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Valvular interstitial cell transformation : implications for aortic valve calcification Boroomand, Seti 2014

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     Valvular Interstitial Cell Transformation: Implications for Aortic Valve Calcification by Seti Boroomand B.Sc., Shahid Beheshti (Melli) University, Iran, 1999     A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Pathology and Laboratory Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  June 2014   © Seti Boroomand, 2014 ii  Abstract Aortic valve stenosis (AVS) involves the transformation of valvular interstitial cells (VIC) into an osteoblastic phenotype. Such valvular disease is mostly associated with both thickening and calcification of the valve cusps, which is accompanied by inflammation and remodeling of the tissue. This process is mediated by the VIC that carry out an impressive array of functions throughout the calcification process. For this dissertation, I hypothesized that in AVS, VIC transform from a myofibroblast phenotype to osteoblast-like cells and that the canonical Wnt and TGFβ pathways and vitamin D3 interactively and collaboratively contribute to these phenomena. In order to test this hypothesis, I established an in vitro model of calcification by culturing human primary VIC in a pro-calcification conditioned medium. Calcified cells display several molecular characteristic features of human AVS, including increased levels of alkaline phosphatase and the formation of calcium nodules. These changes increased over time and peaked at 28 days of treatment. To define possible mechanisms of AVS, I first characterized human VIC in regards to the process of calcification. I showed for the first time in vitro that these VIC express bone specific markers, the characteristic of normal osteoblasts. To determine the factors involved in osteoblastic transformation in this model, I examined WNT3A and TGFβ, known to be involved in normal bone formation. Both calcified human aortic valve tissues and VIC express excess WNT3A and TGFβ1. Adding WNT3A and TGFβ1 to the VIC cultures increased the levels of cell mineralization. Further, the addition of DKK1, the WNT3A antagonist, decreased VIC calcification in vitro. By using various combinations of WNT3A, TGFB1 and DKK1, I made the novel observation that the suppression of DKK1 by TGFB1 allowed WNT3A to drive calcification in VIC in vitro.  Finally, I examined the role vitamin D3 iii  that is associated with vascular calcification in rats. Vitamin D3 can up-regulate VIC calcification in vitro, however its mechanism of action appears to be independent of the Wnt and TGFβ pathways. In conclusion, the canonical Wnt and TGFβ pathways function interactively through DKK1 to transform VIC to osteoblast-like cells and vitamin D3 promotes this process in an independent manner.  iv  Preface This dissertation contains chapters which are based on in preparation manuscripts: A version of chapter 3 is in preparation [Boroomand S, Walker D, Rahmani M, Allahverdian S, Kaur J, Samra A, Elliott M, Seidman M, McManus B. Canonical WNT activation increases valvular interstitial cell associated calcification via TGFβ mediated down-regulating DKK1, submitted February 2014].  A version of Chapter 4 is in preparation [Boroomand S, Meredith A, Luo Z, Samra A, McManus B. 1α, 25(OH) 2D3 regulates the valvular interstitial cell calcification independent from the TGFβ and the canonical WNT pathways].  All the figures in Chapter 1 (Introduction) are made and prepared by myself. In all manuscripts and chapters, I was responsible for designing and performing the experiments, as well as writing the manuscripts. Co-authors Drs. David Walker, David Granville and Michael Seidman assisted with scientific guidance as well as proof-reading the manuscript. Dr. Bruce McManus helped formulate the research plan, the experimental approach, and the overall focus on mechanisms of valve disease. He also reviewed and revised all documents pertinent to this thesis.  Ethics approval was obtained from the UBC/Providence Research Ethics Board, certificate numbers B05-0185, H05-50208, B09-0220, H09-01571.  v  Table of Contents Abstract .......................................................................................................................................... ii Preface ........................................................................................................................................... iv Table of Contents ...........................................................................................................................v List of Tables ..................................................................................................................................x List of Figures ............................................................................................................................... xi List of Abbreviations and Acronyms ....................................................................................... xiii Acknowledgements .................................................................................................................... xvi Dedication ................................................................................................................................. xviii CHAPTER 1: INTRODUCTION .........................................................................................................1 1.1 Aortic Valve Stenosis ..................................................................................................... 1 1.1.1 Metastatic Calcification .............................................................................................. 2 1.1.2 Dystrophic Calcification ............................................................................................. 2 1.1.3 Characteristics of Aortic Valve Calcification ............................................................. 2 1.1.4 Pathogenesis of Aortic Valve Stenosis ........................................................................ 3 1.1.5 Etiology and Pathogenesis of Aortic Valve Stenosis .................................................. 5 1.2 Structural and Functional Characteristics of Normal vs. Calcified Aortic Valves ......... 6 1.3 Valvular Interstitial Cell ................................................................................................. 9 1.4 Wnt Signaling Pathway................................................................................................. 12 1.4.1 Wnt in Normal Bone Development ........................................................................... 12 1.4.2 Wnt in Vascular Calcification ................................................................................... 13 1.4.3 Wnt in Valvular Calcification ................................................................................... 15 vi  1.4.4 Dickkopf1 (DKK1), an WNT3A Antagonist .............................................................. 17 1.5 TGFβ Signaling in Calcification of Valves .................................................................. 19 1.6 1, 25 (OH)2 D3 (Vitamin D3)....................................................................................... 22 1.6.1 1, 25 (OH)2 D3 in Valvular Calcification ................................................................ 22 1.7 Thesis Rationale, Hypothesis and Specific Aims ......................................................... 25 CHAPTER 2: HUMAN VALVULAR INTERSTITIAL CELLS: A MODEL TO STUDY AORTIC VALVE CALCIFICATION ...............................................................................................................28 2.1 Introduction ................................................................................................................... 28 2.2 Materials and Methods .................................................................................................. 31 2.2.1 Cell and Tissue Procurement .................................................................................... 31 2.2.2 Antibodies ................................................................................................................. 32 2.2.3 Isolation of Human Valvular Interstitial Cells ......................................................... 33 2.2.4 Cell Culture Treatment ............................................................................................. 33 2.2.5 Western Blots ............................................................................................................ 34 2.2.6 Histochemical Staining for Alkaline Phosphatase .................................................... 35 2.2.7 Von Kossa Staining for Calcium ............................................................................... 35 2.2.8 Immunocytochemistry ............................................................................................... 36 2.2.9 Alkaline Phosphatase Activity Assay ........................................................................ 36 2.2.10 Statistical Analysis ................................................................................................ 37 2.3 Results ........................................................................................................................... 38 2.3.1 Calcium Nodules Are Formed in Vascular SMC and VIC ....................................... 38 2.3.2 Alkaline Phosphatase and Von Kossa Staining Are Strongly Positive in Calcified VIC ................................................................................................................................... 41 vii  2.3.3 Alkaline Phosphatase Activity Is Amplified in Calcified VIC ................................... 44 2.3.4 Osteogenic Treatment Induces Osteoblast Transformation in VIC .......................... 46 2.3.5 Bone Regulators Are Overexpressed in Calcified VIC ............................................. 49 2.4 Discussion ..................................................................................................................... 51 CHAPTER 3: CANONICAL WNT ACTIVATION INCREASES VALVULAR INTERSTITIAL CELL ASSOCIATED CALCIFICATION VIA TGFΒ-MEDIATED DOWN-REGULATION OF DKK1 ........54 3.1 Introduction ................................................................................................................... 54 3.2 Materials and Methods .................................................................................................. 57 3.2.1 Tissue Procurement .................................................................................................. 57 3.2.2 Antibodies ................................................................................................................. 57 3.2.3 Histology and Immunohistochemistry....................................................................... 58 3.2.4 Isolation of Human Valvular Interstitial Cells ......................................................... 58 3.2.5 Tissue and Cell Culture ............................................................................................ 59 3.2.6 OsteoImage Mineralization Assay ............................................................................ 60 3.2.7 Alkaline Phosphatase Activity Assay ........................................................................ 60 3.2.8 Western Blots ............................................................................................................ 60 3.2.9 Immunocytochemistry on VIC ................................................................................... 61 3.2.10 Statistical Analysis ................................................................................................ 62 3.3 Results ........................................................................................................................... 63 3.3.1 WNT3A and TGFβ1 Are Up-regulated in AVS ......................................................... 63 3.3.2 Exogenous WNT3A Induces an Osteoblast Phenotype in Cultured Human VIC ..... 68 3.3.3 Canonical Wnt Signaling Regulates Human VIC Calcification ............................... 71 3.4 Discussion ..................................................................................................................... 79 viii  CHAPTER 4: 1Α, 25(OH)2D3 REGULATES VALVULAR INTERSTITIAL CELL CALCIFICATION .. ........................................................................................................................................................82 4.1 Introduction ................................................................................................................... 82 4.2 Materials and Methods .................................................................................................. 85 4.2.1 Tissue Procurement .................................................................................................. 85 4.2.2 Isolation of Human Valvular Interstitial Cells ......................................................... 85 4.2.3 Treatment of Cell Cultures........................................................................................ 86 4.2.4 Histology and Immunohistochemistry....................................................................... 86 4.2.5 Western Blots ............................................................................................................ 87 4.2.6 Alkaline Phosphatase Activity Assay ........................................................................ 88 4.2.7 OsteoImage Mineralization Assay ............................................................................ 88 4.2.8 Statistical Analysis .................................................................................................... 89 4.3 Results ........................................................................................................................... 90 4.3.1 VDR Expression in Primary Valvular Interstitial Cells of Human Aortic Valves .... 90 4.3.2 Vitamin D3 and the Phenotype Shift of VIC ............................................................. 94 4.3.3 Vitamin D3 Increases the Expression of Bone Specific Markers ............................. 97 4.3.4 Vitamin D3 Can Induce VIC Calcification ............................................................... 99 4.3.5 The Role of Vitamin D3 in Calcification of VIC Is Independent of β-catenin and SMAD2 ................................................................................................................................ 101 4.4 Discussion ................................................................................................................... 104 CHAPTER 5: OVERALL SYNOPSIS .............................................................................................109 5.1 Discussion ................................................................................................................... 109 5.2 Conclusion .................................................................................................................. 113 ix  5.3 Future Directions ........................................................................................................ 116 Bibliography ...............................................................................................................................118  x  List of Tables Table 1. Proportion and intensity scores for WNT3A, TGFβ1 and DKK1 immunohistochemical staining in human aortic valve cusps. ........................................................................................... 64 Table 2. In vivo proportion and intensity scores for VDR immunohistochemical staining in human aortic valve cusps. ............................................................................................................. 92  xi  List of Figures Figure 1. Examples of normal and calcific valves. ......................................................................... 4 Figure 2. Artistic representative of gross anatomy of the aortic valve. .......................................... 7 Figure 3. Artistic representation of an aortic valve cusp in different orientations. ........................ 8 Figure 4. Potential pathway underpinning calcification of aortic valve. ...................................... 11 Figure 5. Overview of the canonical Wnt signaling and a model of DKK1 interaction with the canonical Wnt pathway. ................................................................................................................ 18 Figure 6. Overview of the TGFβ pathway. ................................................................................... 21 Figure 7. Overview of the normal vitamin D3 pathway. .............................................................. 24 Figure 8. Calcium nodule formation in different cell type at various timepoints. ........................ 40 Figure 9. Alkaline phosphatase and von Kossa staining in VIC and vSMC. ............................... 42 Figure 10. Alkaline phosphatase localization in VIC. .................................................................. 43 Figure 11. Alkaline phosphatase activity in VIC, vSMC and WPMY-1 cells at different timepoints. ..................................................................................................................................... 45 Figure 12. Expression of osteoblast markers in vSMC................................................................. 47 Figure 13. Expression of osteoblast markers in VIC. ................................................................... 48 Figure 14. Expression of pro-osteogenic mediators (WNT3A and TGFβ1) in calcified VIC. ..... 50 Figure 15. Immunohistochemistry of human aortic valves with different degrees of calcification for WNT3A, TGFβ1, and DKK1. ................................................................................................. 65 Figure 16. Cell calcification and alkaline phosphatase activity levels in cell treated with TGFβ1, WNT3A or both. ........................................................................................................................... 69 Figure 17. Expression of RUNX2 and osteopontin in treated VIC. ............................................. 70 xii  Figure 18. LiCl mechanism of action. .......................................................................................... 72 Figure 19. Calcification levels in WNT stimulated cell cultures. ................................................. 73 Figure 20. Effect of DKK1 treatment on calcification and alkaline phosphatase activity levels in WNT3A treated cells. ................................................................................................................... 74 Figure 21. Nuclear translocation of β-catenin treated with TGFβ1 in human VIC. ..................... 77 Figure 22. Cells treated with TGFβ1 express less DKK1. ............................................................ 78 Figure 23. Working model of the interaction between the Wnt and TGFβ pathways in valvualr calcification. .................................................................................................................................. 81 Figure 24. Vitamin D receptor (VDR) is present in human aortic valve and valvular interstitial cells. .............................................................................................................................................. 91 Figure 25. Vitamin D receptor (VDR) is present in human aortic valve and valvular interstitial cells. .............................................................................................................................................. 93 Figure 26. Vitamin D changes the morphology of the VIC. ......................................................... 95 Figure 27. αSMA protein expression. ........................................................................................... 96 Figure 28. Vitamin D shifts VIC phenotype toward osteoblast-like cells. ................................... 98 Figure 29. Vitamin D increases alkaline phosphatase activity and calcification levels in vitro. 100 Figure 30. Vitamin D3 treatment does not alter the expression of pSMAD2 and AXIN2 in VIC...................................................................................................................................................... 103 Figure 31. Putative mechanism of action of signaling pathways involved in valvular calcification...................................................................................................................................................... 