ENDOGLIN IS DISPENSABLE FOR BLOOD VESSEL

MORPHOGENESIS BUT IS REQUIRED FOR ENDOCARDIAL

CUSHION FORMATION IN THE DEVELOPING MOUSE EMBRYO

by

Gregory A. Anderson

A thesis submitted in conformity with the requirements for the degree

of Master of Science

Graduate Department of Medical Biophysics

University of Toronto

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While these forms may be included Bien que ces formulaires in the document page count, aient inclus dans la pagination, their removal does not represent il n'y aura aucun contenu manquant. any loss of content from the thesis. Canada Endoglin is dispensable for blood vessel morphogenesis but is required for endocardial cushion formation in the developing mouse embryo

Gregory A. Anderson

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

Science

Graduate Department of Medical Biophysics, University of Toronto, 2008

ABSTRACT

Endoglin (Eng) is an ancillary receptor for the transforming growth factor beta

(TGF-P) superfamily expressed primarily on endothelial cells that can bind the ligands

TGF-31, TGF-33, activin, bone morphogenetic protein (BMP)2, BMP7, BMP9 and

BMP10. Loss of endoglin expression leads to lethality in mice at embryonic day 9.5

(E9.5) and is associated with defects in , smooth muscle cell (SMC) recruitment/differentiation, and development. In vitro embryoid body (EB) and

SMC differentiation assays, as well as in vivo chimeric studies show that angiogenic and

SMC differentiation defects are not the primary deficiencies leading to lethality in these mice. Analysis of chimeric embryos and in vitro (AVC) explant assays point to a blockage in endothelial-to-mesenchymal transition (EMT) in the of endoglin null embryos, suggesting that endoglin is first required in cardiac morphogenesis in the developing embryo.

ii Table of Contents Page Abstract ii Table of Contents iii List of Abbreviations and Symbols iv

Chapter 1 Introduction 1.1 Vascular development 2 1.2 4 1.3 The TGF-p superfamily of signaling molecules 7 1.4 Endoglin 10 1.5 Endoglin and human disease 12 1.6 Endoglin and mouse genetic studies 16 1.7 Rationale 18 1.8 Thesis objectives 19 1.9 Attributions 19

Chapter 2 Materials and Methods 2.1 Embryonic stem cell derivation 21 2.2 Embryoid body formation and angiogenic sprouting assay in vitro 21 2.3 Differentiation ofES cells by OP9 co-culture 24 2.4 Chimera aggregations and staining 27 2.5 Determination of donor-derived ES cell contribution to chimeric mice by real-time PCR 28 2.6 Chimera heart histology 30 2.7 Heart EMT analysis 30

Chapter 3 Results 3.1 Endoglin is not required for endothelial differentiation or sprout formation in vitro 34 3.2 Endoglin deficient cells contribute to angiogenesis in vivo 36 3.3 Determination of the involvement of endoglin in smooth muscle cell differentiation in vitro 37 3.4 The role of endoglin in endocardial cushion formation in the developing embryonic heart 41 3.5 Endoglin mutant atrioventricular canals cannot efficiently undergo EMT 45

Chapter 4 Discussion and Concluding Remarks 4.1 Discussion 50 4.2 Concluding remarks 61

Chapter 5 References 64

in List of Abbreviations and Symbols a-SMA Alpha smooth muscle actin

P-gal Beta galactosidase

ALK Activin-like kinase

Ang Angiopoietin

AVC Atrioventricular canal

AVM Arteriovenous malformation

BMP Bone morphogenetic protein

BMPR Bone morphogenetic protein receptor

BSA Bovine serum albumin co2 Carbon dioxide

DAB 3,3 '-Diaminobenzidine

E Embryonic day of development

EB Embryoid body

EC Endothelial cell

ECM Extracellular matrix

EMT Endothelial-to-mesenchymal transition

Eng Mouse endoglin gene

ES cell Embryonic stem cell

FACS Fluorescence activated cell sorting

FISH Fluorescence in situ hybridization

Flk Fetal liver kinase

GFP Green fluorescent protein

iv HHT Hereditary Hemorrhagic Telangiectasia

HRP Horseradish peroxidase

HSC Hematopoietic stem cell

Id Inhibitor of differentiation

ITS Insulin-transferrin-selenium

KO Knock-out (i.e. deletion of gene(s))

LIF Leukemia inhibitory factor

MEF Mouse embryonic fibroblast

OFT Outflow tract

PAI Plasminogen activator inhibitor

PBS Phosphate buffered saline

PCR Polymerase chain reaction

PECAM Platelet/endothelial cell adhesion molecule

P1GF Placental growth factor

RTK Receptor tyrosine kinase

RT-PCR Reverse transcription polymerase chain reaction siRNA Short interfering ribonucleic acid

SM22a Smooth muscle 22 alpha

SMAD Sons-of-mothers against decapentaplegic

SMC Smooth muscle cell

TGF-J3 Transforming growth factor beta

Tie Tyrosine kinase with Ig and EGF homology domains

VEGF Vascular endothelial growth factor VEGFR Vascular endothelial growth factor receptor

WT Wild-type

X-gal 5-bromo-4-chloro-3-indolyl-P-D-galactosidase

VI CHAPTER 1

INTRODUCTION 2

1.1 Vascular development

Critical to the survival of the developing embryo is the uninterrupted development of the cardiovascular system. This organ system is the first to develop during embryogenesis, and its integrity is essential for proper growth and development of the

embryo, as it is necessary to supply oxygen and nutrients to the various tissues of the body, as well as to carry away wastes such as carbon dioxide and uric acid.

Differentiation and coordinated assembly of several distinct cell lineages, including the blood vessels, contractile heart, circulating blood cells, and mesenchymal support cells

are critical to the development of this complex organ system. Cardiovascular

development starts shortly after gastrulation when endothelial cell (EC) precursors termed angioblasts emerge from mesodermal tissue. This process occurs in the extra­ embryonic , and is closely associated with the formation of primitive hematopoietic stem cells (HSCs), which are necessary for formation of the developing blood system. These angioblasts and HSCs are found together in the of the yolk sac, suggesting that they have arisen from a common precursor known as a hemangioblast. In a process known as , these hemangioblast cells coalesce

into a very simple honeycomb network of EC tubes that play a key role in patterning the

in the embryo proper (Coffin et al., 1991; Risau et al., 1988). Once

this primary vascular plexus is established, these vessels are remodeled into a more

mature vasculature with a complex hierarchy in a process known as angiogenesis. This process involves coordinated sprouting, proliferation, and branching of differentiated

ECs, as well as pruning of existing immature blood vessels to form a more mature and

intricate network of blood vessels (Risau et al., 1988). Angiogenesis is required for 3 vascularization of the brain and other organs, such as the kidney, in the developing embryo, and is important in processes such as follicular development, wound healing, and tumour growth in adults (Adams and Alitalo, 2007). The final stages of remodeling of this vascular system occur when mural support cells, such as pericytes and smooth muscle cells (SMCs), are recruited to the vessel wall leading to vessel stabilization and extracellular matrix deposition and elaboration (Beck and D'Amore, 1997; Carmeliet,

2000; D'Amore, 1992).

Stimulation of angiogenic growth is a tightly regulated process in the development of a mammalian embryo, and is dependent upon the controlled expression of a variety of signaling modulators. The most intensely studied molecule that controls blood vessel morphogenesis is vascular endothelial growth factor A (VEGFA), which is part of a large family of potent angiogenic regulators including placental growth factor

(P1GF), VEGFB, VEGFC, VEGFD, and VEGFE (Ferrara et al., 2003; Shibuya, 2006).

These ligands signal through cell surface receptors known as receptor tyrosine kinases

(RTKs) of the VEGF receptor family, specifically VEGF receptor 1 (VEGFR-l/Flt-1),

VEGFR-2/Flk-l, and VEGFR-3/FU-4 (Yancopoulos et al., 2000). Another subfamily of receptor tyrosine kinases, namely the Tie receptors (Tiel and Tie2/Tek), possess potent angiogenic properties as well. Tie2 is able to bind the four members of the angiopoietin

(Ang) family, Angl-4, while the ligand for Tiel remains elusive (Jones et al., 2001).

These VEGFR and Tie receptors have been shown to be almost exlusively expressed on cells of endothelial lineage (Yancopoulos et al., 2000), which explains the effects these signaling modulators have on EC proliferation, vasculogenesis, angiogenesis, anti­

inflammatory processes, and pericyte/SMC association (Ward and Dumont, 2002). The 4 importance of these ligands in vascular development is exemplified by the absolute requirement for tightly regulated levels of their expression, as mice lacking these gene products die during embryogenesis with severe circulatory system defects (Carmeliet et al., 1996a; Dumont et al., 1994; Ferrara et al., 1996; Puri et al., 1999; Sato et al, 1995).

1.2 Heart development

Vascular development is highly coordinated with the differentiation and morphogenesis of the heart so as to achieve unidirectional and continuous blood flow

(Srivastava and Olson, 2000). The process of heart development involves a large number of precisely orchestrated molecular events, the perturbation of which can lead to catastrophic consequences such as congenital heart defects or death. Formation of the heart begins shortly after gastrulation when cardiomyocytes cluster along the ventral midline of the embryo in an area termed the heart field (Figure 1.1, I), which is a region of the early embryo that is competent to respond to inductive signals (Jacobson and Sater,

1988). These cells converge and begin maturing to form a linear heart tube that has contractile motion and is composed of an outer myocardium, an inner endocardial cell lining, and an extensive extracellular matrix (ECM) known as the cardiac jelly (Figure

1.1, II) (Armstrong and Bischoff, 2004). In a process that is similar in all vertebrates, the linear heart tube then undergoes rightward looping which is required for proper orientation of the ventricles with respect to the atria, and for alignment of the heart chambers with the vasculature (Figure 1.1, III) (Srivastava and Olson, 2000). Although the linear heart tube is patterned along its anterior-posterior axis to form the FIGURE 1.1

I II III

Figure 1.1. Schematic representation of mammalian heart formation. I, Cardiomyocytes cluster along the ventral midline of the embryo in a cardiac crescent in the heart field. II, Fusion of the specified cardiomyocytes forms a linear heart tube that is patterened along its anterior-posterior axis. Ill, Formation of the endocardial cushions in the region between the common atria and ventricle (atrioventricular canal) and in the distal region of the conotruncus/outflow tract allows for rightward looping of the linear heart tube to occur. This properly orients the atrial region of the common heart tube posterior to the common ventricle. A, atrium; V, ventricle; CT, conotruncus/outflow tract; LA, left atrium; RA, right atrium; LV, left ventricle; RV, right ventricle; AVC, atrioventricular canal. Adapted from Schroeder et ah, 2003. 6 individual heart chambers (left and right atria and ventricles) of the mature heart, it is not until this looping step has occurred that these chambers become morphologically distinguishable from one another (Schroeder et al., 2003). Proliferation and maturation of these cells within each cardiac chamber then occurs, and is necessary to support the increasing hemodynamic load during embryonic growth. The molecular details of these final maturation steps are largely undefined, though they likely involve the integration of extracellular signals with transcription factor programs, both cardiac-specific as well as those that are ubiquitously expressed.

