Regulation of Activation Phase of Angiogenesis by Trancription Factors Ets1 and Ets2

REGULATION OF ACTIVATION PHASE OF ANGIOGENESIS BY TRANCRIPTION FACTORS ETS1 AND ETS2

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of the Ohio State University

By Sankha Ghosh, M.Sc.

Graduate Program in Molecular, Cellular and Developmental Biology The Ohio State University 2014

Dissertation Committee:

Dr.Michael C. Ostrowski, Advisor

Dr.Jeffrey Parvin

Dr.Denis Guttridge

Dr. Qianben Wang

Sankha Ghosh

2014

ABSTRACT

Vascular remodeling is a necessary process not only for embryonic development but also for specific physiological and pathological conditions in adult. Several growth factors are required at specific stages in order for the vascular development to occur. Angiogenesis also involves complex crosstalk between several endothelial cell (EC) processes including cell cycle, cell survival and migration. Signaling pathways, stimulated by the growth factors and communication between endothelial cells and extra cellular matrix

(ECM), coordinate these processes. Deregulation of any of the factors or pathways can lead to severe defects in vessel formation.

Transcription factors ETS1 and ETS2 are required for EC functions necessary for embryonic angiogenesis. EC specific deletion of ETS2 with conventional deletion of

ETS1 results in defective vascular branching. Additionally, the double mutant embryos are embryonic lethal. Owing to this lethal nature, the specific gene targets of these factors are yet to be identified. In the current study, we elucidate the effect of endothelial cell specific deletion of Ets1 and Ets2 on angiogenesis and characterize the downstream regulatory pathways.

Inducible Cre-loxp technology was used to specifically delete both Ets1 and Ets2 in endothelial cells after birth. Deletion of Ets1 and Ets2, when restricted to endothelial cells in new born mice (P1-P3), reduced retinal angiogenesis. Similarly, EC infiltration and invasion into matrigel plugs subsided when matrigel admixed with mouse mammary tumor cells was injected into adult mice with inactivated Ets1 and Ets2 specifically in

ECs. Gene expression array performed on RNA, isolated from cultured aortic endothelial cells, comparing the double knockout cells with controls revealed reduced expression of key cell cycle and cell survival regulators in double mutant cells. In addition, both these factors were found to occupy the enhancer regions of the target genes indicating these factors directly regulate expression of the target genes. Recruitment of coactivators

CBP/p300 was diminished in the absence of ETS1 and ETS2. Deletion of Ets1 and Ets2 in cultured aortic EC resulted in altered cell cycle phases with a G2/M phase arrest and increased sensitivity to apoptosis in vitro.

These results demonstrate that deletion of Ets1 and Ets2 in endothelial cells inhibits angiogenesis by blocking cell cycle progression and decreasing cell survival.

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DEDICATION

This work is dedicated to my parents and my brother without their blessings and love I could

not have accomplished this work. I also dedicate this work to my wife Tapahsama for her

continuous support and encouragement through these years.

ACKNOWLEDGMENTS

I express my heartfelt gratitude to my advisor Dr. Michael C. Ostrowski, for his invaluable support, continuous guidance and especially his patience throughout this research work. His useful suggestion, moral support and encouragement to think independently enabled me to complete my work successfully.

I am extremely grateful to Dr. Jeff Parvin, Dr. Dennis Guttridge, and Dr. Qianben Wang for their time, support and useful advice throughout this study.

I thank all past and present members of Ostrowski Lab. I am extremely thankful to Dr.

Haritha Mathsyaraja for being such a friend that she is. I thank Dr. Sudarshana Sharma for his extremely valuable intellectual inputs. I am grateful to Subhasree Balakrishnan for making my time in lab enjoyable. Many thanks go to Heaher Carrey, Jennifer Cabrera

Anisha Mathur and Xin Liu for being such wonderful lab mates.

I would also like to thank Dr. Lianbo Yu, Dr. Dias Kurmashev for their help with bioinformatics. I also thank the CCC Shared Resources and Cores for their essential role in this work.

Last but not the least I thank my family, my wife Tapahsama and my wonderful group of friends here in Columbus for making graduate school and life in Columbus wonderful experience.

VITA

30 November 1980…………………………………… Born – Kolkata, India

May 2005……………………………………………….B.Sc., Zoology University of Calcutta Kolkata, India July 2007………………………………………………..M.Sc., Zoology University of Calcutta Kolkata, India

September 2007 – December 2009……………………..Teaching Assistant April 2010 – August 2014 The Ohio State University.

January 2010 – March 2010…………………………….Research Assistant The Ohio State University.

FIELDS OF STUDY Major Field: Molecular, Cellular and Developmental Biology

TABLE OF CONTENTS

Page

ABSTRACT…………………………………………………………………. ii

DEDICATION………………………………………………………………. iv

ACKNOWLEDGEMENTS………………………………………………… v

VITA…………………………………………………………………………. vi

LIST OF TABLES………………………………………………………….. xi

LIST OF FIGURES………………………………………………………… xiii

LIST OF ABBREVIATIONS………………………………………………. xiii

CHAPTER 1 INTRODUCTION…………………………………………... 1

1.1. Development of Vascular Network…………………………………. 3 1.1.1. Evolution of Circulatory Network…………….……………… 3 1.1.2. Vasculogenesis………………………………………...……… 7 1.1.2.1. Mechanism of Vasculogenesis……….……...………... 7 1.1.2.2. Regulation of Vasculogenesis…..…….……...……….. 10 1.1.3. Angiogenesis……………………………………………...... 13 1.1.3.1. Phases of Angiogenesis…………….....….…………… 13 1.1.3.2. Mediators of Vessel Branching.………………....…..... 21 1.1.3.3. Cellular Processes Involved in Angiogenesis….....…... 33 1.1.3.4. Angiogenesis in Development and Disease……...... 37 1.2. Ets Family of Transcription Factors...………………………………. 39 1.2.1. The ETS Domain………………………..…..………………….. 43 1.2.2. The PNT Domain……...….…………………………………… 44

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1.2.3. The Biological Role of Ets Factors…………………………… 47 1.2.3.1. Ets Factors in Embryonic Development...…...………... 47 1.2.3.2. Ets Proteins in Cancer………………...……...……….. 49

CHAPTER 2 MATERIALS AND METHODS…………………………... 51 2.1. Animal Husbandry………………………………………………….. 51 2.1.1. Transgenic Mice used……………………………....…………. 51 2.1.2. Animal Care…………………………………………………… 52 2.1.3. Mouse Genotyping……………………………………………. 52 2.1.3.1. Tail DNA Preparation………………………………… 52 2.1.3.2. Genotyping Primers and PCR Conditions……...... 52 2.1.4. Animal Procedures……………………………………………. 54 2.1.4.1. Induction of Gene Deletion ………………….……….. 54 2.1.4.2. Eye and Retina Dissection…..………………………… 54 2.1.4.3. Subcutaneous matrigel plug injection………………… 55 2.2. Tissue Culture and Viral Infections…………………………………. 55 2.2.1. Primary Cell Extraction and Culture………………………….. 55 2.2.1.1. Aortic Endothelial Cells………………………...... 55 2.2.2. Lentiviral Infections…………………………………………… 56 2.3. Microarray and Data Analysis……..………………………………… 56 2.4. cDNA Preparation…………………………………………………... 57 2.4.1. RNA Extraction……………………………………………….. 57 2.4.2. Reverse Extraction……………………………………………. 57 2.4.3. Primers Used for real-time PCR………………………...... 58 2.5. Chromatin Immunoprecipitation Assay (ChIP)……………………... 60 2.5.1. Chromatin Preparation………………………………………... 60 2.5.2. Primers Used for real-time PCR………………………...... 60 2.5.3 ChIP-Sequencing and Analysis………………………………... 61 2.6. Quantitative Real Time PCR (qRT-PCR)…………………………… 63 2.6.1. qRT-PCR Reaction Conditions……………………………….. 63 2.6.2. qRT-PCR Analysis……………………………………………. 63 2.7. Western Blot Analysis………………………………………………. 63 2.7.1. Protein Isolation………………………………………………. 63 2.7.2. Western Blot…………………………………………………… 64 2.8. Cell Based Assays…………………………………………………... 64 2.8.1. BrdU Proliferation Assay…………………………………….. 64 2.8.2. Apoptosis Assay………………………………………………. 64 2.8.3. Migration Assays……………………………………………… 65 2.8.3.1. Scratch Wound/Wound Healing Assay………………. 65 2.8.3.2. Single Cell Migration Track Assay……………...... 65 2.8.3.3. Matrigel Tube Formation Assay………………...... 65 2.9. Histology and Immunostaining..……………………………………. 66

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2.10. Imaging and Quantification…………………………………….….. 66

CHAPTER 3 Ets1AND Ets2 IN ACTIVATION PHASE OF ANGIOGENESIS…………………………………………… 67 3.1. Introduction……………...………………………………………….. 67 3.1.1. Ets Factors in Angiogenesis……………..………....…………. 69 3.1.2. Retinal Angiogenesis in Mice ……………….…....…………. 69 3.1.3. Trangenic Alleles Used…………………...………………..…. 73 3.2. Results…………………...………………………………………….. 77 3.2.1. Tie-2-Cre efficiently recombines Ets1/2fl alleles specifically in EC…………………………………………………………….. EC.. 77 3.2.2. Tie-2-Cre; Ets1fl/fl;Ets2fl/fl display defective retinal angiogenesis 79 3…..2.3. Tie-2-Cre; Ets1fl/fl;Ets2fl/fl demonstrate reduced tumor angiogenesis…………………………………………………... 85 3.2.4. Lentiviral-Cre efficiently deletes Ets1/2 in vitro 90 3………….…..…..2.5. Ets1 and Ets2 regulate endothelial cell cycle, cell survival and migration……………………………………………………... 92 3.2.5.1. EC Lacking Ets1 and Ets2 Undergo Increased Apoptosis 92 ……………….3.2.5.2. Loss Of Ets1 And Ets2 Leads To G2/M Phase Arrest... 93 3.2.5.3. EC Specific Ets1/Ets2 Ablation Results in Reduced Migration and Tube Formation In Vitro…….. 97 3.2.5.4. Deletion of Ets1/Ets2 Leads to Increased EC Adhesion. 97 3.3. Discussion.……………....………………………………………….. 102

CHAPTER 4 EC SPECIFIC REGULATORY NETWORK OF ETS1/2. 107 4.1. Introduction………...……………………………………………….. 107 4.1.1. Biological Role of ETS1 and ETS2 as Transcription Factors.... 108 4.1.2……………………………. Coactivators of Ets Factors …………...... …………………….. 109 4.2. Results……………...……………………………………………….. 111 4.2.1. Specific EC processes required for angiogenesis are disrupted in the absence of Ets1 and Ets2……………...…… 111 4.2.2. Loss of Ets1/Ets2 affects expression of genes involved EC cell cycle, cell survival and migration …………………… 114 4.2.3. ETS1 and ETS2 directly regulate specific cell cycle, anti apoptosis and migration genes…………………………… 116 4.2.4. ETS1 and ETS2 recruit coactivators CBP/p300 …………….. 123 4.3. synergistically..…...Discussion.……………....………………………………………….. …………………………… 125

CHAPTER 5 CONCLUSIONS AND FUTURE DIRECTION…..…….... 128 5.1. Conclusions ………………………………………………………… 128 5.2. Future Directions……...……………………………………………. 131 5.2.1. Effect of ETS1/ETS2 on EC processes in vivo..……………… 131 5.2.2. Mechanism of G2/M phase cell cycle arrest……..…….....…... 132 5.2.3. Post transcriptional control of EC specific processes by ETS1/ETS2………………………………………………… 133

BIBLIOGRAPHY...... 136

LIST OF TABLES

Table 2.1. Primers Used for Genotyping Mice and Cells……..……..…...... 53 Table 2.2. UPL Primers Used for cDNA Quantitative Realtime-PCR (qRT- PCR)……………………………………………………….. 59 Table 2.3. UPL Primers Used for ChIP………………...…………………….. 62 Table 3.1. Breeding Strategy and The Genotypes of The Control and Experimental Mice Used for Retinal Angiogenesis Assay……….. 84 Table 3.2. Genotypes of The Control and Experimental Mice Used for Matrigel Plug Assay….…………………………………. 89 Table 4.1. List of Motifs Identified by Analyzing The Genomic Regions Occupied by Both ETS1 and ETS2………...……………………... 122

LIST OF FIGURES

Figure 1.1. Evolution of Circulatory System……………………………..…... 4 Figure 1.2. Vertebrate Circulatory Systems ………………………………….. 6 Figure 1.3. Schematic of Vasculogenesis ……………...…………………….. 9 Figure 1.4. Regulation of Developmental Vasculogenesis…...……………….. 12 Figure 1.5. Two Phases of Angiogenesis ….……………………………..…... 14 Figure 1.6. Molecular Mechanism of Activation phase….………………..…... 17 Figure 1.7. Resoltuion Phase of Angiogenesis…………...……………………. 20 Figure 1.8. Molecular Regulators of Angiogenesis.……...……………………. 31 Figure 1.9. VEGF-VEGFR-2 Signaling Cascade During Vascular Development………………………………………………………Development 32 Figure 1.10. .. Domains……………… of Ets Family Transcription Factors….………………… 41 Figure 1.11. Phylogeny Of Ets Family Proteins…………………..…………….. 42 Figure 1.12. Structure of ETS and PNT Domain …………….…..…………….. 46 Figure 3.1. Development of the superficial vascular plexus in mouse retinas... 72 Figure 3.2. Targeting strategy for Ets1fl allele……………………..………….. 76 Figure 3.3. Cre Recombinase Efficiently Deletes Ets1fl and Ets2fl alleles in vivo………………………………………….. 78 Figure 3.4. Conditional Deletion of Ets1 and Ets2 Attenuates Murine Retinal angiogenesis……………………………………... 81 Figure 3.5. Ets1 and Ets2 are required for Tumor Angiogenesis Progression… 86 Figure 3.6. Cre-recombinase Mediated in vitro deletion of Ets2fl……………. 91 Figure 3.7. Deletion of Ets1 and Ets2 Affects EC Survival and Cell Cycle….. 94 Figure 3.8. Deletionmigration of in Ets1 vitro and Ets2 Affects EC Migration and Adhesion In Vitro……………………………………………………………. 099 Figure 4.1. Biological Processes Essential for Angiogenesis are Disrupted in Ets1/Ets2 Double Mutant cells…………………………….…… 112 Figure 4.2. Deletion Of Ets1 And Ets2 Results In Deregulation Of Genes Genes Involved In Cell Cycle, Cell Survival And Migration...... 115 Figure 4.3. ETS1 and Ets2 are enriched the Enhancer Regions of Specific Cell cycle and Cell Survival Genes……………………… 119 Figure 4.4. ETS1 and ETS2 Recruit Coactivators CBP/P300 to The Enhancer Regions of Specific Genes……………………… 124

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LIST OF ABBREVIATIONS

A Alanine Adamts ADAM metallopeptidase with thrombospondin type Ang Angiopoietin AP-1 Activator protein 1 ATF Activating transcription factor ATP Adenosine-5'-triphosphate BAD Bcl-2-associated death promoter BCL2 B-cell lymphoma 2 BCL-XL B-cell lymphoma-extra large BMK Big MAPK BrDU 5-bromo-2-deoxyuridine BSA Bovine serum albumin ca Constitutively active CAM Chick Allantoic Membrane CD31 Cluster of differentiation molecule 31 Cdc Cell Division Cycle CDKs Cyclin Dependant Kinases ChIP Chromatin Immunoprecipitation cIAP Cellular Inhibitor of Apoptosis Cre Cyclization Recombination CREB cAMP response element binding CXCL12 Chemokine (C-X-C motif) ligand 12 DEPC Diethylpyrocarbonate Di-I-LDL di-I-acetylated-Low Density Lipoprotein DKO Double KnockOut Dll Delta-like DMEM Dulbecco Modified Eagle's Medium DMSO Dimethyl Sulfoxide dn Dominant negative DNA Deoxyribonucleic acid

xiii dNTPs Deoxyribonucleotide DuSPs Dual Specificity Phosphatases E Embryonic Day EC Endothelial Cells ECGS Endothelial Cell Growth Supplement ECM Extra-Cellular Matrix EdMT Endothelial-Mesenchymal Transition EDTA ethylenediaminetetraacetic acid EGF-R Epidermal growth factor-receptor eIF Eukaryotic Initiation Factor ELK1 Ets LiKe gene1 ERF Ets2 repressor factor ERG Ets-related gene ERK Extracellular signal Regulated Kinase ES Embryonic Stem ETS E Twenty Six Etv2 Ets variant gene 2 FACS Fluorescence-activated cell sorting FAK Focal Adhesion Kinase FBS Fetal Bovine Serum FGF Fibroblast derived Growth Factor FGFR Fibroblast derived Growth Factor Receptor fl flox'-containing loxPsites Fsp Fibroblast Specific Protein GFP Green Fluorescent Protein GFs Growth Factors GSEA Gene Set Enrichment Analysis GRB2 Growth factor receptor-bound protein 2 GTP Guanosine-5'-triphosphate H2O2 Hydrogen peroxide H3 Histone 3 H4 Histone 4 HCl Hydrochloric Acid HDAC Histone Deacetylase HGF Hepatocyte Growth Factor HIF Hypoxia Inducible Factor HRP Horse Radish Peroxidase HUVEC Human Umbilical Vein Endothelial Cells IEGs Immediate Early Genes IGF-R Insulin Growth Factor-Receptor ILK Integrin Linked Kinase xiv

IP Immunoprecipitation IRES Internal Ribisome Entry Site JNK c-Jun N-terminal kinases K3Fe(CN) Potassium ferricyanide 6 K4Fe(CN) Potassium ferrocyanide 6 KCl potassium chloride loxP locus of X-ing over P1 Lys Lysozyme MAP3K MAP kinase kinase kinase MAP4K MAP kinase kinase kinase kinase MAPK Mitogen Activated Protein Kinase MAPKK MAP Kinase Kinase MEF MADS box transcription enhancer factor MEFs Mouse Embryonic Fibroblasts MEK1 Mitogen-activated protein kinase kinase MeOH Methanol MgCl2 Magnesium chloride MKP MAPK phosphatases MLC Myosin Light Chain MLCK Myosin Light Chain Kinase MLEC Mouse Lung Endothelial Cells MMP Matrix Mettaloproteinase MMTV Mouse Mammary Tumor Virus MS Mass Spectrometry MSK Mitogen- and Stress-activated protein Kinase Na3VO4 Sodium Vanadate NaCl Sodium Chloride NaHCO3 Sodium Bicarbonate Nuclear Factor Kappa-light-chain-enhancer of activated NF-κB B cells NP40 Nonidet P-40 O/N Overnight PAK p21-Activated Kinase PBS Phosphate Buffered Saline PCR Polymerase Chain Reaction PDGF Platelet Derived Growth Factor PECAM Platelet Endothelial Cell Adhesion Molecule PFA Paraformaldehyde PGK Phospho Glycero kinase xv

PI3K Phosphoinositide 3-kinases PKB Protein Kinase B PLA2 Phospholipases A2 PMSF Phenylmethanesulphonylfluoride PS Penicillin -Streptomycin PTB Phosphotyrosine binding PTEN Phosphatase and tensin homolog PTK Protein tyrosine kinase PyMT Polyoma Middle T-Antigen RNA Ribonucleic acid RSKs Ribosomal S6 Kinases RTKs Receptor Tyrosine Kinases S Serine SA Senescence-Associated SAPK Stress Activated Protein Kinase SDS Sodium Dodecyl Sulfate Ser Serine SH2 Src Homology 2 SHC Src homology 2 domain-containing siRNA Small interfering RNA SMCs Smooth Muscle Cells SOS Son of sevenless SPARC Secreted protein, acidic, cysteine-rich SRF Serum Response Factor TAFs Tumor Associated Fibroblasts TAMs Tumor Associated Macrophages TBS Tris-Buffered Saline TFs Transcription Factors TGF Transforming Growth Factor Thbs Thrombospondin Thr Threonine Tyrosine kinase with immunoglobulin-like and EGF-like TIE domains TNF Tumor Necrosis Factor Tyr Tyrosine uPA Urokinase Plasminogen Activator VEGF Vascular Endothelial Growth Factor VSMCs Vascular Smooth Muscle Cells vWF von Willebrand Factor Y Tyrsoine

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CHAPTER 1

INTRODUCTION

The blood vascular network serves as the principal communication channel between various organs and tissues of the body. It is also critically important in maintaining homeostasis and coordinating wound repair. Thus angiogenesis, the formation of new blood vessels from a preexisting vascular network, is not only essential for embryonic development but is also indispensable for several physiological and pathological conditions including tumor growth

(Risau et al., 1997; Zetter et al., 1998; Carmeliet and Jain, 2011). The process of angiogenesis can broadly be categorized into two distinct phases, the activation phase and the resolution phase. Angiogenic factors, such as hypoxia or vascular endothelial growth factors (VEGFs), induce the activation phase in which endothelial cells (ECs) proliferate and migrate through the degraded extracellular matrix (ECM) components to form new capillary sprouts. In resolution phase, the vessels mature with the reconstitution of the basement membrane and recruitment of perivascular cells (Adams et al., 2007; Tanaka et al.,

2010). The execution of both these phases requires complex crosstalk between angiogenic inducers and inhibitors: signaling pathways that affect the growth, migration, survival of

ECs in the activation phase and their differentiation in the resolution phase (Zetter et al.,

1998; Folkman J, 2003; Cheresh et al., 2008). Vascular endothelial growth factors (VEGFs) have traditionally been considered as one of the most important inducers of both pre- and postnatal vascular development (Hoeben et al., 2004; Adams et al., 2007; Sivaraj et al.,

2013). The cellular responses to VEGF signaling are exerted through the modulation of several intracellular pathways such as the ras/Raf/Mek/Erk pathway involved in EC functions during angiogenesis (Serban D, 2008; Yang et al., 2008; Kim et al., 2010).

