PT, et al. Biological effects of rAAV-caAlk2 coating on structural allograft healing. Mol Ther. 2005 Aug;12(2):212–8.

133. Huang W, Yang S, Shao J, Li Y-P. Signaling and transcriptional regulation in osteoblast commitment and differentiation. Front Biosci. 2007;12:3068–92.

134. Felin JE, Mayo JL, Loos TJ, Jensen JD, Sperry DK, Gaufin SL, et al. Nuclear variants of bone morphogenetic proteins. BMC Cell Biology. 2010 Mar 15;11(1):20.

135. Urist MR. Bone: formation by autoinduction. Science. 1965 Nov 12;150(3698):893–9.

136. Smith DM, Cooper GM, Mooney MP, Marra KG, Losee JE. Bone morphogenetic protein 2 therapy for craniofacial surgery. J Craniofac Surg. 2008 Sep;19(5):1244–59.

140

137. Katagiri T, Yamaguchi A, Komaki M, Abe E, Takahashi N, Ikeda T, et al. Bone morphogenetic protein-2 converts the differentiation pathway of C2C12 myoblasts into the osteoblast lineage. J Cell Biol. 1994 Dec;127(6 Pt 1):1755–66.

138. Nishio Y, Dong Y, Paris M, O’Keefe RJ, Schwarz EM, Drissi H. Runx2-mediated regulation of the zinc finger Osterix/Sp7 gene. Gene. 2006 May 10;372:62–70.

139. Yamaguchi A, Komori T, Suda T. Regulation of osteoblast differentiation mediated by bone morphogenetic proteins, hedgehogs, and Cbfa1. Endocr Rev. 2000 Aug;21(4):393– 411.

140. Xiao Y-T, Xiang L-X, Shao J-Z. Bone morphogenetic protein. Biochem Biophys Res Commun. 2007 Oct 26;362(3):550–3.

141. Hillger F, Herr G, Rudolph R, Schwarz E. Biophysical comparison of BMP-2, ProBMP-2, and the free pro-peptide reveals stabilization of the pro-peptide by the mature growth factor. J Biol Chem. 2005 Apr 15;280(15):14974–80.

142. Bragdon B, Moseychuk O, Saldanha S, King D, Julian J, Nohe A. Bone morphogenetic proteins: a critical review. Cell Signal. 2011 Apr;23(4):609–20.

143. Wozney JM, Rosen V, Celeste AJ, Mitsock LM, Whitters MJ, Kriz RW, et al. Novel regulators of bone formation: molecular clones and activities. Science. 1988 Dec 16;242(4885):1528–34.

144. Canalis E, Economides AN, Gazzerro E. Bone morphogenetic proteins, their antagonists, and the skeleton. Endocr Rev. 2003 Apr;24(2):218–35.

145. Seidah NG. The Proprotein Convertases, 20 Years Later. In: Mbikay M, Seidah NG, editors. Proprotein Convertases [Internet]. Totowa, NJ: Humana Press; 2011 [cited 2013 Apr 1]. p. 23–57. Available from: http://link.springer.com.myaccess.library.utoronto.ca/content/pdf/10.1007%2F978-1- 61779-204-5_3

146. Seidah NG. What lies ahead for the proprotein convertases? Ann N Y Acad Sci. 2011 Mar;1220:149–61.

147. Takahashi S, Hatsuzawa K, Watanabe T, Murakami K, Nakayama K. Sequence requirements for endoproteolytic processing of precursor proteins by furin: transfection and in vitro experiments. J Biochem. 1994 Jul;116(1):47–52.

148. Zhou J-JA. Enhancing Production of Recombinant BMP-2 in Mammalian Cell Culture Systems by Inhibition of Pro-protein Cleavage using 9DR Peptides. 2008 Jul 30 [cited 2012 Dec 3]; Available from: https://tspace.library.utoronto.ca/handle/1807/11175

141

149. Nohe A, Hassel S, Ehrlich M, Neubauer F, Sebald W, Henis YI, et al. The mode of bone morphogenetic protein (BMP) receptor oligomerization determines different BMP-2 signaling pathways. J Biol Chem. 2002 Feb 15;277(7):5330–8.

150. Hassel S, Schmitt S, Hartung A, Roth M, Nohe A, Petersen N, et al. Initiation of Smad- dependent and Smad-independent signaling via distinct BMP-receptor complexes. J Bone Joint Surg Am. 2003;85-A Suppl 3:44–51.

151. Nohe A, Keating E, Knaus P, Petersen NO. Signal transduction of bone morphogenetic protein receptors. Cell Signal. 2004 Mar;16(3):291–9.

152. Groppe J, Greenwald J, Wiater E, Rodriguez-Leon J, Economides AN, Kwiatkowski W, et al. Structural basis of BMP signalling inhibition by the cystine knot protein Noggin. Nature. 2002 Dec 12;420(6916):636–42.

153. Simic P, Vukicevic S. Bone morphogenetic proteins: from developmental signals to tissue regeneration. Conference on Bone Morphogenetic Proteins. EMBO Rep. 2007 Apr;8(4):327–31.

154. Ying S-X, Hussain ZJ, Zhang YE. Smurf1 facilitates myogenic differentiation and antagonizes the bone morphogenetic protein-2-induced osteoblast conversion by targeting Smad5 for degradation. J Biol Chem. 2003 Oct 3;278(40):39029–36.

155. Zhu H, Kavsak P, Abdollah S, Wrana JL, Thomsen GH. A SMAD ubiquitin ligase targets the BMP pathway and affects embryonic pattern formation. Nature. 1999 Aug 12;400(6745):687–93.

156. Schmitt JM, Hwang K, Winn SR, Hollinger JO. Bone morphogenetic proteins: an update on basic biology and clinical relevance. J Orthop Res. 1999 Mar;17(2):269–78.

157. Satija NK, Singh VK, Verma YK, Gupta P, Sharma S, Afrin F, et al. Mesenchymal stem cell- based therapy: a new paradigm in regenerative medicine. J Cell Mol Med. 2009 Dec;13(11- 12):4385–402.

158. Aslan H, Zilberman Y, Arbeli V, Sheyn D, Matan Y, Liebergall M, et al. Nucleofection- based ex vivo nonviral gene delivery to human stem cells as a platform for tissue regeneration. Tissue Eng. 2006 Apr;12(4):877–89.

159. Park J, Ries J, Gelse K, Kloss F, von der Mark K, Wiltfang J, et al. Bone regeneration in critical size defects by cell-mediated BMP-2 gene transfer: a comparison of adenoviral vectors and liposomes. Gene Ther. 2003 Jul;10(13):1089–98.

160. Miyazaki M, Sugiyama O, Zou J, Yoon SH, Wei F, Morishita Y, et al. Comparison of lentiviral and adenoviral gene therapy for spinal fusion in rats. Spine. 2008 Jun 1;33(13):1410–7.

142

161. Blum JS, Barry MA, Mikos AG, Jansen JA. In vivo evaluation of gene therapy vectors in ex vivo-derived marrow stromal cells for bone regeneration in a rat critical-size calvarial defect model. Hum Gene Ther. 2003 Dec 10;14(18):1689–701.

162. Peterson B, Iglesias R, Zhang J, Wang JC, Lieberman JR. Genetically modified human derived bone marrow cells for posterolateral lumbar spine fusion in athymic rats: beyond conventional autologous bone grafting. Spine. 2005 Feb 1;30(3):283–289; discussion 289– 290.

163. Gugala Z, Olmsted-Davis EA, Gannon FH, Lindsey RW, Davis AR. Osteoinduction by ex vivo adenovirus-mediated BMP2 delivery is independent of cell type. Gene Ther. 2003 Aug;10(16):1289–96.

164. Chang SC-N, Lin T-M, Chung H-Y, Chen PK-T, Lin F-H, Lou J, et al. Large-scale bicortical skull bone regeneration using ex vivo replication-defective adenoviral-mediated bone morphogenetic protein-2 gene-transferred bone marrow stromal cells and composite biomaterials. Neurosurgery. 2009 Dec;65(6 Suppl):75–81; discussion 81–83.

