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Investigation of Notch and nitric oxide signaling in the cardiovascular system Chang, Alex Chia Yu 2012

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INVESTIGATION OF NOTCH AND NITRIC OXIDE SIGNALING IN THE CARDIOVASCULAR SYSTEM  by ALEX CHIA YU CHANG B.Sc., The University of British Columbia, 2006  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE STUDIES (Experimental Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2012  © Alex Chia Yu Chang, 2012  ABSTRACT Notch signaling is evolutionarily conserved and regulates various developmental and pathological processes. We are interested in the role of Notch signaling in cardiac valve formation and postnatal vascular remodeling. The heart is the first organ to form in a developing embryo and a common site of congenital defects. Valvuloseptal defects are the most common of the cardiac anomalies seen in the newborn with a prevalence of 1-2% and are often a result of deregulated atrioventricular canal (AVC) formation. The process of endothelial-to-mesenchymal-transition (EndMT) is required for the growth and maturation of the AVC and Notch signaling has been shown to induce this process. In chapter 3, we identify a novel link between Notch and Nitric Oxide (NO) signaling during EndMT. We show that Notchactivation directly induces GUCY1A3 and GUCY1B3, the soluble guanylyl cyclase that forms the heterodimeric NO receptor, during EndMT. We also show that Notch-induced expression and secretion of the TGFβ family member, Activin A, results in increased NO production via a PI3-kinase/Akt signaling mechanism. Paracrine activation of NO signaling by Activin A contributes to early onset of EndMT in the developing AVC. Functional arteries are essential for restoring blood flow and tissue regeneration in response to hypoxia, ischemia, or wound healing. Arterial obstruction can cause distal tissue ischemia and requires rapid reperfusion to limit tissue necrosis. Ischemia recovery requires two processes: angiogenesis (capillary sprouting) and arteriogenesis (expansion of existing vessels secondary to mechanical stress or chemical stimuli). Arteriogenesis involves two phases: a rapid vasodilatory phase followed by vascular expansion and remodeling. In chapter 4, we examine the relationship of endothelial Notch signaling and arteriogenesis using a hindlimb ischemia model. As the link between Notch and NO signaling unfolded in the heart study, we turned our attention to the NO pathway in the initial phase of arteriogenesis – vasodilation. The data presented in this dissertation defines a novel link between Notch signaling and NO signaling in the cardiovascular system and may help to explain Notch-induced EndMT in other pathologies.  ii  PREFACE The work described in Chapter 3 has been published in Developmental Cell (see below for full citation). The work described in Chapter 4 is unpublished but has been written up as a short report for submission. Many people collaborated in generation of this data and three collaborative publications that are not included in my thesis study are listed below. Alex Chia Yu Chang and Aly Karsan conceptualized the experiments as well as reviewed the manuscripts. The projects were led by Alex Chia Yu Chang who prepared all written work and figures, and performed experiments with exceptions of those listed below: Chapter 3: Kyle Niessen and YangXin Fu performed the microarray study. Validations and corresponding figures were generated by Alex Chia Yu Chang. YangXin Fu and Michele Fournier helped with the Flag-CSL chromatin immunoprecipitation experiments. Audi Setiadi performed and optimized Notch conditioned medium experiments. Justin Smrz helped with some of the AVC explants and Victoria Garside (Dr. Pamela Hoodless Lab) performed in situ hybridization and AVC explant immunofluorescent staining experiments. Chapter 4: Michelle Ly performed all the hindlimb ischemia surgeries. Katherine Lu performed immunofluorescent staining on murine collateral vessel cryosections and analyzed the protein expression using a “tumor mapping” algorithm written by Alastair Kyle (Dr. Andrew Minchinton Lab). Megan Fuller and Justin Smrz assisted with animal experiments. Chapter 5: Figure 5.1 is a modified version of a figure created by Dr. Aly Karsan. The following are the list of manuscripts I have published during my graduate school career. For each publication, I have summarized the major findings and have indicated my contribution to the published data. Fu Y, Chang A, Chang L, Niessen K, Eapen S, Setiadi A, and Karsan A. (2009) Differential regulation of TGFbeta signaling pathways by notch in human endothelial cells. J Biol Chem 284(29): 19452-62.  iii  •  This peer-reviewed article showed that Notch pathway synergistically interacts with TGFbeta pathway through Smad3 to induce EndMT.  •  I was involved in generating all the data for Figure 4 and parts of Figures 7 and 8.  Vrljicak P, Chang AC, Morozova O, Wederell ED, Niessen K, Marra MA, Karsan A and Hoodless PA. (2010) Genomic Analysis Distinguishes Phases of Early Development of the Mouse Atrio-Ventricular Canal. Physiological Genomics 40(3): 150-7. •  This peer-reviewed article showed temporal distribution of mesenchymal genes during the EndMT process and of specific Notch and TGFβ targets.  •  I was involved in generating parts of Figures 1, 3 and 4.  Fu Y*, Chang AC* (co-first author), Fournier M, Chang L, Niessen K, and Karsan A. (2011) RUNX3 maintains the mesenchymal phenotype after termination of the Notch signal. (*co-author) J Biol Chem 286(13):11803-13. •  This peer-reviewed article identified RUNX3 as a novel Notch target in the developing AVC.  •  I was involved in generating all the data for Figure 7 and parts of Figures 1, 2, and 5.  Chang AC, Fu YX, Garside VC, Niessen K, Chang L, Fuller M, Setiadi A, Smrz J, Kyle A, Minchinton A, Marra M, Hoodless P, and Karsan A (2011). Notch initiates the endothelial-to-mesenchymal transition in the atrioventricular canal through autocrine activation of soluble guanylyl cyclase. Dev Cell 21(2): 288300. •  This peer-reviewed article identified GUCY1A3, GUCY1B3 and INHBA as novel Notch targets and showed that Notch signaling can activate the nitric oxide pathway to promote EndMT in the developing AVC and has been modified and presented as Chapter 3 in this dissertation.  iv  •  I wrote this paper. I was involved in generating data for every figure in this manuscript. This dissertation uses data that was established in this manuscript.  The use of mouse models was approved by the UBC Animal Care and Ethics Committee (protocols: A07-0717 and A07-0197). The use of biohazardous materials and chemicals were approved by the UBC Biosafety Committee (Protocol B11-0098).  v  TABLE OF CONTENTS Abstract.......................................................................................................................... ii  Preface .......................................................................................................................... iii  Table of Contents ......................................................................................................... vi  List of Figures .............................................................................................................. ix  List of Abbreviations.................................................................................................... xi  Acknowledgements.................................................................................................... xiii  Dedication ................................................................................................................... xiv  Chapter 1 – Introduction ............................................................................................... 1  1.1 Notch Signaling ................................................................................................................. 1  1.2 Nitric Oxide Signaling ....................................................................................................... 5  1.2.1 Distribution and function of the NOS proteins ............................................................... 7  1.2.2 NO Receptors: soluble Guanylyl Cyclases .................................................................... 8  1.3 Cardiac development ........................................................................................................ 8  1.4 Vascular development..................................................................................................... 13  1.4.1 Vasculogenesis and angiogenesis in embryonic development ................................... 13  1.4.2 Arteriogenesis in embryonic development................................................................... 15  1.5 Notch and NO pathways in the cardiovascular system ............................................... 16  1.5.1 Notch signaling and the cardiovascular system .......................................................... 16  1.5.2 NO signaling and the cardiovascular system .............................................................. 18  1.6 Mutations of the Notch pathway in human diseases ................................................... 19  1.6.1 Aortic valve disease .................................................................................................... 19  1.6.2 Alagille syndrome ........................................................................................................ 20  1.6.3 Cerebral autosomal dominant arteriopathy with subcortical infarcts and leukoencephalopathy (CADASIL) ......................................................................................... 21  vi  1.7 Aim of the study............................................................................................................... 23   Chapter 2 – Materials and methods ........................................................................... 25  2.1 Reagents........................................................................................................................... 25  2.2 Cell culture, gene transfer and RNA interference ......................................................... 26  2.3 Immunoblotting................................................................................................................ 27  2.4 In situ hybridization ......................................................................................................... 28  2.5 RNA collection and RT-PCR ........................................................................................... 29  2.6 Brightfield imaging and immunofluorescent imaging ................................................. 29  2.7 Chromatin immunoprecipitation .................................................................................... 30  2.8 Mouse models .................................................................................................................. 31  2.9 Atrioventricular canal (AVC) explant assay .................................................................. 31  2.10 Analysis of AVC explants and cardiac cushions........................................................ 33  2.11 Hindlimb ischemia ......................................................................................................... 33  2.12 Aortic ring myograph .................................................................................................... 34  2.13 β-Galactosidase detection ............................................................................................ 34  2.14 Data analysis .................................................................................................................. 35   Chapter 3 – Notch signaling activates nitric oxide pathway in a cell nonautonomous fashion to drive ENDMT in developing AVC ........................................ 37  3.1 Introduction ...................................................................................................................... 37  3.2 Results .............................................................................................................................. 39  3.2.1 GUCY1A3 and GUCY1B3 are novel Notch targets ..................................................... 39  3.2.2 Notch-induced sGC drives EndMT .............................................................................. 43  3.2.3 Notch activates NO synthesis in a cell non-autonomous fashion. ............................... 46  3.2.4 Notch-induced Activin A activates eNOS and promotes EndMT. ................................ 52   vii  Chapter 4 – Notch induction of nitric oxide participates in the initial vasodilatory response during postnatal arteriogenesis ................................................................ 59  4.1 Introduction ...................................................................................................................... 59  4.2 Results .............................................................................................................................. 60  4.2.1 Endothelial Notch activation is not required to maintain blood pressure in the adult .. 60  4.2.2 Endothelial Notch activation is required for vasodilation post-ischemia ...................... 63  4.2.3 Endothelial Notch activation is required for NO generation and signaling post-ischemia ............................................................................................................................................. 68   Chapter 5 – Summary, perspectives and future directions ..................................... 71  5.1 Summary of Notch and the NO pathway in the developing AVC ................................ 71  5.2 Summary of Notch and the NO pathway in post-natal arteriogenesis ....................... 77   References ................................................................................................................... 81  Appendix A. Primers and viral sequences used. ..................................................... 97  Appendix B. Published RBPj binding sites. ............................................................. 99   viii  LIST OF FIGURES Figure 1.1 – Structure of Notch receptor and ligands. .................................................................................. 2  Figure 1.2 – Simplified view of Notch signaling. ........................................................................................... 3  Figure 1.3 – Nitric Oxide signaling. ............................................................................................................... 6  Figure 1.4 – Schematic of embryonic heart development. ......................................................................... 10  Figure 1.5 – EndMT in the developing AVC................................................................................................ 11  Figure 2.1 – Quantification of EndMT in AVC explants .............................................................................. 32  Figure 2.2 – Schematic diagram of aortic ring myograph setup. ................................................................ 36  Figure 3.1 - Evaluation of Notch candidate target gene expression. .......................................................... 41  Figure 3.2 - Notch signaling induced GUCY1A3 and GUCY1B3 expression. ............................................ 42  Figure 3.3 - Endothelial Notch inhibition blocks Gucy1b3 expression in AVC............................................ 44  Figure 3.4 - Pharmacological inhibition of Notch-Gucy1b3 axis blocks AVC EndMT. ................................ 45  Figure 3.5 - Lentiviral inhibition of Gucy1b3 blocks AVC EndMT. .............................................................. 47  Figure 3.6 - In vivo inhibition of sGC blocks early EndMT. ......................................................................... 48  Figure 3.8 - In vivo Notch inhibition blocks nitric oxide production in the AVC. .......................................... 51  Figure 3.9 - Notch activation induces INHBA expression and Activin A release. ....................................... 53  Figure 3.10 - Notch activation induces Inhba expression in vivo. ............................................................... 54  Figure 3.11 - Notch-induced Activin A activates eNOS via the PI3K/Akt pathway. .................................... 56  Figure 3.12 - PI3K/Akt-eNOS axis is important for AVC EndMT. ............................................................... 57  Figure 3.13 - Activin A is important for AVC EndMT. .................................................................................. 58  Figure 4.1 – Endothelial Notch inhibition results in decreased NO but no change in mean arterial pressure. .............................................................................................................................................. 61  Figure 4.2 – Longitudinal weight monitoring of EC-Notch inhibited mice. .................................................. 62  Figure 4.3 – Long-term endothelial Notch inhibition does not affect vasodilatory response. ..................... 64  Figure 4.4 – Impaired postnatal arteriogenesis in VEtTAxTetOS-dnMAML-GFP mice. ............................. 65  ix  Figure 4.5 – Inhibition of Notch signaling does not affect vessel remodeling at early times post-ischemia. ............................................................................................................................................................. 66  Figure 4.6 - Inhibition of Notch signaling does not affect expression of NO receptor Gucy1b3. ................ 67  Figure 4.7 – Notch is required for NO signaling following HLI. ................................................................... 70  Figure 5.1 - Proposed Model of Notch-sGC-NO signaling in developing AVC. .......................................... 76  Figure 5.2 – Proposed model of role of Notch-NO signaling during arteriogenesis ................................... 78   x  LIST OF ABBREVIATIONS ADAM  a disintegrin and metalloprotease domain  ANK  ankyrin  AVC  atrioventricular canal  BH4  tetrahydrobiopterin  cAMP  3’, 5’ cyclic adenosine monophosphate  CSL  C promoter-binding factor 1, suppressor of hairless, and lag-1  CDK  cyclin dependent kinase  cGMP  3’, 5’ cyclic guanosine monophosphate  ChIP  chromatin immunoprecipitation  DAPT  N-[N-(3, 5-difluorophenacetyl)-l-alanyl]-S-phenylglycine t-butyl ester  DLL  delta-like  DNA  deoxyribonucleic acid  DSL  delta/serrate/lag-2  EC  endothelial cell  EGF  epidermal growth factor  EndMT  endothelial-mesenchymal transformation  FAD  flavin adenine dinucleotide  FMN  flavin mononucleotide  HES  hairy enhancer of split  HEY  hairy/enhancer of split-related with YRPQ motif  HLI  hindlimb ischemia  HMEC  human microvascular endothelial cell  HUVEC  human umbilical vein endothelial cell  ITCH  itchy E3 ubiquitin protein ligase homolog  JAG  jagged  LNR  lin-12/notch repeats  MAML  master-mind like xi  NADPH  nicotinamide adenine dinucleotide phosphate  NICD  notch intracellular domain  NLS  nuclear localization signal  NO  nitric oxide  NOS  nitric oxide synthase  PDE  phosphodiesterase  PDZ  post synaptic density protein (PSD95), Drosophila disc large tumor suppressor (DlgA) and zonula occludens-1 protein (Zo-1)  PEST  proline-glutamine-serine-threonine  PKG  cGMP-dependent protein kinase  RAM  RBPJ interacting domain  RBPJ  recombination signal-binding protein 1 for J-kappa  RNA  ribonucleic acid  ROS  reactive oxygen species  RT-qPCR  reverse transcription – quantitative polymerase chain reaction  SAGE  serial analysis of gene expression  sGC/GUCY  soluble guanylyl cyclase  SMA/Acta2  smooth muscle actin, actin alpha 2  SMC  smooth muscle cell  TAD  transactivation domain  TSS  transcription start site  VASP  vasodilator-stimulated phosphoprotein  xii  ACKNOWLEDGEMENTS First I would like to thank my family for their determined support. Thanks to my father, Daniel Chang, who encouraged me to pursue my passion for research and supported me along the way. Thanks to my mother, Sylvia Chang, who constantly cheered me up despite up and downs. Thanks to my siblings, Serena and Andrew, who endured my long work hours, incomprehensible scientific short talks, and supported me with their humor and encouragements. Special thanks to my wife Shirley for all her support and being part of a big accomplishment in my life. I would like to acknowledge my supervisor Dr. Aly Karsan. Without his guidance and trust, I would not be at where I am today. Thank you for providing me the opportunity to be part of a wonderful lab, giving me the freedom to explore and discover, and teaching me the passion to ask fundamental questions. I would also like to acknowledge all members of the Karsan lab, past and present, which have made my PhD journey wonderful. Special thank you goes to Dr. Kyle Niessen and Dr. Yangxin Fu for helping to get started in the lab and providing thoughtful discussion and feedbacks. Big thanks to Dr. Pavle Vrljicak, Sam Lee, Victoria Garside and Alastair Kyle for technical expertise and interesting discussions. Thank you to Gordon Robertson for guiding me into the field of bioinformatics. To my graduate committee members, Drs. Pamela Hoodless, Ismail Laher and Steven Jones, thank you for your dedication and insight into my research project. Financial support for my graduate studies was provided by a PhD Studentship from University of British Columbia.  