MAPKs activate transcription factors, such as ETS1 and ETS2, to regulate gene expression during angiogenesis. ETS1 and ETS2 belong to a family of transcription factors that share a highly conserved DNA-binding domain and a core DNA-binding consensus sequence. A considerable number of in-vitro studies suggest involvement of ETS1 in EC differentiation and function through regulation of the expression of vascular endothelial growth factor receptor 2 (VEGFR2) and Tie2, a receptor tyrosine kinase for angiopoietin (Ang) (Hashiya et al., 2004; Dejana et al., 2007). Depletion of ETS2 by small interfering RNA impedes the induction of the expression of aminopeptidase N (APN/CD13), a potent regulator of angiogenesis (Petrovic et al., 2003). Surprisingly, while only deletion of Ets2 is embryonic lethal, neither gene is essential for the development of embryo proper (Bories et al., 1995;

Muthusamy et al., 1995; Barton et al., 1998; Yamamoto et al., 1998).

Despite the growing body of evidence implicating ETS1 and ETS2 as important mediators of angiogenesis, due to the embryonic lethal nature of double mutant embryos, specific effector cellular processes and gene targets of this pathway remain relatively unknown. The aim of the current project was to decipher the downstream regulatory network of ETS1 and

ETS2 during angiogenesis. Recent evidence from our lab has demonstrated that EC specific

deletion of Ets2 in combination with global Ets1 knockout results in embryonic lethality with severe vascular defects. This phenotype can be rescued by the presence of a single copy of either Ets1 or Ets2 indicating the overlapping nature of their function (Wei et al.,

2009). In addition, dysregulated expression of mmp9 and bcl2l1 indicated cell survival and migration as the potential downstream effector processes of ETS1 and ETS2 regulatory network. Since these processes are essential for the activation phase of angiogenesis, we hypothesized that ETS1 and ETS2 regulate the activation phase of angiogenesis by modulating expression of genes involved in cellular processes such as cell survival, cell cycle and migration.

1.1. Development of Vascular Network

1.1.1. Evolution of Circulatory Network

Sustenance of life requires energy and procurement of energy necessitates all animals to perform tasks to capture and metabolize nutrients, acquire and distribute oxygen to aid metabolism and excrete metabolic waste and undigested material. To perform these tasks efficiently, different species have employed different strategies. Diploblastic animals and some of the early triploblastic animals, such as flatworms, lack any circulatory system. They obtain oxygen/nutrients and remove carbon dioxide by diffusion. However, diffusion is slow and useful only over small distances. Further increase in body size required the evolution of a circulatory system (Figure 1.1.) to provide efficient transportation to and from each cell in the body.

Figure 1.1. Evolution of Circulatory System

Adpated from Monahan-Earley R., 2013

Most triploblastic animals consist of fluid filled cavity, lined with mesoderm-derived epithelium (mesothelium), termed the coelom (Perez-Pomares et al., 2009). Coelomic cavities are devoid of a pumping system. Instead, transpiration is performed with the help of cilia lining the mesothilial cells or by the contraction of the mesothelial cells. Since coelomic cavities tend to be compartmentalized, they function in the local circulation of fluid (Munoz-Chapuli et al, 2005). In contrast, blood vascular systems have evolved to provide transport throughout the body of segmented animals. Blood vascular systems follow one of two principal designs: open or closed. Invertebrates display both open to closed vascular system. In arthropods and non-cephalopod mollusks, circulatory system is ‗open‘ in that the circulating fluid (hemolymph), a combination of blood and interstitial fluid, is not enclosed in vessels. Conversely, in all vertebrates blood never leaves the closed network of vessels, called ‗closed‘ circulatory system. In vertebrate system, transport is mediated by a central muscular pump. Amphibians, reptiles and fish exhibit different stages of circulatory system evolution. For example, fish have a single circulatory system consisting of an undivided two chambered heart whereas amphibians have a three chambered heart with double circuit blood flow (Figure 1.2.).

Figure 1.2. Vertebrate Circulatory Systems

Adpated from boundless.com

In mammals and birds, the heart is completely divided into four chambers with two atria and two ventricles. The separation of oxygenated blood from deoxygenated blood improves the efficiency of double circulation. The four-chambered heart of birds and mammals evolved independently from a three-chambered heart.

Blood vessel development in vertebrates can be divided into two distinct mechanisms; vasculogenesis, the de novo formation of the primitive vascular plexus and angiogenesis, the orchestrated remodeling of the primary vascular network to give rise to a mature vascular network.

1.1.2. Vasculogenesis

Vasculogenesis denotes de novo blood vessel formation during embryogenesis.

Angioblasts (angiogenic progenitor cells) migrate to sites of vascularization, differentiate into endothelial cells, and coalesce to give rise to the vascular primordia of the embryo

(Risau and Flamme, 1995; Carmeliet P, 2003).

1.1.2.1. Mechanism of Vasculogenesis

Circulatory system starts to develop soon after gastrulation. In embryo, earliest blood vascular structures appear in the form of blood islands which are aggregations of mesenchymal cells called hemangioblasts. The hemangioblasts are derived from mesoderm adjacent to the extraembyronic endoderm (Gonzalez-Crussi et al., 1971). They are common precursor to both endothelial cells and hematopoietic cells. Within the blood islands, the peripherally located angioblasts differentiate further to form endothelial cells.

The more internally situated cells give rise to hematopoietic precursors that would become the blood cells (Figure 1.3.).

Angioblasts first appear in the extra embryonic tissues where they develop together with the hematopoietic progenitor cells. Within the embryo, angioblasts develop unaccompanied without the concomitant differentiation of hematopoietic cells to ultimately aggregate directly into the dorsal aorta or cardinal vein. This process is mediated by vascular endothelial growth factor (VEGF), NOTCH and Sonic hedgehog

(SHH) (Pardanaud and Dieterlen-Lievre, 1993; Wilkinson et al., 2010; Kofler et al.,

2011) (Figure 1.4.). Extraembryonic vasculogenesis leads to the formation of a primitive vascular plexus whereas in the embryo proper, solitary angioblasts migrate and coalesce to form capillaries. By the two somite stage, the extra- and intra-vasculature fuse with each other (Risau and Flamme, 1995). Vasculogenesis continues as angioblasts differentiate into endothelial cells and begin to develop a vascular lumen. As lumenization occurs, endothelial cells form tight junctions. A basal lamina is deposited along the basolateral surface. The association of pericytes with the basement membrane marks vessel maturation. Later in development, the vascular plexus connects to the developing heart.

Figure 1.3. Schematic of Vasculogenesis

Adpated from Patel-Hett S., 2013

1.1.2.2. Regulation of Vasculogenesis

Signaling involved in regulation of vasculogenesis is poorly understood. However, studies performed on in vivo models suggest distinct roles of several growth factors.

VEGFs have long been established as the most crucial growth factor regulating vascular development. The VEGF family includes VEGF-A and its other related family members,

VEGF-B, VEGF-C, VEGF-D, endocrine gland VEGF (EG-VEGF), VEGF-E, VEGF-F,

VEGF-b and placental growth factor (PlGF) (Patel-Hett et al., 2013). Heterozygous

VEGF mice (VEGF+/-) lacking only one allele are embryonic lethal. VEGF+/- embryos show abnormal blood vessel development as a result of excessive EC death (Carmeliet et al., 1996; Ferrara et al., 1996). This phenotype worsens with the deletion of both copies of VEGF. VEGF-/- embryos lack a complete dorsal aorta even at embryonic day 8.5

(E8.5) and contain only a few ECs. By E9.5 the mutant embryos become necrotic.

VEGF family members interact mainly with three tyrosine kinase receptors, VEGFR-1

(Flt-1), VEGFR-2 (KDR in humans and Flk-1 in mouse) and VEGFR-3 (Flt-4). Genetic studies reveal the combined growth factor-receptor function in vascular development.

Mice lacking Flk-1 are embryonic lethal (~E9.5) due to defective hemangioblast differentiation and failure to initiate vasculogenesis indicating their involvement in hemangioblast differentiation (Shalaby et al., 1995) (Figure 1.4.). Deficiency of Flt-1 also results in embryonic lethality (Fong et al., 1995). However the vascular defects observed in these embryos are due to EC overgrowth, indicating that Flt-1 negatively regulates

VEGF dependent EC division. VEGFR-3 mutant embryos show major cardiovascular defect resulting in death at E9.5 (Patel-Hett et al., 2013). 10

Binding of VEGF ligands to their respective receptors initiate several signaling cascades that activate transcription factors (TFs) involved in vascular development.

Phosphorylation of VEGFR-2 Tyr1054 and Tyr1059 residues are required for maximal receptor activation (Kendall et al., 1999). Src activation through the recruitment of TsAd to phosphorylated receptor enhances endothelial cell migration and vascular permeability.

Binding of PLCγ to the receptor activates PKC to activate endothelial cell proliferation and migration through PKD or the MAPK pathways (Mechtcheriakova et al., 1999;

Mechtcheriakova et al., 2001; Hofer and Schweighofer, 2007; Wang et al., 2008).

Recuitment of SHB (Src Homology 2 Domain Containing Adaptor Protein B) stimulates endothelial cell migration or survival through the PI3K and AKT/PKB pathway. FAK,

Paxillin and IQGAP are also activated by phosphorylation of VEGFR-2 to promote endothelial cell motility (Figure 1.9.).

Fibroblast growth factors (FGF), specifically FGF2 appear to be crucial for the induction of angioblasts and hematopoietic cells from lateral and paraxial plate mesoderm (Cox and

Poole, 2000). Among the hedgehog family of morphogens, Indian hedgehog (Ihh) is potentially involved in blood island development in yolk sac. Ihh deficient mice exhibit

~50% embryonic lethality during mid gestation (Byrd et al, 2002). Ihh-/- mice have defective yolk sac formation in that the blood vessels are smaller and fewer in number.

Cell surface integrin receptors are another class of molecule important for vascular development. These receptors mediate cell-cell and cell-extra cellular matrix (ECM) adhesion required for EC survival and vessel branching (Cheresh et al., 2008; Velazquez et al., 2002).

Figure 1.4. Regulation of Developmental Vasculogenesis

(a) Hemangioblasts give rise to both hematopoietic lineage and angioblasts. (b) NOTCH signaling mediates arterial-venous specification by promoting arterial cell fate (light red cells). (c) VEGF, NOTCH and SHH mediates dorsal aorta and cardinal vein formation. In extra embryonic vascular development, angioblasts fuse to form the primitive vascular plexus. Adpated from Chung A., 2013

1.1.3. Angiogenesis

Angiogenesis is the process of new blood vessel formation from preexisting vasculature.

It is essential in development, reproduction and wound repair. High energy requirement during embryo development necessitates extensive angiogenesis. In adult, the blood vessels remain at a quiescent stage. However, during specific physiological conditions endothelial cells revert to a more active state to form new blood vessels in response to angiogenic factors. Pathological conditions, such as cancer and inflammatory disorders, also elicit similar response from the quiescent endothelial cells to promote excessive vessel growth or abnormal vascular remodeling (Potente et al., 2011).

1.1.3.1. Phases of Angiogenesis

The process of angiogenesis involves two distinct stages for mature vascular network formation, the activation phase and the resolution phase. During the activation phase, the basement membrane degrades resulting in increased vascular permeability. ECs migrate through the basement membrane and undergo proliferation to form the capillary lumen.

The newly formed vessels are ensheathed by pericytes and vascular smooth muscle cells

(VSMCs) to provide integrity and stability in the resolution phase (Figure 1.5.).

Figure 1.5. Two Phases of Angiogenesis

Adpated from Goumans M., 2003

Activation phase

In a quiescent vessel, the basement membrane and mural cells surround the ECs preventing them detaching and migrating out of the vessel (Eble et al., 2009). However, the presence of an angiogenic signal, such as VEGF, VEGF-C, FGF released by hypoxic, tumor or inflammatory cells, initiates the breakdown of the matrix and detachment of the pericytes. The proteolytic degradation of the basement membrane is mediated by matrix metalloproteases (MMPs), which not only helps the vessels dilate but also releases the pro angiogenic factors stored in the matrix (Arroyo and Iruela-Arispe, 2010). ECs loosen their junction and start to migrate towards the ECM surface in response to integrin signaling.

In order to provide a directional guidance to the migrating ECs and prevent them from migrating en masse towards the signal, on endothelial cell, called the tip cell, is selected to lead the nascent sprout following the guidance cues (Figure 1.6.). Endothelial cells surrounding the tip cells assume the role of ‗stalk cells‘, which divide to extend the stalk.

The NOTCH pathway regulates the specification of both the tip and the stalk cells

(Eilken and Adams, 2010 and Phng and Gerhardt, 2009). Stalk cells exhibit higher

NOTCH activity as well as the NOTCH ligand Jagged1 (JAG1) expression as compared to the tip cells. In contrast, the tip cells express higher levels of NOTCH ligand DLL4.

NOTCH upregulates VEGFR1 in stalk cells with a corresponding downregulation of

VEGFR2 to render the stack cells less responsive to the sprouting activity of VEGF

(Phng and Gerhardt, 2009).

Tip cells are equipped with filopodia to guide the sprout in properly folowing environmental cues. Migrating endothelial cells express receptors for axon-guidance cues, including ephrin receptors (EPH); neuropilins (NRPs) and PLEXIN-D1, which binds semaphorins; ROBO4, and UNC5B, which binds netrin proteins (Figure 1.6.).

Filopodia formation is regulated by CDC42, which is activated by VEGF (De Smet et al.,

2009). In comparison, stalk cells exhibit fewer filopodia and are more proliferative which is required to establish a vascular lumen. They also form junctions with neighboring cells and produce basement membrane components to provide integrity to the sprout. NOTCH activity is essential for stalk cell activity. The Notch-regulated ankyrin repeat protein

(NRARP) allows stalk cells to proliferate. It is also responsible for maintaining the stability of the nascent vessel connections (Phng et al., 2009).

Once formed, the sprout continues to grow in a highly directional manner guided by the environmental cues sensed until it comes in contact with other ECs. On contact with other tip cells, tip cells lose their motile phenotype and fuse with each other to add perfused neovessels to the existing network (Figure 1.6.). Although the exact mechanism of anastomotic process is poorly understood, other cell types may play significant roles during vessel fusion. Myeloid cells, specifically macrophages appear to act as cellular chaperones that support vessel anastomosis (Fantin et al., 2010). VE-cadherin containing junctions promote the adhesions between tip cells. After the connection is set up, the new vessel must undergo a maturation phase, termed resolution phase, to attain aquiescent stage in order to continue to be functional (Carmeliet et al., 2011).

Figure 1.6. Molecular Mechanism of Activation Phase

Adpated from Carmeliet P., 2011

Resolution Phase

The maturation of nascent vasculature is important for its integration to the vascular network to develop a vascular pattern that caters to the needs of the local tissues. The maturation process involves recruitment of other cell types, increased formation of cell junctions as well as deposition of a basement membrane (Jain, 2003).

Recruitment of mural cells (pericytes and VSMCs) to neovessels is a necessary component of vessel stabilization. Mural cells are indispensable for normal development, homeostasis and organ function. Several growth-factor families, such as Platelet derived growth factors (PDGFs), angiopoietins and transforming growth factor β (TGF-β), contribute to the recruitment process (Jain, 2003). Differentiation, proliferation and migration of mural cells are stimulated by TGF-β signaling. In absence of functional

TGF-β receptor 2 (TGFBR2), endoglin, or activin receptor-like kinase 1 (Alk1) development of mural cells is impaired leading to fragile vessel formation in mice

(Pardali et al., 2010). Endothelial cells release PDGF-B to communicate with platelet- derived growth factor receptor-β (PDGFR-β) expressed by mural cells to promote proliferation and guide migration of the mural cells (Gaengel et al., 2009; Hellberg et al.,

2010) (Figure 1.6.). Absence of either PDGF-B or PDGFR- β results in pericyte deficiency, vessel leakage, micro-aneurysm formation, and bleeding. Mural cells also produce Angiopoietin-1 (ANG1), which is required for vessel stabilization by aiding in pericyte adhesion and tightening of endothelial cell junctions to prevent leakage. Lack of ephrinB2 leads to erratic mural cells migration and defective vasculature (Pitulescu and

Adams, 2010). VSMCs also require NOTCH signaling for proper arterial differentiation

(Gridley, 2010) (Figure 1.6.).

In mature vessels, endothelial cells attain a quiescent state by assuming a phalanx phenotype. Since mature vessels need to adjust their shape and function to account for changing tissue oxygen demands, phalanx ECs express oxygen sensors and hypoxia- inducible factors (HIFs). HIFs allow the ECs to adapt to changes in oxygen demand and optimize blood flow (Fraisl et al., 2009 and Majmundar et al., 2010).

Figure 1.6. Resoltuion Phase of Angiogenesis

Adpated from Potente M., 2011

1.1.3.2. Mediators of Vessel Branching

Angiogenesis is a highly orchestrated multistep process that requires critical control and coordination of EC behavior. Several signaling pathways control EC behavior during vascular remodeling. As discussed below, every step of this process is tightly regulated by growth factors that stimulate different intracellular signaling pathways leading to activation of transcription factors in order to regulate expression of genes required for proper execution of angiogenesis.

Cell Types Regulating Angiogenesis

Apart from ECs and pericytes, which are directly involved in the angiogenesis process, fibroblast, immune cells and tumor/epithelial cells regulate different aspects of vascular remodeling (Figure 1.8.).

