University of Southern California Dissertations and Theses

Proliferation and maturation events in second heart field cells during cardiovascular development activated by the Delta like ligand-4 mediated notch signaling

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Proliferation and maturation events in second heart field cells during cardiovascular development activated by the Delta like ligand-4 mediated notch signaling

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Content PROLIFERATION AND MATURATION EVENTS IN SECOND
HEART FIELD CELLS DURING CARDIOVASCULAR
DEVELOPMENT ACTIVATED BY THE DELTA LIKE LIGAND-4
MEDIATED NOTCH SIGNALING

by

Prashan De Zoysa

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
DEVELOPMENT, STEM CELLS, AND REGENERATIVE MEDICINE

August 2021

ii

Acknowledgments

I wouldn’t have been where I am today if not for numerous individuals who helped me
throughout this wonderful journey of graduate work officially known as the PhD in Development,
Stem Cells and Regenerative Medicine.

One of the most important advices I received during my first year of PIBBS (Programs in
Biomedical and Biological Sciences) is that it’s not the field of the PhD that I end up with which
matters, but the environment of the lab I join which would be my home away from home for the
next 5-6 years or so. Going to lab every day, I reminisce how valuable that piece of information
is. For, my mentor, Dr. Ram Kumar Subramanyan is everything I could have expected to be in a
mentor and more. He is supportive, brilliant, encouraging, enthusiastic and also firm at the same
time. He is a terrific role model, not only as a mentor but also as a manager who has taught me
valuable attributes both in and outside the lab. He always pushed me to strive to be the best
version of myself. I believe that working under his supervision for the last few years has
prepared me well for any career in my future.

Similarly, I want to thank my thesis committee members including chair Dr. Ellen Lien and Dr.
Young Kwon Hong and my own PI Dr. Ram Kumar Subramanyan for continually giving me
encouragement, support and constructive criticism in improving my research. A special thanks
also goes to my previous committee chair, Dr. Henry Sucov who was instrumental in mentoring
me and guiding my projects in the right direction and was always willing to give me advice in
making my research projects better.

A lab doesn’t become whole without the people in it. Hence, I want to thank my lab members
whose presence made it enjoyable to come to work every day. Special thanks go to Omar
Toubat, with whom I’ve had numerous discussions about the science behind the experiments
we conduct, techniques and even general camaraderie. Also, the presence of Drayton Harvey
and Jongkyu Choi in the lab has been wonderful both in helping with running experiments as
well as the cheerful enthusiasm they bring to the lab.

Additionally, I want to mention specific individuals from collaborating labs who helped me with
unique techniques. Susana Cavallero with X-gal staining, Hua Shen with Insitu hybridization and
Binyun Ma with Cell culture experiments have all been helpful to me in successfully completing
my projects.
Furthermore, a special thanks goes to my friend Marilena Elani Melas for her invaluable
friendship during our times as graduate students pursing PhDs at USC. Similarly, the building
manager at Norris Ezralow Tower, Alfred Ascencio has always been a great friend and a
supporter as well.

Last but not least, I would like to thank my family. Especially parents (Justin Sepala De Zoysa
and Mangalika Kulasinghe De Zoysa), sister (Madhushani De Zoysa Guruge) and fiancée (Fleur
Lobo). I am immensely thankful for my mother for being a pillar of my strength and always
believing in me even when I wouldn’t believe in myself. My fiancée and life partner, Fleur Lobo
has been another pillar of support who has helped me both personally and professionally to
achieve the best. In fact, I am very fortunate to share many life moments of personal and
professional happinesses and successes with her.

iii

Table of Contents

Acknowledgements ..................................................................................................................... ii
List of Figures ............................................................................................................................ vi
List of Tables ........................................................................................................................... viii
A bstr ac t …… ……… …… ………… ……… ……… … ………… ……… ……… … ………… ……… ……i x

Chapter 1 ............................................................................................................................. 1
Introduction ............................................................................................................................. 1
1.1 Early Heart Development ................................................................................................. 1

1.2 Arch Artery Development ................................................................................................. 1

1.3 Congenital Heart Defects ................................................................................................. 1

1.4 Notch Signaling ................................................................................................................. 2

1.5 Second Heart Field Progenitor Cells (SHF) ................................................................... 3

1.6 Notch signaling ins SHF cells .......................................................................................... 3

Chapter 2 ............................................................................................................................. 5
Methods .................................................................................................................................... 5
2.1 Mice ................................................................................................................................... 5

2.2 Tissue analysis and histology ......................................................................................... 5

iv

2.3 Thoracic Oran and Cell Culture ....................................................................................... 5

2.4 Fgf8 Promoter and Enhancer analysis ........................................................................... 5

2.5 India ink Injection .............................................................................................................. 6

2.6 Human umbilical arterial endothelial cell (HUAEC) tube formation assay .................. 6

Chapter 3 ............................................................................................................................. 7
Dll4 expression in the developing heat and the arch arteries ...................................... 7
3.1 Dll4 is expressed at relevant sites and time-points during OFT formation in the
developing heart ................................................................................................................. 7

3.2 Pharyngeal arch arteries express Dll4 at appropriate time points during their re-
organization ......................................................................................................................... 8

Chapter 4 ........................................................................................................................... 16
Dll4 mediated notch signaling in SHF cells is required for the proper cardiac
development .......................................................................................................................... 16
4.1 Introduction ..................................................................................................................... 16

4.2 Dll4 expression in SHF is required for appropriate development of SHF- derived
RV and OFT ...................................................................................................................... 16

4.3 Dll4 expression is required for SHF cell proliferation to maintain an adequate
progenitor cell pool ........................................................................................................... 17

4.4 Dll4-mediated Notch signaling regulates Fgf8 expression in SHF ............................ 18

4.5 Genetic synergy between Dll4 mediated Notch and Fgf8 signals in SHF
proliferation ........................................................................................................................ 19

4.6 Discussion ....................................................................................................................... 20

v

Chapter 5 ........................................................................................................................... 38
Haploinsufficiency of Dll4 in Second Heart Field cells is the ideological
mechanism for the cardiac defects seen in Adams-Oliver syndrome ...................... 38
5.1 Introduction ..................................................................................................................... 38

5.2 Haploinsufficiency of Dll4 in SHF disrupts OFT alignment ...................................... 38

5.3 Partial conditional loss of Dll4 expression leads to reduction I SHF progenitor cell
pool .................................................................................................................................... 39

5.4 Discussion ....................................................................................................................... 40

Chapter 6 ........................................................................................................................... 46
Dll4 mediated notch signaling in SHF cells is required for the proper cardiac
development .......................................................................................................................... 46
6.1 Introduction ..................................................................................................................... 46

6.2 Dll4 expression in SHF progenitors is required for the appropriate development of
the aortic arch ................................................................................................................... 46

6.3 Dll4 mediates vessel maturation via the expression of Hey1 ..................................... 48

6.4 Dll4 expression in PAA is required for arterial specification and development of
smooth muscle coat ......................................................................................................... 48

6.5 Discussion ....................................................................................................................... 49

Concluding Remarks ................................................................................................. 57

References........................................................................................................................ 63

vi

List of Figures

Fig 3.1
Dll4 is expressed by SHF progenitor cells and SHF-derived structures in the developing heart ....
................................................................................................................................................. 10

Fig 3.2
Dll4 is expressed at appropriate time-points by SHF progenitor cells and SHF-derived structures
in the developing heart .............................................................................................................. 12

Fig 3.3
Dll4 is expressed by SHF progenitor cell-derived pharyngeal arch artery endothelium as well as
in the dorsal aorta ..................................................................................................................... 14

Fig 4.1
Dll4 expression in SHF is required for appropriate development of SH-derived RV
and OFT.................................................................................................................................... 24

Fig 4.2
Knockout of Dll4 in SHF cells leads to reduced size of SHF-derived RV and OFT .................... 26

Fig 4.3
Knockout of Dll4 in SHF cells leads to DORV with an obligatory VSD by E14.5 ........................ 28

Fig 4.4
Dll4 expression is required for SHF cell proliferation to maintain an adequate progenitor
cell pool..................................................................................................................................... 29

Fig 4.5
Dll4 expression in SHF cells is required for SHF progenitor cell proliferation ............................ 31

Fig 4.6
Dll4 expression in SHF cells is required to maintain expression of key SHF-related proteins .... 32

Fig 4.7
Impact of loss of Dll4 in SHF on expression of SHF-specific molecules .................................... 34

Fig 4.8
Dll4-mediated notch signaling regulates Fgf8 expression in SHF .............................................. 35

Fig 4.9
Dll4-mediated notch signaling regulates Fgf8 expression in SHF .............................................. 37

Fig 5.1
Haploinsufficiency of Dll4 in SHF results in misalignment of OFT.............................................. 42

vii

Fig 5.2
Haploinsufficiency of Dll4 in SHF leads to reduced size of RV and foreshortened OFT ............. 43

Fig 5.3
Dll4 expression is required for SHF cell proliferation to maintain an adequate progenitor cell
pool ........................................................................................................................................... 44

Fig 6.1
Dll4 expression in SHF is required for the maturation of the aortic arch .................................... 52
Fig 6.2
Haploinsufficiency of Dll4 in SHF leads to development defects in the pharyngeal arch arteries
................................................................................................................................................. 53

Fig 6.3
Dll4 expression is required proper Hey1 expression and other downstream changes ............... 54
Fig 6.4
Dll4 expression in SHF is required for arterial identity in pharyngeal arch arteries .................... 56

viii

List of Tables

Table 1
Primer Information .................................................................................................................... 58

Table 2
Antibody, Vector information, Cell Lines and Chemicals ........................................................... 61

ix

Abstract

During cardiogenesis, progenitor cells from the anterior second heart field (SHF) contribute to
the developing right ventricle (RV) and outflow tract (OFT). Animal studies have suggested a
role for notch pathway in OFT maturation and ventricular compaction. Whether notch pathway
plays a role in early cardiac progenitor cell biology remains to be elucidated. Delta like ligand-4
(Dll4), the arterial specific notch ligand, is expressed by murine SHF progenitor cells from E8.5
to E12.5, time points that are crucial in SHF biology. I show that Dll4-mediated notch signaling is
critically required for SHF proliferation such that Dll4 knockout results in proliferation defects of
SHF cells leading to a reduced SHF progenitor cell pool. Reduction in SHF cells available for
incorporation into the developing heart leads to underdeveloped RV and OFT causing a
spectrum of defects that vary depending on the timing and extent of loss of Dll4. Mechanistic
analyses demonstrate that Dll4-mediated notch signaling maintains Fgf8 expression in SHF by
transcriptional regulation at the promoter level. This is further confirmed by in vitro and in vivo
studies involving exogenous supplementation of Fgf8 in ex-vivo organ cultures to rescue the
proliferation defect observed in mutant SHF as well as utilizing combined heterozygous
knockout of Dll4 and Fgf8 in SHF cells to demonstrate genetic synergy between these two
pathways in OFT alignment. Hence, my results establish a novel role for Dll4-mediated notch
signaling in SHF progenitor biology. Dll4-signaling maintains Fgf8 expression and SHF
proliferation to ensure an adequate pool of progenitor cells for incorporation into the developing
OFT. Adams-Oliver syndrome in which cardiac defects account for one fourth of the cases and
in particular OFT alignment defects is characterized by heterozygous loss of function mutation
in Dll4. The mechanism underlying this genotype-phenotype correlation has not yet been
established. With my knowledge of the crucial role Dll4-mediated notch signaling plays on SHF
progenitor cell proliferation, I hypothesized that depletion of SHF progenitor pool of cells due to
partial loss of Dll4 is responsible for the OFT alignment defects seen in Adams-Oliver syndrome
clinically. My murine model provides a molecular mechanism to explain the cardiac defects
observed in Adams-Oliver syndrome and establishes a novel clinical role for Dll4-mediated
notch signaling in SHF progenitor biology. When SHF progenitors mature to assume
endocardial and myocardial cell fates, as mentioned previously Dll4 expression is initially
required for the early development of the heart after which Dll4 expression subsides and taken
over by another notch ligand, Jagged1. A subset of these SHF progenitors also matures to form
the pharyngeal arch artery (PAA) endothelium. Dll4 was originally identified as an arterial
endothelial-specific notch ligand that plays an important role in blood vessel maturation, but its
role in aortic arch maturation has not been studied to-date secondary to the early lethality
observed in Dll4 knockout mice. I show that, unlike in SHF-derived endocardium and
myocardium, Dll4 expression persists in SHF-derived arterial endothelial cells. Dll4 expression
maintains arterial identity in the PAAs and plays a critical role in the maturation and re-
organization of the 4
th
pharyngeal arch artery, in particular. Haploinsufficiency of Dll4 in SHF
leads to highly penetrant aortic arch artery abnormalities, similar to those observed in the clinic,
primarily resulting from aberrant reorganization of bilateral 4
th
pharyngeal arch arteries. Hence, I
show that SHF progenitors that assume an arterial endothelial fate continue to express Dll4 and
the resulting Dll4-mediated notch signaling transitions from an early proliferative to a later
maturation role during aortic arch development. Overall, my work shows the versatility of notch
ligand Dll4 in acting both as a proliferative signal or a maturation signal even within the same
cell type depending on what those cells eventually mature in to.

1

Chapter 1
Introduction

1.1 Early Heart Development

In the developing mouse embryo, the heart fields develop at around E7 as bilateral fields of two
mesodermal cell progenitors in the lateral plate mesoderm. By E7.5, the cranial segment of
these bilateral field fuse in the midline to form the cardiac crescent made up of two different
progenitor cells, namely the First Heart Field (FHF) and Second Heart Field (SHF) (Vincent and
Buckingham, 2010; Luxan et al., 2016; Lin, et al., 2012). The FHF cells continually migrate to
the midline and fuses to form the primitive heart tube, by E8 (Luxan et al., 2016). This tube will
be continually grown and elongated with more SHF cells being added from the arterial and
venous poles. By E8.5, the heart tube will loop, positioning the venous poles dorsally to the
arterial poles containing four distinct segments, atrium, atrioventricular canal, ventricles and
outflow tract (OFT) (Luxan et al., 2016). In this segmented heart, whereas atria are developed
with equal contributions from FHF and SHF cells, Left Ventricle becomes a sole product of FHF
cells while the Right Ventricle (RV) and OFT are exclusively developed from the contributions
from SHF cells (Kelly et al., 2001, Mjaatvedt et al., 2001, Rochais et al., 2009, Waldo et al.,
2001).

1.2 Arch Artery Development

In the developing embryo, the blood exits the heart through a common vessel, the Outflow tract
leading to the aortic sac before being circulated dorsally and laterary through multiple
pharyngeal arch arteries to the twin Dorsal aortae which combine caudally to form a single
dorsal aorta (Hiruma et al., 2002; Priya et al., 2018). Between E9.5 to E14.5, the developing
pharyngeal arch arteries and the dorsal aorta rearrange in a complex manner to form the
mature aortic arch arteries that is seen in the adult heart (Hiruma et al., 2002; Priya et al., 2018).
Between these pharyngeal arch arteries, the first and second degenerate without forming a
structure while the third give rise to the right and left carotid arteries. Right and left fourth
pharyngeal arch arteries contribute to the part of right subclavian artery and aortic arch
respectively. Similarly, right and left sixth pharyngeal arch arteries contribute to the right and left
pulmonary arteries (Hiruma et al., 2002; Priya et al., 2018). The left sixth pharyngeal arch artery
additionally contribute to the ductus arteriosus as well. Finally, the right dorsal aorta give rise to
the part of the right subclavian artery while the left dorsal aorta give rise to the aortic arch and
the descending aorta (Hiruma et al., 2002; Priya et al., 2018).

1.3 Congenital Heart Defects

Congenital Heart Defects (CHD) are the most common type of birth defect in the United States,
occurring n 1% of all infant births and accounting for a third of all major congenital defects
(Simeone et al., 2014; Neeb et al., 2013). Out of these, 25% of the newborns with a CHDs have
a critical CHD (CCHD) requiring surgical intervention before one year of age (Simeone et al.,

2

2014). Due to the elevated number of incidences involving CHDs and the requirement for
surgical intervention, nationally, the associated costs for CHD incidences run in to more than 5
billion annually (Simeone et al., 2014). CHDs come in many shapes and forms. However,
among these, outflow tract anomalies account for 30% of all the CHDs (Neeb et al., 2013).
Specifically, major OFT structural vascular defects includes Persistent Truncus Arteriosus
(PTA) also referred to as Common Arterial Trunk wherein there is only a single undivided OFT
exiting the heart instead of the properly developed Aorta and Pulmonary Artery coming out of
the Left and Right Ventricle (RV) respectively; Double Outlet Right Ventricle (DORV), in which
the properly divided aorta and pulmonary vessels both exit the right ventricle (RV);
Transposition of the Great Arteries (TGA), in which the aorta and pulmonary arteries while fully
septated, come out of the incorrect chamber, namely aorta arising from the RV and the
pulmonary artery arising from the left ventricle (LV) and Overriding Aorta (OA), which is an OFT
alignment defect where the aorta is shifted slightly to the right over a Ventricular Septal Defect
(VSD) in a improperly divided heart receiving mixed blood from both ventricles (Neeb et al.,
2013). Furthermore, Overriding Aorta is usually associated with a combination of heat defects
known as Tetralogy of Fallot (TOF) which also includes Pulmonary stenosis (PS) where the
pulmonary artery is narrowed, Right Ventricular Hypertrophy where the muscular wall of the
right ventricle is abnormally thickened along with a VSD (Neeb et al., 2013).

