Nfatc1 in Cardiac Valve Development and EPDC Invasion

University of Cincinnati

Date: 11/30/2010

I, Michelle D Combs , hereby submit this original work as part of the requirements for the degree of Doctor of Philosophy in Developmental Biology.

It is entitled: NFATc1 in cardiac valve development and EPDC invasion

Student's name: Michelle D Combs

This work and its defense approved by:

Committee chair: Katherine Yutzey

Committee member: Walter Keith Jones

Committee member: Jeff Molkentin

Committee member: James Wells

Committee member: Yi Zheng

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Last Printed:1/13/2011 Document Of Defense Form NFATc1 in cardiac valve development and EPDC invasion

A dissertation submitted to the Division of Graduate Studies and Research of the

University of Cincinnati

in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

in the Graduate Program in Molecular and Developmental Biology

of the College of Medicine

2010

Michelle D. Combs

Bachelor of Science, Quincy University, 1996

Committee Chair: Katherine E. Yutzey, Ph.D.

W. Keith Jones, Ph.D.

Jeffery D. Molkentin, Ph.D.

James M. Wells, Ph.D.

Yi Zheng, Ph.D. Abstract

Congenital malformations are the most common cause of death in infancy in the United

States. Of these birth defects, most are malformations of cardiac valvuloseptal structures and a significant number are coronary vessel malformations. Therefore, identifying molecular mechanisms that regulate cardiac valve and coronary artery development is of great clinical importance. Cardiac valve morphogenesis begins with growth of endocardial cushions in the atrioventricular canal and outflow tract regions of the looping heart. After growth, endocardial cushions are remodeled into thin leaflets, characteristic of mature heart valves. Nuclear Factor of Activated T-cells cytoplasmic 1

(NFATc1) is a transcription factor necessary for heart valve development. The studies detailed here demonstrate that Vascular Endothelial Growth Factor A (VEGF)/ NFATc1 pathway function promotes endocardial cushion growth, while Receptor Activator of

NFκB (RANKL)/ NFATc1 pathway function is associated with valve remodeling. These studies further demonstrate that NFATc1 serves as a nodal point in the transition from endocardial cushion growth to valve remodeling via ERK1/2 or JNK1/2 copathway activation.

In the course of these studies NFATc1 expression by PE, epicardium, and

EPDCs was discovered. During heart looping, PE cells migrate onto the myocardium and proliferate to form the epicardium. A subset of epicardial cells undergo epithelial-to- mesenchymal transformation (EMT) and invade the subepicardium and myocardium as epicardium-derived cells (EPDCs). EPDCs differentiate into coronary endothelial and

ii smooth muscle cells, as well as adventitial fibroblasts that produce the fibrous matrix.

Studies detailed in this dissertation demonstrate that conditional loss of NFATc1 expression in EPDCs in mice causes embryonic death by E18.5 with reduced coronary vessel and fibrous matrix penetration into myocardium. These studies further demonstrate that RANKL/NFATc1 pathway components are expressed in EPDCs and loss of NFATc1 in EPDCs causes loss of CtsK expression in the myocardial interstitium in vivo. Likewise, RANKL treatment induces CtsK expression in PE-derived cell cultures via a calcineurin-dependent mechanism. In chicken embryo hearts, RANKL treatment increases the distance of EPDC invasion into myocardium, and this response is calcineurin-dependent. Together, these data demonstrate a critical role for the RANKL/

NFATc1 signaling pathway in promoting invasion of EPDCs into myocardium by induction of extracellular matrix-degrading enzyme gene expression.

iii

Table of Contents

ABSTRACT ii

TABLE OF CONTENTS 1

LIST OF FIGURES AND TABLES 6

CHAPTER 1. Introduction

Overview 12

Overview of Heart Development 12

Valve Development 14

Coronary Vessel and Fibrous Matrix Development 19

NFATs 25

Calcineurin and NFATs in Heart Development 27

VEGF/ NFATc1 29

RANKL/ NFATc1 30

Experimental Rationale 33

References 36

Figure Legends 56

Figures 60

CHAPTER 2. VEGF and RANKL regulation of NFATc1 in heart valve development

Abstract 68

Non-Standard Abbreviations and Acronyms 68

Introduction 70

Materials and Methods 73

Results 74

Discussion 85

Acknowledgements 88

Sources of Funding 88

Disclosures 88

References 89

Figure Legends 95

Figures 99

Supplementary Methods 107

References 114

Supplemental Figure Legends 116

Supplemental Figures 119

CHAPTER 3. NFATc1 promotes epicardium-derived cell (EPDC) invasion into myocardium

Summary 125

Introduction 126

Materials and Methods 128

Results 135

Discussion 146

Acknowledgements 151

References 152

Figure Legends 160

Figures 165

Supplemental Figure Legends 172

Supplemental Figures 175

CHAPTER 4. Summary and Discussion

Major Findings 182

NFATc1 in Cardiac Valve Maturation 182

NFATc1 in EPDC Invasion 184

Experimental Limitations and Alternative Approaches 186

Calcineurin Inhibitors 191

A Broad View of NFATc1-regulated Mechanisms 192

References 194

Figure Legend 201

Figures 202

APPENDIX I. Heart valve development: regulatory networks in development and disease

Abstract 205

Introduction 206

Overview of Valve Development 206

Endocardial Cushion Formation and EMT 211

Growth of Endocardial Cushions and Valve Primordia 216

Diversification of Valve Cell Types 223

Heart Valve ECM Maturation and Organization 227

Heart Valve Development and Disease 230

Conclusions and Perspectives 232

Acknowledgements 233

Sources of Funding 234

Disclosures 234

References 235

Figure Legends 264

Tables and Figures 266

APPENDIX II. Transcriptional regulation of heart valve progenitor cells

Abstract 273

Introduction 274

Overview of Endocardial Cushion Formation and Valve Remodeling 274

Transcriptional Regulation of Valve Development 277

Twist1 278

Tbx20 279

Msx1 and Msx2 281

NFATc1 282

Sox9 284

Scleraxis 286

Overall Conclusions and Future Perspectives 287

References 289

Tables 303

LIST OF FIGURES AND TABLES

CHAPTER 1.

Figure 1. Graphical overview of heart development 60

Figure 2. Stratified ECM compartments are evident in mature semilunar and atrioventricular valves 61

Figure 3. Schematic representation of embryonic AV valve development 62

Figure 4. Schematic overview of coronary vessel and fibrous matrix development 63

Figure 5. Illustration depicting the structure of human NFATc1, NFATc2, NFATc3,

and NFATc4 mRNA transcripts 64

Figure 6. Model of VEGF/NFATc1 pathway signaling in human pulmonary valve

endothelial cells 65

Figure 7. Model of RANKL/NFATc1 pathway signaling in osteoclasts 66

CHAPTER 2.

Figure 1. E10.5 NFATc1-/- mouse ECC endothelial and mesenchymal cells exhibit

decreased proliferation 99

Figure 2. VEGF treatment of ECC cells induces NFATc1 nuclear localization 100

Figure 3. VEGF-induced proliferation of ECC cells is dependent upon calcineurin

signaling 101

Figure 4. NFATc1, RANKL, and CtsK mRNA expression in developing chick AVC 102

Figure 5. RANKL treatment of ECC cells induces NFATc1 nuclear localization and

increased CtsK expression via a Cn-dependent mechanism 103

Figure 6. Ligand-specific effects on ECC cell proliferation and CtsK expression.

RANKL inhibits VEGF-induced ECC cell proliferation 104

Figure 7. JNK1/2 activation is not seen in E11.5 mouse ECCs, but is

detected in E12.5 mitral and tricuspid valve endothelial cells in vivo 105

Figure 8. VEGF-induced ECC cell proliferation is MEK1-ERK1/2-dependent.

RANKL-induced CtsK expression and RANKL inhibition of

VEGF-induced cell proliferation is JNK1/2 dependent 106

Supplemental Figure 1. NFATc1-positive cells co-express endothelial markers,

and VEGF induces proliferation of endothelial cells in cultured ECCs. 119

Supplemental Figure 2. Expression of endothelial and mesenchymal markers by

cultured ECC cells over time mimics gene expression observed in maturing ECCs/

mitral valves in vivo 120

Supplemental Figure 3. OPG does not inhibit VEGF-induced cell proliferation

and sFlt1 does not inhibit RANKL-induced CtsK expression 121

Supplemental Figure 4. Percent diphosphorylated ERK1/2 positive cells is significantly reduced in cultures treated with U0126 compared to controls 122

Supplemental Figure 5. Model of NFATc1 function in the transition from

ECC growth to valve remodeling 123

CHAPTER 3.

Figure 1. NFATc1 is expressed by the PE and coronary vessels and colocalizes

with WT1 in the epicardium and EPDCs 165

Figure 2. WT1-Cre(+);NFATc1(fl/fl) embryos have reduced NFATc1 positive epicardial cells and EPDCs in addition to increased myocardial compaction 166

Figure 3. WT1-Cre(+);NFATc1(fl/fl) embryos lack interstitial fibrous matrix and have reduced penetration of Collagen1a1-expressing cells 167

Figure 4. WT1-Cre(+);NFATc1(fl/fl) embryos have reduced investment of activated fibroblasts, and reduced intramyocardial vessel penetration 168

Figure 5. RANKL/NFATc1 pathway components are expressed in mouse and chick embryos during EPDC invasion 169

Figure 6. WT1-Cre(+);NFATc1(fl/fl) embryos have reduced CtsK expression in the myocardial interstitium 170

Figure 7. RANKL increases EPDC migration distance and CtsK expression via a calcineurin/ NFAT-dependent mechanism 171

Figure S1. NFATc1 is expressed by PE, epicardium, EPDCs, and coronary vessels in chick demonstrating a conserved expression pattern in vertebrates 175

Figure S2. NFATc1 mRNA is expressed in epicardial cells and EPDCs of chick and mouse 176

Figure S3. An intact epicardium is apparent with systemic loss of NFATc1 177

Figure S4. Control and WT1-Cre(+);NFATc1(fl/fl) embryos are grossly indistinguishable until E17.5 when WT1-Cre(+);NFATc1(fl/fl) hearts show

signs of failure. WT1-Cre(+);NFATc1(fl/fl) hearts lack fibrillar collagen deposition

in myocardium and die at late embryonic stages 178

Figure S5. Coronary endothelial cell differentiation is apparent with loss of

NFATc1 expression in EPDCs 179

Figure S6. Cultured chick PE cells express EPDC markers 180

CHAPTER 4.

Figure 1. E10.5 NFATc1 -/- mouse epicardial cells exhibit decreased proliferation 202

Figure 2. Primary avian PE cells with nuclear NFATc1 coexpress CtsK in vitro 203

APPENDIX I

Table 1. Genetic lesions in ECM components of endocardial cushions/ valve primordia 266

Table 2. Genetic lesions in ECM components of cardiac valves 267

Figure 1. Stratified ECM compartments are evident in mature SL and AV valves 268

Figure 2. Model for regulatory interactions that control endocardial cushion formation (A) and EMT (B) 269

Figure 3. Model for regulatory interactions that control growth of endocardial cushions/ valve primordial 270

Figure 4. Model for regulatory interactions controlling AV valve stratification and lineage diversification 271

APPENDIX II

Table 1. Gene expression and associated human disease of transcription factors involved in heart valve progenitor cell development 303

Table 2. Transcription factors and their downstream targets involved in heart valve development 304

Chapter 1

Introduction

Overview

Cardiac abnormalities are the most common birth defects, occurring at a rate of 1 in every 100 live births (Lloyd-Jones et al., 2010). Malformations in cardiac valvuloseptal structures necessitate the most invasive surgeries during the first year of life in the

United States and are associated with increased risk of valve disease in adults (Lloyd-

Jones et al., 2010). Coronary artery malformations occur in 1.3% of the population and can lead to myocardial infarction, arrhythmia, or sudden death (Kayalar et al., 2009).

The coronary vessels and cardiac fibrous matrix are epicardium-derived structures

(Reese et al., 2002). For these reasons, better understanding of molecular mechanisms that govern cardiac valve and epicardium-derived structure development may have far- reaching consequences in improving public health. Previous work demonstrated that the transcription factor Nuclear Factor of Activated T-cells cytoplasmic 1 (NFATc1) is necessary for cardiac valve remodeling during development. In this dissertation,

NFATc1 is identified as a nodal point in the transition from endocardial cushion growth via Vascular Endothelial Growth Factor A (VEGF)/ NFATc1 pathway function to valve remodeling via Receptor Activator of NFκB Ligand (RANKL)/NFATc1 signaling. In addition a novel role for NFATc1 in EPDC invasion into myocardium is demonstrated.

Overview of heart development

“You change your life by changing your heart.” - Anonymous

During gastrulation, cardiac progenitor cells occupy the anterior half of the primitive streak and migrate in an anteriolateral fashion within the lateral plate mesoderm to form bilateral heart-forming fields (Nakajima et al., 2009; Snarr et al.,

2008). The lateral plate mesoderm splits into two definitive precursor pools, the somatic mesoderm that will contribute to the inner lining of the body wall, limbs, and dermis; and the splanchnic mesoderm that will form the heart myocardium and endocardium, the mesothelial lining of the organs, the proepicardium, and contributes to other organs such as the head musculature (Larsen, 2001; Gilbert, 2003; Lie-Venema et al., 2007;

Manner et al., 2001; Nathan et al., 2008). Within the splanchnic mesoderm cells of the heart-forming regions begin differentiating into endocardial and myocardial progenitors of the primary heart field as folding of the embryo brings these regions together into a cardiac crescent (Figure 1) (Snarr et al., 2008; Srivastava and Olson, 2000). Further embryonic folding brings the lateral regions of the cardiac crescent together where they fuse in a cranial to caudal fashion at the midline (Fishman and Chien, 1997). This fusion event forms the primitive heart tube consisting of an inner layer of endocardium and an outer layer of myocardium separated by an acellular extracellular matrix (ECM) called the cardiac jelly (Snarr et al., 2008). Just after formation, the primitive heart tube begins to beat (Nakajima et al., 2009). This heartbeat, which begins so early in development, must continue unceasingly throughout the life of the organism.

The heart tube when formed, is attached along the dorsal axis to the dorsal mesocardium of the foregut (Larsen, 2001; Snarr et al., 2008). This tissue is resorbed at all but the cranial-most and caudal-most aspects to leave the heart tube suspended in

the pericardial cavity. At this point, the heart tube is arranged with the primitive outflow tract (bulbus cordis/ conotruncus) and ventricle in a superior position to the primitive atria and inflow tract (sinus venosus) (Larsen, 2001). Cell migration into the heart from the outflow and inflow tract regions by cells of the second heart field and high rates of proliferation cause the heart tube to elongate and loop simultaneously (Dyer and Kirby,

2009; Larsen, 2001). The second heart field is a group of cells derived from splanchnic mesoderm and situated at the medial and caudal most aspects of the bilateral heart- forming regions (Dyer and Kirby, 2009). These cells migrate into the heart tube and contribute to formation of the outflow tract, right ventricle, interventricular septum, small portions of the left ventricle, atria, and sinus venosus (Black, 2007). Cardiac looping continues until the presumptive atria are positioned atop the ventricles forming the basic four-chambered heart structure (Larsen, 2001). Further cardiac development consists mainly of maturation of valvuloseptal structures and formation of coronary vessels and fibrous matrix by epicardium-derived cells (EPDCs).

Valve development

The 4-chambered vertebrate heart has aortic and pulmonic semilunar (SL) valves at the arterial pole as well as mitral and tricuspid valves separating the atria and ventricles. The coordinated opening and closing of the heart valves occurs approximately three billion times in an average human lifespan and is required for unidirectional blood flow (Schoen, 2008). The three cusps of each SL valve and the 2

(mitral) or 3 (tricuspid) leaflets of the atrioventricular (AV) valves consist of complex

stratified connective tissue (Rabkin-Aikawa et al., 2005; Schoen, 2008). The valve leaflets are ensheathed in endocardial endothelial cells with intervening valve interstitial cells (VIC) that function in homeostasis and disease (Hinton et al., 2006; Rabkin-Aikawa et al., 2004). The valves are stratified into extracellular matrix (ECM) layers rich in elastin (ventricularis of SL/atrialis of AV), proteoglycan (spongiosa) and collagen

(fibrosa), oriented relative to blood flow (Figure 2) (Hinton et al., 2006; Combs and

Yutzey, 2009a). The most obvious difference between the AV and SL valves is the presence of supporting chordae tendineae on the ventricular aspect of the tricuspid and mitral valves. However, comparable supporting connective tissue is present in the aortic and pulmonic roots and hinge regions of the SL valves (Hinton et al., 2008; Hinton et al.,

2006). Morphogenetic and structural differences also exist among the individual mural and septal AV valve leaflets, but, in general, the molecular mechanisms of valve development are conserved among AV and SL valve leaflets. Extensive conservation of valve developmental mechanisms also has been observed among vertebrate species including chicken, mouse, and human.

The first evidence of valve development is swellings called endocardial cushions that appear in the atrioventricular canal (AVC) and outflow tract (OFT) regions of the looping heart (embryonic day [E]3 chick, E9.5 mouse, E31 to E35 human) (Figure 3)

(Wessels and Sedmera, 2003; Fishman and Chien, 1997; Martinsen, 2005; Moorman et al., 2003). Endocardial cushion formation is induced by myocardial production of signaling molecules, such as BMP2 and BMP4, that inhibit expression of chamber- specific genes in the AVC and OFT, while increasing synthesis of ECM components

(Appendix I Figure 2A) (Harrelson et al., 2004; Lyons et al., 1990; Ma et al., 2005;

Plageman and Yutzey, 2004; Combs and Yutzey, 2009a). This increased ECM or

“cardiac jelly” deposition between the myocardium and endocardium, along with the

hydrophilic nature of the ECM proteoglycans, causes the tissue to protrude or swell into

the interior lumen of the heart forming the endocardial cushions (Camenisch et al.,

2000; Henderson and Copp, 1998; Markwald et al., 1975). Even at this early stage,

endocardial cushions act as physical barriers that prevent the backflow of blood through

the primitive heart tube (Schroeder et al., 2003). Signaling molecules originating from

both the myocardium and endocardium of the AVC and OFT are necessary for proper

endocardial cushion formation and epithelial-to-mesenchymal transformation (EMT) of

endocardial endothelial cells (Krug et al., 1985). EMT occurs as a subset of endocardial

cushion endothelial cells break connections with neighboring cells and migrate into the

cardiac jelly to populate the endocardial cushions with mesenchymal cells. Proliferation

and maintenance of the valve endothelial layer is dependent upon VEGF, while Notch1,

TGFβ, and Wnt/β-catenin signaling promote endothelial cell delamination and EMT

(Appendix I Figure 2B)(Combs and Yutzey, 2009a).

After EMT, the endocardial cushions and subsequent valve primordia undergo growth via cell proliferation and continued ECM synthesis (Armstrong and Bischoff,

2004; Hinton et al., 2006; Martinsen, 2005). Previous studies demonstrate that multiple signaling mechanisms regulate proliferation of VIC including Wnt/ β-catenin, TGFβs,

BMPs, FGF4, Shp2/ ERK1/2, and EGFs (Appendix I Figure 3)(Combs and Yutzey,

2009a). Work detailed in this dissertation demonstrates that VEGF/ NFATc1 signaling

promotes proliferation of valve endothelial cells during endocardial cushion growth. The

AVC valve primordia are part of a larger mass of tissue called the septum intermedium that is formed via fusion of the endocardial cushions at E4.5 in chicks, E11.5 in mice, and E37 to E42 in humans (Martinsen, 2005; Moorman et al., 2003; Webb et al., 1998;

Wessels and Sedmera, 2003). Septum intermedium tissue contributes to the membranous ventricular septum and fibrous continuity overlying the ventricular septum adjacent to the valve primordia that form the septal tricuspid and mitral valve leaflets

(Martinsen, 2005). The OFT endocardial cushions also fuse and contribute to the formation of the aortic and pulmonary valve leaflets and supporting structures (Qayyum et al., 2001).

During fetal stages of the chicken (E14), mouse (E16.5 to 17.5), and human (20 to

39 weeks gestation), the valve primordia are remodeled into thin valve leaflets. Studies detailed in this thesis demonstrate that RANKL/ NFATc1 signaling in valve endothelial cells inhibits VEGF-induced cell proliferation and induces expression of ECM remodeling enzymes associated with valve maturation. The characteristic trilaminar architecture of mature valve ECM begins to emerge during remodeling and is associated with differential gene expression by VIC. Notch1 and elastin are expressed by VIC on the surface of the valve exposed to unidirectional pulsatile blood flow, while

Wnt ligands induce periostin expression on the side of the valve away from flow (Figure

2 and Appendix I Figure 4) (Combs and Yutzey, 2009a). Fibrillar collagen deposition is also apparent in the fibrosa layer away from blood flow (Aikawa et al., 2006; Hinton et al., 2006; Kruithof et al., 2007). Additional specialized ECM compartments are the

proteoglycan-rich spongiosa layer, characterized by BMP induced aggrecan expression, as well as the tenascin-rich chordae tendineae and supporting structures (Lincoln et al.,

2004, 2006; Combs and Yutzey, 2009a). Together, these ECM compartments are required for normal valve structure and function, with dysregulation leading to disease.

The emerging evidence for activation of valve developmental pathways during adult valve disease pathogenesis has potentially important implications in the treatment of human cardiovascular disease (Rabkin et al., 2001; Rabkin-Aikawa et al., 2004;

Rabkin-Aikawa et al., 2005; Rajamannan et al., 2003). In the normal adult valve, the valve endothelial cells and VIC are relatively quiescent with little or no synthetic activity or cell proliferation (Aikawa et al., 2006; Hinton et al., 2006). The most common types of valve disease are myxomatous, characterized by insufficiency and inappropriate ECM production, and stenotic, with leaflets that are thickened, stiff and mineralized (Freeman and Otto, 2005; Rabkin-Aikawa et al., 2005). Increased cell proliferation and activation of VIC with increased synthetic activity is associated with both types of valve pathogenesis (Aikawa et al., 2007; Hinton et al., 2006; Rabkin-Aikawa et al., 2004;

Wirrig and Yutzey, 2010; Cole et al., 1984). It is not known whether VIC that express genes related to valve development represent a dedifferentiated cell type or whether there is a relatively undifferentiated cell population in normal adult valves. Recent studies have begun to define distinct types of VIC, that may have specific roles in valve pathogenesis, and these may be related to diversified cell types seen during development (Liu et al., 2007). Alternatively, cells from extracardiac origins, such as mesenchymal or hematopoietic stem cells, may populate the adult valves and could

contribute to disease pathology or have valve regenerative potential (Visconti et al.,

2006). There is initial evidence that the increased ECM production and VIC activation/ proliferation in valve pathogenesis is related to developmental pathways (Aikawa et al.,

2007; Garg et al., 2005; Paruchuri et al., 2006; Rajamannan et al., 2005; Chakraborty et al., 2010; Wirrig and Yutzey, 2010). Further studies are necessary to rigorously test this hypothesis and develop therapeutic strategies that prevent inappropriate activation of developmental pathways by valve endothelial cells and VIC.

Currently, a valve stem cell population has not been identified, however, the reactivation of developmental mechanisms in adult valve endothelial cells and VIC suggests that these cells may have regenerative potential. The detailed analysis of regulatory pathways that control valve development also has implications in valve tissue engineering. In general, efforts directed toward generating engineered valves do not take into account the diversity of VIC or their abilities to generate ECM with distinct structural characteristics (Schoen, 2005). The application of recent research into valve developmental mechanisms to the generation of engineered valves will likely improve the long-term function of these tissue constructs and could lead to improved therapeutics or replacement strategies. Likewise, manipulation of known valve developmental mechanisms could be applied to the treatment and management of the most common types of valve disease.

Coronary vessel and fibrous matrix development

The coronary vessels supply blood to the heart itself, therefore, every

cardiomyocyte must be in contact with a coronary vessel for nutrients and gas exchange

(Reese et al., 2002). The structural anatomy of coronary vessels does not differ from that of other blood vessels throughout the body. Coronary arteries and veins consist of a tunica intima, tunica media and tunica adventitia (Forbes, 2001). The tunica intima forms the innermost vessel lining and is comprised of a single layer of squamous endothelial cells bounded by connective ECM and internal elastic membrane. The tunica media overlays the tunica intima and is composed of vascular smooth muscle cells and elastic fibers, while the outermost layer, the tunica adventitia, is composed mainly of an external elastic membrane and collagen-rich ECM (Forbes, 2001). Arteries branch into arterioles and then branch further into capillaries to deliver oxygenated blood to the heart muscle. Arterial capillaries interface with venous capillaries which communicate to larger venules and finally veins to carry deoxygenated blood back into the right atrium and out to the lungs for reoxygenation (Forbes, 2001). Two exceptions to this are the pulmonary arteries, which carry deoxygenated blood to the lungs and the pulmonary veins that carry oxygenated blood from the lungs to the left atrium (Forbes,

2001).

The cardiac fibrous matrix is a collagen-rich ECM scaffold required for proper cardiac function (Lunkenheimer et al., 2006). The netted arrangement of cardiomyocytes within the fibrous matrix has been compared anatomically to the structure of the tongue where each cardiomyocyte is in contact with one or more of its neighbors in a 3-dimensional architecture (Lunkenheimer et al., 2006). The fibrous matrix serves as an anchor point to which cardiomyocytes can establish strong

connections through specialized structures called costameres (Samarel, 2005). The fibrous matrix is also “stress tolerant” in that it is able to be deformed by cardiomyocyte contraction and transmit this force to neighboring cardiomyocytes as part of the cardiac cycle (Samarel, 2005). The physical demands placed on the network of fibrous matrix and cardiomyocytes that make up the ventricles require that it have three characteristics necessary for cardiac function (Lunkenheimer et al., 2006). Firstly the network must be

“self-bracing” so that fixed points may be established in the network around which other cells can contract. Second, the network is “self-restraining” so that there are set limits to the amount of deformation that can occur in any one area. Lastly, the network must retain its proper shape throughout the cardiac cycle and in this manner properly regulate blood flow (Lunkenheimer et al., 2006).

Development of the coronary vasculature and fibrous matrix begins with formation of the proepicardium (PE/ PEO) at E9.25 in mouse, HH14 in chick and 24 dpc in humans (Figure 4) (Manner et al., 2001; Reese et al., 2002; Manner and Ruiz-Lozano,

2008). The PE is an outgrowth of the pericardial mesothelium covering the septum transversum and/or sinus venosus region of the developing embryo (Lie-Venema et al.,

2007; Manner et al., 2001). The inductive signals that initiate PE anlagen formation are not well characterized, however, studies indicate that BMP4 expression in the lateral plate mesoderm upregulates GATA4 transcription in the mesenchyme surrounding the septum transversum and liver bud (Rojas et al., 2005). Loss of GATA4 expression in these cells prevents PE outgrowth and expression of PE marker genes such as Tbx18 and Wilmʼs Tumor 1 (WT1) (Watt et al., 2004). WT1 expression by PE cells is required

for PE development; however, loss of Tbx18 expression in these cells does not affect

epicardium formation (Schlueter et al., 2006). Studies in chick indicate that BMP2 and

BMP4 expression by sinus venosus myocardium and PE anlagen respectively are

required for maintenance of PE marker gene expression, while FGF family members

regulate PE cell proliferation and PE villi outgrowth (Schlueter et al., 2006; Torlopp et

al., 2010).

The epicardium is formed when PE cells migrate onto the looping heart and

proliferate to envelop the myocardium (Figure 4). Migration of PE cells onto the heart

occurs by three main mechanisms and differs by species (Manner and Ruiz-Lozano,

2008). In organisms with an enclosed pericardial space, such as mice and dogfish, PE

villi bud off and form vesicles that float freely through the pericardial fluid and onto the

myocardium (Manner and Ruiz-Lozano, 2008). In organisms with an open pericardial

space, such as avians and amphibians, PE cells migrate onto the heart via an ECM

bridge that forms at the dorsal side of the AVC (Manner and Ruiz-Lozano, 2008). The

third method for PE migration occurs in all species and consists of direct migration of PE

cells over the sinus horns and onto the myocardium (Manner and Ruiz-Lozano, 2008).

Epicardium formation begins at the dorsal aspect of the heart near the AVC and

proceeds ventrally and cranio-caudally until the entire myocardium is covered by an

epithelial epicardial sheet by E10.5 in mouse and HH21 in avians (Dong et al., 2008).

Studies in mice indicate that multiple factors are important for PE cell migration (TGFβ1-

3, retinoic acid, BMPs, Angiopoietin1, Erythropoietin) and adhesion (α4-integrin,

VCAM1, Connexin 43) during epicardium formation (Olivey and Svensson, 2010). As

epicardium cells contact the myocardium, an ECM is secreted between the epicardium and myocardium. This ECM-filled compartment is called the subepicardial space

(Manner and Ruiz-Lozano, 2008).

Epicardium-derived cells (EPDCs) are the major source of cells for formation of the coronary vessels and fibrous matrix (Lie-Venema et al., 2007; Reese et al., 2002). The epicardium forms as a polarized epithelium and epicardial cells maintain a high mitotic index from E10.5-E11.5 in mouse (Wu et al., 2010). This allows for parallel cell division in which both cells remain in the epicardium as well as perpendicular cell division where one cell remains in the epicardium and the sister cell delaminates from the epicardium and enters the subepicardial space. Cells that delaminate from the epicardium undergo

EMT and become mesenchymal EPDCs (Figure 4) (Reese et al., 2002; Manner and

Ruiz-Lozano, 2008). EMT that occurs to form EPDCs is mechanistically comparable to

EMT events that occur in other tissues such as endocardial cushions during development (Olivey and Svensson, 2010; Reese et al., 2002; Manner et al., 2001).

Studies in mouse and chick have determined that FGFs, VEGF, and EGFs promote epicardial EMT, however, the role of TGFβ family members in this process is not yet fully understood (Manner and Ruiz-Lozano, 2008; Olivey and Svensson, 2010). Once formed, EPDCs invade the subepicardial space and myocardium where they will eventually differentiate into cells of the coronary vasculature and interstitial fibroblasts

(Figure 4) (Reese et al., 2002; Manner and Ruiz-Lozano, 2008).

Coronary vasculogenesis begins as angioblasts from the proepicardial villi and

EPDCs invade the subepicardial space and myocardium (Majesky, 2004). Studies

highlighted in this dissertation indicate that RANKL/ NFATc1 signaling is important for

ECM degradation and EPDC invasion during this process. Beginning in the subepicardial space, angioblasts and EPDCs differentiate into endothelial cells that then assemble into tubules (Tomanek, 2005; Zeini et al., 2009; Reese et al., 2002; Majesky,

2004). Studies in chick and mouse indicate that VEGF is an important regulator of this process (Majesky, 2004). Vasculogenesis continues until the exterior of the heart is covered by a coronary vascular plexus (Zeini et al., 2009; Majesky, 2004). Coronary plexus capillaries invade the aortic root beginning at E13.5 in mouse, HH32 in chick, and 32-38 dpc in humans to establish blood flow through the forming vessels (Dong et al., 2008; Tomanek, 2005). This initiates further angiogenesis and vascular pruning throughout the forming vascular tree in the subepicardium and myocardium (Dong et al.,

2008; Majesky, 2004). Concurrently, EPDCs are recruited to reinforce the forming vessels with coronary smooth muscle cells (Majesky, 2004). Smooth muscle cell recruitment is dependent upon PDGF-B production by coronary endothelial cells (Dong et al., 2008). Coronary vessel growth and maturation proceeds throughout gestation and into the postnatal period (Tomanek, 2005; Dong et al., 2008).

Interstitial (adventitial) fibroblasts comprise as much as 70% of the cells of the ventricular wall (Lie-Venema et al., 2007). Histological studies in quail-chick chimera embryos indicate that the majority of interstitial fibroblasts are of epicardial origin, however, further study of interstitial fibroblast patterning and differentiation has been hindered by lack of immunohistochemical markers (Wessels and Perez-Pomares, 2004;

Lie-Venema et al., 2007). EPDCs that will differentiate into interstitial fibroblasts enter

the subendocardium and myocardium with EPDCs of the coronary vasculature

(Wessels and Perez-Pomares, 2004; Lie-Venema et al., 2007). Research detailed in

this thesis indicates that pre-interstitial fibroblast EPDCs, like pre-vascular EPDCs,

require RANKL/ NFATc1 signaling for invasion into myocardium (Reese et al., 2002;

Wessels and Perez-Pomares, 2004; Lie-Venema et al., 2007). Once in place interstitial

fibroblasts secrete ECM components of the cardiac fibrous matrix required for proper

heart function (Lie-Venema et al., 2007; Lunkenheimer et al., 2006).