Unilateral blood flow in the developing heart begins with rhythmic contractions and localized extracellular expansion within the regionally restricted endocardial cushions that act as primitive valves in both the atrioventricular canal (AVC) and the outflow tract (OFT). Endocardial cushion cell transformation occurs at the AV boundary to initiate formation of the mitral and tricuspid valves and somewhat later in the OFT to form the aortic and pulmonary valves (Markwald et al., 1977). These endocardial cushions form when soluble signals from the myocardium reach specific cells of the endocardial layer of the cushion, causing these cells to transdifferentiate into mesenchymal cells and invade the underlying ECM by a process known as endothelial to mesenchymal transition (EMT) (de la Pompa et al., 1998; Ranger et al., 1998). The number of gene expression pathways involved in endocardial cushion formation is immense, as shown by the growing number of single gene mutations associated with defective endocardial cushion formation and subsequent defects. Several members of the transforming growth factor beta (TGFP) superfamily have been shown to be important for endocardial cushion formation in both avian and mammalian species 7

(Bourdeau et al., 1999; Brown et al., 1999; Camenisch et al., 2002; Ma et al., 2005;

Mercado-Pimentel et al., 2007), suggesting that they are major players in this highly complex interplay of precisely regulated signals required for cardiac development.

1.3 The TGF-P superfamily of signaling molecules

The TGF-(3 superfamily is a large family of cytokines necessary for diverse cellular processes in most living organisms. The ligands of this superfamily include transforming growth factors (TGFs), activins, and bone morphogenetic proteins (BMPs).

These signaling molecules have a wide range of temporal and spatial expression patterns, and are necessary for various developmental processes including controlling the formation of the neural tube, limbs, cartilage, bone, and sexual organs (Massague et al.,

1994), as well as being important for several cellular processes, including differentiation, migration, and wound repair (Mummery, 2001). Three isoforms of the TGF-P ligand exist: TGF-pi, 2, and 3, of which TGF-pi is the most important for blood vessel development and remodeling (Harradine and Akhurst, 2006). All three of these isoforms are secreted from cells as a high molecular weight latent complex in which the C-terminal mature homodimer is non-covalently associated with a dimer of its N-terminal pro- region; cleavage of this TGF-P dimer precursor then occurs in the extracellular milieu to activate the ligand (Annes et al., 2003).

The signal transduction pathways mediated by the TGF-P superfamily have been characterized in vitro and in vivo. Chemical cross-linking studies have identified three receptor types on the basis of molecular weight as type I (TpRI), type II (TpRII), and type III (TpRIII) on most mammalian cells in culture (Mummery, 2001). TGF-P binds 8 directly to TpRII (Lin et al., 1992) and facilitates the recruitment of TpRI to form a stable

complex at the cell surface (Ebner et al., 1993). The autophosphorylated and

constitutively active TpRII then phosphorylates serine/threonine residues on the

cytoplasmic tail of TpRI, causing it to become active (Massague et al., 1994). In mammals, the TGFP type II receptor family consists of five members: TPRII, BMPRIIA,

BMPRIIB, Activin (Act)RIIA, and ActRIIB, while the TGFp type I receptor family

consists of seven members: activin receptor-like kinase (ALK)l-7 (Harradine and

Akhurst, 2006), suggesting that these receptors mediate multiple signals (Figure 1.2).

Once activated, the type I receptor is able to phosphorylate and activate downstream

signalling molecules known as the SMAD intracellular transcription factors (Shi and

Massague, 2003). With respect to TGFP signalling, the most relevant type I and II

receptors are TpRII, ALK1, and ALK5. When recruited into a complex with TPRII and

activated, ALK5 will interact with and phosphorylate SMAD2 and SMAD3 which then

form a complex with SMAD4 and shuffle to the nucleus. This SMAD complex then up-

regulates the transcription of target genes such as plasminogen activator inhibitor 1 (PAI-

1), connexin 37, and SM22a, all of which are necessary for extracellular matrix

deposition and the resolution phase of angiogenesis (Ota et al., 2002). Conversely, when

ALK1 is recruited into the TpRII complex and is activated, it interacts with and

phosphorylates a set of SMADs normally associated with BMP signalling, namely

SMAD1, SMAD5, and SMAD8 (Oh et al., 2000). These SMADs then complex with

SMAD4 and shuffle to the nucleus to upregulate the transcription of a different set of FIGURE 1.2

Figure 1.2. The receptors for the TGF-P superfamily mediate many complex signalling pathways in endothelial cells. The type II receptors (ActRIIA, ActRIIB, T|3RII, BMPRII) bind their respective ligands, thus activating downstream SMAD cascades within the cell. These two different SMAD pathways (SMAD2/3 and SMAD 1/5/8) interact with the common SMAD4 and shuffle to the nucleus, where they upregulate the transcription of different target genes. 10

target genes, namely inhibitor of differentiation 1 (Id-1), Id-2, SMAD6 and SMAD7, all

of which are necessary for cellular migration, proliferation, and the activation phase of

angiogenesis (Ota et al., 2002). Thus, it appears that these two competing signalling pathways counter-balance one another and control the angiogenic state of the endothelial

cell.

There are several factors that add complexity to the TGF-p signalling paradigm;

for instance, in vitro biochemical data shows that ALK5 must be recruited into a stable

complex with TPRII and ALK1 in order for proper ALK1 signalling through SMAD1/5/8

(Goumans et al., 2003). In vivo transgenic mouse data, on the other hand, show that

ALK1 and ALK5 do not have overlapping expression patterns in mouse tissues (Seki et

al., 2006), which suggests alternate interpretations for the in vitro data. In addition, the

role that endoglin, a receptor for the TGF-P superfamily, plays in modulating these

signaling pathways is complex and the use of different experimental systems has lead to

conflicting data in terms of the contribution of endoglin to the potentiation of TGF-P

signals through the various ALK pathways (Lebrin et al., 2004; Pece-Barbara et al.,

2005). This supports the need for a developmentally relevant in vivo system to tease out

the role played by endoglin in early mammalian development.

1.4 Endoglin

Endoglin is a 180 kD homodimeric type I transmembrane protein that is expressed

predominantly on ECs from early gestation through to the adult stage (Gougos and

Letarte, 1988a), though its expression is also found on activated monocytes, tissue

macrophages, erythroid precursors, and synctiotrophoblast of term (Gougos and 11

Letarte, 1988b; Gougos et al., 1992; Lastres et al., 1992). The endoglin protein was

identified in Dr. Letarte's laboratory (Quackenbush and Letarte, 1985), and was

subsequently cloned, sequenced, and characterized as a disulfide-linked homodimer

(Gougos and Letarte, 1990). Endoglin was mapped to chromosome 9q34->qter in humans by fluorescence in-situ hybridization (FISH) analysis (Fernandez-Ruiz et al.,

1993), and its genomic sequence revealed 14 exons (McAllister et al., 1994). Endoglin was also shown to be highly expressed on endocardial cushion tissue during

early human cardiac development (Qu et al., 1998). Initial biochemical studies of

endoglin demonstrated its role as an ancillary receptor of the ligands TGF-pi and TGF-

P3, but not TGF-P2 (Cheifetz et al., 1992). In addition to these two ligands, endoglin has

also been shown to bind activin, bone morphogenetic protein (BMP)-2, BMP-7, BMP-9,

and BMP-10 (Barbara et al., 1999; David et al., 2007; Scharpfenecker et al., 2007), though the biochemistry and functional significance of these interactions has not been

experimentally explored to the same extent as for TGFpi. Further biochemical analyses

determined that mammalian cells express two isoforms of endoglin: L-endoglin (L-Eng), the full-size and commonly expressed isoform, and S-Eng, a short isoform of L-Eng

(Bellon et al., 1993). L-Eng contains 47 amino acids in its cytoplasmic tail and is

constitutively phosphorylated, whereas S-Eng only contains 14 amino acids in its

cytoplasmic tail and is unphosphorylated (Bellon et al., 1993). Characterization of the promoter of the endoglin gene demonstrated that endoglin expression is inducible by

TGF31, and has stronger inducibility in ECs compared to other cell types such as

epithelial cells (Graulich et al., 1999; Rius et al., 1998), thus suggesting that the endoglin 12 promoter may be useful for future gene therapy scenarios that would effectively target

ECs specifically.

Endoglin is considered to be an ancillary receptor for the TGFP superfamily, much like TpRIII/betaglycan, as neither have signalling properties of their own. Though endoglin and betaglycan share a high degree of amino acid similarity (Cheifetz et al.,

1992), their function appears to be quite different, as betaglycan is able to bind all three isoforms of TGF-p. The role that endoglin plays in TGFp signalling is very complex and not fully understood, but what has been demonstrated is that endoglin is able to bind the

TpRII either in the presence or absence of its ligand, TGFpi or TGFp3, and can then help to recruit the TpRI into the complex (Guerrero-Esteo et al., 2002). Importantly, in vitro studies have shown that the TpRI that can be recruited by endoglin can be either ALK1 or

ALK5, which, as described above, promote opposing downstream SMAD signalling cascades that lead to the activation phase or resolution phase of angiogenesis, respectively. Thus, the earliest biochemical analyses supported a role for endoglin in fine- tuning the balance between ALK1 and ALK5 signalling in ECs, thereby controlling the activation state of these cells (Goumans et al., 2002) (Figure 1.3).