165. Dai KR, Xu XL, Tang TT, Zhu ZA, Yu CF, Lou JR, et al. Repairing of goat tibial bone defects with BMP-2 gene-modified tissue-engineered bone. Calcif Tissue Int. 2005 Jul;77(1):55–61.

166. Kagiwada H, Yashiki T, Ohshima A, Tadokoro M, Nagaya N, Ohgushi H. Human mesenchymal stem cells as a stable source of VEGF-producing cells. J Tissue Eng Regen Med. 2008 Jun;2(4):184–9.

167. Delcombel R, Janssen L, Vassy R, Gammons M, Haddad O, Richard B, et al. New prospects in the roles of the C-terminal domains of VEGF-A and their cooperation for ligand binding, cellular signaling and vessels formation. Angiogenesis. 2012 Dec 20;

168. Ferrara N, Henzel WJ. Pituitary follicular cells secrete a novel heparin-binding growth factor specific for vascular endothelial cells. Biochem Biophys Res Commun. 1989 Jun 15;161(2):851–8.

169. Bluteau G, Julien M, Magne D, Mallein-Gerin F, Weiss P, Daculsi G, et al. VEGF and VEGF receptors are differentially expressed in chondrocytes. Bone. 2007 Mar;40(3):568–76.

170. Beamer B, Hettrich C, Lane J. Vascular Endothelial Growth Factor: An Essential Component of Angiogenesis and Fracture Healing. HSS J [Internet]. 2009 Sep 9 [cited 2010 Feb 5]; Available from: http://www.ncbi.nlm.nih.gov/pubmed/19763695

171. Deckers MML, van Bezooijen RL, van der Horst G, Hoogendam J, van Der Bent C, Papapoulos SE, et al. Bone morphogenetic proteins stimulate angiogenesis through osteoblast-derived vascular endothelial growth factor A. Endocrinology. 2002 Apr;143(4):1545–53.

143

172. Zelzer E, Glotzer DJ, Hartmann C, Thomas D, Fukai N, Soker S, et al. Tissue specific regulation of VEGF expression during bone development requires Cbfa1/Runx2. Mech Dev. 2001 Aug;106(1-2):97–106.

173. Ferrara N, Gerber H-P, LeCouter J. The biology of VEGF and its receptors. Nat Med. 2003 Jun;9(6):669–76.

174. Dai J, Rabie ABM. VEGF: an essential mediator of both angiogenesis and endochondral ossification. J Dent Res. 2007 Oct;86(10):937–50.

175. Houck KA, Ferrara N, Winer J, Cachianes G, Li B, Leung DW. The vascular endothelial growth factor family: identification of a fourth molecular species and characterization of alternative splicing of RNA. Mol Endocrinol. 1991 Dec;5(12):1806–14.

176. Claffey KP, Senger DR, Spiegelman BM. Structural requirements for dimerization, glycosylation, secretion, and biological function of VPF/VEGF. Biochim Biophys Acta. 1995 Jan 5;1246(1):1–9.

177. Lee S, Jilani SM, Nikolova GV, Carpizo D, Iruela-Arispe ML. Processing of VEGF-A by matrix metalloproteinases regulates bioavailability and vascular patterning in tumors. J Cell Biol. 2005 May 23;169(4):681–91.

178. Cross MJ, Dixelius J, Matsumoto T, Claesson-Welsh L. VEGF-receptor signal transduction. Trends Biochem Sci. 2003 Sep;28(9):488–94.

179. Parker MW, Xu P, Li X, Vander Kooi CW. Structural basis for selective vascular endothelial growth factor-A (VEGF-A) binding to neuropilin-1. J Biol Chem. 2012 Mar 30;287(14):11082–9.

180. Houck KA, Leung DW, Rowland AM, Winer J, Ferrara N. Dual regulation of vascular endothelial growth factor bioavailability by genetic and proteolytic mechanisms. J Biol Chem. 1992 Dec 25;267(36):26031–7.

181. Rini BI, Small EJ. Biology and clinical development of vascular endothelial growth factor- targeted therapy in renal cell carcinoma. J Clin Oncol. 2005 Feb 10;23(5):1028–43.

182. Whitaker GB, Limberg BJ, Rosenbaum JS. Vascular endothelial growth factor receptor-2 and neuropilin-1 form a receptor complex that is responsible for the differential signaling potency of VEGF(165) and VEGF(121). J Biol Chem. 2001 Jul 6;276(27):25520–31.

183. Karumanchi SA, Bdolah Y. Hypoxia and sFlt-1 in Preeclampsia: The “Chicken-and-Egg” Question. Endocrinology. 2004 Nov 1;145(11):4835–7.

184. Colnot CI, Helms JA. A molecular analysis of matrix remodeling and angiogenesis during long bone development. Mech Dev. 2001 Feb;100(2):245–50.

144

185. Gerber HP, Vu TH, Ryan AM, Kowalski J, Werb Z, Ferrara N. VEGF couples hypertrophic cartilage remodeling, ossification and angiogenesis during endochondral bone formation. Nat Med. 1999 Jun;5(6):623–8.

186. Mishima Y, Lotz M. Chemotaxis of human articular chondrocytes and mesenchymal stem cells. J Orthop Res. 2008 Oct;26(10):1407–12.

187. Zelzer E, Olsen BR. Multiple roles of vascular endothelial growth factor (VEGF) in skeletal development, growth, and repair. Curr Top Dev Biol. 2005;65:169–87.

188. Furumatsu T, Shen ZN, Kawai A, Nishida K, Manabe H, Oohashi T, et al. Vascular endothelial growth factor principally acts as the main angiogenic factor in the early stage of human osteoblastogenesis. J Biochem. 2003 May;133(5):633–9.

189. Samee M, Kasugai S, Kondo H, Ohya K, Shimokawa H, Kuroda S. Bone morphogenetic protein-2 (BMP-2) and vascular endothelial growth factor (VEGF) transfection to human periosteal cells enhances osteoblast differentiation and bone formation. J Pharmacol Sci. 2008 Sep;108(1):18–31.

190. Peng H, Usas A, Olshanski A, Ho AM, Gearhart B, Cooper GM, et al. VEGF Improves, Whereas sFlt1 Inhibits, BMP2-Induced Bone Formation and Bone Healing Through Modulation of Angiogenesis. Journal of Bone and Mineral Research. 2005;20(11):2017–27.

191. Liu TM, Lee EH. Transcriptional Regulatory Cascades in Runx2-Dependent Bone Development. Tissue Eng Part B Rev. 2012 Nov 14;

192. Ducy P, Zhang R, Geoffroy V, Ridall AL, Karsenty G. Osf2/Cbfa1: a transcriptional activator of osteoblast differentiation. Cell. 1997 May 30;89(5):747–54.

193. Karsenty G, Ducy P, Starbuck M, Priemel M, Shen J, Geoffroy V, et al. Cbfa1 as a regulator of osteoblast differentiation and function. Bone. 1999 Jul;25(1):107–8.

194. Otto F, Thornell AP, Crompton T, Denzel A, Gilmour KC, Rosewell IR, et al. Cbfa1, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell. 1997 May 30;89(5):765–71.

195. Schroeder TM, Jensen ED, Westendorf JJ. Runx2: a master organizer of gene transcription in developing and maturing osteoblasts. Birth Defects Res C Embryo Today. 2005 Sep;75(3):213–25.

196. Inada M, Yasui T, Nomura S, Miyake S, Deguchi K, Himeno M, et al. Maturational disturbance of chondrocytes in Cbfa1-deficient mice. Dev Dyn. 1999 Apr;214(4):279–90.

197. Stock M, Otto F. Control of RUNX2 isoform expression: the role of promoters and enhancers. J Cell Biochem. 2005 Jun 1;95(3):506–17.

145

198. Ducy P, Karsenty G. Two distinct osteoblast-specific cis-acting elements control expression of a mouse osteocalcin gene. Mol Cell Biol. 1995 Apr;15(4):1858–69.