Similarly, congenital arch abnormalities involve a wide spectrum of malformations involving the
disordered embryogenesis of branchial arch arteries affecting 1-2% of the general population
(Priya et al., 2018). However, unlike OFT abnormalities, these arch artery defects could be
asymptotic only being diagnosed when the patient is imaged for a different clinical reason or
symptomatic where the patient displays clinical manifestations linked to the underlying defect
such as shortness of breath and difficulty in swallowing (Priya et al., 2018). Some of the most
common arch artery malformations are Hypoplastic aortic arch which is defined as the
narrowing of the aorta that can happen at any segment of it, be it proximal, distal or the isthmus
as well as Interrupted Aortic Arch, where the aorta doesn’t form completely leaving a
discontinuation in a portion of the aortic arch (Priya et al., 2018). Depending on where the
discontinuation of the aortic arch happens, the interrupted artic arch is classified as Type A
(disruption after the left subclavian), Type B (disruption between the left carotid and left
subclavian) and Type C (disruption before the left carotid, between the innominate artery and
left subclavian) (Priya et al., 2018). Aberrant formation of the subclavian arteries is another form
of arch malformation where the left and right subclavian arteries do not originate at their usual
locations in the aortic arch (Priya et al., 2018). On the other hand, the arch defect right sided
aortic arch forms when the aortic arch courses to the right of the trachea where as normally it
courses to the left (Priya et al., 2018). Finally, another complicated arch abnormality, a vascular
ring arises when a right sided aortic arch and an aberrant left subclavian artery forms
concomitantly in essence forming a ring around the trachea and esophagus which could further
leads to the constriction of the said structures (Priya et al., 2018)

1.4 Notch Signaling

Notch signaling is an evolutionary conserved signaling pathways that plays an important role in
cell fate specification, development, differentiation and patterning in numerous cell types in the
body. The most extensively characterized Notch signaling pathway is known as the canonical
Notch signaling pathway which involves a notch transmembrane receptor interacting with a
notch ligand from a neighboring cell (Andersson et al., 2011). This interaction leads to the
proteolytic cleavage of the notch transmembrane domain followed by the internalization of the

3

notch intracellular domain (Notch ICD) (8). Notch ICD then translocates to the nucleus where it
interacts with CBF1/Suppressor of Hairless/LAG-1 (CSL) family DNA binding proteins
(Andersson et al., 2011). This formed complex then binds to the DNA and activate cell specific
target genes depending on the tissue type (de la Pompa and Epstein, 2012). In mammals, there
are four notch receptors (Notch 1-4) and five Notch ligands (Delta Like Ligand 1, 3, 4 and
Jagged 1 and 2) (Andersson et al., 2011). Notch signaling regulates numerous developmental
processes in the developing embryo including Brain, Esophagus, Pancreas, Prostate and many
other organs (Andersson et al., 2011). Similarly, mutations in the components of the notch
signaling pathway be it on the ligands or the receptors leads to variety of defects in the said
organs (Andersson et al., 2011). Research work by multiple groups have shown previously that
notch signaling plays an important role in cardiac development as well (Luxan et al., 2016;
MacGrogan, et al., 2016). Hence, when there are mutations in certain notch components, it
leads to cardiac developmental defects (Luxan et al., 2016; MacGrogan, et al., 2016). Some of
the well-known defects include bicuspid aortic valve disease caused by Notch 1 mutations and
the Alagille syndrome cause by heterozygous mutation of notch ligand Jagged1 (McKellar et al.,
2007; Garg, 2016; Li et al., 1997; Luxan et al., 2016)

1.5 Second Heart Field Progenitor Cells (SHF)

As previously mentioned, at the onset of heart development, two progenitor cell types play
prominent roles (Vincent and Buckingham, 2010; Luxan et al., 2016; Verzi et al., 2005).
Different components of the fully developed heart have origins in these different cell types
(Vincent and Buckingham, 2010; Verzi et al., 2005). Accordingly, the right ventricle and the
outflow tract has its roots on the Second Heart Field (SHF) cells (Vincent and Buckingham,
2010; Verzi et al., 2005). The cells that comprise the second heart field which is also known as
anterior heart field, reside in the pharyngeal mesoderm of the developing embryo and are being
progressively added to the arterial pole of the developing heart at the time of cardiac looping
(Verzi et al., 2005). Identification of this unique cell type giving rise to right ventricle and the
outflow tract of the developing heart is evident by the restricted expression of certain genes and
transgenes only in those structures where mutations in some of the key genes such as mef2c,
Islet1, Nkx2.5, Hand2, and Foxh1, appear to selectively affect the development of the said
structures (Verzi et al., 2005). Hence, to understand the proper boundary and other lineages
derived from this SHF population of cells, researchers have used regulatory elements from the
mouse mef2c gene to direct the expression of Cre recombinase exclusively in the second heart
field and its derivatives in transgenic mice (Verzi et al., 2005).

1.6 Notch signaling in SHF cells

Previous studies have looked in to Notch singling’s affects on cardiac development (High et al.,
2007; High et al., 2009). Specifically, High et all have looked at Jagged1 mediated notch
signaling and how Jagged1 knockout in SHF cells affect cardiac and arch artery development.
These mutant mice show a variety of OFT and arch artery defects. However, the phenotypes
they saw was incompletely penetrant compared to Notch knockout phenotypes indicating notch
signaling could be acting through other ligands other than Jagged1 to cause more severe and
completely penetrant cardiac and arch artery phenotypes. Furthermore, during ventricular
chamber development, Sequential Notch ligand–receptor activation is required for the
cardiomyocytes to move from a more proliferative phase helped by Dll4-notch signaling to a
more maturation phase with help by Jagged1-notch signaling (D’Amato et al., 2016). This

4

transition is facilitated by the concomitant expression of Mfng, a Fringe family of
glycosyltransferases that helps in notch ligand selectivity, Dll4 in the case of cardiomyocyte
proliferation (D’Amato et al., 2016).

To this end, Delta Like Ligand 4 (Dll4) is a possible candidate because of its unique requirement
in arterial development (Benedito and Duarte, 2005). However, due to early embryonic lethality
of whole body Dll4 knockouts, no study has specifically looked in to the affects of Dll4 in cardiac
and arch artery development (Duarte et al., 2004).

Hence, the work that follows tried to look at the effects of Dll4 on cardiac and arch artery
development by using cardiac specific Cre lines to knock out Dll4 only in the SHF cells. Our
work has revealed a novel mechanism of notch signaling in SHF in which when notch signaling
transitions from an early proliferative to a late maturation signaling system, that transition is not
only ligand specific as previously described (D’Amato et al., 2016), but also cell specific.

5

Chapter 2
Methods

2.1 Mice

All animal experiments were carried out under protocols approved by the Institutional Animal
Care and Use Committee of the University of Southern California Islet1-Cre (Cai et al., 2003)
and Mef2c-AHF-Cre (Verzi et al., 2005), Wnt1-Cre (Jiang et al., 2000), Rosa26-
tdTomato (R26RtdT) (Madisen et al., 2010), and Notch Reporter (Nowotschin et al.,
2013) mouse lines have been previously described. In all Cre lines, the Cre gene was
maintained on the paternal side to eliminate risk of germline transmission. Dll4
F/F
mice were
generated in Duarte lab and previously reported (Benedito and Duarte, 2005; Duarte, et al.,
2004; Koch, et al., 2008). Fgf8
F/F
mice were received from Moon lab and have also been
reported (Park, et al., 2006). Dll4-F2-LacZ mice were a kind gift from Joshua Wythe (Wythe, et
al., 2013). Embryo dissection was carried out by standard methods. Genotyping was
undertaken using standard PCR techniques and specific primers used are listed in
Supplemental Table 1.

2.2 Tissue analysis and histology

Immunofluorescence (IF), In situ hybridization (ISH), X-gal staining and Hematoxylin-Eosin
stains were performed using standard techniques. The antibodies used for staining are listed in
Supplemental Table 2. Standard validation techniques included deletion of primary or secondary
antibody or use of blocking peptide to validate antibody specificity, as appropriate. The Dll4
probe used for in situ hybridization has been previously described (Benedito and Duarte, 2005).
Fgf8, Fgf10 and Mef2c ISH were undertaken using RNAscope protocol (Advanced Cell
Diagnostics, Newark, CA). SHF proliferation and apoptosis was assessed by counting number
of double-positive cells in multiple high-power fields in control and mutant sections. Area in
sections positive for β-galactosidase staining was analyzed using ImageJ and normalized to
control. In all cases experiments were repeated in multiple sections of multiple embryos from
different litters with littermate controls. Two-tailed t-test was used to compare significant
differences at a p value <0.05.

2.3 Thoracic Organ and Cell Culture

Embryos were harvested at E9.5. Thoracic region of the embryo was dissected by removing the
head up to the level of pharynx and the lower trunk below the level of the thorax. Thoracic
organs were cultured in Dulbecco’s Modified Eagle Media (DMEM) supplemented with 10%
Fetal Bovine Serum (FBS) and 1% Penicillin/Streptomycin and varying doses of recombinant
Fgf8 (NOVUS/NBP2-35033) for 8 hours. The organs were then cryoembedded and analyzed.
For cell culture, commercially available cell lines were authenticated, lack of contamination
confirmed, and cultured in same medium as above.

2.4 Fgf8 Promoter and Enhancer analysis

6

1kb segments encompassing the 6bp putative binding site of RBPjk in the Fgf8 promoter and
enhancer regions were cloned using primers shown in Supplemental Table 1. The PCR
products were cloned into a promoter-less (Promega/E1771) or an enhancer-less
(Promega/E1761) luciferase vector as appropriate. 293T or HeLa cells were cultured in the
presence of Notch inhibitors (DAPT 30ng/µL, MilliporeSigma/D5942-5MG, or SAHM1 20ng/µL,
MilliporeSigma/491002-1MG) to quench endogenous Notch activity. Cells were co-transfected
with the luciferase construct and NICD expression vector (Addgene/3XFlagNICD1). 24h later,
luminescence was measured using a standard luminometer.
2.5 India ink Injection

For India ink injections, embryos were dissected in cold PBS at E10.5 and E12.5. Chest wall
and pericardial tissues were carefully dissected to expose the heart and glass micropipettes
were used to inject ink into the primitive ventricle and OFT. Embryos were incubated in 4%
paraformaldehyde overnight at 4 C. Whole mount bright field imaging was performed, with
frontal and right and left lateral images to evaluate arch artery organization.

2.6 Human umbilical arterial endothelial cell (HUAEC) tube formation
assay

HUAEC were treated with control, Dll4 or Hey1 siRNA (50nM) as appropriate. Dextran-coated
Cytodex 3 microcarriers were coated with HUAEC, resuspended in fibrinogen, and added to
wells of a 24-well plate containing thrombin to allow clotting. 1ml of endothelial basal media
containing 2% fetal bovine serum (FBS) with or without recombinant VEGF (2.5ng/ml) was
added to each well, and dermal fibroblasts were plated on top of the clot. Beads were
photographed at 5× magnification after 3-5 days in culture.

Large portions of this wok (Chapter 3) is published in Development (De Zoysa et al., Development
1 September 2020; 147 (17): dev185249)
7

Chapter 3
Dll4 Expression in the developing heart and the arch arteries

3.1 Dll4 is expressed at relevant sites and time-points during OFT
formation in the developing heart

I began by evaluating Dll4 expression using multiple modalities in the developing heart at
various embryonic stages and into the neonatal period (Fig 3.1, Fig 3.2). There was good
correlation between Dll4 protein (IF) and transcript (ISH) expression. As a complementary
technique, I used stable transgenic founder mouse lines in which the non-coding region (F2) in
the third intron of Dll4 drives a minimum promoter LacZ reporter (F2-LacZ) (Wythe, et al., 2013).
LacZ expression serves as a surrogate for Dll4 expression in these animals. This enhancer
element was identified specifically for activity in the arterial endothelial elements, and as such, I
found that LacZ expression was strongly observed in arterial endothelial cells and OFT
endocardium and faithfully phenocopied Dll4 protein expression in this regard, as previously
reported (Wythe, et al., 2013). Although expression was also observed in the ventricles and
SHF progenitor cells, the level of expression was significantly lower compared to IF or ISH. This
difference is likely because of lower levels of expression of this particular enhancer in these
cells.

Dll4 expression was discernable in the pharyngeal mesodermal region as early as E8.5 (Fig 3.2
A-A”). Between E9.5-11.5, at a time of intense SHF proliferation and incorporation into the
developing heart, robust expression of Dll4 was observed in the area of the SHF and developing
OFT. By E9.5, Dll4 was broadly expressed in the region of SHF progenitors in the pharyngeal
mesoderm on both transverse (Fig 3.1 A, A’, B, Fig 3.2 B, B’) and sagittal (Fig 3.1 C, C’)
sections. Dll4 expression in this region was confirmed by x-gal staining in F2-LacZ mice (Fig 3.1
D-F’, Fig 3.2 G, G’). Similarly, at E10.5, strong Dll4 expression could be demonstrated in the
region of SHF progenitors by IF (Fig 3.1 G, G’, H, I, I’, Fig 3.2 C, C’), ISH (Fig 3.1 M, M’, N, O,
O’) and x-gal staining (Fig 3.1 J, J’, K, L, L’). To specifically evaluate Dll4 expression in SHF
cells, I lineage traced SHF by crossing Mef2c-AHF-Cre (Verzi, et al., 2005) mice with Rosa26-
tdTomato (R26R,tdT) mice and stained sections for Dll4. At E9 (Fig 3.1 R-R’’’) and E10.5 (Fig
3.1 S-S’’’), tdT positive cells in the pharyngeal mesoderm co-expressed Dll4 confirming SHF
expression. Complementarily, I co-stained sections with Islet1, which is specifically expressed
by SHF progenitor cells at this time point. There was significant overlap between Islet1 and Dll4
expression in the pharyngeal mesoderm (Fig 3.1 P-P’’’, Fig 3.2 Q-Q’’’) confirming that Dll4 is
indeed expressed by SHF progenitor cells. Double staining with the endothelial specific marker,
CD31, and Dll4 showed that a distinct set of endothelial cells (presumably of arterial origin) in
this region also expressed Dll4 (Fig 3.1 T-T’’’). In contrast, airway epithelium marked by Nkx2-1
did not express Dll4 (Fig 3.2 R-R’). By E11.5, Dll4 expression was still seen in pharyngeal
mesodermal region of SHF cells (Fig 3.2 E, E’, I, I’), but was lost at later embryonic time-points
(Fig 3.2J). I then evaluated the expression of Jagged1, the other Notch ligand of relevance in
OFT development (High, et al., 2009), in SHF progenitors. Jagged-1 expression was barely
detectable in Islet1 positive SHF progenitor cells in the pharyngeal mesoderm at E10.5 (Fig
3.1Q-Q’’’), whereas there was more robust expression seen in atrial and ventricular
myocardium.
8

The developing OFT, which is derived from SHF progenitors, also displayed strong Dll4
expression. At E9.5, both IF (Fig 3.1 B, B’, C, C’, Fig 3.2B) and x-gal staining (Fig 3.1 E, E’, F,
F’, Fig 3.2G) confirmed Dll4 expression in the OFT. By E10.5, there was robust Dll4 expression
in the OFT endocardium and a lower, yet detectable, expression in the OFT myocardium by all
three modalities (Fig 3.1 H, H’, I, I’, K, K’, L, L’, N, N’, O, O’, Fig 3.2 H, H’). By E11.5, OFT
expression was weaker (Fig 3.2 F, F’).

Atrial and ventricular endocardium also showed robust Dll4 expression up to E12.5. SHF-
derived RV endocardium expressed Dll4 from E9.5 through E12.5 (Fig 3.1 A, A’’, D, D’’, G, G’’,
J, J’’, M, M’’, Fig 3.2C, C’’, Fig 3.2 E, E’’, J, J’). At earlier time-points, Dll4 expression was
present, but much less robust, in RV myocardium (Fig 3.1 A’’, G’’, Fig 3.2A’’), and myocardial
expression was mostly lost by E11.5 (Fig 3.2E’’). Endocardial expression was reduced
beginning E14.5 (Fig 3.2 K, L) and, by birth, Dll4 expression was observed only in coronary
arterial elements (Fig 3.2 M, M’). Such a temporal variability in the expression of Notch ligands
in the developing ventricle has been previously reported (de la Pompa and Epstein, 2012). In all
the sections evaluated, Dll4 was also expressed in the dorsal aorta as expected, but not in the
adjacent cardinal vein, confirming specificity of the signal observed (Fig 3.1 D, D’, E, G, G’, Fig
3.2H).

3.2 Pharyngeal arch arteries express Dll4 at appropriate time points
during their re-organization

I studied Dll4 protein (IF) and transcript (ISH) expression in PAA at relevant time points during
their formation and re-organization (Fig 3.3). As an additional technique, I used stable
transgenic founder mouse lines in which the non-coding region (F2) in the third intron of Dll4
drives a minimum promoter LacZ reporter (F2-LacZ) (Wythe, et al., 2013). LacZ expression
serves as a surrogate for Dll4 expression in these animals. This enhancer element was
identified specifically for activity in arterial endothelial elements, and as such, this model is
particularly relevant to study Dll4 expression in developing arch arteries.

At E9.5, Dll4 expression is evident in transverse sections of the fourth PAA extending out from
the OFT (Fig 3.3 A, A’) and in the developing dorsal aorta in sagittal sections (Fig 3.3 B, B’). Dll4
expression is also evident in the fourth PAA in coronal sections (Fig 3.3 C, C’). Whole mount x-
gal staining in F2-LacZ mice reveals LacZ signals indicating Dll4 expression in 3
rd
, 4
th
and 6
th

PAA bilaterally (Fig 3.3 D1, D2). Transverse, sagittal and coronal sections of these mice confirm
Dll4 expression in the same areas as demonstrated by IF (Fig 3.3 E-G’). Dll4 protein expression
persists in bilateral 3
rd
, 4
th
and 6
th
PAA, dorsal aorta and intersomitic vessels at E10.5 as well
(Fig 3.3 H-J’). Insitu hybridization reveals Dll4 mRNA expression in 4
th
PAA and dorsal aorta in
transverse (Fig 3.3 K, K’) and sagittal (Fig 3.3 L, L’) sections. Similarly, whole mount x-gal
staining and sections in F2-LacZ mice confirms Dll4 expression in 3
rd
, 4
th
and 6
th
PAA bilaterally
(Fig 3.3 M1-P’). At both these stages, Dll4 expression is also observed in SHF progenitors in the
pharyngeal mesoderm as was previously shown.

To confirm the activity of Notch receptor in these cells, I crossed F2-LacZ mice with a Notch
reporter line and studied LacZ expression by IF and x-gal staining. 3
rd
, 4
th
and 6
th
PAA express
Dll4 and demonstrate Notch activity at E9.5 (Fig 3.3 Q, QL1-QR3) and E10.5 (Fig 3.3 R, R’, R1-
9

R3). Finally, to confirm Dll4 expression in vascular elements, I stained E10.5 sections for the
endothelial specific marker, CD31, and Dll4. I was able to confirm co-expression of Dll4 and
CD31 in dorsal aorta (DA in Fig 3.3 S3), but lack of Dll4 expression in adjacent CD31-positive
cardinal vein (V in Fig 3.3 S3), as anticipated.

