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Expression Analysis of CTHRC1 in the Murine Embryo during
Mid-facial Development
by
Dr. Laurene Dao-Pei Yen
A thesis submitted in conformity with the requirements
for the degree of Masters of Science in Dentistry (Orthodontics)
Department of Orthodontics
University of Toronto
© Copyright by Laurene Dao-Pei Yen, 2014
ii
Expression Analysis of CTHRC1 in the Murine Embryo during Mid-
facial Development
Dr. Laurene Dao-Pei Yen
Masters of Science in Dentistry (Orthodontics)
Department of Orthodontics
University of Toronto
2014
Abstract
Collagen Triple Helix Repeat Containing 1 (CTHRC1) has been shown to regulate collagen
expression, cell migration and interact with the Wnt/planar cell polarity pathway. Objective:
Determine CTHRC1 expression in the midface of mouse embryos during craniofacial
development. Methods: qPCR and immunohistochemistry were used to determine the
expression of Cthrc1 at embryonic (E) days of development E8.5-E13.5. Results: CTHRC1
expression was in the notochord and neural tube at E8.5, mesenchyme of the midface at E9.5-
E10.5 and areas of cartilage formation at E11.5-E13.5. Conclusions: The expression of
CTHRC1 in the developing craniofacial region suggests a role of CTHRC1 in migration of
cranial neural crest cells and in chondrocyte proliferation and differentiation. We speculate the
functions of CTHRC1 during craniofacial development are via its interactions with the Wnt/PCP
pathway, shown previously to play a significant role during craniofacial development. Support:
Donald G. Woodside Fund and NSERC.
iii
Acknowledgments
I would like to thank my supervisor, Dr. Siew-Ging Gong for your guidance and support
throughout my thesis. Thank you for the constant encouragement, especially during the writing
of my thesis.
I would also like to express my sincere thanks to the members of my committee, Dr. Bernhard
Ganss and Dr. Bryan Tompson for their scientific guidance and advice throughout my MSc
degree.
I would like to thank the Donald G. Woodside Fund for funding part of my project.
A special thanks to Mr. James Holcroft, for his help with the quantitative PCR portion of my
thesis, and Feryal Sarraf, for her technical expertise in the use of the microscopes and digital
cameras in the histology lab.
Thank you to my husband, Dr. Alexander Unterberger¸ for his constant support and
encouragement throughout my degree. I could not have done it without you!
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Table of Contents
Abstract …ii
Acknowledgments…iii
List of Figures…viii
Chapter 1: Introduction…1
1.1 Development and morphogenesis of the vertebrate face…1
1.1.1 Role of neural crest cells in craniofacial development…1
1.1.2 Formation and migration of cranial neural crest cells…2
1.1.2.1 Induction and specification of CNCCs and NCCs…2
1.1.2.2 Delamination…3
1.1.2.3 Migration…3
1.1.2.4 NCC positional identity…5
1.1.2.5 NCC differentiation and proliferation…6
1.1.3 Facial prominences and CNCC derivatives of the face…6
1.1.4 Facial development and facial prominences…7
1.2 Craniofacial developmental anomalies…10
1.3 Proteins and genes involved in formation of craniofacial structures…11
1.3.1 Fibroblast growth factors and bone morphogenetic proteins…12
1.3.2 Sonic hedgehog…13
1.3.3 Wnt pathways…14
1.3.3.1Wnt pathways and craniofacial development…17
v
1.3.3.2 Wnt5A…18
1.4 Collagen Triple Helix Repeat Containing 1(Cthrc1)…20
1.4.1 Background…20
1.4.2 CTHRC1 structure…21
1.4.3 Expression pattern of Cthrc1…22
1.4.4 Possible functions of Cthrc1…22
1.4.4.1 Cthrc1 role in collagen deposition…23
1.4.4.2 Role of Cthrc1 in osteogenesis, osteoclastic bone formation and bone
remodeling…24
1.4.4.3 Role in cell motility and tissue repair…27
1.4.5 Genes and signalling pathways involved with CTHRC1…28
1.4.5.1 Cthrc1 activation of Wnt/PCP pathway…28
1.4.5.2 Cell-specific action of Cthrc1…30
1.5 Rationale for study…31
1.6 Hypothesis/Aims…31
Chapter 2: Materials and Methods…32
2.1 Embryos…32
2.2 RT-qPCR…32
2.2.1 RNA isolation…32
2.2.2 Reverse transcription of RNA samples…33
2.2.3 Quantitative PCR…33
vi
2.3 Histological processing and Paraffin Embedding…34
2.4 Immunohistochemistry…34
2.5 Documentation and analysis of Cthrc1 expression…35
Chapter 3: Results…36
3.1 Quantitative expression of Cthrc1 mRNA transcripts during midface development…36
3.2 Expression of CTHRC1 protein…37
3.2.1 CTHRC1 peptide competition assay…37
3.2.2 Spatial expression of CTHRC1 in the midface at different developmental
stages…38
3.2.3 CTHRC1 expression in the mandible and developing tooth bud…43
Chapter 4: Discussion…45
4.1 Summary of Cthrc1 expression during mouse embryo midface development…45
4.1.1 Summary of qPCR mRNA levels of Cthrc1 mRNA during E8-5-E13.5…45
4.1.2 Summary of immunolocalization experiment results of CTHRC1 protein from E8.5-
E13.5…46
4.2 Involvement of Cthrc1 in midface development…47
4.2.1 Role of Cthrc1 in cell migration…47
4.2.2 Role of Cthrc1 in epithelial/mesenchyme interactions…48
4.2.3 Role of Cthrc1 in regulation of collagen formation and deposition…50
4.2.4 Role of Cthrc1 in bone formation…53
4.2.5 Summary of roles of Cthrc1 in midface development…55
vii
4.3 Future Directions for Cthrc1 studies…56
4.3.1 Midface development in Cthrc1 knockout models…56
4.3.2 Cthrc1 expression in mouse models specifically targeting effects on midface
development…57
4.3.3 Colocalization studies of CTHRC1 with other proteins with a known function in
midface development…57
4.4 Conclusions…58
References…59
Copyright Acknowledgements…68
viii
List of Figures
Figure 1: Migration and skeletal fates of the CNCCs…4
Figure 2: A schematic diagram of the pharyngeal arches…7
Figure 3: The canonical Wnt signaling pathway…15
Figure 4: The non-canonical Wnt signaling pathway…17
Figure 5: Summary of WNT5A functions during cartilage development and disease….20
Figure 6: The structure of the CTHRC1 protein…21
Figure 7: A model of selective activation of the Wnt/PCP pathway by CTHRC1…30
Figure 8: Temporal expression profile of Cthrc1 mRNA across E8.5-E13.5…36
Figure 9: Peptide competition assay on consecutive coronal sections through the anterior part of
the midface in an E13.5 embryo…37
Figure 10: CTHRC1 peptide expression in E8.5…38
Figure 11: CTHRC1 protein expression in E10.5 embryos…39
Figure 12: CTHRC1 protein expression in E11.5…40
Figure 13: CTHRC1 protein expression at E12.5…41
Figure 14: CTHRC1 protein expression at E13.5…42
Figure 15: CTHRC1 protein expression in tooth bud formation…44
1
Chapter 1
Introduction
Development of the craniofacial region is a complex process. It is regulated by different genes,
proteins and signalling pathways. In this chapter, the development and morphogenesis of the
vertebrate face and the involvement of neural crest cells will be discussed. These topics will be
followed by the sequence of formation of the facial structures and a discussion of craniofacial
developmental anomalies.
1.1 Development and morphogenesis of the vertebrate face
1.1.1 Role of neural crest cells in craniofacial development
Development and morphogenesis of the vertebrate face is a complex and highly orchestrated
process. A major population of cells that contribute to the craniofacial region is the neural crest
cells (NCCs) that form along the entire length of the developing embryo. NCCs are transient
migratory cells that develop at the border between the neural plate (NP) and the epidermis. NCCs
delaminate from their site of origin along the dorsal neural tube and migrate ventrally along
different pathways. NCCs contribute to many different cell types and tissues in the body, such as
the enteric nervous system, melanocytes, connective tissues, and myofibroblasts which line
blood vessels, neurons and glia of the peripheral nervous system, pigment cells, endocrine cells,
cardiac structures, smooth muscle cells and tendons (1). NCCs in the rostral region of the
developing embryo are known as cranial neural crest cells (CNCCs). In contrast to trunk NCCs,
CNCCs can give rise to osteoblasts and chondrocytes (2) and are the major contributor to
structures of the facial skeleton, cartilages and bones of the jaws, middle ear and neck, cranial
nerves, smooth muscles, dermis and facial connective tissues (3).
During the migration of the CNCCs and after their arrival, their interactions with the surrounding
ectoderm, endoderm and neuroectoderm are integral for normal development of the face (4-6).
This multi-step process is regulated both temporally and spatially and is made up of the
coordinated action of numerous genes and signalling pathways (7). Due to this high level of
regulation, craniofacial development is susceptible to perturbations, resulting in facial
dysmorphologies. Many birth defects associated with craniofacial malformations can be
attributed to the defects in the generation, proliferation, migration and differentiation of CNCCs
2
(8). Understanding the complex and sequential mechanisms of craniofacial development and
how they relate to the formation of facial structures is essential to the orthodontic specialty.
1.1.2 Formation and migration of cranial neural crest cells
The development of the NCCs and CNCCs can be divided into a number of stages:
a) Induction
b) Delamination
c) Migration
d) Differentiation
1.1.2.1 Induction and Specification of CNCCs and NCCs
Induction of NCCs is a multistep process resulting in a molecular cascade of events involved in
establishing their migratory and multipotent characteristics. The origin of NCCs can be traced to
the border of the NP in a region of ectoderm situated between the NP and the non-neural
ectoderm (NNE). Immediately beneath the ectoderm there is a layer of mesoderm, and together
with the NP and NNE, these tissues are collectively believed to contribute to the induction of the
NC. Induction of NCCs has been shown as early as during gastrulation during embryonic
development (9).
Induction involves a complex set of extracellular signals which transform the fate of cells along
the medio-lateral and anterior-posterior axes of the embryo (10). The signals that position the
NCCs along the axes are released from the NP, the epidermis and the lateral mesoderm. For
complete induction of NCCs, there must be a transformation of the naive ectoderm into the
neural crest. A precise combination of extracellular signalling molecules operate to activate the
expression of transcription factors that define the neural crest territory and control subsequent
neural crest development (11). Molecules such as wingless-type MMTV integration site
(WNTs), Notch, bone morphogenetic proteins (BMPs) and fibroblast growth factors (FGFs) are
secreted from the adjacent epidermis and underlying mesoderm and separate the NNE
(epidermis) from the NP during neural induction (12-14). The induction of the expression of
regulatory transcription factors (Msx1/2, Pax3/7, Zic1, Dlx3/5, Hairy2, Id3, Ap2) specify the NP
border that subsequently trigger the expression of NC specifiers, a second set of transcription
factors (Snail2, FoxD3, Sox9/10, Twist, cMyc, and Ap2). NC specifiers are proposed to
3
ultimately control neural crest behavior, from epithelial-mesenchymal transition (EMT) and
delamination to migration and differentiation (15).
One theory proposes that NCCs are specified during gastrulation (9) whereas the second, more
classical, theory postulates that NCCs are specified during the time of neural tube closure. After
specification, contact-mediated signalling between tissues in the dorsal neural tube results in
EMT in neural ectoderm cells at the neuroectoderm and NNE border (1).
1.1.2.2 Delamination
After induction and specification at the neuroepithelium, NCCs undergo EMT and leave their
site of origin through a delamination process. Delamination is the splitting of the NCCs from
their surrounding tissues whereas EMT is a series of events at the molecular level which
orchestrates a change from an epithelial to mesenchymal phenotype (11, 16). EMT is a multi-
step process in which the NCCs lose their apico-basal polarity and disassemble intercellular
adhesion complexes required for epithelial formation (16), allowing the NCCs to separate from
the neuroepithelium and ectoderm (11). Contact-mediated signaling between tissues in the dorsal
neural tube stimulates cells at the neural/non-neural border to undergo EMT(17). This results in a
highly invasive phenotype characteristic of NCCs, behaviour which is also shared with
metastatic cells (18).
In the cranial region, CNCCs delaminate all at once, whereas the trunk NCCs delaminate
progressively, leaving the neuroepithelium one by one after neural tube closure (19).
1.1.2.3 Migration
The CNCCs follow migratory pathways that are conserved among vertebrate species. CNCC
migration is directed along well-defined routes that end in the ventral part of the brain and
branchial arches (20). Interactions between the CNCCs and their local environment are critical in
CNCC directional and collective migration (21). A number of cellular mechanisms have been
shown to operate during the directed migration of the NCCs. NCCs become polarized in the
direction of their migration with a tail at the back of the cell and filopodia and lamellipodia at the
side towards their migration. A mechanism known as contact inhibition of locomotion (CIL)
exists when two migrating NCCs make contact and they retract their protrusions and change
direction (22, 23). The non-canonical WNT planar cell polarity (PCP) signalling pathway
4
(section 1.3.3) and cell-cell contact are crucial in controlling the polarity of migrating NCCs (8).
Even though the NCCs experience CIL when they contact each other, they also tend to migrate
in large groups, more so than what would be predicted given the CIL phenomenon. This is a
result of a phenomenon called co-attraction, which allows the collective migration of NCCs
despite low cell adhesion and dispersion due to CIL (14). NCCs have also been shown to move
around barriers introduced into the migration path and can re-target their direction (24).
The migration of CNCCs proceeds along a well-defined route, ending in the ventral part of the
brain and the pharyngeal arches (PA) (25). NCCs from the diencephalon and anterior
mesencephalon migrate into the frontonasal process (FNP) and NCCs from the posterior
mesencephalon and hindbrain migrate to the PAs (8). CNCCs in the hindbrain region are
compartmentalized in 7 distinct regions called rhombomeres (r). At first, the CNCCs migrating
to the PAs migrate as a continuous wave but then they split into three distinct segregated streams
(8). CNCCs from the posterior mesencephalon, r1 and r2 fill the first PA (PA1) and NCCs from
r4 fill pharyngeal arch 2 (PA2) (Figure 1 ) (3, ). In the post-otic hindbrain, NCCs from r6-r8
colonize PA3-6 (21). The subpopulations of NCCs that migrate to and target the PAs migrate in
stereotypical streams, maintaining a spatial segregation. This segregation has an important
impact on craniofacial patterning and the early anteroposterior patterning of NCCs by
establishing the segmental pattern in which the pharyngeal region of the vertebrate head is
formed (26, 27).
5
Figure 1: Migration and skeletal fates of the CNCCs. A) Embryo showing the colonization of the
head and pharyngeal arches by fore-, mid- and hindbrain NCCs. There are 7 segments in the
hindbrain known as rhombomeres (r) from which crest cells emigrate in three major paths to the
pharyngeal or branchial arches (BA), coded in blue, yellow and green. B) A skull drawing
showing comparative contributions of the NCC populations to cranial skeletal elements of the
human. The major bones are coded to match the origin of the contributing migratory neural crest
streams. Reproduced from Gong, 2014. (3, 25).
To allow proper positioning in the PA and the proper assembly of structures, the subpopulations
of NCCs are guided by a complex set of cues to which they respond to locally during their
migration (28). From the dorsal neural tube into the craniofacial region the NCCs follow well-
marked paths by communication with the surrounding neural, facial and pharyngeal epithelia and
cephalic mesoderm (17). Examples of signalling molecules that guide the NCCs include: FGF-2
and FGF-8, which function in a chemotactic manner (29); Ephrins, which are ligands for Eph
receptors (30); and semaphorin proteins (31).
1.1.2.4 NCC positional identity
The positional identity of NCCs is regulated by the homeodomain (HD) transcription factors of
the homeobox gene family (Hox) (32, 33). The HD expression is induced and maintained in
NCCs in later developmental stages via signals from the surrounding local environment (8).
Anteroposterior NCC positional identity is thought to be acquired pre-migration (34, 35);
however, it is not permanent and there is some degree of plasticity. For example, intrinsic
molecular programs of the NCCs can be switched to new programs if exposed to ectopic
environmental cues (34, 36, 37). The involvement of the Hox genes was first shown in a mouse
model where targeted inactivation of Hox2a resulted in homeotic transformation of PA2
elements into PA1-like skeletal elements (38). Previous experiments by Santagati et al. (28) and
Pasqualetti et al. (39) in Xenopus demonstrated that the skeletal pattern of mandibular and hyoid
crest is not irreversibly committed before migration of NCC. However, positional information
has to be maintained throughout the post-migratory states in order to provide information about
size, shape and orientation of PA2 skeletal elements.
6
Dorso-ventral axis patterning information is regulated by the NCCs through the Dlx homeobox
code. The involvement of Dlx genes has been investigated mainly through loss of function
mutations in the mouse (40). Namely, the partitioning of PA1 is achieved with the two Dlx
combinations; Dlx1/2 for the maxilla and Dlx 1/2/5/6 for the mandibular process. Mutations of
Dlx1 or Dlx2 primarily affect the maxilla and mutations of Dlx 5 affect the mandibular process
(41-43).
It is clear the diversity of NCC-derived skeletal elements among vertebrates arises from
modification of the levels, timing or spatial expression of HD factors. These factors play a
crucial role in spatial identity and modifying the expression of pattern genes can result in
aberrant development of structures (8).
1.1.2.5 NCC differentiation and proliferation
Once the NCCs have arrived in the facial prominences, they begin to proliferate and form the
facial structures. Many factors regulate the proliferation of NCCs, such as the Sonic hedgehog
(SHH) protein and WNT ligands. Disruption of these factors can affect the process of
craniofacial morphogenesis and cause aberrant proliferation and morphogenesis of NCCs (4).
There are two theories as to how NCCs determine which structures to make once they migrate to
their final destination. The first theory is that the NCCs have an intrinsic program that is
activated once they leave the neural tube. This intrinsic program contains information for
molecular patterning for the formation of facial structures. The second theory is that the NCCs
acquire facial patterning information from the surrounding tissue at their destination (44). The
dominant theory is that of the latter theory, that the NCCs retain multipotency into the later
stages of embryonic development (1).
1.1.3 Facial prominences and CNCC derivatives of the face
During the migration towards their final destinations ventrally, extensive interactions occur
between the CNCCs and the surrounding mesoderm and overlying ectoderm and endoderm. The
CNCCs in each facial prominence have distinctive molecular signatures and interact with
different epithelia during development (1).
