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RASIP1 REGULATES VASCULAR TUBULOGENESIS RASIP1 REGULATES VASCULAR TUBULOGENESIS

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RASIP1 REGULATES VASCULAR TUBULOGENESIS - PPT Presentation

APPROVED BY SUPERVISORY COMMITTEEOndine Cleaver PhDThomas Carroll PhD ChairMelanie Cobb PhDEric Olson PhDToMy familyACKNOWLEDGEMENTI am extremely grateful toDr Ondine Cleaver who has been the greatest ID: 878962

cell rasip1 formation vascular rasip1 cell vascular formation figure cells lumen ecs endothelial vessels embryos blood expression arhgap29 adhesion

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1 RASIP1 REGULATES VASCULAR TUBULOGENESIS
RASIP1 REGULATES VASCULAR TUBULOGENESIS APPROVED BY SUPERVISORY COMMITTEE Ondine Cleaver, Ph.D. Thomas Carroll, Ph.D. (Chair) Melanie Cobb, Ph.D. Eric Olson, Ph.D. To My family ACKNOWLEDGEMENT I am extremely grateful to Dr. Ondine Cleaver, who has been the greatest mentor and best friend to me. I feel super lucky to be able to join her laboratory. It was in th is most friendly and encouraging family - like environment that she has created for everyone of us I realized just how much I enjoy the challenges and rewards of scientific research. Being a Cleaver lab member has been one of the most cherished memories of my life. My deepest g ratitude also goes to my committee members, Drs. Thomas Carroll (Chair) , Melanie Cobb and Eric Olson. Their great insights, passion s in science, invaluable advice and unconditional support have always kept me on the right track and driven me to pursue high er goals. I also want to thank them fo r their efforts in creating the best graduate school and the best research department for all of us. They are my role models in my scientific career. I want to

2 thank Dr . Michael Haberland . The treï
thank Dr . Michael Haberland . The treendous aount of „ cra zy’ ideas and amazing hands - on experiences he shared with me in molecular biology will be a life - long benefit. I’d like to thank all current and past eers in the C leaver lab, particularly Diana Chong, Alethia Villasenor, Brian Fairbanks and Stephen Fu f or their great help in developing ideas and carrying out experiments. Also thanks to all other lab mates Stryder Meadows, Peter Fle t cher, Yeon Koo, Lelani Marty, David Barry for their support and friendship. I also want to thank all the technical and administrative staff s of the graduate school, the Genetics & Development program, and the Molecular Biology Department for their daily support. Last but not least, I want to give my greatest thanks to my wife Qiuping Chen ( 陈秋萍 ), my parents Xin Zheng ( 郑欣 ) a nd Bailing Xu ( 许百伶 ) , my sister Duo Xu ( 许多 ) and all my family for their endless love. They are pillars of my life. RASIPA REGULATES VASCULAR TUBULOGENESIS by Ke Xu ( 许可 ) DISSERTATION Presented to the Faculty of the Graduate School of Biomedical Scien

3 ces The University of Texas Southwes
ces The University of Texas Southwestern Medical Center at Dallas In Partial Fulfillment of the Requirements For the Degree of DOCTO R OF PHILOSOPHY The University of Texas Southwestern Medical Center at Dallas Dallas, Texas May, 2011 vi Copyright by Ke Xu , 2011 All Rights Reserved vii RASIPA REGULATES VASCULAR TUBULOGENESIS Ke Xu, Ph.D. The University of Texas Southwestern Medical Center at Dallas, 2011 Mentor: Ondine Cleaver, Ph.D. Cardiovascular function depends on patent blood vessel formation by endothelial cells (ECs). However very little is known about the mechanisms underlyin vascular „tuuloenesis’. This study identifies Rasip1 as a unique, endothelial - specific regulator of Rho GTPase signaling, which is essential for endothelial lumen morphogenesis . We found that Rasip1 is strongly expressed in vascular endothelial cells throughout development across species . Similar to the well - characterized vascular markers VEGFR2 and PECA M, Rasip1 is specifically expressed in angioblasts prior to vessel formation, in the initial embryonic vascular plexus, in the gr

4 owing blood vessels during angiogenesis
owing blood vessels during angiogenesis and in the endothelium of mature blood vessels into the postnatal period. Rasip1 expres sion is undetectable in VEGFR2 null embryos, which lack endothelial cells, suggesting that Rasip1 is endothelial - specific. viii Ablation of Rasip1 both in vitro and in vivo strongly affects vascular integrity. Specifically, siRNA - mediated reduction of Rasip1 s everely impairs angiogenesis in endothelial cell cultures, and morpholino knock down experiments demonstrate that Rasip1 is required for embryonic vessel formation in frog embryos . Mice lacking Rasip1 fail to form patent lumens in all blood vessels, including the early endocardial tube. Rasipl null angioblasts fail to properly localize the polarity determinant Par3 and display defective cell polarity, resulting in mislocalized junctional complexes and loss of adhesion to extracellular matrix (ECM). D epletion of either Rasip1 or its binding partner RhoGAP Arhgap29 in cultured ECs blocks in vitro lumen formation, fundamentally alters the cytoskeleton and reduces integrin - dependent adhesion to ECM. These d efects result from increased RhoA/ROCK/myosin II

5 activity and blockade of Cdc42 and Rac
activity and blockade of Cdc42 and Rac1 signaling. Together, our work identify Rasip1 as a novel endothelial factor that plays an essential role in vascular tubulogenesis . ix TABLE OF CONTENTS Abstract ................................ ................................ ................................ ..................... vii List of publications ................................ ................................ ................................ .... xi List of figures ................................ ................................ ................................ ........... xii List of tables ................................ ................................ ................................ ............. xv List of abbreviations ................................ ................................ ................................ xvi Chapter I: Introduction ................................ ................................ .......................... 1 Vasculogenic tubulogenesis: cord - to - tube transition ................................ ............... 3 Angiogenic tubulogenesis: lumen extension ................................ .........

6 ................... 5 Cellular mecha
................... 5 Cellular mechanism of vascular tubulogenesis ................................ ........................ 7 Vascular tubulogenesis: mechanistic heterogeneity ................................ .............. 13 Chapter II: Materials and methods ................................ ................................ ..... 19 Chapter III: Rasip1 is important for vascular morphogenesis ....................... 35 Identification of Rasip1 expression in murine endothelial cells ........................... 39 Rasip1 is expressed in vascular endothelial during vascular plexus formation ... 40 Rasip1 during late embryogenesis ................................ ................................ .......... 43 Rasip1 expression is absent in vascularless embryos ................................ ............ 45 Rasip1 is required for angiogenesis in cultured ECs ................................ ............. 47 Rasip1 is required for embryonic blood vessel formation ................................ ..... 49 Discussion ................................ ................................ ................................ ................. 55 Ch

7 apter IV: Rasip1 directs vascular tubulo
apter IV: Rasip1 directs vascular tubulotenesis ................................ .......... 60 Rasip1 is essential for cardiovascular development ................................ .............. 62 x Blood vessel tub ulogenesis requires Rasip1 ................................ ........................... 62 Rasip1 binding partners ................................ ................................ ........................... 66 Rasip1 and Arhgap29 are required for in vitro lumen formation .......................... 69 Rasip1 and Arhgap29 are required for activation of Cdc42 and Rac1 during lumen formation ................................ ................................ ................................ ................... 70 Rasip1 and Arhgap29 regulate EC architecture via repression of RhoA ............. 72 Failure of EC - ECM adhesion in vitro in the absence of Rasip1 or Arhgap29 ..... 76 Rasip1 is required for proper in vitro endothelial - ECM adhesion ........................ 80 Rasip1 is required for establishment of endothelial apicobasal polarity .............. 83 Discussion ................................ ...............

8 ................. ......................
................. ................................ ................. 85 Chapter V: Summary ................................ ................................ ............................ 96 References ................................ ................................ ................................ ................ 98 xi LIST OF PUBLICATIONS 1) Ke Xu , Anastasia Sacharidou, Stephen Fu, Diana C. Chong, Brian Skaug, Zhijian J. Chen, George E. Davis and Ondine Cleaver. Blood vessel tubulogenesis requires Rasip1 regulation of GTPase signaling. Development Cell . 2011; 20(4): 526 - 539. 2) Jian Xie, Tao Wu, Ke Xu , Ivan K. Huang, Ondine Cleaver, and Chou - Long Huang. Endothelial - Specific Expression of WNK1 Kinase Is Essential for Angiogenesis and Heart Development in Mice . The American Journal of Pathology . 2009; 175(3):1315 - 27. 3) Ke Xu , Diana Chong, Scott Rankin, Aaron Zorn and Ondine Cleaver. Rasip1 is required for endothelial cell motility, angiogenesis and vessel formation. Developmental Biology . 2009; 329(2):269 - 79. 4) Ondi ne Cleaver, Dan Quiat, Ke Xu , Alethia Villasenor. Morphogenesis of blood vessels during mouse vasculogenesis (Ab

9 stract) . Developmental Biology . 2007
stract) . Developmental Biology . 2007; 306(1):445. 5) YuRui Zhao, Quan Zheng, Kenneth Dakin, Ke Xu , Manuel L. Martinez, and Wen - hong Li. New Caged Coumarin Fluorophores with Extraordinary Uncaging Cross Sections Suitable for Biological Imaging Applications. J. Am. Chem. Soc . 2004; 126(14) pp4653 – 4663 (Article). 6) Ying - Jie Wang, Wen - hong Li, Jing Wang, Ke Xu , Ping Dong, Xiang Luo, and Helen L. Yin. Cr itical role of PIP5KI  87 in InsP 3 - mediated Ca 2+ signaling. Journal of Cell Biology . 2004; 167(6) 1005 - 1010. xii LIST OF FIGURES Figure 1.1. Development of the vascular systems ................................ ................... 2 Figure 1.2. Vasculogenic tubulogenesis: cord - to - tube transition ........................... 4 Figure 1.3. Angiogenic tubulogenesis: lumen extension ................................ ........ 6 Figure 1.4. Junctional contacts in epithelial and endothelial tubes ........................ 9 Figure 1.5. Vascular tubulogenesis: Cell hollowing vs. Cord hollowing ............ 1 2 Figure 1.6. Vascular tubulogenesis: mechanistic he terogeneity .......................... 1 5

10 Figure 3.1. The regulation of Ras a
Figure 3.1. The regulation of Ras activity ................................ ............................. 37 Figure 3.2. Expression of Rasip1 in vascular endothelium during early embryogenesis ................................ ................................ ................................ ........... 4 1 Figure 3.3. Vascular expression of Rasip1 in embryonic organs and tissues ...... 44 Figure 3.4. Expression of Rasip1 is restricted to VEGFR2 - dependent endothelium ................................ ................................ ................................ ............... 47 Figure 3.5. Rasip1 ablation in MS1 cells by transient siRNA transfection hinders endothelial tube formation and migration ability ................................ .................... 48 Figure 3.6. Conservation of Rasip1 sequence across species ............................... 50 xiii Figure 3.7. Expression of Xenopus tropicalis Rasip1 transcripts in frog embryos by in situ hybridization marks the developing embryonic blood vessels .............. 52 Figure 3.8. Rasip1 knowdown in frog embryos results in failure of blood vessel formation ...........

11 ..................... ..................
..................... ................................ ................................ .................... 54 Figure 4.1. Rasip1 gene targeting scheme ................................ ............................. 6 1 Figure 4 . 2 . Rasip1 is essential for mammalian vascular morphogenesis ............ 63 Figure 4 . 3 . Rasip1 is essential for vascular tubulogenesis in all blood vessels 65 Figure 4 . 4 . Rasip1 and Arhgap29 are required for in vitro EC lumen formation 68 Figure 4.5. Rasip1 and Arhgap29 are required for regulation of small GTPase signaling ................................ ................................ ................................ ..................... 7 1 Figure 4.6. Rasip1 and Arhgap29 regulate EC architecture and tubulogenesis .. 73 Figure 4.7. Rasip1 and Arhgap28 regulate EC archite cture through Rho family GTPases ................................ ................................ ................................ ..................... 75 Figure 4.8. Rasip1 and Arhgap29 are required for maturation of endothelial – ECM adhesion ................................ ................................ ............

12 .................... ........... 78
.................... ........... 78 Figure 4.9. Rasip1 - / - angioblasts remain cuboidal and fail to adhere to surrounding ECM ................................ ................................ ................................ ...... 8 1 xiv Figure 4.10. Rasip1 - / - angiobloasts display defective cell polarity and fail to localize junctional proteins to cord periphery ................................ ......................... 84 Figure 4.11. Rasip1 regulates EC contractility and adhesion by modulating Rho family small GTPases ................................ ................................ ............................... 86 Figure 4.12. The balance betwee n EC - EC and EC - ECM adhesion is critical to vascular cord to tube transition ................................ ................................ ................ 88 xv LIST OF TABLES Table 2.1. Primers used in making in situ probes ................................ .................. 24 Table 2.2. Primers for generating and genotyping Rasip1 null allele .................. 29 xvi LIST OF ABBREVIATIONS AJ Adhesion Junction BMP Bone Morphog

13 enetic Protein BSA Bovine Seru
enetic Protein BSA Bovine Serum Albumin cDNA Complementary DNA DIG Digoxigenin DNA Deoxyribonucleic Acid EC Endothelial Cell EGFP Enhanced GFP EST Expressed Sequence Tag FA Focal Adhesion FB Fibrillar Adhesion FBS Fetal Bovine Serum Flk1 Fetal Liver kinase 1 FN Fibronectin FX Focal Complex GAP GTPase Activating Protein GDP Guanosine 5’ - diphosphate xvii GEF Guanine Exchange Factor GFP Green Fluorescent Protein GTP Guanosi ne Triphosphate HPRT Hypoxanthine - guanine Phosphoribosyltransferase ISV Intersomitic/Intersegmental Vessel KDR Kinase insert Domain Receptor MO Morpholino PBS Phosphate Buffered Saline PCR Polymerase Chain Reaction PECAM Platelet Endothelial Cell Adhesion Molecule PFA paraformaldehyde RT - PCR Reverse Transcriptase - PCR siRNA small interfering RNA TGFβ Transforming Growth Factor beta Tie2 Tyrosine kinase with immunoglobulin - like and EGF - like domain 2 TJ Tight Junction VE GF

14 Vascular Endothelial Growth Factor
Vascular Endothelial Growth Factor xviii VEGFR2 VEGF Receptor 2 TEM Transmission Electron Microscopy 1 CHAPTER I Introduction The cardiovascular system is the first functional organ system to form in the developing embryo s across all vertebrate species, providing tissues with nutrient and gas exchange required for life . Defects in the structure and/or function of the cardiovascular system lead to early e mbryonic lethality . T he vasculature emerges from aggregation of angioblasts . Angioblasts are endothelial precursors that arise from mesodermal cells that differentiate either within blood islands, structures composed of hematopoietic cells (blood cell prec ursors) surrounded by a mantle of angioblasts, or within embryonic tissues as scattered cells in both extraembryonic and embryonic tissues . Vessels subsequently form via v asculogenesis , or the coalescence of individual aniolasts “ in situ ” to for solid priitive vascular „cords’, which then undero tuuloenesis ( Drake and Fleming, 2000 ; Risau and Flamme, 1995 ) . These lumenless, linear ag gregates then transform into tubes to carry blood. The first ves