10

11

Fig 3.1
Dll4 is expressed by SHF progenitor cells and SHF-derived structures in the
developing heart.
Representative images of E9.5 and E10.5 embryos are shown. Dll4 protein expression
(immunofluorescence, IF) was studied in E9.5 transverse (A and B) and sagittal fixed-frozen
sections (C). Dll4 is expressed in the pharyngeal mesoderm (PM) in the SHF progenitor cell
region (A and C, and higher magnification of upper boxed area of A in A’, and C in C’).
Transverse sections demonstrate expression in RV endocardium and myocardium (A, and
higher magnification of lower boxed area of A in A”) and developing OFT (B, and higher
magnification of boxed area of B in B’). X-gal staining in Dll4-F2-LacZ embryos was used as
a complementary method to assess Dll4 expression (D-F). Comparable sections show
staining in the PM (arrowheads in D’ and F’), developing RV (D, and higher magnification of
lower boxed area of D in D’) and OFT (E, and higher magnification of boxed area of E in E’).
E10.5 embryos also demonstrate a very similar pattern of Dll4 expression on IF (G-I) and x-
gal staining in Dll4-F2-LacZ embryos (J-L). Dorsal aorta (DA in G’ and J’) expresses Dll4,
while adjacent cardinal vein (V) does not. Dll4 transcript expression was evaluated by in situ
hybridization (ISH) (M-O). Comparable sections again show staining in PM, developing RV
and OFT. Transverse sections were co-stained with Islet1 and Dll4 (P) or Jagged1 (Q) at E9.
Higher magnification of boxed area in P and Q are shown as Islet1 expression (P’, Q’), Dll4
expression (P’’), Jagged1 expression (Q’’) and merged image (P’’’, Q’’’) to demonstrate
robust expression of Dll4, and lack of expression of Jagged1 in the PM. Transverse sections
were stained for Dll4 in Mef2c lineage traced embryos at E9 (R) and E10.5 (S). Higher
magnification of boxed area in R and S are shown as Mef2c expression (R’, S’), Dll4
expression (R’’, S’’) and merged image (R’’’, S’’’) to demonstrate co-localization of Dll4 on
SHF expressing cells in the PM. Transverse sections of E10.5 embryo were co-stained for
Dll4 and vascular endothelial (CD31) marker (T). Higher magnification of boxed area in T is
shown as CD31 expression (T’), Dll4 expression (T’’) and merged image (T’’’) to
demonstrate co-localization of Dll4 and CD31 expression in (arterial) endothelial elements in
the pharyngeal mesoderm.
The boxed regions of A’, D’, G’, J’, M’, P’ and Q’ in transverse sections and C’, F’, I’, L’ and
O’ in sagittal sections indicate the region occupied by SHF progenitor cells.
A, Aorta; DA, Dorsal Aorta ; ISV, Inter-somitic Vessels; PM, Pharyngeal Mesoderm; V, Vein.
Scale Bars: 50µm (R’-S’’’), 100µm (A’-C’, G’-I’, M’-O’, P’-Q’’’, R T’-T’’’), 150µm (D’-F’, J’-L’),
250µm (A-C, G-I, M-O, P, Q, S, T), 300µm (D-F, J-L)
12

13

Figure 3.2
Dll4 is expressed at appropriate time-points by SHF progenitor cells and SHF-derived
structures in the developing heart.
Dll4 protein expression at various embryonic time-points (immunofluorescence, IF) is shown
in A-F. Transverse section of E8.5 embryo shows Dll4 expression in SHF progenitor cell
region in the pharyngeal mesoderm (A, and higher magnification of upper boxed area of A in
A’), and RV endocardium and myocardium (A, and higher magnification of lower boxed area
of A in A’’). Transverse section of E9.5 embryo shows Dll4 expression in SHF progenitor cell
region in the pharyngeal mesoderm (B, and higher magnification of boxed area of B in B’).
Transverse section of E10.5 embryo shows Dll4 expression in SHF progenitor cell region in
the pharyngeal mesoderm (C, and higher magnification of upper boxed area of C in C’), and
RV endocardium and myocardium (C, and higher magnification of lower boxed area of C in
C’’). Sagittal section of E10.5 embryo shows Dll4 expression in SHF territory in the
pharyngeal mesoderm (D, and higher magnification of boxed area of D in D’). Transverse
section of E11.5 embryo shows weaker, but discernable, Dll4 expression in SHF progenitor
cell region in the pharyngeal mesoderm (E, and higher magnification of upper boxed area of
E in E’). By E11.5, RV expression is largely restricted to the endocardium and lost in
myocardium (E, and higher magnification of lower boxed area of E in E’’). More cranial
transverse section of E11.5 embryo shows persistent, Dll4 expression in OFT endocardium
and, to a significantly lower degree, myocardium (F, and higher magnification of boxed area
of F in F’). X-gal staining in Dll4-F2-LacZ embryos was used as a complementary method to
assess Dll4 expression (G-M). Transverse sections of E9.5 embryo (G) show staining in PM
(arrowheads in magnified upper boxed area of G in G’). Transverse sections of E10.5
embryo (H) show OFT endocardium being strongly positive (Magnified boxed area of H in
H’). Sagittal section of E11.5 embryo shows more restricted x-gal staining in PM (I, and
arrowhead in higher magnification of boxed area of I in I’), and in developing OFT. By E12.5,
transverse sections demonstrate that RV expression is largely restricted to the endocardium
(J, and higher magnification of boxed area of J in J’). Transverse sections (K, L) of E14.5
embryos show scattered stains in RV endocardium, with prominent staining in developing
coronary arterial endothelial elements. By post-natal day 3, transverse sections demonstrate
expression restricted to coronary arteries in the RV wall (M, and higher magnification of
boxed area of M in M’). Dll4 transcript expression was evaluated by in situ hybridization
(ISH). Representative images of transverse and sagittal sections of E10.5 embryos showing
absence of stains with sense strand ISH (N-P) confirming specificity of the signal shown in
Fig1. Transverse sections of an E10.5 embryo were co-stained for Dll4 and Islet1 expression
(Q). Higher magnification of boxed area in Q is shown as Dll4 expression (Q’), Islet1
expression (Q’’) and merged image (Q’’’) to demonstrate co-localization of Dll4 and Islet1
expression in SHF progenitor cells in the PM. The insets in Q’-Q’’’ are further magnified
views of the respective boxed regions in each image. Transverse section of E10.5 embryo
for the lung bud marker Nkx2-1 shows Nkx2-1 expression in the pharynx in the PM (R, and
higher magnification of boxed area of R in R’).
The boxed regions of A’, B’, C’, E’, G’, Q’ and R’ in transverse sections and D’ and I’ in
sagittal sections indicate the region occupied by SHF progenitor cells.
Scale Bars: 50µm (A’, A’’), 100µm (A, B’-F’, N’-R’), 150µm (G’-J’, L, M’), 250µm (B-F, N-R),
300µm (G-J, K, M)
14

15

Fig 3.3
Dll4 is expressed by SHF progenitor cell-derived pharyngeal arch artery endothelium
as well as in the dorsal aorta.
Representative images of E9.5 and E10.5 embryos are shown. Dll4 protein expression
(immunofluorescence, IF) was studied in E9.5 transverse (A), sagittal (B) and coronal (C)
fixed-frozen sections. Dll4 is expressed in the pharyngeal mesoderm in the SHF progenitor
cell region and the cardiac OFT connecting to the 4
th
pharyngeal arch artery and then to the
dorsal aorta (A and higher magnification of upper boxed area of A in A’). Sagittal sections
also show that Dll4 is expressed in the aortic sac from the OFT as well as the dorsal aorta (B
and higher magnification of upper boxed area of B in B’). Coronal sections show 3
rd
and 4
th

pharyngeal arch arteries expressing Dll4 (C and higher magnification of upper boxed area of
C in C’). X-gal staining in Dll4-F2-LacZ embryos was used as a complementary method to
assess Dll4 expression both in whole mount (D) and sections (E). Whole mount lacZ staining
reveals X-gal signals in the 3
rd
, 4
th
and 6
th
pharyngeal arch arteries bilaterally (Arrowheads in
D1 and D2). Sections show X-gal staining pattern similar to IF in transverse (E), sagittal (F)
and coronal (G) sections. E10.5 embryos also demonstrate Dll4 expression on IF in the
pharyngeal mesoderm, 4
th
pharyngeal arch artery and developing dorsal aorta (DA) and
intersomitic vessels (H-J). Dll4 transcript expression evaluated through in situ hybridization
(ISH) (K-L) also showed similar expression pattern. Cardinal vein (V) adjacent to DA (H’)
does not express Dll4 as expected. X-gal staining in Dll4-F2-LacZ embryos at E10.5 shows
Dll4 expression in the bilateral 3
rd
, 4
th
and 6
th
pharyngeal arch arteries in whole mount
examination (arrowheads in M1 and M2). Sections (N-P) confirm expression pattern seen
with IF. Arrowheads in P’ represent the bilateral 3
rd
, 4
th
and 6
th
pharyngeal arch arteries.
Coronal sections of Dll4-F2-LacZ and Notch reporter transgene positive embryos were then
evaluated by LacZ immunostaining. At E9.5, LacZ positivity and Notch activity are seen in
both the left (Q, QL1-Ql3) and right (Q, QR1-QR3) 3
rd
and 4
th
pharyngeal arch arteries. This
co-localization was evident at E10.5 as well (R1-R3). Arrowheads in R’ and R3 represent the
3
rd
, 4
th
and 6
th
pharyngeal arch arteries. There is co-localization of Dll4 and CD31 expression
in (arterial) endothelial elements in the pharyngeal mesoderm and dorsal aorta (DA).
Adjacent cardinal vein is CD31-positive (S2), but Dll4-negative (S1) confirming specificity of
Dll4 signal.
Wholemount magnification: x3 (D1, D2, M1, M2)
Scale Bars: 50µm (C’, R1-R3), 75µm (G’), 100µm (A’, B’, H’, H’’, I’, J’ K’, K’’, L’, QL1-QL3,
QR1-QR3, S1-S3), 150µm (E’, F’, N’, O’, P’, R’), 250µm (A-C, H, I-L, Q, S ), 300µm (E-G, N-
P, R).

Large portions of this wok (Chapter 3) is published in Development (De Zoysa et al., Development
1 September 2020; 147 (17): dev185249)
16

Chapter 4
Dll4 mediated notch signaling in SHF cells is required for the proper
cardiac development

4.1 Introduction

The role played by Notch pathway in cardiac progenitor cell biology remains to be elucidated.
Delta-like ligand-4 (Dll4), the arterial-specific Notch ligand, is expressed by second heart field
(SHF) progenitors at time-points crucial in SHF biology. Dll4-mediated Notch signaling is
critically required for maintaining an adequate pool of SHF progenitors, such that Dll4 knockout
results in reduction in proliferation and increase in apoptosis. Reduced SHF progenitor pool
leads to an underdeveloped right ventricle (RV) and outflow tract (OFT). In its most severe form,
there is severe RV hypoplasia and poorly developed OFT resulting in early embryonic lethality.
In milder form, the OFT is foreshortened and misaligned resulting in double outlet right ventricle.
Dll4-mediated Notch signaling maintains Fgf8 expression by transcriptional regulation at the
promoter level. Combined heterozygous knockout of Dll4 and Fgf8 demonstrates genetic
synergy in OFT alignment. Exogenous supplemental Fgf8 rescues proliferation in Dll4 mutants
in ex-vivo culture. These results establish a novel role for Dll4-mediated Notch signaling in SHF
biology. More broadly, the model shown here provides a platform for understanding oligogenic
inheritance that results in clinically relevant OFT malformations.

4.2 Dll4 expression in SHF is required for appropriate development of
SHF-derived RV and OFT

Global knockout of Dll4 is embryonically lethal due to vascular maturation arrest (Duarte, et al.,
2004). No mutant survived past E10,5, with the majority dying even earlier. The few mutants
that survived to E10.5 were severely underdeveloped and demonstrated arrested cardiac
development and a very poorly developed OFT (Fig 4.2 A, A’), precluding detailed analysis of
cardiac-specific effects. To circumvent this early mortality, I conditionally knocked out Dll4
expression in SHF using specific Cre lines. I used both Islet1-Cre (more global SHF expression,
Cai, et al., 2003) and Mef2c-AHF-Cre (anterior SHF-specific expression, Verzi, et al., 2005)
lines. Efficient recombinase-mediated loss of Dll4 in the SHF was confirmed at E9.5 in Mef2c-
AHF-Cre,Dll4
F/F
embryos, which showed loss of expression in the pharyngeal mesoderm (Fig
4.2 B, B’) and SHF-derived RV, but persistent robust expression in the LV (Fig 4.2 B, B”).

Depending on the time and extent of Dll4 knockout, a spectrum of phenotypic defects was
observed in mutant embryos. Homozygous deletion of Dll4 in a more extensive population of
cells (Islet1-Cre,Dll4
F/F
) resulted in complete lack of RV and poorly developed hearts at E10.5
(Fig 4.2 C, C’). There was early embryonic lethality such that, by E10.5, only 13% mutants (25%
predicted by Mendelian inheritance) could be recovered. No live embryo was recovered by
E14.5 (Fig 4.1M). More restricted loss of expression in anterior SHF only (Mef2c-AHF-

17

Cre,Dll4
F/F
)

also resulted in recovery of fewer embryos at E14.5 than predicted (6% vs. predicted
25%). All six mutants recovered live at E14.5 displayed Double Outlet Right Ventricle (DORV,
Fig 4.1 A’, B’, Fig 4.3B). These embryos had a large ventricular septal defect (VSD, arrow in Fig
4.1A’). The OFT was appropriately septated with distinct aortic and pulmonary valves (Fig 4.3B
19-22, and 34-36). However, the aortic valve originated from the RV (arrowhead in Fig 4.1B’) at
the same level as the pulmonary valve (side-by-side orientation). The aortic valve connected to
the aorta and the pulmonary valve to the pulmonary artery appropriately. This implies that
Mef2c-AHF-Cre-mediated Dll4 knockout impacts OFT alignment without any impact on
septation. I interpret these findings to indicate that more extensive knockout of Dll4 in SHF
results in a severe reduction in SHF-derived RV and OFT resulting in early embryonic lethality,
but anterior SHF-specific loss leads to less severe reduction in the size of these SHF-derived
structures. The resultant shorter OFT is incapable of expanding to align itself appropriately over
the developing RV and LV, resulting in DORV. All of these conditional mutants showed cardiac
inflow tract (venous pole) development that was appropriate for embryonic stage, implying that
loss of Dll4 mediated by these cre lines did not result in inflow tract defects. Similarly, lung
development was also appropriate for age in all mutants examined.

I then examined mutant embryos in the Mef2c-AHF-Cre background at earlier time-points. At
E9.5, the RV appeared to be slightly smaller in mutants (Fig 4.2 D, D’, E) and the OFT was
foreshortened (Fig 4.2 F, F’, G). By E10.5, this difference was clearly evident on both whole
mount evaluation (Fig 4.2 H, H’) and sections (Fig 4.1 C, C’). In addition, the OFT was also
much shorter and paucicellular (asterisk in Fig 4.1H’ compared to Fig 4.1H) in the mutants. I
then proceeded to label the SHF-derived structures by breeding in the Rosa26-LacZ
(R26R,LacZ) gene into the mutant crosses in order to quantitate this reduction. At E10, x-gal
stained hearts were examined for the size of the LacZ-positive RV and OFT. Whole mount
examination confirmed that mutant hearts had a smaller RV (Fig 4.1 D, D’, F) and shorter,
narrower OFT (Fig 4.1 I, I’, K) compared to controls. Similarly, evaluation of sections showed
that the mutant RV was 50% smaller than control RV (Fig 4.1 E, E’, G) indexed to the size of the
respective LV. The OFT was also 50% shorter in length (Fig 4.1 J, J’, L).

Global heterozygous loss of Dll4 demonstrates incompletely penetrant haploinsufficiency
resulting in vascular maturation defects and embryonic lethality. I therefore sought to evaluate if
heterozygous loss of Dll4 in these two Cre backgrounds had any phenotypic consequence. Of
the 14 Islet1-Cre,Dll4
F/wt
embryos examined at E14.5, 6 (43%, Fig 4.1M) demonstrated DORV
(Fig 4.1 A’’, B’’, Fig 4.3C) with a large VSD and aorta that arose from the RV. The aortic valve
was appropriately located caudal to the pulmonary valve, implying that the sub-pulmonary
conus was well developed in these mutants (Fig 4.3C 15-18 and 30-34). The remaining 8
embryos had normal cardiac anatomy. In contrast, all 38 Mef2c-AHF-Cre,Dll4
F/wt
mice examined
at E14.5 had normal anatomy. These mice were born alive and grew and reproduced normally.
These results indicate that Dll4 expression is required for appropriate development of SHF-
derived RV and OFT. The observed phenotypes range from complete lack of RV and OFT
following more extensive knockout, to a fully penetrant DORV following anterior SHF-specific
knockout, to an incompletely penetrant DORV with partial loss and normal heart development in
the setting of partial loss of Dll4 in a more restricted pool of SHF cells.

4.3 Dll4 expression is required for SHF cell proliferation to maintain
an adequate progenitor cell pool

18

The observed mutant phenotypes suggest that there is a reduction in the number of SHF
progenitor cells that are incorporated into the developing heart in Dll4 mutants. One potential
mechanism to explain this finding would be an inadequate pool of SHF progenitors available for
incorporation. To directly test this hypothesis, I fate-mapped SHF cells (Mef2c-AHF-
Cre,R26R,LacZ) in the pharyngeal mesodermal region. Both in transverse (Fig 4.4 A-C) and
sagittal (Fig 4.4 D-F) sections, the area occupied by LacZ positive SHF progenitor cells was
significantly reduced in mutants (by 67% and 50%, respectively). This would imply that the
reduction in SHF-derived structures in the heart is due to a reduction in the size of SHF
progenitor pool. I studied SHF proliferation to explain this reduction in SHF progenitor pool. To
this end, control and mutant embryos were stained for Islet1 to mark SHF cells and pHH3 to
identify proliferating cells. Double-positive cells in the region of the pharyngeal mesoderm were
counted in controls and mutants. At E9.5 (Fig 4.4 G-I), Dll4 knockout resulted in a 51%
reduction in proliferating SHF cells, whereas there was no change in proliferation of Islet1
negative, non-SHF cells. This proliferation defect persisted to E10.5 (Fig 4.5 A-C), wherein a
72% reduction in proliferating SHF cells was observed. Given that SHF proliferation was
impacted by E9.5, I studied apoptosis in SHF cells a day later by double staining for Islet1 and
TUNEL. Dll4 knockout resulted in a significant increase in SHF progenitor cell apoptosis at
E10.5 (Fig 4.4 J-L). Taken together, these results indicate that Dll4 expression in SHF maintains
SHF cell proliferation during the crucial time period between E9-11 when these cells are being
actively incorporated into the developing heart. Loss of Dll4 expression results in reduced
proliferation of SHF cells and their subsequent apoptotic loss. These events lead to a significant
reduction in the pool of progenitor cells available for incorporation into the developing heart,
which in turn leads to a reduction in the size of SHF-derived RV and OFT, and the resultant
phenotypes described above.

