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Non-canonical Tgf-beta signaling in craniofacial development

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Non-canonical Tgf-beta signaling in craniofacial development

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NON-CANONICAL TGF-BETA SIGNALING PATHWAY
IN CRANIOFACIAL DEVELOPMENT

by

Jieun Kim

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CRANIOFACIAL BIOLOGY)

May 2009

Copyright 2009 Jieun Kim

ii
DEDICATION

This thesis in on the surface a summary of my research these past several years,
but upon closer look, it is much more than that. It represents the challenging
journey I have been fortunate to experience on so many different levels. I have
learned and grown so very much through it all; and most importantly, although
this is the end of this journey, it is in many ways just the beginning of “my
journey.” I am sincerely thankful to all of you for helping me achieve so much,
and laying the foundation for me to pursue further.

To my Dad in heaven and Mom, with Love.

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ACKNOWLEDGEMENTS

I am grateful for the good fortune I have had as a graduate student.
I would like to express my sincerest gratitude to my advisor Dr. Vesa
Kaartinen. I am very lucky to have him as my mentor. He has always been there
with an open mind, his office door always open to walk through. With thoughtful
patience, he has always listened to what I say, or sometimes try to say, always
giving me advice. He has never hesitated in giving me his guidance, enthusiasm
and encouragement.
I also deeply want to thank all the members of my dissertation committee:
Drs. Malcolm Snead, Saverio Bellusci, Wei Shi and Yang Chai. They gave me
positive and cheerful encouragement as well as generous, thoughtful and
excellent advice.
I would like to thank everyone from Dr. Vesa Kaartinen’s lab and the
Saban research institute, especially Marek Dudas, Andre Nagy, Somyoth
Sridurongrit (Ott), Wai-Yee Li and Penny Thomas. Because of them, my time
here was that much more enjoyable
Finally, I also want to thank James, my two brothers, Hungjung and
Yeounjung and all my family for all their support.

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Table of Contents

Dedication --------------------------------------------------------------------------------

Acknowledgements --------------------------------------------------------------------

List of Tables -----------------------------------------------------------------------------

List of Figures ----------------------------------------------------------------------------

Abstract -------------------------------------------------------------------------------------

Chapter I. Introduction ----------------------------------------------------------------
1-1. Embryonic craniofacial development ---------------------------------------
1-2. Neural crest -----------------------------------------------------------------------
1-3. Several signaling pathways involved in the craniofacial development
1-4. Tgf-β signaling ---------------------------------------------------------------------
1-5. Tgf-β ligands and receptors ------------------------------------------------------
1-6. Canonical Tgf-β signaling pathway -------------------------------------------
1-7. Regulation of Tgf-β signaling pathway ---------------------------------------
1-8. Non-canonical Tgf-β signaling pathway -------------------------------------
1-9. Chaper I References -------------------------------------------------------------

Chapter II. Unconventional Tgf-ß receptors combination -----------------
2-1. Summary ---------------------------------------------------------------------------
2-2. Introduction ------------------------------------------------------------------------
2-3. Materials and Methods ----------------------------------------------------------
2-3-1. Preparation of chimeric Tgf-β receptor constructs -------------------
2-3-2. Cell culture, co-transfection of constructs and treatment ----------
2-3-3. Western blot analyses ------------------------------------------------------
2-3-4. Whole-mount embryo analyses ------------------------------------------
2-4. Results ------------------------------------------------------------------------------
2-4-1. Expression of Alk5 and type II receptors ------------------------------
2-4-2. Alk5 can form functional signaling complexes with unconventional
type II receptors in vitro -----------------------------------------------------
2-5. Discussion ------------------------------------------------------------------------
2-6. Chapter II References ----------------------------------------------------------

Chapter III. Rac1 is indispensable in the craniofacial and cardiac
neural crest, but is not required for NCC migration in vivo
3-1. Summary --------------------------------------------------------------------------
3-2. Introduction -----------------------------------------------------------------------
3-3. Materials and methods ----------------------------------------------------------
3-3-1. Mouse breeding, genotyping and embryo collection ----------------
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3-3-2. Cell death and proliferation assay ---------------------------------------
3-3-3. Whole-mount embryo analysis -------------------------------------------
3-3-4. Neural tube culture -----------------------------------------------------------
3-3-5. Isolation of neural crest cells from 1st pharyngeal arch ------------
3-3-6. Histology and TEM analysis -----------------------------------------------
3-4. Results ------------------------------------------------------------------------------
3-4-1. Neural crest-specific Rac1 mutants -------------------------------------
3-4-2. Rac1/Wnt1-Cre mutants display severe craniofacial defects -----
3-4-3. Depletion of Rac1 in the neural crest leads to severe outflow
tract defects and embryonic death --------------------------------------
3-4-4. NCCs lacking Rac1 show defective cell migration and
spreading in vitro -------------------------------------------------------------
3-4-5. NCC survival, proliferation or rhombomere identity is not
impaired in Rac1/Wnt1-Cre mutants ------------------------------------
3-4-6. Rac1 is required for appropriate patterning and
differentiation of postmigratory NCCs in vivo --------------------------
3-4-7. The blistering cyst of the mutant embryos is caused by the
absence of the anchoring fibrils in the basement membrane ------
3-5. Discussion --------------------------------------------------------------------------
3-5-1. Epithelial blisters and mid-facial clefting --------------------------------
3-5-2. Rac1 deficiency and neural crest cell migration ----------------------
3-6. Chapter III References ------------------------------------------------------------

Chapter IV. Conditional and conventional mutants for Trim33 – the
Trim33 conventional knockout mice are embryonic lethal
4-1. Summary ----------------------------------------------------------------------------
4-2. Introduction --------------------------------------------------------------------------
4-3. Materials and Methods -----------------------------------------------------------
4-3-1. Construction of the targeting vector ---------------------------------------
4-3-2. Screening of correctly targeted embryonic stem cells ----------------
4-3-3. Generation of mutant mice -------------------------------------------------
4-3-4. Reverse transcription – PCR -----------------------------------------------
4-4. Results and Discussion -----------------------------------------------------------
4-4-1. Trim33 expression pattern during mouse embryogenesis ----------
4-4-2. Conventional and conditional knockout alleles for Trim33 -----------
4-4-3. The Trim33 null knockout cause embryonic lethality around E9.5 --
4-4-4. The Trim33 conditional knockout during E9 to E15 cause severe
embryonic axis formation defect and failure of neural tube closure
4-4-5. Male mosaic EIIa-cre mice can be efficiently used to segregate
the floxed alleles ---------------------------------------------------------------
4-6. Chapter IV References -----------------------------------------------------------

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Chapter V. Conclusions ------------------------------------------------------------
5-1. Alk5 forms functional signaling complexes with unconventional type II
receptors ------------------------------------------------------------------------------
5-2. Rac1 is indispensable in the craniofacial and cardiac neural crest,
but is not required for NCC migration in vivo --------------------------------
5-3. Trim33 conditional and conventional knockout mice were generated
- the Trim33 conventional knockout mice are embryonic lethal -----------
5-4. Chapter V References ------------------------------------------------------------

Bibliography ------------------------------------------------------------------------------

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LIST OF TABLES

Table 1-1. Endogeneous Fgf/Fgfr expressed during the development of the
craniofacial structures and their possible roles in developmental
disorders --------------------------------------------------------------------------- 17
Table 1-2. Tgf-β ligands --------------------------------------------------------------------- 22
Table. 4-1 Trim family proteins ------------------------------------------------------------- 84

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LIST OF FIGURES

Fig. 1-1. Facial formation in the early embryo stage -----------------------------
Fig. 1-2. Palatal formation in the mouse embryo -----------------------------------
Fig. 1-3. Tooth formation in the mouse embryo -------------------------------------
Fig. 1-4. Schematic chick and mouse skulls -----------------------------------------
Fig. 1-5. Regulatory steps in neural crest formation ------------------------------
Fig. 1-6. Fate map of neural crest cells along the body axis --------------------
Fig. 1-7. Cranial neural crest cells and their differentiation capacity ----------
Fig. 1-8. Reciprocal interaction between epithelium and mesenchyme
during tooth development --------------------------------------------------

Fig. 1-9. Canonical Tgf-β signaling pathway ---------------------------------------
Fig. 1-10. Rac1 in non-canonical Tgf-β signaling pathway ----------------------
Fig. 2-1. Conventional Tgf-β receptor combination -------------------------------
Fig. 2-2. Endogenous co-expression of Alk5 and type-II receptors ---------
Fig. 2-3. Activation of ALK5 by two different type II receptors in vitro -------
Fig. 3-1. Whole-mount in situ hybridization of with Rac1 RNA probe
with E10 embryos -----------------------------------------------------------

Fig. 3-2. Craniofacial defects in Rac1/Wnt1-Cre mutant embryos -----------
Fig. 3-3. Abnormal patterning of pharyngeal arch arteries and
outflow tract defects in Rac1/Wnt1-Cre embryos ---------------------

Fig.3-4. NCCs deficient in Rac1 failed to migrate in vitro, spread poorly and
showed a reduced amount of focal adhesion complexes -----------
Fig. 3-5. NCCs deficient in Rac1/Wnt-Cre migrate normally in vivo ---------

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Fig. 3-6. Normal apoptosis, cell proliferation and rhombomere
identity in Rac1/Wnt1-Cre mutants ----------------------------------------

Fig. 3-7. Abnormal patterning of cranial ganglia and smooth
muscle cells in Rac1/Wnt1-Cre ----------------------------------------------

Fig. 3-8. Abnormal connection between the basal lamina and the
underlying NC-derived mesenchyme in Rac1/Wnt1-Cre mutants ---

Fig. 4-1. Nucleic and amino acid sequences of hTIF1α (a) and hTIF1γ (b) -
Fig. 4-2. Defective Trim33 function cause abnormal hematopoiesis
in zebrafish (A) and human (B) -------------------------------------------
Fig. 4-3. TIF1γ as a branch in the TGFb-activated Smad pathway -----------
Fig. 4-4. Ectodermin is required for the formation of ectoderm
(A) by preventing Tgf-β and Bmp signaling in the ectoderm area (B)