By the end of fourth week of human embryonic development, most of the CNCCs have reached
their final destinations. Within the pharyngeal arch is a center mass of mesodermal cells,
7
surrounded by CNCCs and externally covered by ectoderm and internally bordered by endoderm
(Figure 2) (45). Continued interactions between surface ectoderm, CNCCs, mesoderm and
pharyngeal endoderm are needed for the proper development of the facial structures (46, 47). For
example, coordinated interactions between CNCCs and the mesoderm cells result in the
differentiation of myoblast and skeletal precursors to form the musculature and skeleton of the
facial region (48, 49). Signals that emanate from the CNCCs instruct and inform mesodermal
cells to differentiate into myoblast precursors and how to organize themselves around the
developing skeletal elements.
1.1.4 Facial development and facial prominences
The major portion of the face in the human is formed starting between the fourth and eighth
weeks of development. The face is derived from buds of tissue—called facial prominences—that
undergo symmetrical and asymmetrical growth. Appropriate fusion events are required to result
in mature facial structures (50). These facial primordia appear in the fourth week of development
around the primordial stomodeum through inductive influences of organizing centers located in
the prosencephalon and rhombencephalon (51).
Figure 2: A schematic diagram of the pharyngeal arches. (A) The arches are indicated by
different colors. The first arch is divided into maxillary (1a, orange) and mandibular (1b, yellow)
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component. The second arch (2, green); third arch (3, purple); fourth arch (4, pink) and the sixth
arch (6, blue). B. A coronal cut through the human embryo showing the tissue contribution to
each arch. The arches (1,2,3,4) are composed of a core of neural crest (nc, yellow) and
mesoderm (mes, green) surrounded by both surface ectoderm (se, pink) and pharyngeal
endoderm (pe, purple). Reproduced from Cordero et al. 2010 (1).
At about the 4th
to 5th
week range (E9.5 in mice) of human development, CNCCs are migrating
and arriving at the facial prominences (52). A total of seven prominences in the face will
eventually be present ventral to the forebrain the FNP, two lateral nasal processes (LNP), two
maxillary processes and two mandibular processes. The FNP surrounds the ventrolateral part of
the forebrain which give rise to the optic vesicles of the eyes and also will give rise to the
forehead, middle of the nose, upper lip, philtrum and primary palate (53). The lateral region of
the FNP, also known as the median nasal process (MNP), will fuse with the LNPs and the
maxillary processes to create the alae and columnellae of the nose. Initially, around the 5th
week
of embryonic development, a pair of placodes form ventral to the developing forebrain. Further
growth and proliferation of the placodes lead to a deepening, resulting in the formation of nasal
pits which are surrounded by the lateral nasal process (LNP) and medial nasal processes (MNP)
(51). These nasal placodes are bilateral oval thickenings of the surface ectoderm and develop on
the inferolateral parts of the FNP. Mesenchyme in the margins of the placodes proliferates to
produce horseshoe shaped elevations known as the MNP and LNP (Figure 2).
The maxillary and mandibular processes are derivatives of the first pharyngeal arch and give rise
to the upper and lower jaws, respectively. The maxillary processes enlarge and grow medially
towards each other due to proliferation of the mesenchyme within the processes. This causes the
MNPs to move towards the median plane and each other. The LNPs are separated from the
maxillary processes by the cleft of the nasolacrimal groove. Merging of the LNP with the
maxillary process starts around the 6th
week of development. Between the 7th
and 10th
weeks of
development the MNPs merge with each other and with the maxillary and lateral nasal processes
resulting in a continuity of the upper jaw, lip and separation of the nasal pits from the
stomadeum. When the MNPs merge they form the intermaxillary segment which is made up of
the philtrum of the upper lip, the premaxillary part of the maxilla and the primary palate (55).
The lateral parts of the upper lip, most of the maxilla and secondary palate arises from the
maxillary processes (51). The secondary palate begins to form in the early 6th
week from the
9
mesenchymal projections of the maxillary processes. The palatal shelves are outgrowths of
NCC-derived mesenchyme covered by a layer of oral and surface epithelia on their outer surface.
Within the palatal shelves is NCC derived mesenchyme. For the palate to form, the maxillary
processes expand and undergo rotation to a horizontal position. The palatal shelves must have
outgrowth in order to fuse. As the jaws grow, the tongue becomes relatively smaller and moves
inferiorly, allowing the lateral palatine processes to elongate and ascend to a horizontal position.
These palatal processes fuse in the median plane as well as with the nasal septum and primary
palate (55).
The maxillary processes merge with the mandibular prominences and the mesenchyme from the
second pair of PA invades the primitive lips and cheeks and will later differentiate into the facial
muscles of expression which are supplied by the facial nerve, a nerve of the second PA. The
mesenchyme in the first PA differentiates into the muscles of mastication which are innervated
by the trigeminal nerve from the first PA.
At the 9th
-12th
week of development the nasal septum develops in a downward growth fashion
from the fused MNPs and begins fusing with the palatine processes. Bone will gradually develop
in the primary palate and extends from the maxillae and palatine bones into the lateral palatine
processes to form the hard palate. In the midfacial region, NCCs condense within the single
median frontonasal process to form the pre-cartilaginous nasal capsule, which undergoes
chondrogenesis to form the nasal septum. The nasal septum has been proposed to play the role of
pacemaker or growth center for the subsequent growth of the face and the skull. The nasal
septum contributes to the overall changes in morphology observed during the development of the
facial skeleton (56). To date, the fundamental mechanisms involved in the initiation, growth,
boundary setting and differentiation of the cartilaginous and skeletal structures of the frontonasal
region are not well characterized.
Tooth formation also occurs around this stage. At 37 days of embryonic development in humans,
a thickened band of epithelium forms in the upper and lower jaws around the mouth. The
primary epithelial bands give rise to the dental lamina and the vestibular lamina. Localized
thickening of the epithelium, also known as placodes, form within the epithelial bands (57).
Beneath the epithelium lies an embryonic connective tissue called ectomesenchyme into which
NCCs have migrated. Epithelial outgrowths descend into the ectomesenchyme at the sites of
10
deciduous teeth and ectomesenchymal cells proliferate and accumulate around these outgrowths.
Within the first twelve days of development in mice, the epithelium possesses the potential to
induce tooth formation but after 12 days of development this potential is assumed by the
ectomesenchyme such that the ectomesenchyme can elicit tooth formation form different
epithelia. Experiments have shown that first arch ectomesenchyme combined with foot
epithelium can induce enamel organ formation from the plantar epitheliulm (58). If the epithelial
enamel organ is recombined with skin mesenchyme, the organ loses its dental characteristics and
assumes those of the epidermis. These experiments indicate that odontogenesis is initiated by
factors from the epithelium initially and then later the ectomesenchyme.
1.2 Craniofacial developmental anomalies
Due to the highly regulated nature of growth and fusion of the facial processes, craniofacial
development is extremely susceptible to genetic and environmental perturbation that results in
craniofacial malformations. There are many opportunities for congenital facial anomalies to
occur. Anomalies result mainly from maldevelopment of the neural crest tissue which give rise
to the connective tissue and skeletal primordia of the face (51).
Facial dysmorphologies are a component of 75% of birth defects (7). Additionally, perturbances
in the critical and highly regulated roles of CNCC specification, delamination, migration and
differentiation during craniofacial development is best illustrated when abnormalities occur in
craniofacial birth defects such as Treacher Collins syndrome, cleft lip and palate, 22q11.2
microdeletion syndrome and CHARGE syndromes.
One of the most common craniofacial malformations is clefting of the lip and palate and clefting
of the secondary palate alone. Cleft of the primary lip and palate has an incidence of 1/700 and
clefting of the secondary palate alone has an incidence of 1/1500. Environmental factors such as
a large tongue or ankylosis can cause clefting of the palate because the tongue will not descend,
preventing the palatal shelves from fusing (1). Genetic factors, such as mutations in transcription
factors like TBX 22 (59) and the gene Wntb9, play a role in secondary palatal clefting (60). In
mammals, Wnt signalling is critical for the proliferation of CNCCs within the maxillary
processes (61). Transforming growth factor (TGF-α and TGF-β) and epithelial growth factor
receptor (EGFR) play a role in the fusion process of the epithelial cells on the palatal processes
of the secondary palate. Egfr-/- mice have midline defects with an elongated primary palate,
11
clefting of the palate and micrognathia (62). Any disruption in the timing or extent of growth of
the facial prominences can result in facial malformations.
Treacher Collins syndrome is an example of abnormal regulation and proliferation of CNCCs. It
is characterized by downward slanting eyes, micrognathia, conductive hearing loss,
underdeveloped zygoma, drooping part of the lateral lower eyelids, and malformed or absent
ears. The syndrome is caused by a mutation in TCOF1, which encodes the Treacle protein. A
decrease in the Treacle protein results in a significant reduction in the number CNCCs, due to
apoptosis and a decrease in migration and proliferation (63), and results in severe craniofacial
hyperplasia and dysplasia.
In summary, early development of the craniofacial region starts with the highly regulated process
of CNCC development and their eventual contribution to the major structures of the craniofacial
region. During the formation of the CNCCs, interactions between the surrounding surface
ectoderm, neuroectoderm and endoderm are crucial for normal development of the face (1) and
under strict molecular regulation. Many genes have been characterized to be significant players
in the regulation of CNCC development and morphogenesis.
1.3 Proteins and genes involved in formation of craniofacial structures
A number of genes, proteins and specific signalling pathways have been shown to be involved at
different stages of craniofacial formation. During early craniofacial development, epithelial-
mesenchymal interactions are especially crucial in determining the types of tissues and extent of
outgrowth of the different structures. Patterning and differentiation events later in development
are also regulated by a series of molecules with specific functions. A review of the major
signalling pathways and specific molecules with known roles in regulating some of the major
events in craniofacial development will be discussed. For example, members of the BMP family,
Wnt/β-catenin pathways and FGFs are necessary for NCC generation and survival (64-66). In the
final part of this section, a description of Cthrc1, the focus of the current study, will be
conducted.
Early on, the prosencephalic organizing center, located at the rostral end of the notochord
beneath the forebrain, induces the visual, inner ear apparatuses and the upper third of the face.
12
The rhombencephalic center induces the middle and lower thirds of the face (52). The forebrain
establishes many signalling centres in the ectoderm that covers the frontonasal zone. Interactions
between CNCCs and epithelia from the forebrain and facial ectoderm are required for proper
development of the FNP (67). The forebrain provides SHH signalling which imprints on the
forebrain ectoderm, controlling differential cell proliferation, and transforming the FNP into the
nasal region. The nasal placodes provide morphogenetic information to the mesenchyme of the
lateral nasal prominence and act as a signalling center for the lateral nasal skeleton, inducing
cartilage and bone (68).
Many genes involved in signal transductions, transcription regulation, binding and catalytic
activities have been identified in the developing first PA. Examples are Msx-1 and 2, Pax8,
Bmp4, 5 and 7, Fgfr11 and Tgf-β. Reciprocal epithelial and mesenchymal signalling events in the
first PA induce transcription factors that function to differentiate oral versus aboral and
rostrocaudal surfaces and patterning of skeletal elements (68). An example of the signalling
between the endoderm, mesoderm and epithelium is that of TBX1, a T-box transcription factor.
Tbx1 is expressed in the endoderm, mesoderm and epithelium of the palatal shelves and FNP
(69). Decreased Tbx1 expression in the endoderm in PA development and epithelia of the palatal
shelves results in secondary NCC abnormalities (70). Tbx1 knockout mice show defects in the
PAs and pouches (70).
The secreted peptide, Endothelin I, signals from the facial epithelia and mesoderm to the
intervening CNCCs that form the PA1 skeleton. Mice with a homozygous mutation in EdnI have
malformed bones of the jaw and throat and defective musculature (71). Recent genetic studies in
humans have indicated that mutations in the Endothelin pathway may play a role in
Waardenburg-Shah syndrome and Hirschprung disease which are caused by defects in NCC
behaviour.
Epithelial-mesenchymal interactions have been shown to play a key role in tooth development.
Examples of genes that are expressed in the NCC-derived ectomesenchyme which initiate tooth
formation are Lim-homeobox genes, Lhx-6 and Lhx-7. It has been shown that Fgf8 signalling
from the epithelium induces the expression of the Lim-homeobox genes in the mesenchyme.
Fgf8 from the epithelium also induces Pax-9 expression in the mesenchyme (57).
13
1.3.1 Fibroblast growth factors and bone morphogenetic proteins
FGFs and BMPs have many roles during craniofacial and pharyngeal skeletal morphogenesis.
The roles of FGF include serving as survival factors for CNCCs (72, 73), directing CNCCs to
adopt the ectomesenchymal fate (74), acting as chemoattractants to promote lateral migration of
endodermal cells for segmentation of the pharyngeal endoderm into pouches and the correct
patterning of the CNCC-derived skeletal elements (26). For example, a Fgf8 conditional
inactivation in the ectoderm of PA1 in a mouse model results in NCC apoptosis and absence of
most PA1 skeletal elements (73). FGF signalling is also involved in NCC spatial identity and the
establishment of anteroposterior and dorsoventral polarity of PAs (75, 76).
BMPs are osteogenic agents that induce differentiation of mesenchymal cells toward an
osteoblastic lineage and stimulate differentiation and functions of osteoblasts during bone
modeling (77, 78). BMPs are critical in regulating bone formation (79) and play an integral role
during craniofacial and pharyngeal skeletal morphogenesis, such as patterning of the maxilla and
mandible (8), mesenchyme proliferation (80) and the pathogenesis of craniofacial synostosis
(81).
At early stages of development, FGF8 and BMP4 pattern the maxilla-mandibular region and
define it from the premandibular domain (82) before the CNCCs arrive. The Fgf8 expression
domain is induced by SHH signalling from the endoderm and delimited by Bmp4 which is
expressed on both sides of adjacent Fgf-8 expressing ectoderm (82-84). BMP4 and FGF8 control
the location of incoming NCCs by activating the patterning genes Dlx1, Barx1 and Msx1. These
epithelial-mesenchymal interactions are needed for specifying the identity of the pre-mandibular
and maxilla-mandibular regions. Fgf8 also has a role in anteroposterior and dorsoventral PA
patterning and left-right symmetry of the craniofacial skeleton (85).
1.3.2 Sonic hedgehog
SHH signalling has many roles in craniofacial development. Disruption of SHH signalling in
chick, mouse and zebrafish models results in severe head skeleton abnormalities due to defects in
CNCC survival, proliferation and patterning (86, 87). If SHH is absent in the foregut endoderm,
development of Meckel’s cartilage is prevented as well as some associated PA1 structures, due
14
to NCC apoptosis (88). SHH also has a late-stage role in NCCs to promote differentiation into
cartilage and establishing skeletal polarity in the mediolateral axis of the embryo (89, 90).
SHH signalling from facial ectoderm is involved in the specification of NCC spatial identity
(91). Excess SHH results in supernumerary Meckel’s cartilage. SHH also has a late-stage role in
NCCs to promote differentiation into cartilage and establishing skeletal polarity in the
mediolateral axis of the embryo (89, 90).
A signalling center called the frontonasal ectodermal zone (FEZ) exists in the ectoderm
overlying the FNP. It regulates the growth and dorsoventral polarity of the upper beak in birds
(91). Mice have two FEZ in the left and right MNP (92). When the FEZ is grafted ectopically it
can reprogram the fate of underlying NCCs (91), showing an epithelial-mediated patterning
instruction. FGF8 and SHH are both expressed in the FEZ facial epithelia and they promote
cartilage outgrowth by inducing BMP4 expression the underlying NCCs (92). It has been
proposed that the persistence of FGF8 expression in the facial ectoderm of the duck, but not the
chick, is what contributes to the distinct morphology of the duck beak by inducing it to produce
more cartilage (93).
SHH is also involved in epithelial-mesenchymal interactions during tooth initiation. SHH is
expressed in the dental ectoderm of mice at E11. Shh knockout mice have little development of
facial processes. Addition of SHH soaked beads to oral ectoderm can induce local epithelial
proliferation to produce invaginations like those of tooth bud formation, implicating Shh in tooth
initiation (57).
1.3.3 Wnt pathways
In vertebrates, Wnt proteins have roles in regulating body axis specification, patterning of germ
layers and tissues, cell adhesion, cell migration, cell proliferation, cell differentiation,
morphogenetic movements during embryogenesis and organogenesis, and growth and metastasis
of cancers/tumours (94). WNTs are secreted glycoproteins that interact with Frizzled (Fz)
proteins on the cell surface. Fz proteins are a family of seven transmembrane G protein-coupled
receptor proteins and they mediate cell signalling from the cell surface to cytoplasm (95). The
Wnt signalling pathway can be divided into two subsets: the β-catenin-dependent canonical
pathway and the β-catenin-independent non-canonical pathways.
15
In the β-catenin-dependent canonical pathway, Wnt proteins combine and interact with an
Fz/low density lipoprotein receptor (LRP) complex on the surface of target cells (Fig. 3). This
interaction leads to stabilization of β-catenin proteins (96) and subsequent signalling cascades. In
the resting state β-catenin in the cytoplasm is phosphorylated. During activation, the accumulated
β-catenin translocates to the nucleus and interacts with transcriptional factors to activate
transcription of target genes (97). This canonical Wnt pathway is involved in cell fate, regulation
of body axis, patterning of neuroectoderm and amplification of neural progenitors (94).
Figure 3: The canonical Wnt signalling pathway. In resting cells, β-catenin is assembled in a
multiprotein complex with CK1α, GSK3β, APC and Axin and is primed for phosphorylation by
GSK3β. Phosphorylated β-catenin interacts with β-Trcp and is degraded by ubiquitation (A). In
16
the activated state, Wnt proteins act on the Fz/LRP complex on the surfaces of the target cells.
Upon Wnt-Fz signaling, Wnt-Fz and LRP coordinate Dvl activation, which results in recruitment
of axin to the plasma membrane. Activated Dvl dissociates the multiprotein complex which leads
to the inactivation of GSK3β, which can no longer phosphorylate β-catenin. Excess free
cytoplasmic β-catenin translocates to the nucleus and binds to TCF/LEF transcription factors,
causing transcriptional activation of target genes (B). Reproduced from Miao et al., 2013 (95).
The β-catenin-independent non-canonical Wnt signalling pathways has two categories: the
Wnt/Ca2+
pathway and the PCP pathway (Figure 4) (95). These pathways are involved in cell
proliferation, cell adhesion and cell differentiation, among other biological roles (98). The
Wnt/Ca2+
pathway leads to the release of intracellular calcium through G-proteins and also
involves activation of phospholipase C and protein kinase C (PKC) (99, 100). The non-canonical
Wnt-PCP pathway regulates cytoskeleton organization via actin polymerization through
activation of small GTPases (Figure 4). It also regulates the establishment of polarity within the
plane of the epithelium and allows cells to obtain directional information during embryogenesis
(101). Fzd activates JNK and directs asymmetrical cytoskeletal organization and coordinated
polarization of cells within the plane of epithelial sheets (102, 103). The Wnt-PCP pathway also
controls convergent extension movements during gastrulation and is involved in the directional
migration of cells in the developing palatal shelves (104).