115    xiii  List of Abbreviations and Acronyms  αSMA  Alpha-smooth muscle actin AVA  Aortic valve area AVC  Aortic valve calcification AVS  Aortic valve stenosis BMP  Bone morphogenic protein BMP2  Bone morphogenic protein 2 BSA  Bovine serum albumin DAPI  4', 6-diamidino-2-phenylindole DKK1  Dickkopf 1 DMEM Dulbecco’s modified eagle’s medium DVL  Dishevelled ECM  Extra cellular matrix FBS  Fetal bovine serum FZD  Frizzled GSK3β Glycogen synthase kinase 3-beta H & E  Hematoxylin and eosin HEK  Human embryonic kidney HRP  Horseradish peroxidase IHC  Immunohistochemistry IIAM  International institute for advancement of medicine LDL  Low density lipoprotein xiv  LEF  Lymphoid enhancer factor LiCl  Lithium chloride LRP 5/6 Low density lipoprotein receptor-related protein MCDB -131 Molecular and cellular developmental biology-131 MOSLB Modified oncogene science lysis buffer MSC  Mesenchymal cell NR  Nuclear receptor OPG  Osteoprotegerin OPN  Osteopontin pNNP  p-nitrophenyl phosphate PBS  Phosphate buffer saline REB  Research ethics board RFU  Relative fluorescent unit  R-SMAD Receptor-regulated SMAD smGM  Smooth muscle growth medium SAVR  Surgical aortic valve replacement SE  Standard error SMC  Smooth muscle cell SEM  Standard error of the mean TCF  T-cell factor TGFβ  Transforming growth factor beta TGFβ1  Transforming growth factor beta-1 TNFα  Tumor necrosis factor-alpha xv  U/L  Unit/liter vSMC  Vascular smooth muscle cell VDR  Vitamin D receptor VIC  Valvular interstitial cell  xvi  Acknowledgements I would like to thank many people who helped me along the way, in particular my supervisor, Dr. Bruce McManus, who supported me from the beginning and provided me with the opportunity to fulfill my dream in pursuing a PhD. It has been a challenging but rewarding experience. I would like to thank the chair of my advisory committee, Dr. David Walker, without your support this thesis seemed impossible.  I would like to thank the members of my advisory committee for their valuable support and constructive advice through the years: Drs. David Granville, Darryl Knight, and Bruce Verchere. To my fellow laboratory mates and good friends at the Centre, Lisa Ang, Anna Meredith, Dorota Stefanowicz, Amrit Samra, Sima Allahverdian, Maziar Rahmani, Mitra Esfandiarei and my childhood friend, Faranak Mahmoodi. You made the last few years the most memorable time of my life.  I would like to thank my family for the endless support and encouragement, especially my mother, Manizheh, you taught me the value of learning and education and you were always my role model in every aspect. My aunts: Minoo, Shahnaz, Jamileh and my only uncle, Ali, you are always in my heart. To other family members: Dada, Kianoush, Nazli, Shaheen, Amir, farinaz and Nezhaadi. I would also like to thank my little son, Arya, you are my inspiration, and last but not least my little furry friend, Mickey. I am also grateful to my colleagues at the Centre for Heart and Lung Innovation for their willingness to help throughout the years: Elizabeth Walker, Zongshu Luo, Jon Carthy, Farshid S. Garmaroudi, David Lin, David Marchant, Jaspreet Kaur, and Michael Seidman. I would like to acknowledge the members of Heart Pathology Registry at the Centre for Heart and Lung xvii  Innovation throughout the years with special thanks to Liz Matzke, Adrian Fung, and Joanne Kwan.  xviii  Dedication To my mother and grandmother Manizheh and Nooshafarin. Everything I have I owe to you.  To my son, Arya, everything I do is for you.  1  Chapter 1: INTRODUCTION  1.1 Aortic Valve Stenosis Aortic valve stenosis (AVS) is the most common cause of acquired valvular heart disease, often leading to aortic valve replacement in developed countries. The greatest risk factors for AVS are aging and consumption of a high-calorie diet [3, 4]. The prevalence of AVS has been estimated at 2-3% in patients ≥65 years of age and increasing to 4% in those ≥75 years of age. Mortality for untreated symptomatic severe AVS is as much as 50–60% at two-year post-diagnosis in high-risk patients [5, 6]. Surgical aortic valve replacement (SAVR) is currently the preferred treatment for patients with severe symptomatic AVS [7]. Without surgery, prognosis is extremely poor, with a 3-year survival rate <30%. However, 33% of all patients aged ≥75 years with severe AVS are declined as candidates for surgery due to a high risk of operative mortality [8, 9]. While mortality for untreated symptomatic severe AVS is high, even those patients who ultimately undergo surgical valve replacement are still at a high risk of morbidity and mortality from the procedure itself [5-9]. Valvuloplasties, not requiring open heart surgery, have become more common in cases of valve leakage, whereby complications associated with artificial valves and anticoagulant therapy can be avoided. Advances in less invasive catheter-based valve techniques seem to be as good as conventional ones. In the treatment of AVS, artificial valve replacement via catheter has become almost routine for patients having an excessive risk for open heart surgery [7, 10, 11]. As the population ages, the prevalence of aortic stenosis inevitably rises. Aortic stenosis in aging adults, and the management questions it poses, will be increasingly common. Demographic projections indicate that AVS prevalence will increase more 2  than three-fold by 2050 in North America alone, placing an ever larger burden on health services and emphasizing the urgent need for improvements in disease prevention and treatment to curtail spiraling social and economic costs [12, 13]. 1.1.1 Metastatic Calcification Metastatic calcification is a form of calcification that occurs in normal tissue due to elevated serum level of calcium (hypercalcemia) when calcium phosphorus metabolism is impaired. It may happen throughout the body, hence the term “metastatic”, however it mostly affects interstitial tissues in kidney, lungs and gastric mucosa [14]. The alkaline components excreted from these tissues make them more susceptible to calcification [15]. 1.1.2 Dystrophic Calcification Dystrophic calcification is a form of calcium deposition that occurs in areas of necrosis [14]. This is the form of calcification observed in atherosclerotic plaques and aging or damaged heart valves. In dystrophic calcification, calcium binds to the phospholipid in the cell membrane. Calcium binds to the phosphate group and forms calcium phosphate. This compound is similar to the hydroxyapatite seen in regular bone growth. Calcium phosphate deposition forms microcrystals which propagate and lead to more calcification. Ongoing dystrophic calcification can result in organ dysfunction [16]. 1.1.3 Characteristics of Aortic Valve Calcification Clinically, aortic valve calcification is a progressive disorder characterized by calcification of the aortic valve leaflets in the absence of rheumatic heart disease [17]. The 3  disease spectrum ranges from mild valve thickening often without obstruction of blood flow known as aortic valve sclerosis to severe calcification with impaired leaflet motion termed aortic stenosis. Thus aortic valve calcification is divided, on a functional basis, into aortic sclerosis, in which the leaflets do not obstruct left ventricular outflow, to aortic stenosis, in which cusps stiffen and obstruction of left ventricular outflow is present by virtue of calcification (Figure 1). 1.1.4 Pathogenesis of Aortic Valve Stenosis Aortic valve stenosis (AVS) is the end stage of valvular calcification when the cusps are thickened and stiff enough to cause obstruction in blood flow (Figure 1B). The risk factors most closely linked to AVS are also those that have been associated with atherosclerosis, such as aging, male gender, smoking, hypertension, elevated lipoproteins and diabetes [18]. Beyond these shared risk factors, several similar pathological patterns are also observed between AVS and atherosclerosis [19, 20]. In both conditions the pathological lesion is characterized by a more permeable endothelium, activation of inflammatory processes, extracellular matrix remodeling and mineralization [21, 22]. In many pathological events involving chronic inflammation, calcification of the soft tissue has been reported. Both atherosclerosis and valvular calcification are associated with chronic inflammation and concomitant calcification of the soft tissue. However in AVS, calcification is more so the end stage of the disease, whereas in atherosclerosis calcification can begin at any point of plaque development process and the sequence of events in plaque formation does not appear to determine calcium deposition or progression of calcification as it does in valvular disease [23].   4    Figure 1. Examples of normal and calcific valves. The normal aortic valve has three diaphanous cusps with open commissures and with no evidence of calcium deposit in the sinuses of Valsalva (▼) (A). A calcified aortic valve is identified by the calcium deposits on the sinuses of Valsalva (*) (B and C). Calcified aortic valve disease is defined, on a functional basis, into aortic stenosis, in which cusps stiffen and obstruction to left ventricular outflow is present (B) and aortic sclerosis (C), in which the thickened leaflets do not obstruct left ventricular outflow despite accompanying degrees of calcification.               5  1.1.5 Etiology and Pathogenesis of Aortic Valve Stenosis The most common cause of AVS is aortic sclerosis, when the valve leaflets thicken as a result of fibrosis, leading to calcification and progression to AVS with significant obstruction that resembles atherosclerosis. This process may be accompanied by low density lipoprotein (LDL) deposition, and active inflammation of the valve tissues. The most common cause of AVS in younger adults is congenital bicuspid aortic valve, a condition that affects men more than women [24]. Rheumatic fever is the most common cause of AVS in developing countries and affects all age groups[25].                     The main subject of this thesis is the form of AVS that is described as degenerative aortic valve stenosis which develops slowly and is not diagnosed in patients until they are in their 70s to 90s. Currently the pathophysiological mechanisms underlying AVS are still not fully understood. Previously, it was believed that simple passive accumulation of calcium in the cusps of valves causes disease, however recent studies demonstrate that calcification occurs as a result of an active inflammatory process that in turn results in activation of valvular interstitial cells and eventually their transformation into an osteoblast-like phenotype [26, 27]. The hallmark of this inflammatory process in aortic valve pathogenesis is the formation of lesions including mostly macrophages, T lymphocytes and mast cells [26, 28, 29]. Oxidized LDL also accumulates in aortic valve lesions and this accumulation is mediated partly by extracellular matrix (ECM) proteoglycans and valvular interstitial cells (VIC) [30]. This process eventually leads to the transformation of VIC from that of a myofibroblast into that of an osteoblast-like cell. The precise mechanisms of this process remain unknown. 6  1.2 Structural and Functional Characteristics of Normal vs. Calcified Aortic Valves  Normal aortic valves consist of three semilunar and diaphanous cusps, with open commissures and no evidence of calcium deposits on the cusp surface, sinuses of Valsalva, nor in the nodule or body of Arantius (Figures 2 and 3). Aortic valve cusps are composed of three morphologically distinct tissue layers: the fibrosa, spongiosa, and ventricularis [31] (Figures  3 A and B). The endothelium covering the valves forms a continuum with the aortic endothelium (Figure 3B). The ventricularis layer faces the left ventricle and is composed of a dense network of elastic fibers aligned radially and a collagenous component oriented in a circumferential direction (Figure 3). The layer facing the aorta is the fibrosa and consists mainly of collagen fibers. These fibers are also arranged in a circumferential direction (Figures 3 A and B). Between the fibrosa and ventricularis is the spongiosa, a loosely organized tissue, containing collagen, elastin and proteoglycans, populated by myofibroblasts known as valvular interstitial cells (VIC) [17]. In summary, all three layers are composed of collagen fibers, elastic sheets, proteoglycan matrix and valvular interstitial cells, together providing the necessary biochemical and mechanical properties for proper valve function [31-33].  In this regard, type I collagen fibers are arranged circumferentially while sheets of elastin are oriented radially.       7           Figure 2. Artistic representative of gross anatomy of the aortic valve. The aorta is laid open here to show the semilunar cusps of the aortic valve. The valve has three cusps (left, posterior and right) that surround the orifice of the aorta (aortic opening) distal to the left ventricle. They are similar in structure to those of pulmonary valves,but the cusps are larger, thicker, and stronger; the lunulae are more distinct, and the noduli or corpora Arantius are thicker and more prominent. Opposite the valves the aorta has slight bulbous dilatations called aortic sinuses (sinuses of Valsalva), which are larger than those at the origin of the pulmonary artery from the right ventricle.           8  A.           B.          Figure 3. Artistic representation of an aortic valve cusp in different orientations. The cusps of normal aortic valves are very smooth and thin. At the apex of the free edge of each leaflet there is a fibrous nodule named nodulus Arantius (A). A radial cross section of the aortic valve shows the three different layers: the fibrosa on the aortic side, the spongiosa in the middle and the ventricularis on the ventricular side of the valve. An endothelial layer covers the underlying tissue (B). Leaflets making up the aortic valve have distinctive fibrous layers: the superficial aortic layer (fibrosa) contains large collagen fibers oriented in a circumferential direction. The central layer is also formed of collagen fibers and in the same direction but are smaller in diameter.  This layer (spongiosa) also contains a loose connective tissue of proteoglycans and frequent VIC. The last ventricular layer (ventricularis) is made of elastic sheets oriented, for the most part, in the radial direction (B). 9  1.3 Valvular Interstitial Cell Within all three layers of healthy aortic valve tissues there are interstitial myofibroblast-like cells that exhibit characteristics in common with both fibroblasts and smooth muscle cells [34]. Immunohistochemical studies of aortic valve tissues demonstrate that these valvular interstitial cells (VIC) are not isolated to a specific layer but are present throughout the three layers of valve tissue [35]. VIC maintain the structural integrity of the healthy leaflet. In normal valves VIC exhibit a quiescent phenotype, but they are activated with the onset of injury and/or remodeling of valves.  In fact there are five different VIC phenotypes in aortic valve leaflets with specific functions essential for normal physiology and  pathological processes. These phenotypes are referred to as mesenchymal, quiescent, activated, progenitor, and osteoblastic VIC [36]. VIC exhibit plasticity properties and may transform from one phenotype to another during  pathological processes [37].  Among  VIC phenotypes, activated VIC express alpha smooth muscle actin (αSMA) and play active roles in wound healing and valve repair processes including proliferation, migration, matrix remodeling and creation of abnormal hemodynamic or mechanical forces [38]. Although VIC function in aortic valves is not fully understood, it is believed that they may play an important role in wound healing through ECM deposition by virtue of capabilities such as ECM synthesis, contractility and calcification in response to insult [34, 35] .  Although the anatomy and histology of heart valves is well described (Figure 3), the cellular and molecular processes that regulate valve structure and function in health and disease still remain unclear. Valvular diseases primarily involve thickening and calcification of the valves and are associated with chronic inflammation and remodeling of the tissue [17, 39]. These 10  pathogenic processes are most likely mediated by VIC as they are the most prevalent cell type in the valvular interstitum and carry out an array of functions characteristic of wound healing following injury; as such they may play an integral part in the reparative processes [40]. VIC play a major role during an inflammatory response. They are capable of synthesizing and secreting several cytokines, chemokines and other inflammatory mediators [21, 41, 42]. Some of these inflammatory mediators secreted by VIC may recruit leukocytes during the inflammation and repair process [21, 43-45]. The venerable response-to-injury hypothesis suggests that valvular thickening is a consequence of valvular remodeling and repair processes. VIC produce and may respond to a variety of growth factors such as TGFβ and WNT3A necessary for cell proliferation, migration and differentiation [46-48]. The diverse biological processes such as contractile capability, cell communication and secretory properties that VIC are capable of have been shown to be critical for proper function of aortic valves [49]. Dysregulation of these normal cellular processes leads to chronic inflammation and calcification that eventually cause valve stenosis [35, 46, 50, 51]. Thus, by nature of their ability to modulate the inflammatory response and invoke a number of reparative processes through the secretion of some mediators such as WNT3A and TGFβ1, VIC represent a key cell type in understanding injury, inflammation and repair events in valve disease. A potential pathway that underpins aortic valve calcification is summarized in Figure 4.          11        Figure 4. Potential pathway underpinning calcification of aortic valve. 1) Monocyte and macrophages infiltrate endothelium and release cytokines that act on valvular myofibroblasts (VIC), which in turn promote proliferation and matrix remodeling. 2) LDL that is taken into the tissue is oxidatively modified and taken up by macrophages to form foam cells. 3) Foam cells express TGFβ and other cytokines and chemokines that act on valvular myofibroblasts. 4) A sub-population of VIC in the fibrosa layer of the valve becomes activated in response to these cytokines and synthesizes ECM. 5) A subset of valvular myofibroblasts transform into an osteoblast-like phenotype that is capable of calcium nodule formation.         12  1.4 Wnt Signaling Pathway WNT protein, derived from Drosophilia Wingless and the mouse Int-1 genes, is a member of a larger family of cysteine-rich secreted glycoproteins [52]. This family of proteins that activates cell surface receptor-mediated signaling pathways, plays a role in a variety of processes that involve embryonic cell fate, proliferation, migration, adhesion, survival, apoptosis, cell differentiation and bone growth [53-56]. WNT signaling pathways have been classified as “canonical” on the basis of their ability to inhibit β-catenin degradation and “non-canonical” when β-catenin degradation is permitted. WNT3A protein binds the Frizzled receptor and its co-receptor LRP5/6 (lipoprotein receptor-related proteins 5 or 6) to activate an intracellular signaling cascade that controls β-catenin degradation [55]. In the absence of WNT ligand, β-catenin is targeted for ubiquitin-mediated degradation [57]. In the presence of WNT ligand stimulation, the canonical signaling pathway triggers a series of phosphorylation events that lead to the accumulation of cytosolic β-catenin, which then translocates to the nucleus where it binds the T-cell factor (TCF) or lymphoid enhancer binding factor (LEF) transcription factors to initiate transcription of target genes. The canonical Wnt pathway is summarized in Figure 5. 