NFATs

The Nuclear Factor of Activated T-cells (NFAT) family of transcription factors has five

members NFATc1 (NFAT2/ NFATc), NFATc2 (NFAT1, NFATp), NFATc3 (NFAT4,

NFATx), NFATc4 (NFAT3), and NFAT5 (TonEBP) each characterized by a Rel

homology DNA binding domain (Aramburu et al., 2006; Hogan et al., 2003; Jin et al.,

2003). NFAT factors are evolutionarily related to NFκB proteins, the link between them being represented by NFAT5 (Serfling et al., 2004). This factor is found in Drosophila and humans, where it functions in regulating cellular response to hypertonic stress

(Hogan et al., 2003; Serfling et al., 2004). Unlike NFAT5, NFATc1-c4 activation and nuclear localization is controlled by Ca2+ and the calcium-responsive phosphatase

calcineurin (Hogan et al., 2003; Vihma et al., 2008). The calcineurin-regulated NFATs

contain an N-terminal transactivation domain and regulatory region (Figure 5) (Hogan et

al., 2003; Serfling et al., 2004). The regulatory region is a conserved ~300 amino acid

sequence with four types of serine-rich sequence motifs (Hogan et al., 2003).

Calcineurin binds to NFATs and dephosphorylates three of the four serine-rich sequence motif types thereby causing a conformational change in NFAT structure that unmasks a nuclear localization sequence, also located in the regulatory region (Figure

5) (Hogan et al., 2003; Okamura et al., 2004). Though not obligate dimers, intranuclear

NFATs commonly homodimderize and heterodimerize with other NFATs or unrelated proteins, such as AP-1 and GATA factors, to promote DNA binding and activate target gene transcription (Molkentin et al., 1998; Rao et al., 1997). NFAT nuclear localization is opposed by NFAT export kinases. GSK3, CK1, and DYRK1A have been identified as constitutive NFAT export kinases; however, it is likely that other NFAT export kinases remain to be identified (Hogan et al., 2003; Macian, 2005; Okamura et al., 2004; Arron et al., 2006). Cytosolic NFATs are phosphorylated by maintenance kinases from the

Mitogen Activated Protein Kinase (MAPK) family to inhibit nuclear translocation

(Macian, 2005).

Activation of NFAT proteins occurs via the calcium-calmodulin-activated protein phosphatase calcineurin (PP2B) (Molkentin, 2004; Hogan et al., 2003). The calcineurin molecule is composed of three proteins: the catalytic subunit calcineurin A, and two calcium-binding regulatory subunits calcineurin B and calmodulin (Molkentin, 2004).

Ca2+ influx into the cell cytosol triggers Ca2+ binding to calmodulin and calcineurin B to activate calcineurin (Molkentin, 2004; Crabtree, 1999). Calcineurin in turn activates

NFATs inducing nuclear translocation. Multiple endogenous and pharmacological inhibitors antagonize calcineurin/ NFAT activation. Calcineurin-binding protein 1

(CABIN1/ CAIN), A-kinase anchor protein (AKAP79/ AKAP5), calcineurin homologous

protein (CHP) and the calcipressins (MCIP1-4) are endogenous inhibitors of calcineurin activation (Macian, 2005; Rothermel et al., 2003; Nilsson et al., 2008). DSCR1 (MCIP1/

ADAPT78) is a direct downstream target of NFATc1 in endocardial cushion/ cardiac valve endothelial cells and along with DYRK1A has been implicated in the etiology of cardiac malformations associated with Downʼs syndrome (Lange et al., 2004; Wu et al.,

2007; Arron et al., 2006). Pharmaceutical inhibition of calcineurin is mediated by

Cyclosporin A and FK506 (tacrolimus), while more specific inhibition of calcineurin/

NFAT binding may be achieved via VIVIT peptide (Nilsson et al., 2008).

Calcineurin and NFATs in heart development

Multiple studies in chick and mouse demonstrate that calcineurin and NFAT function is not necessary for heart tube formation and cardiomyocyte differentiation (Schulz and

Yutzey, 2004). However, calcineurin/ NFAT activation are necessary for subsequent cardiac development. NFATc3 and NFATc4 are expressed by cardiomyocytes and mouse embryos lacking both NFATc3 and NFATc4 transcription die at E10.5 of cardiac failure due to metabolic and mitochondrial dysfunction in cardiomyocytes (Bushdid et al., 2003). Genetic deletion of NFATc3 or NFATc4 alone does not produce this phenotype, highlighting the potential for functional redundancy and compensation among NFAT factors (Bushdid et al., 2003). Because NFATs also have important roles in angiogenesis, NFATc3-/-; NFATc4-/- embryos also have peripheral vessel disorganization (Graef et al., 2001). Germline calcineurin B mutation in mouse embryos results in embryonic death at E10.0-E11 (Graef et al., 2001). These embryos exhibit

vascular patterning defects similar to those seen in NFATc3; NFATc4 double mutants

(Graef et al., 2001). Mice with a genetic deletion of NFATc2 are protected from calcineurin-induced pathological cardiac hypertrophy, however, cardiac development is apparently normal in these animals (Bourajjaj et al., 2008). NFATc2-/-; NFATc3-/-;

NFATc4-/- mice have inhibited endocardial cushion EMT suggesting that these NFATs function in the myocardium to promote EMT during valvulogenesis (Chang et al., 2004).

NFATc1 is a key regulator of immune, bone, blood vessel, cardiac valve and

EPDC-derived structure development (Combs and Yutzey, 2009b; de la Pompa et al.,

1998; Macian et al., 2000; Nilsson et al., 2008; Ranger et al., 1998; Combs submitted manuscript). NFATc1 has four isoforms, which occur as a result of variable splicing

(Figure 5)(Vihma et al., 2008). The NFATc1 gene is a direct downstream target of

NFATc1 itself enabling sustained activation via autoamplification (Asagiri et al., 2005).

In both mouse and chick embryos, cardiac NFATc1 is expressed by endocardial cushion endothelial cells and colocalizes with expression of its regulatory phosphatase calcineurin (de la Pompa et al., 1998; Liberatore and Yutzey, 2004). NFATc1 null mice have normal endocardial cushion formation, however, these cushions fail to elongate and remodel causing embryonic lethality at E12.5-E14.5 (de la Pompa et al., 1998;

Phoon et al., 2004; Ranger et al., 1998). Mice with a genetic deletion of calcineurin B specifically in endothelial cells phenocopy NFATc1-/- heart defects, suggesting that calcineurin/ NFATc1 interaction in endothelial cells is vital to endocardial cushion elongation and remodeling (Chang et al., 2004). Previous studies demonstrated that pharmaceutical inhibition of calcineurin activity by Cyclosporin A at E10.5 inhibits

coronary vascular plexus formation, however, the downstream targets of calcineurin necessary for this process were not identified (Zeini et al., 2009). It was also shown that Cyclosporin A treatment at E11.0, the initiation of endocardial cushion elongation and remodeling, prevented further valve development. The phenotype of these mice mirrored that of NFATc1-/- and calcineurin B endothelial nulls discussed previously

(Chang et al., 2004).

VEGF/ NFATc1

The human VEGF-A (hereafter referred to as VEGF) gene consists of 8 exons that are alternatively spliced to yield 5 common VEGF isoforms. These isoforms (121, 145, 165,

189, 206) function identically, but differ in their ECM binding and solubility. VEGF has the ability to bind VEGF receptors VEGFR-1(Flt-1), VEGFR-2 (Flk-2/KDR), Neuropilin-1 and Neuropilin-2, however, VEGFR-2 is the primary mediator of VEGF signaling and is expressed in endocardial cushion endothelial cells during elongation (Byrne et al., 2005;

Miquerol et al., 1999). VEGF is a potent endothelial cell-specific mitogen that must be tightly regulated for proper embryonic development and endocardial cushion formation.

Decreased VEGF signaling prior to endocardial cushion formation causes impaired endothelial cell proliferation and prevents endocardial cushion development (Armstrong and Bischoff, 2004; Stalmans et al., 2003). Conversely, threefold overexpression of

VEGF in the cardiovascular system prior to endocardial cushion formation causes hyperproliferation of endothelial cells and prevents EMT in the cushions (Miquerol et al.,

2000). The onset of endocardial cushion growth (E10.5) is marked by a sharp increase

in VEGF expression in the endothelial cells of the AVC and OFT regions (Miquerol et al., 1999). Work by Chang et al. demonstrates that calcineurin/ NFAT activation in the myocardium inhibits VEGF signaling to maintain a balance between cell proliferation and EMT during endocardial cushion growth (Chang et al., 2004). Experiments in adult human pulmonary valve endothelial cells show that VEGF treatment induces cell proliferation and migration via a calcineurin/ NFATc1 dependent mechanism (Figure 6)

(Jang et al., 2010; Johnson et al., 2003). Endocardial cushion endothelial cells also express Downʼs Syndrome Critical Region 1 (DSCR1). DSCR1 is a negative regulator of calcineurin activation and a downstream target of VEGF/ NFATc1 signaling in human umbilical vein endothelial cells. (Abe and Sato, 2001; Lange et al., 2004) Endocardial cushion endothelial cells of wild type mice express DSCR1 during endocardial cushion growth; however, DSCR1 expression is lost in endocardial cushion endothelial cells of

NFATc1-/- mice (Lange et al., 2004).

RANKL/ NFATc1

Receptor Activator of NFkB Ligand (RANKL/ TNF-related activation induced cytokine) is a TNF superfamily member that is essential for immune and skeletal development as well as homeostasis (Takayanagi, 2005). RANKL signaling is well studied in bone models where it stimulates osteoclast differentiation and function via a calcineurin/

NFATc1-dependent mechanism. Osteoclasts are vital to bone formation and homeostasis (Negishi-Koga and Takayanagi, 2009). These osteolytic cells degrade bone mineral and ECM for invasion of vascular cells and new bone formation (Karsenty

et al., 2009; Negishi-Koga and Takayanagi, 2009). Osteoclast differentiation begins

when RANKL, produced by osteoblasts, binds to its cognate receptor RANK on the

bone marrow-derived monocyte/ macrophage precursor cell (pre-osteoclast) surface

thereby activating TRAF6, c-Fos, and Ca2+ signaling pathways (Figure 7)(Takayanagi,

2005). TRAF6 recruitment initiates NFκB, Akt, and MAPK activation (Takayanagi,

2005). NFκB and c-Fos copathway signaling induces NFATc1 transcription in the pre- osteoclast cell (Sitara and Aliprantis, 2010). Ca2+ oscillations, needed to activate calcineurin, are induced via ITAM domain-harboring immunoreceptor stimulation of phospholipase Cγ (Takayanagi, 2005). Activation of NFATc1 by calcineurin is necessary and sufficient for osteoclast differentiation and function (Takayanagi et al.,

2002). NFATc1 induces transcription of a spectrum of genes required for osteoclast function and survival including NFATc1, Cathepsin K (CtsK), matrix metalloproteinase

(MMP) MMP9, and MMP14 while suppressing transcription of the osteoclast inhibitor osteoprotegrin (OPG) (Sitara and Aliprantis, 2010). Autoamplification of NFATc1 by binding its own promoter is seen in osteoclasts and endocardial cushion endothelial cells (Takayanagi, 2005; Combs and Yutzey, 2009b; Asagiri et al., 2005). This behavior is not exhibited by other NFAT family members and is thought to be essential for optimal

NFATc1 function (Takayanagi, 2005).

Use of calcineurin inhibitors, such as Cyclosporin A and FK506, to inhibit pathological bone resorption in humans can lead to further decreases in bone density

(Koga et al., 2005; Sitara and Aliprantis, 2010). This outcome was surprising until it was discovered that calcineurin/ NFAT function also regulates bone formation by osteoblasts

(Koga et al., 2005; Sitara and Aliprantis, 2010). NFATc1 regulates transcription of genes vital to osteoblast function including collagen I (Sitara and Aliprantis, 2010).

Expression of constitutively active NFATc1 in murine osteoblasts leads to an osteopetrotic phenotype with osteoblast overgrowth and hyperproliferation as well as increased osteoclast activity (Winslow et al., 2006). Together these studies demonstrate an important role for NFATs and particularly NFATc1 in bone growth and homeostasis.

Although traditionally associated with immune and skeletal function, emerging evidence suggests that calcineurin/ NFATc1 activation promotes ECM remodeling and cell migration/ invasion in diverse tissues. CtsK functions in ECM degradation for remodeling and cell invasion during the course of normal development; however, overactivation is also correlated with pathological conditions such as Cherubism, myxomatous cardiac valve disease, and cancer metastasis (Liu et al., 2003; Quintanilla-

Dieck et al., 2008; Rabkin et al., 2001). In the lung, NFATc1 and Ctsk are expressed by smooth muscle cells, which are responsible for physiologic and pathogenic ECM remodeling (Singh et al., 2010). Mice with genetic deletion of CtsK have impaired physiologic lung ECM remodeling and increased fibrosis after injury (Buhling et al.,

2004). Following balloon injury, lung smooth muscle cells migrate via an NFATc1- dependent mechanism and antagonism of NFATs by treatment with VIVIT peptide prevents airway ECM remodeling (Karpurapu et al., 2010). NFATc1 activation is also associated with increased breast cancer cell motility and metastasis (Seifert et al.,

2009). NFATc1-induced breast cancer cell invasion is blocked by treatment with

Cyclosporin A or Akt inhibition (Seifert et al., 2009; Yoeli-Lerner et al., 2005). Increased expression of the RANKL/ NFATc1 target CtsK is also correlated with an invasive phenotype in melanoma tumors (Quintanilla-Dieck et al., 2008). Together, these data suggest that RANKL/ NFATc1 signaling is a common mechanism for physiological and pathological ECM remodeling and cell invasion.

Experimental rationale

Cardiac valvulogenesis is a complex process that begins with the formation of endocardial cushions in the atrioventricular canal and outflow tract regions of the looping heart. The endocardial cushions are populated as overlying endothelial cells delaminate, undergo EMT, and invade the extracellular matrix ECM (Person et al.,

2005). During EMT, endocardial cushion endothelial cells are highly proliferative, and increased mitotic index is a feature of both endothelial and mesenchymal cells of the developing valves (Hinton et al., 2006; Person et al., 2005). After outgrowth, the ECM of the valves is remodeled into a highly organized, trilaminar architecture characteristic of mature cardiac valves. During ECM remodeling, valve interstitial cell proliferation is decreased and endothelial cells express ECM remodeling enzymes (Lange and Yutzey,

2006; Lincoln et al., 2004). It is clear that precise regulation of valve cell cycle, growth, and remodeling is required for normal valve development, as alterations in these processes are linked to valve defects (Armstrong and Bischoff, 2004; Chang et al.,

2004; Person et al., 2005).

Previous work demonstrated that NFATc1 is required for endocardial cushion elongation and remodeling (Ranger et al., 1998; de la Pompa et al., 1998).

Subsequently, Lange and Yutzey found that DSCR1 expression is lost in endocardial cushion endothelial cells of NFATc1-/- mice (Lange et al., 2004). As stated previously,

DSCR1 is a downstream target of VEGF/ NFATc1 signaling in human umbilical vein endothelial cells (Abe and Sato, 2001). It was known that VEGF is highly expressed in endocardial cushion endothelial cells during endocardial cushion growth and VEGF treatment is able to induce proliferation of adult pulmonary valve endothelial cells in vitro

(Miquerol et al., 1999, Johnson et al., 2003). These data led to the hypothesis that

VEGF/ NFATc1 signaling promotes endocardial cushion endothelial cell proliferation during endocardial cushion growth. However, it was not known if NFATc1-/- mice have decreased endocardial cushion cell proliferation or if VEGF can induce NFATc1 activation and promote cell proliferation in endocardial cushion cells during development.

Lange and Yutzey demonstrated that CtsK expression is lost in endocardial cushion endothelial cells of NFATc1-/- mice and implicated RANKL/ NFATc1 signaling in endocardial cushion remodeling (Lange and Yutzey, 2006). RANKL induces NFATc1 activation and calcineurin/ NFATc1-dependent ECM remodeling enzyme expression in osteoclasts. These data led to the hypothesis that NFATc1 may be part of a molecular switch that regulates the transition from endocardial cushion growth via VEGF/ NFATc1 signaling to endocardial cushion remodeling via RANKL/ NFATc1 signaling. It was not known if RANKL could induce NFATc1 activation and CtsK expression in endocardial

cushion cells. Importantly, if NFATc1 was found to regulate both endocardial cushion growth and remodeling, how then would NFATc1 target gene specificity be achieved?

These questions led to the studies detailed in Chapter 2 of this dissertation.

In the process of collecting data on NFATc1 regulation of endocardial cushion growth and remodeling, NFATc1 expression by PE, epicardium, and EPDCs was discovered. Cells of the PE and epicardium are highly proliferative during PE migration and epicardium formation (Wu et al, 2010). Proliferation of epicardium cells declines as

EPDCs undergo EMT and invade the subepicardial space and myocardium. At the same time during development, endocardial cushion endothelial cells undergo cell proliferation followed by EMT and endocardial cushion invasion. Expression of NFATc1 in both cell populations combined with the functional similarities in these processes led to the hypothesis that NFATc1 promotes epicardium cell proliferation during epicardium formation and regulates ECM remodeling enzyme expression for EPDC invasion into myocardium. The molecular mechanism for EPDC invasion into myocardium and the effects of loss of NFATc1 in epicardium and EPDCs during heart development had not been previously investigated. It is these questions, which are investigated in Chapter 3 of this dissertation.

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Figure legends

Figure 1. Graphical overview of heart development.

A) Within the splanchnic mesoderm cells of the heart-forming regions begin differentiating into endocardial and myocardial progenitors of the primary heart field as folding of the embryo brings these regions together into a cardiac crescent. B) Lateral regions of the cardiac crescent come together and fuse in a cranial to caudal fashion to create the primitive heart tube. The heart tube consists of an inner layer of endocardium and an outer layer of myocardium separated by an acellular extracellular matrix (ECM) called the cardiac jelly. At this point, the heart tube is arranged with the primitive outflow tract (bulbus cordis/ conotruncus) and ventricle in a cranial position to the primitive atria and inflow tract (sinus venosus). C) Cell migration into the heart from the outflow and inflow tract regions by cells of the second heart field and high rates of proliferation cause the heart tube to elongate and loop simultaneously. Valve development begins as endocardial cushions that will become the AV valves (AVV) form in the AVC and those that will become the semilunar valves form in the conotruncal (CT) region. D) Cardiac looping continues until the presumptive atria are positioned cranially to ventricles forming the basic four-chambered heart structure. Color coding represents the location of cells that contribute to major cardiac structures over time including neural crest contributions to the septum that divides aortic and pulmonary roots. Days indicate timing of events in human embryonic development. A = atrium, Ao = aorta, DA = ductus arteriosus, LA = left atrium, LCC = left common carotid, LSCA = left subclavian artery,

LV = left ventricle, PA = pulmonary artery, RA = right atrium, RCC = right common

carotid, RSCA =right subclavian artery, RV =right ventricle, V =ventricle; III, IV, VI = aortic arch arteries. Adapted from Srivastava and Olson (Srivastava and Olson, 2000).

Figure 2. Stratified ECM compartments are evident in mature semilunar and atrioventricular valves. A) Schematic representation of one of three valve cusps of the aortic or pulmonic semilunar valve with fibrosa (F), spongiosa (S) and ventricularis

(V) layers indicated. B) Schematic representation of one atrioventricular valve leaflet with atrialis (A), spongiosa (S), and fibrosa (F) layers indicated. The mitral valve has two leaflets, whereas the tricuspid valve has three leaflets, all of which are supported by chordae tendineae (CT). The direction of pulsatile blood flow is indicated for both semilunar and atrioventricular valves (arrow). (Combs and Yutzey, 2009a)

Figure 3. Schematic representation of embryonic AV valve development.

A) Endocardial cushions form as swellings of ECM in the AV region of the looping heart. B-C) Endocardial cushions undergo growth and remodeling. D) The mature valve consists of a thin valve leaflet (leaflet) connected to the ventricular papillary muscles (pap. muscle) by chordae tendineae (tendon). AV-myo = atrioventricular myocardium, atr = atria, epi = epicardium, vent = ventricle. Adapted from Wessels and

Sedmera (Wessels and Sedmera, 2003).

Figure 4. Schmatic overview of coronary vessel and fibrous matrix development.

A-B) The epicardium is formed when PEO/ PE cells migrate onto the looping heart and

proliferate to envelop the myocardium. C) ECM proteins are secreted into the subepicardial space between the epicardium and myocardium. D) Cells that delaminate from the epicardium undergo EMT and become mesenchymal EPDCs. E) Once formed, EPDCs invade the subepicardial space and myocardium. F-G) EPDCs differentiate into coronary smooth muscle cells, coronary endothelial cells and interstitial fibroblasts. Adapted from Reese et al. (Reese et al., 2002).

Figure 5. Illustration depicting the structure of human NFATc1, NFATc2, NFATc1, and NFATc4 mRNA transcripts. Exons are indicated with Roman numerals and depicted as boxes with intervening intronic sequences represented as lines. Intron size is indicated below each line. Codons are shown as colored boxes with the size of the related protein coding region specified above each exon. White exon regions represent untranslated sequence. Asterisks indicate rare variants. NES = nuclear export signal,

NLS = nuclear localization signal, DBD = DNA binding domain, RHD = Rel homology domain, Cn = calcineurin A. Adapted from Vihma et al. (Vihma et al., 2008).

Figure 6. Model of VEGF/NFATc1 pathway signaling in human pulmonary valve endothelial cells. In cardiac valve endothelial cells, VEGF activates calcineurin causing dephosphorylation of NFATc1. NFATc1 translocates to the cell nucleus where it binds DNA in complexes with other transcription factors such as AP-1 to promote cell proliferation. DSCR1 is an endogenous calcineurin inhibitor expressed by valve endothelial cells. Adapted from Armstrong and Bischoff (Armstrong and Bischoff,

2004).

Figure 7. Model of RANKL/NFATc1 signaling in osteoclasts.

Osteoclast differentiation begins when RANKL binds to RANK receptor on the bone

marrow-derived monocyte/ macrophage precursor cell (pre-osteoclast) surface thereby

activating TRAF6, c-Fos, and Ca2+ signaling pathways. NFκB and c-Fos copathway signaling induces NFATc1 transcription in the pre-osteoclast cell. NFATc1 undergoes autoamplification by binding to its own promoter to activate transcription. NFATc1 cooperates with multiple transcription factors to induce transcription of a spectrum of genes including CtsK, NFATc1, MMP9 and MMP14. Adapted from Takayanagi

(Takayanagi, 2005).

Chapter 2

VEGF and RANKL regulation of NFATc1 in heart valve development*

Michelle D. Combs, Katherine E. Yutzey

Division of Molecular Cardiovascular Biology

Cincinnati Childrenʼs Hospital Medical Center ML7020

240 Albert Sabin Way

Cincinnati, OH 45229

*Featured as the cover article in

Circulation Research 2009 September 11; 105(6): 565-574

Abstract

Rationale: Nuclear Factor of Activated T-cells cytoplasmic 1 (NFATc1) activity in

endocardial cushion (ECC) endothelial cells is required for normal ECC growth and

extracellular matrix (ECM) remodeling during heart valve development.

Objective: The mechanisms of NFATc1 activation and downstream effects on cell

proliferation and ECM remodeling enzyme gene expression were examined in NFATc1

mutant mice and chick ECC explants.

Methods and Results: NFATc1-/- mice display reduced proliferation of ECC endothelial and mesenchymal cells at embryonic day 10.5, while myocardial cells are unaffected.

Vascular Endothelial Growth Factor A (VEGF) activates NFATc1 and promotes ECC cell proliferation via the regulatory phosphatase, Calcineurin (Cn), and MAPK-ERK

Kinase 1 - Extracellular signal-Related Kinase 1/2 (MEK1-ERK1/2)-dependent signaling.

As ECCs mature, Receptor Activator of NFκB Ligand (RANKL) and the ECM remodeling enzyme, Cathepsin K (CtsK), are expressed by ECC endothelial cells. RANKL inhibits

VEGF-induced cell proliferation while causing increased expression of CtsK via

Cn/NFATc1 and c-Jun N-terminal Kinase 1/2 (JNK1/2)-dependent signaling.

Conclusion: These data support a novel mechanism for the transition from ECC growth

to remodeling in which NFATc1 promotes a sequential pattern of gene expression via

cooperation with ligand-specific cofactors such as MEK1-ERK1/2 or JNK1/2.

Non-Standard Abbreviations and Acronyms: NFATc1 (Nuclear Factor of Activated

T-cells cytoplasmic 1), ECC (endocardial cushion), ECM (extracellular matrix), VEGF

(Vascular Endothelial Growth Factor A), Cn (calcineurin), MEK1-ERK1/2 (MAPK-ERK

Kinase 1 - Extracellular signal-Related Kinase 1/2), RANKL (Receptor Activator of NFκB

Ligand), CtsK (Cathepsin K), JNK1/2 (c-Jun N-terminal Kinase 1/2), EMT (epithelial to mesenchymal transformation), E (embryonic day), CsA (Cyclosporin A), HPVEC

(human pulmonary valve endothelial cell), BrdU (bromodeoxyuridine), HH (Hamburger and Hamilton stage), sFlt1 (soluble Flt receptor chimera), OPG (Osteoprotegerin), BSA

(bovine serum albumin), ICLSM (immunofluorescence and confocal laser scanning microscopy), pHH3 (phosphohistone H3), ISH (in situ hybridization), pJNK1/2

(phosphorylated JNK1/2), dpERK1/2 (diphosphorylated ERK1/2), PI3K

(Phosphoinositide 3-Kinase), MMP9 (Matrix Metalloproteinase 9), IHC

(immunohistochemistry), AVC (atrioventricular canal), Myo (myocardium), Endo (ECC endothelial cells), Mes (mesenchymal cells), MV (mitral valve), TV (tricuspid valve), NT

(neural tube)

Introduction

Congenital malformations of cardiac valves affect 1-2% of the population; however, the molecular mechanisms that govern valve development are still not completely understood (Garg et al., 2005). Cardiac valvulogenesis is a complex process that begins with the formation of endocardial cushions (ECCs) in the atrioventricular canal and outflow tract regions of the looping heart. The ECCs are populated as overlying endothelial cells delaminate, undergo epithelial to mesenchymal transformation (EMT), and invade the extracellular matrix (ECM) (Person et al., 2005).

During EMT, ECC endothelial cells are highly proliferative, and increased mitotic index is a feature of both endothelial and mesenchymal cells of the developing valves (Hinton et al., 2006; Person et al., 2005). After outgrowth, the ECM of the valves is remodeled into a highly organized, trilaminar architecture characteristic of mature cardiac valves.

During ECM remodeling, valve interstitial cell proliferation is decreased and endothelial cells express ECM remodeling enzymes (Lange and Yutzey, 2006; Lincoln et al., 2004).

It is clear that precise regulation of valve cell cycle, growth, and remodeling is required for normal valve development, as alterations in these processes are linked to valve defects (Armstrong and Bischoff, 2004; Chang et al., 2004; Person et al., 2005). While much is known about signaling mechanisms regulating ECC formation and EMT, mechanisms governing the transition from growth to remodeling in the developing valves are relatively uncharacterized.

Nuclear Factor of Activated T-cells cytoplasmic 1

(NFATc1/NFAT/NFATc/NFAT2), a transcription factor of the NFAT family, functions in

development and homeostasis of the brain, skeleton, immune system, and heart

(Macian et al., 2000). In both mouse and chick embryos, cardiac NFATc1 expression is

specific to ECC endothelial cells and overlaps with expression of its regulatory

phosphatase Calcineurin (Cn) (Chang et al., 2004; de la Pompa et al., 1998; Liberatore and Yutzey, 2004). NFATc1 null mice, or those lacking NFATc1 expression specifically in endothelial cells, have normal ECC formation and EMT, however, these ECCs fail to grow and remodel, resulting in lethality at embryonic day (E)12.0-E14.5 (de la Pompa et al., 1998; Ranger et al., 1998). Mice lacking expression of Cnb1 specifically in endothelial cells or chick embryos treated with the Cn inhibitor Cyclosporin A (CsA), just prior to cushion growth, phenocopy NFATc1-/- mouse models, thereby illustrating the importance of the Cn-NFATc1 interaction for valve growth and remodeling (Chang et al.,

2004; Liberatore and Yutzey, 2004). A necessary spatiotemporal window for Cn-

NFATc1 activation in ECC endothelial cells has been defined, however, upstream effectors and intersecting pathways of NFATc1 in ECC endothelial cells during this critical period were not previously identified.

Vascular endothelial growth factor A (VEGF/VEGFA/VEGF165) is critical for

development and maintenance of heart, lung, and vascular tissues (Cleaver and Melton,

2003; Ferrara et al., 2003; Miquerol et al., 1999). VEGF levels must be tightly controlled

during cardiac valve morphogenesis, as VEGF signaling maintains the ECC endothelial

cell layer during ECC formation, but is also a potent inhibitor of EMT (Armstrong and

Bischoff, 2004; Dor et al., 2001). VEGF is expressed by myocardium and ECC

endothelial cells during ECC formation and growth, however, expression is extinguished

during ECC remodeling (Miquerol et al., 1999). While it is known that under or

overexpression of VEGF in ECC endothelial cells disrupts ECC morphogenesis, the role

of VEGF in post-EMT ECC growth is not known (Dor et al., 2001). VEGF/NFATc1

signaling has been implicated in homeostasis of valve endothelial tissue, as VEGF

treatment of adult human pulmonary valve endothelial cells (HPVECs) increases cell

proliferation (Johnson et al., 2003). However, VEGF regulation of NFATc1 in ECC cells

during development has not been demonstrated.

Receptor Activator of NFκB Ligand (RANKL/TRANCE/TNFSF11/OPGL/ODF) is a

member of the TNF family of signaling molecules that is best known for its role in

promoting osteoclast differentiation and production of ECM remodeling enzymes, such

as Cathepsin K (CtsK) (Takayanagi, 2005; Theoleyre et al., 2004). In myxomatous and

diseased human mitral valves containing high levels of fragmented collagen and elastin,

CtsK expression is upregulated by interstitial cells (Rabkin et al., 2001). Consistent with

RANKL/NFATc1 pathway activity during valve remodeling, CtsK is normally expressed by murine valve endothelial cells at E13.5 (Lange and Yutzey, 2006). However, CtsK is

not expressed in NFATc1 null embryos (Lange and Yutzey, 2006). Conserved

spatiotemporal expression of RANKL/NFATc1-related genes among vertebrates has not

been previously demonstrated. Likewise, the ability of RANKL to promote CtsK

transcription in a Cn-dependent manner in ECC cells has not been tested.

The present study examines the relationship of VEGF and RANKL signaling

mechanisms in the regulation of NFATc1 in cardiac valve growth and remodeling. In

isolated avian ECC cultures, VEGF signaling promotes cell proliferation via activation of

Cn/NFATc1 together with MAPK-ERK Kinase1-Extracellular signal-Related Kinase1/2

(MEK1-ERK1/2) copathways. In vivo, NFATc1 is necessary for normal ECC growth, as mice lacking NFATc1 expression have decreased ECC endothelial cell proliferation at

E10.5. In addition, RANKL/NFATc1 signaling is conserved among avian and mammalian embryos and RANKL acts during valve remodeling to promote CtsK expression via activation of Cn/NFATc1 and Jun N-terminal Kinases1/2 (JNK1/2) signaling. These data suggest NFATc1 plays a central role in the transition from ECC cell proliferation, in response to VEGF signaling, to ECM remodeling enzyme production, in response to RANKL signaling. VEGF/NFATc1 signaling promotes cell proliferation and not CtsK expression via cooperation with MEK1-ERK1/2-dependent cofactors, while RANKL/NFAT signaling inhibits cell proliferation and increases CtsK expression via cooperation with JNK1/2-dependent cofactors.