199. Li Y-L, Xiao Z-S. Advances in Runx2 regulation and its isoforms. Med Hypotheses. 2007;68(1):169–75.

200. Mundlos S, Otto F, Mundlos C, Mulliken JB, Aylsworth AS, Albright S, et al. Mutations involving the transcription factor CBFA1 cause cleidocranial dysplasia. Cell. 1997 May 30;89(5):773–9.

201. Ogawa E, Maruyama M, Kagoshima H, Inuzuka M, Lu J, Satake M, et al. PEBP2/PEA2 represents a family of transcription factors homologous to the products of the Drosophila runt gene and the human AML1 gene. Proc Natl Acad Sci USA. 1993 Jul 15;90(14):6859–63.

202. Harada H, Tagashira S, Fujiwara M, Ogawa S, Katsumata T, Yamaguchi A, et al. Cbfa1 isoforms exert functional differences in osteoblast differentiation. J Biol Chem. 1999 Mar 12;274(11):6972–8.

203. Sudhakar S, Li Y, Katz MS, Elango N. Translational regulation is a control point in RUNX2/Cbfa1 gene expression. Biochem Biophys Res Commun. 2001 Nov 30;289(2):616– 22.

204. Banerjee C, Javed A, Choi JY, Green J, Rosen V, van Wijnen AJ, et al. Differential regulation of the two principal Runx2/Cbfa1 n-terminal isoforms in response to bone morphogenetic protein-2 during development of the osteoblast phenotype. Endocrinology. 2001 Sep;142(9):4026–39.

205. Komori T, Yagi H, Nomura S, Yamaguchi A, Sasaki K, Deguchi K, et al. Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell. 1997 May 30;89(5):755–64.

206. Liu W, Toyosawa S, Furuichi T, Kanatani N, Yoshida C, Liu Y, et al. Overexpression of Cbfa1 in osteoblasts inhibits osteoblast maturation and causes osteopenia with multiple fractures. J Cell Biol. 2001 Oct 1;155(1):157–66.

207. Quack I, Vonderstrass B, Stock M, Aylsworth A, Becker A, Brueton L, et al. Mutation Analysis of Core Binding Factor A1 in Patients with Cleidocranial Dysplasia. The American Journal of Human Genetics. 1999 Nov;65(5):1268–78.

208. Lian JB, Stein GS, Javed A, van Wijnen AJ, Stein JL, Montecino M, et al. Networks and hubs for the transcriptional control of osteoblastogenesis. Rev Endocr Metab Disord. 2006 Jun;7(1-2):1–16.

209. Sato M, Morii E, Komori T, Kawahata H, Sugimoto M, Terai K, et al. Transcriptional regulation of osteopontin gene in vivo by PEBP2alphaA/CBFA1 and ETS1 in the skeletal tissues. Oncogene. 1998 Sep 24;17(12):1517–25.

146

210. Benson MD, Aubin JE, Xiao G, Thomas PE, Franceschi RT. Cloning of a 2.5 kb murine bone sialoprotein promoter fragment and functional analysis of putative Osf2 binding sites. J Bone Miner Res. 1999 Mar;14(3):396–405.

211. Kern B, Shen J, Starbuck M, Karsenty G. Cbfa1 contributes to the osteoblast-specific expression of type I collagen genes. J Biol Chem. 2001 Mar 9;276(10):7101–7.

212. Jiménez MJ, Balbín M, Alvarez J, Komori T, Bianco P, Holmbeck K, et al. A regulatory cascade involving retinoic acid, Cbfa1, and matrix metalloproteinases is coupled to the development of a process of perichondrial invasion and osteogenic differentiation during bone formation. J Cell Biol. 2001 Dec 24;155(7):1333–44.

213. Thirunavukkarasu K, Halladay DL, Miles RR, Yang X, Galvin RJ, Chandrasekhar S, et al. The osteoblast-specific transcription factor Cbfa1 contributes to the expression of osteoprotegerin, a potent inhibitor of osteoclast differentiation and function. J Biol Chem. 2000 Aug 18;275(33):25163–72.

214. Komori T. Runx2, a multifunctional transcription factor in skeletal development. J Cell Biochem. 2002;87(1):1–8.

215. Drissi H, Pouliot A, Koolloos C, Stein JL, Lian JB, Stein GS, et al. 1,25-(OH)2-vitamin D3 suppresses the bone-related Runx2/Cbfa1 gene promoter. Exp Cell Res. 2002 Apr 1;274(2):323–33.

216. Gilbert L, He X, Farmer P, Rubin J, Drissi H, van Wijnen AJ, et al. Expression of the osteoblast differentiation factor RUNX2 (Cbfa1/AML3/Pebp2alpha A) is inhibited by tumor necrosis factor-alpha. J Biol Chem. 2002 Jan 25;277(4):2695–701.

217. Prince M, Banerjee C, Javed A, Green J, Lian JB, Stein GS, et al. Expression and regulation of Runx2/Cbfa1 and osteoblast phenotypic markers during the growth and differentiation of human osteoblasts. J Cell Biochem. 2001;80(3):424–40.

218. Lee M-H, Kim Y-J, Yoon W-J, Kim J-I, Kim B-G, Hwang Y-S, et al. Dlx5 specifically regulates Runx2 type II expression by binding to homeodomain-response elements in the Runx2 distal promoter. J Biol Chem. 2005 Oct 21;280(42):35579–87.

219. Dobreva G, Chahrour M, Dautzenberg M, Chirivella L, Kanzler B, Fariñas I, et al. SATB2 is a multifunctional determinant of craniofacial patterning and osteoblast differentiation. Cell. 2006 Jun 2;125(5):971–86.

220. Hassan MQ, Tare RS, Lee SH, Mandeville M, Morasso MI, Javed A, et al. BMP2 commitment to the osteogenic lineage involves activation of Runx2 by DLX3 and a homeodomain transcriptional network. J Biol Chem. 2006 Dec 29;281(52):40515–26.

221. Isenmann S, Arthur A, Zannettino ACW, Turner JL, Shi S, Glackin CA, et al. TWIST family of basic helix-loop-helix transcription factors mediate human mesenchymal stem cell growth and commitment. Stem Cells. 2009 Oct;27(10):2457–68. 147

222. Kim S, Koga T, Isobe M, Kern BE, Yokochi T, Chin YE, et al. Stat1 functions as a cytoplasmic attenuator of Runx2 in the transcriptional program of osteoblast differentiation. Genes Dev. 2003 Aug 15;17(16):1979–91.

223. Zaidi SK, Sullivan AJ, Medina R, Ito Y, van Wijnen AJ, Stein JL, et al. Tyrosine phosphorylation controls Runx2-mediated subnuclear targeting of YAP to repress transcription. EMBO J. 2004 Feb 25;23(4):790–9.

224. Yoshida CA, Furuichi T, Fujita T, Fukuyama R, Kanatani N, Kobayashi S, et al. Core- binding factor beta interacts with Runx2 and is required for skeletal development. Nat Genet. 2002 Dec;32(4):633–8.

225. Hong J-H, Hwang ES, McManus MT, Amsterdam A, Tian Y, Kalmukova R, et al. TAZ, a transcriptional modulator of mesenchymal stem cell differentiation. Science. 2005 Aug 12;309(5737):1074–8.

226. Pelletier N, Champagne N, Stifani S, Yang X-J. MOZ and MORF histone acetyltransferases interact with the Runt-domain transcription factor Runx2. Oncogene. 2002 Apr 18;21(17):2729–40.

227. Ducy P, Starbuck M, Priemel M, Shen J, Pinero G, Geoffroy V, et al. A Cbfa1-dependent genetic pathway controls bone formation beyond embryonic development. Genes Dev. 1999 Apr 15;13(8):1025–36.

228. Drissi H, Luc Q, Shakoori R, Chuva De Sousa Lopes S, Choi JY, Terry A, et al. Transcriptional autoregulation of the bone related CBFA1/RUNX2 gene. J Cell Physiol. 2000 Sep;184(3):341–50.

229. Zheng H, Guo Z, Ma Q, Jia H, Dang G. Cbfa1/osf2 transduced bone marrow stromal cells facilitate bone formation in vitro and in vivo. Calcif Tissue Int. 2004 Feb;74(2):194–203.