17
Figure 4: The non-canonical Wnt signalling pathway. The non-canonical Wnt signalling pathway
includes the Wnt/Ca2+
signalling pathway and the planar cell polarity Wnt pathway. In Wnt/Ca2+
,
certain Wnt-Fz interactions activate the cytoplasmic protein Dvl, which increases the level of
cytosolic Ca2+
. Subsequent activation of Ca2+
-sensitive enzymes CamKII, PKC and others
induces the target gene expression and corresponding biological effects. In the Wnt/PCP
pathway, activated Dvl activates the small GTPases Rho and Rac, which leads to the activation
of Jun kinase (JNK) and rho kinase (ROK). Subsequently, transcription factors, such as the AP1
family, are activated and target gene expression is induced. Reproduced from Miao et al., 2013
(95).
1.3.3.1 Wnt pathways and craniofacial development
The Wnt/β-catenin signalling pathway components are highly conserved and are crucial for the
patterning and morphogenesis of many developmental processes, including face, limb, skeleton,
central nervous system, skin and ectodermal appendages like teeth and hair (105, 106). In
18
craniofacial development, Wnt/β-catenin is crucial for NCC induction and migration and for
proper fusion of facial prominences (66, 107, 108).
Studies in the chick and mouse have shown that Wnt signalling in specific regions of the facial
prominences correlate with outgrowth and fusion of processes (61, 109). In particular, mouse
studies have demonstrated that the LNPs were the primary region of Wnt-β-catenin signalling
(61). Mice with a loss-of-function (LOF) or gain-of-function (GOF) mutation for β-catenin
resulted in aberrant Wnt signalling in the ectoderm and embryos showed facial dysmorphologies
at the crucial stage of fusion and growth of prominences of mid-face development. Between E9.5
and E12.5 (7), LOF mutants of β-catenin had a hypoplastic facial morphology, e.g.,
underdeveloped mandibular processes, and a narrow FNP which resembled a beak at birth. GOF
mutants had a lack of controlled directional growth in their facial prominences, resulting in
increased size of the maxillary and mandibular processes and improper fusion of the processes.
GOF mutants also had ectopic cartilage throughout the head region resulting in malformations of
the head. Both the GOF and LOF mutants had alterations in the shape of their face, which would
affect the underlying cranial skeleton.
The proper growth of facial prominences is needed for cartilage formation as the mesenchymal
cells beneath the ectoderm require proliferation signals to establish a critical number of
progenitor cells (110). The progenitor cells respond to localized and intrinsic patterning signals
from the ectoderm to form cartilage of the correct shape and size (110). The strictly canonical
WNTs (2B, 7A, 9B and 16) and WNT4 and WNT6 (canonical and non-canonical Wnt ligands)
are expressed in the epithelia while WNT5A, 5B and 11 are limited to mesenchyme (109). These
data demonstrate that ectodermal Wnt/β-catenin signalling plays a crucial role in the formation
and patterning of the craniofacial skeleton.
1.3.3.2 Wnt5A
Mutations in the human Wnt pathway genes have been shown to have an effect on the
craniofacial and limb skeleton (111). For example, a recessive missense mutation of WNT5A or
WNT5A receptor ROR2 results in micrognathia, clefting and rhizomelic limb shortening seen in
Robinow syndrome.
19
Wnt5a was exclusively expressed in the mesenchyme in the facial region of developing chicken
embryos (109). Of the WNTs found in the chicken embryo face (WNT5A, WNT5B and
WNT11), 5A was the most abundant and was found in all facial prominences. At stage 21 of
chicken embryo development there are strong WNT5A signals throughout the mesenchyme of
the maxillary and mandibular prominences, lateral nasal processesand lateral edges of the
frontonasal mass, with the highest expression in Meckel’s cartilage.
In the non-canonical pathway, WNT5A binds to Fz receptors or can bind to the Ror2 receptor
(112). When WNT5A binds these receptors the JNK/ PCP pathway or calcium signalling
pathways are activated, leading to changes in actin cytoskeleton, cell polarity and cell movement.
In development, WNT5A is expressed in cartilage blastema, which may promote
chondrogenesis. However, in excess it induced rapid loss of the cartilage matrix due to induction
of metalloproteinase and aggrecanase enzymes. WNT5A regulates matrix stability but not the
initial steps of chondrogenesis. It keeps canonical signalling low in cartilage blastema and in this
manner, promotes chondrogenesis and cartilage differentiation (113). See Figure 5 for the roles
of WNT5A in development.
Some other Wnt ligands of significance to craniofacial development are WNT3A and WNT9B.
A deletion of Wnt3a causes death upon birth and these embryos have mandibular defects. Wnt9b
is involved in lip fusion and its targeted deletion results in cleft lip in animal models. A deletion
of Wnt5a leads to truncation of the upper and lower jaws (113) (114).
20
Figure 5: Summary of WNT5A functions during cartilage development and disease. A) During
normal development, there is synthesis of WNT5A in the cartilage blastema and by the newly
differentiated chondrocytes. The secreted WNT5A acts back on the same cells to inhibit
canonical activity. The net result of reducing canonical activity is to promote chondrocyte
differentiation and matrix secretion. Reproduced from Hosseini-Farahabadi et al. (113).
1.4 Collagen Triple Helix Repeat Containing 1 (Cthrc1)
1.4.1 Background
Collagen Triple Helix Repeat Containing 1 (Cthrc1) was originally discovered in a screen for
novel sequences induced in a rat arterial injury model, being expressed in adventitial cells of
remodelling arteries and dermal fibroblasts during skin wound healing (115). Cthrc1 was also
identified in a microarray study of the midface region of the mouse embryo at E10.5 (Gong
unpublished data, 2006). Cthrc1 mRNA was upregulated in the MNP compared to the LNP,
suggesting a possible critical role in the development of the midface. It is for this reason as well
as information that will be discussed in the following section that the CTHRC1 gene and protein
was chosen as a gene of interest for a role in development of the midface.
21
1.4.2 CTHRC1 Structure
CTHRC1 protein is a 30 kDa secreted N-glycoprotein containing a short collagen motif with an
NH2-terminal peptide for extracellular secretion, a short collagen triple helix repeat of 36 amino
acids and a COOH terminal globular domain (Figure 6). The helix repeat comprises 12 Gly-X-Y
repeats, a domain believed to be responsible for trimerization of the protein and protection from
cleavage (116). The normal active form of CTHRC1 is an N-glycosylated trimer anchored on the
cell surface (94).
Figure 6: The structure of the CTHRC1 protein. Adapted from Lindner (117).
CTHRC1 is exclusive to vertebrates and is highly conserved throughout evolution, showing little
homology to other currently known proteins (115). Its short collagen-like motif is similar to the
collagen domains present in the C1q/tumour necrosis factor-α related proteins, (118). CTHRC1
is thought to belong to the adiponectin/complement factor family (119) that all contain the
conserved collagen domain with the 12 GLY-x-y repeats and a globular domain in the C-
terminal half. Adiponectin is a protein hormone that modulates a number of metabolic processes,
including glucose regulation and fatty acid oxidation (120). Similar to adiponectin, CTHRC1
exists in monomeric, dimeric and trimeric forms (55). Under reducing conditions in Western blot
analysis, CTHRC1 exists as a 28 kDa protein (119). The biological activity of CTHRC1 is
restricted to the highly conserved 200 amino acids at the C-terminal region (94) which contains
an N-glycosylation site that stabilizes the CTHRC1 protein by decreasing its turnover rate.
Upregulation of CTHRC1 is seen to coincide with its increased glycosylation rate in certain cells
22
such as human oral squamous cell carcinoma (121). N-glycosylation also promotes tethering of
CTHRC1 to the cell membrane, which promotes actin polymerization and cell polarity (94).
1.4.3 Expression Pattern of Cthrc1
During development, Cthrc1 mRNA expression has been identified in visceral endoderm,
developing kidney and the heart of mouse embryos, with specific abundance in the cartilage
primordia, growth plate cartilage (excluding the hypertrophic zone), bone matrix and periosteum
(122). Prior to neural crest migration (E8.5 in mice), Cthrc1 mRNA was seen in the notochord
and the floor plate of the anterior ventral neural tube. By E9.5 transcripts were seen in somites,
branchial arches, otic placode and the hindbrain-midbrain junction with the somatic expression
becoming more pronounced by E10.5. Expression of the gene was observed in developing bone
formed via endochondral and intramembranous ossification, such as skull bones, ribs, vertebrae
and cartilage primordia (122). Cthrc1 transcripts have been shown to be expressed prominently
in chondrocytes during E14.5 of mouse development (119). Although chondrocytes from
condensing mesenchyme contained high levels of CTHRC1 protein and mRNA, hypertrophic
chondrocytes no longer expressed the mRNA. At much later time points (E18.5) developing
incisors show CTHRC1 in the dentin line and at the interface of epithelial ameloblasts and
underlying mesenchyme.
Postnatally, the expression of Cthrc1 in adult tissues has been shown at low levels and restricted
to basal expression in bone, brain and mature bone-resorbing osteoclasts. However, Cthrc1
expression surges in pathological states such as arterial injury, skin wounds or cancer (115, 122,
123). During development, Cthrc1 transcripts are prominently expressed in chondrocytes (119).
In mouse pups, high levels of CTHRC1 protein expression were seen in chondrocytes of the
resting and proliferating zone of the growth plate (122). Conversely, non-proliferating
chondrocytes in adult articular cartilage and fibrous cartilage of the meniscus did not express
CTHRC1.
1.4.4 Possible Functions of Cthrc1
Several studies have suggested that Cthrc1 plays important roles in collagen regulation, postnatal
bone remodeling, and cell migration. The following section will review the literature on the
23
involvement of Cthrc1 in these processes and discuss the possible role of Cthrc1 in the processes
in development.
1.4.4.1 Cthrc1 Role in collagen deposition
The localization of CTHRC1 to tissues rich in collagen, such as cartilage (collagen type II), bone
matrix (collagen type I) and skin (collagen type I and III), and to active sites of collagenous
matrix deposition suggests a role for Cthrc1 in modulating collagen matrix synthesis. The role of
Cthrc1 in collagen deposition has mainly been seen in studies in vascular remodelling after
injury. Cthrc1 was first discovered in a screen for differentially expressed sequences in balloon-
injured versus normal arteries where it was expressed by fibroblasts of the remodeling adventitia
and by smooth muscle cells of the neointima and was shown to decrease or inhibit collagen
matrix deposition (115). In vitro overexpression of CTHRC1 caused a dramatic reduction in
collagen type I mRNA, procollagen protein levels and collagen deposition (115, 124).
Transgenic mice that constitutively overexpress Cthrc1 under a CMV promoter have brittle
bones, caused by a reduction in collagenous bone matrix (124). The blood vessels of Cthrc1
transgenic mice had widespread cartilaginous metaplasia of the tunica media which suggests that
Cthrc1 causes the shifting of type I collagen expression profile typical of bone to a collagen type
II pattern typically found in cartilage. This shift to the subtype of collagen found in cartilage
would further support findings showing chondrocytes of the growth plate of developing bones
with abundant expression of Cthrc1 mRNA (122).
Vascular remodeling after injury is controlled by TGF-β signalling that mediates negative
aspects of vessel repair such as neointimal lesion formation, smooth muscle cell proliferation,
increased collagen deposition and lumen narrowing (125). Cthrc1 expression patterns overlap
significantly with those of TGF-β family members in calcified tissues and cartilaginous matrix,
as well as developing bone and cartilage (122, 124), periosteum, osteocytes (126), developing
skull bones, ribs, vertebrae, cartilage primordia, hyptertrophic and proliferative zones of growth
plate chondrocytes and all zones of endochondral ossification (122). CTHRC1 interacts with the
TGF-β signalling pathway to modulate collagen regulation (59). CTHRC1 can be regulated by
TGF-β and conversely, may also regulate TGF-β responsiveness and affect TGF–β target genes
(124). For example, Cthrc1 mRNA levels are increased in response to TGF-β but CTHRC1
protein is also a cell type-specific inhibitor of TGF-β which impacts collagen type I and III
24
deposition (61), neointimal formation and differentiation of smooth muscle cells (122, 124). In
transgenic mice that have upregulated Cthrc1 gene expression, TGF–β signalling is reduced in
smooth muscle cells (124).
CTHRC1’s signalling interactions with TGF-β have been shown to be via activation of Smad 2/3
complexes (127). The promoter region of Cthrc1 contains a putative Smad binding site (128) . It
has been suggested that the activation of Cthrc1 transcription could be regulated by TGF-β
signalling through Smad proteins (123). Different studies have shown that CTHRC1 regulates
extracellular collagen deposition by inhibiting phosphorylation of Smad2/3 activation via
inhibition of TGF-β signalling. Phosphorylated Smad 2/3 increases expression of CTHRC1 that
in turn inhibits the deposition of extracellular collagen controlled by Smad 2/3 phosphorylation
(123). Smooth muscle cells that overexpressed CTHRC1 protein had reduced levels of phospho-
Smad 2/3 (122). Cthrc1 transcript induction by TGF-β, the CTHRC1 inhibition of TGF-β
sensitive reporters and the reduction of phospho-Smad 2/3 levels in vivo provide evidence that
these two proteins are working as antagonists to maintain balance in extracellular matrix
components. The mechanism by which CTHRC1 disrupts TGF-β signalling, leading to a
reduction in collagen is not clear. Since CTHRC1 is a secreted protein, it is likely to function as a
ligand in a signalling pathway with downstream effects on collagen promoter activity (124).
1.4.4.2 Role of Cthrc1 in osteogenesis, osteoclastic bone formation and bone remodelling
The skeletal system fulfills mechanical, supportive, and protective roles in animals (129). Its
formation is controlled by tightly regulated programs of cell proliferation, differentiation,
survival, and organization (130). The initiation of the skeletal system is from mesenchymal
condensations, in which skeletal precursor cells, also known as osteochondral progenitors, give
rise to either chondrocytes to form the cartilage or osteoblasts to form the bone.
Bone is a mineralized connective tissue consisting by weight of 28% type 1 collagen and 5%
noncollagenous structural matrix proteins, synthesized by osteoblasts. These constituents
accumulate as the uncalficied matrix, osteoid, that acts as a scaffold for the deposition of apatite
crystals of bone to make up the remaining 67% of bone. This mineral is in the form of small
plates which lodge in the holes and pores of collagen fibrils (57). Some osteoblasts become
trapped in the bone matrix and are then referred to as osteocytes. The osteoclast is a
25
multinucleated cell which resides against the bone surface where they can bind and secrete
enzymes to demineralize the bone matrix and degrade the organic matrix.
The difference in the initial fate choices of osteochondral progenitors to either chondrocytes or
osteoblasts determines whether ossification is endochondral or intramembranous (57).
Intramembranous ossification is the direct laying down of bone into the primitive connective
tissue (mesenchyme). During intramembranous ossification, bone develops directly within the
soft connective tissue whereby osteoblasts differentiate from the mesenchyme and begin to
produce bone matrix (57). In contrast, the initiation of endochondral ossification requires the
presence of cartilage as a precursor (131). During endochondral ossification, condensation of
mesenchymal cells gives rise to cartilage cells. This cartilage will eventually be replaced by
bone. Cartilage consists of collagen of which the main type is collagen type II, in contrast to
bone which is primarily collagen type I. Perichondrium forms around the periphery of the
cartilage which gives rise to a cartilage model that will eventually be replaced by bone through
the action of osteoblasts. Prechondrogenic mesenchymal condensations in the developing mouse
face are seen as early as E8.5 and 9.5 and cartilage primordia forms at E11.5-14.5 (132, 133).
Overall, collagen deposition is critical in both endochondral and intramembranous bone
formation as the collagen matrix acts as a scaffold in endochondral bone formation and is a
component of the bone material in intramembranous bone formation and makes up the
components of cartilage.
Bone mass is regulated by a process of continual remodelling, which is based on the action
between osteoblastic bone formation and bone resorption, and is coordinated by mediation of
many signalling pathways such as parathyroid hormone, TGF-β and BMPs (134). Bone turnover
rates of 30%-100% in childhood are common and slow down in adulthood (57).
CTHRC1 has been shown to be involved in postnatal bone formation mainly through effects on
osteoblasts. CTHRC1 is expressed in bone tissue in vivo, and is a gene identified as a
downstream target of bone morphogenetic protein-2 (BMP-2) in osteochondroprogenitor-like
cells (119). Micro-computed tomography and bone histomorphometry analyses showed that
Cthrc1-null mice have low bone mass due to decreased osteoblastic formation and conversely,
transgenic mice overexpressing Cthrc1 under an osteoblastic promoter displayed high bone mass
due to an increase in osteoblastic bone formation (135). Cthrc1 osteoblast-specific
26
overexpressing and null mice also had increased and decreased levels of osteoblast specific
genes, ALP, Col1a1 and Osteocalcin, respectively, with no change in osteoclast numbers. In
vitro, CTHRC1 stimulated differentiation and mineralization of osteoprogenitor cells with a
particular acceleration of osteoblast proliferation (135). CTHRC1 has been shown to be induced
by BMP2 and is not required for skeletal development, though it is required for bone
maintenance homeostasis (135, 136). In addition to TGF-β, CTHRC1 has also been shown to be
regulated by BMP-4 (115, 128), suggesting that CTHRC1 may act as one of the downstream
targets for BMP-Smad signalling and that its expression in growth plate cartilage and bone
matrix controls collagen deposition though regulation of Smad2/3 and TGF-β signaling (122,
124).
In addition to transgenic and knockout studies, in vitro studies have further supported a role of
CTHRC1 in bone formation through its effects on osteoblasts via osteoclasts. Osteoclasts placed
on a substrate containing hydroxyapatite had upregulated expression of Cthrc1 mRNA compared
to the dentin slices, suggesting the CTHRC1 protein production is closely linked with osteoclast
attachment to calcified tissue (119). The same result was observed when osteoclasts were placed
in an environment with high extracellular calcium and phosphate. Cthrc1 expression increased in
a high-turnover state (RANKL injections in vivo) and decreased in conditions associated with
suppressed bone turnover (aging and after alendronate treatment) (119). BrdU incorporation
assays showed that CTHRC1 stimulates osteoblast proliferation in vitro and in vivo. Colony-
forming unit (CFU) assays in bone marrow cells showed CTHRC1 stimulates differentiation of
osteoprogenitor cells and osteoblasts (136). Experiments using ST2 cells, a stromal line derived
from mouse bone marrow showed that CTHRC1 targets stromal cells to stimulate osteogenesis
by binding to a putative cell surface receptor on ST2 cells to stimulate osteoblastic differentiation
as well as recruitment in order to promote bone formation (119).