1.5 Endoglin and human disease

Critical evidence that endoglin has a highly important role in vascular development is revealed by the identification of human ENDOGLIN (ENG) as the gene mutated in the disease Hereditary Hemorrhagic Telangiectasia type 1 (HHT1)

(McAllister et al., 1994). HHT is an autosomal dominant vascular disorder that has variable penetrance and expressivity, and affects approximately 1 in 10,000 13

FIGURE 1.3

Vessel Maturation New Vessel Formation (resolution phase of angiogenesis) (activation phase of angiogenesis)

Figure 1.3. Endoglin potentiates TGF-P signalling through both the ALK1 and the ALK5 pathways. Upon binding of TGF-pi or -P3, the TpRII recruits a TpRI into the complex which can be either ALK1 or ALK5. If ALK1 is recruited to the complex, this allows binding of endoglin, which potentiates downstream signalling of SMAD1/5/8 with SMAD4, thus upregulating the transcription of genes that lead to the activation phase of angiogenesis. If ALK5 is recruited to the TpRII complex, this also allows binding of endoglin; this is a separate signalling pathway that signals through SMAD2/3 along with SMAD4, which upregulates the transcription of genes that lead to the resolution phase of angiogenesis. 14

individuals worldwide (Guttmacher et al., 1995), though a higher prevalence rate is found

in some geographically isolated areas. Over 150 mutations in ENG have been identified

(Abdalla and Letarte, 2006) and clinical manifestations of this disease are quite variable both within and between families. These manifestations have been attributed to a breakdown in capillary structures that lead to blood shunting between major arteries and

veins (Braverman et al., 1990) that can lead to life-threatening complications such as

internal hemorrhages and stroke. For example, spontaneous recurrent nosebleeds from

telangiectases in the nasal mucosa is the presenting sign in greater than 90% of HHT patients (Assar et al., 1991), and the severity of these nosebleeds typically increases with

age and may require blood transfusions to combat anemia (Abdalla and Letarte, 2006).

Telangiectases can also develop in the gastrointestinal tract of older patients (Plauchu et

al., 1989), and AVMs in the liver are seen in up to 40% of HHT patients (Buscarini et al.,

2004). Up to 48% of HHT1 patients will present with a pulmonary AVM, which is

caused by a direct connection between the and vein (Letteboer et al.,

2006) that leads to hypoxemia, stroke, and brain abscess (Ference et al., 1994; Kjeldsen

et al., 2000; Shovlin et al, 1995). HHT2, a disease similar to HHT1 but with a somewhat

lessened incidence of pulmonary AVMs and a higher incidence of liver AVMs, is

attributed to a mutation in the ALK1 gene (Buscarini et al., 2004). In both cases, though

the diseases are autosomal dominant, biochemical data suggests that the mechanism of

action is loss of function due to haploinsufficiency, and not dominant gain of function

(Abdalla et al., 2000; Pece et al., 1997). As such, loss-of-function mouse models are

appropriate for investigation of the HHT disease phenotype. 15

In addition to being involved in HHT1, recent research demonstrated a role for endoglin in the pathogenesis of preeclampsia (Venkatesha et al., 2006). Preeclampsia is a pregnancy-specific condition that is characterized by the onset of hypertension and proteinuria in the third trimester of gestation; it is a complicating factor in 5% of pregnancies worldwide, and is a major cause of maternal, fetal, and neonatal mortality, especially in developing countries (Sibai et al., 2005). The disappearance of disease symptoms following delivery of the baby has pointed to the central role of the placenta in this disease. The clinical manifestations of preeclampsia, including vasoconstriction, organ ischemia, and increased vascular permeability suggest that widespread EC dysfunction is the major problem (Sibai et al., 2005). It has been shown that the level of circulating soluble endoglin (sEng) in the serum of pregnant women is a prognostic indicator of disease state (Levine et al., 2006; Venkatesha et al., 2006), thus suggesting that angiogenesis is disturbed in preeclamptic . Just how important circulating levels of sEng will be in determining the risk factor for preeclampsia in pregnant women remains to be determined, but it appears to be the best prognostic indicator identified thus far for this disease (Levine et al., 2006).

The role of endoglin in angiogenesis has prompted several groups to analyze its contribution to the progression of various cancers. The aim of several antiangiogenic therapeutic strategies is to induce tumour regression and to inhibit its metastatic spread by blocking the needs of nutrients and oxygen exchange to these neoplastic cells

(Fonsatti et al., 2003). It has long been established that ECs of the tumour vasculature undergo a more rapid proliferation than quiescent ECs in the vasculature of normal tissues (Denekamp, 1990), and as such, this intriguing therapeutic approach aims to 16 selectively target proliferating ECs within tumour-associated blood vessels. In solid tumours, endoglin has been shown to be expressed on the ECs of intratumoral blood vessels and on tumour stromal components (Fonsatti et al., 2001), in sarcomas of different histotypes (Fonsatti et al., 2001), and in ovary (Henriksen et al., 1995) and breast carcinomas (Fonsatti et al, 2000). In a melanoma study, it was shown that endoglin was present on the ECs of 25% and 34% of primary and metastatic lesions, respectively (Altomonte et al., 1996). Overall, these studies suggest that microvessel density as tested by endoglin staining is a good prognostic indicator for cancer progression as well as sEng levels in serum. And although no anti-angiogenic therapies using antibodies to endoglin or small molecule inhibitors of endoglin have been tested in clinical trials, this remains a very active area of research, and may yield some exciting results in the years to come.

1.6 Endoglin and mouse genetic studies

The important functional role of endoglin in the cardiovascular system was confirmed with analysis of mice bearing targeted mutations in the endoglin gene. Three independent laboratories deleted the endoglin gene in mice {Eng'1'), and though they all focused on different aspects of the phenotype of these mice, all of the groups described overlapping and complementary defects in the cardiovascular system. While all groups concluded that vasculogenesis occurred properly and early gastrulation of the embryos was normal, development arrested between E9.5 and E10.5 of gestation. Bourdeau et al.

(1999) described defects in angiogenesis in the embryo proper that resulted in fragile blood vessels susceptible to hemorrhage, as well as defects in heart development; 17 specifically, endocardial cushion formation in the atrioventricular canals of these embryos was abrogated. Arthur et al. (2000) focused more specifically on the yolk sacs of the embryos, and noted that yolk sac angiogenesis and hematopoiesis were disrupted, which led to growth retardation and overall fragility of the Eng1' embryos. Li et al.

(1999) focused their studies on a completely different aspect of development: they described defects in vascular smooth muscle cell (SMC) recruitment and differentiation to newly forming blood vessels in the embryos, and concluded that these defects pre­ empted defects in angiogenesis and heart development. These complementary and yet somewhat opposing papers sparked much debate over the primary cellular role of endoglin in mammalian development. However, the problem in determining what the primary defect is in these Eng1" embryos is apparent when one takes into account the fact that these three processes of angiogenesis, vascular SMC differentiation, and heart development are all closely temporally spaced during mammalian development and they are all dependent upon one another. As such, it is reasonable to propose that if heart development is altered, this will have downstream effects on both angiogenic processes and SMC differentiation, and vice versa. In order to determine which of these developmental processes is the primary defect in embryos devoid of endoglin, experiments need to be designed that address each process in an individual manner, without the confounding influence of the other factors.

In addition to characterizing the developmental defects seen in embryos lacking endoglin expression, both Bourdeau et al. (2001) and Arthur et al. (2000) described several features of adult endoglin heterozygous (Eng+I~) mice that conformed to clinical manifestations of HHT1. It was suggested that these mice could serve as a mouse model 18 of the disease, and clearly indicated that in addition to its essential role in development, much still had to be determined about the role of endoglin in human disease.

1.7 Rationale

Our understanding of the molecular mechanisms that control the differentiation and assembly of the cell lineages involved in cardiovascular development has grown significantly through the characterization of signaling modulators such as TGF-p* and their respective receptors. Endoglin is an ancillary transmembrane receptor for the TGF-P superfamily that is expressed in ECs from early gestation to the adult stage. In humans, the critical role of endoglin in vascular development is shown in patients with HHT1, preeclampsia, and various forms of cancer.

Mice bearing a homozygous null mutation in endoglin (Eng~'~) die at mid- gestation due to potential defects in angiogenesis, SMC differentiation, and heart development. However, the specific role of endoglin in these three processes needs to be clarified, due to their close temporal spacing during mammalian development.

Determining the primary defect may point to a functional role for endoglin in early developmental processes. As such, experiments need to be devised that look at each of these individual aspects of the function of endoglin in an independent manner so that the primary developmental defect seen in these Eng1' mice can be deciphered. As more information is gathered about endoglin and its role in TGF-P signalling, the greater our ability will be to provide more effective strategies for patients who suffer from HHT1, preeclampsia, and various forms of cancer. 19

1.8 Thesis objectives

The purpose of the work presented was to determine the relative contribution of defects in angiogenesis. SMC differentiation, and cardiac development to the lethal phenotype of endoglin deficient mice. My hypothesis is that defective heart development is the primary defect seen in endoglin deficient mice. To address these objectives we used in vitro and in vivo strategies that allow analysis of each of these cellular processes individually. In order to perform these experiments, several new embryonic stem (ES) cell lines were derived that are wild-type (Eng+I+), heterozygous for endoglin (Eng+I), or endoglin null (Eng '). An in vitro embryoid body assay and analysis of in vivo chimeric mouse embryos addressed whether angiogenesis requires normal endoglin expression.

Furthermore, a smooth muscle progenitor cell assay was employed to investigate whether

SMC differentiation is affected by loss of endoglin expression, and an in vivo chimeric mouse assay and an in vitro EMT assay to determine whether or not the lack of endoglin negatively affects cardiac development.

1.9 Attributions

The aggregations cited in Table 3.1 were performed in the Sunnybrook

Transgenic Facility by either Sharon Karamath or Gillian Sleep. All histology slides shown in Figure 3.4 were cut in the Sunnybrook Histology Laboratory by Petia

Stefanova. The embryos shown in Figure 3.4A and B were collected and stained in collaboration with Amna Karabegovic. CHAPTER 2

MATERIALS AND METHODS 21

2.1 Embryonic stem cell derivation

The Eng+/+ and Eng+/~ embryonic stem (ES) cells (clones 1B-62 and 1B-63, respectively) were generously provided by Dr. Michelle Letarte (The Hospital for Sick

Children, Toronto, Ontario, Canada). The Eng+~ ES cells were previously generated

(Bourdeau et al., 1999) by gene targeting the parental wildtype 129/Ola-derived E14 ES

cell lines, replacing 609 base pairs (bp), including exon 1 of Eng and its initiation codon, with the E. coli LacZ gene and a neomycin resistance cassette, while leaving the endoglin promoter intact. Thus, endoglin expressing cells can be identified following staining with

X-gal (5-bromo-4-chloro-3-indolyl-|3-D-galactosidase). I generated Eng1' ES cells from

Eng1' clone 1B-63 by selection with high concentration G418 (5.0 mg/mL) for 10 days,

changing media every second day (Mortensen et al., 1992). Multiplex polymerase chain reaction (PCR) was used to screen resistant colonies for Eng'1' as previously described

(Bourdeau et al., 1999). Independently derived Eng'1' clones (1F-10 and 2E-10) and

Eng+/~ clones (1G-10 and 2A-9) (that had gone through G418 selection but did not undergo homologous recombination) were used for all subsequent embryoid body

experiments, ES-cell-differentiation experiments, and chimera aggregations (Figure 2.1).