4.4 Dll4-mediated Notch signaling regulates Fgf8 expression in SHF

I sought to identify the molecular mechanisms that act downstream of Dll4-mediated Notch
signaling to regulate SHF proliferation. I evaluated expression levels of various molecules with
relevance to SHF biology. Fgf8 is a key regulator of SHF proliferation and maturation (Ilagan, et
al., 2006; Park, et al., 2006, Fischer, et al., 2002). I studied Fgf8 expression in SHF of control
and mutant embryos at mRNA and protein level, by co-staining for Mef2c and Islet1,
respectively. Fgf8 mRNA and protein levels were markedly reduced in the pharyngeal
mesoderm of mutant embryos (without significant change in areas outside SHF territory such as
neural tube) at both E9.5 (Fig 4.6 A-B’’’ and E-F’’’) and E10.5 (Fig 4.7 A-B’’’). Fgf10, another
important molecule in SHF maturation (Watanabe, et al., 2012), was also significantly reduced
in SHF at mRNA (Fig 4.6 C-D’’’) and protein (data not shown) levels, but not in atrial wall (non-
SHF derived tissue). Hand2 is an important specification marker of RV myocardium
(Tsuchihashi, et al., 2011), and this also showed markedly reduced expression in RV of mutants
(Fig 4.6 G, G’, H,H’). There was no change in the very low basal expression level in the LV
(non-SHF-derived, Fig 4.6 G’’,H’’). There was no difference in the expression levels of
molecules in other pathways of relevance in SHF biology, such as Bmp4 (Liu, et al., 2004) or
Mlc2v (Franco, et al., 1999) (Fig 4.7 C-F). In order to evaluate potential rotation abnormality, I
stained for the sub-pulmonary rotation marker Sema3C. The extent of Sema3C expression in
the developing OFT and its regionalization was not impacted in the mutants compared to
controls (Fig 4.7 G-N). Lastly, left-right asymmetry regulators like Nodal and Pitx2 were also not
altered in mutant embryos (data not shown). These results would imply that the primary
molecular defect underlying the DORV phenotype observed in the mutants is reduced SHF

19

progenitor cell proliferation and incorporation into the developing OFT. Alternative mechanisms,
such as defective OFT rotation, while possible, appear less likely to be the primary defect.

I chose to further pursue Fgf8 regulation given its important role in SHF biology, in particular its
role in SHF progenitor cell proliferation (Ilagan, et al., 2006; Park, et al., 2006). Ligand binding to
Notch receptors results in proteolytic cleavage and release of the Notch intracellular domain
(NICD). NICD binds to downstream molecules and assembles a transcriptional machinery
comprised of proteins such as RBPjk and master-mind-like (MAML). This complex activates
transcription of Notch target genes within the nucleus. TGGGAA is the putative consensus
binding sequence for RBPjk, the essential transcription factor in Notch signaling (Del Bianco, et
al., 2010; Castel, et al., 2013). I studied the mouse chromosome 10 upstream of the Fgf8
transcriptional start site and identified two putative RBPjk binding sites 989 and 4410 bases
upstream of 5’UTR (Fig 4.8A). I cloned 1KB regions around these binding sites as well as a
control 1KB region not including either site into a promoter-less luciferase expression vector. It
has been previously demonstrated that the 3
rd
large intron of the Fgf8 gene has significant
enhancer activity (Gemel, et al., 1999). I therefore evaluated this intron as well and identified the
consensus sequence for RBPjk binding within this intron. I cloned two 1KB regions of this intron,
one with and one without this binding site. These were sub-cloned into an enhancer-less
luciferase expression vector. I then performed luciferase assay in two different cell lines using
two different methods to quench basal Notch activity. 293T cells were treated with DAPT, a γ-
secretase inhibitor, and then transfected with the various luciferase expression vectors with and
without NICD expression vector to induce Notch activity. Luciferase expression was increased
2.5-fold from baseline only when the promoter 1 construct was co-transfected with NICD,
indicating that Notch signaling regulates Fgf8 expression at the promoter level (Fig 4.8B).
Similarly, HeLa cells also showed a 2.4-fold increase in luciferase expression when promoter 1
construct was co-transfected with NICD (Fig 4.9A). Using another inhibitor of Notch protein
assembly to quench basal activity, SAHM1, I was able to reproduce a 2-fold increase in
luciferase expression in 293T cells with promoter 1 construct (Fig 4.9B). To confirm the
specificity of the binding sequence in promoter 1, I created two site-directed mutant clones of
promoter 1 (Fig 4.8A), both of which lost luciferase activity confirming the veracity of the putative
binding site (Fig 4.8C).

4.5 Genetic synergy between Dll4-mediated Notch and Fgf8 signals in
SHF proliferation

These results thus far indicate that Dll4-mediated Notch signaling in SHF regulates Fgf8
expression to maintain SHF proliferation. Loss of Dll4 leads to loss of Fgf8 expression and a
reduction in SHF proliferation and progenitor pool of cells. To confirm that Fgf8 was the key
downstream molecular pathway that impacted SHF proliferation, I evaluated whether
replenishing Fgf8 could rescue the loss in SHF proliferation in an in vitro model system. I
dissected the thoracic area of E9.5 embryos and cultured them for 8 hours in vitro in the
presence of increasing doses of exogenous recombinant Fgf8. Cultured ‘organs’ were then
sectioned and stained for Islet1 and pHH3 to discern the degree of SHF proliferation.
Exogenous Fgf8 supplementation led to a small, statistically insignificant, increase in double-
positive proliferating SHF cells in the pharyngeal area of thoracic organs from control embryos
(Fig 4.8 D-D’’, G, Fig 4.9C-D’’). Thoracic organs from mutant embryos exhibited a greater than
4-fold reduction in SHF proliferation compared to controls under baseline culture conditions (Fig
4.8 E-E’’, G). Supplementation with Fgf8 led to a dose-dependent and significant increase in
SHF proliferation in mutant organs (Fig 4.8 F-F”, G, Fig 4.9E-E’’). At 100ng/µL of exogenous

20

Fgf8, there was no difference in the number of proliferating SHF cells in mutant organs
compared to controls. These data further support the hypothesis that the reduction in SHF
proliferation observed with loss of Dll4 expression is primarily due to loss of Fgf8 expression.

I then studied genetic synergy between these two pathways in vivo. I hypothesized that because
Dll4 and Fgf8 pathways impacted SHF proliferation, compound partial loss of both of these
proteins would have a more penetrant SHF phenotype. Mef2c-AHF-Cre,Dll4
F/wt
embryos
displayed normally developed hearts at E14.5 (Fig 4.8 I, I’ and M). Partial loss of Fgf8 in SHF
(Mef2c-AHF-Cre,Fgf8
F/wt
) resulted in an incompletely penetrant phenotype. Of the 7 embryos
evaluated, only one (14%) demonstrated mal-alignment of the aortic valve (Fig 4.8 K, K’). In two
(28%) other mice, a very shallow VSD with a normal OFT was encountered and the remainder
of the embryos was normal (Fig 4.8 J, J’ and M). In contrast, 10 of the 12 (83%) embryos with
concomitant partial loss of both proteins (double heterozygous Mef2c-AHF-Cre,Dll4
F/wt
,Fgf8
F/wt
)
displayed DORV (Fig 4.8 L, L’ and M) confirming genetic synergy between these two pathways.
There was a gradation in the severity of the phenotypes observed. The VSD in Fgf8
heterozygotes was very shallow and there was only a slight displacement of the aortic valve
towards the RV. The double heterozygotes showed deeper VSD and the aortic valve was more
prominently overriding the ventricular septum, reminiscent of the clinically encountered
tetralogy-type DORV. The aortic valve was still more caudal in location compared to the
pulmonary valve in these mutants. In contrast, the Dll4 knockout embryos had an even larger
VSD and the aortic valve was completely displaced over the RV and located at the same level
as the pulmonary valve (Fig 4.1B’ compared to Fig 4.8L’).

4.6 Discussion

This study evaluates the biological role of Dll4 expression in SHF progenitors and demonstrates
that Dll4 expression is required for progenitor cells to proliferate and ensure the availability of an
adequate pool of cells for incorporation into the developing heart. Such a pro-proliferative role
for Dll4 has been suggested in other progenitor beds as well. Dll4 is expressed by retinal
progenitor cells and serves as the major Notch ligand to expand the progenitor pool (Luo, et al.,
2012). Dll4 is also expressed in a subset of neural progenitors in the spinal cord and its
expression is required for inter-neuronal subset specification (Rocha, et al., 2009). Dll4 signaling
is required to ensure early commitment to T cell lineage and to maintain an adequate pool of T
cell progenitors (Hozumi, et al., 2008, Billiard, et al., 2012, Yu, et al., 2015). In the context of
cardiomyocytes, following initial cardiomyocyte specification, endocardial Dll4-Notch1 signaling
promotes cardiomyocyte proliferation, while subsequent patterning requires downregulation of
Dll4 expression later in gestation (D'Amato, et al., 2016). Thus, Dll4 serves as the primary Notch
ligand that expands cells immediately following their early commitment to ensure that an
adequate pool of cells is available for differentiation into their ultimate cell fate.

Notch receptor and ligands are expressed widely at different time-points and are thought to play
an important role in heart development. My work show that during early time-points of heart
development (E8.5-E10.5), Dll4, but not Jagged1, is expressed by SHF progenitor cells. As SHF
cells mature to form the OFT and RV, they continue to express Dll4. Prior studies have shown
that the expression of myocardial cell specific factors in the developing cardiomyocyte
suppresses Dll4 expression (D'Amato, et al., 2016). This evaluation confirms these findings
demonstrating that by E11.5, Dll4 expression is lost in the myocardium and is primarily
restricted to the endocardium. D’Amato et al showed by RNA analysis that as early as E9.5, Dll4
mRNA is restricted to the endocardium alone. The protien analysis suggests continued

21

expression, albeit weak, in the myocardium up to E11.5. This discrepancy may relate to
experimental differences, or may represent residual protein translated from mRNA expressed
earlier in development. This current study also provides additional insights into the role of Notch
signaling in OFT development. High et al. utilized dominant-negative mastermind-like protein to
knockout signaling by all Notch receptors in the SHF (High, et al., 2009). They observed similar
cardiac phenotypes including DORV, VSD and, occasionally, common arterial trunk. Using
Jagged1 knockout, they demonstrate that Notch signaling regulates endothelial-mesenchymal
transition and maturation between E12.5-13.5 within an adequately formed OFT, a process they
elegantly recapitulate in vitro. Their study did not evaluate a particular role for Notch in SHF
progenitor cell biology. The results demonstrate that during early stages of cardiogenesis, Notch
signaling is primarily mediated by Dll4 and plays a distinct and novel role in maintaining SHF
proliferation. Loss of Dll4 results in reduced pool of SHF cells leading to a foreshortened OFT,
which also results in a fully penetrant DORV phenotype, as observed by High et al. my results
also show that Fgf8 is the primary mediator of Notch signaling in SHF, similar to the
observations of High et al. However, I did not notice any septation defects in Dll4 mutants unlike
the common arterial trunk phenotype observed by High et al, implying that Dll4 and Jagged1
mediated Notch signaling pathways likely diverge at some downstream level. The results from
these two studies would imply that Notch signaling is crucial in OFT development, but is
orchestrated by different ligands at different time-points, The earlier effect mediated by Dll4
primarily regulates SHF proliferation, while the later role mediated by Jagged1 regulates more
specific maturation effects, such as EMT. Such a differential effect of Notch signaling has also
been shown in the context of cardiomyocyte development (D’Amato et al., 2016).

Neural crest knockout of Dll4 has no phenotype (data not shown) consistent with lack of
expression of Dll4 in neural crest cells. Neural crest-specific knockout of Notch signaling
primarily resulted in defects in pharyngeal arch patterning and pulmonary artery stenosis, and
rarely VSD (High, et al., 2007). There were no OFT alignment defects reported. Taken together
with prior observations, these results would imply that SHF-expressed Dll4 signals via SHF-
expressed Notch receptors to mediate SHF progenitor cell biology. Such signaling by SHF cells
into other SHF cells has been described in the context of Fgf8, wherein Fgf8 secreted by SHF
acts on Fgfr also expressed by SHF cells (Park, et al., 2008). Dll4 and Notch have generally
been thought to interact in trans, such that membrane-bound Dll4 on one cell interacts with
Notch expressed on the adjacent cell. Whether Dll4-Notch signaling in SHF also represents
trans interaction or cis interaction remains to be elucidated.

These data also show that Dll4-mediated Notch signaling regulates Fgf8 expression. The
importance of Fgf8 pathway in heart development is well established (Frank, et al., 2002;
Macatee, et al., 2003; Park, et al., 2006; Park, et al., 2008). Generally, it is believed that the first
enhancer segment located in the third intron of Fgf8 serves as the primary regulator of Fgf8
expression. I show here that Notch signaling regulates Fgf8 expression at the promoter level.
Given the importance of Fgf8 in SHF biology, two distinct sites of regulation allow for
redundancy and the ability to further modify expression through multiple mechanisms. Around
E9 in mice, as SHF progenitors are actively proliferating, Dll4 regulates proliferation through
multiple pathways. Fgf8 levels begin to fall by E9.5, and concomitantly, there is reduction in
SHF proliferation. The data would therefore suggest that the primary mechanism by which Dll4
regulates SHF proliferation is via Fgf8 expression. As I have shown, other molecules such as
Fgf10 are also reduced when Dll4 expression is lost. These changes in other molecules may, in
part, explain some of the earlier reduction observed in SHF proliferation. This may also underlie
the observation that the phenotype seen in homozygous Dll4 knockout is more penetrant and
severe than the phenotype in Dll4 and Fgf8 double heterozygotes.

22

Notch pathway mutations have been implicated in a variety of congenital heart defects.
Mutations in the Notch ligand, Jagged1, is thought to be causative in Alagille syndrome, which is
characterized by biliary malformations, pulmonary artery stenosis and rarely OFT defects.
Recently, heterozygous deleterious mutations in Dll4 have been implicated in Adams-Oliver
syndrome. This is a rare genetic disease characterized by aplasia cutis congenita, terminal
transverse limb defects and cutis marmorata. CHD is encountered in about 20% of these
patients, and include VSDs, or DORV/tetralogy type defects (Meester, et al., 2015; Nagasaka,
et al., 2017). In a large study of whole genome sequencing or targeted resequencing of the
Dll4 gene with a custom enrichment panel in independent families with Adams-Oliver syndrome,
nine heterozygous mutations in Dll4 were identified, including two nonsense and seven
missense variants (Meester, et al., 2015). All of these mutations resulted in loss of Dll4 function.
Similar to these clinical reports, heterozygous loss of Dll4 in Islet1-Cre background in the study
resulted in about a 40% incidence of DORV/tetralogy type defects. Thus, my study is the first
demonstration of the molecular basis underlying the clinical CHD finding in this syndrome.
De novo mutations in single genes have been shown to contribute to approximately 10% of all
severe CHD (Zaidi, et al., 2013), implying that the majority of CHD lacks an identifiable
monogenic etiology. Interaction between mutations in two distinct genes can potentiate or
suppress the impact of these mutations in isolation. There is growing evidence to suggest that
such complex and co-existing oligogenic mutations may underlie a larger proportion of CHD
(Akhirome, et al., 2017, Granados-Riveron, et al., 2012, Jin, et al., 2017). It is conceivable that
heterozygous mutations may be inherited from parents who could be silent carriers, but the
convergence of these mutations in the offspring would result in CHD not observed in either
parent. In a recent study by Gifford et al, compound heterozygosity in Mkl2, Myh7, and Nkx2-5
genes inherited by the offsprings of clinically unaffected parents (who carried only one or two of
the mutations) resulted in non-compaction cardiomyopathy (Gifford, et al., 2019). Similarly,
double heterozygous mutations in dynein family of proteins have been implicated in heterotaxy
(Li, et al., 2016). With particular reference to tetralogy-type defects, Topf et al sequenced twelve
genes implicated in the SHF transcription network in 93 non-syndromic tetralogy patients (Topf,
et al., 2014). Concomitant heterozygous mutations in Hand2 and Foxc1 were found to be
functionally significant in their cohort of patients. My data showing genetic synergy between Dll4
and Fgf8 pathways serves as a potential model to study compound heterozygosity in DORV.
While the more severe phenotype observed in Islet1-Cre mediated Dll4 mutants may be due to
more widespread gene loss, it could also have resulted from compound heterozygosity, given
that the Islet1-Cre line I used is a knock-in and, therefore, a functional Islet1 heterozygote. The
incomplete penetrance of phenotypic defects in heterozygous mice may also be leveraged to
study the impact of other environmental teratogenic events in a genetically permissive
background. Thus, the mouse models I used have broad relevance for further evaluating the
impact of genetic mutations in OFT anomalies.
The spectrum of phenotypic defects observed in this mutants also bears resemblance to the
DORV spectrum seen in the clinical setting. The most severe form of defect seen with Dll4
homozygous loss in either cre background is not viable, and as such, could explain the lack of
Dll4 homozygous mutations in the clinical setting. The milder forms of defects seen with
heterozygous loss of Dll4 in Islet1-Cre background or the Dll4/Fgf8 double heterozygotes in
Mef2c-AHF-Cre background are highly reminiscent of the tetralogy-type DORV or tetralogy of
Fallot encountered in children. This would suggest that one molecular mechanism underlying
DORV/Tetralogy is a later and more regional loss of proliferative signals in SHF. This allows
SHF-derived structures to develop early in gestation, however, the RV and, in particular, OFT
are hypoplastic resulting in OFT mal-alignment. The degree of mal-alignment would vary
between over-riding the septum (as in tetralogy) to originating primarily from RV with aortic-

23

mitral discontinuity (as seen in DORV). The variability in the thickness of the conus in the OFT
that I observed is also frequently encountered in clinical DORV/tetralogy and has relevance in
the surgical approach to correct these lesions. Whether and how these subtle phenotypic
variations impact long-term outcomes in children remains to be elucidated.