The impact of CTHRC1 on bone mass was examined in mice with ovariectomies.
Ovariectomised mice are estrogen deficient and develop bone loss secondary to bone resorption.
(135). Wild-type mice showed a 47% trabecular bone loss after ovariectomy while transgenic
mice overexpressing Cthrc1 showed only 38% trabecular bone loss, suggesting a stimulatory
effect of CTHRC1 on bone formation.
27
More recently it has been shown that CTHRC1 is secreted by resorbing active osteoclasts and
acts as a coupling agent between bone resorption and formation. In vitro experiments showed
that Cthrc1 expression stimulated osteogenic differentiation of osteoblasts, measurable by
increased alkaline phosphatase and osteocalcin and had bone formation stimulating activity
comparable to that of BMP-2 and FGF-1(119). Mice containing a systemic or osteoclast specific
knockout of Cthrc1 were born appearing grossly normal but with absence of Cthrc1 mRNA.
There was no difference in mRNA levels between the two types of knockout mice, indicating
that mature osteoclasts are the major source of CTHRC1 in vivo. Both types of knockout mice
had low bone mass and abnormalities similar to osteoporosis. Histomorphometirc analysis
showed decreased osteoid surface and bone formation rate compared to control mice. These
results concluded that CTHRC1 protein is produced and secreted by mature osteoclasts and acts
on osteoblastic cells to stimulate osteoblastic differentiation and recruitment which promotes
bone formation and maintains bone mass and trabecular structure through regulation of bone
formation.
From these studies the common theme is that Cthrc1 plays a role in postnatal bone formation
due to regulation of collagen deposition as well as having an effect on osteoblasts. Taken with
results from Cthrc1 transgenic and knock out mouse experiments, evidence of a role in collagen
deposition and in vitro studies on osteoblast proliferation, CTHRC1 seems to have an anabolic
effect on bone formation though collagen matrix deposition and regulation of osteoclasts and
osteoblasts.
1.4.4.3 Role in cell motility and tissue repair
CTHRC1 is also shown to be involved in cell migration (137). Inhibition of Cthrc1 in vitro
decreases cell migration, while overexpression increases cell migration (115, 128). CTHRC1
enhances migration of smooth muscle cells and fibroblasts in a scratch wound assay, enabling
CTHRC1 overexpressing cells to migrate into the wound area significantly faster than control
cells (115, 122). Enhanced expression of CTHRC1 is involved in vascular remodelling; in rat
fibroblasts it promotes cell migration and inhibits collagen I synthesis in these cells (124). Under
normal conditions in blood vessels, CTHRC1 resides in the cytoplasm of smooth muscle cells in
an unprocessed form, despite the presence of a signal peptide. Under conditions of vascular
injury, CTHRC1 is released from the smooth muscle cells and cleavage of the N-terminal
28
propeptide results in a molecule with an increased ability to inhibit collagen matrix deposition
(138). The ability of CTHRC1 to reduce collagen matrix deposition correlates with increased cell
migration.(139). This limitation of collagen matrix deposition has been suggested as one
mechanism by which CTHRC1 promotes cell migration (128). CTHRC1 expression is also
linked to cellular proliferation, such as Schwann cell proliferation (140) and enhanced
proliferation of osteoblasts (135).
CTHRC1 has the same positive effect on migration in tumour cell lines and has been found in
increased levels in highly metastatic tumours, which suggests it plays a role in cancer
progression by promoting cell migration (128). Cthrc1 transcripts and protein are increased in
malignant melanoma, cancers of the GI tract, lung, breast, liver and pancreatic cancers when
compared to normal tissues (141) and are highly active in degrading extracellular matrix proteins
in several of these malignant tumours (128, 142, 143). A significant increase of CTHRC1 has
been observed in patients with metastatic bone cancer compared to non-metastatic tumours
(121). A germline mutation in Cthrc1 was identified in patients with Barrett’s esophagus and
esophageal adenocarcinoma (143). Cthrc1 was among 48 genes that are significantly associated
with risk of recurrence of patients with stage II/III colon cancer (144).
CTHRC1 expression has been used as a cancer prognostic indicator. Lack of CTHRC1 has been
demonstrated in non-invasive stages of melanoma in contrast to enhanced expression in primary
invasive melanomas and metastatic melanomas (142). Inhibition of Cthrc1 using short
interfering RNA showed decreased invasion of melanoma cell lines and patients with bone
metastases had a significant increase in CTHRC1 stromal expression compared to patients
without.
In conclusion, the current literature supports a role for Cthrc1 in cell migration, collagen
deposition and bone formation.
1.4.5 Genes and signalling pathways involved with CTHRC1
1.4.5.1 Cthrc1 Activation of Wnt/PCP pathway
Cthrc1 is also suggested to interact with components of the Wnt/PCP pathway in the non-
canonical Wnt pathway by forming a CTHRC1-Wnt-FZD/Ror2 complex to selectively activate
the Wnt/PCP pathway (94). The normal active form of CTHRC1 is an N-glycosylated trimer
29
anchored on the cell surface. CTHRC1 is a positive regulator of the non-canonical Wnt
signalling and simultaneously inhibits the canonical branch (94). Results from Yamamoto et al.
(94) suggest that CTHRC1 is a Wnt cofactor protein that selectively activates the Wnt/PCP
pathway by stabilizing an extracellular ligand-receptor interaction. In vitro, CTHRC1 has been
shown to interact with multiple extracellular components of Wnt signalling including both
canonical and non-canonical Wnt proteins, Fzd proteins and the Wnt/PCP co-receptor Ror2 but
not with the canonical Wnt co-receptor LRP6 or the PC component Vangl2 (94, 139) (Figure 7).
The CTHRC1 protein is thought to positively regulate the Wnt/PCP pathway by promoting
interaction between the Wnt ligand and the Fz receptor complex (94). In conditions where
appropriate receptors are available, Wnt proteins may activate the canonical or non-canonical
pathways but when CTHRC1 is present the interaction of Wnt proteins with the Fzd/Ror2
complex is enhanced and the selective activation of the Wnt/PCP pathway in this environment
suppresses the canonical pathway (145).
Through the Wnt/PCP pathway, CTHRC1 may regulate cell motility in embryogenesis and
cancer cell migration and invasiveness in adult tissues (128). It has also been speculated that the
Wnt/PCP pathway activation by CTHRC1 contributes to the promotion of cell motility because
the PCP pathway regulates actin polymerization through GTPase signalling, which alters cellular
morphology and increase cellular motility (94). This Wnt/PCP pathway plays an important role
in controlling cell polarity and movement and has been implicated in chondrocyte maturation
and cartilage formation during ontogenesis (146). The Wnt/PCP pathway regulates chondrocyte
maturation and cartilage formation as genetic alterations in the pathway are associated with
chondroplasia, dysregulation of collagen deposition and changes in cartilage morphology (94,
139, 147), thus further supporting a role for Cthrc1 in collagen deposition and cartilage
formation (139).
30
Figure 7: A model of selective activation of the Wnt/PCP pathway by CTHRC1. CTHRC1 is
cell-surface anchored and enhances the interaction of Wnt proteins, Fzd proteins and Ror2
complex to activate the Wnt/PCP pathway (94). The interaction of Wnt proteins with Fzd/Ror2 is
selectively enhanced in the presence of Cthrc1. Adapted from Yamamoto et al.(94)
1.4.5.2 Cell-specific action of Cthrc1
Cthrc1’s cell-specific behaviour is well characterized, with different actions or effects depending
on the tissue it is expressed in. CTHRC1 inhibits TGF-β signalling in smooth muscle cells but
not in endothelial cells (124). There are differing results on the effect of CTHRC1 on Col1a1
expression—upregulation of CTHRC1 in osteoblasts results in increased expression of Col1a1
whereas in PAC1 cells, a smooth muscle cell line, upregulated CTHRC1 produced reduced
mRNA levels of Col1a1 and collagen deposition (115, 124). High levels of CTHRC1 were also
associated with reduced collagen deposition in smooth muscle cells but increased collagen
deposition in osteoblasts (123, 135). These results suggest CTHRC1 function may differ between
cell types and that it may work through its own signalling pathway that is functional in a cell-
type specific manner (135).
In summary, many roles of CTHRC1 are supported by various studies in the literature.
Transgenic mouse studies have suggested CTHRC1 has a possible role in bone formation,
specifically in osteoblast regulation and collagen matrix deposition. In disease states, CTHRC1 is
associated with increased cell motility and migration and alterations in collagen matrix
deposition. These roles in cell migration, collagen matrix deposition and regulation of bone
formation have been demonstrated in vitro as well as in postnatal animals. Cell migration and
motility, bone formation and regulation of collagen expression are all important steps in
31
embryogenesis and therefore the involvement of CTHRC1 in these processes during
development is highly probable. The role of CTHRC1 in cell migration, collagen matrix
deposition and bone formation makes it a gene of interest to study in craniofacial formation as
these processes are crucial in midface development.
1.5 Rationale for Study
Many craniofacial abnormalities can be attributed to defects in the generation, proliferation,
migration and differentiation of cranial NCCs and epithelial-mesenchymal induction interactions.
Any perturbation in these processes can result in craniofacial malformations and developmental
anomalies of the facial structures. Although existing studies describe the involvement of a
number of well-known genes in midfacial development, there still remain a great number of
candidate genes whose role in development remains unexplored.
Given the evidence that the CTHRC1 protein and mRNA play roles in cell migration, collagen
deposition, and bone formation, we speculate that it plays such roles in the early and later events
during the development of the midface.
1.6 Hypothesis/Aims
The hypothesis is that CTHRC1 protein has an expression pattern in the developing mouse
embryo midface indicative of a role in craniofacial development of the mouse with respect to cell
migration, bone formation and collagen deposition.
The aims of this study are to perform a temporal and spatial analysis of CTHRC1 expression in
at crucial time points of the developing midface in mouse embryos.
32
Chapter 2
Materials and Methods
2.1 Embryos
Timed pregnant wild-type CD-1 mice were obtained from the Toronto Centre for
Phenogenomics. Embryos were removed from the uteri of mice at embryonic (E) stages E8.5,
E9.5, E10.5, E11.5, E12.5 and E13.5. Embryos were isolated from the uteri with the aid of a
Nikon SMZ800 dissecting microscope, at a temperature of 4°C. Whole embryos were dissected
for time points E8.5 and E9.5 and the head was dissected from time points E11.5, E12.5 and
E13.5. Approximately five embryos were used for each time point of immunohistochemical
staining with the CTHRC1 antibody.
Another group of eight embryos at stages E8.5, E9.5, E10.5, E11.5 and E13.5 were used for
reverse transcriptase quantitative polymerase chain reactions (RT-qPCR). The whole embryo
was collected for stage E8.5. The head was dissected from the body of the embryos of stage
E9.5, E10.5, E11.5 and E13.5. Tissues were stored at -80°C.
2.2 RT-qPCR
To obtain a temporal profile of the expression of the Cthrc1 gene mRNA, RT-qPCR was
performed on RNA samples from embryos at stages E8.5, E9.5, E10.5, E11.5 and E13.5.
2.2.1 RNA isolation
Tissues of embryos harvested at specific embryonic stages, as described above, were
homogenized in TRI Reagent (Sigma Aldrich) for 5 minutes. Chloroform was added to samples,
which were shaken for 15 seconds. Tissue samples were centrifuged at 12,000g for 15 minutes at
4°C. The aqueous phase was removed and mixed with isopropanol, incubated for 10 minutes at
room temperature and then centrifuged at 12,000g for 8 minutes at 4 °C. The supernatant was
removed to isolate the RNA pellet, which was washed with 75% ethyl alcohol (ETOH) and
centrifuged at 7500 g for 5 minutes at 4 °C. The ETOH supernatant was removed and the RNA
pellet was resuspended in distilled H20. Samples were run on a 2% agarose gel and stained with
33
ethidium bromide to determine the integrity of the RNA; specifically, to determine the presence
of the small (2 kb) and large (5 kb) ribosomal RNA.
2.2.2 Reverse transcription of RNA samples
Reverse transcription of the RNA from each time point sample was performed using the Maxima
First Strand cDNA Synthesis Kit for RT-qPCR (#K1641; Thermo Fischer Scientific). A reaction
mix was made with the sample template RNA, H20, Maxima Enzyme Mix and 5X reaction mix.
Reaction samples were incubated for one cycle of 10 minutes 25°C, 30 minutes 72°C, 5 minutes
85°C and then stored at 4°C.
2.2.3 Quantitative PCR
Primers for Cthrc-1 were designed using Ensembl software on the Cthrc1 gene
(http://useast.ensembl.org/Mus_musculus/Gene/Sequence?g=ENSMUSG00000054196;r=15:390
76932-39087121). Primers were selected that skipped the intron between exons 1 and 2 (3,323
bases). The primers spanned basepairs 62-279 of the Cthrc1 gene, producing an amplified
fragment of 218 basepairs. The primer sequences were:
Forward primer: 5’-T G C T G C T G C T A C A G T T G T C C-3’
Reverse primer: 5-’T C C C T T T T C C C C T T T G A A T C-3’.
All reactions were performed in a 96-well plate using the iTaq™ Universal SYBR® Green
Supermix (Biorad). GAPDH was used as the reference gene.
PCR reactions of cDNAs from tissue samples at each time point sample were run in duplicate at
two different dilutions (1:5 and 1:10), for quality control. PCR conditions were as follows: 40
cycles of 94°C, 58°C and 72°C to allow for denaturing of the cDNA, binding of the primer and
extension of the primer, respectively. cDNA quantities were determined using the CFX96
Touch™ Real-Time PCR Detection System and CFX manager software 3.0 (Biorad) and plotted
on a graph using the formula: 2∆CT(target)
/ 2∆CT(reference)
. “∆CT target” was the change in cycle
threshold value for the tissues at each time point compared to E8.5 and “∆CT reference” was the
change in cycle threshold values of GAPDH at the corresponding time points compared to E8.5.
The values from this ratio were plotted using Excel software (Microsoft Office 2007).
34
2.3 Histological Processing and Paraffin Embedding
Embryos were transferred to Bouin’s fixative (Polysciences, Inc.) to incubate for 24 hours,
followed by storage in 70% ETOH at room temperature (20°C). Prior to paraffin embedding the
embryos were stored in 80% ETOH for 1 hour, 100% ETOH for 1 hour, followed by incubation
in methyl benzoate overnight. Afterwards, embryos were transferred to toluene for 2 hours.
All embryos were processed for paraffin sectioning according to Gong (2001)(148). Sections
were mounted on Superfrost slides (Fisherbrand). Embryos were oriented such that they were cut
in a coronal, transverse or sagittal plane. The choice of orientation for different time points
depended on the need to capture the pattern of CTHRC1 expression in specific midfacial
structures at different stages of development in the midface. Therefore, embryos at E8.5 and E9.5
were oriented only in sagittal or coronal planes and E10.5-13.5 were oriented in the coronal or
transverse planes.
2.4 Immunohistochemistry
Immunohistochemical reactions using a rabbit polyclonal antibody to Cthrc1 (ab85739; Abcam)
were conducted on tissue sections at different embryonic stages according to Gong (2001)(148).
To ensure specificity of the α-CTHRC1 antibody, a peptide competition assay was performed to
rule out false positive results. Transverse sections of embryos at E13.5 were selected based on a
distinct pattern of expression. The Cthrc1 peptide (ab101727; Abcam) was incubated with the
primary antibody at 5x the primary antibody concentration used for the immunohistochemistry at
4°C for 24 hours prior to the staining. The staining was then carried out as described in Gong
(2001)(148), with additional slides receiving the peptide-blocked primary antibody instead of the
primary antibody alone. Subsequent steps for the staining of these slides were as described in
Gong (2001)(148). The signal detection was compared in the sections stained with the peptide-
blocked primary antibody and the primary antibody alone.
The sections on the slides were deparaffinised and rehydrated with washes of 100% ETOH, 95%
ETOH and 70% ETOH. Tissue sections were washed for 30 minutes in phosphate buffered
saline (PBS). Sections were treated with 0.3% H202 for 20 minutes to quench endogenous
peroxidase activity and washed for 10 minutes with PBS. Antigen retrieval was conducted by
treating sections with 0.01 NaCitrate buffer (pH 6.0) for 10 minutes at 90°C, followed by washes
35
for 10 minutes in PBS. Tissues were then incubated in a blocking solution of 1.5X normal goat
serum (Zymed), 0.01% Saponin and 0.1% Bovine Serum Albumin (Sigma) to prevent non-
specific binding of the secondary antibody. The tissue sections were subsequently incubated with
the primary antibody at a 1:100 dilution in PBS overnight at 4°C.
The next day, tissue sections were washed with PBS for 30 minutes and incubated with a
polyclonal goat-anti-rabbit biotinylated IgG (R&D Systems) at a dilution of 3:100 for 40 minutes
at room temperature. The tissues were washed for 30 minutes with PBS and then incubated with
Vectastain Elite ABC kit (Vector Laboratories) for 1 hour. Lastly, sections were washed in PBS
for 30 minutes and the antibody signal was detected using a DAB Peroxidase Substrate Kit
(Vector Labs). The sections were then rehydrated and coverslips were placed using Permount
mounting media (Fischer Scientific). Negative controls were performed concurrently, without the
presence of the primary antibody.
2.5 Documentation and analysis of Cthrc1 expression
Images of immunohistochemically stained histological sections were taken using Spot Advanced
software and the Spot Diagnostic Camera attached to an Olympus BX51 camera at 10x and 4x
magnification. Signal localization was determined by images taken with the digital camera and
observations were recorded for each developmental stage. Location as well as spatial distribution
of the signal was recorded.
36
Chapter 3
Results
3.1 Quantitative expression of Cthrc1 mRNA transcripts during midface development
Analysis of Cthrc1 expression in the midface tissues of embryos at E8.5, E9.5, E10.5, E11.5 and
E13.5 developmental stages by quantitative polymerase chain reaction (qPCR) revealed a
specific temporal pattern. There was an increase of Cthrc1 mRNA from time points E8.5 to E9.5.
At time point E10.5 there was a decrease in mRNA levels from the E9.5 time point. At E11.5
there was an increase of the Cthrc1 transcripts back to a level similar to what was seen at E9.5.
At the last time point of E13.5 the highest level of Cthrc1 transcript increase was seen, as
compared to E8.5 (Figure 8).
Figure 8: Temporal expression profile of Cthrc1 mRNA across E8.5-E13.5. RNAs were
extracted from tissues of embryos from each time point, reverse transcribed to produce the
corresponding cDNA and amplified by qPCR. Cthrc1 mRNA levels were compared to levels at
time point E8.5.