2.2 Embryoid body formation and angiogenic sprouting assay in vitro

ES cells were cultured on a murine embryonic fibroblast (MEF) feeder layer

(MEFs were growth arrested by Y-irradiation) in media supporting the growth of ES cells

and composed of Dulbecco's modified Eagle's medium (Sigma-Aldrich, St. Louis, MO)

supplemented with 15% fetal bovine serum (FBS; Gemini Bio-Products, Woodland, CA),

2 mM Glutamax (Gibco, Grand Island, NY), 100 U/mL penicillin (Gibco, Grand Island, 22

FIGURE 2.1

DS E9 F9 G9 A10 610 C10 010 E10 F10 G10

476bp 9B ' MriMMB-^^fHMHl * v llM^mtoM&L ^ J|^^H i|piip:llp:'MM 'vp *— 3Mt>p

W WW A

F8 r« 06 H6 EB AS 010 E10

Figure 2.1. Multiplex PCR gels of independently derived ES cell clones following selection in high G418. (A) and (B) represent two independent experiments. The ladder on the left hand side represents 100 bp increments, increasing from bottom to top. The band at 300 bp indicates the WT endoglin allele, while the band at 476 bp represents the mutant LacZ allele. * represents the Eng'~ clone selected from each experiment, while ** represents the Eng+I~ clone selected from each experiment. The clone numbers are written at the top. 23

NY), 100 (xg/mL streptomycin (Cibco, Grand Island, NY), non-essential amino acids

(Gibco, Grand Island, NY), 1 mM sodium pyruvate (Gibco, Grand Island, NY), 2- mercaptoethanol (Sigma-Aldrich, St. Louis, MO), and 2,000 U/mL recombinant leukemia

inhibitory factor (LIF; Chemicon International, Harrow, UK). The ES cells were grown at

37°C with 5% CO2, and splitting of the cells was performed every second or third day using IX trypsin-EDTA (Sigma-Aldrich, St. Louis, MO). Differentiation of the ES cells was induced by removing LIF from the culture medium, aggregating the cells into

embryoid bodies (EBs) by placing 20 uL drops of 1200 cells/drop on the lid of a non­

adherent 96-well tissue culture dish, and placing the lid over sterile mineral oil (day 0).

The drops were left hanging on the lid for 4 days at 37°C and 5% CO2. After 4 days, each

individual EB was plated into one well of an 8-well glass culture slide (Becton Dickinson

(BD) Falcon, Franklin Lakes, NJ). The EBs were cultured for a further 6 days, with the

media being changed on the third day.

The cultured EBs were washed with phosphate-buffered saline (PBS; Sigma-

Aldrich, St. Louis, MO) and fixed with cold acetone:methanol (1:1) for 5 minutes. After being washed with PBS twice, the EBs were then treated with 0.3% hydrogen peroxide

for 30 minutes at 4°C. The EBs were then washed twice with PBS, blocked for 60

minutes at room temperature in PBS containing 1% goat serum (Sigma-Aldrich, St.

Louis, MO) and 0.2% bovine serum albumin (BSA; Gibco, Grand Island, NY) ("blocking

solution"), and incubated with a monoclonal rat anti-mouse PECAM-1 antibody (BD

Pharmingen, Franklin Lakes, NJ) diluted in blocking solution overnight at 4°C. The

following day the EBs were washed 3 times for 10 minutes each with PBST [PBS with

Ca2+ and Mg2+ (Sigma-Aldrich, St. Louis, MO) containing 0.05% Tween-20 (Sigma- 24

Aldrich, St. Louis, MO)], and incubated with a biotinylated secondary goat anti-rat IgG antibody (Vector Laboratories, Burlingame, CA) in blocking solution for 60 minutes at room temperature. The EBs were then washed 3 times for 10 minutes each with PBST,

and then incubated with streptavidin-horseradish peroxidase (DakoCytomation, Glostrup,

Denmark) for 30 minutes at room temperature. Following 4 washes with PBS, a

chromogen substance that is a substrate for horseradish peroxidase (DAB kit from Vector

Laboratories, Burlingame, CA) was added to the EBs for 5 minutes at room temperature

and rinsed with distilled water. After mounting with Entellan® (EMD Chemicals,

Darmstadt, Germany), the slides were analyzed and photographed using a transmitted

light Leica DM L microscope with DC camera attachment (Leica, Heerbrugg,

Switzerland) (Figure 2.2). The EBs were then assessed for sprouting angiogenesis based

on the following categories: I, no angiogenic sprout formation from vasculogenic region;

II, fewer than 10 angiogenic sprouts from vasculogenic region; III, greater than 10

angiogenic sprouts from vasculogenic region. Scoring was performed by a blinded

observer on two separate days.

2.3 Differentiation of ES cells by OP9 co-culture

ES cell/OP9 differentiation co-cultures were performed using a modified method

of Nakano et al (1994). 5xl05 undifferentiated ES cells were seeded onto green

fluorescent protein (GFP)+ OP9 stromal cell monolayers (a generous gift from Dr. Juan

Carlos Zufiiga-Pfiucker) in 10-cm dishes for 5 days in a media supporting the growth of

OP9 cells (minimum essential alpha medium with L-glutamine (Gibco, Grand Island,

NY) supplemented with 20% FBS (HyClone, Logan, Utah), 100 U/mL penicillin (Gibco, 25

FIGURE 2.2

In vitro ES cell differentiation ES Cells Gen* 0418

Eng*h Eng*l- Eng-I- J

Hanging Drop EB Culture OPS Co-Culture 1 Rk-1+ Cels

Microscope Sfida e I e e 0P9 Co-Culture e e e e

Quantitative 1 Quantitative analysis of EC analysis of differentiation SMC and vascular differentiation network formation

Figure 2.2. In vitro ES cell differentiation overview. Heterozygous ES cells were obtained after gene targeting of a wild-type ES cell source (Bourdeau et al., 1999), and mutant ES cells were obtained following selection of heterozygous cells in high concentrations of G418. All three ES cell lines were then used for subsequent in vitro analysis by hanging drop EB culture, or OP9 coculture as described. 26

Grand Island, NY), 100 ug/mL streptomycin (Gibco, Grand Island, NY), and 2- mercaptoethanol (Sigma-Aldrich, St. Louis, MO) ("OP9 media"). After 5 days of co- culture, cells were harvested using IX trypsin-EDTA, made into a single-cell suspension by vigorous pipetting, and stained with a phycoerythrin-conjugated rat anti-mouse Flk-

1/VEGFR2 antibody (BD Pharmingen, Franklin Lakes, NJ) for 45 minutes on ice. After washing with, and resuspending the cells in PBS supplemented with 2% FBS (Gemini

Bio-Products, Woodland, CA), Flk-1+ progenitor cells were sorted by fluorescence activated cell sorting (FACS) using a BD FACSDiva or BD FACSAria cell sorter. 4xl05

Flk-1+ cells were then plated onto a fresh OP9 monolayer in a 6-well dish (BD Falcon,

Franklin Lakes, NJ) and cultured for a further 4 days in OP9 media.

The cultured cells were then washed with PBS (Sigma-Aldrich, St. Louis, MO), and fixed with 4% paraformaldehyde (Sigma-Aldrich, St. Louis, MO) for 10 minutes at

4° C. The cells were then washed twice with PBS and then treated with hydrogen peroxide solution [50% methanol, 10% hydrogen peroxide (from 30% stock; Sigma-

Aldrich, St. Louis, MO), 40% PBS] for 30 minutes at 4° C. The cells were then washed twice with PBS, blocked with blocking solution [PBS supplemented with 1% goat serum

(Sigma-Aldrich, St. Louis, MO) and 0.2% bovine serum albumin (BSA; Gibco, Grand

Island, NY)] for one hour at room temperature, and then incubated with a primary monoclonal antibody diluted in blocking solution [mouse anti-mouse a-smooth muscle actin (Sigma-Aldrich, St. Louis, MO)] overnight at 4° C. The following day the cells were washed 3 times for 10 minutes each with PBS containing Ca2+ and Mg + (Sigma-

Aldrich, St. Louis, MO) and 0.05% Tween-20 (Sigma-Aldrich, St. Louis, MO) ("PBST") and then incubated with a horseradish peroxidase conjugated goat anti-mouse IgG 27 antibody diluted (Bio-Rad Laboratories, Hercules, CA) in blocking solution for 60 minutes at room temperature. The cells were then washed 3 times for 10 minutes each with PBST, and then visualized with a chromogen substance that is a substrate for horseradish peroxidase (DAB kit from Vector Laboratories, Burlingame, CA) for 5 minutes each. The wells were analyzed and photographed using an inverted Leica DM IL microscope with DC camera attachment (Leica, Heerbrugg, Switzerland) (Figure 2.2).

2.4 Chimera aggregations and staining

Chimeric mice were produced by aggregation of wild-type host embryos (8-cell morulas) with ES cells (ES-cell ^--> embryo aggregation). In our chimeras, the donor cells that normally express endoglin are stably labeled with P-galactosidase, thus allowing them to be distinguished from the wild-type host cells following X-gal staining.

All chimeric embryos were dissected at E10.5 and kept in pH 7.3 PBS on ice until all embryos were collected. The embryos were then fixed in PBS supplemented with glutaraldehyde (Sigma-Aldrich, St. Louis, MO), EGTA (Bio-Shop Canada Inc.,

Burlington, ON, Canada), magnesium chloride (Sigma-Aldrich, St. Louis, MO), and formaldehyde (Sigma-Aldrich, St. Louis, MO) for 30 minutes at room temperature. The embryos were then washed 3 times for 5 minutes each in PBS supplemented with magnesium chloride (Sigma-Aldrich, St. Louis, MO), deoxycholate (Sigma-Aldrich, St.

Louis, MO), and Nonidet P40 (Roche Diagnostics, Indianapolis, IN) ("wash buffer"), and then stained with X-gal solution [wash buffer supplemented with X-gal (Biosynth AG,

Switzerland), potassium ferrocyanide (Sigma-Aldrich, St. Louis, MO), and potassium ferricyanide (Sigma-Aldrich, St. Louis, MO)] for 2 - 4 hours at 37° C. The embryos were 28 then washed in PBS for 5 minutes, and then fixed in 10% formaldehyde for 2 - 3 hours at room temperature. The embryos were then rinsed in PBS, and observed and photographed using a Leica M stereoscope with DC camera attachment (Leica,

Heerbrugg, Switzerland) (Figure 2.3).