Fig. 4-5. Breeding scheme to get the Trim flox and Trim null knockout mice
Fig. 4-6. Trim33 protein expression in the craniofacial area at E14 (A)
and thorax at E17 (B) --------------------------------------------------------
Fig. 4-7. Trim33 targeting vector and screening of ES colonies ---------------
Fig. 4-8. Generation of mice carrying the floxed (Trim33
FX
) and
knockout (Trim33
KO
) Trim33 alleles ----------------------------------------

Fig. 4-9. Development of embryos deficient in Trim33 is arrested
at the early somitogenic stage ---------------------------------------------

Fig. 4-10. The deletion of Trim33 function after neurulation causes
the twisted body axis and neural tube closure failure --------------

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ABSTRACT

Craniofacial malformations including cleft lip, cleft palate and
craniosynostosis are among the most common birth defects in humans. A Tgf-β
signaling pathway has been shown to be important during craniofacial
development. Detailed mechanisms of the canonical Tgf-β signaling pathway in
vitro have been well established, and a significant amount of information has
been accumulated about a role of the canonical Tgf-β signaling pathway in
craniofacial development. However, the canonical Tgf-β signaling pathway alone
is not enough to explain all of the published findings of different craniofacial
phenotypes in mice harboring mutations in genes encoding Tgf-β signal
transduction components in vivo. Therefore, we hypothesized that a non-
canonical Tgf-β signaling pathway also acts as an essential player in craniofacial
development. To this end, we studied a function of the non-canonical Tgf-β
signaling pathway in craniofacial development using three different approaches.
First, we discovered that non-conventional Tgf-β receptor combinations can also
act as functional receptor complexes. Second, we studied a role of Rac1, as a
possible downstream mediator of Tgf-βs, in craniofacial development using the
tissue-specific knockout mouse model system. We found out that Rac1 is an
essential factor in the craniofacial development, and an important regulator of
neural crest cell behavior. Third, to find out a function of Trim33, which was
recently identified as a Smad4-independent regulator of Tgf-β signaling, we

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generated the Trim33 conditional knockout mouse line. We discovered that
Trim33 is required for the early embryonic development. This study shows that
the non-canonical Tgf-β signaling pathway is also essential factor in craniofacial
development.

1
Chapter I
Introduction

1-1. Embryonic craniofacial development
The cranium is a unique structure in vertebrates. It plays a vital role in
protecting its contents, the brain. In addition, it is important in communication
with the outside world having several well-formed sensory organs. Formation of
the face tends to be among the most complicated processes in embryonic
development (Wilkie and Morriss-Kay 2001). Facial structures are derived from
several primitive facial primordia including the frontonasal process, lateral nasal
processes and paired maxillary and mandibular processes. The facial primordia
arrange to form the stomodeum, which is the future mouth and develop facial
organs such as the nose, upper and low jaw, palate, teeth, a part of the eye and
a part of ear by growth and delicate fusion of each facial primordium. The facial
primordia are composed of epithelial layers derived from the ectoderm and
endoderm and the mesenchyme derived from the mesoderm just like many other
organs. However, an additional mesenchymal component, the neural crest,
makes the facial development unique and rather sophisticated. Facial
development begins at embryonal day 7.5-8.0 (E7.5-E8) (early developmental
week 4 in human) with the neural crest cell induction. At E8.5 (late
developmental week 4 in human) the 1
st
pharyngeal arch develops maxillary and
mandibular components. Around E9 (still late developmental week 4 in human),
2
the otic pit can be clearly seen and the 3
rd
pharyngeal arch starts to be visible. At
E9.5, with the appearance of the forelimb bud, the forebrain vesicle is
subdivided. Since the neural crest cell induction, the neural crest cells keep
migrating along the lateral-ventral pathway. When the neural crest cells arrive to
the anterior aspects of the face, around E10 (early developmental week 5 in
human), the nasal processes start to form. With the swelling of maxillary
processes, the lateral and medial nasal processes get apposed and fuse at
E11.5. Figure 1-1 shows the overall process of facial formation (Dudas and
Kaartinen 2005).

Fig.1-1 Facial formation in the early embryo stage. Frontonasal process (yellow),
lateral nasal process (orange), two maxillary (green) and two mandibular (blue)
are required for the formation of facial structure. Each primordial grow and
arrange to form stomodeum. The fusion between frontonasal process and lateral
nasal processes give rise to form the nose structure. The fusion of nasal process
and maxillary forms the upper lip, maxillary and palate. In addition, the fusion of
two mandibular forms the lower lip and mandible. [Reproduced from (Dudas and
Kaartinen, 2005)
3

The palate formation takes place from E12 to E15 to separate the oral
cavity from the nasal cavity (Hilliard et al., 2005). Palatal shelves grow bilaterally
out from the maxillary processes, elevate above the tongue and eventually form
a contact around E14. The palate formation is completed with the epithelial
fusion of two palatal shelves (Dudas et al., 2007). Figure 1-2 depicts an overall
scheme of palate formation (Alappat et al., 2005).

Fig. 1-2 Palatal formation in the mouse embryo. A) Frontal view of an E11.5
head showing bilateral palatal shelves projecting internally from the maxillary
primordial. (B and C) the paired palatal shelves at E12.5 (B) and E13.5 (C) are
vertically oriented on either side of the tongue. (D) At E14.5 the shelves are
horizontally oriented above the dorsum of the tongue and are fused medially to
form a closed palate. Abbreviations: M, molar tooth bud; P, palate; PS, palatal
shelf; T, tongue [Reproduced from (Zhang et al., 2005)].

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Tooth formation starts around E11.5 (6
th
developmental week in human),
with the thickening the oral ectoderm (Jernvall and Thesleff 2000; Zhang et al.,
2005) and continues through the early and late bud stages, cap and bell stages,
and is eventually finished during the postnatal life. Figure 1-3 shows shows early
stages of tooth development (Alappat et al., 2005).

Fig. 1-3 Tooth formation in the mouse embryo. Dental lamina stage (E11.5): the
oral epithelium thickens locally to form the molar and incisor tooth germs. (C and
D) Early bud stage (E12.5): the epithelial thickening invaginates into the
subjacent mesenchyme which condenses around the epithelial bud. (E and F)
Late bud stage (E13.5):increased proliferation of the dental epithelium causes it
to invaginate further into the dental mesenchyme; (G and H) Cap stage (E14.5):
differential proliferation within the dental epithelium causes a population of the
dental mesenchyme, the dental papilla, to be surrounded by the convoluting
dental epithelium. Abbreviations: DE, dental epithelium;DM, dental
mesenchyme; DP, dental papilla; EK, enamel knot [Reproduced from (Zhang et
al., 2005)].

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Skull is also another important component in the craniofacial development
(Sasaki et al., 2006). It is composed of multiple bones, and can be subdivided
into the neurocranium and viscerocranium depending of the germ layer origin.
While the neurocranium is mainly derived from the mesoderm and protects the
brain, viscerocranium is derived from the neural crest cells and contributes to the
development of the facial bones and connective tissue. Figure 1-4 shows a
diagram of the skull and its cellular origins (Noden and Trainor 2005).

Fig. 1-4 Schematic chick and mouse skulls. The color showes the contributions
of neural crest, paraxial and lateral mesoderms to the cranial skeleton
[Reproduced from (Noden and Trainor 2005)].

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Congenital craniofacial developmental defects are very common. Morever,
so far, more than 150 human craniofacial syndromes have been identified
(Chacon GE et al., 2007). Craniofacial malformations can be caused by defects
in multiple developmental processes, such as induction or migration of neural
crest cells and outgrowth or fusion of facial primordia. Moreover, both
environmental factors, e.g. alcohol, cigarette smoke or other toxins and genetic
factors can be responsible for disturbances in normal craniofacial development
(Young et al., 2000). Due to the advances in molecular biology and gene
knockout technology, many genes responsible for normal craniofacial
development have been identified, such as signaling related genes [Fgfs (Nie et
al., 2006; Rice et al., 2004; (Sasaki et al., 2006), Tgf-βs (Kaartinen et al., 1995;
Dudas et al., 2006), Wnts and Bmp (Dudas et al., 2004; Nie et al., 2006)], and
transcription factors [Msx (Tribulo et al., 2003) and Dlx (Depew et al., 2002)
genes] .