Materials and methods

E10.5 NFATc1 mouse embryos were collected and genotyped as previously described (Ranger et al., 1998). Proliferative indices of ECC cells were determined via quantification of bromodeoxyuridine (BrdU) labeled nuclei versus total nuclei in paraffin embedded mouse heart sections. Explanted superior and inferior atrioventricular ECCs, isolated from Hamburger and Hamilton stage 25 (HH25; E4.5) chicken embryos,

(Hamburger and Hamilton, 1951) were maintained in culture for 2–7 days with one or more of the following treatments added to the culture media: VEGF (R&D Systems)

50ng/mL, soluble Flt receptor chimera (sFlt1) (R&D Systems) 50ng/mL, RANKL (R&D

Systems) 800ng/mL, Osteoprotegerin (OPG) (R&D Systems) 1ug/mL, Cyclosporin A

(CsA) (Novartis) 1ug/mL, Bovine Serum Albumin (BSA) (Sigma) 100-800ng/mL, U0126

(Promega) 10uM, SP600125 (Calbiochem) 2.5uM, and/or DMSO (Sigma) 0.005-0.01%.

Real Time RT-PCR for CtsK expression and in situ hybridization for NFATc1, RANKL,

and CtsK mRNA were performed as previously described (Ehrman and Yutzey, 1999;

Lincoln et al., 2006; Shelton and Yutzey, 2007). An expanded materials and methods are included in the supplementary materials section.

Results

NFATc1-/- mouse embryos have decreased proliferation of ECC endothelial cells in vivo

During ECC formation, endothelial cells populate the ECC mesenchyme by undergoing EMT and migrating into the cardiac jelly. ECCs then enter a growth period

(mouse E10.5-E13.5) during which endothelial and mesenchymal cells of the ECC are highly proliferative (Hinton et al., 2006; Lincoln et al., 2004). In order to determine if

NFATc1 is necessary for ECC endothelial cell proliferation, the proliferative index of

ECC endothelial cells was examined in NFATc1+/+, +/- and -/- E10.5 mouse embryos.

Pregnant NFATc1+/- females were injected intraperitoneally with BrdU labeling solution.

The proliferative index of endothelial cells overlying ECCs along with ECC

mesenchymal cells and ventricular myocytes was assessed for at least six embryos of

each genotype isolated from separate litters. Adjacent sections were labeled with MF20

antibody in order to visualize the myocardial boundaries of atrioventricular ECCs (Figure

1A-C). Wild type (NFATc1+/+) and heterozygous (+/-) mice have 32% and 30% of ECC

endothelial cell nuclei labeled with BrdU respectively, while NFATc1 null (-/-) littermates have a significantly lower (21%) percentage of BrdU labeled nuclei (Figure 1D).

Decreased proliferation also was observed in ECC mesenchymal cells that arise from

ECC endothelial cells via EMT (Figure 1D) (Lincoln et al., 2004). The proliferative defect observed is not due to general embryo failure, as no differences were detected in the mitotic index of ventricular myocytes among genotypes (Figure 1D). It is important to note that cardiac myocytes do not express NFATc1. This suggests that Cn/NFATc1 signaling is necessary during ECC formation and growth for normal ECC cell proliferation. No difference in the number of apoptotic cells were observed in ECCs of

NFATc1+/+, +/-, or -/- E10.5 mice, as determined by anti-Cleaved Caspase3 labeling

(data not shown). These results indicate that the hypoplastic ECC phenotype observed in NFATc1-/- embryos at E13.5 is likely due to decreased proliferation of ECC endothelial and endothelial-derived mesenchymal cells as early as E10.5, when these embryos are grossly indistinguishable from NFATc1+/+ and +/- littermates.

VEGF treatment increases ECC cell proliferation via Calcineurin/NFATc1 and MEK1-

ERK1/2 signaling.

VEGF regulates endothelial cell adhesion, cell cycle, and inflammatory cell recruitment during development and throughout postnatal life (Ferrara et al., 2003). The ability of VEGF to increase NFATc1 nuclear localization and cell proliferation was examined in ECC cells isolated from HH25 (E4.5) chicken embryos and placed in cell culture. The avian ECC culture system was used because HH25 chick embryos have

much larger ECCs than murine embryos at the same stage in development, large numbers of synchronously staged ECCs can be collected at one time, and cultured ECC explants can be treated with cytokines or inhibitors to manipulate developmental pathways without the confounding effects of myocardial or systemic interactions (Hinton et al., 2006; Lincoln et al., 2006; Shelton and Yutzey, 2007). These explants contain valve progenitor cells as well as precursors of septum intermedium (Person et al.,

2005). NFATc1-positive ECC cells are endothelial as indicated by co-expression with endothelial markers such as Sox17 and VEGFR2 (Supplemental Figure 1A-A”; data not shown) (Cleaver and Melton, 2003; Lefebvre et al., 2007). NFATc1-positive cells from non-dissociated ECC explants exist as clusters of Sox17 or VEGFR2 positive endothelial cells surrounded by Smooth Muscle α-Actin positive, MF20, and Sox17 negative mesenchymal cells. Myocyte contamination of cultures was not detected by immunofluorescence and confocal laser scanning microscopy (ICLSM) or Real Time

RT-PCR. Isolated atrioventricular ECCs were treated with VEGF, soluble Flt1 receptor chimera (sFlt1, a VEGF inhibitor), Cyclosporin A (CsA, a Cn inhibitor), VEGF+sFlt1,

VEGF+CsA, or BSA (as a vehicle control). Nuclear localization NFATc1 of was evaluated by ICLSM.

Cultures treated with VEGF contained significantly more cells with nuclear

NFATc1 labeling than control cultures (76% vs. 16%), indicating that VEGF promotes

NFATc1 nuclear localization (Figure 2A-B, G). In contrast, cultures treated with

VEGF+sFlt1 or sFlt1 alone had a comparable number of cells containing nuclear

NFATc1 as control cultures, indicating that increased NFATc1 nuclear localization was a

specific effect of VEGF treatment (Figure 2C,E, G). ECC cells treated with VEGF+CsA or CsA alone also showed low level NFATc1 nuclear localization comparable to controls, demonstrating that NFATc1 nuclear accumulation following VEGF treatment is

Cn-dependent (Figure 2D, F-G). It is important to note that nuclear size and morphology in all cultures of this study were comparable and apparently normal with no evidence of toxicity. Since NFATc1 is predominantly expressed by endothelial cells of the ECC in vivo and in culture, these results demonstrate that VEGF treatment promotes NFATc1 nuclear localization in ECC endothelial cells.

To determine if VEGF induces proliferation of ECC cells, HH25 chick ECCs were cultured and treated with VEGF, VEGF+sFlt1, VEGF+CsA, sFlt1, CsA, or BSA. The proliferative index was calculated as the percent of total nuclei labeled with the M-phase marker anti-phosphohistone H3 (pHH3) antibody. Cultures treated with VEGF had a significantly higher percentage of pHH3 positive cells (4.4%) than BSA treated controls

(2.3%) (Figure 3A-B, G). Proliferation indices from cultures treated with a combination of VEGF+sFlt, VEGF+CsA or sFlt1 alone were comparable to BSA treated controls, indicating that the increased proliferation is a specific effect of VEGF treatment and is

Cn-dependent (Figure 3C-E, G). Cultures treated with CsA alone had a significantly lower percentage of pHH3 positive nuclei, which could be due to the ability of CsA treatment to block endogenous Cn/NFAT signaling and affect cell proliferation in the absence of added VEGF (Figure 3F-G). Endogenous expression of VEGFR2, the main receptor for VEGF signaling, is specific to ECC endothelial cells and it was determined that VEGR2 is localized to clusters of endothelial cells in culture (data not shown)

(Armesilla et al., 1999; Miquerol et al., 1999). Similarly, the majority of pHH3 positive

cells co-express endothelial markers such as Sox17 and NFATc1 in VEGF-treated

cultures (Supplemental Figure 1B-C). Together, these data indicate that VEGF

increases proliferation of ECC endothelial cells in a Cn-dependent manner.

RANKL increases CtsK gene expression via Cn/NFATc1

In osteoclasts, RANKL/NFATc1 signaling promotes bone ECM remodeling by

inducing expression of NFATc1 target genes, including the ECM remodeling enzyme

CtsK (Takayanagi, 2005). RANKL/NFATc1 pathway components are expressed by

murine ECC endothelial cells during physiological valve remodeling and by human ECC

cells during pathological remodeling, suggesting that this pathway has a role in normal

valve development and human disease mechanisms (Lange and Yutzey, 2006; Rabkin

et al., 2001; Steinmetz et al., 2008). Since species-specific differences between mouse

and chicken in TGFβ signaling molecules have been noted during ECC formation, we sought to determine that the spatiotemporal expression patterns of NFATc1, RANKL, and CtsK during valve growth and remodeling are conserved in chicken embryos

(Person et al., 2005). In situ hybridization (ISH) revealed that in chicken embryos,

NFATc1 is expressed by ECC endothelial cells throughout growth and remodeling

(chicken E4.5-E14) (Figure 4A-B; data not shown). In contrast, RANKL and CtsK are absent in ECC but are expressed later during valve remodeling (E7), as detected by ISH

(Figure 4C-F), or Real Time RT-PCR (data not shown), which correlates with data from mouse models (Lange and Yutzey, 2006). Previous studies indicate that, in mouse

embryos, VEGF expression is upregulated in atrioventricular canal cells at E10.5, during

ECC growth, and is depleted by E14.5 (Dor et al., 2001; Miquerol et al., 1999). Similarly in avian embryos, VEGF expression is upregulated in the atrioventricular canal during

ECC growth (Rodgers et al., 2006). Therefore, VEGF and NFATc1 are expressed during ECC growth, while RANKL/NFATc1 pathway components are not expressed until

ECC remodeling in both chicken and mouse model systems. These findings suggest that VEGF/NFATc1 signaling during ECC growth and RANKL/NFATc1 signaling during valve remodeling are conserved mechanisms controlling valve development among vertebrates.

To examine the ability of RANKL to induce NFATc1 nuclear localization in ECC cells, avian ECCs were cultured and treated with RANKL, Osteoprotegerin (OPG, a soluble RANKL inhibitor), CsA, RANKL+OPG, RANKL+CsA, or BSA. The percentage of cells with nuclear NFATc1 was determined by ICLSM. Cells treated with RANKL had a significantly higher percentage of cells containing nuclear NFATc1 (37%) than BSA treated controls (16%), indicating that RANKL promotes NFATc1 nuclear localization in

ECC cells (Figure 5A-B, G). Cells treated with RANKL+OPG or OPG alone had

NFATc1 nuclear localization comparable to controls, indicating that increased nuclear

NFATc1 is a specific effect of RANKL treatment (Figure 5C, E, G). ECC cells treated with RANKL+CsA or CsA alone also showed NFATc1 nuclear localization comparable to controls, demonstrating that NFATc1 nuclear accumulation induced by RANKL is Cn- dependent (Figure 5D, F-G). Together, these results indicate that RANKL treatment promotes NFATc1 nuclear localization in cultured ECC cells.

To determine if RANKL induces expression of the ECM remodeling enzyme

CtsK in ECC cells, avian ECCs were explanted and cultured for 7 days with the

aforementioned treatments added to media. In these experiments, addition of RANKL

to cultured ECCs resulted in 7.1 fold higher CtsK mRNA expression than BSA treated

controls, as detected by Real Time RT-PCR (Figure 5H). Treatment with RANKL+OPG

or OPG alone yielded CtsK expression levels comparable to controls, indicating that

increased CtsK expression is a specific effect of RANKL treatment (Figure 5H).

Treatment with RANKL+CsA significantly inhibited the induction of CtsK expression, while CsA treatment alone had expression comparable to BSA (Figure 5H). For all Real

Time RT-PCR experiments, GAPDH and β-actin mRNA levels prior to normalization were comparable among culture groups, indicating that cultures contained a similar number of live cells at collection. Taken together, these results indicate that RANKL promotes expression of CtsK via Cn signaling in ECC cells.

The results obtained for 7-day cultures are comparable to those obtained when cells were maintained in culture for 4 days. However, no induction of CtsK mRNA expression was observed at 48 hours, even though RANKL-induced NFATc1 nuclear localization at this time point (Figure 5B, G; data not shown). Expression of markers associated with ECC maturation were assessed for ECC explants maintained in culture for 2 days versus those cultured for 7 or 10 days. While NFATc1 expression remained comparable among all cultures, CtsK expression increased over the culture period. This increase in CtsK mimics gene expression of ECCs and valves in vivo (Supplemental

Figure 2A-B). Similarly, expression of Periostin and Versican in ECC cultures increased

over time, while Scleraxis mRNA levels remain relatively unchanged. This pattern of gene expression closely resembles gene transcription in vivo (Supplemental Figure 2A-

B). Together, these data suggest that cultured ECC cells have a pattern of gene expression consistent with maturing valves in vivo and that RANKL-induced CtsK expression in ECC cells is not only ligand-dependent, but is time-dependent as well.

Cn/NFATc1 activation is a nodal point in RANKL and VEGF signaling

The specificity of VEGF and RANKL induction of cell proliferation and ECM remodeling enzyme expression was examined. In order to determine if VEGF/NFATc1 signaling can induce CtsK transcription, HH25 avian ECC cells were cultured in the presence of VEGF, VEGF+sFlt1, VEGF+CsA, sFlt1, or CsA. None of these treatment groups expressed increased Ctsk mRNA compared to BSA treated controls (Figure 6A).

To determine if RANKL/NFATc1 signaling promotes ECC cell proliferation, ECCs were cultured in the presence of RANKL, RANKL+OPG, RANKL+CsA, OPG, CsA, or BSA. In these experiments, none of the treatment groups exhibited increased proliferation compared to BSA treated controls (Figure 6C). These results show that downstream effects of NFATc1 activation in ECC cells are ligand-dependent.

To examine the signaling hierarchy and crosstalk at the level of VEGF and

RANKL receptors upstream of NFATc1, ECC explants were cultured and treated with

VEGF+OPG (RANKL inhibitor) or RANKL+sFlt (VEGF inhibitor). These data demonstrated that VEGF-induced ECC cell proliferation does not require RANK receptor function (Supplemental Figure 3A), and likewise, RANKL-induced CtsK mRNA

expression does not require VEGF receptor function (Supplemental Figure 3B).

Therefore, VEGF and RANKL signaling act independently with separable downstream effects in ECCs; however, both VEGF-induced cell proliferation and RANKL-induced

CtsK expression are Cn/NFATc1-dependent.

RANKL inhibits VEGF-induced proliferation of ECC cells

The above results are consistent with a mechanism whereby VEGF activation of

NFATc1 promotes ECC proliferation, followed by RANKL activation of NFATc1 to induce

ECM remodeling enzyme expression in maturing valves. Therefore, experiments were performed to examine the response of ECC cells in the presence of both VEGF and

RANKL signals concurrently. Cells treated with VEGF+RANKL together had a proliferative index comparable to BSA treated controls (2.6% vs. 2.3%) (Figure 6D).

This is in contrast to VEGF-treated cultures that had a significantly higher proliferative index than BSA treated controls (Figure 6D). These results indicate that RANKL treatment of ECC cells inhibits VEGF-induced ECC cell proliferation. In addition, treatment of ECC cells with RANKL alone significantly inhibits cell proliferation compared to control cultures (Figure 6C-D), consistent with RANKL inhibition of endogenous ECC cell proliferation mechanisms. Similarly, ECC cells were cultured in the presence of VEGF+RANKL to determine the effects on CtsK transcription. Addition of VEGF with RANKL to cultures does not significantly inhibit RANKL-induced CtsK expression (Figure 6B) and VEGF alone does not affect CtsK expression. Taken together, these results indicate there is crosstalk in the signaling pathways that regulate

NFATc1, whereby RANKL inhibits VEGF-induced proliferation of ECC cells while activating CtsK transcription.

VEGF and RANKL require MEK1-ERK1/2 and JNK1/2 signaling, respectively, to induce proliferation and CtsK expression.

RANKL stimulates ECM remodeling enzyme production via co-activation of

Cn/NFATc1 and JNK1/2 pathways in osteoclasts (Takayanagi, 2005). JNK1/2 activation in cardiac valves was examined in vivo via immunohistochemistry on E12.5 mouse heart sections with anti-phosphorylated JNK(Thr183/Tyr185) antibody. JNK1/2 activation was detected in mitral and tricuspid valve endothelial cells consistent with

RANKL and JNK1/2 activity during valve ECM remodeling in vivo (Figure 7). To determine if RANKL-induced CtsK expression requires JNK1/2 signaling in ECC cells, avian ECC explants were cultured for 7 days in the presence of RANKL+DMSO,

RANKL+SP600125 (a JNK1/2 inhibitor), SP600125, or DMSO (vehicle control). ICLSM was used to determine that SP600125 treatment of cultured ECC cells significantly decreased phosphorylated JNK(Thr183/Tyr185) expressing ECC cells, while RANKL- induced NFATc1 nuclear localization was not significantly altered (Supplemental Figure

4E-H and data not shown). Real Time RT-PCR demonstrated that SP600125 treatment significantly decreased RANKL-induced CtsK expression (Figure 8C). Therefore,

RANKL-induced CtsK expression in ECC cells is JNK1/2-dependent.

In vascular endothelial cells, VEGF stimulation of VEGFR2 activates Cn/NFAT and ERK1/2 copathways together to promote gene transcription (Ferrara et al., 2003;

Schweighofer et al., 2007). In order to determine if ERK activation is specifically required for VEGF-mediated effects downstream of NFATc1 in ECC cells, ECCs were cultured in the presence of VEGF+DMSO, VEGF+U0126 (a MEK1-ERK1/2 inhibitor),

U0126, or DMSO (vehicle control). ICLSM was used to determine that U0126 treatment of cultured ECC cells significantly decreased diphosphorylated ERK1/2 expressing ECC cells, while VEGF-induced NFATc1 nuclear localization was not significantly altered

(Supplemental Figure 4A-D and data not shown). VEGF and DMSO control treated cells had mitotic indices of 4.0% and 2.6% respectively, as determined by pHH3 immunoreactivity (Figure 8A). Addition of U0126 to cultures either alone or in combination with VEGF blocked the effects of VEGF treatment, and presumably endogenous VEGF signaling on cell proliferation, resulting in a significantly decreased mitotic index of 1.3% for both, compared to control cultures (Figure 7A). In contrast,

MEK1-ERK1/2 inhibition had no effect on RANKL-induced CtsK expression in cultured

ECC cells. As a specificity control, treatment with Phosphoinositide 3-Kinase (PI3K) inhibitor LY294002 had no effect on either VEGF-induced proliferation or RANKL- induced CtsK expression in ECC cells (Figure 7D; data not shown). These data demonstrate that VEGF-induced ECC cell proliferation is MEK1-ERK1/2-dependent, however, RANKL-induced CtsK expression does not require MEK1-ERK1/2 activity.

Conversely, JNK1/2 inhibition with SP600125 did not significantly alter VEGF- induced ECC cell proliferation, demonstrating that VEGF-induced ECC cell proliferation is JNK1/2-independent (Figure 8B). To determine if RANKL inhibition of VEGF-induced

ECC cell proliferation requires JNK1/2 activation, ECC cells were treated with VEGF,

RANKL, and SP600125. These experiments showed that RANKL-mediated inhibition of

VEGF-induced ECC cell proliferation is JNK1/2-dependent (Figure 8E). Interestingly,

JNK1/2-dependent signaling has not previously been associated with maturation of ECC cells. Overall, these results show that, in conjunction with Cn/NFATc1, MEK1-ERK1/2 activation is necessary to achieve VEGF-induced ECC cell proliferation, while JNK1/2 activation is necessary for RANKL-induced CtsK expression and for RANKL-mediated inhibition of ECC cell proliferation.

Discussion

During heart valve morphogenesis, ECCs transition from growth, characterized by high cell proliferation, to remodeling, during which the ECM is stratified and mature valve leaflets become apparent (Hinton et al., 2006). Investigation into the role of

NFATc1 in valve maturation supports a model whereby VEGF/NFATc1/ERK1/2 signaling promotes ECC cell proliferation during ECC growth, and

RANKL/NFATc1/JNK1/2 signaling inhibits VEGF-induced cell proliferation, while promoting CtsK expression, during valve remodeling. These data also support a novel mechanism for the transition from ECC growth to remodeling in which NFATc1 promotes a sequential pattern of gene expression via cooperation with ligand-specific cofactors MEK1-ERK1/2 and JNK1/2 (Supplemental Figure 5).

NFATc1-/- mouse embryos exhibit decreased proliferation of ECC endothelial and mesenchymal cells at E10.5. Prior to this time, ECCs are apparently normal, demonstrating that NFATc1 is not required for ECC formation and EMT (Chang et al.,

2004; de la Pompa et al., 1998; Ranger et al., 1998). VEGF is an upstream activator of

Cn/NFATc1 and requires MEK1-ERK1/2 activation in promoting proliferation of cultured

ECC cells. In vivo ECC growth is characterized by nuclear localization of NFATc1 in endothelial cells in addition to expression of VEGF and VEGFR2 (Chang et al., 2004;

Dor et al., 2001; Rodgers et al., 2006). VEGF signaling must be tightly regulated for normal valvulogenesis to occur, as VEGF is necessary for endothelial proliferation and maintenance, as well as being a potent inhibitor of EMT during initial formation of the

ECCs (Dor et al., 2001; Rodgers et al., 2006). In the atrioventricular canal myocardium,

VEGF expression is negatively regulated by NFATc3/c4, however, NFATc1 has not been shown to regulate VEGF transcription in the ECC (Chang et al., 2004). By E14.5 in mouse, valve remodeling has begun and expression of VEGF and VEGR2 are lost in valve endothelial cells, supporting a model whereby loss of VEGF signaling in the ECC endothelial cells is associated with the transition from growth to remodeling during valvulogenesis (Dor et al., 2001; Miquerol et al., 1999).

The RANKL/NFATc1 pathway is conserved among vertebrates and is active in endothelial cells of remodeling valves. This study is the first to report that RANKL induces CtsK expression via co-stimulation of Cn/NFATc1 and JNK1/2 pathways in

ECC cells. In the skeletal system, NFATc1 is a key regulator of osteoclast differentiation and function in response to RANKL signaling (Takayanagi, 2005). Upon

RANKL binding in osteoclasts, the RANK receptor recruits adaptor molecules that co- stimulate NFAT and JNK1/2 pathways ultimately leading to NFATc1/AP1-mediated activation of ECM remodeling enzymes such as Ctsk and Matrix Metalloproteinase 9

(MMP9) (Takayanagi, 2005). Expression of RANKL/NFATc1 pathway components,

RANKL, CtsK and MMP9 are associated with increased pathogenic ECM remodeling and calcification of human valves suggesting this pathway may play a role in valve maturation and disease (Rabkin et al., 2001; Steinmetz et al., 2008). In contrast, VEGF stimulation of NFATc1 in HPVECs induces endothelial cell proliferation, which implicates NFATc1 in normal homeostasis of the valve endothelium (Johnson et al.,

2003).

The work presented here and elsewhere demonstrates that NFATc1 participates in complex regulatory interactions during valve development. In developing osteoclasts and endothelial cells, NFATc1 forms complexes with other NFATs as well as unrelated transcription factors such as Elks, GATAs and AP1, to bind DNA (Crabtree and Olson,

2002; Graef et al., 2001; Macian et al., 2000). Ligand-specific responses to Cn/NFATc1 activation occur through selective co-stimulation of NFATc1 partners in T-cells, where genes associated with increased immune response are targeted by NFATc1/AP1 complexes, while genes associated with dampened immune response are activated by

NFATc1 in the absence of AP1 (Hogan et al., 2003). JNK1/2 signaling is important for outflow tract development, but its role in valve development has not been previously reported (Craig et al., 2008). MEK1-ERK1/2-activated transcription factors are necessary for EMT and ECC cell proliferation (Krenz et al., 2005; Stevens et al., 2008).

Together these data suggest NFATc1 plays a role in regulating the transition from ECC growth to valve remodeling via partnership with ligand-specific cofactors to elicit gene expression. Further interrogation of NFATc1 and NFATc1 costimulatory pathway

functions in valve maturation and homeostasis may reveal new therapeutic targets for prevention and treatment of congenital valve defects and disease.

Acknowledgments

We are indebted to Alexander Lange for his continued insight and technical support, to Kristen Lipscomb-Sund and Timothy Mead for editorial comments, and to

Christina Alfieri, Joy Lincoln, Elaine Shelton and Heather Evans-Anderson for their technical assistance.

Sources of funding

American Heart Association Great Rivers Affiliate Predoctoral Fellowship Award

#715107B

NIH/NHLBI SCCOR in Pediatric Cardiology #P50HL074728

Disclosures

None

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Figure legends

Figure 1. E10.5 NFATc1 -/- mouse ECC endothelial and mesenchymal cells

exhibit decreased proliferation. A-B) Immunohistochemistry (IHC) of NFATc1 +/+ (A)

and NFATc1-/- (B) mouse embryo AVC sections shows anti-BrdU labeled (brown) ECC endothelial (arrows) and mesenchymal cells (arrowheads) and hematoxylin (blue) stained nuclei. C) IHC of NFATc1+/+ mouse embryo atrioventricular canal (AVC) section shows MF20-reactive myocardium (Myo), and non-reactive ECC endothelial cells (Endo; arrows) and mesenchymal cells (Mes; arrowheads). D) Quantification of percent BrdU positive AV endothelial, mesenchymal and myocardial cells in six embryos for each genotype (n=6). *P≤0.01.

Figure 2. VEGF treatment of ECC cells induces NFATc1 nuclear localization. A-F)

ICLSM of avian ECC cultures treated with BSA (A), VEGF (B), VEGF+sFlt (C),

VEGF+CsA (D), sFlt (E), or CsA (F) and labeled with anti-NFATc1 antibody (green) and

ToPro3 iodide nuclear stain (blue) shows cells with nuclear NFATc1 labeling (arrows) and cells lacking nuclear NFATc1 labeling (arrowheads). G) Quantification of percent cells with nuclear NFATc1 labeling in three independent experiments for each treatment

(n=3). *P≤0.01.

Figure 3. VEGF-induced proliferation of ECC cells is dependent upon Calcineurin signaling. A-F) ICLSM of ECC cultures treated with BSA (A), VEGF (B), VEGF+sFlt

(C), VEGF+CsA (D), sFlt (E), or CsA (F) and labeled with anti-pHH3 antibody (pink) and

ToPro3 (blue) shows M-phase ECC cells (arrows). G) Quantification of percent mitotic

cells in three independent experiments for each treatment (n=3). *P≤0.01.

Figure 4. NFATc1, RANKL, and CtsK mRNA expression in developing chick AVC.

In situ hybridization of E5 and E7 chick heart sections. A-Bʼ) NFATc1 mRNA is

expressed by endothelial cells of ECC and mitral valves (arrows). C-Fʼ) RANKL and

CtsK transcripts are absent in E5 ECCs (C+E), however, RANKL and CtsK mRNAs are expressed by E7 mitral valve (MV) endothelial cells (arrows) (D,Dʼ,F,Fʼ).

Figure 5. RANKL treatment of ECC cells induces NFATc1 nuclear localization and increased CtsK expression via a Cn-dependent mechanism. A-F) ICLSM of

ECC cultures treated with BSA (A), RANKL (B), RANKL+OPG (C), RANKL+CsA (D),

OPG (E), or CsA (F) and labeled with anti-NFATc1 antibody (green) and ToPro3 (blue) shows cells with nuclear NFATc1 labeling (arrows) and cells lacking nuclear NFATc1 labeling (arrowheads). G) Quantification of percent cells with nuclear NFATc1 labeling in three independent experiments for each treatment (n=3). H) Quantification of fold difference in CtsK transcript for treated ECC cultures. Real Time RT-PCR samples were run in triplicate and normalized to GAPDH mRNA for three independent experiments per treatment (n=3). *P≤0.01 and #P≤0.05.

Figure 6. Ligand-specific effects on ECC cell proliferation and CtsK expression.

RANKL inhibits VEGF-induced ECC cell proliferation. A-B) Quantification of fold

difference in CtsK transcript for treated ECC cultures. Real Time RT-PCR samples were run in triplicate and normalized to GAPDH mRNA for three independent experiments per treatment (n=3). C-D) Quantification of percent pHH3 positive nuclei in treated ECC cultures for at least three independent experiments per treatment (n=3-6).

*P≤0.01.

Figure 7. JNK1/2 activation is not seen in E11.5 mouse ECCs, but is detected in

E12.5 mitral and tricuspid valve endothelial cells in vivo. IHC of E11.5 and E12.5 control mouse sections was performed using anti-phosphorylated JNK(Thr183/Tyr185)

(pJNK) antibody. A-Aʼ) pJNK positive cells are not detected in E11.5 ECCs

(arrowhead). B-Bʼ) pJNK positive endothelial cells (arrows in Bʼ) are detected in both mitral (MV) and tricuspid (TV) valves (arrowheads) at E12.5. C) No primary antibody control E12.5 AVC section. D) Positive control E11.5 neural tube (NT) section. Arrows indicate pJNK positive cells.

Figure 8. VEGF-induced ECC cell proliferation is MEK1-ERK1/2-dependent.

RANKL-induced CtsK expression and RANKL inhibition of VEGF-induced cell proliferation is JNK1/2 dependent. A,B,E) Quantification of percent pHH3 positive nuclei in treated chick ECC cultures for three independent experiments per treatment

(n=3). C-D) Quantification of fold difference in CtsK mRNA expression as determined

by Real Time RT-PCR. Samples were run in triplicate and normalized to GAPDH transcript for three independent experiments per treatment (n=3). *P≤0.01 and #P≤0.05.

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Supplementary methods

Chicken and mouse embryo collection

Fertilized white leghorn chicken eggs (CBT Farms, MD and Charles River Laboratories,

CT) were incubated at 38°C under high humidity. Embryos were collected at

Hamburger and Hamilton (HH) stages 25, 26, 30 and 36 corresponding to E4.5, 5, 7 and 10 days, respectively (Hamburger and Hamilton, 1992). NFATc1 heterozygous mutant mice were obtained from Dr. Laurie Glimcher (Ranger et al., 1998). Mouse embryos were generated via timed matings with observation of a copulation plug designated as E0.5. Embryos were collected at E10.5 and genotyping for NFATc1 mutation was performed by PCR using primers designed for the wild type and targeted alleles as described in Ranger et al. (Ranger et al., 1998). All animal procedures were approved and performed in accordance with institutional guidelines.

Immunohistochemical analysis and quantification of cell proliferation

Pregnant female mice were injected on E10.5 intraperitoneally with 10μL/g body weight

Bromodeoxyuridine (BrdU) Labeling Solution (Zymed). Females were sacrificed two hours post injection and embryos were collected, fixed, and processed for immunohistochemistry as previously described (Bushdid et al., 2003). Proliferation of endocardial cushion (ECC) endothelial and mesenchymal cells was determined as described in Bushdid et al., except that the proliferative index for ECC endothelial cells was determined by the number of BrdU positive ECC endothelial cell nuclei divided by the total number of endothelial cell nuclei within a 5μm section (Bushdid et al., 2003).

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The proliferative index for ECC mesenchymal cells was determined as the number of

BrdU positive ECC mesenchymal cell nuclei divided by the total number of ECC mesenchymal cell nuclei per section. The proliferative index for ventricular myocytes was determined as the number of BrdU positive myocyte nuclei divided by the total number of myocyte nuclei per section. At least five comparable heart sections were analyzed per embryo for six embryos of each genotype. For MF20 antibody labeled sections, MF20 antibody (Iowa Hybridoma Bank) was diluted 1:200 in 0.5% blocking solution (Pierce) and incubated on sections overnight at 4°C. For anti-phosphorylated

JNK(Thr183/Tyr185) labeled sections, E11.5 and E12.5 mouse embryos were collected into ice cold 1x phosphate-buffered saline (PBS) and fixed in 4% paraformaldehyde overnight at 4 °C. Embryos were dehydrated and paraffin embedded as previously described (Bushdid et al., 2003). 5μm mouse sections were deparaffinized, rehydrated, and primary antibody applied per manufacturerʼs protocol (Abcam). This procedure included antigen retrieval with Tris-EDTA pH9.0 buffer. Washing steps were performed using 1x Tris-buffered saline with 0.01% Tween 20 (TBST). Anti-phosphorylated

JNK(Thr183/ Tyr185) antibody (Abcam) was diluted in 1:400 in 0.5% blocking solution

(Pierce). For MF20 and pJNK labeling, secondary antibody application and HRP detection was performed according to manufacturerʼs instructions (using TBS) with the

Pierce Ultra-sensitive ABC Kit and DAB Enhanced Metal Substrate Kit (Pierce).

