CCCTC-Binding Factor (Ctcf) Is Required for Mouse Cardiovascular Development

CCCTC-binding Factor (Ctcf) is Required for Mouse Cardiovascular Development

Anna Roy

A thesis submitted in conformity with the requirements for the degree of Master of Science

Graduate Department of Molecular Genetics University of Toronto

CCCTC-binding Factor (Ctcf) is Required for Mouse Cardiovascular Development

Anna Roy

Master of Science Department of Molecular Genetics University of Toronto

2016

Abstract

Disruption of transcriptional programs controlling development of the cardiovascular system can cause congenital cardiovascular disease. These programs are tightly regulated by chromatin regulators that alter gene expression via formation of active or repressive chromatin and by mediating long-range chromatin interactions. However, how the process of long-range chromatin interactions, facilitated by chromatin mediators, regulate cardiovascular development is not understood. One potential highly conserved chromatin regulator of cardiovascular development is the CCCTC-binding factor (CTCF). Through conditional deletion of Ctcf in mouse cardiac and endothelial progenitors, I found that

Ctcf is essential for cardiovascular development. Inactivation of Ctcf in cardiac progenitors induces severe cardiac defects by embryonic day 12.5, while inactivation of

Ctcf in endothelial progenitors causes extraembryonic vascular defects by E9.5-10.5.

Future investigations in the mechanistic involvement of Ctcf in cardiovascular development will help determine how the process of long range chromatin interactions guide developmental transcriptional programs and function of CV systems.

ii Acknowledgement I would like to sincerely thank my supervisor, Dr. Paul Delgado-Olguin, for all the help and time he has provided me throughout my graduate studies. I appreciate that he was always available when I needed help, for better or for worse. He was an encouraging and motivating mentor that pushed me to give my best and to always improve. His support and guidance has driven me to become an independent researcher and person. In addition, I thank my committee members, Dr. Ian Scott and Dr. Marc Meneghini, for their amazing feedback and support throughout my graduate journey. I want to thank every member of the Delgado-Olguin Lab and 10th floor SickKids PCGRL lab members: Dr. Lijun Chi, Laura Caporiccio, Abdalla Ahmed, Sandra Vuong, and Dr. Bo (Peter) Li. I specifically want to thank Dr. Lijun Chi and all the help she has given me. No matter the time of day or even if she was in the middle of an experiment, she would not hesitate to help me out. No words can describe how grateful I am to have the assistance of Dr. Chi during my graduate career. I want to acknowledge my appreciation for all the animal husbandry work Laura Caporiccio has done for me. My project could not have run smoothly without her. I thank Dr. Bo (Peter) Li for all the career related advice given to me and for accompanying me to the UofT Karolinska exchange program in Stockholm; we had a great time there. To all my lab members, thank you for the company and memories you have provided me over these past few years. You have all been like a second family to me and it was a pleasure to share my graduate experience with you all. I also treasure the love and support given to me by my parents, brothers and my best friend, Anindita (Annie) Bose, throughout my studies. I am grateful to have a caring friend like Annie, who has been through all my ups and downs in life and provided countless memories filled with laughter. Lastly, I thank the my affiliations, The Hospital for Sick Children; Research Institute and the Department of Molecular Genetics at University of Toronto for giving me this opportunity of a lifetime to explore, learn and grow in the field of Science.

iii Table of Contents

Acknowledgement ...... iii Table of Contents ...... iv List of Abbreviations ...... vi List of Figures ...... vii Chapter 1 – Introduction ...... 1 1.1 Heart Development ...... 1 1.2. Molecular regulation of the developing heart ...... 3 1.3. Vasculature Development: ...... 4 1.4. Molecular regulation of vascular development ...... 7 1.5. Transcriptional regulation by chromatin modifiers ...... 8 1.6. Histone modifiers regulating cardiovascular development ...... 9 1.7. Does the process of long-range chromatin interaction modulate CVS develoment and function? ...... 11 1.8. The function of CCCTC-binding factor, Ctcf ...... 12 1.9. Role of Ctcf in development ...... 13 1.10. Ctcf and cardiovascular development ...... 14 1.11. Thesis Objectives ...... 15 Chapter 2 – Materials and Methods ...... 16 2.1. Mice ...... 16 2.2. Genotyping ...... 16 2.3. Immunostaining ...... 17 2.3.1 Immunofluorescence ...... 17 2.3.2. Whole mount immunostaining of embryos ...... 17 2.4 Histology ...... 17 2.5. Gene Expression Analysis ...... 18 2.6. Microscopy and Imaging ...... 19 Chapter 3 – Results I ...... 20 3.1 Expression of Ctcf in the developing heart...... 20 3.2. Conditional deletion of Ctcf in cardiac lineages...... 20

iv 3.3. Ctcf is essential in cardiomyocytes for embryogenesis and its deletion causes severe cardiac defects ...... 23 3.4. Ctcf-deficient cardiomyocytes did not exhibit decreased proliferation or increased cell death...... 25 3.5. Inactivation of Ctcf induces misregulation of well-characterized cardiac genes ...... 26 3.6. Ctcf deficiency restructures the chromatin interactions of the Irx4 regulatory domain ...... 26 Chapter 4 – Results II ...... 29 4.1 The expression of Ctcf in endothelial cells during vascular development ...... 29 4.2. Conditional inactivation of Ctcf in endothelial progenitors ...... 30 4.3. Ctcf in endothelial progenitors is essential for embryogenesis ...... 32 4.4. Ctcf mutant embryos have narrow cerebral vasculature ...... 33 4.5. Ctcf mutants have yolk sac vascular remodelling defects ...... 34 4.5.1 Ctcf deficiency does not affect cell proliferation or cause increased cell death in yolk vascular endothelium...... 35 4.5.2. Yolk sac vascular defects and hemodynamic force ...... 37 4.6. Ctcf mutant placentas have a reduction in vascular labyrinth area ...... 38 4.6.1 The fetal vasculature of Ctcf mutant placentas do not exhibit cell death or proliferation defects...... 39 4.6.2. Ctcf mutant placentas have misregulation of various vascular regulatory genes .... 39 Chapter 5 – Conclusion ...... 42 5.1 Discussion ...... 42 5.1.1 Ctcf is essential for heart development ...... 42 5.1.2 Ctcf is essential for vascular development ...... 44 5.2. Future Directions ...... 46 5.2.1 Elucidate the gene expression profile of Ctcf-deficient endothelial cells ...... 46 5.2.2 Analyze the genome-wide distribution of Ctcf in developing endothelial cells ... 47 5.2.3 Identify Ctcf mediated long-range chromatin interactions in developing endothelial cells ...... 47 5.2.4 Validate direct targets of Ctcf in vascular development ...... 48 5.3. Conclusion ...... 48 Chapter 6 – References ...... 49

v List of Abbreviations

4C-seq: chromatin conformation capture Hand2: heart and neural crest derivative followed by high throughout sequencing expressed 2 ANG-1: angiopoietin-1 HAT: histone acetyltransferase Baf60c: subunit of BAF complex HDAC: histone deacetylase bFGF: basic fibroblast growth factor Hey1: hairy/enhancer of split related with BMP: bone morphogenetic protein YRPW motif protein 1 BSA: bovine serum albumin Hey2: hairy/enhancer of split related with Cas3: caspase 3 YRPW motif protein 2 CBS: CTCF binding sites HOX: homeobox CD31/pecam1: platelet endothelial cell HOXA: homeobox A cluster genes adhesion molecule1 HRE: hypoxia response element CHD: congenital heart disease IGF2: insulin-like growth factor 2 ChIP-Seq: Chromatin Isl1: ISL LIM homebox immunoprecipitation followed by high KDM: lysine demethylases throughput sequencing Klf: kruppel-like transcription factor COUP-TFII: chicken ovalbumin KMT: lysince methyltransferases upstream transcription factor II LCR: locus control region CT3: cardiac troponin T Mef2c: myocyte enhancer factor 2C CTCF: CCCTC-binding factor MeOH: methanol CVD: cardiovascular disease MESP1: mesoderm posterior 1 homolog CVS: cardiovascular system MRF: myogenic regulatory factors DAPI: 4’,6-diamidino-2-phenylindole MyoD: myogenic differentiation 1 DEPC: diethylpyrocardbonate Nkx2.5: NK2 homeobox 5 Dll4: delta-like ligand 4 OFT: outflow tract DUB: deubiquitylases PBS: phosphate buffered saline E: embryonic day PCR: polymerase chain reaction Elf-1: E74-like factor 1 PFA: paraformaldehyde EMT: epithelial-to-mesenchymal pHH3: phosphorylated histone H3 transition PLGF: placental growth factor eNOS: endothelial nitric oxide synthetase PTM: post-translational modifications Er71: ETS-related 71 RT: room temperature Erg: ETS related gene RTK: receptor tyrosine kinase EtOH: ethanol Scl: stem cell leukemia ETS-1: V-Ets Avaian Erythroblastosis SHH: sonic hedgehog Virus E26 Oncogene Homolog 1 Six1: SIX homeobox 1 Ezh2: enhancer of zeste homolog 2 SOX7: Sex Determining Region Y – Box FACs: fluorescently activated cell sorting 7 FGF: fibroblast growth factor Tbx5: T-box-5 Fli-1: friend leukemia integration 1 Tel: translocation-Ets-leukemia or ETV6 transcription factor TF: transcription factors Flk1: fetal liver kinase 1 VEGF: vascular endothelial growth factor FOX-C2: forkhead transcription factor VEGFR: vascular endothelial growth G9a/EHMT2: euchromatin histone-lysine factor receptor N methyltransferase 2 VSD: ventricular septum defects GATA4: GATA binding protein 4 GLP: G9a – like protein