1.4.1 Wnt in Normal Bone Development  Wnt pathway has been demonstrated to be involved in osteoblastogenesis and bone homeostasis. In bone differentiation the canonical Wnt pathway promotes bone formation by facilitating osteoblast differentiation from mesenchymal stem cells [58]. WNT10b has been shown to initiate mesenchymal stem cell differentiation along an osteoblast lineage instead of an adipocyte lineage in developing bone [59, 60]. In osteoblasts the canonical Wnt pathway also 13  promotes bone formation by increasing their proliferation and calcification while inhibiting osteoclastogenesis by increasing the expression of osteoprotegerin (OPG) (an inhibitory agent of osteoclastogenesis) [61]. Wnt/β-catenin pathway promotes osteoblast differentiation in bone development via suppressing the development of bone-matrix-degrading osteoclasts through the production of OPG [61, 62]. Further, the canonical Wnt pathway may promote mineralization by the expression of alkaline phosphatase [63]. The Wnt pathway is one of several regulatory pathways that control the expression of alkaline phosphatase, a specific marker of mineralization, and therefore initiate mineralization [64]. Moreover, the bone morphogenic proteins (BMP) pathway, the major bone formation mechanism, and the canonical Wnt pathway have common targets that can induce osteogenic differentiation from mesenchymal stem cells [65, 66]. It has been suggested that BMP2 stimulates LRP5, resulting in the canonical Wnt pathway activation and osteogenic differentiation [67]. The precise role of non-canonical Wnt pathway in regulating bone metabolism is still unknown [68].  1.4.2 Wnt in Vascular Calcification With aging, the arterial vasculature becomes the second most extensively calcified structure in the body after the skeleton [69]. Once thought to be a passive process, recent data indicate that the process of arterial calcification is an actively regulated process that involves pro-osteogenic signaling [69, 70]. The process of vessel calcification is similar to that occuring in bone mineralization [71]. Both in vivo and in vitro studies have revealed expression of proteins essential for bone formation such as bone morphogenic protein 2 (BMP2),  osteopontin, and osteocalcin also occur in arteries at sites of calcification [72-74].  14  Vascular calcification also may be a result of hypercholesterolemia [75] likely via Wnt/β-catenin signaling pathway co-receptor, LRP5, which is an important regulator of the differentiation of osteoblasts in bone formation  [76] and has also been implicated in cholesterol metabolism [77, 78]. Further, preliminary data indicate that the paracrine signals provided by BMP and Wnt signaling and matrix turnover promote osteogenic differentiation and calcification by vascular smooth muscle cells (vSMC) [79-81]. Thus, the osteogenic gene expression program pathways such as Wnt, Notch, transforming growth factor TGFβ signaling [82-86] are recognized as possible contributors to vascular calcification. For instance LDL-/- male mice on high fat diet develop atherosclerotic plaques with calcification [87].  As seen in bone growth, the expression of BMP2 and MSX2 is observed in the calcification of the plaques, as well as subsequent RUNX2, a bone specific transcription factor [70, 88].  MSX2 is a transcription factor first identified in human femoral osteoblasts that promotes osteoblastogenesis [89].Vascular smooth muscle cells (vSMC) express MSX2 resulting in up-regulation of WNT3A, WNT7A and WNT7B with concomitant increases in alkaline phosphatase activity, as an identified marker of calcification [90]. Another example of vascular calcification occurs in end-stage renal disease, where the excess of calcium and phosphate ions can promote osteogenic differentiation of vascular SMC and increase RUNX2 expression [91, 92]. The mechanisms suggested here by which vascular calcification occurs may also happen in valvular calcification. 15  1.4.3 Wnt in Valvular Calcification WNT is involved in embryogenesis and plays a role in the development of the endocardial cushions, from which valves arise in the developing heart [93]. In addition to its role in embryogenesis and development of heart valves, Wnt signaling plays an important role in osteogenic transformation of mesenchymal stem cells (MSC) in the process of forming normal bone tissue. Therefore, it is not surprising that Wnt signaling may participate in pathological calcification of aortic valves. The activation of the canonical Wnt pathway was first shown to be present in valve calcification when the expression of WNT3A and β-catenin was measured in calcified human aortic and mitral valves. The expression of both WNT3A and β-catenin was up-regulated in calcified valves as compared to normal ones [94, 95]. Further it has been shown that the expression of LRP5, the WNT3A co-receptor, was increased in both calcified aortic and mitral valves along with the over-expression of RUNX2, a bone specific transcription factor. Further studies on LRP5-/- mice revealed that the absence of this WNT receptor in animal models can reduce aortic valve calcification [96]. In addition, the expression of β-catenin, the transcriptional regulator of the canonical Wnt pathway, is up-regulated in calcified aortic valves in both rabbit and mouse models [77, 97]. It is worth mentioning that recent evidence suggests there is a direct regulation of RUNX2 by the canonical Wnt signaling pathway, in particular RUNX2 acts as a target gene of the canonical Wnt pathway in a mouse model [95, 98].  The mechanism by which Wnt pathway is activated in the process of calcification is linked to the bone morphogenic protein 2 (BMP2). 16  BMP2 reflects the alternative signaling cascade that is implicated in both normal and pathological calcification [99]. BMP2 can stimulate the expression of MSX2, which in turn mediates the release of WNT proteins and consequently plays a role in vascular and valvular calcification [90, 100].            17  1.4.4 Dickkopf1 (DKK1), an WNT3A Antagonist Dickkopf (DKK) is a secreted protein, rich in cysteine that is involved in embryonic development. DKK1 was originally identified as an embryonic head development inducer and was shown to act as the WNT antagonist that blocks the activation of the canonical Wnt pathway in many cell types and vertebrate species [101]. DKK1 is known as a negative regulator of WNT [102, 103] that does not bind or interact with WNT, but rather binds to the WNT receptor LRP [104, 105]. In the developmental process of bone formation it has been demonstrated that inhibition of DKK1 allows the Wnt pathway to inhibit osteoclast differentiation. Osteoclasts are the cells that resorb bone matrix, and thus if DKK1 inhibits mesenchymal cell differentiation into osteoclasts, it will indirectly promote bone mass production via osteoblast differentiation [106].  DKK1 blocks the Wnt pathway via binding and modulating the WNT co-receptor, LRP5/6. By binding LRP5/6 and its co-receptor Kremen, DKK1 inhibits formation of the WNT-LRP5/6-Frizzled complex and thereby hinders activation of the Wnt pathway (Figure 5) [105].    18    Figure 5. Overview of the canonical Wnt signaling and a model of DKK1 interaction with the canonical Wnt pathway.  In the absence of WNT ligand (A), cytosolic β-catenin is phosphorylated by GSK-3β and targeted for degradation by the ubiquitin pathway (proteosomal destruction). Upon WNT ligand binding (B), a series of phosphorylation events is initiated which culminates in the phosphorylation and inactivation of GSK-3β (deactivation of the destruction complex). This in turn leads to the accumulation of cytosolic β-catenin, which translocates to the nucleus where it binds the TCF family of transcription factors to initiate transcription of target genes. WNT3A is negatively regulated by DKK1. DKK1 binds to LRP6 and acts as a specific WNT antagonist. DKK1 and LRP6 can form a complex with Kremen, a single transmembrane proteins that is a high-affinity receptor for DKK1 (C). The LRP6/DKK1/Kremen complex rapidly endocytosis leading to the inhibition of Wnt/β-catenin signaling [107]. (Modified from [108]) LRP, lipoprotein receptor-related protein; APC, adenomatous polyposis coli; GSK, glycogen synthase kinase; TCF, T-cell factor; DKK1, Dickkopf1.     19  1.5 TGFβ Signaling in Calcification of Valves Transforming growth factor β (TGFβ) signaling is involved in regulating many cell processes including cell development, fate, growth, differentiation and apoptosis [109]. When TGFβ binds the type II receptor, it activates the TGFβ type I receptor and forms a complex. This complex phosphorylates the receptor-specific SMADs, SMAD2/3 which then with SMAD4 (co-SMAD) make a complex that moves to the nucleus and facilitates the transcription of target genes (Figure 6) [109, 110]. There are numerous reports demonstrating the direct and indirect involvement of TGFβ in various biological processes including wound healing, angiogenesis, bone formation, and skin formation as well as in multiple pathologies such as inflammatory and fibrotic diseases and tumor development [111]. The study of transgenic mouse models lacking TGFβ1 has revealed the crucial role of this molecule in immune system development [112]. In lung fibrosis TGFβ1 increases the ECM synthesis. TGFβ1also induces the expression of α-smooth muscle actin (α-SMA) and collagen type I synthesis and deposition in myofibroblasts [113].  TGFβ1 participates in the pathogenesis of many cardiovascular diseases such hypertension, atherosclerosis, restenosis, hypertrophy, and heart failure [114-116]. There is also evidence that TGFβ plays a pivotal role in valvular diseases, including aortic valve fibrosis and calcification [117, 118]. More precisely in the porcine VIC cell model TGFβ1 induces the transformation of VIC into an osteoblast phenotype and is therefore implicated in valvular calcification [119]. In vitro studies of porcine VIC have demonstrated that the addition of TGFβ1 20  to cultured porcine VIC induces the formation of calcium nodules and expression of osteoblast markers [120]. Thus TGFβ is of paramount interest in this study.  Morphogenic signals provided by both TGFβ and canonical Wnt pathways are necessary for proper tissue and organ development [121]. In addition to morphogenic signals the activation of control pathways that regulate overall cell and tissue growth are required. The Hippo pathway, a well conserved pathway in mammals, is one such pathway that regulates tissue growth [122, 123] and acts as a link between TGFβ and canonical Wnt pathways. In the presence of Hippo pathway activity, the nuclear components, TAZ/YAP are sequestered in the cytosol, where they suppress the activity of both TGFβ and canonical Wnt pathways [121].  Further, DKK1 is known to play a role as a link between the canonical Wnt and TGFβ pathways. In pathological context, activation of the canonical Wnt pathway stimulates fibroblasts in culture and induces fibrosis in vivo [124-126]. Over-expression of DKK1 in animal models of skin fibrosis induced by TGFβ, prevents the fibrosis and highlights a key role for the interaction of both canonical Wnt and TGFβ pathways through DKK1 in the pathogenesis of fibrotic diseases [127].         21    Figure 6. Overview of the TGFβ pathway. After TGFβ undergoes activation, it binds to the type II receptor (II). Binding of TGFβ results in the formation or stabilization of a complex of the type I and II receptors (I/II), and the type II kinase phosphorylates (P) and activates the type I receptor. The activated type I receptor kinase phosphorylates receptor-specific SMADs, which, for the TGF-β pathway, include SMAD2 and SMAD3. This step can be inhibited by Smad7. Phospho-SMAD2 and 3 form complexes with the co-SMAD (SMAD4) and move into the nucleus, where they may interact with other transcription factors or co-activators and co-repressors to regulate transcription.  22  1.6 1, 25 (OH)2 D3 (Vitamin D3) Since its discovery in the 1920s, vitamin D was believed to be exclusively involved in bone formation and maintenance of skeletal integrity [128].  Vitamin D is not a true vitamin, but rather a secosteroid hormone, which can be produced endogenously or acquired by dietary consumption. Vitamin D has an endocrine mechanism of action and is sequentially synthesized in the skin, liver and kidney (Figure 7). However, the major source of vitamin D in the body is synthesis in the skin in the form of cholecalciferol (Vitamin D3) [129, 130]. This precursor of vitamin D undergoes two sequential hydroxylation processes in liver and kidney respectively to produce the active form of vitamin D, 1,25-dihydroxycholecalciferol (calcitriol). Calcitriol is transported in the circulation to the target cells where it has both genomic and non-genomic effects [131]. Genomic effects of vitamin D are mediated via binding of the nuclear vitamin D receptor (VDR) [132]. Vitamin D is thought to regulate 3% of the genome, and has numerous effects, including enhancement of cellular differentiation, proliferation and immune responses [132, 133], in addition to its classical role in mineral homeostasis. Non-genomic effects of active vitamin D are poorly understood [134]. What is known of the vitamin D pathway is summarized in Figure 7. 1.6.1 1, 25 (OH)2 D3 in Valvular Calcification In addition to the role of vitamin D and calcium in the maintenance of skeletal health, there is also evidence that vitamin D plays a critical role in pathological calcification [135, 136]. Evidence supports that vitamin D might be involved in the process of vascular calcification. The 23  effects of hypervitaminosis D have been well characterized in animal models. Excess amounts of vitamin D administrated to rats can rapidly produce calcium overload in the circulation and in the major sites of calcium deposit were on the medial elastic fibers near the lumen of the arteries. Toxic doses of vitamin D resulted in calcification leading to destruction of elastic fibers and arterial stiffness [137]. In addition, widespread soft tissue calcification is also detected after calcitriol treatment [138].  Of relevance here in these observations, there is a significant association of vitamin D receptor polymorphism with aortic valve stenosis. The B allele of the vitamin D receptor is more common in patients with calcified aortic valves [139].  The BsmI polymorphism of the vitamin D receptor gene is a marker of bone mineral mass [140, 141]. The individuals with the B allele have less bone mineral mass and more rapid bone mineral loss with advancing age, than those with the b allele [141, 142]. Whether other genes regulating calcium homeostasis are involved in the pathogenesis of calcific aortic valve is a question to be answered.          24       Figure 7. Overview of the normal vitamin D3 pathway. Vitamin D may be taken up from the diet but is mostly produced in the skin from sun exposure in form of cholecalciferol. From vitamin D3, two enzymatic activation steps are required to produce the biologically active form of vitamin D [1,25(OH)2 D3] first in liver and then in kidney. In vitamin D target cells, 1,25(OH)2 D3 translocates to the nucleus and binds to the vitamin D receptor (VDR) [143].                  25  1.7 Thesis Rationale, Hypothesis and Specific Aims Once thought to be caused by a passive process involving calcium attaching to the valve leaflet, there is growing evidence to support that atherosclerotic risk factors and other factors can induce valve myofibroblast cells to transform to an osteoblast-like phenotype. Examination of human calcified aortic valve tissue has shown that several pathologic pathways are involved. A growing number of laboratories are demonstrating that osteogenic gene programs in calcifying cardiovascular tissue involve complex signaling pathways similar to bone biology. However, the exact molecular mechanisms involved in valvular calcification have not been elucidated.  For this dissertation, I have examined the effects of a canonical WNT ligand, WNT3A, TGFβ1, and vitamin D3 on calcification of primary human valvular interstitial cells, in vitro and in vivo. More precisely, I studied the role of WNT3A, TGFβ1 and vitamin D3 in the process of a phenotype shift of VIC from myofibroblast to that of an osteoblast-like cell in vitro.  In Chapter 2, in the process of establishing a better model for calcification, I have tested different human cell types in an osteogenic environment and only vascular SMC and aortic VIC calcified to formed calcium nodules. However, VIC calcification was significantly more extensive than what I observed in vascular SMC. Since the main goal of this study is to improve understanding of valvular calcification and since VIC are the most abundant cells in aortic valves, I pursued the rest of the study using the in vitro VIC model. Moreover, calcified VIC in vitro (Chapter 2) expressed WNT3A, TGFβ1 and VDR, which further supported the choice of this model. The fact that all these agents mentioned above are pro-osteogenic and have been shown to be involved in calcification, encouraged me to further investigate the role of these three 26  pro-osteogenic factors in the process of VIC calcification individually and in combination with each other. Through the investigation discussed in Chapter 3, I determined the role of the canonical Wnt and TGFβ pathways, in valvular interstitial cells during the process of calcification, characteristic of aortic valve stenosis. Wnt and TGFβ pathways are involved in the calcification of bone and also the calcification occurring in atherosclerotic plaques. In fact both Wnt and TGFβ pathways have been implicated in the calcification of AVS. In this chapter I examined the role of DKK1 as the potential link between the Wnt and TGFβ pathways. As discussed above vitamin D overdoses have been shown to play a role in calcification of soft tissue. It is also evident that vitamin D is involved in normal calcification. In Chapter 4 I examined whether vitamin D plays a role in calcification of valvular interstitial cells. Additionally I tested the potential interaction between vitamin D and the canonical Wnt and TGFβ pathways.  The overarching hypothesis of this thesis is that Wnt plays an important role in cardiovascular disease pathogenesis. For this thesis, I hypothesized that the canonical Wnt pathway plays a pivotal role in transformation of valvular interstitial cells to an osteoblast-like phenotype and that TGFβ and vitamin D contribute to this process interactively and additively, respectively.  The specific aims for the present thesis study were:   27  1) To characterize VIC in an osteogenic environment.  2) To determine the role of Wnt signaling pathway in calcification of VIC and contribution of TGFβ in the activation of the canonical Wnt pathway in VIC calcification.    3) To determine the effect of vitamin D in the process of VIC calcification and the potential of a link between vitamin D and Wnt and/or TGFβ pathways. 28  Chapter 2: HUMAN VALVULAR INTERSTITIAL CELLS: A MODEL TO STUDY AORTIC VALVE CALCIFICATION  2.1  Introduction  Calcification is a common process in different disease states, which implies that such deposition of calcium is abnormal or at least adverse. The deposition of calcium in normal tissues due to the disturbance in calcium phosphorus metabolism is a form of abnormal calcification known as metastatic calcification such as calcification  seen in kidney or lungs [14]. Another form of calcification, known as dystrophic calcification, occurs in damaged tissue such as calcification seen in arteries and atherosclerotic plaques [14]. Of note, while hypercalcemia does not initiate pathological or dystrophic calcification, it can contribute to it [144]. Valvular calcification is a form of pathological dystrophic calcification. Valvular calcification is the main cause of heart valve failure that leads to valve replacement and is the most common valve disease in developed countries, yet the mechanism of initiation and progression of the disease remains to be elucidated. While the simplest circumstance in which one might study calcification is the normal calcification of bone, little information is available from this process. There are few animal or cell culture models available for the study of aortic valve calcification, however they all have significant limitations. While some species such as swine can naturally develop calcified aortic valve lesions others such as rabbits and mice need external stimulation such as genetic mutations and hypercholesterolemic diet to develop the disease [145-147]. Conducting experiments on these in vivo models may be costly, but especially represent a good step after the work we have undertaken. All in vivo models have their own place in investigations, however for our purposes, the in vitro model of VIC to be described was deemed very useful and informative. 