xiii  DEDICATION  … to all my teachers  xiv  CHAPTER 1 – INTRODUCTION 1.1 Notch Signaling Notch signaling was first identified in Drosophila, where Notch mutants displayed a “notching” wing phenotype, in early 1900’s. The Notch receptor was subsequently cloned and characterized as a transmembrane receptor (Wharton, Johansen et al. 1985). Since the identification and cloning of the receptor, Notch signaling has been carefully characterized to be involved in many biological processes such as angiogenesis, somitogenesis, heart development, neuronal development, and hematopoietic expansion and differentiation (Iso, Hamamori et al. 2003; Gridley 2007; Lewis, Hanisch et al. 2009; Bigas, Robert-Moreno et al. 2010; Liu, Sato et al. 2010; Tao, Chen et al. 2010; Kato 2011). Notch activation requires the binding of a Notch ligand (on the signaling cell) to a Notch receptor (on the receiving cell). Notch receptors are translated as 200-300 kDa precursors, processed in the trans-Golgi by Furin (a process known as S1 cleavage), and presented on the cell surface as noncovalently linked heterodimers (Blaumueller, Qi et al. 1997; Logeat, Bessia et al. 1998; Bush, diSibio et al. 2001). Furin modification occurs at the S1 cleavage site resulting in an extracellular fragment (containing ligand binding epidermal growth factor (EGF)-like repeats and a transmembrane domain) and an intracellular fragment (containing nuclear localization signals (NLS)) that associates with calcium ion (Figure 1.1)(Blaumueller, Qi et al. 1997). In mammals, there are four Notch receptors (termed Notch1-4) and five Notch ligands (Delta-like (Dll) 1, 3, 4 and Jagged (Jag) 1, 2) (Figure 1.1). Notch activation involves three steps: ligand binding, two proteolytic cleavages releasing the intracellular domain of Notch (NICD), and subsequent translocation of NICD into the nucleus (Figure 1.2). Upon binding of the ligand, the Notch receptor undergoes a conformational change where the extracellular cleavage site becomes accessible. The extracellular proteolytic cleavage termed S2 is mediated by a disintegrin and metalloproteinase 10 and/or 17 (ADAM10 and/or ADAM17) (Wen, Metzstein et al. 1997; Brou, Logeat et al. 2000) and the liberated extracellular portion of Notch is endocytosed  Figure 1.1 – Structure of Notch receptor and ligands. In mammals, there are four Notch receptors (Notch 1 through Notch 4) and five Notch ligands (Jag1/2, Dll1/3/4). Both Notch and Notch ligands are transmembrane proteins. Unlike the ligands, Notch receptors are expressed on cell surfaces as heterodimers stabilized through calcium interactions. There are 29-36 epidermal growth factor-like (EGF) repeats in the extracellular domain of Notch responsible for ligand interaction. Three Lin-12/Notch (LNR) repeats have been shown to be involved in regulating receptor activity. The Notch intracellular domain (NICD) does not contain a DNA binding domain. NICD localizes to downstream target promoters by interacting with RBPJ through a RBPJ interacting domain (RAM). Protein-protein interaction is mediated by Ankyrin (ANK) repeats. Two nuclear localization signal (NLS) domains, a transactivation domain (TAD), and a PEST domain are responsible for NICD nuclear localization, activity and stability respectively. All Notch ligands contain a conserved Delta/Serrate/Lag2 (DSL) domain. The extracellular portion of Notch ligands bind to Notch receptors through EGF repeats. The extracellular portion of Jag1/2 also contains a cysteine-rich domain and a von Willebrand factor Type C domain. Intracellular domains of Jag1 and Dll1 contain a PDZ motif.  2  Figure 1.2 – Simplified view of Notch signaling. Notch signaling activation involves two cells: a ligand presenting cell and a receptor expressing receiving cell. Upon ligand binding, Notch receptors on the receiving cell undergo conformational changes resulting in exposure of proteolytic sites. Proteolytic cleavages release the intracellular domain of Notch resulting in translocation into the nucleus, binding to DNA-binding partner RBPJ/CSL and recruitment of co-activators such as MAML to activate transcription of downstream target genes.  3  into the ligand signaling cell (Parks, Klueg et al. 2000; Morel, Le Borgne et al. 2003). The remaining Notch receptor, resulting from loss of the extracellular portion, undergoes another conformational change and undergoes a second proteolytic (S3) cleavage mediated by the γ-secretase complex composed of presenilin-1, presenilin-2, Pen-2, Aph-1 and nicastrin (De Strooper, Annaert et al. 1999; Struhl and Greenwald 2001; Francis, McGrath et al. 2002; Hu, Ye et al. 2002). This proteolytic cleavage releases the Notch intracellular domain (NICD) which subsequently translocates to the nucleus (De Strooper, Annaert et al. 1999; Okochi, Steiner et al. 2002). The NICD contains 2 nuclear NLS (Stifani, Blaumueller et al. 1992), and upon release from the membrane tether, NICD translocates into the nucleus and binds to DNA-interacting RBPJ (Recombination signal-Binding Protein 1 for J-Kappa) through the RBPJ interacting (RAM) and the ankyrin (ANK) repeat domains (Kurooka, Kuroda et al. 1998; Beatus, Lundkvist et al. 2001; Zweifel, Leahy et al. 2003; Ehebauer, Chirgadze et al. 2005; Ong, Cheng et al. 2006). RBPJ is also known as CSL: CBF1 (C Promoter-Binding Factor 1), Suppressor of Hairless, and Lag-1. In the absence of NICD, RBPJ forms a repressor complex with either the silencing mediator of retinoid and thyroid hormone receptors/nuclear receptor co-repressor/histone deacetylase 1 (SMRT/NcoR/HDAC1) or the CBF1-interacting corepressor/histone deacetylase 2/Sin3A-associated protein 30 kDa (CIR/HDAC2/SAP30) (Kao, Ordentlich et al. 1998; Hsieh, Zhou et al. 1999). The binding of NICD causes the displacement of the repressor complex bound to RBPJ and recruits coactivators such as MAML (Master-mind Like) and the histone acetyltransferase (HAT) complex through the ANK and the transactivator domain (TAD) (Hsieh, Henkel et al. 1996; Kurooka, Kuroda et al. 1998; Tani, Kurooka et al. 2001; Wu, Sun et al. 2002). This activation results in transcription of target genes such as the hairy enhancer of split (HES) and hairy/enhancer of split-related with YRPW motif (HEY, also called HESR, CHF, HRT) family proteins (Davis and Turner 2001; Iso, Chung et al. 2002). RBPJ has been shown to bind to a (C/T)(A/G)TG(A/G/T)GA(A/G/T) motif (Pursglove and Mackay 2005); most validated targets possess a more stringent (C/T)(A/G)TGGGAA binding sequence. Validated RBPJ binding sites are listed in Appendix B. Previous work in this lab has  4  identified novel Notch targets: ACTA2 (also known as smooth muscle actin, SMA), SNAI2, SMAD3, and RUNX3 (Noseda, Fu et al. 2006; Niessen, Fu et al. 2008; Fu, Chang et al. 2009; Fu, Chang et al. 2011). Notch signaling is terminated by two modifications: phosphorylation and subsequent ubiquitination. Besides acting as a coactivator, MAML can recruit cyclin dependent kinase 8 (CDK8) which hyperphosphorylates serine residues of the PEST domain in NICD (Fryer, White et al. 2004). Following phosphorylation, F-box and WD repeat domain containing 7 (FBW7/SEL10) is recruited and ubiquitinates NICD thus targeting it for proteasomal degradation (Fryer, White et al. 2004). Alternatively, others have shown that NICD phosphorylation occurs through glycogen synthase kinase 3 beta (GSK3β) followed by NICD ubiquitination by the ITCH and c-CBL E3-ubiquitin ligases (Qiu, Joazeiro et al. 2000; Foltz, Santiago et al. 2002; Jehn, Dittert et al. 2002; Espinosa, Ingles-Esteve et al. 2003). 1.2 Nitric Oxide Signaling Nitric oxide (NO) is a multifunctional gaseous molecule that has a very short half-life. NO has multiple roles depending on the concentration and source. NO can act as a neurotransmitter, a neuromodulator, an anti-thrombotic agent, a pro- or anti-inflammatory agent, a proliferative, or a cytotoxic agent. But most importantly, the NO pathway has been well established to induce a vasodilatory response in the vasculature (Murad 2006; Yetik-Anacak and Catravas 2006). Conversely, NO dysregulation has been implicated in various cardiovascular diseases such atherosclerosis, as well as in cancer (Napoli and Ignarro 2001; Munzel, Daiber et al. 2005; Fukumura, Kashiwagi et al. 2006; YetikAnacak and Catravas 2006). Biologically, NO is synthesized by nitric oxide synthases (NOS) (Ignarro, Harbison et al. 1986; Murad 1986). Vasoconstriction of vessels is initiated by the influx of calcium ions into smooth muscle cells (SMC). Calcium ions can be released from the sarcoplasmic reticulum within a cell (Maclennan and Zvaritch 2011) or can enter from the extracellular space (Hill, Zou et al. 2001; Fuchs, Dietrich et al. 2010; Zholos, Johnson et al. 2011). In the presence of high concentrations, calcium binds to calmodulin and this binding activates myosin light chain kinase (MLCK) by phosphorylation of Ser19. This activation allows interaction between MLCK and actin filaments and results in SMC contraction. In the vasculature, NO 5  VEGF, insulin, IGF1, shear stress, estrogen  Endothelial Cell  CaM P-Akt  HSP90 P-eNOS L-Arg + O 2  NO + L-Citrulline Ca 2+  Cav -1  Ca 2+ NO  Smooth Muscle Cell  PKG GTP  GUCY1A3/1B3  Actin phosphorylation  cGMP  PDEII  Figure 1.3 – Nitric Oxide signaling. In the cardiovascular system, NO synthesis occurs in endothelial cells. Upon stimulation with external stimuli, activation of the PI3K/Akt results in phosphorylation of eNOS at Ser1177. This phosphorylation activates the eNOS complex and results in the synthesis of NO through catalysis of a reaction involving L-Arginine and oxygen. Vascular smooth muscle vasodilation is achieved when NO diffuses into a neighboring cell, binds to sGC and amplifies the signal through synthesis of the second messenger cGMP. Increased concentration of cGMP activates cGMPdependent kinase PKG which in turn blocks calcium influx channels and activates downstream pathway to dephosphorylate actin filaments. NO signaling is shut off by phosphodiesterases that hydrolyze the second messenger cGMP.  6  acts as a vasodilator and is synthesized in the endothelium of the vasculature. NO diffuses into the SMC and binds to the soluble Guanylyl Cyclase (sGC) heterodimer. The sGC heterodimer consists of an alpha and a beta subunit. This binding of NO causes a conformational change of sGC and activates cyclase activity to generate second messenger 3’, 5’ cyclic guanosine monophosphate (cGMP). Increased cGMP levels activate cGMP-dependent protein kinase (PKG or PRKG) to block calcium channels and dephosphorylate myosin light chain kinase, thus releasing actin filaments (Hofmann, Ammendola et al. 2000). In addition, PRKG-dependent phosphorylation of actin capping protein vasodilator stimulated protein (VASP) has been shown to be important in cell morphology and cell adhesion (Kwiatkowski, Gertler et al. 2003). PRKG also serves as a negative feedback regulator of the NO pathway where activation of downstream phosphodiesterase 5 (PDE5) hydrolyzes cGMP thus turning the pathway off (Essayan 2001; Rybalkin, Yan et al. 2003). 1.2.1 Distribution and function of the NOS proteins There are three NOS proteins: endothelial NOS (eNOS or NOS3), inducible NOS (iNOS or NOS2) and neuronal NOS (nNOS or NOS1). NOS2 was originally cloned from macrophages and studies showed that NO generated through this enzyme functions to kill pathogens (Lowenstein, Glatt et al. 1992). NOS1 was cloned from neurons (Bredt, Hwang et al. 1991), and NOS3/eNOS was cloned from the vascular endothelium (Marsden, Heng et al. 1993). In general, all three NOS proteins dimerize and form a zinc-thiolate cluster with a zinc ion that is tetrahedrally coordinated to 2 CXXXXC motifs (1 motif per monomer) (Raman, Li et al. 1998). The N-terminal, also known as the oxygenase domain, binds tetrahydrobiopterin (BH4), molecular oxygen and the NO substrate L-Arginine. The C-terminal, also known as the reductase domain, consists of a nicotinamide adenine dinucleotide phosphate (NADPH) binding domain, a flavin mononucleotide domain (FMN) and a flavin adenine dinucleotide (FAD) domain (Forstermann, Closs et al. 1994). Since this dissertation studies endothelial (EC) biology, eNOS will be the focus from here on in. In EC, eNOS localizes to the caveolae through (1) irreversible myristoylation (addition of a 14 carbon chain) on N-terminal glycine and (2) post-translational palmitoylation (addition of a 16 carbon 7  chain) at Cys15 and Cys26 (Dudzinski and Michel 2007). Prolonged agonist stimulation causes depalmitoylation of eNOS and releases eNOS into the cytosol (Dudzinski and Michel 2007). In the presence of increased intracellular calcium, the eNOS activator calmodulin bind to eNOS and allows better electron transfer from the reductase domain to the oxygenase domain (Dudzinski and Michel 2007). eNOS activity is regulated by phosphorylation at three activating serine sites (Ser1177, Ser635, and Ser617) and two inhibitory sites (Thr495 and Ser116) (Dudzinski and Michel 2007; Mount, Kemp et al. 2007). To generate NO, the enzyme needs to cycle twice. First, NOS hydroxylates L-Arginine into Nω-hydroxyl-L-arginine. Second, an oxidation reaction using an oxygen molecule generates NO and L-Citrulline (Martasek, Miller et al. 1998). This two-step synthesis requires the cofactor BH4 and mutation of the BH4 binding site (Cys331Ala) abolishes NOS activity (Forstermann 2010). Disruption of this process may lead to generation of reactive oxygen species (ROS) and is termed endothelial dysfunction. 1.2.2 NO Receptors: soluble Guanylyl Cyclases There are three classes of guanylyl cyclases: multiple transmembrane class, membrane receptor class, and soluble class. Of the three classes of guanylyl cyclases, NO signaling is propagated through the soluble GC (sGC) heterodimer (Baker and Kelly 2004; Yamagami and Suzuki 2005). The other two classes of guanylyl cyclases do not bind to NO but can also induce vasodilation when bound to natriuretic peptides (Garbers and Lowe 1994). The sGC heterodimer is composed of one alpha subunit (GUCY1A2 or GUCY1A3) and one beta subunit (GUCY1B3) though the functions of the alpha subunits are complementary (Mergia, Friebe et al. 2006). GUCY1A3/1B3 is expressed in most tissues, whereas GUCY1A2/1B3 is predominantly found in the brain (Giuili, Scholl et al. 1992; Mergia, Russwurm et al. 2003). In addition to blood pressure regulation, sGC proteins have also been shown to promote endothelial cell survival and migration (Pyriochou, Beis et al. 2006; Pyriochou, Zhou et al. 2007). 1.3 Cardiac development Congenital heart defects occur in ~2% of newborns each year. Of these, valvular defects are the most prominent (Hoffman and Kaplan 2002). In both human and mice, the heart is the first organ to develop in the embryo. Cardiac development is initiated by differentiation of cardiac progenitors, which 8  are specified in the primitive streak at embryonic day (E) 7.0. At E7.0, myocardial markers (Nkx2.5, Gata4, eHand) are induced by the Bmp2 and Noggin signaling in the cardiac progenitors (Schlange, Andree et al. 2000). Lineage tracing studies of Cited2, Medp1 and Mesp2 have shown that cardiac progenitor cells arise from mesoderm at E7.0 (Kitajima, Takagi et al. 2000; Schlange, Andree et al. 2000). Cardiac progenitors then undergo a mesenchymal transition and migrate out from the primitive streak, moving laterally on both sides, to form the left and right heart fields at E7.5. Anterior-medial fusion of the heart fields establishes the cardiac crescent at E8.0 which fuses along the embryonic midline giving rise to the primitive heart tube at E8.5. At E9.0, the heart tube loops rightward and compartmentalization of atria, atrioventricular canal (AVC), ventricle, and outflow tract (OFT) begins (Figure 1.4). For the heart to deliver sufficient oxygen and nutrients to the growing embryo, the organ needs to develop into a functional four-chambered heart and establish unidirectional blood flow. This septation of the heart requires the formation of the septum and the heart valves to ensure unidirectional blood flow. The mature valve leaflets are enclosed in a sheath of endocardial cells with intervening valve interstitial cells. The valves are stratified into three extracellular matrix layers: an atrialis or a ventricularis layer (enriched with elastin), a spongiosa layer (enriched with proteoglycan) and a fibrosa layer (enriched with collagen) (Hinton, Alfieri et al. 2008). The valvulogenic process in the OFT and AVC forms four sets of heart valves: the mitral valve (regulates blood flow from left atrium to left ventricle), the tricuspid valve (prevents blood backflow between right atrium and ventricle), the aortic valve (regulates blood flow from the left ventricle into the aorta) and the pulmonary valve (regulates blood flow from the pulmonary vein into the right ventricle) (Snarr, Kern et al. 2008). Lineage tracing experiments have shown that the aortic leaflet in the mitral valve and the septal leaflet in the tricuspid valve are derived from the inferior and superior AVC; moreover, the mural leaflet in the mitral valve and the antero-superior and posterior inferior leaflets in the tricuspid valve are derived from lateral AVCs (de Lange, Moorman et al. 2004; Snarr, Kern et al. 2008). However, less is known about the exact contribution to the aortic and pulmonary valves during OFT fusion (Combs and Yutzey 2009).  9  RA  LA  OFT AVC RV  LV  LA RA RV  LV  Figure 1.4 – Schematic of embryonic heart development. E8.5 (Day 21 for humans) linear heart tube is depicted with the inner lining of endothelium (blue) and the outer layer of myocardium (peach). Compartmentalization begins after heart looping at E9.5 (Day 28 for humans) and eventually gives rise to the four mature chambers at E12.5 (Day 50 for humans). RA = right atrium; LA = left atrium; RV = right ventricle; LV = left ventricle; OFT = outflow tract; AVC = atrioventricular canal.  10  A  E8.5  E9.5  E10.5  E11.5  B Notch Activation EC  Migrating Mesenchyme  TGFbeta2 MYO  Figure 1.5 – EndMT in the developing AVC. (A) Representative micrographs of embryonic hearts from various developmental stages (the images are from www.mouseatlas.com). AVCs have been marked with red dotted lines. (B) Schematics representing cross-section through the AVC during EndMT. Upon Notch activation, endocardial cells in the AVC lose cell-cell contact and acquire a spindle-like morphology and invade and migrate into the cardiac cushion. EC = Endocardial Cells; MYO = Myocardium.  