2.5 Determination of donor-derived ES cell contribution to chimeric mice by

real-time quantitative genomic PCR

Real-time PCR primers were created based on DNA sequence data that would amplify a region of the mutant LacZ allele (donor cells). The Tie2 allele was used as a baseline genome, and the percent contribution of samples were quantitated relative to this. The primer sequences were as follows:

LacZ allele forward: TATCTCTGGATACCGGATAAG

LacZ allele reverse: TGTAAAACGACGGGATCATCG

Tie2 allele forward: AAGAGCGAGTGGACCATGCGA

Tie2 allele reverse: AGGAGCAAGCTGACTCCACAG

Chimera embryo yolk sacs or tail clippings were digested in PBS containing pH

8.0 Tris (Sigma-Aldrich, St. Louis, MO), pH 8.0 EDTA (BioShop, Burlington, ON),

0.5% sodium dodecyl sulfate (Gibco, Grand Island, NY), and proteinase K (Sigma-

Aldrich, St. Louis, MO) overnight at 55° C. The quantity of DNA in each sample was quantified using an UltroSpec 3100 (Fisher Scientific, Ottawa, ON) spectrophotometer, and 100 ng of DNA was used for real-time PCR for each sample. The amount of donor cell DNA was determined by comparing the amount of LacZ amplicons in each sample 29

FIGURE 2.3

In vivo chimera analysis

ES Cells WHd-type Mating

£ng+/-or£hjr-/- WT WT

X / I Pseudo-pregnant recipient female

E10.5 Chimeras I Compare Big* and Big* contribution to: - Embryonic vasculature . Extraembryonic vasculature -Heart

Figure 2.3. In vivo chimeric mouse experimental overview. Heterozygous or mutant endoglin ES cells were aggregated with an 8-cell morula from a wild-type cross. The resulting aggregate was injected into a pseudo-pregnant female recipient mouse, and chimeric embryos were collected at El0.5. 30 to a standard curve created from a dilution series of pooled DNA from digested yolk sacs of Eng' ' embryos. The amount of host cell DNA was determined by comparing the amount of Tie2 amplicons in each sample to a standard curve created from a dilution series of pooled DNA from digested yolk sacs of Eng1' embryos. The amount of LacZ amplicons in each sample relative to the amount of Tie2 amplicons in each sample determined the percent donor cell contribution.

2.6 Chimera heart histology

Following whole mount X-gal staining, serial sections of the embryonic chimeras in the transverse plane or sagittal plane, as indicated, were made through the atrioventricular canals of the embryonic hearts. The sections were then counter-stained with nuclear fast red, analyzed, and photographed using a transmitted light Leica DM IL microscope with DC camera attachment (Leica, Heerbrugg, Switzerland).

2.7 Heart endothelial to mesenchymal transition (EMT) analysis

Timed matings between Eng+/~ mice were established with the morning of a vaginal plug defined as embryonic day E0.5. For all analyses, in order to minimize slight differences in developmental age of wild-type (Eng+I+), heterozygote (Eng+/), and mutant

(Eng'") embryos, at least 9 embryos of each genotype were analyzed (3 embryos from 3 litters). Atrioventricular canal explant cultures were established using a method described by Camenisch et al (2002), based on a method first published by Bernanke and Markwald

(1982). The entire atrioventricular canal and adjacent myocardium from the embryos were dissected and explanted onto a collagen gel [type I collagen from rat tail (Sigma- 31

Aldrich, St. Louis, MO), 10X Media 199 (Gibco, Grand Island, NY), and sodium bicarbonate (Sigma-Aldrich, St. Louis, MO)] that had been hydrated for at least 30 minutes [OPTI-MEM media (Gibco, Grand Island, NY) supplemented with 1% FBS

(Gemini Bio-Products, Woodland, CA), 0.01% insulin-transferrin-selenium (ITS; Gibco,

Grand Island, NY), 100 U/mL penicillin (Gibco, Grand Island, NY), and 100 ug/mL streptomycin (Gibco, Grand Island, NY)]. The explants were allowed to attach for 12 hours at 37° C and 5% C02, at which point they were bathed in IX Media 199 (Gibco,

Grand Island, NY) supplemented with 1% FBS, 0.01% ITS, 100 U/mL penicillin and 100

(xg/mL streptomycin. The explants were cultured at 37° C and 5% CO2 for 24 and 48 hours, as indicated, and examined and scored for mesenchymal cell invasion and photographed using an inverted Leica DM IL microscope with DC camera attachment

(Leica, Heerbrugg, Switzerland) (Figure 2.4). Migrating mesenchymal cells were identified by focusing the microscope on the surface of the gel and then gradually adjusting to focus on increasing depths within the gel. Migrating mesenchymal cells were identified and scored according to their characteristic spindle-shaped appearance below the gel surface (Runyan and Markwald, 1983), before the genotype of the respective embryo was known, according to the following rationale: 0-0 invaded mesenchymal cells; 1 - 1-20 invaded mesenchymal cells; 2 - 21-50 invaded mesenchymal cells; 3 — 51-

99 invaded mesenchymal cells; 4 - >100 invaded mesenchymal cells. 32

FIGURE 2.4

fir vivo EMT assay

Heterozygous Mating

Eng*l- Eng+I- I

E9.5 Chimeras

Atrioventricular canals removed (dashed line) and explanfed onto type I hydrated collagen gel

Invasion migration Visualize and quantify migration of mesenchymal cells Ink) collagen gel (EMT)

Figure 2.4. Endothelial-to-mesenchymal transition (EMT) experimental overview. Heterozygous endoglin mice are mated, and at E9.5 chimeric embryos are retrieved, and their atrioventricular canals (AVCs) are dissected. The AVCs of the three possible endoglin genotypes are placed lumen down onto a hydrated type I collagen gel, and mesenchymal cell invasion/migration is quantified. CHAPTER 3

RESULTS

33 34

3.1 Endoglin is not required for endothelial differentiation or sprout formation

in vitro

Previous studies showed that lack of endoglin in developing Eng'1' embryos leads to defects in angiogenesis, both in the yolk sac and the embryo proper (Arthur et al.,

2000; Bourdeau et al., 1999; Li et al., 1999). However, whether this is the primary defect during development that leads to embryonic death or whether it is secondary to heart development defects or support cell recruitment/differentiation defects has yet to be determined. Thus, to assess the role of endoglin in angiogenesis in an in vitro setting, we utilized a model of differentiation in which embryoid bodies (EBs) are formed for each of the endoglin ES cell lines via the hanging drop method, and then transferred to tissue culture-treated 8-well culture slides in order to observe and score endothelial sprouting potential. ECs and their vascular networks were identified by staining with the pan- endothelial marker PECAM-1/CD31.

Eng++, Eng+~, and Eng" EBs were all able to form vascular sprouts and an intricate vascular network in vitro (Figure 3.1, A-F). The EBs and their respective vascular networks were scored by a blinded observer based on the following criteria: I, no angiogenic sprout formation from vasculogenic region; II, fewer than 10 angiogenic sprouts from vasculogenic region; III, greater than 10 angiogenic sprouts from vasculogenic region. When scored, no distinction in sprout forming ability can be made between the three genotypes of ES cells (Figure 3.1G). Thus, endoglin does not appear to mediate the ability of EBs to form vascular structures, arguing that the angiogenic potential of ES cell-derived vascular progenitors, at least in an in vitro setting, is not diminished in the absence of endoglin. 35

FIGURE 3.1

X IX IXX Embryold Body Category

Figure 3.1. Endoglin is not necessary for angiogenic sprouting from embryoid bodies (EBs). (A), (D) Eng+I+ EBs show characteristic angiogenic sprouting and network formation after culture. (B), (E) Eng+/~ EBs can also undergo angiogenic sprouting and network formation after culture. (C), (F) Eng'~ EBs show no deficits in their ability to (Figure 3.1 cont.) form angiogenic sprouts and networks after culture, suggesting that endoglin expression is not necessary for angiogenesis in the absence of circulation. All samples stained with PECAM-1. (A) - (C) 100X magnification; (D) - (F) 200X magnification. (G) When scored by a blinded observer, no observable difference between the EBs generated from the three endoglin ES cell lines were documented in their ability to form angiogenic sprouts and angiogenic networks in vitro (I: no angiogenic sprout formation from vasculogenic region; II: fewer than 10 angiogenic sprouts forming from vasculogenic region; III: greater than 10 angiogenic sprouts forming from vasculogenic region). Error bars are S.E.M. for each group. 36

3.2 Endoglin deficient cells contribute to angiogenesis in vivo

In order to overcome the embryonic lethal phenotype that endoglin deletion entails, and to study the ability of Eng"1" cells to contribute to cardiovascular lineages in an in vivo setting, we used chimeric mice. Analysis of mosaic, or chimeric embryos is one of the most powerful scientific experimental tools to gain insights into complex developmental processes. A chimera is an organism composed of cells of more than one genotype (usually WT and mutant). Analysis of the behaviour of the mutant cells compared to the WT cells provides information about the normal function of the mutated gene. Chimera analysis has been used extensively in the nematode worm, fruit flies, zebrafish, chicks, and mice to dissect a number of developmental processes, including cranial neural tube morphogenesis (Chen and Behringer, 1995), eye and nasal development (Quinn et al., 1996), vasculogenesis and hematopoiesis (Shalaby et al.,

1997), and in cardiovascular development (Puri et al., 1999). Since the null allele of endoglin was produced by insertion of the LacZ gene into the first exon (Bourdeau et al.,

1999), both Eng+/" and Eng''' donor cells are stably labeled with P-galactosidase, thus allowing them to be distinguished from host tissue following X-gal staining (cells will stain a bright blue colour). Since Eng+I' embryos develop normally (Bourdeau et al.,

1999), comparing the X-gal staining pattern of Eng+I' and Eng'1' chimeric embryos gives us the ability to assess the developmental potential of endoglin deficient cells in a partially WT environment. Use of chimeras also allows us to circumvent the embryonic lethal phenotype of Eng ' embryos, due to the fact that the WT cells can make up for cellular deficiencies conferred by the mutant donor cells. 37

Several litters of endoglin mutant and control chimeras were analyzed at embryonic day (E)10.5 by X-gal staining (Table 3.1). E10.5 embryos that were generated from the aggregation of wild-type and Eng+I~ ES cells were developmentally normal and displayed a varying degree of donor cell contribution to tissues (Figure 3.2, A, E, I, M) whereas El0.5 embryos that were created from the aggregation of wild-type and Eng''

ES cells were of one of three phenotypes: (I) mosaicism of <20% donor cells (Figure

3.2, B, F, J, N); (II) mosaicism of 20-50% donor cells (Figure 3.2, C, G, K, O); (III) mosaicism of >50% donor cells (Figure 3.2, D, H, L, P). Type III embryos recapitulated the previously reported endoglin knock-out (KO) phenotype (Bourdeau et al., 1999), displaying severe growth delay, improper looping of the heart, and pericardial edema.