Fibroblasts, a principal component of stroma, control angiogenesis not only by releasing factors required to regulate angiogenesis but also by remodeling the ECM. The matrix sequesters protein of both pro- and antiangiogenic nature such as MMP9, a pro- angiogenic factor (Egeblad and Werb, 2002) and Thrombospondin1 (Thbs1) that functions to inhibit angiogenesis by sequestering VEGF (Good et al., 1990). Fibroblasts function to secrete several such ECM components including thrombospondin (Jaffe et al.,

1983). Fibroblasts also secrete several cytokines that promote blood vessel sprouting

(Figure 1.8.). Fibroblast-derived matrix proteins, such as Collagen alpha-1 (I) (Col1A1), procollagen C-endopeptidase enhancer 1 (PCOLCE), and secreted protein acidic and rich in cysteine (SPARC), have been shown to be necessary not only for EC lumen formation in the fibrin gel bead assay but providing the required stiffness of the matrix using

rheology as well (Newman et al., 2011). Furthermore, during tumor angiogensis, fibroblast specific deletion of Ets2 limits the growth of primary breast tumors with reduced angiogenesis (Wallace et al., 2013).

Like fibroblasts, immune cells such as macrophages also secrete potent pro-angiogenic cytokines and growth factors as well as a broad array of angiogenesis-modulating enzymes necessary for proper execution of angiogenesis (Figure 1.8.). Role of immune cells in angiogenesis is specifically prominent in tumor angiogenses. TNF-α and TGF-β released by tumor associated macrophages as well as several extracellular proteases that stimulate VEGF-A expression to induce angiogenesis (Torisu et al., 2000). Conditional ablation of the Ets2 oncogene in mouse macrophages decreases the frequency of lung metastases in mouse breast cancer models. In this model, macrophage specific- Ets2 represses anti-angiogenic genes, such as Thbs1, to coordinate tumor angiogenesis

(Zabuawala et al., 2010). In addition, the CSF1-ETS2 signaling axis in myeloid cells promotes metastatic tumor progression. Over expression of CSF1 inducible miRs in macrophages increase angiogenesis and tumor cell proliferation in a melanoma matrigel plug model (Mathsyaraja, H. unpublished data).

Tumor/epithelial cells also regulate angiogenesis in a manner similar to fibroblasts.

Hypoxia caused by inefficiently vascularized tumors can lead tumor cells to act as a source for secreted factors that aid blood vessel sprouting and hence angiogenesis

(Fukumura et al., 1998) (Figure 1.8.). Tumor cell derived MMPs enhance the release of pro-angiogenic factors from the surrounding stroma creating a favorable environment for

angiogenesis (Ito et al., 2007). Tumor cells also stimulate fibroblasts and immune cells to release growth factors and cytokines that act as mitogens for angiogenesis.

Extra Cellular Matrix and Matrix Metalloproteinases (MMPs)

The Extra Cellular Matrix (ECM) an intricate network of macromolecules occupying the space surrounding most of the cells in multicellular organisms. The matrix is composed of a variety of proteins and polysaccharides, such as collagen IV, laminins, fibronectins, heparin sulphates and SPARC, secreted locally by cells (Figure 1.8.). Due to its dynamic nature, ECM supports various functions such as providing support, promoting cellular communication and guiding migratory cells. The matrix can also serve as a reservoir of various factors required for stimulating different processes including activators and repressors of angiogenesis. Owing to its ability to act as a scaffold, ECM helps maintain the quiescent state of the mature vessels. It not only prevents the ECs to migrate physically, but the antiproliferative properties of the ECM components prevent the onset of neovessel formation. However, physiological changes that necessitate angiogenesis trigger activation of MMPs, which degrade the basement membrane. Remodeling of the

ECM enables the ECs to move unrestricted and it converts the basement membrane into a pro-angiogenic environment as well with the release of angiogenic factors (Liotta et al.,

1980; Deryugina and Quigley, 2010).

MMPs are a family of enzymes that remodel the ECM in a manner dependent on the local physiological cues (Figure 1.8.). Based on their functional location, MMPs can be subdivided into two groups; the secreted MMPs such as collagenases (MMP-1, MMP-8 and MMP-13), stromelysins (MMP-3, MMP-10 and MMP-11), gelatinases (gelatinase A

or MMP-2; gelatinase B or MMP-9) (Pepper, 2001) and membrane-type MMPs (MT-

MMPs) such as MT1-MMP (Pepper, 2001). Macrophages, neutrophils and mast cells initiate angiogenesis by MMP9-mediated activation of VEGF (Du et al., 2008; Heissig et al., 2010). MT1-MMPs, released by tip cells mediate basement membrane degradation and promote EC migration and invasion (Hiraoka et al., 1998; Galvez et al., 2002;

Oblander et al., 2005). In contrast, MMPs also prevent excess and unwanted branching by cleaving plasma proteins, matrix molecules, or other proteases (Nyberg et al., 2005).

However, given the destructive nature, the function of MMPs must be tightly regulated to prevent excessive degradation of the ECM. For instance, loss of the inhibitor PAI-1 leaves too little support for sprouting (Blasi and Carmeliet, 2002).

Growth Factors

Vascular endothelial growth factors play a surprisingly predominant role in regulating the angiogenic process. Absence of either VEGF (VEGF-A) or its receptor VEGFR-2 causes attenuation of vascular development (Carmeliet, 2003). Selection of tip cells is largely dependent on surrounding VEGF gradient which upregulates DLL4 expression in tip cells. DLL4 activates NOTCH in stalk cells that reduces VEGFR-2 expression. Without

VEGFR-2 stalk cells become unresponsive to angiogenic signals of VEGF ensuring selection of only one tip cell. The effect of VEGF on ECs is dependent on the source and location. Vessel branching is promoted by matrix-bound VEGF whereas the soluble form of VEGF stimulates vessel enlargement. VEGF released by ECs themselves are required for maintaining vascular homeostasis (Lee et al., 2007). However, paracrine VEGF, released by tumor cells, myeloid cells etc, increases vessel branching (Stockmann et al.,

2008). Unlike VEGFR-2, VEGFR-3 is only required during early embryogenesis

(Tammela and Alitalo, 2010). Function of VEGFR-1 (Flt-1) in angiogenesis is yet to be elucidated. Since VEGFR-1 loss enhances vessel growth, it appears to function as decoy to sequester and moderate the level of free VEGF available to activate VEGFR-2 (Fischer et al., 2008).

During the resolution phase, ECs release PDGF-B to chemoattract the pericytes expressing PDGF receptor-β (PDGFR-β) (Hellberg et al., 2010) (Figure 1.8.). The ablation of PDGF-B results in insufficient pericyte recruitment leading to vessel leakage and bleeding. PDGFR-β-hypomorph mice exhibit blood–brain barrier (BBB) defects and neurodegenerative damage owing to insufficient number of pericytes (Quaegebeur et al.,

2010). Quiescent ECs in mature vessels depend on pericytes for their survival. Pericytes releases VEGF that confer protection to ECS against VEGF withdrawal. During tumor angiogenesis, VEGF inhibits PDGFR-β signaling in mural cells and trigger abnormal vessel growth.

Fibroblast growth factors (FGFs) can induce angiogenesis by activating the receptors on

ECs directly or by stimulating the release of angiogenic factors from other cell types

(Beenken and Mohammadi, 2009). FGFs stimulate EC proliferation and migration as well as production of collagenase and plasminogen activator (Gospodarowicz et al.,

1989; Terranova et al., 1985). FGF signaling is also involved in maintaining vascular integrity. Loss of FGF signaling leads to fragmented vessel as a consequence of dissociation of adherens junctions and tight junctions in ECs (Murakami et al., 2008).

Among the fibroblast growth factors, bFGF and FGF1 have both angiogenic and arteriogenic properties and FGF9 stimulates angiogenesis during bone repair.

The human angiopoietin (ANG) family consists of two receptors, TIE-1 and TIE-2, and three ligands, ANG-1, ANG-2 and ANG-4. The angiopoietins- Ang1 and Ang2 are

75KD-secreted molecules that share a high degree of homology. ANG-1 is a TIE-2 agonist, and ANG-2 acts as a competitive ANG-1 antagonist in a context-dependent manner. TIE-1 has no identified ligand and is thought to act as negative regulator of TIE-

2 (Augustin et al., 2009). ANG-1, expressed by the mural and tumor cells, is primarily involved during the quiescent state of the vessels. ANG-1 promotes vessel tightness by enhancing mural cell coverage and basement deposition (Saharinen et al., 2008) (Figure

1.8.). In response to angiogenic stimuli, tip cells release ANG-2, which antagonizes

ANG-1 and TIE-2 signaling. As a result, vascular permeability is increased through detachment of mural cell to support endothelial cell sprouting (Augustin et al., 2009).

TIE-2 deficient mice are embryonic lethal. The mutant embryos display abnormal vasculature with reduced vessel branching and sprouting angiogenesis (Sato and Rifkin,

1989; Dumont et al., 1992).

The transforming growth factor-β (TGF-β) family is a group of highly conserved cytokines (Massague, 1990). The TGF-β family ligands bind type II receptors facilitated by Endoglin (ENG), a type III receptor. The stimulated type II receptors phosphorylate type I receptors, such as activin receptor-like kinase (ALK 1, also known as ACVRL1) to activate the downstream Smads (Driesche et al., 2003). Absence of TGF-β receptors

ALK-1, TGFR-1 (ALK-5), TGFR-2 or ENG results in poor arteriovenous formation

(Beenken and Mohammadi, 2009). However, due to inconsistent results both pro- and anti-angiogenic properties have been attributed to TGF-β. This appears to be, at least partially, due to the context dependent functions of TGF-β family members (Carmeliet and Jain, 2011). At low doses, the transforming growth factor-β 1 (TGF-β1) favors activation phase by upregulating angiogenic factors and proteinases. In contrast, higher dose stimulates resolution phase by inhibiting EC growth and recruitment of VSMCs

(Carmeliet, 2003). TGF-β1 deficient mice die due to defective formation of yolk sac vasculature and hematopoietic system (Goumans et al., 2003).

Signaling Pathways

VEGF receptors carry multiple tyrosine residues with differing function. Some tyrosine residues are autophosphorylation sites, some act as docking sites and others regulate intrinsic kinase activity (Eichmann et al., 1997; Patel-Hett et al., 2013). When VEGF binds to its receptors, it leads to homo- or heterodimerization to activate the receptor kinase activity or autophosphorylation to stimulate downstream signaling. Tyrosine (Y)

1175 (or Y1173 in mouse) represents a docking site for phospholipase C-gamma (PLC-γ)

(Figure 1.9.). The activation of PLC-γ/ protein kinase C (PKC) pathway by the phosphorylation of Y1175 results in the activation of the c-Raf-MEK-MAP-kinase cascade, which promote endothelial cell proliferation, and release of calcium (Lyttle et al., 1994; Xia et al., 1996; Takahashi et al., 2001). Mice carrying a phenylalanine residue in place of Y1173 die at E9.5. The mutant mice display severely defecting vascular development (Sakurai et al., 2005). B-Raf knockout mice are embryonic lethal with extra-embryonic vascular deformities (Wojnowski et al., 1997). Deletion of MEK1 in

mice results in embryonic lethality at E10.5 due to abnormal placental vasculature

(Giroux et al., 1999). In addition, over-expression of dominant negative (dn) MEK1 in human umbilical vein EC (HUVEC) inhibits ERK1/2 activation compromising EC survival (Mavria et al., 2006). Mice with EC specific knockout of ERK2 in combination with conventional deletion of ERK1 are embryonic lethal. The double knockout embryos exhibit defective EC proliferation and migration leading to poor vascular differentiation

(Srinivasan et al., 2009).

Phosphoinositide 3-kinase (PI3K) is also activated by the phosphorylation of Y1175/1173 residue of VEGFR2. Akt/PKB pathway, stimulated by activated PI3K, mediates endothelial cell survival and migration as well as vascular permeability and angiogenic processes through its activation of endothelial nitric oxide synthase (eNOS) (Dayanir et al., 2001; Fujio and Walsh, 1999; Olsson et al., 2006; Fukumura et al., 2001) (Figure

1.9.). PLC-γ/PKC also activates protein kinase D (PKD)-dependent phosphorylation and nuclear export of histone deacetylase 7 (HDAC7) which regulates migration and proliferation of ECs (Wang et al., 2008). Y951 residue in VEGFR2 is a docking site for

T-cell specific adaptor (TsAd), which associates with the cytoplasmic tyrosine kinas Src to regulate EC migration and vascular permeability (Matsumoto et al., 2005) (Figure

1.9.).

The NOTCH signaling pathway is an evolutionary conserved, short-range communication transducer that operates in various cell types to regulate a broad range of cellular processes (Andersson et al., 2011). Notch receptors are large single pass type I transmembrane proteins. Four different NOTCH receptors exist in mammals (NOTCH 1-

4) that are bound by five ligands of the Jagged (Jagged 1 and Jagged 2) and Delta-like

Ligands (DLL1, DLL3 and DLL4) (Radtke et al., 2013). As discussed earlier, NOTCH with the antipodal expression of two of its ligands, DLL4 and JAG1 strongly influences tip and stalk cell specification. Excess NOTCH signaling in ECs reduce the number of filopodia and the cells are excluded from the tip position (Jakobsson et al., 2010). DLL4 in endothelial cells also upregulates PDGFR-β in NOTCH+ mural cells to enhance vessel maturation (Benedito et al., 2009). NOTCH signaling in stack cells works in a negative feedback loop mechanism as it upregulates its own inhibitor, NOTCH-regulated ankyrin repeat protein (NRARP) (Phng et al., 2009). This negative NOTCH regulation enables the ECs to dynamically switch between the tip and the stalk cell phenotypes (Eilken and

Adams, 2010; Phng and Gerhardt, 2009). Genetic studies also support the importance of

NOTCH signaling in angiogenesis. NOTCH1 deficient mice are embryonic lethal at

E11.5 exhibiting defective vessel remodeling. Double mutant mice lacking both

NOTCH1 and -4 show more severe vascular deformity including poor formation of large vessels (Krebs et al., 2000). Constitutively active expression of NOTCH in ECs similarly leads to embryonic lethality as a result of abnormal vascular development (Uyttendaele et al., 2001). Deletion of either JAG1 or a single copy of DLL4 results in embryonic death

(Xue et al., 1999; Krebs et al., 2004).

Since angiogenesis is highly regulated by the interaction between vascular cells and surrounding tissue, ECs require mechanism to communicate with the surrounding matrix.

Integrins are glycosylated, heterodimeric transmembrane receptors that mediates the intercellular and cell-matrix interaction (Desgrosellier and Cheresh, 2010; Hodivala-

Dilke, 2008). Integrins act as signal hubs owing to their ability to communicate with multiple molecules and provide bi-directional signal transfer. The communication between ECs and pericytes is mediated by integrin to aid in vessel maturation. Growth factors, such as VEGF, FGF, or their ligands can promote vessel growth by binding to integrins (Figure 1.9.). IntegrinαVβ3 stimulates the RAS pathway in ECs to increase

ERK1/2 activation, which regulates EC survival and migration (Hood et al., 2003).

Expression of zymogen protease required by the invading tip cells is also upregulated under the influence of integrins.

Semaphorins are a family of secreted or transmembrane glycoproteins. Class 3 semaphorins consist of 7 member designated semaphorin A-G. While Sema3C appears to have a pro-angiogenic effect by increasing integrin activity (Banu et al., 2006), Sema3A and Sema3F inhibit EC proliferation, migration and survival and limit angiogenesis

(Bielenberg et al., 2004; Kessler et al., 2004; Miao et al., 1999). Netrins, a group of secreted proteins, bind to either the deleted in colorectal cancer (DCC) or uncoordinated-

5 (UNC5) receptors. Ablation of UNC5 in mice causes embryonic lethality at E12.5 due to poor capillary branching (Lu et al., 2004). However, owing to opposing results, the overall effect of netrins on angiogenesis is yet to be delineated.

Figure 1.8. Molecular Regulators of Angiogenesis

Adpated from Weis M., 2011

Figure 1.9. Function of VEGF-VEGFR-2

Signaling Cascade During Vascular Development

Adpated from Patel-Hett S., 2013

1.1.3.3. Cellular Processes Involved in Angiogenesis

Angiogenesis is a complex multistep process. It involves stimulation of quiescent ECs to become highly proliferative and migratory and form neovessels. This angiogenic switch, the conversion of quiescent to active ECs, is also dependent on sell survival. The following section details the importance of these cellular processes in angiogenesis.

Cell Proliferation/Cell Cycle

Cell proliferation, at its core, is the increase in cell number, which involves cell growth followed by cell division, together constituting the cell cycle. During each cycle, cells must replicate its genetic material (interphase) and segregate the sister chromatids into two genetically identical daughter cells (mitosis or M phase). The synthetic (S) phase of interphase where DNA replication takes place is separated from M phase by gap phases

(G1 and G2). In addition, some cells, such as ECs in mature vessels, enter a quiescent stage called G0 (Gap zero), which is either a stage separate from interphase or an extended G1. Cell cycle progression is mediated by Cyclins and Cyclin Dependent

Kinases (CDKs). These kinases when associated with the respective Cyclins act as switches to activate specific protein at appropriate stages of cell cycle through phosphorylation. The entire process is perused by checkpoints at certain stages. Any abnormality causes a cell cycle arrest and may lead to apoptosis.

In adults, the differentiated endothelial cells (ECs), lining the inner surface of mature blood vessels, adopt the quiescent (G0) state. However, they enter active cell cycle in response to specific physiological and pathological conditions requiring increased nutrient and oxygen supply (Risau 1997; Zetter 1998; Carmeliet 2011; Potente 2011).

This plasticity of mature ECs is essential since of cell division is required to elongate the newly formed sprout and develop the vascular lumen. GFs like VEGF and FGF stimulate

EC proliferation essential for both vasculogenesis and angiogenesis (Takahashi et al.,

2001).

Cell Survival/Apoptosis

Apoptosis is a form of cell death following a programmed sequence of events leads to the characteristic morphological changes, such as DNA fragmentation, cell shrinkage and blebbing, without the release of harmful substances. This process is crucial in developing and maintaining a healthy population of cells by eliminating the unwanted ones.

Maintaining EC survival by preventing apoptosis is an essential component of vessel branching. EC survival is regulated by growth factors and cell-matrix adhesion (et al.,

2002). VEGF promotes EC survival. VEGFR2 and the PI3K/Akt have been identified as the signal transduction pathways as essential for mediating EC survival induced by

VEGF (Fujio and Walsh, 1999; Gerber et al., 1998). Hypoxia-induced reduction of

VEGF expression leads to regression of retinal capillaries due to enhanced EC apoptosis in neonatal rats (Alon et al., 1995). Targeted deletion of Ets1 and Ets2, transcription factors downstream of VEGF-RAS-MAPK pathway, in EC results in embryonic lethality at E13.5-E15.5 as a result of reduced EC survival (Wei et al., 2009). Similarly in tumor angiogenesis, vessel regression caused by EC apoptosis is observed as a result of VEGF inhibition (Benjamin et al., 1998; Jain et al., 1998; Benjamin et al., 1999; Shaheen et al.,

1999). VEGF was shown to induce the expression of antiapoptotic proteins such as Bcl-

2, survivin, and XIAP (Gerber et al., 1998; Nor et al., 1999; Tran 1999). Another growth

factor, bFGF can inhibit EC apoptosis induced by radiation or growth factor deprivation.

Similar to VEGF, bFGF also upregulates the expression of the antiapoptotic proteins such as Bcl-2 and surviving (Fuks et al., 1994; Karsan et al., 1997; O‘Connor et al., 2000).

The importance of the ECM as a cell survival factor is demonstrated by a phenomenon called anoikis in which ECs rapidly undergo apoptosis in the absence of any ECM interaction mediated by integrins (Meredith et al., 1993; Stromblad et al., 1996). Studies disrupting the ανβ3-mediated EC adhesion to extracellular matrix, demonstrated inhibition of tumor- and growth factor–induced angiogenesis in vivo due to increased induction of

EC apoptosis leading to in neovessels but not in quiescent ECs (Brooks et al., 1994). The ligation state ανβ3-integrin affects p53 activity and the Bax cell death pathway to influence EC survival (Stromblad et al., 1996). Furthermore, association of α5β1-, the

ανβ3-, and the α1β1-integrin with Shc can mediate cell survival and cell cycle progression via the Ras/MAPK/ERK pathway (Wary et al., 1996).