vi List of Figures Chapter 1 Figure 1-1 – Mouse cardiac development------2 Figure 1-2 – Mouse vascular development------6 Figure 1-3 – Transcriptional regulation by chromatin modifiers------11 Chapter 3 Figure 3-1 – Ctcf is present in cardiomyocytes in the developing and adult heart------20 Figure 3-2 – Inactivation of Ctcf in cardiac lineages------22 Figure 3-3 – Ctcf in cardiomyocytes is essential for embryonic development------23 Figure 3-4 – Ctcf deficient cardiomyocytes cause severe heart defects at E12.5------24 Figure 3-5 – Ctcf mutants at E12.5 do not have cell death or proliferation defects------25 Figure 3-6 – Ctcf mutant hearts have downregulation of well-characterized cardiac genes------27 Figure 3-7 – Ctcf organizes the IrxA cluster in to distinct regulatory domains------28 Chapter 4 Figure 4-1 – Ctcf is present in mouse embryonic and adult endothelial cells ------29 Figure 4-2 – Inactivation of Ctcf in endothelial progenitors------31 Figure 4-3 – Ctcf in endothelial cells is essential for embryonic development------32 Figure 4-4 – Ctcf mutant embryos do not exhibit vascular defects in the trunk and heart at E10.5------33 Figure 4-5 – Ctcf mutants have narrow cerebral vasculature------34 Figure 4-6 – Ctcf mutants have yolk sac vascular remodelling defects------36 Figure 4-7 – Endothelial specific Ctcf mutant yolk sac show no change in cell death or proliferation at E9.5------37 Figure 4-8 – Ctcf mutant yolk sac has no significant change in eNOS levels in E9.5-----38 Figure 4-9 – Ctcf mutant placental labyrinth have a reduction in vascular area at E10.5------40 Figure 4-10 –Endothelial specific Ctcf mutant placentas have no cell death or proliferation defects------41 Figure 4-11- Endothelial specific Ctcf mutant placentas at E9.5 have an increase in expression of vascular regulatory genes------42

vii Chapter 1 – Introduction The cardiovascular system (CVS) is one of the first systems to develop in mammalian embryos (1, 2, 3). It is comprised of a beating heart and a hierarchical vascular network that spreads throughout the whole body, as well as extraembryonic tissues like the placenta and yolk sac during development (1, 2, 3). The development of the CVS is crucial for the maintenance of embryogenesis and is regulated by various transcriptional programs, patterned and organized in a spatiotemporal manner. (1-4). Understanding the developmental regulation of CVS is fundamental to comprehending congenital cardiovascular diseases, specifically congenital heart disease (CHD) (1, 4). CHD arise from misregulation of developmental transcriptional programs and affect approximately 1% of live newborns worldwide (1, 4). It is known that chromatin regulators tightly regulate these CVS transcriptional programs (1, 5). Thus, understanding the involvement of chromatin regulators in cardiovascular development will potentially help comprehend the origins of congenital cardiovascular diseases (1, 5). One potential epigenetic regulator involved in cardiovascular development is the CCCTC-binding factor (CTCF). In my thesis, I have started an examination of the requirement of CTCF in mouse cardiovascular development. In this chapter, I will give an overview of mouse cardiovascular development, the different types of transcriptional programs involved, and provide a rational for investigating the chromatin regulator, CTCF, in mouse cardiovascular development.

1.1 Heart Development During early stages of mouse embryonic development, a cluster of cells in the blastula form a structure known as the primitive streak at embryonic day (E) 6.5 (1, 2). The cells in the primitive streak undergo a series of cell specification, division and migration via epithelial-to-mesenchymal transition (EMT), to give rise to the three germ layers (ectoderm, mesoderm and endoderm) (1, 2, 5, 6, 7). This event is known as gastrulation (1, 2, 5, 6, 7). It is the mesodermal layer that gives rise to components of the heart, vasculature, and skeletal muscle (7). Around embryonic day (E) E6-7.5, a subset of mesodermal cells from the anterior pharyngeal mesoderm, give rise to a common cardiac progenitor cell population (Fig 1.1, 1, 2). This population differentiates into two cardiac

1 progenitor cell populations (1,2). The cardiac progenitor cells from the anterior mesoderm make up the cardiac crescent, also known as the first heart field (FHF) (1,2). The cardiac progenitor cell population from the pharyngeal mesoderm, known as the second heart field (SHF), situates anteriorly and medially of the cardiac crescent (1,2,8). During E8.0, cells of the cardiac crescent migrate together to form a linear heart tube (Fig 1.1, 1, 2, 8). This linear heart tube enlarges in size at the anterior and posterior ends. This heart tube acts as the foundation to form the future heart. At the same time, cells from the secondary heart field migrate towards the venous and arterial poles of the linear tube (1,2,8). The heart tube undergoes rightward looping by E8.5, and the venous pole shifts anteriorly at E10.5 to form primitive atria and ventricles (Fig 1.1, 1, 2, 8). This is analogous to the heart sitting on itself and in turn causing chamber formation. Morphologically, the four separate chambers of the heart are established after septation has occurred at E15 (Fig 1.1, 1, 2, 8). The first heart field helps further compose the left ventricle and parts of the left and right atria (1, 2, 8). The secondary heart field contributes to structuring the right ventricle, parts of the left and right atria and the outflow tract (OFT) (1,2). The OFT undergoes subsequent septation to create the aorta and pulmonary artery of the developing heart (1,2).

Figure 1.1. Mouse cardiac development. Image modified from (2). First heart field (cardiac crescent) represented in red, and second heart field represented in blue are formed at E7.5. The first and second heart fields merge to form a linear heart tube at E8.0, and undergo rightward looping at E8.5. The venous poles moves anterior to form the future heart chambers at E10.5. At E15.5, septation of the heart forms the four separate chambers of the heart.

2 1.2. Molecular regulation of the developing heart The process of heart development is comprised of the complex organization of cell signalling, communication, and tight gene regulation of the cardiogenic transcriptional programs (2). The earliest known cardiac progenitors are observed during gastrulation, after the expression of mesoderm posterior 1 homolog, Mesp1, in a subpopulation of mesodermal cells (9, 10). Mesp1 expressing multipotent progenitors at E6.75 give rise to the first and second heart fields (9, 10). These heart fields merge together to form the cardiac crescent and linear cardiac tube at the embryonic midline (9, 10). Active and repressive signals from major signalling pathways such as Bone Morphogenetic Protein (BMP), Fibroblast Growth Factor (FGF), Wnt, Sonic Hedgehog (SHH) and Notch signalling, regulate cell survival, proliferation, differentiation and cell fate of cardiac progenitors in the FHF and SHF (11). Signals from these pathways initiate a series of cascading events to stimulate Mesp1 expressing progenitors to trigger the activation of key cardiac transcription factors (TF) including Gata4, Nkx2.5, Tbx5, Isl1 and Baf60c, a chromatin remodeller, in the cardiac tube (1, 4, 12-13). Gata4 in association with Baf60c, subunit of the BAF chromatin-remodelling complex, physically activates Nkx2.5 in both FHF and SHF to regulate the gene expression of downstream targets responsible for cardiac differentiation (2, 4). This indicates that TFs do not work alone but can affiliate with chromatin mediators to induce cardiac differentiation. Cardiac transcription factors can give rise to different cardiac lineages comprised of cardiomyocytes and endothelial cells in the FHF and cardiomyocytes, endothelial cells and smooth muscle cells in SHF (10). These cardiac lineages make up different structures of the heart. These TFs are critical for cardiac development, as deficiency induces severe cardiac defects and embryonic lethality (1, 2, 4). Mouse embryos null for Gata4 have defective heart looping, septation and hypoplastic ventricles (14). Inactivation of Nkx2.5 causes arrested heart development during cardiac looping, resulting in a single ventricle with thin myocardium and reduced ventricular trabeculation (15, 16). In cells of the FHF, Tbx5 interacts with Gata4 and Nkx2.5 to regulate cardiac progenitor cell differentiation, chamber formation and ventricular septation (17-19). As the linear heart tube forms, Tbx5 has a gradient expression, concentrated in the cells of the left ventricle (17-19). This Tbx5 expression becomes exclusively restricted to the left ventricle as the heart

3 matures and is maintained in adult hearts (17-19). Haploinsufficiency of Tbx5 causes heart malformations consisting of atrial septum defects (ASDs), ventricular septum defects (VSDs) and cardiac conduction system defects (17-19). In cells of the SHF, expression of Isl1, activated by Wnt/β-catenin signalling, is a common marker for cardiac progenitors and mediates cell survival, proliferation and differentiation in cardiogenesis (20, 21). Isl1 null mice do not undergo cardiac looping and lack the formation of SHF derived cardiac structures such as outflow tract, atria and right ventricle (20, 21). The expression of the SHF transcription factor, Hand2, along with Gata4, interacts with the Mef2c transcription factor (4, 22). Mef2c, controlled by Notch signalling, induces cardiac ventricular formation (4, 22). Loss of Hand2 or Mef2c in mice impairs ventricular formation (23, 24). Cardiac transcription factors controlled by Notch, Wnt, FGF, BMP, and SHH signalling pathways mediate cardiogenic transcription programs during mouse heart development.

1.3. Vasculature Development: As the beating heart pumps replenished and nourishing blood, the vasculature acts as a bridge for the blood to travel from the heart to the body. The development of the vasculature begins at E7.5 (25, 26). Mesodermal precursors migrate from the primitive streak to the embryo proper and extraembryonic tissues, such as the yolk sac and placenta (Fig 1.2A, 25, 26). The yolk sac is the first place for vasculogenesis, the de novo formation of blood vessels (25, 26). In the yolk sac, mesodermal precursors aggregate to form blood islands, that later give rise to the different components of the vasculature (Fig1.2A, 25, 26). The cells in the center of the blood island form hematopoietic blood cells, while the cells radially form endothelial cells, lining the blood vessels (25, 26). The endothelial cells in the yolk sac subsequently form the vascular primary plexus and undergo extensive remodelling (25-27). Vascular remodelling is comprised of angiogenesis, the formation of new blood vessels branching from preexisting vessels, vascular pruning and organization of large and small blood vessels into a hierarchical vascular network during E8.5-9.5 (25 -27). At the same time, mesodermal precursors that migrate to the embryo proper differentiate to form the vascular primary plexus (26). The

4 primary plexus undergoes arterial and venous specification and vascular remodelling to become an organized vascular network that runs through all the developing organs (26).

