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Evaluation of the role of ANGPTL4 in tendon vascularization Mousavizadeh Ahmadabadi, Seyed Rouhollah 2015

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Evaluation of the Role of ANGPTL4 in Tendon Vascularization  by Seyed Rouhollah Mousavizadeh Ahmadabadi MSc, Imam Hossein University, 2006  A THESIS SUBMITTED IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in The Faculty of Graduate and Postdoctoral Studies (Experimental Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  December 2015  © Seyed Rouhollah Mousavizadeh Ahmadabadi, 2015 ii  Abstract The mechanisms that regulate angiogenic activity in injured or mechanically loaded tendons are poorly understood. In this study we hypothesised that repetitive stretching of tendon cells alters the expression and release of angiogenic factors which may promote tendon vascularization.  In order to examine the effects of repetitive stretching on the expression of angiogenic genes, primary human tendon cells were subjected to cyclic stretching. Cyclic stretching of two-dimensional primary tenocyte cell cultures increased the expression of VEGF, bFGF and Cox-2. But, by extending the time course, VEGF, bFGF and Cox-2 were progressively downregulated. Angiogenic profiling of tendon cells by qPCR array identified a number of other genes (ANGPTL4, FGF-1, TGFα, VEGF-C and SPHK1) that responded to tensile loading in a similar pattern. Further experiments revealed that cyclic stretching of human tendon fibroblasts stimulated the expression and release of ANGPTL4 protein via TGF-β and HIF-1α signalling. ANGPTL4 promoted the angiogenic activity of endothelial cells. Angiogenic activity was also increased following injury and following ANGPTL4 injection into mouse patellar tendons, whereas the patellar tendons of ANGPTL4 knock out mice displayed reduced angiogenesis following injury. In human rotator cuff tendons, there were both spatial and quantitative associations of ANGPTL4 with tendon endothelial cells.  The experiments described in this thesis have shown that ANGPTL4 may assist in the regulation of vascularity in injured or mechanically loaded tendon. TGF-β and HIF-1α are two signalling pathways that modulate the expression of ANGPTL4 by tendon fibroblasts and which could, in future, be manipulated to influence tendon healing or adaptation.   iii  Preface Portions of the introductory text were modified from previously written abstracts for conferences, an application for a WorkSafe BC scholarship and my published manuscripts. I performed the literature review and wrote the manuscripts. Dr. Alexander Scott and Vincent Duronio participated in revision and editing. A version of chapter 3 has been published as “Mousavizadeh R, Khosravi S, Behzad H, McCormack RG, Duronio V, Scott A. (2014) Cyclic strain alters the expression and release of angiogenic factors by human tendon cells. PLoS One. 13;9(5):e97356.”.  Versions of Chapters 4 and 5 have been submitted for publication.  A version of Appendix A has been published as “Mousavizadeh R, Backman L, McCormack RG, Scott A. (2014) Dexamethasone decreases substance P expression in human tendon cells: an in vitro study. Rheumatology (Oxford)”. I and Dr. Ludvig Backman from Umea University participated in this project. Dr. Backman performed the protein quantification for substance P and participated in writing the manuscript. I conducted the experimental design, performed the assays, reviewed the literature and wrote the manuscript under the supervision of Dr. Scott. The main portion of chapter 2 was modified from my published and submitted manuscripts as explained for following chapters. I formulated the hypotheses and experimental design, performed the assays, analyzed the data and generated the figures. The zymography was conducted by Shahram Khosravi. Alex Lu performed the image processing and immunostaining of mouse tendon tissues. Alexandra Kobza and Attia Riaz assisted with some of the qPCR tests. Dr. iv  Gholamreza Safaee assisted in harvesting mouse tendons and prepararing figures. Mazyar Ghaffari also assisted with the dual luciferase assay. Dr. Aly Karsan and Dr Eleanor Jones provided HUVEC cells and plasmids respectively. Dr. Hayedeh Behzad assisted with experimental design. Dr. Giada Marino performed the in vitro ANGPTL4 cleavage assay. Dr. Vincent Duronio and Dr. Scott provided overall guidance, logistical support and the facilities for the project. They also participated in the literature review and in the writing of manuscripts.  The use of orthopaedic autograft material (semitendinosis tendon) for isolating tendon cells was approved by the UBC Clinical Research Ethics Board (certificate number H10-00220), and each patient provided written informed consent. The primary HUVEC cells were isolated from normal umbilical cords under a UBC approved human ethics certificate (certificate number H03-50102). The UBC Animal Care Committee has examined and approved the experimental protocols for the breeding (certificate number A14-0053) and use of animals (certificate number A12-0092).          v  Table of contents  Abstract .......................................................................................................................................... ii Preface ........................................................................................................................................... iii Table of contents ........................................................................................................................... v List of tables................................................................................................................................... x List of figures ................................................................................................................................ xi List of abbreviations .................................................................................................................. xiii Acknowledgements ................................................................................................................... xvii Chapter 1. Introduction ............................................................................................................... 1 1.1. Research aims .................................................................................................................................... 1 1.2. Human tendon .................................................................................................................................... 2 1.2.1. Structure ...................................................................................................................................... 2 1.2.2. Matrix .......................................................................................................................................... 3 1.2.3. Vascular system .......................................................................................................................... 4 1.2.4. Cell population ............................................................................................................................ 5 1.3. Tendon injuries .................................................................................................................................. 6 1.3.1. Terminology ................................................................................................................................ 6 1.3.2. Epidemiology .............................................................................................................................. 7 1.4. Tendon healing ................................................................................................................................... 8 1.4.1. Inflammatory phase..................................................................................................................... 8 1.4.2. Proliferative phase....................................................................................................................... 9 1.4.3. Remodeling phase ..................................................................................................................... 10 vi  1.5. Overuse tendinopathy ...................................................................................................................... 11 1.5.1. Etiology ..................................................................................................................................... 11 1.5.2. Histopathology .......................................................................................................................... 12 1.5.3. Cytokines and growth factors ................................................................................................... 13 1.5.4. MMPs ........................................................................................................................................ 15 1.6. Role of neovascularization in tendinopathy ..................................................................................... 16 1.7. Mechanotransduction ....................................................................................................................... 17 1.8. Mechanobiology of tendon cells ...................................................................................................... 19 1.9. Mechanical stimulus - effects on gene regulation ............................................................................ 20 1.10. Substance P (SP) ............................................................................................................................ 27 1.11. TGF-β signaling pathway .............................................................................................................. 27 1.12. Function of TGF-β in tendon tissue ............................................................................................... 30 1.13. Role of HIF-1 in tendon injury ...................................................................................................... 31 1.14. Angiopoietin-like 4 ........................................................................................................................ 33 1.14.1. Function of ANGPTL4 ........................................................................................................... 33 1.14.2. Regulation of ANGPTL4 ........................................................................................................ 34 1.15. Specific aims and hypotheses ........................................................................................................ 36 1.15.1. Study I: Evaluation of the effect of cyclic stretching on the expression of angiogenic factors by tendon cells ......................................................................................................................................... 36 1.15.2. Study II: Investigation of the mechanisms that regulate ANGPTL4 ...................................... 36 1.15.3. Study III: Characterizing the function of ANGPTL4 protein in tendon tissue ....................... 37 Chapter 2. Materials and methods ............................................................................................ 38 vii  2.1. Animal studies ................................................................................................................................. 38 2.2. Primary tendon cell culture .............................................................................................................. 38 2.3. Ethics ............................................................................................................................................... 39 2.4. Mechanical stimulation of tendon cells ........................................................................................... 40 2.5. Flow cytometry ................................................................................................................................ 40 2.6. Cell proliferation .............................................................................................................................. 41 2.7. Endothelial cell tube formation assay .............................................................................................. 41 2.8. Gene expression analysis ................................................................................................................. 42 2.9. Zymography ..................................................................................................................................... 50 2.10. Immunoblotting .............................................................................................................................. 50 2.11. Enzyme-Linked Immunosorbent Assay (ELISA) .......................................................................... 52 2.12. TGF-β luciferase assay .................................................................................................................. 52 2.13. Immunohistochemistry .................................................................................................................. 52 2.14. Image Processing ........................................................................................................................... 53 2.15. In vitro ANGPTL4 cleavage assay ................................................................................................ 55 2.16. In vitro scratch assay ...................................................................................................................... 55 2.17. Statistics ......................................................................................................................................... 56 Chapter 3. Cyclic stretching alters the expression and release of angiogenic factors by human tendon cells ..................................................................................................................... 57 3.1. Background ...................................................................................................................................... 57 3.2. Results .............................................................................................................................................. 58 3.2.1. Cell viability of tendon cells after applied cyclic stretching ..................................................... 58 viii  3.2.2. Increased angiogenic activity of factors released by repetitively stretched tendon cells .......... 59 3.2.3. Cyclic Stretching Upregulates the Expression of Angiogenic Factors in Tendon Cells ........... 61 3.2.4. Cyclic stretching increases the release of the ANGPTL4 and MMP-2 proteins ....................... 64 Chapter 4. TGF-β and HIF 1α modulate the expression of ANGPTL4 in response to cyclic stretching ..................................................................................................................................... 67 4.1. Background ...................................................................................................................................... 67 4.2. Results .............................................................................................................................................. 68 4.2.1. HIF-1α modulates the expression of ANGPTL4 ...................................................................... 68 4.2.2. ANGPTL4 induced by cyclic stretching is mediated by TGF-β activity .................................. 71 4.2.3. ANGPTL4 is induced by both HIF-1α and TGF-β ................................................................... 73 Chapter 5. Role of ANGPTL4 in tendon vascularization ....................................................... 76 5.1. Background ...................................................................................................................................... 76 5.2. Results .............................................................................................................................................. 77 5.2.1. Angiogenic activity of ANGPTL4 promotes tendon vascularization ....................................... 77 5.2.2. Lack of ANGPTL4 has no effect on gene markers for tendon cells and their progenitors ....... 85 5.2.3. MMPs are able to cleave recombinant ANGPTL4 protein ....................................................... 86  Discussion.................................................................................................................................... 88 Chapter 6. .................................................................................................................................... 88 6.1. Cyclic stretching modulates angiogenic factors in tendon cells ...................................................... 88 6.2. ANGPTL4 activity is increased in cyclically stretched tendon fibroblasts and promotes angiogenesis in injured tendon ................................................................................................................ 91 Chapter 7. Conclusions and perspectives ................................................................................. 96 Bibliography ................................................................................................................................ 98 ix  Appendices ................................................................................................................................. 115 Appendix A. Substance P and Its Regulatory Pathway in Response to Glucocorticoids 115 A.1. Material and Methods ............................................................................................................... 116 A.1.1. Cell culture ........................................................................................................................ 116 A.1.2. Experimental condition and substances for experimental design ..................................... 117 A.1.3. RNA isolation, reverse transcription, and qPCR .............................................................. 118 A.1.4. Immunocytochemistry (ICC) ............................................................................................ 120 A.1.5. Immunoblotting ................................................................................................................. 120 A.1.6. SP enzyme immunoassay (EIA)........................................................................................ 121 A.1.7. Statistical analysis ............................................................................................................. 121 A.2. Results ....................................................................................................................................... 122 A.2.1. Dexamethasone down-regulates SP expression (TAC1 mRNA) ...................................... 122 A.2.2. Dexamethasone reduces SP production ............................................................................ 122 A.2.3. Dexamethasone reduces SP expression through the glucocorticoid receptor ................... 122 A.2.4. Dexamethasone reduces the expression of interleukins .................................................... 123 A.2.5. Dexamethasone inhibits the expression of SP induced by IL-1β ...................................... 123 A.2.6. Dexamethasone inhibits the expression of SP induced by cyclic strain............................ 123 A.3. Discussion ................................................................................................................................. 125 Appendix B. Tendon cell characterization .......................................................................... 128 B.1. Methods..................................................................................................................................... 128 B.2. Result ........................................................................................................................................ 130  x  List of tables Table 1-1. Classification of tendon injuries .................................................................................... 7 Table 1-2. Cytokines and their effects in tendon tissue. ............................................................... 14 Table 1-3. Mechanical stimulation of human tendon cells. .......................................................... 21 Table 2-1. Gene symbols and description of angiogenic factors analyzed by Human Angiogenesis RT² Profiler™ PCR Array ..................................................................................... 43 Table 2-2. Oligonucleotide sequence of primers and amplicon sizes of selected angiogenic genes........................................................................................................................................................ 48 Table 2-3. Oligonucleotide sequence of primers and amplicon sizes of selected mouse genes. .. 49 Table A-1 Oligonucleotide sequence of primers and amplicon sizes ......................................... 