11  At E9.0, the AVC is comprised of a monolayer of endothelium and several layers of myocardium. The myocardium secretes extracellular matrix into the interstitial space and this extracellular matrix is often referred as the cardiac jelly. During septation and valvulogenesis, the endothelium undergoes endothelial-to-mesenchymal-transition (EndMT) and cells populate the cardiac jelly. EndMT is the process by which endocardial cells loose cell-cell junctions, transitions into mesenchymal cells and acquire a more migratory phenotype. Next, the cushions grow and the fusion of aorticopulmonary septum, outflow septum, interventricular septum, atrioventricular septum and primary atrial septum divides the heart into four chambers (Nakajima, Yamagishi et al. 2000). The valve primordia continue to grow and elongate and eventually give rise to the mature valves (Hinton, Alfieri et al. 2008). The ErbB signaling pathway and hyaluronic acid have been shown to play a key role in regulating cardiac jelly formation. Hyaluronic acid is a glycosaminoglycan that forms the majority of the cardiac jelly (Day and Prestwich 2002) and deletion of HA-synthase 2 in mice results in absence of cardiac jelly and causes a block of EndMT (Camenisch, Spicer et al. 2000; Camenisch, Schroeder et al. 2002). At E9.5, endocardial cells of the AVC and OFT are activated to undergo EndMT by signals from both the myocardium and other endocardial cells (Figure 1.5). Various pathways tightly regulate this developmental process (Armstrong and Bischoff 2004; Srivastava 2006; Niessen and Karsan 2008), but for the focus of this dissertation work, we examined the Notch pathway in AVC EndMT. Previous work from our lab has shown that Notch activation induces SNAI2, a transcriptional repressor, and facilitates EndMT by down regulating EC adherens junction protein VE-cadherin (Niessen, Fu et al. 2008). This downregulation of VE-cadherin releases these Notch-activated endocardial cells allowing them to invade and migrate into the cushions. Moreover, we have also shown that RUNX3, a Notch target induced during EndMT, can also induce the expression of SNAI2 after Notch signaling is turned off thus ensuring the downregulation of VE-cadherin in mesenchymal cells (Fu, Chang et al. 2011). Although these findings show an important participation of Notch pathway during EndMT, a direct link to a migratory pathway is lacking. 12  1.4 Vascular development In the developing embryo, formation of the vascular network is essential for nutrient, oxygen and waste exchange to sustain the growing embryo. The vasculature is formed in an orchestrated manner and disruption of this process leads to embryonic lethality. The vasculature is composed of EC, that line the inner vessel walls, and SMC/mural cells which surround the endothelial cells. There are three independent mechanisms that contribute to overall vasculature formation: vasculogenesis, angiogenesis and arteriogenesis. Vasculogenesis is the process by which clusters of EC assemble into the primary capillary plexus, a network with uniform vessel diameter. This process is thought to be limited to embryonic development (Flamme, Frolich et al. 1997; Ferguson, Kelley et al. 2005). Angiogenesis is the formation of new blood vessels from an existing vascular network. The primitive plexus formed through vasculogenesis is then remodeled into a network of blood vessels with various diameters through angiogenesis (Carmeliet 2000). During this process, vessels are pruned and new vessels are formed by either intussusception or sprouting (Carmeliet 2000). During angiogenic sprouting, the extracellular membrane is degraded, EC sprout from the existing endothelial tube, proliferate, migrate and form new EC tube structures. Mural cells are then recruited and stabilized by the deposition of extracellular matrix. This process forms both arteries and veins (Carmeliet 2003). The artery/vein specification and markers are discussed below. Arteriogenesis refers to an increase in the diameter of existing arteries. This remodeling process involves the thickening of the surrounding medial layer comprised of vascular SMC. In the adult, the established vasculature is relatively quiescent, with the exception of during menstruation (Becker and D'Amato 2007) and in pathological conditions such as tumor growth (Patenaude, Parker et al. 2010), ischemia in the cardiovascular system and wound healing (Carmeliet 2003). 1.4.1 Vasculogenesis and angiogenesis in embryonic development Vasculogenesis occurs spatially and temporally prior to angiogenesis during development and can be subdivided into two categories: extraembryonic and intraembryonic. The first vascular structure formed extraembryonically is the yolk sac (Ferguson, Kelley et al. 2005). The mesodermal precursors 13  from the primitive streak migrate to the yolk sac and form blood islands, differentiate into angioblasts, and form endothelial-lined cords filled with primitive blood cells. These endothelial cords then fuse with one another to form the primary capillary plexus of the yolk sac (Ferguson, Kelley et al. 2005). Similarly, in the embryo proper, angioblasts line up along the midline followed by specification and lumen formation giving rise to the dorsal aorta and the cardinal vein (Drake and Fleming 2000). Recent findings showed that murine embryonic aorta lumen formation is accomplished by series of myosin/actin rearrangements emanating from adherens junction VE-cadherin to create mechanical forces that pulls apart individual endothelial cells (Strilic, Kucera et al. 2009). Normally, arteries deliver oxygenated blood to the body away from the heart with high flow rates. Veins return the low-oxygenated blood back to the pulmonary system and into the lungs for reoxygenation. It was thought that the endothelial structures were first formed and the hemodynamic forces from the circulation induced arterial-venous specification. Refuting this view, studies have shown angioblasts with specific arterial markers (EphrinB2 (Wang, Chen et al. 1998), Neuropilin1 (Herzog, Kalcheim et al. 2001), Notch4 and Dll4 (Villa, Walker et al. 2001), and CD44 (Wheatley, Isacke et al. 1993)) and venous markers (EphrinB4 (Wang, Chen et al. 1998), Neuropilin2 (Herzog, Kalcheim et al. 2001), and COUP-TFII (You, Lin et al. 2005)) migrating from the mesoderm prior to establishment of circulation suggesting that the arterial/venous fate is predetermined. During angiogenesis, vascular endothelium is activated by angiogenic factors to delaminate and invade the extracellular matrix in a dose-dependent manner. At the site of angiogenic sprouting, special sets of EC are activated and begin to present filopodial protrusions to explore the local environment. During this process, cells that receive the angiogenic signal become the tip cell and activate Notch signaling to ensure other endothelial cells remain as stalk cells. The objective is to have the tip cell lead the way and form vessels in hypoxic regions, whereas stalk cells proliferate and form the walls of the lumen of the new vasculature. One of the most characterized angiogenic cytokines is vascular endothelial growth factor (VEGF). Tip cells are characterized by the ability to sense VEGF concentrations through their expression of vascular endothelial growth factor receptor 2 (VEGFR2) (Gerhardt, Golding et al. 14  2003). Endothelial stalk cells proliferate and elongate the endothelial sprout guided by the tip cells. One of the mechanisms to ensure only a few tip cells are licensed while the stalk cells remain stalk cells is through VEGF-Dll4 signaling. In the tip cells, binding of VEGF induces the expression of Dll4 ligands which can then activate Notch signaling in the neighboring stalk cells (Lawson, Vogel et al. 2002). This activation of Notch signaling in the stalk cells causes a transcriptional repression of VEGFR thus preventing the stalk cell from becoming a tip cell (Liu, Shirakawa et al. 2003; Suchting, Freitas et al. 2007). Examination of heterozygous deletion of Dll4 in mice demonstrated that disruption of this tip cellstalk cell balance resulting in a dramatic increase in the number of angiogenic sprouts yielding collapsed and unperfused vessels (Suchting, Freitas et al. 2007). Similarly, use of a neutralizing antibody against Dll4 in tumors showed an increase in tip cells as well as non-perfused vasculature (Ridgway, Zhang et al. 2006). Genetic mutations altering either the bioavailability of VEGF (Carmeliet, Ferreira et al. 1996; Ferrara, Carver-Moore et al. 1996; Damert, Miquerol et al. 2002; Duan, Nagy et al. 2003) or VEGF receptors (Fong, Rossant et al. 1995; Shalaby, Rossant et al. 1995; Dumont, Jussila et al. 1998) results in embryonic lethality due to vascular malformation defects. The vessel is stabilized by the recruitment of vascular smooth muscle cells (VSMC, for large vessels) and/or pericytes (for microvasculature) (Carmeliet 2003). In the embryo, intersomitic vessels are derived from angiogenic sprouts from the dorsal aorta (Coffin and Poole 1988). 1.4.2 Arteriogenesis in embryonic development Throughout the process of vascular development, vessels expand and the increase in diameter results in increased pressure on the vessel wells. To sustain such an increase in pressure, vessels increase the thickness of the vascular SMC wall to ensure vascular integrity (Carmeliet 2000). Currently it is assumed that signals that induce recruitment and growth of vascular SMC are involved in arteriogenesis. In the absence of vascular SMC, vascular structure is lost and hemorrhage bleeding is observed in murine embryos (Hungerford and Little 1999; Sorensen, Adams et al. 2009). Vascular SMC development initiates at major arteries at E9.5 through E11.5 as observed by staining with a vascular SMC marker α−smooth muscle actin (ACTA2/SMA) (Takahashi, Imanaka et al. 1996). 15  1.5 Notch and NO pathways in the cardiovascular system 1.5.1 Notch signaling and the cardiovascular system Notch1 null animals die at E10.5 with cardiovascular defects and lack of angiogenesis (Krebs, Xue et al. 2000; Grego-Bessa, Luna-Zurita et al. 2007). Specifically, absence of cellularization in the AVC, blocked tuberculation in the ventricles, and disrupted vasculature coupled with hemorrhage suggests Notch1 having multiple roles during cardiovascular development (Krebs, Xue et al. 2000; Grego-Bessa, Luna-Zurita et al. 2007). Notch2 null mice die with vascular and neuronal defects but with no obvious cardiac defects (McCright, Lozier et al. 2002), whereas, Notch2 hypomorphic mice suffer kidney developmental defects (McCright, Gao et al. 2001). Although Notch4 null animals are viable and fertile (Krebs, Xue et al. 2000), Notch1 heterozygous/Notch4 null animals show enhanced cardiovascular defects. These mice display disrupted vasculature with intersomitic hemorrhage, growth retardation and pericardial effusions suggesting overlapping roles in endothelial signaling (Krebs, Xue et al. 2000). Notch1/Notch2 double null animals were reported to have disrupted left-right asymmetry through the regulation of Nodal (Krebs, Iwai et al. 2003). Endothelial-targeted Notch1 knockdown animals die at E10.5 due to cardiovascular failure as evidenced by pericardial and intersomitic hemorrhage (Limbourg, Takeshita et al. 2005). However, the presence of a primary vascular plexus in the yolk sac suggests ECNotch1 inhibition does not block vasculogenesis but is required for vascular remodeling (Krebs, Xue et al. 2000; Limbourg, Takeshita et al. 2005; Grego-Bessa, Luna-Zurita et al. 2007). Conversely, global overexpression of the Notch1 intracellular domain results in growth arrest with abnormalities including lack of neural tube closure, disorganized somites and disrupted vasculature (Liu and Lobe 2007). With respect to Notch ligands, Jag1 null animals die in the embryonic stage at E10.5 due to massive hemorrhage, collapsed vasculature and failure to remodel the vascular plexus (Xue, Gao et al. 1999). However, unlike Notch1 mutants, Jag1 null embryos do not show defects in somite segmentation suggesting ligand-specific Notch functions. Endothelial-targeted Jag1 null animals die in the embryonic period at around E10.5 due to failure in vascular smooth muscle development (High, Lu et al. 2008). In contrast to Jag1, Jag2 null animals die perinatally due to craniofacial morphogenetic defects but no 16  cardiovascular defects (Jiang, Lan et al. 1998). As for the Delta-like ligands, a Dll3 knockout animal has not been reported. Dll1 null animals show disrupted epidermis formation but lack any cardiac defects (Estrach, Cordes et al. 2008). Dll4 null animals showed haploinsufficiency and displayed vascular remodeling defects such failure to remodel the vascular plexus, growth retardation and pericardial effusions (Krebs, Shutter et al. 2004). Disruption of Notch cofactors has also been examined. Rbpj null animals die at E10.5 with cardiovascular defects (Krebs, Shutter et al. 2004; Grego-Bessa, Luna-Zurita et al. 2007). The vascular defects of Rbpj null animals were similar to the Notch1/Notch4 double mutants with sever hemorrhage (Krebs, Shutter et al. 2004; Grego-Bessa, Luna-Zurita et al. 2007). In EC-targeted Rbpj null embryos severe growth retardation and defects in vascular remodeling were observed (Oka, Nakano et al. 1995; Krebs, Shutter et al. 2004). Inhibition of Notch signaling by overexpressing a pan Notch inhibitor, dominant-negative MAML (dnMAML), in the endothelium causes embryonic lethality at E10.5 due to cardiovascular defects (High, Lu et al. 2008; Fu, Chang et al. 2009). Targeting of Notch-induced genes has also implicated the importance of Notch signaling in cardiac development. Hey2 null animals display a phenotype resembling Tetralogy of Fallot, a condition defined by ventricular septal defect, overriding aorta, infundibular pulmonary stenosis, and often right ventricular hypertrophy (Krantz, Smith et al. 1999; Donovan, Kordylewska et al. 2002; McElhinney, Krantz et al. 2002; Kamath, Spinner et al. 2004). Hey1/Hey2 and Hey1/HeyL double knockout animals showed cardiac cushion defects (Fischer, Schumacher et al. 2004; Fischer, Steidl et al. 2007). From these studies we can conclude that most embryonic lethality observed in Notch knockout studies that are due to cardiovascular defect occurs at E10.5, a developmental stage immediately after AVC EndMT. Disruption of the Notch pathway has also implicated disruption of angiogenesis and arteriogenesis as evidenced by hemorrhage. It is also very clear that ligand specific activation and expression of Notch downstream targets are tightly regulated and play a critical role in cardiovascular development. 17  1.5.2 NO signaling and the cardiovascular system eNOS null animals have been reported to exhibit hypertension and some embryos exhibit bicuspid aortic valves (Huang, Huang et al. 1995; Shesely, Maeda et al. 1996; Tsutsui, Shimokawa et al. 2006; Zholos, Johnson et al. 2011). It is interesting to note that the bicuspid aortic valves observed in eNOS null animals has also been reported in humans with a NOTCH1 mutations generating truncated transcripts (Garg, Muth et al. 2005). In endothelial cells, CAV1 protein binds to eNOS and localizes at cell surface caveolae structures. Interestingly, Cav1 null animals show various cardiovascular abnormalities including pulmonary hypertension and cardiac hypertrophy but no bicuspid aortic valves (Razani, Engelman et al. 2001). These studies suggest loss of eNOS protein results in hypertension and bicuspid aortic valves but disruption of eNOS localization does not affect eNOS function. Besides regulating the NO-dependent vasodilatory response, sGC proteins have also been shown to play an important role in platelet aggregation (Hofmann, Ammendola et al. 2000). In Gucy1a2 and Gucy1a3 null animals, Mergia et al. did not observe the predicted hypertensive phenotype (Mergia, Friebe et al. 2006). Instead, measuring sGC activity revealed compensation effect between the two alpha subunits and the animals appear to be normal (Mergia, Friebe et al. 2006). The same group next showed that Gucy1b3 null and SMC-targeted Gucy1b3 null animals exhibited hypertension, impaired gastrointestinal tract peristalsis and platelet coagulation defects (Friebe, Mergia et al. 2007; Dangel, Mergia et al. 2010; Groneberg, Konig et al. 2010; Groneberg, Konig et al. 2011). Although these studies did not report any cardiac defects, whether Gucy1b3 null animals exhibit cardiac defects is yet to be determined. Work in zebrafish (Pyriochou, Beis et al. 2006) has shown that inhibition of sGC blocks angiogenesis and cell migration, which suggested to us that activation of sGC might promote migration and invasion during EndMT. Downstream of sGC the NO signaling pathway is propagated through PRKG. Prkg null animals exhibited abnormal vascular smooth muscle function and increased mean systemic arterial blood pressure (Pfeifer, Klatt et al. 1998; Wegener, Nawrath et al. 2002). Vasp null animals did not exhibit any abnormalities in the adult heart and no obvious defects in smooth muscle function were observed 18  (Aszodi, Pfeifer et al. 1999). However, platelets derived from Vasp null animals exhibited enhanced collagen-induced platelet aggregation and were less sensitive to cGMP/cAMP aggregation block (Zucker 1989; Aszodi, Pfeifer et al. 1999). Taken together, these studies suggest that the hypertensive phenotype observed is due to disruption of NO synthesis and NO pathway activation in a context dependent manner. Disruption of sGC or VASP proteins in platelets may explain the aggregation defect. 1.6 Mutations of the Notch pathway in human diseases 1.6.1 Aortic valve disease In humans, aortic valve disease has been found to be caused by mutation in the Notch1 gene resulting in bicuspid aortic valve leaflets (Garg, Muth et al. 2005). Normal aortic valves should have three rather than two leaflets (Cripe, Andelfinger et al. 2004), and the most common clinical outcome is calcification of a bicuspid aortic valve (Garg, Muth et al. 2005). In the general population, 1-2% newborns are born with congenital heart defects, with bicuspid aortic valve being the most prevalent defect (Hoffman and Kaplan 2002). Bicuspid aortic valves tend to result in calcification and this is the third leading cause of heart disease in adults (Hoffman and Kaplan 2002). In some bicuspid aortic valve patients, mutations at position 4515 in the Notch1 locus result in a premature stop codon in the extracellular domain of Notch1 (Garg, Muth et al. 2005). Currently, the actual mechanism of truncated Notch1 causing bicuspid aortic valves is unknown. It has been suggested that the truncated mRNA may be degraded by the non-sense mRNA decay pathway (Frischmeyer, van Hoof et al. 2002). On the other hand, whether haploinsufficiency of Notch1 or truncated Notch1 protein acting as a dominant-negative can cause the bicuspid aortic valve disease remain to be elucidated. Calcification of the aortic valve is thought to be caused by loss of endothelial Notch which represses the expression of osteogenic genes. Gene expression analysis reveals Notch pathway components are highly expressed in the arterial endothelial cells. Using an in vitro system it was demonstrated that Notch1 activation induces expression of Hey1 and Hey2 which represses Runx2 activity by physically interaction (Garg, Muth et al. 2005). Runx2 is a member of the runt related transcription factor family and encodes a nuclear protein containing Runt DNA binding domain. Runx2 has been linked to valvular calcification where it regulates the 19  expression of osteogenic genes such as osteopontin and osteocalcin (O'Brien, Kuusisto et al. 1995; Ducy, Zhang et al. 1997; Rajamannan, Gersh et al. 2003). Moreover, our lab showed that another Runx member, Runx3, is a novel Notch target in vivo in the AVC and induction of Runx3 in endothelial cell lines can induce the mesenchymal marker Slug even after Notch signaling is turned off (Fu, Chang et al. 2011). Whether Notch1 mutations found in human contributes to decreased Runx3 expression (leading to bicuspid valve disease) and/or lowered expression of Hey1 and Hey2 resulting in higher expression of osteogenic genes through Runx2 remain to be elucidated. 