Close
About
FAQ
Home
Collections
Login
USC Login
Register
0
Selected
Invert selection
Deselect all
Deselect all
Click here to refresh results
Click here to refresh results
USC
/
Digital Library
/
University of Southern California Dissertations and Theses
/
Jagged1 functions downstream of Twist1 in the specification of the coronal suture and the formation of a boundary between osteogenic and non-osteogenic cells
(USC Thesis Other)
Jagged1 functions downstream of Twist1 in the specification of the coronal suture and the formation of a boundary between osteogenic and non-osteogenic cells
PDF
Download
Share
Open document
Flip pages
Contact Us
Contact Us
Copy asset link
Request this asset
Transcript (if available)
Content
JAGGED1 FUNCTIONS DOWNSTREAM OF TWIST1 IN THE SPECIFICATION OF
THE CORONAL SUTURE AND THE FORMATION OF A BOUNDARY BETWEEN
OSTEOGENIC AND NON-OSTEOGENIC CELLS
by
Hai-Yun Yen
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
(BIOCHEMISTRY AND MOLECULAR BIOLOGY)
December 2009
Copyright 2009 Hai-Yun Yen
ii
Acknowledgements
I would like to show my gratitude to my advisors Dr. Robert Maxson and Dr. Louis
Dubeau. They have given me the freedom to explore and grow as a graduate student and
as a researcher. I could not have done my work without their supports and guidance. I
would also like to thank my committee members, Dr. Pradip Roy-Burman and Dr. Gage
Crump, for their insightful discussions and suggestions.
People in the Maxson lab and Dubeau lab have been very helpful in various ways. I
would like to give special thanks to Nancy Wu for her kindness and for helping me with
her outstanding performance of animal surgery and to Mamoru Ishii for sharing his lab
skills, experience and knowledge with me. I would also like to thank Youzang Yen,
Linda Liu, Mandy Ting, Paul Roybal and Chris Schafer for making this lab a rewarding
and stimulating place to work. Jingjing Sun is such an optimistic person and fun to be
around. I am truly grateful for her friendship.
My family has always given me support and had confidence in me. I especially
appreciate my parents and grandmother for encouraging my education and for believing
in my potential. I thank them for raising me to be an independent thinker and a
responsible person. Last but not least, I would like to express my appreciation to my
boyfriend, Cliff who always cheers me up and makes me smile. His support, caring, and
friendship have made it all possible.
iii
Table of Contents
Acknowledgements ii
List of Tables v
List of Figures vi
Abstract viii
Chapter1 1
Jagged1 functions downstream of Twist1 in the specification of the coronal
suture and the formation of a boundary between osteogenic and non-
osteogenic cells
Introduction 1
1. Suture biology 1
2. Craniosynostosis 4
3. Genes of interest in craniosynostosis 8
Materials and Methods 13
Results 17
1. The Notch ligand Jagged1 is required in cranial suture
development 17
2. Jagged1 is a downstream effector of Twist1 in suture
development 37
Conclusion 52
1. Sutural cells identity was changed in early development of the
skull 52
2. Notch signaling in suture patterning 53
3. Notch signaling in Drosophila wing imaginal disc is comparable
to that in coronal suture 54
4. Notch signaling interacts with other major signaling pathways
during suture formation 57
5. Epistasis model of Twist1 and Notch signaling in coronal suture 58
Chapter 1 References 61
iv
Chapter 2 70
Inactivation of Brca1 in ovarian granulosa cells is associated with increases
in circulating estrogen levels and bone mineral density
Introduction 70
Materials and Methods 74
Results 77
1. Absence of a functional Brca1 in granulosa cells increases
serum estrogen levels in mice 77
2. Fshr-Cre is expressed in pituitary gland and brain 79
3. Ovaries carrying a Brca1 mutation are responsible for the
increase of serum estradiol 81
4. Phenotypes suggested that mutant mice were exposed to high
levels of estradiol after estrus induction and synchronization 81
5. Long-term exposure to elevated estrogen increases bone density
in mutant mice 85
Conclusion 89
Chapter 2 References 92
Bibliography 96
Appendix 108
1. ALP section staining 108
2. ALP whole-mount staining 109
3. LacZ section staining 110
4. Tissue preparation (OCT embedding) 111
5. Immunoperoxidase staining 112
6. Immunofluorescence staining 113
7. Section in situ hybridization (from Eva Chan) 114
8. RNA probe 116
v
List of Tables
Table 1: Sutural synostosis 5
Table 2: Craniosynostosis syndromes 6
Table 3: Signaling pathways and craniosynostosis 7
Table 4: Offspring obtained from crossing Mesp1-Cre;Jagged1
cko/+
males
and Jagged1
cko/+
or Jagged1
cko/cko
females 22
Table 5: Offspring obtained from crossing Dermo1-Cre;Jagged1
cko/+
males
and Jagged1
cko/+
or Jagged1
cko/cko
females 23
Table 6: Offspring obtained from crossing Wnt1-Cre;Jagged1
cko/+
males
and Jagged1
cko/+
or Jagged1
cko/cko
females 25
Table 7: Percentage of sections with boundary crossing neural crest in
coronal mid-suture mesenchyme 44
Table 8: Penetrance of craniosynostosis 46
Table 9: Craniosynostosis index (CI) 47
Table 10: Measurements of the size of retrotympanic process 48
Table 11: Middle ear ossicle malformations in Twist1
+/-
;Jagged1
+/-
cross at
P21 50
Table 12: Craniosynostosis index (CI) for coronal sutures 51
vi
List of Figures
Figure 1: Mouse skull structure 3
Figure 2: The expression of Jagged1 and Twist1 in the prospective mouse
coronal suture at embryonic stage (E) 14.5 19
Figure 3: Mesp1-Cre mediated conditional inactivation of Jagged1 in
mesoderm causes abnormal coronal suture phenotype at E14.5 21
Figure 4: Dermo1-Cre mediated conditional inactivation of Jagged1 causes
craniosynostosis 24
Figure 5: Jagged1 is not required in the neural crest for coronal suture
development 26
Figure 6: Cell-mixing at mesoderm-neural crest boundary in the coronal
suture of Mesp1-Cre;Jagged1
cko/cko
embryos 28
Figure 7: Inactivation of Jagged1 in mesoderm causes an expansion of
Notch2 expression in the prospective coronal suture 30
Figure 8: The expression of Hes1, a downstream target of Notch signaling,
was consistent with that of Notch2 in Dermo1-Cre;Jagged1
cko/cko
mutant mice 31
Figure 9: Notch ligand, Dll1, was expressed in the mid-sutural
mesenchyme 32
Figure 10: RTK signaling pathway is not altered in Jagged1 mutant
embryos at E14.5 34
Figure 11: Both BMP and Wnt signaling pathways were affected in
Dermo1-Cre;Jagged1
cko/cko
mutants 36
Figure 12: Twist1 regulates Notch signaling pathway in coronal suture at
E12.5 38
Figure 13: Jagged1 exhibits a specific expression pattern in coronal suture
at E13.5 40
Figure 14: Heterozygous loss of Twist1 caused boundary defects in coronal
suture at E14.5 41
Figure 15: Twist1 expression was not influenced by Jagged1 dosage 42
vii
Figure 16: Genetic interaction between Twist1 and Jagged1 and its effect
on craniosynostosis 45
Figure 17: The size of retrotympanic process (RTP) of the squamosal bones
was reduced in Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mutants 49
Figure 18: The malformations of middle ear ossicles 50
Figure 19: Notch signaling in Drosophila wing disc (from González et al.,
2006) 55
Figure 20: Notch signaling in mouse coronal suture 56
Figure 21: Signaling pathways in coronal suture 58
Figure 22: Epistasis model of Twist1 and Notch signaling in coronal suture 59
Figure 23: Conditional knockout of Brca1 in granulosa cells resulted in an
increase of serum estradiol 78
Figure 24: Fshr-Cre is expressed in pituitary gland and brain 80
Figure 25: Ovarian granulosa cells are responsible for increased serum
estradiol in mutant mice 82
Figure 26: Analysis of cell proliferation in uterus after PMSG treatment 83
Figure 27: Analysis of cell proliferation in uterus after PMSG and hCG
treatments 84
Figure 28: Vascular wall dilation in mutant ovaries after PMSG and hCG
treatments 86
Figure 29: There is no difference in the number of blood vessels in corpus
luteum between wildtype and mutant after PMSG and hCG
treatments 87
Figure 30: High trabecular bone mass in mutant female mice 88
viii
Abstract
In this dissertation, I would like to address how the evolutionarily conserved Notch
signaling molecules regulate the formations of murine cranial bones and sutures.
Mutations in the Notch ligand, JAGGED1, cause Alagille syndrome, which has
craniosynostosis as a feature. The expression pattern of Jagged1 at mouse coronal suture
suggested that it might play a role in establishing boundary between osteogenic and non-
osteogenic cells. Tissue-specific knockout of Jagged1 in mouse mesoderm affected the
expression of downstream Notch signaling at sutural cells and resulted in
craniosynostosis. Immunostaining results also implied that the boundary between
presumptive cranial bones has been established by Notch signaling at early stage, while
the opposing osteogenic fronts of the bones are still far away from each other. I further
demonstrated the genetic interactions between Notch signaling and Twist1 which is an
important pathogenic gene in regulating cranial sutures morphogenesis. Twist1 regulates
Notch signaling in sutural mesenchyme and maintain suture patency. The phenotypic
studies of mouse skulls and middle ear ossicles indicated that Jagged1 interacts
functionally with Twist1 in several distinct developmental settings. This work reveals a
molecular network that controls cranial development, and establishes a new model of
boundary formation at developing cranial suture.
1
Chapter 1
Jagged1 functions downstream of Twist1 in the specification of the coronal suture
and the formation of a boundary between osteogenic and non-osteogenic cells
Introduction
Tissue patterning is one of the fundamental subjects regarding embryonic development.
The effect of boundaries on pattern formation was studied in many systems as diverse as
Drosophila wing disc (Bray, 1998; Major and Irvine, 2005; Buceta et al, 2007) and the
mammalian hindbrain (Kiecker and Lumsden, 2005). Previous studies in our lab showed
that loss of boundary integrity in cranial coronal suture was accompanied by
craniosynostosis (Merrill et al., 2006; Ting et al., 2009). Studies in sutural biology help us
to understand cell behaviors during boundary formation. Moreover, we could use it as a
model system to elucidate boundary establishment.
1. Suture biology
1.1. Skull functions, structures and development
Skull provides basic structure of the head. The mammalian skull gives protection to the
fragile brain, supports sensor organs and makes mastication possible. It has to perfectly
coordinate with other organs during development. So the brain would have enough space
to grow, the eyes and ears would be positioned in the right places and we can breathe and
masticate properly. The correct patterning of the skull is so important, not just for
aesthetic reasons but for optimizing the survival ability of an organism.
2
Skull can be separated into neurocranium and viscerocranium anatomically.
Neurocranium surrounds the brain and viscerocranium makes up the bones of the face.
We could further subdivide neurocranium into skull vault (calvaria) and skull base. The
mammalian calvaria covers the "upper part of the head" (it is the definition of "calvaria”
in Latin) and protects brain from impact injury. It is composed of flat bones including
two-paired frontal, two-paired parietal and one unpaired interparietal bone. These bones
are developed by intramembranous ossification. In contrast to endochondral bones
formation that is a process occurs mostly in long bones, intramembranous bones are
formed directly from mesenchymal cells without the mediation of a cartilage phase.
The mammalian skull vault is embryologically derived from mesoderm and neural crest
cells. Cell distribution and migration in the skull have been studied by using Cre/R26R
labeling system and DiI injection (Jiang, X., et al, 2002; Chai and Maxson, 2006). The
frontal bones are neural crest-derived while the parietal bones and coronal suture
mesenchyme are mesoderm-derived.
At E8.5, cranial neural crest cells emigrate from the caudal forebrain, midbrain, and
prorhombomere A of the hindbrain. The cells are migrating rostrally and are then located
at frontonasal and first branchial arch at E9.5. In the meanwhile, a clear boundary
between neural crest- and mesoderm-derived mesenchymal cells is established.
Beginning at this stage, the subectodermal cranial mesenchymal cells migrate upward
toward the top of the head and start condensation. At E12.5, the condensed frontal and
parietal bone primordia can be detected by staining with alkaline phosphatase (ALP), an
3
early osteoblast differentiation marker (Ishii et al, 2003). The presumptive coronal sutures
are ALP negative and are clearly distinguishable in the developing skull.
1.2. Suture functions and development
The flat bones of the skull vault are connected by fibrous tissue called sutures. The
following diagrams show the names and the positions of the six cranial sutures in the
mouse skull.
In the mammalian, these rigid joints are found only in the skull (Persson, 1995). Sutures
play important roles in controlling skull bone development and improving skull
architecture (Rice, 2008). Sutures are able to perform the following functions:
1) Sutures serve as growth centers during embryonic and early postnatal skull
development. It has been suggested that cranial sutures are the major growth sites
of the skull bones. (Opperman, 2000)
2) To maintain the elasticity of the skull and allow deformation of the skull during
childbirth.
Sagittal
Interfrontal
Occipitalinterparietal (OIP)
Lambdoid
Coronal
Interparietal bone
Parietal
bone
Frontal
bone
Squamosal
Fig. 1. Mouse skull structure
4
3) To absorb mechanical stress including impact loading, mastication and internal
pressures.
The timing of normal suture closure is different between human and mouse. Most human
calvarial sutures fuse in the forth decade of life, except for the metopic (interfrontal)
suture which closes at early age (9 months to 2 years). In the mouse, all calvarial sutures,
except for the posterior section of the frontal suture and the OIP suture, remain patent for
life (Aviv et al., 2002).
The human brain grows rapidly in the first year of life. It is important that the growth of
skull vault be coordinated with the expanding growth of the brain during the early years.
The skull vault growth and suture formation are closely related. As mentioned above,
sutures are the major growth sites of the skull bones and open sutures allow the skull to
develop into its normal shape. Premature fusion of the suture, also called
craniosynostosis, results in craniofacial asymmetry and increased intracranial pressure
when the growing brain presses against a rigid skull.
2. Craniosynostosis
2.1. Classification of craniosynostosis
The word “carnio-syn-osto-sis” is combined by the following words. Cranio- is from
Greek “kranion” which means “skull”; syn- is from Greek “sun” which means
“together”; osteo- is from Greek “osteon” which means “bone” and -osis is a suffix
5
denoting a diseased condition. Taken together, this word indicates a disease with
abnormal fusion of the skull bones.
Primary craniosynostosis, which is defined as synostosis without an underlying
abnormality of the brain or metabolic defect, occurs in 1 in 2500 live births (Wilkie,
1997). It is defined by the suture(s) involved and whether it is syndromic or
nonsyndromic.
1) Sutural synostosis:
Synostosis in different suture is associated with specific skull phenotypes. The
following table indicates skull shapes that are characteristic for different types of
sutural synostosis (Kabbani and Raghuveer, 2004; Morriss-Kay and Wilkie, 2005).
Table 1 Sutural synostosis
Sutural synostosis Skull shape Prevalence
Sagittal
Scaphocephaly
-A narrow head that is
excessively long.
40-55%
Bilateral coronal
Brachycephaly
-The skull is excessively
wide and short from front
to back.
Unilateral coronal
Plagiocephaly
-The opposite forehead
bulges significantly.
20-25%
Metopic
(Interfrontal)
Trigonocephaly
-Triangular shaped
forehead.
5-15%
Lambdoid
The skull around the
involved suture is
flattened.
0-5%
2) Nonsyndromic or syndromic:
6
More than 50% of the craniosynostosis is nonsyndromic (Morriss-Kay and Wilkie,
2005). Nonsyndromic craniosynostosis occurs sporadically and usually affects a
single suture. The pathogenesis of nonsyndromic craniosynostosis is largely
unknown. There were only a few cases indicated that FGFR3 and TWIST1 might be
involved (Boyadjiev, 2007).
Syndromic means that craniosynostosis is just one feature of the condition. Over 100
syndromic craniosynostosis have been described. The following is a list of the most
common craniosynostosis syndromes and their responsible genes (Rice, 2008; Jenkins
et al, 2007; Warman et al., 1993).
Table 2 Craniosynostosis syndromes
Syndromes Gene mutation Suture phenotypes
Apert
FGFR2
constitutive
activation
Coronal synostosis. Widened
sagittal and metopic sutures.
Crouzon
FGFR2
constitutive
activation
Coronal, sagittal and lambdoid
synostosis.
Pfeiffer
FGFR1 and 2
constitutive
activation
Coronal and sagittal synostosis
in Type I. Multi-sutural
synostosis in Type II.
Saethre-Chotzen
TWIST1,
haploinsufficient
Coronal synostosis.
Carpenter
RAB23
haploinsufficient
Coronal, sagittal and lambdoid
synostosis.
Craniofrontonasal
EPHRINB1
haploinsufficient
Coronal synostosis.
Muenke
FGFR3
constitutive
activation
Coronal synostosis.
Boston-type
MSX2
gain of function
Multi-sutural synostosis
7
2.2. Genetic basis of Craniosynostosis
Not too long ago, the genetic cause of craniosynostosis was totally unknown. The first
gene identified in craniosynostosis was MSX2 (Jabs et al., 1993). For over a decade now,
we know quite a number of genes that cause craniosynostosis (Morriss-Kay and Wilkie,
2005; Cohen, 2006). These genes are components of several important signaling
pathways including BMP, RTK, Hedgehog, Wnt and Notch pathways. In the following
table, I listed the signaling pathways and gene components that are known to associate
with craniosynostosis. Some of the genes have confirmed mutations in human patients
(gene names in upper cases) and some of them only have mice models (upper and lower
cases).
Table 3 Signaling pathways and craniosynostosis
Signaling
pathways
Genes Human syndromes References
TGFBR1, TGFBR2 Loeys-Dietz Loeys et al., 2005
MSX2 Boston-type Jabs et al., 1993
FIBRILLIN-1 (FBN1) Shprintzen-Goldberg Sood et al., 1996
BMP
Gdf6 - Settle et al., 2003
FGFRs 1-3
Apert, Crouzon,
Pfeiffer, Muenke,
Jackson-Weiss
Hajihosseini, 2008
TWIST1 Saethre-Chotzen Howard et al., 1997
EPHRINA4 (EFNA4) Nonsyndromic Merrill et al., 2006
EFNB1 Craniofrontonasal Twigg et al., 2004
EphA4 - Ting et al., 2009
Dusp6 - Li et al., 2007
Pdgfr alpha
- Moenning et al.,
2009
RTK
Nell-1 - Zhang et al., 2002
Hedgehog RAB23 Carpenter Jenkins et al., 2007
Wnt Axin2 - Yu et al., 2005
Notch JAGGED1 Alagille Kamath et al., 2002
8
3. Genes of interest in craniosynostosis
In this work, I was focusing on the roles of Twist1 and Jagged1 genes in
craniosynostosis. In the literature, there is little connection between these two genes. I
would demonstrate that Twist1 and Jagged1 work cooperatively in suture formation.
3.1 Twist1
Recent studies from our laboratory have showed the developmental basis of Saethre-
Chotzen syndrome, caused by heterozygous loss of function of TWIST1 (Merrill et al.,
2006; Ting et al., 2009). Twist1 is a transcriptional factor containing a basic helix-loop-
helix domain. It was first identified in Drosophila as a gene crucial for proper gastrulation
and mesoderm formation (Thisse et al., 1988). Members of the Twist family have been
identified in human, mouse, frog, lancelet, nematode, leech, zebrafish, jellyfish and
chicken (Castanon and Baylies, 2002)
Mutations of Twist1 gene in different organisms revealed its functions in early
mesodermal patterning and osteogenesis. I listed some of the important observations or
phenotypes in the animal models, and the features of human patients with Twist1 gene
mutations:
1) Drosophila: Mesodermal cell fate commitment (Castanon et al., 2001). Muscle
fiber development (Lavato et al., 2005).
2) Mouse: Model of Saethre-Chotzen syndrome (el Ghouzzi et a., 1997).
Twist1
-/-
mice die at E11.5 due to a failure of cranial neural tube closure. Twist1
+/-
9
mice exhibits the following phenotypes—Craniosynostosis, polydactyly,
abnormal head cartilage development, misalignment of upper and lower incisors,
delayed growth of the retrotympanic process (RTP) of the squamosal bone and
thoracic skeletal abnormalities (Bourgeois et al., 1998).
3) Human: Saethre-Chotzen syndrome (OMIM:101400)
The common feature of the syndrome—craniosynostosis, ptosis of the eyelids,
deviated nasal septum, brachydactyly, and partial soft tissue syndactly and
radioulnar synostosis.
Previous studies from our lab showed that Twist1 mutants have a defect in the boundary
between neural crest and mesoderm in the coronal suture. (Merrill et al., 2006; Ting et al.,
2009). This boundary coincides with the boundary between osteogenic and non-
osteogenic compartments within the suture. The ephrin-Eph signaling was found to be
altered in Twist1
+/-
mice. Reduced dosage of Twist1 and EphA4 results in inappropriate
targeting of migratory osteogenic cells to the coronal suture. It was suggested that this
pathfinding defect is a key cause of craniosynostosis in Twist1 and EphA4 mutants. The
role of Twist1 in cranial mesenchymal cell migration is consistent with its role in tumor
metastasis (Vernon and LaBonne, 2004). Other studies suggested that Twist1 affects
osteoblast differentiation through its interaction with FGFR signaling (Rice et al., 2000)
and Runx2 (Yousfi et al.,2002).
10
3.2 Jagged1
Jagged1 serves as a ligand for Notch signaling pathway. Constituents of the Notch
pathway in vertebrates include the membrane receptors, Notch1-4, and membrane bound
ligands, Jagged1-2 and Dll1, 3 and 4 (Kopan and Ilagan, 2009). After ligand activation,
the intracellular domain (NICD) of Notch receptor is released by proteolysis and
translocated to the nucleus. Within the nucleus, NICD replaces repressors from the DNA
binding protein, CSL, and recruits a coactivator to form a transcription complex that
modulates the expression of downstream genes such as Hes and Hey family genes (Bray,
2006). Notch signaling is critical for a variety of developmental processes, including
developmental boundary formation (such as its function in the Drosophila wing disc) and
cell type specification (Lai, 2004; Bolós et al., 2007).
In humans, heterozygous loss of function of JAGGED1 results in Alagille syndrome, a
multi-organ disorder characterized by impaired development of intrahepatic bile ducts, as
well as defects in the heart, eye, kidney, face, and skull (Alagille, 1987; Oda et al., 1997;
Krantz et al., 1998; Emerick et al., 1999; Kamath et al., 2004). Synostosis of the coronal
suture occurs at low frequency (Kamath et al., 2002). Mutations in NOTCH2, a receptor
for JAGGED1, also cause many of the defects of Alagille syndrome, though
craniosynostosis has not been reported to be among them (McDaniell et al, 2006).
In contrast to humans with JAGGED1 mutations, Jagged1
+/-
mice develop normally,
except for eye dysmorphologies (iris coloboma and cornel opacity). However,
homozygous mutants die at E10.5 with defects in embryonic and yolk sac vascular
11
remodeling (Xue et al., 1999). Jagged1/Notch2 compound mutants exhibit a set of
deficiencies similar to those of humans affected with Alagille syndrome, although these
mutants do not have craniosynostosis or other craniofacial defects (McCright et al.,
2002). Recently, conditional Jagged1 alleles were created, and several studies have
documented conditional phenotypes in different tissues:
1) Inactivation of Jagged1 with Foxg1-Cre in inner ear causes loss of outer hair cells
(Kiernan et al., 2006; Brooker et al., 2006).