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ECC cell cultures and treatments

Prefused superior and inferior atrioventricular canal (AVC) ECCs were harvested from embryonic chicken hearts at HH25, as previously reported by Lincoln et al. (Lincoln et al., 2006), with the exception that the ECC cells were not dissociated before being placed into one well of a two-well collagen coated chamber slide. Six hearts were used for each experimental condition in duplicate. For RNA isolation in all experiments except where noted, ECC cells were incubated for 7 days. Recombinant human (rh)

VEGF-165 (R&D Systems) 50ng/mL, rhVEGFR1/Flt1/Fc Chimera (sFlt1)(R&D Systems)

50ng/mL, Cyclosporin A (CsA) (Novartis) 1μg/mL, rhRANKL (R&D Systems) 800ng/mL, rhOPG (R&D Systems) 1μg/mL, BSA (Sigma) 100-800ng/mL (as a vehicle control),

U0126 (Promega) 10μM, SP600125 (Calbiochem) 2.5μM, LY294002 (Calbiochem)

25μM, or Dimethyl sulfoxide (DMSO) (Sigma) 0.005-0.01% (as a vehicle control) was added to culture media at the time of dissection and replenished after 3 days. For immunofluorescence, ECCs were incubated 1-7days in M199 media (Cellgro) containing 10% Fetal Bovine Serum (Hyclone), 1% Penicillin-Streptomycin (Gibco), and

0.1% Chick Embryo Extract (Sera Labs International). For nuclear localization and proliferation studies, media was replaced after 24 hours with EGM-2MV-Microvascular

Endothelial Cell Medium-2 (Cambrex) with all SingleQuot additives except VEGF and incubated another 24h. For examination of VEGF and RANKL regulation of NFATc1 nuclear localization, ECCs were treated with rhVEGF-165 50ng/mL, sFlt1 50ng/mL, CsA

1μg/mL, BSA 100ng/mL for 20 minutes or rhRANKL 800ng/mL, rhOPG 1μg/mL, CsA

1μg/mL, BSA 800ng/mL for 30 minutes. Cultures with inhibitors (sFlt1, CsA, or OPG)

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were pre-treated for 2 hours prior to addition of rhVEGF or rhRANKL. To validate U0126 inhibition of VEGF-induced ERK1/2 activation and SP600125 inhibition of RANKL- induced JNK1/2 activation, ECCs were treated with rhVEGF-165 50ng/mL, U0126

(Promega) 10μM, DMSO 0.01% for 20 minutes or rhRANKL 800ng/mL, SP600125

(Calbiochem) 2.5μM, DMSO 0.005% for 30 minutes. Cultures with inhibitors (U0126 or

SP600125) were pre-treated for 2 hours prior to addition of rhVEGF or rhRANKL.

Cultures used to determine proliferation were treated for 24 hours with rhVEGF, rhRANKL, sFlt1, rhOPG, CsA or BSA. For each experiment, samples were collected in biological duplicate and data were collected from 3-6 independent experiments for each condition.

Immunofluorescence and scanning laser confocal microscopy

ECC cultures were fixed and prepared for confocal microscopy as detailed in Evans-

Anderson et al. (Evans-Anderson et al., 2008), with the following exception; fixed cultures were incubated for 10 minutes in 0.3% Triton-X in PBS and then washed 3x5 minutes with PBS before being incubated for 1hour in blocking solution (1% BSA, 0.1% cold water fish skin gelatin, 0.1% Tween-20, 0.05% NaN3 /PBS). Mouse monoclonal anti-NFATc1 (Santa Cruz) (1:100), rabbit polyclonal anti-phosphohistone H3 (Ser10)

(pHH3) (Upstate) (1:100), guinea pig anti-Sox 17 (generous gift from Dr. Jeffrey

Whitsett) (1:1000), mouse monoclonal anti-Cathepsin K (Santa Cruz) (1:100), rabbit polyclonal anti-Flk1 (Santa Cruz) (1:200), mouse monoclonal anti-MAP kinase, activated

(diphosphorylated ERK1/2) (Sigma clone MAPK-YT) (1:10,000), or rabbit polyclonal

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anti-phosphorylated JNK(Thr183/Tyr185) (Abcam) (1:400) primary antibodies were diluted in 1:1 blocking solution/PBS and incubated on slides overnight at 4ºC.

Corresponding Alexa-goat anti-mouse-488, Alexa-goat anti-guinea pig-488, Alexa- donkey anti-rabbit-488, Alexa-goat anti-guinea pig-568 and/ or Alexa-goat anti-rabbit-

568 (Molecular Probes) secondary antibodies were applied at a concentration of 1:200 in PBS for 1-3 hours. ToPro3 iodide nuclear stain (Molecular Probes)(1:1000 in PBS for

10minutes) was used to label cell nuclei. Immunofluorescence was detected using a

Zeiss LSM 510 confocal microscope and images were obtained using Zeiss LSM version 3.2 SP2 software. Ten random microscopic fields were imaged for each experimental condition and all images were captured in parallel using identical confocal laser settings, constant PMT filters and integration levels. Percent cells with nuclear

NFATc1 fluorescence was determined by dividing the number of cells with nuclear

NFATc1 labeling by the total number of NFATc1 positive cells per microscopic field.

The percent pHH3 positive cells was determined by dividing the number of pHH3 positive nuclei by total nuclei in a microscopic field. Data were collected from 3-6 independent experiments with biological duplicates for each condition.

RNA isolation and Real Time RT-PCR

ECC cell cultures were collected in 200μL Trizol reagent (Invitrogen), total RNA isolated, and cDNA generated using SuperScript II (Invitrogen) per manufacturerʼs protocol. 600ng of cDNA in Power SyberGreen Master Mix (ABI) was subjected to

Quantitative Real Time PCR (MJ Research, Opticon 2) analysis using the following

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20pmol primers: CtsK- 5ʼ-AAAGCAGTACAACGGCAAGG-3ʼ and 5ʼ-

GAGCTCACATCTTGGGGAAG-3ʼ; NFATc1- 5ʼ-CTCTGGAGAGCCCTAGAATTGA-3ʼ

and 5ʼ-CGCAGAAGTTTCCTTTCCTG-3ʼ. The identity of the CtsK and NFATc1 PCR

products was confirmed by sequencing. Amplification reactions were performed as 95º

5 minutes, (94º 30 seconds, 64º 30 seconds, 72º 30 seconds, plate read) x35 cycles,

72º 10 minutes, melting curve from 65º-95º read every 1º and hold 10 seconds, then 10º

5 minutes. Primers and reaction conditions used for GAPDH, β-actin, Periostin,

Versican, and Scleraxis were previously reported (Lincoln et al., 2006; Shelton and

Yutzey, 2007, 2008). Samples were run in triplicate and gene expression levels were determined as previously described by Lincoln et al. (Lincoln et al., 2006), with the standard curve for each primer set generated with HH34 whole heart cDNA. All reported values were normalized to corresponding GAPDH levels (Lincoln et al., 2006).

Gene expression levels were also confirmed by normalization to β-actin. For each

experiment, samples were collected in biological duplicates and data were collected

from three independent experiments for each condition.

In situ hybridization

The chicken CtsK/ JTAP1 (GenBank accession #NM_204971) and RANKL (GenBank

accession #NM_001083361) sequences were amplified from HH34 whole heart cDNA

using the following primers: CtsK- 5ʼ-AAAGCAGTACAACGGCAAGG-3ʼ and 5ʼ-

GAGCTCACATCTTGGGGAAG-3ʼ and RANKL- 5ʼ-ACACGCCCTTTGAAAATCAG-3ʼ

and 5ʼ-AATGCCCCAAAGTAAGTTGC-3ʼ. Sequences of CtsK (891bp) or RANKL

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(630bp) were ligated into pGEM-T vector (Promega) and confirmed by DNA sequencing. Vectors containing CtsK and RANKL sequence inserts were linearized with NcoI and DIG-labeled riboprobes were prepared using SP6 polymerase as previously described (Ehrman and Yutzey, 1999; Shelton and Yutzey, 2007). NFATc1 plasmid was a generous gift from Dr. D.W. Benson. Chicken NFATc1 sequence was generated using primers: 5ʼ-CTCTGAGTATTACCTGCCTCCA-3ʼ and 5ʼ-

CAATGAACAGCTGTAGCGTGAG-3ʼ and ligated into pBluescript SK+ vector. Vector containing the NFATc1 (1216bp) sequence was linearized with BamHI and DIG-labeled riboprobe was prepared as described above using T7 polymerase. In situ hybridization of chick tissue sections was performed as described in Shelton and Yutzey et al.

(Shelton and Yutzey, 2007). Proteinase K (20μg/mL) was applied to E5 sections for 5 minutes and E7 for 8 minutes. 4-Nitro blue tetrazolium chloride/ 5-Bromo-4-chloro-3- indolyl-phosphate solution (Roche) was applied to all sections in parallel with developing times of 30 minutes for CtsK, 45 minutes for RANKL and 60 minutes for NFATc1 chick heart sections.

Statistical analysis

Statistical significance was determined by Studentʼs t-test with p≤0.01 or p≤0.05 as indicated. Data are reported as a mean with standard error of the mean (SEM).

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References

Bushdid, P.B., Osinska, H., Waclaw, R.R., Molkentin, J.D., and Yutzey, K.E. (2003).

NFATc3 and NFATc4 are required for cardiac development and mitochondrial function.

Circ Res 92, 1305-1313.

Ehrman, L.A., and Yutzey, K.E. (1999). Lack of regulation in the heart forming region of avian embryos. Dev Biol 207, 163-175.

Evans-Anderson, H.J., Alfieri, C.M., and Yutzey, K.E. (2008). Regulation of cardiomyocyte proliferation and myocardial growth during development by FOXO transcription factors. Circ Res 102, 686-694.

Hamburger, V., and Hamilton, H.L. (1992). A series of normal stages in the development of the chick embryo. 1951. Dev Dyn 195, 231-272.

Lincoln, J., Alfieri, C.M., and Yutzey, K.E. (2006). BMP and FGF regulatory pathways control cell lineage diversification of heart valve precursor cells. Dev Biol 292, 292-302.

Ranger, A.M., Grusby, M.J., Hodge, M.R., Gravallese, E.M., de la Brousse, F.C., Hoey,

T., Mickanin, C., Baldwin, H.S., and Glimcher, L.H. (1998). The transcription factor NF-

ATc is essential for cardiac valve formation. Nature 392, 186-190.

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Shelton, E.L., and Yutzey, K.E. (2007). Tbx20 regulation of endocardial cushion cell proliferation and extracellular matrix gene expression. Dev Biol 302, 376-388.

Shelton, E.L., and Yutzey, K.E. (2008). Twist1 function in endocardial cushion cell proliferation, migration, and differentiation during heart valve development. Dev Biol

317, 282-295.

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Supplemental Figure Legends

Supplemental Figure 1. NFATc1 positive cells co-express endothelial markers, and VEGF induces proliferation of endothelial cells in cultured ECCs.

Immunofluorescence with confocal laser scanning microscopy (ICLSM) of cultured ECC cells. Aʼ-Aʼʼ) Coexpression of the endothelial marker Sox17 (pink) and NFATc1 (green) in cultured ECC cells (arrows). B-C) Coexpression of M-phase marker pHH3 (pink) and endothelial markers Sox17 (green) (B) and NFATc1 (green) (C) induced by VEGF treatment (arrows). D) Coexpression of Sox17 (pink) and CtsK (green) in ECC cells

(arrows). ToPro3 iodide nuclear stain was used to visualize cell nuclei (blue).

Supplemental Figure 2. Expression of endothelial and mesenchymal markers by cultured ECC cells over time mimics gene expression observed in maturing

ECCs/ mitral valves in vivo. A) Expression of endothelial (NFATc1, CtsK) and mesenchymal (Periostin, Versican, Scleraxis) marker mRNAs in 2, 7, and 10 day cultured ECC cells. B) Expression of endothelial (NFATc1, CtsK) and mesenchymal

(Periostin, Versican, Scleraxis) marker genes in avian ECCs/ mitral valves isolated at

E4.5, E7, and E10. Fold change in mRNA expression was determined by Real Time

RT-PCR and normalized to GAPDH levels. Data presented is representative of 3 independent experiments performed in biological duplicate with PCR performed in triplicate.

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Supplemental Figure 3. OPG does not inhibit VEGF-induced cell proliferation and sFlt1 does not inhibit RANKL-induced CtsK expression. A) Quantification of percent pHH3 positive nuclei in treated chick ECC cultures for three independent experiments per treatment (n=3). B) Quantification of fold difference in CtsK mRNA expression for treated avian ECC cultures was determined by real time RT-PCR.

Samples were run in triplicate and normalized to GAPDH mRNA expression for three independent experiments per treatment (n=3). *p≤0.01

Supplemental Figure 4. Percent diphosphorylated ERK1/2 positive cells is significantly reduced in cultures treated with U0126 compared to controls.

Percent phosphorylated JNK positive cells is significantly reduced in cultures treated with SP600125 compared to controls. A) Quantification of percent anti- diphosphorylated ERK1/2 (dpERK) positive cells per total cell nuclei for three independent experiments per treatment (n=3). *p≤0.01 B-D) Representative images showing anti-dpERK positive cells (arrows) in VEGF+DMSO (B), VEGF+U0126 (C), and

DMSO (D) treated ECC cultures. Green= dpERK and blue= ToPro3 iodide nuclear stain. E) Quantification of percent anti-phosphorylated JNK(Thr183/Tyr185) (pJNK) positive cells per total cell nuclei for three independent experiments per treatment (n=3).

*p≤0.01 F-H) Representative images showing anti-pJNK positive cells (arrows) in

RANKL+DMSO (F), RANKL+SP600125 (G), and DMSO (H) treated ECC cultures.

Green=pJNK and blue= ToPro3 iodide nuclear stain.

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Supplemental Figure 5. Model of NFATc1 function in the transition from ECC growth to valve remodeling. During ECC growth, VEGF/ NFATc1/ MEK1-ERK1/2 signaling promotes cell proliferation. As remodeling begins, RANKL/ NFATc1/ JNK1/2 signaling increases CtsK mRNA expression while inhibiting cell proliferation. These data support a novel mechanism for the transition from ECC growth to remodeling in which

NFATc1 promotes a sequential pattern of gene expression via cooperation with ligand- specific cofactors MEK1-ERK1/2 and JNK1/2.

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Chapter 3

NFATc1 promotes epicardium-derived cell (EPDC) invasion into myocardium*

Michelle D. Combs1, Caitlin M. Braitsch1, Alexander W. Lange2, Jeanne F. James1

Katherine E. Yutzey1*

1 Division of Molecular Cardiovascular Biology, Cincinnati Childrenʼs Hospital Medical

Center ML7020, 240 Albert Sabin Way, Cincinnati, Ohio, 45229, USA.

2 Division of Pulmonary Biology, Cincinnati Childrenʼs Hospital Medical Center, 240

Albert Sabin Way, Cincinnati, Ohio, 45229, USA.

*Submitted for publication to Development October 21, 2010

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Summary

Epicardium-derived cells (EPDCs) contribute to formation of coronary vessels and fibrous matrix of the mature heart. Nuclear Factor of Activated T-cells cytoplasmic 1

(NFATc1) is expressed in cells of the proepicardium (PE), epicardium, and EPDCs in mouse and chick embryos. Conditional loss of NFATc1 expression in EPDCs in mice causes embryonic death by E18.5 with reduced coronary vessel and fibrous matrix penetration into myocardium. In osteoclasts, calcineurin-mediated activation of NFATc1 by Receptor Activator of NFκB Ligand (RANKL) signaling induces Cathepsin K (CtsK) expression for extracellular matrix degradation and cell invasion. RANKL/NFATc1 pathway components also are expressed in EPDCs and loss of NFATc1 in EPDCs causes loss of CtsK expression in the myocardial interstitium in vivo. Likewise, RANKL treatment induces CtsK expression in PE-derived cell cultures via a calcineurin- dependent mechanism. In chicken embryo hearts, RANKL treatment increases the distance of EPDC invasion into myocardium, and this response is calcineurin- dependent. Together, these data demonstrate a critical role for the RANKL/NFATc1 signaling pathway in promoting invasion of EPDCs into myocardium by induction of extracellular matrix-degrading enzyme gene expression.

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Introduction

During cardiac morphogenesis cells of the PE migrate onto and envelop the myocardium to form the epicardium (Gittenberger-de Groot et al., 2010; Lie-Venema et al., 2007; Reese et al., 2002). A subset of epicardial cells undergo epithelial to mesenchymal transformation (EMT) and invade the subepicardial space and myocardium as EPDCs. EPDCs contribute to the formation of coronary vessels and the collagen-rich extracellular matrix (ECM) support scaffold for the cardiomyocytes called the fibrous matrix. EPDC invasion into subepicardium and myocardium is vital to proper coronary vessel and fibrous matrix patterning; however, molecular mechanisms that regulate this process remain largely unknown.

The PE is an outgrowth of extracardiac mesothelial cells overlying the septum transversum (mouse)/sinus venosus (chick) that migrates onto the myocardium to form the epicardium. Within the polarized epithelium of the epicardium, the orientation of cell division differentiates cells that will remain in the epicardium from cells that delaminate from the epicardium and undergo EMT to become mesenchymal EPDCs (Wu et al.,

2010). Epicardium cell EMT is mechanistically comparable to other EMT events, including in the endocardial cushions, with shared expression of signaling and cell adhesion marker genes (Olivey and Svensson, 2010; Reese et al., 2002). The PE and its derivative epicardium and EPDCs express transcription factors including Wilmʼs

Tumor 1 (WT1) and Tbx18 (Gittenberger-de Groot et al., 2010; Lavine and Ornitz, 2009;

Lie-Venema et al., 2007; Olivey and Svensson, 2010; Wu et al., 2010). After invasion of

EPDCs into myocardium, expression of WT1 and Tbx18 is downregulated as the cells

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differentiate into coronary smooth muscle, endothelia, and interstitial fibroblasts. In the

mature heart, EPDC derivatives are present in the ECM-rich fibrous matrix that provides

cardiac connective tissue as well as in the smooth muscle and endothelial cells of the

coronary vasculature (Gittenberger-de Groot et al., 2010; Lavine and Ornitz, 2009; Lie-

Venema et al., 2007; Reese et al., 2002).

NFATc1 is a transcription factor of the NFAT family that is regulated by calcium

(Ca2+) and the phosphatase calcineurin (Crabtree and Olson, 2002). During cardiac

development, NFATc1 is expressed by endocardium as well as endocardial cushion and

valve endothelial cells (de la Pompa et al., 1998; Ranger et al., 1998). Mice lacking

NFATc1 expression have normal endocardial cushion formation and EMT, but lack ECM

remodeling and CtsK expression necessary for valve maturation, causing embryonic lethality at embryonic day (E)12.5-E14.5 (Combs and Yutzey, 2009b; de la Pompa et

al., 1998; Lange and Yutzey, 2006; Ranger et al., 1998). NFATc1 is also required in

osteoclasts where it promotes ECM remodeling and invasion (Aliprantis et al., 2008;

Negishi-Koga and Takayanagi, 2009). To stimulate osteoclast function, RANKL binds to

its cognate receptor RANK leading ultimately to activation of calcineurin and nuclear

translocation of NFATc1 (Negishi-Koga and Takayanagi, 2009; Sitara and Aliprantis,

2010). NFATc1 promotes expression of an array of potent ECM-degrading enzymes

including CtsK (Negishi-Koga and Takayanagi, 2009). NFATc1-dependent ECM

degradation and osteoclast invasion are necessary for vascularization of bone and

collagen deposition for new bone formation (Motyckova and Fisher, 2002; Raggatt and

Partridge, 2010). Likewise, EPDC invasion is dependent upon ECM-remodeling.

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However, the role of NFATc1 signaling in EPDC invasion and maturation has not been investigated previously.

Materials and methods

Chicken and mouse embryo collection

NFATc1-/- and NFATc1(fl/fl) mouse lines were obtained from Dr. Laurie Glimcher of

Brigham and Womenʼs Hospital, Boston, Massachusetts (Aliprantis et al., 2008; Ranger et al., 1998). WT1-Cre mice were obtained from Dr. William Pu of Childrenʼs Hospital

Boston, Boston, Massachusetts (Zhou et al., 2008). GATA5-Cre mice were originally generated by the lab of Dr. John Burch and were obtained from Drs. Pilar Ruiz-Lozano of University of California San Diego and Vesa Kaartinen of University of Michigan

School of Dentistry, Ann Arbor, Michigan (Merki et al., 2005). Mouse embryos were generated via timed matings with observation of a copulation plug designated as E0.5.

Embryos that were alive and morphologically comparable to littermates were collected at E10.5, E11.5, E12.5, E13.5, E14.5 and E17.5. Genotyping for NFATc1 mutation and/or WT1-Cre expression was performed by PCR using primers and methods previously described (Aliprantis et al., 2008; Ranger et al., 1998; Zhou et al., 2008).

Postnatal genotyping was performed using DNA isolated from tail clips at postnatal day

(P)21. Fertilized white leghorn chicken eggs (Charles River Laboratories, CT, USA) were incubated at 38°C under high humidity and chicken embryos were sacrificed at E4,

E5, E7 and E14. All animal procedures were approved and performed in accordance with institutional guidelines.

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Chicken whole heart and PE cell culture

E7 chick whole hearts were collected and labeled with 25μM carboxyfluorescein diacetate succinimidyl ester (CFSE)(Invitrogen Life Tech, CA, USA) in M199 media for 1 hour. Hearts were cultured 18 hours in 0.01% BSA-coated chamber slides containing culture media (M199 (Cellgro Mediatech, Inc, VA, USA), 10% fetal bovine serum

(Hyclone Thermo Fisher Scientific, MA, USA), 1% chick embryo extract (Sera Labs

International, West Sussex, UK), and 1% penicillin-streptomycin (Invitrogen)). The culture media contained the following treatments; 800ng/mL recombinant human (rh)

RANKL (R&D Systems, MN, USA), 1μg/mL rhOPG (R&D), 1μg/mL Cyclosporin A (CsA)

(Novartis International AG, Basel, Switzerland), 800ng/mL BSA (Sigma-Aldrich, MO,

USA)(vehicle control). Four independent experiments were performed in biological triplicate for each treatment group (n=4).

Aggregated PE cells, identified based on cell morphology and morphological landmarks, were dissected from PE/atrioventricular groove region using tungsten needles (Schulte et al., 2007). The cell aggregates from 4 embryos were transferred to a 0.01% rat-tail collagen-coated chamber slide. Cells were cultured 4 days in culture media (described above) with treatments described above with a media change on day

3 of culture. Cells were collected, RNA isolated and Real Time RT-PCR for CtsK expression was performed as previously described (Combs and Yutzey, 2009b). Four independent experiments were performed in biological duplicate for each treatment type

(n=4).

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Immunofluorescence and laser scanning confocal microscopy (ICLSM)

Mouse and chick embryos were collected, fixed, dehydrated and paraffin-embedded as previously described (Shelton and Yutzey, 2007). 5μm sections were deparaffinized, rehydrated and antigen retrieval was performed using Antigen Unmasking Solution

(#H3300; Vector Labs, CA, USA). Sections were prepared for ICLSM as previously described using the following primary antibodies: mouse monoclonal anti-NFATc1

(1:100)(BD Biosciences, CA, USA), rabbit polyclonal anti-Wilmʼs Tumor 1

(1:100)(EMD/Calbiochem, CA, USA), mouse monoclonal anti-Cathepsin K

(1:100)(Santa Cruz Biotech., Inc., CA, USA), rabbit polyclonal anti-Cx40 (1:100)(Santa

Cruz), rabbit polyclonal anti-Col1a1(1:100)(Millipore, MA, USA) and Alexa Fluor conjugated secondary antibodies: goat anti-mouse-488, goat anti-rabbit-488, donkey anti-rabbit-568 (Invitrogen) (Combs and Yutzey, 2009b).

Preparation and processing of chick heart sections for ICLSM with goat polyclonal anti-Tbx18 antibody (Santa Cruz)(1:250) was performed as previously described (Christoffels et al., 2009). Antigen unmasking was performed under pressure for 3 minutes with Antigen Unmasking Solution (#H-3300 Vector Labs) per manufacturerʼs guidelines. Primary antibody labeling and tyramide signal amplification

(TSA) was performed with Renaissance TSA Fluorescence Systems Tyramide Signal

Amplification Kit (Perkin Elmer, MA, USA) per manufacturerʼs instructions.

Cultured primary chick PE cells were collected and processed for ICLSM as previously described (Combs and Yutzey, 2009b) using antibodies and dilutions listed

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previously. Cultured chicken whole hearts were collected and paraffin embedded as above with the exception that d-Limonene (Hemo-D, Fisher Scientific, PA, USA) was used instead of xylene. 5μm sections were deparaffinized with Hemo-D and rehydrated.

Sections were blocked using ImmunoPure ABC Ultra-Sensitive Peroxidase Mouse IgG

Staining Kit (Fisher Scientific) according to manufacturerʼs instructions. Primary antibody was applied overnight at 4°C (anti-CtsK 1:100)(Santa Cruz). Secondary antibody was applied per manufacturerʼs protocol. Alexa Fluor conjugated tertiary antibody, streptavidin anti-568 (Invitrogen)(1:100) was applied for 30 minutes.

For each independent experiment, immunofluorescence was detected using a

Zeiss LSM 510 confocal microscope and images were captured using Zeiss LSM version 3.2 SP2 software in parallel using identical confocal laser settings, constant

PMT filters and integration levels.

Quantification of ventricular Col1a1 immunofluorescence, percent myocardial penetration and EPDC migration

For quantification of the area of Col1a1 immunofluorescence per ventricular area, pictomicrographs of right ventricle, left ventricle, apex, interventricular septum, and atrioventricular valves were generated from at least 3 comparable non-consecutive sections of six WT1-Cre(+);NFATc1(fl/fl) embryos and six control littermates using methods described above for ICLSM. Ventricular area (in pixels) was measured using

Image J64 software. The pixel area above 42 fluorescent units in the green channel

(fluorescent pixels) in the outlined region was measured via Image J64. The fluorescent

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area was divided by the ventricular area to determine fluorescence per ventricular area.

The fluorescent area per ventricular area for control embryos was set to 1 to determine

the fold change for WT1-Cre(+);NFATc1(fl/fl) embryos.

To determine percent myocardial penetration of Col1a1 fluorescence, ICLSM pictomicrographs of right ventricle and apex were obtained as described above.

Ventricular and apex myocardial wall diameters were measured in an upper (basal), middle, and lower (apical) region of each image using Image J64 software. The images were converted to grayscale using Adobe Photoshop software then converted to a binary (black and white) using Image J64. The distance from epicardium to the deepest point of Col1a1 reactivity (distance penetration) along the ventricle/apex myocardial wall diameter was measured for the upper, middle and lower points on each image. The distance of penetration divided by the total ventricular/apex diameter was calculated as the percent myocardial penetration.

For quantification of EPDC migration in treated chicken embryo whole hearts, images were collected and a separate binary image was made as described above.

The distance each CFSE-labeled cell migrated away from the epicardium into the apex myocardium was measured in two comparable non-serial sections for each treatment group per independent experiment using Image J64 software. Average distance of

EPDC migration was determined by the total distance of cell migration divided by total number of cells that migrated away from the epicardium for each treatment group. Four independent experiments were performed in biological duplicate (n=4).

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Immunohistochemical analysis for intramyocardial vessel penetration

Immunohistochemistry (IHC) was performed using ImmunoPure ABC Ultra-Sensitive

Peroxidase Mouse IgG Staining Kit (Fisher Scientific) according to manufacturerʼs instructions. The following primary antibodies were applied overnight at 4°C; mouse monoclonal anti-NFATc1(1:100)(BD Biosciences), mouse monoclonal anti-

SMA(1:200)(Sigma-Aldrich) rabbit polyclonal anti-Cx40 (1:100)(Santa Cruz), mouse monoclonal MF20(1:200)(Developmental Studies Hybridoma Bank, IA, USA). Antigen retrieval was performed using Antigen Unmasking Solution (#H3300; Vector Labs).

Horseradish peroxidase(HRP) detection was performed using 3,3-

Diaminobenzidine(DAB) Enhanced Metal Substrate Kit (Pierce Biotech, Inc, IL, USA) per manufacturerʼs protocol.

Pictomicrographs were obtained with an Olympus BX51 microscope using Digital

Camera Systems Spot software version 4.5 and filter and exposure settings were unchanged for comparison between WT1-Cre(+);NFATc1(fl/fl) and control animals. The distance from the epicardium to the closest vessel wall (vessel penetration) was measured, and percent intramyocardial penetration was calculated as vessel penetration divided by ventricular diameter. Coronary vessels 2.33μm2 or greater were used for quantification. Images for quantification were collected from at least four non- consecutive comparable sections from six WT1-Cre(+);NFATc1(fl/fl) embryos and six control littermates (n=6 for each genotype).

Mouse embryo RNA isolation and Real Time RT-PCR

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E14.5 mouse ventricle and forelimb tissue was collected in 200μL Trizol reagent

(Invitrogen), RNA isolated, and 1μg total RNA was used for cDNA synthesis with

SuperScript II (Invitrogen) per manufacturerʼs protocols. Real Time PCR was performed

using Taqman gene expression assays (Applied Biosystems Life Tech, CA, USA) for

SMA (Assay ID: Mm01546133_m1) and CtsK (Assay ID: Mm00484039_m1) on

StepOnePlus Real-Time PCR System (Applied Biosystems Life Tech, CA, USA) per

manufacturerʼs instructions. β2-microglobulin (Assay ID: Mm00437762_m1) was used

for normalization, and these results were confirmed by normalization with β-actin (Assay

ID: Mm02619580_g1). Data were collected for four WT1-Cre(+);NFATc1(fl/fl) embryos

and four WT1-Cre(-);NFATc1(fl/wt) control littermates (n=4 for each genotype).

In situ hybridization

In situ hybridization using digoxygenin-labeled antisense RNA probes for mouse

NFATc1, RANKL, RANK and chick NFATc1, RANKL and CtsK were performed as

previously described (Combs and Yutzey, 2009b; Lange and Yutzey, 2006).

Statistical analysis

Statistical significance was determined by Studentʼs t-test with p≤0.01 as indicated.

Data are reported as a mean with standard error of the mean (SEM).

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Results

NFATc1 is expressed in the PE, epicardium, EPDCs and mature coronary vessels in vivo.

NFATc1 expression in the epicardium and epicardium-derived structures was visualized by immunofluorescence with confocal laser scanning microscopy (ICLSM) of

E10.5, E11.5, E14.5 and E17.5 mouse embryos as well as E4, E7 and E14 chick embryo heart sections. During epicardial formation and proliferation, NFATc1 protein is expressed in mouse PE cells as well as cells of the septum transversum and endocardium at E10.5 (Figure 1A,Aʼ). In E4 chick, NFATc1 protein is localized to epithelial cells of the PE, liver bud, epicardium and sinus venosus (Figure S1A). As development proceeds, NFATc1 is expressed by a subset of cells of the epicardium and

EPDCs of the subepicardial space, as they begin to invade the myocardium, at E11.5-

E14.5 in mouse and E7 in chick (Figure 1B,Bʼ,D and Figure S1B). NFATc1 also is expressed by endothelial cells of coronary vessels and aortic valve endocardium at

E17.5, just prior to birth (Figure 1C,Cʼ and Figure S1C). This pattern of NFATc1 protein expression by cells of the epicardium and EPDCs is consistent with mRNA expression in both chick and mouse hearts as determined by in situ hybridization (ISH) (Figure S2).