1.6.2 Alagille syndrome Alagille syndrome is an autosomal dominant disorder that affects the heart, kidney, liver and other parts of the body. Alagille syndrome patients are commonly afflicted with neonatal jaundice and impaired development of intrahepatic bile ducts (resulting in cholestasis), abnormalities in the eye, heart, skeleton, and a characteristic facial phenotype with broad forehead, pointed mandible and bulbous tip of the nose (Li, Krantz et al. 1997). Alagille syndrome has been mapped to mutations at the Jag1 locus and mutation of Jag1 accounts for 94% of such patients (Warthen, Moore et al. 2006). Moreover, mutation of the Notch2 locus has also been implicated (McDaniell, Warthen et al. 2006). Molecular studies showed that JAG1 mutations in Alagille syndrome patients result in the expression of a dominant negative form of JAG1 protein that acts to inhibit Notch signaling (Boyer, Crosnier et al. 2005; Boyer-Di Ponio, WrightCrosnier et al. 2007). The most common cardiovascular defect in Alagille syndrome patients is peripheral pulmonic stenosis. In less frequent cases, Alagille syndrome has been associated with cardiac cushion defects (Eldadah, Hamosh et al. 2001; McElhinney, Krantz et al. 2002). Furthermore, 13% of Alagille syndrome patients exhibit Tetralogy of Fallot (Krantz, Smith et al. 1999; McElhinney, Krantz et al. 2002; Kamath, Spinner et al. 2004). However, Jag1 null animals or Notch2 hypomorphic animals that lack ankyrin repeats do not recapitulate the Alagille syndrome phenotype (Hamada, Kadokawa et al. 1999; Xue, Gao et al. 1999). Heterozygous Jag1 mice displayed eye defects while the Jag1-null mice died at E10 due to vascular defects (Xue, Gao et al. 1999). Heterozygous Notch2 mice with a disruption of the ankyrin repeat sequences die prior to E11.5 (Hamada, Kadokawa et al. 1999). Notch 2 animals lacking 20  EGF repeats displayed kidney defects and myocardial hypoplasia (McCright, Gao et al. 2001). Moreover, mice with a genotype of Jag1-null and Notch2-hypomorphic allele developed jaundice, impaired intrahepatic bile duct development, and abnormalities of the eye, heart and kidney, thus reproducing the Alagille syndrome phenotype (McCright, Lozier et al. 2002). The mechanism behind the need for extra Notch2 insufficiency in the mouse model to recapitulate the human Alagille syndrome is unknown. Possible explanations may include a compensatory effect by expression of other Notch ligands or the fact that Jag1 is expressed at higher levels in the mouse compared to the human. 1.6.3 Cerebral autosomal dominant arteriopathy with subcortical infarcts and leukoencephalopathy (CADASIL) CADASIL is an autosomal dominant progressive disorder of arterial vessels in the brain resulting in increased incidence of repeated stroke, migraine, headaches, and white matter lesions that may result in cognitive loss or psychiatric symptoms such as depression in some patients (Joutel, Corpechot et al. 1996; Joutel, Vahedi et al. 1997; Joutel and Tournier-Lasserve 1998; Kalimo, Viitanen et al. 1999; Joutel, Andreux et al. 2000; Joutel 2011). CADASIL results from a progressive loss of vascular SMC and increased vascular fibrosis leading to the narrowing of the lumen of the arteries in the brain (Chabriat, Vahedi et al. 1995). Histologically, vessels from CADASIL patients are often characterized by accumulation of granular osmophilic deposits in the medial layer (Ruchoux, Chabriat et al. 1994; Mayer, Straube et al. 1999). Genetic mapping in CADASIL patients revealed that a missense point mutation in the first 5 EGF repeats in the Notch3 locus accounted for 95% of CADASIL cases (Joutel, Corpechot et al. 1996; Joutel, Vahedi et al. 1997; Dichgans, Ludwig et al. 2000). This Notch3 mutation causes a gain or loss of a cysteine residue, resulting in an odd number of cysteine residues in the EGF repeat region (Dichgans, Ludwig et al. 2000). The mutated EGF repeats are thought to result in a conformational change and cause accumulation of the ectodomain of Notch3 (Joutel, Andreux et al. 2000). Accumulation of the 210 kDa ectodomain of Notch3 in the vascular smooth muscle plasma membrane has been observed in CADASIL patients (Joutel, Andreux et al. 2000).  21  Notch3 expression is restricted to vascular SMC. Animals overexpressing mutated Notch3 (Arg90Cys) transgene under the control of the SM22alpha promoter recapitulate the CADASIL phenotype (Ruchoux, Domenga et al. 2003). In the Notch3-Arg90Cys transgenic animals, the CADASIL phenotype was observed around 10 to 12 months (Ruchoux, Domenga et al. 2003). In a more recent study, the Notch3-Arg170Cys knock-in mouse also recapitulated the CADASIL phenotype with typical granular osmophilic material deposition, developed brain histopathology, but also motor defects such as staggering gait and limb paresis (Wallays, Nuyens et al. 2011). CADASIL Notch3 mutant proteins have been shown to be expressed on cell surfaces, form multimers through sulfhydryl crosslinking, and maintain the ability to bind to Notch ligands (Haritunians, Boulter et al. 2002; Opherk, Duering et al. 2009; Duering, Karpinska et al. 2011). Notch3 activation has been shown to activate cell survival and inhibition of this pathway by accumulation of the ectodomain of Notch3 may explain the loss of vascular smooth muscle cells in CADASIL patients (Wang, Prince et al. 2002; Takahashi, Adachi et al. 2010). CADASIL Notch3 mutant proteins have been shown to recycle at a much slower rate and are retained in endoplasmic reticulum which blocks cell proliferation (Takahashi, Adachi et al. 2010). These findings suggest that accumulation of the ectodomain of Notch3 results in the loss of vascular smooth muscle cells surrounding the vessels both by inhibiting cell proliferation and inducing cell apoptosis. Thus one could speculate that loss of vascular smooth muscle cells may result in loss of vascular integrity potentially resulting in hemorrhage or stroke. As previously discussed, Notch3-null mice are viable and fertile and do not exhibit cardiovascular development defects or CADASIL phenotypes (Krebs, Xue et al. 2003). In parallel, it has been demonstrated that the Notch3-Arg90Cys mutation can still activate the Notch pathway (Monet, Domenga et al. 2007). These results suggest that the Notch3 mutation does not result in the expression of a dominant negative but that the accumulation of the ectodomain of Notch3 may block or activate other pathways by binding to receptors or signaling molecules, or may have a general toxic effect.  22  1.7 Aim of the study As discussed previously, Notch signaling in the endothelium plays a crucial role in cardiovascular development. The aim of this dissertation is to identify downstream pathways that participate in cardiovascular development. As alluded to in the previous sections, in this thesis we present data showing a novel link between Notch and NO signaling. Previous reports have shown that disruption of key Notch members result in cardiovascular defects that lead to embryonic lethality. In this dissertation, we used a tetracycline inducible binary transgenic mouse system. In this system, a driver mouse strain expressing the transactivator protein (tTA) under the regulation of the VE-cadherin promoter (VEtTA) is crossed with a responding strain where the expression of the pan Notch inhibitor dominant-negative MAML fused with GFP is regulated under the TetOS promoter (TetOS-dnMAML-GFP). In the absence of tetracycline, the tTA protein binds to the TetOS promoter, inducing the expression of dnMAML-GFP in the endothelial cells, and thus blocking Notch activation. The advantage of this binary transgenic mouse system is that when animals are treated with tetracycline or doxycycline in the drinking water, tetracycline or doxycycline can bind to the tTA protein and prohibit it from binding to the TetOS promoter thus allowing normal Notch activation. As shown in this dissertation, this binary transgenic model can be used to inhibit endothelial Notch activation at any stage of development rendering the ability to bypass embryonic lethality as seen in Notch1 null animals. Although a handful of genes have been identified as Notch targets, the contribution of individual targets to the Notch phenotype is still lacking. In Chapter 3, we sought to identify novel Notch targets by using a microarray genetic screen in the hope of identifying novel Notch targets. This exercise led to the identification of the soluble nitric oxide receptors, GUCY1A3 and GUCY1B3 as novel Notch targets. Next, we hypothesized that Notch signaling activates the NO signaling pathway by (a) upregulating sGC proteins and/or (b) increasing NO biosynthesis in the developing AVC. Notch signaling has also been shown to play an important role in postnatal arteriogenesis (Limbourg, Takeshita et al. 2005; Limbourg, Ploom et al. 2007). Postnatal arteriogenesis is the expansion of existing vessels secondary to mechanical stress or chemical stimuli and is recognized to be the main 23  repair mechanism following ischemic injury (Carmeliet 2000). As supported by findings in Chapter 3, we used our inducible transgenic system and examined the Notch-NO axis during the initial phase of arteriogenesis using a hindlimb ischemia (HLI) model.  24  CHAPTER 2 – MATERIALS AND METHODS 2.1 Reagents Rabbit anti-GUCY1B3, rabbit anti-Hey2, mouse anti-phospho-Vasp-Ser239 and rabbit antiphospho-VASP-S157 were acquired from Abcam (Cambridge, MA). Rabbit anti-p-eNOS-S1177, rabbit anti-eNOS, rabbit anti-pAkt, rabbit anti-Akt, rabbit anti-cleaved Notch1 antibody and LY294002 inhibitor were acquired from Cell Signaling Technologies (Beverly, MA). Rat anti-RBPJ (RBPjk-T6719) was acquired from the Institute of Immunology, Tokyo (Hamaguchi et al., 1992). Rabbit anti-nitrotyrosine, protein A & protein G agarose beads, and diaminorhodamine 4M-AM (DAR4M-AM) were acquired from Chemicon/Millipore (Billerica, MA). Rat anti-mouse CD31/PECAM antibody and type-I collagen were acquired from BD Sciences (Franklin Lakes, NJ). Rabbit anti-GFP, X-gal, 1 Kb and 100 bp DNA ladders, SuperScript II reverse transcriptase, Trizol and Alexa Fluor secondary antibodies were acquired from Invitrogen (Carlsbad, CA). DAPI (4’,6-diamidino-2-phenylindole), mouse monoclonal antibody against the FLAG epitope (M2), rabbit IgG, mouse IgG, goat anti-mouse horse radish peroxidase (HRP), goat anti-rat HRP, rat anti-goat HRP, rat anti-mouse HRP, 1H-[1,2,4]oxadiazolo-[4,3-a]quinoxalin-1-one (ODQ), NωNitro-L-arginine methyl ester hydrochloride (L-NAME), Insulin/Transferrin/Selenium (ITS) supplement, and mouse anti-tubulin were purchased from Sigma-Aldrich (St. Louis, MO). The Griess reagent system and Fugene HD transfection reagent were obtained from Promega (Madison, WI). Neutralizing anti-INHBA antibody, Cyclic-GMP assay kit and human Activin A (INHBA) Quantikine ELISA kit were purchased from R&D systems (Minneapolis, MN). Human IGF2 ELISA kit was obtained from Diagnostic Systems Laboratories (Webster, Texas). All primers were acquired from Integrated DNA Technologies (San Diego, CA). BAY41-2272 compound was purchased from Alexis/Enzo Life Sciences (Farmingdale, NY). N-[N(3,5-difluorophenacetyl)-l-alanyl]-S-phenylglycine t-butyl ester (DAPT) and Akt inhibitor Triciribine were acquired from Calbiochem/EMD Chemicals (Gibbstown, NJ). Rabbit anti-smooth muscle actin (Acta2) was acquired from NeoMarkers/Termo Scientific (Fremont, CA). Bradford reagents were acquired from Bio-Rad (Mississauga, Ontario). Complete protease inhibitor cocktail tablets were acquired from Roche (Mississauga, Ontario). TransIT-293 transfection reagent was acquired from Mirus (Madison, WI). 25  Western Lightning® Plus-ECL was acquired from Perkin Elmer (Waltham, MA). Bioflex – Scientific Imaging Films were acquired from InterScience (Markham, ON). Pre-stained protein ladders were acquired from Fermantas/Thermo Scientific (Burlington, ON). DIG labels, Boehringer blocking reagent, anti-DIG antibody, BM Purple substrate, T7 and SP6 RNA polymerases were acquired from Roche Applied Science (Indianapolis, IN). pDrive cloning kit, mini- and midi-prep extraction kits and gel extraction kits were acquired from Qiagen (Toronto, ON). 2.2 Cell culture, gene transfer and RNA interference Human microvascular endothelial cells (HMEC) were provided by Centers for Disease Control and Prevention (Atlanta, GA) (Ades, Candal et al. 1992) and cultured in MCDB 131 medium (SigmaAldrich, St Louis, MO) supplemented with 10% heat-inactivated calf serum (HyClone, Logan, Utah), 10 ng/mL of epidermal growth factor (Sigma-Aldrich, St. Louis, MO) and 100 U each of penicillin and streptomycin (Sigma-Aldrich, St. Louis, MO). Human umbilical vein endothelial cells (HUVEC) were isolated as previously described (Karsan, Yee et al. 1997) and cultured in MCDB 131 medium (SigmaAldrich, St Louis, MO) supplemented with 10% heat-inactivated calf serum and 10% heat-inactivated fetal calf serum (HyClone, Logan, Utah), 20 ng/mL of epidermal growth factor (Sigma-Aldrich, St. Louis, MO) and 100 U each of penicillin and streptomycin. The retroviral producing PhoenixTM-Ampho cell line and lentiviral producing HEK293T cell line were obtained from Dr. Gary Nolan (Stanford University, Palo Alto, CA) and cultured in DMEM medium (Sigma-Aldrich, St Louis, MO) supplemented with 10% heatinactivated calf serum and 100 U each of penicillin and streptomycin. HMEC were transduced using the retroviral vectors MIY, MIY-Jag1, MIY-Dll4, MIY-NICD, pLNCX, pLNCX-FLAG-RBPJ, MSCVNeo, and MSCVNeo-NICD with virus produced using PhoenixTM-Ampho. HMEC were infected with lentivirus produced from HEK293T transiently transfected with pLentiloxshRBP-A, pLentilox-shRBP-B, pLentilox-shGucy1B3 #1, pLentilox-shGucy1b3 #2, pLentilox-Inhba and pLentilox–shRandom. Briefly, constructs were transiently transfected into either the PhoenixTM-Ampho or HEK293T using TransIT®-LT1 Transfection Reagent (Mirus Bio, Madison, WI) according to manufacturer’s instructions. Retroviral and lentiviral supernatants were collected and filtered through a 26  0.45 μm filter, supplemented with 8 μg/mL Polybrene (Sigma-Aldrich, St. Louis, MO) and fresh medium before applying to HMEC. The virus producing cells were replenished with fresh medium and this procedure was repeated once over the next 24 hours. The pLNCX and MSCVNeo transduced cells were then selected for Neomycin resistance using 300 mg/mL G418 (Invitrogen, Carlsbad, CA) for 48 hours. The MIY and pLentilox transduced cells were sorted by fluorescent activated cell sorting for yellow fluorescent protein (YFP) or green fluorescent protein (GFP) using a FACS-440 flow-sorter (BD, Franklin Lakes, NJ). The pLNC-dnAkt construct was a generous gift from Dr. Issei, Komuro (Chiba University, Japan). The MSCV-dnMAML construct was a generous gift from Dr. Warren Pear (University of Pennsylvania, Philadelphia, PA). To generate conditioned medium, HMEC transduced with MSCVNeo or MSCVNeo-NICD were serum starved for 16 hours and conditioned medium was passed through a 0.45 μm filter before applying on receiving HMEC. All cells were starved in serum-free MCDB medium for 24 h prior to collection of conditioned medium. Conditioned medium was filtered with a 0.45 μm filter and applied onto parental HMEC. 2.3 Immunoblotting Cell lysates were collected in RIPA buffer (50 mM HEPES pH7.4, 150 mM NaCl, 10% glycerol, 1.5 mM MgCl2, 1 mM EGTA, 1 mM sodium vanadate, 10 mM sodium pyrophosphate, 10 mM NaF, 1% Triton X-100, 1% sodium deoxycholate, 0.1% SDS, 1 mM PMSF and 1 X protease inhibitor cocktail) as previously described (Fu, Chang et al. 2009). Briefly, Cells are washed once with phosphate-buffered solution (PBS) and lysed with RIPA buffer on ice. Cell lysates were collected into an eppendorf tube and sonnicated using a microtip sonicator. Samples were then centrifugated at 13,000 rpm for 10 minutes and the supernatant cell lysates were transferred into a new eppendorf tube and protein concentrations were determined using Bio-Rad DC Protein Assay System (Bio-Rad Laboratories, Hercules, CA). Samples (5080 μg) are denatured in 1X loading dye at 100°C for 5 minutes and loaded onto a 7.5% or 10% SDSPAGE gel. Electrophoresis was accomplished by applying 100-120 V current through the SDS-PAGE gel. The protein samples were then transferred onto nitrocellulose membrane (Bio-Rad Laboratories, Hercules, CA), blocked in 5% skim milk/PBS/0.1% Tween for 1 hour at room temperature, incubated with 27  primary antibody in 5% skim milk/PBS/0.1% Tween at 4°C overnight, washed three times with PBS/Tween, incubated with secondary antibody (HRP) in 5% skim milk/PBS/0.1% Tween at room temperature for 1 hour, washed three times with PBS/Tween, and developed by enhanced chemiluminescence (PerkinElmer Life Sciences, Boston, MA). Membranes were probed using the following antibodies: 1:1000 rabbit anti-N1IC, 1:3000 rabbit anti-eNOS, 1:3000 rabbit anti-p-eNOS-S1177, 1:3000 rabbit anti-GUCY1B3, 1:3000 rat anti-RBPJ, 1:3000 rabbit anti-pAkt, 1:3000 rabbit anti-Akt, and appropriate secondary antibody (1:10000 HRP-conjugated IgG, Sigma-Aldrich, St. Louis, MO). 2.4 In situ hybridization In situ probes were designed to be 300-500 bp cDNAs that target the 3’UTR, and were PCR amplified and cloned into pDrive vector using pDrive cloning kit (Qiagen). Clones were isolated and plasmids were prepared using mini-prep kits (Qiagen, Toronto, ON) and sent for sequencing (McGill University and Genome Quebec Innovation Centre, McGill University, Montreal, QC). Confirmed vectors were then linearized using restriction enzyme digest and in situ probes were made using DIG labels and T7 or SP6 RNA polymerase (Roche Applied Science, Indianapolis, IN). The DIG-labeled probes were isolated in RNA-free centri-spin columns and stored in -80°C for use. E10.5 hearts were isolated and fixed in 4% PFA at 4°C overnight. The embryonic hearts were then washed twice in PBT (PBS, 0.1% Tween-20), incubated in 6% H2O2 in PBT at room temperature for 1 hour, and washed twice with PBT. The embryonic hearts were then treated with 10 μg/mL proteinase K in PBT for 25 minutes, rinsed once with 2 mg/mL glycine in PBT followed once with PBT. The embryonic hearts were then post-fixed for 20 minutes in 4% PFA/0.2% Glutaraldehyde in PBT, washed twice with PBT, and rinsed once and incubated in hybridization mix (50% formamide, 1.3X SSC, 5 mM EDTA, 50 μg/mL Yeast RNA, 0.2% Tween-20, 0.5% CHAPS, 100 μg/mL Heparin) for 1 hour at 65°C. To the mixture, 1 μL of DIG-labeled RNA probe (1 μg/mL) was added and the samples were immediately placed on 65°C tube stand and left rocking overnight. Samples were then rinsed twice with pre-warmed (65°C) hybridization mix, washed twice with hybridization mix for 30 minutes each, washed 10 minutes in pre-warmed hybridization mix/MABT (100 mM Maleic acid, 150 mM NaCl pH7.5, 0.1% Tween-20) in 1:1 ratio, washed twice with MABT, then 28  incubated: 15 minutes in MABT, 1 hour in MABT/2% Boehringer blocking reagent (BBR), 1 hour in MABT/2% BBR/20% heat-inactivated serum, and overnight at 4°C in MABT/2% BBR/20% heatinactivated serum with 1:2000 dilution of anti-DIG antibody. For immunodetection, hearts were washed three times with MABT and incubated in Boehringer Mannheim (BM) purple substrate (Roche Applied Science, Indianapolis, IN) at 37°C until color developed. The reaction was terminated with several PBS washes and samples were embedded in OCT (Sakura, Japan) for cryosectioning. 2.5 RNA collection and RT-PCR Cultured cells or C57BL/6J embryonic hearts were isolated to study gene expression. Total RNA was isolated by Trizol (Invitrogen) isolation according to manufacturer’s instructions. Generation of first strand cDNA was previously described (Noseda, McLean et al. 2004). Briefly, 5 μg RNA samples were DNase treated before synthesis of cDNA using SuperScript II reverse transcriptase reagent (Invitrogen, Carlsbad, CA). RT-qPCR was carried out using the SYBR green (Applied Biosystems, Foster City, CA) method on an Applied Biosystems 7900HT with primers listed in Appendix A. All primers targeting transcripts spanned exon-exon junctions designed using the Roche Probe Library website (Roche Applied Science, Indianapolis, IN). 