Overall, the results from the qPCR revealed that expression of Cthrc1 mRNA peaked at E9.5
relative to E8.5, followed by a decrease in expression at E10.5 and then a steady increase after
37
E11.5. The results confirmed the presence of Cthrc1 mRNA in the midface during these time
points of embryonic development.
3.2 Expression of CTHRC1 protein
The spatial expression of CTHRC1 protein was assayed at six different time points of embryonic
development: E8.5, E9.5, E10.5, E11.5, E12.5 and E13.5. Tissue sections of the sagittal,
transverse and coronal planes through the developing craniofacial region were used for the
expression analysis. A distinct spatial pattern of expression of CTHRC1 was clearly observed at
each time point (Figures 10-15).
3.2.1 CTHRC1 peptide competition assay at E13.5 to confirm specificity of primary CTHRC1 antibody
A peptide competition assay of the CTHRC1 antibody revealed the specificity of the antibody for
its antigen. Compared to the distinct expression of CTHRC1 in the nasal septum and nasal
capsule (Figure 9C; see below for more details of expression), no staining was observed in the
areas of CTHRC1 expression on a consecutive tissue section of the nasal capsule and septum
where the antibody was blocked with the CTHRC1 peptide (Figure 9A). The negative control
(absence of primary antibody) also showed no staining (Figure 9B), similar to that of the blocked
primary antibody. These results indicate that the primary antibody, raised against the CTHRC1
peptide, was specific to the protein with little cross-reactivity or non-specific binding.
Figure 9: Peptide competition assay on consecutive coronal sections through the anterior part of
the midface in an E13.5 embryo. A) Peptide block at 5X antibody concentration showing no
signal. B) Negative control staining with goat anti-rabbit secondary antibody showing minimal
38
background signal. C) CTHRC1 primary antibody (1:100) and secondary goat anti-rabbit
antibody showing a distinct pattern of CTHRC1 expression in the nasal septum (asterisks) and
nasal capsule (arrow).
3.2.2 Spatial expression analysis of CTHRC1 in the midface at different developmental stages
At the earliest observed time point of E8.5, CTHRC1 expression was limited to the midline of
the embryo, most strikingly in the notochord and neural tube (Figure 10). In the midline,
expression was observed in the notochord, extending from the rostral end and along the length of
the dorsal side of the embryo (asterisks, Figure 10A and C). In addition, CTHRC1 expression
was also seen in the neuroectoderm in the area of the ventral brain vesicle and at the rostral end
of the embryo (arrows, Figure 10B and D). Expression of the protein was observed in a small
area of the oral ectoderm (open arrowhead, Figure 10D).
Figure 10: CTHRC1 protein expression in E8.5 of coronal (A, B and D) and sagittal (C) sections.
A) Asterisk shows CTHRC1 localized to the notochord (nc) and arrow shows CTHRC1
expression in neuroectoderm (ne). B) Arrow shows CTHRC1 expression in neuroectoderm (ne).
C) Sagittal section showing CTHRC1 localized (asterisks) to notochord (nc) along rostral-caudal
length of the embryo. D) CTHRC1 expression in the neuroectoderm (arrow) and oral ectoderm
(open arrowhead).
39
By E9.5 and E10.5, CTHRC1 protein was exclusively expressed in the mesenchyme (Figure 11).
Expression was present in the facial prominences, specifically the mesenchyme of the
frontonasal and medial nasal processes and the maxillary and mandibular processes (Figure 11 A
and B). In the more anterior or ventral region of the medial portion of the midface, e.g., the
developing frontonasal region, CTHRC1 was widely expressed throughout the whole area,
including the medial-most or central portion of the FNP (Figure 11B and C).
Another consistent pattern of CTHRC1 expression in the craniofacial region was the
mesenchymal expression of CTHRC1 adjacent to the ectoderm and neuroectoderm. CTHRC1
mesenchymal expression in the frontonasal and medial nasal processes was adjacent to the
ectoderm of the respective facial processes (Figure 11A and C). Also, in certain areas, CTHRC1
protein was strongly expressed in mesenchymal tissues immediately adjacent to the
neuroectoderm of the telencephalic brain vesicles (arrowhead, Figure 11D).
Figure 11: CTHRC1 protein expression in E10.5 embryos, in coronal (A, B and D) and sagittal
(C) sections. A) CTHRC1 protein expression in mesenchyme of the frontonasal process (FNP)
40
(arrows). B) CTHRC1 expression in the frontonasal process (FNP, arrow), maxillary process
(Mx, tailed arrow) and mandibular process (Md, open arrow). C) Expression in FNP adjacent to
ectoderm (arrow) and in ventricle of heart (H). D) Distinct expression in mesenchyme next to
neuroectoderm (MeN, arrowhead) and mesenchyme in Md process (open arrow). Figures A, B
and D are sections from three E10.5 representative embryos, with A and B sections being more
ventral and D located more dorsally.
By stage E11.5 of embryonic craniofacial development, expression of the CTHRC1 protein was
observed in a distinctive pattern in areas of cartilage formation and deposition. As cellular
condensations are initiated in different parts of the developing craniofacial region, CTHRC1
expression was clearly expressed in many of these structures. The maxilla at this stage presents
with a nasal septal cartilage analgen, a rod like structure that is flanked with two wing-like
cartilaginous processes in the developing nasal capsule. In the maxilla and forming snout of the
mouse, distinct expression was observed to localize at the forming nasal septum and nasal
capsule (Figure 12 A and B).
Figure 12: CTHRC1 protein expression in E11.5 through transverse sections of the maxilla nasal
septum and nasal capsule. A) CTHRC1 expressed in areas of the nasal capsule (nc) and nasal
septum (ns) (arrows). B) CTHRC1 in nasal capsule (nc) and nasal septum (ns). A, B = 4x and
10X magnifications, respectively.
ns
nc
41
CTHRC1 was still strongly expressed in the nasal septum and other areas of cartilage formation
of the midface at the later time points of E12.5 and E13.5. With further development of the nasal
cartilaginous areas, the staining of CTHRC1 appeared as an almost complete ring-like pattern
with a central rod staining, within which contains the bilateral nasal cavities (Figures 13A, 13B
and 14).
Figure 13: CTHRC1 protein expression at E12.5 in coronal sections of the nasal capsule and a
transverse section of the mandible. A) A coronal section of the snout showing CTHRC1
expression in the nasal capsule (nc). B) A coronal section showing CTHRC1 expression in the
42
nasal septum (ns). C) A transverse section showing CTHRC1 expression in Meckel’s cartilage
(mc).
At E13.5, the same pattern of expression of the CTHRC1 protein was seen in the maxilla and
mandible, but the signal became more distinct and covered a larger surface area as the regions of
cartilage formation and deposition increased with further development of the midface.
Expression was also seen in other areas of cartilage formation in the mouse embryo head, such as
in the developing bones of the calvarium (Figure 14A).
Figure 14: CTHRC1 protein expression at E13.5 in sections of head (A), the maxilla (C and D)
and midface including mandible (D). A) CTHRC1 expression in Meckel’s cartilage (arrow, mc)
and bones of skull (arrow, c). B) More dorsal transverse section of midface and mandible
showing CTHRC1 expression in nasal septum (ns) and nasal capsule bones (nc). C) More
anterior section of CTHRC1 expression in nasal septum (ns) and nasal capsule bones (nc). D)
Arrows showing Cthrc1 expression in nasal capsule (arrow, nc) and bones of maxilla (arrow,
Mx).
mc
nc
M
x
c
43
3.2.3 CTHRC1 expression in the mandible and developing tooth bud
In the developing mandible of embryos at stages E9.5, E10.5 and E11.5, the pattern of CTHRC1
expression was consistent with the pattern seen at E10.5 in the maxillary process and frontal
nasal process. CTHRC1 expression was localized to the mesenchyme and was always adjacent to
areas of the ectoderm. For instance, the protein was expressed in embryos at stages E9.5 and 10.5
in areas of outgrowth, with a higher expression in the anterior and ventral portion of the facial
processes (Figure 11 A and C) and the oral half of the mandibular process (open arrow, Figure
11B). As cartilaginous structures are laid down in the mandible, the expression of CTHRC1 was
clearly evident. One structure is Meckel’s cartilage, a bilateral cartilaginous bar across the
mandibular process. A strong CTHRC1 signal was localized in the area corresponding to
Meckel’s cartilage (Figure 13C and 14A). The distinct pattern of CTHRC1 expression in
Meckel’s cartilage was only observed at later time points, specifically E12.5 (Figure 13C) and
E13.5 (Figure 14A). CTHRC1 expression was also observed at time point E12.5 lateral to the
Meckel’s cartilage in an area that most likely represents the membranous ossification of the body
of the mandible (arrow, Figure 13C).
CTHRC1 also appeared to be expressed in areas of tooth formation. Initially, its expression was
generally throughout the mesenchyme with a stronger signal near the epithelial thickenings on
the margins of the stomodeum, which corresponds to the formation of the dental placode (Figure
11B, 11D and 15A). At E11.5, the formation of what is equivalent to the cap stage of tooth
development has started in the mandible. CTHRC1 expression at this time was seen in the
mesenchyme directly below areas of epithelia corresponding to the enamel organ (Figure 15B
and C). This distinct pattern of mesenchymal expression in the area corresponding to the dental
papilla continued until the last observed time of E13.5 (Figure 15D).
44
Figure 15: CTHRC1 protein expression in tooth bud formation. A) E10.5: Mandible, arrows
showing CTHRC1 expression in mesenchyme of the mandibular process (Md) near the
ectoderm. B) E11.5: Tongue (t) and Mandibular process (Md), arrow indicating CTHRC1
expression in mesenchyme near tooth buds. C). E11.5: Arrows pointing to CTHRC1 expression
in mesenchyme near tooth bud. D) E13.5: Arrows showing expression in mesenchyme adjacent
to developing tooth.
45
Chapter 4
Discussion
This chapter of the thesis will outline the possible roles for Cthrc1 in midface development by
drawing conclusions based on the data from the Results section in combination with evidence
from the literature. The results will be briefly summarized followed by a discussion of proposed
roles for Cthrc1 in the development of the midface, future directions and concluding remarks.
4.1 Summary of Cthrc1 expression during mouse embryo midface development
4.1.1 Summary of qPCR mRNA levels of Cthrc1 mRNA during E8.5–E13.5
Overall, our data revealed that Cthrc1 has a quantitative expression profile that changed across
development, with these changes correlating closely with some of the major changes during early
midfacial development. There was an increase in Cthrc1 mRNA levels from E8.5–E9.5, a stage
when the CNCC begin to migrate from the neural tube (149) towards the facial prominences
located ventrally (150). At E10.5 there was a decrease from the E9.5 mRNA levels with levels
still about that of E8.5. This interval corresponds to CNCCs arriving at the locations where they
will begin to interact with surrounding tissue to begin the process of condensation of cell types
(57). At E11.5 there was an increase of the Cthrc1 transcripts back to a level similar to E9.5. At
E11.5 there has been condensation of cell types, such as mesenchyme, that will start developing
into chondrocytes or other specific cell types. At the last time point of E13.5 the highest level of
Cthrc1 mRNA transcript increase was seen compared to E8.5. At this time, proliferation and
differentiation of tissue such as cartilage is starting in the midface of the embryo. Overall,
increases in Cthrc1 mRNA were seen at E9.5, with a relative decrease at E10.5 and then a steady
increase from E11.5 onwards.
It is important to note that for the time point of E8.5, mRNA was extracted from the whole
embryo due to difficulty in obtaining enough materials from only the heads of the small embryo.
For the remaining time points the head of the embryo was dissected from the body and processed
for qPCR. The levels of mRNA at E8.5 may not reflect a true comparison to the other time points
46
because it was measured from the whole embryo rather than the head. This would most likely
lead to an overestimation of the mRNA level at E8.5 in the head as all tissues in the embryo were
processed.
4.1.2 Summary of immunolocalization experiments results of CTHRC1 protein from E8.5-13.5
The immunolocalization experiments, performed by immunohistochemistry (IHC), in this study
provided information regarding the spatial expression of the CTHRC1 protein at the time points
E8.5, E9.5, E10.5, E11.5, E12.5 and E13.5, allowing visualization of the type of tissue and
boundaries of CTHRC1 expression. The spatial expression patterns of CTHRC1 suggest that it
has roles in cell migration, epithelial-mesenchyme interactions, collagen formation and bone
formation. Expression at E8.5 of CTHRC1 at the notochord, oral ectoderm and neuroectoderm
suggest a role in cell migration and signalling of NCCs towards the structures or prominences to
which they will migrate. This corresponds to previous results from Durmus et al. (122) who
showed Cthrc1 mRNA expression in the notochord at E8.5. At E9.5 and 10.5, the close
association of CTHRC1 protein in the epithelial-mesenchyme border supports a role of CTHRC1
in epithelial-mesenchymal interactions and signalling between the two tissues during formation
of midface structures. The presence of CTHRC1 in the medial nasal process of the FNP supports
results from the microarray experiment screening genes expressed in the medial nasal and lateral
nasal process (Gong, unpublished results 2006). CTHRC1 protein expression was also seen in
the heart which corresponds to previous mRNA expression studies of Cthrc1 mRNA in the
mouse embryo (122). CTHRC1 expression patterns at E11.5 in areas of condensation of
mesenchyme for future cartilage and bone formation such as the area of the future nasal septum,
nasal capsule suggest a role of CTHRC1 in cell condensations. Expression was also seen in the
mandibular process in the mesenchyme but was concentrated to areas adjacent to the epithelium
of developing tooth buds at the placode and bud stage of tooth development, suggesting a role in
epithelial mesenchyme interactions. At E12.5 and E13.5 CTHRC1 expression suggests a role in
chondrocytic differentiation, as it was present in the nasal septum and capsule, a role in
intramembranous ossification due to its expression in developing bones of the calvarium and the
area adjacent to Meckel’s cartilage, and a role in epithelial-mesenchyme interactions due to its
expression in mesenchyme corresponding to the dental papilla.
47
4.2 Involvement of Cthrc1 in midface development
This section of the discussion will examine the possible roles for Cthrc1 in the development of
the midface in the mouse embryo based on previous studies from the literature as well as the
results presented in this thesis. Known events of craniofacial development will be correlated to
the expression patterns seen with the qPCR and IHC results and from this, possible roles for
Cthrc1 will be discussed.
4.2.1 Role of Cthrc1 in cell migration
The previously documented role of Cthrc1 in cell migration (128, 137), cancer progression and
vessel injury (83, 115) via its ability to reduce collagen matrix deposition supports a possible role
for it in cell migration during embryogenesis. NCCs need to migrate in order to reach their target
tissues from the neural tube (8).
Overall, a role for CTHRC1 in cell migration is supported by the spatial expression pattern in the
midface during early embryonic development when CNCCs are known to migrate from the
neural crest to the facial prominences. The upregulation of Cthrc1 mRNA as assayed by RT-
qPCR during this time point also supports an increase in CTHRC1 protein in preparation for its
role in cell migration. During early embryonic development, around E8.5 (day 17-19 in human
embryonic development), NCCs are still located in the neural tube, but about to begin migration
to various structures in the body or head (68). Our spatial analysis of CTHRC1 showed a change
in expression in the embryonic midline at E8.5, to the mesenchyme in the facial prominences by
E9.5 (approximately day 22 in human embryo development) (50, 149). During this stage in the
embryonic mouse development the CNCCs are beginning to migrate (149) ventrally towards the
facial prominences (52, 150). The increase in Cthrc1 mRNA levels from E8.5–9.5 in
combination with this dramatic change in expression pattern from the embryo midline to facial
prominences suggest that CTHRC1 protein may play a role in the promotion and emigration of
the CNCCs.
A possible role for Cthrc1 in mediating migration of CNCC is corroborated by other studies
showing the involvement of CTHRC1 with Wnt ligands. Recent studies have shown that the
non-canonical Wnt–PCP pathway plays a major role in neural crest migration (151). PCP
signalling controls CIL between NCCs by localizing different PCP proteins at the site of cell
48
contact during collision and locally regulating the activity of Rho GTPases (151). The PCP
pathway regulates actin polymerization which can alter cellular morphology and cellular
motility. Vertebrate PCP signalling is regulated by Wnt proteins and CTHRC1 has been shown
to co-precipitate in vitro with Ror2, the receptor for WNT5A, and non-canonical Wnt proteins
(94). One study supporting a possible role in cell migration for CTHRC1 during development
was Yamamoto et al. (94) where a knockout of Cthrc1 alone did not produce a phenotype;
however, upon introduction of a heterozygous Vangl2 mutation, a gene involved in the migration
of groups of cells during vertebrate embryogenesis, abnormalities characteristic of a PCP mutant
were expressed. Abnormalities included a shortened body axis, open neural tube and
misorientation of sensory hair cells of the cochlea. These defects may be due to failure of NCC
migration, indicating a possible role for Cthrc1 in the migration of these cells. Based on both in
vivo and in vitro results (94) it is possible that CTHRC1 promotes cell migration through
activation of the Wnt/PCP pathway.
Another interesting finding from our study was the localization of CTHRC1 to midline structures
such as the notochord at E8.5. The notochord is located in the embryonic midline, ventral to the
neural tube, exists transiently in the vertebrate and has a role in patterning of the neural tube
(152). The proximity of the CTHRC1 protein expression in the notochord to the location of
CNCCs in the neural crest in our immunolocalization experiments suggests an association and
possible interaction between the CTHRC1 protein and CNCCs at this time point. If there is an
association of CTHRC1 with NCCs with respect to migration or guidance, this colocalization to
the midline adjacent to the area where NCCs reside would support this.
4.2.2 Role of Cthrc1 in epithelial/mesenchyme interactions
Epithelial-mesenchymal tissue interactions (EMI) are critical in the initiation of cell
differentiation and morphogenesis in virtually every organ in the vertebrate body (153). During
development, the mesenchymal cells beneath the facial ectoderm receive proliferation signals for
the appropriate amount of time to establish a critical number of progenitor cells (110). EMI
immediately precede condensation of either prechondrogenic mesenchyme during
chondrogenesis or they initiate transition from osteogenic mesenchyme to preosteoblasts in
osteogenesis in craniofacial skeletal development (153). Several molecules such as BMP-4,
BMP-2, Msx-1 and TGFβ are known to be involved in EMI which form the structures of the
49
developing face (8, 80, 153). CTHRC1 is a downstream target of BMP-2 and is known to be
regulated by TGF-β and BMP-4 (115, 124, 128, 135). The association of CTHRC1 with
signalling molecules known to participate in epithelial-mesenchymal interactions, in addition to
the spatial expression seen in the mouse midface in this study supports a role for CTHRC1 in the
EMI process. There was a consistent pattern of expression in the mesenchyme of the FNP,
maxillary and mandibular processes concentrated in areas adjacent to the epithelia and
neuroectoderm at E10.5 and E11.5. EMIs are crucial for specifying the identity of the pre-
mandibular and maxilla-mandibular regions (82). These signalling interactions co-ordinate the
outgrowth of the facial primordia from buds of undifferentiated mesenchyme into the intricate
series of bones and cartilage structures that, together with muscle and other tissues, form the
adult face (154). SHH signalling promotes the differentiation of NCCs into cartilages (89, 90)
and based on our results showing CTHRC1 colocalization to areas of cartilage formation there is
a possibility of interactions between the SHH in the epithelium and CTHRC1 in the underlying
mesenchyme during differentiation of chondrocytes.