1-2. Neural crest
Neural crest is a unique, transient cell population found only in
developing vertebrate embryos. After formation of the three germ layers, the
neural crest is derived from the dorsal ectoderm and contributes to development
of some mesenchymal tissues (Bronner-Fraser and Fraser 1988; Anderson
1989). So, neural crest is also called “ectomesenchyme” (Breau et al., 2008).
Two most striking characteristics of neural crest cells are their high capacity of
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migration (Bronner-Fraser M. 1993) and differentiation (Le Douarin et al., 2004)
into a variety of cell types. Derived from the dorsal ectoderm, neural crest cells
migrate to many distant locations in the developing embryo including the skin,
cranium, face and heart and even the enteric system. Neural crest cells destined
to a certain location differentiate appropriately to a corresponding cell- or tissue-
type. For instance, neural crest cells migrating to the skin differentiate to pigment
cells (Richardson and Sieber-Blum 1993), and neural crest cells that arrive to the
craniofacial area differentiate to the connective tissue (Noden and Trainor 2005;
Chai et al., 2000). Similarly, neural crest cells surrounding the aortic arch arteries
become smooth muscle cells (Jiang et al., 2000). Due to this fascinating
differentiation capacity, the neural crest is also considered as a migrating stem
cell population (Thomas et al., 2008; Le Douarin et al., 2008).
Neural crest induction starts with the expression of specifier genes in the
border between the neural plate and non-neural ectoderm at the early
gastrulation. Several specifier genes have been identified, e.g., the homeobox
transcription factors, Msx, Dlx3 and Dlx5 or other transcription factors e.g., Pax3,
Pax6 and Zic (Meulemans and Bronner-Fraser 2004; Huang and Saint-Jeannet
2004; Sato et al., 2005). The specifier genes are induced by extracellular
signaling molecules from surrounding tissues, such as the non-neural ectoderm
and the underlying dorsal mesoderm including the prechordal/paraxial mesoderm
and notochord. These surrounding tissues secret several extracellular signaling
molecules such as Bmp, Fgf, Wnt and Notch/Delta signaling molecules to induce
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the neural plate border specifier genes (Monsoro-Burq et al., 2005). The ultimate
goal of these extracellular molecules is to establish a Bmp signaling gradient in
the dorsal ectoderm. Usually, the continuous dorsal ectoderm composed of the
neural plate and the non-neural ectoderm express high concentrations of Bmp
molecules. On the other hand, Bmp antagonists contribute to a development of a
concentration gradient, so that the neural plate has a relatively low level of Bmp
signaling activity (Wilson and Edlund 2001). This activity increases progressively
along the dorsal ectoderm. The neural crest is induced in the area having the
intermediate level of Bmp signaling activity (Liem et al., 1995; Trainor 2005). Due
to the heterogeneity of a composition and concentration of several extracellular
signaling molecules, the neural crest induction occurs heterogeneously, which in
turn results in the heterogeneity of a neural crest cell population.
Once the neural plate border is determined with the appropriate specifier
genes expression, the neural crest cell population is determined. Several
transcription factors, such as Snail/Slug, FoxD3, Sox9, Ap2, Id, and c-Myc are
known as the neural crest specifier genes (Nieto 2002; Cano et al., 2000;
Cheung and Briscoe 2003; Bellmeyer et al., 2003). Snail/Slug and Sox9 are
usually used as neural crest specific markers. On the other hand, c-Myc and Id
have an essential function in maintaining the neural crest’s multipotent state by
inhibition of the immature differentiation. The neural crest specifier genes act in
the stimulation of a number of several downstream targets, which trigger the
neural crest cells to induce an activation of several cell adhesion molecules
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including integrins, cadherins and NCAM (Neural Cell Adhesion Molecule).
Changes on specific neural crest cell surface molecules induce the
reorganization of the surrounding extracellular matrix. Also, the reorganized
extracellular matrix acts as a new stimulus to neural crest cells themselves
(Perris and Lofberg 1986). Finally, the neural crest cells gain a motile phenotype,
delaminate from the neural ectoderm and make the environment permissive for
neural crest cell migration. This process called the epithelial-mesenchymal
transition (EMT) takes place at E7.6 in the mouse (early developmental week 4 in
human) (Kang and Svoboda 2005). Cadherin molecules are one of the key
components involved in the EMT of neural crest (Taneyhill 2008). In the initial
stage of the neural crest induction, E-cadherin is expressed in the dorsal
ectoderm. During the neural crest induction progress, the E-cadherin expression
is maintained in the non-neural ectoderm, while its expression is downregulated
in the neural plate. Instead, N-cadherin substitutes the E-cadherin in the neural
plate. In the border between the neural and non-neural ectoderm another
cadherin molecule, Cadherin-6B is upregulated and folds this region to a convex
shape. Subsequently, the N-cadherin and Cadherin-6B expression is
downregulated and the neural crest cells are delaminated from the border
between neural plate and non-neural ectoderm. With the delamination of the
neural crest cell population, the neural plate still maintaining the N-cadherin
expression becomes discontinuous from the non-neural ectoderm, and forms the
neural tube. Delaminated neural crest cells having a low level of cell adhesion
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molecules keep secreting signals to the extracellular matrix and vice versa.
However, the proper expression of cell adhesion molecules is important for an
appropriate migration direction and differentiation to correct cell types. For
example, N-cadherin is downregulated during neural crest cell migration, but is
highly expressed in neural crest cells that are differentiating into dorsal root
ganglia (DRG). Moreover, the extracellular matrix is important for the neural crest
cell migration and differentiation. Figure 1-5 shows the overall step and
regulatory genes in the neural crest formation (Sauka-Spengler and Bronner-
Fraser, 2008).

Fig. 1-5. Regulatory steps in neural crest formation. [Reproduced from (Sauka-
Spengler and Bronner-Fraser, 2008)].
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The neural crest cell migration takes place practically on all axial levels
along the entire rostral-caudal body axis. From the dorsal-ventral axis view, there
are two main neural crest cell migration pathways, i.e., dorsal-lateral and ventral.
Depending on their origin, the neural crest cells are subdivided into several
groups. Cranial neural crest cells, which give rise to many components of
craniofacial skeletomuscular system, are derived from the most rostral part of the
embryo, from the midbrain to hindbrain. Cardiac neural crest cells are derived
from the posterior hindbrain–anterior somatic (somites 1-3) region. This cell
population contributes the smooth muscle cell layer of the aortic arch arteries and
the septum to separate the aorta from the pulmonary artery. Trunk neural crest
cells have their origin at the more rostral region (from somite 6 throught the tail);
most of them differentiate to peripheral nerves. An origin of Vagal/Sacral neural
crest cells overlapps with the origin with the trunk neural crest cells. This neural
crest cell population differentiates to the enteric nervous system. The neural crest
cells migrating dorso-laterally stay close to the ectoderm and differentiate to
pigment cells, melanocyte.s Most of the other neural crest cells having a ventral
migration pathway differentiate to a variety of different cell types, such as the
peripheral nervous system and enteric nervous system (Le Douarin et al., 2004).
Figure 1-6 shows the different neural crest cell population aroused from different
body axis and its different differentiation capacity (Le Douarin et al., 2004). The
cranial neural crest cell population possesses a unique ability to differentiate to
cartilage, bone and muscle as well as the pigment cells and peripheral nervous
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tissue. While the connective tissue in the trunk part is derived from the
mesoderm, the most of the craniofacial connective tissue is derived from the
cranial neural crest cells. Cranial neural crest cells can also be subdivided into
several groups depending on their origin on the rostral-caudal body axis (Trainor
and Krumlauf 2000).

Fig. 1-6. Fate map of neural crest cells along the body axis. [Reproduced from
(Le Douarin et al., 2004)].
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The developing embryonic head is composed of three primary brain
vesicles, forebrain, midbrain and hindbrain. The developing hindbrain has a
transiently segmented structure, called the rhombomeres (in total there are 8
rhombomeres). The cranial neural crest cells are derived from each
rhombomere, except the rhombomeres 3 and 5. Neural crest cells from each
rhombomere display characteristic migration patterns and destinations, and there
is very little if any mixing taking place between neural crest cells with different
origins suggesting that each population has their own guiding cues for migration.
So far, several molecules, such as Eph-ephrin, Semaphorin-Neuropilin and Slit-
Robo have been identified as important guidance and cell-sorting molecules
functioning in neural crest cell migration. By expressing different sets of Eph-
ephrin, Semaphorin-Neuropilin and Slit-Robo molecules, neural crest cells form
segregated migration streams and maintain their identities during migration.
Cranial neural crest cells derived from the midbrain and rhombomere 1 and 2
migrate ventrally and contribute for the formation of the 1
st
pharyngeal arch. The
1
st
pharyngeal arch gives rise to a part of a frontal nasal process, and the
maxillary and madibular part, which become the main facial primordia. Most of
the craniofacial structures including a part of skull such as frontal bone, nasal
bone, jaw bones, teeth and the middle ear bones incus and malleus are derived
from this population of cranial neural crest cells (Santagati and Rijli 2003). The
trigeminal ganglia innervating for the eye, jaw and teeth are also derived from
this population of cranial neural crest cells. The cranial neural crest cells that
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originate from the rhombomere 4 migrate to the 2
nd
pharyngeal arch. This cell
population contributes to a part of the neck region such as the hyoid cartilage
and facial nerves. The 3
rd
, 4
th
pharyngeal arch are composed of cranial neural
crest cells derived from the rhombomeres 6-8, which contribute to the formation
of the hyoid cartilages, thymus, parathyroid and thyroid glands. Figure 1-7 shows
the genes regulating the cranial neural crest cells and their differentiation
capacity (Noden and Trainor, 2005).