15 sels consist of a relatively simple and
sels consist of a relatively simple and h omogeneous endothelial cell EC network of vessels, often termed a „plexus’. Susequently, the coplexity of the vasculature increases draatically 1 2 Figure 1.1. Development of the vascular systems (adapted and redrawn from ( Carmeliet, 2005 ) ). During vasculogenesis, angioblast derived ECs form a primitive vascular plexus de novo. This initial network is then undergo a process called angiogenesi s including the remodeling, sprouting and pericyte/smooth muscle cells (PC/SMC) stabilizing of the vessels to form a mature vasculature containing a complex network of large and small vessels. Lymphangiogenesis initiates from veins. 3 as new vessels sprout from pre - existing vessels via sprouting angiogenesis ( Carmeliet, 2000 ; Risau, 1997 ) . Sprouting extends existing lumens and rearrange the initially simple, net - like, primary tubular network , via remodeling angiogenesis , into a complex hierarchical system , which includes specialized ECs, such as arteries and veins. As these vessels mature and stabilize, they become ensheathed by smooth muscle cells and pericytes. Howeve

16 r, they continue to grow coordinately
r, they continue to grow coordinately with organs and tissues, providing the tissues they perfuse with the nutrients and oxygen required for viability ( Figure 1.1 ) . Vasculogenic tubulogenesis : cord - to - tube transition Formation of a cohesive seamless and contiguous network of blood vessels to carry blood is essential for proper cardiovascular function. However, while a growing understanding is emerging regarding the specification, patterning and sprouting of blood vesse ls, the mechanisms underlying the cellular morphogenesis and molecular pathways that direct vascular tubulogenesis are only beginning to be unraveled. Tubulogenesis is a fundamental process that is essential for the development of many tubular organs. Duri ng vasculogenesis, the angioblast - formed linear vascular cords undergo a morphogenic change during which the angioblasts transition from a cuboidal to a flattened shape to encircle a 4 Figure 1.2. Vasculogenic tubulogenesis: cord - to - tube transition . ( A ) Morphogenic changes of angioblasts from single cells to linear cords and patent tubes. ( B ) TEM images and cartoons showing two angioblasts before and after cord - to - tube transi

17 tion. Junctional molecules are represen
tion. Junctional molecules are represented as blue bars. asterisk in TEM region : vascular lumen between two ECs. 5 central lumen ( Figure 1.2 A ). This step is fundamental to establishing a functional circulatory system. Dramatic cellular changes take place during this process, including the re - localization/distribution of junctional molecules to cell periphery allowing the opening of the initial luinal „slit’ etween 2 - 3 ECs ( Figure 1.2B ). Angiogenic tubulogenesis : lu men extension Similar to de novo lumen and tube formation in initial embryonic vessels, lumens must also form in new angiogenic sprouts. However, the mechanisms are likely to differ to some extent, as initial vessels start as solid cords with no connection s to existing lumens, while angiogenic vessels grow from pre - existing tubes, with pre - existing lumens ( Figure 1.3 ). Much work has recently focused on sprout formation and the role of VEGF and Notch signaling during this process. The distal growing end of a nioenic vessels consist of endothelial „tip’ cells, which extend long filopodia and migrate in a manner similar to axonal growth cones, sensing their microenvironment. The

18 proximal end of these vessels, by contr
proximal end of these vessels, by contrast, consists of „stalk’ cells that ay eit her form a trailing cord or quickly become lumenized ( Gerhardt et al., 2003 ; Holderfield and Hughes, 2008 ) . The process that contr ols tip cell growth is dependent on a remarkable interplay of selective Notch ligand activation, as well as the establishment of well - controlled VEGF gradients ( Chappell et al., 2009 ) . To date, however, relatively little 6 Figure 1.3. Angi ogenic tubulogenesis: lumen extension . 7 attention has been focused on the timing, mechanics or molecular regulation of lumen formation in stalk cells. Comparing and contrasting lumen formation in different types of vessels will reveal whether similar mechanisms apply to all vessels or whether locally dis tinct modes of lumen formation will reflect inherent heterogeneity of vessels. Cellular mechanisms of vascular tubulogenesis As a tubular organ, the endothelial vasculature shares a lot of common features with epithelial tubes found in a number of other o rgans. A considerable amount of work has been carried out to elucidate epithelial tubulogenesis during the past several decades. Epith

19 elial tubes are present in many organs
elial tubes are present in many organs including lung, kidney, salivary glands and pancreas. These tubes are generally com posed of a sheath of cuboidal or columnar epithelial cells, with defined apical membranes facing a central lumen, their lateral edges interfacing with each other closely, and their basal membranes making up the tube periphery ( Figure 1.4A ). Similarly, ECs of functional vessels consist of a luminal (apical) membrane facing the f l owing blood and an abluminal (basal) membrane in contact with the basement membrane. The principal difference between epithelial and endothelial tubes is that the junctional contacts (lateral membranes) between ECs consist of a much smaller contact area ( Figure 1.4B ). 8 Figure 1. 4 . Junctional contacts in epithelial and endothelial tubes. Compared to lumen forming epithelial cells, endothelial cells display much smaller juncti onal contacts/Lateral membranes. 9 Nonetheless, a number of analogies between the two tissue types can be made. Depending on the types of organs and microenvironments, most epithelial tubulogenesis processes fall into one or a combination of several of the following catego

20 ries: wrapping (i. e. vertebrate primary
ries: wrapping (i. e. vertebrate primary neurulation), budding (i.e. lung), cell hollowing (i.e. Drosophila trachea), cord hollowing (i.e. zebrafish gut), cavitation (i.e. salivary gland), and cell division and intercalation (i.e. zebrafish neural tube) ( Andrew and Ewald, 2010 ; Lubarsky and Krasnow, 2003 ) . Among these mechanisms, at least three have been implicated in endothelial lumen formation: budding, cell hollowing and cord hollowing ( Iruela - Arispe and Davis, 2009 ) . (1) Budding By definition, budding consists of formation and extension of a tube via growth from a pre - existing tube. During lung development, the pulmonary epithelium repeatedly buds and extends finger - like projections, until it becomes a continuous, highly branched, tubular tree. During blood vessel formation, sprouting angiogenesis is essentially synonymous with budding ( Figure 1.3 ).ECs along the wall of a blood vesse l become locally activated, degrade the su rrounding basement membrane, cells migrate out, with tip cells at the leading front of the growing vessel invading surrounding tissue. As angiogenic sprouting gives rise to patent vessels, it can thereby be conside red one mechanism for lumen

21 10 formation. Howeve, what proce
10 formation. Howeve, what processes specifically drive stalk cells to organize, extend and maintain the parental lumen remain unclear. (2) Cell hollowing: vacuole fusion In contrast to budding where new lumens extend directly from pre - existing lumens, cell hollowing represents a different cellular mechanism whereby new lumens emerge intracellularly. In this case, lumens initiate as multiple small vesicles or larger vacuole s, that fuse to produce a central lumen, which in turn becomes connected with similar lumens in adjacent cells ( Figure 1.5A ). This type of lumen formation has been observed in Drosophila terminal tracheal cells ( Guillemin et al., 1996 ; Levi et al., 2006 ) , as well as in ECs. Indeed, until recently vascular lumen formation h ad primarily been studied in vitro, and a large body of work using live imaging of ECs in 3D matrices has demonstrated that intraendothelial lumen formation occurs by cell hollowing via vacuole fusion ( Davis et al., 2000 ; Iruela - Arispe and Davis, 2009 ; Sacharidou et al., 2010 ) . Vacuole fusion has also been observed in vivo , during formation of intersegmental vessels in zebrafish ( Kamei et al., 2006 ) . Together, these o

22 bservations suggested that cell hollowi
bservations suggested that cell hollowing was an important mechanism in both endothelial and non - endothelial tubulogenesis. 11 Figure 1. 5 . Vascular tubulogenesis: Cell hollowing vs. Cord hollowing. 12 (3) Cord hollowing: vacuole fusion and cell rearrangement A third type of mechanism by which some lu mens form is termed cord hollowing. In this process, lumenal space s generated extracellularly between ECs, even as they remain joined peripherally ( Figure 1.5B ). Expansion of apical membranes to create an intervening extracellular space can be achieved eit her by addition of new lumenal membrane, or by removal or clearance of junctions from the cord center. The result of either process is to build up the net surface area of the lumenal membrane. Addition of new lumenal membrane du ring lumen formation has be en shown in some cases to occur via directed exocytosis of vesicles, which fuse with and expand the lumen at the cord center. This type of vectorial vacuole fusion has been observed in cultured MDCK epithelial cells ( Lipschutz et al., 2000 ; Lubarsky and Krasnow, 2003 ; Vega - Salas et al., 1988 ) . It has also been shown in endotheli

23 al cells and suggested to be dependent o
al cells and suggested to be dependent on Rab7 - directed centripetal transport of vacuoles ( Zovein et al., 2010 ) . Interestingly, both cell polarity and proper control of cell adhesion appear to provide a critical framework for this process, as loss of β1 interin disrupts Par3 localization resultin in luen failure. 13 Vascular tubulogenesis: mechanistic heterogeneity Tubulogenesis has fascinated developmental and cell biologists for decades and recent efforts have been directed towards understanding it in vertebrate blood vessels ( Hogan and Kolodziej, 2002 ; Stuart et al., 1995 ; Zeeb et al., 2010 ) . As the number of studies has increased, our understanding of the underlying mechanisms has evolved. Until recently, cell hollowing was considered a common, if not predominant, mode of de novo endothelial lumen formation. Clear live imaging of ECs cultured in 3D matrices provided strong support for vacuole - based lumen formation ( Davis et al., 2000 ) . Cells were shown to generate intracellular vacuoles that would align at the cell center and fuse with each other to create lumens. Similarly, vacuole fusion has been observed in the growing vessels of vert

24 ebrate embryos ( Blum et al., 2008 ; K
ebrate embryos ( Blum et al., 2008 ; Kamei et al., 2006 ; Liu et al., 2011 ; Wang et al., 2010 ) ( Figure 1.6A ). Live imaging of growing zebrafish intersegmental vessels (ISVs) identified fusing vacuoles during lumen fo rmation ( Kamei et al., 2006 ) . This study examined at high resolution the dev elopment of ISVs, which were thought to assemble stereotypically with three ECs in a head - to - tail cord along myotomal boundaries ( Isogai et al., 2003 ) . Two photon live cell imaging identified vacuoles during angiogenic sprouting and suggested intra - cellular fusion of endothelial vacuoles at the center of ISV ECs suggesting cell hollowing. 14 Also required for ex pansion of extracellular space and tubulogenesis is the de - adhesion and/or redistribution of junctional molecules at the cord center. In this case, cells within a cord create a lumen by de - adhering from each other locally at the luminal membrane, but remai ning tethered at the lumen periphery. This differential adhesion thereby alters EC shape and rearranges ECs relative to each other. Alternately, ECs might also redistribute existing junctions to the periphery, away from the lumen. Such junctional redistrib ution

25 has been observed in the gut epithelium
has been observed in the gut epithelium of zebrafish ( Horne - Badovinac et al., 2001 ) and more recently during vasculogenesis in mouse ( Xu et al., 2011 ) ( Figure 1.6 C ). To date no definitive experimental evidence has clearly dist inguished lumenal membrane expansion, versus either de - adhesion or clearance of junctions, however it is very possible that a number of cellular phenomena occur coordinately during lumen formation. Indeed, both directed vacuolar transport and junctional re distribution have been observed to occur concurrently in the arterioles of late gestation mouse embryos ( Zovein et al., 2010 ) . These vessels arise first as cords of cuboidal ECs, which then open a central lumen following redistribution of junctional molecules to the cord periphery (away from the center) and Rab7 - directed vesicle transport to the lumenal membrane ( Figure 1.6 B ) . It will be of great interest to assess whether both mechanisms apply more globally to forming vessels. 15 Figure 1. 6 . Vascular tubulogenesis: mechanistic heterogeneity. ( A ) Vacuole fus ion. ( B ) integrin induced cell polarity formation. ( C ) Rasip1 - Rho regulated integrin - acto - myosin activi

26 ty. ( D ) Zebrafish cardinal vein forma
ty. ( D ) Zebrafish cardinal vein formation. ( E ) Repulsive signal mediated Drosophila heart tube formation. ( F ) Acto - myosin and negative charges mediated v ascular slit initiation. 16 The existence of a variety of mechanisms is highly likely, however, given the endothelial heterogeneity that is known to exists in different vascular beds. One example involves lumen formation during initial arteriovenous segregati on in zebrafish ( Herbert et al., 2009 ) ( Figure 1.6D ). High - resolution imaging of zebrafish vascular morphogenesis showed that the primary axial dorsal aorta (artery) arises first by vasculogenic aggregation of angioblasts. Secondarily, the caudal ve in develops via selective ventral sprouting and migration of angioblasts from the dorsal aorta, a process they show to be regulated by EphrinB2 - EphB4 signaling. Interestingly, the caudal lumen forms via angioblasts aggregating into a partially fored and „ open’ vessel, which then rapidly fills with lood accuulated at the interface of the two vessels. Upon this „fillin’ of the caudal vein, a cardiovasculature with functional circulation is then rapidly established. The cellular mechanisms underlying

27 this „open’ vessel foration reï
this „open’ vessel foration reain to e examined, but suggest the probability of a variety of mechanisms underlying tubulogenesis. Yet another diverent echanis for “lood vessel” luen foration was reported during Drosophila heart morphogenesis ( Medioni et al., 2008 ; Santiago - Martinez et al., 2008 ) ( Figure 1.6E ). In flies, the heart represents an open - ended and contractile endolyph „vessel’ of sorts, with tuular orpholoy and a distinct internal lumen. In these studies, formation of the cardiovascular lumen by two parallel rows of myoendothelial cells is shown to be modulated by membrane 17 repulsion. S lit - Robo signaling downregulates E - cadherin at the lumenal cell - surface and prevents fusion between apposed cells. Instead, the two cells form junctions only at their dorsal - and then ventral - most regions, resulting in the formation of an internal central lumen, enclosed by two rows of cells. Even more interestingly, distinct from typical epithelial tubulogenesis process es , the fly heart lumen formed by this mechanism is created by the basal rather than apical cell surfaces, suggesting that fundamental diff erences ca

28 n occur between different types of epit
n occur between different types of epithelial and endothelial tube forming cells with respect to cell polarity features. A similar lumenal repulsion mechanism has also been suggested during mouse dorsal aortae formation. Following cord hollowing, observations by Lammert and colleagues showed that accumulation of negatively charged sialomucins along apposed lumenal membranes results in the initial opening of slit - like spaces between ECs ( Strilic et al., 2010 ; Strilic et al., 2009 ) ( Figure 1.6F ). Specifically, they show that sialic acids of lumenal glycoproteins create repulsive electrostatic fields that result in membranes moving away from each other at the cord center. Neutralizing these charges with injection of cationic protamine sulfates inhibits normal lumen formation. Taken together, these many divergent examples of lumen f ormation mechanisms suggest that different vessels may form via a range of different 18 cellular mechanisms. It is not completely unexpected, as endothelium is known to display a high level of heterogeneity across different tissues ( Aird, 2003 ) . We propose that distinct or even yet to e discovered „novel’ echaniss of luen forma

29 tion will likely be identified as vascul
tion will likely be identified as vascular beds of differ ent organs are more extensively examined and understood. 19 CHAPTER II Materials and methods Isolation of Rasip1 sequence A pYX plasmid containing mouse Rasip1 cDNA piece (1047 – 3170bp, spanning exons 4 through 7) was obtained from OpenBiosystems ( BC072584 ). For making longer in situ probes, the full - length coding region (2886bp) of Rasip1 was amplified from E8.5 mouse cDNA by RT - PCR, using primer set P1 (Table 2.1). Xenopus tropicalis Rasip1 partial coding region sequence (1151bp, exon2 - exon7) wa s cloned by RT - PCR using primer set P2 . The amplified fragment s w ere subcloned into pGEM - T - Easy Vector (Promega) by TA cloning. Mouse strains VEGFR2 null embryos were generated by mating Flk1(VEGFR2) - lacZ heterozygous males and females (kindly provide d by Drs. Janet Rossant and Eli Keshet). Embryos were dissected manually in ice cold PBS. Embryos lacking blood vessels were identified visually, by the absence of yolk sac blood vessels, 19 20 and genotypes (of either embryos or adults) were confirmed by PCR, u sing primers to lacZ (Table 2.1), which yield a 630bp