During embryonic development, the hierarchical vascular network also develops in the placenta (28). The placenta acts as the interface where nutrient and gas exchange occur between the mother and the developing fetus (28). The placenta is derived from the trophectoderm, the outer cell layer in the blastocyst stage at E3.5 (Fig1.2 B, 28). After implantation of the blastocyst into the maternal uterine wall at E4.5, the mural trophectoderm, not in contact with the inner cell mass, gives rise to trophoblast giant cells (Fig1.2B, 29). The polar trophectoderm, cells in contact with the inner cell mass, differentiate into the extraembryonic ectoderm and the ectoplacental cone (Fig 1.2B, 29). At E8.0, the allantois is formed by the budding of extraembryonic mesoderm from the posterior end of the embryo (28, 29). The allantois is a site of vasculogenesis and grows to meet the placental chorion, in a manner known as the chorio-allantoic attachment (28, 29). The allantois contains precursors essential for the development of the umbilical cord, connecting the mother to the embryo (28, 29). This process causes villi formation and ruffling or folding of the chorion (28). This creates space for the vasculature to form from the allantois and invade into the placenta at E9.5-10.5 (28). At the same time, the maternal blood vessels in the placenta lose their endothelial lining, which is replaced by placental syncytiotrophoblast cells (28, 29). Syncytiotrophoblast cells act as interface for gas and nutrient exchange between the maternal vasculature and the fetal vasculature in the labyrinth layer of the placenta (28). The development of the placenta is crucial to maintaining proper embryogenesis (28, 29).

5 Figure 1.2. Mouse vascular development. Image modified from (25, 28). A. Vascular development in yolk sac and embryo proper. In the yolk sac, endothelial progenitors aggregate to form blood islands. Inner cells in the blood island become hematopoietic cells, while outer cells differentiate into endothelial cells lining blood vessels and later primary plexus. Further differentiation and remodelling of blood vessels form a complex network of yolk sac vascular network. In the embryo proper, endothelial progenitors aggregate to form primary plexus that differentiate and remodel into mature vascular networks. B. Vascular development in the mouse placenta. The trophectoderm layer at E3.5 later gives rise to the trophoblast cells that differentiate into the extraembryonic ectoderm and ectoplacental cone to form the placenta at E8.0. The allantois budding from the posterior end of the embryo extends to meet the placental chorion in what is known as chorio-allantoic attachment at E8.0-8.5. The allantois undergoes de novo formation of blood vessels and as a results, the chorion ruffles to make space for vasculature expansion into the placental labyrinth region at E9.5-10.5.

6 1.4. Molecular regulation of vascular development Similar to cardiac development, multiple transcription factors and signalling pathways mediate endothelial cell differentiation in vascular development. One essential pathway for early embryonic vascular development is the vascular endothelial growth factor (VEGF) signalling pathway (30). The VEGF signalling pathways are mediated by three receptor tyrosine kinases (RTKs): vascular endothelial growth factor receptor 1 (Vegfr1), Vegfr2, Vegfr3 (31), These receptors are induced by five ligands, the vascular endothelial growth factor A (VegfA), B, C, D, and placenta growth factor (Plgf) (31). Mesodermal progenitor cells in the posterior primitive streak are induced by basic fibroblast growth factor (bFGF) and BMP4 signalling, along with ETS-related 71 (Er71) and forkhead transcription factor (Fox)-C2 transcription factors, to express Vegfr2, also known as fetal liver kinase 1 (Flk1) (26, 32). Flk1-expressing mesoderm is a multipotent progenitor population that gives rise to a broad spectrum of mesodermal cell lineages consisting of endothelial, hematopoietic, cardiomyocyte, smooth muscle, and skeletal muscle cells (26). Flk1-expressing progenitors are restricted to endothelial and hematopoietic lineages via Er71, Gata1, 2, and stem cell leukemia (Scl) transcription factors (26, 32). Er71 is apart of the Ets family, consisting of Ets-1, Elf-1, Fli-1, Tel and Erg and is important in activating essential endothelial genes by binding to their enhancers (33). Er71 is a central regulator of endothelial cell specification and inhibits Flk1 positive progenitors from becoming cardiogenic mesoderm (32). Flk1 positive cells migrate from the primitive streak to the yolk sac to form blood islands and subsequent primary plexus (26, 32). VegfA signalling in Flk1 expressing cells further mediate the development of the vascular and hematopoietic pathways (26, 32, 33). Mouse embryos null for Flk1 are embryonic lethal at E9, lack blood vessels and hematopoietic cells (32). VegfA haploinsufficiency leads to embryonic lethality at E10-11 with reduced vascular blood vessel size (32). The maturation of the primary plexus, mediated by Ets and Kruppel-like transcription factor (Klf) proteins, undergo endothelial cell differentiation, specification and migration to form the primitive vasculature (33). In culture, endothelial cells lacking Ets1 and Erg have a reduction in endothelial cell migration and blood vessel tube development (33). Notch signalling pathway further differentiates the primitive vascular endothelium into arterial and venous specification (32, 33). The direct

7 downstream targets of Notch, Hairy/enhancer-of-split related with YRPW motif protein 1 (Hey1) and Hey2, along with FOXC1, 2 mediate the specification of arterial endothelium (26, 32). Venous endothelium is specified by the suppression of Notch signalling by chicken ovalbumin upstream transcription factor II (Coup-TFII) and the Sox family factors, Sox7, 18 (26, 32). The maturation of the vasculature, along with VegfA and Notch signalling, requires Tie2-Ang1 signalling (34). Tie2, a member of the receptor tyrosine kinase family, with its angiopoietin-1 (Ang-1) ligand, is expressed in the vascular endothelium and maintains the stabilization and remodelling of developing blood vessels (34). Ang-1 null mice dies at E12.5, with ruptured blood vessels, decreased vascular complexity containing reduced branching and number of endothelial cells (34). The positioning, migration and sprouting of blood vessels are mediated by VegfA attractant gradients that direct the vascular pathway (34). Notch signalling further control blood vessels branching and morphogenesis (32). Activation of Notch signalling inhibits blood vessel branching, while loss of Delta-like ligand 4 (Dll4), a Notch ligand, causes an increase in ectopic vessel branching (26, 32). Notch and VegfA signalling are central mediators for the activation of vascular transcription factors responsible for cell survival, proliferation and differentiation of vascular lineages during development (26, 32).

1.5. Transcriptional regulation by chromatin regulators The previous sections highlight various signalling pathways in cardiovascular development mediating the activation of key transcription factors involved in the differentiation of cardiovascular lineages. Chromatin regulators tightly regulate and coordinate transcription factors in cardiovascular gene expression programs by altering chromatin structure (Fig 1.3, 5). The chromatin structure consists of the packaging and storing of DNA into 4 pairs of histone core proteins (1). The histone core consists of H2A, H2B, H3, and H4 molecules (1). Histone modification factors, or histone modifiers, are a subset of chromatin regulators that can chemically modify histones (1, 5). Chromatin regulators or transcription factors can read these chemically modified histones to help reveal or enclose chromatin from the histone core (1, 5). This allows DNA accessibility or inaccessibility for transcription factors to mediate active or repressive gene expression, respectively (Fig 1.3, 1, 5). Chromatin regulators can also mediate long

8 range-chromatin interactions that enable distant regulatory elements to act on gene promoters and create chromatin transcriptional hubs (35). This permits inclusive or exclusive transcription of genes by the transcriptional machinery (35). The function of chromatin regulators in cardiovascular transcriptional programs will be further addressed in the sections below.

1.6. Histone modifiers regulating cardiovascular development Histone modifiers are essential in cardiovascular development (1). Histone modifiers are proteins that chemically alter histones through post-translational modifications (PTMs), such as phosphorylation, methylation, acetylation and ubiquitylation (1, 36). Histone modifiers that add or remove such PTMs consist of the following; Histone acetyltransferases (HATs), histone deacetylases (HDACs), which add or remove acetyl groups on conserved lysines in histone proteins, lysine methyltransferases (KMTs) and lysine demethylases (KDMs), which add or remove methyl groups on lysine amino groups, deubiquitylases (DUBs) and ubiquitylation enzymes (E1, E2, E3), which add or remove ubiquitin, respectively (1, 36, 37). The combination of histone modifications by these proteins form a distinct “histone code” (38). Proteins can read this code to instruct active or repressive gene transcription and subsequently regulate gene expression (38). Chromatin regulators can induce long-range chromatin interactions to further elicit the instructions presented by the histone code (1, 39). The process of long-range chromatin interactions mediated by chromatin regulators can traffic the accessibility of the transcriptional machinery (1, 39). Our lab and others have shown that histone modifiers are central regulators of both cardiac and vascular development (36, 37, 40-42). The histone methyltransferase enhancer of zeste homolog 2 (Ezh2), part of the polycomb repressive complex 2 (PRC2), represses gene expression by trimethylating lysine 27 of histone H3 (H3K27me3) (42). Ezh2 is crucial for heart development (42). Deletion of Ezh2 by Nkx2.5-Cre-mediated homologous recombination in cardiac progenitor cells causes embryonic lethality at late gestation with hypertrabeculation and ventricular septal defects (42). Repressive marks set by Ezh2 during development persist into adulthood (40). Deletion of Ezh2 in the anterior heart field cardiac progenitors induces postnatal right ventricular cardiac hypertrophy and

9 fibrosis (40). The gene encoding the SIX homeobox 1 (Six1) transcription factor is expressed in cardiac progenitors but is repressed during cardiomyocyte differentiation by Ezh2 mediated repression (40). Loss of Ezh2 disrupts proper cardiac gene expression via activation of Six1-mediated cardiac genes, resulting in adult-onset of cardiac hypertrophy (40). This is indicative of histone modifiers having essential functions in the developing and adult heart. Ezh2 also controls vascular development (41). Ezh2 inactivation in endothelial progenitor cells causes embryonic lethality at E13.5-14.5 with vascular rupture, suggestive of poor vascular integrity (41).

The repressive histone modifiers, Glp and G9A, which methylate H3 lysine 9 (H3K9me), can also regulate cardiovascular development (43). Combined Glp and G9a knockdown in mouse cardiomyocytes induces severe cardiac defects such as atrioventricular septal defects and leads to lethality at neonatal stages (43). Current work from my lab has found that the expression of G9a in endothelial progenitors is essential for embyogenesis. G9a mutant embryos are inviable at E15.5, with vascular defects consisting of reduced vascular branching in the placental labyrinth (Work in progress, Lijun Chi). Preliminary data show G9a in endothelial progenitors may target the Recombination signal binding protein for immunoglobulin kappa J region, Rbpj, transcription factor. Rbpj is an important regulator of endothelial gene expression via Notch signalling (Work in progress, Lijun Chi). Further analyses on the mechanistic basis of G9a in endothelial progenitors are under way.

10 Figure 1.3. Transcriptional regulation by chromatin modifiers. Image modified from (44). Chromatin modifiers can regulate cellular transcriptional programs by controlling gene expression at the histone level and/or via long-range chromatin interactions.