230. Zhang X, Yang M, Lin L, Chen P, Ma KT, Zhou CY, et al. Runx2 overexpression enhances osteoblastic differentiation and mineralization in adipose--derived stem cells in vitro and in vivo. Calcif Tissue Int. 2006 Sep;79(3):169–78.

231. Byers BA, Pavlath GK, Murphy TJ, Karsenty G, García AJ. Cell-type-dependent upregulation of in vitro mineralization after overexpression of the osteoblast-specific transcription factor Runx2/Cbfal. J Bone Miner Res. 2002 Nov;17(11):1931–44.

232. Byers BA, García AJ. Exogenous Runx2 expression enhances in vitro osteoblastic differentiation and mineralization in primary bone marrow stromal cells. Tissue Eng. 2004 Dec;10(11-12):1623–32.

233. Gersbach CA, Byers BA, Pavlath GK, García AJ. Runx2/Cbfa1 stimulates transdifferentiation of primary skeletal myoblasts into a mineralizing osteoblastic phenotype. Exp Cell Res. 2004 Nov 1;300(2):406–17.

148

234. Gersbach CA, Le Doux JM, Guldberg RE, García AJ. Inducible regulation of Runx2- stimulated osteogenesis. Gene Ther. 2006 Jun;13(11):873–82.

235. Phillips JE, Guldberg RE, García AJ. Dermal fibroblasts genetically modified to express Runx2/Cbfa1 as a mineralizing cell source for bone tissue engineering. Tissue Eng. 2007 Aug;13(8):2029–40.

236. Hirata K, Tsukazaki T, Kadowaki A, Furukawa K, Shibata Y, Moriishi T, et al. Transplantation of skin fibroblasts expressing BMP-2 promotes bone repair more effectively than those expressing Runx2. Bone. 2003 May;32(5):502–12.

237. Yang S, Wei D, Wang D, Phimphilai M, Krebsbach PH, Franceschi RT. In vitro and in vivo synergistic interactions between the Runx2/Cbfa1 transcription factor and bone morphogenetic protein-2 in stimulating osteoblast differentiation. J Bone Miner Res. 2003 Apr;18(4):705–15.

238. Lee S-J, Kang S-W, Do H-J, Han I, Shin DA, Kim J-H, et al. Enhancement of bone regeneration by gene delivery of BMP2/Runx2 bicistronic vector into adipose-derived stromal cells. Biomaterials. 2010 Jul;31(21):5652–9.

239. Long F. Building strong bones: molecular regulation of the osteoblast lineage. Nat Rev Mol Cell Biol. 2012 Jan;13(1):27–38.

240. Nakashima K, Zhou X, Kunkel G, Zhang Z, Deng JM, Behringer RR, et al. The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell. 2002 Jan 11;108(1):17–29.

241. Bouwman P, Philipsen S. Regulation of the activity of Sp1-related transcription factors. Mol Cell Endocrinol. 2002 Sep 30;195(1-2):27–38.

242. Milona M, Gough JE, Edgar AJ. Expression of alternatively spliced isoforms of human Sp7 in osteoblast-like cells. BMC Genomics. 2003 Nov 7;4:43.

243. Gao Y, Jheon A, Nourkeyhani H, Kobayashi H, Ganss B. Molecular cloning, structure, expression, and chromosomal localization of the human Osterix (SP7) gene. Gene. 2004 Oct 27;341:101–10.

244. Hong SHH, Lu X, Nanes MS, Mitchell J. Regulation of osterix (Osx, Sp7) and the Osx promoter by parathyroid hormone in osteoblasts. J Mol Endocrinol. 2009 Nov;43(5):197– 207.

245. Nakashima K, de Crombrugghe B. Transcriptional mechanisms in osteoblast differentiation and bone formation. Trends Genet. 2003 Aug;19(8):458–66.

246. Kim Y-J, Kim H-N, Park E-K, Lee B-H, Ryoo H-M, Kim S-Y, et al. The bone-related Zn finger transcription factor Osterix promotes proliferation of mesenchymal cells. Gene. 2006 Jan 17;366(1):145–51. 149

247. Wang B, Huang S, Pan L, Jia S. Enhancement of bone formation by genetically engineered human umbilical cord-derived mesenchymal stem cells expressing osterix. Oral Surg Oral Med Oral Pathol Oral Radiol. 2012 Jul 19;

248. Kaback LA, Soung DY, Naik A, Smith N, Schwarz EM, O’Keefe RJ, et al. Osterix/Sp7 regulates mesenchymal stem cell mediated endochondral ossification. J Cell Physiol. 2008 Jan;214(1):173–82.

249. Zhang C, Cho K, Huang Y, Lyons JP, Zhou X, Sinha K, et al. Inhibition of Wnt signaling by the osteoblast-specific transcription factor Osterix. Proc Natl Acad Sci USA. 2008 May 13;105(19):6936–41.

250. Cheng S-L, Shao J-S, Charlton-Kachigian N, Loewy AP, Towler DA. MSX2 promotes osteogenesis and suppresses adipogenic differentiation of multipotent mesenchymal progenitors. J Biol Chem. 2003 Nov 14;278(46):45969–77.

251. Lee M-H, Kwon T-G, Park H-S, Wozney JM, Ryoo H-M. BMP-2-induced Osterix expression is mediated by Dlx5 but is independent of Runx2. Biochem Biophys Res Commun. 2003 Sep 26;309(3):689–94.

252. Celil AB, Campbell PG. BMP-2 and insulin-like growth factor-I mediate Osterix (Osx) expression in human mesenchymal stem cells via the MAPK and protein kinase D signaling pathways. J Biol Chem. 2005 Sep 9;280(36):31353–9.

253. Xing W, Singgih A, Kapoor A, Alarcon CM, Baylink DJ, Mohan S. Nuclear factor-E2- related factor-1 mediates ascorbic acid induction of osterix expression via interaction with antioxidant-responsive element in bone cells. J Biol Chem. 2007 Jul 27;282(30):22052–61.

254. Maehata Y, Takamizawa S, Ozawa S, Kato Y, Sato S, Kubota E, et al. Both direct and collagen-mediated signals are required for active vitamin D3-elicited differentiation of human osteoblastic cells: roles of osterix, an osteoblast-related transcription factor. Matrix Biol. 2006 Jan;25(1):47–58.

255. Nishimura R, Hata K, Matsubara T, Wakabayashi M, Yoneda T. Regulation of bone and cartilage development by network between BMP signalling and transcription factors. J Biochem. 2012 Mar;151(3):247–54.

256. Lu X, Gilbert L, He X, Rubin J, Nanes MS. Transcriptional regulation of the osterix (Osx, Sp7) promoter by tumor necrosis factor identifies disparate effects of mitogen-activated protein kinase and NF kappa B pathways. J Biol Chem. 2006 Mar 10;281(10):6297–306.

257. Zambetti GP, Horwitz EM, Schipani E. Skeletons in the p53 tumor suppressor closet: genetic evidence that p53 blocks bone differentiation and development. J Cell Biol. 2006 Mar 13;172(6):795–7.

150

258. Zhu J, Shimizu E, Zhang X, Partridge NC, Qin L. EGFR signaling suppresses osteoblast differentiation and inhibits expression of master osteoblastic transcription factors Runx2 and Osterix. J Cell Biochem. 2011 Jul;112(7):1749–60.

259. Yoshida CA, Komori H, Maruyama Z, Miyazaki T, Kawasaki K, Furuichi T, et al. SP7 inhibits osteoblast differentiation at a late stage in mice. PLoS ONE. 2012;7(3):e32364.

260. Tu Q, Valverde P, Chen J. Osterix enhances proliferation and osteogenic potential of bone marrow stromal cells. Biochem Biophys Res Commun. 2006 Mar 24;341(4):1257–65.

261. Lai Q-G, Yuan K-F, Xu X, Li D, Li G-J, Wei F-L, et al. Transcription factor osterix modified bone marrow mesenchymal stem cells enhance callus formation during distraction osteogenesis. Oral Surg Oral Med Oral Pathol Oral Radiol Endod. 2011 Apr;111(4):412–9.