30 PCR fragment. Embryos and Histology
PCR fragment. Embryos and Histology CD1 mice embryos were collected from pregnant females (E7.5 through E15.5) after dissection in ice - cold PBS buffer and fixed in 4% PFA in PBS solution overnight at 4  C with gentle rocking. The amnion was removed during dissection for better probe penetration. Embryos were washed three times in PBS for 5 min, and dehydrated using a series of ethanol washes. Embryos were then stored in 75% ethanol at – 20  C. Postnatal t issue was collected and fixed in a similar manner. For wax sectioning of embryos following in situ hybridization, the embryos were fixed and dehydrated as described above. Embryos were rinsed twice in 100% ethanol for 5 min, twice in xylene at RT for 10 min, then a mixture of 1:1 paraplast : xylene at 60  C for 10 min, then a series of 100% paraplast at 60  C (McCormick Scientific). The embryos were then embedded and sectioned with a Biocut 2030 microtome. For examination, the sections were placed o n glass slides, deparaffinized in xylene twice for 5 min each and mounted on SuperfrostPlus glass slides (Fisher) using Permount (Fisher). 21 DIG - labeled RNA probes Proe synthesis was carried out

31 at 37˚C for 2 h ou rs: 1µg linearized
at 37˚C for 2 h ou rs: 1µg linearized plasmid, 2.0µl DIG - RNA lab eling mix (Roche), 2.0µl 10X transcription buffer (Roche), 1.5µl Placental ribonuclease inhibitor (Promega), 1.0µl T3/T7 RNA polymerase (Roche), RNase free water to a final volume of 20µl. DNA template was removed using 2µl RQ1 DNase I (Promega), at 37  C f or 15 min. The probes were then purified with Micro Bio - spin columns (Bio - RAD). 10x hybridization stock solution was prepared at a concentration of 10µ/l in „prehy’ solution: 50% Formamide (Fisher), 5xSSC (pH 4.5), 50µg/ml Ribonucleic acid from Torula y east, Type VI (Sigma), 1% SDS, 50µg/ml Heparin (Sigma). Stock solution is stored at – 80  C. Whole mount in situ hybridization Whole mount in situ hybridization in mouse embryos was carried out usin a protocol adapted fro D. Wilkinson’s Method ( Wilkinson, 1999 ) . Briefly, e mbryos stored in 75% ethanol at – 20˚C were rehydrated in stepwise fashion to PBST. Then, t he embryos were treated with 10µ g/ml pro teinase K (time treated varied with age of tissue; 2min - 30min), fixed in a 0.2% gluteraldehyde/4% PFA solution, and pre - hybridized at 65  C for 1 hour. The samples were

32 transferred into 22 hybridizat
transferred into 22 hybridization mix, containing 1µg/ml Dig - labeled probes described above . The in situ hybridization post - hybridization washes and antibody incubation were carried out using a Biolane HTI automated incubation liquid handler (Holle & Huttner). Color development was carried out using BM purple solution (Roche). Frog in situ hybri dization was carried out using a similar standard in situ hybridization protocol ( Costa et al., 2003 ) . Zebrafish in situ s were performed as previously described ( Neumann et al., 2009 ) . In situ hybridization on sections Paraffin sections (on glass slides) were washed 3x3min in PBS, followed by a 10min treatment with 15  g/ml proteinase K. Sections were then rinsed in PBS, fixed in 4% PFA for 5 min, and incubated for 10min in acetylation solution: mix of 2.66ml Triethanolamine, 350  l HCl, 750  l acetic anhydride and 200ml water. Prehybridization was carried out in plastic slide mailers (Fisher) containing hybri dization buffer at RT for 1 hour. Slides were then transferred to a humidified chamber (humidified with 50% formamide/5xSSC) for probe hybridization (probe at 1  g/ml) with 100  l probe/sli

33 de (covered with glass coverslips) at 68
de (covered with glass coverslips) at 68°C overnight. 23 Slides were was hed post - hybridization in 2xSSC at 72°C just long enough to allow coverslips to separate. Then slides were rinsed in 0.2xSSC at 72°C and RT for 1x1min, respectively, then MBST buffer at RT (100mM Maleic acid, 150mM NaCl, pH7.5, 0.1% Tween20). Slides were i ncubated in blocking solution (2% blocking reagent (Roche) and 5% heat - inactivated sheep serum in MBST) for 1 hour at RT. Anti - D IG alkaline phosphatase conjugated antibody was applied on slides in a chamber humidified with MBST (250  l of 1/4000 anti - D IG an tibody (Roche)), covered with parafilm and incubated at 4°C overnight. Slides were washed for 3x30min in MBST after antibody incubation, and treated in NTMT (100mM NaCl, 100mM Tris, pH9.5, 50mM MgCl2, 0.1% Tween20) for 3x5min. Color reaction was carried ou t using BM purple as described above. For microscopic examination, slides were sealed and coverslipped using Permount (Fisher). 24 Table 2.1. Primers used in making in situ probes Primer Sequence P1 Mouse Rasip1 in situ probe 5’ - ATGCTATCTGG TGAACGAAAG - 3’ 3’ - TCAAGGTGTCGAAGCCACCG - 3’ P2 Xen

34 opus tropicalis Rasip1 in situ prob
opus tropicalis Rasip1 in situ probe 5’ - ATTAAGGGAAAGAGAAGAAAGCATCT - 3’ 5’ - GCATACAGTGTCTTGGTCAGATAATATAC - 3’ P3 VEGFR2 in situ probe 5’ - GACGGAGAAGGAGTCTGTGC - 3’ 5’ - GGGACAGGACCACTTCCAT - 3’ P4 Primers to LacZ cassette 5’ - GGTGGCGCTGGATGGTAAGC - 3’ 5’ - CGCCATTTGACCACTACC - 3’ β - Galactosidase reaction E mbryos (or isolated organs) were fixed in 5mM EGTA (pH 8.0), 0.2% gluteraldehyde, 2mM MgCl 2 and PBS solution for 5 min on ice. After fixation, embryos were rinsed 3 times for 5 min in PBS. 50 mM Potassium Ferrocyanide (K 4 Fe(CN) 6  3H 2 O) and Potassium Ferricyanide (K 3 Fe(CN) 6 ) solutions, stored at RT in dark, were used to make lacZ staining solution: 20 mM K 4 Fe(CN) 6  3H 2 O, 20 mM K 3 Fe(CN) 6 , 2 mM MgCl 2 , 0.02% NP - 40, add wate r or 1xPBS to 500 µl. 25 Staining solution was warmed to 37  C before adding X - Gal (Growcells) to avoid X - Gal precipitation. 4 µl of 100 mg/ml X - Gal stock (in dimethyl formamide) was then added to the lacZ staining solution. Embryos were placed in staining sol ution and color reaction was allowed to develop at 37  C overnight. When staining was evaluat

35 ed to be optimal, embryos were washed wi
ed to be optimal, embryos were washed with PBS 3 times for 5 min each, post - fixed in 4%PFA overnight, and transferred to 80% glycerol for viewing. siRNA transfecti on siRNAs obtained from Idt - DNA were transfected as per Idt - DNA’s standard protocol . Briefly, for the 24 - well format, 1.25µl of 20µM dicer substrate siRNA was diluted in 50µl of Opti - MEM I Reduced Serum Medium (Invitrogen) for each well . 1 µl of Lipofectam ine 2000 (Invitrogen) transfection reagent was diluted in another 50µl of Opti - MEM I Reduced Serum Medium. After 5 minutes incubation at RT, the diluted siRNA and the transfection reagent were combined together, and incubated for 20 minutes at RT. MS1 cell s (ATCC) were plated on a 24 - well plate with a density of 5  10 4 cells/well in 400µl DMEM containing 10% FBS and without penicillin/streptomycin. The pre - mixed 100µl transfection complexes were then added drop - wise on top of the cells. After gentle mixing by rocking the plate back and forth, the cells were incubated at 37˚C in a 5% CO2 26 incubator prior to following assays. siRNAs obtained from Invitrogen, and Dharmacon were transfected into cultured endothelial cells using stand

36 ard protocol as previously described (
ard protocol as previously described ( Koh et al., 2008 ) . Endothelial cell assays „Tue - foration assays’ we re carried out in a 96 - well plate. 50µl of Matrigel (BD Matrigel 354234) was thawed on ice and plated on the bottom of each well. ECs cultured in one well of a 24 - well plate (90% confluency) were trypsinized, plated in one Matrigel coated well of a 96 - well plate and cultured at 37˚C. When usin wild type cells, the anioenic areation of ECs (or „tues ’ ) starts to occur within a few hours. For better viewing, cells were stained with - AM (Cell Biolabs) before microscopic examination. Quantification of angiogenic branchpoints was accomplished by counting observable branchpoints within 8 representative areas within each plated well. „Wound - healin’ assays were carried out 72 hours post - transfection. Briefly, the cell monolay er is scratched using a sterile P200 pipet tip to create a „cell - free’ area (the wound). The cells were then washed once with DPBS. Cells are then allowed to recover and irate into the „cell - free’ area. Iaes were 27 acquired immediately after scratching a nd rinsing, and also a

37 fter an overnight incuation at 37˚C
fter an overnight incuation at 37˚C for coparison of wound width. Distance irated was calculated as half of the total change in width. Morpholino (MO) knockdown of Xenopus Rasip1 Xenopus tropicalis embryos were injected with 16ng Rasip1 - MO (Gene - tools) into 1 cell at the 2 - cell stage for assessment of vascular defects using in situ hybridization, or into both cells for assessment of transcript knockdown by RT - PCR. Embryos were allowed to develop to stage 32, then fixed in preparat ion for in situ hybridization. Morpholino - injected embryos were fixed in MEMFA (0.1M MOPS pH7.4, 2mM EGTA, 1mM MgCl 2 , 4% PFA), transferred to 100% ethanol and stored at - 20  C. For evaluation of transcript knockdown efficiency, embryos were allowed to devel op to either stage 25/26 or 29/30 and frozen directly on dry ice for RT - PCR. Generation of Rasip1 null allele The mouse 129sv genomic DNAs (provided by UTSW transgenic core) were used as template for the generation of a conditionally null allele by 28 amplif ying a region of DNA flanking exon 3 and two additional homologous arms 5’ and 3’ usin Takara’s LA Taq polyerase. The three suclones were clon

38 ed into a neomycin/thymidine kinase dou
ed into a neomycin/thymidine kinase double selection vector pGKneo - TK2 - Floxed - Flip (kindly provided by Thomas C arroll). The exon 3 containing subclone was inserted into a multiple cloning site flanked by loxP sites. Neomycin resistance cassette was designed being flanked by Flp recombinase target (FRT) sites for removing if necessary. This targeting vector was elec troporated into mouse eryonic ste cells y UTSW transenic core. 5’ and 3’ external proes for Southern blot screening were PCR amplified and subcloned into pGEM - T - Easy vector (Invitrogen). SphI digested purified genomic DNA from each clones was screene d by Southern blot using both probes for homologous recombination. The correct clone was injected into C57BL6/J blastocysts for generating Rasip1 targeted mouse line. After germ line transmission confirmation, the animals were bred with Sox2 - Cre mice (kind ly provided by Thomas Carroll) to generate a Rasip1 null allele. Litters were genotyped by PCR using primers listed in the Supplemental Experimental Procedures. The wild type, floxed, and null alleles give bands of 165 base pairs, 247 base pairs, and 331 b ase pairs, respectively. 29

39 Table 2.2 . Primers for generating and
Table 2.2 . Primers for generating and genotyping Rasip1 null allele Primer Sequence P5 5’ hooloous ar 5’ - TCGGTACCCAGTTGGCATCGTGCTTCTA - 3’ 3’ - GCGTCGACAGGGGATGTGTACTGCGTTC - 3’ P 6 3’ hooloous ar 5’ - TCTTAATTAAGGCTTGCTCTTTCACAGGACCCTGG - 3’ 5’ - TCTTAATTAATATGACAGCGGGACAGAGTGGGC - 3’ P 7 Rasip1 Exon3 5’ - GTCCCTGCAGGCTCTCTGTTCACCTATTTCTTACCAAGG - 3’ 5’ - GCTCCTGCAGGCCACGGCCACACAAACGACAAGAA - 3’ P8 5’ Southern lot proe 5’ - GTCCAATACGCATAACCTGTTCT - 3’ 5’ - CGCCATTTGACCACTACC - 3’ P9 3’ Souther lot proe 5’ - CAGCTCTGCAATGACTTGGA - 3’ 5’ - CAAAACCAAAACCCAACCTG - 3’ P10 WT and floxed allele genotyping 5’ - GGGGTGACAGTGGAACACA - 3’ 5’ - CGGTGGGAAGAATGGAGATA - 3’ P11 Null allele genotyping 5’ - ATGGTATGCCTGCCATTTGT - 3’ 5’ - ACGACTGTGCCTTCATGTTG - 3’ 30 Cell Culture Human umbilical vein ECs (HUVEC, PCS100 - 010) and mouse pancreatic islet EC line MILE SVEN 1 (MS1, CRL - 2279) were obtained from ATCC (or Lonza) and cultured per ATCC’s standard protocols. 293AD cells (kindly provided ï

40 ¢y Eric Olson) were rown in Dulec
¢y Eric Olson) were rown in Dulecco’s Modified Eale Mediu (DMEM, GIBCO) with 10% Fetal Bovine Serum (FBS). Establishment of MS1 Rasip1 - FLAG cell line C - terminal FLAG tagged Rasip1 cassette was inserted into pEN_TTmiRc2 and then sub - cloned into pSLIK - Neo (kindly provided by Zhijian Chen) by gateway cloning. 10 µ g Rasip1 - FLAG expressing pSLIK vector was transfected into 293AD cells together with 7.5 µ g of each of the two packaging plasmids pLp1, and pLp2, and 5 µ g of the vesicul ar stomatitis virus (VSV) G envelope plasmid pLp - VSVG diluted in Opti - MEM (Invitrogen). The medium was changed 12 hours post - transfection, and the viral particles were harvested and filtered through 45 micron micro filters upon applying to MS1 ECs for infe ction. The infected MS1 cells were selected with 4mg/ml G418 (Sigma), and the expression of Rasip1 - FLAG were analyzed by western blot and immunofluorescent staining using a monoclonal anti - M2 antibody (Sigma). 31 Tandem Affinity Purification of Rasip1 Complex MS1 Rasip1 - FLAG cells were cultured with the presence of 1µg/ml doxycyclin for 3 days to induce the expression of exogenous Rasip1 - FLAG fusion protein. T

41 he cell lysate from ten 10cm dishes were
he cell lysate from ten 10cm dishes were incubated with 50µl Anti - FLAG M2 Affinity Gel (Sigma A2220) as per Sia’s standard protocol. The ound proteins were eluted with 200µl 0.2mg/ml FLAG peptide (Sigma F3290). The elutes were then incubated with 5µg Rasip1 antibody (Abcam ab21018) or IgG agarose (Sigma A0919, for control) at 4C for 1 hour. The Rasip1 an tibody bound protein complex were then incubated with 50µl rec - Protein G Sepharose 4B (Zymed 10 - 1241) at 4C for 1 hour, followed by elution with 40µl 0.1mg/ml human RASIP1 immunizing peptide (aa951 - 962, UTSW Peptide Synthesis Core Facility). The final elut es were resolved by SDS - PAGE and processed by silver staining using Invitroen’s SilverQuest kit (LC6070). Specific ands were excised and subjected to Mass Spectrometry as previously published ( Xu et al., 2009b ) . Immunofluorescent Staining Cells cultured on glass coverslips were washed twice in PBS and fixed in 4% Paraformaldehyde (PFA) at RT for 10 min, followed by permeablization in PBSN (0.1% NP40 in PBS) at RT for 15 min. The cells were then blocked in 32 CAS - Block (Invitrogen) for 1 hour a t 4C, followed by primary and secondary antibodies