However, type I and type II embryos clearly show that endoglin deficient cells have the ability to contribute to angiogenic sprouting in various tissues in the embryo and the fetal component of the placenta. Moreover, Eng+/~ and Eng''' donor cells have the ability to undergo intial angiogenic sprout formation in both the brain and intersomitic region

(arrows in Figure 3.2E, I, G, K). These results corroborate our in vitro EB assay, in which the angiogenic potential of Eng'' ES cells appears to be equivalent to Eng+/~ counterparts.

3.3 Determination of the involvement of endoglin in smooth muscle cell

differentiation in vitro

A previous study reported that loss of endoglin leads to impaired smooth muscle cell (SMC) development during embryogenesis (Li et al., 1999). The authors suggested that this was the primary defect seen in endoglin knockout embryos, as loss of SMC development leads to impaired angiogenesis in the embryo, and could thus account for 38

Table 3.1. Embryos retrieved for all E10.5 chimera dissections. Amount of mutant donor cell contribution to Eng'~ embryos was determined by real-time PCR. Type I = 1-19% chimerism, type II = 20-50% chimerism, type III = 51-100% chimerism.

Embryo Genotype C0 Value (LacZ) Co Value (Tie2) % Contribution Classification 1-32 Eng+/- N/A N/A N/A N/A 33 Undetectable 2.51 <1 Type I 34 Undetectable 9.22 <1 Type I 35 3.00 11.39 26 Type II 36 Undetectable 9.04 <1 Type I 37 Undetectable 4.56 <1 Type I 38 5.57 5.92 94 Type III 39 3.33 5.81 57 Type III 40 5.67 10.84 52 Type III 41 2.76 4.40 63 Type III 42 Undetectable 9.79 <1 Type I 43 Undetectable 7.38 <1 Type I 44 Undetectable 3.02 <1 Type I 45 4.57 8.39 55 Type III 46 Undetectable 8.38 <1 Type I 47 Undetectable 14.19 <1 Type I 48 Undetectable 5.64 <1 Type I 49 Undetectable 4.01 <1 Type I 50 Eng-/- 9.86 15.30 64 Type III 51 Undetectable 14.07 <1 Type I 52 Undetectable 23.59 <1 Type I 53 9.11 20.66 44 Type II 54 Undetectable 7.85 <1 Type I 55 Undetectable 10.10 <1 Type I 56 Undetectable 11.94 <1 Type I 57 Undetectable 11.53 <1 Type I 58 Undetectable 47.62 <1 Type I 59 Undetectable 47.20 <1 Type I 60 45.41 84.21 54 Type III 61 Undetectable 56.33 <1 Type I 62 Undetectable 48.69 <1 Type I 63 Undetectable 58.61 <1 Type I 64 13.55 65.46 21 Type II 65 Undetectable 42.45 <1 Type I 66 32.42 40.62 80 Type III 67 Undetectable 93.63 <1 Type I 68 Undetectable 21.88 <1 Type I 39

FIGURE 3.2

Figure 3.2. Endoglin heterozygous and mutant cells are able to contribute to angiogenic sprouting in the developing chimeric embryo at El0.5. Chimeric mice created from the aggregation of wild-type (WT) and Eng ES cells develop normally (A) and display heterozygote donor cell contribution to angiogenic sprouting in the head region (E), intersomitic region (I), and the placenta (M). Chimeric mice created from the aggregation of WT and Eng'1' ES cells display one of three phenotypes. Type I embryos (n=25) contained less than 20% mutant donor cell contribution and recapitulated the WT phenotype in that they developed normally (B), and had little to no donor cell contribution to angiogenic sprtouting in the head (F) or intersomitic region (J), and 40

(Figure 3.2 cont.) varying contribution to the placenta (N). Type II embryos (n=3) contained 20-50% mutant donor cell contribution and they developed normally (C), and displayed mutant donor cell contribution to angiogenic sprouting in the head (G), intersomitic region (K), and placenta (O). Type III embryos (n=8) contained greater than 50% mutant donor cell contribution and recapitulated the endoglin knock-out phenotype in that they were developmentally delayed (D), and had a high amount of mutant donor cell contribution to the whole body which lead to poor growth of the head (H), and a developmentally delayed heart and pericardial edema (L). The placentas of type III embryos were normal and showed a high amount of mutant donor cell contribution (P). All embryos are stained for P-galactosidase expression with X-gal. Arrows indicate donor cell contribution to the indicated tissues. Scale bars are 1mm. 41 both the angiogenic defects and heart development defects seen in these embryos. In order to determine whether or not endoglin deficient ES cells can form SMCs in vitro, we have employed the OP9-coculture system as described in materials and methods. Briefly, after 5 days of culture on an OP9 stromal cell layer, each of the three endoglin ES cell lines were sorted for Flk-1+ (VEGFR2+) mesodermal progenitor cells via fluorescence activated cell sorting (FACS). These Flk-1+ cells were re-plated onto a fresh OP9 stromal cell layer, where they were cultured for a further 4 days. After the final culture period, the cells were stained with an antibody against a-smooth muscle actin (a-SMA) to detect if the Flk-1+ progenitor cells have the ability to form SMCs. As can be seen in Figure 3.3A,

Eng+,+ ES cells are able to differentiate into SMCs to the same degree as Eng+/~ (Figure

3.3B) and Eng'1' (Figure 3.3C) ES cells in vitro using the OP9-coculture system. To confirm that the staining was not due to background staining of the underlying OP9 stromal cells, a plate containing just these cells was also stained, and was found to not express SMA (Figure 3.3D). Thus, endoglin expression is not necessary for differentiation of progenitor cells into SMCs in an in vitro culture system, where the influence of circulatory factors has been removed.

3.4 The role of endoglin in endocardial cushion formation in the developing

embryonic heart

Endoglin mutant embryos have previously been shown to have defects in heart development; in particular, rightward looping of the linear heart tube does not occur due to defective endocardial cushion formation in the atrioventricular canal (Bourdeau et al.,

1999). However, it has not previously been established whether or not this heart defect is 42

FIGURE 3.3

Figure 3.3. Endoglin mutant cells can differentiate into smooth muscle cells (SMCs) in vitro. Flk-1+ progenitor cells were sorted from ES cell-OP9 coculture samples for each of the three endoglin ES cell genotypes, cultured on fresh OP9 stromal cells, and stained for a-smooth muscle actin (a-SMA). Eng+/+ ES cells were able to differentiate into SMCs (A), as were the Eng+/~ ES cells (B) and the Eng~'~ ES cells (C). OP9 stromal cells were also stained with the antibody to a-SMA (D) to show that they do not express this marker. All panels are shown at a magnification of 200X. 43 the primary cause of circulatory failure in these embryos, or whether it is secondary to circulatory deficiencies caused by loss of angiogenic potential or mural cell differentiation and recruitment. Since our in vitro assays and in vivo chimera data suggest that neither angiogenesis nor SMC differentiation is defective in these animals, we further pursued the idea that heart development may be the primary developmental roadblock seen in Eng~'~ embryos.

We analyzed both sagittal and transverse sections through the atrioventricular canals (AVCs) of the hearts of these animals. Chimeras obtained from Eng+I~ ES cells show that these heterozygous donor cells can contribute quite readily to the endocardial cushions of the AVC in the developing hearts of these embryos (Figure 3.4E, arrow), with no developmental defects being conferred to the animal (Figure 3.4A). Chimeras obtained from Eng" ES cells, on the other hand, display a wide range of heart phenotypes: type III (n=8) embryos show characteristic signs of growth retardation and heart looping defects (Figure 3.4B), and transverse sections through the AVCs of their hearts show that the endocardial cushions have failed to form. In all the type III embryos, the lack of cushion formation was associated with a high amount of mutant donor cell contribution (Figure 3.4F, arrowhead). In contrast, type I embryos (n=25) are developmentally normal (Figure 3.4C), and sagittal sections through the AVCs of their hearts show that very few mutant donor cells have contributed to the endocardial cushions or the surrounding heart tissue (Figure 3.4G, arrow). Finally, type II embryos

(n=3) are developmentally normal (Figure 3.4D), and sagittal sections through the AVCs of their hearts show that while X-gal+ donor cells can contribute to the endocardial layer 44

FIGURE 3.4

Figure 3.4. Endoglin mutant cells cannot contribute to the endocardial cushions of the developing embryonic heart. Chimeric mice obtained from the aggregation of WT and Eng+/~ ES cells are developmentally normal (A), and transverse sections through the atrioventricular canal (AVC) of their hearts show heterozygous donor cell contribution to the endocardial cushions (E, arrow). Type III chimeric embryos obtained from the aggregation of WT and Eng'' ES cells display characteristic growth delay, linear heart tube, and pericardial edema (B), and transverse sections through the AVCs of these hearts show that the endocardial cushions have not formed properly (F, arrowhead), due to the high amount of mutant donor cell contribution. Type I chimeric embryos obtained from the aggregation of WT and Eng'1' ES cells are developmentally normal (C), and sagittal (Figure 3.4 cont.) sections through the AVCs of their hearts show that there is little to no mutant donor cell contribution to the heart or the endocardial cushions (G, arrow). Type II chimeric embryos obtained from the aggregation of WT and Eng'1' ES cells are developmentally normal (D), and sagittal sections through the AVCs of their hearts show that while mutant donor cells can contribute to the overlying of the AVC, there is no mutant donor cell contribution to the endocardial cushions themselves (H, arrow). All embryos are stained with X-Gal for (3-galactosidase expression. Scale bars are lmm. 45

of their AVCs, no mutant derived cells are present in the endocardial cushions themselves

(Figure 3.4H, arrow). We infer from these findings that, while endoglin-deficient cells

can contribute to the endothelial cell lineage as well as participate in early patterning of

the vascular tree, these cells are not tolerated in the developing atrioventricular canal.

3.5 Endoglin mutant atrioventricular canals cannot efficiently undergo EMT

The lack of contribution of Eng-/- cells to the atrioventricular region in chimeric

embryos suggests an essential role for endoglin in the process of EMT, which is

necessary for the formation of the endocardial cushions. If the cells of the endocardium

do not undergo EMT, invade the underlying extracellular matrix, and differentiate into

mesenchymal cells, then the endocardial cushions do not form and the heart cannot

undergo proper rightward looping to form the mature four-chambered heart. In order to

test the ability of the embryonic hearts from the three endoglin genotypes to undergo this

process of EMT, we have used an in vitro EMT assay, which was first described by

Bernanke & Markwald (1982). In this assay, the AVCs are dissected from the hearts of

the embryos and placed face down on a hydrated collagen I lattice. If EMT can occur, the

endocardial cells from the AVCs should spread out across the top of the gel, and then

differentiate into mesenchymal cells and invade the gel. By quantitatively determining

the ability of the Eng+I+, Eng"1', and Eng'1' embryonic hearts to undergo the process of

EMT in vitro, we can determine whether interference with EMT is the underlying defect

in the failure of endoglin-null hearts to properly form the endocardial cushions.