2) Tg Alfp-Cre;Jagged1
loxP/dDSL
adult mice show biliary abnormalities (Loomes et al.,
2007).
3) Inactivation of Jagged1 with Le-Cre mimics the eye defects in Jagged1
+/-
mice
(Le et al., 2009).
4) Inactivation of Jagged1 with Pdx1-Cre causes pancreatic ductal malformation
(Golson et al., 2009).
The consequences of inactivation of Jagged1 in tissues of the skull vault have not yet
been investigated.
There were three reasons that prompted us to investigate the relationship between
Jagged1 and Twist1 genes in suture formation. First, Previously studies showed that
EphA4 is a downstream effector of Twist1 (Ting et al., 2009). However, EphA4
-/-
mice
showed lower penetrance and less severe phenotype of craniosynostosis than that of
Twist1
+/-
mice, suggesting that there may be additional genes downstream of Twist1,
acting in parallel with EphA4 in the maintenance of the osteogenic-non-osteogenic
boundary and in the pathophysiology of craniosynostosis. Second, Twist1 is normally
12
activated both in osteogenic compartment (bones) and non-osteogenic compartment
(coronal suture mesenchyme). Therefore, Twist1 could not define the osteogenic-non-
osteogenic boundary solely according to its expression pattern. It is more likely that
Twist1 cooperates with other factors that are able to establish a boundary between
osteogenic and non-osteogenic compartments. Third, Jagged1 is a good candidate to
work with Twist1 in the suture. Mutations in Jagged1 causes Alagille syndrome, which
has craniosynostosis as a feature, and Notch signaling is also known to play an important
role in boundary determination.
Here I provide evidence that the Notch ligand, Jagged1 is required in cranial suture
formation and it functions cooperatively with Twist1 to maintain the boundary during
suture development.
13
Materials and Methods
Mouse mutants and genotyping
All genetically modified mice used in this study were as described previously: Twist1
(Chen and Behringer, 1995), Wnt1-Cre (Danielian et al., 1998), Mesp1-Cre (Saga et al.,
1999), Dermo1-Cre (Yu et al., 2003), and R26R (Soriano, 1999).
Jagged1 conventional knockout mice were obtained from Dr. Yang Chain (Center for
Craniofacial Molecular Biology, Los Angeles, CA) and were originally obtained from Dr.
Thomas Gridley (Xue et al., 1999.).
Jagged1 floxed mice were obtained from Dr. Neil Segil (House Ear Institute, Los
Angeles, CA) and were originally obtained from Dr. Julian Lewis (Brooker et al., 2006).
The genotyping conditions were as follows:
Jagged1
Primers:
JGK01: tct cac tca ggc atg ata aac c (22)
JGK02: taa cgg gga ctc cgg aca ggg (21)
SOL1: tgg atg tgg aat gtg tgc gag (21)
PCR program: 94°C. 5 min.; 94°C. 1 min.; 58°C. 1 min.; 72°C. 1 min. 30sec.; Goto #2 34
times; 72°C. 10 min.; 4°C forever.
Expected product size: JGK01+JGK02 for wildtype band ~400 bp;
JGK01+SOL1 for mutant band ~240 bp
Twist1
Primers:
TwB: gcc tgt ttt cta tga ccg ct (20)
14
Twi3: aat cca tct tgt tca atg gcc gat c (21)
PCR program: 94°C. 4 min.; 94°C. 1 min.; 60°C. 1 min.; 72°C. 2 min.; Goto #2 37 times;
72°C. 7 min.; 60°C 10 min.; 4°C forever.
Expected product size: TwB+Twi3 for mutant band ~1 Kb
Jagged1 flox
Primers:
Jag1-Commont: ga act cag gac agt gct c (19)
Jag1-DEL: ata gga ggc cat gga tga ct (20)
Jag1-FX: gtt tca gtg tct gcc att gc (20)
PCR program: 94°C. 3 min.; 94°C. 30 sec.; 59.6°C. 45 sec.; 72°C. 1 min.; Goto #2 34
times; 72°C. 10 min.; 4°C forever.
Expected product size: Jag1-Com+Jag1-FX for wildtype band ~390 bp;
mutant band ~390 bp
Jag1-Com+ Jag1-DEL for deleted band ~3300 bp
Cre
Primers:
Cre-forward: ctc tgg tgt agc tga tga tc (20)
Cre-reverse: taa tcg cca tct tcc agc ag (20)
PCR program: 94°C. 5 min.; 94°C. 30 sec.; 55°C. 1 min.; 72°C. 1 min.; Goto #2 34
times; 72°C. 5 min.; 4°C forever.
Expected product size: ~330 bp
R26R
Primers:
R26Rcom: aaa gtc gct ctg agt tgt tat (21)
R26Rwt: gga gcg gga gaa atg gat atg (21)
R26RKO: gcg aag agt ttg tcc tca acc (21)
15
PCR program: 94°C. 5 min.; 94°C. 30 sec.; 53°C. 45 sec.; 72°C. 1 min.; Goto #2 34
times; 72°C. 10 min.; 4°C forever.
Expected product size: R26Rcom+ R26Rwt for wildtype band ~650 bp;
R26Rcom+ R26RKO for knockout band ~350 bp
Skull preparation
The heads of postnatal day 21 mice were skinned and stained with Alizarin Red S
(80mg/l in 1% KOH) for about one week to reveal mineralized bone. The skulls were
then cleared
and stored in 100% glycerol.
The heads of newborn mice were stained with Alcian blue and Alizarin Red S fully
according to the procedure described previously (McLeod, 1980).
Histochemical staining for alkaline phosphatase (ALP) and β-galactosidase activity
The ALP staining procedures for whole-mount and tissue sections were performed as
described previously (Ishii et al., 2003), with minor modifications. (See Appendix 1 and
2 for complete protocols) Briefly, E14.5 embryo heads for whole-mount staining were
fixed in 4% PFA overnight and washed with PBS three times for 10min. Samples were
stored at 70% EtOH at 4°C for two days and bisected sagittally. Brains and skins were
removed for clear illustration. The specimens were then stained with 0.01% BCIP and
0.025% NBT in NTMT solution.
Cre-activated β-galactosidase activity in Mesp1-Cre;R26R, Mesp1-
Cre;R26R;Jagged1
cko/cko
, Dermo1-Cre;R262, Wnt1-Cre;R26R embryos were detected by
X-gal staining on 10-µm cryostat sections as described previously (Ishii et al., 2003)
(Appendix 3). Cryostat sections of E14.5 embryos derived from a cross between Wnt1-
16
Cre;Twist1
+/-
;Jagged1
+/-
males and R26R females were stained with X-gal followed by
immunostaining for Jagged1 as described below.
Immunostaining and in situ hybridization
The heads of E12.5, E13.5 and E14.5 embryos were fixed with 4% paraformaldehyde and
embedded in HistoPrep (Fisher Scientific) (Appendix 4). Transverse frozen sections were
cut in a cryostat at 10-µm thickness.
Immunoperoxidase staining was performed by using streptavidin-biotin-peroxidase
method (Histostain-SP Kit, Zymed), followed by Diaminobenzidin tetrahydrochloride
(DAB) substrate (Zymed) with or without hematoxylin counterstain according to the
manufacturer's instructions (Appendix 5). Primary antibodies were purchased from the
following companies and used at the indicated dilutions: goat anti-Jagged1 (Santa Cruz,
sc-6011, 1:800), rabbit anti-Notch2 (sc-5545, 1:500), goat anti-Dll1 (sc-12531, 1:500),
goat anti-Hes1 (sc-13842, 1:500), rabbit anti-β-catenin (Sigma, C2206, 1:400) and rabbit
anti-P-Erk1/2 antibody (Cell Signaling, #4376, 1:400).
Indirect immunofluorescence staining was performed by using rabbit anti-P-Smad1/5/8
antibody (Cell Signaling, #9511, 1:100) and Rhodamin Red-X goat anti-rabbit IgG
secondary antibody (Invitrogen, R6394, 1:100). The cell nuclei were revealed by co-
staining with DAPI (Appendix 6).
Section in situ hybridization was carried out as described previously (Chen et al., 2007)
(Appendix 7). Digoxigenin-labeled Twist1 antisense RNA probe was generated from
“Twist N’ pBSK” plasmid (linearized with Bam HI, using T7 promoter) as reported by
Ishii et al. (Ishii et al., 2003). (Appendix 8).
17
Results
1. The Notch ligand Jagged1 is required in cranial suture development
1.1 Jagged1 and Twist1 expression patterns overlapped in coronal suture
mesenchyme
Mutations in JAGGED1 are responsible for Alagille syndrome, which has premature
suture fusion as a feature (Kamath et al., 2002). In this study, the authors identified two
patients with classical features of Alagille syndrome (tetralogy of Fallot, jaundice,
paucity of bile ducts and butterfly vertebrae in Patient A; jaundice, paucity of bile ducts,
pulmonary stenosis, and an atrial septal defect in Patient B), who also had unilateral
coronal synostosis. Mutations in JAGGED1 gene were identified in both of the patients
(2504del5 and E553X, respectively) and no other mutations in major craniosynostosis
genes including FGFR1-3 and TWIST1 were detected. Although it is possible that there
are other craniosynostosis genes involved, these results intrigued us and led us to
examine the role of the Notch ligand, Jagged1 in cranial suture formation.
Previous studies from our laboratory have demonstrated that the boundary between
osteogenic and non-osteogenic cells in mouse coronal suture becomes evident at
embryonic stage (E) 14.5 (Merrill et al., 2006; Ting et al., 2009). In these studies,
boundary defects could not be found at earlier stages in the transgenic mouse embryos
carrying mutations in Twist1 or EphA4 genes. Therefore, I examined the expression of
Jagged1 at E14.5 when the boundary is first evident at suture. Immunostaining revealed a
specific expression pattern of Jagged1 protein in the coronal suture (Fig. 2A). The
prospective parietal bone (PB) was enclosed by two bands and the lower band was across
18
the coronal suture ectocranial to the prospective frontal bone (FB) (Fig. 2A, arrow). This
pattern perfectly matches the boundary of osteogenic cells during frontal and parietal
bones development. It suggests that Jagged1 might form a boundary and provide
guidance for the developing bones.
Mutation in TWIST1 causes Saethre-Chotzen syndrome, which is characterized by a
broad set of malformations including craniosynostosis. The expression pattern of Twist1
during mouse coronal suture development was reported previously (Johnson et al., 2000;
Yoshida et al., 2005). Resembling in situ hybridization results were obtained and showed
that Twist1 was expressed in the prospective frontal and parietal bones and as well as in
the sutural cells (Fig. 2B, arrow). It was also apparent in a layer of cells outside the
prospective bone layer (Fig. 2B, EL). From previous studies in our lab, we learned that
osteogenic precursor cells migrate within this ectocranial layer by using DiI labeling
(Ting et al., 2009).
The specific expression patterns of Jagged1 and Twist1 indicated that they may have
distinctive roles during cranial development. However, it is interesting that their
expressions overlap significantly in the suture mesenchyme. Therefore, a genetic
interaction between these two genes is suggested as well. In section two, I would
demonstrate the results of an epistatic relationship between Twist1 and Jagged1.
19
Fig. 2. The expression of Jagged1 and Twist1 in the prospective mouse coronal
suture at embryonic stage (E) 14.5.
(A) Jagged1 antibody staining was performed on transverse frozen sections of
E14.5 mouse head as depicted. An adjacent section was stained for the early
osteoblast marker, alkaline phosphatase (ALP) with nuclear fast red counterstain. A
superimposition of the Jagged1 immunostaining and ALP is shown for clarity.
Jagged1 was expressed at mid-sutural mesenchyme (arrow) and established a
boundary line for the developing parietal bone (PB) at this stage. (B) In situ
hybridization analysis of the suture marker and craniosynostosis gene, Twist1.
Twist1 was expressed in the prospective bones and in the sutural cells (arrow). Note
that Jagged1 and Twist1 are both expressed in the prospective coronal suture
(arrows), as well in an ectocranial cell layer (EL). CS, prospective coronal suture;
EL, ectocranial layer; FB, prospective frontal bone; PB, prospective parietal bone.
Scale bars: 50 μm.
PB
FB
CS
A
PB
FB
B
PB FB
FB PB
Superimposed Superimposed
CS
CS
PB
FB
EL
EL
20
1.2 Jagged1 is required in mesoderm and not in neural crest for coronal suture
development
The expression pattern of Jagged1 suggested that it might function in providing guidance
for the developing bones and the specification of sutural cells. Mice heterozygous for a
conventional Jagged1 knockout allele are viable, but have no detectible defects in skull
vault development (Xue et al., 1999 and our own observations). Jagged1
-/-
mice,
however, showed defects in vasculature remodeling at early embryonic stage. The
embryos die from hemorrhage at E10.5, precluding an examination of Jagged1 function
in the development of the skull vault and sutures at later stages. I therefore used a tissue-
specific conditional knockout approach (Cre-LoxP) to delete Jagged1 in either
mesoderm-derived or neural crest-derived cells and examined their effects on cranial
development. Jagged1 cko mice were generated by Brooker et al. (Brooker et al., 2006).
The mice carry LoxP sites flanked exons encoding the DSL (Delta-Serrate-Lag2) domain,
required for binding to Notch receptors.
According to immunostaining results, Jagged1 was expressed at mesoderm-derived
coronal suture mesenchyme at E14.5. It is known that Mesp1-Cre induces recombination
in all mesoderm-derived tissues (Saga et al., 1999). By crossing to R26R reporter mice
line, Mesp1-Cre activity was confirmed in prospective parietal bone, suture, and in
tissues between the epidermis and prospective bones (Fig. 3A). The attempt to get
Mesp1-Cre;Jagged1
cko/cko
newborn or adult mice were not successful. Of 90 newborn or
P21 pups examined, none of the genotype Mesp1-Cre;Jagged1
cko/cko
was obtained,
suggesting that homozygous inactivation of Jagged1 in mesoderm causes lethality during
21
embryogenesis (Table 4). This outcome is consistent with the known requirement of
Jagged1 in vascular development (Xue et al., 1999). There were few Mesp1-
Cre;Jagged1
cko/cko
embryos able to survive to E14.5. Examination of the developing skulls
of such embryos by whole mount staining for the osteoblast marker, alkaline phosphatase
(ALP), revealed a narrowing of the prospective coronal suture, consistent with early
suture fusion phenotype (Fig. 3B,C, arrow) (Ting et al., 2009).
Fig. 3. Mesp1-Cre mediated conditional inactivation of Jagged1 in mesoderm
causes abnormal coronal suture phenotype at E14.5.
(A) The R26R allele served as an indicator of Mesp1-Cre activation in E14.5
embryos. X-gal staining showed that the Mesp1-driven Cre recombination was
specifically presented in the prospective coronal suture (arrow), parietal bone, and
dermis. (B,C) Mesp1-Cre;Jagged1cko/cko embryos died around E13.5-E15.5. I
examined the morphology of the developing coronal suture at E14.5 by means of
whole mount ALP staining. Mesp1-Cre;Jagged1cko/cko mutants exhibited a
narrowing of the prospective coronal suture (CS). CS, prospective coronal suture;
FB, prospective frontal bone; PB, prospective parietal bone. Scale bars: 50 μm in
A; 500 μm in B,C.
Mesp1-Cre;R26R
FB
PB
FB
PB
A
CS
Mesp1-Cre;Jagged1
cko/cko
PB FB
Wildtype
PB FB
CS
B C
CS
22
Table 4
Offspring obtained from crossing Mesp1-Cre;Jagged1
cko/+
males and
Jagged1
cko/+
or Jagged1
cko/cko
females*
*Out of 47 pups examined at P0 and 43 pups at P21, no Mesp1-Cre;Jagged1
cko/cko
mice
were obtained.
Mesp1-Cre;Jagged1 cko newborn
Mesp1-Cre; Mesp1-Cre; Mesp1-Cre;
Genotypes Jagged1
+/+
Jagged1
+/+
Jagged1
cko/+
Jagged1
cko/+
Jagged1
cko/cko
Jagged1
cko/cko
Total
number of pups 6 7 10 16 8 0 47
Mesp1-Cre;Jagged1 cko P21
Mesp1-Cre; Mesp1-Cre; Mesp1-Cre;
Genotypes Jagged1
+/+
Jagged1
+/+
Jagged1
cko/+
Jagged1
cko/+
Jagged1
cko/cko
Jagged1
cko/cko
Total
number of pups 6 4 12 16 5 0 43
23
As an alternative to Mesp1-Cre, we used Dermo1-Cre to inactivate Jagged1 (Yu et al.,
2003). We were able to recover pups at postnatal day 21 (P21) with the genotype
Dermo1-Cre;Jagged1
cko/cko
(Table 5). Cells carrying the recombined allele were located
preferentially in the mesoderm, and very few LacZ-positive cells were observed
occasionally in frontal bone (neural crest-derived) area (Fig. 4A). Alizarin Red stains of
skulls of Dermo1-Cre/+;Jagged1
cko/cko
mutants revealed synostosis of the coronal sutures
with a penetrance of 86% (n=14) (Fig. 4B,C). The punctate lacZ staining of Dermo1-
Cre;R26R embryos indicates that Cre may not be activated in 100% of the mesodermal
cells (Fig. 4A), and this could reduce the penetrance of the synostosis phenotype. Dermo1
is also known as Twist2, which is a possible craniosynostosis gene. Dermo1-Cre mice
harbor a Cre recombinase knock-in allele. Therefore, it is possible that heterozygous
mutation of Dermo1 contributes to synostosis phenotype. However, Dermo1-
Cre/+;Jagged1
cko/+
mice have no detectible suture defects, suggesting there was no
significant influence of Dermo1 gene in suture formation. Misalignments of upper and
lower incisors as a consequence of craniosynostosis were also noted (Fig. 4D,E).
Table 5
Offspring obtained from crossing Dermo1-Cre;Jagged1
cko/+
males and
Jagged1
cko/+
or Jagged1
cko/cko
females
Dermo1-Cre;Jagged1 cko P21
Dermo1-Cre; Dermo1-Cre; Dermo1-Cre;
Genotypes Jagged1
+/+
Jagged1
+/+
Jagged1
cko/+
Jagged1
cko/+
Jagged1
cko/cko
Jagged1
cko/cko
Total
20 17 36 29 28 14 144
Number of pups
24
Fig. 4. Dermo1-Cre mediated conditional inactivation of Jagged1 causes
craniosynostosis.
(A) The Dermo1-driven Cre activity was detected by X-gal staining at E14.5. The
LacZ-positive cells were located preferentially in the mesoderm, and very few of
them were observed occasionally in frontal bone (neural crest-derived) area. (B,C)
Dermo1-Cre;Jagged1cko/cko mice exhibited synostosis of coronal sutures at
postnatal day (P) 21 (arrows). The skulls were stained with Alizarin Red S to reveal
mineralized bones. (D,E) Misalignments of upper and lower incisors as a
consequence of craniosynostosis were observed in Dermo1-Cre;Jagged1cko/cko
mutant mice at P21. CS, prospective coronal suture; FB, prospective frontal bone;
PB, prospective parietal bone; SS, prospective sagittal suture. Scale bars: 50 μm in
A; 1 mm in B,C.
Dermo1-Cre;Jagged1
cko/cko
Wildtype
PB
FB
SS
CS
B C
Dermo1-Cre;R26R A
FB PB
FB PB
CS
D E Dermo1-Cre;Jagged1
cko/cko
Wildtype
25
Results with Mesp1-Cre and Dermo1-Cre suggested that Jagged1 is required in the
mesoderm to maintain the suture in an open, non-ossified configuration. To test whether
Jagged1 also has a role in the neural crest-derived portion of the skull and in the coronal
suture, we made use of Wnt1-Cre. The R26R marker allele revealed, as expected from
previous results (Merrill et al., 2006), that Wnt1-Cre-mediated recombination took place
with high efficiency in the prospective frontal bone and meninges (Fig. 5A). We
produced mice with the genotype, Wnt1-Cre;Jagged1
cko/cko
(Table 6). Such mice were
viable at least through P21. Their skulls showed no evidence of synostosis of the coronal
suture (Fig. 4B,C). However, ossification defects were apparent in the frontal bone (Fig.
5B, arrow).
Table 6
Offspring obtained from crossing Wnt1-Cre;Jagged1
cko/+
males and
Jagged1
cko/+
or Jagged1
cko/cko
females
Wnt1-Cre;Jagged1 cko P21
Wnt1-Cre; Wnt1-Cre; Wnt1-Cre;
Genotypes Jagged1
+/+
Jagged1
+/+
Jagged1
cko/+
Jagged1
cko/+
Jagged1
cko/cko
Jagged1
cko/cko
Total
12 16 32 35 14 16 125 Number of pups
26
Fig. 5. Jagged1 is not required in the neural crest for coronal suture
development.
(A) Wnt1-Cre-mediated recombination took place in the prospective frontal bone
and meninges (M) (B,C) Wnt1-Cre;Jagged1
cko/cko
mice show normal coronal sutures,
but an ossification defect was apparent in the frontal bone (arrow). CS, prospective
coronal suture; FB, prospective frontal bone; M, meninges; PB, prospective parietal
bone. Scale bars: 50 μm in A; 1mm in B,C.
Wnt1-Cre;R26R
PB
FB
A
CS
M
FB
PB
Wildtype Wnt1-Cre;Jagged1
cko/cko
B C
PB
FB
SS
CS
27
Our previous findings demonstrated that the boundary between osteogenic and non-
osteogenic cells in the coronal suture has a crucial role in craniosynostosis. The mixing of
mesoderm-derived and neural crest-derived cells was observed at the boundary by using
either Wnt1-Cre;R26R or Mesp1-Cre;R26R labeling system in synostosis embryos
(Twist1
+/-
and EphA4
+/-
) (Merrill et al., 2006; Ting et al., 2009). Together with the well-
documented role of Notch signaling in boundary formation, these findings prompted us to
test for boundary defects within the coronal sutures of Jagged1 conditional mutants. To
examine the cell mixing phenotype, I crossed R26R into Mesp1-Cre;Jagged1
cko/cko
mice
and examined the distribution of lacZ expressing cells in the coronal suture. Mesp1-Cre
here serves both the conditional knockout and the labeling system. As is evident in Fig.
6B, lacZ positive cells crossed from the mesoderm compartment to the crest compartment
in substantial numbers. Thus, Jagged1 is required in the mesoderm to maintain the
integrity of the boundary between osteogenic and non-osteogenic cells in the coronal
suture.
28
Fig. 6. Cell-mixing at mesoderm-neural crest boundary in the coronal suture of
Mesp1-Cre;Jagged1
cko/cko
embryos.
(A) The R26R allele served as an indicator of Mesp1-Cre activation in E14.5
embryos. In control embryos, Mesp1-Cre recombination was restricted in the
prospective coronal suture (arrow), parietal bone, and dermis. A diagram was
shown in the bottom panel for clarity. (B) Inactivation of Jagged1 in mesoderm by
Mesp1-Cre caused boundary defects. Note the lacZ positive cells in the frontal
bone territories (arrowheads). CS, prospective coronal suture; FB, prospective
frontal bone; PB, prospective parietal bone. Scale bars: 50 μm.