Before the CNCCs arrive at their destination in the facial prominences, ectodermal Fgf8
prefigures the prospective oral cavity. This FGF8 expression is induced by SHH signalling from
the endoderm and is delimited by BMP-4 expressed on both sides of the adjacent Fgf8-
expressing endoderm (82-84). The ectodermally derived FGF8 and BMP4 control the
regionalization of incoming NCCs through the activation of specific patterning genes such as
Dlx1in the cases of FGF8 while BMP-4 induces Msx1 expression in the underlying mesenchyme
(8, 82). CTHRC1 has been shown to be regulated by BMP-4 (115, 128). Bmp4 mediates the
activity of proliferation zones in the FNP and between the FNP, maxillary and nasal prominences
(93) and its expression in the FNP is crucial for the final morphology of the midface in chick
embryos (155, 156). BMP-4 expression in NCCs of the mesenchyme in the FEZ is induced by
Shh and Fgf8 and promotes cartilage outgrowth (92, 157). Therefore the evidence from our study
showing mesenchymal spatial expression of CTHRC1 in proximity to the epithelia in both
maxilla and mandible prominences in addition to evidence that BMP-4 is already involved in
epithelial-mesenchymal supports a role for CTHRC1 in epithelial-mesenchyme interactions.
CTHRC1 expression was first seen in the mesenchyme adjacent to the epithelium in the facial
processes of E9.5 and E10.5 embryos in our study. β-catenin mouse mutants with upregulated
Wnt/β-catenin in the ectoderm during facial morphogenesis showed normal facial development
50
at E9.0. Shortly after, when the facial prominences began to enlarge, the nasal processes,
maxillary and mandibular processes of these mice also had an increase in overall size compared
to wild-type and altered expression of fgf8 and Shh, Bmp2 (7). Interactions of CTHRC1 with
members of the Wnt signalling pathway, CTHRC1 as a downstream target of BMP-2 and FGF8,
and BMP4 having a possible connection with CTHRC1 signalling, is further evidence that
CTHRC1 plays a critical role in mediating critical interactions between the ectoderm and
mesenchyme that leads to the proper shape and size of the face.
The proximity of CTHRC1 expression to the epithelium destined to become the enamel organ
supports a role in induction of this tissue which will go on to form the enamel and dentin of the
tooth. This concentration and proximity of the CTHRC1 expression in the mesenchyme of the
facial prominences to the epithelium and ectoderm in addition to the fact that CTHRC1 is known
to interact with other molecules involved in epithelial mesenchyme interaction would support its
role in this process. In the case of tooth development, BMP-4 is a regulator of CTHRC1 and
when it is expressed in the ectoderm of the facial processes it induces the expression of Msx1 in
the underlying mesenchyme (8) which is important in tooth formation. Bmp4 and Msx1 act in a
positive feedback loop to drive sequential tooth formation. BMPs, FGFs, SHH and Wnt
pathways are all repeatedly used throughout tooth development (158). Our results show a
localization of CTHRC1 in the mesenchyme starting at E9.5 and remains until our last time point
of E13.5. Therefore at these time points there may be epithelial-mesenchyme interactions driving
tooth formation from the placode stage to the tooth bud stage.
4.2.3 Role of Cthrc1 in regulation of collagen formation and deposition
The stages of chondrogenesis in the craniofacial area have been divided into a) epithelial-
mesenchymal interactions preceding condensation of prechondrogenic mesenchyme, b)
condensation c) prechondroblasts differentiating into chondroblasts, d) deposition of
extracellular matrix and e) terminal differentiation and mineralization depending on the cartilage
type (153). Condensations are initiated by alteration of the mitotic activity and aggregation of
cells and are a product of the preceding stage of EMI (153). Our spatial expression analyses of
embryos at later developmental stages support important roles for CTHRC1 in a number of these
critical stages of chondrocytic condensation and differentiation. By E11.5 (approximately day 33
of human development), sites of mesenchymal condensations in the facial prominences are
51
initiated and these areas will eventually differentiate into chondrocytes or other specific cell
types. The CTHRC1 pattern of expression in our experiments at this time most likely represents
the beginning of condensations of chondrogenic mesenchyme which is verified in our results by
CTHRC1 expression in areas of future cartilage formation. The qPCR experiment shows a
dramatic increase in the levels of Cthrc1 mRNA between E10.5 and E11.5. The increase may
reflect an increased production of CTHRC1 protein for its upcoming role in condensation of
chondrogenic cells. Our results did not show expression in the area of the future nasal septum at
E9.5 or E10.5 but a distinct pattern was seen at E11.5. This may be interpreted as condensation
not having started yet or if it has, CTHRC1 protein expression may not be strong enough to
produce a visible signal with IHC or its expression has not started yet in this area of
condensation. These areas will form cartilage and may go on to ossify via endochondral
ossification. This is the first time point in the IHC experiments to show expression of CTHRC1
in the exact position of future cartilage formation. In our experiments, the CTHRC1 expression
at E11.5 in the nasal septum and capsule marks chondrogenic condensation for the maxilla,
preceding the cell differentiation into chondrocytes prior to cartilage formation. At E12.5 and
E13.5 there is also a significant expression of CTHRC1 in the area of future Meckel’s cartilage.
Meckel’s cartilage does not form the bones of the mandible, which form from intramembranous
ossification, but will go on to form the sphenomandibular ligament, and malleus and incus bones
of the ear via endochondral ossification. Meckel’s cartilage forms in the 5th
week of human
development which corresponds to E14.5 in the mouse. Therefore at E12.5 and 13.5
condensations of chondrogenic precursors cells would be taking place in Meckel’s cartilage,
slightly later than the condensations in the maxilla.
BMP-4 is another growth factor that has been shown to be upregualted during chondrogenic
condensation, acting as an inducer of condensations (159). It has been shown to be present in
condensing mesenchyme in facial processes of mice (160). BMP-2 is also present in condensing
chondrogenic and osteogenic mesenchyme in the embryonic chick (161). CTHRC1 is a
downstream target of BMP-2 and is regulated by BMP-4 (115, 128) therefore one mechanism by
which CTHRC1 regulates chondrogenic condensation may be through these two molecules.
These condensations mark the onset of selective gene activity that will enable cell differentiation,
for example in prechondrogenic condensations there is a 7-fold increase in mRNA for type II
collagen, illustrating the importance of condensation for the initiation of cell differentiation (153,
52
162). Similarly in our results we see a near 6-fold increase in Cthrc1 mRNA at E11.5 relative to
E8.5. This increase in mRNA indirectly reflects a future increase in the protein. In the case of
CTHRC1, the amount of mRNA may be increasing at this time in preparation for differentiation
of chondrocytes or to start producing collagen. Since CTHRC1 is known to promote collagen
type 2 deposition, the protein expression in the cartilage anlagen of the nasal capsule and nasal
septum as well as Meckel’s cartilage in addition to the increase in mRNA transcripts supports a
role for CTHRC1 in chondrocyte differentiation. The duration of condensation can vary as it is a
transient stage on the way to overt cell differentiation.
Our results also support a role for CTHRC1 in collagen deposition. At E12.5 and 13.5, a strong
pattern of CTHRC1 protein expression was seen in the cartilage anlagen of the nasal septum and
nasal capsule. Like E11.5, this distinct pattern mirrors exactly the future structure of the nasal
septum and capsule, supporting a role for CTHRC1 in collagen deposition. The fact that
CTHRC1 shifts the collagen expression pattern to that of collagen type II supports a role for
CTHRC1 in collagen deposition for the cartilage of this area. The situation would not be the
same for bone deposition since CTHRC1 inhibits expression of type I collagen mRNA and
matrix deposition (122, 123).
The exact mechanism by which CTHRC1 regulates collagen deposition is not known but there
are several possibilities based on current studies in the literature. CTHRC1 inhibits type I
collagen deposition via inhibition of TGF-β (138) (124) and collagen deposition is regulated by
TGF-β activation of Smad 2/3 complexes. Therefore, a role of CTHRC1 in collagen matrix
deposition for bone formation would likely be inhibitory. Transgenic studies show that CTHRC1
overexpression resulted in brittle bones when overexpressed in all cell types (124) but when only
overexpressed in osteoblasts, these mice had increased bone mass. Although CTHRC1 inhibits
TGF-β, certain members of the TGF-β family such as Activin are known to enhance
condensation size and stimulate chondrogenesis (163). This suggests that CTHRC1 may have a
positive role in bone formation, just not via collagen deposition. In the developing face the TGF-
βs and BMPs regulate maxillary mesenchymal cell proliferation and extracellular matrix
synthesis (164). However, the exact mechanism by which CTHRC1 regulates collagen matrix
deposition through TGF-β is not entirely clear
53
Another mechanism by which CTHRC1 may regulate collagen deposition and chondrogenesis is
by its interaction with Wnt ligands. It has been suggested that CTHRC1 interacts with
components of the Wnt/PCP pathway by forming a CTHRC1-Wnt-Fzd/Ror2 complex to
promote activation of this pathway (94). The WNT5A ligand also binds the Ror2 receptor and
activates PCP pathway (165). WNT5A is expressed at high levels in mandibular chondrogenesis
in the chicken embryo in Meckel’s cartilage (113). Results from this thesis also show that
CTHRC1 is expressed in Meckel’s cartilage. WNT5A regulates collagen matrix stability, thereby
promoting chondrogenesis via suppression of the canonical Wnt pathway and promoting the PCP
pathway in cartilage blastema which is needed for chondrogenesis (113). The spatial expression
pattern of WNT5A in the chicken embryo at stage 24 in the mesenchyme of the frontonasal
mass, lateral nasal and maxillary process has a similar pattern as what was seen for CTHRC1 at
stage E10.5 in the mouse embryo with respect to distribution near the epithelia (109). Based on
the similarities in spatial expression of WNT5A and CTHRC1 in Meckel’s cartilage and the
mesenchyme of the midface, the role of WNT5A in promoting collagen matrix stability and the
proposed role of CTHRC1 stabilizing the Wnt/Ror2 interaction, it is possible that CTHRC1
regulates collagen deposition through an interaction with WNT5A.
Overall the results from the IHC and qPCR data of this study support a role for CTHRC1 in
chondrogenic condensation, as seen at E10.5 and 11.5 in the maxilla and E12.5 in Meckel’s
cartilage as well as differentiation of chondrocytes. The results also support a positive role for
deposition of collagen type II since the CTHRCI expression is seen in areas of future cartilage
formation. However, since past studies show that CTHRC1 actually downregulates expression of
type I collagen mRNA and inhibits its deposition, it may have a negative role for this type of
collagen. The exact mechanism by which CTHRC1 regulates collagen deposition and
chondrogenic condensation is not known but several possibilities have been put forth in the
preceding paragraphs.
4.2.4 Role of Cthrc1 in bone formation
Our data of a possible function for CTHRC1 in cartilage formation would support a role in
endochondral bone formation, where cartilage is replaced by bone. We speculate that CTHRC1,
being highly expressed at sites of intramembranous ossification in the craniofacial region, e.g.,
54
body of the mandible and calvarium, plays a direct role in osteogenesis in the developing
craniofacial region.
Studies have shown via transgenic and Cthrc1 null mice that CTHRC1 is a positive regulator of
osteoblastic bone formation postnatally (135). Even though these studies were done on postnatal
mice, the positive effect on osteoblasts would warrant further investigation into the effects of
CTHRC1 in osteoblasts during development. Our results show the presence of CTHRC1 protein
in areas of intramembranous ossification in the midface, mandible and skull of E12.5 and E13.5
mouse embryos. At E13.5, CTHRC1 protein expression was seen in the forming bones of the
skull in the posterior calvarium which forms by intramembranous ossification. This presence of
CTHRC1 in areas of intramembranous ossification in addition to previous studies in postnatal
mice supports a role for CTHRC1 in osteoblastic bone formation and regulation in the embryo.
Another function of CTHRC1 in bone formation in development may be coupling of bone
formation and resorption. CTHRC1 has been shown to couple bone formation and resorption. It
is secreted by osteoclasts in postnatal mice (119). CTHRC1 secreted by osteoclasts targets
stromal cells to stimulate osteogenesis and is increased in high bone turnover states and
decreased in conditions associated with suppressed bone turnover. An osteoclast specific deletion
of Cthrc1 induced osteopenia due to reduced bone formation (166). These studies as well as the
expression of CTHRC1 from the experiments in this thesis, support a role for CTHRC1 as a
stimulatory signal for stromal cells to stimulate osteogenesis at E12.5 and E13.5 in areas of
intramembranous ossification.
Again, the mechanism by which CTHRC1 is involved in bone formation is unclear and can only
be speculated on based on the current studies and the results of this thesis. It is known that
CTHRC1 is regulated by BMP-4, BMP-2 and TGF-β, all of which have known roles in bone
formation as well as midface development. CTHRC1 is a downstream target of BMP-2 in
osteochondroprogenitor-like cells and is also present in bone cells in vivo. Other candidates in
the mechanism of CTHRC1 regulation of bone formation are Wnt ligands. We previously
discussed the involvement of WNT5A in chondrogenesis and its possible interaction with
CTHRC1. In vitro studies have shown that when combined with WNT3A, a bone anabolic
protein, CTHRC1 was capable of stimulating chemotaxis of bone-marrow derived stromal cells
(119), which is consistent with previous studies that show cross-talk between the Wnt pathway
55
and CTHRC1 (94). Therefore, CTHRC1 may interact with WNT3A to stimulate chemotaxis of
stromal cells involved in bone formation. Wnt3a is known to control the fate of both
mesenchymal cells and NCCs in the craniofacial processes (167) and regulates palatal fusion in
animal models (168). Also, targeted deletions of Wnt3a in mouse models caused death up birth
due to mandibular defects (109) however at this time no studies have shown WNT3A protein
expression in the developing midface.
During craniofacial growth, CTHRC1 may play a role in bone formation by being directly
involved in regulating the differentiation of mesenchyme into preosteoblast cells. In vitro studies
showed that forced Cthrc1 expression in osteoblastic cells stimulated osteoblast differentiation
(166) and CTHRC1 protein stimulated differentiation and mineralization of osteoprogenitor cells
and osteoblasts (135). Interestingly, differentiation of preosteoblasts precedes condensation
which amplifies the number of osteogenic cells in contrast to chondrogenesis where
condensation precedes the appearance of chondrogenic cells (153). It is thus logical to infer that
activation of differentiation to the osteogenic pathway is occurring in osteoblasts just prior to and
around the time of osteogenic condensation in the areas lateral to Meckel’s cartilage and in the
bones of the maxilla and skull.
Although there are few studies on the effect of CTHRC1 on bone formation, the present
literature and results from this thesis support a role for CTHRC1 in osteogenic condensation,
osteoblastic bone formation, coupling of bone resorption and formation and differentiation of
osteoblasts in the developing midface. No studies to date have looked at differences in
differentiation and mineralization of theses cell types in the developing midface during
craniofacial formation in the mouse embryo in the absence or presence of CTHRC1. Most
transgenic and knockout studies have focused on the absence or presence of gross abnormalities
in Cthrc1 transgenic and knockout mice at birth, without looking at changes in osteogenesis
during development. Further studies need to be done of mice embryos during development
especially in the area of the midface to determine the exact role of CTHRC1 in bone formation
during this crucial time of craniofacial formation.
4.2.5 Summary of role of Cthrc1 in midface development
In summary, the results from this thesis support four main roles for CTHRC1 in midface
development of the mouse embryo at critical stages in midface development; a role in collagen
56
formation of the midface, a role in bone formation of the midface, a role in epithelial-
mesenchyme interactions and lastly a role in cell migration of CNCCs to the facial prominences.
Measurement of the Cthrc1 mRNA transcripts and analysis of spatial expression of the CTHRC1
protein in the developing midface allowed us to draw these conclusions in addition to previous
studies supporting these roles.
4.3 Future Directions for Cthrc1 studies
The results from this thesis show the temporal and spatial patterns of expression of Cthrc1
mRNA transcripts and CTHRC1 protein at critical stages of midface development in the murine
embryo. These results show changes in the CD1 wild-type mouse during normal development.
4.3.1 Midface development in Cthrc1 knockout models
Several knockout mouse models for Cthrc1 have been generated (94, 119, 122, 135, 169). Most
of these studies have not looked at changes in midface development during embryogenesis.
Therefore, future directions of this project would be to study in greater detail the formation of the
midface in Cthrc1 knockout mice compared to wild type mice. Future studies involving the use
of Cthrc1 knockout models can address the effects of an absent Cthrc1 gene specifically on
craniofacial growth and development. For example, studies looking at the amount and quality of
collagen formation in the midface in knockout mice compared to wild-type animals could
evaluate the role of Cthrc1 in chondrogenic condensation and collagen deposition. An
examination of the architecture of midfacical structures such as snout length, size of nasal
capsule and presence of abnormalities such as clefting, and changes in structures requiring
epithelial-mesenchyme interactions such as teeth would give more information of the role of
Cthrc1 during craniofacial development. The focus until now has been on postnatal changes after
Cthrc1 knockout or upregulation in transgenic mice, with any observations on the effects in
embryos being on a gross-anatomical level. Studies on changes in the midface of mice embryos
at the histological and molecular level as a result of Cthrc1 knockout will give more detailed
information as to its role in midface development.
57
4.3.2 Cthrc1 expression in mouse models specifically targeting effects on midface development
The expression of Cthrc1 mRNA and protein during development (122) and postnatally (119,
124, 135) has been shown in several studies, including this thesis. Numerous mouse models with
deletions, mutations or overexpression of specific genes resulting in craniofacial malformations
have been mentioned in the literature (170-173). For example, deletions of the Wnt3a gene
results in mandibular defects, deletion of Wnt9b results in cleft lip and deletion of Wnt5a results
in the truncation of upper and lower jaws. Deletions of the Tcof1 gene produce a mouse model of
Treacher-Collins syndrome (63, 174). IHC and qPCR of the midface region for Cthrc1 mRNA
and protein in these mouse models compared with what is seen in wild-type mice would show
any changes in Cthrc1 expression during specific syndromes. This may help further support or
disprove already known roles of Cthrc1 during midface development.