2.6 Brightfield imaging and immunofluorescent imaging All tissue samples were fixed in 4% paraformaldehyde (PFA) overnight, dehydrated in 30% sucrose/Phosphate Buffered Solution (PBS), and embedded in OCT. Ten μm cryosections were collected and stored at –80°C until staining. For AVC explants, samples were fixed in 24 wells using 4% PFA for 5 minutes. For immunofluorescence staining that requires antigen retrieval (NICD staining), sections were immersed into pre-heated sodium citrate buffer (10 mM Sodium Citrate, 0.05% Tween 20, pH6.0) and heated for 30 minutes and left to cool before commencing immunofluorescence staining. Slides were then washed with PBS and incubated overnight with primary antibody (dilution range from 1:100-400) in staining buffer (4% calf serum/0.1%Triton X-100/PBS). Slides were washed with PBS and incubated at room temperature in the dark with secondary antibodies (1:300) in staining buffer for one hour. Slides were then washed with PBS and nuclei were stained with DAPI (1 μg/mL) and mounted with ProLong 29  Gold anti-fade (Invitrogen). All micrographs were acquired using an Axiovert S100 Zeiss microscope. Images were captured using Northern Eclipse software and quantifications performed using ImageJ (NIH). 2.7 Chromatin immunoprecipitation HMEC were transduced with pLNCX or pLNCX-FLAG-RBPJ, and the ChIP assay was performed as previously described (Noseda, Fu et al. 2006). Briefly, HMEC DNA was cross linked in 1% formaldehyde in serum free medium for 5 minutes at room temperature. Cross linking was terminated by adding glycine (125 mM final) for 5 minutes. Cells were washed twice with PBS, lysed in 1 mL of Lysis Buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100 and 1X protease inhibitor cocktail), scraped and transferred into eppendorf tubes and sonnicated with a cup-horn sonicator (10 minutes, 30 seconds on, 30 seconds off). Samples were centrifuged at 15,800 rcf for 10 minutes and cross-linked DNA was transferred into a new eppendorf tube. DNA concentration was determined by GeneQuant pro spectrophotometer (Fisher Scientific, Ottawa, ON). Anti-Flag M2 affinity agarose or protein A/G beads (50 μL) were pre-washed with Lysis Buffer and 50-150 μg sample DNA was added and incubated overnight at 4°C with shaking. Agarose beads were gently centrifuged (2 minutes at 850 rcf) and supernatant that contains unbound, non-specific DNA were removed. Samples were washed once with Low Salt immune Complex Wash Buffer (20 mM Tris-HCl pH 8, 150 mM NaCl, 2 mM EDTA and 1% Triton X-100), once with High Salt immune Complex Wash Buffer (20 mM Tris-HCl pH 8, 500 mM NaCl, 2 mM EDTA and 1% Triton X-100), once with LiCl immune Complex Wash Buffer (10 mM Tris-HCl pH 8, 0.25 M LiCl, 1 mM EDTA and 1% Triton X-100), twice with TE Buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) and eluted with 400 μL Elution Buffer (1 % SDS, 0.1 M NaHCO3). In parallel, 1% of input DNA was resuspended in 400 μL Elution Buffer. All samples were then treated with 16 μL of 5 M NaCl and heated at 65°C overnight to de-cross link DNA. The samples were then treated with RNaseA and proteinase K (8 μL of 0.5 mM EDTA, 16 μL of 1 M Tris-HCl pH 6.5, 2 μL of 10 mg/ml protease K, 2 μL of RNaseA, and 2 μL of 10 mg/ml glycogen) at 37°C for 2 hours.  ChIP DNA was recovered using standard  phenol/chloroform extraction, ethanol precipitated, and resuspended in 50 μL of H2O. ChIP pull-down was 30  evaluated using RT-qPCR with primers designed to amplify RBPJ binding sites in human GUCY1A3, GUCY1B3, INHBA and HEY1 promoters using primers listed in Table S1. Binding enrichment to the relevant promoter was calculated using the percentage of ChIP DNA (relative to input chromatin) normalized to the vector control. 2.8 Mouse models Male and female C57BL/6J mice weighing between 25 to 35 g were provided by Jackson Laboratories (Bar Harbor, ME) and used for RT-qPCR and AVC explants. TetOS-dominant negative MAML1 fused with GFP (dnMAML-GFP) transgenic mice were generated in-house by Linda Chang and maintained on a CD1 background (Fu, Chang et al. 2009). The VE-cadherin-tTA (VEtTA) strain was a gift from L. Benjamin, Harvard Medical School (Boston, MA) (Meadows, Iyer et al. 2009) and maintained on the FVB/NJ background. Endothelial Notch inhibition was accomplished by crossing VEtTA mice with TetOS-dnMAML-GFP mice, and removing tetracycline from the drinking water. Suppression of dnMAML expression was achieved by administering either tetracycline (50 μg/ml) or doxycycline (50 μg/ml) in the drinking water of the pregnant females. All animal protocols were approved by the Animal Care Committee of the University of British Columbia (Vancouver, British Columbia). 2.9 Atrioventricular canal (AVC) explant assay C57BL/6J mice were used for all AVC explant experiments. AVC explants assays were performed as previously described (Niessen, Fu et al. 2008). Briefly, AVCs were manually dissected in cold sterile PBS using 26G needles. 24-well plates were pre-layered with Type I collagen and soaked with AVC medium (1% FBS/ 0.5% ITS/DMEM) prior to performing explants. Explants were allowed to attach overnight and cultured for 48 hours with various inhibitors or activators in AVC medium: γ-secretase inhibitor (DAPT, 10 μM), sGC inhibitor (ODQ, 10 μM), NOS inhibitor (L-NAME, 50 μM), NO-independent sGC activator (BAY41-2272, 10 μM), or anti-Activin A neutralizing antibody (0.5 mg/mL). Numbers of AVC explants are indicated in corresponding figures. Explants were fixed with 4% PFA for 5 minutes then stained with anti-Acta2/SMA antibody and DAPI. All immunofluorescence micrographs were acquired using an Axiopan2 Zeiss microscope. 31  Figure 2.1 – Quantification of EndMT in AVC explants (A) Representative micrographs of AVC explants viewed by brightfield microscopy, Acta2 immunofluorescence (green) or DAPI fluorescence (blue) are shown. Edges of AVC explants are outlined in red. (B) Raw and normalized (to AVC area) DAPI readings are shown on the left, with a graphical representation of the normalized number of DAPI-positive pixels relative to distance from the AVC margin shown on the right. (C) Comparison between the area covered by DAPI fluorescence and Acta2 immunofluorescence (to confirm EndMT) shows very high correlation.  32  2.10 Analysis of AVC explants and cardiac cushions The number and distance of migrating cells were analyzed as previously described (Niessen, Fu et al. 2008; Chang, Fu et al. 2011) and is depicted in Figure 2.1. Briefly, all DAPI and SMA images were first inverted using ImageJ. Using an in-house algorithm (Niessen, Fu et al. 2008; Fu, Chang et al. 2011), all the pixels in acquired images were converted from a 0 to 255 gray scale to a 1 to 254 gray scale. The actual explants were manually colored in black (value = 255). The amount of cell invasion and migration was quantified by measuring the number of positively stained pixels to the nearest pixel of the explant. The results are expressed as the number of positively stained pixels over distance (pixels) away from the explant. For normalization, the result is divided by the size of the AVC explant, and the distance is converted from pixels into microns. For cellularization of AVC cushions, the nuclear count function in ImageJ was used to quantify number of nuclei in the cushion. Nuclei counts were normalized to cushion surface area and expressed as nuclei number over surface area (μm2). 2.11 Hindlimb ischemia Endothelial Notch inhibition was achieved by crossing VEtTA to TetOS-dnMAML-GFP. All of the animals were housed with doxycycline (50 μg/ml) supplemented water to suppress dnMAML-GFP expression until 6 weeks of age. All animals were subjected to 2 weeks of Notch inhibition and were used for femoral ligation experiments at 8 weeks of age. Littermate controls were used in parallel. Briefly, mice were anesthetized under isoflurane and from the inguinal ligament the artery was ligated distal to the third arterial branch and proximal to the fourth, the latter being just proximal to a major bifurcation evident to the naked eye (Limbourg, Korff et al. 2009). Mice received meloxicam analgesic for 2 days following surgery. Blood flow in the ischemic and non-ischemic legs was monitored with a laser Doppler perfusion scanner (PeriScan PIM II, Perimed; Las Vegas, NV) at 37°C prior to surgery and on postoperative days 0, 1, and 2. The level of reperfusion was calculated by taking the ratio of perfusion of the ischemic leg over the non-ischemic leg. At treatment endpoints, mice were perfused with 2% Heparin in PBS, in situ fixed with 2% PFA, and both the ischemic and non-ischemic thighs were excised. The 33  tissues were then fixed with 4% PFA overnight at 4°C, washed once in PBS, decalcified with EDTA solution (14% EDTA) overnight at 4°C, and dehydrated in 30% sucrose PBS overnight at 4°C. The semimembranous muscle with the collateral arteries were excised from the femoral artery and embedded in OCT. Ten μm transverse cryosections of ischemic femoral artery, non-ischemic femoral artery, ischemic collateral artery and non-ischemic collateral artery were collected for each animal through the length of the vessel. 2.12 Aortic ring myograph Contractility studies of VEtTAxTetOS-dnMAML-GFP and littermate control aortas were performed as previously described (Liang, Tan et al. 2008). Briefly, the thoracic dorsal aortas were trimmed of surrounding tissues and cut into 2 mm rings. At least four aortic rings from each animal were used. Aortic rings were mounted on tungsten wires (diameter 40 μm) and placed in individual 5 mL wire myograph chambers (Multi Myograph Model 610 M, Danish Myo-tech, Aarhus, Denmark). The chambers were filled with physiological salt solutions, maintained at 37 °C, and aerated with 95% oxygen and 5% carbon dioxide. The baseline tension was gradually adjusted to 5 milli-Newton over 60 min. Bathing solution was changed at 15 min intervals. Once the aortic rings reache equilibration, they were challenged twice (15 min each) with 80 mM potassium containing physiological salt solutions to ensure sample viability (Red treatments in Figure 2.2). Basal NO-dependent vasodilatory response was measured by treating phenylepherine (blue treatments in Figure 2.2) contracted aortic rings with acetylcholine (light orange treatments in Figure 2.2). This step was repeated twice. NO-independent vasodilatory response was measured by treating phenylepherine contracted aortic rings with a NO donor SNP (green treatment in Figure 2.2). Concentration-response curves were generated for all treatments described above. 2.13 β-Galactosidase detection For whole mount embryonic heart staining, E10.5-E12.5 whole hearts were manually dissected using a 26G needle and fixed in 2% PFA for 5 minutes. The hearts were then washed twice in β-gal wash solution (2 mM MgCl2, 0.01% sodium deoxycholate, 0.02% NP-40 in PBS pH7.4) and incubated for 30 minutes. The hearts were then incubated with β-gal staining solution (1 mg/ml X-gal, 5 mM potassium 34  ferrocyanide, 5 mM potassium ferricyanide in β-gal wash solution) at 37°C for a minimum of 4 hours before being frozen in OCT for cryosectioning. 2.14 Data analysis All data are shown as the mean ± SEM of multiple experiments unless otherwise stated. All statistical analyses were performed using a two-tailed Student t-test unless otherwise stated and were considered to be statistically significant at P < 0.05.  35  Figure 2.2 – Schematic diagram of aortic ring myograph setup.  36  CHAPTER 3 – NOTCH SIGNALING ACTIVATES NITRIC OXIDE PATHWAY IN A CELL NON-AUTONOMOUS FASHION TO DRIVE ENDMT IN DEVELOPING AVC 3.1 Introduction During cardiac development, the linear heart tube loops rightward at embryonic day (E) 8.5 to initiate the establishment of a four–chambered structure (E9.0-9.5) consisting of paired left and right atria, and ventricles. In order to separate the chambers, swellings composed of extracellular matrix in the atrioventricular canal (AVC) and the outflow tract, referred to as cardiac cushions, become populated by mesenchymal cells generated from the overlying endocardium. This process called endothelial-tomesenchymal transformation (EndMT) commences at E9.5 in the AVC resulting in endocardial cells (EC) delaminating from the monolayer and invading the underlying matrix (Camenisch, Molin et al. 2002). Following remodeling, the cushions eventually develop into the cardiac valves and the membranous septum of the adult heart (Eisenberg and Markwald 1995; Niessen and Karsan 2008). Several signaling pathways coordinate to initiate and potentiate EndMT with the Notch pathway being a central player in the process (Timmerman, Grego-Bessa et al. 2004; Niessen and Karsan 2008). Notch proteins are a family of transmembrane receptors (Notch1-4 in mammals) that are activated upon binding by Notch ligands: Jagged (Jag) 1 and -2, and Delta-like (Dll) 1, -3 and -4. Upon activation, Notch receptors undergo proteolytic processing and the intracellular domain of Notch (NICD) is released from its membrane tether. NICD then translocates to the nucleus, binds to the DNA-binding co-repressor RBPJ (recombination signal binding protein for immunoglobulin kappa J region), and recruits the coactivator MAML1 (Mastermind-like 1), to derepress and activate transcription of downstream targets such as the HES and HEY family of transcription factors (Iso, Kedes et al. 2003). Deletion of Notch1 or Rbpj in mice results in malformation of the cardiac cushions (Timmerman, Grego-Bessa et al. 2004). Deletion of the Notch target gene Hey2 or double deletion of Hey1 and Hey2, or Hey1 and HeyL results in various congenital heart defects including cardiac cushion defects (Donovan, Kordylewska et al. 2002; Fischer, Schumacher et al. 2004; Fischer, Steidl et al. 2007). Mutations of Notch1 are associated with non37  syndromic valvular disease such as bicuspid aortic valves, as well as with more severe cardiac anomalies (Garg, Muth et al. 2005). Mutations in Jag1 have been implicated in Alagille syndrome, an entity that comprises a spectrum of defects including pulmonary artery stenosis, Tetralogy of Fallot and additional valvular anomalies (Li, Krantz et al. 1997; Oda, Elkahloun et al. 1997; Eldadah, Hamosh et al. 2001). Nitric oxide (NO) has also been shown to play a role in cardiac function (Fukumura, Kashiwagi et al. 2006; Rastaldo, Pagliaro et al. 2007), blood pressure homeostasis (Sessa 2005), and epithelialmesenchymal transformation (Vyas-Read, Shaul et al. 2007). NO is generated by nitric oxide synthases (NOS), of which there are three isoforms: endothelial (eNOS, or NOS3), inducible (iNOS, or NOS2) and neuronal (nNOS, or NOS1). External signals (shear stress, calcium influx, or growth factors) culminate in phosphorylation events that can either activate (Ser116 and Ser1177) or inhibit (Thr495) eNOS (Mount, Kemp et al. 2007). Similar to NOTCH1 mutations in human (Garg, Muth et al. 2005), eNOS knockout mice also display bicuspid aortic valves and other major cardiac anomalies (Huang, Huang et al. 1995; Feng, Song et al. 2002; Tsutsui, Shimokawa et al. 2006; Aicher, Urbich et al. 2007). Of the three classes of type III guanylyl cyclases (GC), NO signaling is propagated through the soluble GC (sGC) heterodimer (Baker and Kelly 2004; Yamagami and Suzuki 2005), which consists of an alpha subunit (GUCY1A2 or GUCY1A3) and a beta subunit (GUCY1B3). GUCY1A3/1B3 is expressed in most tissues, whereas GUCY1A2/1B3 is predominantly found in the brain (Giuili, Scholl et al. 1992; Mergia, Russwurm et al. 2003). Activation of sGC leads to production of the second messenger 3’, 5’ cyclic guanosine monophosphate (cGMP), which results in smooth muscle cell relaxation by blocking calcium influx and dephosphorylation of myosin light chains (Ignarro, Harbison et al. 1986; Murad 1986). In addition to blood pressure regulation, sGC proteins have also been shown to promote endothelial cell survival and migration (Pyriochou, Beis et al. 2006; Pyriochou, Zhou et al. 2007). In this study we demonstrated that Notch induces activation of the NO-sGC axis through an autocrine loop during EndMT in the AVC. The promoters of both GUCY1A3 and GUCY1B3 are bound by RBPJ and transcription is induced upon Notch activation. Inhibition of Notch specifically in endocardial cells in vivo inhibits EndMT and decreases Gucy1b3 expression. Similarly, sGC inhibition blocks EndMT in AVC 38  explants and decreases cushion cellularization at E9.5 and E10.5 in vivo. Concomitant with upregulation of the GUCY genes, Notch induces Activin A, which in turn activates PI3K and Akt in a paracrine fashion followed by downstream eNOS activation and NO production. Here we show that Notch signaling increases NO production simultaneously with induction of the NO receptor sGC and this pathway contributes to EndMT in the AVC. 3.2 Results 3.2.1 GUCY1A3 and GUCY1B3 are novel Notch targets To identify novel Notch targets in the AVC endocardium during heart development, we compared two microarray datasets with AVC transcriptome libraries. Transcripts induced by at least two-fold in NICD-transduced human umbilical vein endothelial cells (HUVEC), and in Dll4-activated HUVEC (Harrington, Sainson et al. 2008), were compared with mRNA expressed in the AVC at E9.5, E10.5 and E11.5 as determined by Serial Analysis of Gene Expression (SAGE) (Siddiqui, Khattra et al. 2005; Vrljicak, Chang et al. 2010). Of the 24 candidate genes identified (Figure 3.1 A), 11 transcripts were upregulated in human EC transduced with Notch ligands (Jag1 or Dll4) or NICD compared to the empty vector control using reverse transcription-quantitative polymerase chain reaction (RT-qPCR) (Figure 3.1 B). Of the 11 validated candidates, 5 have been reported to be associated with heart development, EndMT or are regulated by Notch signaling (HEY1, JAG1, SDC1, SDC2, and SNAI2 (Fischer, Schumacher et al. 2004; Fischer, Steidl et al. 2007; High, Lu et al. 2008; Niessen, Fu et al. 2008; Arrington and Yost 2009)). Of the remaining 6 candidates, GUCY1A3 and GUCY1B3 were selected for further study because both genes are highly induced by Notch signaling (Figure 3.1 B) and heterodimerize to form the soluble NO receptor, sGC. RT-qPCR showed that Notch activation by ligandinduced activation (Jag1, Dll4) or enforced expression of NICD increased GUCY1A3 mRNA levels 316 ± 43, 18 ± 1.5 and 313 ± 43.8-fold respectively, and GUCY1B3 mRNA levels by 154 ± 3.2, 13.4 ± 1.7 and 282 ± 37.2-fold, respectively (Figure 3.2 A). As a positive control, we confirmed that the known Notch target HEY1 was also induced by Notch activation (Figure 3.2 A). Induction of GUCY1A3 and GUCY1B3  39  mRNA was abolished when Jag1- or Dll4-expressing EC were treated with the γ-secretase inhibitor, DAPT (Figure 3.2 B), thus confirming Notch-dependent induction. To determine whether the ligand-expressing or Notch-activated cells induce sGC, ligandexpressing (Jag1 or Dll4, YFP+) were cocultured with parental EC (YFP-), flow-sorted after 48 hours into YFP+ (ligand-expressing cells) and YFP- (parental cells) populations, and each population analyzed by RT-qPCR. As noted by induction of ACTA2 and HEY1 (Figure 3.2 C), Notch activation occurs mainly in the parental cells in this coculture system. The major induction of GUCY1A3 and GUCY1B3 mRNA levels also occurred in the YFP- parental cells (Figure 3.2 D); consistent with the notion that GUCY1A3 and GUCY1B3 are induced by Notch activation. By training a motif scanner GADEM (Li 2009) with previously published RBPJ binding sites (Appendix B), we analyzed the human and mouse GUCY1A3 and GUCY1B3 promoters (from 1.5 kb upstream of the transcriptional start site [TSS] to the end of intron 1) and identified several putative RBPJ binding sites (Figure 3.2 E). Five putative RBPJ binding sites were identified in the human GUCY1B3 promoter, and three of these sites were conserved in the mouse, while only one conserved putative RBPJ binding site was noted between the human and murine GUCY1A3 promoters. Chromatin immunoprecipitation (ChIP) followed by qPCR with primers flanking the RBPJ elements demonstrated that RBPJ bound all three conserved RBPJ consensus motifs in the human GUCY1B3 promoter (-1065: 6.7 ± 2.1 fold; -37: 2.6 ± 0.4 fold; +247: 3.4 ± 0.9 fold) as well as the single conserved RBPJ consensus motif in the human GUCY1A3 promoter (-944: 4.4 ± 1.7 fold) (Figure 3.2 F) in EC transduced with Flagtagged RBPJ. As a positive control, we showed RBPJ binding to the human HEY1 promoter located 180 bp upstream of the TSS (Figure 3.2 F). Immunoblotting of EC activated by NICD, Jag1 or Dll4 showed that Notch activation induced GUCY1B3 protein (Figure 3.2 G), and the induction was abolished when RBPJ was knocked down (Figure 3.2 H). Thus, GUCY1A3 and GUCY1B3 are direct Notch targets activated by an RBPJ-dependent mechanism.  