Mesp1-Cre;Jagged1
cko/cko
;R26R B Mesp1-Cre;R26R
FB
PB
FB
PB
A
CS
FB
PB
FB
FB
PB
FB
PB
PB
CS
29
1.3 Inactivation of Jagged1 in mesoderm affects the expression patterns of Notch2
and Hes1 in coronal suture
Jagged1 functions as a ligand for Notch signaling. Among the four known Notch
receptors, mutations in NOTCH2 were found in some cases of the Alagille syndrome
patients who had no detectable JAGGED1 mutations (McDaniell et al, 2006). Notch2 also
works as a genetic modifier with Jagged1 in the mice model of Alagille syndrome
(McCright et al., 2002). In addition, it is known to function cooperatively with Jagged1
in the development of bile ducts (Lozier et al., 2008) and lens fiber cells (Saravanamuthu
et al., 2009), and has been shown to be negatively regulated by Jagged1 (Yuan et al.,
2006). This potential functional relationship between Jagged1 and Notch2 led us to
assess the influence of Jagged1 on the expression of Notch2.
In wild type embryos at both E13.5 and E14.5, Notch2 was expressed in the prospective
frontal and parietal bones in comparison with ALP staining of adjacent sections and was
excluded from the coronal suture (Fig. 7A). In Mesp1-Cre and Dermo1-Cre;Jagged1
cko/cko
mutants, Notch2 was expanded into the suture (Fig. 7B,D, arrows). This expansion did
not occur in Wnt1-Cre;Jagged1
cko/cko
mutants (Fig. 7C, arrowheads). These results can be
correlated with the occurrence of the synostosis phenotypes as in Fig. 3-5. It suggests that
loss of Jagged1 in sutural cells might change their identities and therefore obliterate the
boundary between osteogenic and non-osteogenic cells.
30
Fig. 7. Inactivation of Jagged1 in mesoderm causes an expansion of Notch2
expression in the prospective coronal suture.
Analysis of Notch2 expression by immunostaining at E13.5 and E14.5 embryos.
(A,C) Notch2 was expressed in the prospective bone territories and was excluded
from the coronal suture (arrowheads) in both wildtype and Wnt1-Cre;Jagged1
cko/cko
embryos. (B,D) Conditional knockout of Jagged1 in mesoderm by Mesp1-Cre or
Dermo1-Cre resulted in an expansion of Notch2 expression (arrows). CS,
prospective coronal suture; FB, prospective frontal bone; PB, prospective parietal
bone. Scale bars: 50 μm.
PB
FB
CS
B
Mesp1-Cre;Jagged1
cko/cko
FB
PB
CS
FB
PB
A Wildtype
CS
FB
PB
CS
PB
FB
D Dermo1-Cre;Jagged1
cko/cko
FB
PB
CS
PB
FB
CS
C Wnt1-Cre;Jagged1
cko/cko
FB PB
CS
E13.5
E14.5
E13.5
E14.5
CS
31
We also checked the expression of Hes1, a downstream effector of Notch signaling
(Jarriault et al., 1995), as an indicator of Notch signaling activity. Hes1 exhibited a
similar expression pattern to that of Notch2 in wildtype and Dermo1-Cre;Jagged1
cko/cko
mutants, respectively (Fig. 8A,B), suggesting that inactivation of Jagged1 resulted in an
increase in Notch signaling in the sutural mesenchyme.
Fig. 8. The expression of Hes1, a downstream target of Notch signaling, was
consistent with that of Notch2 in Dermo1-Cre;Jagged1
cko/cko
mutant mice.
(A,B) Hes1 acts as an indicator of Notch signaling activity. Hes1 immunostaining
confirmed that this downstream target exhibited a similar expression pattern to that
of Notch2 in wildtype and Dermo1-Cre;Jagged1
cko/cko
embryos, respectively. CS,
prospective coronal suture; FB, prospective frontal bone; PB, prospective parietal
bone. Scale bars: 50 μm.
FB
PB
Wildtype
CS
Dermo1-Cre;Jagged1
cko/cko
PB FB CS
B A
E14.5
32
In addition to Jagged1, there are other ligands that might have influences on the Notch
signaling in the coronal suture. I checked the expression of Dll1,2 and 4 and found that
Dll1 protein was present in the bones and sutural cells (Fig. 9A). There was no change in
Dll1 expression in the Dermo1-Cre;Jagged1
cko/cko
mutants (Fig. 9B). Thus in the absence
of Jagged1, Dll1 might be available to serve as a ligand for Notch2.
Fig. 9. Notch ligand, Dll1, was expressed in the mid-sutural mesenchyme.
(A,B) Dll1 immunostaining studies were performed in coronal suture at E14.5. Dll1
protein was present in the prospective bones and sutural cells (arrows). No
significant difference between wildtype and mutant was found at this stage. CS,
prospective coronal suture; FB, prospective frontal bone; PB, prospective parietal
bone. Scale bars: 50 μm.
Dermo1-Cre;Jagged1
cko/cko
PB
FB PB
FB
B A Wildtype
CS
CS
PB FB
PB
FB
E14.5
Superimposed Superimposed
33
1.4 Crosstalk between Notch and other signaling pathways in the developing suture
There are several signaling pathways that are known to be involved in craniosynostosis. I
examined whether loss of Jagged1 function in mesoderm affects the activities of the
RTK, BMP, and Wnt/β-catenin pathways in the coronal suture. These pathways are
known to function during osteogenic differentiation, and mutations in the pathway
components cause craniosynostosis (Morriss-Kay and Wilkie, 2005).
1) RTK (receptor tyrosine kinase) pathway
Previous work in our lab identified ephrin-A2 proteins was expressed in a single layer of
cells on the ectocranial side of the prospective frontal bone (Fig. 10A). In Twist1 mutants,
its expression is restricted to the anterior (closer to eyes) and does not extend into the
coronal suture region (Merrill et al., 2006). In Dermo1-Cre;Jagged1
cko/cko
mutants, the
pattern of ephrin-A2 was similar to that in the wildtype (Fig. 10B). I also examined P-
Erk1/2, a well-characterized downstream effector of RTK activation. Previous studies
showed an altered expression of P-Erk1/2 in EphA4 mutants, which exhibit
craniosynostosis and defects in the osteogenic-non-osteogenic boundary (Ting et al.,
2009). In wildtype embryos, P-Erk1/2 was expressed in the prospective bones and
osteogenic fronts, but not in the suture. In Dermo1-Cre;Jagged1
cko/cko
mutants, P-Erk1/2
expression expanded toward suture, but cannot be detected in the mid-suture
mesenchyme (Fig. 10C,D). The expression coincided with the ALP expression and did
not resemble the patterns of Notch2 and Hes1 in the mutants.
34
Fig. 10. RTK signaling pathway is not altered in Jagged1 mutant embryos at
E14.5.
(A,B) Ephrin-A2 protein was expressed in a single layer of cells on the ectocranial
side of the prospective frontal bone. The expression extended into coronal suture
region (arrowheads) both in the wildtype and mutants. (C,D) P-Erk1/2 acts as a
downstream effector of RTK activation. Similar expression patterns of P-Erk1/2
were found in wildtype and mutant. CS, prospective coronal suture; FB, prospective
frontal bone; PB, prospective parietal bone. Scale bars: 50 μm.
PB FB
PB
FB
Wildtype Anti-
P-Erk1/2
CS
CS
PB
FB
PB FB
Anti-
ephrinA2
CS
CS
Superimposed Superimposed
Dermo1-Cre;Jagged1
cko/cko
35
2) BMP pathway
BMP pathway is involved in the specification and differentiation of osteogenic cells.
Mutations in BMP signaling components, including TGFBR1, TGFBR2, Msx2 and Gdf6,
lead to abnormal cranial suture fusion. I found that the distribution of P-Smad1/5/8
expressing cells was altered in Dermo1-Cre;Jagged1
cko/cko
mutants in a manner similar to
the changes described previously in Twist1-EphA4 mutants (Ting et al., 2009). Whereas
in wild type sutures, P-Smad1/5/8 positive cells were concentrated in the osteogenic
fronts; in mutants, they were distributed throughout the suture (Fig. 11A,B).
3) Wnt pathway
Canonical Wnt pathway also plays a role in cranial suture formation (Yu et al., 2005).
The expression of β-catenin, an essential downstream effector of Wnt signaling, was
altered in Dermo1-Cre;Jagged1
cko/cko
mutants (Fig. 11C,D). Few β-catenin positive cells
were apparent in the sutural mesenchyme of wildtype embryos. In Dermo1-
Cre;Jagged1
cko/cko
mutants, ectopic expression of β-catenin was detected in the coronal
sutural mesenchyme.
Together, these results suggested that Jagged1 acts in parallel with or downstream of
RTK signaling, and upstream of canonical BMP and Wnt/β-catenin signaling.
36
Fig. 11. Both BMP and Wnt signaling pathways were affected in Dermo1-
Cre;Jagged1
cko/cko
mutants.
(A,B) The distribution of P-Smad1/5/8 expressing cells was altered in the coronal
suture of Dermo1-Cre;Jagged1
cko/cko
mutants. The signals were distributed
throughout the suture in the mutants whereas in the wildtype, they became more
concentrated in the osteogenic fronts. (C,D) Ectopic expression of β-catenin was
detected in the coronal sutural mesenchyme in the mutants (arrow). In wildtype, it
was restricted to the prospective bone area. CS, prospective coronal suture; FB,
prospective frontal bone; PB, prospective parietal bone. Scale bars: 50 μm.
Wildtype Dermo1-Cre;Jagged1
cko/cko
Anti-
P-Smad1/5/8
PB
FB
Wildtype Dermo1-Cre;Jagged1
cko/cko
PB
FB
Anti-
-Catenin
CS
CS
CS CS
PB
FB
PB FB
37
2. Jagged1 is a downstream effector of Twist1 in suture development
2.1 The Notch signaling is altered in Twist1 mutants
There are a couple of facts that suggest a genetic interaction between Jagged1 and Twist1
genes. First, the overlapped Twist1 and Jagged1 expression in the coronal suture (Fig. 2).
Second, the similar synostosis phenotypes in patients with Twist1 or Jagged1 mutations,
and in their transgenic mice models as well. Heterozygous loss of Twist1 in mice exhibits
more sever suture phenotypes than Jagged1 homozygous mutants, I therefore first
assessed the influence of Twist1 on the expression of Jagged1, Notch2 and Hes1. The
analysis was performed at several stages of coronal sutural development, including E12.5
and E13.5 prior to any boundary defects in Jagged1 or Twist1 mutant embryos (Figs 12
,13), as well as E14.5 when such defects are first evident (Fig. 14).
At E12.5, the osteogenic fronts of frontal and parietal bones, which were shown by the
expression of ALP in adjacent section, are far from each other and the cells in between
could be osteogenic or non-osteogenic. The exact position of the suture mesenchymal
cells is hard to be identified. At this stage, Jagged1 immunostaining revealed that
Jagged1 protein was located in the frontal and parietal bone territories and its expression
extended beyond the ALP domain in the prospective parietal bone, into an area of
prospective osteogenic tissue and further into prospective coronal suture area (Fig. 12A).
Notch2 expression overlapped with ALP activity and was detectable but very weak in the
prospective coronal suture (Fig. 12E).
38
In Jagged1
+/-
embryos, Jagged1 expression was downregulated (Fig. 12B), indicating
heterozygous loss of Jagged1 was significantly correlated with Jagged1 protein
expression. Jagged1 expression changed little and seems to be weaker in Twist1
+/-
mutant
embryos at this stage (Fig. 12C), but Notch2 expression clearly expanded into the sutural
mesenchyme (Fig. 12G). Combination Twist1
+/-
;Jagged1
+/-
mutants exhibited a reduction
in Jagged1 activity and an expansion of Notch2 activity similar to that of Twist1
+/-
embryos (Fig. 12D,H).
Fig. 12. Twist1 regulates Notch signaling pathway in coronal suture at E12.5.
The expression of Jagged1 (A-D) and Notch2 (E-F) proteins was examined at
E12.5. (A) Jagged1 expression extended from the parietal bone into the coronal
suture, and then further into the frontal bone. The expression changed little but was
downregulated in other genotypes (B-D). Notch2 was expressed predominantly at
bone area in the wildtype (E), whereas its expression expanded into the sutural
mesenchyme in the Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mutants (G,H, arrows). CS,
prospective coronal suture; FB, prospective frontal bone; PB, prospective parietal
bone. Scale bars: 50 μm.
Jagged1
+/-
PB
FB
Twist1
+/-
FB
Twist1
+/-
;Jagged1
+/-
PB
FB
FB
PB
D C B
PB
FB
PB
FB PB
H G F
Wildtype
PB
A
PB
E
FB
Anti-
Jagged1
Anti-
Notch2
FB
39
In wildtype embryos at E13.5, Jagged1 expression in the parietal bone territory divided
into two bands of cells flanking the prospective parietal bone (Fig. 13A, arrows). The
lower band was continuous across the coronal suture and joined to the upper band
ectocranial to the developing frontal bone. Notch2 was expressed in the prospective bone
territories and was excluded from the prospective coronal suture (Fig. 13E). Hes1 showed
a similar but relatively weaker expression compared with Notch2 in both the frontal and
parietal bones and was also excluded from the prospective coronal suture (Fig. 13I).
These patterns of Jagged1, Notch2 and Hes1 were similar in Jagged1
+/-
mutants (Fig.
13B,F,J), but significantly different in Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mutants at E13.5.
Jagged1 expression in the endocranial band reduced significantly in Twist1
+/-
embryos
(Fig. 13C, arrowhead). The superimposed image showed that loss of the lower band
seemed to affect the distribution of ALP-expressing cells. The expression of Jagged1 was
largely lost in Twist1
+/-
;Jagged1
+/-
mutants (Fig. 13D, arrowheads). In Twist1
+/-
and
Twist1
+/-
;Jagged1
+/-
mutants, both Notch2 and Hes1 expanded into the coronal suture
(Fig. 13G,H,K,L).
40
Fig. 13. Jagged1 exhibits a specific expression pattern in coronal suture at E13.5.
(A) Immunostaining showed a specific pattern of Jagged1, in which two bands flanked
the prospective parietal bone (arrows) and the lower band was across the coronal suture
and joined to the upper band ectocranial to the developing frontal bone. (C) The lower
band was almost lost in Twist1
+/-
mutants (arrowhead). (D) The signals in double mutants
were largely reduced. Notch2 (E-H) and Hes1 (I-L) were also examined. Expansion of
the Notch2- or Hes1-positive cells was observed in the sutures of Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mutants (G,H,K,L, arrows). CS, prospective coronal suture; FB, prospective
frontal bone; PB, prospective parietal bone. Scale bars: 50 μm.
PB
Jagged1
+/-
PB
Twist1
+/-
Twist1
+/-
;Jagged1
+/-
FB
FB PB
FB PB FB
PB
PB
FB
PB
PB
FB
FB
PB FB PB
FB
FB
FB PB
PB
Superimposed Superimposed Superimposed Superimposed
G F E
D C B A
J I
H
K L
CS CS CS
CS
CS CS CS CS
CS CS CS CS
FB
Wildtype
PB
PB
FB PB FB
FB
PB FB
Anti-
Jagged1
Anti-
Hes1
Anti-
Notch2
41
Similarly at E14.5, Notch2 and Hes1 continued to be expressed in the osteogenic layer in
wildtype and Jagged1
+/-
embryos (Fig. 14E,F,I,J). In Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
compound mutants, Notch2 and Hes1 expanded significantly into the Jagged1 domain in
the midsutural mesenchyme (Fig. 14G,H,K,L). Taken together, these data suggested that
Twist1 controls the distribution of Jagged1-, Notch2- and Hes1-expressing cells.
Moreover, the expansion of Notch2 presages changes in ALP activity that accompany
craniosynostosis at E12.5 and E13.5. The expression of Twist1 was also examined and no
difference was found in Jagged1
+/-
mutant mice compared to wildtype (Fig. 15A,B).
Fig. 14. Heterozygous loss of Twist1 caused boundary defects in coronal suture
at E14.5.
The expression of Jagged1 (A-D), Notch2 (E-F) and Hes1 (I-L) was examined in
the coronal suture. (A-D) The embryos with the indicated genotypes also carried
Wnt1-Cre;R26R transgenic alleles. X-gal staining revealed the boundary crossing
neural crest-derived cells in the sutural mesenchyme in the Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mutants (C,D, arrowheads). Notch2 (E-H) and Hes1 (I-J) displayed
similar expression patterns in the suture. Heterozygous loss of Twist1 caused
expansion of the two genes in the suture (G,H,K,L, arrows). CS, prospective
coronal suture; FB, prospective frontal bone; PB, prospective parietal bone. Scale
bars: 50 μm.
FB
PB
PB
FB
FB
PB
PB
FB
FB
PB
FB
PB
PB
PB
FB
FB
PB
PB
FB PB
FB
PB FB
PB
FB PB
Wildtype Jagged1
+/-
PB
FB
Twist1
+/-
FB
PB
Twist1
+/-
;Jagged1
+/-
PB
FB
FB
PB
PB FB
FB
PB
Superimposed Superimposed Superimposed Superimposed
CS
CS
CS
CS
CS
CS
CS
CS
CS
CS
CS CS
A
K J I
H G F E
D C B
L
FB
FB
Anti-
Jagged1
Anti-
Hes1
Anti-
Notch2
42
Fig. 15. Twist1 expression was not influenced by Jagged1 dosage.
(A,B) Heterozygous loss of Jagged1 did not affect the Twist1 expression in coronal
suture at E14.5. CS, prospective coronal suture; FB, prospective frontal bone; PB,
prospective parietal bone.
Wildtype PB
FB
PB
FB
Jagged1
+/-
CS
CS
43
2.2 Reduced Jagged1 dosage exacerbates the osteogenic-non-osteogenic boundary
defect in Twist1 mutant mice
Homozygous loss of Jagged1 in mesodermal cells (Dermo1-Cre;Jagged1
cko/cko
mice)
resulted in premature suture fusion and led to cell mixing at mesoderm-neural crest
boundary (Fig. 6B). By using Wnt1-Cre;R26R reporter system, we could trace the cell
population of the prospective frontal bone (neural crest-derived) in the wildtype or
mutant embryos, and also observe the intactness of the mesoderm-neural crest boundary.
As in Fig. 14C and D, a small number of lacZ positive cells detached from continuous
cell population and crossed into the coronal suture compartment in both Twist1
+/-
and
Twist1
+/-
;Jagged1
+/-
mutants at E14.5. Previous studies showed that the crossing
boundary phenotype was not found at earlier stage in Twist1 mutant embryos (Merrill et
al., 2006).
To determine whether there was an increase in the severity of the boundary defect in
Twist1
+/-
;Jagged1
+/-
compared with Twist1
+/-
mutants, I observed the boundary in serial
transverse sections across the coronal suture in several embryos and presented the results
by counting the percentage of sections with crossing boundary phenotype in each
genotypes (Table 7). In Twist1
+/-
;Jagged1
+/-
mutants, there was a significant increase in
the proportion of the coronal suture over which boundary defects were evident. Thus
reduced Jagged1 function in the context of the heterozygous Twist1 mutant further
compromised the osteogenic-non-osteogenic boundary.
44
Table 7
Percentage of sections with boundary crossing neural crest
in coronal mid-suture mesenchyme*
* All of the embryos with indicated genotypes carried Wnt1-Cre;R26R alleles as well. 10-
µm coronal sections through the heads of E14.5 embryos were collected and stained with
X-gal followed by Jagged1 immunostaining as seen in Fig. 13A-D. Twenty sections from
similar level of the heads were selected for each embryo and the percentage of sections
with boundary crossing neural crest in coronal mid-suture mesenchyme (as indicated by
arrowheads in Fig. 13C,D) were calculated. Adjacent sections were stained for ALP for
determining presumptive coronal sutures. All pairwise comparisons (two-tailed Student's
t-test) among means were significant different (P<0.05), except for Wildtype vs.
Jagged1
+/-
.
2.3 A significant genetic interaction between Jagged1 and Twist1
The results of immunostaining and boundary defect implied that Jagged1 and Twist1
might function together in regulating suture development. To test this possibility, I
crossed Jagged1
+/-
heterozygous mice with Twist1
+/-
heterozygous mice, and compared
the phenotypes of their offspring at P21. Homozygous mutants are not available since
Jagged1
-/-
mice at E10.5 and Twist1
-/-
mice die at E11.5. As mentioned earlier, there is no
detectable suture phenotype in Jagged1 heterozygous mutants. In Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mice, synostoses were exhibited not only in the coronal sutures but also in
other types of sutures (Fig. 16). Synostoses were found to occur in the following five
45
cranial sutures -- coronal, squamosal, lambdoid, occipitointerparietal (OIP), and
interfrontal sutures. In Twist1
+/-
;Jagged1
+/-
mice, the penetrance of synostosis was
increased in all five sutures in comparison to Twist1
+/-
mice (Table 8). The total
penetrance reached 100% in the heterozygous compound mutants.
Fig. 16. Genetic interaction between Twist1 and Jagged1 and its effect on
craniosynostosis .
The skulls of P21 mice were stained with Alizarin Red S. (A-D) Synostoses of
coronal (arrows) and lambdoid (arrowhead) sutures were observed in Twist1
+/-
and
Twist1
+/-
;Jagged1
+/-
mice. The compound mutants also showed sever fusion
phenotypes in occipitointerparietal (OIP) (E,F, arrows) and squamosal (G,H,
arrows) sutures. CS, prospective coronal suture; FB, prospective frontal bone; LS,
lambdoid suture; OIP, occipitointerparietal suture; PB, prospective parietal bone;
SQ, squamosal suture. Scale bars: 1 mm.
Twist1
+/-
;Jagged1
+/-
Twist1
+/-
PB
FB
SS
LS
OIP
IF
PB
FB
CS
SS
LS
OIP
IF
Wildtype Twist1
+/-
;Jagged1
+/-
CS
PB
SQ
LS
Wildtype
PB
FB
SS
CS
LS
A Jagged1
+/-
B
C
F
D
LS
OIP
PB
LS
SQ
E
G
H
Wildtype Twist1
+/-
;Jagged1
+/-
46
*The percentage of postnatal day 21 mice with suture fusion phenotypes was examined.
The synostosis phenotypes of five different sutures (totally eight sutures), including left
coronal, right coronal, left squamosal, right squamosal, left lambdoid, right lambdoid,
occipitointerparietal (OIP) and interfrontal suture, were examined under microscope on
alizarin red S-stained skulls.
**L+R: Left+Right sutures
Table 8
Penetrance of craniosynostosis*
47
Table 9
Craniosynostosis index (CI)*
Penetrance represents the percentage of gene carriers who show a certain phenotype.
From the result, we are not be able to tell the severity of the phenotype exhibited in the
carrier. Therefore, I used a quantitative assessment, the craniosynostosis index (CI)
(Oram and Gridley, 2005) to measure the severity of the synostosis phenotypes (Table 9).
The CI scores were given to the five sutures mentioned above (For further details of the
method, please see Table 9). The CI for each individual suture was increased
significantly in Twist1
+/-
;Jagged1
+/-
mutants compared with other genotypes. Whereas the
sum score was 3.8±2.3 for Twist1
+/-
mice, it was 10.1±2.6 for Twist1
+/-
;Jagged1
+/-
compound heterozygous mutants. These results are consistent with a strong genetic
interaction between Twist1 and Jagged1 in the developing skull vault.
* The Craniosynostosis index (CI) is a scoring method adopted from the following paper
-- Oram and Gridley, 2005. Eight sutures as indicated in Table 5 were scored for each
skull. The degree of fusion was observed under the microscope and a score was given to
each suture according to the “score index” (0: unfused; 1: <50% fused; 2: ≥50% fused
and 3: 100% fused.) Two scores for left and right sutures of the same type were added
together. For example, the maximum score given to two coronal sutures in each skull is
six. The mean score±s.e.m. for each type of sutures were calculated separately. The sum
score of the eight sutures represents the severity of craniosynostosis in each genotype.