42 labeling in CAS - Block. The stained cov
labeling in CAS - Block. The stained coverslips were mounted using VECTASHIELD HardSet Mounting Media with DAPI (Vector Labs H - 1500) and examined by epi - or confocal fluorescence microscopy. For whole mount immunofluorescence, the mouse embryos were fixed in 4% PFA at 4C for 1 hour, and primary antibody staining was carried out at 4 º C overnight. For sectioned mouse embryo immunofluorescence, the tissues were processed as previously described ( Villase nor et al., 2008 ) . If necessary, antigen retrieval was done using Retriever 2100 in their Buffer B. The sections were then blocked in CAS - Block (Invitrogen) for 1 hour at 4C, followed by primary and secondary antibodies labeling in CAS - Block in a humid chamber. The mounting and imaging conditions are the same as described above. EC lumen and tube formation in 3D collagen matrices HUVECs were suspended as single cells within 3.75 mg/ml collagen type I matrices and allowed to undergo EC morphogenesis as previously described ( Koh et al., 2008 ) . Cultures were fixed at indicated time points with 3% glutaraldehyde for several hours. In some cases, cultures were stained with 0.1% Toludine Blue containing 30% methanol and de - staine

43 d with MilliQ H 2 O, prior to 33
d with MilliQ H 2 O, prior to 33 photography, visu alization and analysis. Some 3D collagen gels were also extracted to examine protein expression or protein - protein interactions. 3D EC culture pull - down assays EC Cultures were lysed at the indicated time - points using cold detergent lysis buffer and incubated with S protein agarose beads as described ( Koh et al., 2008 ) . In separate experiments, supernatants were incubated with GST - PAK - PBD or GST - RhoA - PBD protein beads (in the absence or presence of 100 nM GDP) for 90 in at 4˚C to assess the deree of Cdc42/Rac1 or RhoA activation. The beads were washed four times with washing buffer. Bound active Cdc42/Rac1 or RhoA proteins were detected by Western blots. Flow Cytometry HUVE Cs detached from the dish by 50mM EDTA (pH8.0) treatment were washed with and resuspended as 10 7 /ml in labeling buffer (Ca 2+ free PBS containing 0.5% BSA). After incubation at 37C for 15 min with or without 5mM MnCl 2 , the cells were labeled with primary antibodies at 1:50 for 30 min on ice. Cells were then washed twice and labeled with fluorophore conjugated secondary 34 antibody in labeling buffer at 4C for 30 m

44 in. After final wash (twice in chilled
in. After final wash (twice in chilled PBS), cells were res uspended in 500µl 1% PFA and subjected to FACS analysis (UTSW Flow Cytometry Facility). Microscopy, Imaging and Statistical Analysis for in vitro Lumen Formation Visualization and image acquisition of EC lumen formation was performed using an inverted microscope (CKX41: Olympus) and real time - lapse imaging of ECs undergoing lumen and tube formation was done using a Nikon TE2000 - E system with a temperature - controlle d chamber. Image analysis was performed using MetaMorph software. Statistical analysis of EC vasculogenesis was performed using SPSS 11.0 software. Statistical significances were assessed by a paired - samples t test. 35 CHAPTER III Rasip1 is important for vascular morphogenesis Most of the molecular mechanisms responsible for blood vessel formation are not yet well understood. For decades, much attention was given to the role of vascular endothelial growth factor (VEGF) and its influence on EC migration and proliferation ( Ferrara et al., 2003 ; Yancopoulos et al., 2000 ) . Recently, however, discovery of a host of endothelial „uidance’ cues, which either attract or repel ECs an

45 d shape indi vidual blood vessels, has b
d shape indi vidual blood vessels, has broadened our understanding of how cell - cell signaling influences the morphogenesis of individual vessels and the vascular network as a whole. These signaling molecules include the Eph - ephrins ( Kuijper et al., 2007 ) , bone morphogenetic proteins (BMPs) ( Lebrin et al., 2005 ; Moser and Patterson, 2005 ; Park et al., 2006 ) , transforming growth factors (TGF  s) ( Lebrin et al., 2005 ) , Notch and Notch ligands ( Roca and Adams, 2007 ) and many others. In addition, a number of cell - autonomous factors have also recently been shown to be critical for proper EC behavior and blood ves sel formation. Many of these factors, such as small GTPases Ras, Rho, Rac, Cdc42, Pak and their many effectors/modulators ( Fryer and Field, 2005 ; Garnaas et al., 2008 ; Gitler et al., 2003 ; Kranenburg et al., 2004 ; Merajver and Usmani, 2005 ; 35 36 Tan et al., 2008 ) are already known to drive basic cell processes such as cell migration, cell proliferation an d establishment of cell polarity. Despite recent advances, the molecular mechanisms underlying much of blood vessel formation in vivo remain unclear, and elucidation of both extracellular

46 signaling events and cell autonomous re
signaling events and cell autonomous regulatory signaling cascades w ill advance our understanding of vascular specification and patterning, in both normal and pathological conditions ( Coultas et al., 2005 ) . Many Ras family members and their regulators have been implicated in vascular development ( Gitler et al., 2003 ; Henkemeyer et al., 1995 ; Tan et al., 2008 ) , including EC migration ( So snowski et al., 1993 ; Tan et al., 2008 ) , capillary tube assembly ( Connolly et al., 2002 ) , angiogenesis ( Aitsebaomo et al., 2004 ; Fryer and Field, 2005 ; Kranenburg et al., 2004 ; Merajver and Usmani, 2005 ) , blood vessel homeostasis ( Komatsu and Ruoslahti, 2005 ) and vascular permeability ( Serban et al., 2008 ) . Ras molecules are small GTPases widely shown to function as molecular switches coordinating multiple cellular behaviors like growth, proliferation, migrati on and differentiation. Ras GTPases cycle between the GTP - bound (active) and GDP - bound (inactive) states, under the influence of GAPs ( G TPase A ctivating P roteins), and GEFs ( G TPase E xchange F actors) ( Figure 3.1 ). 37 Figure 3.1. The regulation of Ras activity (Adapted from Figure

47 15 - 14, The Molecular Biology of the
15 - 14, The Molecular Biology of the Cell., 4 th edition. 2002) “ GTPase - activating proteins (GAPs) inactivate Ras by stimulating it to hydrolyze its bound GTP; the inactivated Ras remains tightly bound to GDP. Guanine nucl eotide exchange factors (GEFs) activate Ras by stimulating it to give up its GDP; the concentration of GTP in the cytosol is 10 times greater than the concentration of GDP, and Ras rapidly binds GTP once GDP has been ejected … The Ras GAPs maintain most of the Ras protein (~95%) in unstimulated cells in an inactive GDP - bound state. ” 38 Ras family prot eins have been shown to activate signaling cascades downstream of VEGF ( Cross et al., 2003 ; K ranenburg et al., 2004 ; Roberts et al., 2004 ) . VEGF stimulation of ECs i ncreases the amount of activated Ras , while dominant negative Ras constructs inhibit VEGF - induced endothelial proliferation, migration and assembly ( Meadows et al., 2001 ) . However, although the Ras pathway proteins have been implicated in vascular development their exact role is not well understood. Six years ago , Mitin and colleagues reported the identification of a novel Ras - interacting protein, Ra

48 sip1/Rain, which displays the characteri
sip1/Rain, which displays the characteristics of an endomembrane Ras effector ( Mitin et al., 2004 ) . Their experiments showed that Rasi p1 possesses a Ras - associating domain (RA), homologous to the RA domains of other Ras effectors, and that Rasip1 preferentially binds to the GTP - loaded form of Ras, both in vitro and in vivo . Its enrichment in adult lung and high expression in tr ansformed EC lines suggested the possibility that Rasip1 is expressed by ECs ( Mitin et al., 2006 ) . In an effort to discover unknown regulators of blood vessel development, we performed a microarray screen that transcriptionally profiled embryonic aortal ECs ( data not shown ). Among numerous EC - enriched transcripts, we identified Rasip1 . Here, we show that expression of Rasip1 is strikingly restricted to the endothelium of the developing vasculature, in both frog and mouse, and we 39 demonstrate that Rasip1 is essential for proper endothelial cell angiogenic assembly and migration, both in vivo and in vitro . We propose that Rasip1 plays important roles du ring vasculogenesis and angiogenesis, possibly regulating the function of Ras proteins in ECs. Identification of Rasi

49 p1 expression in murine endothelial cell
p1 expression in murine endothelial cells To identify sequences enriched in the embryonic dorsal aortae, we carried out Affymetrix microar ray screening of aortal ECs from E8.25 mouse embryos ( data not shown ). dChip ( Li and Wong, 2001 ) and Genespring software analysis was used to compare array data (to non - vascular array sets) and extract endothelial - enriched sequences. We initially identified Rasip1 as an EST ( AI853551 ) showing 50 - fold enrichment in ECs over other tissues. A longer clo ne was acquired commercially (OpenBiosystems), allowing production of Dig - labeled antisense probes encompassing the region from exon 4 through exon 12 ( ~2000bp) of the Rasip1 transcript. The Rasip1 genomic structure has been previously described ( Mitin et al., 2004 ) , however no developmental expression or function has been reported . 40 Rasip1 is expressed in vascular endothelium during vascular plexus formation (E7.5 - E10.0) Using in situ hybridization, we characterized embryonic expression of Rasip1 in mouse embryos and found it to be principally expressed in vascular endothelium. At E7.0 Rasip1 is initially detected in the parietal yolk sac, in a punctate

50 ring of cells (data not shown). Soon the
ring of cells (data not shown). Soon thereafter, at E7.5, expression expands to scattered cells of the extraembryonic yolk sac blood islands ( Fig ure 3 . 2 A ). At E8.0, individual cells expressing Rasip1 within the extraembryonic mesoderm can be observed at increasingly ventrolateral locations, in regions previously described as containing angioblasts ( Drake et al., 2000 ; Ferkowicz and Yoder, 2005 ) . The punctate appearance of Rasip1 expression in extraembryonic tissues at this stage suggests that these cells are angioblasts ( Fig ure 3 . 2 B ), as it closely resembles that of vascular endothelial growth factor receptor 2, VEGFR2 (or Fl k1/KDR ), and Tal1 both established markers for early angioblasts ( Drake and Fleming, 2000 ) . At E8.25 - E8.5, Rasip1 is strongly expressed throughout the embryonic and extraembryonic endothelium in a pattern recognizable as the primary vascular plexus, including the endocardium, the forming dorsal aortae and the primordia of the cardinal veins ( Fig ure 3 . 2 C,D ). During th ese stages, vasculogenesis of the principal embryonic blood vessels is occurring and major 41 Figure 3.2. Expression of Rasip1 in vascular endotheliu

51 m during early embryogenesis. ( A - F )
m during early embryogenesis. ( A - F ) Whole mount in situ hybridization showing whole stained embryos. Note expression in both scattered angioblasts (thin arrows), forming dorsal aortae (black arrowheads), intersomitic vessels (ISVs) (red arrowheads) and yolk sac vessels (thick arrows). ( G - I ) Transverse sections of in situ hybridizations showing endothelial - specific expression of Rasip1 in ( G ) dorsal aortae, ( H ) yolk sac vessels, and ( I ) heart endocardium. ( J - L ) Comparison of Rasip1 expression with that of the vascular markers VEGFR2 and PECA M, in E9.5 embryos. Note overall similarity of expression, especially in the ISVs (red arrowheads) and trunk vessels. Note difference in intensity of vascular staining in distinct regions, such as the cephalic vessels (red arrows). al, allantois; e, endoca rdium; en, endoderm; m, myocardium . The scale bars represent 200  m in all panels except J - L, where they represent 50  m. 42 vessels are taking shape (i.e. parallel dorsal aortae in Fig ure 3 . 2 D ) ( Walls et al., 2008 ) . As embryogenesis continues, Rasip1 expression continues to be expressed in developing blood vessels. After embryonic turn

52 ing, at E8.75, expression is evident in
ing, at E8.75, expression is evident in all large and small blood vessels, including the spro uting intersomitic/intersegmental vessels (ISVs) ( Fig ure 3 . 2 E ). Expression of Rasip1 within ISVs of later embryos ( Fig ure 3. 2 E,F ) suggests a role not only during vasculogenesis, but also during extension of vascular sprouts, or angiogenesis. Transverse sec tions through E8.5 embryonic tissues reveal that expression is restricted to the endothelium in all tissues examined, including the dorsal aortae ( Fig ure 3 . 2 G ) and yolk sac vessels ( Fig ure 3 . 2 H ). In addition, Rasip1 is expressed within the endothelium of t he endocardium, but not in myocardium ( Fig ure 3 . 2 I ). We compared expression of Rasip1 with that of other known vascular markers, such as VEGFR2 and PECAM ( Fig ure 3 . 2 J - L ), and found that Rasip1 outlined almost identical vascular structures in the developing embryo. For instance, expression of all three markers was observed in aortae, ISVs, endocardium and vessels of the lateral plate and head mesoderm. Of note, different vascular beds appeared t o express these three vascular markers with varying intensity. For instance, the head plexus of E9.5 emb

53 ryos expressed 43 VEGFR2 more
ryos expressed 43 VEGFR2 more robustly, while Rasip1 was more strongly expressed than either VEGFR2 or PECAM in the ISVs and endocardium ( Fig ure 3 . 2 J - L ). T hese differences reveal surprising endothelial heterogeneity at early stages of vascular development. Nonetheless, overall expression analysis suggests that Rasip1 is primarily restricted to vascular endothelium. Rasip1 during late embryogenesis (E10.5 - bi rth) Analysis of Rasip1 transcripts later during development reveals their expression in established vessels. Expression could indeed be detected in the blood vessels of various organs throughout midgestion stages ( Fig ure 3 . 3 ). Specifically, we found Rasip1 strongly expressed in the vessels of all embryonic organs and tissues examined, including heart, lung , head, limb bud, pancreas, spleen and stomach ( Xu et al., 2009a ) . When compared to the expression of VEGFR2 and PECAM , we found that Rasip1 generally marked identical vascular beds, with only slight variations in expressi on intensity . Expression of Rasip1 in later embryonic vessels, after their formation via either vasculogenesis or angiogenesis, implies that it has a mainte

54 nance functi on in mature vessels. Indee
nance functi on in mature vessels. Indeed, Rasip1 continued to be expressed in the endothelium of vessels into postnatal stages ( Xu et al., 2009a ) and was detected in adult organs, particularly in the highly vascularized lung ( Mitin et al., 2004 ) . 44 Figure 3.3 . Vascular expression of Rasip1 in embryonic organs and tissues. A, D columns) Whole mount β - galactosidase staining using Flk1 ( VEGFR2 ) - lacZ embryos. Whole mount in situ hybridization of Rasip1 ( B, E columns) and PECAM ( C, F columns). ( A - F ) Hearts. ( A’ - F’ ) Lungs. ( A” - C” ) Heads. ( D” - F” ) Limb buds. Note similarity of expressio n of Rasip1 in most vessels, as marked by VEGFR2 and PECAM expression (black arrowheads). Expression of all three vascular markers can be observed in the endocardium of the ventricle trabeculae in the heart (A - C) and of the coronary vasculature (arrows, D - F). Expression of all three markers is evident in the proxial ECs of the early lun uds (A’ - C’), althouh PECAM is not expressed in the ECs of the most distal tips of the buds at E10.5 (red arrowheads), while both VEGFR2 and Rasip1 are obse