Fig. 1-7 Cranial neural crest cells and their differentiation capacity. [Reproduced
from (Noden and Trainor, 2005)].
15
1-3. Several signaling pathways involved in the craniofacial development
Though the cranial neural crest cells are the main cellular resource in the
facial formation, involvement of the three other germ layers in the facial
development should not be depreciated. The craniofacial development also
shares one of the common mechanisms in oranogenesis, i.e., epithelial and
mesenchymal interaction. From the beginning of the neural crest cell induction,
the neural crest cells keep receiving signals from the epithelium derived from the
ectoderm and endoderm and from the mesondermally derived mesenchyme and
vice versa. During neural crest cell induction, the Bmp antagonists, Fgfs and
Notch/delta from mesoderm or Wnts from the non-neural ectoderm are good
examples in these interactions. After the neural crest cell induction, those
signaling factors keep involved in the facial primordia outgrowth and patterning.
Among several signaling pathways, Fgf, Hedgehog, Endothelin and Pdgf
signaling recruit the epithealil-mesenchymal interaction as a crucial mechanism
in facial primordia development.
More than 23 soluble Fgf ligands and 4 distinct receptors have been
identified. Fgf signaling promotes proliferation and differentiation of the cranial
neural crest cells into chondrocytes. Fgf-2, 4, 8 expressed in the epithelium of the
facial primordia stimulate their outgrowth and the differentiation by binding to
mesenchymally expressed Fgf receptors including Fgfr-1, 2, and 3. Any defects
in these genes leads to craniofacial malformations. It is known that a missense
mutation of the Fgfr-2 causes the human Apert’s syndrome with craniosynostosis
16
and severe craniofacial defects. Another receptor mutation, the constitutive
activation of Fgfr3 causes the human Crouzon’s syndrome displaying
craniosynostosis and the midfacial hypotrophy. The misexpression of Fgf4 leads
to an abnormal tooth development. For the palatogenesis, the Fgf signaling
pathway is also important acting in an indirect epithelial-mesenchymal
interaction. Fgf10 expressed in the palatal mesenchyme stimulates the
mesenchyme itself by inducing several gene expression related to cell
proliferation with the Shh signal secreted from the palatal epithelium. The Shh
expression is induced by Fgfr2, which is activated by the Fgf10 ligand from
mesenchyme. Disruption in this Fgf10/Fgfr-2 epithelial-mesenchymal interaction
leads to a formation of cleft palate. Table 1-1 shows the craniofacial malformation
caused by the defective Fgf signaling pathway (Nie et al., 2006).
Hedghog signaling is involved in many aspects of the craniofacial
development. From the three hedgehog ligands (Sonic, Indian and Desert
hedgehogs), the Sonic hedgehog is the most extensively studied in this context
(Calloni et al, 2007 and Haworth et al., 2007). During facial primordia
development, Shh ligand is expressed in the epithelium and the Shh receptor,
Patched is expressed in the mesenchyme. Shh signaling in the mesenchyme is
triggered by an interaction between Shh and Patched, which inactivates Patched
and leads to an activation of another transmembrane receptor, Smoothened.
This in turn leads to an activaton of several transcription factors, in the Gli family,
i.e., Gli-1, 2 and 3. Gli-2 and 3 are expressed in the facial mesenchyme and the
17
knockout of these genes causes the facial abnormalities. Similarly, mutations in
Patched cause the human Gorlin’s syndrome. The knockout of Shh ligand leads
to a very severe form of holoprocencephaly.
Endothelin and Pdgf signaling in facial development utilize a similar
mechanism of epithelial-mesenchymal interaction. The ligand, Endothelin-1 and
Pdgf are produced in the epithelium and receptors expressed in the
mesenchyme are activated by ligands secreted from the epithelium.

Table 1-1. Endogeneous Fgf/Fgfr expressed during the development of the
craniofacial structures and their possible roles in developmental disorders.
[Reproduced from (Nie et al., 2006)].
18
Tgf-β signaling is also an important player in the facial development
(Kaartinen et al., 1995 and Dudas et al., 2006; Ito et al., 2003) . Tgf-β superfamily
consists of Tgf-β 1, 2, 3 (transforming growth factor), Bmps (bone morphogenic
proteins), GDFs (growth and differentiation factors) and Activins/Inhibins. Most of
the ligands and receptors of the Tgf-β superfamily are expressed in the
developing facial primordia. Members of the Tgf-β superfamily function in
outgrowth of facial processes, chondrogenesis, patterning and apoptosis. With
the help of the gene knockout technology, many functions of the members of the
Tgf-β superfamily have been identified. Tgf-β 2 knockout mice have defects in
the maxillary and mandibular development and cleft palate with variable
penetrance. Tgf-β3 knockout mice display cleft palate with 100% penetrance
(Kaartinen et al., 1995). Bmps form a subfamily in the Tgf-β superfamily. Even
though there is a functional redundancy between Bmps, several Bmp knockouts
display craniofacial malformations. Especially Bmps are important in formation of
the craniofacial skeletal structures including skull and teeth (Nie et al., 2006;
Dudas et al., 2004). Bmp4 null mice die too early functional analysis in the
craniofacial skeletal system (Winnier et al., 1995). However, Bmp4 haplo-
insufficiency leads to a frontal and nasal bone hypotrophy. Also, Bmp4 is one of
the inducers of the tooth formation (Ohazama et al., 2005). Bmp4 is expressed in
the oral ectoderm around E10.5 and acts as a main factor in the ectodermal
signaling center. Subsequently, tooth formation is continued with well-organized
epithelial-mesenchymal interactions as shown in Figure 1-8 (Thesleff 2006).
19

Fig. 1-8 Reciprocal interaction between epithelium and mesenchyme during tooth
development. [Reproduced from (Thesleff 2006)].

1-4. Tgf-β signaling
In 1982, the Tgf-β was discovered as a growth factor, which was able to
induce “transformation” of normal untransformed cells (Anzano et al., 1982). It
was found out that Tgf-β secreted from “transformed” cells promoted
“transformation” of normal cells via autocrine or paracrine mechanisms. Even
though Tgf-β was originally discovered as a potentially transforming “cancer
causing” gene, it was subsequently discovered that that Tgf-βs played severeal
important functions by regulating cell proliferation, differentiation, migration and
maintenance of the cellular homeostasis. In addition, the Tgf-β could act
multifunctionally even in a same cell. It could promote or inhibit cell proliferation
by a context depending manner For instance, in a pre-malignant state, Tgf-β
20
acts as a tumor suppressor by inhibiting cell proliferation. However, when the
cells enter the malignant state, Tgf-β promotes the epithelial-mesenchymal
transition (EMT) and becomes as one of the tumor progression factors by
facilitating cell invasion (Bachman and Park 2005; Xu et al., 2009; Battaglia et
al., 2009)

1-5. Tgf-β ligands and receptors
Twentynine structurally related Tgf-β ligands have been identified
(Massagué 1998). According to their sequence homologies and functional
characters, these 29 ligands are subdivided into several sub-groups, such as
Tgf-βs, Activins, Bmps, and Gdfs. Each subfamily is also composed of several
members such as 3 members in Tgf-β subfamily, more than 8 members in Bmp
family and more than 4 member in Activin family. Table 1-2 shows the sequence
homologies and each main function. In Tgf-β subfamily, three different Tgf-β
isoforms, called Tgfb1, -2 and -3, have been discovered. They all show very
similar signaling characteristics in vitro. However, in vivo they all display distinct,
both spatially and temporally tightly controlled expression patterns. Tgf-βs act as
important regulatory factors during embryogenesis. Due to the development of
the gene knockout technology, many functions of members of the Tgf-β
superfamily in embryogenesis have been identified (Chang et al., 2001). While
Tgf-β 1 has been shown to be an essential factor in embryonic haematopoiesis
and vasculogenesis (Dickson et al., 1995), Tgf-β 2 was shown to play an
21
important role in the formation of craniofacial, heart and urogenital structures
(Sanford et al., 1997). Mice deficient in Tgf-β3 suffer from cleft palate and died
soon after the birth (Kaartinen et al., 1995). To conclude, defective Tgf-β
signaling causes a number of severe developmental defects including
craniofacial and heart malformations. Several different receptors for members of
the Tgf-β superfamily have been identified (type I, type II and type III receptors)
(de Caestecker 2004). Among them, the type I and type II receptors are the main
receptors participating in the formation of the receptor complexes to transduce
the Tgf-β signaling. As a co-receptor, the type III receptor functions as a regulator
or modifier of Tgf-β signaling (López-Casillas et al., 1991). For example, by
binding to the ligands, type III receptors could regulate the concentration of
available ligand or the localization of ligands or trafficking of ligands. So far, 7
different type I receptors (Alk1 – Alk7) (Franzén et al., 1993; ten Dijke et al.,
1993) and 5 different type II receptors (BmpR-II, ActR-IIA, ActR-IIB, TgfbRII,
AmhR) have been identified (Liu et al., 1995; Lin et al., 1992).

22

Table 1-2. Tgf-β ligands. [Reproduced from (Massagué J. 1998)].
23
1-6. Canonical Tgf-β signaling pathway
Tgf-β signaling pathway is initiated by the binding of Tgf-β ligands to their
specific cell surface receptors. Usually, the Tgf-β ligands function as a
homo/hetero-dimer form (Sun et al., 1995; Wrana et al., 1992). There are two
ways for ligands to bind their receptors. Tgf-βs and Activins bind first to a
homodimer of type II receptors, which are constitutively active serine/threonine
kinases. Next, type II receptor-ligand complexes bind to a homodimer of type I
receptors, form a heterotetrameric complex and activate the serine/threonice
kinase activity of type I receptors. Bmps and Gdfs, in turn bind to type I and type
II receptors at a same time and induce the formation of heterotetrameric receptor
complex. In both cases, type I receptors are activated by type II receptors. Type I
receptors contain the called GS domain between the transmembrane and kinase
domains, which has a highly conserved SGSGSG amino acid sequence. Also,
type I receptors have the called L45 loop region in their kinase domain, which is
responsible for the specific recognition of a downstream signal transducer, Smad
(Shi and Massagué 2003). When the type II receptor activate the type I receptor,
the serine residues in the GS domain in the type I receptor are phosphorylated
by the type II receptor and activate the kinase domain of type I receptor. It is
known that there are specific binding combinations between the type I and type II
receptors as well as between the ligand and receptor complexes. For example,
heterodimeric Tgf-βs bind to TgfβRII, which binds and activates Alk5 (TgfR-I).
Bmp2, in turn, binds to the BmpRII and Alk3 (BmpR-IA) complex. However,
24
there are several pieces of evidence suggesting that the rules to assembly these
receptor combinations are not that strict. For example, binding of Gdf11 to the
activin type II receptors (ActR-IIA or ActR-IIB), leads to an activation of Alk5
(TgfR-I). These non-conventional Tgf-β receptor combinations will be discussed
in Chapter II. The activated type I receptors induce a recruitment of selected
specific R-Smad proteins and activate the R-Smad proteins by phosphorylation.
So far, 8 kinds of Smad proteins have been identified. According to their function,
the Smad proteins are subdivided into three groups, such as receptor-specific
Smads (R-Smad), Co-Smad and inhibitory Smads (I-Smad) having a inhibition
function of Tgf-β signaling (Massagué J. et al., 2005). Smads 1, 5 and 8 are
activated by the Alk1, Alk2 (ActR-1), Alk3 (BmpR-1A) and Alk6 (BmpR-1B).
Smads 2 and 3 are activated by Alk4 (ActR-1B) or Alk5 (TgfR-1). Activated type I
receptors activate R-Smad proteins by phosphorylation of the serine domain in
the SSXS sequence in the C-terminal region (Wu G. et al., 2000; Wu JW. et al.,
2001). The activated R-Smad proteins bind to the Co-smad (Smad 4) and form
either heterodimeric or heterotrimeric Smad protein complexes. The activated R-
Smad proteins undergo the conformational changes and expose the nuclear
location sequence (NLS) (Xiao et al., 2000). Subsequently, the Smad protein
complexes accumulate to the nucleus. The Smad protein complex plays an
important role in the regulation of gene expression with several transcription
factors or co-transcription factors. Figure 1-9 shows the general canonical Tgf-β
signaling pathway (Schmiere and Hill, 2007).
25

Fig. 1-9. Canonical Tgf-β signaling pathway. [Reproduced from (Schmiere and
Hill, 2007)].