262. Vaes B, Van’t Hof W, Deans R, Pinxteren J. Application of MultiStem(®) Allogeneic Cells for Immunomodulatory Therapy: Clinical Progress and Pre-Clinical Challenges in Prophylaxis for Graft Versus Host Disease. Front Immunol. 2012;3:345.

263. Byers BA, García AJ. Exogenous Runx2 expression enhances in vitro osteoblastic differentiation and mineralization in primary bone marrow stromal cells. Tissue Eng. 2004 Dec;10(11-12):1623–32.

264. Zheng H, Guo Z, Ma Q, Jia H, Dang G. Cbfa1/osf2 transduced bone marrow stromal cells facilitate bone formation in vitro and in vivo. Calcif Tissue Int. 2004 Feb;74(2):194–203.

265. Lee J-S, Lee J-M, Im G-I. Electroporation-mediated transfer of Runx2 and Osterix genes to enhance osteogenesis of adipose stem cells. Biomaterials. 2011 Jan;32(3):760–8.

266. NanoDrop User Manual [Internet]. Available from: http://memphys.dk/sites/default/files/files/basic_page/%3Cem%3EEdit%20Basic%20page %3C/em%3E%20UV-Vis%20spectroscopy/nd-1000-v3.7-users-manual-8.5x11.pdf

267. Stratagene. AdEasyTM XL Adenoviral Vector System INSTRUCTION MANUAL [Internet]. Available from: http://www.genomics.agilent.com/files/Manual/240010.pdf

268. Luo J, Deng Z-L, Luo X, Tang N, Song W-X, Chen J, et al. A protocol for rapid generation of recombinant adenoviruses using the AdEasy system. Nat Protoc. 2007;2(5):1236–47.

269. Wahlfors J, Loimas S, Pasanen T, Hakkarainen T. Green fluorescent protein (GFP) fusion constructs in gene therapy research. Histochemistry and Cell Biology. 2001 Jan 22;115(1):59–65.

270. Galipeau J, Li H, Paquin A, Sicilia F, Karpati G, Nalbantoglu J. Vesicular stomatitis virus G pseudotyped retrovector mediates effective in vivo suicide gene delivery in experimental brain cancer. Cancer Res. 1999 May 15;59(10):2384–94.

151

271. Tomasz M, Lipman R, Chowdary D, Pawlak J, Verdine GL, Nakanishi K. Isolation and structure of a covalent cross-link adduct between mitomycin C and DNA. Science. 1987 Mar 6;235(4793):1204–8.

272. Peel SAF, Hu ZM, Clokie CML. In search of the ideal bone morphogenetic protein delivery system: in vitro studies on demineralized bone matrix, purified, and recombinant bone morphogenetic protein. J Craniofac Surg. 2003 May;14(3):284–91.

273. Pfaffl MW. A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 2001 May 1;29(9):e45.

274. Leng P, Ding C, Zhang H, Wang Y. Reconstruct large osteochondral defects of the knee with hIGF-1 gene enhanced Mosaicplasty. The Knee. 2012 Dec;19(6):804–11.

275. Gheisari Y, Soleimani M, Azadmanesh K, Zeinali S. Multipotent mesenchymal stromal cells: optimization and comparison of five cationic polymer-based gene delivery methods. Cytotherapy. 2008;10(8):815–23.

276. Madeira C, Mendes RD, Ribeiro SC, Boura JS, Aires-Barros MR, da Silva CL, et al. Nonviral Gene Delivery to Mesenchymal Stem Cells Using Cationic Liposomes for Gene and Cell Therapy. BioMed Research International [Internet]. 2010 Jun 24 [cited 2013 Jun 27];2010. Available from: http://www.hindawi.com/journals/bmri/2010/735349/abs/

277. Clements BA, Incani V, Kucharski C, Lavasanifar A, Ritchie B, Uludağ H. A comparative evaluation of poly-L-lysine-palmitic acid and Lipofectamine 2000 for plasmid delivery to bone marrow stromal cells. Biomaterials. 2007 Nov;28(31):4693–704.

278. Baksh D, Yao R, Tuan RS. Comparison of proliferative and multilineage differentiation potential of human mesenchymal stem cells derived from umbilical cord and bone marrow. Stem Cells. 2007 Jun;25(6):1384–92.

279. Kim A. Development and Characterization of a Controlled Expression System for Osteogenic Genes. [Toronto]: University of Toronto; 2011.

280. Gysin R, Wergedal JE, Sheng MH-C, Kasukawa Y, Miyakoshi N, Chen S-T, et al. Ex vivo gene therapy with stromal cells transduced with a retroviral vector containing the BMP4 gene completely heals critical size calvarial defect in rats. Gene Ther. 2002 Aug;9(15):991– 9.

281. Bergelson JM, Cunningham JA, Droguett G, Kurt-Jones EA, Krithivas A, Hong JS, et al. Isolation of a common receptor for Coxsackie B viruses and adenoviruses 2 and 5. Science. 1997 Feb 28;275(5304):1320–3.

282. Liu G, Shu C, Cui L, Liu W, Cao Y. Tissue-engineered bone formation with cryopreserved human bone marrow mesenchymal stem cells. Cryobiology. 2008 Jun;56(3):209–15.

152

283. Rust PA, Costas C, Cannon SR, Briggs TWR, Blunn GW. Characterisation of cryopreserved cells freshly isolated from human bone marrow. Cryo Letters. 2006 Feb;27(1):17–28.

284. Liu G, Zhou H, Li Y, Li G, Cui L, Liu W, et al. Evaluation of the viability and osteogenic differentiation of cryopreserved human adipose-derived stem cells. Cryobiology. 2008 Aug;57(1):18–24.

285. Kotobuki N, Hirose M, Machida H, Katou Y, Muraki K, Takakura Y, et al. Viability and osteogenic potential of cryopreserved human bone marrow-derived mesenchymal cells. Tissue Eng. 2005 Jun;11(5-6):663–73.

286. McGinley L, McMahon J, Strappe P, Barry F, Murphy M, O’Toole D, et al. Lentiviral vector mediated modification of mesenchymal stem cells & enhanced survival in an in vitro model of ischaemia. Stem Cell Res Ther. 2011;2(2):12.

287. Mumaw J, Jordan ET, Sonnet C, Olabisi RM, Olmsted-Davis EA, Davis AR, et al. Rapid Heterotrophic Ossification with Cryopreserved Poly(ethylene glycol-) Microencapsulated BMP2-Expressing MSCs. Int J Biomater [Internet]. 2012 [cited 2013 Jul 10];2012. Available from: http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3296315/

288. Gao D, Critser JK. Mechanisms of cryoinjury in living cells. ILAR J. 2000;41(4):187–96.

289. Haack-Sørensen M, Kastrup J. Cryopreservation and revival of mesenchymal stromal cells. Methods Mol Biol. 2011;698:161–74.

290. Yue B, Lu B, Dai KR, Zhang XL, Yu CF, Lou JR, et al. BMP2 gene therapy on the repair of bone defects of aged rats. Calcif Tissue Int. 2005 Dec;77(6):395–403.

291. Wang Y, Mostafa NZ, Hsu CYM, Rose L, Kucharki C, Yan J, et al. Modification of human BMSC with nanoparticles of polymeric biomaterials and plasmid DNA for BMP-2 secretion. J Surg Res. 2013 Jul;183(1):8–17.

292. Leighton M, Kadler KE. Paired basic/Furin-like proprotein convertase cleavage of Pro- BMP-1 in the trans-Golgi network. J Biol Chem. 2003 May 16;278(20):18478–84.

293. Byun H, Halani N, Gou Y, Nash AK, Lozano MM, Dudley JP. Requirements for mouse mammary tumor virus Rem signal peptide processing and function. J Virol. 2012 Jan;86(1):214–25.

294. Pulkki MM, Myllymaa S, Pasternack A, Lun S, Ludlow H, Al-Qahtani A, et al. The bioactivity of human bone morphogenetic protein-15 is sensitive to C-terminal modification: characterization of the purified untagged processed mature region. Mol Cell Endocrinol. 2011 Jan 30;332(1-2):106–15.