Pairwise comparisons among four genotypes were analyzed by two-tailed Student's t-test
and significant differences are marked with (§) for P<0.05 or (§§) for P<0.000001.
** L+R: Left+Right sutures
48
Jagged1 and Twist1 function cooperatively in tissues other than cranial sutures. It has
been reported that Twist1
+/-
mutants exhibited polydactyly of the hind feet. Heterozygous
loss of Jagged1 in Twist1
+/-
mutants increases the penetrance of such phenotype from
5.6% to 13.6%. Dr. Gage Crump also helped me to identify the abnormal phenotypes of
retrotympanic process (RTP) of the squamosal bones and the middle ear ossicles.
Delayed ossification of the RTP was observed in Twist1
+/-
and Twist1
+/-
;Jagged1+/- mice
at both newborn and P21 stage (Fig. 17). The size of RTP was measured at P21 and a
significant reduction in the size of RTP was found in Twist1
+/-
;Jagged1
+/-
mutants when
compared with Twist1
+/-
mice
(Table 10).
Table 10
Measurements of the size of retrotympanic process*
* The photos of retrotympanic process (RTP) of the squamosal bones and hemacytometer
were taken under the microscope under the same conditions. Pixels of the selected area
on the photos were counted by Adobe Photoshop. Measurement was extrapolated by the
known size of hemacytometer square. All pairwise comparisons (two-tailed Student's t-
test) among means were significant different (P<0.05), except for Wildtype vs. Jagged1
+/-
. (n=5 for each genotype and each mouse has two RTPs)
Genotype Wildtype Jagged1
+/-
Twist1
+/-
Twist1
+/-
;Jagged1
+/-
Size of RPT (mm
2
) 0.88±0.22 0.81±0.15 0.59±0.12 0.38±0.23
49
Fig. 17. The size of retrotympanic process (RTP) of the squamosal bones was
reduced in Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mutants. Alizarin red staining of the
mutant mice showed a reduced calcified area (arrows) at the RTP at both P0 (A-D)
and P21 (E-H).
The middle ear contains three tiny bones known as the ossicles: malleus, incus, and
stapes. Among the three of them, malleus looked normal in each genotype. There were
four types of middle ear ossicles malformations found in the mutant mice. Two of them
suggest the genetic interaction between Twist1 and Jagged1. The penetrance of extra
process and fragmentation of incudomalleal joint of the incus were increased in Twist1
+/-
;Jagged1
+/-
mice, compared to Twist1
+/-
mice (Fig. 18A-C) (Table 11). However, the
other two phenotypes including bobble-shaped incudomalleal joint of incus (Fig. 18D)
and stapes with small or filled lumen (Fig. 18E-H) only occurred in Jagged1
+/-
mutants.
Wildtype Jagged1
+/-
A
D
C B
Twist1
+/-
;Jagged1
+/-
Twist1
+/-
E H G F
P0
P21
50
Fig. 18. The malformations of middle ear ossicles. (A) a normal incus from a
wildtype P21 mouse. Incudes with extra process (B) or with a broken incudomalleal
joint (C) were found in Twist1
+/-
and Twist1
+/-
;Jagged1
+/-
mutants. (D) The bobble-
shaped incudomalleal joint phenotype was only found in Jagged1
+/-
mutants. (E,F)
Stapes were smaller and with a closed lumen in Jagged1
+/-
mutants at P21. (G,H)
The newborn skulls stained with Alizarin Red and Alcian Blue showed that the
stapedial artery, a small artery that passes through the lumen of the stapes was
missing (arrowhead).
Table 11
Middle ear ossicle malformations in Twist1
+/-
; Jagged1
+/-
cross at P21
Wildtype Jagged1
+/-
A D C B
Twist1
+/-
;Jagged1
+/-
E H G F
Twist1
+/-
;Jagged1
+/-
Wildtype Wildtype Jagged1
+/-
Jagged1
+/-
Stapes
extra process of the fragnemtation of the bobble-shaped
incudomalleal joint incudomalleal joint incudomalleal joint
No. of affected No. of affected No. of affected No. of affected
Wildtype 18 0 (0%) 0 (0%) 3 (17%) 0 (0%)
Jagged1
+/-
20 0 (0%) 0 (0%) 15 (75%) 5 (25%)
Twist1
+/-
16 6 (38%) 2 (13%) 0 (0%) 0 (0%)
Twist1
+/-
;Jagged1
+/-
13 9 (69%) 3 (23%) 0 (0%) 1 (8%)
Incus
small or no lumen
Phenotypes
n
51
We next used the conditional Jagged1 allele, together with Wnt1-Cre or Mesp1-Cre, and
the conventional Twist1 allele, to compare the strength of the interaction between
Jagged1 and Twist1 in neural crest and mesoderm. From data presented in Table 12, mice
with the genotype Mesp1-Cre;Jagged1
cko/+
;Twist1
+/-
exhibited a significantly greater
craniosynostosis index than mice with the Twist1 heterozygous knockout alone. In
contrast, when Wnt1-Cre was used in place of Mesp1-Cre, there was no indication of an
interaction between Twist1 and Jagged1. These data suggest that Jagged1 is required in
mesoderm for its interaction with Twist1.
* The skulls were collected at postnatal day 21 and stained with Alizarin Red S. The
Craniosynostosis index (CI) was calculated as in Table 3. All pairwise comparisons (two-
tailed Student's t-test) were significant different (P<0.05), except for the following: (1)
Mesp1-Cre vs. Mesp1-Cre;Jagged1
cko/+
, (2) Wnt1-Cre vs. Wnt1-Cre;Jagged1
cko/+
, and (3)
Wnt1-Cre;Twist1
+/-
vs. Wnt1-Cre;Jagged1
cko/+
;Twist1
+/-
. (P>0.1)
wildtype mutants
Table 12
Craniosynostosis index (CI) for coronal sutures*
52
Conclusion
Jagged1 functions in boundary formation and is responsible for Alagille syndrome,
which has craniosynostosis as a feature (Kamath et al., 2002). I demonstrated the roles of
the Notch signaling in craniosynostosis and in the formation of a boundary in the coronal
suture. Jagged1 exhibits a specific expression pattern in the coronal suture. Therefore, by
using conditional gene targeting in mice, I showed that inactivation of Jagged1 in the
suture mesodermal cells resulted in craniosynostosis. The molecular signals in the sutural
cells changed at E12.5 and E13.5 prior to the stage when cell-mixing at the osteogenic-
non-osteogenic boundary in the coronal suture can be detected (Merrill et al., 2006). I
also found that Twist1 controls Jagged1 expression in the prospective coronal suture and
Twist1 and Jagged1 exhibit a strong genetic interaction. Our results thus link Alagille and
Saethre-Chotzen syndromes at the pathophysiological level.
1. Sutural cells identity was changed in early development of the skull
It has been suggested that the time of fusion of the coronal suture in Apert syndrome
skulls is at 15 weeks of gestation according to measurement of the distance between the
mineralization centers of the frontal and parietal bones in comparison with the
progressive separation of these centers in a series of fetal skulls (Mathijssen et al., 1999).
Earlier work from our lab showed that heterozygous loss of Twist1 function causes
defects in the boundary between osteogenic and non-osteogenic cells in the coronal
suture as early as E14.5 (Merrill et al., 2006). Recent study indicated that mice carrying
the Apert Fgfr2S252W mutant allele exhibit osteogenic differentiation of the sutural
53
mesenchyme at E13.5 (Holmes et al., 2009). My observation sets back the time of
detection of a change in sutural cells in a craniosynostosis disorder to E12.5 with the
finding of upregulation of Notch2. At E13.5, a corresponding increase in the activity of
the Notch effector, Hes1 was also detected.
2. Notch signaling in suture patterning
Jagged1 and Notch2 have a complementary expression pattern in the wildtype suture.
Interestingly, this complementary pattern of Notch ligand and receptor can be observed in
many other tissues, including Drosophila leg segmentation (de Celis et al., 1998), chick
limb (Vargesson et al., 1998), mouse dentate gyrus of the brain (Stump et al., 2002),
mouse kidney glomeruli (McCright et al., 2001), mouse small intestine (Schröder and
Gossler, 2002) and human corneal epithelial cells (Ma et al., 2007). In mouse coronal
suture, the expression of Jagged1 and Notch2 implied their patterning roles in suture
formation. At E13.5 and E14.5, Jagged1 was expressed in the sutural mesenchyme and
the cells immediately adjacent to the osteogenic territory, suggesting it may maintain a
boundary between the osteogenic and non-osteogenic compartments of the coronal
suture. Loss of Jagged1 in mesoderm-derived sutural cells resulted in boundary defects
(cells across boundary) and craniosynostosis.
Ectopic expressions of Notch2 and Hes1 were found in the early development of coronal
sutural cells in the mutants. Notch signaling has been reported to modulate osteogenesis
in many types of cells. Increased Notch signaling can promote osteogenic differentiation
in MC3T3-E1 osteoblastic cells and primary human bone marrow mesenchymal stem
54
cells (hMSCs) (Tezuka et al., 2002; Nobta et al., 2005). Recent study also suggested that
Notch signaling can induce osteogenic differentiation of human aortic smooth muscle
cells (HASMCs) through upregulation of Msx2 (Shimizu et al., 2009). In addition, Hes1
can serve as a coactivator for Runx2 in rat ROS17/2.8 osteoblastic cells (McLarren et al.,
2000) and MC3T3-E1 cells (Suh et al., 2008). Therefore upregulation of Notch signaling
in the coronal sutures of Jagged1 mutants is consistent with the subsequent change in
sutural cells to an osteogenic identity. Conversely exclusion of Notch activity from
sutural cells may serve to maintain such cells in a non-osteogenic state.
Mutation in NOTCH2 has been reported in some cases of Alagille syndrome (McDaniell
et al., 2006). Patients with NOTCH2 mutations have a spectrum of defects that are similar
to those with JAGGED1 mutations. However, these defects do not include
craniosynostosis. Notch2 mutant mice also don’t have detectable suture abnormality
(McCright et al., 2002). According to the observation that Notch2 is upregulated in the
mutant sutural cells, this result is predictable. It is likely that gain of function rather than
loss of function mutations in Notch2 would cause craniosynostosis. Thus far germline
gain of function mutations in NOTCH2 have not been reported.
3. Notch signaling in Drosophila wing imaginal disc is comparable to that in coronal
suture
Notch pathway functions during diverse developmental and physiological processes. Its
functions can be classified into three categories: 1) lateral inhibition (for example,
differentiation of neural precluster cells); 2) lineage decisions (ex., neuron-glia fate
55
determination) and 3) boundaries/inductive (ex., D/V boundary of Drosophila wing
imaginal disc) (Bray, 2006). A major function of compartment boundaries is to establish
specialized border cells between the two compartments (Irvine and Rauskolb, 2001;
Blair, 2003). These border cells separate two compartments and provide morphogens that
influence their fate. The Notch pathway has a prominent role in the development of such
cells in several developmental settings. For examples, Notch signaling mediates
segmentation of mammalian hindbrain (Kiecker and Lumsden, 2005) and dorsal-ventral
boundary formation of the wing imaginal disc (Micchelli et al., 1997; Irvine and
Rauskolb, 2001). In the wing disc, an explicit model for boundary formation has been
proposed. Notch is first expressed broadly on both sides of the boundary as indicated in
the figure below. Subsequently, through the actions of the dorsally expressed selector
gene, Ap, the Notch ligand Serrate (i.e. Jagged1) and the Fringe glycosyl transferase are
induced dorsally. Fringe alters the glycosylation of Notch in dorsal cells, making it more
sensitive to Delta but insensitive to Serrate. The end result is activation of Notch in a
narrow band of cells at the D/V boundary.
(González et al., 2006)
Fig. 19. Notch signaling in Drosophila wing disc
56
We suggest that in the coronal suture, the Notch pathway functions in a manner
analogous to its activity in the D/V boundary of the wing disc, with some differences. At
E12.5, Notch2 is expressed broadly in the frontal, parietal and coronal suture territories.
By E14.5, expression, and activity as assessed by Hes1, are excluded from the boundary
region in the suture. Jagged1 expression begins broadly but becomes localized in a
narrow band of sutural cells, which do not express Notch2 or Hes1. The cells that express
Jagged1 may serve as border cells. Loss of Jagged1 function results in expansion of
Notch2 and Hes1 into the sutural domain consistent with Jagged1 autonomously
repressing Notch2 expression . The result is an absence of border cells (Jagged1
expressing cells) and loss of integrity of osteogenic-non-osteogenic boundary, which
coincides with the neural crest-mesoderm lineage boundary. It is interesting that in
Drosophila, active Notch signaling is required for border cell development. In the coronal
suture, it is the inverse. The Notch pathway is inactive in the sutural (border) cells, and
active in the flanking osteogenic regions.
Drosophila wing disc Mouse coronal suture
border
cells
Fig. 20. Notch signaling in mouse coronal suture
57
4. Notch signaling interacts with other major signaling pathways during suture
formation
As indicated previously in introduction, there are several signaling pathways involved in
suture formation. I examined whether Notch signaling interacts with RTK, Bmp and Wnt
signaling pathways. Molecular epistasis experiments did not identify cross regulatory
interactions between Jagged1 and ephrin-Eph signaling in the coronal suture. Similarly
no change in the phosphorylation status of the Erk1/2 was found in Jagged1 mutant
sutures. Thus Jagged1 is not likely to be upstream of the RTK pathways. Whether Erk1/2
signaling controls Jagged1 remains to be determined.
I also detected an increase in β-catenin-expressing cells in sutures of Dermo1-
Cre;Jagged1
cko/cko
mutants. Liu and colleagues showed that inactivation of the Wnt
pathway inhibitor, Axin2, results in craniosynostosis, and that a deficiency of β-catenin
rescues this phenotype (Liu et al., 2007). These findings, together with our demonstration
that β-catenin is upregulated in Jagged1 mutant sutures raises the possibility that a key
function of Jagged1 is to exclude β-catenin/Wnt activity from sutural cells and thus
maintain such cells in a non-osteogenic state.
I noted a change in the distribution of P-Smad1/5/8 expressing cells in Jagged1 mutants
as well. This change was similar to a change in Twist1-EphA4 mutants described
previously (Ting et al., 2009). In wildtype sutures, P-Smad1/5/8 positive cells were
concentrated in the osteogenic fronts; in mutants, they were distributed throughout the
suture. These cells may be osteogenic cells that are located ectopically in the sutural
58
domain instead of contributing to the frontal or parietal bones. Interestingly, an
interaction between Notch intracellular domain (NICD) and Smad1 was also observed in
myogenic differentiation (Dahlqvist et al., 2003) and Cos7 cell line (Takizawa et
al.,2003) , suggesting a possible regulation mechanism.
5. Epistasis model of Twist1 and Notch signaling in coronal suture
Twist1
+-/
;Jagged1
+/-
compound mutants exhibit more sever phenotypes, including
craniosynostosis, polydactyly, abnormalities of RTP and middle ear ossicles, than
Twist1
+/-
mice do. It suggests a significant genetic interaction between Twist1 and
Jagged1 genes. At coronal suture, combine with the notch signaling model mentioned
earlier, I propose the following model to explain the molecular network of control suture
formation. (Figure 22. Epistasis model of Twist1 and Notch signaling in coronal suture)
Fig. 21. Signaling pathways in coronal suture
59
In wildtype sutures, Twist1 positively regulates Jagged1 in sutural mesenchyme. Jagged1
represses Notch2. Jagged1 and Dll1 in the suture could potentially signal through Notch2
in the bone primordium. In the Twist1 mutant, Jagged1 is reduced in the sutural
mesenchyme (dotted lines signify reduced activity), resulting in an increase in Notch2
and Hes1 expression, and thus to a mis-specification of sutural mesenchyme cells to an
osteogenic fate.
Finally, previous results from our lab suggested an interaction between Twist1 and EphA4
(Ting et al., 2009). In combined with my results, Twist1 may be placed at the top of a
Fig. 22. Epistasis model of Twist1 and Notch signaling in coronal suture
60
regulatory hierarchy, controlling two independent pathways, ephrin-Eph and
Jagged1/Notch. Ephrin-Eph functions in the guidance of osteogenic cells along the
ectocranial mesenchyme to their destinations in the developing frontal and parietal bones.
A failure of this process results in mis-migration of osteogenic precursor cells into the
coronal suture. Jagged1/Notch functions in the initial specification of sutural cells and in
the boundary between the osteogenic and non-osteogenic compartments in the coronal
suture. Heterozygous loss of Twist1 function results in downregulation of both EphA4
and Jagged1 and consequent changes in osteogenic precursor migration and the
osteogenic-non-osteogenic boundary in the coronal suture. Genetic combinations of
EphA4
-/-
or Jagged1
+/-
with Twist1
+/-
exacerbate these defects, as expected. It is interesting
that the Twist1 expression pattern in the developing suture is broader than either Jagged1
or EphA4; thus additional regulatory influences must act on Jagged1 and EphA4 to
produce their highly focal expression patterns.
61
Chapter 1 References
Alagille, D., Estrada, A., Hadchouel, M., Gautier, M., Odièvre, M. and
Dommergues, J. P. (1987). Syndromic paucity of interlobular bile ducts (Alagille
syndrome or arteriohepatic dysplasia): review of 80 cases. J. Pediatr. 110, 195-200.
Aviv, R. I., Rodger, E., Hall, C. M. (2002). Craniosynostosis. Clin. Radiol. 57, 93-102.
Blair, S. S. (2003). Lineage compartments in Drosophila. Curr. Biol. 13, R548-551.
Bolós, V., Grego-Bessa, J. and de la Pompa, J. L. (2007). Notch signaling in
development and cancer. Endocr. Rev. 28, 339-363.
Bourgeois, P., Bolcato-Bellemin, A. L., Danse, J. M., Bloch-Zupan, A., Yoshiba, K.,
Stoetzel, C. and Perrin-Schmitt, F. (1998). The variable expressivity and incomplete
penetrance of the twist-null heterozygous mouse phenotype resemble those of human
Saethre-Chotzen syndrome. Hum. Mol. Genet. 7, 945-957.
Boyadjiev, S. A. (2007). International Craniosynostosis Consortium. Genetic analysis of
non-syndromic craniosynostosis. Orthod. Craniofac. Res. 10, 129-137.
Bray, S. (1998). Notch signalling in Drosophila: three ways to use a pathway. Semin.
Cell Dev. Biol. 9, 591-597.
Bray, S. J. (2006). Notch signalling: a simple pathway becomes complex. Nat. Rev. Mol.
Cell Biol. 7, 678-689.
Brooker, R., Hozumi, K. and Lewis, J. (2006). Notch ligands with contrasting
functions: Jagged1 and Delta1 in the mouse inner ear. Development 133, 1277-1286.
Buceta, J., Herranz, H., Canela-Xandri, O., Reigada, R., Sagués, F. and Milán, M.
(2007). Robustness and stability of the gene regulatory network involved in DV boundary
formation in the Drosophila wing. PLoS One 2, e602.
Carver, E. A., Oram, K. F. and Gridley, T. (2002). Craniosynostosis in Twist
heterozygous mice: a model for Saethre-Chotzen syndrome. Anat. Rec. 268, 90-92.
Castanon, I. and Baylies, M. K. (2002). A Twist in fate: evolutionary comparison of
Twist structure and function. Gene 287, 11-22.
Castanon, I., Von Stetina, S., Kass, J., Baylies, M.K. (2001). Dimerization partners
determine the activity of the Twist bHLH protein during Drosophila mesoderm
development. Development 128, 3145--3159.
62
Chai, Y. and Maxson, R. E. Jr. (2006). Recent advances in craniofacial morphogenesis.
Dev. Dyn. 235, 2353-2375.
Chen, Y. H., Ishii, M., Sun, J., Sucov, H. M., Maxson, R. E. Jr. (2007). Msx1 and
Msx2 regulate survival of secondary heart field precursors and post-migratory
proliferation of cardiac neural crest in the outflow tract. Dev. Biol. 308, 421-437.
Chen, Z. F. and Behringer, R. R. (1995). twist is required in head mesenchyme for
cranial neural tube morphogenesis. Genes Dev. 9, 686-699.
Cohen, M. M. Jr. (2006). The new bone biology: pathologic, molecular, and clinical
correlates. Am. J. Med. Genet. A. 140, 2646-2706.
Connerney, J., Andreeva, V., Leshem, Y., Mercado, M. A., Dowell, K., Yang, X.,
Lindner, V., Friesel, R. E. and Spicer, D. B. (2008). Twist1 homodimers enhance FGF
responsiveness of the cranial sutures and promote suture closure. Dev. Biol. 318, 323-
334.
Dahlqvist, C., Blokzijl, A., Chapman, G., Falk, A., Dannaeus, K., Ibâñez, C. F.,
Lendahl, U. (2003). Functional Notch signaling is required for BMP4-induced inhibition
of myogenic differentiation. Development 130, 6089-6099.
Dahmann, C. and Basler, K. (1999). Compartment boundaries: at the edge of
development. Trends Genet. 15, 320-326.
Danielian, P. S., Muccino, D., Rowitch, D. H., Michael, S. K. and McMahon, A. P.
(1998) Modification of gene activity in mouse embryos in utero by a tamoxifen-inducible
form of Cre recombinase. Curr. Biol. 8, 1323-1326.
de Celis, J. F., Tyler, D. M., de Celis, J., Bray, S. J. (1998). Notch signalling mediates
segmentation of the Drosophila leg. Development 125, 4617-4626.
el Ghouzzi, V., Le Merrer, M., Perrin-Schmitt, F., Lajeunie, E., Benit, P., Renier, D.,
Bourgeois, P., Bolcato-Bellemin, A. L., Munnich, A. and Bonaventure, J. (1997).
Mutations of the TWIST gene in the Saethre-Chotzen syndrome. Nat. Genet. 15, 42-46.
Emerick, K. M., Rand, E.B., Goldmuntz, E., Krantz, I. D., Spinner, N. B. and
Piccoli, D. A. (1999). Features of Alagille syndrome in 92 patients: frequency and
relation to prognosis. Hepatology 29, 822-829.
Golson, M. L., Loomes, K. M., Oakey, R. and Kaestner, K. H. (2009). Ductal
malformation and pancreatitis in mice caused by conditional Jag1 deletion.
Gastroenterology 136, 1761-1771.e1.
63
González, A., Chaouiya, C. and Thieffry, D. (2006). Dynamical analysis of the
regulatory network defining the dorsal-ventral boundary of the Drosophila wing imaginal
disc. Genetics 174, 1625-1634.
Hajihosseini, M. K. (2008). Fibroblast growth factor signaling in cranial suture
development and pathogenesis. Front. Oral Biol. 12, 160-177.
Holmes, G., Rothschild, G., Roy, U. B., Deng, C. X., Mansukhani, A. and Basilico, C.
(2009). Early onset of craniosynostosis in an Apert mouse model reveals critical features
of this pathology. Dev. Biol. 328, 273-84.
Howard, T. D., Paznekas, W. A., Green, E. D., Chiang, L. C., Ma, N., Ortiz de Luna,
R. I., Garcia Delgado, C., Gonzalez-Ramos, M., Kline, A. D. and Jabs, E. W. (1997).
Mutations in TWIST, a basic helix-loop-helix transcription factor, in Saethre-Chotzen
syndrome. Nat. Genet. 15, 36-41.