26
1-7. Regulation of Tgf-β signaling pathway
There are several ways to regulate the Tgf-β superfamily signaling. Tgf-βs
are secreted as inactive precursors containing the latency associated peptide
(LAP) and the mature Tgf-β peptide (Saharinen et al., 1996). With the function of
integrins or several proteases in the extracellular matrix including matrix-
metalloporteinases (MMP) (Wilkins-Port and Higgins 2007), plasmin or
thrombospondin, active Tgf-βs are released. In contrast, Bmps are secreted as
active forms directly. With the function of Bmp antagonists, the levels of active
Bmps are regulated. The Bmp antagonists, such as noggin, chordin or sclerostin,
bind to Bmp ligands directly and prevent them from binding to their receptors
(McMahon JA. et al., 1998). Regulation of Smad protein levels is also important
in control of Tgf-β signaling. As negative feedback regulators, Smad 6 and 7 are
induced by the Tgf-β signaling and inhibit it by down-regulation of the R-Smad
activity. Smurf (Smad – Ubiquitination – regulatory factor) is another factor
regulating the Smad proteins targeting them for degradation (Zhu et al., 1999) So
far, two Smurf proteins are identified. While the Smurf2 interacts with every kinds
of R-Smad proteins and participates in the overall Tgf-β signaling, the Smurf1
interacts with only Bmps related R-Smad, Smad 1, 5 and 8 and regulate
especially the Bmp signaling (Inoue and Imamura, 2008).

27
1-8. Non-canonical Tgf-β signaling pathway
Even though the Smad proteins are key mediators of Tgf-β signaling,
there are several lines of evidence suggesting that the Tgf-β signaling can also
take place without canonical intervention of Smad proteins (Zhang 2009), which
is called non-canonical Tgf-β signaling pathway. Smad-independent pathway,
which is totally absence of involvement of Smad protein, is one of the cases of
non-canonical Tgf-β signaling pathway. It is known that the Tgf-β signaling
pathway is essential in the craniofacial development, especially in the palate
formation. Dudas et al. suggested that the Tgf-β3 in the palatal epithelium acts as
an essential factor in the palatal shelves’ fusion by activation of Alk5/Smad
pathway (Dudas 2004a). However, Yang et al. showed that the Smad4 deficient
mice in their epithelium do not have any defect in the palate formation (Yang et
al., 2005). This result shows that the Smad4 is not an essential factor in the
palate formation and suggests that there can be other factor responsible for the
palatal fusion under the downstream of the Tgf-β3, Alk5/Smad2, 3 pathway. As
one of the non-canonical Tgf-β signaling pathway, it is also known that cells
deficient Smad4 induce MAPK signaling pathway by the action of Tgf-β ligands
without any Smad protein intervention (Yu et al., 2002; Xu et al., 2008). Recently,
Xun et al. suggested that p38 MAPK is also involved the tooth and palatal
development (Xun et al., 2008). The presence of non-canonical Tgf-β signaling is
rather well established, however, its biological function, particularly in vivo, is still
poorly understood. Several evidences showed that the Rac1 is also one of
28
the intermediators in the non-canonical Tgf-β signaling pathway (Fig. 1-10)
(Janda et al., 2002; Wang et al., 2006; Nawshad et al., 2005). However, the
Rac1 function in the craniofacial development is not yet clear. In chapter III, the
Rac1 function in the craniofacial development, especially in the neural crest will
be discussed.

Fig. 1-10. Rac1 in non-canonical Tgf-β signaling pathway. [Reproduced from
(Nawshad et al., 2005)].
29
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39
Chapter II
Non-conventional Tgf-βreceptors combination

2-1. Summary
As one of the essential factors in craniofacial development, Tgf-β signaling
is triggered by a binding of ligands to their cognate type I and type II receptors.
Upon ligand binding these two types of receptors form a functional
heterotetrameric complex. It is known that there are some certain preferred
receptor combinations between the type I and type II receptors. On the other
hand, several pieces of evidence, especially in the craniofacial development,
suggest that rules to form these receptor combinations are not that strict. To
further address this question, we first confirmed that, in developing craniofacial
tissues, there are several different kinds of type I and type II receptors expressed
at a same time. To identify functional non-conventional Tgf-β receptor
complexes, we co-transfected Alk5 and different type II expression vectors
having a specific binding domain for an artificial heterodimerizer into cells that
lack the endogenous Tgf-β type II receptor. By using the heterodimerizer, we
demonstrated that a non-conventional Tgf-β receptor complex between Alk5 and
Activin type II receptor-b phosphorylates the Smad2 proteins just like the
conventional Alk5/TgfbRII receptor combination complex. This study shows that
a non-conventional Tgf-β receptor combination can also act as a functional
receptor complex.
40
2-2. Introduction
The transforming growth factor-β (Tgf-β) signaling is initiated when ligands
are bound to their receptors. As one of the largest growth factor super-families,
Tgf-β super-family is composed of more than 29 different ligands (Massagué
1998). Based on their functional and regulatory characteristics, the Tgf-β super-
family can be divided into two groups, i.e., the Tgf-β/Activin sub-family and the
Bmp (bone morphogenetic proteis)/GDF (growth and differentiation factor)
subfamily (Newfeld et al., 1999). According to the sequence and evolutionary
relationship, these two groups can be further subdivided to smaller subfamilies,
such as Tgf-βs, Activins, Inhibins, Bmp2, Bmp5/6/7 and Gdf5/6/7, ect. Three
different types of Tgf-β receptors have been identified, i.e., type I and type II and
type III receptors (de Caestecker 2004). As an accessory receptor, the type III
receptor (López-Casillas et al., 1991), which is also called betaglycan, does not
participate to the Tgf-β signaling pathway directly. Instead, it acts as a regulatory
cell membrane protein by facilitating the Tgf-β ligand binding to the type I or type
II receptors (Lewis et al., 2000). The type I and type II receptors are the main
receptors transmitting Tgf-β signals. The Tgf-βs and Activins bind to the type II
receptors, which are constitutively active serine/threonine kinases. The ligand-
type II receptor complex in turn binds to the type I receptor and activate the
cytoplasmic kinase domain of type I receptor. However, Bmps bind to the type II
and type I receptors at a same time. Similar to their ligands, the type I and type II
receptors act as homo-dimers. Therefore, the ligand binding induces a
41
formation of heterotetrameric receptor complexes (Wrana et al., 1992). So far, 7
different kinds of type I receptors (de Caestecker 2004 and ten Dijke et al., 1994)
and 5 different kinds of type II receptors have been identified (de Caestecker
2004). Since the first cloned receptor was shown to bind Activin, it was named as
Activin receptor-like kinase (ten Dijke et al., 1993), and subsequently all the type
I receptors have been named as Alks, from Alk1-Alk7. The type I receptors have
also been named based on ligands they bind. For example, another name of
Alk2 is ActRI (Activin type I receptor) and Alk5 is also called TgfβRI (Tgf-β type I
receptor). Five different type II receptors have been named based on their ligand
binding affinities, such as, BmpR-II (Bmp type II receptor), ActR-II (Activin type II
receptor) and TgfbRII (Tgf-β type II receptor), etc. Figure 2-1 from Joan
Massague’s review paper (Massagué J. 1998) summarizes conventional
combinations of ligand, type I and type II receptor complexes.

Fig. 2-1 Conventional Tgf-β receptor combination. [Reproduced from (Massagué
J. 1998)].
42
However, many studies have accumulated evidence that the ‘rules’ for these
conventional combinations are not that strict. Chen et al. showed that the type I
and type II receptors can form heteromeric complexes without any presence Tgf-
β ligands, even though the Tgf-β signaling pathway cannot be triggered (Chen et
al., 1995). Pannu et al. reported that the formation of type I and type II receptor
complex does not follow strict the 1:1 receptor ratio rule (Pannu et al., 2004). The
receptor preference of ligands is not that strict, either. In addition, functional
combinations of type I and type II receptosr are also variable. As one of the key
elements in normal craniofacial development, the function of Tgf-β signaling
pathway has been studied. Interestingly, phenotypes of the neural crest specific
Tgfbr2 and Alk5 mutants demonstrate some striking differences. Therefore, in
this study, we examined whether Alk5 could be activated by type II receptors
other than the Tgf-β type II receptor.