4.3.3 Colocalization studies of CTHRC1 with other proteins with a known function in midface development
Many studies exist showing the role of certain genes and proteins in craniofacial development.
Examples are Wnt ligands (109) and Flrt 2 and Flrt3 (175). A colocalization study of CTHRC1
protein expression with other well-known proteins in craniofacial development would allow
comparisons between expression patterns and may help elucidate further roles of CTHRC1 in
midface development by similarities or dissimilarities in the expression patterns. For example,
our results of CTHRC1 expression at E10.5 in the mesenchyme of the facial processes show a
markedly similar expression as Wnt5A mRNA in the mesenchyme of maxillary and mandibular
processes of stage 24 chicken embryos (109). Wnt5A has a role in promoting chondrogenesis
and also has a role in outgrowth of the facial prominences and palate formation (104, 176). The
same comparison could be done at the level of mRNA transcript expression. This would show
any similarities in gene expression patterns with other well-known players in craniofacial
development such as Bmps or Fgf genes.
58
4.4 Conclusions
The results obtained so far are consistent with the original hypothesis of this thesis that Cthrc1
plays a regulatory role in cell migration, collagen formation and bone formation of the midface.
Based on our results and studies in the literature there is also evidence for Cthrc1 having a role in
epithelial-mesenchyme interactions during midface formation in the developing craniofacial
region.
59
References
1. Cordero DR, Brugmann S, Chu Y, Bajpai R, Jame M, Helms JA. Cranial neural crest cells on the move: their roles in craniofacial development. Am J Med Genet A. 2011;155a(2):270-9. 2. Kuratani S. Cephalic neural crest cells and the evolution of craniofacial structures in vertebrates: morphological and embryological significance of the premandibular-mandibular boundary. Zoology (Jena). 2005;108(1):13-25. 3. Gong SG. Cranial neural crest: Migratory cell behavior and regulatory networks. Exp Cell Res. 2014. 4. Cordero D, Marcucio R, Hu D, Gaffield W, Tapadia M, Helms JA. Temporal perturbations in sonic hedgehog signaling elicit the spectrum of holoprosencephaly phenotypes. J Clin Invest. 2004;114(4):485-94. 5. Creuzet S, Couly G, Le Douarin NM. Patterning the neural crest derivatives during development of the vertebrate head: insights from avian studies. J Anat. 2005;207(5):447-59. 6. Sandell LL, Trainor PA. Neural crest cell plasticity. size matters. Adv Exp Med Biol. 2006;589:78-95. 7. Reid BS, Yang H, Melvin VS, Taketo MM, Williams T. Ectodermal Wnt/beta-catenin signaling shapes the mouse face. Dev Biol. 2011;349(2):261-9. 8. Minoux M, Rijli FM. Molecular mechanisms of cranial neural crest cell migration and patterning in craniofacial development. Development. 2010;137(16):2605-21. 9. Basch ML, Bronner-Fraser M. Neural crest inducing signals. Adv Exp Med Biol. 2006;589:24-31. 10. Steventon B, Carmona-Fontaine C, Mayor R. Genetic network during neural crest induction: from cell specification to cell survival. Semin Cell Dev Biol. 2005;16(6):647-54. 11. Theveneau E, Mayor R. Neural crest delamination and migration: from epithelium-to-mesenchyme transition to collective cell migration. Dev Biol. 2012;366(1):34-54. 12. Meulemans D, Bronner-Fraser M. Gene-regulatory interactions in neural crest evolution and development. Dev Cell. 2004;7(3):291-9. 13. Huang X, Saint-Jeannet JP. Induction of the neural crest and the opportunities of life on the edge. Dev Biol. 2004;275(1):1-11. 14. Mayor R, Theveneau E. The neural crest. Development. 2013;140(11):2247-51. 15. Stuhlmiller TJ, Garcia-Castro MI. Current perspectives of the signaling pathways directing neural crest induction. Cell Mol Life Sci. 2012;69(22):3715-37. 16. Thiery JP, Sleeman JP. Complex networks orchestrate epithelial-mesenchymal transitions. Nat Rev Mol Cell Biol. 2006;7(2):131-42. 17. Noden DM, Trainor PA. Relations and interactions between cranial mesoderm and neural crest populations. J Anat. 2005;207(5):575-601. 18. Radisky DC, LaBarge MA. Epithelial-mesenchymal transition and the stem cell phenotype. Cell Stem Cell. 2008;2(6):511-2. 19. Ahlstrom JD, Erickson CA. New views on the neural crest epithelial-mesenchymal transition and neuroepithelial interkinetic nuclear migration. Commun Integr Biol. 2009;2(6):489-93. 20. Trainor PA. Craniofacial birth defects: The role of neural crest cells in the etiology and pathogenesis of Treacher Collins syndrome and the potential for prevention. Am J Med Genet A. 2010;152a(12):2984-94. 21. Kulesa PM, Fraser SE. In ovo time-lapse analysis of chick hindbrain neural crest cell migration shows cell interactions during migration to the branchial arches. Development. 2000;127(6):1161-72. 22. Carmona-Fontaine C, Matthews HK, Kuriyama S, Moreno M, Dunn GA, Parsons M, et al. Contact inhibition of locomotion in vivo controls neural crest directional migration. Nature. 2008;456(7224):957-61.
60
23. Teddy JM, Kulesa PM. In vivo evidence for short- and long-range cell communication in cranial neural crest cells. Development. 2004;131(24):6141-51. 24. Kulesa PM, Lu CC, Fraser SE. Time-lapse analysis reveals a series of events by which cranial neural crest cells reroute around physical barriers. Brain Behav Evol. 2005;66(4):255-65. 25. Le Douarin NM, Dupin E. The neural crest in vertebrate evolution. Curr Opin Genet Dev. 2012;22(4):381-9. 26. Graham A. Deconstructing the pharyngeal metamere. J Exp Zool B Mol Dev Evol. 2008;310(4):336-44. 27. Kuratani S, Matsuo I, Aizawa S. Developmental patterning and evolution of the mammalian viscerocranium: genetic insights into comparative morphology. Dev Dyn. 1997;209(2):139-55. 28. Santagati F, Minoux M, Ren SY, Rijli FM. Temporal requirement of Hoxa2 in cranial neural crest skeletal morphogenesis. Development. 2005;132(22):4927-36. 29. Kubota Y, Ito K. Chemotactic migration of mesencephalic neural crest cells in the mouse. Dev Dyn. 2000;217(2):170-9. 30. Davy A, Soriano P. Ephrin-B2 forward signaling regulates somite patterning and neural crest cell development. Dev Biol. 2007;304(1):182-93. 31. Lalani SR, Safiullah AM, Molinari LM, Fernbach SD, Martin DM, Belmont JW. SEMA3E mutation in a patient with CHARGE syndrome. J Med Genet. 2004;41(7):e94. 32. Hunt P, Whiting J, Nonchev S, Sham MH, Marshall H, Graham A, et al. The branchial Hox code and its implications for gene regulation, patterning of the nervous system and head evolution. Development. 1991;Suppl 2:63-77. 33. Lumsden A, Krumlauf R. Patterning the vertebrate neuraxis. Science. 1996;274(5290):1109-15. 34. Couly G, Grapin-Botton A, Coltey P, Ruhin B, Le Douarin NM. Determination of the identity of the derivatives of the cephalic neural crest: incompatibility between Hox gene expression and lower jaw development. Development. 1998;125(17):3445-59. 35. Hunt P, Clarke JD, Buxton P, Ferretti P, Thorogood P. Segmentation, crest prespecification and the control of facial form. Eur J Oral Sci. 1998;106 Suppl 1:12-8. 36. Trainor PA, Ariza-McNaughton L, Krumlauf R. Role of the isthmus and FGFs in resolving the paradox of neural crest plasticity and prepatterning. Science. 2002;295(5558):1288-91. 37. Trainor P, Krumlauf R. Plasticity in mouse neural crest cells reveals a new patterning role for cranial mesoderm. Nat Cell Biol. 2000;2(2):96-102. 38. Gendron-Maguire M, Mallo M, Zhang M, Gridley T. Hoxa-2 mutant mice exhibit homeotic transformation of skeletal elements derived from cranial neural crest. Cell. 1993;75(7):1317-31. 39. Pasqualetti M, Ori M, Nardi I, Rijli FM. Ectopic Hoxa2 induction after neural crest migration results in homeosis of jaw elements in Xenopus. Development. 2000;127(24):5367-78. 40. Depew MJ, Simpson CA, Morasso M, Rubenstein JL. Reassessing the Dlx code: the genetic regulation of branchial arch skeletal pattern and development. J Anat. 2005;207(5):501-61. 41. Qiu M, Bulfone A, Martinez S, Meneses JJ, Shimamura K, Pedersen RA, et al. Null mutation of Dlx-2 results in abnormal morphogenesis of proximal first and second branchial arch derivatives and abnormal differentiation in the forebrain. Genes Dev. 1995;9(20):2523-38. 42. Depew MJ, Liu JK, Long JE, Presley R, Meneses JJ, Pedersen RA, et al. Dlx5 regulates regional development of the branchial arches and sensory capsules. Development. 1999;126(17):3831-46. 43. Acampora D, Merlo GR, Paleari L, Zerega B, Postiglione MP, Mantero S, et al. Craniofacial, vestibular and bone defects in mice lacking the Distal-less-related gene Dlx5. Development. 1999;126(17):3795-809. 44. del Barrio MG, Nieto MA. Overexpression of Snail family members highlights their ability to promote chick neural crest formation. Development. 2002;129(7):1583-93. 45. Graham A. Development of the pharyngeal arches. Am J Med Genet A. 2003;119a(3):251-6.
61
46. Couly G, Creuzet S, Bennaceur S, Vincent C, Le Douarin NM. Interactions between Hox-negative cephalic neural crest cells and the foregut endoderm in patterning the facial skeleton in the vertebrate head. Development. 2002;129(4):1061-73. 47. Ruhin B, Creuzet S, Vincent C, Benouaiche L, Le Douarin NM, Couly G. Patterning of the hyoid cartilage depends upon signals arising from the ventral foregut endoderm. Dev Dyn. 2003;228(2):239-46. 48. Rinon A, Lazar S, Marshall H, Buchmann-Moller S, Neufeld A, Elhanany-Tamir H, et al. Cranial neural crest cells regulate head muscle patterning and differentiation during vertebrate embryogenesis. Development. 2007;134(17):3065-75. 49. Grenier J, Teillet MA, Grifone R, Kelly RG, Duprez D. Relationship between neural crest cells and cranial mesoderm during head muscle development. PLoS One. 2009;4(2):e4381. 50. Rossant J, Tam PPL. Mouse Development
Patterning, Morphogenesis, and Organogenesis. San Diego: Academic Press [Imprint]
Elsevier Science & Technology Books; 2002. Available from: http://myaccess.library.utoronto.ca/login?url=http://books.scholarsportal.info/viewdoc.html?id=/ebooks/ebooks0/elsevier/2009-12-02/2/9780125979511Available from: http://myaccess.library.utoronto.ca/login?url=http://www.sciencedirect.com/science/book/9780125979511. 51. Moore KL, Persaud TVN. Before we are born : essentials of embryology and birth defects. 6th ed. Philadelphia: Saunders; 2003. xv, 448 p. p. 52. Kaufman M. The Atlas of Mouse Development. Revised Edition ed. London: Academic Press Limited; 1992. 53. Helms JA, Cordero D, Tapadia MD. New insights into craniofacial morphogenesis. Development. 2005;132(5):851-61. 54. Proffit WR, Fields HW, Sarver DM. Contemporary orthodontics. 5th ed. St. Louis, Mo.: Elsevier/Mosby; 2013. xiii, 754 p. p. 55. Dixon A, Hoyte D, Ronning O. Fundamentals of Craniofacial Growth. Didier D, editor. New York: CRC Press; 1997. 56. Pavlov MI, Sautier JM, Oboeuf M, Asselin A, Berdal A. Chondrogenic differentiation during midfacial development in the mouse: in vivo and in vitro studies. Biol Cell. 2003;95(2):75-86. 57. Nanci A, Ten Cate AR. Ten Cate's oral histology : development, structure, and function. 8th ed. St. Louis, Mo.: Elsevier; 2013. xiii, 379 p. p. 58. Kollar EJ, Baird GR. Tissue interactions in embryonic mouse tooth germs. II. The inductive role of the dental papilla. J Embryol Exp Morphol. 1970;24(1):173-86. 59. Braybrook C, Warry G, Howell G, Mandryko V, Arnason A, Bjornsson A, et al. Physical and transcriptional mapping of the X-linked cleft palate and ankyloglossia (CPX) critical region. Hum Genet. 2001;108(6):537-45. 60. Lan Y, Ryan RC, Zhang Z, Bullard SA, Bush JO, Maltby KM, et al. Expression of Wnt9b and activation of canonical Wnt signaling during midfacial morphogenesis in mice. Dev Dyn. 2006;235(5):1448-54. 61. Brugmann SA, Tapadia MD, Helms JA. The molecular origins of species-specific facial pattern. Curr Top Dev Biol. 2006;73:1-42. 62. Miettinen PJ, Chin JR, Shum L, Slavkin HC, Shuler CF, Derynck R, et al. Epidermal growth factor receptor function is necessary for normal craniofacial development and palate closure. Nat Genet. 1999;22(1):69-73.
62
63. Dixon J, Jones NC, Sandell LL, Jayasinghe SM, Crane J, Rey JP, et al. Tcof1/Treacle is required for neural crest cell formation and proliferation deficiencies that cause craniofacial abnormalities. Proc Natl Acad Sci U S A. 2006;103(36):13403-8. 64. Anderson RM, Stottmann RW, Choi M, Klingensmith J. Endogenous bone morphogenetic protein antagonists regulate mammalian neural crest generation and survival. Dev Dyn. 2006;235(9):2507-20. 65. Mayor R, Guerrero N, Martinez C. Role of FGF and noggin in neural crest induction. Dev Biol. 1997;189(1):1-12. 66. Garcia-Castro MI, Marcelle C, Bronner-Fraser M. Ectodermal Wnt function as a neural crest inducer. Science. 2002;297(5582):848-51. 67. Hu D, Helms JA. The role of sonic hedgehog in normal and abnormal craniofacial morphogenesis. Development. 1999;126(21):4873-84. 68. Sperber GH, Guttmann GD, Sperber SM. Craniofacial embryogenetics and development. 2nd ed. Shelton, CT: People's Medical Pub. House USA; 2010. 250 p. p. 69. Zoupa M, Seppala M, Mitsiadis T, Cobourne MT. Tbx1 is expressed at multiple sites of epithelial-mesenchymal interaction during early development of the facial complex. Int J Dev Biol. 2006;50(5):504-10. 70. Walker MB, Trainor PA. Craniofacial malformations: intrinsic vs extrinsic neural crest cell defects in Treacher Collins and 22q11 deletion syndromes. Clin Genet. 2006;69(6):471-9. 71. Kimmel CB, Ullmann B, Walker M, Miller CT, Crump JG. Endothelin 1-mediated regulation of pharyngeal bone development in zebrafish. Development. 2003;130(7):1339-51. 72. Szabo-Rogers HL, Geetha-Loganathan P, Nimmagadda S, Fu KK, Richman JM. FGF signals from the nasal pit are necessary for normal facial morphogenesis. Dev Biol. 2008;318(2):289-302. 73. Trumpp A, Depew MJ, Rubenstein JL, Bishop JM, Martin GR. Cre-mediated gene inactivation demonstrates that FGF8 is required for cell survival and patterning of the first branchial arch. Genes Dev. 1999;13(23):3136-48. 74. Blentic A, Tandon P, Payton S, Walshe J, Carney T, Kelsh RN, et al. The emergence of ectomesenchyme. Dev Dyn. 2008;237(3):592-601. 75. Grigoriou M, Tucker AS, Sharpe PT, Pachnis V. Expression and regulation of Lhx6 and Lhx7, a novel subfamily of LIM homeodomain encoding genes, suggests a role in mammalian head development. Development. 1998;125(11):2063-74. 76. Tucker AS, Yamada G, Grigoriou M, Pachnis V, Sharpe PT. Fgf-8 determines rostral-caudal polarity in the first branchial arch. Development. 1999;126(1):51-61. 77. Abe E, Yamamoto M, Taguchi Y, Lecka-Czernik B, O'Brien CA, Economides AN, et al. Essential requirement of BMPs-2/4 for both osteoblast and osteoclast formation in murine bone marrow cultures from adult mice: antagonism by noggin. J Bone Miner Res. 2000;15(4):663-73. 78. Hughes FJ, Collyer J, Stanfield M, Goodman SA. The effects of bone morphogenetic protein-2, -4, and -6 on differentiation of rat osteoblast cells in vitro. Endocrinology. 1995;136(6):2671-7. 79. Bandyopadhyay A, Yadav PS, Prashar P. BMP signaling in development and diseases: a pharmacological perspective. Biochem Pharmacol. 2013;85(7):857-64. 80. Lan Y, Jiang R. Sonic hedgehog signaling regulates reciprocal epithelial-mesenchymal interactions controlling palatal outgrowth. Development. 2009;136(8):1387-96. 81. Dudas M, Sridurongrit S, Nagy A, Okazaki K, Kaartinen V. Craniofacial defects in mice lacking BMP type I receptor Alk2 in neural crest cells. Mech Dev. 2004;121(2):173-82. 82. Shigetani Y, Nobusada Y, Kuratani S. Ectodermally derived FGF8 defines the maxillomandibular region in the early chick embryo: epithelial-mesenchymal interactions in the specification of the craniofacial ectomesenchyme. Dev Biol. 2000;228(1):73-85. 83. Haworth KE, Healy C, Morgan P, Sharpe PT. Regionalisation of early head ectoderm is regulated by endoderm and prepatterns the orofacial epithelium. Development. 2004;131(19):4797-806.