55 rved in this p opulation. This heterogen
rved in this p opulation. This heterogeneity is also observed in the cephalic vessels at E10.5, where VEGFR2 is robustly expressed in the most mediolateral/distal vessels of the mesencephalon, while Rasip1 and PECAM are expressed at lower levels (red arrows, A” - C”). Rasi p1 is expressed in the vessels of the developing limb buds, including the interdigit vessels (white arrows, D” - F”). a, atria; , ronchus; r, ranchial arches; t, trachea; te, telencephalon; v, ventricle. The scale bars represent 100  m in A - F’ and 250  m in A” - F”. 45 Rasip1 expression is absent in vascularless embryos To definitively test whether Rasip1 is restricted to vascular endothelium, we assessed its expression in VEGFR2 mutant embryos that lack vascular endothelium ( Shalaby et al., 1995 ) . First, we compared Rasip1 expression ( Fig ure 3 . 4 B ) to that of Flk1(VEGFR2) - lacZ ( Fig ure 3 . 4 A ) in VEGFR2 +/ - heterozygous mice, which display no detectable abnormalities, and found both outlined the developing vasculature as expected. However VEGFR2 - / - homozygotes, which lack all blood vessels, exhibited no trace of Rasip1 expression by in

56 s itu hybridization ( Figure 3. 4 C ).
s itu hybridization ( Figure 3. 4 C ). Rasip1 and VEGFR2 are expressed in similar vascular domains in VEGFR2 heterozygotes, from E8.25 to E9.0, while in VEGFR2 null emb ryos we observed no Rasip1 expression in any embryonic region. These findings suggest that Rasip1 is expressed exclusively in VEGFR2 - dependent cell types. Given that VEGFR2 is primarily expressed in and required for ECs ( Shalaby et al., 1995 ; Yamaguchi et al., 1993 ) , it is likely that Rasip1 is also expressed exclusively in those cell types, but not in other non - vascular mesodermal or mesenchymal cell populations. We propose that Rasip1 is a novel, and during embryoge nesis, largely specific marker of embryonic blood vessels throughout development. 46 Figure 3. 4. Expression of Rasip1 is restricted to VEGFR2 - dependent endothelium. ( A, B, A’, B’ ) Comparison of Flk1 ( VEGFR2 ) - lacZ staining and Rasip1 expression. Note overall similarity of expression. Rasip1 expression closely resembles Flk1 ( VEGFR2 ) - lacZ expression at both E8.5 ( A, B ) and E9.0 ( A’, B’ ). ( C, C’ ) VEGFR2 - / - null embryos, lacking all endothelium. Embryos in

57 ( C - C” ) have been stained by in
( C - C” ) have been stained by in s itu hybridization for Rasip1 expression and allowed to develop same length of time as wildtype embryos in B - B” . Note complete lack of Rasip1 expression in these mutants. A” - C”) Sections throuh eryos in A’ - C’ showin presence of aortae and perineural vascular plexus in wildtype embryos, while these vascular structures are missing in VEGFR2 - / - embryos. al, allantois; g, gut tube; h, heart; hd, head; da or black arrowheads , dorsal aortae; n, neural tube; ys, yolk sac . The scale bars represent 200   (A’ - C’) and 50  m (A - C, A’’ - C’’). 47 Rasip1 is required for angiogenesis in cultured ECs To identify the potential role of Rasip1 in ECs, we used an in vitro siRNA approach to knock down endogenous Rasip1 expression in cultured mouse MS1 ECs ( Fig ure 3 . 5 A ). To examine whether reduction of Rasip1 levels might have a functional impact on MS1 cell behavior, we carried out both in vitro tube - formation and wound - healing assays. These assays allowed evaluation of the effect of Rasip1 knockdown on endothelial angiogenesis and cell migration, respectively. Striking

58 ly, si Rasip1 - treated MS1 cells, in wh
ly, si Rasip1 - treated MS1 cells, in which Rasip1 transcript levels wer e signifi cantly reduced , almost completely lost the ability to form plexus - like vascular structures when cultured on Matrigel ( Fig ure 3 . 5 B,D - F ). This observation supports the notion that Rasip1 is required for endothelial function. To evaluate the effect, we quanti fied the number of branch points created by the coalescence of ECs into cords/tubes, and found that these were reduced by over 85%. In addition, ablation of Rasip1 function in MS1 cells also dramatically decreased EC migration ability. Using an in vitro sc ratch assay, the „healin rate’ of a scratch „wound’, across a onolayer of ECs, was sinificantly reduced. While unmanipulated or siHPRT transfected cells were able to heal the wound following overnight incubation, si Rasip1 transfected cells migrated only about 50% the distance over that same timeframe ( Fig ure 3 . 5 D’ - F’,D” - F” ). This effect is not likely to be indirect, such as a result of decreased cell proliferation, 48 Figure 3.5. Rasip1 ablation in MS1 cells by transient siRNA transfection hinders endothelial tube formation

59 and migration ability. ( A ) Semi - qu
and migration ability. ( A ) Semi - quantitative RT - PCR (30 cycles) shows the knockdown of Rasip1 at the mRNA level. ( B, D - F ) si Rasip1 treated MS1 cells fail to for “tues” when plated on Matrigel. B) Quantification shows that formation of linear structures (tubes or cords) as measured by counting branching points in the vascular plexus, is decreased by approxiately 85%. C, D’ - F’, D” - F ”) Knockdown of Rasip1 inhibits endothelial cell migration. si Rasip1 - treated cells show slow healin rate in “scratch assay”. While untreated cells irate quickly into the cell free area (D’, D”), si Rasip1 - treated cells migrate less than half the distance durin the sae tie period (F’, F”). C) Quantification of endothelial iration in the scratch assay was determined by relative unit distance measured in images (as shown). 49 since the timeframe of the healing study is too short to allow significant prol iferation within the gap (wound healing carried out overnight). T hese results indicate that Rasip1 function is required in cultured ECs, for both angiogenic coalescence and migration, and is therefore likely to play impor

60 tant roles in blood vessel developm ent
tant roles in blood vessel developm ent. Rasip1 is required for embryonic blood vessel formation Bioinformatic comparison of Rasip1 sequences revealed that it was highly conserved across many different species, from human to lower vertebrates, including frog. The DILute domain of Rasip1, for instance, displayed almost 85% identity at the amino acid level between mouse and Xenopus trop icalis ( Figure 3. 6 ). This high level of similarity suggested Rasip1 might also be expressed in the vessels of other species, such as frog, and might play a conserved role in vessel formation. Thus, to assay Rasip1 function during embryonic vessel formation in vivo , we examined its expression and function in Xenopus tropicalis embryos. This tractable model system provided us with an avenue for in vivo assays. RT - PCR was used to amplify a fragment containing approxim ately 1900bp of the Rasip1 coding region from Xenopus tropicalis cDNA. Using this construct, we generated Dig - labeled probe for in situ hybridization and compared 50 Figure 3.6. Conservation of Rasip1 sequence across species. Conservation at the prot ein level in the DILute domain of RAS

61 IP1, between mouse, human and frogs. Con
IP1, between mouse, human and frogs. Consensus between mouse and frog protein sequences of DILute domain is 83.3%. Similar conservation is observed in other domains. 51 Rasip1 expression to known vascular markers ( Fig ure 3 . 7 A - H ). We note that erg ( Fig ure 3 . 7 A,B ), VE - Cadherin ( Fig ure 3 . 7 C ), and msr ( Fig ure 3 . 7 D ) expression patterns outline the early vasculature in Xenopus tropicalis , during both vasculogenesis and angiogenesis, as previously described in Xenopus laevis ( Baltzinger et al., 1999 ; Devic et al., 1996 ) . Similarly, we found that Xenopus Rasip1 is expressed in the developing vasculature throughout development, including early expression in emerging angioblasts (stage 25) and during vessel coalescence ( Fig ure 3 . 7 E - G ). In addition, Rasip1 is expressed strongly during angiogenesis, in sprouting ISV s ( Fig ure 3 . 7 G ), and remains expressed after initiation of heart beat (st.34) and blood circulation ( Fig ure 3 . 7 H ). To determine if Rasip1 function is required for vessel development in vivo , we targeted the sequence in Xenopus tropicalis embryos with antis ense Rasip1 morpholino oligos ( Rasip1 - MO

62 ) designed to inhibit splicing of Rasip
) designed to inhibit splicing of Rasip1 transcripts ( Fig ure 3 . 8 A,B ). RT - PCR analysis with primers spanning the exon - intron boundary confirmed that Rasip1 splicing, and hence expression of mature transcripts, was eff ectively abolished in embryos radially injected with 30ng of Rasip1 - MO ( Fig ure 3 . 8 C,D ). To specifically examine the effect of Rasip1 knockdown on embryonic blood vessel development, we injected the Rasip1 - MO (16 ng), into one side of the embryos at the 2 - cell stage and then assayed them by in situ hybridization at stage 32 with the endothelial marker msr ( Fig ure 3 . 8 E - I ). 52 Figure 3.7. Expression of Xenopus tropicalis Rasip1 transcripts in frog embryos by in situ hybridization marks the developing embryonic blood vessels (in all panels anterior is to the left). ( A - D ) Vascular markers reveal the embryonic vasculature at stages indicated, including angioblasts (white arrowheads in A, B) and developing blood vessels, such as the posterior cardinal vein (black arrow, B - D). ( A ) vegfr2 ; ( B ) erg , vascular ETS factor; ( C ) ve - cad, vascular endothelial cadherin; ( D ) msr , vascular G - protein coupled

63 receptor. ( E - H ) Rasip1 initially
receptor. ( E - H ) Rasip1 initially marks scattered angioblasts ( E , st.25), but progressively marks aggregating flank vessels ( F , st.28). ( G ) As vessels form, Rasip1 marks all embryonic frog vessels examined, including the flank plexus (red arrow), endocardium (red arrowhead), cardinal veins (black arrow) and ISVs (black arrowheads). ( H ) Expression of Rasip1 declines slightly in vessels as they mature (including data not shown). The scale bars represent 250  m. 53 We found that blood vessel development was severely inhibited in Rasip1 - MO injected embryos as detected by msr staining . Most strikingly, we found that the posterior cardinal vein failed to form on the injected side in 80% of injections. In addition, we noted a significant reduction in both the number of ECs, as marked by msr , along the flank of the embryo, and the organiz ation of these cells into the vitelline plexus (reduction of the branching points from 23 to 4) ( Fig ure 3 . 8 F , J ). The uninjected side, in contrast, remained unaffected and displayed normal vascular structures, including a normal posterior cardinal vein ( Fig ure 3 . 8 E , G ). We also observed that sprouting ISVs

64 failed to appear, which was not surpri
failed to appear, which was not surprising as they originate from the posterior cardinal vein ( Fig ure 3 . 8 H , J ). Sectioning of injected embryos revealed the distinct absence of the posterior cardinal vein and ISVs on the injected side ( Fig ure 3 . 8 I ). Quantification of these observations showed that the cardinal vein and ISVs were lost over 90% of the time, while the plexus EC branching was reduced by over 80%. These experiments provide evidence that Rasip1 is r equired in vivo for proper vessel development. 54 Figure 3.8 . Rasip1 knockdown in frog embryos results in failure of blood vessel formation. ( A, B ) Schematic of morpholino targeting the splice junction of Xenopus tropicalis Rasip1 sequence. ( C ) Schematic of microinjection of Rasip1 morpholino injections into blastomeres of early cleavage stage embryo. ( D ) Semi - quantitative RT - PCR shows the knockdown of Rasip1 transcript expression in radially injected embryos (24 cycles) . ( E - H ) msr in situ hybridization of injected embryo : uninjected ( E , G ) or injected ( F , H ) sides. ( I ) Transverse sections through injected embryos show that cardinal veins and ISVs are lost o

65 n the injected side . ( J ) Quantif ic
n the injected side . ( J ) Quantif ication of the t otal number of ISVs and flank plexus branch points from the injected versus uninjected sides of MO - injected embryos. Y - axis for ISVs is to left; Y - axis for flank branch points is to right. The scale bars represent 100  m ( E , F ), and 50  m ( I - H ), respectively. 55 Discussion In this chapter, w e show that Rasip1 is specifically expressed in the endothelium of the developing blood vessels of both mouse and frog embryos. Additionally, we demonstrate that this expression initiates early, in angioblasts prior to their aggregation into vessels, and c ontinues in established vessels, which grow and remodel via angiogenesis. We also demonstrate that Rasip1 is fundamentally required in ECs, both in vitro and in vivo . Knocking down Rasip1 in cultured ECs inhibits their migration and coalescence into vessel - like structures, while knocking down Rasip1 in amphibian embryos results in failure of vessel development. These findings establish Rasip1 as a novel and robust marker of embryonic ECs, throughout their specification and differentiation, and as a likely i mportant regulator of vascular development.

66 (1) Rasip1 is a novel endothelial
(1) Rasip1 is a novel endothelial marker Few vascular genes have proven useful as specific markers of the endothelium. VEGFR2, PECAM, Tie2 and VE - Cadherin are the most frequently used markers available to date, in t hat they are highly enriched in ECs and are often used as specific markers. However, they are not completely endothelial specific, as they are often transiently expressed in other tissues, at some point 56 during development. For instance, VEGFR2 is also dete cted in hematopoietic cells ( Yamaguchi et al., 1993 ) , PECAM is also in macrophages ( Lee, 1991 ) , Tie2 is found in mesenchymal cells of heart outflow trac ts ( Kisanuki et al., 2001 ) , VE - Cadherin is also expressed in liver hematopoietic stem cells ( Kim et al., 2005 ) and both VEGFR2 and PECAM are also found in lymphatic vessels ( Enholm et al., 2001 ) . In addition, most other commonly used endothelial markers such as Dll4, Egl7, ephrin - B2, EphB4, Jagged1, Notch1 and many more, are widely ex pressed in a number of other organs and tissues ( Conway et al. , 2001 ; Eichmann et al., 2005 ; Torres - Vazquez et al., 2003 ) . Our study adds Rasip1 to the short list of useful endothelial - enriche

67 d sequences, which can be used to study
d sequences, which can be used to study the developing cardiovascular system. Lik e VEGFR2 , Rasip1 transcript levels are highly enriched in embryonic ECs. This is in contrast to PECAM and Tie2 whose transcript levels are lower, and thus difficult to visualize by in situ hybridization. While we have assayed commercially available antibod ies to RASIP1 protein, both in vitro and in vivo , none have proven useful for either immunofluoresence or immunohistochemistry. Until effective antibody reagents become available, assays for Rasip1 transcripts will be useful for studies ranging from examin ation of early angioblast specification to vasculogenesis and angiogenic vessel formation. Given its 57 expression conservation across species, in both frog and mouse, it will very likely be more widely applicable to vascular studies in other species as well. (2) Rasip1 is essential for endothelial cell function Knockdown of Rasip1 levels in cultured ECs reveals a critical role for Rasip1 in basic cellular functions, such as cell motility and angiogenesis. When Rasip1 function is reduced, both these basic endothelial behaviors are severely abrogated. si Rasip1 - treated cell