Irvine, K. D. and Rauskolb, C. (2001). Boundaries in development: formation and
function. Annu. Rev. Cell Dev. Biol. 17, 189-214.
Ishii, M., Merrill, A. E., Chan, Y. S., Gitelman, I., Rice, D. P., Sucov, H. M., Maxson,
R. E. Jr. (2003). Msx2 and Twist cooperatively control the development of the neural
crest-derived skeletogenic mesenchyme of the murine skull vault. Development 130,
6131-6142.
Jabs, E. W., Müller, U., Li, X., Ma, L., Luo, W., Haworth, I. S., Klisak, I., Sparkes,
R., Warman, M. L., Mulliken, J. B., et al. (1993). A mutation in the homeodomain of
the human MSX2 gene in a family affected with autosomal dominant craniosynostosis.
Cell 75, 443-450.
Jarriault, S., Brou, C., Logeat, F., Schroeter, E. H., Kopan, R. and Israel, A. (1995).
Signalling downstream of activated mammalian Notch. Nature 377, 355-358.
Jenkins, D., Seelow, D., Jehee, F. S., Perlyn, C. A., Alonso, L. G., Bueno, D. F.,
Donnai, D., Josifova, D., Mathijssen, I. M., Morton, J. E., Orstavik, K. H,. Sweeney,
E., Wall, S. A., Marsh, J. L., Nurnberg, P., Passos-Bueno, M. R. and Wilkie, A. O.
(2007). RAB23 mutations in Carpenter syndrome imply an unexpected role for hedgehog
signaling in cranial-suture development and obesity. Am. J. Hum. Genet. 80, 1162-1170.
Jiang, X., Iseki, S., Maxson, R. E., Sucov, H. M. and Morriss-Kay, G. M. (2002).
Tissue origins and interactions in the mammalian skull vault. Dev. Biol. 241, 106-116.
Johnson, D., Iseki, S., Wilkie, A. O. and Morriss-Kay, G. M. (2000). Expression
patterns of Twist and Fgfr1, -2 and -3 in the developing mouse coronal suture suggest a
key role for twist in suture initiation and biogenesis. Mech. Dev, 91, 341-345.
64
Kabbani, H and Raghuveer, T. S. (2004). Craniosynostosis. Am. Fam. Physician. 69,
2863-2870.
Kamath, B. M., Stolle, C., Bason, L., Colliton, R. P., Piccoli, D. A., Spinner, N. B.
and Krantz, I. D. (2002). Craniosynostosis in Alagille syndrome. Am. J. Med. Genet.
112, 176-180.
Kamath, B. M., Spinner, N. B., Emerick, K. M., Chudley, A. E., Booth, C., Piccoli,
D. A. and Krantz, I. D. (2004). Vascular anomalies in Alagille syndrome: a significant
cause of morbidity and mortality. Circulation 109, 1354-1358.
Kiecker, C. and Lumsden, A. (2005). Compartments and their boundaries in vertebrate
brain development. Nat. Rev. Neurosci. 6, 553-564.
Kiernan, A. E., Xu, J. and Gridley, T. (2006). The Notch ligand JAG1 is required for
sensory progenitor development in the mammalian inner ear. PLoS Genet. 2, e4.
Kopan, R. and Ilagan, M. X. (2009). The canonical Notch signaling pathway: unfolding
the activation mechanism. Cell 137, 216-233.
Krantz, I. D., Colliton, R. P., Genin, A., Rand, E. B., Li, L., Piccoli, D. A. and
Spinner, N. B. (1998). Spectrum and frequency of jagged1 (JAG1) mutations in Alagille
syndrome patients and their families. Am. J. Hum. Genet. 62, 1361-1369.
Lai, E. C. (2004). Notch signaling: control of cell communication and cell fate.
Development 131, 965-973.
Le, T. T., Conley, K. W. and Brown, N. L. (2009). Jagged 1 is necessary for normal
mouse lens formation. Dev. Biol. 328, 118-126.
Li, C., Scott, D. A., Hatch, E., Tian, X. and Mansour, S. L. (2007). Dusp6 (Mkp3) is a
negative feedback regulator of FGF-stimulated ERK signaling during mouse
development. Development 134, 167-176.
Liu, B., Yu, H. M. and Hsu, W. (2007). Craniosynostosis caused by Axin2 deficiency is
mediated through distinct functions of beta-catenin in proliferation and differentiation.
Dev. Biol. 301, 298-308.
Loeys, B. L., Chen, J., Neptune, E. R., Judge, D. P., Podowski, M., Holm, T., Meyers,
J., Leitch, C. C., Katsanis, N., Sharifi, N., et al. (2005). A syndrome of altered
cardiovascular, craniofacial, neurocognitive and skeletal development caused by
mutations in TGFBR1 or TGFBR2. Nat. Genet. 37, 275-281.
65
Lovato, T. A. L., Benjamin, A. R., Cripps, R. M. (2005). Transcription of Myocyte
enhancer factor-2 in adult Drosophila myoblasts is induced by the steroid hormone
ecdysone. Dev. Biol. 288, 612-621.
Lozier, J., McCright, B. and Gridley, T. (2008). Notch signaling regulates bile duct
morphogenesis in mice. PLoS One 3, e1851.
Ma, A., Boulton, M., Zhao, B., Connon, C., Cai, J., Albon, J. (2007). A role for notch
signaling in human corneal epithelial cell differentiation and proliferation. Invest.
Ophthalmol. Vis. Sci. 48, 3576-3585.
Major, R. J. and Irvine, K. D. (2005). Influence of Notch on dorsoventral
compartmentalization and actin organization in the Drosophila wing. Development 132,
3823-3833.
Mathijssen, I. M., van Splunder, J., Vermeij-Keers, C., Pieterman, H., de Jong, T.
H., Mooney, M. P. and Vaandrager, J. M. (1999). Tracing craniosynostosis to its
developmental stage through bone center displacement. J. Craniofac. Genet. Dev. Biol.
19, 57–63.
McCright, B., Lozier, J. and Gridley, T. (2002). A mouse model of Alagille syndrome:
Notch2 as a genetic modifier of Jag1 haploinsufficiency. Development 129, 1075-1082.
McCright, B., Gao, X., Shen, L., Lozier, J., Lan, Y., Maguire, M., Herzlinger, D.,
Weinmaster, G., Jiang, R. and Gridley, T. (2001). Defects in development of the
kidney, heart and eye vasculature in mice homozygous for a hypomorphic Notch2
mutation. Development 128, 491-502.
McDaniell, R., Warthen, D. M., Sanchez-Lara, P. A., Pai, A., Krantz, I. D., Piccoli,
D. A. and Spinner, N. B. (2006). NOTCH2 mutations cause Alagille syndrome, a
heterogeneous disorder of the notch signaling pathway. Am. J. Hum. Genet. 79, 169-173.
McLarren, K. W., Lo, R., Grbavec, D., Thirunavukkarasu, K., Karsenty, G. and
Stifani, S. (2000). The mammalian basic helix loop helix protein HES-1 binds to and
modulates the transactivating function of the runt-related factor Cbfa1. J. Biol. Chem.
275, 530-538.
McLeod, M. J. (1980). Differential staining of cartilage and bone in whole mouse fetuses
by alcian blue and alizarin red S. Teratology 22, 299-301.
Merrill, A. E., Bochukova, E. G., Brugger, S. M., Ishii, M., Pilz, D. T., Wall, S. A.,
Lyons, K. M., Wilkie, A. O. and Maxson R. E. Jr. (2006). Cell mixing at a neural
crest-mesoderm boundary and deficient ephrin-Eph signaling in the pathogenesis of
craniosynostosis. Hum. Mol. Genet. 15, 1319-1328.
66
Micchelli, C. A., Rulifson, E. J. and Blair, S. S. (1997). The function and regulation of
cut expression on the wing margin of Drosophila: Notch, Wingless and a dominant
negative role for Delta and Serrate. Development 124, 1485-1495.
Moenning, A., Jäger, R., Egert, A., Kress, W., Wardelmann, E. and Schorle, H.
(2009). Sustained platelet-derived growth factor receptor alpha signaling in osteoblasts
results in craniosynostosis by overactivating the phospholipase C-gamma pathway. Mol.
Cell. Biol. 29, 881-891.
Morriss-Kay, G. M. and Wilkie, A. O. (2005). Growth of the normal skull vault and its
alteration in craniosynostosis: insights from human genetics and experimental studies. J.
Anat. 207, 637-653.
Nobta, M., Tsukazaki, T., Shibata, Y., Xin, C., Moriishi, T., Sakano, S., Shindo, H.
and Yamaguchi, A. (2005). Critical regulation of bone morphogenetic protein-induced
osteoblastic differentiation by Delta1/Jagged1-activated Notch1 signaling. J. Biol. Chem.
280, 15842-15848.
Oda, T., Elkahloun, A. G., Pike, B. L, Okajima, K., Krantz, I. D., Genin, A., Piccoli,
D. A., Meltzer, P. S., Spinner, N. B., Collins, F. S. and Chandrasekharappa, S. C.
(1997). Mutations in the human Jagged1 gene are responsible for Alagille syndrome.
Nat. Genet. 16, 235-242.
Opperman, L. A. (2000). Cranial sutures as intramembranous bone growth sites. Dev.
Dyn. 219, 472-485.
Oram, K. F. and Gridley, T. (2005). Mutations in snail family genes enhance
craniosynostosis of Twist1 haplo-insufficient mice: implications for Saethre-Chotzen
Syndrome. Genetics 170, 971-974.
Rice, D. P. (2008). Developmental anatomy of craniofacial sutures. Front. Oral Biol. 12,
1-21.
Rice, D. P., Aberg, T., Chan, Y., Tang, Z., Kettunen, P. J., Pakarinen, L., Maxson,
R. E. and Thesleff, I. (2000). Integration of FGF and TWIST in calvarial bone and
suture development. Development 127, 1845-1855.
Saga, Y., Miyagawa-Tomita, S., Takagi, A., Kitajima, S., Miyazaki, J. and Inoue, T.
(1999). MesP1 is expressed in the heart precursor cells and required for the formation of
a single heart tube. Development 126, 3437-3447.
Saravanamuthu, S. S., Gao, C. Y. and Zelenka, P. S. (2009). Notch signaling is
required for lateral induction of Jagged1 during FGF-induced lens fiber differentiation.
Dev. Biol. 332, 166-176.
67
Schröder, N. and Gossler, A. (2002). Expression of Notch pathway components in fetal
and adult mouse small intestine. Gene Expr. Patterns. 2, 247-250.
Settle, S. H. Jr., Rountree, R. B., Sinha, A., Thacker, A., Higgins, K. and Kingsley,
D. M. (2003). Multiple joint and skeletal patterning defects caused by single and double
mutations in the mouse Gdf6 and Gdf5 genes. Dev. Biol. 254, 116-130.
Shimizu, T., Tanaka, T., Iso, T., Doi, H., Sato, H., Kawai-Kowase, K., Arai, M.and
Kurabayashi, M. (2009). Notch signaling induces osteogenic differentiation and
mineralization of vascular smooth muscle cells: role of Msx2 gene induction via Notch-
RBP-Jk signaling. Arterioscler. Thromb. Vasc. Biol. 29, 1104-1111.
Slater, B. J., Lenton, K. A., Kwan, M. D., Gupta, D. M., Wan, D. C. and Longaker,
M. T. (2008). Cranial sutures: a brief review. Plast. Reconstr. Surg. 121, 170e-178e.
Sood, S., Eldadah, Z. A., Krause, W. L., McIntosh, I. and Dietz, H. C. (1996).
Mutation in fibrillin-1 and the Marfanoid-craniosynostosis (Shprintzen-Goldberg)
syndrome. Nat. Genet. 12, 209-211.
Soriano, P. (1999). Generalized lacZ expression with the ROSA26 Cre reporter strain.
Nat. Genet. 21, 70-71.
Stump, G., Durrer, A., Klein, A. L., Lütolf, S., Suter, U. and Taylor, V. (2002).
Notch1 and its ligands Delta-like and Jagged are expressed and active in distinct cell
populations in the postnatal mouse brain. Mech. Dev. 114, 153-159.
Suh, J. H., Lee, H. W., Lee, J. W. and Kim, J. B. (2008). Hes1 stimulates
transcriptional activity of Runx2 by increasing protein stabilization during osteoblast
differentiation. Biochem. Biophys. Res. Commun. 367, 97-102.
Takizawa, T., Ochiai, W., Nakashima, K. and Taga, T. (2003). Enhanced gene
activation by Notch and BMP signaling cross-talk. Nucleic. Acids Res. 31, 5723-5731.
Thisse, B., Stoetzel, C., Gorosotiza-Thisse, C. and Perrin-Schmidt, F. (1988).
Sequence of the twist gene and nuclear localization of its protein in endomesodermal
cells of early Drosophila embryos. EMBO J. 7, 2175–2183.
Tepass, U., Godt, D. and Winklbauer, R. (2002). Cell sorting in animal development:
signalling and adhesive mechanisms in the formation of tissue boundaries. Curr. Opin.
Genet. Dev. 12, 572-582.
Tezuka, K., Yasuda, M., Watanabe, N., Morimura, N., Kuroda, K., Miyatani, S. and
Hozumi, N. (2002). Stimulation of osteoblastic cell differentiation by Notch. J. Bone.
Miner. Res. 17, 231-239.
68
Ting, M. C., Wu, N. L., Roybal, P. G., Sun, J., Liu, L., Yen, Y. and Maxson R. E. Jr.
(2009). EphA4 as an effector of Twist1 in the guidance of osteogenic precursor cells
during calvarial bone growth and in craniosynostosis. Development 136, 855-864.
Twigg, S. R., Kan, R., Babbs, C., Bochukova, E. G., Robertson, S. P., Wall, S. A.,
Morriss-Kay, G. M. and Wilkie, A. O. (2004). Mutations of ephrin-B1 (EFNB1), a
marker of tissue boundary formation, cause craniofrontonasal syndrome. Proc. Natl.
Acad. Sci. U.S.A. 101, 8652-8657.
Vargesson, N., Patel, K., Lewis, J. and Tickle, C. (1998). Expression patterns of
Notch1, Serrate1, Serrate2 and Delta1 in tissues of the developing chick limb. Mech.
Dev. 77, 197-199.
Vernon, A. E. and LaBonne, C. (2004). Tumor metastasis: a new twist on epithelial-
mesenchymal transitions. Curr. Biol. 14, R719-721.
Warman, M. L., Mulliken, J. B., Hayward, P. G. and Müller, U. (1993). Newly
recognized autosomal dominant disorder with craniosynostosis. Am. J. Med. Genet. 46,
444-449.
Wilkie, A. O. (1997). Craniosynostosis: genes and mechanisms. Hum. Mol. Genet. 6,
1647-1656.
Xue, Y., Gao, X., Lindsell, C. E., Norton, C. R., Chang, B., Hicks, C., Gendron-
Maguire, M., Rand, E. B., Weinmaster, G. and Gridley, T. (1999). Embryonic
lethality and vascular defects in mice lacking the Notch ligand Jagged1. Hum. Mol.
Genet. 8, 723-730.
Yoshida, T., Vivatbutsiri, P., Morriss-Kay, G., Saga, Y. and Iseki, S. (2008). Cell
lineage in mammalian craniofacial mesenchyme. Mech. Dev. 125, 797-808.
Yoshida, T., Phylactou, L. A., Uney, J. B., Ishikawa, I., Eto, K. and Iseki S. (2005).
Twist is required for establishment of the mouse coronal suture. J. Anat. 206, 437-444.
Yousfi, M., Lasmoles, F. and Marie, P. J. (2002). TWIST inactivation reduces
CBFA1/RUNX2 expression and DNA binding to the osteocalcin promoter in osteoblasts.
Biochem. Biophys. Res. Commun. 297, 641-644.
Yu, H. M., Jerchow, B., Sheu, T. J., Liu, B., Costantini, F., Puzas, J. E., Birchmeier,
W. and Hsu, W. (2005). The role of Axin2 in calvarial morphogenesis and
craniosynostosis. Development 132, 1995-2005.
69
Yu, K., Xu, J., Liu, Z., Sosic, D., Shao, J., Olson, E. N., Towler, D. A. and Ornitz, D.
M. (2003). Conditional inactivation of FGF receptor 2 reveals an essential role for FGF
signaling in the regulation of osteoblast function and bone growth. Development 130,
3063-3074.
Yuan, Z. R., Kobayashi, N. and Kohsaka, T. (2006). Human Jagged 1 mutants cause
liver defect in Alagille syndrome by overexpression of hepatocyte growth factor. J. Mol.
Biol. 356, 559-568.
Zhang, X., Kuroda, S., Carpenter, D., Nishimura, I., Soo, C., Moats, R., Iida, K.,
Wisner, E., Hu, F. Y., Miao, S., et al. (2002). Craniosynostosis in transgenic mice
overexpressing Nell-1. J. Clin. Invest. 110, 861-870.
70
Chapter 2
Inactivation of Brca1 in ovarian granulosa cells is associated with increases in
circulating estrogen levels and bone mineral density
Introduction
According to the American Cancer Society, Ovarian cancer represents the second leading
gynecologic cancer, following cancer of the uterine corpus, and causes more deaths per
year than any other female reproductive cancers. It is estimated that there will be 21,550
new cases and 14,600 deaths from ovarian cancers in the US in 2009. Unfortunately, the
death rate has not been reduced in the last 50 years.
There are three main types of ovarian tumors based on the cell of origin: epithelial
tumors, germ cell tumors, and stromal tumors. The majority (85% to 95%) of ovarian
tumors are epithelial and it is also the most lethal among ovarian neoplasms. To date,
researchers have relatively little knowledge about the causes of ovarian cancers.
Suggested risk factors for ovarian cancer include: age, obesity, family history of breast
and ovarian cancer, prolonged use of unopposed estrogens and inherited mutations in
BRCA1 or BRCA2 genes. BRCA1 and BRCA2 genes are the most well known inherited
genetic factors that increase the possibility of developing epithelial ovarian cancer.
In the early 90’s, the breast cancer susceptibility gene BRCA1 was cloned using linkage
analysis of family with multiple cases of breast and ovarian cancers. (Hall, 1990) The
lifetime ovarian cancer risk for women with BRCA1 mutations was estimated to be 57%.
71
(King et al, 2003). Since then, Knudson’s two hit hypothesis of tumorigenesis has
become a popular model to explain the mechanism of breast and ovarian cancer, i.e. loss
of heterozygosity (LOH) of the BRCA1 gene in cancer cells. On a basis of this
hypothesis, scientists found the roles of Brca1 in cellular DNA repair, genomic stability,
and checkpoint control (Deng, 2006).
However, there are a number of facts that implied BRCA1 is not compatible with the
traditional role of a tumor suppressor gene and other mechanisms may be applied. First,
loss of Brca1 does not provide an advantage in the survival of cancer cell lines, which is
unusual for classic tumor suppressor genes. The number of available breast cancer cell
lines is small and only very few of them have identified BRCA1 mutations (Elstrodt et al.,
2006). Second, BRCA1 and p53, a classic tumor suppressor gene, have similar functions
in DNA repair and cell cycle regulation. Unlike mutations of p53 that predisposes to both
hereditary and sporadic cancers, somatic mutations in the BRCA1 are rarely associated
with sporadic breast or ovarian cancer (Futreal, 1994). Third, germline mutation in
BRCA1 predisposes carriers mostly to breast and ovarian cancers. LOH model cannot
easily explain the association of ubiquitously expressed BRCA1 with the development of
tissue-specific breast and ovarian tumors. Fourth, prophylactic oophorectomy in women
with germline BRCA1 mutations prevents occurrence of breast cancer and peritoneal
cancer, suggesting the involvement of secreted molecules in BRCA1-related tumor
development (Kauff et al., 2002; Rebbeck et al., 2002).
72
Our previous study suggested a cell-nonautonomous mechanism of Brca1 in tumor
development (Chodankar et al., 2005). We conditional knocked out Brca1 in ovarian
granulosa cells by using Fshr-Cre and resulted in the development of cystic tumors in the
ovaries and uterine horns. Those tumors only carried normal Brca1 suggesting that
mutation of Brca1 caused the tumor indirectly.
The cell-nonautonomous phenomenon implied that secreted molecules are responsible for
tumor formation. Recently, more and more studies focus on the link between Brca1 and
estrogen signaling pathways. Estrogens are stimulated by pituitary-secreted
gonadotrophins, FSH and leutenizing hormone (LH). Estradiol is the most potent
estrogen and is mainly responsible for estrogen action. Other estrogens, i.e. estrone and
estriol, are far less active. The biological functions of estrogens are mediated by binding
to estrogen receptors (ERs), ERα or ERβ. ERs belong to the nuclear receptor
superfamily, a family of ligand-regulated transcription factors. Brca1 negatively regulates
estrogen receptor signaling (Fan et al, 1999) and estrogen biosythesis (Hu et al., 2005),
suggesting inactivation of Brca1 potentially increases estrogenic effects. The regulatory
relationship between Brca1 and estrogen signaling is consistent with the fact that Brca1
and estrogen have both been associated with an increased risk of ovarian cancer (Lacey et
al, 2006).
In this study, I demonstrated that conditional inactivation of Brca1 in ovarian granulosa
cells resulted in an increase in serum estrogen levels of the mice strongly predisposed to
ovarian and uterine epithelial tumors. The elevated estrogen levels are associated with
73
several physical consequences including increased cell proliferation in uterus and blood
vessels dilation. Most importantly, mutant mice were suggested to be exposed to
persistent higher estrogen levels compared with wildtype. It is supported by the evidence
of increased bone density of the mutant mice.
74
Materials and Methods
Mouse mutants and genotyping
The mice used in this study were as described previously: Brca1 flox (Xu et al.,1999);
Fshr-Cre (Chodankar et al., 2005); R26R (Soriano, 1999).
The genotyping conditions were as follows:
Cre
Primers:
Cre-forward: ctc tgg tgt agc tga tga tc (20)
Cre-reverse: taa tcg cca tct tcc agc ag (20)
PCR program: 94°C. 5 min.; 94°C. 30 sec.; 55°C. 1 min.; 72°C. 1 min.; Goto #2 34
times; 72°C. 5 min.; 4°C forever.
Expected product size: ~330 bp
Brca1 flox
Brca1 floxed exon11 primer A: ctg ggt agt ttg taa gca tcc (21)
Brca1 floxed exon11 primer B: caa taa act gct ggt ctc agg c (22)
PCR program: 94°C. 5 min.; 94°C. 30 sec.; 58°C. 1 min.; 72°C. 1 min.; Goto #2 34
times; 72°C. 5 min.; 4°C forever.
Expected product size: Wildtype band~470bp
Floxed band~530bp
R26R
Primers:
R26Rcom: aaa gtc gct ctg agt tgt tat (21)
R26Rwt: gga gcg gga gaa atg gat atg (21)
R26RKO: gcg aag agt ttg tcc tca acc (21)
75
PCR program: 94°C. 5 min.; 94°C. 30 sec.; 53°C. 45 sec.; 72°C. 1 min.; Goto #2 34
times; 72°C. 10 min.; 4°C forever.
Expected product size: R26Rcom+ R26Rwt for wildtype band ~650 bp;
R26Rcom+ R26RKO for knockout band ~350 bp
PMSG and hCG treatments
Virgin females were injected intraperitoneally with 5 IU of Pregnant Mare Serum
Gonadotropin (PMSG) (Sigma, G4877) for 48hrs, followed by 5 IU of human chorionic
gonadotropin (hCG) (Sigma, C8554) for 24 hrs, if applicable.