63
84. Haworth KE, Wilson JM, Grevellec A, Cobourne MT, Healy C, Helms JA, et al. Sonic hedgehog in the pharyngeal endoderm controls arch pattern via regulation of Fgf8 in head ectoderm. Dev Biol. 2007;303(1):244-58. 85. Albertson RC, Yelick PC. Fgf8 haploinsufficiency results in distinct craniofacial defects in adult zebrafish. Dev Biol. 2007;306(2):505-15. 86. Ahlgren SC, Bronner-Fraser M. Inhibition of sonic hedgehog signaling in vivo results in craniofacial neural crest cell death. Curr Biol. 1999;9(22):1304-14. 87. Washington Smoak I, Byrd NA, Abu-Issa R, Goddeeris MM, Anderson R, Morris J, et al. Sonic hedgehog is required for cardiac outflow tract and neural crest cell development. Dev Biol. 2005;283(2):357-72. 88. Brito JM, Teillet MA, Le Douarin NM. An early role for sonic hedgehog from foregut endoderm in jaw development: ensuring neural crest cell survival. Proc Natl Acad Sci U S A. 2006;103(31):11607-12. 89. Eberhart JK, Swartz ME, Crump JG, Kimmel CB. Early Hedgehog signaling from neural to oral epithelium organizes anterior craniofacial development. Development. 2006;133(6):1069-77. 90. Wada N, Javidan Y, Nelson S, Carney TJ, Kelsh RN, Schilling TF. Hedgehog signaling is required for cranial neural crest morphogenesis and chondrogenesis at the midline in the zebrafish skull. Development. 2005;132(17):3977-88. 91. Hu D, Marcucio RS, Helms JA. A zone of frontonasal ectoderm regulates patterning and growth in the face. Development. 2003;130(9):1749-58. 92. Hu D, Marcucio RS. Unique organization of the frontonasal ectodermal zone in birds and mammals. Dev Biol. 2009;325(1):200-10. 93. Wu P, Jiang TX, Shen JY, Widelitz RB, Chuong CM. Morphoregulation of avian beaks: comparative mapping of growth zone activities and morphological evolution. Dev Dyn. 2006;235(5):1400-12. 94. Yamamoto S, Nishimura O, Misaki K, Nishita M, Minami Y, Yonemura S, et al. Cthrc1 Selectively Activates the Planar Cell Polarity Pathway of Wnt Signaling by Stabilizing the Wnt-Receptor Complex. Developmental Cell. 2008;15(1):23-36. 95. Miao CG, Yang YY, He X, Li XF, Huang C, Huang Y, et al. Wnt signaling pathway in rheumatoid arthritis, with special emphasis on the different roles in synovial inflammation and bone remodeling. Cell Signal. 2013;25(10):2069-78. 96. Logan CY, Nusse R. The Wnt signaling pathway in development and disease. Annu Rev Cell Dev Biol. 2004;20:781-810. 97. Krause U, Gregory CA. Potential of modulating Wnt signaling pathway toward the development of bone anabolic agent. Curr Mol Pharmacol. 2012;5(2):164-73. 98. De A. Wnt/Ca2+ signaling pathway: a brief overview. Acta Biochim Biophys Sin (Shanghai). 2011;43(10):745-56. 99. Sheldahl LC, Park M, Malbon CC, Moon RT. Protein kinase C is differentially stimulated by Wnt and Frizzled homologs in a G-protein-dependent manner. Curr Biol. 1999;9(13):695-8. 100. Kuhl M, Sheldahl LC, Park M, Miller JR, Moon RT. The Wnt/Ca2+ pathway: a new vertebrate Wnt signaling pathway takes shape. Trends Genet. 2000;16(7):279-83. 101. Dale RM, Sisson BE, Topczewski J. The emerging role of Wnt/PCP signaling in organ formation. Zebrafish. 2009;6(1):9-14. 102. Karner C, Wharton KA, Carroll TJ. Apical-basal polarity, Wnt signaling and vertebrate organogenesis. Semin Cell Dev Biol. 2006;17(2):214-22. 103. Karner C, Wharton KA, Jr., Carroll TJ. Planar cell polarity and vertebrate organogenesis. Semin Cell Dev Biol. 2006;17(2):194-203. 104. He F, Xiong W, Yu X, Espinoza-Lewis R, Liu C, Gu S, et al. Wnt5a regulates directional cell migration and cell proliferation via Ror2-mediated noncanonical pathway in mammalian palate development. Development. 2008;135(23):3871-9.
64
105. Cadigan KM, Nusse R. Wnt signaling: a common theme in animal development. Genes Dev. 1997;11(24):3286-305. 106. Grigoryan T, Wend P, Klaus A, Birchmeier W. Deciphering the function of canonical Wnt signals in development and disease: conditional loss- and gain-of-function mutations of beta-catenin in mice. Genes Dev. 2008;22(17):2308-41. 107. Brault V, Moore R, Kutsch S, Ishibashi M, Rowitch DH, McMahon AP, et al. Inactivation of the beta-catenin gene by Wnt1-Cre-mediated deletion results in dramatic brain malformation and failure of craniofacial development. Development. 2001;128(8):1253-64. 108. Niemann S, Zhao C, Pascu F, Stahl U, Aulepp U, Niswander L, et al. Homozygous WNT3 mutation causes tetra-amelia in a large consanguineous family. Am J Hum Genet. 2004;74(3):558-63. 109. Geetha-Loganathan P, Nimmagadda S, Antoni L, Fu K, Whiting CJ, Francis-West P, et al. Expression of WNT signalling pathway genes during chicken craniofacial development. Dev Dyn. 2009;238(5):1150-65. 110. Chai Y, Maxson RE, Jr. Recent advances in craniofacial morphogenesis. Dev Dyn. 2006;235(9):2353-75. 111. Koay MA, Brown MA. Genetic disorders of the LRP5-Wnt signalling pathway affecting the skeleton. Trends Mol Med. 2005;11(3):129-37. 112. Kikuchi A, Yamamoto H, Sato A, Matsumoto S. New insights into the mechanism of Wnt signaling pathway activation. Int Rev Cell Mol Biol. 2011;291:21-71. 113. Hosseini-Farahabadi S, Geetha-Loganathan P, Fu K, Nimmagadda S, Yang HJ, Richman JM. Dual functions for WNT5A during cartilage development and in disease. Matrix Biol. 2013;32(5):252-64. 114. Person AD, Beiraghi S, Sieben CM, Hermanson S, Neumann AN, Robu ME, et al. WNT5A mutations in patients with autosomal dominant Robinow syndrome. Dev Dyn. 2010;239(1):327-37. 115. Pyagay P, Heroult M, Wang Q, Lehnert W, Belden J, Liaw L, et al. Collagen triple helix repeat containing 1, a novel secreted protein in injured and diseased arteries, inhibits collagen expression and promotes cell migration. Circulation Research. 2005;96(2):261-8. 116. Chen YL, Wang TH, Hsu HC, Yuan RH, Jeng YM. Overexpression of CTHRC1 in hepatocellular carcinoma promotes tumor invasion and predicts poor prognosis. PLoS One. 2013;8(7):e70324. 117. Lindner V. [cited 2014 May 4]. Available from: http://sackler.tufts.edu/Faculty-and-Research/Faculty-Research-Pages/~/media/Sackler/Page%20Images/Faculty%20Research%20Page%20Images/Lindner%20Fig%202.jpg. 118. Wang Y, Xu A, Knight C, Xu LY, Cooper GJ. Hydroxylation and glycosylation of the four conserved lysine residues in the collagenous domain of adiponectin. Potential role in the modulation of its insulin-sensitizing activity. J Biol Chem. 2002;277(22):19521-9. 119. Takeshita S, Fumoto T, Matsuoka K, Park KA, Aburatani H, Kato S, et al. Osteoclast-secreted CTHRC1 in the coupling of bone resorption to formation. J Clin Invest. 2013;123(9):3914-24. 120. Diez JJ, Iglesias P. The role of the novel adipocyte-derived hormone adiponectin in human disease. Eur J Endocrinol. 2003;148(3):293-300. 121. Liu G, Sengupta PK, Jamal B, Yang HY, Bouchie MP, Lindner V, et al. N-glycosylation induces the CTHRC1 protein and drives oral cancer cell migration. J Biol Chem. 2013;288(28):20217-27. 122. Durmus T, LeClair RJ, Park KS, Terzic A, Yoon JK, Lindner V. Expression analysis of the novel gene collagen triple helix repeat containing-1 (Cthrc1). Gene Expression Patterns. 2006;6(8):935-40. 123. LeClair R, Lindner V. The Role of Collagen Triple Helix Repeat Containing 1 in Injured Arteries, Collagen Expression, and Transforming Growth Factor beta Signaling. Trends in Cardiovascular Medicine. 2007;17(6):202-5. 124. LeClair RJ, Durmus T, Wang Q, Pyagay P, Terzic A, Lindner V. Cthrc1 is a novel inhibitor of transforming growth factor-beta signaling and neointimal lesion formation. Circulation Research. 2007;100(6):826-33.
65
125. Lindner V. Vascular repair processes mediated by transforming growth factor-beta. Z Kardiol. 2001;90 Suppl 3:17-22. 126. Janssens K, ten Dijke P, Janssens S, Van Hul W. Transforming growth factor-beta1 to the bone. Endocr Rev. 2005;26(6):743-74. 127. Ghosh AK. Factors involved in the regulation of type I collagen gene expression: implication in fibrosis. Exp Biol Med (Maywood). 2002;227(5):301-14. 128. Tang L, Dai DL, Su M, Martinka M, Li G, Zhou Y. Aberrant expression of collagen triple helix repeat containing 1 in human solid cancers. Clinical Cancer Research. 2006;12(12):3716-22. 129. Yang Y. Skeletal morphogenesis during embryonic development. Crit Rev Eukaryot Gene Expr. 2009;19(3):197-218. 130. Toy J, Yang JM, Leppert GS, Sundin OH. The optx2 homeobox gene is expressed in early precursors of the eye and activates retina-specific genes. Proc Natl Acad Sci U S A. 1998;95(18):10643-8. 131. Caetano-Lopes J, Canhao H, Fonseca JE. Osteoblasts and bone formation. Acta Reumatol Port. 2007;32(2):103-10. 132. Olsen BR, Reginato AM, Wang W. Bone development. Annu Rev Cell Dev Biol. 2000;16:191-220. 133. Wright E, Hargrave MR, Christiansen J, Cooper L, Kun J, Evans T, et al. The Sry-related gene Sox9 is expressed during chondrogenesis in mouse embryos. Nat Genet. 1995;9(1):15-20. 134. Harada S, Rodan GA. Control of osteoblast function and regulation of bone mass. Nature. 2003;423(6937):349-55. 135. Kimura H, Kwan KM, Zhang Z, Deng JM, Darnay BG, Behringer RR, et al. Cthrc1 is a positive regulator of osteoblastic bone formation. PLoS One. 2008;3(9):e3174. 136. Kimura H, Kwan KM, Zhang Z, Deng JM, Darnay BG, Behringer RR, et al. Cthrci is a positive regulator of osteoblastic bone formation. PLoS ONE. 2008;3(9). 137. Kim JH, Baek TH, Yim HS, Kim KH, Jeong SH, Kang HB, et al. Collagen triple helix repeat containing-1 (CTHRC1) expression in invasive ductal carcinoma of the breast: the impact on prognosis and correlation to clinicopathologic features. Pathol Oncol Res. 2013;19(4):731-7. 138. Leclair RJ, Wang Q, Benson MA, Prudovsky I, Lindner V. Intracellular localization of Cthrc1 characterizes differentiated smooth muscle. Arteriosclerosis, Thrombosis, and Vascular Biology. 2008;28(7):1332-8. 139. Kelley MW. Leading Wnt down a PCP Path: Cthrc1 Acts as a Coreceptor in the Wnt-PCP Pathway. Developmental Cell. 2008;15(1):7-8. 140. Apra C, Richard L, Coulpier F, Blugeon C, Gilardi-Hebenstreit P, Vallat JM, et al. Cthrc1 is a negative regulator of myelination in schwann cells. Glia. 2012;60(3):393-403. 141. Kharaishvili G, Cizkova M, Bouchalova K, Mgebrishvili G, Kolar Z, Bouchal J. Collagen triple helix repeat containing 1 protein, periostin and versican in primary and metastatic breast cancer: An immunohistochemical study. Journal of Clinical Pathology. 2011;64(11):977-82. 142. Turashvili G, Bouchal J, Baumforth K, Wei W, Dziechciarkova M, Ehrmann J, et al. Novel markers for differentiation of lobular and ductal invasive breast carcinomas by laser microdissection and microarray analysis. BMC Cancer. 2007;7(55). 143. Orloff M, Peterson C, He X, Ganapathi S, Heald B, Yang YR, et al. Germline mutations in MSR1, ASCC1, and CTHRC1 in patients with Barrett esophagus and esophageal adenocarcinoma. JAMA - Journal of the American Medical Association. 2011;306(4):410-9. 144. O'Connell MJ, Lavery I, Yothers G, Paik S, Clark-Langone KM, Lopatin M, et al. Relationship between tumor gene expression and recurrence in four independent studies of patients with stage II/III colon cancer treated with surgery alone or surgery plus adjuvant fluorouracil plus leucovorin. J Clin Oncol. 2010;28(25):3937-44. 145. Park M, Moon RT. The planar cell-polarity gene stbm regulates cell behaviour and cell fate in vertebrate embryos. Nat Cell Biol. 2002;4(1):20-5.
66
146. Li Y, Dudley AT. Noncanonical frizzled signaling regulates cell polarity of growth plate chondrocytes. Development. 2009;136(7):1083-92. 147. Kudryavtseva E, Forde TS, Pucker AD, Adarichev VA. Wnt signaling genes of murine chromosome 15 are involved in sex-affected pathways of inflammatory arthritis. Arthritis and rheumatism. 2012;64(4):1057-68. 148. Gong SG. Characterization of olfactory nerve abnormalities in Twirler mice. Differentiation. 2001;69(1):58-65. 149. Mort RL, Hay L, Jackson IJ. Ex vivo live imaging of melanoblast migration in embryonic mouse skin. Pigment Cell Melanoma Res. 23. England2010. p. 299-301. 150. Avery JK. Oral development and histology. 2nd ed. New York Stuttgart ; New York: Thieme Medical Publishers ; Georg Thieme Verlag; 1994. x, 422 p. p.
151. Mayor R, Theveneau E. The role of the non-canonical Wnt-planar cell polarity pathway in neural crest migration. Biochem J. 2014;457(1):19-26. 152. Stemple DL. Structure and function of the notochord: an essential organ for chordate development. Development. 2005;132(11):2503-12. 153. Hall BK, Miyake T. Divide, accumulate, differentiate: cell condensation in skeletal development revisited. Int J Dev Biol. 1995;39(6):881-93. 154. Francis-West P, Ladher R, Barlow A, Graveson A. Signalling interactions during facial development. Mech Dev. 1998;75(1-2):3-28. 155. Abzhanov A, Protas M, Grant BR, Grant PR, Tabin CJ. Bmp4 and morphological variation of beaks in Darwin's finches. Science. 2004;305(5689):1462-5. 156. Wu P, Jiang TX, Suksaweang S, Widelitz RB, Chuong CM. Molecular shaping of the beak. Science. 2004;305(5689):1465-6. 157. Abzhanov A, Tabin CJ. Shh and Fgf8 act synergistically to drive cartilage outgrowth during cranial development. Dev Biol. 2004;273(1):134-48. 158. Lan Y, Jia S, Jiang R. Molecular patterning of the mammalian dentition. Semin Cell Dev Biol. 2014;25-26:61-70. 159. King JA, Marker PC, Seung KJ, Kingsley DM. BMP5 and the molecular, skeletal, and soft-tissue alterations in short ear mice. Dev Biol. 1994;166(1):112-22. 160. Jones CM, Lyons KM, Hogan BL. Involvement of Bone Morphogenetic Protein-4 (BMP-4) and Vgr-1 in morphogenesis and neurogenesis in the mouse. Development. 1991;111(2):531-42. 161. Thesleff I, Vaahtokari A, Vainio S, Jowett A. Molecular mechanisms of cell and tissue interactions during early tooth development. Anat Rec. 1996;245(2):151-61. 162. Hall BK, Miyake T. The membranous skeleton: the role of cell condensations in vertebrate skeletogenesis. Anat Embryol (Berl). 1992;186(2):107-24. 163. Jiang TX, Yi JR, Ying SY, Chuong CM. Activin enhances chondrogenesis of limb bud cells: stimulation of precartilaginous mesenchymal condensations and expression of NCAM. Dev Biol. 1993;155(2):545-57. 164. Mukhopadhyay P, Greene RM, Pisano MM. Expression profiling of transforming growth factor beta superfamily genes in developing orofacial tissue. Birth Defects Res A Clin Mol Teratol. 2006;76(7):528-43. 165. Kikuchi A, Yamamoto H, Sato A, Matsumoto S. Wnt5a: its signalling, functions and implication in diseases. Acta Physiol (Oxf). 2012;204(1):17-33. 166. Takeshita S. An osteoclast-derived coupling factor. Osteoporosis International. 2011;22:S516-S7. 167. Yamaguchi TP, Takada S, Yoshikawa Y, Wu N, McMahon AP. T (Brachyury) is a direct target of Wnt3a during paraxial mesoderm specification. Genes Dev. 1999;13(24):3185-90.
67
168. Wan XZ, Qin WX, Tan N, Yao GF, Tan YT, Kuang WX, et al. CTHRC1 gene is overexpressed in human hepatocellular carcinomas and promotes the metastasis of human hepatocellular carcinoma cell MHCC97L. [Chinese]. Tumor. 2007;27(6):476-9+83. 169. Stohn JP, Perreault NG, Wang Q, Liaw L, Lindner V. Cthrc1, a novel hormone involved in regulation of metabolism. FASEB Journal. 2013;27. 170. Mishina Y, Snider TN. Neural crest cell signaling pathways critical to cranial bone development and pathology. Exp Cell Res. 2014. 171. Shukla V, Coumoul X, Wang RH, Kim HS, Deng CX. RNA interference and inhibition of MEK-ERK signaling prevent abnormal skeletal phenotypes in a mouse model of craniosynostosis. Nat Genet. 2007;39(9):1145-50. 172. Dab S, Sokhi R, Lee JC, Sessle BJ, Aubin JE, Gong SG. Characterization of esophageal defects in the Crouzon mouse model. Birth Defects Res A Clin Mol Teratol. 2013;97(9):578-86. 173. Kamiya N, Ye L, Kobayashi T, Mochida Y, Yamauchi M, Kronenberg HM, et al. BMP signaling negatively regulates bone mass through sclerostin by inhibiting the canonical Wnt pathway. Development. 2008;135(22):3801-11. 174. Dixon J, Brakebusch C, Fassler R, Dixon MJ. Increased levels of apoptosis in the prefusion neural folds underlie the craniofacial disorder, Treacher Collins syndrome. Hum Mol Genet. 2000;9(10):1473-80. 175. Gong SG, Mai S, Chung K, Wei K. Flrt2 and Flrt3 have overlapping and non-overlapping expression during craniofacial development. Gene Expr Patterns. 2009;9(7):497-502. 176. Yamaguchi TP, Bradley A, McMahon AP, Jones S. A Wnt5a pathway underlies outgrowth of multiple structures in the vertebrate embryo. Development. 1999;126(6):1211-23.
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