1.7. Does the process of long-range chromatin interaction modulate CVS development and function? Chromatin regulators that mediate the process of long-range chromatin interactions allow three-dimensional organization of chromatin structures, regulate gene expression by controlling enhancer to promoter interactions, and can coordinate transcriptional programs by creating boundaries between different transcriptional activities or histone modifications and topological domains (Fig 1.3, 35, 46, 47, 48). One instances of three-dimensional organization of chromatin is seen with the chromatin regulator, CCCTC-binding factor, CTCF and the lamina-associated domains (LADs) (48). Flanking CTCF binding sites at LADs allow CTCF along with lamina-based proteins to arrange transcriptionally inactive chromatin into loops and bind to the nuclear envelope away from the active nuclear center (48). Long-range chromatin interactions can create chromatin loops that allow simultaneous transcription of a group of genes (46, 47). This is observed in the β-globin locus control region (LCR) in erythroid cells. The

11 LCR located next to the actively expressing β -goblin genes are present in active chromatin loops to bring the genes physically closer. This allows efficient transcription of the β -goblin genes and results in high gene expression levels (39, 47, 49, 50). Chromatin loops can also repress sets of genes by excluding these genes out into chromatin loops from the prospective transcriptional machinery (46, 47). Chromatin regulators, like CTCF, can act on boundaries of active and repressive histone modifications and can separate these domains by the process of long-range chromatin interactions (48). Interruptions of such process of long-range chromatin interactions by chromatin regulators can disrupt these histone domains and cause gene misregulation (48). As long- range chromatin interaction act on genes in varying distances, this adds another level of complexity to gene regulation. As such, understanding chromatin regulator mediated long-range chromatin interaction is important to comprehending the transcriptional regulation involved in development processes, like the CVS. However, it is not known if gene regulation mediated by the process of long-range chromatin interaction is important for guiding CVS development and function. It is not known which genes in CV development are sought to undergo long-range chromatin interactions or how the process of long-range chromatin interaction regulates the transcriptional programs controlling CV development. Addressing these questions will help understand the transcriptional regulation involved in the development and function of CVS.

1.8. The function of CCCTC-binding factor, Ctcf The CCCTC-binding factor, Ctcf, is an important mediator of long-range chromatin interactions (48, 51, 52). Ctcf is a transcriptional regulator, first discovered in the upstream transcriptional start site of the chicken c-myc gene (51, 53). There, it recognizes CCCTC repeat sequences and acts as an insulator (51, 53). An insulator prevents the interaction of distal regulatory elements with the promoter (51, 53). Ctcf is a ubiquitously expressed transcription factor. It consists of 11 highly flexible zinc fingers used in a combinatorial fashion to bind to DNA and multiple protein partners (48, 51). This allows Ctcf to perform various functions in the genome (48, 51). For example, Ctcf can act as barrier element, protecting actively transcribing genes by blocking the spread of repressive heterochromatic marks set by histone modifiers (48, 51). Ctcf is also known

12 to mediate long-range chromatin interactions (48, 51). It can bring together distal regulatory elements located in varying genomic distances to gene promoters (48, 51). The interaction of distal regulatory elements creates chromatin loops that distinguish transcriptionally active or repressive chromatin hubs (35, 48, 51). Ctcf can form homodimers or interact with other protein partners, such as cohesins to mediate long- range chromatin interactions (35, 48, 51). The function of CTCF in long-range chromatin interactions can be exhibited in the insulin-like growth factor 2, IGF2, and H19 locus, which is regulated by a ∼2.4 kilobase (kb) imprinting control region (ICR) in germ cells (54, 55). The ICR located between the two genes is 90kb away from the Igf2 gene and near the 5’ flanking region of H19 gene (54, 55). The ICR is differentially methylated. In the maternally derived chromosome, the ICR is unmethylated and H19 is expressed, whereas in the paternally derived chromosome, the ICR is methylated and Igf2 is expressed (54, 55). These imprinting genes share a common set of enhancers acting downstream of the H19 gene (54, 55). In the maternal allele, the unmethylated ICR allows for the binding of CTCF (48, 52, 54, 55). This creates a boundary in which enhancer activation of the Igf2 promoter is blocked and the Igf2 gene is silenced, to allow expression of the H19 gene (48, 52, 54, 55). The silencing of the Igf2 is performed by CTCF mediated chromatin loops in the maternal chromosome and binding of the N- terminus domain of CTCF to the PRC2 (54, 55). PRC2 tethers to CTCF and helps place repressive H3K27me3 marks to silence the Igf2 gene (54, 55). In the paternal allele, CpG methylation of the ICR prevents the binding of CTCF (48, 52, 54, 55). This allows the enhancer activation of Igf2 gene and repression of H19 by hypermethylation of the H19 promoter (48, 52, 54, 55). The regulation of IGF2/H19 imprinting by CTCF is an exceptional example, displaying the multifaceted roles of CTCF, which acts as an insulator, can mediated long-range chromatin interactions, and can interact with histone modifiers.

1.9. Role of Ctcf in development Ctcf is essential for embryogenesis (35, 48, 51-52). Ctcf deficient mice typically die around the peri-implantation stage (E3.5), but can survive up to E4.5-5.5 with the assistance of the maternal contribution of Ctcf (56). The function of Ctcf is crucial in

13 various developmental processes (35, 48, 51-52). In limb development, Ctcf is required for cell survival of mesenchymal cells and its deficiency causes severe apoptosis and shortening of the limbs (57). In anterior-posterior patterning of the embryonic body axis, the HOX genes, organized into HOXA-D clusters, are important regulators (58). Ctcf coordinates the arrangement of the HOXA gene cluster (58). Ctcf is required for chromatin looping of the HOXA genes and induces PRC-mediated silencing of these genes (58). Deletion of Ctcf causes improper activation of genes in the HOXA cluster (58). In T and B lymphocyte development, Ctcf facilitates proper gene organization and communication during V(D)J recombination of lymphocyte immunoglobins (Ig) and T cell receptors, via long range-chromatin interactions (48, 51, 52, 59). In muscle development, Ctcf regulates the myogenic regulatory factors (MRFs), MyoD and myogenin, to initiate myogenic differentiation by enabling interaction of MyoD and myogenin with muscle-specific promoters (60). However, while the function of Ctcf is known in many developmental processes, little is known about its function in regulating the transcriptional programs that control mouse CV development. Understanding the function of Ctcf in CV development will help determine how the process of long-range chromatin interactions guides development and function of CV systems.

1.10. Ctcf and cardiovascular development Knowledge of the role of Ctcf in regulating cardiovascular transcriptional programs is limited to its function in restraining retinal angiogenesis (61). Ctcf knockdown in mouse neonatal retinas resulted in ectopic growth of blood vessels. It was revealed that Ctcf binds between the hypoxia response element (HRE) and the proximal promoter of vascular endothelial growth factor (VEGF) to prevent upstream enhancer activation of Vegf and restrain growth of retinal angiogenesis (61). Numerous studies have revealed that CTCF Binding Sites (CBSs) flank key genes encoding transcription factors involved in diseases, including cardiovascular disease (62). However, the function of Ctcf and the role of CTCF Bindings Sites in flanking genes encoding transcription factors associated with heart disease are not known. It can be postulated that perhaps Ctcf functions in the transcriptional regulation of CV development and function.

14 1.11. Thesis Objectives In my thesis work, I started to investigate the requirement of Ctcf in CV development. Ctcf is a transcriptional regulator essential for embryogenesis. It is known to function in many developmental processes. However the knowledge of its function in CV development is limited to its ability to restrain neonatal retinal angiogenesis in mice by acting as an insulator. It is known to flank genes encoding transcription factors associated with cardiovascular disease. This suggests that Ctcf may function in the transcriptional regulation of CV development. For my thesis, I hypothesized that Ctcf is required for mouse cardiovascular development. I sought to uncover the requirement of Ctcf in cardiac and vascular development. To accomplish my thesis objectives, I took advantage of mouse models that specifically inactivates Ctcf in cardiac progenitors and in endothelial progenitor cells. I found that the expression of Ctcf in both the heart and vascular endothelium is essential for embryogenesis. Inactivation of Ctcf in early cardiac progenitors caused embryonic lethality at E12.5 with severe heart defects. In collaboration with Dr. Manzanares’ Lab, it was revealed that Ctcf regulates the expression of key cardiac development regulators including the Irx4 gene, involved in ventricular formation. Inactivation of Ctcf in endothelial progenitor cells caused embryonic lethality at E10.5 with vascular defects in the embryo proper and in extraembryonic tissues. Although the mechanistic basis of Ctcf and its regulation of transcriptional programs in vascular development via long range-chromatin interactions remain to be addressed. The overall goal of my thesis will focus on the following two objectives:

1. To elucidate the requirement of Ctcf in mouse heart development.

2. To elucidate the requirement of Ctcf in mouse vascular development.

15 Chapter 2 – Materials and Methods

2.1. Mice Mice strains utilized are the following: Ctcf fl/fl (63), Nkx2.5;cre (64), Tie2;cre (65), and ROSA26mT/mG (66). All animal experiments were approved by the Animal Care Committee and followed the guidelines of the Toronto Centre for Phenogenomics. Matings were set up and vaginal plugs were checked the following day. Each day after the presence of a vaginal plug is considered one embryonic day. Pregnant mice according to the desired stage were euthanized using CO2. All embryos were dissected in cold 1xPBS, fixed in 4% Paraformaldehyde (PFA) and stored in 4˚C overnight. The fixed embryos were dehydrated in a methanol (MeOH) series: 25% MeOH/PBS, 50% MeOH/PBS , 75% MeOH/PBS and 100% MeOH/PBS for 15 min in room temperature (RT) and stored in -20˚C until processed for frozen and histological sectioning and staining. Before processing, embryos were rehydrated with PBS by reversing the methanol series washes.

2.2. Genotyping Mouse tail clips, ear notches and yolk sacs were used for genotyping. Tissues were digested with 300µl of 50mM NaOH at 95˚C for 10-30min, until fully digested. Digestion was stopped with 100µl of 0.5µm tris HCl, and the samples were vortexed. 1µl of digested DNA was used for genotyping by polymerase chain reaction (PCR). PCR reactions were prepared in a total volume of 20µl PCR mix, containing 2µl of 2mm of dNTP, 2µl of 10x PCR buffer, 2µl of forward and reverse primer mix, 0.5µl of Taq and DEPC water up to 20µl. The following thermal cycler conditions were used to amplify floxed alleles of Ctcf are the following: 94°C 3min, 94°C 30sec, 63°C 30sec, 72°C for 1min, repeat steps 2-4 for 35 cycles, then 72°C for 5min, and 12°C infinite hold. Thermal cycler conditions used to amplify Cre promoter are as follows: 95°C 2min, 95°C 40sec, 55°C 50sec, 72°C 1min, repeat steps 2-4 for 35 cycles, and 72°C for 10min. The following primers were used: Ctcf; forward CTAGGAGTGTAGTTCAGTGAGGCC, reverse GCTCTAAAGAAGGTTGTGAGTTC, and Cre; forward ATCCGAAAAGAAAACGTTGA, reverse ATCCAGGTTACGGATATAGT.