29  There are few cell culture models available to study valvular calcification [148, 149]. Even though these cell culture models do not exactly mimic the tissue structure and lack blood flow and hemodynamic effects existing in tissue, still they are the most time and cost effective models to study aortic valve calcification. Among those, porcine valvular interstitial cells are the most commonly used cells in the scientific literature to study aortic valve calcification [147]. Porcine VIC grow fast and easily form calcium nodules in culture in about 2-3 weeks. However since the ultimate goal is to understand the etiology of the disease in human tissues, it seems more relevant to use human cell models if possible. Currently, cultured human vascular SMC are the most commonly used in vitro model to study vascular calcification [150-152]. Human vascular SMC are an in vitro model system that has been shown to mimic some of the features seen in human vascular calcification in vivo [153, 154]. Although human valvular interstitial cells might be a more appropriate model system, an immortal cell line is not commercially available. Further, VIC are the most abundant cell type in valve tissue and are known to be actively involved in the normal function of the valve as well as valvular diseases including valve calcification [34, 155, 156]. VIC exhibit myofibroblast characteristics and are known to undergo a phenotype shift to osteoblast-like cells in the process of valvular calcification [35, 36]. Therefore human VIC might be justifiably utilized as an in vitro model for studying valvular calcification. Indeed it has been shown that on a certain substrate (glycerol) and in a certain environment, human VIC form calcium nodules. Identifying how human VIC behave in an osteogenic environment in vitro is crucial to a better understanding of valvular calcification in vivo.  30  In the current study we aimed to compare primary human vascular SMC with VIC in the process of calcification in vitro. We demonstrate that human VIC likely make a better in vitro model for studying and understanding valvular calcification.                   31  2.2 Materials and Methods 2.2.1 Cell and Tissue Procurement Human aortic valve leaflets from explanted hearts taken at the time of transplantation were used from the Cardiovascular Pathology Registry at St. Paul's Hospital, Vancouver, British Columbia, Canada. Normal human aortic valve leaflets were obtained from the International Institute for the Advancement of Medicine (IIAM) program. For this study, only valves from male patients aged 47-52 years (n=3) with no history of heart valve diseases were acquired for in vitro experiments. Since male gender is one of the risk factors for aortic valve stenosis [21] all the subject materials were taken from middle aged males. Human medial vascular smooth muscle cells (vSMC), a generous gift from Dr. David Granville (HLI, UBC, Vancouver), were maintained in Smooth Muscle Growth Medium-2 (SmGM-2) and supplemented with FBS and growth factors (Lonza; product #CC-3181 and CC-4149) and used for experiments between passages 3-8. Human prostate stromal fibroblast cell line WPMY-1 and HEK-293 cells were obtained from the American Type Culture Collection (ATCC, Rockville, MD). All cell types mentioned above were cultured and maintained in Dulbecco's Modified Eagle Medium (DMEM). DMEM containing 10% fetal bovine serum (FBS) and 100 U/mL penicillin/streptomycin and used for experiments between passages 6-12.  Isolated VIC from human aortic valve tissue were cultured in Molecular and Cellular Developmental Biology 131 medium (MCDB-131), (Life Technologies; product #10372019) containing 10% FBS and 100U/ml penicillin/streptomycin. All cell types were maintained in a humidified incubator at 37°C with 5% CO2 to be used for experiments [157]. Seventy percent 32  confluent cultures were treated with osteogenic condition medium for 6, 24, 48, or 96 h and for 1, 2, 3 and 4 weeks if they maintained viability. Morphological changes were documented photographically with a Nikon 50i series upright microscope equipped with a digital camera prior to performing other functional assays, histochemical staining or collecting cells for molecular analysis. Alkaline phosphatase activity assays were also performed on these cultured cells as described below as a marker of calcification. Von Kossa staining for calcium was performed to confirm calcification. Protein studies were carried out to identify certain cellular markers. All experiments were performed in triplicate and repeated a minimum of 3 independent times. This study was approved by the University of British Columbia/Providence Health Care Research Ethics Board and conforms to the principles outlined in the Declaration of Helsinki for use of human tissues or subjects. 2.2.2 Antibodies Antibodies used in this study include anti-WNT3A (Abcam, Inc., Toronto, ON; product #ab28472), anti-TGFβ1 (Abcam; product #ab64715) and anti-alkaline phosphatase (abcam; product #75699, USA). anti-osteopontin (Sigma-Aldrich, Canada, Oakville, ON; product #07264), anti-RUNX2 (Abcam; product #ab76956), horseradish peroxidase (HRP) conjugated goat anti-mouse (Santa Cruz Biotechnology, Inc., Dallas, TX, USA) and goat anti-rabbit (Santa Cruz; product #sc-2030) and anti-rabbit Alexa-fluor488 conjugated secondary antibody (Invitrogen, product #S-11223, USA) 33  2.2.3 Isolation of Human Valvular Interstitial Cells Human valvular interstitial cell isolation and culture was established by a modification of the enzymatic dispersion technique [158, 159]. Briefly, valve tissues were freshly obtained as described above and immediately washed in MCDB-131 medium. Tissue was minced with a sterilized blade. Enzyme I (2.5 mg/mL collagenase II, Roche; product #11088882001) was applied to the exposed media (MCDB-131 + 10% FBS) and tissue followed by 1 h incubation at 37°C to separate the cells from extracellular matrix (ECM). The tissue was broken down by pipetting a few times and then transferred to a 15 ml tube. Cells were spun down and 3 ml of fresh collagenase was added to the cells and incubated for 24 hours with pipetting at regular intervals to further disperse cells. A panel of antibodies against cytoskeletal elements was used to phenotype the isolated VIC by immunohistochemistry. The valve cells demonstrated positive staining for smooth muscle α-actin, vimentin and desmin, but were negative for smooth muscle myosin heavy chain, suggesting these cells display a myofibroblast-like phenotype in culture [157]. 2.2.4 Cell Culture Treatment Cultured VIC were maintained in MCDB-131 growth medium supplemented with 10% FBS + 200 mg/ml L-glutamine. Osteogenic condition medium was added to half of the cells. The osteogenic medium consists of 50 g/ml ascorbic acid (Sigma-Aldrich, product # A4403-100G, USA) 10 mM glycerol-2 phosphate (Sigma-Aldrich, product # G9891, USA) and 10 mM dexamethasone (Sigma-Aldrich, product # D8893, USA)  [160]. Fresh medium plus osteogenic 34  treatments were added to the previously treated cells every other day for 1, 2, 3, or 4 weeks. Cell morphology and calcium nodule formation were monitored and recorded at each time point. 2.2.5 Western Blots Both untreated cells and cells treated with the reagents from above were washed twice with ice-cold PBS, and then cell lysates were prepared by adding 80 µl modified oncogene science lysis buffer [(MOSLB): (50 mM sodium pyrophosphate, 50 mM NaF, 50 mM NaCl, 5 mM EDTA, 5 mM EGTA, 100 µM Na3VO4, 10 mM)] to the cells and incubating them on ice for another 5 minutes. Cell lysates were then scraped into Eppendorf tubes and spun at 4°C for 10 minutes at 3500rpm. Supernatants containing cell lysate were collected in another tube and stored at -80°C for subsequent experiments. Protein concentration was determined (Bio-Rad Protein Assay, USA) and the extracted protein (25-50 mg) was fractionated by electrophoresis on 12% SDS-polyacrylamide gels, transferred to nitrocellulose membranes (GE Biosciences, Quebec, CA; product #CA27376-991), and blocked with PBS containing 0.1% Tween-20 and 5% non-fat dry milk for 1 hour. The membrane was then incubated with specific primary antibodies; anti-osteopontin (Sigma, product #07264, USA), anti-RUNX2 (abcam; product #ab76956, USA), and Anti-WNT3a (abcam; product #ab-28472, USA), TGFβ1 (abcam; product #ab64715, USA)  overnight at 4°C, followed by secondary antibody binding for 1 hour at room temperature according to manufacturer’s protocol (Bio-Rad, USA). The immunoblots were visualized with an enhanced chemiluminescence (SYNGENE, CHEMI GENIUS2 Bio-imaging system, USA) detection system according to the manufacturer’s protocol (Amersham Pharmacia Biotech, USA). 35  2.2.6 Histochemical Staining for Alkaline Phosphatase Alkaline phosphatase localization was performed on cells as a marker of calcification [161]. Briefly, cells were fixed in paraformaldehyde 4% solution for 15 minutes and then incubated in Naphthol-Tris working solution. Dissolved 5mg naphthol AS-MX (Sigma-Aldrich; product #855, USA) in 0.25 ml N,N dimethylformamide (Sigma-Aldrich; product #D4551) was added to 0.25 ml Tris-hydrochloric acid buffer. The mixture was diluted in 25 ml distilled water and 25 ml Tris buffer. Then 30 mg Fast Red (Sigma-Aldrich; product #201286) was added to the solution and filtered with 2 mm filter paper). After filtering the buffer in 2mm filter paper, the working buffer was added to the cells for an hour. The pink to red coloration was indicative of the alkaline phosphatase enzyme activity in the cells. Cells were washed in distilled water. 2.2.7 Von Kossa Staining for Calcium Von Kossa staining was carried out on paraformaldehyde (Fisher; product #F79-500) fixed cells according to the procedure described by Sheehan DC and Hrapchak BB, and as referenced by the American Master Tech von Kossa stain procedure [162]. Briefly cultured cells were fixed in 4% paraformaldehyde solution for 15 minutes. The cells placed in silver nitrate (Sigma-Aldrich; product #209139, USA) 2% solution for 60 minutes, under a 100-watt incandescent lamp. These slides were then rinsed in distilled water, and placed in 5% sodium thiosulfate solution for 2 to 3 minutes. Then they were rinsed in tap water, and placed in nuclear fast red for 5 minutes. Calcium dissolves in acid, whereas urates or phosphates salts do not [163]. The equipment used for microphotography was a Nikon 50i series upright microscope equipped 36  with a digital camera. Images of the cells presented in this manuscript are originals, without any physical or digital alterations. 2.2.8 Immunocytochemistry Some of the cultured cells were fixed in vitro in 4% formaldehyde for 20 minutes. These cells were then permeabilized with 0.1% Triton X-100 for 20 min, blocked for 30 min with 1% serum free blocking reagent (DAKO; product #X0909, Denmark), and incubated overnight at 4°C with primary anti-alkaline phosphatase (abcam; product #75699, USA) at a concentration of 1:100 in 1% BSA. Following primary antibody binding, cells were incubated with anti-rabbit Alexa-fluor488 conjugated secondary antibody (Invitrogen; product #S-11223, USA) at a concentration of 1:250 in 1% BSA for 1 hour at room temperature in the dark. Cells were then counterstained for 5 min with DAPI Nucleic Acid Stain (Invitrogen; product #R37606, USA) to visualize nuclei and then cover-slipped with Slow Fade Gold antifade reagent (Invitrogen; product #S-36936 USA). Images were captured using a Leica AOBS SP2 confocal microscope as we have previously described [164, 165]. 2.2.9 Alkaline Phosphatase Activity Assay Alkaline phosphatase activity in the cultured cells was measured by SensoLyte® pNPP Alkaline Phosphatase Assay Kit Colorimetric (AnaSpec, product #72146, USA). This assay colorimetrically detects alkaline phosphatase activity using pNPP (p-Nitrophenyl phosphate) phosphatase substrate [166, 167]. Cells were grown and treated as described above. The assay was performed according to the manufacturer’s protocol for biological samples. The intensity of 37  color corresponding to the alkaline phosphatase activity was read using VersaMax ELISA Microplate Reader (Molecular Devices, Sunnyvale, CA, USA). 2.2.10 Statistical Analysis Data are expressed as the mean ± standard error (SE) of at least three independent determinations. Differences between means were determined using a one-way analysis of variance (ANOVA) with pair-wise comparison by the Tukey-Kramer method. Differences were considered to be significant when the p-value was less than 0.05. Analyses were performed using the Graphpad Prism 6 software package (www.graphpad.com).           38  2.3 Results 2.3.1 Calcium Nodules Are Formed in Vascular SMC and VIC To study cell behavior in this osteogenic environment a variety of cell types including VIC, vSMC, HEK-293, and WPMY-1 were cultured in their appropriate growth media. At 70% confluence half of the cultures from each cell type were treated with osteogenic medium for 1, 2, 3 or 4 week duration. The osteogenic medium was refreshed every 48 hours. Morphological changes and calcium nodule formation were monitored and cultures photographed at each time point. The HEK-293 cell growth rate was higher than other cell types and they grew to become multi-cell layered. Before the first time point (week 1) at 90-100% confluent cells started detaching from the culture dish surface and therefore did not persist for the next feeding time. The human prostate stromal fibroblast (WPMY-1) proliferation rate was lower than HEK293 cells and higher than vSMC and VIC. WPMY-1 only grew as single layers of cells yet by week 2 cells became thinner and underwent cell death in both control and treated groups. Calcium nodule formation was not observed in either the WPMY-1 or HEK293 groups, thus they were not deemed proper culture models for further experiments. Human vSMC had a longer lifespan and only grew to form a single cell layer [168]. Therefore they survived through the third week. Pre-nodules of calcium were formed by week 3 of the experiment in the treated cell group, confirming that vSMC can calcify in the osteogenic environment. 39  Human primary VIC that exhibit myofibroblast characteristics [157] also only grew into a single layer with a low growth rate especially in the osteogenic medium [36, 169]. In this study primary VIC survived longer than all of the other cell types. In addition mature calcium nodules are formed in the VIC cultures as early as the week 2 time point. VIC calcified earlier and more extensively than any of the other types of cells used (Figure 8 A, B and C).             40    Figure 8. Calcium nodule formation in different cell type at various timepoints. Primary human VIC were cultured in normal and osteogenic growth media and maintained for 4 weeks (A). Mature calcium nodules were formed at week 3 in treated cells (arrows). Primary human vSMC were cultured and maintained in normal and osteogenic growth media for 3 weeks (B). Calcium pre-nodules started to form in treated cells at week 3 (arrows). Stromal prostate fibroblasts were cultured and maintained in normal and osteogenic media. Cells started to die by end of the week 2. Calcium nodules did not form in any of the cultures (C).    41  2.3.2 Alkaline Phosphatase and Von Kossa Staining Are Strongly Positive in Calcified VIC To demonstrate the presence of cell calcification VIC and vSMC were stained for alkaline phosphatase and with von Kossa for calcium. There was alkaline phosphatase localization in vSMC at week 3 in the treated group. However von Kossa staining for calcium was negative in untreated vSMC and less intensive in the treated vSMC than VIC at week 3 time point.   On the other hand VIC demonstrated positive localization of alkaline phosphatase in treated cells at week 2 which increased in intensity at week 3 and week 4 respectively. Similar to alkaline phosphatase staining, von Kossa staining was positive at week 3 and increased at week 4 in treated VIC (Figure 9 A and B).  We further confirmed the expression of alkaline phosphatase with immunohistochemistry staining for alkaline phosphatase in VIC (Figure 10).       42                  Figure 9. Alkaline phosphatase and von Kossa staining in VIC and vSMC. Primary human VIC were grown in normal and osteogenic growth media and maintained for 4 weeks (A). Alkaline phosphatase staining (red) and von Kossa for calcium (brown) were positive at week 3 and week 4 in treated cells. Primary vSMC were cultured and maintained in normal and osteogenic growth media for 3 weeks (B). Alkaline phosphatase staining is slightly positive at week 3 in treated cells.    43                   Figure 10. Alkaline phosphatase localization in VIC. Primary human VIC were cultured in normal and osteogenic growth media and maintained for 4 weeks. Immunohistochemistry staining was performed for alkaline phosphatase (red) and nuclei (blue). Alkaline phosphatase staining is positive and increases over time, consistent with histochemical staining in Figure 9. (Scale bar= 70μm)  44   2.3.3 Alkaline Phosphatase Activity Is Amplified in Calcified VIC As a functional phenotypic marker of osteoblasts we measured the alkaline phosphatase activity in WPMY-1, vSMC and VIC at weekly time points. Alkaline phosphatase activity in WPMY-1 did not increase for any existing timepoints of treated cells compared to their control group (Figure 11C) In the second and third weeks of growing vSMC alkaline phosphatase activity was slightly increased in treated cells compared to untreated (Figure 11 B). In VIC alkaline phosphatase activity started to increase in the treated group as early as week 1 and continued to increase at subsequent timepoints. In week 4 there was a dramatic increase in alkaline phosphatase activity in treated VIC, verifying cell calcification that matched the mature calcium nodule formation in cultures observed in the previous experiment (Figure 11 A). 45   Figure 11. Alkaline phosphatase activity in VIC, vSMC and WPMY-1 cells at different timepoints.  Human VIC, vSMC and WPMY-1 were cultured in regular and osteogenic media for different timepoints. (VIC for 1, 2, 3 or 4 weeks, vSMC for 1, 2 or 3 weeks and WPMY-1 for 1 or 2 weeks). Alkaline phosphatase activity in VIC was increased significantly at week 2, week 3 and week 4 in treated VIC compared to untreated group and earlier timepoints. (* indicates p-Value < 0.001, significant difference in time and  indicates p-value < 0.01, significant difference in treatments) (A). Alkaline phosphatase activity in vSMC was increased significantly at week 2 and week 3 in treated cells compared to untreated and week 1. (* indicates p-value < 0.001, significant difference in time and  indicates p-value < 0.01, significant difference in treatments) (B). Alkaline phosphatase activity was slightly increased at week 2 comparing to week 1(C). (n=3 for each experimental condition). Error bars represent mean ± SEM. W e e k  1 W e e k  2 W e e k  3 W e e k  405 01 0 01 5 0V ICU/LU n tre a te dO s te o g e n ic  M e d ia**A . A lk a lin e  P h o sp h a ta se  A c t iv ityW e e k  1 W e e k  2 W e e k  3051 01 52 02 5a S M CU/LU n tre a te dO s te o g e n ic  M e d iaB . A lk a lin e  P h o sp h a ta s e  A c tiv ity**W e e k  1 W e e k  202468W P M Y -1U/LU n tre a te dO s te o g e n ic  M e d iaC . A lk a lin e  P h o sp h ta s e  A c t iv ity *    * vSMC 46  2.3.4 Osteogenic Treatment Induces Osteoblast Transformation in VIC To determine whether valvular myofibroblasts were shifting phenotype to that of osteoblasts, we also assessed the expression of the bone specific transcription factor RUNX2 and osteonectin and osteopontin, as osteoblast markers [170-174]. vSMC and VIC were incubated in osteogenic media for up to 3 and 4 weeks respectively. The expression of osteoblast markers were tested in untreated and treated groups of both cell types. vSMC incubated in osteogenic treatment showed less increase in expression of osteopontin at week 3 than VIC (Figure 12 A, B and C). However, RUNX2 and osteonectin were not detected in any groups of vSMC at any time point. Conversely, VIC incubated in osteogenic medium exhibited significantly increased amounts of RUNX2, and osteopontin at week 4. Osteonectin was also expressed significantly in treated VICs (Figure 13 A, B, C and D).      * 47        Figure 12. Expression of osteoblast markers in vSMC. Human vSMC were grown and incubated in regular and osteogenic media. The expression of RUNX2 and osteopontin increased at week 3 compare to week 1 and 2; however, it was not significant increase between the untreated and treated groups at week 3 for both RUNX2 and osteopontin. Osteonectin was not detectable in vSMC (n=3 for each experimental condition). (* indicates p-value < 0.01, significant difference in time). Error bars represent mean ±SEM. (The representative blots are from different wells of the same gel). W e e k  1 W e e k  2 W e e k  301234Densitometry Unit(Normalized to GAPDH)U n tre a te dO s te o g e n ic  M e d iaA . R U N X 2  P r o te in  E x p r e ss io nW e e k  1 W e e k  2 W e e k  30 .00 .51 .01 .52 .02 .5Densitometry Unit(Normalized to GAPDH)U n tre a te dO s t o g e n ic  M e d iaB . O s te o p o n tin  P r o te in  E x p r e s s io nC. * 48        Figure 13. Expression of osteoblast markers in VIC.  Human VIC were incubated in regular and osteogenic media for 1, 2, 3, or 4 weeks. Expression of RUNX2, osteopontin and osteonectin significantly increased at week 4 compared to untreated and earlier timepoints (n=3 for each experimental condition). (* indicates p-value < 0.001, significant difference in time;  indicates p-value < 0.01, significant difference in treatments). Error bars represent mean ± SEM. (The representative blots are from two gels run side by side). W e e k  1 W e e k  2 W e e k  3 W e e k  401234Densitometry Unit(Normalized to GAPDH)U n tre a te dO s te o g e n ic  M e d iaA . R U N X 2  P r o te in  E x p r e ss io n*W e e k  1 W e e k  2 W e e k  3 W e e k  401234Densitometry Unit(Normalized to GAPDH)U n tr a te dO s te o g e n ic  M e d iaB . O s te o p o n tin  P r o te in  E x p r e s s io n*W e e k  1 W e e k  2 W e e k  3 W e e k  401234Densitometry Unit(Normalized to GAPDH)U n tre a te dO s te o g e n ic  M e d iaC . O s te o n e c t in  P r o te in  E x p r e s s io n*D. 49  2.3.5 Bone Regulators Are Overexpressed in Calcified VIC To examine whether ossification and bone growth regulators are involved in VIC and vSMC calcification we assessed the expression of pro-calcification mediators in the process of VIC and vSMC calcification. We examined the expression of TGFβ1, BMP2, WNT3A, and TNFα in both cell types. The expression of WNT3A and TGFβ1 was significantly increased in osteogenic-treated VIC at week 4 as compared to untreated VIC at the same time point. VIC in other time points (week 1, 2, and 3) did not express WNT3A or TGFβ1. Further, in vSMC no increased expression of the osteogenic agents mentioned above was observed in either treated or untreated groups (Figure 14 A, B and C).          50  Week 1Week 2Week 3Week 40 .00 .51 .01 .52 .02 .5V ICDensitometry Unit(Normalized to GAPDH)U n tre a te dO s te o g e n ic  M e d iaA . W N T 3 A  P r o te in  E x p r e s s io n*Week 1Week 2Week 3Week 4.00 .51 .01 .5V ICDensitometry Unit(Normalized to GAPDH)U n tre a te dO s te o g e n ic  M e d iaB . T G F  1  P r o te in  E x p r e s s io n*         Figure 14. Expression of pro-osteogenic mediators (WNT3A and TGFβ1) in calcified VIC. Human VIC were incubated in regular and osteogenic media for 1, 2, 3, or 4 weeks. The expression of WNT3A and TGFβ1 increased significantly in treated cells at week 4 timepoint (n=3 for each experimental condition). (* indicates p-value < 0.001, significant difference in time;  indicates p-value < 0.01, significant difference in treatments). Error bars represent mean ± SEM.  C. 51  2.4 Discussion In this study, the valvular interstitial cell model was found to be superior to other cell types for the study of valvular calcification.  We compared human VIC to vSMC in the process of cell calcification in vitro for the first time. vSMC have previously been shown to be prone to calcification and were therefore used by others as an in vitro model to study the vascular calcification process [175]. In order to study valvular calcification, pigs, rabbits and mice on a Western hypercholesterolemic diet program have been used[118]. To study valvular calcification in vitro however, porcine VIC have been commonly utilized in previous studies [149, 176-178]. Even though animal cell models are commonly used to study many human diseases, they carry limitations. Our study is the first on primary human VIC that has characterized primary human valvular cells in normal culture and in an osteogenic environment both phenotypically and functionally and therefore offers valuable information for the study of valvular calcification. In this work we first attempted to grow and maintain a variety of human cell types in osteogenic media to monitor and compare the cellular responses. Each cell type behaved differently in the osteogenic environment. Only vSMC and VIC produced calcium nodules yet the degree of calcification differed between them. Previous reports have identified that apatite is the predominant mineral deposits in the calcium nodules formed in the vSMC and VIC cultures [179, 180]. In addition, ultrastructural studies of these nodules demonstrated that the mineral deposits found in vSMC and VIC morphologically are similar to deposits found in bone, cartilage and teeth [181, 182]. Finally, it has been shown that calcification initially occurs predominantly extracellularly in association with collagen fibrils and not related to necrotic debris [149, 180]. 52  The components of the osteogenic media were previously shown to promote calcification separately and the combination of these component likely enable the optimal conditions for cells to calcify [147, 183, 184]. VIC phenotype was significantly altered in the osteogenic media and in addition VIC formed mature calcium nodules as early as week2. Alkaline phosphatase staining and von Kossa staining for calcium were positive by week3. While these responses were also seen in vSMC, they were minimal and were diminished. Further, alkaline phosphatase activity increased in treated VIC as early as week 2 and was several-fold higher than the level observed in vSMC. These observations are consistent with previous studies in an animal cell model of porcine VIC, where VIC isolated from pigs showed more susceptibility to calcification in vitro. However, this is the first time the calcification process of several cell types from human have been compared in an in vitro model. To confirm the phenotype shift toward osteoblast-like cells, the expression of osteoblast markers were examined in both calcified human vSMC and VIC. Even though treated vSMC expressed osteopontin and the bone specific transcription factor RUNX2, the osteoblast specific marker osteonectin was not observed. In VIC, however, both RUNX2 and osteonectin were expressed along with osteopontin in treated cells and increased significantly in the treated group compared to controls at week 4. Osteonectin is an early marker of pre-osteoblast cells [185] and our finding fact it is expressed earlier in VIC indicates that human VIC shift phenotype and calcify earlier than human vSMC. Our observations suggest that the VIC model may represent a significantly more appropriate model to investigate valvular calcification. There are several mediators involved in the process of calcification and osteoblast transformation. In light of the advantages of the VIC model, we decided to test for the expression 53  of some of the known mediators responsible for normal calcification and bone development in this model. We tested for the presence of WNT3A, TGFβ1, BMP2, and TNFα [97, 186-188]. WNT3A and TGFβ1expression were both detected in treated cells at week 4, whereas none of these mediators were observed in vSMC (negative data not shown).  The observations in this study led us to the conclusion that human VIC may provide a better model for the study of valvular calcification. Besides the fact that VIC are the most abundant cells in human aortic valves and have been shown to be involved in all the normal and pathological events in the valve tissue, we have shown that these cells can be induced to calcify in culture in a simple-constituted osteogenic medium. Therefore in spite of the labor involved in acquiring and culturing human VIC, they are the most relevant and valuable model to study valvular calcification in vitro especially in light of the absence of a transformed cell line from commercial sources. On the basis of these observations we have used this model to investigate the regulation of this transformation to osteoblast cells (Chapter 3) and the possible contribution of vitamin D3 to this process (Chapter 4).      54  Chapter 3: CANONICAL WNT ACTIVATION INCREASES VALVULAR INTERSTITIAL CELL ASSOCIATED CALCIFICATION VIA TGFΒ-MEDIATED DOWN-REGULATION OF DKK1  3.1 Introduction  Aortic valve stenosis (AVS) is the third leading cause of adult heart failure and the most common form of acquired valvular disease in developed countries. Its prevalence is rapidly increasing due to an aging population [3]. The most common and effective treatment for AVS is surgical replacement of the stenotic valve, a procedure associated with the inherent risks of open heart surgery [4]. Further while novel percutaneous valve replacement techniques are being developed and implemented, they remain costly, are still associated with a high rate of complications, and are not appropriate for all patients [189, 190].  AVS is characterized by thickening and subsequent calcification of the valve cusps. Calcification is a central feature of AVS and contributes substantially to cusp thickness, rigidity, rapid disease progression, and poor survival outcomes [191]. Calcification of aortic valves is now believed to be an active process [69, 192], with features in common with both pathological and normal bone calcification [98, 193, 194]. The molecular mechanisms leading to pathological calcification of the valve tissue are not completely understood. However, recent studies have associated the development of AVS with a response to injury characterized by the accumulation and activation of valvular interstitial cells (VIC) that shift from a myofibroblast phenotype into an osteoblast-like phenotype [22, 192, 195-197]. The precise molecular pathways involved in the development of this phenotype shift are mostly unknown, but examination of human calcified aortic valve tissue has revealed that several pathways may be involved, including the bone 55  morphogenic protein (BMP), transforming growth factor β (TGFβ), and Wnt/βcatenin pathways [96, 187, 198-201].  As valve calcification progresses, the VIC-derived osteoblast-like cells express alkaline phosphatase and the bone specific transcription factor, RUNX2[69, 202-204]. The process is also associated with increased levels of TGFβ [205], particularly the TGFβ1 isoform [206]. Elevated levels of TGFβ1 in calcified aortic valves also precedes the increased collagen deposition associated with fibrotic valves and the decreased levels of α-smooth muscle actin (αSMA) associated with loss of a myofibroblast phenotype in VIC, suggesting that TGFβ1 in fact plays an active role in altering VIC phenotype to that of an osteoblast like cell [187, 206-208]. The TGFβ signaling pathway is composed of the TGFβ ligands, TGFβ type I and II receptors, and the SMAD signal transducers [209]. Upon activation, the receptor complex phosphorylates the receptor-regulated SMAD proteins (R-SMADs), including SMAD2 and SMAD3. Activated R-SMADs interacts with SMAD4, inducing translocation of the complex to the nucleus, binding to specific DNA elements and regulation of transcription of target genes [210-212].  Additionally implicated in aortic valve calcification is the canonical Wnt pathway [213-219]. Wnt protein ligands interact with the frizzled (FZD) family transmembrane receptors and low-density lipoprotein receptor-related proteins 5 and 6 (LRP5 and LRP6) co-receptors [220, 221]. Signal is then transduced through dishevelled proteins (DVL), resulting in the stabilization and nuclear translocation of β-catenin. β-catenin then binds to and activates T-cell factor and lymphoid enhancer factor (TCF/LEF) transcription factors [222]. Wnt signaling is regulated by a variety of endogenous molecules, such as the Dickkopf (DKK) protein family and DKK1 in particular, which functions as a Wnt antagonist [105, 223]. DKK1 levels are themselves 56  regulated in part through the TGFβ signaling pathway [84] and DKK1 regulation is known to be an indirect regulatory pathway involved in fibrosis [127, 224, 225]. Given these observations, we hypothesized that activation of TGFβ signaling may be a critical regulatory event in aortic valve calcification, leading to down-regulation of DKK1 allowing subsequent activation of the Wnt pathway.                   57  3.2 Materials and Methods 3.2.1 Tissue Procurement Pieces of human aortic valve tissue were obtained via the Cardiovascular Tissue Registry at the James Hogg Research Centre/Centre for Heart Lung Innovation, St. Paul’s Hospital and University of British Columbia, Vancouver, British Columbia, Canada. Donor hearts used in this study were explanted at the time of cardiac transplantation in male patients aged 47-52 years with no history of clinically apparent ischemic disease. The reason for choosing male gender for the experiments is because male gender is one of the risk factors for aortic valve stenosis [21]. Immunostaining was performed in cases of normal (n=7), moderate (n=7), and severe (n=10) calcification, as defined echocardiographically by aortic valve area (AVA) [1]. Normal human aortic valve cusps were obtained through the International Institute for the Advancement of Medicine (IIAM) program. All procedures utilizing human tissue were approved by the Research Ethics Board (REB) of the University of British Columbia and Providence Health Care Research Institute and conformed to the principles outlined in the Declaration of Helsinki for use of human tissue and subjects. 3.2.2 Antibodies Antibodies used in this study include anti-WNT3A (Abcam, Inc., Toronto, ON; product number: ab28472), anti-TGFβ1 (Abcam; product #ab64715) and anti-DKK1 (Abcam; product number: ab88334). anti-osteopontin (Sigma-Aldrich, Canada, Oakville, ON; product number: 07264), anti-RUNX2 (Abcam; product #ab76956), anti-β-catenin (BD Biosciences, Mississauga, ON; product #610153), horseradish peroxidase (HRP) conjugated goat anti-mouse (Santa Cruz 58  Biotechnology, Inc., Dallas, TX, USA) and goat anti-rabbit (Santa Cruz; product #sc-2030), and Alexa Flour 488 conjugated anti-mouse (Life Technologies, Burlington, ON; product #A11001). 3.2.3 Histology and Immunohistochemistry Histology and immunohistochemistry were performed as previously described [161]. Briefly, formalin fixed paraffin embedded tissue sections were cut at 4 µm thickness and either stained with hematoxylin and eosin (H&E) or von Kossa calcium stain, or deparaffinized for immunohistochemistry. Antigen retrieval was performed for 15 min using citrate buffer and slides were blocked with Protein Block Serum Free blocking solution (Dako North America, Inc., Cupertino, CA, USA; product #X0909) followed by incubation at  4°C overnight with primary antibodies anti-WNT3A (Abcam, product #ab28472), anti-TGFβ1 (abcam, product #ab64715) or anti-DKK1 (abcam, product #ab88334). Secondary antibody incubation was performed for 1 h at room temperature with horseradish peroxidase (HRP) conjugated goat anti-mouse or goat anti-rabbit (Santa Cruz Biotechnology, Inc., Dallas, TX, USA; product #sc-2005 and sc-2030). Slides were counterstained with hematoxylin, cover-slipped and evaluated by brightfield light microscope. Semi-quantitative scoring of staining intensity for von Kossa and immunohistochemistry was performed in a blinded fashion by a cardiovascular pathologist in accord with established guidelines [2]. 3.2.4 Isolation of Human Valvular Interstitial Cells Human valvular interstitial cell isolation and culture was established by a modification of the enzymatic dispersion technique [158, 159]. Briefly, valve tissues were freshly obtained as described above and immediately washed in MCDB-131 medium. Tissue was minced with a 59  sterilized blade. Enzyme I (2.5 mg/mL collagenase II, Roche, product #11088882001) was applied to the exposed media (MCDB-131 + 10% FBS) and tissue followed by 1 h incubation at 37°C to separate the cells from extracellular matrix (ECM). The tissue was broken down by pipetting a few times and transferred to a 15 ml tube. Cells were spun down and 3 ml of fresh collagenase was added to the cells and incubated for 24 hours with pipetting at regular intervals to further disperse cells. A panel of antibodies against cytoskeletal elements was used to phenotype the isolated VIC by immunohistochemistry. The valve cells demonstrated positive staining for smooth muscle α-actin, vimentin and desmin, but were negative for smooth muscle myosin heavy chain, suggesting these cells display a myofibroblast-like phenotype in culture [157]. 3.2.5 Tissue and Cell Culture Valvular interstitial cells (VIC) were isolated from non-calcified valves by enzymatic digestion as described above. Isolated VIC were cultured in MCDB-131 medium (Gibco) containing 15% fetal bovine serum and 100U/mL penicillin/streptomycin. Cells were maintained in a humidified incubator at 37°C with 5% CO2 and used for experiments between passages 2 and 4 as previously published [157]. Cell phenotype was confirmed via immunostaining for myofibroblast markers [157]. Cultured VIC grown for 3-4 days to 70% confluence were treated with recombinant human WNT3A (100 ng/mL; R&D Systems, Inc., Minneapolis, MN; product #5036-WN-010), TGFβ (1ng/mL, BioLegend, Inc., SanDiego, CA; product #580702) and/or DKK1 (10µg/mL; R&D Systems; product #5439-DK-010), separately and in combination, for up to 96 hours. Recombinant proteins were dissolved in PBS prior to dilution, and control 60  conditions utilized culture with equivalent amounts of vehicle only. Some cultures were treated with 10mM LiCl.  3.2.6 OsteoImage Mineralization Assay The OsteoImage Bone Mineralization Assay (Lonza Group, Ltd., Switzerland; product #PA-1503) was used to quantify mineralization in vitro; this assay is based on specific binding of a fluorescent dye to hydroxyapatite (Ca10(PO4)6(OH)2) [226, 227]. Treated cells were fixed in ethanol for 20 minutes, assayed according to the manufacturer’s protocol, and evaluated on a GENios fluorescent plate reader (Tecan Systems, Inc., San Jose, CA, USA) at 492nm excitation and 520nm emission. 3.2.7 Alkaline Phosphatase Activity Assay Alkaline phosphatase activity in cultured cells was measured by SensoLyte® pNPP Alkaline Phosphatase Assay Kit Colorimetric (AnaSpec, Inc., Fremont, CA, USA; product #72146). This assay colorimetrically detects alkaline phosphatase activity using pNPP (p-Nitrophenyl phosphate) phosphatase substrate [166, 167]. The assay was performed according to the manufacturer’s protocol for biological samples. The intensity of color corresponding to the alkaline phosphatase activity was read using VersaMax ELISA Microplate Reader (Molecular Devices, Sunnyvale, CA, USA).  3.2.8 Western Blots Cells were washed twice with ice-cold phosphate buffered saline (PBS), lysed in 80 µL “modified oncogen science lysis buffer” (MOSLB): (50 mM sodium pyrophosphate, 50 mM 61  NaF, 50 mM NaCl, 5 mM EDTA, 5 mM EGTA, 100 µM Na3VO4, 10 mM HEPES, 0.1% Triton X-100) on ice for another 5 min then scraped into Eppendorf tubes and spun at 4°C for 10 min at 3500 rpm. The supernatants were collected into a fresh tube and stored at -80°C. Protein concentration was determined utilizing the Bio-Rad Protein Assay (BioRad Laboratories, Inc., Mississauga, ON; product #500-0006). From this, 30 mg of protein was fractionated by 12% SDS-polyacrylamide gels, electrophoresis, transferred to nitrocellulose membranes (GE Biosciences, Quebec, CA; product #CA27376-991), and blocked with PBS / 0.1% Tween-20 / 5% non-fat dry milk for 1 h. The membrane was then incubated with specific ready-to-use primary antibodies at 1:500 dilution: anti-osteopontin (Sigma-Aldrich, Canada, Oakville, ON; product #07264), anti-RUNX2 (Abcam, Inc., Toronto, ON; product #ab76956), anti-DKK1 (Abcam, product #ab88334), and anti-β-catenin (BD Biosciences, Mississauga, ON; product #610153)  overnight at 4°C, followed by secondary antibody binding with horseradish peroxidase (HRP) conjugated goat anti-mouse or goat anti-rabbit (Santa Cruz Biotechnology, Inc., Dallas, TX, USA; product #sc-2005 and sc-2030) for 1 h at room temperature. Immunoblots were developed with SuperSignal West Femto Chemiluminescence Substrate as per the manufacturer’s directions (GE Biosciences; product #34095) and visualized on a Chemi Genius2 imaging system (Syngene / Synoptics Ltd., Fredrick, MD, USA).  