68 s, but not si HPRT - treated cells, dis
s, but not si HPRT - treated cells, display a reduction in their propensity to aggregate and form cords or vessels, in a matrigel angiogenesis assay. In addition, whereas untreated or control siRNA - treated ECs will normally migrate actively across tissue culture - treated plastic in vitro , such as in a „wound healin’ assay, cells lackin Rasip1 function fail to irate. Given that these basic cellular behaviors are likely to comprise the foundations of vessel formation, we predicted that endothelial cells in emerging bloo d vessels would also require Rasip1 function. Indeed, reduction of Rasip1 in vivo leads to a dramatic failure of embryonic vessel formation. Using a MO - based approach, we knocked down endogenous expression of Rasip1 in Xenopus tropicalis embryos, which led to a clear failure of the posterior cardinal vein and associated ISVs to form. These results demonstrate that Rasip1 is required for the 58 proper formation of vascular structures that develop via both vasculogenesis (posterior cardinal vein) and angiogenesi s (ISVs). Interestingly, vessels of the flanking vitelline plexus, within the lateral plate mesoderm, while disorganized when

69 Rasip1 function is reduced, were not co
Rasip1 function is reduced, were not completely abrogated. The basis for the difference in the response of these different vascula r beds to the absence of Rasip1 is unclear. However, it has been proposed that inherently different populations of hematopoietic, and perhaps endothelial, cells arise within these different embryonic regions ( Kau and Turpen, 1983 ; Maeno et al., 1985 ) . It is possile that they ay represent „priitive’ (ventral/flank) versus „definitive’ (dorsal lateral plate/somite) ECs, and that these two populations have a differential requirement for Rasip1 function. (3) Rasip1 is not required for angioblast specifica tion Significantly, despite disruption of vessels when Rasip1 function is knocked down in frog embryos, angioblasts still emerge within the mesoderm. This finding indicates that Rasip1 plays a role sometime after initial angioblast specification. This obse rvation is supported by the timing of Rasip1 expression initiation during vessel development in frogs, as angioblast specification is conveniently separated in time from the process of vessel formation via vasculogenesis. In frogs, angioblasts are specifie d within the mesoderm of l

70 ate 59 neurula stage embryos (s
ate 59 neurula stage embryos (st.18 - 22), many hours prior to vessel formation that occurs at the late tailbud stage (st.30 - 32). We find that Rasip1 expression in frogs initiates later (st.22) than VEGFR2 (st.18) ( Cleaver et al., 1997 ) , thus displaying a marked delay and appearing distinctly later than the earliest known angioblast markers. In addition, when Rasip1 function is knocked down using MOs, we observe that early angioblasts em erge relatively normally as assayed by flk1 expression at st.20/21. We therefore suggest that Rasip1 is likely to function in angioblasts and ECs following their initial specification, possibly during their migration, cord formation or tubulogenesis. 60 CHAPTER IV Rasip1 directs vascular tubulogenesis Cardiovascular function depends on patent blood vessel formation by endothelial cells (ECs). However the mechanisms underlying vascular „tuuloenesis’ are only einnin to e unraveled. Followin our pre vious studies described in Chapter III, we show that endothelial tubulogenesis requires the Rasip1 and its binding partner the RhoGAP Arhgap29. Mice lacking Rasip1 fail to form patent lumens in all blood

71 vessels, including the early endocardial
vessels, including the early endocardial tube. Rasip l null angioblasts fail to properly localize the polarity determinant Par3 and display defective cell polarity, resulting in mislocalized junctional complexes and loss of adhesion to extracellular matrix (ECM). Similarly, d epletion of either Rasip1 or Arhg ap29 in cultured ECs blocks in vitro lumen formation, fundamentally alters the cytoskeleton and reduces integrin - dependent adhesion to ECM. These defects result from increased RhoA/ROCK/myosin II activity and blockade of Cdc42 and Rac1 signaling. This stud y identifies Rasip1 as a unique, endothelial - specific regulator of Rho GTPase signaling, which is essential for blood vessel morphogenesis. 60 61 Figure 4.1. Rasip1 gene targeting scheme. ( A ) Diagram showing the vector design for generation of the Rasip1 null allele. Exon 3 is deleted, resulting in a truncated, non - functional protein. ( B - C ) Confirmation of the deletion of Rasip1 by Southern blot ( B ) and RT - PCR ( C ). S, SphI cutting site; TK, HSV thymidine kinase; P1 - P3, genotyping primers. 62 Rasip1 is essential for cardiovascular development In the p

72 revious study, we showed that Rasip1
revious study, we showed that Rasip1 was expressed exclusively in ECs of murine and amphibian embryos throughout embryogenesis ( Xu et al., 2009a ) . To examine whether Rasip1 might regulate vasculogenesis in higher vertebrates, we generated mice lacking Rasip1 function ( Figure 4. 1 ). Heterozygous Rasip1 mice were phenotypically normal and viable, while the null mutation was embryonic lethal. Homozygous null embryos appeared grossly normal at E8.25, but were dead by E10.5 ( Xu et al., 2011 ) . At E9.5, Rasip1 - / - mutant embryos dis played growth retardation and wid espread edema and their yolk sac appeared smooth, lacking the hierarchical vascular network evident in heterozygous controls ( Figure 4.2 A - D ). No significant differences were found in placental tissues (data not shown). Blood vessel tubuloge nesis requires Rasip1 To understand the origins of the observed cardiovascular failure in Rasip1 - / - mutants, we sought to characterize developing blood vessels. A Flk1 - LacZ allele was bred into the Rasip1 null line ( Shalaby et al., 1995 ) . Initial angioblast numbers and distribution were normal ( Xu et al., 2011 ) , indicating that Rasip1 is not r

73 equired for angioblast specification, pr
equired for angioblast specification, proliferation, or patterning. However by 63 Figure 4.2 . Rasip1 is essential for mammalian vascular morphogenesis . ( A - D ) E9.5 littermate mouse embryos showing defects in Rasip1 - / - embryos ( B ) and yolk sacs ( D ). ( E - H ) Whole mount Flk1 - lacZ beta - galactosidase staining of Rasip1 +/ - ( E , G ) and Rasip1 - / - ( F , H ) yolk sacs ( E , F ) and embryonic heads ( G , H ), showing narrow Rasip1 - / - blood vessels that fail to remodel. ( I - L ) Connexin40 (Cx40 or Gja5) ( I , J ) or EphB4 - lacZ ( K , L ) in littermate E9.5 Rasip1 +/ - and Rasip1 - / - yolk sacs ( I , J ) or embryos ( K , L ) showing failed arterial ( I , J ) and venous ( K , L ) differentiation in Rasip1 - / - vessels. Scale bars: 500  m ( A - D ), 10 0  m ( E, F ), 200  m ( G - L ) . 64 E9.5, mutant vessels failed to remodel from an initial plexus into their typical hierarchical array of large and small vessel s, both in the yolk sac and embryonic tissues ( Figure 4. 2E - H ). In addition, arteriovenous differentiation failed in Rasip1 - / - vessels, as assessed by the arterial marker Connexin40 (Gj

74 a5) and EphB4 - LacZ ( Wang et al., 19
a5) and EphB4 - LacZ ( Wang et al., 1998 ) ( Figure 4. 2I - L ). Together, these results showed that development of embryonic blood vessels was severely impaired in the absence of Rasip1. Remodeling and arteriovenous differentiation defects are hallmark effects of hemodynamic failure, as both processes depend on blo od flow ( le Noble et al., 2004 ) . Therefore, to identify possible vessel occlusions in Rasip1 mutants that would hinder blood circu lation, we examined the first major embryonic vessels, the paired dorsal aortae. In both Rasip1 +/ - and Rasip1 - / - embryos, E8.0 - 8.5 pre - aortal angioblasts aligned normally in two parallel rows, however they appeared distinctly narrower in mutant embryos , often displaying visible occlusions ( Figure 4. 3 A,B ). Indeed, sections revealed that patent, continuous lumens were globally absent in all blood vessels, including those of the dorsal aortae ( Figure 4. 3 C - F ), endocardium ( Figure 4. 3 G , H ), yolk sac ( Figure 4 . 3 I - L ), vitelline artery, inflow tracts and allantois (data not shown). Absence of vascular tubes was evident at all stages examined, from the onset of cord formation at the 1somite (1S) stage, and

75 throughout the entire time frame of norm
throughout the entire time frame of normal tubulogenesis ( 2S to 65 Figure 4.3 . Rasip1 is essential for vascular tubulogenesis in all blood vessels. ( A - L ) Flk1 - LacZ beta - galactosidase staining of littermate E8.5 (6 somite stage) Rasip1 +/ - and Rasip1 - / - embryos and tissues, in whole mount or section. ( A , B ) Ventral views of E8.5 embryonic paired dorsal aortae. Anterior is up. ( C - F ) Transverse sections of E8.5 embryos, through the trunk region, posterior to the heart. ( E , F ) Closeup views, showing open lumens in Rasip1 +/ - ;Flk1 - lacZ embryos ( C , E ), but lumenles s cords in Rasip1 - / - ;Flk1 - lacZ embryos ( D , F ). Sections through hearts of E8.5 ( G , H ), showing absence of a lumen in the endocardium of Rasip1 - / - ;Flk1 - lacZ embryos ( H ). Yolk sacs in whole mount ( I , J ) or section views ( K , L ). Vascular tubes are absent in all Rasip1 - / - vessels examined. Arrows, dorsal aortae. Arrowheads, yolk sac vessels. ys: yolk sac. Scale bars: 1 00 µ m ( A, B, I, J ), 50 µ m ( C - H , K , L ). 66 6S) ( Xu et al., 2011 ) . We examined developmental intermediates at each somite stage (from 1S to 12S, a

76 t E8.0 - 8.5), as well as at E9.0, E9.5
t E8.0 - 8.5), as well as at E9.0, E9.5 and E10.5. Defects did not result from failed proliferation, as similar numbers of ECs were present in Rasip1 +/ - versus Rasip1 - / - aortae and we found no proliferation defects ( Xu et al., 2011 ) . Given that failure of tubulog enesis occurred at the onset of vessel formation, prior to the onset of blood flow, and that EC proliferation was not affected, we reasoned that loss of Rasip1 resulted in morphogenetic failure due to cellular defects intrinsic to mutant angioblasts. Rasi p1 binding partners While Rasip1 had previously been shown to bind Ras GTPases by in vitro pull down assays ( Mitin et al., 2004 ) , its physiological binding partners in endothelium and its downstream signaling pathways have remained unknown. To elucidate the molecular mechanism of Rasip1 function during vasculogenesis and lumen formation, we sought to determine its binding partners by two - step affinity purification and mass spectrometry, using a tetracycline inducible stable endotheli al cell line (MS1 Rasip1 - FLAG ) ( Xu et al., 2011 ) . Amongst several putative Rasip1 bindi ng partners identified ( Figure 4.4A ) were two proteins of particular interes

77 t due to their previous implication in v
t due to their previous implication in vascular development: non - muscle myosin heavy chain IIA (NMHCIIA) an d a RhoA - specific GAP, Arhgap29 . 67 NMHCIIA is an important motor protein in the actomyosin complex. NMHCIIA and MLC (myosin light chain) together comprise myosin II, which modulates diverse processes such as cell structure, adhesion, contractility and motility ( Conti and Adelstein, 2008 ; Even - Ram and Yamada, 2007 ) . NMHCIIA often exerts its regulatory control by acting as a scaffold for signaling molecules, such as kinases and Rho GTPase guanine nucleotide exchange factor s (GEFs) ( Even - Ram et al., 2007 ) . Recently, it was reported that i n VEGF haploinsufficient embryos, NMHCIIA failed to be recruited to the luminal membrane of dorsal aortic ECs, which failed to form vascular lumens ( Strilic et al., 2009 ) . NMHCIIA was proposed to regulate actomyosin tension of the cell membrane, driving lumen expansion. In Rasip1 - / - failed vessels and Rasip1 siRNA depleted cells ( Xu et al., 2011 ) , levels of NMHCIIA at the EC - EC interface appear unchanged, suggesti ng that recruitment of NMHCIIA to the cell membrane was Rasip1 - independent. We therefore reasoned

78 that NMHCIIA activity and function, rat
that NMHCIIA activity and function, rather than levels, were likely altered in Rasip1 - / - mutants. Arhgap29 (PARG1), the other identified Rasip1 binding partne r was known to be expressed in fish vasculature ( Gomez et al., 2009 ) and to possess a GAP domain for RhoA ( Myagmar et al., 2005 ) . In mouse embryos, we found that Arh gap29 protein was present in the ECs of the paired aortae during vasculogenesis, in a man ner analogous to that reported in fish ( Figure 4.4 B ). In 6 8 Figure 4.4. Rasip1 and Arhgap29 are required for in vitro EC lumen formation. ( A ) Silver staining of the affinity - purified Rasip1 complex. Marked bands were identified by mass spectrometry. ( B ) Arhgap29 protein in E8.5 mouse embryonic aortic ECs, ventral view. Arrows, C ) Rasip1 and Arhgap29 expression in cultured EC (MS1). Arrowheads, Rasip1/Arhgap29 co - localization in punctae. Nuclei, DAPI (blue). ( E ) Lu men formation is blocked in siRasip1 - /siArhgap29 - treated HUVECs cultured in 3D collagen matrices. Arrowheads, visible EC lumen. ( F ) Quantification of lumen formation in control - or siRasip1 - /siAhgap29 - treated HUVECs. Error bars represent standard deviatio n

79 . 69 addition, Rasip1 and Arhg
. 69 addition, Rasip1 and Arhgap29 were present together in cytoplasmic punctae in cultured ECs (~50% rate of overlap) ( Figure 4.4 C ). Rasip1 and Arhgap29 are required for in vitro lumen formation Overlap of Rasip1 and Arhgap29 in ECs, both in vivo and in vitro, suggested that they were likely to function together during blood vessel development. To assess the requirement for Rasip1 and Arhgap29 during in vitro tubulogenesis, we used a HUVEC 3D matrix assay that mimics the developmental process of vasculoge nesis ( Kamei et al., 2006 ; Sacharidou et al., 2010 ) . To determine whether Rasip1 or Arhgap29 function was required for EC lumen formation in vitro , siRNA - de pleted cells were assessed for their lumen formation ability . Strikingly, EC lumen formation was completely blocked in siRNA - treated cells. Movies of depleted cells showed that although ECs extended cellular processes, these would snap back and rarely maintain connections to the surrounding matri x ( Xu et al., 2011 ) . In addition, cells would dynamically make and break contacts with each other, and preliminary lumenal structures within i ndividual ECs initiated but quickly collapsed. In contr

80 ast, the control ECs would generate and
ast, the control ECs would generate and sustain lumens, maintain steady adherence to the surrounding matrix, and interconnect to form multicellular lumenal structures ( Figure 4. 4D , E ). 70 Rasip1 and Arhgap 29 are required for activation of Cdc42 and Rac1 during lumen formation Previous in vitro studies have shown that vascular lumen formation depends on proper regulation of the Rho family of monomeric G proteins (Cdc42/Rac1/RhoA). Specifically, Cdc42 and Rac1 activation were shown to be required for lumen formation in 3D collagen matrix assays, while RhoA activation had the opposite effect and suppressed it ( Bayless and Davis, 2002 , 2004 ; Iruela - Arispe and Davis, 2009 ; Nobes and Hall, 1995 ; Sacharidou et al., 2010 ) . We thus examined Rasip1 and Arhgap29 regulation of Rho GTPase activity using this same assay system and found markedly decreased levels of activated Cdc42 and Rac1 levels in siRNA - treated HUVECs ( Figure 4.5A ). In addition, we examined a kinase cascade downstream of Cdc42 that was recently shown to be required for in vitro EC lumen formation ( Koh et al., 2008 ; Koh et al., 2009 ) . As expected, signaling downstream of Cdc42 and Rac1 wer