16 2.3. Immunostaining 2.3.1 Immunofluorescence Frozen sections (4-8µm) on glass slides were fixed in 4% PFA for 5 min and washed in 1x PBS for 3x 5 min each. Slides were blocked for 15min with 5% goat serum and 0.1% triton x-100 in 1x PBS, and incubated with primary antibodies diluted in blocking buffer at 4˚C overnight in a humidified chamber. Slides are washed in 1xPBS 3x 10 min each and incubated with secondary antibodies, diluted in blocking buffer, for 1h at room temperature in a dark humidified chamber. Slides are washed in 1xPBS, 3x 5min each and then once with 1xPBS with 0.05% tween 20 for 5min. Slides were then mounted with VectaShield mounting medium with DAPI (Life Technologies). The following antibodies and their respective dilutions were used: Ctcf (67) (1/300), phosphorylated histone H3 (Santa Cruz, SC-8656-R, 1:100), cleaved caspase 3 (Cell Signaling, 9661, 1:100), CT3 (Development Studies Hybridoma Bank, 1:200), CD31/pecam-1 (BD Pharmingen, 553370, 1:100), and GFP (GeneScript, A01694, 1:1000).

2.3.2. Whole mount immunostaining of embryos Embryos were fixed in 4% PFA overnight and washed in 1xPBS for 3x 5min each before they were permeabilized and blocked for 1h at RT in PBT (1xPBS with 0.2% Triton-X-100) with 0.1% BSA and 2% goat serum. Embryos were then incubated with primary antibodies (Pecam-1) diluted in blocking buffer (0.1% BSA and 2% normal goat serum) at 4˚C overnight. Embryos were washed 1xPBS for 5x 5 min each, and blocked in PBT with 2% normal goat serum for 1 h at room temperature. Embryos were incubated with secondary antibodies diluted in 0.1% BSA and 2% goat serum for 1 hour at room temperature in the dark. Embryos were washed in 5x 1xPBS for 5min each. Finally, embryos were cleared by immersion in 1:1 glycerol to PBS for 3h at 4˚C, and then in 80% glycerol in PBS for 1 h at 4˚C before imaging.

2.4 Histology Samples collected from E9.5 to adult stages were fixed in 4% PFA overnight at 4˚C, washed with 1xPBS for 30min twice at 4˚C, and placed in 70% Ethanol (EtOH) and stored at 4˚C. Embryos were then dehydrated at RT in an Ethanol series: 85% EtoH 2x

17 for 30min each, 95% EtOH 2x for 30min each, 100% EtOH 4x for 45 min each, 100% xylene for 5 min, washed in xylene 3x for 10min each. Next, the embryos were placed in 50% xylene:wax for 30 min at 60˚C and at RT over night, and then placed back at 60˚C for 30min. The xylene:wax was replaced with wax 2x for 1 hour and with new wax for 2 hours at 60˚C. After the dehydration process, embryos were embedded in paraffin wax trays. Wax blocks were then mounted on histology cassettes. The wax block was sectioned on a microtome at a thickness of 4-8µm. The sections were put on water and then on glass slides, which were then incubated for 30min at 37˚C and then at 60˚C for 4 hours to overnight. The slides were prepared for staining with hematoxylin and eosin. Slides were placed in the following solutions: xylene 2x for 10 min, an EtOH series consisting of 100% EtOH 2x for 2min each, 90% EtOH for 2min, 70% EtOH for 2min, 50% for 2min, 30% for 2min, then tap water 3x quickly. The slides were then stained with hematoxylin for 10min, washed under tap water 3x, placed in 0.5% acid alcohol for 5 sec, washed under tap water 3x, placed in 1% lithium carbonate for 5 sec, washed under tap water 3x, and then subjected to the EtoH series washes, of 30%, 50% and 70% for 1 min each. Next, the slides were stained with 3% Eosin (diluted Eosin in 705 EtOH) for 10 min, then washed with 90% EtOH for 1min, 100% EtOH 2x for 1min and placed in xylene x2 for 5 min. Lastly, the stained slides were mounted with Permount mounting media and left to dry in the fume hood.

2.5. Gene Expression Analysis Freshly dissected placentas and yolk sacs of E9.5 embryos were collected in Trizol Reagent (Invitrogen) and RNA was extracted, using the Zymo Research Direct™RNA MiniPrep kit. cDNA was synthesized from RNA using SuperScript® VILO cDNA synthesis kit (Invitrogen). The cDNA template, 1µl from original cDNA synthesis reaction, was used in 10µl qPCR reactions. For qPCR, BIO-RAD SsoAdvanced™ Universal SYBR® Green Supermix was used in 384-well plates. The qPCR program was ran on the CFX384 Touch™Real-Time PCR Detection System using the provided 3AMPMELT program with an annealing temperature of 61.6°C. Thermal cycler conditions were as followed: 95°C 30sec, 95°C 15sec, 61.6°C 15sec, 72°C 30sec, plateread, repeat 55x, and a melt curve conditions of: 95°C 10sec, 55°C 31 sec, 55°C

18 5sec + 0.5°C /s Ramp 0.5°C/s, plateread, repeat 80x. Data was analyzed using BIO-RAD CFX manager software and relative gene expression to housekeep gene was acquired. The following primers were used: Kdr, forward CTGTGAACGCTTGCCTTATG, reverse AGTCGCTGTCTTGTCAATTCC; VegfA, forward ATCTTCAAGCCGTCCTGTG, reverse TCTCCTATGTGCTGGCTTTG; Erg, forward GCCTCCCAATATGACCACAA, reverse ATTCTTTCACCGCCCACTC; eNOS, forward GCACCCAGAGCTTTTCTTTG, reverse TGACACAATCCCTCTTTCCG.

2.6. Microscopy and Imaging Whole mount embryo and immunofluorescence imaging were done using the Nikon SMZ1500 microscope and the Nikon Eclipse Ni microscope, respectively. Quantification and image analysis were done using Cell Counter and Angiogenic Analyzer plugin tools from the ImageJ program.

19 Chapter 3 – Results I

3.1 Expression of Ctcf in the developing heart. To uncover where Ctcf may be acting during heart development, I detected Ctcf protein in sections of E9.5 and 11.5 embryos, postnatal day (P) 2 and adult hearts by immunofluorescence (fig 3.1). This analysis revealed nuclear staining of Ctcf ubiquitously in cardiomyocytes and non-cardiomyocytes in heart sections. Cardiomyocytes were marked by staining for Cardiac Troponin T (CT3), a cardiac marker. Ctcf was detected throughout development and in adult hearts.

Figure 3.1. Ctcf is present in cardiomyocytes in the developing and adult heart. (A) Immunofluorescence of Ctcf (red), cardiac troponin T (CT3) (green) and DAPI (blue) in right ventricle (RV) heart sections at E9.5, E11.5, P2, and adult. Scale 50µm.

3.2. Conditional deletion of Ctcf in cardiac lineages. To uncover the requirement of Ctcf in heart development, I took advantage of a conditional Ctcf mouse allele. Cre-mediated recombination was used to inactivate Ctcf in cardiac progenitors, followed by observation of morphological defects in mutant progeny (fig 3.2A). The allele used to inactivate Ctcf integrated two loxP sites flanking exons 3 to 12, which encode the zinc fingers. To inactivate Ctcf in cardiac lineages, I used a

20 transgenic line expressing Cre driven by the Nkx2.5 promoter. Nkx2.5 is a marker of early cardiac progenitors expressed at E7.5. Cre-mediated deletion of Ctcf exons 3 to 12 will occur in Nkx2.5 expressing cardiac progenitors.

To confirm absence of Ctcf protein in mutant cardiac progenitors, I performed immunostaining for Ctcf and CT3 in mutant and control heart sections (fig 3.2B). Loss of Ctcf protein signal was detected predominately in mutant cardiomyocytes, while Ctcf protein signal persisted in mutant CT3 negative non-cardiomyocytes.

21 Figure 3.2. Inactivation of Ctcf in cardiac lineages. A. LoxP sites (orange) flank Ctcf exons 3 to 12. Nkx2.5-driven Cre excises loxP-flanked exons, in early cardiac progenitors. B. Immunostaining of Ctcf (red), and CT3 (green) in right ventricle heart sections. Nuclei were stained with DAPI (blue). Arrows indicate Ctcf and CT3 double positive cardiomyocytes. C. Quantification of Ctcf and CT3 double positive

22 cardiomyocytes in control (Ctcf fl/+) and mutant (Ctcffl/fl;Nkx2-5-cre). Bars represent the mean +/- S.D. of three biological replicates. Scale 50µm. * p<0.05.

3.3. Ctcf is essential in cardiomyocytes for embryogenesis and its deletion causes severe cardiac defects The ablation of Ctcf in early cardiac lineages causes embryonic lethality at E12.5. No viable mutant embryos were observed after this stage (fig 3.3). These Ctcf mutants have cardiac defects that arise at E11.5, becoming severe by E12.5, and consist of pericardial edema, a hypoplastic heart and hemorrhaging of the ventricles (fig 3.4A,B,G,F). To observe the internal cardiac structure of these mutants, I performed histological analysis. Ctcf mutants have thin ventricular walls with a decrease in the number of cells making up the ventricular walls (fig 3.4. D, E, F, J, K, L).