In summary, Dll4-mediated Notch signaling plays a crucial in role in early SHF progenitor cell
proliferation, primarily via regulation of Fgf8 expression. Dll4 expression is required to maintain
an adequate pool of SHF cells that contribute to the RV and OFT in the developing heart. Loss
of Dll4 results in a spectrum of OFT abnormalities. In their most severe forms, there is extreme
cardiac under-development and early embryonic lethality. Milder forms represent clinically
relevant CHD and, apart from providing a molecular mechanism for such clinical phenotypes,
also provide a platform to study more complex oligogenic inheritance patterns.

24

25

Fig 4.1
Dll4 expression in SHF is required for appropriate development of SHF-derived RV
and OFT.
Dll4 expression was conditionally knocked out in SHF progenitor cells using Islet1- or Mef2c-
mediated cre expression. H&E stained transverse sections of E14.5 embryos show a
normally developed heart in cre-negative littermate controls (A, B). Dll4 homozygous
knockout driven by Mef2c-AHF-Cre (Mef2c-Cre,Dll4
F/F
) (A’ and B’) and Dll4 heterozygous
knockout driven by Islet1-Cre (Islet1-Cre,Dll4
F/wt
) (A’’ and B’’) show a large ventricular septal
defect (arrow in A’ and A’’) and double outlet right ventricle (arrowhead in B’ and B’’). H&E
stained transverse sections of E10.5 control (C, H) and Mef2c-Cre,Dll4
F/F
mutant (C’, H’)
embryos demonstrate hypoplastic RV and a foreshortened and paucicellular OFT (asterix in
H’) in mutants. SHF-derived structures were identified in developing heart by lineage tracing
using the R26RLacZ mice crossed into Mef2c-AHF-Cre line. X-gal stained whole mount,
transverse and sagittal sections of E10 control (D, E, I, J) and mutant (D’, E’, I’, J’) embryos
confirm hypoplastic RV and shorter and narrower OFT. Area (Mean, SEM) of the LacZ
positive RV in whole mount (6 control and 7 mutant) embryos (F) and LacZ-positive
ventricular wall within the entire ventricular wall was measured and normalized to control
embryo in transverse sections (57 control and 50 mutant sections, G). This shows a 50%
reduction in size of SHF-derived RV in mutants (p<0.0001 by 2-tailed t-test) by both
methods. Length (Mean, SEM) of LacZ-positive OFT in whole mount embryos (6 control and
7 mutant, K) and LacZ-positive OFT normalized to control embryo in sagittal sections (10
control and 9 mutant, L) was measured. This shows a 40-50% reduction in SHF-derived
OFT length in mutants (p<0.0001 by 2-tailed t-test). Table in M indicates number and
phenotypes of embryos recovered amongst the different genotypes shown. First two rows
denote the number of embryos recovered, the percentage recovery and the expected
percentage recovery based on Mendelian inheritance.
LV, Left Ventricle; OFT, Outflow Tract; RV, Right Ventricle; A, Aortic valve, P, Pulmonary
valve, arrow - VSD
Whole mount magnification: x30 (D, D’, I, I’); Scale Bars: 150µm (C, C’, H, H’, E, E’, J, J’),
300µm (A-B’’)

26

27

Figure 4.2
Knockout of Dll4 in SHF cells leads to reduced size of SHF-derived RV and OFT.
Whole mount examination of an E9 heterozygous embryo (Dll4
wt/Lacz
) (A) and a Dll4 null
mutant (Dll4
LacZ/Lacz
) (A’) showing severely underdeveloped RV and OFT in the mutants in
addition to an arrested dorsal aorta (DA). Tissue-specific knockout of Dll4 was confirmed by
Dll4 staining in transverse sections of E9.5 Mef2c-AHF-Cre,Dll4
F/F
mutant embryos (B). Dll4
expression is lost in the pharyngeal mesoderm region (B, and higher magnification of upper
boxed area of B in the boxed region of B’), as well as the majority of the RV (B, and higher
magnification of lower boxed area of B in B’’). Dll4 expression is still preserved in the first-
heart field derived LV (B, and higher magnification of lower boxed area of B in B’’). Whole
mount examination of E10.5 control (C) and Islet1-Cre mediated Dll4 homozygous knockout
(Islet1-Cre,Dll4
F/F
) (C’) embryos. Mutant embryos show lack of adequate-sized RV (asterisk
in C’) and severely foreshortened OFT compared to control. H&E stained transverse
sections of E9.5 embryos show a normally developed RV (D) and elongated OFT (F) in
littermate controls. Dll4 homozygous knockout driven by Mef2c-AHF-Cre (Mef2c-Cre,Dll4
F/F
)
leads to significantly smaller RV (D’) and a severely foreshortened OFT (F’). Area (Mean,
SEM) of the right ventricular wall was measured and normalized to control embryo in
transverse sections (138 control and 132 mutant, E). This shows a 40% reduction in size of
RV in mutants (p<0.0001, by 2-tailed t-test) compared to controls. Length (Mean, SEM) of
the OFT was measured and normalized to control embryo in 12 control and 9 mutant
transverse sections (G). This also shows a 40% reduction in the OFT length in mutants
(p<0.0001, by 2-tailed t-test) compared to controls. Whole mount examination of E10.5
control (H) and Mef2c-Cre,Dll4
F/F
mutant (H’) embryos. Mutant embryos show hypoplastic
SHF-derived RV (elliptical area in H’ compared to H).
LV, Left Ventricle; OFT, Outflow Tract; RV, Right Ventricle; DA, Dorsal Aorta
Whole mount magnification: x30 (A, A’, C, C’, H, H’); Scale Bars: 100µm (B’,B’’), 150µm
(D,D’,F,F’), 250µm (B)

28

Figure 4.3
Knockout of Dll4 in SHF cells leads to DORV with an obligatory VSD by E14.5.
Serial H&E stained transverse sections of E14.5 embryos of cre-negative littermate controls
(A), Dll4 homozygous knockout driven by Mef2c-AHF-Cre (Mef2c-Cre,Dll4
F/F
) (B) and Dll4
heterozygous knockout driven by Islet1-Cre (Islet1-Cre,Dll4
F/wt
) (C) show appropriate
ventricular septum and outflow tract alignment in the control and DORV phenotype in both
mutants.
LV, Left Ventricle; OFT, Outflow Tract; RV, Right Ventricle; A, Aortic valve, P, Pulmonary
valve, arrow - VSD
300µm (A1-C36)

29

30

Figure 4.4
Dll4 expression is required for SHF cell proliferation to maintain an adequate
progenitor cell pool.
SHF cells were lineage traced by crossing the R26RLacZ mice into Mef2c-Cre line.
Transverse and sagittal sections of control (A, A’, D, D’) and Mef2c-AHF-Cre,Dll4
F/F
mutant
(B, B’, E, E’) E10 embryos were x-gal stained. LacZ-positive area (Mean, SEM) within the
pharyngeal mesodermal region (boxed region) was measured in transverse sections (38
control and 55 mutant) and normalized to control embryo (C). Mutants demonstrate a 67%
reduction in the SHF cell progenitor pool size compared to the controls (p < 0.0001 by 2-
tailed t-test). LacZ-positive area (Mean, SEM) in SHF region (boxed region) was measured
in sagittal sections (109 control and 89 mutant) and normalized to control embryo (F).
Mutants demonstrate a 50% reduction in the SHF cell progenitor pool size compared to the
controls (p < 0.0001 by 2-tailed t-test). Transverse sections of E9.5 control (G) and Mef2c-
AHF-Cre,Dll4
F/F
mutant (H) embryos were co-stained for Islet1 and pHH3 expression to study
SHF proliferation. Higher magnification of boxed area in G and H are shown as Islet1
expression (G’, H’), pHH3 expression (G’’, H’’) and merged image (G’’’, H’’’). Islet1 and
pHH3 double-positive cells and cells positive for pHH3, but negative for Islet1 were counted
separately in 21 control and 23 mutant fields within the boxed regions of G’’’ and H’’’ (I,
Mean, SEM) showing a 51% reduction in proliferating SHF cells in mutants compared to
controls (p<0.0001 by 2-tailed t-test) whereas there was no difference in proliferating non-
SHF cells (p>0.05). Transverse sections of E10.5 control (J) and Mef2c-AHF-Cre,Dll4
F/F
mutant (K) embryos were co-stained for Islet1 and TUNEL expression to study SHF
apoptosis. Higher magnification of boxed area in J and K is shown as Islet1 expression (J’,
K’), TUNEL expression (J’’, K’’) and merged image (J’’’, K’’’). Double-positive cells were
counted in 21 control and mutant fields each within the boxed region of J’’’ and K’’’ (L, Mean,
SEM) showing an 11-fold increase in apoptosis in SHF in mutants compared to controls
(P<0.0001 by 2-tailed t-test).
Scale Bars: 100µm (G’-H’’’, J’-K’’’), 150µm (A-B’, D-E’), 250µm (G, H, J, K)

31

Figure 4.5
Dll4 expression in SHF cells is required for SHF progenitor cell proliferation.
Transverse sections of E10.5 control (A) and Mef2c-AHF-Cre,Dll4
F/F
mutant (B) embryos
were co-stained for Islet1 and pHH3 expression to study SHF proliferation. Higher
magnification of boxed area in A and B are shown as Islet1 expression (A’, B’), pHH3
expression (A’’, B’’) and merged image (A’’’, B’’’). Islet1 and pHH3 double-positive cells and
cells positive for pHH3, but negative for Islet1 were counted separately in 28 control and 26
mutant fields within the boxed regions of A’’’ and B’’’ (C, Mean, SEM) showing a 72%
reduction in proliferating SHF cells in mutants compared to controls (p<0.0001 by two-tailed
t-tests), whereas there was no difference in proliferating non-SHF cells (p>0.05).
Scale Bars: 100µm (A’-B’’’), 250µm (A,B)

32

33

Figure 4.6
Dll4 expression in SHF cells is required to maintain expression of key SHF-related
proteins.
Transverse sections were evaluated for Mef2c and Fgf8 transcript expression at E9 in
control (A) and Mef2c-AHF-Cre,Dll4
F/F
mutant (B) by RNAscope. Higher magnification of
boxed area in A and B are shown as Mef2c expression (A’, B’), Fgf8 expression (A’’, B’’),
and merged image (A’’’, B’’’) to demonstrate the reduced expression of Fgf8 transcripts in
the mutants compared to the controls in the PM. Similarly, transverse sections were
evaluated for Mef2c and Fgf10 transcript expression at E9 in control (C) and Mef2c-AHF-
Cre,Dll4
F/F
mutant (D). Higher magnification of boxed area in C and D are shown as Mef2c
expression (C’, D’), Fgf10 expression (C’’, D’’), and merged image (C’’’, D’’’) to demonstrate
that the PM in mutants have decreased expression of Fgf10 transcripts. Transverse sections
of control (E) and Mef2c-AHF-Cre,Dll4
F/F
mutant (F) E9.5 embryos were co-stained for Islet1
and Fgf8 protein expression. Higher magnification of boxed areas in E and F are shown as
Islet1 expression (E’, F’), Fgf8 expression (E’’, F’’) and merged image (E’’’,F’’’) showing
reduced expression of Fgf8 in SHF region. Transverse sections of control (G) and Mef2c-
AHF-Cre,Dll4
F/F
mutant (H) E11.5 embryos were stained for Hand2 protein expression.
Higher magnification of boxed areas in G and H show the RV and LV in control (G’,G’’) and
mutant (H’,H’’). Hand2 expression is lost in the mutant RV compared to controls. There is no
change in the low basal level expression seen in LV.
LV, Left Ventricle; RV, Right Ventricle
Scale Bars: 50µm (E’-F’’’), 100µm (A’-D’’’,E, F, G’-H’’), 250µm (A-D, G-H)

34

Figure 4.7
Impact of loss of Dll4 in SHF on expression of SHF-specific molecules.
Transverse sections were evaluated for Mef2c and Fgf8 transcript expression at E10 in
control (A) and Mef2c-AHF-Cre,Dll4
F/F
mutant (B) by RNAscope. Higher magnification of
boxed area in A and B are shown as Mef2c expression (A’, B’), Fgf8 expression (A’’, B’’),
and merged (A’’’, B’’’) to demonstrate the reduced expression of Fgf8 transcripts in the
mutants compared to the controls in the PM. Transverse sections of control (C, E) and
Mef2c-AHF-Cre,Dll4
F/F
mutant (D, F) E11.5 embryos were stained for BMP4 (C, D), and
Mlc2v (E, F) protein expression. C’, D’, E’, F’ are magnified views of boxed areas in C, D, E
and F respectively. The expression levels of these molecules were comparable between
controls and mutants. Transverse sections of control (G, H , I and J) and Mef2c-AHF-
Cre,Dll4
F/F
mutant (K, L, M and N) E10.5 embryos were stained for Semaphorin3C
expression. Higher magnification of boxed areas in G, H, I, J, K, L, M and N are shown in G’,
H’, I’ J’, K’, L’, M’ and N’ demonstrating similar expression levels and regionalization of
Semaphorin3C in the OFT of mutants compared to controls.
Scale Bars: 100µm (A’-N’), 250µm (A-N)

35

36

Figure 4.8
Dll4-mediated notch signaling regulates Fgf8 expression in SHF.
Schematic representation (A) of the mouse chromosome 10 around region of the Fgf8 gene
(E - denotes exon). Putative RBPjk binding sites are indicated with asterisk. Constructs
cloned for luciferase assay are shown as black boxes. Promoter 3 and Enhancer 2 were
used as negative controls. 293T cells were treated with DAPT to quench basal Notch
activity. They were then transfected with various luciferase expression vectors with (empty
bars) or without NICD (solid bars) expression vector. Luciferase activity was measured in
triplicate wells (Mean, SEM) 24h later with 8 experimental repeats (B). The experiment was
then repeated three times in triplicate after mutating the putative RBPjk binding site of
Promoter 1. Mutation of putative binding sites led to loss of luciferase activity (C). Thoracic
regions were dissected in control (D-D’’) and Mef2c-AHF-Cre,Dll4
F/F
mutant (E-F’’) embryos
at E9.5 and cultured in vitro. Mutant organs were cultured with (F-F’’) or without (E-E’’)
exogenous recombinant Fgf8. Sections were then co-stained for Islet1 and pHH3 expression
to study SHF proliferation. Representative images are shown as Islet1 expression (D’, E’,
F’), pHH3 expression (D’’, E’’, F’’) and merged image (D, E, F). Double-positive cells were
counted in multiple fields (23 untreated control, 23 Fgf8 100ng/µL control, 40 Fgf8 500ng/µL
control, 37 Fgf8 untreated mutant, 7 Fgf8 100ng/µL mutant, and 14 Fgf8 500ng/µL mutant
sections, G, Mean, SEM) showing a significant reduction in SHF proliferation in mutant
organs compared to control (p < 0.0001 between Fgf8 untreated control and mutant, p >
0.05 between Fgf8 untreated controls and Fgf8 treated mutants by two-tailed t-tests). For
quantification purposes, the boxed regions in D’, E’ and F’ were used as area occupied by
SHF progenitor cells. Exogenous Fgf8 supplementation had no significant impact on control
embryos, but fully rescued proliferation defect seen in mutant embryos. Compound
heterozygotes were analyzed by H&E staining of transverse sections of E14.5 embryos to
demonstrate genetic synergy between Dll4-mediated Notch and Fgf8 signaling in SHF
maturation. Cre-negative control embryos showed fully septated ventricles (H) and an aortic
valve normally aligned over the left ventricle (H’). Heterozygous knockdown of Dll4 driven by
Mef2c-AHF-Cre (Mef2c-Cre,Dll4
F/wt
) also demonstrated normal phenotype (I, I’).
Heterozygous knockdown of Fgf8 driven by Mef2c-AHF-Cre (Mef2c-Cre,Fgf8
F/wt
) showed a
low incomplete penetrance of cardiac defects. The majority of the embryos showed normal
phenotype (J, J’). 14% of the embryos had a shallow VSD (arrow in K) and a slightly mal-
aligned aorta mildly over-riding the ventricular septum (arrowhead in K’). Double
heterozygous knockdown of Dll4 and Fgf8 driven by Mef2c-AHF-Cre (Mef2c-Cre,
Dll4
F/wt
,Fgf8
F/wt
) showed high penetrance of DORV. 83% of the embryos studied showed
VSD (arrow in L) and a prominent over-riding of aorta with greater than 50% aorta arising
from the RV (arrowhead in L’). Table in M indicates number and phenotypes of embryos
recovered amongst the different genotypes shown. First two rows denote the number of
embryos recovered, the percentage recovery and the expected percentage recovery based
on Mendelian inheritance.
Scale Bars: 50µm (D-F’’), 300µm (H-L’)

37

Figure 4.9
Dll4-mediated notch signaling regulates Fgf8 expression in SHF.
HeLa cells were treated with DAPT to quench basal Notch activity. They were then
transfected with various luciferase expression vectors with (empty bars) or without (solid
bars) NICD expression vector. Luciferase activity was measured in triplicate wells (Mean,
SEM) 24h later (A). 293T cells were treated with SAHM1, a competitive inhibitor of
downstream Notch assembly, to quench basal Notch activity. They were then transfected
with various luciferase expression vectors with (empty bars) or without (solid bars) NICD
expression vector. Luciferase activity was measured in triplicate wells (Mean, SEM) 24h later
in 4 experimental replicates (B). Thoracic region was dissected in control (C, D) and Mef2c-
AHF-Cre,Dll4
F/F
mutant (E) embryos at E9.5 and cultured in vitro with varying doses of
exogenous recombinant Fgf8. Sections were then co-stained for Islet1 and pHH3 expression
to study SHF proliferation. Representative images are shown as Islet1 expression (C’, D’,
E’), pHH3 expression (C’’, D’’, E’’) and merged image (C, D, E). See quantification in Fig 5F.
For quantification purposes, the boxed regions in D’, E’ and F’ were used as the region
occupied by SHF progenitor cells.
Scale Bars: 50µm (C-E’’).