81 e impaired in the absence of Rasip1 or
e impaired in the absence of Rasip1 or Arhgap29, as the active (phosphorylated) form of key GTPase effector kinases, such as pPak4, pSrc, pB - Raf, pC - Raf and pErk, were reduced in siRNA - depleted ECs ( Figure 4.5B ). Interestingly, although cell morphology and behavior were similarly affe cted by absence of either Rasip1 or 71 Figure 4.5. Rasip1 and Arhgap29 are required for regulation of small GTPase signaling . ( A ) C dc 42, R ac 1 and R ho A activities and ( B ) reduction of kinase signaling downstream of Cdc42/Rac1 pathway in siRNA - treated HUVECs in 3D cultures at 24hrs. 72 Arhgap29, some differences in downstream signaling reduction were reproducible (pSrc). Rasip1 and Arhgap29 regulate EC arc hitecture via repression of RhoA In contrast to the reduced activities of Cdc42 and Rac1, we found that RhoA activation was upregulated in the absence of either Rasip1 or Arhgap29 ( Figure 4.5A ). This finding was consistent with the notion that RhoA antagonizes lumen formation and must be kept at bay ( Bayless and Davis, 2002 ) . Because activated RhoA was shown to suppress vascular lumen maintenance via cytoskeletal modulation (

82 Bayless and Davis, 2004 ) , we examined
Bayless and Davis, 2004 ) , we examined the EC cytoskeleton in the absence of either Rasip1 or Arhgap29. Strikingly, individua l knockdowns of Rasip1 or Arhgap29 lead to nearly identical defects in EC architecture ( Figure 4.6A - C ). At higher resolution, we observed a significant increase in actin stress fiber assembly (a process dependent on RhoA) in the absence of either Rasip1 or Arhgap29 ( Figure 4.6 D - 3F ). Moreover, we assessed „stailized’ icrotuules and found a sharp decrease in acetylated tuulin in Rasip1 or Arhgap29 depleted cells ( Figure 4.6 G - 3J ). This effect was also surprisingly detectable in vivo in E9.5 Rasip1 - / - whole embryo lysates ( Figure 4.6 K ). Rasip1 and Arhgap29 therefore are both independently required for cellular structure and contractility of ECs. 73 Figure 4.6. Rasip1 and Arhgap29 regulate EC architecture and tubuloge nesis . ( A - F ) Immunofluorescent phall oidin staining showing gross cell morphology and stress fibers in siRNA - treated HUVECs. ( G - I ) „Stailized’ icrotuule shown y acetylated - tubulin staining in siRNA - treated HUVECs. Insets, high magnification view of microtubules. Nucl

83 ei, DAPI (blue). ( J, K ) Acetylated
ei, DAPI (blue). ( J, K ) Acetylated α - tubulin level is down regulated in siRNA treated ECs in vitro ( J ) and in vivo in E9.5 Rasip1 - / - embryos ( K ), n = 2. Error bars represent standar d deviation. 74 We next examined possible Rasip1 modulation of myosin II. We reasoned that myosin II activity was likely to be altered, due to four findings: 1. Rasip1 binds both a RhoA - specific GAP (Arhgap29) and NMHCIIA, which is RhoA - dependent ( Figure 4.4 A ); 2. activated RhoA levels were increased in Rasip1/Arhgap29 - depleted cells ( Figure 4.5A ); 3. stress fiber assembly (known to be RhoA dependent) was significantly increased in Rasip1/Arhgap29 - depleted cells ( Figure 4.6D - F ); and 4. Rasip1/Arhgap29 - depleted cells display dramatically decreased acetylated tubulin ( Figure 4.6G - J ) and NMHCIIA is known t o suppress tubulin acetylation ( Even - Ram et al., 2007 ) . We thus examined Rasip1 and Arhgap29 modulation of RhoA - dependant myosin II via the regulatory molecules Rho - kinase (ROCK) and the ROCK substrate Myosin Light Chain (MLC) ( Matsumura, 2005 ; Rikitake and Liao, 2005 ) . Phosph orylated MLC (pMLC) is required for non - muscle myosin II activ

84 ity and regulation of the actomyosin co
ity and regulation of the actomyosin complex. Indeed, we found significantly increased levels of pMYPT (T696) (downstream target of ROCK1/2) and pMLC (T18/S19) in the absence of either Rasip1 or Arhgap29 ( Figure 4.7A ). These data further support the hypothesis that myosin IIA activity, not localization, depends on Rasip1 and Arhgap29. In addition, hyperactivation of ROCK/MLC/myosin IIA, and the manifested cytoskeletal alterations, suggest that an important function of Rasip1/Arhgap29 is 75 Figure 4.7. Rasip1 and Arhgap29 regulate EC architecture and tubulogenesis through Rho family GTPases. ( A ) Western blots showing RhoA and ROCK activity (pMYPT, T696), phosphorylation of myosin light chain (pMLC, T18/S19) in siRNA - treated HUVECs, n = 3. ( B ) Dominant negative (DN) RhoA rescues in vitro lumenless phenotype in 3D collagen matrices in siRasip1 - /siArhgap29 - treated HUVECs. ( C ) Loss of Rasip1, which causes a lumenless EC phenotype in 3D collagen ma trices, is rescued by siRNA knockdown of RhoA. Error bars represent standar d deviation. 76 to suppress ROCK, MLC and thus non - muscle myosin IIA activity via RhoA, during endothelial morphogenesis. T

85 o confirm whether Rasip1 suppression of
o confirm whether Rasip1 suppression of RhoA signaling played a role during EC lumen formation, we knocked down expression of RhoA in the background of Rasip1 or Arhgap29 loss - of - function. While siRasip1 and siArhgap29 treated cells were blocked in their ability to form lumens in vitro (as shown above), this failure was reversed upon co - expression of dominant negative RhoA ( Figure 4.7B ). By contrast, dominant negative Cdc42 or Rac1 constructs ( Bayless and Davis, 2002 ) had no effect. These results were further confirmed with co - siRNA treatments to simultaneously knockdown expression of RhoA and either Rasip1 or Arhgap29 ( Figure 4.7C ). Thus while s iRhoA alone had no effect on in vitro lumen formation, it largely rescued the absence of either Rasip1 or Arhgap29. Failure of EC - ECM adhesion in vitro in the absence of Rasip1 or Arhgap29 As RhoA and NMHCIIA are both known to regulate cell adhesion, we examined both EC - EC and EC - ECM adhesion in Rasip1/Arhgap29 - depleted cells. Flow cytometry indicated neither PECAM nor VE - cadherin levels were altered in the absence of Rasip1 or Arhgap29 ( Xu et al., 2011 ) . In addition, despite irregular 77 borders, rel

86 atively normal levels of the adhesion mo
atively normal levels of the adhesion molecules ZO - 1, β - catenin and PECAM were found at cell - cell interfaces of Rasip1 or Arhgap29 depleted cells ( Xu et al., 2011 ) , suggestin g at least grossly that EC - EC adhesion was not reduced. By contrast, we found that siRNA - treated ECs exhibited significantly reduced adhesion to several t ypes of ECM ( Xu et al., 2011 ) . To assess the molecular machinery that regulates cell adhesion to ECM, we examined focal adhesions, which are structures essential for adhesion of cells to m atrix during both embryonic development and blood vessel formation ( Bohnsack and Hirschi, 2003 ; Goody and Henry, 2010 ) . Focal adhesions first appear as adhesion complexes (FXs), which mature into focal adhesions (FAs) and then into fibrillar adhesions (FBs) ( Tomar and Schlaepfer, 2009 ) . An excellent indicator of immature FX is phosphorylated paxillin (pPaxillin), which is recruited to nascent FXs, but lost as th ey mature ( Zaidel - Bar et al., 2007 ) . We found that pPaxillin (Y118) positive FXs were expanded in siRasip1 or siArhgap29 depleted cells compared to control siRNA - treated cells ( Figure 4 .8A - C ), especially at the cell periphery ( Figu

87 re 4 .8 D - 4F ). In addition, total lev
re 4 .8 D - 4F ). In addition, total levels of pPaxillin (Y118) were significantly increased in the absence of Rasip1 or Arhgap29, both in vitro and in vivo ( Figure 4 .8 G ), indicating a likely increase in immature FXs. Of n ote, it has been reported that in plated cells, FXs are born at the cell periphery and mature 78 Figure 4.8. Rasip1 and Arhgap29 are required for maturation of endothelial - ECM adhesion. ( A - F ) Immature focal adhesions shown by phosphorylated Paxilli n (Y118) staining in siRNA - treated HUVECs. ( G ) Phosphorylated Paxillin (Y118) is up regulated in siRNA treated ECs in vitro and in vivo in Rasip1 - / - embryos, n = 2. ( H ) Flow cytometry showing decrease in activated β 1 integrin (9EG7) in the absence of Rasip1 or Arhgap29. MnCl 2 (5mM) treatment reveals total surface levels of integrins are unchanged. ( I - K ) Mature fibrillar adhesions shown by activated β 1 integrin (9EG7) expression in siRNA - treated HUVECs. Insets, high magnification view of 9EG7 + FBs. Nucl ei, DAPI (blue). Error bars represent standard deviation. 79 into FBs at the center of cell ( Tomar and Schlaepfer, 2009 ) , suggesting that in

88 the absence of either Rasip1 or Arhga
the absence of either Rasip1 or Arhgap29, FX are favored over FAs/FBs and maturation of adhesions is inhibited. We then assessed mature focal adhesions (FBs), which contain clustered or „activated’ interins, ut lack paxillin ( Morgan et al., 2009 ) . Given the central role of β1 interin in adhesion of ECs to ECM, as well as in blood vessel formation ( Drake et al., 1992 ; Zovein et al., 2010 ) , we examined activated β 1 usin an „ activated’ state - specific antibody, 9EG7 ( Bazzoni et al., 1995 ) . In the absence of either Rasip1 or Arhgap29, activated β 1 integrin levels were reduced 70% and 80% respectively ( Figure 4 .8 H , black columns). To reveal total cell β 1 integrin levels, we artificially activated β 1 using manganese ( Lenter et al., 1993 ) and found these unchanged ( Figure 4 .8 H , white columns). These results suggest that Rasip1 is required for activation, rather than levels, of β 1 integrin. We further examined the reduction of activated β 1 integrin in Rasip1/Arhgap29 depleted ECs using immunofluorescent staining with 9EG7. Control HUVECs displayed FBs as linear tracts of clustered β 1 integrin in the central region of the cell

89 ( Figure 4 .8 I ). B y contrast, siRasi
( Figure 4 .8 I ). B y contrast, siRasip1 - or siArhgap29 - treated cells lacked all FB organization into tracts, showing only reduced and punctate β 1 foci ( Figure 4 .8J, K ). Thus, activated β 1 levels and distribution were 80 severely affected in the absence of Rasi p1 or Arhgap29, althouh total β 1 remained unchanged ( Xu et al., 2011 ) . We also found that activated α vβ 3 integrins were affected in a similar manner, suggest ing that Rasip1 and Arhgap29 work together to regulate EC adhesion to the surrounding ECM, likely via multiple integrins, and in the absence of either molecule the dynamic maturation of adhesion conta cts is compromised, reducing EC - ECM adhesion. Rasip1 is required for proper in vivo endothelial - ECM adhesion To validate our in vitro findings that Rasip1 regulates EC adhesion, we examined the interface between adjacent embryonic ECs (EC - EC), as well as between ECs and surrounding ECM (EC - ECM), in Rasip1 +/ - and Rasip1 - / - embryos. Given that tight junctions (TJs) and adherens junctions (AJs) components play critical roles in epithelial lumen formation ( Lampugnani et al., 2010 ) , we first examined in vivo EC - EC

90 junction s. While claudin - 5 and ZO -
junction s. While claudin - 5 and ZO - 1 (TJs) highlighted narrow peripheral junctions between wild type aortal ECs at E8.5 ( Figure 4.9 A, C ), TJs were found in clusters at the center of Rasip1 - / - cords ( Figure 4.9 B, D ). Similarly, VE - cadherin (AJs) was normally only at periphe ral junctions ( Figure 4.9E ), however it was ectopically distributed in Rasip1 - / - cords ( Figure 4.9 F ). E8.5 mutant aortic ECs also remained distinctly cuboidal, similar to β 1 - depleted ECs ( Zovein et al., 2010 ) ( Figure 4.9C - F , and data not shown). 81 Figure 4.9. Rasip1 - / - angioblasts remain cuboidal and fail to adhere to surrounding ECM. ( A - N ) Confocal immunofluorescence microscopy of 3 - 6 somite Rasip1 +/ - ( A, C, E, G, I, K, M ) and Rasip1 - / - ( B, D, F, H, J, L, N ) dorsal aortae (Flk1 - EGFP, A, B, G, H, I - N ) in transverse sections: ( A, B ) claudin - 5, ( C, D ) ZO - 1 and laminin, ( E, F ) VE - cadherin and collagen IV, and ( G, H ) fibronectin. Nuclei, DAPI (blue) . ( O ) Aver age dorsal aortae diameter at 6 - 8 somite stage, n=30. Arrows, EC - EC junctions in Rasip1 +/ - embryos. Arrowheads, mislocalized EC - EC junctions. Scal

91 e bars, 25 µ m. Error bars represent s
e bars, 25 µ m. Error bars represent standard deviation. 82 We found the opposite situation when we examined adhe sion of ECs to the surrounding matrix. Aortic cords normally undergo tubulogenesis in immediate contact with both collagen IV and fibronectin (FN), which are secreted by the surrounding trunk mesenchyme, as well as laminin secreted by the underlying endode rm. While Rasip1 - / - aortic ECs remained tightly associated with the endoderm, mutant ECs lost adherence to the surrounding mesoderm ( Figure 4.9D, F, H, L and N ). This selective failure of EC - ECM adhesion was observed at all stages after 3S during vasculogenesis. At 3S, both wildtype and mutant cord ECs were in contact with mesenchymal cells in a nearly identical manner ( Figure 4.9I, J ). However, between 4 to 6S, an intervening space progressively expanded ( Figure 4.9K - N ). Indeed, while ECs remained in tight contact with the endoderm, a highly reproducible and growing cavity appeared between the mesenchyme and the Rasip1 - / - aortic cord ECs. Because of the failure of the aortae to acquire patency, the diameter of these cords was reduced compared to wi ld type, with most of them under 5

92 µ m ( Figure 4.9 O ). Together, these d
µ m ( Figure 4.9 O ). Together, these data suggest the interesting possibility that the overlying mesenchyme provides structural support, via integrins, to the expanding aortic endothelial tubes. 83 Rasip1 is required for esta blishment of endothelial apicobasal polarity The presence of TJs in the center of Rasip1 - / - cords, however, suggested EC polarity defects. We examined initiation of lumen formation in wild type cords and found early clearance of TJs (ZO - 1) at the cord center, at 2S ( Figure 4.10A - A” ). Thus, ECs rapidly refine their apicobasal character during lum en formation, with junctional ZO - 1 segregating from apical PODXL. In contrast, Rasip1 - / - cords displayed a distinct failure to clear the apical membrane of TJs as lumen formation initiated ( Figure 4.10B - B” ). This ectopic localization of ZO - 1 persisted in m utant cords in older embryos ( Xu et al., 2011 ) . Aberrant apical localization of the no rmally junctional ZO - 1 molecules was particularly striking upon co - staining with the apical marker PODXL. In wild type ECs, the two quickly segregated during lumen formation, with ZO - 1 at the cord periphery and PODXL along internal surfa