2-3. Materials and Methods
2-3-1. Preparation of chimeric Tgf-β receptor constructs.
The Smad2 expression vector was obtained from Rik Derynck. The
cytoplasmic domain of Alk5, TgfbRII and ActR-IIB were prepared by the following
method. The total mRNA was isolated from E10 mouse embryo by using the
RNeasy Mini kit from Qiagen. The total cDNA was synthesized by using the
Omniscript RT Kit (Qiagen) with the isolated total mRNA. The cytoplasmic
domains of Alk5, TgfbRII and ActR-IIB were amplified by PCR reaction with the
43
synthesized cDNA as a template. The oligomer sequences for PCR reaction are
followed. For Alk5, the nucleotide 541-1543 (accession # is L26110). For TgfbRII
the nucleotide 901-2027 (accession # is NM_009371). For ActR-IIB the
nucleotide 602-1651 (accession # is NM_007397). The PCR fragment from Alk5
cytoplasmic domain was subcloned into the pC
4
-R
H
E plasmid and the PCR
fragments of TgfbRII and ActR-IIB cytoplasmic domains were subcloned into the
pC
4
M-F2E plasmid. The pC
4
-R
H
E and pC
4
M-F2E plasmid vectors were from
ARGENT
TM
Regulated Heterodimerization Kit (ARIAD Pharmaceuticals, Inc.) The
pC
4
-R
H
E plasmid vector expresses the FRB* domain under a control of the
hCMV promoter. A PCR fragment of the cytoplasmic domain of Alk5 was fused
in-frame to the FRB* domain. The pC
4
M-F2E vector has 2 copies of FKBP
domains following the myristoylation signal under the hCMV promoter. The PCR-
generated cytoplasmic domains of TgfbRII and ActR-IIB were fused in-frame to
the FKBP domain. An artificial heterodimerizer, AP21967, can be used to induce
a complex formation between the FRB* and FKBP. The myristoylation domain of
pC
4
M-F2E is responsible for dodging the protein into the cytoplasmic membrane.
All constructs were sequenced to verify that the open reading frame was retained
and that no mutations were created during PCR reactions.

2-3-2. Cell culture, co-transfection of constructs and treatment
The DR26 cell line, which is defective for the endogeneous TgfβRII was
obtained from Joan Massague. The cells were cultured with DMEM/F-12 medium
44
(Invitrogen) containing 10% of FBS (Invitrogen) and Penicillin/Streptomycin
(Sigma) at 37°C, 5% of CO
2
. For transfections, the DR26 cells were plated onto
24-well plate and cultured with DMEM/F-12 medium containing 10% of FBS
without antibiotics and grown for 24 hrs. With Opti-MEM I reduced serum
medium (Invitrogen), the DNAs were diluted and mixed together. As a positive
control, 200ng of the Smad2 vector, 400ng of the Alk5 vector and 400ng of the
TgfβRII vector were mixed with 2.0µl of Lipofectamine 2000 (Invitrogen) in a final
volume of 100µl of Opti-MEM I reduced serum medium. As an experimental
sample, 200ng of the expression Smad2 vector, 400ng of Alk5 vector and 400ng
of ActR-IIB were mixed with 2.0µl of Lipofectamine 2000 in a final volume of
100µl of Opti-MEM I reduced serum medium. After adding the DNA mixture, the
cells were washed with PBS and incubated with 0nM, 50nM, 500nM
concentrations of heterodimerizer, AP21967 to induce the heterodimerization for
40min. After 48 hours the transfected cells were washed with PBS, lysed with 2X
laemmli sample buffer and harvested.

2-3-3. Western blotting analyses
The 10% acrylamide gesl were prepared with the 30% of acrylamide mix
from Bio Rad and TEMED from invitrogen. The western blot apparatus was from
Bio Rad, and the protein molecular marker was from Invitrogen. The proteins
were separated in the polyacrylamide gel with 110voltage for 1.5hr and
transferred to the PVDF membranes from Amersham. The membranes were
45
blocked with 5% milk incubated with primary antibodies (anti-phospho-Smad2
(Upstate biotech), anti-Smad2 (Santa Cruz biotech), anti-HA (Santa Cruz; for the
detection of Alk5, TgfβRII and ActRIIB proteins).

2-3-4. Whole-mount embryo analyses
For the whole-mount RNA in situ hybridization, embryos were fixed with
4% of buffered formaldehyde for 12 hrs and dehydrated with 30%, 50%, 70%,
100% of ethanol. Antisense RNA probes were synthesized with NTP DIG RNA
labeling mix from Roche Applied Science. Probes specific for Alk5, ActR-IIA,
ActR-IIB, BmpR-II and TgfbRII were used. After pre-hybridization without probe
for 1hr at 70°C, the embryos were incubated with RNA probes for overnight at
70°C. After washing out the remaining unbound RNA probe, the embryos were
incubated with DIG antibody for overnight at 4°C. After 5-6hrs washing out with
TBSTL (Tris buffered saline solution with levamisole), the color was developed
with the BM purple AP substrate solution (Roche) at 37°C.

2-4. Results
2-4-1. Expression of Alk5 and type II receptors
The whole mount in situ hybridization with RNA probes showed that the
Alk5 (type I receptor) and the type II receptors, ActR-IIA, ActR-IIB, BmpR-II and
TgfbRII are expressed in the first pharyngeal arch at E10.5 (Figure 2-2).

46

Fig. 2-2 Endogenous co-expression of Alk5 and type-II receptors.

2-4-2. Alk5 can form functional signaling complexes with unconventional
type II receptors in vitro.
To further explore the possibility that Alk5 could be activated by several
different type II receptor kinases in vivo, we made use of the rapamycin-based
regulated heterodimerization system (Stockwell and Schreiber, 1998 and Rivera
et al., 1996). This methodology is based on a finding that the non-
immunosuppressive rapamycin analogue AP21967 induces heterodimerization
between an FKBP12 protein domain and a mutated version of a large PI3K
homolog FRAP called FRB*, without interfering with the activity of endogenous
FRAP (Pollock et al., 2000). We constructed a mammalian expression vector
encoding the FRB*-ALK5 cytoplasmic domain fusion protein (FRB*-ALK5cyt). In
addition, we generated vectors containing a myristoylation sequence, two FKBP
domains, and either the TGFβRII kinase domain (Myr-FKBP-TGFβRII-kin) or
ACVRIIB kinase domain (Myr-FKBP-ACVRIIB-kin) (Figure 2-3 A–B). We
expected that these constructs could associate conditionally upon addition of the
heterodimerizer AP21967, which would subsequently lead to the phosphorylation
47
of a downstream signal transducer, Smad2, as shown in the schematic model
(Figure 2-3 B). The constructs were transfected into Tgfbr2-deficient DR26 cells
together with the Smad2 expression vector. After 24 h, the cellular extracts were
prepared and analyzed for Smad2 phosphorylation. In the absence of the
dimerizer, Smad2 phosphorylation was undetectable. Addition of the dimerizer to
cells co-transfected with Myr-FKBP-TGFβRII-kin and FRB-ALK5cyt induced
strong Smad2 phosphorylation, consistent with the earlier findings of Stockwell
and Schreiber. They showed that this system can be used to investigate the
mechanisms of Smad activation resulting from formation of a complex between
cytoplasmic domains of TGFβRI and TGFβRII (Stockwell and Schreiber, 1998).
Moreover, Myr-FKBP-ACVRIIB-kin and FRB-ALK5cyt fusion proteins, in the
presence of the heterodimerizer, could also stimulate Smad2 phosphorylation,
albeit more than 10-fold less efficiently than TGFβRII and ALK5 fusion proteins
under identical conditions (Figure 2-3 C).

48

Fig. 2-3 Activation of ALK5 by two different type II receptors in vitro.
Regulated heterodimerization was used as a model system to test whether ALK5
may also be activated by ACVRIIB in addition to canonical TGFβRII. (A)
Expression constructs for type II receptors contained N-terminal myristoylation
signal (Myr), two copies of FK506 binding domains (FKBP), and regions
coding for kinase domains of ACVRIIB (construct IIA in panel A) and
TGFβRII (construct IIT in panel A), respectively. The ALK5 cytoplasmic
domain was fused to FKBP-rapamycin binding domain (FRB*, see Materials
and methods; construct I in panel A). (B) Heterodimerizing agent (AP21967,
Ariad Pharmaceuticals; brown in the scheme) can be used to induce regulated
heterodimerization of chimeric type I and type II receptor fusion proteins
expressed in cell culture. (C) Western blot analysis shows that the kinase domain
of ACVRIIB was able to activate the ALK5 fusion protein in Tgfbr2-deficient
DR26 cells co-transfected with the Smad2 expression vector, albeit less
efficiently than the corresponding TGFβRII domain, as demonstrated by
phosphorylation of the effector protein Smad2. Left panel-exposure 5 min; right
panel—exposure 20 min.
49
To conclude, these experiments provide direct biochemical evidence that ALK5
activation can be induced not only the kinase activity of TGFβRII, but also by the
other type II receptor kinases such as ACVRIIB, provided that they are properly
associated with each other.

2-5. Discussion
The defects in calvaria of the Alk5/Wnt1-Cre mice are analogous with
those found in the corresponding Tgfbr2/Wnt1-Cre mutants (Ito et al., 2003). In
both mutants, defects occurred not only in bones derived from the neural crest,
but also in mesenchyme-derived structures. Because the role of TGF-β signaling
in morphogenesis of cranial bones has already been studied in detail in
Tgfbr2/Wnt1-Cre mutants (Sasaki et al., 2006), this study does not address
calvarial development. Interestingly, facial phenotypes of Alk5 mutant mice are
far more severe than those seen in Tgfbr2/Wnt1-Cre embryos. To explain this
phenotypic difference, we considered the possibility that during facial
morphogenesis, ALK5 could become activated also by the other type II
receptors. For example, GDF11, which is a ligand closely related to TGF-βs, has
been shown in cell culture studies to bind ACVRIIA as well as ACVRIIB, and to
activate Smad2 possibly via ALK5 (Mazerbourg et al., 2004 and Oh et al., 2002).
Gdf11 is strongly expressed both in maxillary and mandibular primordia at E10–
E11, and in the pre-fusion palatal MEE. Moreover, we show that its putative
50
binding partner Acvr2B is strongly expressed in the developing facial
prominences. These findings are in concordance with earlier studies
demonstrating that both Gdf11 and Acvr2B null mutants exhibit cleft palate with
variable penetrance (Ferguson et al., 2001, McPherron et al., 1999 and Matzuk
et al., 1995). Consequently, we utilized the regulated heterodimerization system
in a cell culture model to show that, at least in this simulated set-up, the kinase
domain of ACVRIIB can activate the intracellular domain of ALK5, albeit less
efficiently than that of TGFβRII. Therefore, GDF11 signaling via ACVRIIB and
ALK5 in the neural crest derived facial mesenchyme, may account for different
phenotypes we see in Alk5 mutants, when compared to corresponding Tgfbr2
mutants. Although not detected in our experiments, it is possible that ALK5 may
also form signaling complexes with additional type II receptors, and/or transmit
signals mediated by more members of the TGF-β superfamily ligands than
currently known. Thus, widespread and uniform deletion of Alk5 in the
ectomesenchymal cells would be reflected in a complex phenotype with
spatiotemporally complex pattern of localized expression changes of various
genes, which is fully consistent with our findings in Alk5/Wnt1-Cre mutants.