Measurement of serum estradiol
Blood samples were collected by cardiac puncture under avertin anesthesia. Serum was
collected following overnight incubation at 4°C and stored at -20°C. Estradiol
concentrations were measured in Dr. Frank Stanczyk’s laboratory by radioimmunoassay
with a preceding extraction step using ethyl acetate:hexane (3:2). The assay sensitivity is
3 pg/ml and the interassay coefficient of variation is 8-10% (Stanczyk et al., 2003).
Transplantation of ovaries under kidney capsule
Ovaries were interchanged between 5-week-old wildtype and mutant littermates. Mice
were anesthetized with an intraperitoneal administration of avertin. Two intact ovaries
from donor mouse were removed and saved in warm PBS temporarily. The kidney from
recipient mouse was exteriorized through a dorsal incision and kept moist with warm
PBS. A small hole was cut in the kidney capsule and the capsule was lifted by a capillary
76
with sealed end. Two ovaries from the donor mouse were inserted under the capsule. The
incisions were closed and the mice were allowed to recover.
Whole-mount X-gal staining
Fshr-Cre-activated β-galactosidase activity in tissues of interest was detected by X-gal
staining as described previously (Ishii et al., 2003). Briefly, samples were fixed in 4%
PFA for 15 min and stained with X-gal staining solution* at 37°C O/N. (* X-gal staining
solution: 1M MgCl
2
20 µl, 5% DOC 20 µl, 2% NP-40 100 µl, 2% X-gal (in DMSO) 500
µl, potassium ferricyanide (FeIII) 0.0825g, potassium ferrocyanide (FeII) 0.105g in 1x
PBS)
BrdU labeling and immunostaining
Mice were given a single intraperitoneal injection of 5-bromo-2'-deoxyuridine (BrdU)
/PBS (200µg/g of body weight) two hours before sacrifice. Uteri were collected and fixed
with 4% paraformaldehyde and embedded in HistoPrep (Fisher Scientific). Transverse
frozen sections were cut in a cryostat at 10-µm thickness. BrdU signals were detected by
BrdU Staining Kit (Invitrogen, #93-3943) with hematoxylin counterstain according to the
manufacturer's instructions.
Immunofluorescence staining was performed by using rabbit anti-von Willebrand Factor
antibody (Sigma, F3520, 1:100) and Rhodamin Red-X goat anti-rabbit IgG secondary
antibody (Invitrogen, R6394, 1:100). The cell nuclei were revealed by co-staining with
DAPI.
77
Results
1. Absence of a functional Brca1 in granulosa cells increases serum estrogen levels in
mice
Inactivation of the Brca1 gene specifically in mice ovarian granulosa cells led to
development of cystic tumors in ovaries and uterine horns (Chodankar et al., 2005). It
also caused ductal dilatation in mammary glands. These findings suggested that Brca1
may influence tumor development through a cell-nonautonomous mechanism. Given the
fact that 1) estrogen is a well-established risk factor for ovarian and breast cancer (Mørch
et al., 2009) ; 2) granulosa cells play a major role in estrogen production (Stocco, 2008)
and 3) Brca1 negatively regulates aromatase which converts androgens to estrogens (Lu
et al., 2006; Ghosh et al., 2007), I hypothesized that the amount of estrogen released by
granulosa cells is affected in Fshr-Cre;Brca1
flox/flox
mutant mice.
Our previous attempts to detect the difference of serum estrogen levels between wildtype
and Fshr-Cre;Brca1
flox/flox
mutant mice were not successful. It was probably due to the
large variations of estradiol (E2) concentrations between individuals and also a restricted
change of hormone levels under natural estrous cycle conditions. I therefore synchronized
mouse estrus cycle and stimulated follicles growth with Pregnant Mare Serum
Gonadotropin (PMSG) for 48 hours. The mice blood serum was collected and assayed by
radioimmunoassay (RIA) for estradiol concentrations. The results showed that the serum
estradiol level was significantly increased in Fshr-Cre;Brca1
flox/flox
mutant mice after
PMSG stimulation (Fig.23) .
78
Fig. 23. Conditional knockout of Brca1 in granulosa cells resulted in an
increase of serum estradiol. (A) The times of PMSG administration and serum
collection are indicated by arrows. (B) The level of serum estradiol was
significantly increased in 3-month-old mutant mice 48hrs after PMSG stimulation.
(P<0.001) (n=14 for wildtype; n=17 for mutant)
Sacrifice
the mice
5 IU PMSG
24 hr. 24 hr.
follicles growth
(Follicular phase)
Serum
collection
RIA (Dr. Frank Stanczyk lab)
A
B
wildtype mutants
79
2. Fshr-Cre is expressed in pituitary gland and brain
Previous studies suggested that granulosa cells is the only cell type which is thought to
express the follicle-stimulating hormone receptor (Fshr) in females (Griswold et al.,
1995; Gromoll and Simoni, 2005). The expression of Fshr-driven Cre in granulosa cells
was confirmed by Chodankar et al (Chodankar et al., 2005). However, by using
Cre/R26R reporter system, I found that Fshr-driven Cre was also expressed in the
pituitary gland and hypothalamus of the brain, which are both critical for hormone
modulating. As shown in Fig. 24A and B, X-gal staining assay revealed the Cre activity
in anterior and posterior lobe of the pituitary gland and hypothalamus. In addition, blue
cells can be detected in olfactory tubercle, cerebellum and medulla of the brain (Fig.
24B). Pituitary gland and hypothalamus are the major functional units of hypothalamic-
pituitary-axis, which regulates many physiological processes including hormone
releasing. In this case, we could not rule out the possibility that these organs are
responsible for the increase of serum estradiol in mutant mice. Therefore, we performed
the following experiments to demonstrate that ovaries are the cause of the estrogen level
change.
80
Fig. 24. Fshr-Cre is expressed in pituitary gland and brain. (A) Whole mount X-
gal staining shows that Fshr-Cre was expressed in the anterior and posterior lobe of
the pituitary gland. (B) The activities of Fshr-Cre can also be detected in olfactory
tubercle, hypothalamus, cerebellum and medulla of the brain (arrows).
Anterior lobe
Posterior lobe
R26R Fshr-Cre;R26R
cerebellum
hypothalamus
olfactory tubercle
medulla
A
B
81
3. Ovaries carrying a Brca1 mutation are responsible for the increase of serum
estradiol
To examine whether ovary, but not other organs, affects the hormone levels in mutant
mice, we interchanged the ovaries between 5-week-old wildtype and mutant littermates
and transplanted the ovaries under kidney capsule. Female mice become sexually mature
after 6 week of age. Transplant performed at earlier stage prevents the hormone effects in
donor mice. At 3-month-old, the mice were treated with PMSG for 48 hours. Only the
serum from the mice with two functional ovaries under kidney capsule (Fig. 25B,C) was
measured (4 out of 11 at each group). The results were similar to that from the mice
without ovarian transplant (Fig. 23B), indicating mutation of Brca1 in ovarian granulosa
cells results in an increase of estradiol in the mouse serum.
4. Phenotypes suggested that mutant mice were exposed to high levels of estradiol
after estrus induction and synchronization
We observed phenotypes suggested that mutant mice were exposed to higher estradiol
levels after estrus induction and synchronization. It is known that uterine stromal
proliferation positively correlated with circulating estradiol levels (Wood et al., 2007). I
performed BrdU incorporation assay in mice treated with PMSG and found that the
stromal cells proliferation rate was significantly increased in mutant mice (Fig. 26A).
This increased proliferation of stromal cells in the mutants was eliminated after
overiectomy (Fig. 26A). There was no difference in proliferation of luminal and
glandular cells of uterus between wildtype and mutants (Fig. 26B,C), which was
82
Fig. 25. Ovarian granulosa cells are responsible for increased serum estradiol
in mutant mice. (A) Diagram indicates the ovary transplant under kidney capsule
between 5-week-old wildtype and mutant littermates. When mice reached 3 month
of age, they were stimulated with 5IU PMSG for 48 hrs. and then both of the serum
and the kidneys containing transplanted ovaries were collected. (B) The ovaries
look functional under the kidney capsule. Note the blood vessels on the ovary
(arrow). (C) H&E staining shows the functional ovaries after PMSG treatment.
Mature follicles (arrows) and corpora lutea (CL) from previous cycles were
observed. (D) The serum estradiol level was increased in wildtype mice with
mutant ovaries, indicating the ovaries carrying mutant Brca1 in granulosa cells
were responsible for the increase of serum estradiol. (P<0.05) (n=4 for each group)
W
T
MT
A B
WT mice
with MT
ovaries
MT mice
with WT
ovaries
CL
CL
CL
CL
CL
CL
MT mice w/
WT ovaries
WT mice w/
MT ovaries
C
D
MT mice w/
WT ovaries
MT mice w/ WT mice w/
WT ovaries MT ovaries
83
consistent with the estrogen effects on uterine cells in earlier study (Wood et al., 2007). I
also examined cell proliferation in uterus after PMSG and hCG stimulation, which
induced ovulation and luteal formation. The rate of uterine cell proliferation was reduced
in this phase and showed no significant difference between wildtype and mutants (Fig.
27).
Fig. 26. Analysis of cell proliferation in uterus after PMSG treatment. Mice
were treated with PMSG as in Fig. 1A. Two hours before sacrifice, mice were
injected with BrdU to label S-phase cells. The procedure was also performed on
ovariectomized (OVX) mice. (A) Cell proliferation was increased in uterine stromal
cells of mutant mice (*P<0.01), but not in luminal epithelium (B) or glandular
epithelium (C). After ovariectomy, there was no significant difference between
wildtype and mutant. (n=6 for WT and MT; n=4 for OVX WT; n=7 for OVX MT)
*
WT MT OVX WT OVX MT
WT MT OVX WT OVX MT WT MT OVX WT OVX MT
Stromal cells
Luminal epithelium Glandular epithelium
BrdU positive signals (%)
BrdU positive signals (%)
BrdU positive signals (%)
A
B C
84
Fig. 27. Analysis of cell proliferation in uterus after PMSG and hCG
treatments. (A) Mice were treated with PMSG and hCG as indicated. Two hours
before sacrifice, mice were injected with BrdU to label S-phase cells. (B) The BrdU
positive signals in uterine stromal, luminal and glandular epithelium were counted.
There was no significant difference between wildtype and mutant. (n=6 for each
group)
5 IU PMSG 5 IU hCG
Sacrifice
the mice
24 hr. 24 hr. 24 hr.
follicles growth Ovulation
(Follicular phase) (Luteal phase)
A
BrdU
WT MT
WT MT WT MT WT MT WT MT
Stromal cells
Luminal epithelium Glandular epithelium
BrdU positive signals (%)
BrdU positive signals (%)
BrdU positive signals (%)
WT MT
B
85
Estrogen has been shown to induces vascular wall dilation possibly through nitric oxide
signaling pathway (Guo et al., 2005; Leung et al., 2007). After treating with PMSG and
hCG to induce ovulation, I found dilated blood vessels in luteal phase ovaries of the
mutant mice (Fig. 28). There was no difference in the number of blood vessels in the
corpora lutea between wildtype and mutant by detecting vascular endothelial markers,
von Willebrand factor (vWF) (Fig. 29)
5. Long-term exposure to elevated estrogen increases bone density in mutant mice
It is difficult to demonstrate the change in serum estrogen level without estrus induction
and synchronization. Estrogen is known to increase bone density and is widely used to
treat postmenopausal osteoporosis (Lindsay and Tohme, 1990; Felson et al., 1993).
Therefore, to examine whether this increased estrogen levels influence physiological
function under natural conditions, we analyzed the bone density of 6-month-old wildtype
and mutant mice.
Micro-computed tomographic (μCT) analysis of 6-month-old mutant female mice
showed a higher bone mass compared with wildtype (Fig. 30A). The trabecular bone
volume density (BV/TV) measured in the distal femur (Fig. 30B) of mutants was 2-fold
higher than that of wildtype. Detailed trabecular bone parameters were also studied. The
trabecular number (Tb.N), trabecular thickness (Tb.Th) and connectivity density
(Conn.D) were all increased in mutants. These results suggested that mutant mice were
long-term exposed to higher estrogen and therefore exhibited increased bone mineral
density.
86
Fig. 28. Vascular wall dilation in mutant ovaries after PMSG and hCG
treatments. (A) Mice were treated with PMSG and hCG as in Fig. 5. Vascular
dilation was observed in both 3-month and 7-month-old mutant ovaries. (B) H&E
staining of sections from wildtype and mutant ovaries. Note the blood vessels were
dilated in mutants (arrows) compared to those in wildtype (arrowheads).
A
B
7-month-old
3-month-old
Wildtype Mutant
Wildtype
Mutant
CL
CL
CL
CL
CL
CL
CL
87
Fig. 29. There is no difference in the number of blood vessels in corpus luteum
between wildtype and mutant after PMSG and hCG treatments. (A) Anti-Von
Willebrand factor antibody staining revealed the vascular endothelial cells. Note the
dilated blood vessels in the mutant. (B) The number of the blood vessels in the
corpus luteum was not changed in mutant mice.
CL
A
Wildtype Mutant
CL
B
WT MT
88
Fig. 30. High trabecular bone mass in mutant female mice. (A) μCT images of
distal femoral trabecular bone of wildtype (left) and mutant (right) 6-month-old
mice with median BV/TV. (B) μCT analysis of the distal femoral trabecular bone of
wildtype (white) and mutant (black) female mice. BV/TV – trabecular bone volume
density, Tb.N – trabecular number, Tb.Th – trabecular thickness, Conn.D –
connectivity density (n=6 in WT; n=5 in MT; * = p<0.05).
A Wildtype Mutant
B
*
*
*
*
89
Conclusion
We reported earlier that conditional inactivation of Brca1 in ovarian granulosa cells led
to serous cystadenomas in ovaries and uteri (Chodankar et al., 2005). Granulosa cells
play an important role in the regulation of estrus cycle and hormone releasing. The estrus
cycle can be divided into four phases: proestrus, estrus, metestrus and diestrus. Serum
estrogen levels rise during proestrus and peak at the end of this stage. Ovulation occurs
during the following estrus phase. Hao Hong in the lab demonstrated that the ratio of the
estrogen-dominated pre-ovulatory phases to post-ovulatory phases (i.e. the ratio of
proestrus phase/metestrus phase) was increased in Fshr-Cre;Brca1
flox/flox
mutant mice
(paper in press). In addition, the magnitude of this increase is associated with increased
tumor risk.
Longer estrogenic phase suggests higher serum estrogen levels. Consistent with above
data, I showed that the serum estradiol levels were also increased in mutant mice after
estrus synchronization treatment with PMSG. Ovarian transplantation experiments
demonstrated that the estrogen elevation effects were due to the ovaries carrying mutated
Brca1 in granulosa cells. Given that estrogen is a significant factor in increasing the risk
of ovarian cancer, this result provides an evidence that Brca1 is able to control circulating
estrogen levels in the mutant mice with a high predisposition to ovarian and uterine
epithelial tumors.
Despite the big influence of germline BRCA1 mutation on the development of familial
ovarian and breast cancers, somatic mutation of this gene is rarely found in sporadic
90
tumors. According to our cell-nonautonomous model, it is possible that the sporadic
tumor is caused by long-term estrogen stimulation. The tumor itself may not necessarily
carries Brca1 mutation, which was exactly what happened to Fshr-Cre;Brca1
flox/flox
mutant mice. The PCR results demonstrated that the tumors of mutant mice only carried
normal Brca1 gene (Chodankar et al., 2005).
The phenotypes we observed in the mutant mice suggested that the mice were exposed to
higher levels of functional estrogen at both pre-ovulatory phase and post-ovulatory phase.
During the pre-ovulatory, or follicular, phase of the estrus cycle, estradiol acts on uterine
stromal cells to promote cell proliferation. In mutant mice, I demonstrated that the uterine
stromal cells had a higher proliferation rate compared with wildtype. Whether this higher
proliferation rate in stromal cells is associated with tumor formation needs further study.
After ovulation, the luteal phase occurs and is characterized by the formation of corpora
lutea (CL) on the collapsed follicle. CL is a highly vascularized organ. The blood flow is
increased dramatically during early luteal phase (Takasaki et al., 2009). After PMSG and
hCG stimulation, I observed the blood vessels of the ovary were enlarged in mutant mice.
The blood vessels dilatation was likely caused by estrogen, given that estrogen is known
to act as a vasodilator (Mendelsohn and Karas, 1999) and be able to induce vascular wall
dilation through nitric oxide signaling pathway (Leung et al., 2007). This phenotype may
simply reflects the well-established protective role of estrogen in cardiovascular system,
but not be associated with tumor development.
91
In addition to short-term estrogenic effects on reproductive organs including ovaries and
uteri, we also provided an evidence that mutant mice underwent long-term exposure to
higher levels of circulating estrogen. Many reports indicated that estrogen treatments
increase bone mineral density in human (Lindsay and Tohme, 1990; Felson et al., 1993;
Cauley et al., 2003). Consistent with this argument, the bone mass and other bone density
parameters of mutant mice were significantly higher than those of wildtype controls,
suggesting an increased estrogenic effect in mutant mice. These 6-month-old mice were
relatively too young to develop tumors (tumors in the mutant mice were observed at 12 to
20 months of age), but old enough to accumulated estrogenic effects on bone metabolism.
Previous studies have demonstrated that increased bone mineral density is significantly
associated with an increased risk of breast cancer and endometrial cancer. (Persson et al.,
1994; Cauley et al., 1996; Zhang et al., 1997; Hadji et al., 2007). Although the correlation
between bone density and ovarian cancer is not clear, the phenotype we observed in our
mutant mice supports this hypothesis. Therefore, bone mineral density may be a potential
screening parameter for ovarian cancer.
Estrogen is an important risk factor for breast and ovarian cancer. In this study, we
demonstrated that the serum estradiol levels were significantly increased in mice carrying
a Brca1 mutation in their ovarian granulosa cells. Increased estrogen levels contribute to
short-term and long-term effects on mutant mice. This study provides direct evidence that
Brca1 regulates functional circulating estrogen in mice and further supports our cell-
nonautonomous model in Brca1-related tumor development.
92
Chapter 2 References
Cauley, J. A., Lucas, F. L., Kuller, L. H., Vogt, M. T., Browner, W. S. and
Cummings, S. R. (1996). Bone mineral density and risk of breast cancer in older women:
the study of osteoporotic fractures. Study of Osteoporotic Fractures Research Group.
JAMA. 276, 1404-1408.
Cauley, J. A., Robbins, J., Chen, Z., Cummings, S. R., Jackson, R. D., LaCroix, A.
Z., LeBoff, M., Lewis, C. E., McGowan, J., Neuner, J., et al. Women's Health
Initiative Investigators. (2003). Effects of estrogen plus progestin on risk of fracture and
bone mineral density: the Women's Health Initiative randomized trial. JAMA. 290, 1729-
1738.
Chodankar, R., Kwang, S., Sangiorgi, F., Hong, H., Yen, H. Y., Deng, C., Pike, M.
C., Shuler, C. F., Maxson, R. and Dubeau, L. (2005). Cell-nonautonomous induction
of ovarian and uterine serous cystadenomas in mice lacking a functional Brca1 in ovarian
granulosa cells. Curr. Biol. 15, 561-565.
Deng, C. X. (2006). BRCA1: cell cycle checkpoint, genetic instability, DNA damage
response and cancer evolution. Nucleic. Acids Res. 34, 1416-1426.
Easton, D. F., Ford, D. and Bishop, D. T. (1995). Breast and ovarian cancer incidence
in BRCA1-mutation carriers. Am. J. Hum. Genet. 56, 265-271
Elstrodt, F., Hollestelle, A., Nagel, J. H., Gorin, M., Wasielewski, M., van den
Ouweland, A., Merajver, S. D., Ethier, S. P. and Schutte, M. (2006). BRCA1
mutation analysis of 41 human breast cancer cell lines reveals three new deleterious
mutants. Cancer Res. 66, 41-45.
Fan, S., Wang, J., Yuan, R., Ma, Y., Meng, Q., Erdos, M. R., Pestell, R. G., Yuan, F.,
Auborn, K. J., Goldberg, I. D. and Rosen, E. M. (1999). BRCA1 inhibition of estrogen
receptor signaling in transfected cells. Science 284, 1354-1356.
Felson, D. T., Zhang, Y., Hannan, M. T., Kiel, D. P., Wilson, P. W. and Anderson, J.
J. (1993). The effect of postmenopausal estrogen therapy on bone density in elderly
women. N. Engl. J. Med. 329, 1141-1146.
Futreal, P. A., Liu, Q., Shattuck-Eidens, D., Cochran, C., Harshman, K., Tavtigian,
S., Bennett, L. M., Haugen-Strano, A., Swensen, J., Miki, Y, et al. (1994). BRCA1
mutations in primary breast and ovarian carcinomas. Science 266, 120-122.
Ghosh, S., Lu, Y., Katz, A., Hu, Y. and Li, R. (2007). Tumor suppressor BRCA1
inhibits a breast cancer-associated promoter of the aromatase gene (CYP19) in human
adipose stromal cells. Am. J. Physiol. Endocrinol. Metab. 292, E246-E252.
93
Griswold, M. D., Heckert, L. and Linder, C. (1995). The molecular biology of the FSH
receptor. J. Steroid. Biochem. Mol. Biol. 53, 215-218.
Gromoll, J. and Simoni, M. (2005). Genetic complexity of FSH receptor function.
Trends Endocrinol. Metab. 16, 368-373.
Guo, X., Razandi, M., Pedram, A., Kassab, G. and Levin, E. R. (2005). Estrogen
induces vascular wall dilation: mediation through kinase signaling to nitric oxide and
estrogen receptors alpha and beta. J. Biol. Chem. 280, 19704-19710.
Hadji, P., Gottschalk, M., Ziller, V., Kalder, M., Jackisch, C. and Wagner, U.
(2007). Bone mass and the risk of breast cancer: the influence of cumulative exposure to
oestrogen and reproductive correlates. Results of the Marburg breast cancer and
osteoporosis trial (MABOT). Maturitas. 56, 312-321.
Hall, J. M., Lee, M. K., Newman, B., Morrow, J. E., Anderson, L. A., Huey, B. and
King, M.C. (1990). Linkage of early-onset familial breast cancer to chromosome 17q21.
Science 250, 1684-1689.
Hartge, P. (2006). Menopausal hormone therapy and ovarian cancer risk in the National
Institutes of Health-AARP Diet and Health Study Cohort. J. Natl. Cancer Inst. 98, 1397-
1405.
Hu, Y., Ghosh, S., Amleh, A., Yue, W., Lu, Y., Katz, A. and Li, R. (2005).
Modulation of aromatase expression by BRCA1: a possible link to tissue-specific tumor
suppression. Oncogene. 24, 8343-8348.
Hu, Y. (2009). BRCA1, hormone, and tissue-specific tumor suppression. Int. J. Biol. Sci.
5, 20-27.
Kauff, N. D., Satagopan, J. M., Robson, M. E., Scheuer, L., Hensley, M., Hudis, C.
A., Ellis, N. A., Boyd, J., Borgen, P. I., Barakat, R. R., Norton, L., Castiel, M., Nafa,
K. and Offit, K. (2002). Risk-reducing salpingo-oophorectomy in women with a BRCA1
or BRCA2 mutation. N. Engl. J. Med. 346, 1609-1615.
King, M. C., Marks, J. H., Mandell, J. B.; New York Breast Cancer Study Group.