40  Figure 3.1 - Evaluation of Notch candidate target gene expression. (A) Identification of novel Notch targets in the developing AVC. Upregulated genes with minimum of two-fold induction from both the Dll4 microarray and an NICD microarray performed in HUVEC were overlaid with mouse embryonic day E9.5-E11.5 AVC SAGE libraries. (B) RT-qPCR analysis of 24 putative Notch effectors in the AVC using EC transduced with Jag1, Dll4, NICD or empty vector control. Values in the chart represent the fold-change in expression in Notch-activated HUVEC relative to empty vector controls. Genes that are induced by at least one of the Notch ligands as well as by NICD (Notch1), relative to empty vector are highlighted in blue. All results are shown as mean fold-induction ratios ± SEM (n = 3, * P < 0.05).  41  Figure 3.2 - Notch signaling induced GUCY1A3 and GUCY1B3 expression. Expression of novel Notch targets GUCY1A3 and GUCY1B3 identified in our microarray screen were evaluated by RT-qPCR in EC transduced with Jag1, Dll4, NICD or empty vector (A) but blocked in presence of the g-secretase inhibitor, DAPT (10 mM), compared to vehicle DMSO in (B). Data were normalized to the empty vector control and fold-enrichment is shown as mean ± SEM (n = 3, * P < 0.05, ** P < 0.001). (C-D) Notch ligand-receptor signaling induced expression of mesenchymal marker (SMA), Notch target (HEY1), GUCY1A3 and GUCY1B3 were evaluated in sorted Jag1, Dll4 or vector (YFP+) and co-cultured parental EC (YFP-). Data were normalized to the empty vector control and fold-enrichment is shown as mean ± SEM (n = 3, * P < 0.05). (E) Schematic representation of conserved RBPJ binding sites between human and mouse GUCY1A3 and GUCY1B3 promoters. (F) RBPJ occupancy on conserved RBPJ binding sites of human GUCY1B3 and GUCY1A3 promoters were examined by anti-Flag ChIP with IgG ChIP used as a negative control in EC transduced with either Flag-RBPJ or empty vector. qPCR was conducted using primers flanking the conserved RBPJ binding sites. Fold-enrichment was calculated by normalizing to input DNA and then referencing to empty vector anti-Flag ChIP. Values are shown as mean ± SEM (n = 5, * P < 0.05). (G) Cell lysates from EC transduced with Jagged1 (J), Dll4 (D), NICD (N) or empty vector (V) were immunoblotted for GUCY1B3 protein. (H) Notch-activated cells were lentivirally 42 transduced with shRBPJ-A, shRBPJ-B or shRandom (control) and immunoblotted for GUCY1B3 and RBPJ protein. Tubulin expression was used as a loading control.  To demonstrate that Gucy1b3 is a target of Notch in vivo, an EC-specific Tet-inducible binary transgenic mouse model was used where expression of a pan-Notch inhibitor, dominant negative (dn)MAML fused to GFP behind the TetOS promoter, is driven by the VE-cadherin promoter (VEtTA) in the absence of tetracycline (Fu, Chang et al. 2009). To assay Gucy1b3 protein expression during EndMT, EC Notch inhibition was induced at E9.5 by withdrawal of tetracycline from the drinking water and embryos were collected at E10.5 (VEtTAxTetOS-dnMAML-GFP). In VEtTAxTetOS-dnMAML-GFP E10.5 double transgenic hearts, Gucy1b3 protein expression was reduced in both the endocardium and the mesenchyme of the AVC compared to single transgenic littermate controls (Figure 3.3 A-H). Notch inhibition resulted in an approximately two-fold reduction in the intensity of Gucy1b3 staining in the AVC when normalized to DAPI staining intensity (1.9 ± 0.2 vs. 0.8 ± 0.2, P = 0.02) or to cellularity (number of nuclei, 2.5 ± 0.2 vs. 1.4 ± 0.2, P = 0.003). Reduced Gucy1b3 staining was also noted specifically in the endocardium as determined relative to CD31 staining intensity (1.3 ± 0.1 vs. 0.6 ± 0.1, P = 0.005) (Figure 3.3 I). These results suggest that Gucy1b3 expression is regulated by Notch in the developing AVC and that inhibition of EC-Notch signaling blocks Gucy1b3 expression in vivo. 3.2.2 Notch-induced sGC drives EndMT To determine the role of sGC in EndMT, we used an ex vivo AVC explant model in which E9.5 wild-type AVC are cultured on collagen gels and cellular migration and invasion of the matrix is quantified 48 h later (Camenisch, Molin et al. 2002; Niessen, Fu et al. 2008). To ensure that the migrating/invading cells comprised transformed mesenchyme, all explants were also stained with the mesenchymal marker, ACTA2, and the areas of ACTA2 and DAPI staining were correlated (Figure 2.1). Inhibition of sGC activity with [1H-[1, 2, 4] oxadiazolo-[4, 3-a] quinoxalin-1-one] (ODQ; 10 μM) resulted in significant blockade of EndMT (P < 0.05) similar to Notch inhibition (DAPT 10 μM, P < 0.05), compared to vehicle-treated explants (Figure 3.4 A-B). In contrast, administration of the NO-independent sGC activator BAY41-2272 (10 μM) showed marked induction of EndMT (P < 0.05, Figure 3.4 A-B). Additionally, knockdown of  43  Figure 3.3 - Endothelial Notch inhibition blocks Gucy1b3 expression in AVC. GUCY1B3 protein expression was evaluated in E10.5 littermate control (A-D) and VEtTAxTetOS-dnMAML AVC (EH). (I) Intensity of immunofluorescence staining of GUCY1B3 was relative to either DAPI intensity, number of nuclei, or CD31 intensity (only EC staining quantified) and is shown as the mean ratio ± SEM (* P < 0.005) (n= 14 littermate control and 4 VEtTAxTetOS-dnMAML-GFP embryos; three sections per embryo were used for quantification).  44  Figure 3.4 - Pharmacological inhibition of Notch-Gucy1b3 axis blocks AVC EndMT. (A) Quantitative analysis of EndMT in AVC explants from E9.5 wild-type mice treated with BAY41-2272 (10 mM), DAPT (10 mM), ODQ (10 mM), or vehicle after 48 h in culture. Results represent the distance migrated by cells emanating from the edge of the AVC explant normalized to the area of the AVC explant (*P < 0.001). (B) Representative micrographs of bright field and immunofluorescence (SMA [in green] and DAPI [in blue]) are shown.  45  Gucy1b3 in AVC explants using lentiviral transduction with two distinct shRNAs targeting different sequences of Gucy1b3 significantly reduced EndMT compared to a vector control (P < 0.05) (Figure 3.5 A-B). To determine whether sGC activity is required for cushion cellularization in vivo, ODQ (50 mg/kg) was administered to pregnant C57BL/6J mice intraperitoneally 24 h prior to embryo isolation. Cushion cellularization was significantly inhibited at E9.5 when ODQ was administered at E8.5 (vehicle: 43.1 ± 6.6 nuclei/μm2; ODQ: 21.3 ± 6.2 nuclei/μm2, P = 0.02) and at E10.5 with ODQ administered at E9.5 (vehicle: 170.5 ± 4.4 nuclei/μm2; ODQ: 149.4 ± 3.7 nuclei/μm2, P = 0.002), but not at E11.5 when EndMT is complete (vehicle: 270.1 ± 17.5 nuclei/μm2; ODQ: 289.5 ± 14.9 nuclei/μm2) (Figure 3.6 A-B). Taken together these results demonstrate that sGC activation is required for EndMT during the initiation of cushion cellularization. 3.2.3 Notch activates NO synthesis in a cell non-autonomous fashion. Given that sGC is activated by NO, we examined whether Notch also induces NO synthesis in order to activate the sGC pathway in an autocrine fashion. Enforced expression of NICD or co-culture of parental EC with Notch ligand-expressing cells increased NO production compared to vector-transduced cells as measured by accumulation of the NO-reactive dye DAR4M-AM (Figure 3.7 A). Interestingly, NO accumulation was observed not only in Notch-activated cells but also in ligand-expressing or parental EC, suggesting a potential paracrine effect of Notch activation, and this induction was abolished when treated with the pan-NOS inhibitor L-NAME (Figure 3.7 A). Co-culture of parental EC with Notch Jagged1expressing cells also increased NO production as quantified by the Griess reaction (P = 0.002, Figure 3.7 B). To assay for the degree of sGC activation, we measured cGMP concentration in the cell lysates. Cell lysates from co-culture of parental EC with Jag1- or Dll4-transduced EC showed a significant increase in cGMP ( 0.21 ± 0.06 and 0.26 ± 0.08 cGMP (ρmole/mL) / total protein (μg) respectively) compared to cocultures with the empty vector control (0.07 ± 0.02 cGMP (ρmole/mL) / total protein (μg); P = 0.05) (Figure 3.7 C).  46  Figure 3.5 - Lentiviral inhibition of Gucy1b3 blocks AVC EndMT. (A) AVC explants from E9.5 wild-type mice were lentivirally transduced with mshGucy1b3 (#1 & 2) or vector control. Results represent the distance migrated by cells emanating from the edge of the AVC explant normalized to the area of the AVC explant (* P < 0.001). (B) Representative micrographs of bright field and immunofluorescence (SMA [in green] and DAPI [in blue]) are shown.  47  Figure 3.6 - In vivo inhibition of sGC blocks early EndMT. (A) ODQ was (50 mg/kg) administered intraperitoneally 24 h prior to embryo collection. AVCs are outlined red in representative micrographs. (B) Cellularity of the AVC was determined by quantifying the number of nuclei relative to the area of the AVC. Data are shown as mean ± SEM (n = 10, for each time point, * P = 0.05, ** P = 0.002).  48  Figure 3.7 - Notch activation induces NO production in a cell non-autonomous fashion. (A) Co-culture of Notch-ligand expressing (Jag1 or Dll4) or NICD expressing EC with parental EC induces NO as visualized by DAR4M-AM fluorescence where pan NOS inhibitor L-NAME (50 mM) was used as negative control. (B) Mean nitrite levels as measured by the Griess reaction normalized to the total amount of protein in Jag1, Dll4 or vector-transduced EC co-cultured with parental EC treated with either L-NAME (50 mM) or vehicle (n = 5, * P = 0.002). (C) Mean cGMP levels normalized to the total amount of protein in Jag1, Dll4 or vector-transduced EC cocultured with parental EC (n = 7, * P < 0.05). (D-F) Induction of NO and downstream activation in EC treated with NICD conditioned medium. NO pathway activation was assayed by DAR4M-AM staining (D), Griess reaction (E; n = 4, P < 0.05) and cGMP ELISA (F; n = 4, * P < 0.05).  49  To confirm a paracrine effect of Notch on NO synthesis, EC were treated with conditioned medium collected from NICD- or vector-transduced EC. Intracellular NO staining with DAR4M-AM showed accumulation of NO in NICD-conditioned medium-exposed EC over time (Figure 3.7 D), with a significant increase in NO generation as measured by the Griess reaction, compared to vectorconditioned medium-treated controls following 90 and 120 minutes of exposure (1.7 ± 0.1 vs. 1.4 ± 0.1 μM nitrite/μg protein (P = 0.05) and 1.8 ± 0.1 vs. 1.3 ± 0.1 μM nitrite/μg protein (P = 0.05), respectively) (Figure 3.7 E). Cell lysates treated with NICD-conditioned medium also showed a significant increase in sGC activation as determined by an increase in cGMP compared to vector-conditioned medium exposed EC following 90 and 120 minutes of exposure (3.2 ± 1.0 vs. 1.1 ± 0.6 cGMP (ρmole/mL)/total protein (μg) (P = 0.05), and 3.3 ± 0.7 vs. 1.2 ± 0.5 cGMP (ρmole/mL)/total protein (μg) (P = 0.05), respectively) (Figure 3.7 F). Thus, Notch activation induces NO synthesis in a paracrine fashion to activate sGC. To observe NO production in the developing AVC, DAR4M-AM (25 μmole/kg) was administered intraperitoneally into pregnant mice one hour prior to embryo isolation. In the AVC, DAR4M-AM fluorescence was observed in the endocardium and mesenchyme at E10.25 and E10.5 (Figure 3.8 A). In contrast, NO was weakly detectable with DAR4M-AM at E10.25 and E10.5 when Notch signaling was inhibited, by removal of tetracycline from the drinking water 24 h prior in VEtTAxTetOS-dnMAML-GFP double transgenic embryos compared to littermate controls (Figure 3.8 A). Quantification of DAR4M-AM staining relative to the AVC cushion area showed a 2.7 and 5.6-fold decrease of NO in E10.25 and E10.5 VEtTAxTetOS-dnMAML-GFP AVC compared to littermate controls (E10.25: 176 ± 24 vs. 474 ± 70.9; E10.5: 298 ± 103 vs. 53.6 ± 10.7 positive-pixels normalized to AVC area analyzed (P = 0.002)) (Figure 3.8 B). Quantification of DAR4M-AM staining in the endocardium showed a 1.3 and 12.8-fold decrease of NO in E10.25 and E10.5 VEtTAxTetOS-dnMAML-GFP AVC compared to littermate controls (E10.25: 0.41 ± 0.006 vs. 0.30 ± 0.04; E10.5: 0.66 ± 0.35 vs. 0.056 ± 0.02 positive-pixels normalized to endocardial area (P = 0.05)) (Figure 3.8 C). Taken together, these results demonstrate that Notch induces NO production in the developing AVC in a paracrine manner, which is then capable of stimulating the sGC heterodimer formed by Gucy1a3 and Gucy1b3, in Notch-activated endocardial cells. 50  Figure 3.8 - In vivo Notch inhibition blocks nitric oxide production in the AVC. (A) In vivo NO expression was measured by DAR4M-AM fluorescence (25 mmole/kg injected intraperitoneally one hour prior to collecting embryos) in E10.25 and E10.5 AVC from littermate control and VEtTAxTetOS-dnMAML-GFP embryos following 24 h of Notch inhibition (dnMAML induction). Representative sagittal sections are shown where the AVC border is outlined with white dotted lines. Mean DAR4M-AM staining intensity normalized to total AVC area (B) or total EC area (C) shown as the mean ± SEM (n = 4-7 with minimum 3 embryos per group, * P < 0.05).  51  3.2.4 Notch-induced Activin A activates eNOS and promotes EndMT. To identify the paracrine factor responsible for NO induction in the developing AVC that is regulated by Notch, we used gene ontology classifications to select 10 secreted factors as candidate genes that were present both in the EC NICD microarray and the AVC transcriptome datasets (Figure 3.9 A). Using RT-qPCR of NICD-transduced EC, parental EC co-cultured with Notch ligand-expressing (Jag1, Dll4) EC and vector-transduced EC, we validated Inhibin beta A (INHBA), the gene that encodes the subunits of the homodimer Activin A, and Insulin Growth Factor 2 (IGF2), as two factors that were induced by both NICD and ligand activation of Notch. To confirm that the products of these genes were secreted, we measured protein concentrations of Activin A and IGF2 by ELISA in serum-free NICDconditioned medium. We did not detect IGF2 protein, but Activin A was significantly increased in NICDconditioned medium compared to the empty vector conditioned medium (9.11 ± 0.28 vs. 0.05 ± 0.01 ng/mL (P = 0.001), respectively) (Figure 3.9 B). By RT-qPCR we confirmed that ligand-mediated induction of INHBA was Notch-dependent as either Jag1- or Dll4-induced INHBA expression was inhibited by a γsecretase inhibitor (DAPT) (P = 0.005, Figure 3.9 C-D). One conserved putative RBPJ-binding site located in intron 1 of human INHBA was identified (+1018 bp from TSS) and anti-Flag ChIP-qPCR confirmed RBPJ binding to the human INHBA promoter in EC transduced with Flag-RBPJ relative to the empty vector (2.0 ± 0.1 fold enrichment, P = 0.05) (Figure 3.9 E). To assay Inhba expression in vivo, E9.5, E10.5 and E11.5 AVC and whole hearts were harvested for RT-qPCR. mRNA of Inhba and its receptors (Acvr2a, Acvr2b, Acvr1b, and Acvr1c) was detected in the AVC from E9.5 to E11.5, with a significant increase in Inhba expression at E9.5 and E10.5 in the AVC compared to the whole heart (P = 0.05, Figure 3.9 F-H). To confirm that Inhba is a target of Notch in vivo, we performed in situ hybridization on E10.5 VEtTAxTetOS-dnMAML-GFP and littermate controls, which showed a marked decrease in Inhba expression following Notch blockade (Figure 3.10). Collectively, these data indicate that Notch activation drives expression of INHBA and production of the INHBA homodimer, Activin A.  52  Figure 3.9 - Notch activation induces INHBA expression and Activin A release. (A) Potential eNOS activating factors were evaluated by RT-qPCR in Notch-activated EC. EC were transduced with Dll4, Jag1, NICD, or empty vector. Data were normalized to the empty vector control and are shown as mean ± SEM (n = 3, * P < 0.05). (B) Activin A concentrations in empty vector- or NICD-transduced EC conditioned medium were evaluated by ELISA and are shown as mean ± SEM (n = 9, * P = 0.001). (C) Gene expression of INHBA in Notch ligand-activated EC in the presence or absence of DAPT was evaluated by RT-qPCR and data are shown as mean ± SEM (n = 3, * P < 0.0005). (D) Notch ligand-receptor signaling induced expression of INHBA was evaluated in sorted Jag1, Dll4 or vector (YFP+) and co-cultured parental EC (YFP-). Data were normalized to the empty vector control and fold-enrichment is shown as mean ± SEM (n = 3, * P < 0.05). (E) RBPJ occupancy on a conserved putative RBPJ binding site of human INHBA was examined by anti-Flag ChIP with IgG as negative control in EC transduced with either Flag-RBPJ or empty vector. Mean ± SEM of the foldenrichment is shown, determined as a percentage of input DNA and normalized to the empty vector anti-Flag ChIP (n = 3, * P = 0.05). (F-H) RT-qPCR expression analysis of INHBA, INHBB and their cognate receptors evaluated at three time points during AVC development (E9.5, E10.5, and E11.5). Mean fold-enrichment ± SEM of AVC expression relative to expression in the whole heart is shown (n = 3, * P = 0.05).  53  Figure 3.10 - Notch activation induces Inhba expression in vivo. Notch activation is required for Inhba expression (blue stain) in vivo. E10.5 AVCs were examined by in situ hybridization for expression of Inhba in the presence (VEtTAxTetOS-dnMAML) or absence of Notch inhibition (littermate control).  54  To determine whether Activin A is capable of activating eNOS, parental EC were treated with recombinant Activin A. eNOS activation (as determined by anti-phospho-eNOS-Ser1177 immunoblotting) was detected as early as 15 minutes following Activin A stimulation (Figure 3.11 A). Activin A stimulated eNOS activation through PI3K/Akt, as inhibitors targeting either PI3K (LY294002, 10 μM) or Akt (Triciribine, 10 μM) blocked eNOS phosphorylation (Figure 3.11 B). Activin A significantly induced NO accumulation, as determined by the Griess reaction, and this induction of NO was abolished when EC were transduced with a dominant-negative Akt (dnAkt) (P = 0.05, Figure 3.11 C). To determine whether an eNOS-activating factor is secreted from Notch-activated cells, we treated EC with NICD-conditioned medium and immunoblotted cell lysates for phospho-eNOS-Ser1177. We detected eNOS activation as early as 15 minutes following stimulation compared to the empty vector control (Figure 3.11 D). eNOS activation was associated with Akt activation (p-Akt) (Figure 3.11 D). Pretreatment with either LY294002 (10 μM), Triciribine (10 μM) or a neutralizing anti-Activin A antibody (5 ng/mL) blocked the ability of NICD-conditioned medium to activate eNOS (Figure 3.11 E). Furthermore, NICD-conditioned medium on dnAkt transduced EC failed to activate eNOS (Figure 3.11 F) suggesting an Akt-dependent activation. We thus conclude that NICD-conditioned medium activates eNOS via a PI3K/Akt mechanism through the induction and secretion of Activin A. To confirm the direct role of Activin A induced NO in cardiac cushion EndMT, AVC explants treated with either L-NAME (50 μM, P < 0.05), LY294002 (10 μM, P < 0.05), or Triciribine (10 μM, P < 0.05) showed significant inhibition of EndMT compared to vehicle treatment (Figures 3.12 A and B). Moreover, AVC isolated from eNOS-/- mice exhibited pronounced decrease in EndMT compared to wild type AVC (P < 0.05, Figure 3.12 C). Similarly, neutralizing anti-Activin A antibody (5 ng/mL) also inhibited EndMT in AVC explants compared to the IgG control (P < 0.05, Figures 3.13 A and C). Significant EndMT inhibition was also observed in AVC treated with short-hairpin lentivirus targeting Inhba compared to vector control (P < 0.05, Figures 3.13 B and C). These data support a model where Notch-induced Activin A stimulates NO synthesis via eNOS activation to promote EndMT. 