Significantly, NFATc1 is coexpressed with the transcription factors Wilmʼs Tumor 1

(WT1) (Figure 1D) and Tbx18 (Figure S1D-F) known to be expressed by epicardium and

EPDCs (Haenig and Kispert, 2004; Zhou et al., 2008). At E14.5 NFATc1 is localized primarily to the cell cytoplasm of epicardial epithelial cells. This is in contrast to nuclear

NFATc1 localization indicative of activation in EPDCs of the subepicardial space and in

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those invading the myocardium (Figure 1D). Together these data demonstrate that

NFATc1 is expressed by the PE, epicardium, EPDCs, and mature coronary endothelial

cells. In addition NFATc1 activation and nuclear localization occurs in EPDCs invading

the subepicardium and myocardium.

WT1-Cre(+);NFATc1(fl/fl) mice have fewer NFATc1 positive epicardial cells, fewer

NFATc1 positive EPDCs, and reduced or absent myocardial spaces.

NFATc1 function in epicardium and EPDCs was examined in mice with

conditional loss of NFATc1. Systemic loss of NFATc1 does not obviously affect the

initial stages of PE migration or epicardium formation (Figure S3). However, systemic

loss of NFATc1 results in embryonic lethality at E12.5-E14.5 due to defects in heart

valve remodeling (de la Pompa et al., 1998; Ranger et al., 1998). Therefore, mice with

conditional loss of NFATc1 in EPDCs were generated to determine the requirements for

NFATc1 in EPDC maturation and formation of coronary vessels and fibrous matrix.

WT1-Cre mice (WT1-Cre(+)) were bred with NFATc1-flox mice (NFATc1(fl/fl)) for

conditional loss of NFATc1 function in cells of the WT1 lineage (Aliprantis et al., 2008;

Zhou et al., 2008).

WT1-Cre(+);NFATc1(fl/fl) embryos are grossly indistinguishable from control

littermates of the genotypes WT-Cre(+);NFATc1(fl/wt), WT1-Cre(-);NFATc1(fl/fl) or

WT1-Cre(-); NFATc1(fl/wt) throughout development (Figure S4A-C, E-G). At a macroscopic level, the hearts of WT1-Cre(+);NFATc1(fl/fl) embryos are comparable to

controls in size and structure. However, there is some evidence for compromised heart

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function in retention of a W-shaped (bifid) apex beyond E14.5. While WT1-

Cre(+);NFATc1(fl/fl) embryos are collected at the expected Mendelian ratio of 25% at

E13.5-E14.5, WT1-Cre(+);NFATc1(fl/fl) embryos are underrepresented in those collected at E17.5-E18.5 (19%) and comprise only 3% of pups genotyped postnatally

(Figure S4I). In utero echocardiograms demonstrated that the majority of WT1-

Cre(+);NFATc1(fl/fl) embryos lose cardiac contractility and have no detectable heartbeat at E17.5-E18.5 (data not shown). Similar results were obtained with GATA5-

Cre(+);NFATc1(fl/-) mice generated in parallel (data not shown) (Merki et al., 2005).

Like WT1-Cre(+);NFATc1(fl/fl) mice, GATA5-Cre(+);NFATc1(fl/-) mice die prenatally with 0% being present for genotyping at postnatal day (P)21. These data indicate that loss of NFATc1 in EPDCs results in loss of cardiac function and prenatal lethality at

E17.5-E18.5 with variable penetrance.

Successful deletion of NFATc1 expression in epicardial cells/EPDCs of WT1-

Cre(+);NFATc1(fl/fl) mice is demonstrated by lack of anti-NFATc1 antibody labeling of epicardial cells and subepicardial EPDCs, as well as fewer NFATc1-positive intramyocardial EPDCs (Figure 2B, Bʼ). In contrast, NFATc1 positive EPDCs are seen in the subepicardium and scattered throughout the myocardium of control embryos (Figure

2A, Aʼ). NFATc1 protein expression is apparent in non-epicardium- derived tissues such as atrioventricular valve in both WT1-Cre(+);NFATc1(fl/fl) and control embryos as previously described (Figure 2 and Figure S5) (de la Pompa et al., 1998; Ranger et al.,

1998). A few NFATc1-positive cells are noted in the myocardium of WT1-

Cre(+);NFATc1(fl/fl) mice, either from incomplete Cre-mediated recombination or

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NFATc1 expression in WT1 non-expressing cells. These data demonstrate that WT1-

Cre(+);NFATc1(fl/fl) mice have loss of NFATc1 expression specifically in cells of the

WT1 lineage, including EPDCs, and can be used to determine the role of NFATc1 in

EPDC function.

At E17.5 normal ventricular myocardium is characterized by abundant myocardial spaces and blood vessels in both the shallow (outer) and deep (inner) myocardium. In contrast, ventricular myocardium of WT1-Cre(+);NFATc1(fl/fl) embryos is dense with cardiomyocytes, but few myocardial spaces are apparent (Figure 2). Previous studies have indicated a role for epicardium signaling in myocardium compaction (Olivey and

Svensson, 2010). However, a dense compacted myocardium is apparent in hearts of

WT1-Cre(+);NFATc1(fl/fl) embryos at E17.5, indicating that this process is unaffected by loss of NFATc1 function in cells of the WT1-Cre lineage (Figures 2,3 and Figure S4H).

Coronary vessels in WT1-Cre(+);NFATc1(fl/fl) embryos are formed in the atrioventricular canal (AVC), but are reduced in the deep myocardium (Figure 2).

Therefore, loss of NFATc1 expression in cells of the WT1-Cre lineage (EPDCs) results in reduced or absent myocardial spaces with dense myocardium, limited coronary vessel penetration, and prenatal lethality.

Loss of NFATc1 expression in EPDCs results in decreased cardiac fibrous matrix formation.

EPDCs that migrate into the myocardium represent the majority of cells that populate the ventricular wall with interstitial fibroblasts that synthesize the ECM of the

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fibrous matrix (Gittenberger-de Groot et al., 2010; Lie-Venema et al., 2007; Reese et al.,

2002). The formation of cardiac fibrous matrix was examined in E17.5 mouse heart

sections lacking epicardial NFATc1. Trichrome staining demonstrates that the fibrous

matrix (blue) of the heart is essentially absent in WT1-Cre(+);NFATc1(fl/fl) embryos

(Figure 3A2,A4,A6 and Figure S4,H). In addition, the apex in WT1-Cre(+);NFATc1(fl/fl) hearts is bifid, consistent with loss of fibrous matrix collagens in other mouse models

(Figure 3A6 and Figure S4H) (Lincoln et al., 2006). In contrast, abundant fibrous matrix

(blue) is seen throughout the myocardial interstitium in control littermates (Figure

3A1,A3,A5 and Figure S4D). As expected, collagen deposition (blue) in the atrioventricular valves of WT1-Cre(+);NFATc1(fl/fl) embryos is apparently unaffected, indicative of normal NFATc1 function in valves. Together, these data indicate that

NFATc1 function in EPDCs is necessary for formation of the fibrous matrix during cardiac development.

Because Collagen1 is a major component of the cardiac fibrous matrix, anti-

Collagen1a1 (Col1a1) antibody labeling and ICLSM were used to further demonstrate a specific loss of fibrillar Col1a1 in the ventricular myocardium of WT1-

Cre(+);NFATc1(fl/fl) embryos (Ott et al., 2008). WT1-Cre(+);NFATc1(fl/fl) embryos have

Col1a1-positive matrix deposition extending from the surface epicardium into the shallow myocardium, but have significantly reduced Col1a1 reactivity in deeper myocardial interstitium (Figure 3B2,B4,B6). In contrast, control embryos have anti-

Col1a1-positive matrix extending from the epicardium through the inner myocardium to the endothelium of the trabeculae (Figure 3B1,B3,B5). Quantification of percent

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penetration of Col1a1 immunoreactivity in the myocardium demonstrates that, in WT1-

Cre(+);NFATc1(fl/fl) embryos, Col1a1 protein extends only 22.5% of the distance

spanning the ventricular free wall from epicardium to trabeculae, and 17.4% at the apex

(Figure 3D). In contrast, control littermates had Col1a1 penetration of 89.0% and 90.4%

of total ventricular free wall and apex thickness, respectively. Quantification of total

Col1a1 protein per area also demonstrated WT1-Cre(+);NFATc1(fl/fl) embryo hearts

have a significant reduction in Col1a1 protein in the right ventricle, left ventricle, apex,

and interventricular septum compared to control littermates (Figure 3C). Importantly,

atrioventricular valves express high levels of Col1a1 and there is no significant

difference in Col1a1-positive immunofluorescence per atrioventricular valve area among

genotypes. These data demonstrate that loss of NFATc1 expression in EPDCs results

in decreased fibrous matrix synthesis and decreased fibrous matrix penetration into the

myocardium.

Loss of NFATc1 expression in EPDCs results in decreased penetration of coronary

vessels and fewer activated fibroblasts within the myocardium.

EPDC migration into myocardium is required for intramyocardial coronary vessel

development and NFATc1 is expressed in both EPDCs and maturing coronary vessels

(Figure 1 and Figures S1,S5) (Gittenberger-de Groot et al., 2010; Lie-Venema et al.,

2007; Majesky, 2004). EPDC invasion is also required for investment of the ventricular

myocardium with interstitial fibroblasts (Taylor et al., 2003). WT1-Cre(+);NFATc1(fl/fl) embryos were examined in order to determine the effects of loss of NFATc1 expression

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in EPDCs on interstitial fibroblast investment and coronary vessel formation. During

cardiac morphogenesis, EPDC-derived interstitial cells express markers associated with

fibroblast activation, such as Smooth Muscle α-Actin (SMA) (Haudek et al., 2009). IHC

with anti-SMA antibody was used to investigate the distribution of interstitial and

coronary smooth muscle cells in ventricular myocardium of WT1-Cre(+);NFATc1(fl/fl) and control embryos. Deep ventricular areas, towards the trabeculae, in WT1-

Cre(+);NFATc1(fl/fl) embryos have greatly reduced or absent activated fibroblasts and coronary smooth muscle cells (Figure 4B,F). However, both of these cell types are observed in the shallow myocardium of the right ventricular free wall. In contrast, SMA- positive cells are prevalent throughout right ventricular and interventricular septum myocardium of control embryos at E17.5 (Figure 4A,E). Thus, WT1-Cre(+);NFATc1(fl/fl) embryos have reduced interstitial fibroblast and coronary smooth muscle cell penetration into myocardium, while cell differentiation is apparently unaffected. Real

Time RT-PCR demonstrates that, as early as E14.5, WT1-Cre(+);NFATc1(fl/fl) embryos have reduced SMA expression in ventricular myocardium, compared to control littermates, while SMA expression in the forelimbs in unaffected (Figure 4J). Together, these data demonstrate that loss of NFATc1 function in EPDCs results in reduced numbers of activated fibroblasts throughout the myocardium and is consistent with the observed reduction in fibrous matrix deposition.

The majority of differentiated coronary vessel cells arise from EPDCs; therefore,

EPDC invasion into myocardium is required for coronary vessel formation and is vital to proper heart development and myocyte survival (Gittenberger-de Groot et al., 2010;

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Lavine and Ornitz, 2009; Lie-Venema et al., 2007). Coronary vessel formation was assessed in mice lacking NFATc1 expression in EPDCs. IHC and ICLSM in conjunction with antibody labeling with anti-SMA and anti-Connexin 40 (Cx40) was used to detect smooth muscle (SMA) and endothelial (Cx40) cells of the coronary vessels. Coronary vessels containing smooth muscle and endothelial components are apparent in the shallow myocardium of WT1-Cre(+);NFATc1(fl/fl) embryos at E17.5. However, penetrant vessels are not apparent in deeper myocardial areas (Figure 4B,D,F,H). In contrast, coronary vessels comprised of both smooth muscle and endothelial cells are found in both shallow and deep myocardial areas in control littermates (Figure

4A,C,E,G). Quantification of intramyocardial vessel penetration demonstrates that WT1-

Cre(+);NFATc1(fl/fl) embryos have significantly shallower placement of coronary vessels within the myocardium compared to control littermates (Figure 4I). There was no significant difference in the total number of coronary vessels assessed per heart section among genotypes. It is important to note that while location is affected in WT1-

Cre(+);NFATc1(fl/fl) embryos, loss of NFATc1 expression in EPDCs does not prevent differentiation of coronary smooth muscle or endothelial cells (Figure 4 and Figure S5).

In addition, there was no difference in the number of apoptotic cells among genotypes at

E14.5 or E17.5 as determined by TUNEL assay (data not shown). Together, these data demonstrate that loss of NFATc1 in EPDCs leads to reduced penetration of EPDCs into myocardium necessary for coronary vessel and fibrous matrix formation. However, loss of NFATc1 does not appear to affect total coronary vessel number, EPDC cell differentiation, or cell survival.

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WT1-Cre(+);NFATc1(fl/fl) embryos have significantly reduced CtsK expression in the myocardial interstitium in vivo, and RANKL treatment increases CtsK expression in cultured PE cells via a calcineurin-dependent mechanism.

The RANKL/NFATc1 signaling pathway is active in osteoclasts where it is required for cell invasion via induction of the ECM degrading enzyme CtsK (Negishi-

Koga and Takayanagi, 2009). To determine if RANKL/NFATc1 pathway components are expressed in a manner consistent with function in EPDC invasion into the myocardium, in situ hybridization using probes for NFATc1, RANKL, RANK, and CtsK mRNA was performed on mouse and chick heart sections. NFATc1, RANKL, and

RANK mRNAs are expressed by EPDCs in the subepicardium and myocardium of

E13.5 mouse (Figure 5D,E,F). Additionally, NFATc1, RANKL, and CtsK mRNAs are expressed by EPDCs in the subepicardium and myocardium of E7 chick (Figure

5A,B,C). EPDC expression of RANKL/NFATc1 pathway components is seen in addition to previously reported expression by endocardial cushion endothelial cells (Figure 5)

(Combs and Yutzey, 2009b; Lange and Yutzey, 2006). This spatiotemporal expression pattern is consistent with RANKL/NFATc1 function in EPDC invasion of the subepicardial space and myocardium.

NFATc1 function is required for CtsK expression in endocardial cushion endothelial cells and osteoclasts (Lange and Yutzey, 2006; Negishi-Koga and

Takayanagi, 2009). Anti-CtsK antibody labeling and ICLSM was used to determine that

Ctsk expression is greatly reduced or absent in the myocardium of WT1-

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Cre(+);NFATc1(fl/fl) embryos (Figure 6B,D). In contrast, CtsK is abundantly expressed in the ventricular myocardium of control embryos during EPDC invasion at E14.5 (Figure

6A,C). Real Time RT-PCR also demonstrates significantly reduced CtsK transcript levels in ventricles of WT1-Cre(+);NFATc1(fl/fl) embryo hearts, compared to control littermates at E14.5, while CtsK expression in forelimbs is not significantly different among genotypes (Figure 6E). The coexpression of RANKL/NFATc1 pathway components during EPDC invasion, combined with the loss of CtsK expression in mice lacking NFATc1 in EPDCs, is evidence for RANKL/NFATc1 pathway function as a molecular mechanism promoting EPDC invasion into myocardium.

The ability of RANKL activation of NFATc1 to promote CtsK gene expression was determined using cultured primary avian PE-derived cells. PE cells from E4 chick embryos were explanted and subjected to treatment with recombinant human (rh)

RANKL, Cyclosporin A (CsA) a pharmaceutical calcineurin inhibitor, bovine serum albumin (BSA) as a vehicle control, or rhRANKL+CsA. Cultured PE cells maintain expression of epicardial markers such as WT1, Tbx18, and NFATc1 over the 4-day culture period, and few contaminant MF20-positive myocytes were observed (Figure

S6). Treatment of cultured PE cells with rhRANKL significantly increased CtsK mRNA expression as detected by Real Time RT-PCR (Figure 6F). In these experiments, rhRANKL treatment leads to a 5-fold induction of CtsK expression; however, rhRANKL- induced CtsK expression is blocked by addition of the calcineurin inhibitor Cyclosporin A

(CsA). Thus rhRANKL activates CtsK gene expression in cultured PE cells by a calcineurin-dependent mechanism. Addition of CsA alone to PE cells significantly

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reduces CtsK expression below the level of transcription detected in control BSA-treated cells, suggesting that PE-derived cells have a basal level of calcineurin-induced CtsK expression. Together with the observation that CtsK expression and EPDC invasion is reduced in mice with EPDC-specific loss of NFATc1, these data support a mechanism whereby calcineurin-mediated RANKL activation of NFATc1 promotes CtsK expression required for ECM degradation and EPDC invasion into myocardium.

RANKL treatment promotes EPDC invasion into myocardium and CtsK expression via a calcineurin-dependent mechanism in cultured chick whole hearts.

The ability of RANKL to promote NFATc1 activation and EPDC invasion in vivo was examined using cultured chicken embryo whole hearts. Whole hearts were isolated at E7 and the epicardium labeled with CFSE. Whole hearts were then cultured for 18 hours in the presence of rhRANKL, rhOsteoprotegrin (rhOPG) a soluble RANK receptor

(RANK antagonist), CsA, BSA as a vehicle control, rhRANKL+rhOPG, or rhRANKL+CsA. Hearts were then paraffin embedded and sectioned for antibody labeling and ICLSM. EPDC invasion was quantified by measuring the distance CFSE- labeled EPDCs migrated away from the epicardium and into the myocardium. Addition of rhRANKL to the media of cultured hearts significantly increased the distance of EPDC invasion into the myocardium (Figure 7). In these studies, addition of rhRANKL+ CsA to the media inhibited the effects of rhRANKL-induced EPDC invasion, demonstrating that rhRANKL promotes EPDC invasion via a calcineurin-dependent mechanism. Addition of rhRANKL+OPG, CsA alone, or OPG alone significantly decreased EPDC invasion

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compared to BSA treated controls. This suggests a basal level of endogenous

RANKL/NFAT signaling occurs in control hearts that is required for EPDC invasion.

Although rhRANKL increases EPDC migration distance, there was no difference among treatments in the number of EPDCs that migrated away from the epicardium.

Therefore, manipulation of RANKL/NFAT signaling in cultured hearts did not affect epicardium cell EMT or proliferation. These data demonstrate that rhRANKL promotes

EPDC invasion into the myocardium via a calcineurin-dependent mechanism.

In order to determine if manipulation of RANKL/NFAT signaling in EPDCs affects

CtsK expression, ICLSM with anti-CtsK antibody was performed on CFSE-labeled cultured chick hearts subjected to altered RANKL/calcineurin signaling. Addition of rhRANKL to the media of cultured hearts results in increased CtsK expression compared to BSA-treated controls (Figure 7Bʼ). Addition of CsA in combination with rhRANKL restores CtsK immunoreactivity to levels comparable to BSA-treated controls

(Figure 7Dʼ). Addition of CsA alone to cultured hearts results in CtsK protein expression below basal levels (Figure 7Cʼ). Thus, rhRANKL treatment increases CtsK expression via a calcineurin-dependent mechanism in intact hearts. Taken together these data support a mechanism whereby RANKL/NFATc1 signaling promotes EPDC invasion via

CtsK gene induction.

Discussion

Here we demonstrate that loss of NFATc1 in EPDCs results in decreased coronary vessel and fibrous matrix penetration into ventricular myocardium leading to embryonic

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death in mice. We further show that RANKL activation of NFATc1 promotes CtsK

expression and invasion of EPDCs into myocardium. This study is the first to report a

molecular mechanism for promoting EPDC invasion of myocardium necessary for

coronary vessel and fibrous matrix development.

NFATc1 is expressed by PE and epicardial cells that will undergo EMT to

generate EPDCs and also is expressed by endocardial cushion endothelial cells that will

undergo EMT to populate the endocardial cushions (de la Pompa et al., 1998; de Lange

et al., 2004; Lincoln et al., 2004; Ranger et al., 1998). Previous studies demonstrate

that NFATc1 function is not required for endocardial cushion formation or EMT,

however, expression of NFATc1 by endocardial cushion endothelial cells is required for

later processes of valve remodeling (de la Pompa et al., 1998; Lange and Yutzey, 2006;

Ranger et al., 1998). Likewise, data reported here demonstrate that NFATc1 function is

not required for initial stages of epicardium formation or EMT, but is necessary for ECM

remodeling associated with EPDC invasion.

WT1-expressing EPDCs differentiate into coronary endothelial cells, coronary

smooth muscle cells and interstitial fibroblasts (Zhou et al., 2008; Zhou et al., 2010).

Coronary endothelial, smooth muscle, and interstitial fibroblast differentiation is

observed in mice lacking NFATc1 expression in cells of the WT1 lineage. Therefore,

NFATc1 function is not required for differentiation of these cell lineages during heart

development. The observed loss of NFATc1 expression in endothelial cells of the

coronary vessels in WT1(+);NFATc1(fl/fl) embryos demonstrates that WT1-expressing cells contribute to this lineage in vivo. There is some controversy as to the specific

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origins of coronary endothelial cells, and it is possible that the WT1-expressing coronary endothelial cells observed may be from a non-PE origin as suggested by Red-Horse and colleagues (Norden et al., 2010; Red-Horse et al., 2010). Previous studies by Zeini et al demonstrate a requirement for calcineurin-NFAT signaling in endothelial cells for coronary angiogenesis during vascular plexus formation (Zeini et al., 2009). Our data demonstrate that NFATc1 is not specifically required for this process, however, our observations do not preclude the necessity for other calcineurin targets, including other

NFAT family members, in vascular plexus formation. Our studies, taken together with previous work, demonstrate a conserved NFATc1-dependent mechanism for invasion of myocardium by all EPDC progenitor cell types including those of coronary vasculature.

RANKL/NFATc1 pathway function promotes EPDC invasion into myocardium via induction of ECM remodeling enzymes such as CtsK. RANKL/NFATc1 pathway function is required for ECM remodeling by a variety of cell types during development including osteoclasts, cardiac valve endothelial cells, and EPDCs (Combs and Yutzey,

2009a; Negishi-Koga and Takayanagi, 2009). In osteoclasts, NFATc1 activation via

RANKL signaling upregulates several ECM degrading enzymes, including MMP9,

MMP13 and CtsK, which leads to formation of lacunae and cell invasion by ECM remodeling (Karsenty et al., 2009; Sitara and Aliprantis, 2010). Previous studies demonstrate that NFATc1 directly regulates expression of ECM remodeling enzymes

CtsK and ADAMTS9 (Sitara and Aliprantis, 2010). Since mice lacking CtsK expression have no reported cardiac abnormalities, it is likely that multiple ECM remodeling enzymes are regulated by NFATc1 in EPDCs (Chang et al., 2004; Funicello et al., 2007;

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Saftig et al., 1998). This is supported by the observation that loss of NFATc1 in EPDCs or valve endothelial cells leads to ECM remodeling defects resulting in embryonic lethality (de la Pompa et al., 1998; Ranger et al., 1998). Together, these studies indicate that NFATc1 is a major regulator of ECM remodeling and cell invasion in multiple cell types.

CtsK is a cysteine protease linked to physiologic and pathologic cell migration/invasion via ECM degradation (Negishi-Koga and Takayanagi, 2009; Onishi et al., 2010; Rapa et al., 2006). CtsK is a downstream target of calcineurin/NFATc1 activation in multiple cell types including osteoclasts, valve endothelial cells and EPDCs

(Lange and Yutzey, 2006; Negishi-Koga and Takayanagi, 2009). Cathepsins play an important role in ECM degradation and vascular cell invasion during development, wound healing, and cancer metastasis (Lutgens et al., 2007; Onishi et al., 2010; Saftig et al., 1998). The mode of action for CtsK is well described in bone, where osteoclasts secrete CtsK to degrade collagen-rich ECM, forming lacunae through which vasculogenic progenitor cells migrate (Karsenty et al., 2009; Motyckova and Fisher,

2002). EPDC migration through lacunae in the myocardial interstitium has been previously described, but the origin of these intramyocardial spaces is unknown

(Gittenberger-de Groot et al., 2010; Lie-Venema et al., 2007; Reese et al., 2002). Lack of intramyocardial spaces in WT1-Cre(+);NFATc1(fl/fl) embryos suggests that these lacunae could be formed via NFATc1-dependent ECM remodeling during EPDC invasion. Together, these studies support a mechanism whereby remodeling enzymes,

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including CtsK, create spaces in the myocardium necessary for EPDC migration and investment during coronary vessel and fibrous matrix development.

EPDCs are a heterogeneous population of cells that form coronary vessels and fibrous matrix, are necessary for Purkinje fiber differentiation, and support cardiomyoctyes by secreting promitotic factors (Gittenberger-de Groot et al., 2010;

Lepilina et al., 2006; Smart et al., 2007). Recent evidence also indicates that a subset of EPDCs differentiate into cardiomyocytes during development (Cai et al., 2008; Zhou et al., 2008). EPDC dysfunction has been linked to adult cardiac disease, Ebsteinʼs malformation, arrhythmia and cardiomyopathies in humans (Gittenberger-de Groot et al., 2010). It has been demonstrated that adult EPDCs have the potential to reactivate patterns of developmental gene expression and aid in neovascularization and cardiomyocyte survival after ischemic injury (Smart et al., 2007). Because of their regenerative potential, EPDCs hold promise as a source of progenitor cells in adult human hearts and may be useful for therapeutic treatments. Data presented here demonstrate that regardless of terminal cell fate, EPDCs share a common NFATc1- dependent mechanism that is required for invasion into myocardium. Therefore, this signaling mechanism could potentially be exploited in the development of EPDC-based therapies by increasing EPDC migration into myocardium for cardiomyocyte support and formation of EPDC derivatives.

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Acknowledgments

We thank Vicky Moore and Christine Schulte for technical assistance with fetal echocardiography and Dr. Elaine Wirrig for technical assistance with immunofluorescent antibody labeling.

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Figure legends

Figure 1. NFATc1 is expressed by the PE and coronary vessels and colocalizes with WT1 in the epicardium and EPDCs. A-C) ICLSM was performed on mouse heart sections using anti-NFATc1 antibody (green) and ToPro3 iodide nuclear stain (blue). A-

Aʼ) NFATc1 positive cells are apparent in the PE (arrow) at E10.5. Aʼ,Bʼ,Cʼ are magnified views of the boxed regions in A,B,C. B-Bʼ) NFATc1 positive epicardial cells

(arrows) and endocardial cells (arrowhead) at E11.5 are indicated. C-Cʼ) Anti-NFATc1 labeling of coronary endothelial cells (arrow) is shown. D-D”) ICLSM performed on

E14.5 mouse heart section using anti-NFATc1 antibody (green) and anti-WT1 antibody

(red) is shown. D-Dʼ) NFATc1 (green) and WT1 (red) positive cells of the epicardium

(arrow) and EPDCs of the AVC (arrowhead) and right ventricle (*) are indicated. D”)

Merged image with a magnified view of the AVC to highlight colocalization of NFATc1 and WT1 labeling in EPDCs. ST=septum transversum, EC=endocardial cushions,

AoV=aortic valve, PE=proepicardium, Epi=epicardium, Endo=endocardium,

CV=coronary vessel, AVC=atrioventricular canal, RV=right ventricle.

Figure 2. WT1-Cre(+);NFATc1(fl/fl) embryos have reduced NFATc1 positive epicardial cells and EPDCs in addition to increased myocardial compaction. IHC was performed on E17.5 mouse heart sections with anti-NFATc1 antibody (brown). A-

Aʼ) Control WT1-Cre(-); NFATc1(fl/wt) embryo section with NFATc1 positive cells of the epicardium and EPDCs of the subepicardium and myocardium (arrows in Aʼ magnified view of boxed area in A) indicated. NFATc1 positive mitral valve and endocardial cells

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(arrowheads) are shown. Abundant myocardial spaces (*) are seen throughout the

control myocardium as indicated. B-Bʼ) WT1-Cre(+);NFATc1(fl/fl) embryo section with

reduced (black arrow in Bʼ) or absent (red arrow Bʼ) NFATc1 staining of the epicardium

and EPDCs is shown. NFATc1 reactivity is apparent in the mitral valve (arrowheads B

and Bʼ) of WT1-Cre(+);NFATc1(fl/fl) embryo heart section. Note that myocardial spaces are greatly reduced or absent in WT1-Cre(+);NFATc1(fl/fl) myocardium (Bʼ). MV=mitral

valve.

Figure 3. WT1-Cre(+);NFATc1(fl/fl) embryos lack interstitial fibrous matrix and

have reduced penetration of Collagen 1a1 expressing cells. A1-6) Trichrome

staining performed on E17.5 control and WT1-Cre(+);NFATc1(fl/fl) embryo heart

sections. A1,3,5) Fibrillar collagen (blue staining) is seen throughout the myocardium

and tricuspid valve (arrowhead) of control WT1-Cre(-);NFATc1(fl/fl) embryo. A2,4,6)

WT1-Cre(+);NFATc1(fl/fl) embryo lacks blue fibrillar collagen staining in myocardium and has a bifid apex (*). Fibrillar collagen in the tricuspid valve (arrowhead) is indicated.

B1-6) ICLSM performed with anti-Col1a1 antibody (green) and ToPro3 iodide nuclear stain (blue) on E17.5 mouse heart sections. B1,3,5) Col1a1 positive matrix is present throughout the myocardium (bracket) and in the tricuspid valve (arrowhead) of a control

WT1-Cre(+);NFATc1(fl/wt) embryo. B2,4,6) Col1a1 positive matrix extends into the shallow myocardium as indicated by solid bracket. Deeper myocardial areas lack

Col1a1 reactivity (dashed bracket) in WT1-Cre(+);NFATc1(fl/fl) embryo. Col1a1 reactivity in the tricuspid valve (arrowhead) is retained and epicardial blebbing is noted

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(B4 arrow) in the WT1-Cre(+);NFATc1(fl/fl) embryo. C-D) Quantification of fold change

in fluorescence per area (C) and percent myocardial penetration of Col1a1 florescence

(D) was determined in six embryos of each genotype (n=6). *p≤0.01. TV=tricuspid valve,

RV=right ventricle.

Figure 4. WT1-Cre(+);NFATc1(fl/fl) embryos have reduced investment of activated

fibroblasts, and reduced intramyocardial vessel penetration. IHC performed on

E17.5 mouse heart sections using anti-SMA (A,B,E,F) or anti-Cx40 (C,D,G,H)

antibodies (brown). A,E) SMA positive coronary vessels (A arrows) and activated

fibroblasts (B inset arrowheads) in control WT1-Cre(+);NFATc1(fl/wt) embryo are indicated. B,F) SMA positive coronary vessels (B arrows) in shallow but not deep myocardium and lack of intramyocardial activated fibroblasts (F inset arrowhead) are apparent in the WT1-Cre(+);NFATc1(fl/fl) embryo. C,G) Cx40 positive coronary vessels

(C arrows) and IVS vessels (arrowhead G inset) seen in control WT1-

Cre(+);NFATc1(fl/wt) embryo. D,H) Shallow Cx40 positive vessels (D arrows) and lack of IVS vessels (H inset) are depicted in a WT1-Cre(+);NFATc1(fl/fl) embryo. I)

Quantification of percent intramyocardial penetration of coronary vessels was determined in six embryos of each genotype (n=6). J) Real Time RT-PCR quantification of fold change in SMA expression for four E14.5 embryos of each WT1-Cre(-

);NFATc1(fl/wt) control and WT1-Cre(+);NFATc1(fl/fl) genotype (n=4). *p≤0.01.

RV=right ventricle, IVS= interventricular septum.

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Figure 5. RANKL/NFATc1 pathway components are expressed in mouse and chick embryos during EPDC invasion. A-F) In situ hybridization was performed on

E7 chick (A-C) and E13.5 mouse (D-F) heart sections using antisense riboprobes. A,D)

NFATc1 mRNA expression in EPDCs (arrowheads) and valve endothelial cells (arrows) is indicated. B, E) RANK ligand (B,E) and receptor (F) are expressed by EPDCs

(arrowheads) and valve endothelial cells. C) CtsK mRNA is expressed by avian EPDCs

(arrowheads) and valve endothelial cells (arrow).