Apart from cell-matrix communication, cell-cell contact also inhibits apoptosis. VE- cadherin, an adhesive protein present in adherens junctions between endothelial cells, is required for the antiapototic effect of VEGF. Targeted deletion VE-cadherin, results in embryonic lethality at E9.5 due to increased EC apoptosis (Carmeliet et al., 1999).

Platelet EC adhesion molecule-1 (PECAM-1, CD-31) homophilic adhesion can inhibit apoptosis induced by growth factor deprivation (Bird et al., 1999). N-cadherin mediates mural cell-EC adhesion. Embryos lacking N-cadherin die at med gestation exhibiting increased apoptosis (Luo and Radice, 2005).

Cell Migration

The process of cell migration broadly refers to translocation of cells from one location to the other. Coordinated cellular migration is absolutely essential for development and maintenance of multicellular organisms. Migration generally comprises a set of subprocesses, which include polarization, protrusion and adhesion, translocation of the cell body and retraction of the trailing end of a cell.

EC migration is fundamental to angiogenesis. It requires regulation by several factors that culminate into cytoskeleton remodeling to from filopodia that act as sensors to guide the migratory mechanism. Based on the nature of the cues stimulating the movement, EC migration is carried out by 3 major mechanisms, namely chemotaxis, cellular movement directed toward a gradient of soluble chemoattractants such as growth factors; haptotaxis, migration directed toward immobilized ligands; and mechanotaxis, migration caused by mechanical forces. Endothelial cell motility during angiogenesis involves a combination of all three mechanisms (Lamalice et al., 2007). Chemotaxis is initiated in response to growth factors such as VEGF, bFGF, and angiopoietins. VEGF induced activation of the

Rho Family of GTPases stimulate the PI3K-AKT signaling cascade essential for migration (van Nieuw Amerongen et al., 2003). VEGF also activates Cdc42 to regulate stress fiber organization (Lamalice et al., 2004).

Apart from the growth factors, immobilized ECM components can also drive EC migration in a process termed haptotaxis. ECM helps maintain the quiescent state of ECs in mature vessel. However, during the onset of angiogenesis proteases degrade the ECM releasing motogenic signals. EC adherence to EM is mediated by integrins to facilitate

migration. Several signaling pathways responsible for migration are also stimulated by integrins (Klemke et al., 1997). Endothelial cells line the inner wall of blood vessels. Due to blood flow, these cells are always under fluid shear stress. This mechanical stress not only influences migration initiation but modulate various steps of the process including protrusion and adhesion of the leading edge as well as the retraction of the rear of the migratory cells. As an effect of shear stress microtubules elongate toward the direction of the blood flow activating Rac to promote actin polymerization and lamellipodia formation (Li et al., 2005).

1.1.3.4. Angiogenesis in Disease and Development

Angiogenesis is not only a critical process during embryonic and fetal development, it is essential in adult during several physiological processes including wound healing, the menstrual cycle, and pregnancy. Progress and development of specific diseases, such as immunogenic rheumatoid arthritis, psoriasis, ischemic diseases and tumorigenesis, also require formation of new vasculature or the lack thereof.

The Vascular network is the first organ to form and function during vertebrate development underscoring its importance embryogenesis. As the embryo grows, the continual increase in size precludes the use of diffusion as the process for nutrition/gas exchange. Development of a vascular system becomes necessary. As discussed earlier, improper blood vessel remodeling during development results in embryonic lethality.

However, even after embryonic development is complete, the vasculature continues to remodel dynamically. Many events occurring during embryonic vascular development, recapitulate during neovascularization in the adult.

After an injury, the wound is temporarily sealed with the formation of a blood clot and a platelet plug, which secretes VEFGA increasing the local VEGF-A concentration to promote endothelial cell proliferation as repair of injured dermis requires reestablished blood supply (Schultz & Wysocki 2009). In addition, inflammatory cells are recruited to the site of injury to protect against invading pathogens. VEGF-A further increases the secretion of MMP1, MMP2, urokinase-type and tissue-type plasminogen activators (uPA and tPA, respectively), and plasminogen activator inhibitor 1, which together aid the formation of mature granulation tissue required for the final resolution of wound repair

(Bao et al. 2009). Angiogenesis is an essential process during the menstrual cycle as well as to embryo implantation. VEGF-A along with its receptors, and other pro angiogenic factors, such as PlGF and basic fibroblast growth factor (bFGF), are expressed in human endometrium, decidua, and placenta (Smith 2001, Torry & Torry 1997). Luteal function and embryo implantation in rodents and primates are attenuated with the administration of VEGF inhibitors due to defective angiogenesis (Ferrara et al. 1998; Hazzard et al.

2002).

Mechanisms regulating angiogenesis during pathological conditions share several features with physiological angiogenesis. After a tumor crosses a certain size threshold, it requires oxygen more than that can be provided by the local supply. The hypoxic condition generated thereof stimulates angiogenesis. This process will be further detailed in later chapters. Failure to execute normal vasculature formation results in diseases such as coronary artery disease, stroke and Intraocular Neovascular Disorders. VEGF-A is a key mediator of ischemia-induced intraocular neovascularization. Neovascularization in

animal models of retinal ischemia correlates with VEGF expression. (Alon et al. 1995;

Miller et al. 1994).

1.2. ETS Family of Transcription Factors

ETS proteins are a group of evolutionarily related, DNA-binding transcriptional factors unique to metazoans. They are characterized by an evolutionarily conserved DNA- binding domain, termed the ETS domain that recognizes a purine-rich core GGAA/T sequence. The first ETS (E26 transformation-specific sequence) protein identified was a transduced mutant form of Ets1, v-Ets of the E26 avian leukosis virus (ALV). ALV carries v-ets and v-myb and is capable of inducing erythroblastosis in avian species

(LeKang et al., 2008) et al., 1983). Afterwards, 28 different ETS proteins have been identified in humans belonging to 12 subfamilies (Wang et al., 2009) (Figure 1.10.). All of these family members are present in mice except TEL2. The phylogeny tree of the Ets family points toward the duplication of a common ancestral gene early in metazoan development (Figure 1.11.).

Although the ETS domain in highly similar in all ETS proteins, various distinct functional and structural domains are observed outside the Ets domain. Some members carry the ‗Pointed‘ domain near their N-terminal end. Three members of the ternary complex factor (TCF) subfamily harbor the B-box, which enables them to interact with the serum responsive factor (SRF). The OST domain, present only in GABPA, promotes co-factor recruitment (hassle and Richmond, 2001; Kang et al., 2008) (Figure 1.10.). In addition, several posttranslational modifications, such as ubiquitinylation, sumoylation, glycosylation, and acetylation, have also been observed on ETS factors. This overall

diversification of the family members seems to have evolved to provide unique functional properties of individual transcription factors.

Diverse biological functions have been attributed to ETS family proteins. Genetics studies have demonstrated specialized functions ranging from regulating different aspects of embryo development to aging (Hollenhorst et al., 2011). Although most of these functions are mediated by Ets proteins acting as transcription activators, some proteins have been shown to act as repressors also. For example, in mammalian cells, Bcl2l1 is activated by ETS2 (Smith et al. 2000), but repressed by TEL (Irvin et al. 2003).

Figure 1.10. Domains of Ets Family Transcription Factors

Adapted from Hollenhorst P., 2011

1.11 Phylogeny of Ets Family Proteins

The relatedness of ETS domain sequences is demonstrated by a dendogram. The length of each horizontal line indicates estimated evolutionary distance. Adpated from Hollenhorst P., 2011

1.2.1. The ETS Domain

The structure of the evolutionarily conserved ETS domain is a variant of the helix-turn- helix DNA-binding motif. The domain consists of three α-helices (α1-α3) on a small four-stranded, antiparallel β-sheet (β1-β4) scaffold (Figure 1.12.). The α1-β1-β2-α2-α3-

β3-β4 arrangement forms a winged helix-turn-helix (wHLH) architecture (Gaziwala and

Burley 2000). Although the ETS domain recognizes a 9-bp DNA sequence with a 5-bp core, invariant region 5‘-GGA(A/T)-3‘, the actual DNA-protein binding spans over a region of 12 to 15 bp. The three α helices α1-α3, resides in the major groove of DNA.

The two absolutely conserved arginines in α3 provide base contacts to the two guanine

(G) residues of the GGA core. However, there is no other direct contact outside the core region indicating that the flank regions provide specificity for DNA binding (Szymczyna and Arrowsmith, 2000).

Several structural elements are appended to the ETS domain to diversify their DNA binding specificity and in turn their biological functions (Figure 1.12.). ETS1, ETS2,

ETV6, and GABPA, have two C-terminal helices, termed H4 and H5. These helices enhance autoinhibition in ETV6 possibly via a steric mechanism (Green et al., 2010). In

GABPA the helices provides a platform to bind to GABPB1 in order to promote DNA binding (Batchelor et al., 2010). Two more helices, HI-1 and HI-2, join H4 and H5 in

ETS1 and ETS2. The helices provide allosteric autoinhibition of DNA binding

(Hollenhorst et al., 2011).

1.2.2. The PNT Domain

Approximately one third of the Ets family members (11 out of 28) carry a second 80- residue conserved domain, the pointed (PNT) domain (Figure 1.10.). Four α-helices (H2–

H5) along with a short 310- or α-helix (H2‘) constitute the core of this domain. The core structure is similar to that of SAM domain (Figure 1.12.). Similar to the ETS domain, additional helical elements (H0 and H1) are appended to the conserved-core of the PNT domain. The helices provide opportunity for individual members to have unique functions.

In ETS1 and ETS2, the PNT domain acts as the docking site for mitogen-activated protein kinases (MAPKs). More specifically PNT domains facilitate substrate (MAPK;

ERK for ETS1 and ETS2) binding to the transcription factors (Seidel and Graves, 2002).

The helices H2, H4 and H5 allow the PNT domain to interact with its substrate via a three-dimensional surface. MAPK induced phosphorylation of threonine residues immediately preceding the PNT domain enhances the transcriptional activation of ETS1 and ETS2 (Klambt, 1993; and Wasylyk et al, 1997). Phosphorylation of Thr38 located immediately adjacent to ETS1 PNT domain by ERK2 does not change its structural or dynamic properties. However, the phosphorylation promotes the recruitment of the co activator CBP by increasing the affinity of ETS1 for the transcription adaptor zinc- binding domain 1 (TAZ1) domain of CBP by ∼30-fold (Foulds et al., 2004; Hollenhorst et al., 2009; Nelson et al., 2010). Interestingly the same surface of the PNT domain act as the docking sites for both ERK2 and CBP, signifying the importance of the PNT domain

in relaying signal transduction providing an additional source for specificity and unique function.

Figure 1.12. Structure of ETS and PNT Domain

(a) The core ETS domain consisting of three α-helices (red rectangles) on a small four-stranded, antiparallel β-sheet (red arrows). Cyan rectangles represent appended helices. On right is a ribbon diagram of residues 301–441 of ETS1 with appended helices and the core domain. (b) The core PNT domain (green) containing four α-helices (H2– H5) along with a short 310- or α-helix (H2‘) along with appended helices (orange rectangles). On right is the ribbon diagram of residues 29–138 of ETS1 including the PNT domain. Adpated from Hollenhorst P., 2011

1.2.3. The Biological Role of Ets Factors

Ets factors play a significant role in regulating virtually all cellular processes including growth, development, differentiation, survival, and oncogenic transformation (Dittmer and Nordheim 1998; Maroulakou and Bowe 2000; Oikawa and Yamada 2003). Most

ETS family transcription factors are ubiquitously expressed. Gene disruption studies involving most Ets factors results in embryonic or perinatal lethality implying their importance in early development (Bartel et al. 2000; Oikawa and Yamada 2003). The majority of Ets factors act as transcription activators downstream of various signaling pathways. However, at least five members have been identified to act as repressors including Erf, PE1/METS, Elk3/Net, TEL, and TEL2 (Mavrothalassitis and Ghysdael

2000; Gu et al. 2001; Klappacher et al. 2002). Additionally, few members perform as both activator and represseor in a context dependent manner (Sharrocks 2001).

1.2.3.1. Ets Factors in Embryonic Development

ETS-domain proteins contribute to a variety of embryonic development processes. Mice deficient of a functional copy of Ets protein(s) often leads to embryonic lethality or developmental abnormality. Loss of PU.1 in mice results in defective lymphoid cells, especially in B cells and absence of mature macrophages and (McKercher et al., 1996;

Scott et al., 1994). Ablation of Spi-B leads to increased apoptosis of poorly differentiated

B cells (Su et al., 1997). Through mouse chimeras with TEL-/- ES cells, TEL was shown to be essential for the establishment of hematopoiesis of all lineages in the bone marrow

(Wang et al. 1998a). Fli1 knockout mice are lethal at embryonic day 11.5 exhibiting severe hemorrhage in the brain (Hart et al., 2000; Spyropoulos et al., 2000). Mice lacking

Ets1 are viable and fertile, however they show defective B-cell and T-cell apoptosis

(Muthusamy et al. 1995; Barton et al. 1998). Deletion of Ets2 causes embryonic lethality at E8.5. The mutant mice show multiple defects including small ectoplacental cone region and absence of amnion or chorion membranes. However, since the lethality is due to extra embryonic defects, the mice can be rescued by aggregation with tetraploid mouse embryos (Yamamoto et al. 1998).

Several Ets family proteins are involved in lineage development and cell differentiation.

Hematopoietic cell express several Ets proteins, which regulates development and differentiation of the hematopoietic cells. Fli1 regulates the expression of Tie-2/Tek gene, which encodes the receptor of angiopoitin-1 required for proper angiogenesis (Hart et al.,

2000). TEL is also essential for extra embryonic angiogenesis (Wang et al., 1997). AML1 and PU1 regulate expression vav gene required for transition from primitive to definitive hematopoiesis (Okada et al., 1998). PU.1 regulate early myeloid cell differentiation. PU.1 binding sites are found in the promoter regions of several myeloid specific genes including macrophage colony stimulating factor (M-CSF) receptor (c-fms), granulocyte- macrophage colony stimulating factor (GM-CSF) receptor, scavenger receptor, CD11b/

CD18 (Mac-1) among others (Friedman, 2002; Oikawa et al., 1999). Multiple Ets family proteins are expressed in lymphoid cells such as Ets-1, Ets-2, Erg, Fli-1, Elf-1, GABPa,

PU.1 and Spi-B (Anderson et al., 1999). PU.1 seemingly regulates the expression of many B-cell specific genes. Knockout of PU.1 completely abolishes B-cell and macrophage population. Furthermore T-cell development is also delayed in absence of

PU.1 (Spain et al., 1999). Ets proteins, such as ETS1, ETS2, PEA3, ELK1, and GABPA

bind to the control regions of the neural genes such as the Synapsin II (Petersohn et al.,

1995), peripherin (Chang and Thompson, 1996) important for neuronal development.

ETS1 and ETS2 have also been reported to play an important role in osteoblast development and bone formation (Raouf and Seth, 2000). PU.1 collaborate with microphthalmia-associated transcription factor (MITF) to increase target gene expression in osteoclasts (HU et al., 2007). Several Ets factors are also thought to play a crucial role in vascular development as discussed in later chapters.

1.2.3.1. Ets Proteins in Cancer

Deregulation of Ets factors has emerged to be a driving force in neoplastic transformation, metastasis and progression (Seth and Watson, 2005). During cancer progression, ETS genes acquire various mutations leading to altered expression, which affects the regulation of several biological processes. The dysregulation leads to enhancing cell proliferation and inhibiting apoptosis, stimulating cell migration, invasiveness, and angiogenesis. Expression of PEA3 and ETS2 correlates with advancing hepatocellular carcinoma (Iguchi et al., 2000; Ito et al., 2002). Over expression of Ets proteins, such as FLI1, ELF1, PDEF, ETS1, and ETS2, are associated with advanced stages of prostate cancer (Gavrilov et al., 2001). The upregulation of extracellular matrix- degrading proteins including MMP-1, MMP-9, uPA, and the uPA receptor are mediated by enhanced expression of these ETS proteins. The ETS1 and ETS2 are overexpressed in thyroid neoplasia (de Nigris et al., 2000). In breast cancer cells, PEA3, ESE-1/ ESX and

Elf-1 enhance erbB2/HER-2/neu promoter acitivity (Scott et al., 2000).

Development of a tumor depends largely on the surrounding microenvironment. ETS factors act as important modulators of tumor microenvironment. Fibroblasts are an important component of the tumor stroma in that it regulates ECM remodeling, epithelial cell differentiation and angiogenesis (Kalluri and Zeisberg, 2006). Deletion of

Phosphatase and tensin homolog (PTEN) in stromal fibroblasts increased ETS2 expression MMTV-ErbB2 mammary tumor model exhibiting accelerated tumor growth, expanded ECM and increased macrophage infiltration (Trimboli et al., 2009). As stated earlier, Ets2 deletion in fibroblast reduces primary tumor burden. In contrast, macrophage specific deletion of Ets2 decreases frequency of lung metastases (Zabuawala et al., 2010).

CHAPTER 2

MATERIALS AND METHODS

2.1. Animal Husbandry

2.1.1. Transgenic Mice used

The Ets-1 knockout mice were provided by Dr. Muthusamy (The Ohio State University,

Columbus, OH) and described in Muthusamy et al., 1995. The conditional Ets1fl transgenic line was generated in our lab (strategy described in Chapter 3). The conditional Ets2fl transgenic line was generated in our lab (Wei et al, 2009.). A tamoxifen inducible Tie2-Cre-ERTam (Forde et al., 2002) was used to induce Cre expression only in the endothelial cells of post natal mice. The Ets2db transgenic strain was a kind gift from

Dr. Oshima (The Burnham Institute, La Jolla, California). Ets1fl/fl; Ets2fl/fl mice were bred

>10 generations into the FVB/N background. All other animals were maintained on a mixed 129/Sv × FVB/N background. Rosa-loxP transgenic line was purchased from

Jackson laboratory.

2.1.2. Animal Care

Transgenic mice used in the present study were housed in Biomedical Research Tower animal facility at The Ohio State University, Columbus, OH according to the NIH guidelines. All the mice were sacrificed according to animal resources standard operating procedures. Use and care of mice in this study were approved by the Ohio State

University Institutional Animal Care and Use Committee.

2.1.3. Mouse Genotyping

2.1.3.1. Tail DNA Preparation

Approximately 0.2-0.5cm of mouse tail was digested in 200ul of tail lysis buffer (50 mM

KCl, 10mM Tris-HCl (pH8.3), 0.1 mg/ml gelatin, 0.45% NP40, 0.45% Tween20, 1mg/ml

ProteinaseK), 55°C overnight (O/N). The samples were then boiled for 12-15‘ to inactivate ProteinaseK and cooled down on ice. Tail DNA samples were stored at 4oC.

2.1.3.2. Genotyping Primers and PCR Conditions

2μl of each tail DNA sample was used in a 20μl PCR reaction. The PCR program used for genotyping was:

1 cycle at 95°C for 2‘→ 35 cycles of: 95°C (45‘‘), appropriate annealing temperature as listed in Table 2.1. (45‘‘), 72°C (1‘) → 1 cycle of 72°C (10‘). The PCR products were run on a 1.2% agarose gel.