55  Figure 3.11 - Notch-induced Activin A activates eNOS via the PI3K/Akt pathway. (A) EC were treated with either recombinant Activin A (5 ng/mL) or vehicle (0.2% BSA in PBS) for various times and analyzed by immunoblotting cell lysates. (B) Parental EC were treated with recombinant Activin A (5 ng/mL) in the presence of vehicle (DMSO), PI3K inhibitor (LY294002, 10 mM) or Akt inhibitor (Triciribine, 10 mM) for 15 minutes and analyzed by immunoblotting. (C) Mean nitrite levels as measured by the Griess reaction normalized to the total amount of protein in dnAkt or vector-transduced EC treated with Activin A (n = 5, * P = 0.05). (D) Parental EC were treated with conditioned medium collected from either NICD-transduced or empty vector-transduced EC for various times and analyzed by immunoblotting. (E) Parental EC were left untreated (UT), or treated with conditioned medium from either empty vector (V) or NICD (N) transduced EC in the presence of vehicle (DMSO), PI3K inhibitor (LY294002, 10 mM), or Akt inhibitor (Triciribine, 10 mM) for 15 minutes and examined by immunoblotting (left panel). Alternatively, conditioned medium from either empty vector (V) or NICD (N) transduced EC was used to treat parental EC in the presence of neutralizing anti-Activin A antibody or an IgG isotype control antibody (right panel). (E) (F) EC transduced with either dominant-negative (dn) Akt or vector alone were treated with conditioned medium from either empty vector (V) or NICD (N) EC.  56  Figure 3.12 - PI3K/Akt-eNOS axis is important for AVC EndMT. Quantitative analysis of EndMT in AVC explants from E9.5 AVC. (A) Nitric oxide signaling induced EndMT was evaluated by treating AVC with inhibitors L-NAME (50 mM), LY294002 (10 mM), Triciribine (10 mM) or DMSO (vehicle). (B) Representative micrographs of bright field and immunofluorescence (SMA [in green] and DAPI [in blue]) are shown. (C) In parallel, eNOS-/- AVC was evaluated against wild type control. Results represent the distance migrated by cells leaving the edge of the AVC explant normalized to the area of the AVC tissue (* P = 0.05).  57  Figure 3.13 - Activin A is important for AVC EndMT. (A) Activin A effect on EndMT was evaluated by treating AVC with either anti-Activin A neutralizing antibody (0.5 ng/mL) or isotype control IgG. (B) Alternatively, AVCs were lentivirally transduced with mshInhba or vector control. Results represent the distance migrated by cells leaving the edge of the AVC explant normalized to the area of the AVC tissue (*P = 0.05). (C) Representative micrographs of bright field and immunofluorescence (SMA [in green] and DAPI [in blue]) are shown.  58  CHAPTER 4 – NOTCH INDUCTION OF NITRIC OXIDE PARTICIPATES IN THE INITIAL VASODILATORY RESPONSE DURING POSTNATAL ARTERIOGENESIS 4.1 Introduction In the homeostatic adult cardiovasculature system, the closed blood circuitry functions as a logistic organ that extends throughout the body. Functional arteries are essential for restoring blood flow and regeneration in response to hypoxia, ischemia, or wound healing. Arteriogenesis is the expansion of existing vessels secondary to mechanical stress or chemical stimuli and is recognized to be the main repair mechanism upon ischemic injury (Carmeliet 2003). Arterial obstruction can cause distal tissue ischemia and tissue survival requires rapid reperfusion prior to tissue necrosis. Although ischemia can induce capillary sprouting, known as angiogenesis, it is understood that arteriogenesis is required to accommodate the redirected blood flow. Postnatal arteriogenesis is the expansion of existing collateral vessels secondary to mechanical stress or chemical stimuli and is recognized to be the main repair mechanism in ischemic injury (Carmeliet 2000; Carmeliet 2003). Postnatal arteriogenesis has been studied using the hindlimb ischemia (HLI) model. In the HLI model, the femoral artery distal to the origin of the deep femoral artery and proximal to the popliteal artery is ligated and the nearby collateral arteries are examined for arteriogenesis (Limbourg, Korff et al. 2009). The actual arteriogenesis recovery process has been divided into four stages (Scholz, Ito et al. 2000). During the initial vasodilatory phase, obstruction of the main artery causes increased blood flow through the collateral vessels, via NO-dependent vasodilation (Scholz, Ito et al. 2000). Notch signaling has been shown to play an important role in postnatal arteriogenesis, but the mechanism of its action is not completely defined (Limbourg, Takeshita et al. 2005; Limbourg, Ploom et al. 2007). As presented in Chapter 3, we recently have shown that Notch activation induces NO synthesis through the release of a paracrine factor that activates phosphatidylinositol-3’-kinase (PI3K), resulting in phosphorylation and activation of endothelial NO synthase (eNOS) (Chang, Fu et al. 2011). 59  In this study, I examined the effect of EC-Notch inhibition on post-ischemic vasodilation using our VEtTAxTetOS-dnMAML-GFP mouse model. Here I demonstrate that postnatal EC-Notch inhibition results in a significant decrease in basal NO in the mouse aorta, but does not affect the mean arterial pressure (MAP). However, following an ischemic insult, EC-Notch inhibition blocks reperfusion of the ischemic tissue due to lack of NO pathway activation. These findings suggest that Notch activation is required for the initial vasodilatory phase in response to ischemia, but under homeostatic conditions, Notchdependent NO generation is compensated by other mechanisms. 4.2 Results 4.2.1 Endothelial Notch activation is not required to maintain blood pressure in the adult We have previously identified a novel link between Notch and the NO pathway (Chang, Fu et al. 2011). Since targeted deletion of Notch1 results in embryonic death (Timmerman, Grego-Bessa et al. 2004; Fu, Chang et al. 2009), to determine the role of Notch in maintenance of blood pressure in the adult, we conditionally inhibited Notch signaling starting at 6 weeks of age using VEtTAxTetOS-dnMAMLGFP mice (Fu, Chang et al. 2009; Chang, Fu et al. 2011; Fu, Chang et al. 2011). We did not detect a difference in mean arterial pressure between VEtTA-TetOS-dnMAML-GFP and littermate controls (Figure 1A) suggesting that the Notch activation is not critical for basal vascular homeostasis. We also did not detect a difference in body weight between VEtTAxTetOS-dnMAML-GFP and littermate controls (Figure 4.1 B) suggesting that EC-Notch activation is not critical for body weight. To examine whether EC-Notch inhibition can block NO synthesis in adults, we isolated adult aortas and measured NO levels using the Griess reaction. There were significantly lower levels of NO in VEtTAxTetOS-dnMAML-GFP aortas compared to littermate controls (3.0 ± 0.6 versus 9.4 ± 2.7 nitrite (μM) normalized to protein (μg) respectively, P = 0.05) (Figure 4.1 C). This difference in NO was independent of body weight between the VEtTAxTetOS-dnMAML-GFP and littermate controls  60  Figure 4.1 – Endothelial Notch inhibition results in decreased NO but no change in mean arterial pressure. (A) Mean arterial pressure and (B) body weight (g) measurements of VEtTAxTetOS-dnMAML-GFP and littermate controls. Results are expressed as mean ± S.E. (n = 6 per group). (C) NO levels in VEtTAxTetOS-dnMAML-GFP or littermate control aortas were measured using the Griess reaction. Results are normalized to either total protein (μg) or total protein (μg) and body weight (g) and expressed as mean ± S.E. (n = 8 each).  61  35  30 )g (  t h gi e W 25  20 48  50  52  54  56  Age (days) WT - F  KO - F  WT - M  KO - M  Figure 4.2 – Longitudinal weight monitoring of EC-Notch inhibited mice. Weight of animals were monitored every two days post EC-Notch inhibition (at 6 weeks of age). WT = littermate controls; KO = VEtTAxTetOS-dnMAML-GFP mice; F = females; M = males. N = 3-5 per group.  62  (0.07 ± 0.02 versus 0.24 ± 0.06 nitrite (μM) normalized to protein (μg) and body weight (g) respectively, P = 0.05) (Figure 4.1 C). In a longitudinal weight monitoring study in mice up to 18 weeks of age, we did not observe any significant difference in body weight between the VEtTAxTetOS-dnMAML-GFP and littermate control animals suggesting the EC-Notch inhibition has minimal effect on body weight and thus systemic effects were not a contributing factor in the reduced levels of NO observed (Figure 4.2). Although EC-Notch inhibition resulted in an overall decrease in NO, we did not observe defects in vascular contraction in response to phenylepherine, EC-NO dependent vasodilation or NO-independent vasodilation (Figure 4.3 A-C). These results suggest that in adult animals, Notch is important in generating of NO, but inhibition of this pathway may be compensated by other mechanisms to maintain blood pressure and vascular homeostasis. 4.2.2 Endothelial Notch activation is required for vasodilation post-ischemia To evaluate the effect of EC-Notch inhibition in response to ischemia-related vasodilation, we subjected VEtTAxTetOS-dnMAML-GFP and littermate control mice to HLI (Limbourg, Korff et al. 2009). Relative blood flow measurements by laser Doppler revealed comparable perfusion in both VEtTAxTetOSdnMAML-GFP and littermate controls before surgery (Figure 4.4 A-B). Serial readings in littermate controls revealed rapid reperfusion in the ischemic leg immediately after and at 1- and 2-days post-HLI (Figure 4.4 B). However, reperfusion was blunted in VEtTAxTetOS-dnMAML-GFP animals post-HLI (Figure 4.4 B). To confirm that EC-expression of dnMAML-GFP blocked downstream Notch activation, we stained for GFP, NICD and the Notch downstream target Hey2 in the ischemic collaterals. We observed co-localization of GFP staining with CD31 expression in the VEtTAxTetOS-dnMAML-GFP animals, confirming the expression of the dnMAML-GFP transgene specifically in the endothelium (Figure 4.4 C). Notch cleavage was present in both VEtTAxTetOS-dnMAML-GFP and littermate controls as evidenced by presence of NICD form (Figure 4.4 C). However, Notch activity – as determined by upregulation of the Notch target Hey2 – was absent VEtTAxTetOS-dnMAML-GFP indicating that Notch signaling was  63  Figure 4.3 – Long-term endothelial Notch inhibition does not affect vasodilatory response. (A) Concentration-dependent contraction to phenylepherine (PE) normalized to 80 mmol/L potassium chloride with and without L-NAME (n = 5). (B) Concentration-dependent relaxation to acetylcholine (Ach) with and without L-NAME (n = 5). (C) Concentration-dependent relaxation to sodium nitroprusside (SNP) with and without L-NAME. KO = VEtTA x TetOS-dnMAML; WT = littermate controls.  64  Figure 4.4 – Impaired postnatal arteriogenesis in VEtTAxTetOS-dnMAML-GFP mice. (A) Impaired blood flow recovery of ischemic hindlimbs of VEtTAxTetOS-dnMAML-GFP mice shown by Doppler blood flow. (B) Quantitative analysis of blood flow recovery expressed as ischemic to normal laser Doppler ratios in littermate control (day 1 n=20, day 2 n=9) and VEtTAxTetOS-dnMAML-GFP (day 1 n=15, day 2 n=7). Mean ratios were plotted ± S.E. *P < 0.05. (C) Increased Notch activation (NICD staining) in both littermate control and VEtTAxTetOS-dnMAML-GFP surgical collateral arteries; however, Notch signaling propagation was blocked in VEtTAxTetOS-dnMAML-GFP animals (Hey2 staining; arrowheads).  65  Figure 4.5 – Inhibition of Notch signaling does not affect vessel remodeling at early times post-ischemia. (A) Representative micrographs of SMA and CD31 staining of ischemic collaterals. (B) Surface area of VSMC vessel coverage as determined by SMA immunostaining was quantified and is plotted as box-and-whisker plots. P values comparing littermate control and VEtTA-TetOS-dnMAML were calculated by the Mann-Whitney t-test (n = 6 mice per group, total of 32-64 sections per group).  66  Figure 4.6 - Inhibition of Notch signaling does not affect expression of NO receptor Gucy1b3. (A) Representative micrographs of Gucy1b3 staining of ischemic collaterals. (B) Staining quantification was achieved by normalizing the number of positively stained pixels to cell count (DAPI) and are plotted as box-and-whisker plots (n = 6 mice per group, total of 32-64 sections per group).  67  blocked (Figure 4.4 C). These findings suggest that Notch inhibition perturbs the initial vasodilatory phase of arteriogenesis that occurs as a consequence of ischemia. 4.2.3 Endothelial Notch activation is required for NO generation and signaling post-ischemia We next examined the collateral arteries to determine whether there was a difference in VSMC coverage of the vessels at 1- and 2-days following HLI (Figure 4.5 A-B). Quantification of the surface area of VSMC showed no significant differences between VEtTAxTetOS-dnMAML-GFP and littermate controls (Figure 4.5 B) suggesting that VSMC remodeling has not yet occurred at this stage as shown by others (Scholz, Ito et al. 2000). Quantification of Gucy1b3 staining revealed no significant difference between VEtTAxTetOS-dnMAML-GFP and littermate control in the non-ischemic collaterals (1d post-HLI: 86.2 ± 9.2 versus 127.1 ± 17.9; 2 days: 20.3 ± 2.8 versus 26.7 ± 3.9 positive pixels/cell count respectively) and in the ischemic collaterals (1d post-HLI: 125.1 ± 12.3 versus 109.7 ± 23.3; 2d post-HLI: 70.5 ± 18.1 versus 138.6 ± 44.5 positive pixels/cell count respectively) suggesting the surgery did not alter the level of NO receptor (Figure 4.6 A-B). Quantification of nitrotyrosine staining to estimate NO production revealed a significant difference between VEtTAxTetOS-dnMAML-GFP and littermate controls in the ischemic collateral vessels at 2d post-HLI (87.0 ± 22.4 versus 144.7 ± 32.0 positive pixels/cell count respectively, P = 0.02) but not 1d post-HLI (1d post-HLI: 127.4 ± 26.1 versus 126.4 ± 12.6 positive pixels/cell count respectively) (Figure 4.7 A-B). However, this difference in NO between VEtTAxTetOS-dnMAML-GFP and littermate controls was not observed in the non-ischemic limbs (1d post-HLI: 80.0 ± 12.8 versus 63.1 ± 14.5; 2d post-HLI: 30.1 ± 1.8 versus 51.2 ± 11.0 positive pixels/cell count respectively) (Figure 4.7 A-B), suggesting that NO production is secondary to ischemia, and that Notch signaling is required for NO induction in the stressed state. To demonstrate downstream activation of the NO pathway, phosphorylation of Vasp was examined (Butt, Abel et al. 1994). Phospho-Vasp-Ser157 staining in the ischemic collaterals was significantly higher in littermate control than VEtTAxTetOS-dnMAML-GFP (1d post-HLI: 28.2 ± 4.9 versus 12.4 ± 2.7, P = 0.03; 2d post-HLI: 39.7 ± 6.5 versus 14.4 ± 2.6, P = 0.001 positive pixels/cell count respectively) (Figure 4.7 C-D) and was localized to the EC (Figure 4.7 D). This  68  difference in phospho-Vasp-Ser157 was not observed in the non-ischemic collaterals of the contralateral limb, consistent with NO signaling being secondary to ischemia-induced Notch signaling (Figure 4.7 C-D).  69  Figure 4.7 – Notch is required for NO signaling following HLI. Immunofluorescence quantification for nitric oxide (A, nitro-tyrosine) and downstream target phospho-Vasp-Ser157 (C) was achieved by normalizing the number of positively stained pixels to cell count (DAPI) and are plotted as boxand-whisker plots (n = 6 mice per group, total of 32-64 sections per group). Representative micrographs of the immunofluorescent staining are shown in (B) and (D).  70  CHAPTER 5 – SUMMARY, PERSPECTIVES AND FUTURE DIRECTIONS 5.1 Summary of Notch and the NO pathway in the developing AVC In Chapter 3, we show that Notch-induced NO pathway activation results in EndMT. First, we demonstrate that Notch activation directly induces expression of the two components of the heterodimeric sGC, GUCY1A3 and GUCY1B3, through RBPJ binding to the respective promoters. Second, we show that Notch-activated EC secrete Activin A, leading to activation of eNOS and release of NO by a PI3K/Akt-dependent mechanism. Last, we show that activation of the NO pathway contributes to early EndMT in the developing AVC. The overall findings are summarized in Figure 5.1. Recent findings support a role for Akt in promoting EndMT in the AVC (Meadows, Iyer et al. 2009; Feng, Di et al. 2010). It has been reported that the Notch ligands Jag1 and Dll4 have been shown to have opposing effects on angiogenesis, and it is proposed that the distinct spatial expression patterns and opposing functional roles of these two Notch ligands regulates angiogenesis (Iso, Hamamori et al. 2003). While JAG1 and DLL4 appear to have similar effects on GUCY1A3 and GUCY1B3 transcript expression, JAG1 appears to be much more potent in inducing NO whereas prolonged co-culture experiments show that DLL4 seem to be more potent at inducing the INHBA transcript. We also noticed that overexpression of DLL4 in HMEC leads to a decrease in the JAG1 transcript suggesting that there may be JAG1-DLL4 cross-regulation may act through an unknown mechanism. It remains to be investigated whether it is the spatial expression or distinct functional roles of these two ligands that regulates EndMT in the AVC and whether  cross-regulation  occurs.  One  way  to  study  this  difference  may  require  detailed  immunofluorescence studies using highly specific antibodies against individual Notch ligands. In this study, we used motif-scanning programs to identify RBPJ binding sites and validated these using ChIP-qPCR on HMEC overexpressing a Flag-tagged version of RBPJ. We did attempt to perform Rbpj-ChIP from AVC tissue directly; however, we had problems finding a ChIP-grade anti-Rbpj antibody that worked in our hands. To truly identify and validate Rbpj binding sites in vivo, ChIP-grade antibodies are needed for future experiments. It would also be interesting to perform multiple cofactor (such as 71  NICD, MAML and HAT) ChIP-Seq libraries to validate computational derived sites. In this study we used multiple methods to inhibit Notch activation; however, each method comes with its own caveat. First, it has been a very attractive theory to use γ-secretase inhibitors to target Notch-dependent tumors. A recent report comparing a panel of γ-secretase inhibitors against breast tumor cell lines suggests that some proposed γ-secretase inhibitors may exhibit efficacy through inhibiting proteasomal activity rather than inhibiting γ-secretase (Han, Ma et al. 2009). Second, studies conducted by Dr. Iva Kulic from our lab have shown that knocking down of RBPJ may lead to loss of repressive complex and result in a Notchindependent upregulation of Notch targets (unpublished data). Third, although our VEtTAxTetOSdnMAML-GFP animals can inhibit EC-Notch activation, the expression of dnMAML-GFP may also block other pathways that require MAML for downstream activation. Here we took advantage of a previously established AVC explant model to evaluate EndMT (Camenisch, Molin et al. 2002; Niessen, Fu et al. 2008). In AVC development research, the standard assay for measuring AVC EndMT is the ex vivo AVC explant assay. Given the complex composition of cardiac jelly, drawing conclusions from AVC explants that were cultured on type I collagen alone should be cautioned. To assay the robustness of measuring EndMT, we compared the Acta2 staining with previously established DAPI staining (Niessen, Fu et al. 2008). Here we showed that DAPI or Acta2 staining results are highly correlated suggesting our EndMT quantification method was truly measuring migratory/invasive Acta2 positive mesenchymal cells rather than just migrating endothelial cells (Figure 2.1). NO/cGMP signaling was first discovered to be the main mechanism for vasodilation in the vascular SMC (Murad 2006; Yetik-Anacak and Catravas 2006). This was accomplished by showing that activation of PKG blocks calcium influx and phosphorylates and activates myosin light-chain kinase which ultimately dephosphorylates myosin light chain resulting in SMC relaxation. In this dissertation, we have described how activation of the sGC in the AVC endothelium can promote cell migration. Functionally, we have shown that pharmacological activation of sGC induces EndMT and that disruption of the Activin API3K-Akt-eNOS-sGC axis inhibits EndMT. It has been suggested that in tumor endothelial cells, activation 72  of PKG downstream of NO signaling can activate cell migration either through the induction of Ras expression resulting in increased MMP13, or through activation of