51
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Heldin CH, Miyazono K. 1995. Cloning and characterization of a human type II
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Rivera VM, Clackson T, Natesan S, Pollock R, Amara JF, Keenan T, Magari SR,
Phillips T, Courage NL, Cerasoli F Jr, Holt DA, Gilman M. 1996. A humanized
system for pharmacologic control of gene expression. Nat Med. 2, 1028-32.

Sasaki T, Ito Y, Bringas P Jr, Chou S, Urata MM, Slavkin H, Chai Y. 2006.
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proliferation during frontal bone development. Development. 133, 371-81.

Stockwell BR, Schreiber SL. 1998. Probing the role of homomeric and
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dimerizers. Curr Biol. 8, 761-70.

Sun PD, Davies DR. 1995. The cystine-knot growth-factor superfamily. Annu Rev
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ten Dijke P, Ichijo H, Franzén P, Schulz P, Saras J, Toyoshima H, Heldin CH,
Miyazono K. 1993. Activin receptor-like kinases: a novel subclass of cell-surface
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Massagué J. 1992. TGF beta signals through a heteromeric protein kinase
receptor complex. Cell. 71, 1003-1.

54
Chapter III
Rac1 is indispensable in the craniofacial and cardiac neural crest, but is
not required for NCC migration in vivo

3-1. Summary
Craniofacial malformations are among the most common birth defects in
humans. While canonical (Smad-dependent) Tgf-β signaling has been
extensively studied in craniofacial development, much less is known about
parallel non-canonical Tgf-β signaling processes. Previous studies have
suggested that the small Rho-related GTPase Rac1 is a possible mediator of
Smad-independent Tgf-β signals. Based on in vitro studies, it has been proposed
that the main function of Rac1 is to regulate cell migration and attachment by
controlling reorganization of the actin cytoskeleton. Here we deleted Rac1 in the
neural crest cell (NCC) lineage by using the Cre-loxP strategy. The mutant
embryos (Rac/Wnt1-Cre) appeared normal until embryonal day 10 (E10). At E11
the NCC-derived facial structures were grossly rudimentary and displayed blood-
filled cysts, particularly in the developing mandible. Subsequently the mutant
embryos died between E12 and E13 with severe cardiovascular defect.
Surprisingly, the rhombomere identity and NCC migration appeared normal in
Rac1/Wnt1-Cre embryos. However, neural crest-derived structures, e.g., cranial
nerves and pharyngeal arch arteries, displayed defective differentiation.
Moreover, craniofacial NCCs isolated from mutant embryos at E10 demonstrated
55
severe defects in cell attachment and morphology. These results suggest that
Rac1 is a crucial component in craniofacial development playing an important
role in post-migratory NCCs by regulating cell-cell and cell-matrix interactions, as
well as differentiation.

3-2. Introduction
Craniofacial malformations are among the most common birth defects in
humans. During embryogenesis, the craniofacial primordia are derived from the
three germ layers, i.e., ectoderm, endoderm and mesoderm, and from the neural
crest (NC) (Noden and Trainor, 2005; Richman and Lee, 2003; Creuzet et al.,
2005). The neural crest (NC) is formed bilaterally at the neural plate border along
the entire anterioposterior axis (Huang and Saint-Jeannet, 2004; Trainor, 2005).
The NC, which is found only in the developing vertebrates, is subdivided into
cranial, cardiac, vagal and trunk crest depending on the origin of cells on the
rostral to caudal body axis. NC induction is a complex process involving many
signaling pathways e.g., Bmps, Fgfs, Wnts and Notch/Delta (Sasai and De
Robertis, 1997; Nie et al., 2006) (Monsoro-Burq et al., 2003) (Dorsky et al., 1998;
Wu et al., 2005) (Cornell and Eisen, 2005).
The most striking features of neural crest cells (NCC) are their high
migratory capacity and their potential to differentiate to various cell and tissue
types (Le Douarin et al., 2004). In the mouse, this migration starts when the
neural plate folds around the 4-5 somite stage (Nichols, 1986). First, NCCs
56
undergo epithelial-to-mesenchymal transition (EMT), delaminate, migrate ventro-
laterally and differentiate into multiple mesenchymal cell lineages contributing to
the peripheral nervous system, to the connective tissue in the head (Yang et al.,
2000), to formation of the aortico-pulmonary (AP) septum (that separates the
aorta from to pulmonary trunk) and to smooth muscle cells covering the
ascending aorta and aortic arch (Jiang et al., 2000).
NCC migration is accompanied with a down-regulation of cell adhesion
molecules, rearrangement of the extracellular matrix and changes in cell-matrix
interactions. These changes, together with other extracellular signals, e.g.,
growth factors, induce cytoskeletal rearrangements and stimulate formation of
sub-cellular structures, e.g., filopodia and lamellipodia triggering cell migration
(Ridley et al., 2003). Many of these important cellular changes are dependent on
complex functions of small Rho-related GTPases that act as molecular switches
and regulate, rapid assembly and destruction of actin filaments. Other important
functions of Rho-related GTPases include regulation of cell proliferation,
apoptosis and gene expression. The Rho small GTPase protein family is
composed of Rho, Cdc42 and Rac subfamilies (Ridley et al., 2006). Each
member has their own cell-type-specific functions. For instance, in Swiss 3T3
fibroblasts, RhoA, Cdc42 and Rac1 act on the formation of focal adhesion
complexes, filopodia and lamellipodia, respectively. The Rac subfamily is in turn
composed of three members: Rac1, Rac2 and Rac3. Rac1 is expressed
ubiquitously, while Rac2 expression is limited to hematopoietic tissues and Rac3
57
is predominanty expressed in the central nervous system. So far, most of the
Rac1 function has been elucidated through the analysis of cell behavior in vitro.
Rac1 is required for formation of the sub-cellular structures related to cell
migration, such as the lamellipodia, actin stress fibers or focal adhesion
complexes (Guillou et al., 2008). Mouse embryos deficient in Rac1 fail to form
appropriate germ cell layers at die at gastrulation (Sugihara et al., 1998), while
Rac1-deficient neutrophils display defects in inflammatory recruitment, migration
to chemotactic stimuli, and chemoattractant-mediated actin assembly (Glogauer
et al., 2003). Similarly, deletion of Rac1 in endothelial cells causes defects in the
endothelial cell migration with severe defects in angiogenesis (Tan et al., 2008).
One of the most intriguing characteristics of NCCs is their high capacity of
migration; a process where Rac1 has been thought to play a major role. In this
study, we deleted Rac1 function is deleted in NCCs by using the conditional gene
knockout strategy. We discovered that Rac1 is required for appropriate
patterning of facial structures and for proper formation of the AP septum.
However, while NCCs deficient in Rac1 fail to migrate in vitro, they do not
demonstrate notable migratory defects in vivo.

3-3. Materials and Methods
3-3-1. Mouse breeding, genotyping and embryo collection
Rac1
fx/fx
(Glogauer et al., 2003), R26R and Wnt1-Cre mice were
genotyped by PCR as described (Glogauer et al., 2003). Wnt1-Cre mice were
58
obtained by A. McMahon (Harvard University). DNA template for PCR
genotyping was prepared from the yolk sac or tail lysate by using DirectPCR
Lysis Reagents from Viagen Biotech. The mice were maintained 12hrs light/dark
period. Rac1
fx/fx
or Rac1
fx/fx
/R26R+/+ female mice were crossed with
Rac1
fx/+
/Wnt1-Cre male mice during the dark period. The presence of vaginal
plugs was designated as day 0. The plugged females were euthanized by CO2
gas following to the national and institutional guidelines. Embryos at each stage
were collected in the PBS or HBSS on ice. The yolk sacs were saved for
genotyping.

3-3-2. Cell death and proliferation assay
For cell death and proliferation assays, tissues were embedded into
paraffin and sectioned according to standard protocols. Apoptotic cells were
detected with the DeadEnd TM Fluorometric TUNEL system from Promega by
following the manufacturer’s instructions. For a cell proliferation assay, the BrdU
reagent from Amersham was injected (i.p.) to pregnant female mice. After 5
minutes, the females were euthanized and the embryos were collected. BrdU
was detected using the Amersham cell proliferation kit.

3-3-3. Whole-mount embryo analysis
For whole-mount RNA in situ hybridization, collected embryos at each
embryonic stage were fixed with 4% of buffered formaldehyde for 12 hrs and
59
dehydrated with ethanol. Antisense RNA probes were synthesized with NTP DIG
RNA labeling mix from Roche Applied Science. Probes specific for Krox20,
EphA4 and Sox10 were used (Ishii et al., 2005). Whole-mount
immunohistochemistry for neurofilament detection was performed as described in
“Manipulating the mouse embryo (a laboratory manual)” with some modifications.
2H3 antibody from Developmental Studied Hybridoma Bank in University of Iowa
was used. For the color development & detection, SigmaFastTM DAB with Metal
Enhancer Tablet Set was used. Whole-mount X-gal staining was performed as
described in Hogan et al. (Hogan et al., 1994).