119 Table B-1. Oligonucleotide sequence of primers and amplicon sizes of the gene markers for tendon cells. ................................................................................................................................ 129           xi  List of figures Figure 1-1. Tendon Structure .......................................................................................................... 3 Figure 1-2. Large Latent Complex................................................................................................ 29 Figure 1-3. TGF-β Signalling Pathway. ........................................................................................ 29 Figure 1-4. Structure of HIF-1 subunits and their functions. ........................................................ 32 Figure 1-5. Structure and dual function of ANGPTL4. ................................................................ 34 Figure 1-6. Hypothetical function of ANGPTL4 and related pathways in tendon. ...................... 37 Figure 3-1. Flow cytometry histograms of tendon cells following propidium iodide staining. ... 59 Figure 3-2. Increased angiogenic activity of released factors by stretched tendon cells. ............. 60 Figure 3-3 Gene expression array of angiogenic factors expressed by tendon cells after 4 and 12 hours cyclic stretching. ................................................................................................................. 62 Figure 3-4. Real-time quantitative PCR on selected genes........................................................... 63 Figure 3-5. Cyclic stretching increases the expression and release of ANGPTL4 in tendon cell culture. .......................................................................................................................................... 65 Figure 3-6 Zymography of MMP-2 activity. ................................................................................ 66 Figure 4-1. HIF-1 modulates the expression of ANGPTL4 in human tendon cells. .................... 69 Figure 4-2. Correlation of ANGPTL4 and HIF-1α expression in human rotator cuff tendons. ... 70 Figure 4-3. TGF-beta regulates ANGPTL4 expression in human tendon cells. ........................... 72 Figure 4-4. Gene expression array of angiogenic factors in tendon cells after 6 hours incubation with DMOG, PGE2 and TGF-β (mean ± SE; N=3). .................................................................... 74 Figure 4-5. An array of angiogenesis related genes regulated in response to TGF-β and DMOG........................................................................................................................................................ 75 Figure 5-1. In vitro and in vivo angiogenic activity of recombinant ANGPTL4 protein. ............ 79 Figure 5-2. Lack of ANGPTL4 decreases tendon vascularization following tendon injury in an ANGPTL4-defiecient mouse model. ............................................................................................ 80 xii  Figure 5-3. Scratch assay of isolated mouse tendon cells. ............................................................ 81 Figure 5-4. Correlation of ANGPTL4 expression with target genes. ........................................... 82 Figure 5-5. Correlation of ANGPTL4 and CD31 expression in human rotator cuff tendons. ..... 83 Figure 5-6. The scanned images of immunostained human rotator cuff tendons. ........................ 84 Figure 5-7. The expression of selected gene markers in PTs of ANGPTL4-deficient mice (-/-) versus wild type (+/+). .................................................................................................................. 85 Figure 5-8. The cleaved products of ANGPTL4 protein after incubation with MMPs. ............... 87 Figure A-1 Hypothetical pathway regulating SP in response to glucocorticoids ....................... 115 Figure A-2. Dexamethasone decreases TAC1 expression without affecting the mRNA level of the preferred SP receptor (NK1R). ............................................................................................. 125 Figure B-1. The expression of tendon cell markers during different passages. .......................... 130          xiii  List of abbreviations ADAMTS a disintegrin and metalloproteinase with thrombospondin motifs α-SMA alpha smooth muscle actin ATP adenosine triphosphate ARNT aryl hydrocarbon receptor nuclear translocator protein ASTD activity related soft tissue disorder bHLH basic helix-loop-helix bFGF basic fibroblast growth factor BMP bone morphogenetic protein bp base pair COL1A1 collagen, type I, alpha 1 COL3A1 collagen, type III, alpha 1 COL12A1 collagen, type XII, alpha 1 COX Cyclooxygenase CTGF connective tissue growth factor DCN decorin ECM extracellular matrix EDTA ethylenediaminetetraacetic acid EIA enzyme immunoassay ENaC/DEG epithelial sodium channel/degenerin FBMD femur bone mineral density FIH-1 factor inhibiting HIF-1 FN fibronectin xiv  GAGs glycosaminoglycans GDF growth differentiation factor GR glucocorticoid receptor Hz hertz HIF-1 hypoxia-inducible factor 1 HSP heat shock protein HUVEC human umbilical vein cell ICC immunocytochemistry IGF-I  insulin-like growth factor-I IL interleukin ITGA5 integrin, alpha 5 ITGB1 integrin, beta 1 JNK c-Jun N-terminal kinase LAP latency associated peptide  LDH lactate dehydrogenase LLC large latent complex LTBP latent TGF-β-binding protein  LTB4 Leukotriene B4 LOX lysyl oxidase MAPK mitogen-activated protein kinase MAPKKK MAPK kinase kinases MMP matrix metalloproteinases MKX mohawk homeobox xv  NK1R neurokinin 1 receptor ODDD oxygen-dependent degradation domain PAS PER-ARNT-SIM PDGF platelet-derived growth factor PER period circadian protein PGs proteoglycans PGE2 prostaglandin E2 PHD prolyl hydroxylase domain PT ROS patellar tendon reactive oxygen species  Scx scleraxis SIM single-minded protein SP substance p SPHK1 sphingosine Kinase 1 TAC1 Tachykinin, Precursor 1 TAD transactivation domain TAK1 TGF-β-activated kinase 1 TIMP tissue inhibitor of metalloproteinase TGF-β transforming growth factor beta TNF tumor necrosis factor Tnmd tenomodulin THBS1 thrombospondin 1 TRP Transient receptor potential channel xvi  TSP-1 thrombospondin-1 VEGF vascular endothelial growth factor VHL Von Hippel–Lindau tumor suppressor                 xvii  Acknowledgements …Not possible to put in words… I am deeply grateful to my PhD supervisors Dr. Alexander Scott and Dr. Vincent Duronio for their absolute support and their prominent impact on my scientific career and life. I am glad that I had chance to work under their supervision.  I wish to thank my PhD committee members, Dr. Chris Overall, Dr. Michael Underhill and Dr. Michael Hunt, who guided and supported my studies. Their indispensable suggestions led to the elaboration of different steps in this project, and the value I place on their generous support is inexpressible.   The success of my PhD studies also is also partially thanks to my colleagues and friends. My special thanks to Dr. Hayedeh Behzad, Dr. Ludvig Backman (from Umea University), Dr. Gholamreza Safaee, Dr. Alireza Moeen Rezakhanlou, Dr. Mehdi Jafarnejhad, Dr. Sarwat Jamil, Dr. Ronald Wong, Dr. Peng Zhang, Dr. Robert Bell, Mazyar Ghaffari, Payman Hojabrpour, Shahram Khosravi, Alex Lu, Elise Huisman, Eric Pesarchuk and many people in the Jack Bell Research Centre (JBRC) and the Centre for Hip Health and Mobility (CHHM) who provided such a friendly, vivid and dynamic environment, and helped me in my research. I am also grateful to people in Centre for Translational and Applied Genomics (CTAG) and St. Paul's Hospital and the animal facility of JBRC, particularly Kate Orchard, Madeleine Stephens, Stephanie Smith, Josephine Lee and Julie Lorette for their assistance. xviii  I would like to acknowledge Drs. Robert G McCormack and Kirsten Lundgreen, Dr. Aly Karsan and Dr. Eleanor Jones for providing human tendon tissues, HUVEC cells and plasmids, respectively. I am honored to have been the recipient of a research training award from WorkSafe BC for three years which partially provided my stipend and research travel allowances. I am inspired by their commitment to supporting research and improving work-related settings. I am also grateful to the British Society of Matrix Biology and UBC for funding my scientific travels.             1  Chapter 1. Introduction 1.1.Research aims Tendinopathy is a common overuse injury, prevalent among both athletes and workers (Lewis et al. 2009). The disorder is often considered a mechanically driven pathology, as repetitive or forceful loading of tendons are well established risk factors (Seitz et al. 2011). Repetitive strain and shear are thought to induce matrix degeneration in tendon tissue, making the tissue susceptible to damage and eventually to overuse injury (Mehta et al. 2003). However, the mechanisms which precede the development of symptomatic injury have not been fully described but are felt to be multifactorial, with repetitive strain being an important risk factor (Blevins 1997, Arnoczky et al. 2007). To date, there has been very little research aimed at understanding the mechanisms that promote neovascularization in overused tendons. A better understanding of the relationship between repetitive loading and neovascularization, and the contribution of angiogenic factors to the progression of tendon degeneration, can help in the design of novel and effective treatments to limit the advancement of overuse tendon injuries and to improve their clinical outcomes.  The original objective of my PhD project was to determine the factors that promote the proliferation of new blood vessels in response to repetitive stretching of tendon cells. The general goal was the understanding of mechanisms involved in pathogenesis of activity-related soft tissue disorders in order to provide improved understanding that could assist in the future development of new modalities for the disease management.  In my study I have used three complementary research models: 1. In vitro model: identifying molecular factors and regulatory mechanisms that  promote angiogenesis in response to repetitive stretch 2  2. In vivo model: Characterizing the mechanoresponsive genes (identified in model 1) during the course of tendon neovascularization following tissue injury 3. Clinical models: Histological study of human tendon tissue  1.2. Human tendon 1.2.1. Structure Tendons are connective tissues that are specialized for transmitting force from muscle to bone. Their shapes vary based on their anatomical location and force transmission. For example, finger flexors have long and thin tendons to transmit precise and delicate movements while the tendons of quadriceps and triceps brachialis are short and wide to resist and transmit powerful forces (Kannus 2000). The fundamental unit of tendons is the collagen fiber, which consists of a bundle of collagen fibrils. These fibrils are mainly composed of type I collagen which is synthesized by resident fibroblast cells named tenocytes. Tenocytes secrete collagen into the ECM when the polypeptides form a collagen triple helix. The cross-linking of collagen triple helices forms collagen fibrils which then further assemble into microfibrils and then collagen fibrils which appear as visible units under the electron microscope. A bundle of collagen fibrils form a collagen fiber which is visible with light microscopy. Collagen fibers, as structural units of tendon, bear force during tendon elongation. These fibers are surrounded by endotenon which is a sheath of connective tissue. Collagen fibers bundle into a hierarchical organization which make a primary (subfascicle), secondary (fascicle) and tertiary fiber bundles (Figure 1-1.). Tertiary fiber bundles may be covered by an epitenon sheath which is enclosed by a paratenon or a synovial sheath. Tendons with synovial sheaths such as hand tendons are lubricated with synovial fluids which 3  diminish sliding friction. In other tendons like Achilles’s tendon, epitendon and paratenon partial sheaths are responsible for reducing friction.   Figure 1-1. Tendon Structure  1.2.2. Matrix Water constitutes 55-70% of the wet weight of tendon tissue and negatively correlates with the elastic modules of tendon (Kjaer 2004, Birch 2007).  Collagen is the main component of the dry weight of tendon (60-85%) and mostly is collagen type I (95%) with traces of collagen types III, V, XI, XII and XIV.  The main component of tendon matrix is organized collagen fibers which withstand tensile forces.  Elastin fibers, inorganic and ground substances are the remaining major components of tendon matrix and their functions are not as well characterized.  Elastic fibers are 4  2% of the dry weight of tendon and are mainly made of elastin, fibrillins 1 and 2 proteins. These fibers are localized between fascicles to facilitate their sliding (Grant et al. 2013). Inorganic and ground substances include proteoglycans (PGs), glycosaminoglycans (GAGs), glycoproteins, and other small molecules which predominantly are leucine-rich proteins and play a role in lubricating and fibril fusion. The large aggregating PGs (such as versican and aggrecan) and the small leucine-rich proteoglycan (SLRP) are two groups of PGs in tendon tissue which govern water content and collagen fibril assembly of tendon tissue, respectively. A balance between components of tendon matrix determines the mechanical properties of tendon tissue to perform its function in force transmission. The matrix also provide a dynamic microenvironment for resident cells to ensure tissue homeostasis and adaptive responses (Kannus 2000, Kjaer 2004, Birch 2007, James et al. 2008, Screen et al. 2015).   1.2.3. Vascular system  Mature tendons are relatively hypo-vascular (compared to vessel-rich organs like skin, liver, lungs, etc.) due to their limited metabolic activity and unique mechanical function (Pufe et al. 2005). Nutrients for tendon tissue are derived both from synovial fluid diffusion and vascular perfusion; in some tendons such as the flexor tendons of the hand, synovial diffusion may be predominant. However this modest vascular perfusion is essential for tissue homeostasis. Tendon blood vessels originate from three different anatomical areas including surrounding connective tissues (e.g. epitendon, paratenon etc.) , musculotendinous and  osseotendinous junctions  (Fenwick et al. 2002).  Some tendons like the rotator cuff tendons are also vascularized by arteries and arterioles which usually come from the perimysium at the musculotendinous and 5  osseotendinous junctions (Kjaer 2004). The pattern of blood vessels is diverse among different tendons; in some tendons blood vessels extend along the length of tendon while others mostly have curved or branching blood vessels. Vessels insert into epitendon, endotenon and between collagen fibers in tendons covered with sheaths of connective tissue. The insertion sites for some tendons occur at very specific sites along tendon (e.g. the anterior insertion points of blood vessels into the human Achilles). However in other tendons covered with paratenon, blood vessels may insert in any point along the paratenon (Fenwick et al. 2002). Codman and Akerson described a relatively hypovascular zone in supraspinatus tendon which came to be known as “Codmans critical zone”. Hypovascular zones also have been described in a few other tendons such as biceps, Achilles, patella and posterior tibial tendons. Tendon degeneration and rapture more frequently associated with these zones; however there is no strong evidence to suggest that a lack of vascularity is important in the etiology of tendon degeneration (Fenwick et al. 2002).  1.2.4. Cell population The main cell population in tendon tissue is composed of tenocytes and tenoblasts which comprise 90-95% of the resident cells. Tenocytes are the highly elongated fibroblast-like cells that aligned longitudinally between the collagen fibers. These cells synthesise collagen and other factors that modulate tendon matrix (Kannus 2000, Riley 2008). Absence of specific markers for tenocytes makes them difficult to distinguish among other tendon cells; however scleraxis (SCX) and tenomodulin (TNMD) are suggested as potential markers for tenocytes (Shukunami et al. 2006). Tenoblasts have traditionally been identified morphologically – they have a bigger size 6  compared to tenocytes which suggest a higher metabolic activity. These cells have varied morphology from round to spindle shape. Tenoblasts differentiate into tenocytes during tendon aging and slightly lose their metabolic activity and rounded appearance. However tenocytes are also metabolically active and regulate the components of tendon matrix. Tenoblasts may simply represent a more metabolically active tenocyte, rather than a distinct cell population. The other cells in tendon tissue (including synovial cells, chondrocytes, and vascular cells) are associated with tendon sheaths, insertion sites and blood vessels respectively (Kannus 2000).   1.3. Tendon injuries 1.3.1. Terminology Several intrinsic and extrinsic factors can lead to acute or chronic tendon injuries in different site. Achilles, patellar, posterior tibial, elbow, wrist and rotator cuff tendons are the common sites for tendon injuries (Maffulli et al. 2003). A variety of terms pertaining to tendon disorders are referred to in the scientific and clinical literature. The pathological classification of tendon injuries is still a matter of debate. However a classification for the tendon conditions has been adopted based on histological findings and clinical symptoms. Table 1-1 shows the definitions of four types of tendon injuries based on histological and clinical signs.    7  Table 1-1. Classification of tendon injuries Term Definition Tendinopathy A clinical syndrome of tendon pain and thickening, diagnosed according to specific signs and symptoms (Khan et al. 1999). Tendinosis Structural changes in a tendon: most notably, collagen degeneration and extracellular matrix abnormality, with or without, proliferation of nerves and vessels (de Mos et al. 2007).  Activity related soft tissue disorder (ASTD)* An encompassing term which includes tendinopathy, as well as other musculoskeletal soft tissues (paratenon, bursa, muscle, nerve, ligament). Tendon rupture (complete or partial) A macroscopic failure of tendon tissue with catastrophic results.  * Also referred to as cumulative trauma disorder, repetitive strain injury, overuse syndrome, repetitive motion disorder (Rempel et al. 1992).   1.3.2. Epidemiology Tendon disorders are a significant cause of pain and morbidity amongst athletes, workers and the general public. The condition affects numerous tendons throughout the body and can prevent patients from physical activity. Overuse injuries comprise 30% to 50% of all sport injuries (Scott et al. 2006). Tendinopathy is a common overuse injury prevalent among athletes and workers (Kaux et al. 2011). The incidence of musculoskeletal soft tissue injuries is influenced by age and gender. A study by Clayton et al demonstrated that patients with Achilles, patellar, quadriceps or extensor tendon injuries are mostly middle aged; while rotator cuff tears and biceps 8  tendon rupture are predominant in elderly people. The overall age distribution of soft tissue injury in males shows a high peak at the age of 20 to 30. They also showed that the incidence of the soft tissue injuries is higher in males in different age groups; however these gender differences decline with increasing age. The highest incidences of tendon injuries include forearm/hand tendons and Achilles tendon rupture (Clayton et al. 2008). Some population studies in Finland and Hungary suggest an increased risk of tendon injuries in people with O blood type; however in other populations this association was not significant.  The risk of tendon injury is also higher among athletes and sports professionals. Training errors, repetitive movements and overload are the main causes of sport-related tendon injuries (Maffulli et al. 2003).   1.4. Tendon healing After acute tendon injuries, tendons start a healing process which may be categorized into the three following, overlapping stages:  1. Inflammatory phase 2. Proliferative phase 3. Remodeling phase  1.4.1. Inflammatory phase During the first few days after tendon injury, the inflammatory phase of tendon healing begins the repair process. Due to vessel injury, the coagulation cascade, and local upregulation of vascular adhesion molecules, blood cells such as white blood cells and platelets, infiltrate into the 9  tendon tissue to form a hematoma. Entrapped leukocytes and platelets in the fibrin-rich clot release pro-inflammatory mediators, growth factors and chemoattractants at the site of injury which attracts more inflammatory cells including monocytes and lymphocytes. These factors released during clotting and inflammation also stimulate tenocytes and angiogenesis pathways which speed up the next stage of tendon healing. The phagocytic cells such as neutrophils and macrophages are the major cell types which invade the wound (injury) site during the inflammatory phase, and digest necrotic tissues and blood clot (Hope et al. 2007, Yang et al. 2013).    