Figure 3.3. Ctcf in cardiomyocytes is essential for embryonic development. Ctcf fl/fl x Ctcf fl/+;Nkx2.5-Cre mice were crossed and viable embryos at different embryonic stages were collected. n is the number of embryos collected per stage

Figure 3.4. Ctcf deficient cardiomyocytes cause severe heart defects at E12.5. A, B, G, H. Lateral and frontal view of control and Ctcf-deficient embryos at E11.5 and E12.5. G. Ctcf mutants developed pericardial edema (arrow). H,I. Ctcf-deficient hearts were hypo-plastic and formed hemorrhages. C-F, I-L. H&E staining revealed a thinner ventricular wall (arrow) in Ctcf-deficient hearts. Bars represent the mean +/- S.D. of three biological replicates. A, B – 100µm, C-500µm, D-100µm. * p<0.05

24 3.4. Ctcf-deficient cardiomyocytes did not exhibit decreased proliferation or increased cell death. With the reasoning that the thin ventricular walls seen in Ctcf mutant hearts could be a result of defects in proliferation or cell death, cellular analyses on mutant hearts were performed. To assess whether an increase in cell death could result in a thin ventricular walls in mutant hearts, Ctcf control and mutant heart sections at E12.5 were stained for Caspase3 (Cas3), a cell death marker. CT3 was used to identify cardiomyocytes (fig 3.5A). Ctcf mutant hearts did not present increased numbers of Cas3 and CT3 double positive cells (fig 3.5C). It is plausible that the thin ventricular walls in Ctcf mutant hearts are a cause of cell proliferation defects. Ctcf control and mutant heart sections at E12.5 were stained for phosphorylated Histone H3 (pHH3), a proliferation marker, and CT3 (fig 3.5B). Ctcf mutant hearts did not present increased numbers of pHH3 and CT3 double positive cells (fig 3.5D). Thus, Ctcf is not required from proliferation of cardiomyocytes and its deficiency does not cause cardiomyocyte death at E12.5.

Figure 3.5. Ctcf mutants at E12.5 do not have cell death or proliferation defects. A, B. Immunofluorescence for activated caspase 3 (Cas3, green), cardiac troponin 3 (CT3, red), and phosphorylated histone H3 (pHH3, green) in sections of hearts at E12.5. Micrographs show portions of the right ventricle. C, D. Double positive Cas3/CT3, or pHH3/CT3 were quantified. Bars represent the mean +/- S.D. of three biological replicates. N.S. not significant. Scale 50µm.

25 3.5. Inactivation of Ctcf induces misregulation of well-characterized cardiac genes To understand the mechanistic involvement of Ctcf in cardiac development, my collaborators in Dr. Manzanares’ lab analyzed the genome wide expression profile of Ctcf control and mutant hearts using RNA sequencing (68). My collaborators were able to demonstrate that Ctcf mutant hearts at E10.5 had downregulation of cardiac genes, such as Hey2, Mef2c, Hand1, Nkx2.5, Irx4 (Fig 3.6A).

3.6. Ctcf deficiency restructures the chromatin interactions of the Irx4 regulatory domain To determine the involvement of Ctcf in mediating long-range chromatin interactions in cardiac development, the Manzanares’ Lab combined gene expression analysis with Circular Chromatin Conformation Capture (4C) data of E11.5 Ctcf control and mutant hearts. This technique measures all chromatin interactions occurring at a locus of interest (68). It was found that Ctcf deficiency did not induce changes in the chromatin organization of global topologically associated domains but acted on local regulatory domains. Restructuring of chromatin interactions was observed at the Irx4 promoter, part of the IrxA cluster, of Ctcf mutants (Fig 3.7). Decrease in chromatin interactions of Ctcf flanking regions from the Irx1, 2 and Ndufs6, the Irx4 neighbouring gene, to the Irx4 promoter was seen (Fig 3.7A). At the same time, increase in chromatin interactions was observed at the Irx4 promoter region, which normally lies outside of Irx4 regulatory domain (Fig 3.7B). This suggests that loss of Ctcf alters the long-range chromatin interaction involved in regulating the expression of the Irx4 domain by allowing abnormal enhancer activation.

Figure 3.6. Ctcf mutant hearts have downregulation of well-characterized cardiac genes. A. RNA-seq analysis of Ctcf wildtype, heterozygotes and mutant hearts was performed at E10.5. Heatmap displays downregulated genes in blue and upregulated genes in yellow. n=3.

27 Figure 3.7. Ctcf organizes the IrxA cluster into distinct regulatory domains. A. Chromatin interactions of the IrxA cluster using 4C-seq of E11.5 Ctcf control and mutant hearts. Red lines represent promoters of Irx1, Irx2, Irx4 and Ndufs6. Ctcf binding targets of 8-week heart was used as a reference. Arrowheads point to the position of enhancers that interact with Irx1, Irx2 but not Irx4 gene promoters. B. Close up view of yellow box in A. Asterisk and vertical lines denotes reduced interactions of Ctcf binding sites downstream of the Irx4 promoter. New Ctcf binding targets in green (i, ii), and loss of Ctcf binding targets in red (iii, iv).

28 Chapter 4 – Results II In the previous chapter, I discussed the requirement of Ctcf during heart development. In this chapter, I will discuss the requirement of Ctcf in endothelial progenitors during vascular development.

4.1 The expression of Ctcf in endothelial cells during vascular development To determine the expression of Ctcf in endothelial cells during vascular development, immunofluorescence for Ctcf protein and Platelet endothelial cell adhesion molecular1 (Pecam-1), an endothelial marker, was done on mouse sagittal sections of stages E9.5, E11.5, P2 and adult stages (fig 4.1). Ctcf was detected ubiquitously in nuclei and persisted in endothelial cell during vascular development.

Figure 4.1. Ctcf is present in mouse embryonic and adult endothelial cells. Immunofluorescence of Ctcf (red) and pecam-1 (green) in sagittal sections reveal Ctcf expression in embryonic and adult vasculature at E9.5, E11.5, P2 and adult. Scale - 50µm.

4.2. Conditional inactivation of Ctcf in endothelial progenitors To inactivate Ctcf in endothelial cells, I crossed Ctcffl/fl with Tie2-Cre mouse line, which expresses Cre in endothelial progenitors (fig 4.2A). To confirm the accurate deletion of Ctcf in endothelial progenitors cells, I analyzed the presence of Ctcf protein in vascular endothlelium from mutant and control vascular structures of embryos at E10.5 (fig 4.2B). In Ctcf mutants, Ctcf protein signal was lost specifically in Pecam-1 positive endothelial cells, however presence of Ctcf remained in non-endothelial cells. This indicates that Tie2-cre effectively deletes Ctcf in endothelial cells.

30 Figure 4.2. Inactivation of Ctcf in endothelial progenitors. A. LoxP sites (orange) flank Ctcf exons 3 to 12. Tie2 driven Cre excises loxp flanking exons and ablates Ctcf function in endothelial progenitors and its derivatives. B. Immunostaining for Ctcf (red), pecam-1 (green) and DAPI (blue) in vessels sections and quantification of Ctcf and pecam-1 double positive endothelial cells in Ctcf control and mutant. Scale 50µm.

31 4.3. Ctcf in endothelial progenitors is essential for embryogenesis The inactivation of Ctcf in endothelial progenitors is crucial for the development of the embryo (fig 4.3). Inactivation of Ctcf in endothelial progenitors caused embryonic lethality at E10.5. This suggests that the function of Ctcf in endothelium is required for embryogenesis. Ctcf heterozygotes, that have one functional copy of Ctcf, develop normally and are viable after birth. This indicates that one copy of Ctcf is sufficient for proper development, however inactivation of both copies disrupts embryogenesis.

Figure 4.3. Ctcf in endothelial cells is essential for embryonic development. Crossing Ctcf fl/fl x Ctcf fl/+;Tie2-Cre mice and collecting viable embryos at different embryonic stages. n is the number of embryos collected at each stage.

Phenotypic Characterization of Ctcf mutant vasculature The vasculature of the developing embryo encompasses the embryo proper and extraembryonic structures, consisting of the yolk sac and the placenta. In the sections below, I will describe my findings demonstrating the requirement of Ctcf in the development of the embryonic and extraembryonic vasculature.

32 4.4. Ctcf mutant embryos have narrow cerebral vasculature Although Ctcf deletion in endothelial progenitors caused embryonic lethality at E10.5, there were no gross morphological defects observed, aside from stunted embryonic growth (fig4.4A, 4.5A). Quantification of somites, a good indicator for embryonic stage, running along the trunk of E10.5 Ctcf mutant embryos revealed the mutants to be at an equivalent stage to the control. To analyze the embryonic vasculature, I compared different parts of the embryo, including the heart and the trunk, and found no vascular patterning or heart defects (fig 4.4B, C, D, E). I also examined the embryonic head and found no changes in the number of major cerebral vessel branches in Ctcf mutants in both E9.5 and E10.5 (fig 4.5 A, B, C). However, quantification of vessel diameter revealed that the cerebral vessels of the mutants in both the stages were narrow in comparison to the controls (fig 4.5 D).

Figure 4.4. Ctcf mutant embryos do not exhibit vascular defects in the trunk and heart at E10.5. A. Pecam-1 immunostaining of E10.5 control and mutant embryos. B. Close up immunostaining of the embryonic trunk. C. Histological sections of control and Ctcf mutant hearts. D. Close up histological sections of the heart. E. Quantification of vascular branching nodes on the embryonic trunk. OFT, outflow tract., RV, right ventricle., LV, left ventricle. Bars represent the mean +/- S.D. of three biological replicates. N.S. not significant. Scale bar A. 100µm, 50µm., B. 1000µm, 100µm.

Figure 4.5. Ctcf mutants have narrow cerebral vasculature. (A) Whole mount of control (Ctcffl/fl) and mutant (Ctcf fl/fl;Tie2-cre) embryos. Close up embryonic head view stained by immunofluorescence for Pecam-1 (green). (B) Quantification of the major cerebral vessels. (C) Quantification of vascular diameter of major cerebral vessels. Bars represent the mean +/- S.D. of three biological replicates. *p <0.05. Scale (A) 200um, (B) 500um.

4.5. Ctcf mutants have yolk sac vascular remodelling defects The yolk sac is the first site for vasculogenesis and angiogenesis in the embryo. Defects in the yolk sac vasculature cause stunted growth in various mutant models (32, 69, 70, 71). I examined the yolk sac vasculature by immunofluorescence for Pecam-1 in control and Ctcf mutant embryos at E8.5, 9.5 and 10.5 (fig4.6A). I used fluorescent images of the vascular network to create binary tree images to quantify vascular branching by counting the number of vascular nodes using the ImageJ angiogenic

34 analyzer program (fig 4.6B). I also quantified the thickness of blood vessels by measuring diameter (fig. 4.6B). I found that Ctcf mutant yolk sacs had vascular remodelling defects as revealed by the decreased vascular branching and a disorganized vascular network. Ctcf mutant yolk sacs at E8.5 had no changes in vascular branching or thickness than compared to the control (fig 4.6.C, D). At E9.5, mutant yolk sacs had a decrease in vascular branching (fig 4.6 C, D). By E10.5, Ctcf mutants had pale discoloration of the yolk sac, where the vascular branching defects were compensated and were met with a decrease in vascular thickness (Fig 4.6 A, C, D). This indicates that yolk sac remodelling defects in Ctcf mutants initiate at E9.5 and could possibly impair vascular development and as a result cause stunted embryonic growth.