93 ces ( Figure 4.10C - C” ). However, i
ces ( Figure 4.10C - C” ). However, in Rasip1 - / - ECs, the two exhibited significant sustained overlap ( Figure 4.10D - D” ). ZO - 1 was not only ectopically located on apical, but also basal, EC surfaces. Transmission electron microscopy (TEM) studies also revealed failure of the cor d - to - tube transition at early stages of morphogenesis (4S stage), revealing slit - like lumenal spaces and mislocalized c ell - cell junctions ( Figure 4.10E, F ) and confirming that ectopic localization of ZO - 1 represents ectopic junctions. 84 Figure 4.10. Rasip1 - / - angioblasts display defective cell polarity and fail to localize junctional proteins to cord periphery. ( A - D, G - L ) Confocal immunofluorescence microscopy of Rasip1 +/ - ( A, C, G, I, K ) and Rasip1 - / - ( B, D, H, J, L ) aortic ECs in transverse sections : podocalyxin (green), ZO - 1 (red), and Par3 ( G - L, red). Nuclei, DAPI (blue). ( E, F ) Rasip1 null aortic cord angioblasts display mislocalized EC - EC TJs. TEM images of transverse sections of Rasip1 +/ - ( E ) and Rasip1 - / - ( F ) littermates, at 4 - somite stage. ( E, F ) Black arrows, tight junctions; white arrows, slit

94 s/isolated lumens between ECs. ( A”, D
s/isolated lumens between ECs. ( A”, D”, J ) Arrows, ectopic basal TJs; arrowheads, ectopic apical TJs; dotted line, basal endoderm surface. Scale bars, 5 µ m ( A - J ), 20 µ m( K, L ). 85 We examined cell polarity in Rasip1 - / - ECs by assessing the polarity determinant Par3. As EC polarity was recently shown to be disrupted (and Par3 ( Zovein et al., 2010 ) , we asked whether Par3 expression or localization depends on Rasip1. In wild type aortic ECs, like ZO - 1, Par3 was expressed along the entire apical membrane prior to lumen formation, but redistributed junctionally (apicolaterally) as vascular lumens formed ( Figure 4.10G, I and K ). These results were consistent with reports that Par3 localizes to TJs in polarized epithelial cells ( Jaffe et al., 2008 ) . In Rasip1 - / - ECs, by contrast, Par3 initially localized normally to the apical, pre - lumen EC surface, but it subsequently failed to be cleared during lumen formation ( Figure 4.10H, J, and L ) . Par3 could be observed, like ZO - 1, in clusters between apposed ECs along the apical membrane. Interestingly, occasionally we observed proper TJ localization in Rasip1 null ECs, and in these rare instances lum ens

95 formed normally ( Xu et al., 2011 ) .
formed normally ( Xu et al., 2011 ) . However, these isolated lumens were small and formed isolated pockets, but did not connect into functional tubes. D iscussion Here, we identify a critical requirement for Rasip1 during normal blood vessel tubulogenesis. We show that Rasip1 together with Arhgap29, a RhoA - specific GAP, suppresses RhoA signaling and dampens ROCK and non - muscle 86 Figure 4.11. Rasip1 regulates EC contractility and adhesion by modulating Rho family small GTPases. 87 myosin IIA activities in endothelial cells ( Figure 4.11 ). When Rasip1 or Arhgap29 are absent, RhoA activation is elevated, resulting in excessive actomyosin contractility, disruption of cell polarity and consequent failure of lumen morphogenesis. Basal adhesion contacts, between ECs and surrounding ECM, fail to mature (as assessed by lack of integrin activation), resulting in disassociation of endot helium from surrounding mesenchyme. Apical adhesion is also disrupted, as junctional contacts are mislocalized within the central portion of cords, blocking the ability of ECs to open coherent lumens ( Figure 4.12 ). Inappropriate segregation of apicobasal c haracter is evidenc

96 ed by sustained overlap of apical and j
ed by sustained overlap of apical and junctional (lateral) markers in Rasip1 - / - ECs. Additionally, Rasip1/Arhgap29 suppression of RhoA promotes Cdc42 and Rac1 activation, which in turn activates downstream kinases, including Pak4, Src, B - Raf, C - Raf and Erk, required for in vitro EC lumen formation ( Koh et al., 2008 ; Koh et al., 2009 ) . Rasip1 thus represents a unique molecular bottleneck, as it integrates Rho family GTPase signaling to control cell polarity and adhesion, thereby coordinating endothelial morphogenesis and blood vess el tubulogenesis. 88 Figure 4.12. The balance between EC - EC and EC - ECM adhesion is critical to vascular cord to tube transition. 89 (1) Rasip is required for endothelial cell polarity Three key points led us to expect defective cell polarity in Rasip1 null angioblasts. First, Rasip1 regulates endothelial GTPase activity. Second, decades of classical work have demonstrated regulation of cell polarity by Rho GTPases, such as Cdc42 ( Etienne - Manneville and Hall, 2002 ) . Finally, recent findings pointed to the importance of proper cell polarity during vascular lumen formation, both in early embryonic vess

97 els, as well as those formed later durin
els, as well as those formed later during embryogenesis ( Strilic et al., 2009 ; Zovein et al., 2010 ) . Cell polarity defects in Rasip1 null cord ECs we re further revealed by mislocalization of the polarity determinant Par3. Indeed, in the absence of Rasip1, we found a clear failure of Par3 to segregate normally to cell - cell junctions, away from the apical membrane. Par3 is known to associate with the aPK C polarity complex in epithelial cells ( Suzuki and Ohno, 2006 ) and to be important in establishment of arterial cell polarity ( Zovein et al., 2010 ) . We note that while Par3 was previously found to be localized basally in older arterial vessels and reduced in the asence of β 1 integrin ( Zovein et al., 2010 ) , localization of Par3 in early angioblasts of the dorsal aortae was clearly junctional, and levels appeared unchanged in Rasip1 null ECs. It is conceivable that vascuologenic vessels form differently from angiogenic vessels formed later, but that k ey signaling molecules are nonetheless conserved. 90 We note that between clusters of junctional molecules (ZO - 1/Par3/VE - cadherin), we observed slit - like openings, in which apical molecules like podo

98 calyxin/moesin were expressed normally.
calyxin/moesin were expressed normally. We speculate that m uch of the luen foration olecular pathway actually occurs „norally’ in the asence of Rasip1, albeit discontinuously. In Rasip1 null ECs, TJ/AJ localization likely occurs stochastically, such that the morphogenetic process is short - circuited, blocking functional lumen formation and tubulogenesis. (2) Rasip regulation of cell adhesion to ECM via maturation of adhesion contacts The primary defect observed in ECs in the absence of Rasip1 function is failure of proper EC adhesion regulation, both between cell s and with surrounding matrix. Failure of adhesion to matrix was evident both in live cell imaging of in vitro EC lumen formation in siRasip1 - treated cells, as well as in sections of Rasip1 - / - lumenless aortic cords. This EC - ECM failure was particularly st riking in collagen 3D cultures, as ECs extended processes that transiently adhered to the surrounding matrix as they initiated lumen formation, but snapped back and collapsed repeatedly as they lost traction. These behavioral effects can be explained by im paired focal adhesion maturation in Rasip1 - depleted cells. Whether resulting immature f

99 ocal adhesions are merely weaker, or whe
ocal adhesions are merely weaker, or whether 91 outside - in or inside - out signaling via focal adhesion components is compromised, can not be distinguishable from these studi es. It will be interesting to identify additional Rasip1 binding partners, as it can complex with number Rho and Ras family GTPases ( Mitin et al., 2004 ) , and further dissect signaling pathways and their effects on cell adhesion and architecture. An interesting point regarding maturation of endothelial adhesion contacts is that Rasip1 appears to promote engagement of multiple integrins. Specifically, decrease of both β 1 and α v β 3 in siRasip1 - depleted cells, suggests other integrins are likely simil arly regulated. Lumen dependence on multiple integrins is supported by antibody - based knockdowns targeting β 1 ( Drake et al., 1992 ) or α v β 3 ( Drake et al., 1995 ) , which blocked lumen formation in c hick aortae. Redundancy of integrin dependence on Rasip1 during lumen formation likely explains the more profoundly compromised vascular phenotype observed in Rasip1 - / - mice versus EC - specific deletions of single integrins, which have not shown global lume n failure ( Carlson et al., 200

100 8 ; Tanjore et al., 2008 ) . In Rasip1
8 ; Tanjore et al., 2008 ) . In Rasip1 - / - aortae, we suggest all required integrins have been blocked, as Rasip1 controls their activation. One relevant study bypassed the issue of integrin redundancy by examining fibronectin (FN) nul l embryos, as FN is a common ligand for a number of integrins, including β 1 and α v β 3 ( George et al., 1997 ) . Interestingly, the endocardial lumen was absent in FN - / - embryos, and a ortic ECs partially detached 92 from surrounding mesenchyme, similar to Rasip1 null embryos. However, Rasip1 - / - embryos do not phenocopy FN - / - embryos, likely because other matrix components are also required. Together, these data suggest that Rasip1 is critical to integrin function in ECs. (3) Diverse cellular mechanisms drive vascular lumen formation? An unexpected finding in our studies involve d the growing gap between the trunk mesenchyme and the lumenless Rasip1 - / - aortic cords. Our data suggests that Rasip1 - deficient aortic ECs are unable to maintain proper integrin - mediated adhesion to the surrounding mesenchymal ECM and imply that the mesen chyme actively provides support during lumen expansion. We note that while this

101 gap is pronounced surrounding the mutan
gap is pronounced surrounding the mutant aortic cords, similar gaps around other vessels are variable or absent, such as those in the yolk sac. It is therefore possible that di fferent blood vessels create lumens by different cellular mechanisms, but common molecular mechanisms. Indeed other ioloical „tues’ often depend on ultiple spatially - and/or temporally - specific mechanisms for lumen formation. The Drosophila trachea, f or instance, ranges from intracellular vacuole - based lumen generation in fine terminal cells, to multicellular budding and lumen expansion in primary tracheal 93 branches ( Lubarsky and Krasnow, 2003 ) . Therefore, although we present here a mechanism for lumen formation based on angioblast adhesion regulation, w e predict that the highly complex and heterogeneous developing vascular system ( Aird, 2007 ) will exhibit regional differences with respect to the cellular mechanisms employed to generate lumens. As we discussed in Chapter I , controversy has e merged in the field of vascular biology regarding how vascular lumens form. Some observations, both in vitro and in vivo , have suggested that lumen formation occurs via intracellul

102 ar vacuoles, that emerge and fuse ( Da
ar vacuoles, that emerge and fuse ( Davis and Bayless, 2003 ; Kamei et al., 2006 ) . While other observations have favored a mechanism based on angioblast cords rearranging their junctions to the periphe ry and opening up a central lumen based on active cytoskeletal forces ( Jin et al., 2005 ; Strilic et al., 2009 ) . Our in vivo observations of murine aortic ECs reveal no evidence for vacuole fusion, similar to ( Strilic et al., 2009 ) . However, cultured ECs in 3D matrices clearly exhibit vacuo le fusion ( Xu et al., 2011 ) . Of great interest then is that failure of lumen formation in both systems occurs in the absence of Rasip1. This places Rasip1 in a critical regulatory position over the molecular machinery that controls cellular proc esses, such as the cell polarity, cytoskeleton (i.e. cell shape), and localization or robustness of cell junctions and adhesion contacts. Future studies will be aimed at identifying commonalities and differences in different EC types. 94 (4) Rasip1 regulation of GTPase signaling Our data suggests that Rasip1 suppresses RhoA (known to inhibit lumen formation) and promotes activation of Cdc42 and Rac1 (both known positively mediate lumen f

103 ormation). Rescue experiments demonstrat
ormation). Rescue experiments demonstrate that failure of tubulogenesis in the absence of either Rasip1, or its effector Arhgap29, can be fully restored by dominant negative RhoA or siRhoA treatment. As a consequence, we have focused on Rasip1 regulation of RhoA signaling. However, the sharp decrease in Cdc42 and Rac1 activity i n the absence of Rasip1 is of particular interest, as these GTPases play direct roles in assembly and disassembly of junctional components, as well as cytoarchitecture such as filopodia formation ( Popoff and Geny, 2009 ) . Indeed, we see decreased filopodia in the absence of Rasip1 function (data not shown). In addition, Cdc42 has been directly linked to maintenance of EC - EC junctions via VE - cadherin ( Broman et al., 2006 ) , as well as vacuole - based lumen formation ( Bayless and Davis, 2002 ) . However, we identified no relevant effectors for Cdc42 or Rac1 as binding partners in our mass spectrometry screen, therefore it is possible that their regulation by Rasip1 is indirect and occurs via the RhoA pathway. 95 (5) Critical role of actomyosin contractility for vascular tubulogenesis One likely critical effector of vascular tubulogenesis i

104 s the RhoA - specific GAP and Rasip1 bi
s the RhoA - specific GAP and Rasip1 binding partner Arhgap29, which suppress es RhoA signaling specifically in developing blood vessels. Finding that Rasip1 also bound NMHCIIA, which is regulated by RhoA, raises the distinct possibility that a key role of Rasip1 is to suppress actomyosin contractility via suppression of RhoA. This is an appealing model, as angioblasts must dramatically change their shape during lumen formation, transforming from an initially rounded cuboidal morphology, to an exceedingly flattened one. Increased intracellular contractility would inhibit such a shape change. The recent report that NMHCIIA recruitment to the luminal surface of ECs was defective in the lumenless cords of VEGF heterozygotes ( Strilic et al., 2009 ) adds another interesting dimension to the role of contractility during lumen formation. In this case, lack of contractility fails to „end’ the luinal erane and „open up’ the c entral lumen. Together, the data suggests that precisely controlled levels of myosin II activity, and contractility, are required for vascular lumens: too much contractility (as in Rasip1 - / - ECs) or too little contractility (as in VEGF +/ - ECs)

105 both lead to lumen failure. 96
both lead to lumen failure. 96 C HAPTER V Summary Data presented here supports a requirement for Rasip1 function within developing ECs during blood vessel tubulogenesis . Rasip1 is a unique signaling node in endothelial cells. Knockdown of Rasip1 in frog embryos results in failure of the posterior cardinal vein and ISVs to develop and in disorganization of the vitelline plexus. Loss - of - function of Rasip1 by gene targeting in mouse inhibits the formation of patent lumens in all vessels examined, lea ding to embryonic lethality by E10.5. Rasip1 exerts its functions by interacting with two binding partners, nonmuscle myosin heavy chain IIA (NMHCIIA) and a RhoA GAP Arhgap29. Together they mediate acto - myosin activity and integrin mediated EC - ECM adhesion through regulating Rho family small GTPases, RhoA, Rac1 and Cdc42. The failing of this signaling cascade disrupts EC - EC and EC - ECM adhesion balance and the establishment of EC polarity, leading to the collapse of vascular tubes. Our findings therefore identify Rasip1 a unique endothelial specific molecule controlling EC morphogenesis from cord - to - patent blood vessel via cell polarity and adhes

106 ion regulation. This transition is fund
ion regulation. This transition is fundamental to establishing a 96 97 seamless, functional cir culatory system and has been the focus of growing interest. A t this point, multiple questions still remain to be answer ed regarding the underlying molecular a nd cellular mechanisms . For example, does Rasip1 modulate R ho directly downstream of VEGF signalin g ? And if so, w hether is membrane trafficking involved in Rasip1 related pathway s ? How are cells changing to form lumens at the center of newly formed cords? How are the EC - EC adhesion resolved at this place? Are the mechanical forces responsible for „end in’ and „flattenin’ ECs enerated from or with in ECs or from surrounding tissues? How many differ ent layers of EC - ECM cross - talk are important in this regulation? Is anything pumped into the growing cavities prior to blood flow to keep them from collapsi ng? Do different vascular beds share similar mechanism s in their lumen formation? Further studies of Rasip1 and other related molecules/pathways will help clarify these molecular relationships during vascular development. Given the central importance of blood vessels du

107 ring many diseases, such as cancer and
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