3.2.9 Immunocytochemistry on VIC Some of the cultured cells were fixed in vitro in 4% formaldehyde for 20 min. These cells were then permeabilized with 0.1% Triton X-100 (Life Technologies; product #15596) for 20 min, blocked for 30 min with 1% Protein Block Serum Free (Dako North America, Inc., Cupertino, CA, USA; product #X0909), and incubated overnight at 4°C with primary antibody 62  anti-β-catenin (BD Bioscience, product #610153) at a dilution of 1:100 in 1% bovine serum albumin (BSA). Following primary antibody binding, cells were incubated with Alexa Flour 488 conjugated anti-mouse secondary antibody (Life Technologies, product #A11001) at a dilution of 1:250 in 1% BSA for 1 hour at room temperature in the dark. Cells were counterstained for 5 min with NucBlue Fixed Ready Probes Reagent DAPI (Life Technologies, product #R37606) to visualize nuclei and then coverslipped with Slow Fade Gold antifade reagent (Life Technologies, product #S-36936). Examination of cells was performed using a Nikon Eclipse TE300 inverted fluorescence microscope.  3.2.10 Statistical Analysis Data are expressed as the mean ± standard error (SE) of at least three independent determinations. Differences between means were determined using a one-way analysis of variance (ANOVA) with a pairwise comparison by the Tukey-Kramer method. Differences were considered to be significant when the P value was less than 0.05. Analyses were performed using the Graphpad Prism 6 software package (www.graphpad.com).         63  3.3 Results 3.3.1 WNT3A and TGFβ1 Are Up-regulated in AVS Previous studies have reported that WNT3A is expressed in calcified aortic valves [22]. In normal valve tissues, expression of WNT3A is almost undetectable, while expression is increased in AVS tissues (Table 1A, Figure 15A). A similar pattern is seen for TGFβ1, and the increased expression correlates with the degree of calcification (Table 1B, Figure 15B).  In contrast expression of DKK1, a well-studied Wnt pathway inhibitor [102, 223, 228-230], is decreased in tissues with AVS compared to normal valve tissue (Table 1C, Figure 15C). SOST1 [231] and PPARγ [232], two other known WNT3A pathway inhibitors, are not detectable in these tissues (data not shown).     64    Table 1. Proportion and intensity scores for WNT3A, TGFβ1 and DKK1 immunohistochemical staining in human aortic valve cusps.  Human aortic valves categorized by clinical rates for stenosis [1] and by different degree of calcification by von Kossa staining and aortic valve area (AVA) stained for WNT3A (A), TGFβ1 (B) and DKK1 (C) expression. Histological grading for the proportional and intensity of positive staining was evaluated blindly by a cardiovascular pathologist on a semi-quantitative scale who scored the proportion and intensity of the positive staining in accord with established guidelines [2]. Stenotic and more calcified valves expressed more WNT3A (A) and TGFβ1 (B) as compared to normal valves. In contrast, normal valves expressed more DKK1 (C).      65   Figure 15. Immunohistochemistry of human aortic valves with different degrees of calcification for WNT3A, TGFβ1, and DKK1.  A. Representative images of immunohistochemistry staining of human aortic valves with different calcification degree for WNT3A. Arrows indicate the positive staining of VIC for WNT3A. The extent of intensity and/or proportion of WNT3A staining increases with the degree of tissue calcification. (Scale bar for 40X magnification images=20μm and for 10X magnification=50μm).        66   B. Representative images of immunohistochemistry staining of human aortic valve with different calcification degrees for TGFβ1. Arrows indicate the positive staining of VICs for TGFβ1. The extent of intensity and proportion of TGFβ1 staining increases with the degree of tissue calcification. (Scale bar for 40X magnification images=20μm and for 10X magnification=50μm).                 67   C. Representative images of immunohistochemistry staining of human aortic valve with different calcification degrees for DKK1. Arrows indicate the positive staining of VICs for DKK1. In contrast to WNT3A and TGFβ1the extent of intensity and proportion of TGFβ1 staining decreases as the degree of tissue calcification increases to the point that in severely calcified valves DKK1 staining is almost undetectable. (Scale bar for 40X magnification images=20μm and for 10X magnification=50μm).            68  3.3.2 Exogenous WNT3A Induces an Osteoblast Phenotype in Cultured Human VIC To study the possibility that the canonical Wnt pathway plays a role in aortic valve calcification, normal human primary VIC were incubated with human recombinant WNT3A and levels of cell calcification were measured using the Lonza OsteoImage assay. Cells treated for 96 h with WNT3A demonstrated significantly higher levels of cell calcification compared to untreated controls (Figure 16A). Activity of alkaline phosphatase, a calcification/bone metabolism associated enzyme, is significantly increased in cells treated with WNT3A compared to untreated cells (Figure 16B). Additionally, cells incubated with both WNT3A and TGFβ1 showed increased expression of RUNX2 (Figure 17A), bone specific transcription factor, and osteopontin (Figure 17B) a secreted protein expressed by osteoblasts [170-174].              69   Figure 16. Cell calcification and alkaline phosphatase activity levels in cell treated with TGFβ1, WNT3A or both.  VIC (n=3 for each experimental condition) were serum starved 24 h before treatment. VIC were treated with human recombinant proteins WNT3A (100 ng/ml) or TGFβ1 (10 ng/ml) separately and together for 96 h. Calcification was determined using OsteoImage™ fluorescent mineralization assay (RFU=Relative Fluorescent Unit) (A), and alkaline phosphatase activity using pNPP alkaline phosphatase colorimetric kit (B). *** indicates p-value < 0.005. Error bars represent mean ± SEM.  U n tr e a te d T G FW N T 3 A W N T 3 A + T G F01 0 0 0 02 0 0 0 03 0 0 0 04 0 0 0 0RFU (492/520)A . C e ll  C a lc if ic a t io n* * *U n tr e a te d T G FW N T 3 A W N T 3 A + T G F051 01 5U/LB . A lk a lin e  P h o s p h a ta s e A c tiv ity* * *70                   Figure 17. Expression of RUNX2 and osteopontin in treated VIC.  Serum starved VIC (n=3 for each experimental condition) were treated with WNT3A (100 ng/ml) and TGFβ1 (10 ng/ml) separately and together. Protein expression of proteins RUNX2 and osteopontin was determined by Western blot for treated cells at 6 (A) and 96 h (B). RUNX2 expression in treated cells tended to increase with either treatment and tended to be greater with combined treatment of TGFβ1 and WNT3A, although this was not statistically significant (A). Osteopontin expression was also significantly increased in cells treated simultaneously with TGFβ1 and WNT3A (B). *** indicates p-value < 0.001 significant difference in comparison to unstimulated cells (B). Error bars represent mean ± SEM. Densitometry Unit(Normalized to GAPDH)U n tr e a te d T G F1 W N T 3 A T G F1 + W N T 3 A0123A . R u n x 2  P r o te in  E x p r e ss io nDensitometry Unit(Normalized to GAPDH)U n tr e a te d T G F1 W N T 3 A T G F1 + W N T 3 A0123* * *B . O s te o p o n tin  P r o te in  E x p r e s s io nA. RUNX2 Protein Expression 71  3.3.3 Canonical Wnt Signaling Regulates Human VIC Calcification To further test whether the canonical Wnt pathway plays a role in VIC calcification we incubated cultures with LiCl, a stimulus known to activate endogenous Wnt signaling through inhibition of GSK3β[233, 234] (Figure 18). LiCl (10 mM) alone slightly increased levels of calcification, albeit not significantly. The combination of WNT3A and LiCl however, significantly increased calcification compared to controls or either treatment alone (Figure 19). To test for the possible inhibitory effect of DKK1, a known Wnt pathway inhibitor, on cell stimulation we added exogenous human recombinant DKK1 protein to the cultures. The addition of DKK1 to cultures treated with WNT3A induced a significant reduction in calcification and alkaline phosphatase activity compared to cells treated with WNT3A alone (Figure 20).         72  β-catenin APC AXIN GSK-3β P Degraded β-catenin LiCl       Figure 18. LiCl mechanism of action.  LiCl mimics the canonical Wnt pathway by inhibiting the formation of destructive complex responsible for β-catenin degradation.             73              Figure 19. Calcification levels in WNT stimulated cell cultures.  VIC (n=3 for each experimental condition) were serum starved 24 hours before treatment. Cells were treated with human recombinant WNT3A (100 ng/ml) or LiCl (10 mM) separately and together and incubated for 96 h. Cell calcification levels were determined using OsteoImage™ fluorescent mineralization assay (RFU=Relative Fluorescent Unit). * indicates p-value < 0.05 compared to WNT3A treated cells and *** indicates p-value < 0.005 compared to unstimulated cells and cells treated with WNT3A. Error bars represent mean ± SEM. RFU (492/520)u n tre a te d L iC l W N t3 A W N tT 3 A + L iC l05 0 0 01 0 0 0 01 5 0 0 02 0 0 0 0* * **C e ll  C a lc if ic a t io nUntreated            LiCl              WNT3A       WNT3A+LiCl 74                    Figure 20. Effect of DKK1 treatment on calcification and alkaline phosphatase activity levels in WNT3A treated cells.  VIC (n=3 for each experimental condition) were serum starved for 24 hours and treated with human recombinant proteins WNT3A (100 ng/ml) or DKK1 (10 µg/ml) separately and together for 96 h. After incubation, cells were either fixed in EtoH or lysed to determine levels of calcification (A) and alkaline phosphatase activity (B), respectively. Calcification was determined using OsteoImage™ fluorescent mineralization assay (RFU=Relative Fluorescent Unit) (A), and alkaline phosphatase activity using pNPP alkaline phosphatase colorimetric kit (B). Cell calcification was significantly suppressed by treatment with DKK1, (* indicates p-value RFU (492/520)U n tr e a te d D K K 1 W N T 3 A W N T 3 A + D K K 105 0 0 01 0 0 0 01 5 0 0 02 0 0 0 0*A . C e ll  C a lc if ic a t io nU/LU n tr e a te d D K K 1 W N T 3 A W N T 3 A + D K K 1024681 0* * *B . A lk a lin e  P h o sp h a ta s e  A c tiv ity75  < 0.05; A). Alkaline phosphatase activity also declined significantly with DKK1 treatment (*** p-value < 0.005; B). Error bars represent mean ± SEM.                                       76  3.3.4 TGFβ1 Activation of the Canonical Wnt Signaling Pathway is associated with Down-regulation of DKK1 Since TGFβ has a pivotal role in the development of AVS [235-239] we explored a possible role of TGFβ in activation of the canonical Wnt pathway in AVS. TGFβ1 treatment of cultured VIC results in nuclear translocation of β-catenin, consistent with Wnt pathway activation (Figure 21). This was accompanied by a reduction in DKK1 expression (Figure 22), increased cell calcification (Figure 16 A) and alkaline phosphatase activity (Figure 16 B), and increased expression of RUNX2 (Figure 17 A) and osteopontin (Figure 17 B).                             77                      Figure 21. Nuclear translocation of β-catenin treated with TGFβ1 in human VIC.  Immunohistochemistry was performed on cultured VIC (A-D). Cells were stained for β-catenin (A, B), and for nuclei with DAPI (C). Overlay image shows the β-catenin expression in nuclei of cells treated with human recombinant TGFβ1 (D). (Scale bar = 48 µm). Western blot was also performed on nuclear and cytoplasmic fractions of cells to confirm nuclear translocation β-catenin in TGFβ1-treated cells (E).  78                    Figure 22. Cells treated with TGFβ1 express less DKK1. Serum starved VIC (n=3 for each experimental condition) were serum starved for 24 hours and treated with human recombinant TGFβ1 (10 ng/ml) for 96 hours. Western blot analysis of VIC cell lysates indicated a significant decrease in the expression of DKK1 in treated cells (A and B). Protein expression of DKK1 was quantified and normalized to GAPDH (A) and visualized (B) by Western blot analysis, * p-value < 0.05 (A). Error bars represent mean ± SEM.   U n tre a te d T G F0 .00 .20 .40 .60 .81 .09 6 h r sDensitometry Unit(Normalized to GAPDH)A . D K K 1  P r o te in  E x p r e s s io n*B. 79  3.4 Discussion In the present study, the canonical Wnt pathway signaling induces calcification in cultured VIC, with an associated shift to an osteoblast-like phenotype, a phenomenon previously reported [118, 240, 241]. This signaling can be inhibited by the Wnt pathway inhibitor, DKK1. Treatment with TGFβ1 results in down-regulation of DKK1, allowing for Wnt pathway signaling to promote calcification. These findings integrate the previously reported high levels of WNT3A and TGFβ1 associated with AVS, and are also analogous to pathways involved in chondrocyte maturation, skin fibrosis, and liver fibrosis [127, 242, 243].  The calcification we observed was associated with, among other things, increased alkaline phosphatase activity. Such enzymatic activity is associated with active calcification processes, and supports a model in which AVS is an active process [244-249]. This is encouraging for the development of therapies aimed at reversing calcific AVS. In fact, DKK1, the Wnt pathway inhibitor that we found to be expressed in normal aortic valves and down-regulated in calcified valves, is being investigated by others as a potential therapeutic target in other diseases [250, 251], and thus may be a candidate therapeutic target in AVS.  We acknowledge several limitations of our study. First, most of the data are derived from cultured cells. While the material is from primary human culture, not a cell line and not an animal model, the act of culturing cells is known to have phenotypic impact [252]. Additionally, any interactions with other cell types are unlikely to be retained in our culture system. We feel, however, that the data obtained are nonetheless compelling enough to propose an integrated signaling model, potentially leading to more complex in vivo and/or candidate drug studies.  80  Further, much of our data are correlative; we have not strictly documented direct causative actions of many of the pathways implicated. We attempted use of siRNA to confirm the results, but the method has been unsuccessful to date in our cultured VIC. The use of exogenous DKK1 does strongly support the direct role of canonical Wnt signaling in VIC calcification. We did not utilize TGFβ1 signaling pathway inhibitors to confirm the direct effect of that pathway on down-regulation of DKK1; however, a prior report [225] that p38 activation downstream of TGFβ1 directly down-regulates DKK1. Our data lead us to present an integrated model in which TGFβ pathway activation down-regulates DKK1, dis-inhibiting the canonical Wnt signaling pathway, allowing for development of an osteoblastic phenotype, and resulting calcific matrix production (Figure 23). This model does not, however, suggest the native triggers of WNT3A or TGFβ1 expression that lead to AVS. As not all individuals have aortic valve calcification, and there is tremendous variation amongst those that do, some initial stimulus or stimuli must contribute to a switching the healthy valve phenotype (low WNT3A and TGFβ1, high DKK1) to the pro-calcific state (high WNT3A and TGFβ1, low DKK1). Regardless of the trigger, the active, dynamic nature of AVS implied through our data, and the multiple pathways involved, suggest sites for therapeutic intervention that might not only prevent disease progression, but might also contribute to some reversal of the phenotype. It is our hope that our insights will significantly contribute to addressing this significant medical issue.    81   Figure 23. Working model of the interaction between the Wnt and TGFβ pathways in valvualr calcification.                                         In our VIC cultured model increased phosphorylation of SMAD2 (data not shown) upon TGFβ activation may suppress DKK1, a WNT3A inhibitor, allowing the WNT3A signal to be transmitted via canonical WNT receptor through the cell membrane. This will result in cytoplasmic β-catenin accumulation and consequently nuclear translocation of β-catenin where it may increase the transcription of genes fundamental to calcification.  82  Chapter 4: 1Α, 25(OH)2D3 REGULATES VALVULAR INTERSTITIAL CELL CALCIFICATION  4.1 Introduction Aortic valve calcification (AVC) includes a spectrum of disease processes from early alterations in valvular interstitial cell phenotypes to sclerotic thickening and matrix remodeling of the valve leaflets to stenosis and leaflet calcification [253]. AVC was first thought to be a degenerative process resulting from the “wear and tear” of aging [17, 192]. Now AVC is known to be an actively regulated process [197]. However, the cellular events and molecular mechanisms driving AVC, particularly in the early stages of the disease, have yet to be determined. Calcification is a prominent feature of AVC and of course contributes substantially to leaflet thickness, rigidity, and rapid disease progression leading to poor outcomes [191]. Calcification of aortic valves is an active process, in which several of the classical regulators of calcification, initially identified in developing bone, also seem to participate in AVC [241, 254, 255]. As valve stenosis progresses, a subpopulation of valvular myofibroblasts undergoes phenotypic transformation to osteoblast-like cells in vitro. These cells express bone specific markers including: alkaline phosphatase, osteopontin, and RUNX2 (Chapter 2 and 3). The observation that osteoblast-like cells secrete bone specific markers in AVC raises the possibility that osteogenic regulatory molecules involved in bone development could also play a role in development of valvular calcification. Among those regulators participating in bone formation, (vitamin D that is usually prescribed to enhance calcification of bone tissue [256, 257]), may also play a role in calcification of the aortic valve. In fact some case study reports have shown that 83  taking vitamin D supplement increases AVC in a patien with a history of valvular disease [258] and that serum phosphate levels are associated with valvular calcification [202]. 1,25(OH)2D3 (vitamin D3) is a steroid hormone that enhances bone formation by regulating calcium and phosphate metabolism [259]. The activities of vitamin D3 are mediated through a specific vitamin D receptor protein (VDR) [260]. While some have shown that vitamin D administration may induce bone resorption [261], others have provided evidence that administration of vitamin D can stimulate bone deposition [262] confirming the complexity of vitamin D activity in the body.  The role of vitamin D3 is not just limited to regulating bone formation. There is evidence for the expression of VDR in brain, prostate, breast and colon tissues in support of the idea that there are other roles for vitamin D3 other than the calcium deposition in bone tissue [263]. It is not surprising then that there are studies indicating an association of vitamin D with both vascular and valvular calcification in patients with impaired renal function and a history of cardiovascular disease including AVC [264, 265], however the mechanism by which vitamin D might play a role in AVC is still unknown.  At the molecular level in an in vitro study of pulmonary fibroblasts treated with vitamin D3, fibroblast activation by TGFβ1was inhibited. The authors showed that vitamin D inhibited TGFβ mediated activation of fibroblasts and in that process reduced αSMA (α smooth muscle actin) expression, a myofibroblast marker, that was expressed in fibroblasts treated with TGFβ [260].  84  In addition to its effect on the TGFβ pathway it has been shown that the WNT3A induced proliferation and migration of cells in neoplasia of the colon is abrogated by vitamin D [266-268]. These effects of vitamin D on both the WNT3A and TGFβ functions in other tissues suggested to us that possibly vitamin D could play a regulatory role in activating the pathways that we have previously shown to collaborate in VIC transformation to osteoblast-like cells and calcification in vitro. The purpose of this study was to test the hypothesis that vitamin D might induce VIC transformation and calcification process in vitro. This could add significantly to our understanding as to whether and how vitamin D may influence aortic valve calcification through direct effects on these pathways.        