1.4.2. Proliferative phase About 2-3 days after tendon injury, the proliferative or reparative phase of injury begin with proliferation and migration of tenocytes to the injured area, and increased tendon neovascularization. The sources of proliferative tenocytes are from internal and surrounding tissues including endotenon, epitenon, paratenon and synovial sheaths. At this stage phagocytic macrophages begin transforming to become reparative macrophages. The growth factors and cytokines released by these macrophages induce expansion of tendon cells and synthesizing tissue matrix including collagen type III and glycosaminoglycan. Increased release of various factors including bFGF, BMP, CTGF, IGF-I, TGF-β, PDGF, VEGF promote reparative processes including angiogenesis.  PDGF directs macrophages and fibroblasts to the site of injury, while TGF-β stimulates the synthesis of collagen. Some of the induced factors such as VEGF stay upregulated during the next phase of tendon healing. The study of flexor tendon injury shows that indication of new blood vessels appear after 7 days of tendon injury, which is correlated with peak expression of VEGF. MMPs are also players in tendon repair, since they regulate collagen 10  degradation and remodeling. The expression pattern of various MMPs determine the phase of tendon healing. The peak expression of MMP-9 and MMP-13 during the proliferative phase suggests that they play a role in collagen degradation, while MMP-2, MMP-3 and MMP-14 mediate collagen degradation and remodeling at the last phase of tendon healing. The transient upregulation of BMP-12, -13, and -14 promotes differentiation of tenocytes and stimulates the expression of tenogenic markers (Gelberman et al. 1985, Hope et al. 2007, Yang et al. 2013).  1.4.3. Remodeling phase The remodeling phase begins 1-2 months after injury and may take months or years. The synthesis of collagen type I increases and then returns to normal while the deposition of collagen type III diminishes in order to allow tendon fibers to become more aligned and mechanically resilient. The cell populations and glycosaminoglycans also decrease. PDGF and bFGF promote proliferation of tendon fibroblast cells to increase cell populations while IGF-1 and TGF-β induce synthesis of matrix components such as proteoglycan, collagen, and non-collagen proteins. IGF-1 also induces fibroblast proliferation and migration to the site of injury. Some studies showed the beneficial effects of IGF-1 on tendon healing, and induced expression of IGF-1 can improve healing outcomes. TGF-β has broad biological effects on tendon tissue. In addition to its role in matrix synthesis, TGF-β induces the expression of tendon cell markers and is involved in maintenance and differentiation of tendon progenitors. As mentioned before, the expression of MMP-2, MMP-3 and MMP-14 remains unregulated in this phase and plays a critical role in collagen remodeling to promote the formation of mature and normal fibers (Voleti et al. 2012, Yang et al. 2013, Thomopoulos et al. 2015).  11  1.5. Overuse tendinopathy 1.5.1. Etiology Tendinopathy is a common overuse injury, prevalent among both athletes and workers (Lewis et al. 2009). The disorder is often considered a mechanically driven pathology, as repetitive or forceful loading of tendons are well established risk factors (Seitz et al. 2011). Overuse of tendon through repetitive or forceful movements leads to the accumulation of microtrauma which distorts the integrity of collagen fibers and tendon tissue. Also, repetitive strain and shear are thought to induce cell-mediated matrix degeneration in tendon tissue, making the tissue increasingly susceptible to damage and eventually to overuse injury (Mehta et al. 2003). However, the mechanisms which precede the development of symptomatic injury have not been fully described but are felt to be multifactorial, with repetitive strain being an important risk factor. Sedentary lifestyle also contributes to poor tendon healing and tendon degeneration due to low circulation and nutrition of tendon cells. So it seems that a balanced physical activity which promotes collagen synthesis but avoids overuse or overload injury is necessary for tendon health. Some metabolic, inflammatory and genetic disorders such as obesity, rheumatological diseases and Marfan’s syndrome may influence the integrity and strength of collagen fibers and promote tendinopathy. The impact of some conditions on tendinopathy that interfere with microvascularity of tendon such as hypertension, diabetes, hyperuricemia and high serum lipids, emphasize the importance of tendon circulation in tendon health.  It seems that the metabolic state of tendon tissue also is important in the regulation of matrix synthesis and collagen fiber assembly. Therefore, conditions that disturb this regulation may deteriorate the strength and integrity of tendon and lead to tendinopathy. Nutritional conditions and pharmaceutical agents may alter the turnover of matrix production in tendon and change tendon strength. Some evidence suggests a 12  detrimental effect of corticosteroid injection for some tendons such as the Achilles and lateral elbow tendons. Since corticosteroids cannot prevent tendon degeneration and may also reduce the stiffness and strength of tendon, they may in fact exacerbate tendon problems. During maturation and development of tendon, the stiffness and strength of tendon tissue increases; however histological studies have shown that aging has a detrimental impact on tendon tissue. In general, multiple intrinsic and extrinsic factors may simultaneously promote some micro- or macro-level changes in tendon tissue which may accumulate and predispose individuals to tendinopathy (Blevins 1997, Jarvinen et al. 1997, Kannus et al. 1997, Arnoczky et al. 2007, Seitz et al. 2011). In cases of chronic tendinopathy, a lack of inflammatory cellular infiltration in some studies of surgical biopsies has led some to challenge the involvement of inflammation in the etiology of this disorder (Khan et al. 1999). However, several studies have, in both animal models and humans, shown inflammatory reactions in early development of tendinopathy (Backman et al. 1990, Millar et al. 2010). More recently, macrophages and T and B lymphocytes has been detected not only in the early phase, but also in chronic tendinopathy (Schubert et al. 2005, Kragsnaes et al. 2014). Other studies have demonstrated increased levels of macrophage-derived interleukin-1 (IL-1), cyclo-oxygenase (COX)-1, COX-2, and IL-6 (Riley 2008, Millar et al. 2010).  1.5.2. Histopathology  The main histological features of tendinopathy include collagen disorientation and degeneration without inflammation, hyper-cellularity and tenocytes with more round-shaped nuclei, ingrowth of new blood vessels amongst the collagen fibers, and elevated glycosaminoglycans and proteoglycans (Zafar et al. 2009). Animal models have shown mechanical 13  loading causes excessive proliferation of tenocytes, disruption of collagen fibers, and an increase in the noncollagenous matrix. These abnormalities cause tendons to be more prone to microrupture or partial tearing due to the loss of mechanical properties (Wang et al. 2006, Scott et al. 2007, Longo et al. 2009, Andersson et al. 2011). Some in vitro studies have shown that repetitive mechanical loading results in elevated production of factors by tenocytes which are sometimes characterized as inflammatory and/or degenerative (e.g. PGE2, MMP1 or -2, TGFβ) (Khan et al. 2005, Wang et al. 2006, Asundi et al. 2008). It has been reported that sites of subjectively defined pain, clinically palpated tenderness, tendon thickness and increased colour Doppler signal are anatomically associated, indicating a possible association between pain and neurovascular changes resulting from tendon overuse (Lewis et al. 2009, Divani et al. 2010). However, it must be acknowledged that the colour Doppler signal typically associated with tendinopathy may represent not only angiogenesis, but also increased blood flow in vessels which are already present. Angiogenesis may also be accompanied by neurogenesis; i.e, nerves may be proliferating along with neovessels in mechanically loaded tendon tissue increasing the level of substance P and other pain-producing substances in tendon. This histological change could lead to the transition to a symptomatic phase in tendinopathy (Gotoh et al. 1988).  1.5.3. Cytokines and growth factors Damage to tendon due to overuse or tendon rupture alters the expression and release of cytokines, pro-inflammatory factors and growth factors which may influence the course of tendon injury or healing (Millar et al. 2009, Gao et al. 2013). Mechanical stimuli induce tendon cells to release a number of cytokines such as VEGF, TNF-α, IL-6, PGE2, substance P and PDGF which 14  may promote angiogenesis in tendon tissue (Skutek et al. 2001, Skutek et al. 2001, Petersen et al. 2004, Fong et al. 2005, Yang et al. 2005, Backman et al. 2011, Gao et al. 2013, Legerlotz et al. 2013). Tendon fibroblast cells demonstrate endogenous expression of various cytokines and growth factors such as TNFα, IL-1β, IL-6, IL-10, VEGF ,TGF-β, bFGF, and PDGF(Skutek et al. 2001, Schulze-Tanzil et al. 2011).  These cytokines and their role in tendon are summarized in Table 1-2 (Schulze-Tanzil et al. 2011). Table 1-2. Cytokines and their effects in tendon tissue. Cytokine Effects in tendon/on tenocytes IL-1β ECM degradation (MMPs), induction of inflammatory mediators (IL-1β, TNFα, IL-6, COX-2, PGE2), suppression of type I collagen, induction of elastin, cytoskeletal changes TNFα ECM degradation (MMPs), induction of cytokines (IL-1β, TNFα, IL-6, IL-10), suppression of type I collagen, induction of elastin, SOCS1, pro- and anti-apoptotic effects, cytoskeletal changes IL-6 STAT3 phosphorylation, VEGF expression, supports tendon healing, induction of SOCS3 and of IL-10 IL-10 IL-10R1 induction (suppressing macrophage activation) VEGF Neo-angiogenesis, remodeling (MMP expression) TGFβ1 Fibronectin expression, tendon scar formation  15  The induced expression of growth factors in response to physical tension may be involved in the adaptation of connective tissues to increased mechanical loading (Kjaer et al. 2009). Some angiogenic factors may influence the course of degenerative tendon disease. For example, VEGF upregulates tenocyte expression of MMP1, 2 and 3 through induction of transcription factor Ets-1. MMPs are a family of enzymes which are involved in matrix degeneration in tendon tissue. The increased expression of proinflammatory cytokines and their receptors in tendinosis suggests the role of inflammatory response in tendon tissue damage in early stages. However, a lack of inflammatory cells and symptoms related to inflammation in tendinosis could challenge this notion (Dakin et al. 2014). Marked expression of IL-21R in early tendinopathy and its regulation in tendon cells by proinflammatory cytokines may contribute to the tissue damage in tendinopathy. Although IL-21 ligand is not expressed in tendon cells, the IL-21R might be activated by alternative binding ligands (Campbell et al. 2014).  1.5.4. MMPs It is acknowledged that matrix metalloproteinases (MMPs) contribute to the remodeling of the tendon extracellular matrix, and that their balance with MMP inhibitors is necessary for maintaining tendon homeostasis (Magra et al. 2005, Jones et al. 2006). Tendon degeneration is accompanied by changes in the expression and activity of matrix metalloproteinases (MMPs) which are consistent with increased proteolytic degradation of matrices. Several MMPs including MMP1, MMP2, MMP8, MMP12 and MMP14 have activity for the cleavage of fibrillar collagen. These MMPs may have a key role in tendon remodeling during both tendon healing and degeneration (Riley 2008). Mechanical loading has been shown to regulate the expression and 16  activity of MMPs (Archambault et al. 2002, Tsuzaki et al. 2003). Some studies have shown changes in MMP expression or activity in tendon disorders. However, the precise role of MMPs in tendinopathy is not clear. The expression of MMP-2 was upregulated in tendon biopsies taken from Achilles tendinopathy lesions and from ruptured Achilles tendons (Alfredson et al. 2003, Karousou et al. 2008). In isolated tenocytes from rat Achilles tendons, there was a positive correlation between MMP-2 activity and age (Yu et al. 2013). MMP-2 can modulate angiogenesis through cleavage of the FGF receptor and regulation of FGF-2 activity. MMP-2 also binds to integrin αvβ3 on epithelial cells. This interaction induces PI3K/AKT-mediated VEGF expression and subsequent angiogenesis in vitro (Bauvois 2012).  1.6. Role of neovascularization in tendinopathy Numerous studies have shown that tendinopathy often arises in the hypovascular zone of tendon such as the critical zone (Codman) in supraspinatus tendon; however, the actual site of the painful lesion, once it develops, generally shows an increased vascularity. It seems that the sites of subjectively defined pain, clinically palpated tenderness, tendon thickness and neovascularization are anatomically associated; so there may be an association between pain and neurovascular changes resulting from tendon overuse in tendinopathy patients (Pufe et al. 2005, Scott et al. 2008, Divani et al. 2010). Tendinopathic lesions in tendon and paratendon are associated with increased neovascularization and degenerative vascular abnormalities including vascular hyperplasia within tendon. Neovascularization also is an essential part of tendon healing during the proliferative phase; however hypervascularity in chronic tendinopathy is associated with tendon pain.  Several growth factors and cytokines including VEGF, bFGF and TGF-β 17  promote vascularization in tendon tissue but the factors in painful tendon lesions that lead either to ongoing angiogenesis, or a failure to retract new blood vessels, are unknown (Fenwick et al. 2002). Gene expression studies indicate the increased expression of TGF-β, VEGF, PDGF receptor and IGF-I in chronic tendinopathy which may contribute to the hypervascularity of tendon lesions (Riley 2008).  Hypervascularity could potentially disrupt the integrity of load-bearing tendon tissue, which may predispose tendons to tears and to tissue inflammation. In theory, factors that reduce the presence or persistence of new blood vessels or accompanying nerves in chronically painful tendon may relieve pain and improve the biomechanical function of tendon tissue. Indeed, some modalities for tendinopathy such as sclerosant injections and extracorporeal shock-wave therapy may treat painful tendon by targeting blood vessels (Riley 2008). The efficacy of these treatments has not been firmly established, however.  1.7. Mechanotransduction  Live cells are able to sense physical deformations and thereby react to mechanical forces. The mechanism by which cells transform the mechanical stimuli to biological and chemical responses is called mechanotransduction. Mechanical forces alter the conformation and dynamics of different cell components such as cytoskeletal and membrane proteins. These deformation-induced changes can modify the interactions between intracellular proteins and ligands to transduce the mechanical stimuli to biochemical signals (Paluch et al. 2015). The major mechanotransducers in cell membranes include mechanosensitive ion channels, caveolae, and 18  surface receptors. In cell-cell and cell-matrix contacts some proteins such as integrins, focal adhesions, cadherins and gap junctions also contribute to mechanotransduction (Ingber 2006).   In eukaryotic cells, several families of mechanosensitive channels have been identified. Mechano-gated potassium channels including TREK and TRAAK are mostly present in neurones. Stretch and mechanical stress can activate these channels and open them to the flow of potassium (Martinac 2004).  The amiloride-sensitive epithelial sodium channels (ENaC) and degenerins as the subclasses of ENaC/DEG superfamily are pore channels that are permeable to sodium or calcium and play a central role in touch sensation and proprioception (Mano et al. 1999). Transient receptor potential (TRP) channels also contribute to physical sensation.  These chancels are widely expressed in different tissue and are non-specifically permeable to cations such as calcium ions (Chatzigeorgiou et al. 2010).  Some mechanosensitive ion channels such as ENaC and calcium channels are adjacent to integrins (Shakibaei et al. 2003). Integrins are transmembrane proteins which bind to ECM and cytoskeletal proteins. Mechanical forces can transmit from integrins to the ion channels via cytoskeletal components or their direct interactions with the channels (Ingber 2006). Integrins link actin cytoskeleton to ECM proteins and form focal adhesions and complexes. Focal adhesions are an assembly of numerous proteins associated with integrins. Mechanical forces can change the conformation of focal adhesion proteins thereby activating integrins, focal adhesion kinases (FAK), Rho GTPases and some other signalling factors (Schwartz 2010). The signalling cascades triggered by integrins simulate downstream target proteins such as mDial1 and ho-associated kinase (ROCK) which change the dynamics of cytoskeleton filament polymerization. These changes can feed back to the conformation of focal adhesions and ECM protein. In this way, 19  alteration of intracellular filaments triggered by integrin activation promotes a bidirectional effect between integrins, ECM and cytoskeleton to govern some biological processes like cell migration (Ingber 2006).  Activation of signalling pathways by integrins also affects gene expression patterns and leads to changes in cell phenotype and ECM composition (Chiquet et al. 2003). Alteration of ECM composition and structure modifies the interaction between ECM components and some protein complexes. These modifications change the mechanical properties of the ECM, as well as the activity of protein complexes and their bioavailability to the resident cells (Schwartz 2010). In myofibroblasts, integrins transmit contractile forces produced by cytoskeletal fibers and the ECM to the latent TGF-β complex – this contractile force activates the latent complex through conformational changes (Wipff et al. 2008). Integrins also induce proteolytic activation of TGF-β by enhancing the interaction between latent TGF-β and the membrane-bound MMPs (Mu et al. 2002).  1.8. Mechanobiology of tendon cells The strain level measured in tendon tissue is dependent on the type of tendon and the physical activity. Wilson et al recorded the maximum average strain of 5.8% in Achilles tendon during running and strains of 6%-7% were recorded in the patellar tendon during the development of maximal isometric force; 15% peak strain is predicted for the patellar tendon during jumping (Lichtwark et al. 2006, Lavagnino et al. 2008, Couppe et al. 2009). The failure strain of Achilles tendon is 16.1% and 12.8% for fast (10% s-1) and slow (1% s-1) strain rates respectively (Wren et 20  al. 2001). In an in vitro model, the transmission of strain to individual cells in cell culture depends on their adhesion to substrate and usually is less than substrate strain (Wang et al. 2007).  1.9. Mechanical stimulus - effects on gene regulation Tenocytes comprise the main cell population (90%-95%) in tendon tissue, and they may be defined as scleraxis-expressing fibroblasts residing within the extracellular matrix of the tendon, and playing a key role in tendon development, adaption and the response to mechanical loading (Mendias et al. 2012). Tenocytes produce a variety of endogenous cytokines and growth factors which exert both autocrine and paracrine effects (Al-Sadi et al. 2011). Some in vitro studies have shown that repetitive mechanical loading of tendon cells results in an elevated production of soluble factors which are sometimes characterized as inflammatory, catabolic, or anabolic (Almekinders et al. 1993, Wang et al. 2003, Jones et al. 2013). Table 1-3 summarizes the studies of mechanical stimulation of isolated human tendon cells and their results.   