(2003). Breast and ovarian cancer risks due to inherited mutations in BRCA1 and
BRCA2. Science 302, 643-646.
Lacey, J. V. Jr., Brinton, L. A., Leitzmann, M. F., Mouw, T., Hollenbeck, A.,
Schatzkin, A., Leung, S. W., Teoh, H., Keung, W. and Man, R. Y. (2007). Non-
genomic vascular actions of female sex hormones: physiological implications and
signalling pathways. Clin. Exp. Pharmacol. Physiol. 34, 822-6.
94
Lindsay, R. and Tohme, J. F. (1990). Estrogen treatment of patients with established
postmenopausal osteoporosis. Obstet. Gynecol. 76, 290-295.
Lu, M., Chen, D., Lin, Z., Reierstad, S., Trauernicht, A. M., Boyer, T. G. and Bulun,
S. E. (2006). BRCA1 negatively regulates the cancer-associated aromatase promoters I.3
and II in breast adipose fibroblasts and malignant epithelial cells. J. Clin. Endocrinol.
Metab. 91, 4514-4519.
Mendelsohn, M. E. and Karas, R. H. (1999). The protective effects of estrogen on the
cardiovascular system. N. Engl. J. Med. 340, 1801-1811.
Mørch, L. S., Løkkegaard, E., Andreasen, A. H., Krüger-Kjaer, S. and Lidegaard,
O. (2009). Hormone therapy and ovarian cancer. JAMA. 302, 298-305.
Persson, I., Adami, H.O., McLaughlin, J.K., Naessén, T. and Fraumeni, J. F. Jr.
(1994). Reduced risk of breast and endometrial cancer among women with hip fractures
(Sweden). Cancer Causes Control. 5, 523-528.
Rebbeck, T. R., Lynch, H. T., Neuhausen, S. L., Narod, S. A., Van't Veer, L.,
Garber, J. E., Evans, G., Isaacs, C., Daly, M. B., Matloff, E., Olopade, O. I., Weber,
B. L.; Prevention and Observation of Surgical End Points Study Group. (2002).
Prophylactic oophorectomy in carriers of BRCA1 or BRCA2 mutations. N. Engl. J. Med.
346, 1616-1622.
Soriano, P. (1999). Generalized lacZ expression with the ROSA26 Cre reporter strain.
Nat. Genet. 21, 70-71.
Stanczyk, F. Z., Cho, M. M., Endres, D. B., Morrison, J. L., Patel, S. and Paulson, R.
J. (2003). Limitations of direct estradiol and testosterone immunoassay kits. Steroids 68,
1173-1178.
Stocco, C. (2008). Aromatase expression in the ovary: hormonal and molecular
regulation. Steroids. 73, 473-487.
Takasaki, A., Tamura, H., Taniguchi, K., Asada, H., Taketani, T., Matsuoka, A.,
Yamagata, Y., Shimamura, K., Morioka, H. and Sugino, N. (2009). Luteal blood flow
and luteal function. J. Ovarian Res. 2, 1.
Wood, G. A., Fata, J. E., Watson, K. L. and Khokha, R. (2007). Circulating hormones
and estrous stage predict cellular and stromal remodeling in murine uterus. Reproduction
133, 1035-1044
95
Xu, X., Wagner, K. U., Larson, D., Weaver, Z., Li, C., Ried, T., Hennighausen, L.,
Wynshaw-Boris, A. and Deng, C. X. (1999). Conditional mutation of Brca1 in
mammary epithelial cells results in blunted ductal morphogenesis and tumour formation.
Nat. Genet. 22, 37-43.
Zhang, Y., Kiel, D. P., Kreger, B. E., Cupples, L. A., Ellison, R. C., Dorgan, J. F.,
Schatzkin, A., Levy, D. and Felson, D. T. (1997). Bone mass and the risk of breast
cancer among postmenopausal women. N. Engl. J. Med. 336, 611-617.
96
Bibliography
Alagille, D., Estrada, A., Hadchouel, M., Gautier, M., Odièvre, M. and
Dommergues, J. P. (1987). Syndromic paucity of interlobular bile ducts (Alagille
syndrome or arteriohepatic dysplasia): review of 80 cases. J. Pediatr. 110, 195-200.
Aviv, R. I., Rodger, E., Hall, C. M. (2002). Craniosynostosis. Clin. Radiol. 57, 93-102.
Blair, S. S. (2003). Lineage compartments in Drosophila. Curr. Biol. 13, R548-551.
Bolós, V., Grego-Bessa, J. and de la Pompa, J. L. (2007). Notch signaling in
development and cancer. Endocr. Rev. 28, 339-363.
Bourgeois, P., Bolcato-Bellemin, A. L., Danse, J. M., Bloch-Zupan, A., Yoshiba, K.,
Stoetzel, C. and Perrin-Schmitt, F. (1998). The variable expressivity and incomplete
penetrance of the twist-null heterozygous mouse phenotype resemble those of human
Saethre-Chotzen syndrome. Hum. Mol. Genet. 7, 945-957.
Boyadjiev, S. A. (2007). International Craniosynostosis Consortium. Genetic analysis of
non-syndromic craniosynostosis. Orthod. Craniofac. Res. 10, 129-137.
Bray, S. (1998). Notch signalling in Drosophila: three ways to use a pathway. Semin.
Cell Dev. Biol. 9, 591-597.
Bray, S. J. (2006). Notch signalling: a simple pathway becomes complex. Nat. Rev. Mol.
Cell Biol. 7, 678-689.
Brooker, R., Hozumi, K. and Lewis, J. (2006). Notch ligands with contrasting
functions: Jagged1 and Delta1 in the mouse inner ear. Development 133, 1277-1286.
Buceta, J., Herranz, H., Canela-Xandri, O., Reigada, R., Sagués, F. and Milán, M.
(2007). Robustness and stability of the gene regulatory network involved in DV boundary
formation in the Drosophila wing. PLoS One 2, e602.
Carver, E. A., Oram, K. F. and Gridley, T. (2002). Craniosynostosis in Twist
heterozygous mice: a model for Saethre-Chotzen syndrome. Anat. Rec. 268, 90-92.
Castanon, I. and Baylies, M. K. (2002). A Twist in fate: evolutionary comparison of
Twist structure and function. Gene 287, 11-22.
Castanon, I., Von Stetina, S., Kass, J., Baylies, M.K. (2001). Dimerization partners
determine the activity of the Twist bHLH protein during Drosophila mesoderm
development. Development 128, 3145--3159.
97
Cauley, J. A., Lucas, F. L., Kuller, L. H., Vogt, M. T., Browner, W. S. and
Cummings, S. R. (1996). Bone mineral density and risk of breast cancer in older women:
the study of osteoporotic fractures. Study of Osteoporotic Fractures Research Group.
JAMA. 276, 1404-1408.
Cauley, J. A., Robbins, J., Chen, Z., Cummings, S. R., Jackson, R. D., LaCroix, A.
Z., LeBoff, M., Lewis, C. E., McGowan, J., Neuner, J., et al. Women's Health
Initiative Investigators. (2003). Effects of estrogen plus progestin on risk of fracture and
bone mineral density: the Women's Health Initiative randomized trial. JAMA. 290, 1729-
1738.
Chai, Y. and Maxson, R. E. Jr. (2006). Recent advances in craniofacial morphogenesis.
Dev. Dyn. 235, 2353-2375.
Chen, Y. H., Ishii, M., Sun, J., Sucov, H. M., Maxson, R. E. Jr. (2007). Msx1 and
Msx2 regulate survival of secondary heart field precursors and post-migratory
proliferation of cardiac neural crest in the outflow tract. Dev. Biol. 308, 421-437.
Chen, Z. F. and Behringer, R. R. (1995). twist is required in head mesenchyme for
cranial neural tube morphogenesis. Genes Dev. 9, 686-699.
Chodankar, R., Kwang, S., Sangiorgi, F., Hong, H., Yen, H. Y., Deng, C., Pike, M.
C., Shuler, C. F., Maxson, R. and Dubeau, L. (2005). Cell-nonautonomous induction
of ovarian and uterine serous cystadenomas in mice lacking a functional Brca1 in ovarian
granulosa cells. Curr. Biol. 15, 561-565.
Cohen, M. M. Jr. (2006). The new bone biology: pathologic, molecular, and clinical
correlates. Am. J. Med. Genet. A. 140, 2646-2706.
Connerney, J., Andreeva, V., Leshem, Y., Mercado, M. A., Dowell, K., Yang, X.,
Lindner, V., Friesel, R. E. and Spicer, D. B. (2008). Twist1 homodimers enhance FGF
responsiveness of the cranial sutures and promote suture closure. Dev. Biol. 318, 323-
334.
Dahlqvist, C., Blokzijl, A., Chapman, G., Falk, A., Dannaeus, K., Ibâñez, C. F.,
Lendahl, U. (2003). Functional Notch signaling is required for BMP4-induced inhibition
of myogenic differentiation. Development 130, 6089-6099.
Dahmann, C. and Basler, K. (1999). Compartment boundaries: at the edge of
development. Trends Genet. 15, 320-326.
Danielian, P. S., Muccino, D., Rowitch, D. H., Michael, S. K. and McMahon, A. P.
(1998) Modification of gene activity in mouse embryos in utero by a tamoxifen-inducible
form of Cre recombinase. Curr. Biol. 8, 1323-1326.
98
de Celis, J. F., Tyler, D. M., de Celis, J., Bray, S. J. (1998). Notch signalling mediates
segmentation of the Drosophila leg. Development 125, 4617-4626.
Deng, C. X. (2006). BRCA1: cell cycle checkpoint, genetic instability, DNA damage
response and cancer evolution. Nucleic. Acids Res. 34, 1416-1426.
Easton, D. F., Ford, D. and Bishop, D. T. (1995). Breast and ovarian cancer incidence
in BRCA1-mutation carriers. Am. J. Hum. Genet. 56, 265-271
el Ghouzzi, V., Le Merrer, M., Perrin-Schmitt, F., Lajeunie, E., Benit, P., Renier, D.,
Bourgeois, P., Bolcato-Bellemin, A. L., Munnich, A. and Bonaventure, J. (1997).
Mutations of the TWIST gene in the Saethre-Chotzen syndrome. Nat. Genet. 15, 42-46.
Elstrodt, F., Hollestelle, A., Nagel, J. H., Gorin, M., Wasielewski, M., van den
Ouweland, A., Merajver, S. D., Ethier, S. P. and Schutte, M. (2006). BRCA1
mutation analysis of 41 human breast cancer cell lines reveals three new deleterious
mutants. Cancer Res. 66, 41-45.
Emerick, K. M., Rand, E.B., Goldmuntz, E., Krantz, I. D., Spinner, N. B. and
Piccoli, D. A. (1999). Features of Alagille syndrome in 92 patients: frequency and
relation to prognosis. Hepatology 29, 822-829.
Fan, S., Wang, J., Yuan, R., Ma, Y., Meng, Q., Erdos, M. R., Pestell, R. G., Yuan, F.,
Auborn, K. J., Goldberg, I. D. and Rosen, E. M. (1999). BRCA1 inhibition of estrogen
receptor signaling in transfected cells. Science 284, 1354-1356.
Felson, D. T., Zhang, Y., Hannan, M. T., Kiel, D. P., Wilson, P. W. and Anderson, J.
J. (1993). The effect of postmenopausal estrogen therapy on bone density in elderly
women. N. Engl. J. Med. 329, 1141-1146.
Futreal, P. A., Liu, Q., Shattuck-Eidens, D., Cochran, C., Harshman, K., Tavtigian,
S., Bennett, L. M., Haugen-Strano, A., Swensen, J., Miki, Y, et al. (1994). BRCA1
mutations in primary breast and ovarian carcinomas. Science 266, 120-122.
Ghosh, S., Lu, Y., Katz, A., Hu, Y. and Li, R. (2007). Tumor suppressor BRCA1
inhibits a breast cancer-associated promoter of the aromatase gene (CYP19) in human
adipose stromal cells. Am. J. Physiol. Endocrinol. Metab. 292, E246-E252.
Golson, M. L., Loomes, K. M., Oakey, R. and Kaestner, K. H. (2009). Ductal
malformation and pancreatitis in mice caused by conditional Jag1 deletion.
Gastroenterology 136, 1761-1771.e1.
González, A., Chaouiya, C. and Thieffry, D. (2006). Dynamical analysis of the
regulatory network defining the dorsal-ventral boundary of the Drosophila wing imaginal
disc. Genetics 174, 1625-1634.
99
Griswold, M. D., Heckert, L. and Linder, C. (1995). The molecular biology of the FSH
receptor. J. Steroid. Biochem. Mol. Biol. 53, 215-218.
Gromoll, J. and Simoni, M. (2005). Genetic complexity of FSH receptor function.
Trends Endocrinol. Metab. 16, 368-373.
Guo, X., Razandi, M., Pedram, A., Kassab, G. and Levin, E. R. (2005). Estrogen
induces vascular wall dilation: mediation through kinase signaling to nitric oxide and
estrogen receptors alpha and beta. J. Biol. Chem. 280, 19704-19710.
Hadji, P., Gottschalk, M., Ziller, V., Kalder, M., Jackisch, C. and Wagner, U.
(2007). Bone mass and the risk of breast cancer: the influence of cumulative exposure to
oestrogen and reproductive correlates. Results of the Marburg breast cancer and
osteoporosis trial (MABOT). Maturitas. 56, 312-321.
Hajihosseini, M. K. (2008). Fibroblast growth factor signaling in cranial suture
development and pathogenesis. Front. Oral Biol. 12, 160-177.
Hall, J. M., Lee, M. K., Newman, B., Morrow, J. E., Anderson, L. A., Huey, B. and
King, M.C. (1990). Linkage of early-onset familial breast cancer to chromosome 17q21.
Science 250, 1684-1689.
Hartge, P. (2006). Menopausal hormone therapy and ovarian cancer risk in the National
Institutes of Health-AARP Diet and Health Study Cohort. J. Natl. Cancer Inst. 98, 1397-
1405.
Holmes, G., Rothschild, G., Roy, U. B., Deng, C. X., Mansukhani, A. and Basilico, C.
(2009). Early onset of craniosynostosis in an Apert mouse model reveals critical features
of this pathology. Dev. Biol. 328, 273-84.
Howard, T. D., Paznekas, W. A., Green, E. D., Chiang, L. C., Ma, N., Ortiz de Luna,
R. I., Garcia Delgado, C., Gonzalez-Ramos, M., Kline, A. D. and Jabs, E. W. (1997).
Mutations in TWIST, a basic helix-loop-helix transcription factor, in Saethre-Chotzen
syndrome. Nat. Genet. 15, 36-41.
Hu, Y. (2009). BRCA1, hormone, and tissue-specific tumor suppression. Int. J. Biol. Sci.
5, 20-27.
Hu, Y., Ghosh, S., Amleh, A., Yue, W., Lu, Y., Katz, A. and Li, R. (2005).
Modulation of aromatase expression by BRCA1: a possible link to tissue-specific tumor
suppression. Oncogene. 24, 8343-8348.
Irvine, K. D. and Rauskolb, C. (2001). Boundaries in development: formation and
function. Annu. Rev. Cell Dev. Biol. 17, 189-214.
100
Ishii, M., Merrill, A. E., Chan, Y. S., Gitelman, I., Rice, D. P., Sucov, H. M., Maxson,
R. E. Jr. (2003). Msx2 and Twist cooperatively control the development of the neural
crest-derived skeletogenic mesenchyme of the murine skull vault. Development 130,
6131-6142.
Jabs, E. W., Müller, U., Li, X., Ma, L., Luo, W., Haworth, I. S., Klisak, I., Sparkes,
R., Warman, M. L., Mulliken, J. B., et al. (1993). A mutation in the homeodomain of
the human MSX2 gene in a family affected with autosomal dominant craniosynostosis.
Cell 75, 443-450.
Jarriault, S., Brou, C., Logeat, F., Schroeter, E. H., Kopan, R. and Israel, A. (1995).
Signalling downstream of activated mammalian Notch. Nature 377, 355-358.
Jenkins, D., Seelow, D., Jehee, F. S., Perlyn, C. A., Alonso, L. G., Bueno, D. F.,
Donnai, D., Josifova, D., Mathijssen, I. M., Morton, J. E., Orstavik, K. H,. Sweeney,
E., Wall, S. A., Marsh, J. L., Nurnberg, P., Passos-Bueno, M. R. and Wilkie, A. O.
(2007). RAB23 mutations in Carpenter syndrome imply an unexpected role for hedgehog
signaling in cranial-suture development and obesity. Am. J. Hum. Genet. 80, 1162-1170.
Jiang, X., Iseki, S., Maxson, R. E., Sucov, H. M. and Morriss-Kay, G. M. (2002).
Tissue origins and interactions in the mammalian skull vault. Dev. Biol. 241, 106-116.
Johnson, D., Iseki, S., Wilkie, A. O. and Morriss-Kay, G. M. (2000). Expression
patterns of Twist and Fgfr1, -2 and -3 in the developing mouse coronal suture suggest a
key role for twist in suture initiation and biogenesis. Mech. Dev, 91, 341-345.
Kabbani, H and Raghuveer, T. S. (2004). Craniosynostosis. Am. Fam. Physician. 69,
2863-2870.
Kamath, B. M., Spinner, N. B., Emerick, K. M., Chudley, A. E., Booth, C., Piccoli,
D. A. and Krantz, I. D. (2004). Vascular anomalies in Alagille syndrome: a significant
cause of morbidity and mortality. Circulation 109, 1354-1358.
Kamath, B. M., Stolle, C., Bason, L., Colliton, R. P., Piccoli, D. A., Spinner, N. B.
and Krantz, I. D. (2002). Craniosynostosis in Alagille syndrome. Am. J. Med. Genet.
112, 176-180.
Kauff, N. D., Satagopan, J. M., Robson, M. E., Scheuer, L., Hensley, M., Hudis, C.
A., Ellis, N. A., Boyd, J., Borgen, P. I., Barakat, R. R., Norton, L., Castiel, M., Nafa,
K. and Offit, K. (2002). Risk-reducing salpingo-oophorectomy in women with a BRCA1
or BRCA2 mutation. N. Engl. J. Med. 346, 1609-1615.
Kiecker, C. and Lumsden, A. (2005). Compartments and their boundaries in vertebrate
brain development. Nat. Rev. Neurosci. 6, 553-564.
101
Kiernan, A. E., Xu, J. and Gridley, T. (2006). The Notch ligand JAG1 is required for
sensory progenitor development in the mammalian inner ear. PLoS Genet. 2, e4.
King, M. C., Marks, J. H., Mandell, J. B.; New York Breast Cancer Study Group.
(2003). Breast and ovarian cancer risks due to inherited mutations in BRCA1 and
BRCA2. Science 302, 643-646.
Kopan, R. and Ilagan, M. X. (2009). The canonical Notch signaling pathway: unfolding
the activation mechanism. Cell 137, 216-233.
Krantz, I. D., Colliton, R. P., Genin, A., Rand, E. B., Li, L., Piccoli, D. A. and
Spinner, N. B. (1998). Spectrum and frequency of jagged1 (JAG1) mutations in Alagille
syndrome patients and their families. Am. J. Hum. Genet. 62, 1361-1369.
Lacey, J. V. Jr., Brinton, L. A., Leitzmann, M. F., Mouw, T., Hollenbeck, A.,
Schatzkin, A., Leung, S. W., Teoh, H., Keung, W. and Man, R. Y. (2007). Non-
genomic vascular actions of female sex hormones: physiological implications and
signalling pathways. Clin. Exp. Pharmacol. Physiol. 34, 822-6.
Lai, E. C. (2004). Notch signaling: control of cell communication and cell fate.
Development 131, 965-973.
Le, T. T., Conley, K. W. and Brown, N. L. (2009). Jagged 1 is necessary for normal
mouse lens formation. Dev. Biol. 328, 118-126.
Li, C., Scott, D. A., Hatch, E., Tian, X. and Mansour, S. L. (2007). Dusp6 (Mkp3) is a
negative feedback regulator of FGF-stimulated ERK signaling during mouse
development. Development 134, 167-176.
Lindsay, R. and Tohme, J. F. (1990). Estrogen treatment of patients with established
postmenopausal osteoporosis. Obstet. Gynecol. 76, 290-295.
Liu, B., Yu, H. M. and Hsu, W. (2007). Craniosynostosis caused by Axin2 deficiency is
mediated through distinct functions of beta-catenin in proliferation and differentiation.
Dev. Biol. 301, 298-308.
Loeys, B. L., Chen, J., Neptune, E. R., Judge, D. P., Podowski, M., Holm, T., Meyers,
J., Leitch, C. C., Katsanis, N., Sharifi, N., et al. (2005). A syndrome of altered
cardiovascular, craniofacial, neurocognitive and skeletal development caused by
mutations in TGFBR1 or TGFBR2. Nat. Genet. 37, 275-281.
Lovato, T. A. L., Benjamin, A. R., Cripps, R. M. (2005). Transcription of Myocyte
enhancer factor-2 in adult Drosophila myoblasts is induced by the steroid hormone
ecdysone. Dev. Biol. 288, 612-621.
102
Lozier, J., McCright, B. and Gridley, T. (2008). Notch signaling regulates bile duct
morphogenesis in mice. PLoS One 3, e1851.
Lu, M., Chen, D., Lin, Z., Reierstad, S., Trauernicht, A. M., Boyer, T. G. and Bulun,
S. E. (2006). BRCA1 negatively regulates the cancer-associated aromatase promoters I.3
and II in breast adipose fibroblasts and malignant epithelial cells. J. Clin. Endocrinol.
Metab. 91, 4514-4519.
Ma, A., Boulton, M., Zhao, B., Connon, C., Cai, J., Albon, J. (2007). A role for notch
signaling in human corneal epithelial cell differentiation and proliferation. Invest.
Ophthalmol. Vis. Sci. 48, 3576-3585.
Major, R. J. and Irvine, K. D. (2005). Influence of Notch on dorsoventral
compartmentalization and actin organization in the Drosophila wing. Development 132,
3823-3833.
Mathijssen, I. M., van Splunder, J., Vermeij-Keers, C., Pieterman, H., de Jong, T.
H., Mooney, M. P. and Vaandrager, J. M. (1999). Tracing craniosynostosis to its
developmental stage through bone center displacement. J. Craniofac. Genet. Dev. Biol.
19, 57–63.
McCright, B., Gao, X., Shen, L., Lozier, J., Lan, Y., Maguire, M., Herzlinger, D.,
Weinmaster, G., Jiang, R. and Gridley, T. (2001). Defects in development of the
kidney, heart and eye vasculature in mice homozygous for a hypomorphic Notch2
mutation. Development 128, 491-502.
McCright, B., Lozier, J. and Gridley, T. (2002). A mouse model of Alagille syndrome:
Notch2 as a genetic modifier of Jag1 haploinsufficiency. Development 129, 1075-1082.
McDaniell, R., Warthen, D. M., Sanchez-Lara, P. A., Pai, A., Krantz, I. D., Piccoli,
D. A. and Spinner, N. B. (2006). NOTCH2 mutations cause Alagille syndrome, a
heterogeneous disorder of the notch signaling pathway. Am. J. Hum. Genet. 79, 169-173.
McLarren, K. W., Lo, R., Grbavec, D., Thirunavukkarasu, K., Karsenty, G. and
Stifani, S. (2000). The mammalian basic helix loop helix protein HES-1 binds to and
modulates the transactivating function of the runt-related factor Cbfa1. J. Biol. Chem.