Gene Primer ID Sequence Annealing Temperatures °C ETS1I4P2 CAACAACAGCAAGAGCATCC

ETS1E4P2 ACTGTGTGCCCTGGGTAAAG

Ets1 PGKP1 CTAAAGCGCATGCTCCAGACTGCC 55

ETS1 CKO WT CTCTCATTTGCCATCTTTAGC

ETS1 CKO FL GTTTGTTTGTTTGTTTGTTTGTTC

Ets1fl ETS1 CKO KN AACACAATATAACATCTTTTT 50

ETS2-FLOX-R GGAAGAAACGGGAAATCAAA

ETS2-

COMMON-F TGAACTACTGTGTGTGACGAGGA

Ets2 ETS2-KO-R GGATTTTAGCCCAGAAACTTAGA 58

EN3 AATGACAAGACGCTGGGCGG

Ets2db E163 CGTCCCTACTGGATGTACAGCGG 58

YZ101 CCCTGTGCTCAGACAGAAATGAGA

Tie-2 Cre YZ98 CGCATAACCAGTGAAACAGCATTGC 58

Cre WCRE1 CCTGTTTTGCACGTTCACCG

(generic) WCRE3 ATGCTTCTGTCCGTTTGCCG 58

OIMR0316 GGAGCGGGAGAAATGGATATG

OIMR0315 GCGAAGAGTTTGTCCTCAACC

Rosa-loxp OIMR0883 AAAGTCGCTCTGAGTTGTTAT 58

Table 2.1. Primers Used for Genotyping Mice and Cells

2.1.4. Animal Procedure

2.1.4.1. Induction of Gene Deletion

Stock solution of (10 mg ml−1) of tamoxifen was prepared by dissolving 10 mg of tamoxifen free base in a 1.5-ml tube in 250 μl of 100% (vol/vol) ethanol by vortexing at maximum speed until tamoxifen completely dissolves. 750 μl of peanut oil was added and vortex thoroughly to obtain an emulsion.

To induce gene knockout in pups, tamoxifen (diluted 1:10 with peanut oil at the day of injection) was administered using a 1-ml syringe with a 29G hypodermic needle into the stomach of control and experimental littermates (50 μg per day) for 3 days from postnatal day 1 (P1) to P3.

For adult mice, 2 mg of tamoxifen was injected injected intraperitoneally for 5 consecutive days. The mice were also put on a tamoxifen diet after the injections for the course of the experiment.

2.1.4.2. Eye and Retina Dissection

Retinae were harvested from P8 littermates. The pups were sacrificed in accordance with the standard operating procedures. Small blade scissors were used to dissect out eyes out and rinsed in cold phosphate-buffered saline (PBS), and fixed in 4% PFA/PBS

(paraformaldehyde). Retinae were isolated from the fixed eyes with spring scissors under a dissection scope.

2. 1.4.3. Subcutaneous Matrigel Plug Injection

10 week old mice were anesthetized using isoflourane. The mouse was positioned facing downwards. The flank region was cleaned with 70% ethanol and 350ul of ice cold matrigel was injected slowly just below the skin using a 28 ½ G needle.

2.2. Tissue Culture and Viral Infections

2.2.1. Primary Cell Extraction and Culture

2.2.1.1. Aortic Endothelial Cells

Mouse aortic endothelial cells were isolates as described in Huang et al, 2003. Thoracic aortas were isolated from anesthetized mice (3-5 weeks) and cut longitudinally into 1-to

2mm2 pieces. After cleaning the supporting fat tissue around the aorta, the vessel was opened longitudinally and cut into small explants. Four to six explants were placed in fibronectin (50μg/ml in PBS→ 2hrs at 37°C→PBS wash→ stored at 4°C) coated dishes with the inner vessel side down. A small volume of complete EC culture media (DMEM-

F12+20% heat-inactivated FBS+penicillin-streptomycin (PS) +30μg/ml endothelial cell growth supplement (ECGS, Upstate) +10U/ml heparin (Sigma)) was added to the dishes and incubated in 37°C incubator at 5% CO2. Migrating EC were observed within 2-3d.

Explants were removed and discarding when cells were almost confluent (usually by day7). The cells were then trypinized and cultured in complete EC media. Cells from passage 3 were trypsinized and plated into 2-75cm2 and 1-25cm2 flasks. When cells reached ~ 80% confluency they were incubated with 5μg di-I-acetylated-LDL/ml

(Biomedical Technologies, Inc.) O/N. The 25cm2 flask without di-I-Ac-LDL was used as autoflurorescent blank for FACS. For sorting, the cells were trypsinized, resuspended in

FACS buffer (S-MEM lacking Ca2+ + 0.5% BSA+ PS) at a concentration of

1X106cells/ml and sorted. The positively stained population was collected and cultured in complete EC medium.

2.2.2. Lentiviral Infections

5 X 105 EC were cultured O/N and infected with ecotropic lentivirus (pHAGE-IRES-GFP vectors with or without PGK-Cre, kind gift from Dr. Nika Danial at Harvard University) containing 30μg/ml ECGS and 4μg/ml Polybrene. Cells containing lentivirus were centrifuged at 1600rpm for 1hr at RT to facilitate virus infection and incubated at 37°C,

5% CO2 O/N. ~12-14hrs post infection, viral media was removed, and cells were rinsed in PBS. Cells were cultured in fresh complete EC media for 24hrs passaged further.

2.3. Microarray and Data Analysis

Endothelial cells used in this study were isolated from three independent sets of WT,

Ets1-/-;Ets2fl/fl and Ets2fl/fl mice and infected with lentiviral GFP with or without Cre. RNA harvested from control and DKO EC was subjected to gene expression profiling using the

Affymetrix GeneChip Mouse Genome 430 2.0 array. Background correction and quantile normalization was performed to adjust technical bias, and gene expression levels were summarized by RMA method (Irizarry et al., 2003). A filtering method based on percentage of arrays above noise cutoff was applied to filter out low expression genes. A linear model was employed to detect differentially expressed genes. In order to improve

the estimates of variability and statistical tests for differential expression, a variance smoothing method with fully moderated t-statistic was employed for this study (Yu et al.,

2011). The expression data was analyzed by Gene Set Enrichment Analysis (GSEA) as described previously (Subramanian 2005) with gene sets obtained from indicated categories in Toppgene suite (Chen 2007).

2.4. cDNA Preparation

2.4.1. RNA Extraction

EC were resuspended in 1ml of Trizol (Invitrogen) and stored 2-500μl aliquots in RNAse free microfuge tubes at -80oC until use. RNA was extracted according to manufacturer‘s instructions. The RNA pellet was dissolved in 50μl of RNase-free DEPC water for 20 minutes at 55°C. RNA was quantified by OD260, and the purity of was determined by the ratio of OD260 to OD280 (>1.8).

2.4.2. Reverse Transcription

2μg of purified total RNA was reverse transcribed by Superscript III reverse transcriptase

(Invitrogen) with random hexamer primers (Invitrogen). Briefly, RNase-free water and

0.2μg random primers added to RNA sample was heated to 65°C for 5 min and incubated on ice for at least 5min. 5μl 5X First stand buffer (Invitrogen), 1μl 0.1M DTT

(Invitrogen), 10μM dNTPs (Roche), 1μl Superscript III and 1μl RNAse OUT (Invitrogen) were added to the reaction. The RT reaction was carried out in a thermocyler at 25°C for

10 min to allow primers to anneal to RNA, then at 50°C for 60min for reverse transcription, and at 70°C for 15min to inactivate the reaction. The cDNA sample was

diluted to 100μl using. 2μl of each cDNA sample was used for each Real-Time PCR reaction.

2.4.3. Primers used for real-time PCR

Primers for RNA expression analysis were designed using Roche Universal Probe

Library System. For additional specificity, intron-spanning primers were designed. The primer sets used for analysis are listed in Table 2.3.

Gene Left Primer Right Primer Probe #

Anapc5 TGCCGCTTCGGTCACTAT GGCAATCCTAATTGCCTCCT 13

Aspm TGTCTGCCAGAGGGATCAGT CTCTGTGGCTTCGAATTGGT 94

Bcl2l1 CCAGCACTGGGTCCTAAGAG CCAGCACTGGGTCCTAAGAG 26

Birc3 TCTGAACGAGTAAATGGAAGGTT GGTATAGGGCGTCTTGGAAA 6

Birc5 CCGATGACAACCCGATAGA CATCTGCTTCTTGACAGTGAGG 43

Birc6 GTCCCCCAGATACAAGTGACA CGGGTTCCCATTATGTTCC 1

Cdc25c CTGAGCTTGCCTGACGTCTAT CACTTGCAGGTGGGATAGGT 10

E2f7 GGACCACAGCAAATTCAAAAC CGCTCTTCTGTTACGTGAACTTT 21

E2f8 TGGAAGATCTGGATAAAAGCAAG CCAAGCTACTCAGGACATTAGCA 75

Fzr1 CTCGGACAATGGCAAAGAC GGTCCTGAACCTTCTCGATG 3

Mad2l1 CTGACCCCGAGCTCATAAAG ACTGAGCACTTGTACAGCCACT 31

Mad2l1 GCGCCAAGACCTCAACTTT CTCGCGCACATAGAGAATCA 10

Mcl1 GAGCCTGTTGCAGAGACCTT GAGCCTGTTGCAGAGACCTT 2 Mmp3 GATCTAGGTGATGTGCCAGATG CCAAGCTTGCAGAGGTAAGG 52 Mmp9 CTAGGTGAATGCCCCATCC GCAAACTGCAGACTTTGTGG 12

Ranbp1 GGAGGAAGATGAAGAGGAACTTT CAGAAGCTTGACATCTCCAGTG 12

Sphk1 AGCTCTTCCAGAGCCGTGT GCATGGTTCTTCCGTTCG 19

Ube2c GGTGAAACCAAGGAGAAACG GGTGAAACCAAGGAGAAACG 17

Table 2.3. UPL Primers Used for cDNA Quantitative Realtime-PCR (qRT-PCR)

2.5. Chromatin Immunoprecipitation Assay (ChIP)

2.5.1. Chromatin Preparation

ChIP assays were performed as described previously (Hu et al., 2007). Briefly, wild type aortic EC were cultured O/N at a density of 3 X 106 cells per 10-cm dish. Cells were cross-linked with formaldehyde at a final concentration of 1% at 37°C for 10 min before harvest. Soluble chromatin was prepared following sonication with a Branson

250 digital sonifier (Branson Ultrasonics, Danbury, CT) to an average DNA length of

200-1000 bp and precleared with tRNA-blocked Protein G-agarose. ~3X105-cell equivalent of the precleared chromatin was immunoprecipitated with 5 μg anti-ETS1,

20 μg anti-ETS2 antibodies, characterized previously (Wei et al., 2009) and 20 μl anti-

Acetyl-CBP (Lys1535)/p300 (Lys1499) Antibody (Cell Signaling). 10% of the precleared chromatin was set aside as input control. Immunoprecipitation was carried out with overnight at 4 °C. Immune complexes were pulled down using Protein G-agarose, washed, and eluted twice with 250 μl of elution buffer (0.1M NaHCO3, 1%SDS), and reverse cross-linked in 200mM NaCl at 65 °C overnight with 20μg of RNase A

(Sigma). DNA was purified following ProteinaseK treatment (Invitrogen) with the

Qiagen PCR purification kit using the manufacturer's instructions.

2.5.2. Primers used for real-time PCR

Samples were analyzed by real-time PCR using the Roche universal probe library

(Roche Diagnostics, Indianapolis, IN). The primer sets used for analysis are listed in

Table 2.3..

2.5.2. ChIP-Sequencing and Analysis

ChIP-sequencing was performed to combine chromatin immunoprecipitation (ChIP) with high throughput DNA sequencing to identify the binding sites of nuclear proteins in a genome wide manner. Wild type cultured aortic ECS were used to generate the immunoprecipitated and input control libraries using TruSeq DNA Sample Prep Kit v2

(Illumina) following manufacturer‘s direction. The libraries were sequenced on a

HiSeq 2000 system (Illumina).

Resultant sequence reads were mapped to the mm10 assembly of the mouse genome with Bowtie. Duplicate reads were removed, and the remaining unique reads were normalized to for peak calling analysis. Peak calling was performed using HOMER

(Heinz 2010).

Probe Gene Left Primer Right Primer #

Anapc5 TGCCGCTTCGGTCACTAT GGCAATCCTAATTGCCTCCT 13

Aspm TGTCTGCCAGAGGGATCAGT CTCTGTGGCTTCGAATTGGT 94

Bcl2l1 CCAGCACTGGGTCCTAAGAG CCAGCACTGGGTCCTAAGAG 26

Birc3 TCTGAACGAGTAAATGGAAGGTT GGTATAGGGCGTCTTGGAAA 6

Birc5 CCGATGACAACCCGATAGA CATCTGCTTCTTGACAGTGAGG 43

Birc6 GTCCCCCAGATACAAGTGACA CGGGTTCCCATTATGTTCC 1

Cdc25c CTGAGCTTGCCTGACGTCTAT CACTTGCAGGTGGGATAGGT 10

E2f7 GGACCACAGCAAATTCAAAAC CGCTCTTCTGTTACGTGAACTTT 21

E2f8 TGGAAGATCTGGATAAAAGCAAG CCAAGCTACTCAGGACATTAGCA 75

Fzr1 CTCGGACAATGGCAAAGAC GGTCCTGAACCTTCTCGATG 3

Mad2l1 CTGACCCCGAGCTCATAAAG ACTGAGCACTTGTACAGCCACT 31

Mad2l1 GCGCCAAGACCTCAACTTT CTCGCGCACATAGAGAATCA 10

Mcl1 GAGCCTGTTGCAGAGACCTT GAGCCTGTTGCAGAGACCTT 2 Mmp3 GATCTAGGTGATGTGCCAGATG CCAAGCTTGCAGAGGTAAGG 52 Mmp9 CTAGGTGAATGCCCCATCC GCAAACTGCAGACTTTGTGG 12

Ranbp1 GGAGGAAGATGAAGAGGAACTTT CAGAAGCTTGACATCTCCAGTG 12

Sphk1 AGCTCTTCCAGAGCCGTGT GCATGGTTCTTCCGTTCG 19

Ube2c GGTGAAACCAAGGAGAAACG GGTGAAACCAAGGAGAAACG 17

Table 2.4. UPL Primers Used for ChIP

2.6. Quantitative Real-Time PCR (qRT-PCR)

2.6.1. qRT-PCR Reaction Conditions

Each qRT-PCR reaction was set up with 2μl of cDNA (for RNA expression analysis) or

6μl of chromatin (for ChIP analysis) with 10μl of FastStart Universal Probe Library

Master Mix (Roche), 0.1mM of corresponding primer set and 50nM of appropriate probe.

The final 20μl reaction was carried out in Icycler iQ Real-Time Detection system

(BioRad) according to manufacturer‘s instructions. The PCR protocol used was:

1 cycle of 95oC (5‘) → _39 cycles of: 95oC (30‖), 54oC (30‖), 72oC (30‖) → _1 cycle of

72oC (5‘) → _12⁰C (∞).

2.6.2. qRT-PCR Analysis

For RNA expression analysis, the threshold of gene being studied was adjusted by that of a reference ribosomal protein L4 gene. For ChIP analysis, the threshold for the specific gene was adjusted by that of the input. ChIP PCR products were run and analyzed on a 2.5% agarose gel.

2.7. Western Blot Analysis

2.7.1. Protein Isolation

Cells were scraped in modified RIPA buffer (50mM Tris-HCl (pH7.4) + 1% NP-40 +

0.25% Na-deoxycholate `+ 150mM NaCl + 1mM EDTA + 1mM PMSF + 1ugml each of

Aprotinin, Leupeptin, Antipain + 1mM each of PMSF and Na3VO4 ) and lysed on ice for

20‘. The cell suspension was transferred on a microfuge tube and 14,000g at 4⁰C for 20‘.

The supernatant was carefully removed, aliquoted and stored at -80⁰C until use.

2.7.2. Western Blot

Protein concentration was measured by Bradford assay. Samples of equal protein concentration were run on 8-10% SDS-Polyacrylamide gels. The proteins were transferred onto nitrocellulose membrane. The membrane was blocked (5% non-fat dry milk or 5% BSA in 0.05%TBST), incubated with primary antibody O/N at 4⁰C, washed in TBST and incubated with appropriate HRP conjugated secondary antibody for 1hr at

RT. The membrane was then developed using the ECL chemiluminescence system

(Thermo Scientific).

2.8. Cell Based Assays

2.8.1. BrDU Proliferation Assay

Cells were incubated with 3μg/ml BrDU (Sigma) for 2hrs at 37⁰C/5% CO2. The cells were then fixed and BrDU staining was performed as previously described (Braren et al.,

2006). Proliferation was expressed as %BrDU positive to total cells.

2.8.2. Apoptosis Assay

Apoptosis was induced in cultured ECs through serum startvation (0.1% serum, 24 h).

After the induction, both adherent and non adherent cells were harvested, pelleted, washed and pelleted again. Subsequently cells were subjected to PI (Roche) staining or

Annexin V-Pacific Blue/7-AAD (invitrogen/ebioscience) staining followed by fluorescent activated cell sorting (FACS) to assay apoptosis. Labeled cells were analyzed

using a BD LSR II Flow Cytometer. Cells demonstrating DNA content less than that of

G0/G1-phase cells were considered to be apoptotic (sub-G0). For Annexin V-Pacific

Blue/7-AAD staining, cells positive for annexin V were considered to be apoptotic.

2.8.3. Migration Assays

2.8.3.1. Scratch Wound/Wound Healing Assay

EC were cultured to confluent monolayers and incubated with 15ng/ml MitomycinC

(Sigma) for 3hrs at 37⁰C. The cells were then wounded across the well with a 10-μl standard pipette tip, rinsed with PBS to removed floating cells and complete EC media was added. The wounded monolayers were then imaged over 24hrs under the Live

Imaging Zeiss microscope. The wound healing effect was calculated as the number of EC migrating into the wound over different time points.

2.8.3.2. Single Cell Migration Track Assay

24-well dishes were coated with 5μg/mL fibronectin (Sigma) and then overlaid with carboxylate-modified polystyrene fluorescent microspheres (Invitrogen). Treated or untreated control cells were then seeded at low density (4/mm2) in normal growth medium and incubated for a period of 24 h. The ability of the cells to create nonfluorescent tracks was quantified and represented as Average Area Migrated/Cell.

2.8.3.3. Matrigel Tube Formation Assay

The vessel-forming ability of EC was characterized in vitro using a matrigel assay. 110μl

Growth Factor Reduced Matrigel (BD Biosciences) was added per well of a 48-well dish and allowed to solidify at 37⁰C. 4 X 104 EC were then added to the matrigel layer and

allowed to attach for 2hrs at 37⁰C. Complete media was added and cells were maintained in 37⁰C incubator at 5%CO2. Development of vessels was monitored every 4 h for 24 h.

2.9. Histology and Immunostaining

For matrigel plugs, 5 μm sections were prepared from harvested frozen plugs to perform immunostaining. Cultured aortic ECs plated on chamber slides were used for phospho- histone-H3 staining. Primary antibodies used were rat α-mouse CD31 (1:50 dilution; BD

Biosciences) and rabbit α-mouse phospho-Histone H3 (Ser10) (1:100 dilution;

Millipore). Alexa fluor 488 and 594 conjugated secondary donkey α-rat or donkey α- rabbit antibodies (1:250 dilution; Invitrogen) were used for fluorescent detection.

2.10. Imaging and Quantification

A Nikon Eclipse E800 epifluorescence microscope equipped with a Photometrics

Coolsnap camera and Nikon Plan Fluor 4X (N.A. 0.13), 10X (N.A. 0.30), 20X (N.A.

0.50), 40x (N.A. 0.75) and 100x (oil N.A. 1.3) objectives was used to acquire immunofluorescent images. MetaVue software from Molecular Devices was used for image acquisition. An Olympus FV1000 Filter Confocal system using a UPLFLN 40x oil objective (N.A. 1.3) was utilized for confocal microscopy. All images were acquired at room temperature. For retinal vasculature, extent of migration was measured as the radial distance between the optic nerve and the edge of the vasculature towards the retinal periphery. Branch points as well as fiolopodia/sprout number were counted using FIJI

(Johannes 2012). Blood vessel size and branching (matrigel plug sections) were measured

using the ‗connected region‘ plugin and the ‗Analyze skeleton‘ plugin respectively in

FIJI.