Large portions of this wok (Chapter 3) is published in Stem Cells and Development (De Zoysa et al., Stem
Cells Dev 24 April 2021; Epub ahead of print: scd.2021.0058)
38

Chapter 5
Haploinsufficiency of Dll4 in Second Heart Field cells is the
ideological mechanism for the cardiac defects seen in
Adams-Oliver syndrome

5.1 Introduction

Heterozygous loss of function mutation in Delta-like ligand-4 (Dll4) is an important cause of
Adams-Oliver syndrome. Cardiac defects, in particular outflow tract (OFT) alignment defects,
are observed in about one-fourth of patients with this syndrome. The mechanism underlying this
genotype-phenotype correlation has not yet been established. Dll4-mediated Notch signaling is
known to play a crucial role in second heart field (SHF) progenitor cell proliferation. I
hypothesized that depletion of SHF progenitor pool of cells due to partial loss of Dll4 is
responsible for the OFT alignment defects seen in Adams-Oliver syndrome. To demonstrate
this, I studied Dll4 expression by murine SHF progenitor cells around E9.5, a crucial time-point
in SHF biology. I used SHF-specific (Islet1-Cre) conditional knockout of Dll4 to bypass the early
embryonic lethality seen in global Dll4 heterozygotes. Dll4-mediated Notch signaling is critically
required for SHF proliferation such that Dll4 knockout results in 33% reduction in proliferation
and a 4-fold increase in apoptosis in SHF cells, leading to a 56% decline in size of the SHF
progenitor pool. Reduction in SHF cells available for incorporation into the developing heart
leads to underdevelopment of SHF-derived RV and OFT. Similar to the clinical syndrome, 32%
of SHF-specific Dll4 heterozygotes demonstrate foreshortened and misaligned OFT, resulting in
Double Outlet Right Ventricle (DORV). This murine model provides a molecular mechanism to
explain the cardiac defects observed in Adams-Oliver syndrome and establishes a novel clinical
role for Dll4-mediated Notch signaling in SHF progenitor biology.

5.2 Haploinsufficiency of Dll4 in SHF disrupts OFT alignment

Global knockout of Dll4 is embryonically lethal by E9.5 due to vascular maturation arrest
(Duarte, 2004). Interestingly, in the same study, haploinsufficiency of Dll4 was also shown to
disrupt vascular maturation with variable penetrance. In the context of SHF biology, my own
previous work has shown that conditional knockout of Dll4 expression bypasses the early
lethality, allowing evaluation of cardiac development. SHF-specific Dll4 knockout results in a
spectrum of defects extending from severe lack of SHF-derived RV and OFT to a fully penetrant
DORV phenotype (De Zoysa et al., 2020). I, therefore, wanted to study the impact of partial loss
of Dll4 in SHF on OFT development. To that end, I evaluated heterozygous Dll4 mutation in the
Islet1 background (Islet1-Cre,Dll4
F/wt
). Embryos were recovered in expected Mendelian numbers
indicating lack of embryonic lethality. I studied the cardiac phenotype at E14.5 in 28 mutant
embryos compared to 40 cre-negative littermate controls (Fig 5.1). 19 out of 28 mutants had a
normal cardiac phenotype. The remaining nine mutants (32%, Fig 5.1D) demonstrated OFT
alignment defects. There was a gradation noted in the severity of the alignment defect. Some
embryos demonstrated a prominent VSD (Fig 5.1B3) and the aortic valve arose entirely from the

39

right ventricle (Fig 5.1 B4 vs. A4) resembling the clinical DORV. Other mutants displayed a
much shallower VSD (Fig 5.1 C3 compared to B3) and the aorta was over-riding the VSD (Fig
5.1 C4), thus having inflow from both ventricles. This lesion was reminiscent of the clinical
tetralogy-style defect. In both controls and mutants, the OFT was appropriately septated. The
pulmonary valve arose more cranially (indicating a normal sub-pulmonary conus) and exited the
right ventricle (Fig 5.1 A5, B5 and C5). There was normal connection between the posterior
outflow vessel to the systemic circulation and the cranial, anterior vessel to the pulmonary
circulation, indicating appropriate rotation. Thus, this murine model phenocopies the variably
penetrant alignment defect seen in AOS patients. A smaller subset of SHF cells express Mef2c
as they exit the mesoderm to enter the developing heart. Heterozygous loss of Dll4 in Mef2c
expressing SHF cells does not result in a cardiac phenotype (Fig 5.1D) indicating that the timing
and degree of Dll4 loss plays a role in the ultimate cardiac phenotype. Because Islet1- driven
cre recombinase may also be active in a subset of NCC, I wanted to confirm that the observed
defects in Islet1-Cre mice were not neural-crest driven. To that end, I studied the effect of Dll4
loss in Wnt1-Cre mice. Both heterozygous (Fig 5.1 E1,E2) and homozygous (Fig 5.1 F1,F2) loss
of Dll4 in Wnt1-expressing NCC does not result in a discernable cardiac phenotype, consistent
with lack of expression of Dll4 in NCC.

In order to study the developmental defect resulting in the observed cardiac phenotype, I
examined heterozygous mutant embryos at E10.5 when early cardiac assembly is completed. I
used the Islet1-Cre/R26RtdT mice to label the RV and OFT and also injected India ink into RV
of wildtype embryos as a complementary technique to visualize the OFT (Fig 5.2). All embryos
demonstrated appropriate early assembly at this stage. I noticed a gradation in reduction in RV
size and OFT length in mutant embryos. Overall, India ink filled RV was 42% smaller in mutants
compared to controls (Fig 5.2 A, A’, B), while the corresponding tdT-labeled RV was 48%
smaller in mutants compared to controls (Fig 5.2 C, C’, D). Similarly, percentage of ventricular
area occupied by tdT labeled cells (indicating RV portion of the ventricles indexed for LV) was
56% reduced in mutant sections compared to control (Fig 5.2 E, E’, F). The length of the OFT
was 30% reduced in India ink labeled wholemounts of mutants compared to control embryos
(Fig 5.2 G, G’, H) and by 33% in sagittal sections of lineage-traced mutants (Fig 5.2 I, I’, J).
These data indicate that haploinsufficiency of Dll4 in SHF leads to reduction in size of SHF-
derived RV and length of SHF-derived OFT. This foreshortened OFT is unable to align itself
over the developing RV and LV, such that the ensuing phenotype is an alignment defect,
DORV.

5.3 Partial conditional loss of Dll4 expression leads to reduction in
SHF progenitor cell pool

The observed reduction in RV size and OFT length in mutants suggests that there is a reduction
in incorporation of SHF progenitor cells into the developing heart following heterozygous loss of
Dll4. This could result from a reduction in number of SHF progenitors available for incorporation
or from an inability of available SHF cells to migrate into the developing heart. To directly
address this question, I studied the size of SHF progenitor pool. I lineage-traced SHF cells
(Islet1-Cre/R26RtdT) and studied the area occupied by tdT-positive cells in the SHF
mesodermal region (Fig 5.3 A-C). Sagittal sections of mutant embryos showed a 56% reduction
in tdT-positive cells in the SHF mesoderm. This would indicate that the observed reduction in
size of SHF-derived structures in the heart results from a loss of SHF progenitors available for
incorporation into the developing heart. I then studied proliferation in lineage-traced SHF
progenitors. To this end, control and mutant embryos in Islet1-Cre/R26RtdT background were

40

stained for pHH3. Double-positive cells in the region of the pharyngeal mesoderm were counted
in controls and mutants. At E10.5 (Fig 5.3 D-F), heterozygous loss of Dll4 resulted in a 33%
reduction in proliferating SHF cells. Loss of proliferative potential in these early progenitor cells
directs them towards apoptosis, and hence, I studied apoptosis in SHF cells with TUNEL
staining. Partial loss of Dll4 resulted in a 4-fold increase in SHF progenitor cell apoptosis at
E10.5 (Fig 5.3 G-I).

5.4 Discussion

Notch signaling is an evolutionarily conserved signaling pathway that plays important roles in
cell fate specification, development, differentiation and patterning in numerous cell types in the
body. In mammals, there are four transmembrane Notch receptors (Notch 1-4) and five trans-
membrane Notch ligands (Delta-Like Ligand 1, 3, 4 and Jagged 1 and 2) (Andersson et al.,
2011). During canonical Notch signaling, a Notch receptor interacts in trans with its ligand
expressed on a neighboring cell, leading to proteolytic cleavage of the receptor and subsequent
nuclear translocation of the Notch intracellular domain (NICD) (de la Pompa and Epstein, 2012).
NICD forms a complex with CBF1/Suppressor of Hairless/LAG-1 (CSL) family of DNA binding
proteins, which activates transcription of cell-specific downstream effector molecules
(de la Pompa and Epstein, 2012). My own work and others have shown that Notch signaling
plays a variety of different roles in cardiac development, in particular, OFT maturation. Early in
heart development, Dll4 serves as the primary ligand of Notch 1 and plays a proliferative role in
progenitor cells (De Zoysa et al., 2020, D’Amato et al., 2016) similar to the pro-proliferative role
of Dll4 in other progenitor beds, such as retinal (Luo et al., 2012) or neural progenitors (Rocha
et al., 2009). Later in heart development, Dll4 expression wanes and Jagged1 takes over as the
primary Notch ligand. Jagged-mediated Notch signaling regulates subsequent cardiac
patterning (D’Amato et al., 2016) and maturation events such as epithelial-to-mesenchymal
transformation in OFT cushions (High et al., 2009).

Notch pathway mutations have been implicated in several clinical CHDs. With particular
reference to the OFT, Notch mutations have been identified in patients with bicuspid aortic valve
(McKellar et al., 2007, Garg, 2016) and aortic valve calcification (Garg, 2016). Mutations in
Jagged1 are causative in Alagille syndrome, which includes biliary malformations, and
pulmonary artery defects (Li et al., 1997). Animal studies of Dll4 have shown that there is a
dosage-sensitive requirement of Dll4 in arterial maturation, such that over half the embryos with
global Dll4 haploinsufficiency die in early gestation due to a vascular maturation arrest (Duarte,
2004). Likely due to this critical requirement of Dll4 during development, Dll4 mutations have
been infrequently reported in human diseases. Recent evidence suggests a role for Dll4 in
patients with AOS. Initial studies identified mutations in six genes in AOS: Arhgap31, Dock6,
Eogt, Rbpj, Notch1, and Dll4 (Shaheen et al., 2013). Four of these genes (Eogt, Rbpj, Notch1,
and Dll4) play a critical role in the canonical Notch pathway. Whereas autosomal recessive
mutations in Eogt lead to AOS (Shaheen et al., 2013), the other three genes are thought to play
an autosomal dominant role (Stittrich et al., 2014). In a large study of targeted resequencing of
Dll4 gene with a custom enrichment panel in 89 independent families, nine heterozygous loss of
function mutations in Dll4 were identified (Meester et al., 2015). This included two nonsense and
seven missense mutations, all of which were thought to be critical for maintaining structural
integrity of the protein. An additional missense mutation in Dll4 that interferes with its binding to
Notch1 has also been described in a sporadic Japanese patient (Nagasaka et al., 2017). Yet, no
genotype-phenotype correlations have been made in AOS to date (Meester et al., 2015).

41

Almost one quarter of patients with AOS harbor a CHD, with the most common being outlet
VSD or alignment defect, primarily TOF (Hassed et al., 2017), both defects that can result from
derangements in SHF contribution to the developing heart. Based on the current understanding
of Dll4 biology, I submit that the cardiac defects seen in AOS are best explained by a somatic
mutation acquired in mesodermal SHF progenitor cells after the initial steps in embryogenesis
are effectively completed. I have previously shown that knockout of Dll4 in SHF results in RV
and OFT under-development and early embryonic lethality (De Zoysa et al., 2020). The few
embryos that survive to mid-gestation display DORV. I now wanted to model heterozygous Dll4
loss in cardiac OFT-specific progenitor cells to establish a laboratory model of CHD seen in
AOS. Islet1 is expressed early in heart development and globally by SHF progenitor cells
destined to contribute to the developing heart. We, therefore, generated heterozygous
conditional loss of Dll4 in Islet1-expressing SHF cells. Haploinsufficiency of Dll4 had
demonstrable effects on SHF, with a significant reduction in SHF proliferation. SHF progenitors
incapable of proliferation subsequently undergo apoptosis, ultimately leading to a reduction in
pool of SHF progenitors. I have previously shown that Dll4-mediated Notch signaling regulates
Fgf8 and Fgf10 expression in SHF, thereby affecting SHF proliferation (De Zoysa et al., 2020).
Loss of SHF progenitors available for incorporation into the developing heart leads to an
underdeveloped RV and foreshortened OFT, leading to its misalignment. The aortic alignment
defects seen in this model resemble the spectrum of defects seen in the clinic. In milder cases,
the aorta rides across the VSD as seen in TOF, implying inadequate displacement to the LV. In
more extreme cases, the aorta arises completely from the RV resembling TOF-style DORV.
Thus, this model serves as the first molecular demonstration of the genotype-phenotype
correlation of CHD observed in AOS.

The incomplete penetrance of CHD phenotype is consistent with Dll4 biology. The penetrance
of vascular defects in global Dll4 heterozygotes was shown to be strain-dependent (Benedito
and Duarte, 2005), implying that there is likely epistatic regulation of Dll4 function, such that
some individuals are able to overcome the impact of partial loss of Dll4. This mouse model had
a greater prevalence of CHD than reported in the clinic. One potential explanation is that the
Islet1-Cre mice used in this model is a cre recombinase knock-in, in effect acting as
heterozygous loss of Islet1. This oligogenic mutation may underlie the increased penetrance
observed. Alternatively, biological variability driven by the extent and degree of Dll4 loss could
also be in play. Heterozygous loss of Dll4 driven by Mef2c-AHF-Cre, a more restricted SHF
marker, had no phenotype lending further credence to this argument. Lastly, this specific mouse
model does not lend itself to studying associated defects in other organ systems. It has been
suggested that the limb and scalp defects observed in AOS are a result of a more general
vasculopathy (Stittrich et al., 2014). It is well established that Dll4 plays a crucial role in vascular
development and, further, that common mesodermal progenitors drive both vascular and
cardiac development. Future studies aimed at establishing a laboratory model that displays the
multi-organ system defects in AOS should shed more light into these aspects of the disease
process.

In summary, this work has established a laboratory model to explain the CHD observed in AOS
due to heterozygous loss of function of Dll4. my data indicate that a later and more regional loss
of Dll4 in SHF progenitors inhibits their proliferation leading to reduced SHF incorporation in the
developing OFT. A foreshortened OFT is incapable of proper alignment and, therefore, results
in the cardiac alignment defects observed in the clinic. This model would also imply that the RV
in these mutants shares these molecular defects. As such, further studies would help us
understand whether and how this abnormal RV would behave in the long-term in patients who
undergo surgical management of OFT CHD in AOS.

42

Fig 5.1
Haploinsufficiency of Dll4 in SHF results in misalignment of OFT.
Heterozygous conditional loss of Dll4 expression in SHF was achieved using Islet1 -
mediated cre expression. H&E-stained transverse sections of E14.5 cre-negative littermate
control embryos and Dll4 heterozygous mutants (Islet1-Cre,Dll4
F/wt
) show properly
developed tricuspid (T) (A1, B1 and C1) and mitral (M) (A2, B2 and C2) valves. A third of the
mutants demonstrated an outlet VSD (Arrows in B3 and C3) compared to intact septum in
controls (A3). The aortic valve (A) arises entirely from the RV in a subset of mutant embryos
(B4) and over-rides the VSD in the other (C4). In both control and mutant embryos, the
pulmonary valve (P) exits the RV (A5, B5 and C5). D indicates the number and phenotypes
of embryos recovered amongst the different genotypes mentioned. Dll4 heterozygous and
homozygous mutants in Wnt1 background (Wnt1-Cre,Dll4
F/wt
, E1, E2 and Wnt1-Cre,Dll4
F/F
,
F1, F2) demonstrate normal heart phenotype at E14.5.
M, Mitral Valve ; T, Tricuspid Valve ; A, Aortic Valve ; P, Pulmonary Valve
Scale bar - 300µm (A1-C5, E1-F2)

43

Fig 5.2
Haploinsufficiency of Dll4 in SHF leads to reduced size of RV and foreshortened OFT.
Heterozygous conditional loss of Dll4 expression in SHF progenitor cells was achieved using
Islet1-mediated cre expression. Whole mount examination at E10.5 of India ink injected
control wildtype embryos (A) and Islet1 lineage traced control embryos (C) compared to their
respective Dll4 heterozygous mutants (A’ and C’) demonstrates a hypoplastic RV in the
mutants. Area (Mean, SEM) of the RV in India ink injected whole-mount embryos (2 control
and 5 mutant, B) and lineaged traced tdT-positive RV (2 control and 2 mutant, D) was
compared to area of the corresponding LV and normalized to control embryos. This shows
an almost 50% reduction in the RV/LV ratio in mutants (p<0.005 (B) and p<0.05 (D) by
unpaired two-tailed t-test). tdT-positive RV area within the entire ventricular area (10 control,
E and 14 mutant transverse sections, E’) was measured and normalized to control embryo
sections (F). This shows a 56% reduction of the RV in mutants compared to controls
(p<0.0001 by unpaired two-tailed t-test). Length (Mean, SEM) of India ink injected OFT was
measured in whole mount embryos (8 control, G and 14 mutant, G’) and Islet1 lineage-
traced tdT-positive sagittal sections (7 control, I and 5 mutant, I’) and normalized to control
embryos. This shows a 30-33% reduction in OFT length in mutants (p<0.005 (H) and
p<0.0001 (J) by unpaired two-tailed t-test).
Scale Bars: 250µm (E, E’, I, I’)