275, 530-538.
McLeod, M. J. (1980). Differential staining of cartilage and bone in whole mouse fetuses
by alcian blue and alizarin red S. Teratology 22, 299-301.
Mendelsohn, M. E. and Karas, R. H. (1999). The protective effects of estrogen on the
cardiovascular system. N. Engl. J. Med. 340, 1801-1811.
103
Merrill, A. E., Bochukova, E. G., Brugger, S. M., Ishii, M., Pilz, D. T., Wall, S. A.,
Lyons, K. M., Wilkie, A. O. and Maxson R. E. Jr. (2006). Cell mixing at a neural
crest-mesoderm boundary and deficient ephrin-Eph signaling in the pathogenesis of
craniosynostosis. Hum. Mol. Genet. 15, 1319-1328.
Micchelli, C. A., Rulifson, E. J. and Blair, S. S. (1997). The function and regulation of
cut expression on the wing margin of Drosophila: Notch, Wingless and a dominant
negative role for Delta and Serrate. Development 124, 1485-1495.
Moenning, A., Jäger, R., Egert, A., Kress, W., Wardelmann, E. and Schorle, H.
(2009). Sustained platelet-derived growth factor receptor alpha signaling in osteoblasts
results in craniosynostosis by overactivating the phospholipase C-gamma pathway. Mol.
Cell. Biol. 29, 881-891.
Mørch, L. S., Løkkegaard, E., Andreasen, A. H., Krüger-Kjaer, S. and Lidegaard,
O. (2009). Hormone therapy and ovarian cancer. JAMA. 302, 298-305.
Morriss-Kay, G. M. and Wilkie, A. O. (2005). Growth of the normal skull vault and its
alteration in craniosynostosis: insights from human genetics and experimental studies. J.
Anat. 207, 637-653.
Nobta, M., Tsukazaki, T., Shibata, Y., Xin, C., Moriishi, T., Sakano, S., Shindo, H.
and Yamaguchi, A. (2005). Critical regulation of bone morphogenetic protein-induced
osteoblastic differentiation by Delta1/Jagged1-activated Notch1 signaling. J. Biol. Chem.
280, 15842-15848.
Oda, T., Elkahloun, A. G., Pike, B. L, Okajima, K., Krantz, I. D., Genin, A., Piccoli,
D. A., Meltzer, P. S., Spinner, N. B., Collins, F. S. and Chandrasekharappa, S. C.
(1997). Mutations in the human Jagged1 gene are responsible for Alagille syndrome.
Nat. Genet. 16, 235-242.
Opperman, L. A. (2000). Cranial sutures as intramembranous bone growth sites. Dev.
Dyn. 219, 472-485.
Oram, K. F. and Gridley, T. (2005). Mutations in snail family genes enhance
craniosynostosis of Twist1 haplo-insufficient mice: implications for Saethre-Chotzen
Syndrome. Genetics 170, 971-974.
Persson, I., Adami, H.O., McLaughlin, J.K., Naessén, T. and Fraumeni, J. F. Jr.
(1994). Reduced risk of breast and endometrial cancer among women with hip fractures
(Sweden). Cancer Causes Control. 5, 523-528.
104
Rebbeck, T. R., Lynch, H. T., Neuhausen, S. L., Narod, S. A., Van't Veer, L.,
Garber, J. E., Evans, G., Isaacs, C., Daly, M. B., Matloff, E., Olopade, O. I., Weber,
B. L.; Prevention and Observation of Surgical End Points Study Group. (2002).
Prophylactic oophorectomy in carriers of BRCA1 or BRCA2 mutations. N. Engl. J. Med.
346, 1616-1622.
Rice, D. P. (2008). Developmental anatomy of craniofacial sutures. Front. Oral Biol. 12,
1-21.
Rice, D. P., Aberg, T., Chan, Y., Tang, Z., Kettunen, P. J., Pakarinen, L., Maxson,
R. E. and Thesleff, I. (2000). Integration of FGF and TWIST in calvarial bone and
suture development. Development 127, 1845-1855.
Saga, Y., Miyagawa-Tomita, S., Takagi, A., Kitajima, S., Miyazaki, J. and Inoue, T.
(1999). MesP1 is expressed in the heart precursor cells and required for the formation of
a single heart tube. Development 126, 3437-3447.
Saravanamuthu, S. S., Gao, C. Y. and Zelenka, P. S. (2009). Notch signaling is
required for lateral induction of Jagged1 during FGF-induced lens fiber differentiation.
Dev. Biol. 332, 166-176.
Schröder, N. and Gossler, A. (2002). Expression of Notch pathway components in fetal
and adult mouse small intestine. Gene Expr. Patterns. 2, 247-250.
Settle, S. H. Jr., Rountree, R. B., Sinha, A., Thacker, A., Higgins, K. and Kingsley,
D. M. (2003). Multiple joint and skeletal patterning defects caused by single and double
mutations in the mouse Gdf6 and Gdf5 genes. Dev. Biol. 254, 116-130.
Shimizu, T., Tanaka, T., Iso, T., Doi, H., Sato, H., Kawai-Kowase, K., Arai, M.and
Kurabayashi, M. (2009). Notch signaling induces osteogenic differentiation and
mineralization of vascular smooth muscle cells: role of Msx2 gene induction via Notch-
RBP-Jk signaling. Arterioscler. Thromb. Vasc. Biol. 29, 1104-1111.
Slater, B. J., Lenton, K. A., Kwan, M. D., Gupta, D. M., Wan, D. C. and Longaker,
M. T. (2008). Cranial sutures: a brief review. Plast. Reconstr. Surg. 121, 170e-178e.
Sood, S., Eldadah, Z. A., Krause, W. L., McIntosh, I. and Dietz, H. C. (1996).
Mutation in fibrillin-1 and the Marfanoid-craniosynostosis (Shprintzen-Goldberg)
syndrome. Nat. Genet. 12, 209-211.
Soriano, P. (1999). Generalized lacZ expression with the ROSA26 Cre reporter strain.
Nat. Genet. 21, 70-71.
105
Stanczyk, F. Z., Cho, M. M., Endres, D. B., Morrison, J. L., Patel, S. and Paulson, R.
J. (2003). Limitations of direct estradiol and testosterone immunoassay kits. Steroids 68,
1173-1178.
Stocco, C. (2008). Aromatase expression in the ovary: hormonal and molecular
regulation. Steroids. 73, 473-487.
Stump, G., Durrer, A., Klein, A. L., Lütolf, S., Suter, U. and Taylor, V. (2002).
Notch1 and its ligands Delta-like and Jagged are expressed and active in distinct cell
populations in the postnatal mouse brain. Mech. Dev. 114, 153-159.
Suh, J. H., Lee, H. W., Lee, J. W. and Kim, J. B. (2008). Hes1 stimulates
transcriptional activity of Runx2 by increasing protein stabilization during osteoblast
differentiation. Biochem. Biophys. Res. Commun. 367, 97-102.
Takasaki, A., Tamura, H., Taniguchi, K., Asada, H., Taketani, T., Matsuoka, A.,
Yamagata, Y., Shimamura, K., Morioka, H. and Sugino, N. (2009). Luteal blood flow
and luteal function. J. Ovarian Res. 2, 1.
Takizawa, T., Ochiai, W., Nakashima, K. and Taga, T. (2003). Enhanced gene
activation by Notch and BMP signaling cross-talk. Nucleic. Acids Res. 31, 5723-5731
Tepass, U., Godt, D. and Winklbauer, R. (2002). Cell sorting in animal development:
signalling and adhesive mechanisms in the formation of tissue boundaries. Curr. Opin.
Genet. Dev. 12, 572-582.
Tezuka, K., Yasuda, M., Watanabe, N., Morimura, N., Kuroda, K., Miyatani, S. and
Hozumi, N. (2002). Stimulation of osteoblastic cell differentiation by Notch. J. Bone.
Miner. Res. 17, 231-239.
Thisse, B., Stoetzel, C., Gorosotiza-Thisse, C. and Perrin-Schmidt, F. (1988).
Sequence of the twist gene and nuclear localization of its protein in endomesodermal
cells of early Drosophila embryos. EMBO J. 7, 2175–2183.
Ting, M. C., Wu, N. L., Roybal, P. G., Sun, J., Liu, L., Yen, Y. and Maxson R. E. Jr.
(2009). EphA4 as an effector of Twist1 in the guidance of osteogenic precursor cells
during calvarial bone growth and in craniosynostosis. Development 136, 855-864.
Twigg, S. R., Kan, R., Babbs, C., Bochukova, E. G., Robertson, S. P., Wall, S. A.,
Morriss-Kay, G. M. and Wilkie, A. O. (2004). Mutations of ephrin-B1 (EFNB1), a
marker of tissue boundary formation, cause craniofrontonasal syndrome. Proc. Natl.
Acad. Sci. U.S.A. 101, 8652-8657.
106
Vargesson, N., Patel, K., Lewis, J. and Tickle, C. (1998). Expression patterns of
Notch1, Serrate1, Serrate2 and Delta1 in tissues of the developing chick limb. Mech.
Dev. 77, 197-199.
Vernon, A. E. and LaBonne, C. (2004). Tumor metastasis: a new twist on epithelial-
mesenchymal transitions. Curr. Biol. 14, R719-721.
Warman, M. L., Mulliken, J. B., Hayward, P. G. and Müller, U. (1993). Newly
recognized autosomal dominant disorder with craniosynostosis. Am. J. Med. Genet. 46,
444-449.
Wilkie, A. O. (1997). Craniosynostosis: genes and mechanisms. Hum. Mol. Genet. 6,
1647-1656.
Wood, G. A., Fata, J. E., Watson, K. L. and Khokha, R. (2007). Circulating hormones
and estrous stage predict cellular and stromal remodeling in murine uterus. Reproduction
133, 1035-1044.
Xu, X., Wagner, K. U., Larson, D., Weaver, Z., Li, C., Ried, T., Hennighausen, L.,
Wynshaw-Boris, A. and Deng, C. X. (1999). Conditional mutation of Brca1 in
mammary epithelial cells results in blunted ductal morphogenesis and tumour formation.
Nat. Genet. 22, 37-43.
Xue, Y., Gao, X., Lindsell, C. E., Norton, C. R., Chang, B., Hicks, C., Gendron-
Maguire, M., Rand, E. B., Weinmaster, G. and Gridley, T. (1999). Embryonic
lethality and vascular defects in mice lacking the Notch ligand Jagged1. Hum. Mol.
Genet. 8, 723-730.
Yoshida, T., Phylactou, L. A., Uney, J. B., Ishikawa, I., Eto, K. and Iseki S. (2005).
Twist is required for establishment of the mouse coronal suture. J. Anat. 206, 437-444.
Yoshida, T., Vivatbutsiri, P., Morriss-Kay, G., Saga, Y. and Iseki, S. (2008). Cell
lineage in mammalian craniofacial mesenchyme. Mech. Dev. 125, 797-808.
Yousfi, M., Lasmoles, F. and Marie, P. J. (2002). TWIST inactivation reduces
CBFA1/RUNX2 expression and DNA binding to the osteocalcin promoter in osteoblasts.
Biochem. Biophys. Res. Commun. 297, 641-644.
Yu, H. M., Jerchow, B., Sheu, T. J., Liu, B., Costantini, F., Puzas, J. E., Birchmeier,
W. and Hsu, W. (2005). The role of Axin2 in calvarial morphogenesis and
craniosynostosis. Development 132, 1995-2005.
107
Yu, K., Xu, J., Liu, Z., Sosic, D., Shao, J., Olson, E. N., Towler, D. A. and Ornitz, D.
M. (2003). Conditional inactivation of FGF receptor 2 reveals an essential role for FGF
signaling in the regulation of osteoblast function and bone growth. Development 130,
3063-3074.
Yuan, Z. R., Kobayashi, N. and Kohsaka, T. (2006). Human Jagged 1 mutants cause
liver defect in Alagille syndrome by overexpression of hepatocyte growth factor. J. Mol.
Biol. 356, 559-568.
Zhang, X., Kuroda, S., Carpenter, D., Nishimura, I., Soo, C., Moats, R., Iida, K.,
Wisner, E., Hu, F. Y., Miao, S., et al. (2002). Craniosynostosis in transgenic mice
overexpressing Nell-1. J. Clin. Invest. 110, 861-870.
Zhang, Y., Kiel, D. P., Kreger, B. E., Cupples, L. A., Ellison, R. C., Dorgan, J. F.,
Schatzkin, A., Levy, D. and Felson, D. T. (1997). Bone mass and the risk of breast
cancer among postmenopausal women. N. Engl. J. Med. 336, 611-617.
108
Appendix
1. ALP section staining
• Air-diyed frozen sections are fixed in cold Acetone at 4°C for 10 min.
• 1x TBST 3min. x3
• NTMT 3min. x3
• Develop color with staining solution at RT for several min.
• TBST wash 3min. x3
• Nuclear fast red 1min.
• Rinse by ddH2O
• Dehydrate in 75% EtOH, 95% EtOH, 100% EtOH x2, Xylen x3
• Mount with mounting medium
*NTMT (add Tween20 freshly)
100mM NaCl, 100mM Tris-HCl pH9.5, 50mM MgCl2, 0.1% Tween20.
*Staining solution
NTMT lml add NBT(Roche) 5 μl and BCIP(Roche) 3.75 μl.
*10x TBST 100ml (add 0.1% Tween20 freshly)
8g NaCl, 0.2g KCl, 3g Tris, adjust pH to 7.6.
*Nuclear fast red
Kernechtrot (nuclear fast red) 0.2g + aluminium sulfate (Al2(SO4)318H2O) 10g + H2O
200 ml. Heat with stirring till near boiling and cool down O/N. Filter.
109
2. ALP whole-mount staining
• E14.5 Embryo heads were fixed in 4% paraformaldyde (PFA) O/N
• 1x PBS 10min. x3
• Store at 70% EtOH 4°C for 2 days.
• Bisect sagitally and remove brain and skin
• 1x TBST 5min. x2 on ice
• NTMT 5min. x2 on ice
• ½xALP staining solution on ice for 30 to 40min. Monitored under a microscope.
• 1mM EDTA in PBST 4°C
110
3. LacZ section staining
• Sections were fixed in 0.2% glutaraldehyde/ 2mM MgCl2/ PBS 10min. 4°C
• Rince with PBS/2mM MgCl2
• Wash with PBS/2mM MgCl2 10min on ice
• Wash with PBS/2mM MgCl2 / 0.01% DOC/ 0.02% NP-4010min RT
• Stain with X-gal solution 37°C O/N
• Wash with PBS 2min. x3
• Post fix with 4%PFA 1-2 hrs.
• Wash with PBS 10min. x3
• Nuclear fast red 1min.
• Dehydrate in 75% EtOH, 95% EtOH, 100% EtOH x2, Xylen x3
• Mount with mounting medium
*X-gal solution 10ml
1M MgCl2 20 μl, 5% DOC 20 μl, 2% NP-40 100 μl, 2% X-gal (in DMSO) 500 μl,
potassium ferricyanide (FeIII) 0.0825g, potassium ferrocyanide (FeII) 0.105g in 1x PBS
111
4. Tissue preparation (OCT embedding)
• After dissecting, embryo heads are fixed with 4%PFA for 20min.
• Wash with PBS 10min. x3
• 10% sucrose in PBS. Shake on ice till heads sink to the bottom
• 30% sucrose in PBS 4°C O/N
• Embed in OCT medium (HistoPrep, Fisher Scientific, SH75-125D) on dry ice
• Store at -80°C
112
5. Immunoperoxidase staining
• Air-diyed frozen sections are circled with a PAP pen (Zymed) and fixed in 4%
PFA 10min. 4°C
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Slides are treated with Peroxo-block (Zymed 00-2015) for 45sec. for the
elimination of endogenous peroxidase activity
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Blocking solution
Histostain-SP Kit (Rabbit, Bulk), Zymed, #95-6143-B for rabit primary
antibodies
10% Normal Rabbit Serum, Zymed, # 50-061Z for goat primary antibodies
RT 1-2 hrs.
• Primary antibody (dilute in blocking solution) O/N at 4°C
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Biotinylated second antibody from rabbit kit or Rabbit anti-Goat IgG-Biotin
(Zymed, #50-232Zw) RT for 20min.
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Enzyme conjugate solution RT 20min.
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Add DAB substrate mix (Zymed #2020) RT 1-5min. Monitored under a
microscope.
• Wash with dH2O x1, dH2O with 0.1% Tween20 x1
• Hematoxylin 1min.
• Wash with dH2O and put in PBS 20sec.
• Rinse with dH2O
• Dehydrate in 75% EtOH x1, 95% EtOH x1, 100% EtOH x2, Xylen x3
• Mount with mounting medium
113
6. Immunofluorescence staining
• Air-diyed frozen sections are circled with a PAP pen (Zymed) and fixed with 4%
PFA 10min. 4°C
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Blocking solution (Histostain-SP Kit (Rabbit, Bulk), Zymed, #95-6143-B) RT 1-2
hrs.
• Primary antibody (dilute in blocking solution) O/N at 4°C
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Rhodamin Red-X goat anti-rabbit IgG secondary antibody RT 1hr.
• 1x PBS 2min. x2, 1xPBST 2min. x1
• 10 μg/ml DAPI in PBST 1min.
• 1x PBS 2min. x2, 1xPBST 2min. x1
• Mount with ProLong® Gold antifade reagent (Invitrogen)
114
7. Section in situ hybridization (from Eva Chan)
• Air-dried frozen sections are fixed with 4% PFA RT 30 min.
• Wash with DEPC-PBS RT 3min. x3
• Rinse with DEPC-ddH2O 3min. x1
• Pre-hybridize with pre-hybridization solution (Biochain Institute Inc. K2191020)
50°C 3-4hrs
• Heat RNA probe in 95°C 3min. and put it in cooler box
• Add pre-warmed hybridization solution (Biochain Institute Inc.) to RNA probe
(0.4μg/100μl)
• Apply RNA probe mix on slides and cover with Parafilms
• 50°C O/N
• Wash with 2x SSC 50°C 20min. x2
• 1.5x SSC 50°C 20min. x2
• 0.5x SSC 50°C 20min. x1
• Cool down the slides to RT
• Wash with RNase buffer 37°C 15min x1
• Incubate with 10μg/ml RNase in RNase buffer 37°C 30min.
• Wash with 0.2x SSC 37°C 20min. x2
• Wash with 1x TBS RT 3min. x2
• Block with blocking solution RT 2hrs (change new solution every 15min.)
• Incubate with AP-conjugated anti-DIG antibody (Roche) (1:500 in blocking
solution) 4°C O/N
• Wash with 1x TBS/0.5mg/ml levamisole RT 10min. x 6
• Wash with 1x AP buffer (Biochain Institute Inc.) in TBS/0.5mg/ml levamisole RT
30min. x 2
• Incubate with NBT/BCIP in TBS/0.5mg/ml levamisole 4°C O/N
• Stop the substrate reaction by washing with dH2O and stop buffer 10min
• Mount in glycerin mounting media
115
*Stop buffer
5mM EDTA, 20mM Tris, pH8.0
*Glycerin mounting media
10g gelatin + 60ml dH2O, heat to dissolve. Add 70 ml glycerol and 1ml phenol. Heat
with stiring. Aliquot and store at RT. Heat (65°C) before use.
116
8. RNA probe
• 20 μg of linearized plasmid for 1:1 pheno/chroloform extraction
• 2.5 V cold EtOH in -80°C 15min.
• Pellet dissolve in 10 μl DEPC ddH2O.
• UV specc
• mMessage kit (Ambion)
DEPC ddH2O 24 μl + DNA template 4 μl + 10x buffer 4 μl +
10x DIG labeling Mix 4 μl + RNA polymerase 4 μl
• 37°C O/N
• Add 1 μl DNase 37°C 15min.
• Add 60 μl DEPC ddH2O + 50 μl 7.5M LiCl + 2 μl 20mg/ml glycogen
• -20°C O/N
• centrifuge 12000rpm 10min. 4°C
• air dry 10min
• Pellet dissolve in 20 μl DEPC ddH2O
• UV specc
Abstract (if available)
Abstract
In this dissertation, I would like to address how the evolutionarily conserved Notch signaling molecules regulate the formations of murine cranial bones and sutures. Mutations in the Notch ligand, JAGGED1, cause Alagille syndrome, which has craniosynostosis as a feature. The expression pattern of Jagged1 at mouse coronal suture suggested that it might play a role in establishing boundary between osteogenic and non-osteogenic cells. Tissue-specific knockout of Jagged1 in mouse mesoderm affected the expression of downstream Notch signaling at sutural cells and resulted in craniosynostosis. Immunostaining results also implied that the boundary between presumptive cranial bones has been established by Notch signaling at early stage, while the opposing osteogenic fronts of the bones are still far away from each other. I further demonstrated the genetic interactions between Notch signaling and Twist1 which is animportant pathogenic gene in regulating cranial sutures morphogenesis. Twist1 regulates Notch signaling in sutural mesenchyme and maintain suture patency. The phenotypic studies of mouse skulls and middle ear ossicles indicated that Jagged1 interacts functionally with Twist1 in several distinct developmental settings. This work reveals a molecular network that controls cranial development, and establishes a new model of boundary formation at developing cranial suture.
Linked assets
University of Southern California Dissertations and Theses
Conceptually similar
PDF
TWIST1 functions in both mesoderm and neural crest derived cranial tissues to establish and maintain coronal suture patency
PDF
Utilizing zebrafish and mouse models to uncover the underlying genetics of human craniofacial anomalies
PDF
Lineage boundaries and cell migration in the patterning of the mammalian skull
PDF
Elucidating the role of neural crest specific Stat3 signaling in maintaining coronal suture patency during embryonic development
PDF
Homologous cell systems for the study of progression of androgen-dependent prostate cancer to castration-resistant prostate cancer
PDF
Role of cancer-associated fibroblast secreted annexin A1 in generation and maintenance of prostate cancer stem cells
Asset Metadata
Creator
Yen, Hai-Yun
(author)
Core Title
Jagged1 functions downstream of Twist1 in the specification of the coronal suture and the formation of a boundary between osteogenic and non-osteogenic cells
School
Keck School of Medicine
Degree
Doctor of Philosophy
Degree Program
Biochemistry and Molecular Biology
Degree Conferral Date
2009-12
Publication Date
11/20/2010
Defense Date
10/27/2009
Publisher
University of Southern California
(original),
University of Southern California. Libraries
(digital)
Tag
boundary formation,cranial suture,Jagged1,OAI-PMH Harvest,TWIST1
Language
English
Contributor
Electronically uploaded by the author
(provenance)
Advisor
Maxson, Robert E. (
committee chair
), Crump, Gage (
committee member
), Dubeau, Louis (
committee member
), Roy-Burman, Pradip (
committee member
)
Creator Email
haiyunyen@yahoo.com,hyen@usc.edu
Permanent Link (DOI)
https://doi.org/10.25549/usctheses-m2750
Unique identifier
UC1492578
Identifier
etd-YEN-3384 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-278546 (legacy record id),usctheses-m2750 (legacy record id)
Legacy Identifier
etd-YEN-3384.pdf
Dmrecord
278546
Document Type
Dissertation
Rights
Yen, Hai-Yun
Type
texts
Source
University of Southern California
(contributing entity),
University of Southern California Dissertations and Theses
(collection)
Repository Name
Libraries, University of Southern California
Repository Location
Los Angeles, California
Repository Email
cisadmin@lib.usc.edu
Tags
boundary formation
cranial suture
Jagged1
TWIST1