21  Table 1-3. Mechanical stimulation of human tendon cells. Cell Source Mechanical Stimulus Parameters Outcome Ref normal finger flexor repetitive motion with 0.25 strain at 0.17 Hz (10 cycles.min-1), with/without 25 µM indomethacin, or 0.25 strain at 1 Hz (60 cycles.min-1). PGE2↑, LTB4↑in presence of indomethacin, LDH ↔ (Almekinders et al. 1993) normal upper extremities repetitive motion and nonsteroidal anti-inflammatory medication PGE2↑, protein synthesis and PGE2↓ in presence of nonsteroidal anti-inflammatory medication (Almekinders et al. 1995) healthy Achilles and hamstring tendons 10% equibiaxial strain with a frequency of 1 Hz for a total of 120 min/day for 3 days substance P↑, neurokinin-1 receptor↑, MMP3 (Backman et al. 2011, Fong et al. 2013) healthy patellar tendon  cyclic mechanical longitudinal strain frequency of 1 Hz and an amplitude of 5% proliferation↑, HSP 72↔, apoptosis ↑ (Barkhausen et al. 2003) 22  Cell Source Mechanical Stimulus Parameters Outcome Ref semitendinosus and gracilis  tendon  de-tensioning integrin subunit alpha11↓, alpha2 integrin↑, markers for tendon cell differentiation (TNMD, SCX, MKX, COL1A1, COL3A1, COL14A1, DCN, FBMD, CCN2)↓,  pro-inflammatory molecules (COX-1 and COX-2, IL1B, IL6) ↑, COL12A1↔, FN↔, TGFB1↔, (Bayer et al. 2014) healthy semitendinosus and gracilis tendons soft-focused extracorporeal shock wave treatment (0.17 mJ/mm2) cell viability, proliferation and tendon-specific markers (scleraxis and type I collagen) ↑; release of cytokines ( IL-1β, IL-6 and IL-10, TNFα) ↑, TGFβ↑, VEGF↑, MMP-3 and MMP13↔, nitric oxide↓ (de Girolamo et al. 2014) normal hamstring tendon continuous/rest-inserted cyclic equiaxial strain at a frequency of 0.1 Hz for 100 or 1000 cycles collagen type I, III↑; SCXA↑ (Huisman et al. 2014) 23  Cell Source Mechanical Stimulus Parameters Outcome Ref patellar tendon  biaxial strain with magnitude of 5% at a 1 Hz frequency for 30 or 60 min HSP-72↑ (Jagodzinski et al. 2006) patellar tendon   cyclically strain at 4% or 8% at 0.5 Hz for 8 h  collagen type I↑,  collagen type III↔, cell proliferation↑, apoptosis ↑ at 8% strain,  inflammatory factors (IL-2, IL-10, VEGF, IL-6, and TGFβ1)↑,  (Jiang et al. 2012) Achilles tendon uniaxial strain at 5% cyclic strain at 1 Hz for up to 48 h After 24 and 48 h strain: ADAMTS2, ADAMTS4, ADAMTS16, MMP24 and TIMP↑. After 24 h Strain: MMP10,  THBS1, ADAMTS6, and ADAMTSS14 ↑ After 48 h strain: MMP10, MMP11, Lumican, MMP3, and MMP17↓; ADAMTS10↑ (Jones et al. 2013) patellar tendon  uniaxial strain at 4%, 8%, 12% at 0.5 Hz, for 4 hours LTB4 and PGE2↑ at 8% and12%, 5-lipoxygenase ↔ (Li et al. 2004) 24  Cell Source Mechanical Stimulus Parameters Outcome Ref hamstring tendon  10% equiaxial cyclic strain with a frequency of 1 Hz for up to 24 hrs Transient ↑ ANGPTL4, FGF-2, COX-2, SPHK1, TGF-alpha, VEGF-A and VEGF-C;    (Mousavizadeh et al. 2014) flexor digitorum profundus tendon  uniaxial strain with 3.5% elongation at 1 Hz for 1 h/day for up to 5 days matrix proteins (col1a1, fibronectin, biglycan), cytokines and signaling factors (TGF-β1, COX2), and enzymes (MMP27, ADAMTS5) ↑ (Qi et al. 2011) healthy patellar tendon  biaxial cyclic strain with 5% elongation at 1 Hz up to 60 min release of IL-6↑; TGF-beta1, bFGF, and PDGF↔ (Skutek et al. 2001)  healthy  patellar tendon  biaxial cyclic strain with 5% elongation at 1 Hz up to 60 min transient JNK activation and apoptosis (Skutek et al. 2003)  flexor digitorum profundus tendon  cyclic strain (1 Hz, 0.035 strain, 2 h) ATPase activity↑ (Tsuzaki et al. 2005) 25  Cell Source Mechanical Stimulus Parameters Outcome Ref flexor digitorum profundus tendons equibiaxial cyclic strain (1 Hz, 3.5% elongation) for 2 h followed by an 18-h-rest period. IL-1beta, (COX 2), and MMP-3↑; MMP-1↔; ATP release↑ (Tsuzaki et al. 2003) patellar tendon  biaxial cyclic strain (5%, 1 Hz) for up to 60 min nitric oxide  ↑ (van Griensven et al. 2003) patellar tendon  cyclic strain at 4%, 8%, or 12%  with frequency of f 0.5 Hz for 24 hr PGE2, COX-1 and COX-2↑; ↓ COX-2, MMP-1 and PGE2 stimulated by IL-1β, (Wang et al. 2003, Yang et al. 2005) patellar tendon  uniaxial cyclic strain with 8% elongation at 0.1, 0.5 and 1 Hz for 4 hrs PGE2↑, cytosolic phospholipase-A2↑; COX1, COX2 ↑ (frequency dependent); phospholipase-A2 activity ↑ (Wang et al. 2004) patellar tendon  uniaxial cyclic strain, 8%,  (0.5 Hz) for up to 72 hrs phospholipase-A2 activity and α-SMA protein ↑ in a cell orientation dependent (Wang et al. 2004, Wang et al. 2005) 26  Cell Source Mechanical Stimulus Parameters Outcome Ref patellar tendon  uniaxial cyclic strain with 4%, and to 8%, (0.5 Hz, 4 h) cell proliferation ↑, collagen type I and TGF-β1↑, collagen type III↔ (Yang et al. 2004) posterior tibial tendon  cyclic strain of 0% or 5% for 24 hr. Type I collagen and decorin↑ (Chen et al. 2007) supraspinatus tendon cyclic strain with 1% elongation daily for 5 days gap junction protein connexin 43 and intracellular calcium concentration ↑ in presence of steroid  (Triantafillopoulos et al. 2004) human tenocytes cyclic strain (5% at 1Hz) for up to 48 hrs ITGA5 and ITGB1↓;  (Jones et al. 2014) ↓: downregulation; ↑: upregulation; ↔: no changes    27  1.10. Substance P (SP) SP is historically known to be a pain-signalling molecule and since its presence in tendon fibroblasts was confirmed, it has been suggested as a source of pain in tendinopathy. This speculation was further strengthened when the expression of SP was shown to be up-regulated in the state of tendinosis, in which increased nerve sprouting is also seen. This makes the mechanism of locally produced SP on the efferent pain signalling possible, considering the fact that the sprouting nerves do express SP´s preferred receptor, NK-1R (Schubert et al. 2005, Andersson et al. 2011). In the rotator cuff, pain levels are remarkably correlated with tissue levels of SP (Gotoh et al. 1998, Fredberg et al. 2008). Interestingly, although peripheral SP-containing nerves are known to exist and are more extensive in chronically injured tendon, local tendon fibroblasts also appear to be a local source of SP; this finding is in keeping with recent studies demonstrating that SP is locally up-regulated by repetitive overuse in tendon fibroblasts (Andersson et al. 2008, Backman et al. 2011, Backman et al. 2011). SP also regulates the expression of MMPs and TIMP in fibroblast cells and tendon cells (Cury et al. 2008, Fong et al. 2012). SP induces hypercellularity and angiogenesis in tendon tissue (Burssens et al. 2005, Andersson et al. 2011).  1.11. TGF-β signaling pathway Transforming growth factor-β (TGF-β) is a secretory protein released by various cell types as a latent form which, following its activation, modulates extensive biological activities such as cell proliferation and differentiation, apoptosis, inflammation, healing, development, etc. Malfunction of the TGF-β signalling pathway plays a key role in the pathology of many disorders, extending from genetic syndromes to inflammatory and metabolic disorders. TGF-β is produced 28  as three isoforms, TGF-β1, TGF-β2 and TGF-β3, which are secreted into the ECM as a Large Latent Complex (LLC). LLC consist of TGF-β homodimer, Latency Associated Peptide (LAP) and Latent TGF-β-Binding Protein (LTBP) (Figure 1-2.). This complex is activated by various factors such as MMPs, plasmin, thrombospondin-1 (TSP-1), integrins, reactive oxygen species (ROS) and acidic pH which liberate TGF-β from LAP through degradation, denaturation, conformational change or breaking disulfide linkage of LAP (Annes et al. 2003). Binding of active TGF-β dimer to type II receptor promotes recruitment and phosphorylation of type I receptor which induces SMAD phosphorylation and activates SMAD-dependent signalling pathway (Figure 1-3.) (Derynck et al. 2003).  In mammals, seven type I receptors and five type II receptors have been identified which differentially bind to 79 different ligands. Combination of different receptors and ligands define a complex SMAD-mediated responses (Derynck et al. 2003). TGF-β proteins and other members of TGF-β superfamily including BMPs, activins and related protein, also activate other signalling pathways in addition to the SMAD signalling pathway, adding another layer of complexity to the TGF-β related responses. Studies haves shown that TGF-β can activate MAPK pathways through SMAD or some independent mechanisms which differ in slow or rapid activation of MAPK-mediated gene transcription.  TGF-β-activated kinase 1 (TAK1) known as MAPK kinase kinase-7 (MAPKKK-7) is activated by both TGF-β and BMP-4. TAK1, MAPKKK-1 and Rac can activate JNK, p38 MAPK and Erk MAPK pathways. TAK1 also induces NF-κB signalling pathway through phosphorylation of IκB (Derynck et al. 2003).  29    Figure 1-2. Large Latent Complex.   Figure 1-3. TGF-β Signalling Pathway. 30  1.12. Function of TGF-β in tendon tissue The TGF-β superfamily, including Bone Morphogenic Protein (BMP) and TGF-β subfamilies, has a prominent role in tendon development and healing. Induced expression of tendon cell markers such as Scx during tendon development is mediated by TGF-β signalling – this signalling is essential for maintenance of tendon progenitor cells and the recruitment of new cells. Therefore, disruption of the signalling pathway provokes loss of tendon tissue (Pryce et al. 2009).  TGF-β plays a pivotal role in tendon repair and injury due to its diverse functions in collagen deposition, angiogenesis, adhesion and scar formation. TGF-β stimulates collagen synthesis. TGF-β also induces the expression of Scx, a gene which regulates the expression Col1a1 in tenocytes.  Different activities of TGF-β isoforms during tendon healing have been reported. In addition, the expression of TGF-β receptors changes during reparative phases. The TGF-β signalling pathway may also regulate the expression pattern of collagen type I and III during the course of tendon repair (Chan et al. 2008).  The expression levels of TGF- β isoforms and receptors are also altered in chronic tendon lesions, implying a role for the TGF- β pathway in this pathology (Fenwick et al. 2001). It seems that a balance of mechanical forces is essential for normal activity of TGF-β in tendon to support tissue homeostasis. Otherwise, excessive, or lack of, mechanical loading can disturb the physiologic activity of TGF- β and may cause consequent damage (Maeda et al. 2011).    31  1.13. Role of HIF-1 in tendon injury HIF-1 is a well-studied transcription factor that induces an array of genes in response to hypoxic conditions to modulate various biological pathways including angiogenesis, invasion, metastasis, apoptosis and some metabolic pathways. The protein complex belongs to the basic helix-loop-helix (b-HLH) family of transcription factors and comprises a heterodimer of α and β subunits constantly expressed by different cell types. These subunits share some similar domain structures that control their activity and dimerization including b-HLH, PAS named after PER (period circadian protein)-ARNT (aryl hydrocarbon receptor nuclear translocator protein)-SIM (single-minded protein), and transactivation stimulation of transcription (TAD). HIF-1 α also has an oxygen-dependent degradation domain (ODDD) that controls its stability (Figure 1-4.). The activity and stability of HIF-1α is determined by post-translational modifications such as hydroxylation, ubiquitination, acetylation, and phosphorylation; these post-translational modifications are regulated by prolyl hydroxylase domain (PHD), factor inhibiting HIF-1 (FIH-1) and Von Hippel–Lindau tumor suppressor (VHL), respectively, and mitogen-activated protein kinase (MAPK). Phosphorylation activates HIF-1 α whereas other modifications promote inactivation and degradation of the protein in the presence of oxygen (Ke et al. 2006).   32   Figure 1-4. Structure of HIF-1 subunits and their functions.  HIF-1α, as an inducible intracellular transcription factor, is increased in response to hypoxic conditions and modulates angiogenic factors such as VEGF. Tendon is a more oxygen-dependent tissue compared to other joint tissues such as cartilage (Sharma et al. 2005). Factors and conditions that stimulate the metabolic activity of tendon cells may increase the oxygen demand and eventually lead to hypoxic conditions, in part due to the hypovascular nature of the tendon tissue. Torn rotator cuff tendon shows increased HIF-1α and VEGF expression compared to intact tendon tissue, which may cause increased neovascularity. These changes in gene expression could reflect a hypoxic condition in tendon tissue that may also promote the degenerative process and cell apoptosis (Benson et al. 2010, Lakemeier et al. 2010). Cultured tenocytes rapidly upregulate HIF-1α followed by increased apoptotic proteins in response to hypoxia which lead to significant cell death. All VEGF isoforms also were induced by hypoxia. But adding growth factors from PRP to these cells protected them from apoptosis (Liang et al. 2012). Hypoxic conditions are proposed as a cause of induced Nuclear factor-κB (NF-κB) activity in torn rotator cuff tendons – this activity is correlated with increased neaoangiogenesis (Gumina et al. 2013). Hypoxia also induces the expression of inflammatory cytokines, apoptosis and alters collagen matrix regulation in tencoytes. 33  This evidence demonstrates the potential role of hypoxia as an initiator factor in promoting early tendinopathy (Millar et al. 2012). Other stimulus that may induce HIF-1α is mechanical stress. Repetitive mechanical strain induces HIF-1α and VEGF- expression in cultured tendon cells (Petersen et al. 2004).   1.14. Angiopoietin-like 4 1.14.1. Function of ANGPTL4 ANGPTL4 belongs to a superfamily of secreted proteins which regulate angiogenesis. Although there is a structural similarity with angiopoietins, ANGPTL proteins are orphan ligands and they do not bind to the receptor tyrosine kinases Tie 1 or Tie 2. The proteolytic cleavage of native ANGPTL4 protein generates two fragments including an N-terminal coiled-coil fragment (nANGPTL4) and a C-terminal fibrinogen-like domain (cANGPTL4) that have different physiological and pathological functions. The nANGPTL4 fragment is mostly involved in metabolic hemostasis; while cANGPTL4 modulates angiogenesis (Figure 1-5.). Shortly after the discovery of ANGPTL4 as a factor involved in lipid metabolism (known as fasting–induced adipose factor), the function of this protein has been determined in many other metabolic and non-metabolic pathways that modulate energy homeostasis, wound healing, oncogenesis, metastasis, inflammation, lymphangiogenesis and angiogenesis. Recent studies showed that ANGPTL4 provokes the disruption of vascular junction integrity via integrin α5β1-mediated Rac/PAK signaling and the de-clustering and internalization of VE-caderin and claudin-5 which eventually induces vascular leakiness and permeability (Le Jan et al. 2003, Morisada et al. 2006, Gealekman et al. 2008, Hato et al. 2008, Huang et al. 2011). ANGPTL4, induced by hypoxia and high glucose, 34  promotes vessel permeability and angiogenesis in ischemic and diabetic retinopathies (Xin et al. 2013, Yokouchi et al. 2013, Kwon et al. 2015).  Tissue damage due to inflammation in some disorders is meditated by induced expression of ANGPTL4. Lung inflammation due to pneumonia infection and LPS stimulates the expression of ANGPTL4 followed by increased pulmonary tissue leakiness and damage. Therefore silencing or neutralizing ANGPTL4 can protect lung tissue from further damage and improve recovery (Guo et al. 2015, Li et al. 2015). By contrast, ANGPTL4 plays a protective role against inflammation following intake of saturated fat. ANGPTL4 inactivates LPL to block fatty acid uptake into mesenteric lymph node macrophages which decreases foam cell formation and inflammatory gene expression (Lichtenstein et al. 2010).   Figure 1-5. Structure and dual function of ANGPTL4.  1.14.2. Regulation of ANGPTL4 ANGPTL4 is modulated by a variety of factors and receptors, including glucocorticoid receptor (GR), hypoxia inducible factor 1 (HIF-1), prostaglandin E receptor 1, transforming growth factor β (TGFβ) and peroxisome proliferator-activated receptors (PPARs). Different conditions such as fasting, glucocorticoid therapies, hypoxia and treatment with PPAR agonists 35  stimulate ANGPTL4 expression [36-43]. The crosstalk between COX-2-derived prostaglandin E2 (PGE2) and hypoxia synergistically induces ANGPTL4 expression in colorectal cancer, leading to tumour growth by enhancing cell proliferation. Recent findings also suggest the crosstalk of TGFβ and PPAR in stimulation of ANGPTL4; however contrasting results showed either synergistic or antagonistic induction [43-45]. Metabolic conditions that alter the level of circulating free fatty acids and triglycerides would change the synthesis and release of ANGPTL4.  Kersten and colleagues showed that circulating free fatty acids upregulate ANGPTL4 protein in cardiac tissue and immobilised muscles, thereby reducing the local uptake of plasma triglyceride-derived fatty acids (Kersten et al. 2009, Georgiadi et al. 2010, Catoire et al. 2014). High glucose also induces the release of ANGPTL4 from retinal pigment epithelium cells (Yokouchi et al. 2013).  Metabolic conditions such as diabetes and metabolic syndrome increase ANGPTL4 plasma level in association with the inflammatory response (Tjeerdema et al. 2014). The activators of toll like receptors by LPS induces the expression of ANGPTL4 in adipose, muscle and cardiac tissues – this induction is mediated by TLR4 signaling pathways including NF- B and p38 MAPK (Brown et al. 2009).  Inflammation in lung tissue in response to infection with influenza Pneumonia induces the IL6/STAT pathway, thereby promoting the expression of ANGPTL4 (Brown et al. 2009).  36  1.15. Specific aims and hypotheses 1.15.1. Study I: Evaluation of the effect of cyclic stretching on the expression of angiogenic factors by tendon cells The objective of this study was to determine the effect of repetitive stretching on expression and release of angiogenic factors from tendon cells that may affect the vascularity of tendon tissue. I also investigated the regulatory pathways that may govern the expression of mechanoresponsive genes. For this purpose I addressed the following broad question: Which angiogenic factors are induced in response to repetitive stretching? I hypothesized that:  The early response of tendon cells to repetitive stretching leads to upregulation of a specific subset of angiogenic factors.  1.15.2. Study II: Investigation of the mechanisms that regulate ANGPTL4 The overall aim of this study was to investigate the regulatory pathways that modulate ANGPTL4 expression, particularly in response to cyclic stretching. The main question to be addressed by this study was:  How does cyclic stretching of tendon cells regulate ANGPTL4? 37  I hypothesize that: Cyclic stretching increases the expression and release of angiogenic factors through induced activity of transforming growth factor beta (TGF-β) and stabilization of hypoxia inducible factor-1 (HIF-1) (Figure 1-6.).  Figure 1-6. Hypothetical function of ANGPTL4 and related pathways in tendon.  1.15.3. Study III: Characterizing the function of ANGPTL4 protein in tendon tissue Some inter-related in vitro and in vivo models were used to address the following question: What is the function of ANGPTL4 in promoting vascular changes during tendon healing?  I hypothesized that: ANGPTL4 protein promotes angiogenesis following tendon injuries and thus enhances the healing process.  38  Chapter 2. Materials and methods 2.1. Animal studies ANGPTL4 +/+, +/- and -/- mice were generated by inbreeding of ANGPTL4 +/- mice, B6;129S5-Angptl4Gt(OST352973)Lex/Mmucd, obtained from the Mutant Mouse Regional Resource Center  (MMRRC, 032147-UCD). Mice were genotyped acceding to MMRRC protocol. The acute patellar tendon (PT) injury was performed using a 0.30 mm True-Cut Disposable Biopsy Punch (Robbins Instruments, US,  # RBP-030) as previously described (Scott et al. 2011) and wound was sutured. The left uninjured PT was used as control. PTs were harvested after 3 days, 3 weeks and 12 weeks post injury.  5 µg recombinant mouse ANGPTL4 protein (R&D Systems, 4880-AN) reconstituted in 10µl PBS were injected into right PTs of female CD-1 mice (Charles River, St. Constant, Canada) and left PTs were injected with 10µl PBS as control. PTs were harvested 3 days after the injection. All animals were between 8-16 weeks and 25-37 grams at the beginning of the experiments.    2.2. Primary tendon cell culture Primary human tendon cells were isolated from healthy hamstring (semitendinosus) tendons (excess anterior cruciate ligament autograft material) of male and female patients (n=4, mean age 25.75 with SEM ±5.75 years). The tendon biopsies were minced into 3-5 mm pieces and digested by 1.5 mg/mL Collagenase D (Roche Applied Science, Switzerland, #11088866001) for 20 minutes in a shaker incubator (200 rpm) at 37˚C followed by incubation with 0.25% trypsin (TrypLE™, Life Technologies™, USA, #A1217702) for 3 minutes. After washing with PBS, the 39  digested tissues were cultured in high glucose Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum, 2 mm L-glutamine, 100 units/mL penicillin, and 100 μg/mL streptomycin in a humidified incubator containing 5% CO2 at 37°C. After tendon cell adherence to the tissue culture plates, the cultured cells (at 70% confluence) were subcultured 1:3 up to five passages to obtain adequate cells. Primary mouse tendon cells were isolated from harvested tendons from tails. The tendon explants were cultured in same conditions as human tendon cells. Several days after migration of tendon cells from explants, the tissues were discarded and culture cells were expanded. The isolated tendon cells were incubated with DMOG (EMD Millipore, 400091), recombinant human TGF-β1 (R&D Systems, 240-B-002), chetomin (Tocris, 4705) or A-83-01 (Tocris, 2939).  