CHAPTER 3

ETS1 AND ETS2 IN ACTIVATION PHASE OF ANGIOGENESIS

3.1. Introduction

Angiogenesis is the orchestrated vascular remodeling to form new blood vessels from an existing vasculature. Proper execution of this highly complex process requires intricate balance between the angiogenic facilitators and inhibitors. Abnormal vascular development leads to severe pathological condition. Apart from the endothelial cells and the pericytes, which physically form the blood vasculature, other cells, such as fibroblasts, tumor/epithelial cells and immune cells, as well as the extra cellular matrix, regulate angiogenesis. Throughout angiogenesis, several signaling pathways in ECs are stimulated and inhibited by the growth factors released by the cells and EC-ECM adhesion. VEGF appears to be the most potent regulator of angiogenesis being able to modulate several downstream effector pathways. The RAS-MEK-ERK pathway is one of the central signaling pathways downstream of VEGF. Murine knockout models of genes involved in this pathway like B-Raf, Mek1 or Ras-GAP, leads to vascular defects during

embryogenesis (Galabova-Kovacs et al., 2006; Giroux et al., 1999; Henkemeyer et al.,

1995). Being downstream of RAS pathway, Ets factors contribute significantly to regulate cellular processes including cell proliferation, migration and survival required for vascular development.

3.1.1. Ets Factors in Angiogenesis

Several Ets factors including ETS1, FLI1, ERG and TEL1 have been reported to be expressed in the ECs and its precursors. Ets1 regulates expression of Flt1 (VEGFR2), receptor for VEGF (Oikawa and Yamada, 2003). Tie-2/Tek encodes ANG1 reeptor.

Expression of Tie-2/Tek is mediated by Fli1 (Hart et al., 2000). Fli1-/- mice die at E12.5 with defective hematopoiesis. TEL is also essential for vascular development in yolk sac

(Wang et al., 1997). Ablation of Etv2 (Ets Variant 2) results in defective vasculogenesis with significantly reduced number of ECs. Etv2-/- embryos die by E9.5 due to loss of EC- specification. In contrast, when ectopically expressed Etv2 lead to increased blood vessel formation (Lee et al., 2008; Sumanas et al., 2008). Recent studies from our lab demonstrated a significant role of Ets1 and Ets2 in promoting embryonic angiogenesis.

Targeted deletion of Ets2 in EC in conjunction with global Ets1 knockout renders embryonic lethality. The mutant mice show severe vascular defects and enhanced EC apoptosis (Srinivasan et al., 2009). Taken together, these data suggest that Ets factors modulate vascular development temporally in that Fli1 regulates hematopoietic development whereas Etv2 in EC specification and Ets1 and Ets2 in angiogenesis proper.

3.1.2. Retinal Angiogenesis in Mice

In human, an infant is born with completely formed retinal vessels and with regressed hyaloid vasculature. In contrast, mice are born with immature retinal vasculature with persistent hyaloid vessels (Gyllensten LJ and Hellstrom BE, 1954). Retinal vascularization begins in the most superficial (or inner) retinal layers that lie within the ganglion cell layer (Saint-Geniez and D'Amore. 2004). The vasculature grows in radial fashion starting from the central point where optic nerve is located. In human it reaches the periphery just before birth. However, in mice the same is attained in about one week

(Figure 3.1.). The newly formed inner vascular layer then gives rise to vessels formed in the deeper layers. Although other processes of vascularization occur during the development or retinal vasculature, sprouting angiogenesis is the dominant mechanism

(Gariano and Gardner, 2005).

The blood vessel growth in retina has been proposed to be driven by oxygen availability.

Early studies suggested that the expression VEGF in retina is highly driven by hypoxia

(Chan-Ling et al., 1995). Retinal VEGF expression is regulated both spatially and temporally. Two types of microglia, the astrocytes and Müller cells, play a pivotal role in the development of the superficial and deep vascular layers, respectively. VEGF is sequentially expressed by these microglia. In the ganglion cell layer of the retina, the astrocytes act as the source of VEGF, however in the in the internal nuclear layer VEGF expression is carried out by the Müller cells (Stone, J. et al., 1995). As the angiogenic front moves outward, increased VEGF supply behind the front reduces VEGF production to inhibit excessive vessel formation (Saint-Geniez and D'Amore. 2004). Despite, the

strong evidence suggesting an essential role of VEGF in retinal angiogenesis, the specific functions of individual isoforms are yet to be delineated.

Since retinal vascular anatomy is easily visualized, murine retinal angiogenesis has been utilized extensively to study both pathological and physiological angiogenesis. Ease of accessibility of retinal vasculature for imaging remains one of the major advantages.

Since postnatal retinal vascular development is very tightly regulated, it enables reliable detection of any developmental abnormality attributed by gene interruption studies

(Dorrell et al, 2007). Since retinal angiogenesis recapitulates, capsular development in other tissues, mechanism of vessel formation in eye has been utilized to attain valuable insight applicable to general developmental angiogenesis and tumor angiogenis (Rennel et al., 2009; Loges et al., 2009).

Figure 3.1. Development of The Superficial Vascular Plexus in Mouse Retinas

Retinal whole mounts from postnatal day (P) 1 to P8 were stained for endothelial cells with isolectin B4 594 (red). Adpated from Stahl A., 2010

3.1.3. Transgenic Alleles Used

An inducible Cre/loxp system was utilized in junction with floxed alleles of Ets1 (Ets1fl) and Ets2 (Et21fl) to generate transgenic mice with tissue specific deletion of the respective gene. Additionally conventional knockouts of Ets1 (Ets1-) and Ets2 (Ets2db) were also used in this study.

Conditional Ets1 allele: Ets1fl

Mice mutant for both Ets1 and Ets2 are embryonic lethal (Wei et al., 2009). The double mutant mice exhibit defects in vessel branching and reduced vascular complexity. Use of a conditional Ets2 allele containing loxP sites, in combination with global Ets1 knockout and tie-2 cre attributed this defect to EC. However, whether this defect is truly endothelial cell autonomous was uncertain as the Ets1 gene was knocked out globally.

Therefore, a conditional gene targeting strategy was used to enable us to delete Ets1 specifically in ECs either alone or in combination with Ets2. The Ets1fl harbors two loxp sites flanking exon 7 and exon 8 (Figure 3.2.). In presence of Cre recombinase, the loxp site will be rearranged generating an Ets1 allele without the DNA binding domain

(Dittmer, 2003).

Conditional Ets2 allele: Ets2fl

Mice homozygous for the Ets2db allele (deletion of the Ets domain of ets-2) (Yamamoto et al. 1998) are embryonic lethal. The lethality prevents the study the effect of Ets2 on later developmental stages. Moreover, since the gene is knocked out globally, cell specific effect of the deletion on individual processes are hard to dissect. To circumvent this problem a conditional gene targeting strategy was used to generate the conditional

Ets2 allele (Wei et al., 2009). In Ets2fl allele, the exons 3-5 are flanked by the two loxp sites. The pointed domain of Ets-2 is encoded by exon 4 and 5. Thus the deletion of Ets2 exons 3-5 will delete the conserved MAPK phosphorylation site and the pointed domain.

Conventional Ets1 null allele: Ets1-

Ets1 conventional knock out was generated by inserting a neomycin transferase (neo) cassette encompassing exons 3 and 4. The mutant Ets1-/- gene is a null allele. Ets1-/- mice are viable and fertile (Muthusamy et al., 1995). However, they displayed an increased perinatal mortality.

Conventional Ets2 knockout allele: Ets2db

Ets2db/db mice are embryonic lethal before E8.5 from extraembryonic defects (Yamamoto et al. 1998). In the knockout allele, exons 8 to 10, encoding the DNA binding domain, is deleted. The Ets2db knockout allele leads to production of a C-terminal truncated product that lacks nuclear localization signals and DNA binding activity. However, the exons upstream of the Ets domain are fused to the neo transcript thus results in the deletion of a critical portion of the gene but does not in a null allele.

Inducible Cre Transgenic Lines

Transgenic mice expressing Cre-recombinase in specific cells were used to target the conditional alleles in a tissue specific form. To add temporal switch to for cre recombination a tamoxifen inducible Tie2-Cre-ERTam (Forde et al., 2002) was used to induce Cre expression only in the endothelial cells of postnatal mice. In Tie2-Cre transgenic lines, the Cre-recombinase cDNA is inserted downstream of regulatory element of the EC specific murine Tie2 promoter/enhancer region. Rosa-loxP reporter

mediated analysis of the Tie2-Cre transgenic mice demonstrated a pan-endothelial- specific pattern of lacZ staining throughout embryogenesis and adulthood (Kisanuki et al., 2001).

Figure 3.2. Targeting Strategy for Ets1fl Allele

The positions of the PCR primers (arrow head) used to distinguish various alleles are indicated. The dotted line represent a deletion introduced. Cre mediated recombination results in Ets1KO allele without the exons 7 and 8.

3.2. Results

3.2.1. Tie-2-Cre Efficiently Recombines Ets1/2fl Alleles Specifically in EC

As discussed in previous section, a tamoxifen inducible Tie-2cre was utilized to achieve in vivo deletion of Ets1 and Ets2. To determine the efficiency of Cre mediated deletion, primary aortic ECs were isolated from Ets1fl/fl;Ets2fl/fl mice with or without Tie-2Cre and

WT mice. The purity of the isolate EC population was determined by CD-31 staining.

Cre mediated recombination was analyzed both at DNA and at RNA level. PCR genotyping of the genomic DNA isolated from the cells show efficient deletion of both

Ets1fl and Ets2fl genes (Figure 3.3. A). Similarly, comparison of RNA level isolated from both control and mutant cells by qPCR demonstrates robust deletion (Figure3.3. B)

Figure 3.3. Cre Recombinase Efficiently Deletes Ets1fl and Ets2fl Alleles In Vivo

(A) PCR genotyping of aortic ECs isolated from Ets1fl/fl;Ets2fl/fl mice with or without Tie-2Cre and WT mice demonstrating WT, floxed and knockout bands for Ets1 and Ets2 (B) qPCR analysis of Ets1 and Ets2 mRNA harvested from aortic ECs isolate from Ets1fl/fl;Ets2fl/fl mice with or without tie-2Cre.

3.2.2. Tie-2-Cre; Ets1fl/Fl;Ets2fl/Fl Display Defective Retinal Angiogenesis

Previous work from our group demonstrated that mice with genotype Tie2-Cre;Ets1-/-

;/Ets2fl/fl died during mid-gestation due to impaired angiogenesis (Wei et al., 2009). In order to better define the mechanisms underlying the function of the two ETS factors in angiogenesis, we studied the effect of conditional deletion of Ets1 and Ets2 during postnatal retinal angiogenesis, a process characterized by active vessel sprouting

(Garinaro and Gardner, 2005). For these experiments, we generated a conditional Ets1fl.

A tamoxifen inducible transgene Tie2-Cre-ERTam (Forde et al., 2002) was used to effect conditional deletion of the Ets1fl allele and the previously characterized Ets2 fl conditional allele.

To circumvent potential confounding results arising from incomplete recombination in double homozygous mice, we devised breeding strategies that, in addition to the double homozygous (DKO) mice would produce experimental progeny with at least one conventional knockout allele of either Ets1 or Ets2 (Table 3.1.). The breeding strategies would also allow for littermate controls containing at least one functional allele of Ets1 or

Ets2. The pups from the matings were injected with tamoxifen on postnatal days 1–3 to induce temporally regulated recombination of the conditional alleles (Pitulescu et al.,

2010). At postnatal day 8, the retinal vasculature of DKO mice and their littermate controls was visualized by whole mount isolectin B4 staining. This analysis, comparing 8 double-mutant embryos with 6 matched control littermates (Table 3.1.), demonstrated a marked reduction in the extent and complexity of angiogenesis in the DKO mice (Figure

3.4.). Retinal flat mounts demonstrated that the progression of the angiogenic front was

significantly impaired in DKO mice (Figure 3.4.A). While extensive blood vessel branching was apparent in retinas from control littermates at P8, the branching of vessels were decreased ~2.5-fold in DKO mice (Figure 3.4.E). Strikingly, the angiogenic sprouts in the retinas from DKO mice were abnormal, with both the number and length of filopodia significantly reduced compared to controls (Figure 3.4.F and 3.4.G). The phenotypes were identical whether the deletion of Ets1 and Ets2 was limited exclusively to EC (Figure 3.4.B top right panel and Figure 3.4.C top right panel) or were due to a combination of conditional alleles and conventional knockout alleles (Figure 3.4.C, compare the three panels). The presence of only one copy of either Ets1 or Ets2 was sufficient to retain a wild-type vasculature (Figure 3.4.A).

Figure 3.4. Conditional Deletion of Ets1 and Ets2 Attenuates Murine Retinal Angiogenesis (A) Isolectin B4-stained whole mount of entire retina from 8-day-old littermates showing progression of angiogenic fronts after tamoxifen treatment on postnatal days 1–3. n = 6 (control) n = 8 (DKO) (B) Isolectin B4-stained flat mount of single leaves demonstrating the complexity of branching. Top: comparison of Tie-2- cre;Ets1fl/fl;Ets2fl/fl mice with control littermate. The DKO genotype representing mice with conditional double deletion of Ets1/2 specifically in ECs. Bottom: control mice with genotype Tie-2-cre;Ets1fl/+;Ets2 fl/- representing mice with only one copy of Ets1. (C) Representation of sprout number and morphology. (D) Quantitaion of retinal area covered by blood vessels (radial outgrowth), expressed as percentage of control. n = 4 retinae. (E) Quantitation of branch points in a field of view. n = 4 retinae. (F) and (G) Quantitation of (F) filopodial number per sprout and (G) mean length of individual filopodia. n = 219 sprouts (control); n = 125 sprouts (DKO). Scale bars: 500 um (A); 100 um (B); 20 um (C).

Figure 3.4.

continued

Figure 3.4. continued

Littermate Control Experimental Parental Genotype ♂ ♀

1 Tie2-Cre;Ets1fl/+;Ets2+/- Tie2-Cre;Ets1fl/fl;Ets2fl/- Tie2-Cre;Ets1fl/+;Ets2fl/- Ets1fl/-;Ets2fl/+ Tie2-Cre;Ets1fl/-;Ets2fl/-

2 Tie2-Cre;Ets1fl/+;Ets2fl/+ Tie2-Cre;Ets1fl/-;Ets2fl/- Tie2-Cre;Ets1+/-;Ets2+/- Ets1fl/fl;Ets2fl/fl

3 Tie2-Cre;Ets1fl/+;Ets2fl/+ Tie2-Cre;Ets1fl/fl;Ets2fl/fl Tie2-Cre;Ets1fl/+;Ets2fl/+ Ets1fl/fl;Ets2fl/fl

4 Tie2-Cre;Ets1fl/+;Ets2fl/- Tie2-Cre;Ets1fl/-;Ets2fl/fl Tie2-Cre;Ets1+/-;Ets2fl/- Ets1fl/fl;Ets2fl/fl fl/- fl/-

84 Tie2-Cre;Ets1 ;Ets2

5 Tie2-Cre;Ets1fl/+;Ets2fl/fl Tie2-Cre;Ets1fl/-;Ets2fl/fl Tie2-Cre;Ets1+/-;Ets2fl/fl Ets1fl/fl;Ets2fl/fl

6 Tie2-Cre;Ets1fl/+;Ets2fl/+ Tie2-Cre;Ets1fl/fl;Ets2fl/- Tie2-Cre;Ets1fl/-;Ets2+/- Ets1fl/fl;Ets2fl/fl

Table 3.1. Breeding Strategy and The Genotypes of The Control and Experimental Mice Used for

Retinal Angiogenesis Assay

3.2.3. Tie-2-Cre; Ets1fl/Fl;Ets2fl/Fl Demonstrate Reduced Tumor Angiogenesis

To extend these observations, matrigel plug assays were performed to study the effect of the deletion of Ets1 and Ets2 in tumor angiogenesis. For this purpose, 10 week old

Tam fl/fl fl/fll inducible EC-DKO (Tie2-Cre-ER ;Ets1 ;Ets2 ) mice and controls (Table 3.2.) were treated with tamoxifen, and subsequently injected subcutaneously with matrigel plugs containing MVT1 mouse mammary tumor cells. Robust growth of new CD31-positive blood vessels into the matrigel plugs occurred in the control group (Figure 3.5.A). In contrast, both the number of vessels and vessel branching were significantly decreased in the DKO mice (Figure 3.5.B).

Figure 3.5 Ets1 and Ets2 are Required for Tumor Angiogenesis Progression

(A) Staining for CD31 in matrigel plugs: Matrigel, with 5x105 MVT1 cells, was subcutaneously injected into control and DKO mice and harvested after 10 days. Dashed lines indicate the outer margin of the plug. Region in the boxed area is magnified in the next panel of the corresponding genotype. (B) Graphical representation of CD31-positive vessels/field and branching. n =6 plugs. Scale bars: 100 um left most panel, 50 um right three panels (A).

Figure 3.5.

continued

Figure 3.5. continued

Littermate Control Experimental Parental Genotype

♂ ♀

fl/fl fl/fl 1 Tie2- Tie2- Tie2- Ets1 ;Ets2 Cre;Ets1fl/+;Ets2fl/fl Cre;Ets1fl/fl;Ets2fl/fl Cre;Ets1fl/+;Ets2fl/fl

2 Tie2- Tie2- Tie2-Cre;Ets1fl/+;Ets2fl/- Ets1fl/fl;Ets2fl/fl fl/+ fl/fl fl/fl fl/fl Cre;Ets1 ;Ets2 Cre;Ets1 ;Ets2

3 Tie2- Tie2- Tie2- Ets1fl/fl;Ets2fl/fl

89 Cre;Ets1fl/+;Ets2fl/+ Cre;Ets1fl/fl;Ets2fl/fl Cre;Ets1fl/+;Ets2fl/+

Table 3.2. Genotypes of The Control and Experimental Mice Used for Matrigel Plug Assay

3.2.4. Lentiviral-Cre Efficiently Deletes Ets2 In Vitro

Aortic endothelial cells were isolated from mice with genotype Ets1+/+;Ets2 fl/fl and Ets1-/-

;Ets2 fl/fl (Ets1KO) mice and cultured as discussed in chapter 2. The purified EC population was infected with lentivirus expressing Cre-recombinase to obtain Ets2KO (Ets2 single knockout) and DKO (Ets1 and Ets2 double knockout) ECs. Additionally, the cells were also infected with lentivirus without Cre as controls. Lentiviral-Cre mediated deletion analyzed by gene expression and western blot showed ~90% reduction in Ets2 expression levels (Figure 3.6.).

Figure 3.6. Cre-recombinase Mediated In Vitro Deletion of Ets2fl

(A) and (B) RNA expression data and Western blot performed using extracts from cultured aortic endothelial cells of genotype Ets1-/-;Ets2fl/fl infected with GFP or Cre- GFP expressing lentivirus.

3.2.5. Ets1 And Ets2 Regulate Endothelial Cell Cycle, Cell Survival and Migration

Previous studies from our lab demonstrated that in absence of both Ets1 and Ets2, expression of mmp9 and bcl2l1 are dysregulated indicating cell survival and migration as the potential downstream effector processes of ETS1 and ETS2 regulatory network. Since both these processes are essential for proper initiation and maintenance of activation phase of angiogenesis, the data suggested possible involvement of these proteins in regulation of the activation phase. In vitro assays were set up to detect the role of Ets1 and Ets2 in coordinating various processes, such as cell cycle, cell survival, cell proliferation, migration and adhesion, involved in the activation phase.