In our studies, at both 24 and 48 hours, Eng+/+ embryonic hearts had a high degree

of mesenchymal cell invasion (Figure 3.5A, D), as did the Eng+I' embryonic hearts 46

(Figure 3.5B, E). The Eng" hearts, however, had a reduced number of invading mesenchymal cells at 24 hours of culture (Figure 3.5C), but by 48 hours of culture some explants displayed a comparable degree of invasion to control explants (Figure 3.5F).

When quantified (Figure 3.5G), this trend becomes much clearer. These results are consistent with recently published data in the chick heart, in which Mercado-Pimentel et al. (2007) reported that siRNA knock-down of endoglin expression leads to significantly reduced EMT at 18-24 hours post-explantation. As well, Sorensen et al. (2003) have previously reported that no significant difference in EMT exists between Eng+/+ and

Eng" hearts at 48-72 hours post-explantation. These results further corroborate our results here, and point to an early blockage in EMT in the AVCs of Eng'' mice. Taken together with in vitro ES cell differentiation assays, we conclude that it is the heart development defect, not defects in angiogenesis or smooth muscle cell differentiation, that is the primary developmental deficiency conferred by endoglin loss of function. 47

Figure 3.5

Score

Q«notyp«: Eng*t* E/IQ+/- En^f> Eno*/+ Eno+/- EnoJ-

Figure 3.5. Endoglin mutant AVC explants show reduced mesenchymal cell transformation and migration at 24 hours. All images of collagen explant cultures were taken with a plane of focus just beneath the explant itself, to show invading mesenchymal cells. A control Eng+/+ explant shows a typical distribution of mesenchymal cell invasion (Figure 3.5 cont.) at both 24 hours (A) and 48 hours (D). Heterozygote Eng+I~ explants show a similar distribution of mesenchymal cell invasion at both 24 hours (B) and 48 hours (E) to the control explants. Mutant Eng'1' explants show a reduced distribution of mesenchymal cell invasion at 24 hours (C) compared to control and heterozygous explants, but not at 48 hours (F). Quantification of mesenchymal cell invasion (G) shows that Eng'1' AVCs have a reduced amount of mesenchymal cell invasion at 24 hours of 48

(Figure 3.5 cont.) culture. Scoring: 0 = 0 cells invaded; 1 = 1-20 cells invaded; 2 = 21-50 cells invaded; 3 = 51-99 cells invaded; 4 = >100 cells invaded. The dark mass in each picture is the myocardial portion of the explant. Arrows mark selected invading mesenchymal cells in each micrograph. CHAPTER 4

DISCUSSION AND CONCLUDING REMARKS

49 50

4.1 Discussion

Endoglin and the endothelial cell lineage

In this study, we sought to determine the extent to which endoglin expression is necessary for early angiogenesis in a developmental setting. Previous studies by three groups demonstrated that loss of endoglin expression affects angiogenic processes in both the yolk sac and embryo proper during early development (Arthur et al., 2000; Bourdeau et al., 1999; Li et al., 1999). Although endoglin null embryos clearly display defects in angiogenesis, it has not been demonstrated that defective angiogenesis is the primary defect in these embryos. Impaired blood vessel maturation may be secondary to any number of other developmental defects during embryogenesis. Indeed, previous studies have already documented that impaired heart development and contractility directly affects angiogenesis (Huang et al., 2003; Lucitti et al., 2007).

Angiogenesis was studied in both an in vitro and an in vivo system, in order to delineate whether or not loss of endoglin affected EC specification and/or the development of the vasculature. EBs have been used extensively in the literature to study angiogenesis because they faithfully mimic the early stages of vascular development

(Magnusson et al., 2004; Vittet et al., 1996), as have chimeras, due to the range of insight they provide into developmental processes (Chen and Behringer, 1995; Puri et al., 1999;

Quinn et al., 1996; Shalaby et al., 1997). The cultured EBs, which are grown in a system that bypasses circulation influences, are able to form vascular structures independent of the presence of endoglin (compare Figure 3.1 A and C). Similarly, even under the stringent competitive situation that exists in chimeras, differentiation of donor ES cells to

ECs that contribute to many tissues and organs of the body was observed that was 51 independent of the presence of endoglin (Figure 3.2). The fact that endoglin null ES cells were able to contribute to angiogenically sprouting ECs throughout the body of the embryo suggests that endoglin expression is not required in these tissues in the developing embryo. These in vivo experimental results strongly support our in vitro results, and suggest that Eng'ES cells and mesodermal derivatives are normal with respect to their angiogenic potential. These results show that endoglin is not essential for

EC specification during the development of an embryo, as the endoglin deficient ES cells were able to contribute to angiogenic sprout formation in vitro and early blood vessel formation in vivo to the same extent as ES cells that did express endoglin. Other members of the lab have also similarly shown that endoglin null ES cells can contribute to blood vessel formation in the yolk sac of the developing embryo of chimeric mice, as well as to sprouting angiogenesis in the neural tube (data not shown), further supporting the results presented here. Therefore, we conclude that the angiogenic defects that are seen in Eng'1' embryos and have been reported in the literature are secondary to other developmental defects.

Endoglin and the smooth muscle cell lineage

During embryonic development, interactions between the ECs of newly forming blood vessels and mural cells (pericytes and vascular smooth muscle) are essential for proper vascular remodeling and maintenance (Carmeliet et al., 1996b; Hirschi et al.,

1998; Lindahl et al., 1997). Recent work from Gordon Keller's group has shown that

ECs, vascular SMCs, and cardiomyocytes arise from a multipotent Flk-1+ progenitor cell population during development (Kattman et al., 2006). It has also previously been shown 52 that supporting mural cells can also be derived from mesoderm, neural crest, or epicardial cells which then migrate through the body to help form the blood vessel wall (Jiang et al.,

2000; Mikawa and Gourdie, 1996; Topouzis and Majesky, 1996), suggesting that there are several sources of support cell progenitors in an developing mammalian embryo. Li et al. (1999) suggested that Eng'1' embryos have a defect in vascular SMC differentiation and recruitment to newly forming blood vessel walls, and that this defect is likely what leads to their demise at E9.5 of gestation. This conclusion was based on immunohistochemical staining of E9.5 embryos with an antibody to a-SMA. However, the authors did not document any heart defects in these mice, even though this is the developmental timepoint at which endocardial cushion formation and rightward looping of the linear heart tube is occurring. A recent report demonstrated that endoglin is expressed in a subset of cells that are a common precursor to SMCs, ECs, and hematopoietic cells (Ema et al., 2006), which may support the conclusions of Li et al.

(1999) by suggesting that loss of endoglin may negatively affect proper SMC development. As such, to determine whether endoglin specifies SMC differentiation potential, we utilized the OP9-coculture system to derive Flk-1+ cells from ES cells, a system that was first described by Shin-Ichi Nishikawa's group (Hirashima et al., 1999;

Nishikawa et al., 1998).

As shown in Figure 3.3, WT ES cells differentiated in culture stain for a-SMA, and thus are able to form SMCs in vitro. Likewise, Eng1' ES cells and Eng1' ES cells also produce SMA+ cells after differentiation, suggesting that there is no difference in the ability of the three different ES cell lines to form SMCs, at least in an in vitro culture system. These results suggest that endoglin is not required for SMC differentiation from 53

Flk-1+ mesodermal cells, and a differentiation block in this lineage is not the primary defect seen in Eng'1' embryos. Nevertheless, our results do not rule out that functional association between ECs and SMCs is impaired in Eng1' embryos. By histology we can see in chimeric embryos that vascular SMCs have invested the dorsal as well as the

Eng " derived ECs (data not shown), suggesting that when developmental processes are rescued, SMCs are recruited normally to ECs. However, since we could not determine the donor vs. host origin of the SMCs in the chimeric embryos, it is still possible that Eng'1'

SMCs are outcompeted in this context. Further studies on the origin of the SMCs surrounding the blood vessels in the chimeric embryos are necessary to experimentally address this issue. Interestingly, recent work by Seki et al. (2006) has shown that ALK5_/" mice exhibit a defect in the formation of SMC layers around blood vessels, even though the blood vessel lumens form properly. The fact that ALK5 deficiency leads to embryonic lethality argues that angiogenesis does depend upon secondary association with SMCs, and points to a possible role for endoglin in mediating ALK5 signalling during development. Although we have shown that loss of endoglin is not required for differentiation of SMCs from progenitor cells, future experiments will focus on elucidation of the involvement of endoglin in SMC and EC association during angiogenesis. However, we conclude that the reported defect in SMC differentiation of

Eng'1' embryos is secondary to an earlier developmental defect.

Endoglin and endocardial cushion formation

Structural heart defects are the highest reported congenital deficiencies seen in humans, with flaws in the cardiac valves being the most common subtype (Loffredo, 54

2000). The most important step in cardiac development is rightward looping of the linear heart tube: this is necessary for proper orientation of the ventricles with respect to the atria, and is necessary for proper alignment of the atria and ventricles with the circulation.

The endocardial cushions form in the AVC and OFT at a specific time point during development when inductive signals from the myocardium activate ECs in the endocardium to undergo EMT; these cells disengage their cell-to-cell contacts, extend filopodia, and invade the underlying extracellular matrix (Markwald et al., 1979). These invading mesenchymal cells within the newly formed endocardial cushions contribute to the formation of the mitral and tricuspid valves and complete atrial and ventricular septation (Schroeder et al, 2003; Srivastava and Olson, 2000). If EMT does not occur, and mesenchymal cells do not invade the underlying extracellular matrix to form the endocardial cushions, cardiac function is impaired leading to developmental arrest.

While it has been previously reported that Eng'~ mutant embryos have cardiac defects during development (Arthur et al., 2000; Bourdeau et al., 1999), it has never been definitively shown whether or not this is the primary defect causing embryonic death in these animals. In order to address this question both in vivo and in vitro, we analyzed the donor versus host contribution in mutant and control chimeric hearts at E9.5-10.5, and we performed an EMT assay on explanted mutant and control hearts. As expected, developmentally normal chimeric mice created from the aggregation of Eng'~ ES cells and WT ES cells (Figure 3.4A) showed no cardiac defects; transverse sections through the hearts of these embryos showed normal trabeculation of the myocardium (see left- hand side of Figure 3.4E), and proper formation of the endocardial cushions (arrow in

Figure 3.4E). Moreover, the high degree of X-gal staining of these structures indicated 55 that Eng ' cells were able to contribute to the formation of the cushions quite readily.