2.3. Ethics  Because our aim was to examine the potential etiological events of tendinopathy resulting primarily from tensile overload, we elected to use tendon cells from normal (healthy) donors, which necessitated the use of orthopaedic autograft material (semitendinosis tendon). This excess tendon material would otherwise have been discarded. The study was reviewed and approved by the UBC Clinical Research Ethics Board, and each patient provided written informed consent. The primary HUVEC cells were isolated from normal umbilical cords under a UBC approved human ethics certificate.  40  2.4.Mechanical stimulation of tendon cells The isolated human tendon cells were seeded on 6 well BioFlex® Culture Plates coated with collagen type I (Flexcell® International Corporation, USA, # BF-3001C) with a density of 1.2×105 cells per well with high glucose DMEM supplemented with 10% fetal bovine serum, 2 mM L-glutamine, 100 units/ml penicillin, and 100 μg/mL streptomycin. By using FX-4000T™ FlexLink® Starin Unit (Flexcell® International Corporation, USA), isolated tendon cells were exposed to equibiaxial (radial and circumferential) cyclic stretching (1Hz frequency, 10% stretch) for up to 24 hours. This stretch level has previously been shown to be well tolerated by human tendon cells (Backman et al. 2011).  2.5. Flow cytometry The viability of tendon cells after applied cyclic stretching was assessed with propidium iodide staining, as previously described (Scott et al. 2005). The stretched and non-stretched tendon cells were trypsinized and centrifuged along with their supernatant in individual tubes. The cell pellets were washed in PBS and fixed with ice cold 70% ethanol for 1 hour. Then the fixed cells were stained with propidium iodide staining solution [1mL PBS (Ca+2, Mg+2 free, 0.1% glucose), 10 µL RNase A 100 mg/mL and 5 µL propidium iodide 10 mg/mL] and incubated for minimum 30 minutes. The DNA content in suspended cells was analyzed using the FL3 channel on a flow cytometer, Coulter Epics XL- MCL (Beckman Coulter Inc, USA), and cell death was calculated based on cell populations in the sub G1 phase of the cell cycle. 41  2.6. Cell proliferation Conditioned media (DMEM, 5% FBS) of stretched or non-stretched tendon cells were harvested after 24 hours and stored at -80°C. Primary human umbilical vein endothelial cells (HUVECs, kindly provided by Dr. Aly Karsan and isolated as previously described (Karsan et al. 1998)) were seeded in 96 well plates at a density of 2×104 cells/well and incubated with the conditioned media of stretched or non-stretched tendon cells for 24 hours. At this time point cells were assessed with the MTS assay using a modified protocol for CellTiter 96® Aqueous MTS reagent powder (Promega, USA, G1111) and PMS (Sigma, USA, #P9625). 20 µL of MTS/PMS solution were added to the culture media of cultured cells in 96 well plate. After 3 hours’ incubation at 37˚C in a humidified 5% CO2 incubator, the absorbance at 490nm was recorded using an Epoch Microplate Spectrophotometer (BioTek, US).  2.7. Endothelial cell tube formation assay In order to evaluate the angiogenic activity of released factors from tendon cells subjected to cyclic stretching, a tube formation assay was used (modified from (Arnaoutova et al. 2010)). The conditioned media of the stretched and non-stretched tenocytes were harvested after 24 hours and stored at -80°C. Flat bottom 96 well plates were coated with 50 µl of Matrigel (BD Matrigel™ Basement Membrane Matrix, USA, #356231). The HUVEC cells were resuspended in the conditioned media of tendon cells which had been stretched or non-stretched (controls) for 24 hours. Then 100 µl of the resuspended cells (2×104 cells) were added to each well. After 6 hours’ incubation, the tubular networks which form in the matrigel in each well were micrographed using a digital camera (AxioCam ICm 1, Zeiss, Germany) attached to an inverted microscope (Axio 42  Observer.A1, Zeiss, Germany) with 5x objective lens. In some experiments, the tubular networks were stained with Calcein AM (Trevigen, USA, #4892-010-K) and visualized by fluorescence microscopy [18]. Then The TIF format grey-scale images of biological and technical replicates were analyzed with AngioTool software in order to measure the total tube length per field (Zudaire et al. 2011). The angiogenic activity of recombinant human ANGPTL4 (R&D Systems, 4487-AN) was also measured by tube formation. The recombinant protein was added to matrigel and media of endothelial cells at a concentration of 10 µg/ml. After 6 hours incubation the tubular networks of endothelial cells were stained with Calcein AM (Trevigen, USA, #4892-010-K) to be visualized by fluorescence microscopy. Total tube length in micrographs were measured with AngioTool software and data were reported as pixels per field.  2.8.Gene expression analysis Total RNA was extracted and converted to cDNA as explained above in the section titled “cell characterization.” The cDNA templates constructed from RNA samples of stretched tendon cells for 4 and 12 hours and non-stretched cells were used to profile 84 key genes (Table 2-1) involved in modulating the biological processes of angiogenesis with a Human Angiogenesis RT² Profiler™ PCR Array (SA Biosciences, USA, # PAHS-024Z). A three-fold change in gene expression was chosen as a cut off for selecting genes for further analysis. Then conventional qPCR was used to confirm and further analyze the expression pattern of stimulated genes after 1, 2, 4, 6, 8, 12 and 24 hours cyclic stretching. The qPCR data were represented as fold change compared to non-stretched tendon cells.  Gene expression changes from samples harvested at 1, 2, 43  and 4 hours cyclic stretching were grouped as early response genes, and gene expression changes from samples harvested at 6, 8, 12 and 24 hours stretching were grouped as late response genes. We also analyzed the expression of several of the most well-studied angiogenic factors already suspected to play a role in tendons including VEGFs and bFGF. The primer sequences used in conventional qPCR are shown in Table 2-2. The qPCR primers used in this study for detecting mouse genes are listed in Table 2-3. To extract RNA from mouse tendon tissue, mouse patellar tendons were harvested and stored in liquid nitrogen. The frozen tissues were homogenised in a Mikrodismembrator (Sartorius, Germany) and immediately incubated with Trizol followed by chloroform to form a biphasic solution for harvesting the RNA fraction. Total RNA was purified using High Pure RNA Isolation Kit (Roche, Germany, #11828665001) and used as template for cDNA synthesis with High Capacity cDNA Reverse Transcription Kit (Applied Biosystems™, USA, #4368814).  Table 2-1. Gene symbols and description of angiogenic factors analyzed by Human Angiogenesis RT² Profiler™ PCR Array Symbol Description AKT1 V-akt murine thymoma viral oncogene homolog 1 ANGPT1 Angiopoietin 1 ANGPT2 Angiopoietin 2 ANGPTL3 Angiopoietin-like 3 ANGPTL4 Angiopoietin-like 4 ANPEP Alanyl (membrane) aminopeptidase 44  Symbol Description BAI1 Brain-specific angiogenesis inhibitor 1 CCL11 Chemokine (C-C motif) ligand 11 CCL2 Chemokine (C-C motif) ligand 2 CDH5 Cadherin 5, type 2 (vascular endothelium) COL18A1 Collagen, type XVIII, alpha 1 COL4A3 Collagen, type IV, alpha 3 (Goodpasture antigen) CXCL1 Chemokine (C-X-C motif) ligand 1 (melanoma growth stimulating activity, alpha) CXCL10 Chemokine (C-X-C motif) ligand 10 CXCL3 Chemokine (C-X-C motif) ligand 3 CXCL5 Chemokine (C-X-C motif) ligand 5 CXCL6 Chemokine (C-X-C motif) ligand 6 (granulocyte chemotactic protein 2) CXCL9 Chemokine (C-X-C motif) ligand 9 TYMP Thymidine phosphorylase S1PR1 Sphingosine-1-phosphate receptor 1 EFNA1 Ephrin-A1 EFNA3 Ephrin-A3 EFNB2 Ephrin-B2 EGF Epidermal growth factor ENG Endoglin EPHB4 EPH receptor B4 EREG Epiregulin 45  Symbol Description FGF1 Fibroblast growth factor 1 (acidic) FGF2 Fibroblast growth factor 2 (basic) FGFR3 Fibroblast growth factor receptor 3 FIGF C-fos induced growth factor (vascular endothelial growth factor D) FLT1 Fms-related tyrosine kinase 1 (vascular endothelial growth factor/vascular permeability factor receptor) HAND2 Heart and neural crest derivatives expressed 2 HGF Hepatocyte growth factor (hepapoietin A; scatter factor) HIF1A Hypoxia inducible factor 1, alpha subunit (basic helix-loop-helix transcription factor) HPSE Heparanase ID1 Inhibitor of DNA binding 1, dominant negative helix-loop-helix protein ID3 Inhibitor of DNA binding 3, dominant negative helix-loop-helix protein IFNA1 Interferon, alpha 1 IFNB1 Interferon, beta 1, fibroblast IFNG Interferon, gamma IGF1 Insulin-like growth factor 1 (somatomedin C) IL1B Interleukin 1, beta IL6 Interleukin 6  IL8 Interleukin 8 ITGAV Integrin, alpha V (vitronectin receptor, alpha polypeptide, antigen CD51) ITGB3 Integrin, beta 3 (platelet glycoprotein IIIa, antigen CD61) 46  Symbol Description JAG1 Jagged 1 KDR Kinase insert domain receptor (a type III receptor tyrosine kinase) LAMA5 Laminin, alpha 5 LECT1 Leukocyte cell derived chemotaxin 1 LEP Leptin MDK Midkine (neurite growth-promoting factor 2) MMP2 Matrix metallopeptidase 2 (gelatinase A, 72kDa gelatinase, 72kDa type IV collagenase) MMP9 Matrix metallopeptidase 9 (gelatinase B, 92kDa gelatinase, 92kDa type IV collagenase) NOTCH4 Notch 4 NRP1 Neuropilin 1 NRP2 Neuropilin 2 PDGFA Platelet-derived growth factor alpha polypeptide PECAM1 Platelet/endothelial cell adhesion molecule PF4 Platelet factor 4 PGF Placental growth factor PLAU Plasminogen activator, urokinase PLG Plasminogen PLXDC1 Plexin domain containing 1 PROK2 Prokineticin 2 47  Symbol Description PTGS1 Prostaglandin-endoperoxide synthase 1 (prostaglandin G/H synthase and cyclooxygenase) SERPINF1 Serpin peptidase inhibitor, clade F (alpha-2 antiplasmin, pigment epithelium derived factor), member 1 SPHK1 Sphingosine kinase 1 STAB1 Stabilin 1 TEK TEK tyrosine kinase, endothelial TGFA Transforming growth factor, alpha TGFB1 Transforming growth factor, beta 1 TGFB2 Transforming growth factor, beta 2 TGFBR1 Transforming growth factor, beta receptor 1 THBS1 Thrombospondin 1 THBS2 Thrombospondin 2 TIMP1 TIMP metallopeptidase inhibitor 1 TIMP2 TIMP metallopeptidase inhibitor 2 TIMP3 TIMP metallopeptidase inhibitor 3 TNF Tumor necrosis factor TNFAIP2 Tumor necrosis factor, alpha-induced protein 2 VEGFA Vascular endothelial growth factor A VEGFC Vascular endothelial growth factor C 48  Table 2-2. Oligonucleotide sequence of primers and amplicon sizes of selected angiogenic genes.   Target gene Forward primer sequence Reverse primer sequence Amplicon size (bp) ANGPTL4 CTCCCGTTAGCCCCTGAGAG AGGTGCTGCTTCTCCAGGTG 140 Cox-2 CAGGGTTGCTGGTGGTAGGA GCATAAAGCGTTTGCGGTAC 119 FGF-1 GAAGTTTAATCTGCCTCCAGGGAAT CCCCCGTTGCTACAGTAGAG 63 FGF-2 CGGGTGCCAGATTAGCGG GGGTTCACGGATGGGTGT 114 TGFA CCTTGGAGAACAGCACGTC CACATGCTGGCTTGTCCTC 147 SPHK1 CTTCACGCTGATGCTCACTG GTTCACCACCTCGTGCATC 124 VEGFA CCTCCGAAACCATGAACTTT CCACTTCGTGATGATTCTGC 132 VEGFC GCCCCAAACCAGTAACAATC GCTGGCAGGGAACGTCTAAT 109 GAPDH TCTTTTGCGTCGCCAGCCGAG TGACCAGGCGCCCAATACGAC 94 49  Table 2-3. Oligonucleotide sequence of primers and amplicon sizes of selected mouse genes.  *(Ogawa et al. 2005); **(Mendias et al. 2008)Target gene Forward primer sequence Reverse primer sequence Amplicon size (bp) CD31 CAAGCAAAGCAGTGAAGCTG CTAACTTCGGCTTGGGAAAC 146 CD34 ATCCGAGAAGTGAGGTTGGC CAGGGAGCAGACACTAGCAC 156 FLK-1 CTGTGGCGAAGATGTTTTTG TTCATCCCACTACCGAAAGC 163 MMP-3 GGAAATCAGTTCTGGGCTATACGAGG CCAACTGCGAAGATCCACTGAAGAAG 301* MMP-13 TCTTTATGGTCCAGGCGATGA ATCAAGGGATAGGGCTGGGT 82 VEGF-A TTACTGCTGTACCTCCACC ACAGGACGGCTTGAAGATG 189  TNMD TGACTTTAAAAATGGATACACTGGC TCTGCGGGAACCCAAATCAC 171 SCX CCAGCGAAGAACTCATACAGC GGACACCCCTTCTACGTTGT 105**  COL1A2 CGGGATCAGTACGAAAGGGC TGAGCAGCAAAGTTCCCAGTA 178 DCN GCAGTGTTCTGATCTGGGTTTG TGCCTCTGGACTGATTTTGCT 178 Nucleostemin GAGGAATCTGACGAGCCCAA TCTGAGGCTTCAATCACCTTT 101 LOX TACTTCCAGTACGGTCTCCC AGTCTCTGACATCCGCCCTA 151  COL3A1 AGATAAGGGTGAAGGTGGTTCC CCTGGTTCACCATTCTGTCC 135 GAPDH TCACCACCATGGAGAAGGC GCTAAGCAGTTGGTGGTGCA 169 50  2.9. Zymography After 2 hours of cyclic stretching, followed by 6 hours incubation without stretch, the serum-free conditioned media of tendon cells were harvested and stored at -80°C. Zymography assays were carried out following a modified protocol described by Wong et al (Wong et al. 2007). The conditioned medium was concentrated with YM-3 Centricon membranes (Millipore, Billerica, MA) at 7000g for 4 h at 4°C. 5 μg of protein were loaded on a 10% polyacrylamide gel containing 0.1% gelatin (Sigma, Canada) for electrophoresis. Following electrophoresis, the gel was incubated in Triton X-100 exchange buffer including 20 mM Tris–HCl (pH 8.0), 150 mM NaCl, 5 mM CaCl2 and 2.5% Triton X-100. After 30 min incubation followed by three 10 min washes with the incubation buffer (the exchange buffer without Triton X-100), the gel was placed in incubation buffer and incubated overnight at 37°C. Then the incubation buffer was removed and the gel was stained with 0.5% Coomassie blue R250 (Sigma) for 1 hour. The gel was then washed with 30% methanol and 10% acetic acid for 1 hour for de-staining. Gelatinolytic activity of MMP-2 (gelatinase A) was identified as clear areas in the gel at the apparent molecular weight of ~72 kD.  2.10. Immunoblotting  Total protein was harvested in lysis buffer (50 mM Tris-Cl, pH 7.7; 1% Triton X-100; 10% glycerol; 100mM NaCl, 2.5mM EDTA, 10mM NaF) (Hojabrpour et al. 2012) supplemented with cOmplete™ protease inhibitor cocktail (Roche, Germany, #04693124001) and then homogenized by 3 sonication cycles with 25 watts power output at frequency of 23 kHz for 5 seconds on ice and with 10 seconds interval.  The homogenates were spun at 13,000g for 10 min. The protein 51  concentrations of the supernatants were measured using the BCA Protein Assay Kit (Pierce, ThermoScientific, USA, #23225). The protein was heated for 3 min in 5x loading buffer (Fermentas, Lithuania, #R0891). 20 μg of protein from each aliquot was resolved by electrophoresis in a 12% SDS-PAGE gel at a constant voltage of 110 volts. For the detection of secreted ANGPTL4, 20 µg protein of concentrated conditioned media from  stretched and non-stretched tendon cells were used. The stretched cells were subjected to 2 hours of cyclic stretching, followed by 6 hours incubation without stretching and the serum-free conditioned media of tendon cells were harvested and stored at -80°C. The serum free conditioned media of non-stretched tendon cells were used as a control. The conditioned media were concentrated with YM-3 Centricon membranes (Millipore, USA, #4302) at 7000g for 4 h at 4°C. The resolved proteins were transferred to a 0.45 µm nitrocellulose membrane (Biorad, Germany, #162-0115) in cold transfer buffer (25 mM Tris, 192 mM glycine, 20% methanol) at 35 volts for 16h. The membranes were blocked with 5% milk in Tris-buffer saline with 0.5% Tween 20 (TBST), and probed with anti-ANGPTL4 Antibody (Novex®, Life Technologies, USA, #710186) in TBS-T overnight at 4°C, followed by 3 × 10-min TBST washes, followed by Goat Anti-Rabbit IgG, H & L Chain Specific Peroxidase Conjugate (Calbiochem®, USA, # 401315) in TBST. Immunoreactivity was detected using SuperSignal™ West Femto Chemiluminescent Substrate (Thermo Scientific, USA, #PI34095). The density of detected bands was quantified using Image J, and vinculin was used as the reference protein for data normalization.    52  2.11. Enzyme-Linked Immunosorbent Assay (ELISA) The conditioned media of tendon cells cultured in high glucose DMEM supplemented with 10% fetal bovine serum, 2 mM L-glutamine, 100 units/ml penicillin, and 100 μg/ml streptomycin were harvested after 6 hours cyclic stretching and stored at -80°C. The conditioned media of non-stretched tendon cells were used as control. The protein levels of ANGPTL4 in the samples were quantified using a commercial ELISA kit, Human Angiopoietin-like 4 DuoSet (R&D Systems, USA, #DY3485) according to the manufacturer’s protocol.  2.12. TGF-β luciferase assay TGF-β activity in conditioned media of tendon cells was measured using a modified cell-based luciferase assay (Jones et al. 2013). HeLa cells were transfected with CAGA and Renilla plasmids using Metafecten® Pro (Biontex, Germany, #T040). Harvested conditioned media of tendon cells subjected to cyclic stretching were incubated with the transfected cells for 6 hours, and media of non-stretched cells were used as control. The luciferase activity was measured by Dual-Luciferease® Reporter Assay System (Promega, USA, #E1910) according to the kit manual and the luminescence was recorded using an Infinite 200 PRO microplate reader (Tecan, Switzerland). The luminescence of CAGA was normalized to Renilla as a control for transfection efficiency.  2.13. Immunohistochemistry Mouse tendon tissues were fixed in 10% formalin overnight, washed three times with PBS and stored in PBS at 4º. The tissues were dehydrated and embedded in paraffin.  Paraffin-53  embedded tissue sections were deparaffinised in three changes of xylene, and then hydrated through two changes each of 100%, 90%, and 70% alcohol, and then two changes of distilled water. Slides were blocked for endogenous peroxidase activity by two incubations of 20 minutes each in 3% hydrogen peroxide solution at room temperature. Antigen retrieval was performed using a 2% Pepsin in 0.1M HCl solution at 37˚C for 15 minutes. Slides were blocked with 1% BSA + 0.1% Tween in PBS for one hour, and then incubated with anti-CD-31 rabbit primary antibody (Abcam ab28634; 1:100) for one hour. A goat anti-rabbit Alexa 594 fluorescent secondary (Invitrogen A11037; 1:500) was then applied for 30 minutes at room temperature. Finally, slides were incubated with Hoescht (1:10,000) for two minutes to visualize nuclei. All staining procedures were conducted at room temperature in a darkened humidified chamber.  Paraffin-embedded human rotator cuff tendon biopsies were processed at the Provincial Health Services Authority (PHSA) Laboratories, Histology Services, and immunostained with antibodies against CD31 (Abcam ab28634; 1:100), ANGPTL4 (Aviscera Bioscinece, USA, A00309-02; 1:20) and HIF-1α (Novous Biologicals, #NB100-105; 1:50) followed by DAB staining.   2.14. Image Processing Fluorescence micrographs were taken using a Zeiss Axio Observer A.1 with a 10x objective. The exposure time was set at 4096 and 150 ms for CD31 and nuclei respectively. Images from five representative fields of view were taken for each tissue section. For each field of view, images were taken from the fluorescent channels representing CD31 and nuclei. CellProfiler software was used to color and overlay separate channels within a field of view. 54  To quantitate the expression of CD31, a positivity score representing the ratio of stained tissue to total tissue was taken for each CD31 channel image. Staining artefacts and overlapping tissue segments were cropped, and surface area of stained tissue was determined via thresholding using an Otsu method in ImageJ (Schindelin et al. 2012). Total tissue surface area was then determined by outlining the tissue in ImageJ. For human tendon tissue, the tissue slides were scanned using an Aperio Scanscope XT system (Leica Biosystems, USA) and full tissue sections were scored for positivity of DAB chromogen staining using ImageScope software. Sections were outlined, excluding any areas of tissue overlap, and the built-in Positive Pixel Count algorithm was used to count the ratio of positive-to-negative pixels. To quantify the proximity of immunostaining for ANGPTL4 and CD31, tissue images were captured using ImageScope software at a 2.0x magnification. Images were manually aligned, cropped to represent a 4 x 4 mm area of tissue, and subdivided into 64 equal squares. Two trained observers were instructed to mark each square with positive staining. The square was considered to have positive staining if both observers agreed. Where there was a disagreement with one observer noting positive staining and the other not, a third observer was brought in to break the tie. Squares between ANGPTL4 and CD31 staining trials in the same sample were then overlapped, and the number of positive overlaps to mismatches were considered. The percentage of proximity were reported as the ratio of the number of matched areas of CD31 and ANGPTL4 staining to the total number of CD31 staining.  55  2.15. In vitro ANGPTL4 cleavage assay The cleavage of recombinant human ANGPTL4 (R&D Systems, 4487-AN) by MMPs was assessed in Dr. Chris Overall’s lab (Starr et al. 2012). 