3.2.5.1. EC Lacking Ets1 and Ets2 Undergo Increased Apoptosis

Two different in vitro assays were performed to measure the effect of Ets1 and Ets2 on cell survival. After the induction of apoptosis by growth factor starvation, cells were subjected to Propidium iodide (PI) staining followed by flow cytometry. Normal distribution of the cell population during cell cycle was observed in case of control ECs

(Figure 3.7.A). In contrast, number of DKO cells in the sub G0/G1 population representing apoptosis was ~3 fold higher. Moreover, ECs with only Ets1 or Ets2 demonstrated no significant changes in the number of apoptotic cell as compared to wild type ones (Figure 3.7.A). The results of the PI staining assay were confirmed by Annexin

V and 7-Aminoactinomycin D (7-AAD) staining. Similar to PI staining, ~2.5 fold increase in the number of cells in the apoptotic fraction was observed in the absence of functional Ets1and Ets2 (Figure 3.7.B).

3.2.5.2. Loss Of Ets1 And Ets2 Leads To G2/M Phase Arrest

Interestingly, in addition to the increased apoptotic population, G2/M phase population demonstrated ~3 fold increase in the double mutant cells when compared to the controls

(Figure 3.7.A). A higher G2/M population indicated a possible cell cycle arrest. In order to validate this result, ECs were subjected to phospho histone H3 staining. Histone H3 is phosphorylated during mitosis and can be used as a specific immunomarker for cells undergoing mitoses (Hans and Dimitrov, 2001). This experiment demonstrated an obvious increase in the mitotic index (MI), expressed here as %, in the DKO cells. In control cells, the MI ranged between 8 to 10%. In comparison, MI of the DKO cells increased to ~35% (Figure 3.7.C), an increase similar to the one observed in the PI staining assay.

Figure 3.7. Deletion of Ets1 and Ets2 Affects EC Survival and Cell Cycle In Vitro (A) Cultured aortic ECs were serum starved (0.1% FBS) for 24 h. Cells were then harvested and subjected to PI staining followed by FACS. Graphs at right summarize the quantification. The percent apoptosis and M phase is calculated from the subsequent sub- G0/G1 peak and G2/M peak as determined by cell-cycle analysis. Data are plotted as mean percentages ± SD of three independent experiments. (B) Assessment of apoptosis after serum starvation (0.1% FBS, 24 h) of cultured aortic ECs by annexin V/7-AAD staining followed by FACS analysis. ECs in the lower right quadrant are Annexin- positive, early apoptotic cells. The cells in the upper right quadrant are Annexin- positive/7-AAD-positive, late apoptotic cells. Graphs at right demonstrate the quantification. Data are plotted as mean percentages ± SD of annexin V positive cells (both early and late apoptosis) of two independent experiments. (C) Cultured aortic ECs were processed for indirect immunofluorescence using anti-phospho-histone-H3 antibody (red), DAPI was used to stain the DNA. The graphic panel at bottom shows the number of M phase cells for each genotype expressed as mean percentages ± SD of the total number of cells measured from two independent experiments. Scale bars: 50 um (C).

Figure 3.7.

continued

Figure 3.7. continued

3.2.5.3. EC Specific Ets1/Ets2 Ablation Results in Reduced Migration and Tube

Formation In Vitro

In order for sprouting angiogenesis to occur, ECs must be able migrate and invade. An in vitro matrigel tube formation assay was performed with Ets1/Ets2 DKO aortic endothelial cells and controls (3.8.A) to detect the effect of deletion on EC migration.

These assays demonstrated that the control endothelial cells, including single Ets1KO or

Ets2KO cells, formed multicelluar tubular structures 24 hrs after plating, while DKO cells failed to form the mature tube-like structures (Figure 3.8.A). Single cell migration track assay (bead capture assay) was performed to further confirm the migration defect in the

DKO EC. In this assay, migrating EC engulf fluorescent beads leaving non-fluorescent tracks. The average area migrated by a single DKO aortic EC was reduced by ~2-folds compared to the controls (Figure 3.8.B). However, the migration of the single knockout endothelial cells remained unaltered.

3.2.5.4. Deletion of Ets1/Ets2 Leads to Increased EC Adhesion

EC adhesion to ECM is an important aspect of angiogenesis. The extra cellular matrix not only provides physical support, the EC-ECM communication can stimulate various pathways required EC survival and vessel branching. To determine whether Ets1/Ets2 deletion had an effect on EC adherence, a fibronectin based adhesion assay was performed. The read out of the assay was performed by measuring the intensity of bromophenol blue staining of adherent cells at 595nm. The double knockout endothelial

cells adhered with higher affinity to fibronectin at all concentration compared to controls

(Figure 3.8.C).

Figure 3.8. Deletion of Ets1 and Ets2 Affects EC Migration and Adhesion (A) In vitro matrigel tube formation assay: WT, Ets1 KO, Ets2 KO and DKO ECs were plated onto polymerized matrigel layer in a 96-well plate. Tube formation was monitored at 4 hours interval for 24 hours. (B) Migration track assay: The ECs were cultured on plates coated with fibronectin with overlaying fluorescent beads. The ability of the cells to create nonfluorescent tracks was quantified and represented as Average Area Migrated/Cell. Scale bars: (C) The cells were cultured on wells coated with gradually increasing concentration of fibronectin (0-100ug) for 1 hr. Afterwards non adherent cells were removed and adherent cells were stained with bromophenol blue. The absorbance was measured at 595nm. Scale bars: 80 um (A) and (B).

Figure 3.8.

continued

100

Figure 3.8. continued

migrated/cell Average area

101

3.3. Discussion

Angiogenesis is a fundamental process for both embryonic development and specific physiological and pathological condition in adults. Angiogenesis progresses in two distinct phases. The activation phase marks the onset of neo vessel formation in response to angiogenic factors. Later in the resolution phase, the nascent vessel matures with the recruitment of pericytes and the deposition of a basement membrane. A fine balance between pro-angiogenic and anti-angiogenic molecules regulate specific cellular processes required for angiogenesis such as cell cycle, cell migration and cell survival.

As discussed in chapter 1, several signaling pathways are sequentially stimulated and inhibited through the entire process. Dysregulation of a single pathway can lead to angiogenic defects and a severe pathologic condition. In contrast, tumor progression is also highly dependent on angiogenesis to provide nutrition/oxygen for the tumor to grow.

Thus, an extensive knowledge of the mechanism of angiogenesis is essential to treat ischemic diseases or develop tumor angiogenesis specific drugs.

Several TFs have been identified to play an important role in angiogenesis. However, in

Ets factors, such as Ets1, Ets2, Fli1, and Erg, seem to be centrally involved in the regulation of the entire process of vascular development, from endothelial cell differentiation to angiogenesis. Their importance is evidenced by presence of conserved

ETS sites in the promoter or enhancer regions most of the characterized EC genes (Bernat et al., 2006). Additional support comes from the genetic knockout studies. For example,

Fli1 knockout mice exhibit severe hemorrhage in the brain and are embryonic lethal at

E11.5 (Hart et al., 2000; Spyropoulos et al., 2000). Etv2-/- embryos also die by E9.5 with

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poor EC specification (Lee et al., 2008; Sumanas et al., 2008). Although early in vitro studies suggested a role of Ets1 and Ets2 in angiogenesis (Dejana et al., 2007; Hashiya et al., 2004; Lelievre et al., 2001), no vascular phenotype was observed in the murine knock out model of either Ets1 or Ets2 (Muthusamy et al., 1995; Yamamoto et al., 1998). One possible explanation for the lack of a defective vasculature was that these two proteins, being highly structurally similar, are redundant in their functions affecting vascular growth. Our lab has recently shown that absence of both these transcription factors results in embryonic lethality in Ets1-/-;Ets2A72/A72 mice. Similarly, mice with EC specific Ets2 deletion and global Ets1 knockout die before birth. The double mutant mice exhibit severe vascular defect. The data from the single knockout and the double knockout mice, taken together, indicate towards the redundancy of these factors. However, since the Ets1 deletion was conventional, it could not be inferred whether the effect of Ets1 or Ets2 were truly endothelial cell autonomous. Furthermore, owing to lethal nature of the embryos cellular processes downstream of Ets1 or Ets2 regulatory networks could not be identified. In the present study, we used a tamoxifen inducible tie-2Cre and floxed allele of both Ets1 and Ets2 to induce deletion of these TFs specifically in the ECs to show vascular defects in postnatal retinal angiogenesis of P8 mice. Further we identify cell cycle, cell survival and migration as the processes regulated by Ets1 or Ets2 to coordinate angiogenesis.

Murine retinal angiogenesis has long been utilized as a model to study the mechanism of angiogenesis specifically because of the ease with which developmental stages can be observed. In mouse retina, the vasculature starts to develop radially outward at P1 and

103

reaches the periphery at around P8. In our study, we show that in absence of Ets1 or Ets2, the area covered by the retinal angiogenic front was much smaller compared to the control retina. The absence of these transcription factors also rendered the sprouts abnormal. Both the number and complexity of filopodia extended at the protruding front of the tip cells were drastically reduced. To extend these observations, matrigel plug assays were performed to study the effect of the deletion of Ets1 and Ets2 in tumor angiogenesis. The analysis demonstrated that although the matrigel plugs form control mice had substantial formation of new blood vessels, minimal EC invasion and maturation was observed with in mice with the deletion of Ets1 and Ets2. In order to dissect out the mechanism involved in these severe anigiogenic defects, several in vitro assays were performed to identify the dysregulated cellular processes.

ETS1/ETS2 can be modulated by GF induced MAPK or AKT pathway. Effector molecules of the MAPK pathway extracellular signal-regulated kinases 1 and 2 (ERK1 and ERK2) regulate Ets1 or Ets2 function. Absence of ERK1 and ERK2 has been shown to reuce cell proliferation in ECs. Global gene expression array performed on ECs mutant for both ERK1 and ERK2 identified cell cycle as one of the major processes affected by the loss of these kinases (Srinivasan et al., 2009). Taken together, these data suggest that one mechanism by which these kinases regulate cell cycle/proliferation could be mediated by ETS1/ETS2. As expected, the DKO cells exhibited severely compromised cell cycle regulation. The number of cells in the M phase of the cell cycle was significantly higher in the mutant cells as compared to controls suggesting an M phase arrest. Furthermore, our study also demonstrated marked increase in EC apoptosis

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indicating that the increased rate of apoptosis might serve as an exit pathway from the M phase cell cycle arrest in the double mutant cells. Endothelial cell invasiveness was also affected by the loss of these TFs with the DKO cells failing to form three-dimensional tube like structure on matrigel even after 24 hr of culture.

Activation phase of angiogenesis involves the quiescent endothelial cells entering active cell cycle, migration of the tip cells guided by the angiogenic cues and the elongation of the vascular lumen. During this process, once the tip cell is selected, it needs to migrate toward the guidance cues. The filopodia extended by the tip cells senses the environmental guidance cues and the cells migrate accordingly. Lack of sufficient number of filopodia would result in failure of the tip cells to properly migrate. Once the tip cells start migrating, the neighboring stalk cells follow the tip cell and divide to extend the lumen. Cell proliferation is necessary to elongate the newly nascent sprout and develop the vascular lumen. Maintaining EC survival during vessel branching is also essential. Targeted deletion of Ets1 and Ets2, in EC results in embryonic lethality at

E13.5-E15.5 as a result of reduced EC survival (Wei et al., 2009). In short, the activation phase is a highly concerted process where proper execution of specific cellular processes, such as cell cycle, cell survival and migration, is essential. Inhibition of any of these processes would hinder the progress of angiogenesis. In our study, we show that all these processes are dysregulated in absence of ETS1 and ETS2. As a result, the vasculature in

DKO mice fails to develop at the same rate and complexity as that found in control littermates. Furthermore, the vascular defect, observed in mice with targeted deletion of both Ets1 and Ets2 specifically in the endothelial cells, could be rescued by the presence

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of only one copy of either Ets1 or Ets2. Taken together, our data suggest that Ets1 and

Ets2 regulate several endothelial cell functions, such as cell cycle, cell survival and migration, to coordinate the activation phase of angiogenesis in a cell autonomous manner.

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CHAPTER 4

EC SPECIFIC REGULATORY NETWORK OF ETS1/ETS2

4.1. Introduction

Cellular functions need to be able to adapt to the surrounding environment and respond accordingly. Expression of specific genes must be regulated following various internal and external stimuli. By ensuring correct expression of a gene, transcription factors play a central role to many cellular processes. Whether overrepresented or dysfunctional, abnormal transcription factor behavior can lead pathological disorders (Furney et al.,

2006; Boyadjiev and Jabs, et al., 2000). The ETS family of TFs controls a variety of physiologic processes (Hollenhorst et al., 2011; Sharrocks, 2001; Oikawa and Yamada

2003) and is involved in a large number of diseases, particularly cancers (Hsu et al.,

2004; Oikawa, 2004). All ETS proteins share a conserved DNA binding domain, the ETS domain, that recognizes sites harboring a 5‘-GGAA/T-3‘ consensus. These sequences flanking the 5 bp core regions are often conserved for individual ETS members‘ specificity. Another domain, the PNT domain, is also present in about one third of the Ets family members including ETS1 And ETS2. In ETS1/ETS2 the PNT domain acts as a 107

docking site for mitogen-activated protein kinases. ERK1 and ERK2, downstream effector molecules of RAS-MAPK pathway, bind to the PNT domain and phosphorylate

Thr residue and activate the transcription factors (Klämbt, 1993; Yang et al., 1996;

Graves and Petersen, 1998; Petrovic, 2003).

4.1.1 Biological Role of ETS1 and ETS2 as Transcription Factors

Expression of genes involved in a variety of cellular functions, such as cell growth, proliferation, survival, and differentiation, are regulated by ETS1 and ETS2. They are crucial modulators of several biological processes such as tissue remodeling, angiogenesis and tumor metastasis (Sementchenko and Watson, 2000). Significant insight has been gained by studying the murine knockout model of these transcription factors. Ets1 null mice are viable and fertile. However they lack natural killer (NK) cells and exhibit defective B- and T- cell survival (Muthusamy et al., 1995). Ets2db/db (Ets2 allele with deleted the DNA binding domain) mice are embryonic lethal at around E8.5

(Yamamoto et al., 1998). They are severely growth retarded showing several developmental defects, including the absence of amnion and placental architecture defects. However the lethality is due to the extraembryonic defects and mutant embryos can be subjected to tetraploid rescue. The rescued ETS2 null mutants are viable and fertile albeit with a wavy hair phenotype (Yamamoto et al., 1998). The threonine 72 residue gets phosphorylated by ERK1/ERK2. When Thr72 is mutated to alanine, the mutant mice Ets2A72/A72 are viable, fertile and appear to develop normally. However, combining the Ets2A72 allele with the global deletion of Ets1 also resulted in embryonic lethality (Wei et al., 2009). ETS1 and ETS2 also transactivate cdc2 and cyclin D1

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promoters (Wen et al., 1995). ETS1 and ETS2 have been reported to be both pro- apoptotic as well as anti-apoptotic in a context dependent manner.

ETS1/ETS2 are involved in tumorigenesis also. Overexpression of ETS genes has been correlated with a number of human cancers. NIH3T3 cells can be transformed by over expression of ETS1 (Seth and Papas, 1990). Similarly, during tumor progression in prostate and breast cancer ETS2 level goes up. Elevated level of ETS2 in prostate tumor cells stimulates apoptosis. In addition, over expression of ETS2 enhances proliferation of the cells and abolishes their requirement for serum (Seth et al., 1989). In addition to their function in tumor cell, ETS1 and ETS2 have also been implicated to function in tumor stroma. When wild type mammary tumor was transplanted into Ets2A72/A72 mice, rate of tumor growth slowed down significantly (Man et al., 2003). Moreover, fibroblast specific

Ets2 inactivation led to significant reduction in size in MMTV-PyMT mammary tumor model (Wallace et al, 2013). Mice lacking Ets2 specifically in macrophages had fewer and smaller metastatic lesions growing in the lungs in tail-vein injection model

(Zabuawala et al., 2010).

4.1.2. Coactivators of Ets Factors

All members of the Ets family of transcription factors by definition harbor the same DNA binding domain that bind to DNA with a common 5-bp core sequence. Furthermore, in a given cell type more than half of ETs family members can be co-expressed (Hollenhorst et al., 2004). However, genetic knock out studies have led to drastically distinct phenotype as well as redundant functions when specific Ets factor was deleted as discussed earlier. Taken together, these data raise the question of target specificity of

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individual Ets factors. One possible mechanism of achieving specificity could involve the

DNA sequences flanking the core 5-bp region. Since the flanking sequence can be flexible, they can provide enhanced binding only to specific Ets factors. Interaction with other co-regulatory partners seems to be another mechanism to allow combinatorial control of gene expression and specificity of action to ETS-domain proteins (Li et al.,

2000.)

Biochemical and genetic studies have provided evidence that Ets factors form multi protein complexes at enhancers and promoters of target genes (Eisenbeis et al., 1995;

Yang et al., 2000). The CREB-binding protein (CBP) and p300 are two highly structurally similar nuclear proteins that function as transcriptional coactivators by acting as adapters between transcription factors and basal transcription machinery (Goodman and Smolik, 2000). Transcriptional activation by Ets proteins is usually correlated with the recruitment of the co-activator CBP/p300. Activated ERK1/ERK2 binds to the ONT domain of ETS1/ETS2. ERK induced phosphorylation of ETS1 and ETS2 stimulates recruitment of CBP/p300 to enhance transcription of target genes by these transcription factors (Foulds et al., 2004).

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4.2. Results

4.2.1. Specific EC Processes Required for Angiogenesis are Disrupted in The

Absence of Ets1 and Ets2

To initially address the underlying mechanism responsible for angiogenic defects observed in the double mutant mice, low-passage primary mouse aortic EC were used for global gene expression profiling. Expression profiling using the Affymetrix platform was performed with RNA isolated from primary aortic EC representing four genetic groups: wild type, Ets1 KO, Ets2 KO and Ets1/Ets2 DKO. The endothelial cell populations were purified from the aorta of mice of appropriate genotype, and Cre delivered by lentivirus to effect conditional gene deletion. The expression data obtained was analyzed with Gene

Set Enrichment Analysis (GSEA) comparing the expression pattern of the DKO cells with that of the other three genotypes combined (Subramanian et al., 2005). The analysis identified ~2,000 genes differentially expressed in the DKO versus the other genotypes; the top 200 genes differentially expressed in DKO compared to the three controls are represented in the heat maps in Figure 4.1.A. Using stringent criteria (nominal p value <

0.01, FDR q value < 5%) GSEA identified only three fundamental cellular processes affected in EC by the deletion of Ets1 and Ets2: M phase of the cell cycle, cell survival and cell migration (Figure 4.1.B).

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Figure 4.1. Biological Processes Essential for Angiogenesis are Disrupted in Ets1/Ets2 Double Mutant Cells (A) Heatmap of microarray profiling of WT, Ets1 KO, Ets2 KO and DKO endothelial cells. Heatmap was generated using expression levels of 100 upregulated and downregulated genes (as determined by GSEA) comparing DKO endothelial cells with all other indicated genotypes. (B) Gene Set Enrichment Analysis (GSEA), comparing gene expression between DKO endothelial cells with all other genotypes combined (REST), showing key processes that are affected. NES: normalized enrichment score.

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Figure 4.1.

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4.2.2. Loss of Ets1/Ets2 Affects Expression of Genes Involved in EC Cell Cycle, Cell

Survival and Migration

In order to recognize specific genes that might be dysregulated in the DKO cells, the leading edge subsets of these gene sets identified by the GSEA were scrutinized. To verify the expression of these genes contributing to the leading edges, quantitative real- time RT-PCR (q-PCR) was performed on RNA prepared from independently purified sets of aortic EC. This analysis demonstrated expression of genes regulating processes, such as cell survival (Bcl2l1, Mcl1), cell cycle and cell cycle arrest (Ube2c, APC5) and migration (Mmp9, Mmp3), were significantly decreased when both Ets1 and Ets2 were absent (Figure 4.2.).

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Figure 4.2.

Figure 4.2. Deletion of Ets1 and Ets2 Results in Deregulation of Genes Involved in Cell Cycle, Cell Survival and Migration Target gene expression in cultured aortic endothelial cells from double-mutant and control mice as indicated. Gene expression analysis was performed by qPCR on RNA harvested from cultured aortic ECs.