44

45

Fig 5.3
Dll4 expression is required for SHF cell proliferation to maintain an adequate
progenitor cell pool.
SHF cells were lineage traced by crossing R26RtdT mice into Islet1-Cre line. Sagittal
sections of control (A, A’) and Dll4 heterozygous mutant (Islet1-Cre,Dll4
F/wt
) (B, B’) embryos
were evaluated at E10.5. The tdT-positive area (Mean ± SEM) within the SHF mesodermal
region (boxed) was normalized to control embryos (C). Mutants demonstrated a 56%
reduction in the SHF cell progenitor pool size compared to controls (p<0.05). Transverse
sections of Islet1 lineage-traced E10.5 control (D) and Islet1-Cre,Dll4
F/wt
mutant (E) embryos
were stained for pHH3 expression to study SHF proliferation in SHF cells. Higher
magnification of boxed area in D and E are shown as lineage-traced Islet1 expression
through tdT (D’, E’), pHH3 expression (D’’, E’’) and merged image (D’’’, E’’’). Islet1 and pHH3
double-positive cells were counted in 5 control and 5 mutant fields (F, Mean, SEM) showing
a 33% reduction in proliferating SHF cells in mutants (p<0.005). Similarly transverse
sections of Islet1 lineage-traced E10.5 control (G) and Islet1-Cre,Dll4
F/wt
mutant (H) embryos
were stained by TUNEL to study SHF apoptosis. Higher magnification of boxed area in G
and H are shown as lineage-traced Islet1 expression through tdT (G’, H’), TUNEL staining
(G’’, H’’) and merged image (G’’’, H’’’). Double-positive cells were counted in 4 control and 6
mutant fields within the boxed region (I, Mean, SEM) showing a 4-fold increase in apoptosis
in SHF in mutants (p<0.005).
Scale Bars: 100µm (A’, B’, D’-E’’’, G’-H’’’), 250µm (A, B, D, E, G, H)

46

Chapter 6
Delta-like ligand-4 expressed on arterial endothelial cells derived from
second heart field progenitors is crucially required for development of
the aortic arch

6.1 Introduction

Delta-like ligand-4 (Dll4) was originally identified as an arterial endothelial-specific Notch ligand
that plays an important role in arterial specification and blood vessel maturation. The aortic arch
endothelium is a unique cell layer derived from Second Heart Field (SHF) mesodermal
progenitors that express cardiac-specific markers. I show that, unlike in SHF-derived
endocardium and myocardium, Dll4 expression persists in SHF-derived arterial endothelial cells.
Using SHF-specific conditional deletion of Dll4 to circumvent the early lethality from Dll4
knockout, I studied the biological role of Dll4 in aortic arch development. As SHF progenitors
transition from their progenitor state to acquire an endothelial cell fate, Dll4-mediated Notch
signaling switches from providing proliferative to maturation cues. Dll4 expression maintains
arterial identity in the pharyngeal arch arteries and plays a critical role in the maturation and re-
organization of the 4
th
pharyngeal arch artery, in particular. Dll4 expression is required for
endothelial cell interaction with smooth muscle cells, which in this vascular bed are derived from
ectodermal cardiac neural crest. Haploinsufficiency of Dll4 in SHF leads to highly penetrant
aortic arch artery abnormalities, similar to those observed in the clinic, primarily resulting from
aberrant reorganization of bilateral 4
th
pharyngeal arch arteries. In summary, the above data
demonstrate that SHF progenitors that assume an arterial endothelial fate continue to express
Dll4 and the resulting Dll4-mediated Notch signaling transitions from an early proliferative to a
later maturation role in the development of the aortic arch.

6.2 Dll4 expression in SHF progenitors is required for appropriate
development of the aortic arch

The 3
rd
, 4
th
and 6
th
PAA are derived from SHF progenitors. Coronal sections of Mef2c-AHF-
Cre,R262RtdT (Fig 6.1 A, A’) and Islet1-Cre,R262RtdT (Fig 6.1 C, C’) embryos at E10.5 confirm
that PAA endothelial cells express tdT and are hence SHF-derived.

I also studied the Dll4 expression on the SHF derived pharyngeal arch artery endothelium by
costaining Dll4 in transverse sections of SHF-lineage traced E10.5 embryos of both Mef2c-AHF-
Cre,R262RtdT (Fig 6.1 B-B’’’) and Islet1-Cre,R262RtdT (Fig 6.1 D-D’’’) mice. These sections
demonstrate that Dll4 expression is indeed seen in SHF-derived 4
th
PAA endothelial cells.

47

These endothelial cells begin to develop a smooth muscle coat starting at E12.5. Using Wnt1-
Cre,R26RtdT E13.5 embryos, I show that this smooth muscle coat is derived from cardiac
neural crest cells (Fig 6.1 E-E’”).

To study the biological role of Dll4 in SHF progenitors that form the PAA, I employed SHF-
specific Cre-mediated conditional knock-down of Dll4 expression using both Islet1-Cre and
Mef2c-AHF-Cre lines. Homozygous Dll4 knockdown in Islet1-Cre background resulted in very
early embryonic lethality as I have previously shown (De Zoysa et al., 2020), precluding
evaluation of PAA phenotypes. All but one Mef2c-AHF-Cre,Dll4
F/F
embryos recovered at E14.5
demonstrated arch artery defects (Fig 6.1 M). Interestingly, haploinsufficiency of Dll4 in SHF
cells also resulted in highly penetrant arch artery defects in both backgrounds (Fig 6.1 M).

Overall, loss of Dll4 led to a variety of arch artery defects (Fig 6.1M). An isolated right-sided
aortic arch (Fig 6.1 F, F’) was seen in 3 (6.7%) mutant embryos, indicating that perturbation of
PAA re-organization impacts eventual resorption of right dorsal aortic arch in a small subset of
embryos. The majority of embryos revealed 4
th
PAA-related defects. These include a cervical
aortic arch, wherein the aortic arch is derived from the 3
rd
PAA due to inappropriate resorption of
the 4
th
PAA. The aortic arch is observed more cervically in these embryos around the region of
the thymus gland as opposed to more caudally by the atrial appendages in controls (Fig 6.1 G,
G’). Another common anomaly was aberrant right subclavian artery, wherein this artery arose
from the distal arch/proximal descending aorta and traveled posterior to the trachea and
esophagus instead of from the innominate artery, which is the first branch of the aortic arch (Fig
6.1 H, H’). This phenotype results from inappropriate resorption of the right 4
th
PAA. Less
frequently, the mirror image phenotype of an aberrant left subclavian artery was seen in
embryos with right-sided aortic arch (Fig 6.1 I, I’) due to inappropriate resorption of the left 4
th

PAA. Interrupted aortic arch (IAA) was seen in 11% embryos indicating loss of 4
th
PAA. I
observed both IAA type B (aortic arch disruption between the left carotid and left subclavian
arteries) (Fig 6.1 K1’-K4’) and IAA type C (aortic arch disruption between the innominate and left
carotid artery) (Fig 6.1 K1’’-K4’’). Interestingly, 4 of these 5 mice also had aberrant right or left
subclavian arteries indicating that both left and right 4
th
PAA were inappropriately resorbed in
these embryos. In 8 (18%) embryos, an aberrant ductus arteriosus (PDA) was seen defined as
a PDA on the opposite side of the aortic arch (Fig 6.1 J, J’). This represents a defect in
maturation of the 6
th
PAA. Seven (16%) embryos demonstrated a right sided aortic arch with
aberrant left subclavian artery (loss of left 4
th
PAA) along with a left-sided PDA (defect in 6
th

PAA) leading to the clinically well recognized vascular ring phenotype (Fig 6.1 L1’-L3’).
Interestingly, none of the mutant embryos demonstrated any carotid artery abnormalities
implying normal 3
rd
PAA maturation.

I then studied PAA phenotypes earlier in development by whole mount evaluation of India ink-
injected embryos. At E10.5, conditional SHF-Dll4 mutant embryos begin to demonstrate a
spectrum of 4
th
PAA defects. These defects manifest as complete loss (Fig 6.2 B, B’),
hypoplasia (Fig 6.2 C, C’), or inappropriate bifurcation of the 4
th
PAA (Fig 6.2 D, D’) or improper
origin of the 4
th
PAA from the 6
th
PAA (Fig 6.2 E, E’). By E12.5, mutant embryos demonstrate
their final histologic phenotypes. I was able to demonstrate right aortic arches (Fig 6.2 G, G’),
right arch with aberrant left subclavian artery (Fig 6.2 H, H’) and vascular ring (Fig 6.2 I, I”)
phenotypes at this stage.

48

6.3 Dll4 mediates vessel maturation via expression of Hey1
I sought to identify the molecular mechanisms that act downstream of Dll4-mediated Notch
signaling to regulate 4
th
PAA maturation. I stained coronal sections of control and mutant
lineage-traced embryos for CD31 expression (Fig 6.3 A-B6). At E10.5, mutant embryos
demonstrated normal sized 3
rd
PAA compared to controls (Fig 6.3 B1-3 compared to A1-3).
Interestingly, whereas the entire 3
rd
PAA was tdT-positive in control embryos (Fig 6.3 A2)
indicating that it was derived from SHF progenitors, less than half the vessel was tdT-positive in
mutants (Fig 6.3 B2). This would imply that when there is loss of SHF-derived endothelial cells,
the 3
rd
PAA is able to compensate by incorporating endothelial cells derived from non-SHF
source, thereby maintaining an appropriate caliber 3
rd
PAA. In contrast, mutant embryos
showed near complete loss of lumen of 4
th
PAA (Fig 6.3 B4-6), with a much smaller CD31-
positive vessel and no additional contribution by non-SHF-derived, tdT-negative cells. The 4
th

PAA, therefore, is entirely reliant on SHF-derived cells and is unable to compensate for loss of
these cells to maintain its caliber. This would explain the preponderance of 4
th
PAA defects
observed in mutant embryos. The majority of mutant embryos maintained 6
th
PAA size using
SHF-derived endothelial cells. I speculate that Dll4 expression is not required for pulmonary
artery maturation.

Hey1 is a known downstream target of Notch signaling. Previous reports of Hey1 knockout
embryos have shown defective development of the 4
th
PAA, similar to the phenotype I observe
in my mutants (Fujita et al., 2016). I, therefore, evaluated Hey1 expression in PAA in my
mutants. By E9.5, SHF-Dll4 mutant embryos demonstrate near complete loss of Hey1
expression in the 4
th
PAA both on the left (Fig 6.3 DL1-DL3) and right (Fig 6.3 DR1-DR3) sides.
At E10.5, control 4
th
PAA endothelial cells continued to proliferate as demonstrated by pHH3
and CD31 double-positive cells (Fig 6.3 E-E’”). In contrast, mutant 4
th
PAA had no lumen and no
proliferating endothelial cells (Fig 6.3 F-F’’’), suggesting that these arteries sustained a growth
arrest and were headed towards resorption.

I then modeled vascular plexus formation and maturation using tube formation of HUAEC in
fibrin gels (Nakatsu et al., 2003). Using this assay, I measured initial endothelial cell sprouting at
3 days and the complexity of lumenized tubes in terms of branching and anastomosis at 5 days.
In the absence of added factors there was no significant vessel sprouting from HUAEC-coated
beads (Fig 6.3G, H). VEGF (2.5ng/mL) induced robust vessel sprouting at 3 days (Fig 6.3G’)
and formation of complex branching and anastomoses by 5 days (Fig 6.3H’) as previously
reported (Cavallero et al., 2015). When HUAEC were pre-treated with Dll4 or Hey1-specific
siRNA and then coated on to the beads, early vessel sprouting appeared to be comparable in
the presence of VEGF (Fig 6.3 G”, G’”). However, these vessels failed to continue to sprout or
demonstrate any branching or anastomosis by day 5 (Fig 6.3 H”, H’”). Such a maturation arrest
led to stagnation or resorption of these nascent vessels. Thus, my in vitro results confirm that
Dll4 and Hey1 expression in arterial endothelial cells does not impact early vasculogenesis but
rather promotes subsequent maturation of the plexus induced by VEGF leading to branching
and anastomosis.

6.4 Dll4 expression in PAA is required for arterial specification and
development of smooth muscle coat

49

Dll4 is specifically expressed on arterial endothelial cells and its expression is an early event in
arterial identity (Wythe et al., 2013). I, therefore evaluated the expression of arterial and venous
markers in the 4
th
PAA of control and SHF-Dll4 mutants. By E9.5, mutant embryos
demonstrated lack of expression of arterial markers EphrinB2 (Fig 6.4 A” vs. B”) and Neuropilin1
(Fig 6.4 C” vs. D”) in 4
th
PAA compared to controls. It has been shown that in the absence of
induction of arterial markers, vessels express default venous markers. In SHF-Dll4 mutants,
loss of arterial marker expression in 4
th
PAA was associated with aberrant expression of venous
markers such as EphB4 (Fig 6.4 E” vs. F”) and Neuropilin2 (Fig 6.4 G” vs. H”). Thus, Dll4
expression in SHF progenitors is required for SHF-derived 4
th
PAA endothelial cells to express
arterial identity during maturation.

A hallmark of arterial maturation during development is acquiring a smooth muscle cell covering
over endothelial cell layer. Control embryos at E12.5 show normal caliber 4
th
PAA joining the
dorsal aortic arch (Fig 6.4I). Both the 4
th
PAA (derived from SHF progenitors) and the dorsal
aorta (at this level derived from non-SHF mesoderm) express smooth muscle actin coat (Fig
6.4I”) indicating normal arterial maturation. In contrast, the 4
th
PAA is small in caliber with a
normal dorsal aorta in mutant embryos (Fig 6.4J). Because Dll4 expression is only lost in SHF
progenitors in these mutants, only SHF-derived 4
th
PAA lacks smooth muscle coat whereas the
non-SHF derived dorsal aorta has acquired normal smooth muscle coat (Fig 6.4J”). Taken
together, my data would indicate that Dll4 expression is SHF mesodermal progenitors is
required for arterialization of SHF-derived endothelial elements and for them to obtain a smooth
muscle layer as part of the normal arterial maturation process.

6.5 Discussion

During gastrulation, angioblasts are directly specified from lateral plate mesodermal progenitors.
Arteriovenous specification of the angioblasts is genetically determined. Dll4 is a unique,
arterial-specific Notch ligand and is one of the earliest markers of arterial identity (Chong et al.,
2011). Its cognate receptors, Notch1 and 4, are subsequently expressed in the arterial
endothelium, and Dll4-mediated Notch signaling turns on the molecular program that maintains
arterial identity and facilitates maturation of the arterial vasculature. During this maturation
process, nascent arterial endothelial cells interact with smooth muscle cells that are also derived
from mesodermal progenitors. The biological role of Dll4 in arterial precursor cells directly
derived from the mesoderm has been established (Duarte, 2004, Gale et al., 2004).
Haploinsufficiency of Dll4 leads to arteriovenous specification defects and a vascular maturation
arrest resulting in early embryonic lethality. Contrarily, overexpression of Dll4 expands
arterialization of endothelial cells, which also results in failure of appropriate arteriovenous
specification and embryonic lethality (Trindade et al., 2008). Dll4 also plays an important role in
the maturation of arteries at sites of neoangiogenesis in the adult under physiologic and
pathologic conditions (Liu et al., 2012). The ascending aorta, proximal aortic arch and its
branches represent a unique vascular bed during development. Although the endothelial cells
that form these vessels are also mesodermal in origin, they do not undergo direct angioblast
specification from their progenitor state. In contrast, SHF mesodermal progenitors initially
establish a cardiogenic molecular signature with the expression of cardiac-specific molecules
such as Islet-1, Mef2c, Hand2, etc. A subset of SHF progenitors that are incorporated into the
arterial pole of the developing heart subsequently assume an endothelial fate, and begin to

50

express endothelial-specific molecular markers. My study was designed to evaluate the
expression and role of Dll4 as these cells pass through the various molecular signatures.

Notch signaling has been shown to play an important role in development, in general, and heart
development, in particular. In SHF progenitors, I have shown Dll4-mediated Notch signaling
maintains a proliferative phenotype to ensure that an adequate pool of progenitor cells is
available for incorporation into the developing heart (De Zoysa et al., 2020). As the SHF cells
incorporated into the right ventricle and OFT assume endocardial and myocardial cell fates, Dll4
expression wanes and Jagged1 expression begins to predominate. Jagged1-mediated Notch
signaling then controls maturation events such as ventricular compaction in cardiomyocytes
(D’Amato et al., 2016) and endocardial-to-mesenchymal transformation in endocardial cells that
give rise to semilunar valves (High et al., 2009). In contrast to SHF-derived myocardial and
endocardial cells, Dll4 expression persists in SHF-derived arterial endothelial cells (De Zoysa et
al., 2020). My results in this study support a model in which, persistent expression of Dll4 in
SHF cells that assume an arterial endothelial fate, and the resulting Dll4-mediated Notch
signaling, is crucially required for maturation events that govern the re-organization of PAA into
adult aortic arch phenotypes. As in other SHF-derived cell types, Notch signaling transitions
from a proliferation to maturation role in SHF-derived PAA endothelial cells, as well. However,
this transition is not associated with a switch in its primary ligand as observed in the heart such
that, in arterial endothelial cells, the same ligand-receptor interaction also directs arterial
maturation. I speculate that the concomitant expression of other endothelial cell-specific
molecules facilitates such a behavior (Wythe et al., 2013).

Dll4, therefore, maintains its role as an arterial maturation molecule even in cells that have
reached an endothelial fate after transitioning through expression of cardiogenic molecules. I
show that Notch signaling is active in the SHF-derived endothelial cells that express Dll4 and
the physiologic effects of this signaling are discernable well before smooth muscle cells arrive to
the PAA. This would imply that Dll4 expressed in these cells interacts with Notch receptors also
expressed by endothelial cells, and not with Notch expressed by smooth muscle cells derived
from the neural crest. This is consistent with the prior observation that Notch knockdown in the
cardiac neural crest did not phenocopy the defects observed in this current study (High et al.,
2007). Dll4-Notch signaling is generally thought to occur in trans between Dll4 expressed on
one endothelial cell and Notch expressed on a neighboring cell. While that is likely also true in
the SHF, my data are unable to verify whether signaling occurs in cis or trans. Regardless, Dll4-
mediated Notch signaling regulates Hey1 expression, induces arterial identity and supports
continued growth and maturation of nascent blood vessels. These vessels interact with smooth
muscle cells that in this instance are ectodermal in origin, derived from cardiac neural crest
cells. Though the germ cell origin is distinct in this vascular bed, these smooth muscle cells
interact with arterial endothelial cells similar to the interaction seen in other vascular beds where
both cell types are derived from the mesoderm. Loss of Dll4 leads to loss of arterial
specification, resulting in maturation arrest and involution of the developing vessels, with a lack
of recruitment of smooth muscle cells. Interestingly, this phenomenon is most pronounced in the
4
th
PAA. A spectrum of 4
th
PAA defects are observed in mutant embryos, likely related to
biological variability in the timing and extent of cre-mediated Dll4 loss. The 3
rd
PAA, in contrast,
is able to compensate for loss of Dll4 in SHF by incorporating non-SHF derived cells to maintain
growth and maturation. In my model, the 6
th
PAA development does not show a reliance on Dll4

51

expression, which could be due in part to the different cellular interactions that regulate
pulmonary artery maturation compared to aortic maturation (Zhou et al., 2017).