1 µg recombinant protein was incubated with nine different MMPs at a ratio of 10:1 at 37ºC overnight. The cleaved products were confirmed by silver-stained 15% Tris-Tricine SDS-PAGE.  2.16. In vitro scratch assay A modified in vitro scratch assay (Liang et al. 2007) was used to evaluate cell migration in tendon cells that were isolated from ANGPTL4 +/+, +/- and -/- mice. The tendon fibers were pulled out from the mouse tail using a clamp. The tendon cells were isolated and cultured as prescribed before for human tendon cell. The cultured tendon cells were subcultured into 12-well plates at a density of 105 cells/well. A straight scratch was created on the confluent cell monolayer using a p1000 pipet tip. The wells were washed 3 times with PBS to remove debris and detached cells. Then 1 ml of cell culture medium was added to each well. A small dot was placed on the plate with a marker, as a reference point. Using a phase-contrast microscope (Zeiss Axio Observer A.1), the reference point was aligned at the centre of the field of view and the gap between the two sides of scratch was micrographed using a 5x objective lens at different incubation time periods to monitor closure of the scratch. The gap distance in the scratch was measure using Zen lite 2012 software (©Carl Zeiss Microscopy GmbH, 2011) and the migratory distance was calculated by subtracting the distance in each image from the starting distance.  56  2.17.Statistics The qPCR data of the in vitro and in vivo experiments were examined using a one-way ANOVA, followed by Tukey's multiple comparison and paired t-tests, respectively. Paired t-tests were also applied to the data generated by ELISA, tube formation assay and dual luciferase assay. Pearson's correlation coefficients were used to identify the association between target proteins/genes. Descriptive results from Angiogenesis RT² Profiler™ PCR Array were analyzed with Student’s t-test using the Web-Based PCR Array Data Analysis on the SABiosciences website. The other statistical analyses mentioned above were carried out using GraphPad Prism.               57  Chapter 3.Cyclic stretching alters the expression and release of angiogenic factors by human tendon cells 3.1. Background Tendinopathy is a common overuse injury, prevalent among both athletes and workers (Lewis et al. 2009). The disorder is often considered a mechanically driven pathology, as loading of tendons - whether repetitive or forceful - has been established as a risk factor (Seitz et al. 2011). Repetitive strain and shear are thought to induce matrix degeneration in tendon tissue, making the tissue susceptible to further damage and eventually to a symptomatic overuse injury (Mehta et al. 2003). However, the mechanisms that precede the development of symptomatic injury have not been fully described but are felt to be multifactorial, with repetitive strain being an important risk factor (Blevins 1997, Arnoczky et al. 2007).  There is evidence of extensive new blood vessel growth in most types of tendinopathy, including Achilles, patellar and lateral epicondyle tendinopathies, as well as the rotator cuff. Histopathological examination has revealed increased numbers of vessels within and around painful tendons (Matthews et al. 2006, Lakemeier et al. 2010). It has been reported that sites of subjectively defined pain, clinically palpated tenderness, tendon thickness and increased colour Doppler signal are anatomically associated, indicating a possible association between pain and neurovascular changes resulting from tendon overuse (Lewis et al. 2009, Divani et al. 2010). However, it must be acknowledged that the colour Doppler signal typically associated with tendinopathy may represent not only angiogenesis, but also increased blood flow in vessels that are already present. Angiogenesis may be accompanied by neurogenesis, i.e, nerves may be proliferating along with neovessels in mechanically loaded tendon tissue increasing the level of 58  substance P and other pain-producing substances in tendon; this histological change could lead to the transition to a symptomatic phase in tendinopathy (Gotoh et al. 1988). Tenocytes comprise the main cell population (90%-95%) in tendon tissue, and may be defined as scleraxis-expressing fibroblasts residing within the extracellular matrix of the tendon, and playing a key role in tendon development, adaption and the response to mechanical loading (Mendias et al. 2012). Tenocytes produce a variety of endogenous cytokines and growth factors which exert both autocrine and paracrine effects (Al-Sadi et al. 2011). Some in vitro studies have shown that repetitive mechanical loading of tendon cells results in an elevated production of soluble factors which are sometimes characterized as inflammatory, catabolic, or anabolic (e.g. PGE2, TGFβ)(Almekinders et al. 1993, Wang et al. 2003, Jones et al. 2013). Several studies have suggested that such changes in gene expression induced by repetitive stretching of tenocytes could lead to tendinopathy (Arnoczky et al. 2007).  In this study, we investigated the expression and activity of angiogenic factors released by cyclically stretched, scleraxis-expressing cells derived from human tendon tissue.  3.2. Results  3.2.1. Cell viability of tendon cells after applied cyclic stretching Flow cytometry of human tendon cells showed that the cyclic stretching protocol (1Hz frequency, 10% stretching) was well tolerated by tendon cells and did not induce cell death (Figure 3-1). This same cyclic stretching protocol has previously been shown to lead to increased 59  transcription of other factors known to play a role in tendinopathy, such as Substance P (Backman et al. 2011).   Figure 3-1. Flow cytometry histograms of tendon cells following propidium iodide staining. The cell death percentage, as represented by cell populations at Sub G1, showed that applied cyclic stretching (1Hz frequency, 10% stretching) for 24 hours (b) did not induce cell death compared to non- stretched tendon cells (a).  3.2.2. Increased angiogenic activity of factors released by repetitively stretched tendon cells Incubation of HUVEC cells with the conditioned media from 24 hours stretched or non-stretched tendon cells showed that cyclic stretching resulted in the accumulation of angiogenic factors, since media from loaded tendon cells increased the proliferation of HUVEC cells. In addition, the tube formation analysis showed that the released factors from 24 hours stretched tendon cells increased the total length of tubular network of endothelial cells in Matrigel compared to media from non-stretched cells (Figure 3-2). 60    Figure 3-2. Increased angiogenic activity of released factors by stretched tendon cells. Conditioned media from tendon cells subjected to 24 hours stretching (ST) compared to non-stretched cells (NS) increased (a) the proliferation of the endothelial cells (HUVEC) and (b) the tubular network formation by HUVECs in Matrigel (a: T test; b: Paired T test; mean ± SE; **, P < 0.01; N=3). (c) Micrographs of calcein AM-labeled tubular network of HUVECs in Matrigel after incubation with conditioned media of non- stretched (left) and stretched (right) tendon cells. Scale bars, 100 µM.  61  3.2.3. Cyclic Stretching Upregulates the Expression of Angiogenic Factors in Tendon Cells Profiling the expression of angiogenic factors by Human Angiogenesis RT² Profiler™ PCR Array showed that cyclic stretching increased the expression of several angiogenic factors, including some that have already been identified as playing a role in tendinopathy, such as VEGF and COX2, along with some novel genes (Figure 3-3).  Real-time quantitative PCR on selected genes (Figure 3-4.) showed that cyclic stretching within 1-4 hours (early response) transiently upregulated the expression of angiogenic factors including ANGPTL4 (P < 0.001), FGF-2 (P < 0.001), COX2 (P < 0.001), SPHK1 (P < 0.01, TGF-alpha (P < 0.01), VEGF-A (P < 0.001) and VEGF-C (P < 0.01). In fact, the upregulation of these genes reached peak levels within 1-4 hours of cyclic stretching.  By extending the time course (longer than 4 hours), the expression of these genes was observed to return to control levels. The early and transient upregulation of mRNA for angiogenic factors in response to cyclic stretching was significant for all of the above-mentioned genes (Figure 3-4.). The gene expression array showed no increase in the expression of antiangiogenic factors such as BAI1, SERPINF1, THBS1 and 2, TIMP1-3 (Figure 3-3). 62   Figure 3-3 Gene expression array of angiogenic factors expressed by tendon cells after 4 and 12 hours cyclic stretching. The expression of 84 genes involved in angiogenesis in tendon cells was examined by qPCR after 4 and 12 hours cyclic stretching of tendon cells (Mean ± SE; N=4).  63   Figure 3-4. Real-time quantitative PCR on selected genes. The results showed a dynamic response to cyclic stretching for inducing the expression of most angiogenic factors. (Early response: 1-4 hrs cyclic stretching, Late response: 6-24 hrs cyclic stretching). (Mean ± SE; **, P < 0.01; ***, P < 0.001; N=4). 64  3.2.4. Cyclic stretching increases the release of the ANGPTL4 and MMP-2 proteins Western blot analysis of cell lysates showed that cyclic stretching modulated the level of intracellular ANGPTL4 protein. The extracellular levels of ANGPTL4 in response to stretch demonstrated an obvious and statistically significant increase (Figure 3-5.). Zymography analysis showed that cyclic stretching also increased the gelatinase activity of MMP-2 (gelatinase A) in conditioned media of tendon cell culture (Figure 3-6).    65    Figure 3-5. Cyclic stretching increases the expression and release of ANGPTL4 in tendon cell culture. (a) Western blot analysis of ANGPTL4 protein in total protein extracts of tendon cells with no stretching (NS), 2, 4 and 6 hours stretching. (b)  Quantitation of the Western blot bands using vinculin as the loading control. (c) Western blot analysis of ANGPTL4 protein in concentrated serum free conditioned media of tendon cells with no stretching compared with 2 hours cyclic stretching (ST) followed by 6 hours rest. (d) Increased concentration of ANGPTL4 protein in the conditioned media of tendon cells after 6 hours continuous stretching compared with no stretching (Paired T test; mean ± SE; ****, P < 0.0001; N=4).  66   Figure 3-6 Zymography of MMP-2 activity. The concentrated conditioned media of tendon cells resolved on a the zymography gel (10% polyacrylamide gel containing 0.1% gelatin) showed an increased gelatinase activity of MMP-2 in the harvested media from the stretched (ST) tendon cells compared to the non-stretched (NS) cells. There were no other visible bands.      67  Chapter 4. TGF-β and HIF 1α modulate the expression of ANGPTL4 in response to cyclic stretching 4.1. Background Mechanical stimuli induce tendon cells to release a number of cytokines such as VEGF, TNF-α, IL-6, PGE2, substance P and PDGF which may promote angiogenesis in tendon tissue (Skutek et al. 2001, Skutek et al. 2001, Petersen et al. 2004, Fong et al. 2005, Yang et al. 2005, Backman et al. 2011, Gao et al. 2013, Legerlotz et al. 2013). Some studies have shown that several overlapping signalling pathways including Ras/Rho and MAPK signalling, phospholipase C activation, calcineurin-mediated signalling and microRNAs, are activated through stretch-sensitive ion channels at the cell membrane, integrins and integrin-associated proteins and cell-surface receptors (Jaalouk et al. 2009). The mechanisms and regulatory pathways that alter the gene expression and promote vascular changes in overuse tendinopathy are not completely characterized. In this study we examined regulatory pathways that modulate the expression of ANGPTL4, as a mechanoresponsive gene, in response to mechanical stimuli. Several studies have reported increased expression and activity of HIF-1α in response to mechanical stress, and furthermore shown that this increased activity induces the expression of VEGF and thereby promotes angiogenesis (Kim et al. 2002, Petersen et al. 2004). However, the mechanism involved in mechanical activation of HIF-1α is unknown. The expression and activity of TGF-β also are influenced by physical stimuli (Heinemeier et al. 2003, Maeda et al. 2011, Jones et al. 2013). The roles of TGF-β and hypoxia in the modulation of ANGPTL4 expression have been shown in lung and breast cancer, respectively, and have been tied to the promotion of tumour metastasis (Padua et al. 2008, Kim et al. 2011). In this study we examined the potential roles of TGF-β and HIF-1α in the induction of ANGPTL4 expression by cyclic stretching.  68  4.2. Results 4.2.1. HIF-1α modulates the expression of ANGPTL4 Dimethyloxalylglycine (DMOG) is a cell permeable prolyl-4-hydroxylase inhibitor which increases the accumulation of HIF-1α protein (Barrett et al. 2011). The results of gene expression analysis and ELISA showed increased expression of ANGPTL4 mRNA and protein in human tendon cells following DMOG treatment in a dose-responsive manner (Figure 4-1.a, b). Hypoxic conditions also enhanced the release of ANGPTL4 protein from human tendon cells (Figure 4-1.c). Inhibition of HIF-1α by chetomin (Staab et al. 2007) prevented the induced release of ANGPTL4 by cyclic stretching (Figure 4-1.d). Immunostaining of human rotator cuff tendon also showed a strong correlation between the expression of HIF-1α and ANGPTL4 (Figure 4-2.). These data together suggest that the expression and release of ANGPTL4 are regulated by the HIF-1α pathway, and that the induction of ANGPTL4 by cyclic stretching is mediated through this pathway.  69   Figure 4-1. HIF-1 modulates the expression of ANGPTL4 in human tendon cells. DMOG induced ANGPTL4 mRNA (a) and protein (b) in human tendon cells. Hypoxic condition (1% oxygen) also increased the release of ANGPTL4 protein from culture of human tendon cells (c). Induced release of ANGPTL4 from stretched (ST) human tendon cells was diminished by chetomin (100 nM) (d). (Student t-test and one way ANOVA; mean ± SE; *, P < 0.1; **, P < 0.01; ***, P < 0.001; N=4).   70    Figure 4-2. Correlation of ANGPTL4 and HIF-1α expression in human rotator cuff tendons. The scanned images of immunostained human rotator cuff tendons with ANGPTL4 and HIF-1α antibodies (a) showed that the percentage of positive pixels ANGPTL4 staining has a strong correlation with HIF-1α staining (b) (Pearson r= 0.8285; P < 0.0001; N=24). Scale bars, 100 µM. Red and black arrows show blood vessels and separation artefacts, respectively. 71  4.2.2. ANGPTL4 induced by cyclic stretching is mediated by TGF-β activity  Expression and release of ANGPTL4 protein by human tendon cells was blocked by A-83-01 (Figure 4-3.a, b). A-83-01, an inhibitor of Smad2/3 phosphorylation and the TGF-β cascade (Tojo et al. 2005), was also used to study the role of TGF-β pathway in the regulation of ANGPTL4 induced by cyclic stretching. The results demonstrated that blocking the TGF-β pathway by A-83-01 prevented the induced release of ANGPTL4 from tendon cells during cyclic stretching (Figure 4-3.c). To determine the effect of cyclic stretching on the activity of TGF-β released by tendon cells, a cell based luciferase assay was used (Jones et al. 2013). Our data showed increased activity of TGF- β in the conditioned media of stretched tendon cells compared to non-stretched cells (Figure 4-3.d).  72   Figure 4-3. TGF-beta regulates ANGPTL4 expression in human tendon cells. Inhibition of TGF-beta receptor by A83.01 blocked the induced expression of ANGPTL4 mRNA (a) and the release of ANGPTL4 protein (b) in response to TGF-β. A83.01 prevents the induction of ANGPTL4 in response to cyclic stretching (c). TGF-beta activity was increased in the conditioned media of stretched (ST) tendon compared to non- stretched (NS) cells (d). (Student t-test and one way ANOVA; mean ± SE; *, P < 0.1; **, P < 0.01; ***, P < 0.001; N=4)   73  4.2.3. ANGPTL4 is induced by both HIF-1α and TGF-β  We exposed human tendon cells to TGF-β, DMOG (a stabilizer of HIF-1α) or PGE2, and evaluated the expression of 84 genes related to angiogenesis. Our data from the qPCR array showed that several genes were upregulated in response to DMOG, PGE2 and TGF-β (Figure 4-4.). PGE2 had no significant effect on expression of ANGPTL4. Of this set, the only gene induced by both TGF-β and DMOG was ANGPTL4 (Figure 4-5.). This finding accentuates the complementary role of TGF-β and HIF-1α in the regulation of tendon vascularization through ANGPTL4.  74    Figure 4-4. Gene expression array of angiogenic factors in tendon cells after 6 hours incubation with DMOG, PGE2 and TGF-β (mean ± SE; N=3).  75    Figure 4-5. An array of angiogenesis related genes regulated in response to TGF-β and DMOG. A volcano graph of the expression of angiogenesis related genes and the table of upregulated genes following the incubation of human tendon cells with DMOG (100µM) (a) and recombinant TGF-β protein (2.5 ng/ml) (b) for 6 hours showed that ANGPTL4 was the only gene in this array induced by both DMOG and TGF-β (N=3).    76  Chapter 5. Role of ANGPTL4 in tendon vascularization 5.1. Background ANGPTL4 belongs to a superfamily of secreted proteins that regulate angiogenesis. Although there is a structural similarity with angiopoietins, ANGPTL proteins are orphan ligands that do not bind to the receptor tyrosine kinases Tie 1 or Tie 2. Shortly after the discovery of ANGPTL4 as a factor involved in lipid metabolism (known as fasting –induced adipose factor), the function of this protein has been examined in many other metabolic and non-metabolic pathways, including those which modulate energy homeostasis, wound healing, oncogenesis, metastasis, inflammation, lymphangiogenesis and angiogenesis. Recent studies showed that ANGPTL4 provokes the disruption of vascular junction integrity via integrin α5β1-mediated Rac/PAK signalling and the de-clustering and internalization of VE-caderin and claudin-5 which eventually induces vascular leakiness and permeability (Le Jan et al. 2003, Morisada et al. 2006, Gealekman et al. 2008, Hato et al. 2008, Huang et al. 2011). In the current study we characterized the function of the ANGPTL4 protein in tendon vascularization. This study examined the potential role of Angiopoietin-like 4 (ANGPTL4) in the angiogenic response of tendons to repetitive stretching or acute injury. ANGPTL4 was found to stimulate the angiogenic activity of endothelial cells. Angiogenic activity was also increased following injury and following injection of ANGPTL4 into mouse patellar tendons, whereas the patellar tendons of ANGPTL4 knockout mice displayed reduced angiogenesis following injury.   77  5.2. Results 5.2.1. Angiogenic activity of ANGPTL4 promotes tendon vascularization To investigate the angiogenic activity of the ANGPTL4 protein, several in vitro and in vivo approaches were used. In vitro tube formation by endothelial cells on a basement membrane matrix (Matrigel) is a standard technique to evaluate angiogenesis (Arnaoutova et al. 2010). Primary human umbilical vein endothelial cells (HUVECs), which were incubated with full length recombinant human ANGPTL4 protein on matrigel, formed longer tubes compared to controls; this effect was highly significant (p< 0.0001) (Figure 5-1..a). To investigate the in vivo function of ANGPTL4 in tendon tissue, recombinant mouse ANGPTL4 protein was directly injected into the mouse patellar tendon and the expression of several well-characterized angiogenesis markers (including CD31, CD34, KDR/FLK-1 and VEGFA) were evaluated in harvested patellar tendons. Mouse PTs showed significantly increased expression of CD31 protein in tendon tissue injected with recombinant ANGPTL4 protein versus control (Figure 5-1..b). Up-regulation of CD31 and other angiogenesis markers in response to ANGPTL4 protein injection was confirmed by qPCR. The recombinant protein also dramatically induced the expression of MMP-3, whose activity may facilitate tendon remodelling and vascularization (Sahin et al. 2012) (Figure 5-1..c).  Angiogenesis is part of the normal healing process after acute tendon injury (Sharma et al. 2005). To investigate the role of ANGPTL4 in tendon healing, we utilized a patellar tendon injury model in which a biopsy punch is used to induce damage in the tendon tissue and the tendons were harvested 3 weeks after injury. This patellar tendon injury model caused significant up-regulation of ANGPTL4, along with CD31 as an angiogenesis marker, 3 weeks after injury (Figure 5-2). 78  However, in ANGPTL4 -/- mice, the expression of CD31 was significantly altered following injury. Acute tendon injury also significantly induced the expression of MMP-3, but only in wild-type and heterozygous mice and not in ANGPTL4 -/- mice (Figure 5-2.). There was also decreased expression of MMP-3 in the ANGPTL4 -/- mouse (which could explain the decreased migration of isolated tendon cells from -/- mouse tail). Indeed, the isolated cells from wild type mouse tail tendons which were cultured for in vitro scratch/wound healing assay, were significantly faster at filling the gap in the cell monolayer compared to the isolated cells from -/- mouse tail tendons (Figure 5-3.). We also found a moderate to strong correlation between the expression of ANGPTL4 and angiogenesis markers when comparing gene expression in tendon tissue samples from ANGPTL4 +/+ and +/- mice (Figure 5-4.). Finally, we postulated that a correlation betwteen ANGPTL4 and vascularity (indicated by CD31) would be observed in human tendon tissue. Subscapularis and supraspinatus tendons from human rotator cuff were immunostained for CD31 and ANGPTL4 (Figure 5-5..a). Again, a moderate correlation could be seen between the expression of CD-31 and ANGPTL4 (Figure 5-5..b), suggesting an association of ANGPTL4 expression with tendon vascularization. The side-by-side comparison between the scanned images of the serial sections of human rotator cuff tendons which were immunostained with CD31 and ANGPTL4 antibodies showed a similar but not entirely overlapping pattern of staining (Figure 5-6.). Seventy-nine percent of the staining area for CD31 expression was matched with the positive areas of ANGPTL4 staining, supporting a possible role for ANGPTL4 in induction of blood vessel formation.  