153

295. Mason AB, He Q-Y, Halbrooks PJ, Everse SJ, Gumerov DR, Kaltashov IA, et al. Differential effect of a his tag at the N- and C-termini: functional studies with recombinant human serum transferrin. Biochemistry. 2002 Jul 30;41(30):9448–54.

296. Christensen B, Kläning E, Nielsen MS, Andersen MH, Sørensen ES. C-terminal modification of osteopontin inhibits interaction with the αVβ3-integrin. J Biol Chem. 2012 Feb 3;287(6):3788–97.

297. Wen Y, Jiang B, Cui J, Li G, Yu M, Wang F, et al. Superior osteogenic capacity of different mesenchymal stem cells for bone tissue engineering. Oral Surg Oral Med Oral Pathol Oral Radiol. 2012 Jul 26;

298. Lu L-L, Liu Y-J, Yang S-G, Zhao Q-J, Wang X, Gong W, et al. Isolation and characterization of human umbilical cord mesenchymal stem cells with hematopoiesis-supportive function and other potentials. Haematologica. 2006 Aug;91(8):1017–26.

299. Ciavarella S, Dammacco F, De Matteo M, Loverro G, Silvestris F. Umbilical cord mesenchymal stem cells: role of regulatory genes in their differentiation to osteoblasts. Stem Cells Dev. 2009 Oct;18(8):1211–20.

300. Zeddou M, Briquet A, Relic B, Josse C, Malaise MG, Gothot A, et al. The umbilical cord matrix is a better source of mesenchymal stem cells (MSC) than the umbilical cord blood. Cell Biol Int. 2010 Jul;34(7):693–701.

301. Hou T, Xu J, Wu X, Xie Z, Luo F, Zhang Z, et al. Umbilical Cord Wharton’s Jelly: A New Potential Cell Source of Mesenchymal Stromal Cells for Bone Tissue Engineering. Tissue Engineering Part A. 2009 Sep;15(9):2325–34.

302. Karahuseyinoglu S, Cinar O, Kilic E, Kara F, Akay GG, Demiralp DO, et al. Biology of stem cells in human umbilical cord stroma: in situ and in vitro surveys. Stem Cells. 2007 Feb;25(2):319–31.

303. Conconi MT, Burra P, Di Liddo R, Calore C, Turetta M, Bellini S, et al. CD105(+) cells from Wharton’s jelly show in vitro and in vivo myogenic differentiative potential. Int J Mol Med. 2006 Dec;18(6):1089–96.

304. Romanov YA, Svintsitskaya VA, Smirnov VN. Searching for alternative sources of postnatal human mesenchymal stem cells: candidate MSC-like cells from umbilical cord. Stem Cells. 2003;21(1):105–10.

305. Abedin M, Tintut Y, Demer LL. Mesenchymal stem cells and the artery wall. Circ Res. 2004 Oct 1;95(7):671–6.

306. Proudfoot D, Skepper JN, Hegyi L, Bennett MR, Shanahan CM, Weissberg PL. Apoptosis Regulates Human Vascular Calcification In Vitro Evidence for Initiation of Vascular Calcification by Apoptotic Bodies. Circulation Research. 2000 Nov 24;87(11):1055–62.

154

307. Proudfoot D, Skepper JN, Shanahan CM, Weissberg PL. Calcification of Human Vascular Cells In Vitro Is Correlated With High Levels of Matrix Gla Protein and Low Levels of Osteopontin Expression. Arterioscler Thromb Vasc Biol. 1998 Mar 1;18(3):379–88.

308. Kyllönen L, Haimi S, Mannerström B, Huhtala H, Rajala KM, Skottman H, et al. Effects of different serum conditions on osteogenic differentiation of human adipose stem cells in vitro. Stem Cell Res Ther. 2013 Feb 15;4(1):17.

309. Bettiol E, Sartiani L, Chicha L, Krause KH, Cerbai E, Jaconi ME. Fetal bovine serum enables cardiac differentiation of human embryonic stem cells. Differentiation. 2007 Oct;75(8):669–81.

310. Parker E, Shiga A, Davies J. Growing human bone in vitro. Bone Engineering. Davies, JE. Toronto, ON: Squared Incomrporated; 2000. p. 63–80.

311. Aghaloo T, Jiang X, Soo C, Zhang Z, Zhang X, Hu J, et al. A study of the role of nell-1 gene modified goat bone marrow stromal cells in promoting new bone formation. Mol Ther. 2007 Oct;15(10):1872–80.

312. Dilling CF, Wada AM, Lazard ZW, Salisbury EA, Gannon FH, Vadakkan TJ, et al. Vessel formation is induced prior to the appearance of cartilage in BMP-2-mediated heterotopic ossification. J Bone Miner Res. 2010 May;25(5):1147–56.

313. Aubin JE. Osteoprogenitor cell frequency in rat bone marrow stromal populations: role for heterotypic cell-cell interactions in osteoblast differentiation. J Cell Biochem. 1999 Mar 1;72(3):396–410.

314. Gundle R, Joyner CJ, Triffitt JT. Human bone tissue formation in diffusion chamber culture in vivo by bone-derived cells and marrow stromal fibroblastic cells. Bone. 1995 Jun;16(6):597–601.

315. Kuznetsov SA, Krebsbach PH, Satomura K, Kerr J, Riminucci M, Benayahu D, et al. Single- colony derived strains of human marrow stromal fibroblasts form bone after transplantation in vivo. J Bone Miner Res. 1997 Sep;12(9):1335–47.

316. Haynesworth SE, Goshima J, Goldberg VM, Caplan AI. Characterization of cells with osteogenic potential from human marrow. Bone. 1992;13(1):81–8.

317. Krebsbach PH, Kuznetsov SA, Satomura K, Emmons RV, Rowe DW, Robey PG. Bone formation in vivo: comparison of osteogenesis by transplanted mouse and human marrow stromal fibroblasts. Transplantation. 1997 Apr 27;63(8):1059–69.

155

Appendix 1

Materials used in this thesis work are listed below with their respective supplier and catalog number.

Material Supplier Catalog Number AccuPrime™ Taq High Fidelity Invitrogen 12346-086 AD-293 cells Statagene 240085 AdEasy™ XL Adenoviral Vector System Stratagene 240010 Agar BioShop AGR 001.1 Agarose BioShop AGA 001.500 Alexa Fluor 555 Goat Anti-Rabbit IgG Invitrogen A-21428 Alizarin Red Sigma-Aldrich A5533-25G Alkaline buffer solution Sigma-Aldrich A9226 Alkaline phosphatase New England Biolabs M0290 Alpha-MEM Invitrogen 12571-071 Amphotericin B Sigma-Aldrich A2942 Ampicillin Bioshop AMP201 anti-HA antibody Covance MMS-101R anti-Runx2 antibody Abcam ab488 anti-SP7 antibody Abcam ab57355 Ascorbic acid Sigma-Aldrich A4544 BglII restriction enzyme New England Biolabs R014S BJ5183-AD-1 electrocompetent cells Stratagene 200157 BMP2 human cDNA Dr. Bernhard Ganss C2C12 cells ATCC CRL-1772

CaCl2 Sigma-Aldrich C5080 Cell Line Optimization Nucleofector kit Amaxa VCO-1001N Cellytic M Cell Lysis Reagent Sigma-Aldrich C2978 Chloroquine Sigma-Aldrich C6628 DetergentOUT G-Bioscienses GBS10-3000 Dexamethasone Sigma-Aldrich D8893 DH5α competent E. coli cells Invitrogen 18265-017 D-MEM Gibco 11995-040 D-MEM Low Glucose Sigma-Aldrich D 6046 DMSO Sigma-Aldrich D2650 DNase/RNase-Free Distilled Water Invitrogen 10977015 EndoFree Plasmid Maxi Kit Qiagen 12362 Fast Red Violet LB salt Sigma-Aldrich F3381 Fetal bovine serum (FBS) HyClone FuGENE 6 Roche Diagnostics 11815091001 156