4.5.1 Ctcf deficiency does not affect cell proliferation or cell death in yolk vascular endothelium. It could be hypothesized that the vascular remodelling defects observed in the Ctcf mutant yolk sac are a result of increased endothelial progenitor cell death or by reduced cell proliferation. To address this question, I co-stained E9.5 yolk sac sections for Pecam-1 and caspase 3 to analyze cell death (fig 4.7A). No change in endothelial cell death was observed in Pecam-1 positive Ctcf mutant yolk sacs when compared to control (fig 4.7C). To determine if Ctcf deficiency causes endothelial cell proliferation defects, I co-stained yolk sacs for Pecam-1 and phosphorylated histone H3 (fig 4.7B). Similarly, no change in endothelial cell proliferation was observed in Pecam-1 positive Ctcf mutant yolk sacs when compared to control (fig 4.7C). This suggests that Ctcf is not required for survival or maintenance of endothelial cells in the yolk sac.

Figure 4.6. Endothelial specific Ctcf mutants have yolk sac vascular remodelling defects. A. Time lapse of yolk sac vascular development. Whole mount embryo with yolksac and placenta attached at E8.5, 9.5, 10.5 of control (Ctcf fl/fl+, Ctcffl/+:Tie2-cre) and mutant (Ctcf fl/fl;Tie2cre). B. Close up of yolk sacs stained by immunofluorescence for pecam-1 and prospective Binary tree images of yolk sacs generated with ImageJ for quantification. C, D. ImageJ angiogenic analyzer was used to quantify the number of vascular nodes and vessel diameter. Bars represent the mean +/- S.D. of three biological replicates. * p<0.05. Scale 1000µm, 100µm.

Figure 4.7. Endothelial specific Ctcf mutant yolk sacs showed no change in cell death or proliferation at E9.5. (A,B) Immunofluorescence of endothelial cells co-expressing Cas3 and Pecam-1, and pHH3 and Pecam1. (C.) Quantification of Cas3 and pHH3/ pecam-1 double positive endothelial cells. Bars represent the mean +/- S.D. of three biological replicates. N.S. not significant. Scale -100um.

4.5.2. Yolk sac vascular defects and hemodynamic force The discoloration observed in E10.5 Ctcf mutant yolk sac may be an indication of a hematopoietic defect. It is plausible that the yolk sac vascular defects in Ctcf mutants could be secondary to reduced hemodynamic force. It has been shown that hemodynamic force induced by the flow of blood from the initiation of the heartbeat and blood circulation around E8.5, can cause a dragging force on vascular endothelial cells, triggering major vascular remodelling pathways (18). A good indicator of reduced hemodynamic force is downregulation of endothelial nitric oxide synthetase, eNOS, which encodes a mechanosensitive protein (18). Downregulation of eNOS is observed when blood circulation is prevented upon impaired yolk sac vascular remodelling (18).

37 As a first approach to determine if hemodynamic force changes could contribute to the impaired vascular remodelling seen in Ctcf mutant yolk sacs, I performed a qPCR to analyze the levels of eNOS in yolk sacs at E9.5, the first stage at which vascular branching defects are seen (fig 4.8). Although, there was a trend for decreased expression of eNOS in E9.5 Ctcf mutant yolk sacs, the decrease was not statistically significant. In addition, embryonic red blood cells were present in extraembryonic tissues, yolk sac and placenta, at E9.5. Thus, it is possible that vascular defect in the yolk sac at E9.5 may not be secondary to reduced hemodynamic force. However, this needs to be further investigated.

Figure 4.8. Ctcf mutant yolk sac has no significant change in eNOS levels in E9.5. qPCR of eNOS relative to housekeeping gene Pgk1. Bars represent the mean +/- S.D. of three biological replicates.

4.6. Ctcf mutant placentas have a reduction in vascular labyrinth area Stunted embryonic growth can be an indicator for placental defects, which may cause a deficiency in nutrient and gas exchange between the mother and the fetus (28, 72, 73). I examined Ctcf mutant placentas at E9.5 and E10.5 (fig 4.9A). Aside from the pale discoloration of Ctcf mutant placentas at E10.5, no other gross morphological defects were seen. Analysis of the internal placental structures in histological sections uncovered increased sinuses containing enucleated maternal blood cells in Ctcf (fig 4.9B). To examine the fetal vasculature in the placental labyrinth, I tool advantage of a Cre reporter

38 mouse line, Ctcf fl/fl; Rosa mT/mG, which drives expression of GFP in endothelial cells where Cre is activated, once crossed with Ctcf fl/fl;Tie2-Cre. I stained placental cross sections by immunofluorescence for GFP and quantified the fluorescence area by ImageJ (fig 4.9C). I found that Ctcf mutants at E9.5 had a trend towards reduced fetal vascular labyrinth area compared to controls (fig 4.9D). By E10.5, these mutants had a significant decrease in the fetal vascular area (fig 4.9D). The reduction in the fetal placental vascular area may adversely alter the maternal to fetal nutrient and gas exchange and result in impaired embryonic growth.

4.6.1 The fetal vasculature of Ctcf mutant placentas do not exhibit cell death or proliferation defects. To address whether the reduced fetal vasculature in Ctcf mutant placentas is caused by increased cell death or decreased proliferation defects, I co-stained by immunofluorescence for Cas3 and pHH3 with pecam-1 on placental sections at E9.5 and E10.5, respectively (fig 4.10). Quantification of both Cas3 to pecam-1 and pHH3 to pecam-1 double positive endothelial cells revealed no changes in cell death or proliferation in Ctcf mutant placentas in comparison to control placentas (fig 4.10). This indicates that Ctcf in endothelial progenitor cells in the fetal vasculature of the placenta is not required for cell survival or maintenance.

4.6.2. Ctcf mutant placentas have misregulation of various vascular regulatory genes As a first approach to uncover Ctcf targets in endothelial cells, I analyzed the gene expression pattern of several vascular regulators using E9.5 placentas (4.11). I found the upregulation of the following vascular regulatory genes: Kdr (VEGFR2, vascular endothelial growth factor 2), Vegfa (vascular endothelial growth factors, a ligand for kdr) and Erg (ETS-related gene) (fig 4.11). This indicates the possibility that Ctcf may target these endothelial vascular regulators during development of the placental vasculature.

39 Figure 4.9. Ctcf mutant placental labyrinth have a reduction in vascular area at E10.5. A. Whole mount of Ctcf control (Ctcf fl/fl) and mutant (Ctcf fl/fl;Tie2-cre) placentas at E9.5 and E10.5. B. Close up of histological sections of the placental labyrinth. C. Immunofluorescence of placental sections staining for GFP using the RosamT/mG Cre reporter line in control (Ctcf fl/+;Tie2-cre; Rosa mT/mG) and mutant (Ctcf fl/fl;Tie2-cre; Rosa mT/mG). D. Quantification was done by measuring fluorescence area using the ImageJ program. Bars represent the mean +/- S.D. of four-six biological replicates. *p<0.05. Scale bar A; 100µm , B, 50µm, C; 200µm. PL, placenta, UM, umbilical cord, EM, embryo, FB, fetal blood, MB, maternal blood.

40 A.

C. D.

Figure 4.10. Endothelial-specific Ctcf mutant placentas have no cell death or proliferation defects. Immunofluorescence of endothelial cells co-expressing Cas3 and GFP, and pHH3 and GFP at both E9.5 and E10.5. Bars represent the mean +/- S.D. of three biological replicates. Scale -100um

41 Figure 4.11. Endothelial specific Ctcf mutant placentas at E9.5 have an increase in expression of vascular regulatory genes. qPCR of vascular regulators genes, Kdr, Vegfa and Erg. Data are presented as expression relative to Pgk1. Bars represent the mean +/- S.D. of three biological replicates. *P<0.05.

42 Chapter 5 – Conclusion In my thesis, I hypothesized that Ctcf is required for mouse cardiovascular development. To validate my hypothesis, I developed two main aims in my thesis: one, to elucidate the requirement of Ctcf in heart development and two, to elucidate the requirement of Ctcf in vascular development. My results demonstrate that Ctcf is an essential factor for the development of the mouse cardiovascular system.

In this chapter, I will first discuss my results demonstrating that Ctcf is required for cardiac development and second, the requirement of Ctcf in vascular development. Finally, I will discuss future directions to uncover the mechanistic basis for Ctcf in endothelial cells during vascular development.

5.1 Discussion 5.1.1 Ctcf is essential for heart development Ctcf is ubiquitously present in the nuclei of cardiomyocytes throughout development and into adulthood. I found that conditional deletion of Ctcf in cardiac progenitors resulted in predominate loss of Ctcf in the myocardium. Ctcf null embryos are inviable at E12.5, had severe cardiac defects including a small heart, pericardial edema and thin, leaky ventricular walls. This indicates that Ctcf is required to the development of the heart. Heterozygote Ctcf embryos appeared normal and live past E12.5. However, it is possible that proper Ctcf dosage is required for later stages of development or for cardiac maturation after birth. Examining these heterozygotes at later stages will be necessary to assess this possibility. One such example of haploinsufficiency causing cardiac defects is seen with the Nkx2.5, homeobox transcription factor (74). Heterozygous mutation of Nkx2.5 causes atrial septal defects at postnatal stages (74).

To elucidate the mechanistic basis of Ctcf in the transcriptional regulation of cardiac development, I collaborated with Dr. Manzanares’ Lab. First, they found misregulation of various important cardiac regulators, such as Nkx2.5, Irx4, Hand1, and Mef2c, in Ctcf mutant hearts by analyzing the gene expression profile. This suggests that Ctcf in differentiating cardiomyocytes may regulate key cardiac transcriptional programs.