79    Figure 5-1. In vitro and in vivo angiogenic activity of recombinant ANGPTL4 protein. a) Recombinant ANGPTL4 protein (10 µg/ml) induced tube formation by HUVECs in matrigel (N≥3). The images show the micrographs (x5) of tubular network stained with Calcein AM showed the tubular network under florescent microscope. The graph shows the total tube length per micrograph measured by AngioTool software. Scale bars, 100 µM. b) 3 days after injecting 5 µg recombinant ANGPTL4 protein into patellar tendon of CD-1 mouse, expression of CD31 protein was increased compared to control which was injected by PBS with same volume (N=6). The micrographs represent IHC of the patellar tendon tissue immuno-labelled for CD31 and the graph shows the positive pixels for CD31 staining measured by imageScope software. Scale bars, 100 µM. 80  c) The injection of recombinant protein also significantly induced the mRNA expression of endothelial cell markers (CD31, CD34 and FLK), MMP-3 and VEGF-A (Student t-test; mean ± SE; *, P < 0.1; ****, P < 0.0001; N=6).    Figure 5-2. Lack of ANGPTL4 decreases tendon vascularization following tendon injury in an ANGPTL4-defiecient mouse model. The significantly induced expression of ANGPTL4 in PTs of +/+ mice in response to acute injury was associated with increased expression of CD31 and MMP-3 (a). The florescent micrograph of PTs showed a higher expression of CD31 (green colour) in +/+ mice compared to -/- mice (N=70 paired fields) (b). (Student t-test; mean ± SE; ***, P < 0.001; N≥3). Scale bars, 100 µM. 81    Figure 5-3. Scratch assay of isolated mouse tendon cells. 82  a) The micrographs (5x) of the tendon cells isolated from mouse tail of ANGPTL4 +/+, +/- and -/- which were taken 0, 6, 22 and 30 hours after scratching the cell monolayers with a pipet tip. b) Lack of ANGPTL4 in -/- cells decreased tendon cell migration (mean ± SE; N=3).     Figure 5-4. Correlation of ANGPTL4 expression with target genes. a) The linear regression of the expression of ANGPTL4 (y axis) and angiogenesis related genes (x axis) demonstrate a significant correlation with some target genes. b) ANGPTL4 showed a strong correlation with CD31 and a moderate correlation with CD-34, FLK-1, MMP-3 and VEGF. 83    Figure 5-5. Correlation of ANGPTL4 and CD31 expression in human rotator cuff tendons. The scanned images of immunostained human rotator cuff tendons with ANGPTL4 and CD31 antibodies (a) showed that the number of positive pixels ANGPTL4 staining has a moderate correlation with CD31 staining (b) (Pearson r= 0.5723; P < 0.01; N=22). Red and black arrows indicate blood vessels and separation artefacts, respectively.   84   Figure 5-6. The scanned images of immunostained human rotator cuff tendons. The staining pattern for the expression of ANGPTL4 (top) and CD31 (bottom) in serial sections of human rotator cuff tendons were predmominantly concentrated in vascular regions of the tendon, although ANGPTL4 staining is also visible in cells throughout the tendon collagenous matrix (presumably, tenocytes).  Red, black and green arrows indicate blood vessels, separation and folding artifacts, respectively.   85  5.2.2. Lack of ANGPTL4 has no effect on gene markers for tendon cells and their progenitors  To study the role of ANGPTL4 in tendon regeneration, the expression of selected gene markers for tendon cells and tendon progenitors such as tenomodulin, Scleraxis and Nucleostemin was quantified in patellar tendons from ANGPTL4-deficient mice. We also quantified the expression of genes related to tendon matrix such COL1A1, COL3, DCN and LOX. Our qPCR results showed that lack of ANGPTL4 did not have any significant effect on the expression of the selected genes (Figure 5-7.).   Figure 5-7. The expression of selected gene markers in PTs of ANGPTL4-deficient mice (-/-) versus wild type (+/+).  Genes that are involved in matrix synthesis (COL1A2, COL3A1, DCN and LOX), tendon and stem cell markers (SCX, TNMD and Nucleostemin) did not demonstrate significantly different levels of expression in the PTs of ANGPTL4-deficient mice compared to wild type mice (mean ± SE; N≥3). 86  5.2.3. MMPs are able to cleave recombinant ANGPTL4 protein Our Western blot showed that tendon cells produce ANGPTL4 in the full-length form (≈55 kD). Several proteases are able to cleave this form of ANGPTL4 at specific sites to produce N-terminal and a C-terminal fibrinogen-like fragments. The cleavage of ANGPTL4 affects its biological activity (Yin et al. 2009, Lei et al. 2011).  Previously we showed that ANGPTL4 and cyclic stretch induce the expression of MMP-3 and MMP-2 respectively. So we decided to investigate the cross-talk between the MMPs and ANGPTL4. We had the opportunity to evaluate potential cleavage of ANGPTL4 by taking advantage of the expertise available in the laboratory of Dr. Christopher Overall (UBC). The proteolytic cleavages of ANGPTL4 protein were determined by incubating the full length recombinant ANGPTL4 protein with nine different MMPs. Resolving the cleaved product by SDS PAGE showed that MMP-7, MMP-8 and MMP-14 were effective in cleaving the full length protein. MMP-7 was more effective compared to the other MMPs.   87    Figure 5-8. The cleaved products of ANGPTL4 protein after incubation with MMPs.  Different MMPs were incubated with (+) and without (-) full length ANGPTL4 (FL-ANGPTL4) protein. The products were resolved and visualized in silver-stained 15% Tris-Tricine SDS-PAGE along with molecular weight (MW) marker and negative control (NC). 88  Chapter 6. Discussion 6.1. Cyclic stretching modulates angiogenic factors in tendon cells The results of this study build on previous suggestions that angiogenesis may play a role in the pathogenesis of tendon overuse pathology by demonstrating that tendon cells subjected to cyclic stretching display marked angiogenic activity. Tendon cells may therefore be partly responsible for the widespread angiogenesis which is known to occur in chronic tendinopathy (Savitskaya et al. 2011). Alternatively, the angiogenic response of cyclically strained tendon cells may represent an adaptive response as would be seen following physiologic levels of exercise. My approach in this study was to examine the effect of cyclic stretching on the expression of a wide array of angiogenic factors. Gene expression analysis demonstrated that cyclic stretching of primary tendon cells on tissue culture plates increased the mRNA levels of several angiogenic factors including ANGPTL4, COX-2, bFGF (FGF-2), TGFα, VEGF-A, VEGF-C and SPHK1, while no increased expression of antiangiogenic factors (e.g. BAI1, SERPINF1, THBS1 and 2, TIMP1-3) was seen. By extending the time course of cyclic stretching, the expression of these same genes was subsequently observed to be downregulated after 4 hours of strain. This time course suggests that, in response to repeated bouts of cyclic stretching, tendon tissue might be exposed dynamically to repeated, short bursts of angiogenic stimuli which, over time, could lead to an increase in the number of blood vessels in the rotator cuff and associated soft tissues. However, it must be acknowledged that the cyclic stretching regimen used in the current study (equibiaxial stretching of two dimensional cell culture) is rather different than the mechanical conditions which would be experienced by tenocytes in vivo. Therefore, further study in a suitable 89  laboratory model would be required to confirm the physiological or pathophysiological relevance of these findings. The strain level measured in tendon tissue is dependent on the type of tendon and the physical activity. Wilson et al recorded the maximum average strain of 5.8% in Achilles tendon during running and  6%-7% strain were recorded in the patellar tendon during the development of maximal isometric force; 15% peak strain is predicted for the patellar tendon during jumping (Lichtwark et al. 2006, Lavagnino et al. 2008, Couppe et al. 2009). The failure strain of Achilles tendon is 16.1% and 12.8% for fast (10% s-1) and slow (1% s-1) strain rates respectively (Wren et al. 2001). In an in vitro model, the transmission of strain to individual cells in cell culture depends on their adhesion to substrate and usually is less than substrate strain (Wang et al. 2007). Therefore, we reasoned that 10% mechanical stretching of tendon cells is physiologically relevant to the normal stretching of tendon tissue during intense physical activity. In our model, the flow cytometry results showed that 10% stretching of the substrate of tendon cell culture did not induce apoptosis in cultured tendon cells and was well tolerated by tendon cells.    Similar to the cyclic tension model used in the current thesis, a study by von Offenberg Sweeney and colleagues on bovine aortic endothelial cells showed that cyclic strain stimulates MMP-2 activity and expression in a force and time dependent manner (von Offenberg Sweeney et al. 2004). Consistent with this study, we demonstrated that cyclic loading increases release of MMP-2 from tendon cells which resulted in elevated MMP-2 activity in the conditioned media. MMP-2, beyond its role in matrix remodeling and collagen degradation, can potentially modulate angiogenic processes (van Hinsbergh et al. 2008). The role of MMP-2 in tendons is not completely understood and will be an important topic for future study.  90  In this study we showed that the transient upregulation of several angiogenic factors in response to cyclic loading increased the accumulation of factors including ANGPTL4 that may have proliferative effects on HUVEC cells. There is growing evidence that ANGPTL4 induces a pro-angiogenic response which is independent of the effects of VEGF. Recent studies showed that ANGPTL4 provokes the disruption of vascular junction integrity via integrin α5β1-mediated Rac/PAK signaling and the de-clustering and internalization of VE-cadherin and claudin-5 which eventually induce vascular leakiness and permeability (Le Jan et al. 2003, Morisada et al. 2006, Gealekman et al. 2008, Hato et al. 2008, Huang et al. 2011). Our study is the first report showing an induction of ANGPTL4 protein in response to mechanical stimulus. Future studies will attempt to unravel the distribution of this protein in tendon tissue from a larger group of patients with chronic tendinopathy. We also plan to examine the potential mechanisms involved in its mechanical regulation during physiologic levels of exercise. Tenocytes and tenoblasts are the main cell types in tendon tissue. However other cells such as synovial, smooth muscle, and endothelial cells are also present in the tissue (Kannus 2000). In addition, Yanming et al defined a population of tendon stem/progenitor cells which have self-renewal capacity and stem cell phenotypes (Bi et al. 2007). In our study we used a modified protocol used by Backman et al to isolate and culture human tendon cells from human hamstring tendon tissue (Backman et al. 2011). Although the isolated tendon cells were heterogenous and express both stem cell and tendon cell markers (Appendix B.), the selected cell passages (3-5) for our experiments had the highest expression of tendon cells markers (TNMD and SCX) versus stem cell markers (Nucleostemin) which could be due to either a relative increase during culture of the size of population of TNMD/SCX-positive cells, or enhanced differentiation of progenitor cells to the tendon phenotype in these cell passages. These cells also express aggrecan, collagen and 91  decorin proteins. The highest expression of TNMD has been shown in tendon tissue compared to other tissue.  Jelinsky et al showed that the expression of TNMD is reduced when they were grown in two-dimensional culture which suggested cell dedifferentiation (Jelinsky et al. 2010). Although we didn’t compare the expression of cultured tendon cells with tissue and other culture systems, our result showed that the expression of TNMD was increased in subcultured tendon cells during later passages.  In summary, this study demonstrates that cyclic stretching of primary cells isolated from human tendon tissue induces the expression and release of angiogenic factors which can induce endothelial cell proliferation and tube formation. Further work will be required to achieve a more complete understanding of the relationship between repetitive stretching and angiogenesis and its potential contribution to the progression of tendon degeneration or to physiologic adaptation to exercise, particularly with regard to the role of ANGPTL4. Our next studies characterized the role of ANGPTL4 in tendon and revealed the signalling pathways that modulate the expression of ANGPTL4 by tendon fibroblasts during the mechanical stimulation of tendon cells.  6.2. ANGPTL4 activity is increased in cyclically stretched tendon fibroblasts and promotes angiogenesis in injured tendon ANGPTL4 is a factor that modulates cell-matrix communication and matrix degradation and thus may modify tissue matrices and mechanical resistance in tendon tissue, which can contribute to macroscopic and microscopic changes observed in tendinopathy (e.g. disorganized 92  collagen bundles, neovascularization and fragile texture of tissue). This is first report that has shown the induction of ANGPTL4 by cyclic stretching of primary cells in culture. The mechanisms and regulatory pathways that promote vascular changes in overuse tendinopathy are an ongoing topic of study. In these experiments, we found that ANGPTL4 is induced by cyclic stretching and that it regulates tendon vascularity following injury and cyclic stretching. Furthermore, we characterised the angiogenic activity of ANGPTL4, and the pathways that regulate this protein to promote angiogenesis in tendon tissue. We showed that cyclic stretching induces the expression and release of ANGPTL4 through the activities of HIF-1α and TGF-β. ANGPTL4 has multiple functions which include both lipid metabolism and angiogenesis (Hato et al. 2008). The role of ANGPTL4 in lipid metabolism and its inhibitory effect on lipoprotein lipase have been well studied, but its angiogenic activity has remained controversial. In certain conditions, ANGPTL4 inhibits the adhesion, migration, and sprouting of endothelial cells and suppresses vascular leakiness, angiogenesis and tumour metastasis (Ito et al. 2003, Cazes et al. 2006, Galaup et al. 2006). On the other hand, recent studies indicate that the angiogenic activity of ANGPTL4 promotes vessel permeability and induces neoangiogenesis in numerous disorders including ischemic retinopathies, obesity, arthritis and some tumors (Le Jan et al. 2003, Hermann et al. 2005, Gealekman et al. 2008, Ma et al. 2010, Nakayama et al. 2010, Nakayama et al. 2011, Zhang et al. 2012, Xin et al. 2013). The angiogenic activity of ANGPTL4 is independent of VEGF and can stimulate angiogenesis in the presence of factors that inhibit endothelial cell growth and VEGF activity (Le Jan et al. 2003, Gealekman et al. 2008). Although no specific receptor has been identified for ANGPTL4 to convey its angiogenic activity, it has been shown 93  that the protein interacts with integrin 5α1, VE-cadherin, and claudin-5 which disrupts contacts between endothelial cells and promotes blood vessel leakiness (Huang et al. 2011).  To our knowledge, our lab was the first group to report the mechanical stimulation of ANGPTL4 expression and to characterize the function of this protein in tendon vascularization (Mousavizadeh et al. 2014). Kersten’s lab showed that endurance exercise enhances ANGPTL4 protein level in the plasma through metabolic changes and increased release of free fatty acids (Kersten et al. 2009). Although in our study we haven’t determined the effects of free fatty acids and metabolic factors on regulation of ANGPTL4 in tendon cells, our results suggest that TGF-β activity and the HIF-1α pathway mediate the expression and release of ANGPTL4 during cyclic stretching. The roles of TGF-β and hypoxia in the modulation of ANGPTL4 expression have been shown in lung and breast cancer, respectively, and have been tied to the promotion of tumour metastasis (Padua et al. 2008, Kim et al. 2011). TGF-β isoforms and their receptors have broad biological functions and are involved in different processes in tendon tissue including development, growth, healing, adaptation and tissue homeostasis (Fenwick et al. 2001, Klein et al. 2002, Heinemeier et al. 2003, Pryce et al. 2009, Maeda et al. 2011). The expression and activity of TGF-β are influenced by physical stimuli that mediate the altered expression and synthesis of other factors in response to mechanical forces and exercise (Heinemeier et al. 2003, Maeda et al. 2011, Jones et al. 2013). In our cell culture model we found increased activity of TGF-β following cyclic stretching of human tendon cells, which may stimulate the expression and release of ANGPTL4. HIF-1α is another regulator of ANGPTL4 that may become activated in tendon cells by cyclic stretching, even in normoxic conditions. Several studies have shown the increased expression and activity of HIF-1α in response to mechanical stress, which can induce the expression of VEGF and angiogenesis (Kim et al. 2002, Petersen et al. 2004). However, the 94  mechanism involved in the mechanical activation of HIF-1α is unknown. Our study suggests that ANGPTL4 is also induced by increased activity of HIF-1α in response to cyclic stretching. An in situ freezing model of patellar tendon injury demonstrated that HIF-1α stabilization following injury promotes neoangiogenesis via induced expression of VEGF and MMP-3. The biochemical changes and angiogenesis were correlated with deteriorated mechanical properties of tendon tissue in this model (Sahin et al. 2012). Murata et al have shown that MMP-3 and MMP-1 are distinctively regulated by ANGPTL4 in chondrocytes (Murata et al. 2009). MMP-3 plays a crucial role in tissue remodelling and is involved in various normal and pathophysiological processes. The enzymatic activity of MMP-3 promotes the degradation of extracellular matrices and collagen fibers, and induces the activity of other MMPs by protein cleavage. The cleavage of VEGF by MMP-3 and MMP-9 increases the bioavailability of the growth factor and promotes aberrant blood vessel formation (Lee et al. 2005). MMP-3 also induces the proliferation and migration of endothelial cells, thereby promoting angiogenesis and accelerating wound healing after dental pulp injury (Zheng et al. 2009). The altered expression of MMP-3 in acute rotator cuff injuries and tendon healing suggest that it functions in tissue degradation and remodelling (Del Buono et al. 2012). Mathieu et al showed that matrix remodelling during chondrogenesis is influenced by ANGPTL4 which upregulates the expression of several MMPs including MMP-3 (Mathieu et al. 2014). Our in vivo indicate that ANGPTL4 can induce MMP-3 expression during the healing process and following PT injection with recombinant ANGPTL4 protein. Emerging work has shown that ANGPTL4 interacts with ECM proteins such as vitronectin and fibronectin, delaying their proteolytic degradation by MMPs (Goh et al. 2010). The role of ANGPTL4 in matrix degradation and remodelling is likely very 95  complicated and further studies are needed to clarify its effect in tendon remodelling or degeneration.                96  Chapter 7.Conclusions and perspectives  The work in this thesis demonstrates that cyclic stretching induces the expression and release of angiogenic factors from tendon cells, and that these angiogenic factors promote endothelial cell proliferation and tube formation. Further work is required to achieve a better understanding of the relationship between repetitive stretching and angiogenesis and its potential contribution to the progression of tendon degeneration, particularly with regard to the role of ANGPTL4. This is the first study to characterize the function of ANGPTL4 in tendon vascularization and the pathways that regulate the expression of ANGPTL4 in response to cyclic mechanical stretching. My data demonstrated quite convincingly that the angiogenic activity of ANGPTL4 is a key factor in the induction of neovascularization in tendon tissue. Our study also determined the role of TGF-β and HIF-1α in the induction of ANGPTL4 expression, particularly by cyclic stretching. Cyclic stretching stimulates the activity of TGF-β and HIF-1α, which increases the expression and release of ANGPTL4 from human tendon cells.  