85  4.2 Materials and Methods 4.2.1 Tissue Procurement Pieces of human aortic valve tissue were obtained via the Cardiovascular Tissue Registry at the James Hogg Research Centre/ Centre for Heart Lung Innovation, St. Paul’s Hospital and University of British Columbia, Vancouver, British Columbia, Canada. Donor hearts used in this study were explanted at the time of cardiac transplantation in male patients aged 47-52 years with no history of clinically apparent ischemic disease. The reason for choosing male gender for the experiments is because male gender is one of the risk factors for aortic valve stenosis [21]. Immunostaining was performed in cases of normal (n=7), moderate (n=7), and severe (n=10) calcification, as defined echocardiographically by aortic valve area (AVA) [1]. Normal human aortic valve cusps were obtained through the International Institute for the Advancement of Medicine (IIAM) program. All procedures utilizing human tissue were approved by the Research Ethics Board (REB) of the University of British Columbia and Providence Health Care Research Institute and conformed to the principles outlined in the Declaration of Helsinki for use of human tissue and subjects. 4.2.2 Isolation of Human Valvular Interstitial Cells Human valvular interstitial cell isolation and culture was established by a modification of the enzymatic dispersion technique [158, 159]. Briefly, valve tissues were obtained as described above and immediately washed in MCDB-131 medium. Tissue was minced with a sterilized blade. Enzyme I (2.5 mg/mL collagenase II; Roche, product #11088882001, USA) was applied to the exposed media (MCDB-131 + 10% FBS) and tissue followed by 1 hour incubation at 37°C 86  to separate the cells from extracellular matrix. The tissue was broken down by pipetting a few times and transferred to a 15-ml tube. Cells were spun down and 3 ml of fresh collagenase was added to the cells and incubated for 24 hours with pipetting at regular intervals to further disperse cells.  4.2.3 Treatment of Cell Cultures Valve interstitial cells were cultured in MCDB 131 medium (Gibco) containing 15% fetal bovine serum and 100U/ml penicillin/streptomycin. Cells were maintained in a humidified incubator at 37°C with 5% CO2. Cells were used for experiments between passages 2 and 4 [157]. Cultures that had been grown for 3–4 days and were 70% confluent were treated with human recombinant proteins; cultured VICs were treated with 100ng/ml recombinant human WNT3A (R&D Systems; product #5036-WN-010, Minneapolis, MN), 1ng/ml recombinant human TGFβ1 (BioLegend Laboratory; product #580702, Mississauga, ON) and 1µM 1, 25(OH)2D3 (Sigma-Aldrich; product #D1530), separately and in combination. Cells were collected following 6, 24, 48 and 96 hours of treatment for subsequent procedures.       4.2.4 Histology and Immunohistochemistry Histology and immunohistochemistry were performed as previously described [161]. Briefly, formalin-fixed paraffin embedded tissue sections were cut at 4 µm thickness and either stained with hematoxylin and eosin (H&E) or von Kossa calcium stain, or deparaffinized for immunohistochemistry. Antigen retrieval was performed for 15 min using citrate buffer and slides were blocked with Protein Block Serum Free blocking solution (Dako North America, Inc., Cupertino, CA, USA; product #X0909) followed by incubation at 4°C overnight with 87  primary antibodies Anti-VDR (abcam; product #39990). Secondary antibody incubation was performed for 1 h at room temperature with horseradish peroxidase (HRP) conjugated goat anti-rabbit (Santa Cruz Biotechnology, Inc., Dallas, TX, USA; product #sc-2005 and sc-2030). Slides were counterstained with hematoxylin, cover-slipped and evaluated by brightfield light microscope. Semi-quantitative scoring of staining intensity for von Kossa and immunohistochemistry was performed in a blinded fashion by a cardiovascular pathologist in accord with established guidelines [2]. 4.2.5 Western Blots Cells were washed twice with ice-cold phosphate buffered saline (PBS), lysed in 80µL “modified oncogen science lysis buffer” (MOSLB): (50 mM sodium pyrophosphate,. 50 mM NaF, 50 mM NaCl, 5 mM EDTA, 5 mM EGTA, 100 µM Na3VO4, 10 mM HEPES, 0.1% Triton X-100) on ice for another 5 min then scraped into Eppendorf tubes and spun at 4°C for 10 min at 3500 rpm. The supernatants were collected into a fresh tube and stored at -80°C. Protein concentration was determined utilizing the Bio-Rad Protein Assay (BioRad Laboratories, Inc., Mississauga, ON; product #500-0006). Thirty milligrams of protein was fractionated by 12% SDS-polyacrylamide gels, electrophoresis, transferred to nitrocellulose membranes (GE Healthcare Lifesciences, Quebec, CA; product #EP4HYA0010), and blocked with PBS / 0.1% Tween-20 / 5% non-fat dry milk for 1 h. The membrane was then incubated with specific ready-to-use primary antibodies at 1:500 dilution; anti-osteopontin (Sigma-Aldrich, Canada, Oakville, ON; product #07264), anti-RUNX2 (Abcam, Inc., Toronto, ON; product #ab76956), Anti-VDR (abcam; product #39990) and anti-AXIN2 (abcam; product #ab32197), anti-pSMAD2 (Cell signaling; product #3101S) overnight at 4°C, followed by secondary antibody binding with 88  horseradish  peroxidase (HRP) conjugated goat anti-mouse or goat anti-rabbit (Santa Cruz Biotechnology, Inc., Dallas, TX, USA; product #sc-2005 and sc-2030) for 1 h at room temperature. Immunoblots were developed with SuperSignal West Femto Chemiluminescence Substrate as per the manufacturer’s directions (GE Biosciences; product #34095) and visualized on a Chemi Genius2 imaging system (Syngene / Synoptics Ltd., Fredrick, MD, USA).  4.2.6 Alkaline Phosphatase Activity Assay Alkaline phosphatase activity in cultured cells was measured by SensoLyte® pNPP Alkaline Phosphatase Assay Kit Colorimetric (AnaSpec, Inc., Fremont, CA, USA; product #2146). This assay colorimetrically detects alkaline phosphatase activity using pNPP (p-Nitrophenyl phosphate) phosphatase substrate [166, 167]. The assay was performed according to the manufacturer’s protocol for biological samples. The intensity of color corresponding to the alkaline phosphatase activity was read using VersaMax ELISA Microplate Reader (Molecular Devices, Sunnyvale, CA, USA).  4.2.7 OsteoImage Mineralization Assay The OsteoImage Bone Mineralization Assay (Lonza Group, Ltd., Switzerland; product #PA-1503) was used to quantify mineralization in vitro; this assay is based on specific binding of a fluorescent dye to hydroxyapatite (Ca10(PO4)6(OH)2) [226, 227]. Treated cells were fixed in ethanol for 20 min, assayed according to the manufacturer’s protocol, and evaluated on a GENios fluorescent plate reader (Tecan Systems, Inc., San Jose, CA, USA) at 492 nm excitation and 520 nm emission. 89  4.2.8 Statistical Analysis Data are expressed as the mean ± standard error (SE) of at least three independent determinations. Differences between means were determined using a one-way analysis of variance (ANOVA) with a pairwise comparison by the Tukey-Kramer method. Differences were considered to be significant when the P value was less than 0.05. Analyses were performed using the Graphpad Prism 6 software package (www.graphpad.com).          90  4.3 Results 4.3.1 VDR Expression in Primary Valvular Interstitial Cells of Human Aortic Valves To begin, we sought to determine the levels of expression of VDR in human aortic valve cusp tissues showing different degrees of calcification in three groups of control, moderate and severe calcification. Furthermore, since valvular interstitial cells are the predominant cells in aortic valves and known to be responsible for valvular calcification, we also assessed the expression of VDR in human primary valvular interstitial cells in vitro. In vivo VDR was present in both normal and calcified human aortic valves but its expression was significantly elevated with increasing calcification of the diseased tissue (Figure 24, Table 2). VDR was also detected in the human VICs in vitro, using our cultures of primary valve myofibroblasts (Figure 25). The expression of VDR also increased significantly in the cells stimulated with 1, 25(OH)2D3 in a concentration-dependent manner in vitro up to 0.1 µM and decreased with higher concentration of vitamin D3. These cells were treated with 0, 0.05, 0.1, 0.2, and 0.5µM 1, 25(OH)2D3 (vitamin D3) and collected after 48 hours (Figure 25). VDR expression with TGFβ1 and WNT3A was not altered (Data not shown). It has been demonstrated that vitamin D stimulation can activate its nuclear receptor, VDR, and increase its expression up to 10 fold [269].     91   Figure 24. Vitamin D receptor (VDR) is present in human aortic valve and valvular interstitial cells.              Representative sections show immunoreactivity for VDR . Non-calcified (Normal), moderately calcified, and severely calcified human aortic valve tissues.  Arrows indicate positive staining of VDR which is localized in nucleus of valvular interstitial cells. The VDR expression was assessed by an observer blinded to treatment assignment using image-Pro Plus software (Media Cybernetics), Nikon E600 microscope. (Scale bar = 20 μm) 92       Table 2. In vivo proportion and intensity scores for VDR immunohistochemical staining in human aortic valve cusps.            Human aortic valves categorized by clinical rates for stenosis [1] and by different degree of calcification by von Kossa staining and aortic valve area (AVA) stained for VDR expression. Histological grading for the proportional and intensity of positive staining was evaluated blindly by a cardiovascular pathologist on a semi-quantitative scale who scored the proportion and intensity of the positive staining in accord with established guidelines [2]. Although VDR expression was present in most of the tissues the stenotic and more calcified valves expressed more VDR as compared to normal valves.   93               Figure 25. Vitamin D receptor (VDR) is present in human aortic valve and valvular interstitial cells.  Serum starved VIC (n=3 for each experimental condition) were treated with graded concentrations of 1α, 25(OH)2D3 (1µM) for 24 hours. Cell lysates were probed for VDR and normalized to loading control. * p-value <0.05 vs untreated control. Analysis was performed on three independent blots and the error bars represent the mean values ± SEM. 0 M0.050.10.20.5012345V ita m in  DDensitometry (Fold Change)A . V D R  P r o te in  E x p r e s s io n*94   4.3.2 Vitamin D3 and the Phenotype Shift of VIC In order to study the possibility that vitamin D plays a role in shifting VIC phenotype, normal human primary valvular interstitial cells were incubated in vitro with human recombinant TGFβ1 protein and vitamin D separately and in combination for 6, 24, 48 and 96 hours, and the morphology of the cells was monitored photographically at each time point. After 96 hours, untreated cells were large, spread out and stellate in shape with long cytoplasmic extensions (Figure 26 A). However, cells treated for 96 hours with TGFβ1 were smaller compare to untreated cells, narrow, thin and elongated in groups with similar orientation (Figure 26 B). Interestingly cells treated with vitamin D for 96 hours were smaller, rounded or cubical in shape (Figure 26 C). With TGFβ1 and vitamin D3 added together there was mixture of rounded and spindle shaped cells present in the cultures at 96 hours (Figure 26 D). Along with cell shape we also examined the level of αSMA expression, a myofibroblast marker [270-272] in the VIC in vitro that could indicate a phenotype shift. While αSMA was significantly increased in cells treated with TGFβ1 compared to untreated cells, vitamin D treatment alone and with TGFβ1 significantly reduced the level of αSMA in cells (Figure 27).  The significant decrease in αSMA expression levels in VICs treated with vitamin D is consistent with the proposition that vitamin D has shifted the VIC phenotype away from that of a myofibroblast and in this case toward an osteoblast-like cell.   95             Figure 26. Vitamin D changes the morphology of the VIC. Serum starved VICs (n=3 for each experimental condition) were treated with TGFβ1 (10 ng/ml) and 1α,25(OH)2D3 (1µM) for 96 hours. Photomicrograph of cells at 96 hours post treatment; (A) control, (B) 10 ng/ml TGFβ1 (C) 0.1 µM 1α,25(OH)2D3 and (D) combined treatment of TGFβ1 + 1α,25(OH)2D3. (Scale bar= 50μm)    96              Figure 27. αSMA protein expression. Cell lysates were probed for αSMA and normalized to GAPDH as a loading control. * p-value < 0.05 vs untreated controls. Analysis was performed on three independent blots and the error bars are mean values ± SEM.    97  4.3.3 Vitamin D3 Increases the Expression of Bone Specific Markers To further determine whether VIC were shifting away from a myofibroblast phenotype to an osteoblast phenotype, we measured the expression of opsteopontin and RUNX2, both osteoblast markers [170-174] (Chapter 3). Cells incubated with both TGFβ1 and vitamin D3 showed a significant increase in expression of RUNX2 at 6 hours (Figure 28 A). In addition osteopontin was increased in cells treated with vitamin D at 96 hours post treatment (Figure 28 B). The further increase of expression in cells treated with TGFβ1, WNT3A and vitamin D is consistent with the possibility that vitamin D does play a role in shifting VIC phonotype toward that of osteoblast-like phenotype. Therefore the effect of vitamin D is at least parallel with the TGFβ and the canonical WNT pathways in calcifying VIC in vitro.             98                Figure 28. Vitamin D shifts VIC phenotype toward osteoblast-like cells.  Serum starved VIC (n=3 for each experimental condition) were treated with WNT3A (100 ng/ml), TGFβ (10 ng/ml) or 1α,25(OH)2D3 (0.1 µM)separately and together for 6 or 96 h. Expression of proteins RUNX2 and osteopontin were determined by Western blot for treated cells at 6 (A) and 96 (B) h. time points (A) and (B) respectively. * p-value < 0.05;  ** P-value < 0.01; *** p-value < 0.001 versus untreated control. Analysis was performed on three independent blots and the mean values ± SE shown. (Blots are from the non-sequential wells of the same gel).   1 2 3 4 5 6012345Densitometry (Fold Change)A . R U N X 2  P r o te in  E x p r e ss io n*1 . U n tre a te d2 .  T G F  13 . W N T 3 A4 . V it .D5 .  T G F  1 + W N T 3 A6 .  T G F  1 + W N T 3 A + V it.D1 2 3 4 5 601234Densitometry (Fold Change)B .O s te o p o n tin  P r o te in  E x p r e s s io n* * * ** *1 . U n tre a te d2 .  T G F  13 . W N T 3 A4 . V it .D5 .  T G F  1 + W N T 3 A6 .  T G F  1 + W N T 3 A + V it.D99  4.3.4 Vitamin D3 Can Induce VIC Calcification  Our observation that vitamin D3 treatment can increase levels of osteoblast specific marker expression led us to further investigate the role of vitamin D in the calcification process which we have shown can be regulated by an integrated action of the Wnt3A and TGFβ pathways (Chapter 3). Therefore we incubated human primary valvular interstitial cells in vitro with human recombinant WNTA3A and TGFβ1 proteins along with vitamin D3 separately and in combination for 96 h. At 96 h we measured levels of cell calcification using OsteoImage™ assay. Cell calcification was also significantly increased in co-treated VIC with WNT3A, TGFβ1 and Vitamin D3 (Figure 29 A). Additionally, we measured the levels of alkaline phosphatase activity, as another quantitative method to confirm that vitamin D is involved in the VIC calcification process. Indeed alkaline phosphatase activity was significantly increased in cells treated with WNT3A, TGFβ1 and vitamin D simultaneously in contrast to untreated cells (Figure 29 B).        100  Figure 29. Vitamin D increases alkaline phosphatase activity and calcification levels in vitro. VIC (n=3 for each experimental condition) were serum starved 24 hours before treatment. VIC were treated with human recombinant proteins WNT3A (100 ng/ml), TGFβ1 (10 ng/ml) and 1α,25(OH)2D3 (0.1 µM) separately and in combinations for 96 hours. (A) Alkaline phosphatase activity was measured using pNPP alkaline phosphatase colorimetric kit and (B) calcification was determined using OsteoImage™ fluorescent mineralization assay (RFU=Relative Fluorescent Unit). * indicates p-value < 0.05, ** indicates p-value < 0.01 and *** indicates p-value < 0.005. (Error bars represent mean ± SE) 1 2 3 4 5 601 0 0 0 02 0 0 0 03 0 0 0 04 0 0 0 0RFU (492/520)A . C e ll  C a lc if ic a t io n* * *1 . U n tre a te d2 .  T G F  13 . W N T 3 A4 . V it .D5 .  T G F  1 + W N T 3 A6 .  T G F  1 + W N T 3 A + V it.D1 2 3 4 5 6051 01 52 02 5U/L* **B . A lk a lin e  P h o sp h a ta s e  A c tiv ity1 . U n tre a te d2 .  T G F  13 . W N T 3 A4 . V it .D5 .  T G F  1 + W N T 3 A6 .  T G F  1 + W N T 3 A + V it.D101  4.3.5 The Role of Vitamin D3 in Calcification of VIC Is Independent of β-catenin and SMAD2 Since the canonical Wnt and TGFβ pathways are integrated in the calcification of the VIC (Chapter 3), we wished to determine whether and if so how vitamin D interacts with the TGFβ and/or the canonical Wnt pathways in the process of VIC calcification. First we looked at the levels of phosphorylation of SMAD2, a downstream element of the TGFβ pathway, in VIC treated with vitamin D. Surpringly, the expression level of pSMAD2 was decreased in cells treated with vitamin D after 24 hours (Figure 30 A). This result suggests that vitamin D does not act through SMAD phosphorylation of the TGFβ pathway.  To test the effect of vitamin D in activation of β-catenin a downstream element in the Wnt pathway we examined expression levels of AXIN2 in treated VIC. Since AXIN2 is a downstream protein in the WNT3A pathway that participates in formation of the complex responsible for β-catenin degradation in the canonical Wnt pathway, we expected to see some effect if vitamin D3 is working through the canonical Wnt pathway. Interestingly, the level of expression of AXIN2 did not change in cells treated with vitamin D, suggesting that vitamin D does not act through WNT/β-catenin dependent pathway (Figure 30 B). Taking together these results suggest that vitamin D involvement in VIC calcification may occur via an as yet unidentified mechanism that is independent of TGFβ and Wnt.      102                     U n tr e a te d W N T 3 A V it .D W N T 3 A + V it .D0 .00 .51 .01 .52 .02 .5Densitometry Unit(Normalized to GAPDH)A . A X IN 2  P r o te in  E x p r e s s io nU n tr e a te d T G FV it .D T G F+ V it .D0 .00 .51 .01 .52 .02 .5Densitometry Unit(Normalized to GAPDH)B . p S M A D te in  E x p r e s io n103  Figure 30. Vitamin D3 treatment does not alter the expression of pSMAD2 and AXIN2 in VIC.  Serum starved VIC (n=3 for each experimental condition) were treated with human recombinant proteins WNT3A (100ng/ml), TGFβ1 (10ng/ml) and 1α,25(OH)2D3 (0.1 µM) separately and in combinations for 24 hours. Cell lysates were probed for AXIN2 and normalized to GAPDH (A). Same lysates were probed one more time with an antibody specific to pSMAD2, blots were then stripped and re-probed for SMAD2/3 as a loading control (B). ** p-value < 0.01. Analysis was performed on three independent blots and the mean value ± SE shown. (Blots are from non-sequential wells of the same gel).                  104  4.4 Discussion In the current study we have demonstrated that vitamin D3 contributes to the process of the VIC calcification in vitro, when used in conjunction with TGFβ1 and WNT3A (Chapter 3). However, our data also indicate that vitamin D does not appear to act through the TGFβ1 and Wnt pathways but rather seems to function independently from them contributing to calcification in what appears to be an additive fashion. To the best of our knowledge this is the first study that demonstrates simultaneous but independent effects of vitamin D along with TGFβ1 and WNT3A proteins on VIC calcification in vitro. We have shown that in VIC vitamin D3 does produce calcification and that it is enhanced through the involvement of the TGFβ and Wnt pathways. However, the mechanism by which vitamin D produces calcification in VICs remains unknown.  