Similarly, the EMT assay showed that explanted hearts from EngH~ embryos were readily able to undergo EMT and invade the collagen gel at both 24 and 48 hours post- explantation (Figure 3.5B, E, G). In contrast, type I chimeric embryos (Figure 3.4C) have developmentally normal hearts, and sagittal sections through the hearts of these embryos reveal that there is little to no Eng'1' cell contribution to the endocardial cushions of the hearts of these animals, and that WT cells have predominantly formed these hearts (arrow in Figure 3.4G shows no blue staining). Type III chimeric embryos (Figure 3.4B) recapitulate the previously reported Eng'1' (Bourdeau et al., 1999) phenotype and have developmentally delayed hearts (notice the linear heart tube in Figure 3.4B). Transverse sections through the hearts of these embryos reveal that the endocardial cushions have failed to form (arrowhead in Figure 3.4F reveals no endocardial cushion; compare Figure

3.4E to Figure 3.4F), as reported previously for Eng" mutant mice (Bourdeau et al.,

1999). Type II embryos (Figure 3.4D), while developmentally normal, demonstrated some Eng'' cell contribution to the endocardium when sagittal sections are taken through the hearts of these animals. Importantly, however, the endoglin deficient cells were never observed to contribute to the underlying endocardial cushions (arrow in Figure 3.4H).

The EMT assay results also support this data, showing that Eng'1' embryonic hearts do not undergo EMT at the 24-hour time point to the same extent as control hearts (Figure 3.5C,

F, G). Though we observed mutant donor-derived X-gal+ endocardial cells in the overlying endocardium of mutant chimeric hearts, the mesenchymal cells that have invaded the underlying extracellular matrix are X-gal", indicative of WT host contribution. Yet our in vitro EMT data shows that although the endoglin mutant hearts 56 are delayed in their ability to undergo EMT, they are still able to undergo this process given enough time. This finding suggests that endocardial cells lacking endoglin do not contribute to the EMT process, and in vivo fail to form the endocardial cushions, but in vitro these cells are delayed.

Very few of the explanted AVCs from Eng~'~ hearts have the ability to undergo

EMT at 24 hours, suggesting that loss of endoglin expression is directly affecting this process. Not only do these results corroborate our earlier chimeric data which showed that Eng' ' mutant cells could not undergo EMT and invade the extracellular matrix of the forming endocardial cushions, but these results are also supported by a recent report by

Mercado-Pimentel et al. (2007) in which siRNA knockdown of endoglin in chick hearts led to a significant decrease in EMT at 18-24 hours. Similarly, it has been previously shown that endoglin is highly expressed in the endocardial cushions of developing human hearts when these structures are being formed (Qu et al., 1998), suggesting that it plays a role in endocardial cushion development. However, it must be noted that by 48 hours in our studies EMT in Eng'1' hearts is evident, since some of the Eng'1' hearts are able to undergo EMT to the same extent as their Eng+/+ and Eng*1' counterparts. This may be due to the in vitro nature of this assay, in that the culture conditions may be providing extracellular cues to the cultured hearts that could make up for loss of endoglin expression after 48 hours. Alternatively, a small number of Eng'' endocardial cells that have the ability to undergo EMT expand and proliferate quickly between the 24 and 48 hour time point, thus making up for the initial EMT deficiency. We suggest that this delay in EMT is not tolerated in developing mammalian embryos and ultimately leads to embryonic lethality. 57

There are many possible reasons as to why the endoglin mutant endocardial cells are impaired/delayed in their ability to undergo EMT in the mutant chimeric hearts and in the in vitro EMT collagen assay. We cannot attribute this defect to poor migration ability in endoglin deficient cells, since we see that by 48 hours post-explantation some endoglin deficient mesenchymal cells migrate into the gel to an extent comparable to control hearts. It is possible that cell-cell adhesive properties are abrogated in endoglin-null endocardial cells, or that loss of endoglin upregulates the expression of extracellular matrix components. Recently, Jerkic et al. (2006) demonstrated that murine Eng+/- ECs had impaired migratory properties and increased collagen production compared to WT cells in vitro. Though these studies were not performed on endoglin null ECs, these results do suggest that loss of endoglin expression may indeed increase cell-cell adhesion, decrease migration, and/or induce extracellular matrix production of ECs. Further experiments to determine which signalling pathways are interrupted in the absence of endoglin is a priority of our lab.

Based on our in vivo chimeric data and in vitro EMT assay data, we propose a model to explain the defects in endoglin null mutant embryonic hearts. In a normal environment, all of the cells in the endocardium are able to respond to cues from the underlying myocardium, and undergo EMT to form the endocardial cushions of the developing heart (Figure 4.1 A). In the chimeric mice we generated from the aggregation of £«g+/" ES cells and WT ES cells (Figure 4. IB), Eng1' cells are able to undergo EMT 58

FIGURE 4.1

Figure 4.1. Eng" cells cannot undergo EMT to form the endocardial cushions of the AVC. In a normal, WT heart, endocardial cells receive signals from the myocardium, causing them to undergo EMT and invade the cardiac jelly, forming the endocardial cushions of the developing heart in the AVC (A). When chimeras are generated by aggregating WT ES cells with Eng'1' ES cells, this process of EMT is not disturbed, and the heterozygous endoglin donor cells are able to contribute to the formation of the endocardial cushions (B). However, chimeras are generated by aggregating WT ES cells with Eng'1' ES cells, we get two different outcomes. If there are very few endoglin-null donor cells contributing to the developing heart, even though they cannot undergo EMT, WT cells can overcome this deficiency and form the endocardial cushions (C). However, if a high amount of endoglin-null donor cells contribute to the developing heart, the WT cells cannot overcome this block in EMT, and thus the endocardial cushions fail to form (D). 59 and can contribute to the developing endocardial cushions in the AVC during heart development. However, in chimeric mice generated from the aggregation of Eng'1' ES cells and WT ES cells, Eng'1' donor cells are not able to undergo this process of EMT.

Thus, if there are only a small number of Eng1' donor cells contributing to the heart, the wild-type cells can overcome this deficiency of the mutant cells, and can recover proper formation of the endocardial cushions (Figure 4.1C). If, however, there are a large number of Eng'1' donor cells contributing to the heart, the wild-type cells cannot overcome this EMT delay, and the endocardial cushions fail to form (Figure 4.ID). Thus, it appears that endoglin expression is necessary for this process of EMT to occur properly in the heart, and that lack of endoglin expression leads to incomplete heart development and embryonic death.

There are also several potential mediators of endoglin function in the heart. As stated before, endoglin is able to bind TGFpl, TGFp3, BMP2, BMP7, BMP9, and

BMP 10. Previous research has primarily focused on the role that the TGFps play in the function of endoglin in vivo and in vitro. Since neither TGFpi"A nor TGFP37" mice display a heart developmental defect (Dickson et al., 1995; Taya et al., 1999), we hypothesize that endoglin's role in cardiac development may also be mediated through the BMP ligand family. A lot less is known about endoglin's role in BMP signalling, but several studies have demonstrated the essential role of the BMPs in heart development, and this may be the area where endoglin function is found to be required. For instance,

BMP2 is expressed in the myocardium of the AVC (Abdelwahid et al., 2001; Kaartinen et al., 2004; Watanabe et al., 2006), and both traditional and conditional knockout studies of the BMP2 gene in the mouse cardiac lineage have shown that it is important for 60 endocardial cushion formation, including the induction of EMT, extracellular matrix deposition, and the establishment of AVC specificity between the common atrium and ventricle (Ma et al., 2005; Zhang and Bradley, 1996). The type I receptor for BMP2,

ALK3, has also been shown to be indispensable for endocardial cushion formation in conditional KO studies (Gaussin et al., 2002; Park et al., 2006). Thus, it is interesting to speculate on the role that endoglin may be playing in the BMP2-ALK3 signalling complex. It is possible that loss of endoglin downregulates the binding of BMP2 to

ALK3, leading to impaired or reduced downstream signalling through SMAD1/5/8, and thus delaying endocardial cushion formation. Furthermore, BMP9 has recently been shown to signal through the ALK1-SMAD1/5/8 pathway, and loss of BMP9 in ECs leads to a blockage in VEGF-induced angiogenesis (Scharpfenecker et al., 2007). While functional studies have yet to show whether BMP9 is expressed in heart tissue during endocardial cushion formation, it is tempting to speculate that reduced BMP9 signalling in combination with reduced BMP2 signalling in the absence of endoglin may lead to impaired or reduced downstream SMAD1/5/8 activation and transcription of target genes, such as Id-1 and SMAD6, which are necessary for cellular migration and ECM breakdown (Ota et al., 2002). Determination of the functional role of endoglin in both

BMP2 and BMP9 signalling will be crucial to delineating whether reduced signalling via one or both of these ligands is the cause of the delay in EMT in endoglin null endocardial cells. Immunohistological analysis of murine embryos, as well as RT-PCR and in situ hybridization experiments will be critical in determining the temporal and spatial expression pattern of BMP9 in murine embryos, to first determine if it is expressed at a similar time and place to endoglin. While BMP2 has been shown to bind endoglin in in 61 vitro biochemical assays (Barbara et al., 1999), whether this interaction occurs in vivo has not been functionally demonstrated. It is possible that endoglin/BMP2 double heterozygous mice (Eng+/~/BMP2+/~) would display a similar heart defect to either Eng~'~ or BMP2~'~ mice. Creation of this mouse line would be helpful in determining the role of endoglin in BMP2 signalling, and how exactly this affects heart development.

4.2 Concluding Remarks

The characterization of the functional role of endoglin in early mammalian development shows that endoglin expression is indispensable for proper looping and development of the mammalian heart. In this study I demonstrated that defective angiogenesis is not the primary defect affecting Eng'~ embryos, as both an in vitro EB assay and in vivo chimeric mouse assay showed that endoglin-null ES cells could form angiogenic structures in culture and contribute to angiogenic sprouting in developing mice. SMC differentiation was shown to be intact in endoglin-null ES cells, suggesting that defective SMC differentiation is not the primary defect affecting Eng'1' mice.

Endoglin expression is necessary for proper formation of the endocardial cushions during embryonic development, as its expression is necessary for proper EMT to occur in the developing heart. Future studies will be aimed at determining the exact functions of EMT

(e.g. cell adhesion, cell migration, mesenchymal cell differentiation) that endoglin potentiates. The significance of endoglin in both BMP2-ALK3 and BMP9-ALK1 signalling now needs to be determined. Functional in vivo analyses of the spatial and temporal expression patterns of BMP9 and endoglin, as well as double heterozygous endoglin/BMP2 mouse genetic studies are needed to further tease out the full role of 62 endoglin in TGFP superfamily signalling. This increased knowledge about endoglin will further our understanding of the intricacies of TGF-p superfamily signalling in general, and will hopefully lead to an increased understanding of the role endoglin plays in complex diseases such as HHT, preeclampsia, and cancer. CHAPTER 5

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