Gentamicin Sigma-Aldrich G3797 Goat Anti-Rabbit IgG (H+L)-HRP Bio-Rad 170-6515 Goat pAb to Ms IgG (HRP) Abcam ab6789-1 HeLa cells ATCC CMR2-CCL2 HEPES Sigma-Aldrich H7523 HepG2 ATCC 77400 HindIII restriction enzyme New England Biolabs R0104 Hoechst 33342 Invitrogen H3570 HUCPVCs TRT Human BMP2 DuoSet R&D Systems DY355 Human VEGF DuoSet R&D Systems DY293B Immobilon-P membrane Millipore IPVH00010 Immun-Star Western C Chemiluminescent Bio-Rad 1705070 iScript One-Step RT-qPCR with SYBR Green Bio-Rad 170-8893 LB Broth Lennox BioShop LBL 405.1 Kanamycin Sulfate Bioshop KAN201.5 Lipofectamine LTX Invitrogen 15338 Mitomycin C Sigma-Aldrich M4287 N,N-Dimethylformamide Sigma-Aldrich D4254

Na2HPO4 (Sigma, Sigma-Aldrich S7907 NaCl Sigma-Aldrich S7653 Naphthol AS MX-P04 Sigma-Aldrich N5000 NEBuffer 3 New England Biolabs B7003S Neutral Buffer Formalin Sigma-Aldrich HT501128 NOD/SCID mice Charles River Strain code 394 Opti-MEM Medium Invitrogen 31985-062 PacI restriction enzyme New England Biolabs R0547 Phosphate buffered saline (PBS) Gibco 14190-250 Penicillin Sigma-Aldrich P3032 Phoenix-AMPHO cells ATCC CRL-3213 Pierce 660nm Protein Assay ThermoScientific 22660 Platinum pfx DNA Polymerase Invitrogen 11708-013 pmaxGFP Amaxa PmeI restriction enzyme New England Biolabs R0560 P-Nitrophenol (pNP) solution Sigma-Aldrich N7660 P-Nitrophenol phosphate Sigma-Aldrich P5869 Polybrene Millipore TR-1003-G Protease Inhibitor Cocktail Sigma-Aldrich P8340 pShuttle-IRES-hrGFP-2 Stratagene 240082 Qiagen Miniprep Kit Qiagen 27106 QIAprep Spin Miniprep Kit Qiagen 27106

157

QIAquick Gel Extraction Kit Quiagen 27106 QIAquick PCR Purification Kit Qiagen 28106 QuantiChrome Calcium Assay Kit BioAssay Systems DICA-500 QuantiTect Reverse Transcription Kit Qiagen 205311 rhfurin R&D Systems 1503-SE rhBMP2 Dr. Sean Peel Runx2 human cDNA Origene SC302270 Runx2 Rabbit Polyclonal Antibody Santa Cruz Biotech SC-10758 Silver Nitrate BioShop SIL222.25

SP7 human cDNA Open Biosystems Clone ID# 8069055 Stop Solution ELISA R&D Systems DY994 Substrate Solution ELISA R&D Systems DY999 T4 DNA Ligase New England Biolabs M0202S Tris Buffered Saline (TBS) Bio-Rad 170-6435 Tri Reagent Sigma-Aldrich T9424 Triton-X Sigma-Aldrich T8787 Trypsin Invitrogen 25200056 Tween 20 Bio-rad 170-7531 VEGFA human cDNA Open Biosystems Clone ID# 6006890 XhoI restriction enzyme New England Biolabs R0146S X-ray film Clonex 2316-CLMR810 β-glycerophosphate Sigma-Aldrich G9891

158

Appendix 2

In this section the full cDNA sequences of the genes of interest cloned (section 3.3.1.1.) into the pShuttle-IRES-hrGFP-2 are provided.

The cDNA sequences are the actual PCR products (Figure 3.3B-C) ; the blue letters at the beginning and at the end of the sequences are the restriction sites BglII and XhoI respectively; the red letters are the start codon.

The protein sequences include the 3x HA (green letters) fused to the gene which determines the final protein size. HA tag cDNA sequence: tacccatacgatgttccagattacgct; amino acid sequence: YPYDVPDYA. 3x HA molecular weight: 3.27 kDa

Bone Morphogenetic Protein -2 (BMP-2)

BMP2 cDNA Sequence:

AGATCTGatggtggccgggacccgctgtcttctagcgttgctgcttccccaggtcctcctgggcggcgcggctggcctcgttccggagc tgggccgcaggaagttcgcggcggcgtcgtcgggccgcccctcatcccagccctctgacgaggtcctgagcgagttcgagttgcggctg ctcagcatgttcggcctgaaacagagacccacccccagcagggacgccgtggtgcccccctacatgctagacctgtatcgcaggcactc aggtcagccgggctcacccgccccagaccaccggttggagagggcagccagccgagccaacactgtgcgcagcttccaccatgaaga atctttggaagaactaccagaaacgagtgggaaaacaacccggagattcttctttaatttaagttctatccccacggaggagtttatcac ctcagcagagcttcaggttttccgagaacagatgcaagatgctttaggaaacaatagcagtttccatcaccgaattaatatttatgaaat cataaaacctgcaacagccaactcgaaattccccgtgaccagacttttggacaccaggttggtgaatcagaatgcaagcaggtgggaa agttttgatgtcacccccgctgtgatgcggtggactgcacagggacacgccaaccatggattcgtggtggaagtggcccacttggagga gaaacaaggtgtctccaagagacatgttaggataagcaggtctttgcaccaagatgaacacagctggtcacagataaggccattgcta gtaacttttggccatgatggaaaagggcatcctctccacaaaagagaaaaacgtcaagccaaacacaaacagcggaaacgccttaag tccagctgtaagagacaccctttgtacgtggacttcagtgacgtggggtggaatgactggattgtggctcccccggggtatcacgcctttt actgccacggagaatgcccttttcctctggctgatcatctgaactccactaatcatgccattgttcagacgttggtcaactctgttaactct aagattcctaaggcatgctgtgtcccgacagaactcagtgctatctcgatgctgtaccttgacgagaatgaaaaggttgtattaaagaac tatcaggacatggttgtggagggttgtgggtgtcgcCTCGAG

159

BMP2 Protein Sequence.

M V A G T R C L L A L L L P Q V L L G G A A G L V P E L G R R K F A A A S S G R P S S Q P S D E V L S E F E L R L L S M F G L K Q R P T P S R D A V V P P Y M L D L Y R R H S G Q P G S P A P D H R L E R A A S R A N T V R S F H H E E S L E E L P E T S G K T T R R F F F N L S S I P T E E F I T S A E L Q V F R E Q M Q D A L G N N S S F H H R I N I Y E I I K P A T A N S K F P V T R L L D T R L V N Q N A S R W E S F D V T P A V M R W T A Q G H A N H G F V V E V A H L E E K Q G V S K R H V R I S R S L H Q D E H S W S Q I R P L L V T F G H D G K G H P L H K R E K R Q A K H K Q R K R L K S S C K R H P L Y V D F S D V G W N D W I V A P P G Y H A F Y C H G E C P F P L A D H L N S T N H A I V Q T L V N S V N S K I P K A C C V P T E L S A I S M L Y L D E N E K V V L K N Y Q D M V V E G C G C R L E Y P Y D V P D Y A Y P Y D V P D Y A Y P Y D V P D Y A

BMP2 protein size= 425 amino acids (AA); molecular weight= ~ 48.2 kiladaltons (kDa)

Runt-related Transcription Factor 2 (Runx2)

Runx2 cDNA Sequence:

AGATCTGatggcatcaaacagcctcttcagcacagtgacaccatgtcagcaaaacttcttttgggatccgagcaccagccggcgctt cagccccccctccagcagcctgcagcccggcaaaatgagcgacgtgagcccggtggtggctgcgcaacagcagcagcaacagcagca gcagcaacagcagcagcagcagcagcaacagcagcagcagcagcaggaggcggcggcggcggctgcggcggcggcggcggctgcg gcggcggcagctgcagtgccccggttgcggccgccccacgacaaccgcaccatggtggagatcatcgccgaccacccggccgaactcg tccgcaccgacagccccaacttcctgtgctcggtgctgccctcgcactggcgctgcaacaagaccctgcccgtggccttcaaggtggtag ccctcggagaggtaccagatgggactgtggttactgtcatggcgggtaacgatgaaaattattctgctgagctccggaatgcctctgctg ttatgaaaaaccaagtagcaaggttcaacgatctgagatttgtgggccggagtggacgaggcaagagtttcaccttgaccataaccgtc ttcacaaatcctccccaagtagctacctatcacagagcaattaaagttacagtagatggacctcgggaacccagaaggcacagacaga agcttgatgactctaaacctagtttgttctctgaccgcctcagtgatttagggcgcattcctcatcccagtatgagagtaggtgtcccgcct cagaacccacggccctccctgaactctgcaccaagtccttttaatccacaaggacagagtcagattacagaccccaggcaggcacagtc ttccccgccgtggtcctatgaccagtcttacccctcctacctgagccagatgacgtccccgtccatccactctaccaccccgctgtcttcca cacggggcactgggcttcctgccatcaccgatgtgcctaggcgcatttcagatgatgacactgccacctctgacttctgcctctggccttc 160

cactctcagtaagaagagccaggcaggtgcttcagaactgggccctttttcagaccccaggcagttcccaagcatttcatccctcactga gagccgcttctccaacccacgaatgcactatccagccacctttacttacaccccgccagtcacctcaggcatgtccctcggtatgtccgcc accactcactaccacacctacctgccaccaccctaccccggctcttcccaaagccagagtggacccttccagaccagcagcactccatat ctctactatggcacttcgtcaggatcctatcagtttcccatggtgccggggggagaccggtctccttccagaatgcttccgccatgcacca ccacctcgaatggcagcacgctattaaatccaaatttgcctaaccagaatgatggtgttgacgctgatggaagccacagcagttcccca actgttttgaattctagtggcagaatggatgaatctgtttggcgaccatatCTCGAG

Runx2 Protein sequence

M A S N S L F S T V T P C Q Q N F F W D P S T S R R F S P P S S S L Q P G K M S D V S P V V A A Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q Q E A A A A A A A A A A A A A A A A A V P R L R P P H D N R T M V E I I A D H P A E L V R T D S P N F L C S V L P S H W R C N K T L P V A F K V V A L G E V P D G T V V T V M A G N D E N Y S A E L R N A S A V M K N Q V A R F N D L R F V G R S G R G K S F T L T I T V F T N P P Q V A T Y H R A I K V T V D G P R E P R R H R Q K L D D S K P S L F S D R L S D L G R I P H P S M R V G V P P Q N P R P S L N S A P S P F N P Q G Q S Q I T D P R Q A Q S S P P W S Y D Q S Y P S Y L S Q M T S P S I H S T T P L S S T R G T G L P A I T D V P R R I S D D D T A T S D F C L W P S T L S K K S Q A G A S E L G P F S D P R Q F P S I S S L T E S R F S N P R M H Y P A T F T Y T P P V T S G M S L G M S A T T H Y H T Y L P P P Y P G S S Q S Q S G P F Q T S S T P Y L Y Y G T S S G S Y Q F P M V P G G D R S P S R M L P P C T T T S N G S T L L N P N L P N Q N D G V D A D G S H S S S P T V L N S S G R M D E S V W R P Y L E Y P Y D V P D Y A Y P Y D V P D Y A Y P Y D V P D Y A

Runx2 protein size= 550 AA; molecular weight= ~ 60.16 kDa

SP7 Transcription Factor (Osterix)

SP7 cDNA Sequence:

AGATCTGatggcgtcctccctgcttgaggaggaagttcactatggctccagtcccctggccatgctgacggcagcgtgcagcaaattt ggtggctctagccctctgcgggactcaacaactctgggcaaagcaggcacaaagaagccgtactctgtgggcagtgacctttcagcctc caaaaccatgggggatgcttatccagccccctttacaagcactaatgggctcctttcacctgcaggcagtcctccagcacccacctcagg

161

ctatgctaatgattaccctcccttttcccactcattccctgggcccacaggcacccaggaccctgggctactagtgcccaaggggcacag ctcttctgactgtctgcccagtgtctacacctctctggacatgacacacccctatggctcctggtacaaggcaggcatccatgcaggcattt caccaggcccaggcaacactcctactccatggtgggatatgcaccctggaggcaactggctaggtggtgggcagggccagggtgatgg gctgcaagggacactgcccacaggtccagctcagcctccactgaacccccagctgcccacctacccatctgactttgctccccttaatcc agccccctacccagctccccacctcttgcaaccagggccccagcatgtcttgccccaagatgtctataaacccaaggcagtgggaaata gtgggcagctagaagggagtggtggagccaaacccccacggggtgcaagcactgggggtagtggtggatatgggggcagtggggca gggcgctcctcctgcgactgccctaattgccaggagctagagcggctgggagcagcagcggctgggctgcggaagaagcccatccaca gctgccacatccctggctgcggcaaggtgtatggcaaggcttcgcacctgaaggcccacttgcgctggcacacaggcgagaggcccttc gtctgcaactggctcttctgcggcaagaggttcactcgttcggatgagctggagcgtcatgtgcgcactcacacccgggagaagaagttc acctgcctgctctgctccaagcgctttacccgaagtgaccacctgagcaaacaccagcgcacccacggagaaccaggcccgggtcccc ctcccagtggccccaaggagctgggggagggccgcagcacgggggaagaggaggccagtcagacgccccgaccttctgcctcgccag caaccccagagaaagcccctggaggcagccctgagcagagcaacttgctggagCTCGAG

Protein Sequence SP7:

M A S S L L E E E V H Y G S S P L A M L T A A C S K F G G S S P L R D S T T L G K A G T K K P Y S V G S D L S A S K T M G D A Y P A P F T S T N G L L S P A G S P P A P T S G Y A N D Y P P F S H S F P G P T G T Q D P G L L V P K G H S S S D C L P S V Y T S L D M T H P Y G S W Y K A G I H A G I S P G P G N T P T P W W D M H P G G N W L G G G Q G Q G D G L Q G T L P T G P A Q P P L N P Q L P T Y P S D F A P L N P A P Y P A P H L L Q P G P Q H V L P Q D V Y K P K A V G N S G Q L E G S G G A K P P R G A S T G G S G G Y G G S G A G R S S C D C P N C Q E L E R L G A A A A G L R K K P I H S C H I P G C G K V Y G K A S H L K A H L R W H T G E R P F V C N W L F C G K R F T R S D E L E R H V R T H T R E K K F T C L L C S K R F T R S D H L S K H Q R T H G E P G P G P P P S G P K E L G E G R S T G E E E A S Q T P R P S A S P A T P E K A P G G S P E Q S N L L E L E Y P Y D V P D Y A Y P Y D V P D Y A Y P Y D V P D Y A

SP7 protein size= 459 AA; molecular weight= ~ 48.38 kDa

162

Vascular Endothelial Growth Factor A - 121 (VEGF-A121)

VEGF-A121 cDNA sequence:

AGATCTGatggcagaaggaggagggcagaatcatcacgaagtggtgaagttcatggatgtctatcagcgcagctactgccatccaa tcgagaccctggtggacatcttccaggagtaccctgatgagatcgagtacatcttcaagccatcctgtgtgcccctgatgcgatgcgggg gctgctgcaatgacgagggcctggagtgtgtgcccactgaggagtccaacatcaccatgcagattatgcggatcaaacctcaccaaggc cagcacataggagagatgagcttcctacagcacaacaaatgtgaatgcagaccaaagaaagatagagcaagacaagaaaaatgtga caagccgaggcggCTCGAG

VEGF-A121 Protein sequence:

M A E G G G Q N H H E V V K F M D V Y Q R S Y C H P I E T L V D I F Q E Y P D E I E Y I F K P S C V P L M R C G G C C N D E G L E C V P T E E S N I T M Q I M R I K P H Q G Q H I G E M S F L Q H N K C E C R P K K D R A R Q E K C D K P R R L E Y P Y D V P D Y A Y P Y D V P D Y A Y P Y D V P D Y A

VEGF-A121 protein size= 148 AA; molecular weight= ~ 17.4 kDa

163