In this study we also demonstrated in vivo for the first time that levels of VDR expression in human aortic valve tissue increase with increasing degrees of calcification. An earlier clinical study determined which polymorphism of VDR is most abundant in patients with aortic valve calcification [273].We have shown here that the level of VDR expression is up-regulated with increases in the degree of valve calcification in vivo (Figure 22). In the clinical study Greenwood et al. demonstrated that the polymorphism of VDR is derived from expression of the gene with the B allele and is more common in patients with AVC [273]. In our in vitro study we have demonstrated that VDR protein expression also occurs in valvular interstitial cells and that the VDR expression level also increases with vitamin D3 treatment in a dose dependent manner (Figure 23). VDR is a member of nuclear receptor (NR) family and acts as a transcription factor that can regulate the expression of its ligand (vitamin D) gene products [274]. In fact vitamin D binding to VDR can regulate as many as two hundred genes directly or indirectly through 105  vitamin D-VDR dependent pathways [260, 263]. These observations suggested that vitamin D could play an important role in valvular calcification along with numerous different roles in addition to its role in calcium regulation. For example others have shown that VDR-deficient mice develop colon cancer by enhancement of the WNT/β-catenin pathway [268, 275].  In the process of culturing and treating VIC with vitamin D3 we detected interesting changes in morphology that suggested treatment with vitamin D produced a phenotype shift in VIC. Large stellate shaped cells of the control group were transformed into elongated spindle shapes in TGFβ1 treatment which was also accompanied by an increase in αSMA expression consistent with a myofibroblast phenotype. However with vitamin D treatment the stellate shaped VIC changed to a polygonal morphology, accompanied by a decline in αSMA expression. This is consistent with a phenotype shift away from that of a myofibroblast [270, 271, 276] (Figure 23). Valvular interstitial cells are the most abundant cells in aortic valves and exhibit characteristics of myofibroblasts and in fact play an important role in valvular calcification disease [274]. Myofibroblasts are characterized by the presence of smooth muscle cell specific contractile proteins, such as smooth muscle-myosin heavy chains or desmin [157]; however, the presence of αSMA is the most reliable marker of the myofibroblastic phenotype [277]. In our in vitro model the control group expressed αSMA. After 96 hours treatment with TGFβ1 the αSMA expression increases. This indicates that VIC in control and the TGFβ treated group both exhibit a myofibroblast phenotype. However, when cells are treated with vitamin D after 96 hours the αSMA expression decreased significantly. Shape changes and down regulation of αSMA are both consistent with a cell phenotype shifting away from that of a myofibroblast. Simultaneous treatment of VIC with vitamin D and TGFβ significantly lowered levels of αSMA 106  and produced a mixture cell shapes perhaps because the cells did not respond uniformly to TGFβ and vitamin D treatment. The cells responsive to vitamin D decrease the level of αSMA of the cells [260, 278]. To determine the relevance of the above change we examined the expression of osteoblast specific markers in our VIC treated groups. Expression of osteopontin and RUNX2, bone specific markers, significantly increased in VIC treated with vitamin D. These data along with the cell shape changes mentioned above suggest that cell phenotype is in fact shifting to that of osteoblast-like cells. When vitamin D was added along with WNT3A and TGFβ1, the expression of osteopontin and RUNX2 were increased in an apparently additive manner (Figure 25). To the best of our knowledge we are the first to report an up-regulation of bone specific markers in VIC treated with vitamin D. Earlier studies of osteoblast differentiation and maturation in bone also showed that vitamin D along with the activated WNT3A pathway can induce osteoblast differentiation [249, 279].  In accord with our osteopontin and RUNX2 data, levels of alkaline phosphatase activity and cell calcification also increased significantly with combined vitamin D3, TGFβ1 and WNT3A treatment. This observation is also consistent with there having been a transformation of VIC to an osteoblast-like phenotype (Figure 24). Alkaline phosphatase is an early indicator of calcification [207, 280] which was increased in the VIC treatment with vitamin D alone and with TGFβ1 and WNT3A together. All these data strongly suggest that vitamin D, TGFβ1 and WNT3A transform VIC to an osteoblast phenotype that actually produce calcification as seen in AVC. 107  From our study of TGFβ and WNT3A in VIC calcification one might suggest that the phosphorylation of SMAD2, the downstream element of TGFβ and AXIN2 and negative regulator of WNT/β-catenin pathway, might be considered as a potential mechanisms by which vitamin D3 contributes to the induction of VIC calcification [281-284]. We observed no changes in pSMAD2 and AXIN2 expression in VIC treated with vitamin D3, suggesting that vitamin D3 may not interact with the TGFβ and WNT/β-catenin pathways as we hypothesized in the calcification of VIC. The significantly increased VIC calcification observed here in VIC treated with TGFβ, WNT3A and vitamin D3 (Figure 25 A and B) therefore result from an apparent additive effect of vitamin D to our documented calcification by activation of TGFβ and WNT3A pathways in simply calcifying the cells by its known effect on calcium regulation in cultured VIC [187, 285-287]. Based on previous studies on the inhibitory role of vitamin D on the Wnt pathway we anticipated that vitamin D might increase expression of AXIN2 protein. Previously it has been shown that activation of the VDR in the presence of its metabolite ligand, 1α, 25(OH)2 vitamin D3, can suppress the WNT/β-catenin signaling and in some instances can promote malignant cell differentiation [288], and decrease oncogenic cell signaling [274, 279]. However in our VIC calcification model, treatment with vitamin D did not increase AXIN2 expression, suggesting that vitamin D does not affect the Wnt/β-catenin activation by mechanisms known to occur in colon cancer or osteoblast differentiation in regulating bone mass [266, 268, 279].  Another possible mechanism that might be involved in vitamin D mediated VIC calcification is its role in calcium and phosphate hemostasis [289, 290]. In our current study we observed no apparent interaction between the TGFβ, Wnt pathways and vitamin D in the calcification of VIC. Rather 108  this study confirms that the calcification regulatory agents, TGFβ1 and WNT3A independently of vitamin D, significantly increase VIC calcification in an additive fashion. More work is needed to identify the mechanisms by which these regulatory agents drive aortic valve calcification.  This study was observational in the nature. Further exploration is required to identify the possible mechanisms by which vitamin D may induce VIC calcification. We also propose to examine VIC calcification in various conditions where VDR is either over-expressed or inhibited. The data from such experiments may reveal whether vitamin D has a direct effect on VIC transformation from a myofibroblast to an osteoblast phenotype.          109  Chapter 5: OVERALL SYNOPSIS  5.1 Discussion Although once thought to be a passive process of calcium deposition on the cusps, aortic valve calcification (AVC) is now known to be an active process [40] that progresses through the transformation of valvular interstitial cells into osteoblast-like cells that calcify the fibrosa layer of the valve cusps. Several regulatory factors of inflammation have been identified as participants in the process of AVC. Among them the canonical Wnt and TGFβ pathways were shown to play major roles [149, 197, 243, 291]. A third participant that seems to most likely play a role in calcification of aortic valves was vitamin D3 [258].  In the present study, we examined the hypothesis that WNT3A, TGFβ1 and vitamin D3 are involved in the calcification of aortic valves by transforming VIC phenotype into that of osteoblast-like cells. We studied possible mechanisms by which the three candidate players may regulate VIC calcification. Positive scientific reception of AVC as an active inflammatory process has established more interest in studying valve biology and developing possible therapeutic interventions [292]. More studies have been focused on improving the understanding of the biology of those signaling pathways involved in inflammation and calcification processes as well as the biomechanics involved in initiation and progression of AVC. Animal models, specifically transgenic mouse models, have provided essential tools to help achieve this goal [293-295]. While human histopathological data have well demonstrated the inflammatory responses in 110  patients with AVC [296], the mouse models are harnessed to investigate the pathological mechanisms of the disease [188, 297].  Currently available mouse models are not ideal for comprehensive study of AVC, since their aortic valves lack the trilayer leaflet morphology seen in human valves and given their small size mouse valves are undesirable to fully test bio-mechanical impacts of disease [298]. In the current dissertation, to best study the molecular mechanisms of valvular calcification, an in vitro model was required to test our hypothesis that WNT3A and TGFβ1 are key players in the shifting phenotype of VIC from myofibroblast to that of osteoblast-like cells and the possibility that vitamin D is assisting in this process.  There are certain fundamental limitations in utilizing the VIC in vitro model to study valvular calcification. While VIC are the most abundant cells in valve tissue and most likely play critical roles in valvular diseases including calcification, maintaining  them in an artificial culture environment lacks a number of physiological influences. These properties such as blood flow and its hemodynamic effects greatly contribute to the pathophysiological conditions of valvular calcification. Further, the monolayer VIC culture system is representing only VIC- VIC interactions in a 2D system and lacks the interaction of VIC with the ECM as well as the 3D cell-cell communication as exists in the tissue. Additionally, the aortic valve is located in a highly dynamic mechanical environment where it is exposed to blood flow-induced shear stress, pressure loads and mechanical resistance from ECM. Each of these biomechanical stimuli regulates both homeostasis and disease state of the valve [299, 300]. An isolated VIC culture 111  model lacks the biomechanical properties of the native valve and may not integrate all the aspects of the valvular disease. In Chapter 2 we compared calcification in human primary vSMC and VIC in vitro. VIC seemed to calcify more rapidly and formed more calcium nodules compared to vSMC. In vivo, calcified VIC expressed significantly increased amounts of WNT3A and TGFβ1. These observations led us to further investigate the role of Wnt and TGFβ pathways in vitro using VIC (Chapter 3). TGFβ mediated DKK1 down-regulation has been demonstrated to activate the canonical Wnt pathway in liver and lung fibrosis [127]. These observations persuaded us to also examine the involvement of DKK1, the WNT3A antagonist, as a potential link between the canonical Wnt and TGFβ pathways and it seems to be the case. We have shown in VIC that WNT3A promotes calcification and that this process occurs through a β-catenin dependent mechanism. When LiCl, an inhibitor of the β-catenin degradation complex, was added to the process along with WNT3A as the positive control, it enhanced the activation of the Wnt/β-cateniin pathway and subsequently the VIC calcification was significantly increased. We also tested DKK1 as a negative control and found it to inhibit the canonical Wnt pathway and VIC calcification. TGFβ also has been shown to be present in some diseased and calcified aortic valves and to play a role in calcifying porcine VIC [187]. In our study with human VIC adding TGFβ1 to the cultures also produced significantly calcification in vitro. Once TGFβ1 was added to VIC along with WNT3A, it significant enhanced both cell calcification and alkaline phosphatase activity as well as producing an increase in the expression of other osteoblast markers. Our data 112  supported an interaction between the Wnt3a and TGFβ pathway. TGFβ1-treated VIC had more β-catenin nuclear translocation; moreover, significantly lower amounts of DKK1 expression were identified in TGFβ1 treated cells. Less DKK1 expression results in allowing WNT3A signal to transmit into the cell and leads to the transcription of genes necessary for calcification.  Vitamin D plays an important role in the growth and integrity of the normal bone tissue in the body [301]. As it has also been demonstrated that vitamin D is involved in pathological calcification [17, 302, 303], we examined the role of vitamin D3 in aortic valve calcification. Thus it was worthwhile investigating whether vitamin D plays a role in the pathological calcification of VIC in addition to its role in normal bone formation. Having data that suggest that Wnt and TGFβ cooperate in valve calcification, we examined whether vitamin D and the canonical Wnt and TGFβ pathways are integrated in the process of VIC calcification. We confirmed that vitamin D induced VIC calcification in vitro (Chapter 4) as it does in normal in vivo calcification [304]. Addition of vitamin D to the VIC cultures increased cell calcification as well as alkaline phoaphatase activity. Vitamin D also increased the expression of other osteoblast markers in VIC. To determine whether the vitamin D effect on VIC calcification is through the canonical Wnt or TGFβ pathways we tested the expression of AXIN2 and pSMAD2, two key downstream components of the Wnt and TGFβ pathways respectively. No significant changes were observed in the expression of AXIN2 and pSMAD2 supporting that the vitamin D effect does not act through the Wnt or TGFβ pathways and suggesting that they do not interact. The mechanism by which vitamin D induces the transformation of VIC phenotype to that of osteoblast, remains unknown and requires further investigation. 113  5.2 Conclusion  Aortic valve calcification is a complex disease that is characterized by thickening via fibrosis and calcification of valvular cusps which eventually impairs the function of the aortic valves. This pathology is likely a consequence of the contribution of factors such as life style, environment and genetics [21, 305]. The process of AVC in some aspects resembles the development of atherosclerotic plaques.  In the present dissertation WNT3A and TGFβ1 and vitamin D3 were found to play major roles in VIC calcification and transformation when they act together. In human calcified aortic valve, we have shown that WNT3A, TGFβ1 and VDR were elevated and the levels of their expression were increased with the degree of calcification. In contrast, DKK1 expression was decreased in calcified valves. Consistent with our in vivo observations, we confirmed that WNT3A and TGFβ1 are present in calcified VIC and they work interactively toward calcifying valvular interstitial cells in vitro. We determined that TGFβ contribution to the calcification of VIC is perhaps via down-regulating DKK1, which allows WNT3A to act freely toward transforming VIC from myofibroblast phenotype into an osteoblast-like cell.  In the current study, we showed that vitamin D3 was able to enhance VIC calcification and cell transformation. While vitamin D3 induces cell calcification and has additive effects to WNT3A and TGFβ1 outcomes during the process of VIC calcification, the mechanism is not through the Wnt and TGFβ pathway. Vitamin D seems to act independently and through an unknown mechanism that requires more study.  114  In summary, through the work carried out in this dissertation, we have provided novel data suggesting that the integrated canonical Wnt and TGFβ signaling pathways play a key role in regulating VIC phenotype during valvular calcification that goes along with vitamin D in regulating VIC calcification (Figure 31)              115            Figure 31. Putative mechanism of action of signaling pathways involved in valvular calcification.  At the site of either biomechanical and/or chemical injury and secretion of series of inflammatory cytokines and growth factors the fibroblasts and myofibroblasts resident in the aortic valve tissue become activated and start to proliferate, synthesis of extracellular matrix component and secretion of proteins such as WNT3A and TGFβ that facilitate the transformation of myofibroblast phenotype into that of osteoblast. Upon the osteoblast-like cells formation alkaline phosphatase, osteopontin, RUNX2 and other bone specific markers increase and calcium start to deposit that leads to the calcification of the aortic valve tissue. There are exogenous factors such as high levels of vitamin D3 that can help to the process of pathological calcification.   116  5.3 Future Directions A major limitation to the current study is that majority of the key experiments have been performed in vitro in cultured cells. Thus, an important focus of future directions should aim to determine the extent to which these findings translate into the in vivo setting. In this regard, there are a number of animal models and different compounds available to activate or inhibit various aspects of the Wnt and TGFβ signaling pathways, in vivo. The animal studies need to be done under hyper-lipidemic environment. It has been demonstrated that aging LDL receptor knockout mouse models develop moderate aortic valve calcification with the physiological features similar to human. Administration of high carbohydrate diet to LDL receptor knockout mouse is more relevant to human that causes a decrease in aortic valve area, aortic valve leaflet thickening and calcium deposition in the valve. However, mouse aortic valves may not make an ideal study model as they lack the trilayer morphology of the human aortic valve [306]. Also mouse valves are not large enough to test biomechanical impacts on the valve structure and function. In regards to the VIC culture model, we suggest that for future studies the use of 3D culture systems wherein the valvular interstitial cells can interact with each other in three dimensional environment to mimic the VIC condition in the native valve tissue  [307]. The co-culture of VIC and endothelial cells may also be considered in the future experimental design [308]. Finally, the aortic valve disease cannot be reversed when it is diagnosed and valve replacement is the only therapeutic procedure available for these patients. It would be beneficial to understand the cell responses to the focal and regional biomechanical forces in the valve in order to improve the quality of the prostatic valves. We suggest that future in vitro studies should focus on designing models to examine the cell responses to the various mechanical flows present 117  in native valves [309]. Use of devices that can mimic the blood flow in a culture system along with osteogenic conditions would be beneficial to better understand the VIC responses to the chemical and biomechanical stimuli. The mechanisms behind vitamin D3 derived calcification also need to be carefully investigated. In this dissertation we have demonstrated that vitamin D3 acts independent of AXIN2 and pSMAD2. To further elucidate the mechanism of action of vitamin D3 in VIC calcification, we may study the possible interaction of VDR with downstream elements of the canonical Wnt pathway TCF and βcatenin. Further, in the TGFβ pathway, there is a potential role for P38 in interacting with vitamin D that is worthwhile more investigation. In the cultured VIC model studied in this thesis, we may also inhibit TGFβ receptor I to validate the role of DKK1 as a link between the TGFβ and the canonical Wnt pathways. Inhibition of TGFβ receptor I will result in the down-regulation of pSMAD2, which consequently may increase DKK1 expression. Over-expression of DKK1 is expected to then interact with LRP5/6 co-receptor and inhibit the canonical Wnt pathway to contribute to the VIC calcification. 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