Like other vascular beds (Benedito and Duarte, 2005), there is a dosage-sensitive requirement
of Dll4 in SHF-derived arterial endothelial cells. Almost 2/3
rd
of the embryos with heterozygous
loss of Dll4 in SHF demonstrate arch artery defects. In contrast, I have shown that only about
10% of heterozygous SHF Dll4 mutants develop intra-cardiac defects, such as double outlet
right ventricle (De Zoysa et al., 2020), indicating a more crucial role for Dll4 in arterial
development. Islet1-mediated loss of Dll4 had a more penetrant phenotype compared to Mef2c-
driven loss, likely because of the more global expression of Islet1 in SHF. Although Islet1-Cre
also labels a subset of neural crest cells, I have shown that Dll4 is not expressed in the neural
crest (De Zoysa et al., 2021). The arch artery defects seen in these mutants are highly
reminiscent of clinical arch defects. Recent evidence has shown that Dll4 (Meester et al., 2015,
Nagasaka et al., 2017) and Notch pathway (McKellar et al., 2007, Garg, 2016, Li et al., 1997)
mutations are relevant in clinical congenital heart defects. It would be of merit to study the
prevalence of Dll4/Notch mutations in children with isolated arch artery defects.

In summary, these data provide a novel paradigm in which SHF progenitors maintain Dll4
expression as they mature into endothelial cells. Dll4-mediated Notch signaling switches from its
proliferative role in early progenitors to a maturation role in arterial elements. Dll4 expression
induces arterial identity in PAA and regulates events that coordinate PAA re-organization into
the adult aortic arch phenotype.

52

Fig 6.1
Dll4 expression in SHF is required for the maturation of the aortic arch.
Coronal sections of Mef2c-Cre,R26RtdT (A) and Islet1-Cre,R26RtdT (B) embryos at E 10.5
demonstrates that 3
rd
, 4
th
and 6
th
pharyngeal arch artery endothelium is SHF derived.
Similarly transverse sections of Mef2c-Cre,R26RtdT (B) and Islet1-Cre, R26RtdT (D) co-
stained with Dll4 confirm lineage traced cells in 4
th
arch co-express Dll4 (arrows in B’’ and
D’’). Smooth muscle alpha actin stain in Wnt1-Cre,R26RtdT embryos at E13.5 (E)
demonstrates that the smooth muscle medial layer of the aortic arch is neural crest derived.
Conditional loss of Dll4 in SHF was achieved using Islet1 and Mef2c-mediated Cre
recombinase expression. H&E-stained transverse sections of E14.5 cre-negative littermate
control embryos and Dll4 SHF mutants (Mef2c-Cre, Dll4
F/F
, Islet1-Cre,Dll4
F/wt
) show variety
of arch artery defects including right aortic arch (AA, F’), cervical arch (G’), aberrant right
subclavian artery (ARSA, arrowhead in H’), aberrant left subclavian artery (ALSA, arrowhead
in I’) and aberrant ductus arteriosus (PDA, J’). Both Type B (K1’-K4’), and Type C (K1’’-K4’’)
interrupted aortic arch (IAA) were observed (arrowheads in K1’-K4’’ denotes the aorta). A
right aortic arch with aberrant left subclavian artery and a left-sided ductus led to a complete
vascular ring (L1’-L3’). Table in M lists the various arch defects observed in the different
genotypes.
AA, Aortic Arch; RSA, Right Subclavian Artery; LSA, Left Subclavian Artery; PDA, Patent
Ductus Arteriosus; ARSA, Aberrant Right Subclavian Artery; ALSA, Aberrant Left Subclavian
Artery; RCA, Right Carotid Artery; LCA, Left Carotid Artery
Scale bar - 100µm (A’-E’’’), 250µm (A-E), 300µm (F-J’, K1-K4’’, L1-L3’)

53

Fig 6.2
Haploinsufficiency of Dll4 in SHF leads to development defects in the pharyngeal arch
arteries.
Heterozygous conditional loss of Dll4 expression in SHF was achieved using Islet1-mediated
cre expression. Whole mount examination at E10.5 of India ink injected control wildtype
littermate embryos (A) and Dll4 heterozygous mutants (B-E) demonstrates 4
th
arch artery
defects bilaterally - absent (arrowheads in B, B’), hypoplastic (arrowheads in C, C’),
bifurcated (arrowheads in D, D’), and abnormal origin from the 6
th
arch artery (arrowheads in
E, E’). Whole mount examination at E12.5 of India ink injected control wildtype littermate
embryos (F) and mutant embryos (G-I) demonstrate clinically relevant aortic arch defects –
including isolated right aortic arches (G, G’), right arch with aberrant left subclavian artery (H,
H’) and complete vascular ring (I-I’”). Arrowheads in H’ and I’ denotes the aberrant left
subclavian arteries (ALSA).
AA, Aortic Arch; RSA, Right Subclavian Artery; LSA, Left Subclavian Artery; PDA, Patent
Ductus Arteriosus; IA, Innominate Artery; ALSA, Aberrant Left Subclavian Artery; RCA, Right
Carotid Artery; LCA, Left Carotid Artery
Wholemount magnification: x4 (F-I’’); x3 (A-E’)

54

55

Fig 6.3
Dll4 expression is required proper Hey1 expression and other downstream changes.
SHF cells were lineage traced by crossing R26RtdT mice into Islet1-Cre line. Coronal
sections of littermate control (A) and Dll4 homozygoys mutant (Mef2c-Cre,Dll4
F/F
) (B)
embryos were evaluated at E10.5. Mutant embryos maintain the caliber of the 3
rd
pharyngeal
arch artery (B1 vs. A1) although less than half the artery is composed of SHF-derived red
cells compared to the entire artery in controls (B2 vs. A2). Arrowhead in B2 indicate the
tdTomato negative non SHF derived arterial endothelium in the 3
rd
pharyngeal arch artery. In
contrast, the 4
th
pharyngeal arch artery lacks lumen and is very small in mutants (B4 vs. A4),
and all remaining cells are of SHF lineage as in controls (B5 vs. A5). Hey1 expression was
evaluated in the 4
th
pharyngeal arch arteries in littermate control and Dll4 mutants (Islet1-
Cre,Dll4
F/wt
). Hey1 expression is largely absent in both left (DL1-3) and right (BR1-3) sides.
Coronal sections of E10.5 control (E) and Islet1-Cre,Dll4
F/wt
mutant (F) embryos were stained
for pHH3 and CD31 to study endothelial cell proliferation. Higher magnification of boxed area
in E and F demonstrate reduced proliferation in mutants (F’-F’’’) compared to controls (E’-
E’’’). Arrowheads in E’’’ denote the pHH3 and CD31 double positive proliferating endothelial
cells. Invitro analyses of tube formation was performed using Human Umbilical Arterial
Endothelial Cells (HUAEC) in fibrin gels. Representative pictures at 3 (G-G’’’) and 5 (H-H’’’)
days are shown. Tube formation was studied under basal condition (G, H), with addition of
VEGF (2.5ng/ml, G’, H’) and using HUAEC pre-treated with Dll4- (G”, H”) or Hey1-specific
(G’”, H’”) siRNA (50nM).
Wholemount magnification: x5 (G-H’’’)
Scale Bars: 25µm (A1-A6, B1-B6, CL1-CL3, CR1-CR3, DL1-DL3, DR1-DR3, E’-F’’), 50µm
(E, F), 100µm (A, B, C, D)

56

Fig 6.4
Dll4 expression in SHF is required for arterial identity in pharyngeal arch arteries.
Expression of arterial markers EphrinB2 (A, B) and neuropilin1 (C, D) and venous markers
EphB4 (E, F) and neuropilin2 (G, H) were examined in control and Dll4 heterozygous
mutants (Islet1-Cre,Dll4
F/wt
). Transverse sections at the level of distal aortic arch where the
SHF-derived 4
th
pharyngeal arch artery (arrow in I”’ and J”’) meets the non-SHF derived
dorsal aorta (arrowhead in I”’ and J”’) in control (I) and Dll4 heterozygous mutants (Islet1-
Cre,Dll4
F/wt
) (J) was stained for CD31 and smooth muscle alpha actin. Smooth muscle
coverage is observed only around non-SHF-derived dorsal aorta in mutants compared to
both arteries in control (J” vs. I”). Arrowhead in J’’ denotes the lack of expression of Smooth
muscle in SHF derived proximal aorta.
Scale Bars: 25µm (A’-H’’), 50µm (A-H), 250µm (I, J)

57

Concluding Remarks

As previously mentioned in the lierature, Notch signaling is an evolutionary conserved signaling
pathway in many organisms due to its versatility and adaptability. It’s nature of being present in
many cellular systems in diverse organisms from drosophila to humans and diverse biological
processes from development to tumor progression also lends its hand to the extreme
importance it plays in human biology itself (Andersson et al., 2011).

Among other processes, the central role it plays in the cardiac and arch artery development
specifically exemplifies the importance of notch signaling to humans. As previously mentioned,
loss of notch pathway genes has been implicated in a variety of cardiac and arch artery
diseases including bicuspid aortic valves, alagille syndrome, adams-oliver syndrome and many
others (Luxan et al., 2016, Stittrich et al., 2014).

My current work specifically looks at how Delta like ligand-4 (Dll4) mediated notch signaling
impacts cardiac and arch artery development. It shows that when Dll4 mediated notch signaling
acts on Second Heart Field (SHF) cells in the pharyngeal mesoderm, notch signaling acts as a
proliferative signaling initially for the early expansion of the SHF cells. When enough cells are
generated through proliferation, the Dll4 expression recedes, and another notch ligand, Jagged1
takes over in activating notch signaling for the cells’ proper maturation and incorporation in to
the developing heart as cardiomyocytes. On the other hand, in SHF cells which are destined to
migrate in to the pharyngeal arch arteries and develop into mature aortic arches, Dll4 continues
to be the prominent ligand continuing to play a role in activating notch signaling even when the
underlying cells are being transitioned from an early proliferative state in to a more maturation
state.

This finding is a novel concept. While notch signaling is known to play roles both in proliferation
and maturation in the same cell type, current knowledge is that the transition from a more
proliferative to a more maturation state happens mainly due to a change in the ligand that
activates notch signaling itself (D’Amato et al., 2016). However, my work shows that, within the
same progenitor cell type, the same notch ligand has the ability to activate notch signaling both
as a proliferative signal and a maturation signal depending on what the progenitor cells
eventually mature into.

In that sense, my dissertation work has revealed a novel mechanism of notch signaling in SHF
cells in which notch signaling transitions from an early proliferative to a late maturation signaling
system not only through a ligand specific mechanism as previously described (D’Amato et al.,
2016), but also through a cell type specific manner.

58

Table 1
Primer Information

Primers
Primer Sequence
Dll4FF
Dll4-F GTGCTGGGACTGTAGCCACT
Dll4-R TGTTAGGGATGTCGCTCTCC
Dll4-F2-LacZ
LacZ-F ATCCTCTGCATGGTCAGGTC
LacZ-R CGTGGCCTGATTCATTCC
Dll4-F2-LacZ
JDW 24-F ACGGGCCTCTTCCTGTATTT
JDW 25-R AGTGCTGCCTCTGACCTCAT
Mef2c-Cre
Mef2cCre-F GAGCGTACGTGCTGCTTAGA
Mef2cCre-R AATCGCGAACATCTTCAGGT
Islet1-Cre
Islet1Cre Common-F GCCACTATTTGCCACCTAGC
Islet1Cre Mutant allele-R AGGCAAATTTTGGTGTACGG
Isl1Cre Wildtype allele-R CAAATCCAAAGAGCCCTGTC
Fgf8FF
AM28 GTGAGGGATTAGAGAGCG GGTG
AM09 TTGCCTTTGCCGTTGCTCTGC AGGTAG
AM GGTGGTGCAGATGAACTTCAGG
R26RLacZ
LacZ-111FWD TAATAGCGAAGAGGCCCGC
LacZ-611REV CGCCACATATCCTGATCTTCC
Wnt1-Cre
Wnt1-Cre-F CCTCTATCGAACAAGCATGCG

59

Wnt1-Cre-R GCCAATCTATCTGTGACGGC

R26RtdTomato
Common Forward AAGGGAGCTGCAGTGGAGTA
Wildtype Reverse CCGAAAATCTGTGGGAAGTC
Mutant Reverse CGGGCCATTTACCGTAAGTTAT
Notch Reporter
Notch Forward ACGTAAACGGCCACAAGTTC
Notch Reverse AAGTCGTGCTGCTTCATGTG
Promoter 1
Promoter 1-F GAGA-ACGCGT-
GACAACCTTTGAACACCTCTTCTGA
Promoter 1-R GAGA-AGATCT-
GACACTCTTGAAGTTAATGGCCTGA
Promoter 2
Promoter 2-F GAGA-ACGCGT-
TGCCAAATCACTGACAGATAAAA
Promoter 2-R GAGA-AGATCT-
CTTCTGAGTTCCAGGTTTTCCTT
Promoter 3
Promoter 3-F GAGA-ACGCGT-
CAGAGTCTGTGTCACTTTGAACG
Promoter 3-R GAGA-AGATCT-
CGAGTAAATGCCTAAACAGATGG
Enhancer 1
Enhancer 1-F GAGA-ACGCGT-
AGAGGAGAGGTAGCTTGGACCT
Enhancer 1-R GAGA-AGATCT-
AGAGATTCGCAGAGAACAATCC
Enhancer 2
Enhancer 2-F GAGA-ACGCGT-
CAATATGACACTTGGCTGCAAT
Enhancer 2-R GAGA-AGATCT-

60

GTAGAGAGGACCAGCGTTCAGT

Promoter 1 (Mutagenesis)
6 base substitution from
TGGGAA to GTTTCC

Promoter 1 (Mut) -F
AGAAACCTGAGGAAACGGGCCAGATG
GAGGG
Promoter 1 (Mut) -R
AAGGGCCCTGAACCTACC

61

Table 2
Antibody, Vector information, Cell Lines and Chemicals

Antibody, Vector Information, Cell Lines and Chemicals

Antibody

Catalog number

Working
Concentration/
Dilutions
Dll4 (Rabbit polyclonal) Ab7280 (Abcam) 20 μg/ m L
Dll4 (Rabbit polyclonal) PA1-86897 (Thermofisher) 50 μg/ m L
Islet1 (Goat Polyclonal) AF1837 (R&D) 10 μg/ m L
pHH3 (Rabbit polyclonal) 9701S (Cell Signaling) 0.1μ g / m L
Fgf8 (Rabbit Polyclonal) NBP1-41493 (Novus) 20 μg/ m L
Fgf10 (Sheep polyclonal) AF6224-SP (R&D) 10 μg/ m L
CD31 (Goat Polyclonal) AF3628 (R&D) 1μg / m L
CD31 (Rat Monoclonal) 550274 (BD Pharmingen) 1:100
Hand2 (Goat Polyclonal) AF3876-SP (R&D) 20 μg/ m L
Pitx2 (Sheep Polyclonal) AF7388-SP (Novus) 20 μg/ m L
Bmp4 (Rabbit monoclonal) MAB5020-SP (R&D) 6.25μ g / m L
Mlc2v (Rabbit Polyclonal) NBP1-85541 (Novus) 1μg / m L
Semaphorin3C (Rabbit
Polyclonal)
AB214309 (Abcam) 5μg / m L
Jagged1 (Rabbit Polyclonal) PA5-86057 (ThermoFisher) 20 μg/ m L
Nkx2.1 (Abcam) Ab76013 (Abcam) 0.8μ g / m L
Ap2α (Mouse polyclonal) NB100-74359 (Novus) 20 μg/ m L
EphrinB2 (Goat Polyclonal) AF496 (R&D) 1μg / m L
EphB4 (Goat Polyclonal) AF446 (R&D) 1μg / m L
Neuropilin1 (Goat Polyclonal) AF566 (R&D) 10 μg/ m L
Neuropilin2 (Goat Polyclonal) AF567 (R&D) 10 μg/ m L
Hey1 (Rabbit Polyclonal) NBP2-16818 (Novus) 1:100
Mm-Mef2c probe 421011 Assay dependent
Mm-Fgf8-C2 probe 313411-C2 Assay dependent
Mm-Fgf10-C2 probe 446371-C2 Assay dependent
62

Anti-Digoxigenin-POD, Fab
fragments
11207733910 (Roche) 0.5U/mL
Recombinant Mouse FGF-8
Protein
NBP2-35033- 25 μ g
(Novus)
10 0n g/μ L
50 0n g/μ L
VEGF 293-VE-050/CF (R&D) 2.5ng/mL

Vector Information
3XFlagNICD1 20183 (addgene)
pGL3 Luciferase Promoter
Vector

E1761 (Promega)
pGL3 Luciferase Enhancer
Vector

E1771 (Promega)
pGL3 Luciferase Enhancer
Vector

E1771 (Promega)

Cell lines

293T/17 [HEK 293T/17]
CRL-11268 (ATCC)
HeLa
CCL-2 (ATCC)
Human Umbilical Artery
Endothelial Cells

C12202 (PromoCell)

Chemicals

DAPT
D5942-5MG (Sigma-Aldrich)
SAHM1
491002-1MG (EMD Milipore)

63

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Asset Metadata

Creator De Zoysa, Prashan (author)

Core Title Proliferation and maturation events in second heart field cells during cardiovascular development activated by the Delta like ligand-4 mediated notch signaling

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Development, Stem Cells and Regenerative Medicine

Degree Conferral Date 2021-08

Publication Date 07/31/2021

Defense Date 06/11/2021

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag aortic arch,cardiac,Dll4,heart development,Notch signaling,OAI-PMH Harvest,OFT,outfllow tract,PAA,pharyngeal arch artery,second heart field,SHF

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Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Lien, Ching-Ling (Ellen) (committee chair), Hong, Young-Kwon (committee member), Subramanyan, Ram Kumar (committee member)

Creator Email pdezoysa@usc.edu,prashandz@yahoo.com

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