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4.2.3. ETS1 and ETS2 Directly Regulate Specific Cell Cycle, Anti Apoptosis and

Migration Genes

In a complementary analysis, genomic binding patterns of ETS1 and ETS2 were determined in wild-type aortic EC by chromatin immune-precipitation coupled with massively parallel sequencing (ChIP-seq). The resulting sequence reads were aligned to the reference mouse genome (mm10) (Kent et al., 2002) with Bowtie (Langmead et al.,

2009) and subsequently analyzed with HOMER (Heinz et al., 2010). The analysis identified 81,801 ETS1 peaks and 26,792 ETS2 peaks (Figure 4.3.A), and 6,353 genomic regions were found to be bound by both transcription factors. Similar results were obtained when the ChIP-seq data was analyzed with shape based peak identification (data not shown). The genomic regions common to ETS1 and ETS2 binding could be assigned to 6353 ―nearest neighbor‖ protein coding genes: 60% in the intragenic regions

(primarily in introns) and 32% in intergenic locations <100KB distal (Figure 4.3.B). In contrast, only 7% of the shared binding sites were found within 500 bp upstream from the transcription start site. As a test of the enrichment of specific binding sites, the genomic regions occupied by both ETS1 and ETS2 were analyzed for the enrichment of the ETS consensus motif. 12 distinct motifs were identified, with 9 of these 12 contained an ETS- family consensus binding sequence (Table 4.1.).

Circular plots (Circos) were utilized to illustrate the genome wide interaction pattern between the genes differentially expressed in the absence of Ets1 and Ets2 and genomic regions occupied by these transcription factors in tandem (Krzywinski et al., 2009). To better demonstrate the global gene expression changes coordinated by ETS1 and ETS2,

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the expression data of top 2,000 differentially expressed genes identified by the GSEA were represented in the Circos plot along with the global binding pattern of these two transcription factors (Figure 4.3.C).

In order to confirm that genes, differentially expressed in absence of ETS1/ETS2 (as discussed in previous section) are direct targets of ETS1 and ETS2, standard chromatin immunoprecipitation (ChIP) assays were performed on the peak regions identified by

ChIP-seq. Since it was essential to ascertain whether the binding of ETS1 and ETS2 were interdependent, chromatin isolated from the wild type, the double knockout as well as each of the single knockout cells were subjected to ChIP studies. As demonstrated in

Figure 4.3.D, both ETS1 and ETS2 were found to be enriched at the regulatory regions of the genes examined in the wild type chromatin. Absence of either protein did not significantly reduce the binding of the other as shown by the data obtained from the single knockout cells. However, the association was completely abolished when both

ETS1 and ETS2 were deleted.

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Figure 4.3. ETS1 and Ets2 Along with Coactivators CBP/p300 are Enriched in The Enhancer Regions of Specific Cell Cycle and Cell Survival Genes (A) Venn diagram representing the overlap between ETS1 peaks and ETS2 peaks in wild type endothelial cells as detected by ChIP-Seq. (B) Genome-wide distribution of ETS1/2 overlapping binding sites in wild type endothelial cells. (C) Circos plot representing the correspondence between the genomic locations of the peaks and the top 2000 dysregulated genes identified by GSEA. (a) – (d) gene expression pattern of the genes specified in WT, Ets1 KO, Ets2 KO and DKO ECs respectively. (e) – (f) Innermost three circles represents ETS1, ETS2 and overlapping peaks. (D) In vivo chromatin occupancy was examined by ChIP-qPCR in cultured aortic endothelial cells. Anti-Ets1 and anti-Ets2 antibodies were used. Data are plotted as means ± SD of three independent ChIP experiments.

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Figure 4.3.

continued

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Figure 4.3. continued

continued

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Figure 4.3. continued

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Table 4.1. List of Motifs Identified by Analyzing The Genomic Regions Occupied by Both ETS1 and ETS2

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4.2.4. ETS1 and ETS2 Directly Regulate Specific Cell Cycle, Anti Apoptosis and

Migration Genes

The CREB-binding protein (CBP) and p300 are two highly structurally similar nuclear proteins that function as transcriptional coactivators by acting as adapters between transcription factors and basal transcription machinery (Goodman and Smolik, 2000).

Mitogen-activated protein (MAP) kinase induced phosphorylation of ETS1 and ETS2 leads to recruitment CBP/p300 to enhance transcriptional activation (Foulds et al., 2004;

Nelson et al., 2010). In order to determine whether CBP/p300 mediates the regulation of the selected cell cycle and cell survival genes by ETS1 and ETS2, conventional ChIP assays were performed utilizing antibody that recognizes acetylated CBP/p300. Indeed,

CBP/p300 was found to be recruited to each of the ETS1 and ETS2 peaks examined.

Although the recruitment was maintained even with either ETS1 or ETS2 present, deletion of both seemed to significantly affect the recruitment (Figure 4.4.).

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4.4. ETS1 and Ets2 Recruit Coactivators CBP/p300 to The Enhancer Regions of Specific Genes ChIP assays were performed followed by real-time qPCR to show chromatin occupancy by the co-activators CBP/p300. Anti-CBP/P300 acetylation specific antibodies were used. Data are plotted as means ± SD of two independent ChIP experiments.

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4.3. Discussion

The vascular system is essential for both embryonic development and adult life.

Abnormal vascularization can lead to severe pathological conditions including cancer, atherosclerosis, retinopathy and stroke. Following the development of the primitive vasculature in embryo by vasculogenesis, vascular network is rapidly extended and remodeled through angiogenesis. Angiogenesis involves sprouting, vessel branching and vessel maturation (Flamme et al., 1997; Patan, 2004). The entire process is tightly regulated by growth factors and cell-cell as well as cell-ECM communication. A considerable number of studies have examined the role of signaling molecules in angiogenesis. VEGF is considered to be the most crucial growth factor involved in angiogenesis. However, insight about the transcriptional regulation of gene expression required for proper EC function is lacking (De Val and Black, 2009).

Transcriptional activators and repressors bind regulatory sequences within promoters and enhancers to alter gene expression pattern. Several TFs play crucial roles in development and maintenance of the vascular network. However, Ets family transcription factors appear to be more crucially involved in this process than any other TF. In EC, at least 19 different Ets factors are co-expressed. Recent work from our lab demonstrated an overlapping, redundant function of Ets family member ETS1 and ETS2 in regulating embryonic angiogenesis (Wei et al., 2009). However the embryonic lethal nature of the mutant mice ruled out further study of the downstream processes which is crucial in order to better understand the mechanism of aberrant angiogenesis associated with various diseases.

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CBP/p300 are two homologous protein that act as coactivators (Goodman and Smolik,

2000). CBP/p300 possesses intrinsic histone acetyltransferase (HAT) activity. When recruited, they relax the chromatin structure to provide more access to the regulatory region of the transcription factors. Moreover, CBP/p300 are able to recruit basal transcriptional machinery including RNA pol II. It has been demonstrated previously that

ETS1 and ETS2 recruit CBP/p300 to enhance the expression of target genes (Foulds et al., 2004). Recruitment is enabled by the activated ERK induced phosphorylation of

ETS1/ETS2.

In the current study, gene expression analysis comparing the DKO cells with control EC identified cell cycle (specifically M phase of the cell cycle), cell survival and migration as the potential downstream processes of ETS1/ETS2 regulatory network. This data was in accordance with the results obtained from the in vitro studies described in the previous chapter. Gene set enrichment analysis performed on the expression data identified several

ETS1 and ETS2 putative target genes involved in this processes. Among the differentially expressed cell cycle genes, Ube2c is a ubiquitin conjugating enzyme responsible for the destruction of mitotic cyclin and progress through cell cycle.

Knockdown of Ube2c has been shown to induce G2/M phase arrest (Wang et al., 2009;

Shen et al., 2013). Ectopic expression of fused c-terminal fragments of human Aspm in

HeLa cells inhibits spindle assembly leading to mitotic arrest (Higgins et al., 2010).

Similarly, use of Cpd5, a Synthetic Thioalkyl Vitamin K Analogue, inhibits Cdc25c to induce G2/M phase arrest (Tamura et al., 2000). These data suggest that downregulation of these genes in the absence of ETS1/ETS2 could induce the observed M phase arrest.

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The global protein-DNA interaction study revealed that several of the genes dysregulated in DKO cells direct targets of ETS1 and ETS2. Further, the binding of either protein to target gene regulatory region in independent of the other as the enrichment of ETS2 and

ETS1 in Ets1KO and EtsKO cells was similar to that of wild type cells. Since deletion of only Ets1 and Ets2 failed to elicit any observable phenotypic defect, we only focused on the regions bound by both ETS1 and ETS2. Interestingly, the redundant function of these protein appear to be regulated from enhancers as opposed to promoter regions as majority of the co-occupied peaks were found in the inter- and intragenic regions. Taken together, these data suggest the direct regulation of genes specific for cell cycle and cell survival by ETS1 and ETS2 is critical in their regulation of EC function.

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CHAPTER 5

CONCLUSION AND FUTURE DIRECTIONS

5.1. Conclusions

Angiogenesis is essential to satiate the ever growing need for nutrients/oxygen during development as well as in several specific physiological processes in adult. It is a multi step process that involves orchestrated execution of several biological events including vessel sprouting, cell proliferation, directed cell migration etc (Carmeliet and Jain, 2011).

The execution requires complex and highly coordinated crosstalk between angiogenic inducers and inhibitors: signaling pathways that affect the growth, migration, survival and differentiation of ECs (ZEtter, 1998; Folkma, 2003; Cheresh and Stupack 2008).

Delineating these signaling pathways and the corresponding effector genes that mediate

EC functions is paramount in revealing the molecular events that occur during angiogenesis. In the current study, we demonstrate an EC specific and redundant role of

Ets1 and Ets2 that links M phase of endothelial cell cycle with cell survival in regulating angiogenesis. 128

Several previous studies involving overexpression and dominant-negative approaches have implicated different ETS family members such as Ets1, Ets2, Fli1, Erg and Nerf-2 in EC function (De val and Black 2009; Hashiya et al., 2009). Previous genetic and biochemical evidence presented by our lab has demonstrated that during embryonic angiogenesis deletion of Ets1and Ets2 impairs endothelial cell survival in a redundant and cell autonomous fashion. The double knock out mice show impaired angiogenesis and embryonic lethality (Wei et al., 2009). Here we used an inducible Tie2-Cre-ERTam to knock out Ets1 and Ets2 specifically in ECs to look for postnatal vascular defects.

Alterations of postnatal retinal vascularization were observed as phenotypic effects of

EC-specific deletion of these genes. Vascular defects included retarded growth of the angiogenic front, reduced complexity of vascular branching and impaired sprout formation. Interestingly, presence of even one copy of either Ets1 or Ets2 could restore the wild type phenotype. These results argue that the effect of these genes on EC function is redundant and highly cell autonomous. Failure to express functional copies of these proteins attenuated tumor angiogenesis as well. The angiogenic response was severely dampened in the DKO mice as was evident by a marked reduction of EC invasion and maturation. Thus, both developmental and tumor angiogenesis seems to be similarly dependent on the EC specific function of Ets1 and Ets2.

Extracellular signal-regulated kinases 1 and 2 (ERK1 and ERk2) are upstream regulators of ETS1 and ETS2. Previous studies from our lab have shown that Erk1/2 ablation in EC results in reduced EC proliferation. Additionally, global gene expression analysis comparing control and Erk1/2 deleted EC revealed dysregulation of a number of genes

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involved in cell cycle (Srinivasan et al., 2009). Based on these findings, we hypothesized that the endothelial cell cycle specific effect of ERK1/2 could be mediated by ETS1 and

ETS2. Indeed in our current study, abnormal cell cycle appears to be one possible mechanism by which ETS1 and ETS2 regulate EC function. The in vitro assays performed here demonstrate that EC cycle was highly compromised in the DKO cells.

This defect led to an increased M phase DKO EC population disrupting the normal cell cycle phases that are critical in angiogenesis. Furthermore, the deletion of Ets1 and Ets2 promote endothelial cell death, indicating apoptosis is likely the exit pathway for the mutant cells from the induced M phase arrest.

Consistent with the above studies, the GSEA analysis identified the M Phase of the cell cycle and apoptosis among the most important downstream effector processes of ETS1 and ETS2. Several genes involved in cell cycle were differentially expressed in the DKO

ECs, which could account for the M phase arrest. Similarly, the expressions of a number of anti apoptotic genes were also downregulated conferring increased sensitivity to apoptotic signals. Findings from the global DNA-protein interaction study suggested that several of these genes were direct targets of ETS1 and ETS2. Of note, we focused further analysis only on the regions occupied by both ETS1 and ETS2 because deletion of only

Ets1 and Ets2 failed to elicit any observable phenotypic defect. Interestingly, the redundant function of these protein appear to be regulated from enhancers as opposed to promoter regions as majority of the co-occupied peaks were found in the inter- and intragenic regions. In addition, standard ChIP performed on single KO cells confirmed that binding of one protein to these regulatory regions is completely independent of the

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other. In summary, the genetic and biochemical evidence presented here suggests that

Ets1 and Ets2 regulate the cellular machinery responsible for maintaining normal cell cycle and cell survival required for activation phase of angiogenesis.

5.2. Future Directions

In the current study, although we examine cell cycle as a possible mechanism of aberrant angiogenesis due to the loss of Ets1 and Ets2, the exact events that trigger the M phase arrest is yet to be determined. A number of genes were also identified to be downregulated in the DKO EC but were not associated with any ETS1 or ETS2 peaks

(data not shown). One possible explanation might be that these genes are indirect targets of ETS1 and ETS2 and are regulated through miRNAs that are in turn regulated by ETS1 and ETS2. Another interesting avenue to explore would be the processes and genes that are independent targets of only ETS1 or ETS2.

5.2.1. Effect of ETS1/ETS2 on EC processes in vivo

In the present study, we demonstrated how the loss of Ets1 and Ets2 affect the progression of angiogenesis. Retinal angiogenesis model revealed defective and insufficient vasculature with reduced number of tip cell filopodia in the mutant mice as compared to the control littermates. Furthermore, in vitro assays were designed to identify cellular processes dysregulated in the absence of these transcription factors.

However, results from in vitro assays may not be recapitulated in vivo. Thus it is of paramount importance that studies are performed to detect defects in the angiogenic cellular processes in the mice.

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The retinal angiogenesis model, in conjunction with the inducible Cre and floxed alleles of Ets1 and Ets2, can be utilized extensively to achieve these goals. Analysis of EC proliferation can be performed by staining the bromodeoxyuridine (BrdU) labelled cells.

BrdU will be injected into live pups ~3 hrs before they are sacrificed for harvesting the eyes. Phosphorylated histone H3 (pH3) staing of the retinal vasculature along with isoloctin B4 will identify if the endothelial cells experience a cell cycle arrest in vivo.

Similarly cleaved caspase-3 staining of retinal ECs would provide us with insight about apoptotic induction.

5.2.2. Mechanism of G2/M phase cell cycle arrest

At the beginning of the activation phase, the quiescent ECs lining the inner wall of the existing vessels are activated by pro-angiogenic factors and enter active cell cycle.

Increase in the number of stalk cell is crucial in order for the nacent sprout to gow and follow the tip cells. The tip cell migrates toward another sprout to fuse with the tip cell thereof. During proliferation, cellular function cycles between the interphase and the mitotic phase interjected with gap phases. The gap phases are equipped with cellular machinery that scrutinize the progress of the cell through the cycle and detect any error or abnormalities accumulated during the process, the cell cycle checkpoints. The checkpoints operate at stages G1, G2 and M to maintain the integrity of the genome.

During the M phase, the checkpoint machinery functions to detect abnormalities at several steps including chormosome condensation, spindle assembly, spindle checkpoint and chromosome separation. During the metaphase or the spindle checkpoint, separation is prevented until all the chromatids are in place. 132

In the current study, we show DKO cells experience cell cycle arrest at the M phase.

However, at checkpoint the arrest occurs or the cause of the arrest is still to be studied.

However, several of the cell cycle genes tested are involved in spindle formation. Anapc5 is a component of the anaphase promoting complex/cyclosome (APC/C), a cell cycle- regulated E3 ubiquitin ligase that controls progression through mitosis. The APC/C is allowed to degrade cyclin B and securing only after the chromatids align properly at the mitotic plate. However in absence of a functional APC/C mitotic arrest ensues (Zieleke et al., 2008). Ube2c is also an essential component of APC/C (Rape and Kirschner, 2004;

Summers et al., 2008). In addition, Aspm has been proposed to be involved mitotic spindle pole organization (zhong et al., 2005). Although the functional aspects of the genes differentially expressed in double mutant cells suggest that the M phase arrest is possibly due to deregulation of spindle checkpoint, further experiments will be required to dissect out the exact mechanism. Small interfering RNA (siRNA) induced knockdown of individual target genes can be used to determine if absence of one or a combination of these gene are sufficient to recapitulate the phenotypic effect of ETS1/ETS2 knock out.

Moreover, different drugs, such as etoposide, can be used to arrest cell cycle at various stages of M phase to identify a phenocopy. ETS1, ETS2 and a combination of their targets would be added back to reveal whether the phenotype can be rescued and also recognize the exact stage where cell cycle is arrested.

5.2.3. Post-transcriptional control of EC specific processes by ETS1/2 microRNAs (miRNAs) are an abundant class of endogenous small non-coding RNAs which are ~ 22 nucleotides in length that play a critical role in post-transcriptional gene 133

regulation (Lodish et al., 2008, Bartel, 2004). lin‑4 and let‑7 were the first species of miRNA to be discovered. A mature miRNA is ~22 nucleotides long. However, when first transcribed the pri-miRNA may be more than 1 kb long. Th pri-miRNA is cleaved by endonuclease Drosha to generate ∼60–70 nt stem loop intermediate, the pre-miRNA.

This pre-miRNA is then transported to the cytoplasm by Ran-GTP and the export receptor Exportin-5 (Yi et al., 2003; Lund et al; 2004). Drosha processes one end of the mature miRNA in the nucleus whereas the other end is processed in the cytoplasm by the endonuclease Dicer. The mature miRNA mode of action involves the assembly of an

RNA-induced silencing complex or RISC, which identifies targets by perfect (or nearly perfect) complementarity between the miRNA and the mRNA. miRNA possess a 7-8 nucleotide long ‗seed‘ sequence which determines its target specificity during miRNA- mRNA interactions. Once loaded on to the complex the target mRNA is either degraded or prevented from being translated (Lodish et al., 2008). miRNAs play an important role in vasculature development. Dicer hypomorphic mice die at E12,5-E14.5. The mutant embryos exhibit severe defects in both embryonic and extraembryonic vasculature (Yang et al., 2005). Targeted deletion of Dicer specifically in

ECs demonstrates reduced post natal angiogenesis in mice (Suarez et al., 2008). Several individual miRNAs also regulate vascular function. miR-221/222 as negative regulators of angiogenesis (Poliseno et al., 2006). miR-126 is the most abundant microRNA in endothelial cells. miR-126 is important in both pathological and developmental angiogenesis (Fish et al., 2008). In contrast, miR-17-92 cluster acts as negative regulator of angiogensesis (Doebele et al., 2010).

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Since miRNAs are crucially involved in angiogenesis regulation, it would be useful to decipher the role of ETS1/ETS2 in miRNA expression in ECs. To identify the d ifferetially expressed miRs, a miRNA expression analysis comparing WT, Ets1 KO, Ets2

KO and DKO can be performed using the Nanostring platform. Chip-seq data and the

Nanostring data can be compared to identify miRNAs that direct targets of ETS1/ETS2.

In contrast, the Naostring data can be overlapped with the microarray data to recognize putative mRNA targets. Once the miRNAs, potentially involved in angiogenesis, are identified, genetic knockdown studies can be performed to to verify their effect in vascular development.

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