Figure 6. WT1-Cre(+);NFATc1(fl/fl) embryos have reduced CtsK expression in the myocardial interstitium. ICLSM was performed on E14.5 mouse heart sections with anti-CtsK antibody (green). A,C) CtsK expression in control WT1-Cre(+);NFATc1(fl/wt) embryo (arrows) is depicted. B,D) CtsK expression is reduced in a WT1-

Cre(+);NFATc1(fl/fl) embryo (arrow). E) Fold change in CtsK mRNA expression was quantified using Real Time RT-PCR for four E14.5 embryos of each WT1-Cre(-

);NFATc1(fl/wt) control and WT1-Cre(+);NFATc1(fl/fl) genotype (n=4). F) CtsK expression was assessed in primary chick PE cell cultures treated with rhRANKL, CsA, or rhRANKL+CsA. Fold change in CtsK was quantified using Real Time RT-PCR from four independent experiments performed in duplicate (n=4). *p≤0.01. LV=left ventricle.

Figure 7. RANKL increases EPDC migration distance and CtsK expression via a

Calcineurin/NFAT-dependent mechanism. E7 chick whole hearts were treated with

CFSE to label the epicardium then cultured with addition of BSA (A-A”), RANKL (B-B”),

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CsA (C-C”) or RANKL+CsA (D-D”) to media. ICLSM was performed to detect CFSE

(green), CtsK (red) and ToPro3 iodide nuclear stain (blue). CFSE-labeled EPDCs

(arrows) invade the myocardium and express CtsK as indicated. CtsK expression by

CFSE-labeled EPDCs (yellow color on merged section in B”) is indicated (arrowheads).

E) Quantification of average distance of EPDC migration into myocardium in μm for each treatment in four independent experiments (n=4). *p≤0.01.

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Supplemental figures

Figure S1: NFATc1 is expressed by PE, epicardium, EPDCs and coronary

vessels in chick demonstrating a conserved expression pattern in vertebrates.

ICLSM was performed on chick whole embryo and heart sections using anti-NFATc1

(green A-D), anti-WT1 (green E), anti-Tbx18 (green F) and ToPro3 iodide nuclear stain

(blue). A) NFATc1 positive PE and epicardium at E4 are indicated. B) NFATc1

reactivity is apparent in the epicardium and in the EPDCs present in the subepicardium

and myocardium at E7 as shown. C) NFATc1-positive coronary endothelial cells are

seen at E14 in chick embryo heart sections. D-F) E7 chick heart sections demonstrating

comparable patterns of NFATc1 (D), WT1 (E) and Tbx18 (F) expression in epicardial

cells and EPDCs. PE=proepicardium, SV=sinus venosus, LV=left ventricle,

CV=coronary vessel, AVC=atrioventricular canal, Epi=epicardium.

Figure S2. NFATc1 mRNA is expressed in epicardial cells and EPDCs of chick and mouse. In situ hybridization was performed on chick (A-Bʼ) and mouse (C-Dʼ) heart sections using anti-NFATc1 riboprobe (blue). A-Aʼ, C-Cʼ) NFATc1 mRNA is expressed in epicardium (arrow) and EPDCs of E5 chick and E12.5 mouse. Aʼ and Cʼ are magnified views of the boxed regions in A and C. Note NFATc1 expression is apparent in endocardial cushion (EC) endothelial cells (arrowhead) as previously reported. B-Bʼ, D-Dʼ) NFATc1 mRNA expression in epicardium (arrow) and EPDCs of

E7 chick and E14.5 mouse is shown. Bʼ and Dʼ are magnified views of the regions indicated in B and D. Note expression of NFATc1 in mitral valve (MV) endothelial cells

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(arrowhead) as previously reported. EC=endocardial cushion, Epi=epicardium,

MV=mitral valve, LV=left ventricle.

Figure S3. An intact epicardium is apparent with systemic loss of NFATc1.

A-B) Immunohistochemistry was performed on NFATc1+/+ (A) and NFATc1-/- (B) E10.5

mouse embryo heart sections with MF20 antibody. The epicardium (arrow) is apparent

as an MF20-negative epithelium covering the MF20-positive myocardium and is present

in both NFATc1+/+ and NFATc1-/- embryos. Insets are magnified views of boxed

regions in A and B. Epi=epicardium.

Figure S4. Control and WT1-Cre(+);NFATc1(fl/fl) embryos are grossly

indistinguishable until E17.5 when WT1-Cre(+);NFATc1(fl/fl) hearts show signs of

failure. WT1-Cre(+);NFATc1(fl/fl) hearts lack fibrillar collagen deposition in

myocardium and die at late embryonic stages. A,E) Pictomicrographs of E13.5

control WT1-Cre(-);NFATc1(fl/wt) (A) and WT1-Cre(+);NFATc1(fl/fl) (E) hearts. B,F)

E14.5 control WT1-Cre(+);NFATc1(fl/wt) (B) and WT1-Cre(+);NFATc1(fl/fl) (F) embryos

are shown. C,G) E17.5 control WT1-Cre(-);NFATc1(fl/wt) (C) and WT1-

Cre(+);NFATc1(fl/fl) (G) hearts are shown. D,H) Trichrome staining performed on E17.5

control WT1-Cre(-);NFATc1(fl/fl) (D) and WT1-Cre(+);NFATc1(fl/fl) (H) heart sections is depicted. * Denotes bifid apex of WT1-Cre(+);NFATc1(fl/fl) embryos in G-H. I) The percent of the total number of individuals recovered for each genotype is indicated for

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each timepoint. The expected Mendelian ratio for each genotype at each time point is

25%.

Figure S5. Coronary endothelial cell differentiation is apparent with loss of

NFATc1 expression in EPDCs. A-D) ICLSM performed on E17.5 mouse heart

sections using anti-NFATc1 (green), anti-Cx40 (red) antibodies and ToPro3 iodide

nuclear stain (blue). A-B) NFATc1 positive tricuspid valve endothelial cells are seen in

control WT1-Cre(-);NFATc1(fl/fl) (A) and WT1-Cre(+);NFATc1(fl/fl) (B) hearts. C)

NFATc1 and Cx40 expression are colocalized in coronary endothelial cells of control heart (arrow). NFATc1 positive EPDCs are present in the myocardium (arrowheads) in control hearts. D) Coronary endothelial cells (arrow) lacking NFATc1 express Cx40.

Decreased NFATc1 positive EPDCs (arrowheads) are apparent in the myocardium of

WT1-Cre(+);NFATc1(fl/fl) hearts. TV=tricuspid valve, LV=left ventricle.

Figure S6. Cultured chick PE cells express EPDC markers. ICLSM was performed on chick PE cells cultured 4 days. A) Anti-WT1 reactive cells (green and arrow in inset) and WT1 negative cells (arrowhead in inset) are indicated. B) Tbx18 positive cells

(green and arrows in inset) and negative cells (arrowhead in inset) are apparent. C)

NFATc1 positive cells (green and arrows in inset) and negative cells (arrowhead in inset) are shown. D) Low levels of contaminant myocardium are indicated by MF20 cells (green arrow). The vast majority of cells in culture are non-reactive PE cells indicated by arrowheads.

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Chapter 4

Summary and Discussion

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Summary and Discussion

Major Findings

The study detailed in Chapter 2 was the first to describe a model whereby

VEGF/NFATc1/ERK1/2 signaling promotes ECC cell proliferation during ECC growth, and RANKL/NFATc1/JNK1/2 signaling inhibits VEGF-induced cell proliferation, while promoting CtsK expression, during valve remodeling. These data also support a novel mechanism for the transition from ECC growth to remodeling in which NFATc1 promotes a sequential pattern of gene expression via cooperation with ligand-specific cofactors MEK1-ERK1/2 and JNK1/2. The study in Chapter 3 was the first to demonstrate NFATc1 expression in the PE, epicardium, and EPDCs. This study demonstrates that loss of NFATc1 in EPDCs results in decreased coronary vessel and fibrous matrix penetration into ventricular myocardium leading to embryonic death in mice. These data also show that RANKL activation of NFATc1 promotes CtsK expression and invasion of EPDCs into myocardium. This study is the first to report a molecular mechanism for EPDC invasion of myocardium necessary for coronary vessel and fibrous matrix development. Together, these data demonstrate a conserved role for

RANKL/NFATc1 signaling in ECM remodeling for valve development and EPDC invasion during cardiac morphogenesis.

NFATc1 and cardiac valve maturation

The studies detailed here demonstrate that NFATc1 serves as a nodal point in the transition from endocardial cushion growth to valve remodeling. Pediatric valve

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malformations and disease are characterized by aberrant ECM architecture suggesting defective valve remodeling as a primary insult (Hinton et al., 2006). Manipulation of

NFATc1 and NFATc1 copathway signaling holds promise as a means of therapeutic intervention to halt abnormal valve endothelial cell proliferation and/or induce proper

ECM remodeling and thereby restore normal ECM architecture. Further studies are required to determine the signals that initiate RANKL expression in the developing valves as well as to determine the upstream effectors and downstream targets of

NFATc1 copathway activation in order for this goal to be realized.

2.5% of Americans suffer from cardiac valve disease (Lloyd-Jones et al., 2010).

Like pediatric valve disease, adult valve disease is also characterized by reactivation of

ECM remodeling enzyme expression, altered ECM architecture, and endothelial cell dysfunction (Rabkin et al., 2001; Rajamannan, 2010). Previous work demonstrates that adult pulmonary valve endothelial cells also retain the ability to respond to VEGF/

NFATc1 signaling with increased cell proliferation (Johnson et al., 2003; Jang et al.,

2010). The current standard of care for prevention of valve disease progression is limited to control of secondary effectors of valve tissue changes such as blood pressure and lipid metabolism (Stewart, 2009). The work detailed here suggests that restoration of VEGF/ NFATc1 pathway signaling and ERK1/2 copathway signaling may restore the regenerative capacity of adult valve endothelial cells. It is well known that valve disease is often accompanied by ECM degrading enzyme expression, thereby demonstrating the ability of adult valve cells to reactivate gene expression associated with valve remodeling (Rabkin et al., 2001; Chakraborty et al., 2010). This work suggests that

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NFATc1 manipulation may be a useful means of reinitiating and regulating cardiac valve

developmental mechanisms in adult valve cells to restore the proper valve architecture.

Valve replacement surgery with non-autologous tissue, mechanical valve, or pulmonary

autograft (for aortic valve replacement) are the current treatment options for severe

valve dysfunction (U.S. National Library of Medicine et al., 2010). The ability of adult

valve endothelial cells to proliferate and undergo EMT provides evidence of

regenerative potential and plasticity that may be exploited in the treatment of

degenerative valve processes (Johnson et al., 2003; Rajamannan, 2010). Further

studies aimed at manipulating NFATc1 signaling in valve endothelial cells may provide

the means to engineer autologous valve transplants in vitro. It is hoped that a thorough understanding of molecular mechanisms of valve maturation will allow valve endothelial cells to be utilized as a renewable source of progenitor cells for valve repair and homeostasis.

NFATc1 in EPDC invasion

The proepicardium was originally termed the proepicardial organ because of the diverse cell types that arise from this developmental structure (Ratajska et al., 2008).

Further study into the mechanisms of PE-derived structure maturation underscores the heterogeneity of these cells and the intricate interplay of morphogenetic factors involved in orchestrating coronary vessel and fibrous matrix development (Ishii et al., 2010;

Mikawa and Gourdie, 1996; Reese et al., 2002). Surprisingly, these studies demonstrate that NFATc1 promotes EPDC invasion into myocardium regardless of

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terminal cell fate. Coronary heart disease is the leading cause of death in the United

States and worldwide (World Health Organization Media Center, 2008; Xu, 2010). The

current standard of care for severe coronary disease is balloon angioplasty with stent

placement or coronary bypass surgery (Grogan, 2008). EPDCs in the adult mouse

heart maintain the ability to reactivate embryonic gene expression and create new

coronary vessels in vivo (Smart et al., 2007). It has been demonstrated that EPDCs

from the adult human heart are also able to reactive embryonic gene expression and

promote vasculogenesis in vitro (Smart et al., 2009). While lack of NFATc1 does not affect initial stages of PE migration and epicardium formation, additional data collected in this study indicates that genetic loss of NFATc1 results in decreased epicardium cell proliferation at E10.5 (Figure 1). This suggests that manipulation of NFATc1 signaling may promote epicardium cell proliferation, leading to the hope that a patientʼs own

EPDCs may be stimulated to generate new coronary vessels as a treatment for coronary heart disease.

Work by Zeini et al. demonstrates that calcineurin inhibition with Cyclosporin A blocks the ability of epicardium-derived cells to invade matrigel (Zeini et al., 2009). This study along with data presented here suggests that calcineurin/ NFATc1 activation are necessary for EPDC invasion in vitro and in vivo. Data from human studies indicates that upon myocardial damage, cells of the epicardium become activated and migrate to the site of injury (Smart et al., 2009; Smart et al., 2007; Limana et al., 2010). Animal data also demonstrate that endogenous EPDCs, and those harvested from epicardium and expanded in culture, support myocardium repair, revascularization and reduce post-

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infarct fibrosis (Limana et al., 2010; Smart et al., 2009). However, EPDC cell survival and migration to the site of injury is limited (Limana et al., 2010). Data presented here suggests that manipulation of VEGF/NFATc1 signaling may promote epicardium cell proliferation and manipulation of the RANKL/ NFATc1 pathway may increase EPDC migration to the site of injury. Expansion of the epicardium cell population in vivo or in vitro coupled with increased EPDC invasion may significantly improve clinical outcomes associated with EPDC-induced cardiac repair (Pesce et al., 2010).

Experimental limitations and alternative approaches

The experimental approaches described in this dissertation were designed to rigorously test hypotheses by generating specific and quantitative data related to critical molecular and cellular processes in heart development. These studies provide evidence in support of the main hypotheses, however, it is important to recognize experimental limitations, some of which might be overcome by alternative approaches and/or additional experiments.

In both endocardial cushion and EPDC-related experiments, a heterogeneous population of cultured primary avian cells was used to assess changes in CtsK expression resulting from manipulation of RANKL/NFATc1 pathway signaling.

Undissociated superior and inferior endocardial cushions and PE were used for these experiments. In endocardial cushions, only endothelial cells express NFATc1.

Likewise, only a subset of PE cells express NFATc1. Therefore, the contribution of

NFATc1-negative cells to CtsK expression changes assessed by Real Time RT-PCR

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cannot be determined. Human umbilical endothelial cells (HUVECs) or human dermal microvascular endothelial cells (HDMECs) are often used as a substitute for various embryonic endothelial populations. However, these cells do not respond to NFATc1 inhibition in the same way as cardiac valve endothelial cells, thereby precluding their use for these experiments (Armstrong and Bischoff, 2004). Immunofluorescent antibody co-labeling experiments demonstrated that endocardial cushion endothelial cells express NFATc1 and CtsK, and a subset of PE cells co-express NFATc1 and CtsK in vitro (Figure 2). Cell sorting using flow cytometry and/or antibody-coated beads may be useful for enriching the Flk1- or Pecam-positive population of endocardial cushion endothelial cells placed into culture. Further experiments would be necessary to determine if NFATc1-positive PE cells express a unique marker on the external cell membrane by which to identify and sort these cells. An enriched population of NFATc1- positive cultured endocardial cushion or PE cells may be used to assess changes in gene expression induced by RANKL/NFATc1 pathway manipulation.

Another experimental limitation which must be addressed in both endocardial cushion and PE-related experiments is the use of CsA as a calcineurin (Cn) inhibitor.

CsA is commonly used for Cn/NFAT inhibition in vitro and in vivo and has beed employed by previous investigators to study NFAT-dependent processes related to heart valve and coronary vessel development(Chang et al., 2004; Zeini et al., 2009)

Because of its widespread use, CsA modes of action at various doses are well studied.

At the dose used in these experiments (1μg/mL), CsA exerts its action by binding to cyclophilins, which in turn bind to calcineurin and inhibit its ability to dephosphorylate

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substrates such as NFAT proteins(Kiani et al., 2000; Yeo et al., 2007). Addition of CsA

to cultured endocardial cushion and PE cells prevents calcineurin-mediated

phosphatase activity on numerous targets including all calcium-responsive NFATs

(NFATc1, NFATc1, NFATc3, NFATc4). In vivo data collected in conjunction with these

experiments supports the idea that NFATc1 is the relevant calcineurin target necessary

for CtsK expression in both of these cell types. However, the contribution of off-target

effects of CsA administration on CtsK expression was not assessed. Alternative methods for Cn/ NFAT loss of function include, knockdown of NFATc1 transcript with siRNA or use of other Cn/NFAT inhibitors. An attempt was made to transfect cultured endocardial cushion cells with siRNAs directed against NFATc1, however, efficient knockdown of NFATc1 mRNA expression was not achieved. VIVIT is a small cell membrane-permeable peptide that binds to the NFAT interaction domain on calcineurin, while not disturbing calcineurinʼs catalytic domain(Takeuchi et al., 2007). Addition of

VIVIT peptide to cultured endocardial cushion or PE cells would inhibit calcineurin-NFAT interaction without interfering with cyclophilin targets or calcineurin-mediated functions not requiring binding to calcineurinʼs NFAT interaction domain. While this approach does not overcome all experimental limitations, changes in CtsK expression that remain consistent using several different calcineurin inhibitors would provide additional support for calcineurin-NFAT dependent patterns of gene expression.

There are experimental limitations inherent to use of the Cre-LoxP system for conditional deletion of a gene of interest. Timing of Cre activation during development, proper Cre expression in the cell lineage of interest, and the efficiency of Cre-mediated

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Lox-P recombination at the genetic locus of interest are important considerations. Cre- mediated LoxP recombination at the NFATc1 genetic locus occurred as expected as assessed by immunohistochemistry (IHC) for NFATc1 protein. If recombination were less efficient than anticipated, WT1-Cre(+); NFATc1(fl/wt) mice would be bred with

WT1-Cre(-); NFATc1(fl/-) mice and WT1-Cre(+); NFATc1(fl/-) embryos would be used for analysis, thereby eliminating the need for Cre-mediated recombination at one of the two NFATc1 alleles. The WT1-Cre mice have a pgk-neo cassette, which was not removed from the Cre expression construct after ʻknock inʼ into the endogenous WT1 locus. This can alter the timing and expression domain of Cre, causing widespread Cre expression in the early embryo. Embryos used for morphometry and protein analysis were assessed for NFATc1 expression by IHC. It was determined that embryos surviving to the time-points used in this study (E13.5-E18.5) expressed NFATc1 in valve endothelial cells and lack NFATc1 expression in epicardium, EPDCs and EPDC- derivatives. Myocardial cells do not express NFATc1, therefore, Cre expression in a subset of myocytes will not affect experimental outcomes. Further, the phenotype of

WT1-Cre(+); NFATc1(fl/fl) embryos was comparable to that of embryos obtained using another epicardial Cre (GATA5). To further limit Cre activation to a specific developmental window, tamoxifen-inducible WT1-CreERT2 mice could be used.

The experiments detailed in Chapter 3 determined that NFATc1 is necessary for

EPDC invasion; however, these findings do not preclude a role for NFATc1 in epicardial cell proliferation. Epicardial cells undergo proliferation before invading the subepicardial space and myocardium as EPDCs and differentiating into cells of the coronary

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vasculature or cardiac fibrous matrix. Non-myocyte cell proliferation in WT1-Cre(+);

NFATc1(fl/fl) embryos and littermate controls may be assessed using anti- phosphohistone H3 and MF20 antibody labeling. These experiments are currently underway and the results will be incorporated into the manuscript associated with data from Chapter 3.

The embryonic origin of progenitor cells that form coronary vascular endothelium continues to remain elusive. Experiments detailed in Chapter 3 and those performed by

Zhou et al. found that cells of the WT1-Cre lineage contribute to formation of the coronary endothelium(Zhou et al., 2008). Work by Red-Horse et al. demonstrated that cells derived from the sinus venosus form coronary endothelium(Red-Horse et al.,

2010). Because WT1 and NFATc1 are expressed by cells of the sinus venosus and PE, the experiments outlined here can not differentiate between PE-derived and sinus venosus-derived WT1-positive cells and therefore cannot be used to determine the origin of coronary endothelial cells. The answer to this question, while important is beyond the scope of the experiments designed to determine the role of NFATc1 in

EPDCs.

As stated previously, every effort must be taken to assess experimental limitations and interpret experimental outcomes accordingly. Previous attempts to assess the roles of NFATc1 in heart development were hampered by embryonic lethality at E12.5-E14.5 in embryos lacking NFATc1 expression. Experiments detailed in Chapters 2 and 3 of this dissertation were designed to overcome this limitation and were therefore useful in demonstrating novel roles NFATc1 in regulating the transition from AV valve growth to

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remodeling and promoting EPDC invasion into myocardium.

Calcineurin inhibitors

One of the most serious age-related changes that can occur in cardiac valves and blood vessels is calcification. Recent work demonstrates that NFATc1 is a critical regulator of osteogenesis and bone degradation in the skeleton (Winslow et al., 2006; Takayanagi,

2005). Ongoing work by the Yutzey lab demonstrates that cells found within cardiac valves and blood vessels display an osteoblast- and osteoclast-related profile of gene expression (Lincoln et al., 2006; Lange and Yutzey, 2006; Chakraborty et al., 2010).

Pathology specimens show that increased cell proliferation and increased ECM remodeling enzyme expression are associated with calcific lesions in cardiac valves and vessels (Aikawa et al., 2006; Rajamannan et al., 2003; Katsuda and Kaji, 2003).

NFATc1 promotes valve endothelial and epicardium cell proliferation and is associated with ECM remodeling in valves and EPDCs. Therefore, NFATc1 may be associated with osteoblast-like calcification and osteoclast-like ECM remodeling in diseased valve and vessel tissues. Indeed, calcineurin inhibition via Cyclosporin A is correlated with atherosclerosis in humans (Kockx et al., 2010). These associations merit further investigation. Calcineurin/ NFAT inhibitors have been in use for many years as immune suppressant drugs, however, the confounding effects of organ transplant and other serious medical conditions which warrant major immunosuppressant therapy in these patients makes interpretation of the specific effects of calcineurin inhibition on cardiac valve and vessel calcification difficult to assess. However, calcineurin inhibitors are

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finding new uses in cancer and osteoimmune disease treatments (Mancini and Toker,

2009; Sitara and Aliprantis, 2010). A large body of data that encompasses many disease states and pharmaceutical doses may allow for better assessment of the effects of these inhibitors on valve and vessel tissues.

The data presented here further emphasize the importance of NFATc1 function in cardiac development. As stated previously, calcineurin inhibitors such as Cyclosporin A and FK506 are widely used as immune suppressant drugs and are being tested as treatments for bone diseases and cancer (Mancini and Toker, 2009; Sitara and

Aliprantis, 2010). FK506 appears to have a limited ability to cross the placental barrier, however, use of this drug is associated with spontaneous abortion, low birth weight and number of other fetal complications in humans (Tendron et al., 2002). Cyclosporin A readily crosses the placental barrier making use of this drug during pregnancy ill advised (Tendron et al., 2002). However, under special circumstances, low dose

Cyclosporin A is administered throughout pregnancy. These pregnancies must be monitored closely and often result in spontaneous abortion, low birth weight, and other fetal complications in humans (Tendron et al., 2002). The findings presented here underscore the need for caution when administering FK506 and Cyclosporin A to women of childbearing age.

A broad view of NFATc1-regulated mechanisms

The studies detailed in Chapters 2 and 3 in this thesis demonstrate a conserved

NFATc1-regulated mechanism for cell proliferation and ECM remodeling in endocardial

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cushion endothelial cells and PE-derived cells during heart development. Adult valve endothelial and epicardial cells are able to reactivate developmental mechanisms and undergo EMT, leading to the hope that these cells may be used as therapeutic progenitors for valve and coronary vessel repair. While these similarities are striking, it must also be noted that endocardial cushion endothelial cells do not contribute to blood vessel formation during development, while a large number of EPDCs form coronary vessels. Indeed, neoangiogenesis is a pathological feature associated with valve disease, while increased angiogenesis is a long sought-after goal in the treatment myocardial ischemia. Further understanding of the intrinsic similarities and differences between valve endothelium and epicardium, as well as the effects of environmental cues on cell behavior and function, may be the key to harnessing the regenerative potential of both of these cell types. This potential may include the ability to influence epicardium-derived cells to migrate to the site of ischemic injury and form new vasculature rather than fibrotic scar tissue. This knowledge may also make it possible to instruct valve endothelial cells to reconstitute a valve matrix scaffold with VIC that produce and maintain proper ECM architecture. Certainly, identifying conserved mechanisms for cell proliferation and invasion in various tissue types allows researchers to draw upon a much larger body of data than that collected in just one cell type. This speeds progress in all fields of study concerned and may lead to overarching conclusions about cell behavior.

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Figure legends

Figure 1. E10.5 NFATc1 -/- mouse epicardial cells exhibit decreased proliferation.

A-B) Immunohistochemistry (IHC) of NFATc1 +/+ (A) and NFATc1-/- (B) mouse embryo heart sections shows epicardium (arrow) with anti-BrdU labeled (brown) epicardial cells

(arrowheads) and hematoxylin (blue) stained nuclei. C) IHC of NFATc1+/+ mouse embryo heart section shows MF20-reactive myocardium (Myo), and non-reactive epicardium (arrow in magnified inset). D) Quantification of percent BrdU positive epicardial and myocardial cells in six embryos for each genotype (n=6). *P≤0.01.

Figure 2. Primary avian PE cells with nuclear NFATc1 coexpress CtsK in vitro.

A-Cʼ) Immunofluorescence with confocal microscopy of cultured primary avian PE cells treated with 800ng/mL RANKL for 30 min. demonstrates CtsK (green) expression in cells with nuclear NFATc1 (red). A-Aʼ) RANKL treatment induces nuclear localization in

NFATc1-postive cells (arrowheads in Aʼ). NFATc1-negative cells are also present in culture (arrows in A). B-Bʼ) A subset of cells express CtsK in culture (arrowheads in Bʼ), while other PE cells are CtsK-negative (arrowheads in B). C-Cʼ) NFATc1-positive PE cells coexpress CtsK (arrowheads in Cʼ), while NFATc1-negative cells do not express

CtsK (arrows in C). Cell nuclei are stained with ToPro3 iodide (blue).

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Appendix I

Heart Valve Development: Regulatory networks in development and disease*

Michelle D. Combs, Katherine E. Yutzey

Division of Molecular Cardiovascular Biology

Cincinnati Childrenʼs Hospital Medical Center ML7020

240 Albert Sabin Way

Cincinnati, OH 45229

*Published in Circulation Research 2009 August 28; 105(5): 408-421

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Abstract

In recent years, significant advances have been made in the definition of regulatory pathways that control normal and abnormal cardiac valve development.

Here, we review the cellular and molecular mechanisms underlying the early development of valve progenitors and establishment of normal valve structure and function. Regulatory hierarchies consisting of a variety of signaling pathways, transcription factors, and downstream structural genes are conserved during vertebrate valvulogenesis. Complex intersecting regulatory pathways are required for endocardial cushion formation, valve progenitor cell proliferation, valve cell lineage development, and establishment of extracellular matrix (ECM) compartments in the stratified valve leaflets. There is increasing evidence that the regulatory mechanisms governing normal valve development also contribute to human valve pathology. In addition, congenital valve malformations are predominant among diseased valves replaced late in life.

Therefore, the understanding of valve developmental mechanisms has important implications in the diagnosis and management of congenital and adult valve disease.

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Introduction

Defective development of the heart valves occurs in 20-30% of congenital cardiovascular malformations, and the incidence of congenital valve malformations has been estimated as high as 5% of live births (Hoffman and Kaplan, 2002; Pierpont et al.,

2007). Heart valve replacement is the second most common cardiac surgery in the

United States, and the majority of replaced aortic valves have congenital malformations

(Roberts and Ko, 2005; Supino et al., 2004). Developmental defects in valve structure and function occur in several syndromes with identified genetic lesions, including trisomy 21, Noonan, Marfan, Williams and Holt-Oram syndromes (Weismann and Gelb,

2007). Additional isolated gene mutations have been associated with valve development and disease (Garg, 2006; Garg et al., 2005; Kirk et al., 2007). However, in many cases, the underlying causes of valve developmental anomalies and associated dysfunction have not been identified. Here, we review studies of heart valve development and related disease mechanisms in animal models and in tissue culture.

These research efforts provide extensive information on the molecular mechanisms and cellular events that govern the initial formation, maturation and function of heart valves with implications for development of new therapies for valve disease.

Overview of valve development

The four-chambered vertebrate heart has aortic and pulmonic semilunar (SL) valves at the arterial pole as well as mitral and tricuspid valves separating the atria and ventricles. The coordinated opening and closing of the heart valves occurs

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approximately three billion times in an average human lifespan and is required for unidirectional blood flow (Schoen, 2008). The three cusps of each SL valve and the two

(mitral) or three (tricuspid) leaflets of the atrioventricular (AV) valves consist of complex stratified connective tissue (Rabkin-Aikawa et al., 2005; Schoen, 2008). The valve leaflets are ensheathed in endocardial endothelial cells with intervening valve interstitial cells (VIC) that function in homeostasis and disease (Hinton et al., 2006; Rabkin-Aikawa et al., 2004). The valves are stratified into extracellular matrix (ECM) layers rich in elastin (ventricularis of SL/atrialis of AV), proteoglycan (spongiosa) and collagen

(fibrosa), oriented relative to blood flow (Figure 1)(Hinton et al., 2006). The most obvious difference between the AV and SL valves is the presence of supporting chordae tendineae on the ventricular aspect of the tricuspid and mitral valves. However, comparable supporting connective tissue is present in the aortic and pulmonic roots and hinge regions of the SL valves (Hinton et al., 2008; Hinton et al., 2006). Morphogenetic and structural differences also exist among the individual mural and septal AV valve leaflets, but, in general, the molecular mechanisms of valve development are conserved among AV and SL valve leaflets. Extensive conservation of valve developmental mechanisms also has been observed among vertebrate species including chicken, mouse, and human.

The first evidence of valvulogenesis during embryonic development is the formation of endocardial cushions in the AV canal and outflow tract (OFT) of the primitive looped heart tube (Person et al., 2005b; Schroeder et al., 2003). Valve primordia corresponding to individual leaflets and cusps are derived from the

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endocardial cushions, although the precise cushion origins of specific valve components are not well defined. For the AV valves, the septal valve leaflets are derived from the fused inferior and superior endocardial cushions that form in the AV canal of the primitive heart tube, whereas the mural leaflets are derived from mesenchymal cushions that arise laterally in the AV canal after cushion fusion (de Lange et al., 2004). Less is known of how the SL valves arise from the complex arrangement of proximal and distal cushions that form in the OFT. The valve progenitor cells of the endocardial cushions are highly proliferative, whereas little or no cell cycling is apparent later in remodeling and mature valves (Hinton et al., 2006; Lincoln et al., 2004; Rabkin-Aikawa et al., 2004).

The valve primordia continue to grow and elongate into thin fibrous leaflets of the AV valves and cusps of the SL valves, with increased ECM deposition and remodeling

(Hinton et al., 2006). This process differs somewhat for individual valve leaflets. For example, the septal leaflet of the tricuspid valve delaminates from the closely apposed muscular ventricular septum, in contrast to the corresponding mitral valve leaflet that protrudes into the ventricular lumen much earlier in its development (de Lange et al.,

2004; Kruithof et al., 2007; Lincoln et al., 2004; Oosthoek et al., 1998). During late gestation and soon after birth, the valve leaflets become stratified into highly organized collagen-, proteoglycan- and elastin-rich ECM compartments (Hinton et al., 2006;

Kruithof et al., 2007). In mammals, valve maturation and remodeling continues into juvenile stages (Hinton et al., 2008; Hinton et al., 2006; Rabkin-Aikawa et al., 2004).

Cell lineage studies in mice, based on examination of Tie2-Cre expressing cells and their derivatives, demonstrate that the vast majority of the cells present in the

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valves after birth are of endothelial endocardial cushion origin (de Lange et al., 2004;

Lincoln et al., 2004). These studies demonstrate that few, if any, cells of myocardial origin are present in the valve leaflets (de Lange et al., 2004). Likewise, in avians, myocytes are absent from the mature heart valves, with the exception of the mural aspect of the tricuspid valve, which is almost entirely muscle (Lincoln et al., 2004;

Sedmera et al., 1997). Although neural crest and secondary heart field cells are in close proximity to the SL valves, the leaflets themselves are predominantly of endothelial endocardial cushion origin (de Lange et al., 2004; Jiang et al., 2000).