My preliminarily results showed that lack of ANGPTL4 decreases the tendon cell migration. One of the features of tendinopathy is hypercellularity which might be due to induced cell migration and proliferation by mechanical stress. More comprehensive studies may indicate the role of ANGPTL4 in tendon hypercellularity and the mechanisms by which ANGPLT4 modulates cell migration. The potential role of ANGPTL4 in cell migration proposes further work to characterize the function of this protein in endothelial cell migration and angiogenesis. In fact, these studies may determine the mechanism by which ANGPTL4 regulate angiogenesis in tendon tissue.  97  In this study, I also showed that ANGPTL4 induces the expression of MMP-3 and that several MMPs are able to cleave the protein which may alter the biological activity and availability of the protein. Therefore, the cross-talk between MMPs and ANGPTL4 may play a role in the physiological and mechanical properties of tendon tissue. The physiologic role of ANGPTL4 in lipid metabolism and its regulation by free fatty acids supports further investigation into the physiologic effects of exercise on lipid metabolism and the release of free fatty acids that may affect the expression and release of ANGPTL4 in tendon tissue. These future investigations would potentially reveal another pathway that may modulate ANGPTL4 in response to cyclic stretching and exercise. The identification of other mechanoresponsive pathways that modulate ANGPTL4 in tendon tissue, in addition to hypoxia and TGF-β, may accentuate the role of this protein as a key factor in tendon physiology or pathophysiology. Understanding factors and pathways that modulate neovascularization in overuse tendinopathy may lead us to develop new methods for managing and detecting overuse tendon injuries. 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Chen, B. Krishnamachary, P. T. Winnard, Jr., V. Raman, L. Zhen, W. A. Mitzner, S. Sukumar and G. L. Semenza (2012). "HIF-1-dependent expression of angiopoietin-like 4 and L1CAM mediates vascular metastasis of hypoxic breast cancer cells to the lungs." Oncogene 31(14): 1757-1770. Zheng, L., K. Amano, K. Iohara, M. Ito, K. Imabayashi, T. Into, K. Matsushita, H. Nakamura and M. Nakashima (2009). "Matrix metalloproteinase-3 accelerates wound healing following dental pulp injury." Am J Pathol 175(5): 1905-1914. Zudaire, E., L. Gambardella, C. Kurcz and S. Vermeren (2011). "A computational tool for quantitative analysis of vascular networks." PLoS One 6(11): e27385.    115  Appendices Appendix A.  Substance P and Its Regulatory Pathway in Response to Glucocorticoids  The overall aim of this study was to find the potential mechanisms involved in the response of overuse tendon injuries to glucocorticoid, a current treatment for tendon disorders. This study was a collaboration with a research group in Umea University, Sweden. A cell culture model was used to answer the following question:  What is the effect of glucocorticoids in the regulation of angiogenic factors and mechanisms involved in acute pain-relief following corticosteroid injections?  I hypothesize that: Glucocorticoids reduce the expression of SP in human tendon cells through a glucocorticoid receptor (GR)-dependent pathway (Figure A-1).   Figure A-1 Hypothetical pathway regulating SP in response to glucocorticoids  116  A.1. Material and Methods A.1.1. Cell culture Tendon cells were obtained from the midportion of healthy Achilles tendons (Umea University) or the semitendinosus hamstrings tendon from donors undergoing a reconstruction of the anterior cruciate ligament (ACL; University of British Columbia). Each protocol was approved by the local ethics committee (UBC, Umea University), and cell culture was conducted in Canada (hamstrings) or Sweden (Achilles). Written informed consent was obtained according to the Declaration of Helsinki (most recently at the General Assembly in October 2008). Tendon biopsies were washed and minced into 3-5mm pieces, then enzymatically digested at 37°C under constant agitation using collagenase (clostridopeptidase A, Sigma, C-0130) in D-MEM (Dulbecco’s Modified Eagle Medium, Invitrogen, 11960) at a concentration of 1.5 mg/mL. For the hamstring tendon biopsies, this was followed by incubation in 0.25% trypsin for 3min. The digestion product was centrifuged at 800 x g for 5 min and re-suspended and cultured in D-MEM supplemented with 10% fetal bovine serum (FBS; Invitrogen; 16000), 2mM L-glutamine (Invitrogen; 25030), 100units/mL pencillin and 100ug/mL streptomycin (Invitrogen; 15140) at 37°C in a humidified atmosphere of 5% CO2 in air. The media was changed every third day. Confluent cells were harvested using 0.05% trypsin with EDTA (ethylenediaminetetraacetic acid, Invitrogen; 25300) and re-suspended in media and seeded at a 1:3 ratio. Only cells from passages three to five were used for experiments. The culture method and passage number resulted in consistent expression levels of tenocyte markers including scleraxis, tenomodulin, and COL1A2 (appendix b).  117  A.1.2. Experimental condition and substances for experimental design Cells were cultured in 10% fetal bovine serum. Dexamethasone (Sigma; D4902) was used at concentrations from 1 to 400nM for mRNA analysis (6 and 12 h duration), and 1 to 100nM for protein analysis (12 h duration). The glucocorticoid receptor antagonist RU486, (Tocris; 1479), was used at concentrations of 1, 10 and 100nM for 12h simultaneously with dexamethasone. The cell were also treated with recombinant human TGF-β1 (R&D Systems, 240-B-002) and A-83-01 (Tocris, 2939) as the inhibitor of inhibitors of TGF- β signaling (Tocris, 2939) to investigate the function effect of the TGF- β pathway in ANGPTL4 expression.  RU486 and A83.01 were prepared in DMSO, and dexamethasone in absolute ethanol. Controls were incubated for the same durations as experimental conditions with the same volume of vehicle (ethanol/DMSO), which did not exceed 0.1%. The tendon cells also were stimulated with recombinant human IL-1β (R&D Systems; 201-LB) and IL-6 (206-IL) at concentrations of 0, 0.1, 1 and 10 ng/mL for 6 and 12 hours. Tendon cells were seeded on a BioFlex culture plate membrane treated with collagen I (BioFlex; BF-3001C) and were subjected to 10% cyclic strain (ST) with a frequency of 1 Hz for 2 hours per day as previously described (Backman et al. 2011). Before the third bout of cyclic strain, 10 nM dexamethasone was added to culture media of loaded cells and the same volume of vehicle (ethanol) were added to the unloaded cells and loaded cells. Then the tendon cells were incubated for 6 hours.     118  A.1.3. RNA isolation, reverse transcription, and qPCR Primary tendon cells were seeded on 6-well plates at a density of 1.2x105 in triplicates. At the termination of the experiments, total RNA was extracted using an RNA extraction kit (Qiagen; 74106). RNA was reverse transcribed using a High capacity cDNA Reverse Transcription kit (Applied Biosystems; 4368813) and a thermal cycler (Eppendorf Mastercycler EP gradient S, Eppendorf, North America). Quantitative PCR was performed using Sybr green and TaqMan probes for TAC1 and NKR1 respectively and data were normalized with GAPDH and18s rRNA genes. Also the Sybr green probes for IL-1α, IL-1β and IL-6 were used. The primer sequences for TAC1, IL-1α, IL-1β, IL-6 and GAPDH were designed and tested using NCBI/Primer-BLAST and GeneRunner (version 3.05) and synthesized by Life Technologies (Table A-1). Taqman probes for NKR1 and 18s from Life Technologies (#4351372 and #4352930E respectively) were used. The cycle threshold (Ct) values were normalized to the Ct values of housekeeping genes with stable expression (GAPDH and 18s) to obtain the relative gene expression; this value was expressed as fold change compared to controls harvested at each time point.    119   Table A-1 Oligonucleotide sequence of primers and amplicon sizes Target gene Forward primer sequence Reverse primer sequence Amplicon size (bp) TAC1 GATCAAGGAGGAACTGCCGGAGC GGAATCAGCATCCCGTTTGCCCA 109 IL-1α CGCCAATGACTCAGAGGAAGA AGGGCGTCATTCAGGATGAA 120 IL-1β AATCTGTACCTGTCCTGCGTGTT TGGGTAATTTTTGGGATCTACACTCT 78 IL-6 GGTACATCCTCGACGGCATCT GTGCCTCTTTGCTGCTTTCAC 81 GAPDH TCTTTTGCGTCGCCAGCCGAG TGACCAGGCGCCCAATACGAC  94 120  A.1.4. Immunocytochemistry (ICC) Cells cultured on 8-well chamber slides (BD Falcon; 354118) at a density of 1.5x104 cells/well were labelled with rabbit polyclonal antibodies against SP (Serotec, Oxford, UK; 8450-0004) or scleraxis (Abcam; ab58655). Cells were fixed in 2% paraformaldehyde in 0.1M phosphate buffer (pH 7.4) for 5min, washed in phosphate buffer (PBS) 4x3min and blocked in swine normal serum (1:20) for 15min. The culture slides were incubated with primary antibody (SP, concentration of 1:100; scleraxis, concentration of 1:25) for 60min at 37°C. After additional washing and blocking in normal serum the secondary antibody, TRITC- swine anti-rabbit, (Tetramethylrhodamine, Dako; R0156) at a concentration of 1:40, was incubated for 30min at 37°C. Finally, the cells were washed and mounted in Vectashield Hard Set Medium with DAPI (4',6-diamidino-2-phenylindole, Vector Laboratories, Burlingame, CA, USA; H-1500). The slides were examined using a Zeiss Axioskop 2 plus microscope with epifluorescence and an Olympus DP70 digital camera.  A.1.5. Immunoblotting Cells were cultured in 6-well plates at a density of 2x105 cells per well, in triplicates, and at the termination of the experiment, the triplicates were pooled and lysed in RIPA lysis buffer  supplemented with a protease inhibitor cocktail (Sigma; P1860, 1:200). The western blot process was performed as previously described (Backman et al. 2011). Briefly, lysate was separated by electrophoresis at 160 V for 45 min in a 12% TGX gel and subsequently transferred to a polyvinylidene fluoride transfer (PVDF) membrane at 100 V for 60 min. Following blocking, the membrane was incubated with the primary antibody against SP (Santa Cruz; sc-14104) at a concentration of 1:200 and afterwards the secondary antibody, donkey anti-goat HRP (Horseradish 121  peroxidase; Santa Cruz; sc2020) at a concentration of 1:1000. The bands were detected using a chemiluminescent HRP substrate (GE Healthcare, Little Chalfont, Buckinghamshire, UK; RPN2132) and visualized on high-performance chemiluminescence film (GE healthcare; 28-9068-38). The membranes were re-probed with an antibody against beta-actin to confirm equal total protein loading.  A.1.6. SP enzyme immunoassay (EIA) Cells were cultured in 6-well plates at a density of 2x105 cells per well in triplicates, then lysed in a lysis buffer (100 Mm Tris-HCI buffer pH 7.0 containing 1 M NaCl, 2% bovine serum albumin, 4 mM EDTA, 0.2% Triton X-100, 0.02% sodium azide), supplemented with a protease inhibitor cocktail at a concentration of 1:200. The lysates were centrifuged at 13,000 x g at 4°C for 15 min and the supernatant was collected. Protein concentration was determined with Protein Assay Dye Reagent Concentrate (Bio-Rad 500-0006). A total of 500 µg of total protein per sample was used to quantify SP concentration using a commercially available immunoassay kit (Phoenix Pharmaceuticals, CA, USA; EK-061-05) with a linear detection range of 0.07-2.24ng/ml. The absorbance was read at 450nm.  A.1.7. Statistical analysis All experiments were repeated using tendon cells derived from at least three separate patients. Data obtained from qPCR, EIA and Western blot experiments were analyzed with a one way analysis of variance, followed by Tukey's multiple comparison test.   122   A.2. Results A.2.1. Dexamethasone down-regulates SP expression (TAC1 mRNA) SP is encoded by the TAC1 gene. The level of TAC1 mRNA, as measured by qPCR, was significantly down-regulated after dexamethasone exposure in a time dependent manner (Figure A-2.a). This effect was specific to the examined gene, as evidenced by the fact that there was no significant effect of dexamethasone on the expression of NK1R mRNA (Figure A-2.b).   A.2.2. Dexamethasone reduces SP production The production of SP protein was significantly reduced after exposure of dexamethasone at concentrations of 1 and 100nM for 12hours as compared to untreated cells (Figure A-2.c). These data are consistent with the subjective immunocytochemistry evaluation (Figure A-2.d), as well as Western blot results showing that dexamethasone at concentrations of 1 and 100 nM reduced the expression of SP after 12 hours (Figure A-2.e, f).  A.2.3. Dexamethasone reduces SP expression through the glucocorticoid receptor The reduced expression of TAC1, following dexamethasone exposure, is significantly prevented by simultaneous exposure of the glucocorticoid receptor inhibitor, RU486 after 12 hours. At a concentration of 10 µM RU486, the effect of dexamethasone on TAC1 expression was 123  negated (Figure A-2.g). Stimulation of tendon cells with TGF- as an antagonist of TGF-beta pathway, had no significant effect on TAC1 mRNA expression (Figure A-2.h).  A.2.4. Dexamethasone reduces the expression of interleukins The tendon cells exposed to dexamethasone for 6 hours showed reduced expression of IL-1α, IL-1β and IL-6 (Figure A-2.i).  A.2.5. Dexamethasone inhibits the expression of SP induced by IL-1β Stimulation of tendon cells with IL-1β significantly also induced the expression of TAC1 (Figure A-2.j). However dexamethasone prevented the induction of TAC1 expression by IL-1β and inhibited SP expression in presence of IL-1β and IL-6 (Figure A-2.l).   A.2.6. Dexamethasone inhibits the expression of SP induced by cyclic strain The hamstring tendon cells subjected to cyclic loading showed significant upregulation of TAC1 expression compared to the non-strained cells. In contrast, adding dexamethasone at a concentration of 10 nM suppressed the induced expression of TAC1 by cyclic loading (Figure A-2.m). 124   125  Figure A-2. Dexamethasone decreases TAC1 expression without affecting the mRNA level of the preferred SP receptor (NK1R).  qPCR data for TAC1 (a) and NK1R (b) for tendon cells after 6 and 12 hours incubation with increasing concentrations of dexamethasone. EIA (c), ICC (d) and Western blot (e, f) demonstrated reduced SP protein level in the cell lysates of tendon cells after 12 hours incubation with 1 and 100 nM dexamethasone. (g) The reduction of TAC1 expression by dexamethasone was specifically mediated through the glucocorticoid receptor as evident by the abolished effect of dexamethasone when the glucocorticoid receptor antagonist, RU486, at a concentration of 10uM, was simultaneously applied.  The effect of RU486 was shown to be significantly dose-dependent. (h) TGF-beta and the inhibitor of TGF-beta had no significant effect on TAC1 mRNA expression. (i) Dexamethasone after 6 hours incubation reduced the expression of IL-1α/β and IL-6 in tendon cells. Recombinant human IL-1β induced the expression of TAC1 after 6 and 12 hours incubation (j), but the induction by recombinant human IL-6 was not significant (k). (l) Incubation of hamstring tendon cells with dexamethasone for 12 hours at a concentration of 2.5 nM (with or without 1 ng/ml of IL-1β and IL-6) inhibits TAC1 expression. (m) 6 hours incubation of hamstring tendon cells with 10 nM dexamethasone prevented induction of TAC1 in tendon cells subjected to cyclic strain (ST) for 2 hours per day for 3 days (mean ± SE; ns, not significant; *, P < 0.05; **, ** P ≤ 0.01; ***, P ≤ 0.001). (Panels “b, g-m” show the data generated from the hamstring tendon cells; “c-f” are from the Achilles tendon cells and panel “a” is from both types).  A.3. Discussion In the current study we asked whether corticosteroids might influence the expression levels of SP by tendon fibroblasts. Our data confirm that exposure of human cultured tendon fibroblasts to dexamethasone reduces SP mRNA and protein. Our results further show that the reduction of SP following dexamethasone exposure is mediated through the glucocorticoid receptor. The inhibition of SP mRNA by dexamethasone occurs even in presence of the stimulatory effect of IL-126  1β, IL-6 and mechanical loading.  The results of the study could help explain why glucocorticoid treatments lead to acute pain relief effect in patients suffering from tendinopathy.  The etiology of chronic tendon pain is not completely understood, but evidence is accumulating that low grade inflammation probably plays a role, as in osteoarthritis. In the rotator cuff, pain levels are remarkably correlated with tissue levels of SP (Gotoh et al. 1998), and SP expression is also present in the tendons and bursae of people with greater trochanteric pain syndrome (Fearon et al. 2014). Interestingly, although peripheral SP-containing nerves are known to exist and are more extensive in chronically injured tendon, local tendon fibroblasts also appear to be a local source of SP; this finding is in keeping with recent studies demonstrating that SP is locally up-regulated by repetitive overuse in tendon fibroblasts both in vivo and in vitro (Schubert et al. 2005, Backman et al. 2011).  In cases of chronic tendinopathy, a lack of inflammatory cellular infiltration in surgical biopsies has led some to challenge the involvement of inflammation in the etiology of this disorder. However, several studies have shown inflammatory reactions in the early development of tendinopathy (Kvist et al. 1988). More recently, macrophages and T and B lymphocytes have been detected not only in the early phase, but also in chronic tendinopathy. Other studies have demonstrated increased levels of COX-2, and IL-6 (Legerlotz et al. 2012). Evidence thus points in the direction of one or more roles for inflammation during the development of tendinopathy; even if a particular inflammatory cell, activity or cytokine is not detected at a particular time point in a given study it does not necessarily imply that the pathology is devoid of inflammation.  Dexamethasone is a commonly used glucocorticoid.  Ihara et al have shown that dexamethasone down-regulated the preferred SP receptor (NK1R) mRNA in a pancreatic acinar 127  cell line (Ihara et al. 1990). In contrast, our data did not show any significant effect of dexamethasone on NKR1 in tendon cells.  Dexamethasone is an agonist of the glucocorticoid receptor (GR); binding of dexamethasone to the GR facilitates its translocation to the nucleus where trans-repression of transcription factors such as nuclear factor-kB and activator protein 1 by GR negatively regulates inflammatory activity (Tsurufuji et al. 1979, Vandevyver et al. 2013). King et al have shown that GR transactivation by dexamethasone is involved in glucocorticoid-mediated repression of many IL-1β induced inflammatory genes (King et al. 2013). Our results also showed that dexamethasone inhibits the expression of SP induced by IL-1β. Our study therefore supports the notion of a negative regulatory effect of GR on endogenously produced inflammatory mediators, however further studies are needed to examine the crosstalk between GR and other transcription factors in modulation of SP in tendon cells.        128  Appendix B. Tendon cell characterization In this section, I present the data of human tendon cell characterization based on gene expression analysis.  B.1. Methods After each passage, the expression levels of aggrecan, COL1A2, decorin, scleraxis, tenomodulin and nucleostemin were quantified using qPCR in order to characterize the tendon cells’ phenotype. Total RNA was extracted from tendon cells using the RNeasy Plus Universal Mini kit (Qiagen, Germany, #73404) and reverse transcribed to cDNA with a High Capacity cDNA Reverse Transcription Kit (Applied Biosystems™, USA, #4368814). The primer sequences were designed and tested using NCBI/Primer-BLAST and GeneRunner (version 3.05) and synthesized by Life Technologies (Table B-1). The gene expression levels were determined by FastStart Universal SYBR Green Master mix (Roche Applied Science, USA, # 04913914001) using GAPDH as internal control.  129  Table B-1. Oligonucleotide sequence of primers and amplicon sizes of the gene markers for tendon cells. Target gene Forward primer sequence Reverse primer sequence Amplicon size (bp) Agrecan GTGTAAAAAGGGCACAGCCAC ACCAACGATTGCACTGCTCT 478, 360 COL1A2 AGTGTCCACGTCCTCAAAAAGA CAGCAAAGTTCCCACCGAGA 599 Decorin GTCACAGAGCAGCACCTACC TTGTCCAGACCCAAATCAGAACA 378 Scleraxis AAGAAAAGCCAGCGCAGAAAGTTC TCTGCACCTTCTGCCTCAGCAA 320 (Qi et al. 2012) Tenomodulin GAAGCGGAAATGGCACTGATGA TGAAGACCCACGAAGTAGATGCCA 82 Nucleostemin GGGAAGATAACCAAGCGTGTG CCTCCAAGAAGTTTCCAAAGG 98 GAPDH TCTTTTGCGTCGCCAGCCGAG TGACCAGGCGCCCAATACGAC 94 130  B.2. Result Based on the increased expression of COL1A2, scleraxis and tenomodulin along with no increased expression of nucleostemin (associated with pluripotent tendon-derived progenitor cells), the cultured tendon cells from passages 3 to 5 were selected for experiments.  Figure B-1. The expression of tendon cell markers during different passages.  

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