the PKG/PI3K/Akt pathway (Fukumura, Kashiwagi et al. 2006). In a more recent study, it has been shown that activation of NO/cGMP can induce cell de-adhesion by down-regulating integrin affinity (Chigaev, Smagley et al. 2011). Whether these cGMP signals can promote cell migration by activating Ras and down-regulating integrin affinity in the AVC endothelium remains to be tested. The potential pitfall here is that given that most pharmacological inhibitors are non-specific; it would be hard to truly assess the degree of inhibition and side effects in a developing embryo. One potential experiment to evaluate function of sGC in vivo would be to generate a conditional knockout of Gucy1b3 or a transgenic animal overexpressing a dominant-negative form of Gucy1b3 and drive expression under either an EC promoter (to evaluate EndMT initiation) or a mesenchymal promoter (to evaluate mesenchymal migration). In this way, complications from blocking vascular smooth muscle vasodilation can be avoided. Although Gucy1a3 or Gucy1a2 single gene-targeted mice exhibit only a mild phenotype with platelet aggregation defects (Mergia, Friebe et al. 2006), Gucy1b3 targeted mice are hypertensive and demonstrate platelet defects, and impaired peristalsis (Friebe, Mergia et al. 2007; Dangel, Mergia et al. 2010; Groneberg, Konig et al. 2010; Groneberg, Konig et al. 2011). Gucy1a3 and Gucy1b3 proteins have also been shown to play a role in cell migration during angiogenesis in zebrafish (Pyriochou, Beis et al. 2006). Our findings reported here are in keeping with a role for sGC in promoting endothelial migration and invasion in the context of EndMT during AVC development, as seen in angiogenesis. Indeed, the process of angiogenesis requires that EC delaminate and invade the underlying matrix similar to the process of EndMT. eNOS null mice have been shown to exhibit bicuspid aortic valves, lower heart rate, and hypertension (Huang, Huang et al. 1995; Shesely, Maeda et al. 1996; Lee, Zhao et al. 2000; Tsutsui, Shimokawa et al. 2006). Interestingly, NOTCH1 mutations seen in humans are also associated with bicuspid aortic valves (Garg, Muth et al. 2005). Previous studies have shown that various signaling pathways can activate eNOS, in particular, shear stress is a major activator of NO generation (Ziegler, 73  Silacci et al. 1998; Cheng, van Haperen et al. 2005). While it remains controversial whether heart development commences prior to blood flow (DeHaan 1965) or whether blood flow is required to initiate the process (Nonaka, Shiratori et al. 2002; Hove, Koster et al. 2003), it is clear that narrowing at the AVC leads to increased shear stress at the endocardium at that location (Hove, Koster et al. 2003). Shear stress activates eNOS through phosphorylation of residue Ser1177 by Akt (Dimmeler, Fleming et al. 1999; Fulton, Gratton et al. 1999; Michell, Griffiths et al. 1999). Recent studies have suggested that shear stress also activates Notch signaling by inducing Notch receptors and ligands (Wang, Fu et al. 2007; Masumura, Yamamoto et al. 2009). Thus it is tempting to speculate that shear stress induced NO may occur in part through a Notch-dependent mechanism. How then does Notch activate eNOS? Our data suggest that INHBA is a direct target of Notch, and that the INHBA homodimer, Activin A, is increased in NICD-conditioned medium. It should be noted that although we were unable to detect IGF2 protein in our Notch conditioned medium, it is interesting to see both the cytokine (IGF2) and its carrier (IGFBP5) showed up in our expression screen (Figures 3.1 and 3.9). Our studies confirm that Inhba is highly expressed in the AVC at E9.5 and E10.5 compared to the whole heart, and that Activin A appears to be the major paracrine factor in NICD-conditioned medium that activates the PI3K/Akt pathway and eNOS. Members of the TGFβ superfamily are known critical factors in EndMT in the AVC (Nakajima, Yamagishi et al. 2000), and our findings showing that Activin A is important for AVC EndMT is in keeping with a previous study (Moore, Mjaatvedt et al. 1998). However, this is the first demonstration that Notch activates the PI3K/Akt pathway and eNOS in a non-cell autonomous manner through induction of Activin A. A previous study showed that pups from Inhba null and Inhba/Inhbb double null mice develop to term but die immediately after birth with no additional defects suggesting no functional redundancy between the two genes (Matzuk, Kumar et al. 1995). In this study, absence of whiskers and lower incisors and defects in secondary palate suggests craniofacial defects. No heart defects were reported which may suggest compensatory effects of other TGFβ family members or other sources of NO (Matzuk, Kumar et al. 1995). The neural crest is a migratory multipotent population of cells that can give rise to neurons, 74  cartilage and bone, smooth muscle and pigment cells and Notch signaling participates in neural crest formation (Huang and Saint-Jeannet 2004). Interestingly, Slug null animals also exhibit craniofacial defects (Jiang, Lan et al. 1998). Thus with our inducible Notch knockout animal, it would be interesting to block Notch signaling and see whether the Notch-Activin A axis participates in neural crest development which might partially explain the Inhba/Inhbb craniofacial phenotype. However, to fully differentiate the contribution of Slug versus Inhba to craniofacial development, rescue experiments overexpressing Slug or Inhba are needed. VEGF has been reported to act upstream of Notch in the endothelium (Lawson, Vogel et al. 2002; Liu, Shirakawa et al. 2003) as well as during angiogenesis (Takeshita, Satoh et al. 2007), and is one of the Akt activators that results in eNOS phosphorylation (Mount, Kemp et al. 2007). VEGF has been shown to be important for EndMT and AVC development (Armstrong and Bischoff 2004). Based on our results, we propose a model where Notch-activated cells that become EndMT committed begin to lose cell-cell junctions (Niessen, Fu et al. 2008) and increase sGC levels (Figure 5.1). As these cells start to invade and migrate into the cardiac cushion, Notch-induced secretion of Activin A may diffuse into the surrounding endocardium to activate eNOS and induce NO synthesis. NO binding to sGC in the EC committed to mesenchymal transformation then promotes cell migration and invasion.  75  Figure 5.1 - Proposed Model of Notch-sGC-NO signaling in developing AVC.  76  5.2 Summary of Notch and the NO pathway in post-natal arteriogenesis In Chapter 4, we extended our findings to the HLI model to study the role of Notch signaling in arteriogenesis. Deletion of Notch1 or Rbpj in mice results in malformation of the cardiac cushions and leads to embryonic lethality (Timmerman, Grego-Bessa et al. 2004). Heterozygous Notch1 animals and heterozygous Dll1 animals result in defective arteriogenesis following femoral ligation (Limbourg, Takeshita et al. 2005; Limbourg, Ploom et al. 2007). In this study, induction of VE-cadherin promoterdriven dnMAML blocked Notch signaling in the adult endothelium resulted in inhibition of NO generation, and consequently impaired perfusion following ligation of the femoral artery. Interestingly, even though basal NO levels were reduced as determined by quantification of NO in the aorta, there was no measurable difference in mean arterial pressure. Since NO generation was not completely blocked, there are likely other mechanisms to induce NO production. These mechanisms likely involve generation of NO by other NOS isoforms, as well as other mechanisms to activate eNOS. As would be expected from mice in which eNOS has been targeted (Huang, Huang et al. 1995), the remaining NO production when Notch is blocked is sufficient to maintain the basal arterial tone. In contrast, NO generated by eNOS appears to be necessary for vasodilation of collateral arteries after femoral ligation (Mees, Wagner et al. 2007). Our findings suggest that the mechanism of eNOS activation in this model is mediated by Notch, which is activated following arterial occlusion. Taken together, our findings point to a requirement for Notch activation in the post-ischemic vasodilation seen following arterial occlusion, and suggest that this occurs through a mechanism in which eNOS is activated to generate NO. In this current study we did not observe significant tissue remodeling during the vasodilatory phase of arteriogenesis which is in keeping with a previous study (Scholz, Ito et al. 2000). In the ischemic collateral vessels, we observed decreased NO levels by Nitro-tyrosine staining as well as decreased NO downstream activation by phospho-Vasp-Ser157 staining in VEtTAxTetOS-dnMAML-GFP mice. VASP exist as a homo-tetramer (Holt, Critchley et al. 1998; Harbeck, Huttelmaier et al. 2000) and its activity is  77  Figure 5.2 – Proposed model of role of Notch-NO signaling during arteriogenesis  78  tightly regulated at three phosphorylation sites: Ser157, Ser239 and Thr274. All three sites have been shown to be targets of PKG and PKA (Butt, Abel et al. 1994). Studies have shown that PKA preferentially phosphorylates Ser157 (Butt, Abel et al. 1994), whereas, PKG phosphorylates both Ser157 and Ser239 (Butt, Abel et al. 1994; Smolenski, Poller et al. 2000). Phosphorylation of VASP results in loss of filament bundling (Harbeck, Huttelmaier et al. 2000) and anti-capping activities (Barzik, Kotova et al. 2005) and subsequent dissociation from focal adhesion plaques (Smolenski, Poller et al. 2000) and cadherin junctions (Scott, Shewan et al. 2006). Overexpression of VASP increases actin-based stress fibers (Price and Brindle 2000). Thus our observation of decreased NO levels and decreased levels of Vasp activation in VEtTAxTetOS-dnMAMLGFP suggests the loss of reperfusion in VEtTAxTetOS-dnMAML-GFP ischemic legs may be a result of blocked vasodilation with loss of Vasp activation. Our data suggests that endothelial-derived NO is important for arteriogenesis. Previous studies have shown that loss of eNOS does not block arteriogenesis (Ignarro and Napoli 2004; Mees, Wagner et al. 2007; Schaper 2009). Studies showing induction of iNOS expression post-HLI suggest that iNOS also plays a role during arteriogenesis (Ignarro and Napoli 2004; Mees, Wagner et al. 2007; Schaper 2009). However, inhibition of iNOS in eNOS null mice completely blocked arteriogenesis suggesting NO is essential for completion of arteriogenesis (Troidl, Tribulova et al. 2010). Taken together, we propose that Notch signaling contributes to vasodilation at the beginning of arteriogenesis to regulate tissue remodeling at later stages of arteriogenesis (Figure 5.2 A). To test this model, experiments designed to bypass the vasodilatory phase followed with EC-Notch inhibition are needed. So how do the interplay between Notch signaling and NO signaling pathways regulate arteriogenesis? During an ischemic injury, blood is immediately redirected to neighboring vasculature which requires vasodilatory response. As shown in Chapter 4, Notch-eNOS axis can induce this vasodilatory response. Interestingly, increased fluid shear stress (Dimmeler, Fleming et al. 1999) and VEGF has been shown to activate eNOS (Mount, Kemp et al. 2007). As discussed before, VEGF can also activate the Notch pathway thus it would be difficult to differentiate the spatial and temporal 79  expression patterns of all the players during arteriogenesis. To complicate things further, NO synthesized by iNOS seemed to be able to compensate for proper arteriogenesis (Figure 5.2 B) (Ignarro and Napoli 2004; Mees, Wagner et al. 2007; Schaper 2009). Findings from others and the work presented here in Chapter 4 suggests that Notch activation is essential for the vasodilatory response during arteriogenesis, however, the mechanism that allows the transition from a vasodilatory response to tissue remodeling is not well understood (Limbourg, Takeshita et al. 2005; Limbourg, Ploom et al. 2007). In conclusion, the results presented in this dissertation showed that Notch signaling can activate NO pathway through (a) induction of sGC expression and (b) induction of NO synthesis through the secretion of Activin A. Work in Chapter 3 supports a model where disruption of this Notch-NO axis results in a disruption of EndMT in the developing AVC. 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Experiment ChIP  Microarray Screen  Primer Name hGUCY1B3_CSL1_For hGUCY1B3_CSL1_Rev hGUCY1B3_CSL2_For hGUCY1B3_CSL2_Rev hGUCY1B3_CSL3_For hGUCY1B3_CSL3_Rev hGUCY1A3_CSL1_For hGUCY1A3_CSL1_Rev hHEY1CSL1ChIP-For hHEY1CSL1ChIP-Rev hZNF3_neg_ChIP-For hZNF3_neg_ChIP-Rev  Sequence ttcatcatagctctctcggtgctg tggacctgagaagccagatggtat tcctctgtccctgattggttgaag agcatcgagctggagcgagaaca ctccacaccgcagcttcctgag gaacccggagccttcccttc gggaccagattagatggtgtacttgg cccatcatgctgttccattgtg caatcagcggcgcgagaaac gctcacgctttgcctctggttaaa tgggtgacaagagtgagactcc gctggatttcctttgacatgtgtt  Usage ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR ChIP-qPCR  hCCND2_for hCCND2_rev hDMD_for hDMD_rev hGUCY1A3_for hGUCY1A3_rev hGUCY1B3_for hGUCY1B3_rev hIGFBP5_for hIGFBP5_rev hITPR1_for hITPR1_rev hKrt7_for hKrt7_rev hMICAL2_for hMICAL2_rev hMn1_for hMn1_rev hPLVAP_for hPLVAP_rev hRNASET2_for hRNASET2_rev hRND1_for hRND1_rev hSDC1_for hSDC1_rev hSDC2_for hSDC2_rev hSLC1A4_for hSLC1A4_rev hSLIT2_for hSLIT2_rev  ggacatccaaccctacatgc cgcacttctgttcctcacag gcagctgaaacagtgcagac tgcccaccttcattgacac tacaaggtggagaccattggcgat atccagagtgcagtccaattcgca ggaaattgctggccaggttcaagt ttctcctgtggtttctgttcggct ctaccgcgagcaagtcaag gtctcctcggccatctca aaggcatctttggaggaagg ggtgggtagtcatgcatcgt ctgaggctgaagcctggta gggtattccggaggtcgt gaaagttcttgcgcagtgg ggaaggctgaaatgaactgg gacgacgacaagacgttgg gtttgcagggaggtcgtg gcatcaatgccagcttcc tgcattgcttctcactcaaga ggcagaagcctggaactcta tccccaattttagaagcacac gaaaattacacagcctgtttgga cggacattatcgtagtagggaga aggatggaggtccttctgc ccgaggtttcaaaggtgaagt aaacggacagaagtcctagcag aaattgcaaagagaaagccaat tttgcgacagcatttgctac gcacttcatcatagagggaagg catggaggaacttgccactta tccatcagcacaaatacacca  RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR  Start -1142 -1036 -152 -45 309 414 -1021 -913 -180 -4 -5871 -5661  End -1118 -1012 -128 -22 331 434 -995 -891 -160 20 -5849 -5637  452 503 3273 3335 2009 2160 1679 1818 1089 1149 3464 3553 957 1023 3571 3619 4673 4739 382 460 813 853 245 314 933 1020 1039 1081 1190 1228 3106 3151  471 522 3292 3353 2032 2183 1702 1841 1107 1166 3483 3572 975 1040 3589 3638 4691 4756 399 480 832 873 267 336 951 1040 1060 1102 1209 1249 3126 3171  97  Appendix A. Primers and viral sequences used (continued). Secreted Factor Screen  Mouse heart validation  Controls  hPDGFA_qPCR_for hPDGFA_qPCR_rev hIGF2_qPCR_for hIGF2_qPCR_rev hINHBA_qPCR_for hINHBA_qPCR_rev hANGPTL2_for hANGPTL2_rev hDkk3_for hDkk3_rev hFGF13_for hFGF13_rev hGDF11_for hGDF11_rev hLTBP4_for hLTBP4_rev hMTSS1_for hMTSS1_rev hNPFF_for hNPFF_rev hTRIP12_for hTRIP12_rev  acacgagcagtgtcaagtgc attccaccttggccacct acaccctccagttcgtctgt gaaacagcactcctcaacga ctcggagatcatcacgtttg ccttggaaatctcgaagtgc gcccactatgcccactctc ctgcaggcagtctctccat cacatctgtgggagacgaag cccacagtcctcgtcgat ccattgatggcaccaaagat ccacagggatgaggttaaaca accaccgagaccgtcattag agggctgccatctgtctg gctccttccagtgcaggac actgcattcgtccacatcag gcatgttcccgtcatctca gtcatagggcccaggcttag cagcaggaagaccagctctc gtctgggcatcctgtggt ccaaccacaagacgactcaa ggttgtggaactatcacagcag  RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR RT-qPCR  1244 1295 542 623 607 655 1265 1322 638 684 188 227 450 498 2441 2498 1730 1772 242 289 178 225  1263 1312 561 642 626 674 1283 1340 657 701 207 247 469 515 2459 2517 1748 1791 261 306 197 246  mINHBA_qPCR_for mINHBA_qPCR_rev mINHBB_qPCR_for mINHBB_qPCR_rev mACVR2a_qPCR_for mACVR2a_qPCR_rev mACVR2b_qPCR_for mACVR2b_qPCR_rev mACVR1B_qPCR_for mACVR1B_qPCR_rev mACVR1C_qPCR_for mACVR1C_qPCR_rev  atcatcacctttgccgagtc acaggtcactgccttccttg cgagatcatcagctttgcag ggttgccttcattagagacga ccctcctgtacttgttcctactca gcaatggcttcaaccctagt tggctgttcggtttgagc ggccatgtaccgtctggt agagggtggggaccaaac tgcttcatgttgattgtctcg actgcaccttcccacagg ccacatcttctccacaccatc  mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR mouse RT-qPCR  539 595 1597 1659 1167 1221 1142 1198 1138 1181 239 349  558 614 1616 1679 1190 1240 1159 1215 1155 1201 256 369  hs-GAPDH-For hs-GAPDH-Rev mGAPDH_for mGAPDH_rev  cagcaagagcacaagaggaagaga ttgatggtacatgacaaggtgcgg tgcagtggcaaagtggagat tttgccgtgagtggagtcata  RT-qPCR RT-qPCR RT-qPCR RT-qPCR  1131 1266 114 189  1154 1289 133 209  98  APPENDIX B. PUBLISHED RBPJ BINDING SITES. Target Gene Consensus (Barolo, Walker et al. 2000) ASE-site2 tgctgtgggaatat ASE-site3 tatcgtgtgaatat ASE-site4 catcgtgggaagag ASE-site5 ggctgtgagaatat ASE-site6 tgctgtgagaattt ASE-site7 cagcgtgtgaaaaa ASE-site8 atgcgtgggaaatc ASE-site9 tggcgtgggagtca ASE-site2 cggcatgggaaaat ASE-site3 tgtcgtgtgaacat ASE-site4 caccgtgggaagtg ASE-site5 ggctgtgtgaatat ASE-site6 cagtgctgagaatct ASE-site7 cagcgtgtgaaaac ASE-site8 atgcgtgggaattg ASE-site9 gtgtgtgggagcca (Oswald, Liptay et al. 1998) NF-kappa B2 acaagggccgtgggaaatttcctaagcctcgactag Hes-1 gatcgttactgtgggaaagaaagt Bcl-3 ctagtcagcgagtaagtgggaaccgagattccat IL-6 ctagtatcaaatgtgggattttcccatgagtctcat IK-BA ctagtaatcgatcgtgggaaaccccagggaaagaat NF-Kappa B2 1.kappa B ctagtgaaacgtcatgggaattcccccctccgggt Beta-2-Micro ctagtactgaaaatgggaaagtccctttgtaacct IFN-Beta ctagtaagtgaaagtgggaaattcctctgaatagat Angioten.1.kappa B ctagtaccacagttgggatttcccaacctgaccat Invariant chain ctagtgagtgagtggggaatttccagatttgtggt MHC1 ctagtcccagggctggggattccccatctccacat SAA-1 ctagtggaaatgacatggtgggactttccccagggaccaagct TNFA ctagtagtggggtctgtgaattcccgggggtgatt NF-KB2 1.KB-mut ctagtgaaacgtgaggggaattcccccctccgggt NF-KB2 2.KB ctagtggccgagaaggggctttcccggccctgagt A20 ctagtgactttggaaagtcccgtggaaatccccgggcctat Heavy chain ctagtagaatcaaaagggaacttccaaggctgct IG-kappa ctagtctcaacagaggggactttccgagaggccat NF-KB1 ctagtgcgcttcctgggggcttccctaccggctcct (Palmieri, Sasso et al. 1999) 23/24 gttcccacggttcccacgc Hes-1 gtgtgaaacttcccaacga IL-6 KB atgtgggattttcccatg IFNB KB atgtaggaatttcccatg Mad3 ctggggttttcccatg KBF1 agcttgggaattccccac  Organism D. melanogaster D. melanogaster D. melanogaster D. melanogaster D. melanogaster D. melanogaster D. melanogaster D. melanogaster D. virilis D. virilis D. virilis D. virilis D. virilis D. virilis D. virilis D. virilis Mouse Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human Human  99  Appendix B. Published RBPj binding sites (continued). (Lee, Wang et al. 2000) CYP2B1 aggtgtggaatttcccacagtgggtggtgccctccc CYP2B2 aggggtggaatttcccacagtgggtggtgccctccc CYP2B10 agggatgaaatttcccacggtgacttaggccctcca (Noseda, Fu et al. 2006) SMA ctgtctttgctccttgtttgggaagcaagtgggaggagagca SMA ctgtctttgttccttgtttgggaagcaagtgggaggagagca SMA ctgtctttgctccttgtttgggaagcgagtgggaggggatca (Robert-Moreno, Espinosa et al. 2005) Gata2 Seq: -436 to -326 Hes1 Seq: -175 to +13 (Chen, Fischer et al. 1997) ERBB-2 palindrome 1 ctgggagttgccgactcccag ERBB-2 palindrome 2 ctgggagcgcgcttgctcccaa Hes1 gttactgtgggaaagaaagtc (Jarriault, Brou et al. 1995) Hes1 tgaaagttactgtgggaaagaaagtttgggaagtttcacacgag (Ling, Hsieh et al. 1994) EBNA2 - C Promoter gatctggtgtaaacacgccgtgggaaaaaatttatg CD23 gatctcctccttcagccctgtgggaacttgctgctg LMP-1 gatctccggggggcaagctgtgggaatgcggtggcg (Dou, Zeng et al. 1994) pIX agcttgggcgtggcttaagggtgggaaagaatatataa (Niessen, Fu et al. 2008) Snai2 - site 1 (-846) caggaaactggtagatactgagatgg Snai2 - site 2 (-1679) agactgtgtagagtgaaacaagg Snai2 - EMSA site 1 tgtgtgttttgtgggaaatggag Snai2 - EMSA site 2 ggccctttttcccataaaaaaa (Maier and Gessler 2000) Hey1 ctatccatgggaaggggcgcagcgtgggaaaggg (Weng, Millholland et al. 2006) c-myc-promoter tgaggctcctcctcctctttc c-myc-intron 2 cacgggacctgaaaggttct (Fang, Yashiro-Ohtani et al. 2007) Gata3 - exon1a ttccacagggcagtgtcatt Gata3 - exon1b ctcccctgctctgtgtttct Gata3 - EMSA exon 1a tccaactcagtttcacacacctctgat Gata3 - EMSA exon 1a gcggaccggctgggaattacatgtta (Ronchini and Capobianco 2001) cyclin D1 agatgcagtcgctgagattctttggccg cyclin D1 ttgagctgttgctgagattttcgtggtt  Rat Rat Rat Human Chimp Rat mouse mouse Human Human Human Human Epstein-Barr virus Human Epstein-Barr virus Adenovirus Human Human Human Human mouse Human Human Human Human Human Human Human Rat  100  

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