However, there are neural crest-derived melanocytes and dendritic cells of unknown function on the surface of the mature SL and AV valves (Choi et al., 2009; Hinton et al.,

2008; Mjaatvedt et al., 2005; Nakamura et al., 2006). Epicardium-derived cells also have been identified as a source of valve progenitor cells, based on quail-chick transplantation studies (Gittenberger-de Groot et al., 1998). While cell lineage analysis of the chicken proepicardium does not show valve cell investment (Pennisi and Mikawa,

2009), Cre-positive cells are apparent in the developing valves of Tbx18-Cre and WT-

1Cre mice (Cai et al., 2008; Zhou et al., 2008a). However, studies by de Lange et al. demonstrate no investment of epicardial cells in the mature avian valves and conclude that all four valves in mice are almost entirely of endothelial origin (de Lange et al.,

2004). Overall, multiple lines of evidence support the conclusion that the mature valves are derived from endothelial endocardial cushion progenitors with little or no contribution from other cell types.

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Comparison of adult aortic valve leaflet structure and composition demonstrates

similar stratification in humans, sheep, chickens, rabbits, and mice (Hinton et al., 2006).

While hearts with multiple chambers and valves evolved in response to the demands of

separate systemic and pulmonary circulation required for terrestrial life, the molecular

pathways and cellular processes of valve formation have their origins in simpler hearts

that also drive unidirectional fluid flow. Conserved valve cell regulatory mechanisms

consisting of signaling pathways and transcription factors have been reported in ostia

cells of the Drosophila dorsal vessel (Reim and Frasch, 2005; Zeitouni et al., 2007). In zebrafish, endocardial cushions form in the primitive heart tube, although there is some debate regarding whether the cellular events of early endocardial cushion formation are conserved (Beis et al., 2005; Scherz et al., 2008). Recently, high-speed imaging of zebrafish heart valve development demonstrated that the endocardial cushions form initially by invagination of the endocardium, and not an epithelial-to-mesenchymal transition (EMT) of endocardium at the AV canal, as is observed in avians and mammals (Scherz et al., 2008). However, the mature AV valve of the adult zebrafish two-chambered heart is structurally similar to the mammalian AV valves with stratified

ECM and supporting chordae tendineae (Hu et al., 2000). Therefore, the major cellular and molecular events of valve development are largely conserved among animals with hearts composed of multiple chambers.

Since the initial reports of endocardial cushion composition by Markwald et al. in the late-1970s (Markwald et al., 1977; Markwald et al., 1975), the study of heart valve development has expanded to include investigation of a broad spectrum of signaling

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and transcriptional mechanisms that control many aspects of valve development and function. These studies encompass a broad spectrum of approaches and animal model systems with relevance to human congenital and postnatal valve abnormalities. Here we focus on the molecular regulation of valve development in hearts with four chambers, based on human disease mutation analysis, genetic studies in mice, and embryological manipulations in avians.

Endocardial cushion formation and EMT

The first evidence of endocardial cushion formation is swellings that appear in the atrioventricular canal (AVC) and OFT regions of the looping heart (E3 chick, E9.5 mouse, E31-35 human) (Fishman and Chien, 1997; Martinsen, 2005; Moorman et al.,

2003). Endocardial cushion formation is induced by myocardial production of signaling molecules that inhibit expression of chamber-specific genes in the AVC and OFT, while increasing synthesis of ECM components (Figure 2A) (Harrelson et al., 2004; Lyons et al., 1990; Ma et al., 2005; Plageman and Yutzey, 2004). This increased ECM or

ʻcardiac jellyʼ deposition between the myocardium and endocardium, along with the hydrophilic nature of the ECM proteoglycans, causes the tissue to protrude or swell into the interior lumen of the heart forming the endocardial cushions (Camenisch et al.,

2000; Henderson and Copp, 1998; Markwald et al., 1977). Even at this early stage, endocardial cushions act as physical barriers that prevent the backflow of blood through the primitive heart tube (Schroeder et al., 2003). Signaling molecules originating from both the myocardium and endocardium of the AVC and OFT are necessary for proper

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endocardial cushion formation and EMT of endocardial endothelial cells (Figure 2B)

(Krug et al., 1985). EMT occurs as a subset of endocardial cushion endothelial cells break connections with neighboring cells and migrate into the cardiac jelly to populate the endocardial cushions with mesenchymal cells. The processes of endocardial cushion formation and EMT have been extensively studied using in vitro cell culture as well as in vivo model systems.

In general, the regulatory interactions and cellular events of valvulogenesis are conserved in AVC and OFT cushion development. The AVC cushions develop approximately a day earlier than the OFT cushions, and the examination of the OFT cushions is complicated by the presence of neural crest-derived progenitors that form the aorticopulmonary septum (Camenisch et al., 2002; Delot, 2003). Defects in secondary heart field development also preferentially compromise SL, but not AV, valve development related to defects in outflow tract morphogenesis (reviewed in Rochais et al.) (Park et al., 2008; Rochais et al., 2009; Zhang et al., 2008). Many of the molecular regulatory hierarchies that control early stages of valvulogenesis have been defined using AVC explants from mouse or chick embryos due to the larger size and accessibility of cushion tissue. In vivo studies confirm that these interactions also occur in the developing OFT cushions with the exceptions noted below.

Bone Morphogenic Proteins (BMPs) are members of the Transforming Growth

Factor Beta (TGFβ) superfamily and signal predominantly through activation of

SMAD1/5/8 (van Wijk et al., 2007). Data collected using both in vitro primary cell culture and in vivo model systems suggests that BMPs act as the major myocardially-derived

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signals for initiation of endocardial cushion formation and EMT. BMP2 and 4 are expressed in the AVC and OFT myocardium during endocardial cushion morphogenesis in chick and mouse (Nakajima et al., 2000; Somi et al., 2004). Mice lacking myocardial

BMP2 expression fail to express Tbx2 in AVC myocardium, which is necessary for suppression of chamber-specific gene expression and for increased ECM deposition in this region (Ma et al., 2005; Rivera-Feliciano and Tabin, 2006). Studies using mouse

AVC explants demonstrated that BMP2 is sufficient to increase TGFβ2 expression and initiate EMT in AVC endothelial cells (Sugi et al., 2004). The role for BMPs in initiation of EMT is further supported by in vivo analysis of mice lacking myocardial BMP2 expression, which show no AVC endocardial cushion mesenchymal cell formation (Ma et al., 2005; Rivera-Feliciano and Tabin, 2006). Mice lacking expression of BMP- receptor1a in the endocardium also exhibit decreased phospho-SMAD1/5/8 activity and defective EMT in the AVC, further substantiating the requirement for BMP receptor signaling in endocardial cushion endothelial cells during EMT. Aberrant BMP signaling results in downregulation of multiple EMT-related pathways in AVC endocardial cushions, including TGFβ and Notch1, as well as decreased expression of transcription factors such as Twist1 and Msx2 (Ma et al., 2005). Taken together, these studies demonstrate a role for BMPs as important myocardially-derived signals that initiate endocardial cushion formation and EMT.

TGFβs were among the first signaling molecules to be implicated in initiation of endocardial cushion EMT (Brown et al., 1996). TGFβ ligands and receptors are expressed in the AVC and OFT during endocardial cushion formation and EMT in avian

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and murine embryos. In both chick and mouse model systems, TGFβ ligands and

receptors are required for EMT, however, species-specific differences have been noted

(Person et al., 2005b). TGFβ signaling through SMADs 2/3 induces expression of the

transcription factor Slug, which promotes AVC endocardial cushion endothelial cell

activation and invasion during EMT (Romano and Runyan, 1999, 2000). TGFβ activity

has also been associated with increased β-catenin signaling during AVC endocardial

cushion EMT in mice (Liebner et al., 2004). Mice harboring genetic deletion of β-catenin

in cells of the Tie-2 lineage fail to populate AVC endocardial cushions with

mesenchymal cells due to defective EMT (Liebner et al., 2004). In zebrafish,

overexpression of Wnt inhibitors Adenomatous polyposis coli (APC) or Dickkopf1 blocks

AVC endocardial cushion formation, that likely occurs through invagination and not EMT

(Hurlstone et al., 2003; Scherz et al., 2008). Together, these studies suggest that TGFβ

and Wnt/β-catenin signaling are important inducers of endocardial cushion formation,

but the regulatory relationships of these pathways have not been defined.

Notch signaling also plays an indispensable role in endocardial cushion EMT.

The Notch signaling ligand Delta4 and receptors Notch1-4 are expressed by

endocardial cushion endothelial cells of the AVC and OFT prior to and throughout EMT

(Timmerman et al., 2004). In mice that lack expression of Notch1 or the interacting transcription factor RBPJK, the AVC and OFT endocardial cushion swellings are present, but are devoid of mesenchymal cells due to a failure of EMT. In Notch signaling mutants, endocardial cushion endothelial cells extend processes into the cardiac jelly, but they are unable to delaminate and migrate into the ECM. In addition,

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Notch signaling induces expression of the pro-migratory transcription factor Snail in

AVC and OFT endocardial cushion endothelial cells undergoing EMT. Snail directly

represses VE-cadherin promoter activity, thereby allowing activated mesenchymal cells

to break contact with neighboring cells and migrate into the endocardial cushion interior.

Notch signaling also is required for expression of TGFβ2 and multiple TGFβ receptors in

AVC and OFT endocardial cushion endothelial cells, serving to further promote EMT.

Mutations in Notch signaling components in humans are associated with a spectrum of cardiac abnormalities, including defects in tissues derived from AVC and OFT endocardial cushions (Garg et al., 2005; Niessen and Karsan, 2008). These observations suggest that Notch signaling is dispensable for initial ECM deposition during formation of endocardial cushion swellings, but is required for endocardial cushion endothelial cell EMT.

During endocardial cushion formation, the AVC myocardium secretes biologically active adheron-like protein complexes containing ES1, fibronectin, transferrin, ES130, hLAMP1 and other extracellular components to activate adjacent endothelial cells and induce EMT (Krug et al., 1995; Krug et al., 1985; Little et al., 1989; Mjaatvedt et al.,

1991; Rezaee et al., 1993; Sinning, 1997). Proper function of these and other signaling components during AVC endocardial cushion formation and EMT requires the appropriate ECM environment. The endocardial cushion ECM is a hydrated matrix that provides physical support for mechanical function, promotes the invasive phenotype mesenchymal cells, and serves as a scaffold for cell migration (Camenisch et al., 2000;

Schroeder et al., 2003). Disruption of hyaluronan synthase-2 (has2) or versican gene

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expression in mice prevents AVC endocardial cushion formation, and hyaluronan also is

required for mesenchymal cell migration associated with EMT (Camenisch and

McDonald, 2000; Camenisch et al., 2000). Erb receptor activation is coupled to

hyaluronan function in endocardial cushion EMT, as addition of heregulin to has2-/-

AVC explants rescues EMT (Camenisch et al., 2002; Camenisch et al., 2002b).

Furthermore, Erb3-/- null mice exhibit lethality at E13.5 with hypoplastic AVC endocardial cushions due to lack of adequate EMT. Because of its diverse functions,

ECM synthesis must be properly regulated to ensure that the resulting extracellular environment has the appropriate physical and molecular characteristics to support endocardial cushion formation and EMT.

Growth of endocardial cushions and valve primordia

After EMT, the endocardial cushions and subsequent valve primordia undergo growth via cell proliferation and continued ECM synthesis (Armstrong and Bischoff,

2004; Hinton et al., 2006; Martinsen, 2005). The AVC valve primordia are part of a larger mass of tissue called the septum intermedium that is formed via fusion of the endocardial cushions at E4.5 in chicks, E11.5 in mice, and E37-42 in humans

(Martinsen, 2005; Moorman et al., 2003; Webb et al., 1998; Wessels and Sedmera,

2003). Septum intermedium tissue contributes to the membranous ventricular septum and fibrous continuity overlying the ventricular septum adjacent to the valve primordia that form the posterioinferior and septal tricuspid valve leaflets as well as the septal mitral valve leaflet (Martinsen, 2005). The OFT endocardial cushions also fuse and

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contribute to the formation of the aortic and pulmonary valve leaflets and supporting structures (Qayyum et al., 2001). Molecular mechanisms regulating growth of post-EMT endocardial cushions and valve primordia are reviewed below.

Endocardial cushion and valve primordia mesenchymal cell proliferation is both positively and negatively regulated during growth of these structures (Figure 3). BMPs promote growth, as BMP4 mouse mutants display hypocellular AVC and OFT valve primordia that remain unremodeled (Jiao et al., 2003; McCulley et al., 2008). Double mutants for BMP6/7 and those harboring BMPRII mutations have hypoplastic OFT valve primordia with grossly normal AVC, demonstrating an important difference in the local requirements for BMP6/7 signaling (Delot et al., 2003; Kim et al., 2001). Conversely, genetic deletion of the BMP inhibitory SMAD, SMAD6, in mice results in AVC and OFT valve primordia hyperplasia, further illustrating the need for proper levels of BMP signaling to achieve normal valvulogenesis (Galvin et al., 2000). FGFs also promote post-EMT endocardial cushion/valve primordia mesenchymal cell proliferation. FGF4 is expressed throughout the AVC and OFT of chick embryos during growth of endocardial cushions and valve primordia, while FGF receptors 1, 2, and 3 expression is restricted to endocardial cushion/valve primordia cells (Sugi et al., 2003). FGF4 treatment of chick AVC endocardial cushion explants or injection of replication defective retrovirus containing FGF4 coding sequence into chick hearts in ovo increases mesenchymal cell proliferation and results in hyperplastic AVC valve primordia. Conversely, Epidermal

Growth Factor (EGF) signaling inhibits endocardial cushion and valve primordia mesenchymal cell proliferation through antagonism of BMP-mediated activation of

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SMAD1/5/8 in the AVC and OFT (Person et al., 2005b). Therefore, mutations causing

reduced EGF signaling result in hypercellular AVC and OFT valve primordial (Chen et

al., 2000; Krenz et al., 2005). These data demonstrate complex regulation of

mesenchymal cell proliferation during endocardial cushion/valve primordia growth.

In humans, mutations in several genes impinging on the Ras/MAPK pathway,

including Protein Tyrosine Phosphatase Non-receptor type 11 (PTPN11), which

encodes the protein tyrosine phosphatase SHP2, cause Noonan syndrome (Araki et al.,

2009). Noonan syndrome is associated with multiple congenital defects including

cardiac abnormalities in tissues derived from AVC and OFT endocardial cushions

(Fragale et al., 2004). SHP2 promotes Ras/MAPK activation and also acts downstream

of EGF and other growth factor receptors (Fragale et al., 2004; Krenz et al., 2008).

Mice bearing activating Shp2 mutations in Tie-2 expressing cells have increased

ERK1/2 activation and increased proliferation of AVC and OFT endocardial cushion/valve primordia endothelial and mesenchymal cells (Krenz et al., 2008). This phenotype is rescued by genetic deletion of ERK1. It is hoped that further studies of mouse models such as these will lead to better understanding of, and therapies for,

Noonan syndrome.

Canonical Wnt signaling is active in growing AVC and OFT endocardial cushions and valve primordial (Gitler et al., 2003; Hurlstone et al., 2003; Liebner et al., 2004).

Wnt4 and Wnt9b are expressed by endothelial cells of the mouse AVC and OFT

endocardial cushion and valve primordia, while Wnt2, Lef1 and the Wnt inhibitor Fzd2 are expressed in corresponding mesenchymal cells (Alfieri et al., 2010). Wnt9a is

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expressed in chick AVC endocardium, and introduction of replication competent retrovirus expression Wnt9a leads to hypercellular valve primordial (Person et al.,

2005a). The reverse phenotype is seen when avian AVC explants are treated with the

Wnt inhibitor Frzb (Person et al., 2005a). Similarly, mutation of the Wnt signaling inhibitor APC in zebrafish causes increased AVC endocardial cushion and valve primordia mesenchymal cell proliferation (Gitler et al., 2003; Hurlstone et al., 2003;

Liebner et al., 2004). These data suggest that Wnt signaling must be tightly regulated during endocardial cushion/valve primordia growth to maintain proper levels of mesenchymal cell proliferation.

During growth of valve primordia and in cellularized endocardial cushions, mesenchymal cells are distributed throughout the ECM (Hinton et al., 2006). This ECM is rich in hyaluronan, versican, and other basement membrane components, however, differentiating mesenchymal cells also begin to produce collagens 1, 2, 3, 4 and 9 as well as cartilage and tendon-related ECM components such as aggrecan and tenascin

(Chakraborty et al., 2008; Hinton et al., 2006; Lincoln et al., 2006a; Little and Rongish,

1995; Peacock et al., 2008). AVC endocardial cushion explant experiments as well as mouse models demonstrate a role for BMP-regulated transcription factors in maintaining a balance between endocardial cushion/valve primordia mesenchymal cell proliferation and differentiation. Tbx20 and Twist1 are expressed by AVC endocardial cushion/valve primordia cells during growth of these structures and are associated with high levels of valve cell proliferation as well as expression of pro-migratory genes such as periostin, cadherin-11 and matrix-metalloproteinase(MMP)-2 (Shelton and Yutzey, 2007; Shelton

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and Yutzey, 2008). Sox9, another BMP-regulated transcription factor, also promotes cell proliferation and maintenance of proper ECM architecture during endocardial cushion/valve primordia growth (Akiyama et al., 2004; Lincoln et al., 2007). Sox9 mutant embryos have hypocellular AVC and OFT endocardial cushions due to defective proliferation of mesenchymal cells and display dysmorphic valve primordia ECM.

Further, expression of transcription factors Msx1/2 in OFT myocardium and endocardial cushion/valve primordia cells induces expression of BMP4, which negatively regulates

OFT endocardial cushion and valve primordia mesenchymal cell proliferation (Chen et al., 2007). Therefore, Msx1/2 double mutants exhibit hypercellular SL valve primordia.

It is clear that a complex network of transcription factors is necessary to promote proper levels of endocardial cushion/valve primordia mesenchymal cell proliferation and maintain the appropriate ECM architecture during endocardial cushion/valve primordia growth.

Endocardial cushion formation, EMT and growth of endocardial cushions and valve primordia are associated with high levels of endothelial cell proliferation (Hinton et al., 2006; Markwald et al., 1977; Martinsen, 2005). Vascular Endothelial Growth Factor

A (VEGF) is a potent cytokine that promotes endothelial cell proliferation as well as survival (Miquerol et al., 2000). VEGF is highly expressed by myocardium and endocardium prior to endocardial cushion formation, however, endocardial VEGF expression becomes restricted to endothelial cells of the AVC and OFT during endocardial cushion formation, EMT and growth of valve primordial (Dor et al., 2001;

Miquerol et al., 1999). VEGF-receptors 1 and 2 (VEGFR1, 2) are expressed throughout

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the endocardium; however, VEGF and VEGFR expression is absent in mesenchymal endocardial cushion/valve primordia cells. Studies in chick, mouse, and zebrafish demonstrate that VEGF signaling contributes to AVC and OFT endocardial cushion cell proliferation (Dor et al., 2001; Lee et al., 2006; Miquerol et al., 2000). VEGF signaling also inhibits AVC endocardial cushion EMT by promoting maintenance of an endothelial cell phenotype, thereby maintaining a proliferative population of endothelial cells throughout endocardial cushion formation, EMT and endocardial cushion/valve primordia growth. VEGF expression must be strictly controlled during endocardial cushion EMT and endocardial cushion/valve primordia growth, as overexpression inhibits EMT, while underexpression of VEGF results in failure to maintain a proliferative endothelial cell population.

Nuclear Factor of Activated T-cells cytoplasmic 1 (NFATc1) is an NFAT family transcription factor expressed by AV and SL endocardial cushion/valve primordia endothelial cells throughout growth and remodeling (de la Pompa et al., 1998; Lange and Yutzey, 2006; Ranger et al., 1998). NFATc1-/- mouse embryos have normal endocardial cushion formation and EMT, however, AVC endocardial cushions have a reduced proliferative index, and AV and SL valve primordia fail to undergo remodeling, with embryonic lethality by E14.5 (de la Pompa et al., 1998; Ranger et al., 1998; Combs and Yutzey, 2009) In cultured chicken AVC endocardial cushion cells, VEGF promotes endothelial cell proliferation through NFATc1 activation (Combs and Yutzey, 2009).

Likewise, VEGF treatment of human pulmonary valve endothelial cells induces NFATc1- dependent proliferation demonstrating a role for this interaction in adult valve

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homeostasis (Johnson et al., 2003). VEGF and NFATc1 expression are extinguished in

AVC and OFT endocardial cushion mesenchymal cells upon EMT, but are maintained in the overlying endothelial cell layer (de la Pompa et al., 1998; Miquerol et al., 1999;

Ranger et al., 1998). Concomitant with valve remodeling, VEGF expression in AVC valve primordia endothelial cells is downregulated and valve endothelial cell proliferation is greatly diminished, while AV and SL valve endothelial cell NFATc1 expression is maintained (Hinton et al., 2006; Lange and Yutzey, 2006; Miquerol et al., 1999).

Receptor Activator of NFκB Ligand (RANKL), an upstream activator of NFATc1, is expressed in AV and SL valve endothelial cells during the transition from valve primordia growth to remodeling (Lange and Yutzey, 2006; Combs and Yutzey, 2009).

RANKL treatment of primary chicken AVC endocardial cushion cells activates NFATc1 to induce expression of ECM remodeling enzymes, such as Cathepsin K(CtsK), while inhibiting cell proliferation (Combs and Yutzey, 2009). Likewise, RANKL treatment of cultured mouse hearts increases NFATc1 and CtsK transcription (Lange and Yutzey,

2006). CtsK is normally expressed in AV and SL valve endothelial cells during remodeling, however, NFATc1-/- mice lack expression of this proteinase and their valves remain unremodeled (de la Pompa et al., 1998; Lange and Yutzey, 2006; Ranger et al., 1998). These data suggest NFATc1 serves as a nodal point in the transition from growth of valve primordia via endothelial cell proliferation to valve remodeling.

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Diversification of valve cell types

During fetal stages of the chicken (E14), mouse (E16.5-17.5) and human (20-39 weeks), the valve primordia elongate into thin valve leaflets. Valve patterning is evident in differential gene expression on the surface of the valve exposed to unidirectional pulsatile blood flow versus the side of the valve away from flow (Figure 1). Elastin expression is localized to the flow side of the valves, whereas organized collagen fibrils are apparent in the fibrosa layer away from blood flow (Aikawa et al., 2006; Hinton et al.,

2006; Kruithof et al., 2007). Additional specialized ECM compartments are the proteoglycan-rich spongiosa layer as well as the tenascin-rich chordae tendineae and supporting structures (Lincoln et al., 2004, 2006a). Together, these ECM compartments are required for normal valve structure and function, with dysregulation leading to disease (see below). The developmental and molecular mechanisms regulating valve stratification currently are not known. Hemodynamics is often evoked as a driving force in valve development, and there is evidence that blood flow is required for valve maturation in zebrafish (Butcher and Markwald, 2007; Hove et al., 2003). However, it has been particularly difficult to manipulate blood flow in the four-chambered heart in order to determine specific effects on the developing valves, distinct from compromised myocardial function or embryonic viability.

One of the first indicators of valve polarity in mouse and chicken embryos that distinguishes the flow side versus fibrosa side is localized Notch pathway activation and expression of downstream effectors Hey/Hrt/Hesr1 and 2 on the flow side (Mead and

Yutzey unpublished) (del Monte et al., 2007; Garg et al., 2005). Mice lacking Hesr2

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exhibit AV valve thickening and regurgitation after birth, providing evidence for Notch pathway activation in valve leaflet maturation (Kokubo et al., 2004). The role of Notch signaling in establishing polarity of the valves has not been established, but this signaling pathway appears to have multiple roles in valve development and disease

(Garg et al., 2005; Timmerman et al., 2004). An attractive hypothesis is that shear stress on the flow side of the valve promotes localized Notch signaling, thereby initiating valve polarity and stratification, but this has not yet been demonstrated.

There is emerging evidence for diversified cell types in the developing valves that give rise to distinct gene expression profiles associated with ECM compartments (Figure

4). However, the commitment of VIC to fixed lineages has not been unequivocally demonstrated. Likewise, the specific origin of VIC in distinct valve compartments has not been defined by fate mapping or cell lineage analysis of subpopulations or individual valve progenitors in vivo. The examination of the regulatory hierarchies controlling specialized cell types in the valves has been aided by studies of corresponding connective tissue types in other organ systems. Signaling pathways required for cell lineage development in cartilage, tendon and bone are active during valve remodeling

(Lincoln et al., 2006c). For example, transcription factors involved in cartilage and tendon development are localized to subsets of valve progenitor cells and are required for valve differentiation and patterning (Levay et al., 2008; Lincoln et al., 2006a; Lincoln et al., 2007). In addition, the upstream regulators and downstream targets of these transcription factors also are expressed together in the developing valves. Overall, there is increasing evidence that development of distinct ECM compartments with

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specific biomechanical properties in the valves shares molecular regulatory mechanisms with other connective tissue types of similar ECM composition.

The spongiosa layer of the valve leaflets is rich in chondroitin sulfate proteoglycans that provide a compressible ECM similar to cartilage (Schoen, 2008). In addition, the valve leaflets express the transcription factor Sox9 and structural proteins aggrecan, collagen2a1 and cartilage link protein, characteristic of cartilaginous structures (Lincoln et al., 2006a; Lincoln et al., 2007; Wirrig et al., 2007). In contrast, valve supporting structures, including the chordae tendineae, are composed of elastic matrix similar to that observed in tendons, and both express the bHLH transcription factor scleraxis as well as tenascin and collagen14 (Chakraborty et al., 2008; Levay et al., 2008; Lincoln et al., 2006a). In cultured valve progenitor cells, BMP2 treatment promotes expression of Sox9 and aggrecan, whereas FGF4 treatment promotes expression of scleraxis and tenascin (Lincoln et al., 2006a; Zhao et al., 2007). These two pathways antagonize each other in induction of lineage-specific gene expression in the developing valve progenitor cells, as was also observed in the developing limb buds

(Chimal-Monroy et al., 2003; Edom-Vovard et al., 2002). In vivo, Sox9 is required early in proliferation of the endocardial cushion mesenchyme and later in expression of collagen2a1 and cartilage link protein in the differentiated valves (Lincoln et al., 2007).

Likewise loss of scleraxis results in decreased collagen14 expression as well as increased expression of cartilage marker genes and abnormal valve ECM organization

(Levay et al., 2008). Together these studies provide evidence that multipotential valve progenitors of the endocardial cushions differentiate into cells of the valve spongiosa

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layer or supporting apparatus depending on exposure to BMP or FGF signaling, respectively.

Less is known of development of the valve fibrosa layer. During heart valve remodeling, ECM proteins characteristic of fibroblasts and preosteoblast lineages are restricted to the fibrosa layer, oriented away from blood flow (Hinton et al., 2006;

Kruithof et al., 2007; Alfieri et al., 2010) These ECM proteins include osteonectin, periostin, collagens 1 and 3, and fibronectin that contribute to the highly organized collagen matrix, conferring stiffness necessary for valvular sufficiency (Kruzynska-

Frejtag et al., 2001; Norris et al., 2008; Snider et al., 2008). The mature aortic valve fibrosa layer is the usual site of pathologic calcification, and the coexpression of collagen1, osteonectin, and periostin, is characteristic of fibrous connective tissues with the potential to mineralize, such as bone or dermal fibroblasts (Freeman and Otto, 2005;

Murshed et al., 2004). Likewise, cultured VIC express fibrosa markers and can be induced to express osteogenic markers under conditions that also promote mineralization of bone (Liu et al., 2007; Mathieu et al., 2005; Alfieri et al., 2010). Wnt signaling has been implicated in bone lineage development as well as aortic valve calcification (Hu et al., 2005; Rajamannan et al., 2005; Zhou et al., 2008b). Multiple Wnt ligands, including Wnt3a and Wnt7b involved in bone development, are expressed together with the Wnt pathway reporter TOPGAL in remodeling mouse valve leaflets

(Gitler et al., 2003; Alfieri et al., 2010). In addition, Wnt treatment of avian embryo aortic

VIC in culture promotes expression of periostin (Alfieri et al., 2010). Together these analyses provide initial evidence for Wnt regulation of fibrosa layer maturation as well as

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conserved regulatory pathways with osteogenic cell lineages. Further studies are necessary to determine the requirements for Wnt signaling in heart valve stratification and disease mechanisms.

Heart valve ECM maturation and organization

Heart valve development is characterized by increasing complexity and organization of the ECM. The ECM of endocardial cushions prior to EMT is rich in hyaluronan, and the mesenchymal cells in the cushions after EMT express network collagens and MMPs 1, 2, and 13, that promote cell migration (Chakraborty et al., 2008;

Klewer et al., 1998; Shelton and Yutzey, 2007; Shelton and Yutzey, 2008). Electron microscopy studies show high cellularity and relatively unstructured ECM in endocardial cushions and valve primordial (Hinton et al., 2006; Markwald et al., 1977).

Biomechanical studies of avian endocardial cushions demonstrate increased rigidity of the tissue with increased cellularity and collagen deposition over time (Butcher et al.,

2007). Selective degradation of ECM components of endocardial cushions demonstrated that glycosaminoglycans in the cellularized cushions confer elasticity, whereas collagen provides rigidity (Butcher et al., 2007). In the stratified valves, the structurally distinct layers of ECM provide specific biomechanical properties. Elastin fibers, of the ventricularis layer of SL and atrialis layer of the AV valves, confer elasticity to the valve, extending when the valve is open and recoiling when the valve is closed

(Schoen, 2008). The relatively unstructured proteoglycans of the spongiosa layer absorb compressible forces on the leaflets and mediate movements between the highly

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structured elastin fibrils of the ventricularis/atrialis and fibrous collagen of the fibrosa layer (Schoen, 2008). The collagen-rich fibrosa layer provides stiffness and strength to the valve leaflet and is the major structural component of the valves. The collagen composition of the valves changes during maturation of the valve leaflets, with increased mature collagen fibrils at later stages, corresponding to increased structural and functional demands (Aikawa et al., 2006; Chakraborty et al., 2008; Peacock et al.,

2008). Fibrous collagen is the most abundant protein in the mature valves, and the fibrosa layer is predominantly collagen1 fibrils, but collagen3 fibrils also are present

(Hinton et al., 2006; Kruithof et al., 2007; Schoen, 2008). Overall, the precise regulation and organization of the complex layers of the valve ECM is critical for normal valve development, structure and function.

Abnormal expression and distribution of ECM proteins expressed in the valves is associated with developmental valve abnormalities and disease (Tables 1-2). Elastin mutations are associated with Williams syndrome, which includes supravalvular aortic stenosis as well as SL valve disease (Ewart et al., 1993). Loss of elastin in mice leads to arterial abnormalities and perinatal death prior to significant investment in the stratified valves (Li et al., 1998). However, elastin heterozygous mutant mice exhibit aortic valve anomalies, thus demonstrating the importance of elastin in normal valve structure and function (Hinton et al., 2010). Marfan syndrome, which includes aortic valve anomalies in addition to aortic dilation, is caused by mutations in fibrillin-1, also present in aortic valve elastic fibrils (Dietz et al., 1991). Likewise, mice lacking the associated elastic fibril ECM protein fibulin-4 exhibit thickening and calcification of the

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aortic valve (Hanada et al., 2007). Proteoglycan gene mutants have not yet been associated with defects in the stratified valves, but increased synthesis and disorganization of proteoglycans is predominant in pediatric aortic valve disease and adult myxomatous mitral valves (Hinton et al., 2006; Rabkin et al., 2001). It seems likely that additional ECM gene mutations contribute to isolated and familial congenital valve malformations, and ECM genes expressed in the valves are certainly strong candidates in ongoing human genetic analyses.