43 Second, the Manzanares’ Lab found that Ctcf deficiency induced disorganization of the Ctcf mediated long-range chromatin interactions at the Irx 4 promoter. Irx4 has a ventricular specific expression and is targeted by the cardiac transcription factors Nkx2.5 and dHand (75). It is uncertain whether the disorganization of Ctcf-mediated long-range chromatin interactions in the Irx4 locus is the primary cause of the cardiac defects observed in E12.5 Ctcf null embryonic hearts. Irx4 null mutant mice are normal in embryonic and postnatal stages of life, suggesting that other Irx genes compensate for Irx4 deficiency (76). Thus the regulatory function of Ctcf over Irx4 does not completely explain the mutant cardiac phenotype seen. One could speculate the potential for the other misregulated cardiac genes to be direct Ctcf targets. However, further investigation needs to be done to comprehend the mechanistic basis of Ctcf in cardiac development.

5.1.2 Ctcf is essential for vascular development In the second aim of my thesis, I investigated the requirement of Ctcf in vascular development. First, I found that Ctcf is present in the nuclei of endothelial cells during vascular development and persisted into adulthood. I then conditionally inactivated Ctcf, using a Cre recombinase driven by the Tie2 promoter, expressed in endothelial progenitors. As endothelial progenitors differentiate into endothelial and hematopoietic cells, deletion of Ctcf is exhibited in these two populations. Unfortunately, no early markers expressed specifically in endothelial cells are known (77). Ctcf mutants are embryonic lethal at E10.5, demonstrating the necessity of Ctcf in embryogenesis. Ctcf heterozygotes live normally in embryonic and postnatal stages. Aside from the stunted growth at E10.5, no gross morphological defects were observed. Ctcf null embryos had no vascular defects in the trunk, or any obvious heart defects. At a closer look of the embryonic head of Ctcf mutant embryos, I found that they had narrow cranial vasculature. Previous studies have shown that stunted embryonic growth of vascular mutants may be an indicator of vascular defects in extraembryonic tissues, such as the yolk sac and the placenta (28, 32, 69-73).

Next, I examined Ctcf mutant yolk sacs for vascular defects. Ctcf mutant yolk sacs at E9.5 had vascular remodelling defects in which they failed to establish a hierarchical vascular network. This defect persisted in E10.5. Deficiencies in a number of

44 regulators important for vascular development are known to cause yolk sac vascular remodelling defects (78-80). This suggests that Ctcf may target genes involved in yolk sac vascular remodelling.

Ctcf is also inactivated in hematopoietic cells. Ctcf mutants at E10.5 had pale discolored yolk sacs, an indicator for hematopoietic defects. It is known that reduced numbers of hematopoietic cells change the hemodynamic force in the blood vessels. A decrease in hemodynamic force can result in yolk sac vascular remodelling defects. As Ctcf is known to regulate hematopoietic development (49-51, 59, 63), the potential that the Ctcf mutant yolk sac vascular remodelling defects are secondary to reduced hemodynamic force is possible. As a first step to address this issue, I analyzed the expression of endothelial nitric oxide synthetase (eNOS), which is known to be downregulated in mutants in which hemodynamic force is altered by lack of circulation. I analyzed the expression of eNOS in E9.5 yolk sacs, where vascular remodelling defect were first observed. I found that aside from a trend towards a decrease in eNOS levels in the yolk sac, it was not statistical significant. At E9.5, the extraembryonic tissues had presence of blood and could suggest that the vascular remodelling defect at this stage may not be secondary to reduced hemodynamic force. Further analysis of the affect of hemodynamic forces in vascular remodelling in Ctcf mutants needs to be done to resolve this issue. Potential experiments to examine this issue could be to use live imaging techniques to quantify force of blood flow in the yolk sac or by manipulating blood flow to examine if Ctcf mutant yolk sacs still undergo vascular remodelling defects (27).

Placental defects are also known to cause embryonic growth retardation, as they perturb the maternal and fetal nutrient and gas exchange. Ctcf mutant placentas revealed to have a decreased labyrinth expansion and enlarged pockets of maternal blood sinuses at E10.5. It is possible that crosstalk between the maternal and fetal vasculature occur in the labyrinth via regulatory glycoproteins called cytokines (81). Cytokines can be produced and released by endothelial cells to activate receptors and regulate functions of various cells (82, 83). It could be speculated that deficiency of Ctcf in the fetal vasculature perturbs such cytokine-mediated cross talk between the maternal and fetal vasculature and impairs their respective development.

45 As a first approach to identify direct targets of Ctcf in endothelial cells in the placenta, I examined the expression of several regulators of vascular development in E9.5 placentas. I found upregulation of Kdr, VegfA and Erg genes. These are key vascular regulatory genes involved in vasculogenesis and angiogenesis (84, 85). Kdr or VEGF receptor 2 and VegfA are important in the VEGF signalling pathway (84, 85). The VegfA ligand is usually expressed in the epithelium, where Kdr is known to attract VegfA via chemotaxis to promote vascular development (32, 84, 85). Deficiency in Kdr causes embryonic lethality at E9.5, a lack of blood islands, and reduced endothelial and hematopoietic cells (84). On the other hand, haploinsufficiency of VegfA causes yolk sac blood vessel and aorta defects (85). Erg, an ETS related gene, is a transcription factor of the ETS family regulating the Wnt/β-Catenin signalling pathway and required for stability and growth of blood vessels (86). It possible that these three upregulated vascular regulatory genes are direct targets of Ctcf, however further ChIP analysis needs to be done to identify direct targets of Ctcf in endothelial cells in the developing vasculature.

5.2. Future Directions 5.2.1 Elucidate the gene expression profile of Ctcf-deficient endothelial cells I have made progress towards uncovering the global gene expression profile of Ctcf-deficient endothelial cells by performing RNA-sequencing on endothelial cells isolated from Ctcf control and mutant extraembryonic tissues, such as yolk sac and placenta, at E9.5. Taking advantage of the RosamT/mG Cre reporter mouse line, Ctcf fl/fl; RosamT/mG crossed to Ctcf fl/+ ;Tie2-cre, endothelial progenitors have been isolated via fluorescence activated cell sorting (FACs). Approximately 1000-2000 cells are sufficient to isolate RNA and create RNA libraries. From our lab experience, we have previously isolated roughly 5000 to 15,000 endothelial cells from E9.5 extraembryonic tissues. Proper validation of RNA quality via Bioanalyzer and confirmation of Ctcf control and mutant endothelial progenitors by endothelial gene expression analysis has been done. RNA libraries have been made and then sent off for sequencing in The Centre for Applied Genomics (TCAG) at The Hospital for Sick Children. Analysis of the global gene expression data set will be performed in collaboration with Dr. Michael Wilson’s

46 lab (SickKids). Investigating the global expression profile of Ctcf control and mutant endothelial progenitors will help determine potential Ctcf targets in vascular development.

5.2.2 Analyze the genome-wide distribution of Ctcf in developing endothelial cells To identify direct Ctcf targets, the global gene expression profile will be coupled to the analysis of direct Ctcf binding sites uncovered by Chromatin Immunoprecipitation followed by high throughput sequencing (ChIP-seq). Using RosamT/mG transgenic mice crossed to Tie2-cre mice, control endothelial progenitors at E9.5 will be isolated using FACs. A minimum of 100,000 cells is needed to perform ChIP-sequencing. This will be achieved by pooling together endothelial cells from multiple extraembryonic tissues. The analysis of both RNA-seq and ChIP-seq datasets will allow for mapping direct Ctcf targets and identify the transcriptional programs controlled by Ctcf during vascular development.

5.2.3 Determine Ctcf mediated long-range chromatin interactions in developing endothelial cells To determine how Ctcf-mediated long-range chromatin interactions regulate transcriptional programs required for vascular development; chromosome conformation capture coupled with high throughput sequencing (4C, circular chromatin conformation capture) will be performed. The 4C assay detects all the chromatin loops and long-range interactions that occur at one locus of interest (87). This assay will demonstrate all the Ctcf-mediated long-range interactions that occur at the gene of interest and help determine how Ctcf mediated chromatin organization regulate the transcriptional programs required for vascular development. Approximately 10 million cells will need to be isolated to perform 4C-seq (87). Although, the isolation of 10 million endothelial cells from E9.5 extraembryonic tissues may be problematic, pooling of extraembryonic tissues and digestion of the whole tissue can help to circumvent the challenge. Additionally, ex vivo explants of extraembryonic tissues can be cultured to expand the endothelial population (88), and obtain sufficient material from Ctcf control and mutants for 4C-seq. This analysis in combination with global gene expression data and ChIP-seq data will identify Ctcf targets and uncover Ctcf-mediated long range-chromatin interactions in endothelial cells.

47 5.2.4 Validate direct targets of Ctcf in vascular development Once Ctcf target genes are found; their function will be validated in in vitro and in vivo assays. The expression of Ctcf target genes will be analyzed by Western blot and qPCR in Ctcf mutant endothelial cells sorted from Ctcffl/fl;Tie2-Cre; RosamT/mG extraembyronic tissues. Using in vitro assays, rescue experiments on Ctcf mutant endothelial cultures will be done to survey the downstream targets. Overexpression of downstream targets can be performed by transfection of expression vectors in endothelial cells, while knock down of downstream genes can be accomplished by transfecting endothelial cells with siRNAs. Genes that rescue vascular remodelling and expansion of Ctcf mutant cells will be candidates to be validated for in vivo systems. Given the opportunity where mutant mice lines with the downstream gene of interest is available, they will be crossed with Ctcf fl/fl and Ctcf fl/+; Tie2-cre mice, to obtain double heterozygotes or double mutants. These progenies will be assessed for developmental defects. About three litters of double mutants will be examined during the following developmental time points: E8.5-14.5, E18.5, P0 and weaning age. Full or partial rescue of double mutant embryos are considered to survive past E10.5. The extraembryonic tissues of rescued embryos will be examined by immunofluorescence and histology. The expression of Ctcf downstream targets will also be analyzed by qPCR and in situ hybridization. These analyses will help solidify the direct downstream targets of Ctcf and provide further insight into how Ctcf mediated transcriptional regulation in endothelial cells is involved in vascular development.

5.3. Conclusion My research has demonstrated that Ctcf is essential for mouse cardiovascular development. In my thesis, I demonstrated that deficiency of Ctcf in cardiac progenitors caused severe cardiac defects and embryonic lethality at E12.5, while deficiency of Ctcf in endothelial progenitors caused extraembryonic vascular defects and embryonic lethality at E10.5. Further investigations will uncover the mechanisms by which Ctcf involved in long-range chromatin interactions help guide vascular development and function. Uncovering the mechanisms controlling cardiovascular development may also help us to understand the origins of cardiovascular disease.

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