Complex regulation of collagen composition is an important feature of valve maturation and homeostasis. Mutations in multiple collagen genes are associated with connective tissue disorders that include valve dysfunction and disease. Osteogenesis imperfecta is caused by collagen1a1 mutations that can lead to mitral and/or aortic valve insufficiency necessitating replacement, in addition to prevalent skeletal and vascular anomalies (Prockop et al., 1993; Wong et al., 1995). Ehlers-Danlos syndrome is associated with mutations in collagens 3, 5, 11 or tenascin X, and Stickler syndrome is caused by collagen 2 or 11 mutations (Ahmad et al., 1991; Byers, 1994; Kuivaniemi et al., 1997). Both of these syndromes include widespread connective tissue disease, as well as heart valve dysfunction, that can be severe enough to necessitate replacement (Ahmad et al., 2003; McDonnell et al., 2006). Dysregulation of the expression and distribution of fibrous collagen in the valves occurs in valve disease, with increased collagen3 relative to collagen1 fibrils in myxomatous mitral valves (Cole et al., 1984). In mice, targeted mutagenesis of facit collagen genes collagen5a1 and

11a1 results in thickening of SL and AV valves with increased expression of fibrous

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collagens 1 and 3 evident at birth (Lincoln et al., 2006b). Similarly, loss of periostin, which regulates collagen fibrillogenesis, also leads to congenital AV and SL valve anomalies that compromise heart valve structure and function (Norris et al., 2008;

Snider et al., 2008). Overall, a variety of lesions related collagen dysregulation are linked to defects in valve development and also in valve disease.

Heart valve development and disease

There is increasing evidence for a link between congenital valve malformations and late-onset valve disease. The most common valve malformation is bicuspid aortic valve (BAV), which often goes undetected until the valve becomes stenotic and requires replacement late in life (Cripe et al., 2004). Prenatally, there is increasing evidence that aortic valve malformations can lead to more severe congenital heart anomalies including hypoplastic left heart (Hinton et al., 2009; Loffredo, 2000). BAV is heritable, and mutations in the NOTCH1 gene have been associated both with BAV and aortic valve calcification (Cripe et al., 2004; Garg et al., 2005). Aortic valve calcification has been characterized as an osteogenic process with activation of several genes involved in bone mineralization including Runx2 and osteocalcin (Aikawa et al., 2007; Caira et al., 2006; Rajamannan et al., 2003). In developing bone progenitors, Notch1 signaling inhibits mineralization by repressing the transcriptional activity of Runx2, and a related mechanism has been evoked as a protective mechanism in aortic valve disease

(Deregowski et al., 2006; Garg et al., 2005). Increased Wnt signaling, also implicated in valve and bone development and antagonized by Notch signaling, is associated with

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aortic valve disease (Deregowski et al., 2006; Rajamannan et al., 2005). Therefore, signaling pathways involved in normal valve development likely have both positive and negative effects in valve pathogenesis that could be exploited in the treatment of these common conditions.

In the normal adult valve, the VIC are relatively quiescent with little or no synthetic activity or cell proliferation (Aikawa et al., 2006; Hinton et al., 2006). The most common types of valve disease are myxomatous, characterized by insufficiency and inappropriate ECM production, and stenotic, with leaflets that are thickened, stiff and mineralized (Freeman and Otto, 2005; Rabkin-Aikawa et al., 2005). Activation of VIC with increased synthetic activity is observed with both types of valve pathogenesis

(Aikawa et al., 2007; Hinton et al., 2006; Rabkin-Aikawa et al., 2004). It is not known if

VIC can reenter the cell cycle under pathologic conditions. Recent studies have begun to define distinct types of VIC, that may have specific roles in valve pathogenesis, and these may be related to diversified cell types seen during development (Liu et al., 2007).

In addition, bone marrow-derived hematopoietic stem cells have been reported to be present in adult valves, but the function of these cells in valve homeostasis and pathogenesis has not been defined (Visconti et al., 2006). There is initial evidence that the increased ECM production and VIC activation in valve pathogenesis is related to developmental pathways, but further studies are necessary to rigorously test this hypothesis (Aikawa et al., 2007; Garg et al., 2005; Paruchuri et al., 2006; Rajamannan et al., 2005).

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Conclusions and perspectives

Complex regulatory mechanisms that govern normal and abnormal valve development have been defined as a result of the work of many laboratories using a variety of experimental systems. This work has identified conserved regulatory hierarchies involving signaling pathways and transcriptional mechanisms active during both early and late valve development, as well as in other related types of connective tissue. Still, there are many remaining questions to be addressed in the study of valve development. Although the vast majority of cells in the mature valve are of endothelial cushion origin, specific contributions of epicardial- and neural crest-derived cells have yet to be fully defined. In addition, further studies are necessary to map the specific fates of individual endocardial cushion cells in the stratified valves and to determine the plasticity of mature VIC. Likewise little is known of how the common endocardial cushions contribute to specific valve leaflets, especially for the SL valves. In general, individual reports on valvulogenesis have focused on regulatory interactions acting in isolation at specific times and in specific cells of the developing valves. Further studies are necessary to fully define the interactions of these many regulatory pathways in order to have a more complete understanding how valves form during prenatal development and how alterations in these processes lead to valve dysfunction and disease.

Rabkin-Aikawa et al., 2005; Rajamannan et al., 2003). It is not known if VIC that

232

express valve developmental genes represent a dedifferentiated cell type or if there is a relatively undifferentiated cell population in normal adult valves. Alternatively, cells from extra-cardiac origins, such as mesenchymal or hematopoietic stem cells, may populate the adult valves and could contribute to disease pathology or have valve regenerative potential (Visconti et al., 2006). A valve stem cell population has not been identified.

The detailed analysis of regulatory pathways that control valve development also has implications in valve tissue engineering. In general, current efforts directed towards generating engineered valves do not take into account the diversity of VIC or their abilities to generate ECM with distinct structural characteristics (Schoen, 2005). The application of recent research into valve developmental mechanisms to the generation of engineered valves will likely improve the long-term function of these tissue constructs and could lead to improved therapeutics or replacement strategies. Likewise, manipulation of known valve developmental mechanisms could be applied to the treatment and management of the most common types of valve disease.

Acknowledgements

We thank Robert Hinton Jr. as well as Christina Alfieri, Timothy Mead, Santanu

Chakraborty and Jonathan Cheek in the Yutzey lab for communication of results prior to publication.

233

Sources of funding

Work in the author's lab is supported by grants from National Institutes of

Health/National Heart, Lung and Blood Institute, including R01HL82716, and MDC is supported by a pre-doctoral fellowship from the American Heart Association-Great

Rivers Affiliate.

Disclosures

None.

234

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Figure legends

Figure 1. Stratified ECM compartments are evident in mature SL and AV valves.

A) Schematic representation of one of three valve cusps of the aortic or pulmonic SL

valve with fibrosa (F), spongiosa (S) and ventricularis (V) layers indicated. B)

Schematic representation of one AV valve leaflet with atrialis (A), spongiosa (S), and

fibrosa (F) layers indicated. The mitral valve has two leaflets, whereas the tricuspid

valve has three leaflets, all of which are supported by chordae tendineae (CT). The

direction of pulsatile blood flow is indicated for both SL and AV valves (arrow).

Figure 2. Model for regulatory interactions that control endocardial cushion

formation (A) and EMT (B). A) Myocardial BMP2 expression increases hyaluronan

and versican deposition in cushion-forming regions of the AVC and OFT. BMP2

induces Tbx2 transcription in the myocardium, inhibiting chamber-specific gene

expression. VEGF, expressed in endothelial cells, promotes endocardial cushion

endothelial cell proliferation. B) Multiple endocardially-derived signals promote

endocardial cushion EMT. TGFβ signals through Slug to promote EMT, while Notch1 signals through Snail to suppress VE-cadherin (VE-cad) expression and promote EMT.

Wnt/β-catenin signaling increases endocardial cushion EMT. Once the cushions are established, endocardial VEGF expression maintains endothelial cell proliferation and inhibits EMT.

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Figure 3. Model for regulatory interactions that control growth of endocardial

cushions/valve primordia. Cell proliferation in the endothelial cells in the endocardial

cushions is induced by VEGF/NFATc1 and Shp2/ERK1/2 signaling. Mesenchymal cell

proliferation is induced by multiple signaling mechanisms including Wnt/β-catenin,

TGFβs, BMPs, FGF4 and Shp2/ ERK1/2. EGF signaling inhibits mesenchymal cell proliferation.

Figure 4. Model for regulatory interactions controlling AV valve stratification and lineage diversification. Notch1 expression is localized to the flow side of the stratifying valve. In the spongiosa, BMP2 signaling promotes Sox9 expression and deposition of cartilage-related ECM components, such as aggrecan. Wnt signaling in the fibrosa promotes expression of fibroblast/pre-osteoblast-related ECM components, such as periostin. Maturation of valve supporting structures (chordae tendineae) is associated with FGF4 signaling, which induces expression of the tendon-related transcription factor scleraxis and the ECM component tenascin. Although the SL valves do not have chordae tendineae, these signaling pathways also are active in the corresponding regions of the stratified aortic valve cusps and supporting structures.

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Appendix II

Transcriptional regulation of heart valve progenitor cells*

Santanu Chakraborty, Ph.D.; Michelle D. Combs,B.S.; Katherine E. Yutzey, Ph.D.

Division of Molecular Cardiovascular Biology

Cincinnati Childrenʼs Hospital Medical Center ML7020

240 Albert Sabin Way

Cincinnati, OH 45229

*Published in Pediatric Cardiology 2010 April; 31(3): 414-421

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Abstract

The development and normal function of the heart valves requires complex interactions among signaling molecules, transcription factors and structural proteins that are tightly regulated in time and space. Here we review the roles of critical transcription factors that are required for specific aspects of normal valve development. The early progenitors of the heart valves are localized in endocardial cushions that express transcription factors characteristic of mesenchyme, including Twist1, Tbx20, Msx1 and

Msx2. As the valve leaflets mature, they are composed of complex stratified extracellular matrix proteins that are regulated by the transcriptional functions of

NFATc1, Sox9, and Scleraxis. Each of these factors has analogous functions in differentiation of related connective tissue lineages. Together, the precise timing and localized functions of specific transcription factors control cell proliferation, differentiation, elongation and remodeling processes necessary for normal valve structure and function. In addition, there is increasing evidence that these same transcription factors contribute to congenital as well as degenerative valve disease.

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Introduction

The proper development and function of the heart valves are essential for unidirectional blood flow, and abnormal valve development can lead to significant cardiovascular disease. Cardiac valve defects represent approximately 20-30% of all congenital cardiovascular malformations with an incidence as high as 5% of live births

(Hoffman and Kaplan, 2002; Pierpont et al., 2007). In addition, approximately 100,000- inpatient valve procedures are performed in the U.S.A. per year (Thom et al., 2006). In the mature heart, the mitral and tricuspid atrioventricular (AV) valves are localized in the left and right AV junctions, respectively, whereas semilunar valves (SL) are positioned at the roots of the aorta (aortic valve) and pulmonary artery (pulmonic valve) (Schoen,

2008; Srivastava and Olson, 2000). During pre-natal development, valve formation is controlled by complex interactions of signaling molecules and transcription factors that regulate cell proliferation, lineage diversification, differentiation and leaflet remodeling

(Combs and Yutzey, 2009a). Here, we review the transcription factors Twist1, Tbx20,

Msx1, Msx2, NFATc1, Sox9, and Scleraxis and their roles in multiple aspects of valve progenitor cell specification and maturation.

Overview of endocardial cushion formation and valve remodeling

The development of heart valves begins with the formation of endocardial cushions (EC) at atrio-ventricular canal (AVC) and outflow tract (OFT) regions of the primitive heart tube (Combs and Yutzey, 2009a; Person et al., 2005). Endocardial cushion formation is highly conserved in vertebrates and is initiated at embryonic day

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(E) 3 in chick, E9.5 in mouse, and E31-E35 in human (Fishman and Chien, 1997;

Martinsen, 2005; Moorman et al., 2003). The endocardial cushions consist of highly proliferative undifferentiated mesenchymal cells localized between the endothelial endocardium and myocardial cell layers in the OFT and AVC. Endocardial cushion formation is initiated when signaling cues from the AVC and OFT myocardium result in epithelial to mesenchymal transformation (EMT) of adjacent endocardial cells. The transformed mesenchymal cells then migrate into the intervening cardiac jelly to form the endocardial cushions (Markwald et al., 1977; Person et al., 2005). Cell lineage analyses with Tie2-Cre X Rosa26RLacZ reporter mice demonstrate that the majority of cells present in the mature valves are of endothelial endocardial cushion origin (de

Lange et al., 2004; Lincoln et al., 2004). The mesenchymal heart valve progenitor cells in the endocardial cushions are highly proliferative, migratory and undifferentiated, within a loosely organized extracellular matrix (ECM) (Armstrong and Bischoff, 2004; de

Lange et al., 2004; Lincoln et al., 2004). These cells express several transcription factors including Twist1, Tbx20, Msx1 and Msx2 that are associated with mesenchymal precursor populations in a variety of organ systems.

Both myocardial- and endocardially-derived signaling pathways affect endocardial cushion EMT and proliferation of valve progenitor cells. Mice lacking BMP2 in the myocardium or BMPR1a in the endocardium have no AV endocardial cushion mesenchymal cell formation and no expression of the mesenchymal transcription factors Twist1, Msx1 and Msx2 (Ma et al., 2005; Rivera-Feliciano and Tabin, 2006).

BMP signaling also has a likely role in regulation of endocardial cushion cell proliferation

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because mice lacking the BMP inhibitor Smad6 have increased proliferation of endocardial cushion cells (Galvin et al., 2000). In the endocardial endothelial cells,

Notch1 signaling induces expression of the transcription factor Snail that suppresses the expression of VE-cadherin and promotes EMT (Timmerman et al., 2004). Canonical

Wnt/β-catenin signaling is also active in AV and OFT endocardial cushion and is important in EMT and mesechymal cell proliferation (Liebner et al., 2004; Person et al.,

2005). Overall, multiple signaling pathways including those activated by BMPs, Notch, and Wnts control aspects of valve progenitor cell specification, proliferation, migration and differentiation through their actions on specific transcriptional effectors (reviewed in more detail in (Armstrong and Bischoff, 2004; Combs and Yutzey, 2009a)).

Formation of the heart valve leaflets occurs with elongation and ECM remodeling of the endocardial cushions. This process is characterized by decreased cell proliferation and increased deposition and complexity of ECM (Hinton et al., 2006;

Lincoln et al., 2004). The mature valve leaflets are stratified into three layers, the elastin-rich atrialis(AV valves)/ventricularis(SL valves), the proteoglycan-rich spongiosa and highly organized collagen fiber-rich fibrosa (Hinton et al., 2006; Lincoln et al.,

2006b; Rabkin et al., 2001). These diversified matrix compartments share structural and biomechanical properties with other types of connective tissues (Schoen, 2008). There is increasing evidence for conserved regulatory hierarchies of signaling molecules and transcription factors that control both heart valve maturation and differentiation of cartilage, tendon, and bone precursors (Chakraborty et al., 2008; Garg et al., 2005;

Lange and Yutzey, 2006; Lincoln et al., 2006b).

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Transcriptional regulation of valve development

Over the past several years, significant progress has been made in elucidating the transcriptional regulatory hierarchies that control valve development (Table 1).

These studies demonstrate similarities in the regulatory interactions that control endocardial cushion mesenchyme proliferation and gene expression with other mesenchymal progenitor populations in the embryo. A recent microarray gene expression profile of murine E12.5 AV endocardial cushions versus E17.5 remodeled valves identified several transcription factors including Twist1, Tbx20, and Msx1/2 that are expressed in mesenchymal valve progenitor cells (Chakraborty et al., 2008). Later stages of valve development share transcriptional regulatory mechanisms with development of osteoclasts, cartilage, and tendon development, related to ECM remodeling and compartmentalization (Lincoln et al., 2006b). NFATc1 is required in osteoclast differentiation as well as in the transition from proliferation to remodeling of the valve primordia (Combs and Yutzey, 2009b; de la Pompa et al., 1998; Ranger et al.,

1998). In addition, Sox9, required for cartilage precursor differentiation, and Scleraxis, important in development of tendons, are critical for normal ECM compartmentalization in the developing valves (Levay et al., 2008; Lincoln et al., 2006a; Lincoln et al., 2007).

Here we review the individual functions and downstream targets of these transcription factors in endocardial cushion formation and valve leaflet maturation (Table 2).

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Twist1

Twist1, a class II basic-helix-loop-helix (bHLH) transcription factor, was first

identified in Drosophila as a critical regulator of mesoderm formation (Castanon and

Baylies, 2002). In vertebrates, Twist1 promotes cell proliferation and migration in a variety of mesenchymal, embryonic and transformed cells (Barnes and Firulli, 2009). In humans, Twist1 haploinsufficiency causes Saethre-Chotzen (SC) syndrome (OMIM

#10400), characterized by craniofacial abnormalities, and congenital heart defects also are associated with this syndrome (Reardon and Winter, 1994). Twist1 null mouse embryos demonstrate a failure in neural crest migration, hypoplastic limb buds, and vascular defects with lethality by E11.5 (Chen and Behringer, 1995; Soo et al., 2002;

Vincentz et al., 2008). While the initial stages of endocardial cushion formation are apparently normal in Twist1 null embryos (Vincentz et al., 2008), embryonic lethality precludes the detailed assessment of Twist1 function in EC maturation and differentiation.

During heart valve development, Twist1 is expressed throughout the endocardial cushions of the AVC and OFT, and expression is down-regulated in the remodeling valves (Chakraborty et al., 2008; Ma et al., 2005). Endocardial cushion expression of

Twist1 is induced by BMP2 in both chicken and mouse embryos (Ma et al., 2005;

Shelton and Yutzey, 2008). Twist1 gain and loss of function studies in avian endocardial cushion cell cultures demonstrate that Twist1 promotes cell proliferation and migration, while increasing the expression of Cadherin 11 (cdh11), Periostin

(POSTN) and Matrix metalloprotease (Mmp)2 (Shelton and Yutzey, 2008). Twist1 also

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promotes expression of Tbx20, and a Twist1-responsive regulatory element is located in

chicken Tbx20 gene flanking sequences (Shelton and Yutzey, 2008), Horn and Yutzey,

unpublished). In transgenic mice, induced expression of Twist1 in the developing

valves leads to increased cell proliferation, increased expression of Tbx20, prolonged

expression of primitive ECM genes, and abnormal valve remodeling (Chakraborty et al.,

2010). Expression of hyaluronan and proteoglycan link protein1 (Hapln1), collagen type

II, alpha1 (Col2a1), Mmp2 and Mmp13, characteristic of endocardial cushion ECM is increased in the Twist1 transgenic mouse valves, and Col2a1 is a direct downstream target of Twist1 (Chakraborty et al., 2008; Chakraborty et al., 2010). Together, these studies demonstrate a role for Twist1 in promoting EC cell proliferation, migration and primitive ECM gene expression. In addition, the loss of Twist1 is required for the normal progression of valve leaflet stratification and remodeling.

Tbx20

Tbx20 is a member of the Tbx1 subfamily of T-box genes and is expressed in multiple organs, including the heart, of multiple species including Drosophila, Xenopus, avians, rodents, and humans (Plageman and Yutzey, 2005; Stennard and Harvey,

2005). Tbx20 has both activator and repressor functions that are dependent on regulatory element context and interactions with specific cofactors (Plageman and

Yutzey, 2004; Stennard et al., 2003). Human Tbx20 mutations are associated with complex spectrum of developmental and functional cardiac abnormalities including valve defects (Kirk et al., 2007). Mice lacking Tbx20 have reduced myocardial

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proliferation and loss of heart chamber maturation, with embryonic lethality by E10.5

(Cai et al., 2005; Singh et al., 2005; Stennard et al., 2005; Takeuchi et al., 2005). The

early embryonic lethality of these mice prevented analysis of the role of Tbx20 in valve

formation, however global knockdown of Tbx20 with RNA interference provided initial

evidence for Tbx20 function in valve morphogenesis (Takeuchi et al., 2005).

During heart valve development, Tbx20 is strongly expressed in the endocardial cushion mesenchyme in both mouse and chicken embryos (Plageman and Yutzey,

2004; Stennard et al., 2003). Tbx20 gain and loss of function studies performed in chicken AVC explants demonstrated that Tbx20, like Twist1, promotes cell proliferation and migration, while repressing ECM maturation (Shelton and Yutzey, 2007; Shelton and Yutzey, 2008). Tbx20 promotes expression of the ECM remodeling enzymes Mmp9 and Mmp13, while repressing expression of the chondroitin sulfate proteoglycans aggrecan and versican. In addition, Tbx20 expression is induced by BMP signaling, and

Tbx20 is a direct downstream target of Twist1 (see above; (Shelton and Yutzey, 2007;

Shelton and Yutzey, 2008); Horn and Yutzey, unpublished). In cardiomyocytes, Tbx20 promotes cell proliferation through activation and binding to regulatory elements of N- myc and Tbx2 genes (Cai et al., 2005). Corresponding alterations in the expression of these genes also is observed in avian endocardial cushion cells with reduced expression of Tbx20 (Shelton and Yutzey, 2007). Together these studies demonstrate that Tbx20 promotes cell proliferation, while inhibiting maturation of multiple cardiac cell lineages, including endocardial cushion cells. However, the consequences of altered

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Tbx20 function on heart valve development and morphogenesis in vivo are yet to be

determined.

Msx1 and Msx2

Msx1 and Msx2 belong to a subfamily of the Nk-like homeobox genes related to

Drosophila muscle-segment homeobox (msh) (Davidson, 1995). During embryogenesis,

Msx1 and Msx2 are often expressed together in mesenchymal structures, including the limb buds, pharyngeal arches, neural crest, and endocardial cushions of the heart

(Chen et al., 2008; Davidson, 1995). In humans, mutations in MSX1 cause orofacial clefting and tooth agenesis (Blanco et al., 2001; Vastardis et al., 1996), and mutations in

MSX2 cause Boston-type craniosynostosis with premature fusion of skull bones and orofacial bone abnormalities (Jabs et al., 1993). However, no cardiac abnormalities have been reported with mutation of either human MSX1 and MSX2 genes. Similarly mice lacking either Msx1 or Msx2 have no reported cardiac anomalies, but embryos lacking both Msx1 and Msx2 have severe endocardial cushion and conotruncal defects

(Chen et al., 2007; Chen et al., 2008). Interestingly, Msx2 expression is increased in adult human calcific valve disease, and ectopic expression of Msx2 in transgenic mice induces an osteogenic gene program in the valves through increased Wnt signaling

(Shao et al., 2005).

In developing EC, both Msx1 and Msx2 are expressed in endocardial and

mesenchymal cells during EMT in overlapping patterns (Chen et al., 2008). Loss of

both Msx1 and Msx2 leads to a reduction of endocardial cushion formation, while no

endocardial cushion or valve defects are associated with loss of either Msx1 or Msx2

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(Chen et al., 2008; Satokata et al., 2000; Satokata and Maas, 1994). These results support redundant functions for Msx1 and Msx2 during AV valve morphogenesis. In the

Msx1/2 mutant embryos, endocardial expression of Notch1, BMP2/4 and NFATc1 is reduced, and patterning of the AVC myocardium also is abnormal, leading to compromised EMT (Chen et al., 2008). Secondary heart field and neural crest anomalies related to defects in cell proliferation and migration also occur with loss of both Msx1 and Msx2 (Chen et al., 2007). Together the loss of both Msx1 and Msx2 lead to a spectrum of cardiac malformations including double outlet right ventricle

(DORV), pulmonary stenosis, atrial and ventricular septal defects, and hypoplastic ventricles (Chen et al., 2007; Chen et al., 2008). Based on these studies, Msx1 and

Msx2 have overlapping functions in endocardial cushion EMT and also potentially in cell proliferation and survival. Further studies are necessary to define the specific direct downstream targets of these proteins as well as their interacting factors in valve development.

NFATc1

Nuclear Factor of Activated T-cells cytoplasmic 1 (NFATc1/NFAT2/NFATc) is a transcription factor belonging to the NFAT family (Hogan et al., 2003). NFAT factors contain a Rel homology region DNA binding domain and are regulated via dephosphorylation by calcineurin, which promotes nuclear localization. NFATs regulate proliferation, differentiation and homeostasis in numerous cell types during embryogenesis and throughout life (Crabtree and Olson, 2002). Surprisingly, NFAT

282

mutations have not been linked to human disease. Mice lacking NFATc1 expression

experience embryonic lethality by E14.5 due to lack of endocardial cushion growth and

remodeling (de la Pompa et al., 1998; Ranger et al., 1998). In addition, NFATc1

regulates immune cell function and osteoclast differentiation (Takayanagi, 2005). In

osteoclasts, NFATc1 is activated by RANKL signaling and promotes expression of the

ECM remodeling enzyme Cathepsin K (CtsK) (Takayanagi, 2005). This same regulatory interaction occurs during heart valve development (Combs and Yutzey, 2009b; Lange and Yutzey, 2006).

NFATc1 is expressed specifically in endocardial endothelial cells of the primitive heart tube and is restricted to the AVC and OFT endothelial cells at the early stages of endocardial cushion formation (de la Pompa et al., 1998). Endothelial-specific expression of NFATc1 rescues the heart defects in NFATc1-/- mutants, demonstrating the specific requirement for endothelial expression of NFATc1 in normal valve morphogenesis (Chang et al., 2004). EMT is apparently normal in NFATc1 null endocardial cushions. Further studies in NFATc1 null mice and avian endocardial cushion cell cultures demonstrate that NFATc1 is required both for endocardial endothelial cell proliferation and induction of CtsK gene expression (Combs and Yutzey,

2009b; Lange and Yutzey, 2006). NFATc1-dependent endocardial cushion endothelial cell proliferation is induced by VEGF in conjunction with MEK1-ERK1/2 activation. At later stages, RANKL treatment promotes NFATc1 nuclear localization and CtsK expression, while repressing cell proliferation, in conjunction with JNK activation. In vivo, VEGF, NFATc1, and activated ERK1/2 are localized to proliferating endocardial

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cushion endothelial cells, whereas RANKL and CtsK are expressed in the valve primordia during later remodeling stages (Combs and Yutzey, 2009b). Additional

NFATc1 transcriptional targets in the developing valves include its own promoter in an autoregulatory interaction, as well as the calcineurin modulatory enzyme

DCSR1/MCIP1/RCAN1 (Lange et al., 2004; Zhou et al., 2005). NFATc1 also is expressed in adult human pulmonary valve endothelial cells in which VEGF can induce cell proliferation via an NFATc1-dependent mechanism, supporting a role for NFATc1 in adult valve homeostasis (Johnson et al., 2003). Together, these studies demonstrate a complex critical role for NFATc1 in the transition from endocardial cushion growth to remodeling during valve development, as well as in maintenance of the mature valves.

Sox9

Sox9 (SRY-box containing gene 9) is a high mobility group transcription factor expressed in several embryonic tissues including cartilage, where it is required for cell lineage expansion and differentiation (Akiyama et al., 2002). Conditional mutagenesis of

Sox9 in mice demonstrates a requirement for cartilage precursor cell proliferation and differentiation (Akiyama et al., 2002). There is increasing evidence for shared regulatory pathways in heart valve development with development of structurally related connective tissues including cartilage (Lincoln et al., 2006b). Therefore a similar strategy was used in the developing heart valves to demonstrate that Sox9 is required for endocardial cushion cell lineage expansion as well as later differentiation evident in expression of genes associated with cartilage (Lincoln et al., 2007). In humans,

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mutations in SOX9 are associated with campomelic dysplasia (CD), a disease characterized by generalized hypoplasia of endochondral bones and sex reversal with occasional defects in pancreas and/or kidney (Foster et al., 1994; Wagner et al., 1994).

While SOX9 mutations have not been associated with human valve malformations, increased Sox9 expression has been reported with mitral valve calcification in human patients (Caira et al., 2006).

In mouse and chicken embryos, Sox9 is expressed in the endocardial cushions and remodeling valve leaflets, but not in the supporting chordae tendineae (Lincoln et al., 2006a; Lincoln et al., 2007). Loss of Sox9 in mice results in embryonic lethality between E11.5 and E12.5 with hypoplastic ECs (Akiyama et al., 2004). In addition,

NFATc1 is misexpressed in the cushion mesenchymal cells, which may indicate disruption of the endothelial delamination process during EMT (Akiyama et al., 2004).

Tie2Cre mediated loss of Sox9 in the endocardial cushions demonstrates that Sox9 also is required for cell proliferation and expansion of the valve progenitor pool (Lincoln et al.,

2007). In differentiating cartilage progenitor cells, Bmp2 activates expression of Sox9 and the cartilage differentiation marker aggrecan (Chimal-Monroy et al., 2003). This same regulatory interaction is observed in cultured avian endocardial cushion cells

(Lincoln et al., 2006a). Likewise targeted loss of Sox9 with Col2a1Cre in the remodeling valve leaflets in mice results in decreased expression of cartilage-associated proteins,

Col2a1 and Hapln1, further supporting the parallels between valve and cartilage development (Lincoln et al., 2007). In adult mice, heterozygous loss of Sox9 in

Col2a1Cre expressing cells results in thickened valve leaflets and calcification

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characteristic of valve disease (Lincoln et al., 2007). Together these studies demonstrate that Sox9 has critical functions in endocardial cushion formation and valve remodeling and may also serve as a protective factor in calcific valve disease.

Scleraxis

Scleraxis (Scx) is a basic-helix-loop-helix (bHLH) transcription factor expressed in tendons and ligaments as well as in the developing heart valves (Levay et al., 2008;

Lincoln et al., 2006a; Schweitzer et al., 2001). In tendons, Scx is induced at the earliest stages of cell lineage specification, and it also regulates tendon differentiation through activation of structural protein genes including Col14a1 and tenascin (Brent and Tabin,

2004; Schweitzer et al., 2001). In mice, global deletion of Scx, results in severe tendon defects associated with limited use of all paws and back muscles with complete immobility of the tail (Murchison et al., 2007). These mice are viable, but they also have developmental defects in valve formation and progressive valve disease (Levay et al.,

2008). Interestingly, Scleraxis mutations have not been reported in the human population and there are no reports of Scleraxis expression associated with human valve disease.

The chordae tendineae and other supporting structures of the valves share similarities in ECM composition and gene expression with tendon cell lineages (Lincoln et al., 2006a). In both mouse and chicken embryos, Scx expression is induced in the remodeling valves and is predominant in the chordae tendineae of the AV valves (Levay et al., 2008; Lincoln et al., 2006a). In developing tendons, FGF4 promotes expression of

286

Scx and its downstream target tenascin while inhibiting cartilage cell lineage

development (Edom-Vovard et al., 2002). Likewise, in avian endocardial cushion cell

cultures, FGF4 treatment promotes Scx and tenascin expression, while inhibiting

expression of Sox9 and aggrecan (Lincoln et al., 2006a). In mice, genetic deletion of

Scx leads to thickening of the heart valves, decreased expression of Col14a1, and

increased expression of Sox9 (Levay et al., 2008). In addition, mesenchymal cell markers Msx1 and Snai1 persist during the later stages of valve development, which

may be indicative of a failure of these valve progenitors to differentiate (Levay et al.,

2008). Together these studies support a critical role for Scx in the specification and

differentiation of heart valve supporting structures with molecular properties similar to

tendons.

Overall conclusions and future perspectives

Heart valve development is characterized by complex regulation of EMT, cell

proliferation, cell lineage determination, ECM gene expression and morphogenesis.

Each of these processes is regulated by multiple signaling pathways and transcription

factor interactions. Here we discuss the roles of seven transcription factors in specific

aspects of heart valve maturation. Although significant progress has been made in this

area, we are far from understanding all of the molecular regulatory interactions that

control heart valve development. Certainly the complete regulatory networks are more

complex than those presented here. Strikingly, several of the critical regulatory and

structural genes expressed during valve development are associated with genetic

287

lesions that cause congenital valve malformations. These genes and others expressed during valve development are strong candidates in future genetic studies of familial cardiovascular disease. In addition there is increasing evidence that critical regulators of valve development also are expressed during valve pathogenesis in adults. Therefore, the signaling pathways and transcriptional regulators active during valve development represent attractive therapeutic targets in the management and treatment of valve malformations and degenerative disease in a clinical setting.

288

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