3-3-4. Neural tube culture
E9 embryos were collected in HBSS on ice. The trunk segments were cut
out and treated with 0.15% of dispase for 10 min. The segments were gently
triturated with a Pasteur pipette to remove the surrounding tissues from neural
tubes. Isolated neural tubes were washed with HBSS solution containing 10% of
FBS to quench the dispase activity and plated onto fibronectin coated tissue
culture dishes. The explants were cultured in DMEM/F-12 medium (from
invitrogen) containing 10% of FBS (from invitrogen), 1x penicillin & streptomycin
(from Sigma) and 1 X ITS (insulin, transferin and selenium from Invitrogen) at
37°C in 8% CO2 for 48hrs. The cultures were fixed with 4% of buffered
formaldehyde for 5 min at room temperature and stained with X-gal solution for
4-5hrs.
60
3-3-5. Isolation of neural crest cells from 1st pharyngeal arch
The maxillary and mandibular processes were dissected from E10
embryos with microscissors in PBS. The dissected tissues were dissociated by
trypsinazation, and cell single cells suspensions were plated onto the fibronectin
coated dishes, cultured for 3-4 days, and stained for the lacZ activity as outlined
above. For immunofluorescence, the cultured cells were fixed for 5 min with 4%
of buffered formaldehyde. Antibodies specific for FAK (Upstate) and β-
galactosidase, and FITC-Phalloidin were used to detect focal adhesion
complexes, neural crest cell and F-actin, respectively.

3-3-6. Histology and TEM analysis
Embryonic tissues were fixed with 4% buffered formaldehyde for 12 hrs,
dehydrated and embedded in paraffin. Sections were stained with Hematoxylin
and eosin. For TEM analysis, the mandibular tissues were dissected from the
E11 embryos and fixed with 4% of electron microscopy grade formalin with 2.5%
of glutaraldehyde in 0.05% of cacodylate buffer for overnight. After three times
rinses with 0.05% of cacodylate buffer, the mandibular tissues were post-fixed
with 1% of osmium tetroxide in 0.05% of cacodylate buffer for 1 hr. To remove
the phosphate, the tissues were rinsed with distilled water and en bloc stained
with 3% of uranyl acetate in distilled water. With the serial ethanol, the tissues
were dehydrated and cleared with propylene oxide. The tissues were infiltrated
with Epon resin and embedded into the resin with the polymerization at 60ºC for
61
24 hrs. Ultrathin sections and picture recoding were processed at the
“Microscopy & Image Analysis Laboratory” in University of Michigan.

3-4. Results
3-4-1. Neural crest-specific Rac1 mutants
We deleted Rac1 in mouse NCCs by crossing females, which were
homozygous for the floxed Rac1 allele (Rac1
Flox/Flox
) with males, which were
heterozygous both for the floxed Rac1 allele and for the Wnt1-Cre transgene
(Rac1
Flox/+
/Wnt1-Cre
+/-
). The resulting Rac1
Flox/Flox
/Wnt1-Cre
+/-
mice (herein
designated Rac1/Wnt1-Cre) have Rac1 inactivated in the NC lineage, while the
other genetic combinations were used as controls. Wnt1-Cre- induced
recombination leads to efficient ablation of the Rac1 from the neural crest cells
(Figure 3-1).

Fig. 3-1 Whole-mount in situ hybridization of with Rac1 RNA probe with E10
embyros. In the Rac1/Wnt1-Cre mutant embryo, the Rac1 mRNA is
downregulated in the mandibular area (arrow).

Control Rac1/Wnt1-Cre
62
3-4-2. Rac1/Wnt1-Cre mutants display severe craniofacial defects
All Rac1/Wnt1-Cre mutant embryos died before the 13
th
embryonal day
(E13), and displayed severe craniofacial defects including cleft face and
hemorrhaging blisters in the first pharyngeal arch (Figure. 3-2). At E10, the
mutant embryos appeared superficially indistinguishable and pharyngeal arches
that are largely populated by migrating NCCs appeared very similar between
controls and mutants. However, already at this early stage some mutant embryos
displayed epithelial detachment from the underlying mesenchyme in the first
pharyngeal arch (Figure 3-2 D). At E11, the mutant heads appeared wider than
those of controls and abnormal blister-like epithelial detachments could now be
consistently seen in the mandibular arch. Histological analysis showed that
mutant pharyngeal arches contained relatively normal appearing mesenchymal
cells. However, blisters in the mandibular arch were now more prominent and
they could also be found in the mesenchyme of the nasal process. Also, some
mutant embryos showed blood-filled unilateral edema in the mandible. At E12, all
the mutant embryos showed severe mid-facial clefting, which resulted from a
failure of medial nasal processes to fuse. The blistering phenotype, which started
from the mandibular arch at E11, could now be seen consistently seen in the
maxillary processes. The mandible and tongue were small, but the eyes and
whisker follicles developed normally.

63

Fig. 3-2 Craniofacial defects in Rac1/Wnt1-Cre mutant embryos.
At E10, the Rac1/Wnt1-Cre conditional knockout embryos (B) appeared
superficially unaffected when compared to controls (A); however, on histological
sections epithelial detachment from the underlying mesenchyme (arrow in D)
was clearly detectable. At E11 (E-J), the mutant embryos displayed large,
occasionally blood-filled (asterisk in J) blisters in the mandibular arch (arrows in
F, H). Overall, mutant heads appeared wider than those of controls with a wide
separation between the medial nasal processes (F, I; asterisk). Epithelial
detachment could also be seen in the midline region between the two nasal
processes (H; arrowhead). At E12, the Rac1/Wnt1-Cre mutant embryo showed
severe mid-facial cleft (L; asterisk) and the rudimentary maxillary process (N;
asterisk).

Control Control Rac1/Wnt1-Cre Rac1/Wnt1-Cre
64
3-4-3. Depletion of Rac1 in the neural crest leads to severe outflow tract
defects and embryonic death
All Rac1/Wnt1-Cre mutant embryos died before embryonal day 13
suggesting that they suffered either from cardiac or vascular defects. Our
histological examination showed that all mutant embryos displayed a solitary
trunk of the outflow tract, also called as persistent truncus arteriosus or PTA.
(Figure 3-3). Moreover, the distal outflow tract was grossly dysmorphic displaying
dramatically enlarged derivatives of the third PAAs. These bilateral blood-filled
vascular structures were patent leading to the dorsal aorta (DA), which also was
atypically bilateral. However, the connection to the DA seemed to be rather
constricted, which may have further contributed to the distension of these
aberrant vessels. To further characterize the defects observed in the aortic sac
and pharyngeal arch arteries, we performed intracardiac Indian Ink injections at
E10 and E11. Consistent with our histolocigal findings, the third pharyngeal arch
arteries appeared abnormally large, while the 4
th
and 6
th
PAAs appeared patent,
but were abnormally small. Based on these findings, we conclude that the
embryonic lethality was caused by dysmorphic, and dramatically bloated ouflow
tract vessels, which lead to fatal hemodynamic changes.

65

Fig. 3-3. Abnormal patterning of pharyngeal arch arteries and outflow tract
defects in Rac1/Wnt1-Cre embryos. Left lateral view after intracardiac ink
injection at E10 (A, B) and E11 (C, D). At E10, the control embryo displayed a
well-formed third pharyngeal arch artery and still relatively small fourth and sixth
arteries (A), while in the mutant embryo, only the third artery was patent (B). At
E11, the control showed the well-formed 3
rd
, 4
th
and 6
th
arch arteries (C), while in
the mutant (D), the 3
rd
arch artery was very large, and the 4
th
artery appeared
grossly hypoplastic. H&E stained sections (E and F) showed that aortico-
pulmonary septation had failed in the mutant embryo (F) resulting in the common
arterial trunk, which displayed three abnormally shaped valve leaflets. Moreover,
the mutant embryo showed large aberrant vessels bilaterally (asterisks in F).
While the control showed the normal descending aorta (E), the mutant
demonstrated atypical left and right dorsal aorta (Pa, pulmonary artery; Tr,
trachea) (Magnification, 20x)

66
3-4-4. NCCs lacking Rac1 show defective cell migration and spreading in
vitro
Since Rac1 has a well-documented role in cellular migration in vitro, we
first examined the migratory potential of NCCs from Rac1/Wnt1-Cre embryos. To
visualize NCCs we used the R26R reporter assay as described (Soriano, 1999).
We isolated neural tubes from the control (Rac1
Flox/+
/Wnt1-Cre
+/-
/R26R
+/-
) and
mutant (Rac1
Flox/Flox
/Wnt1-Cre
+/-
/R26R
+/-
) embryos at E9.0 and cultured them for
48 hours on fibronectin-coated plates. To visualize cells that had migrated out
from the explant, the cultures were stained for β-galactosidase activity (Figure 3-
4). While the control cultures consistently displayed a large number of positively
staining migratory cells, the mutant cultures showed very few if any NC-derived
blue-staining cells suggesting that Rac1-deficient neural crest cells fail to migrate
on fibronectin in vitro.
Since Rac1 has been shown to play an important role in reorganization of
actin cytoskeleton, we tested whether NCCs lacking Rac1 would spread
normally. We dissected the 1st pharyngeal arches from E10 embryos,
dissociated them into single cell suspension and plated onto fibronectin-coated
plates. The cultures were harvested after 48 hours and stained for β-
galactosidase activity (Figure 3-4 A-D). More than 80% of cells in both mutant
and control cultures were derived from the NC. While the control NCCs spread
well and formed numerous flat membrane protrusions, the mutant NCCs spread
poorly and showed a long and thin morphology. In concordance with the impaired
67
spreading, the mutant cells showed poor attachment. About 80% of control cells,
but only about 20% of mutant cells had attached on fibronectin-coated plates in 5
hours (data not shown). This difference in cell attachment was so consistent that
it allowed reliable identification of a mutant genotype. After 3 days in culture, the
control and mutant NCCs were stained for F-actin and focal adhesion complexes
using phalloidin and FAK antibodies. The NC-identity was confirmed using
immunostaining β-galactosidase. Control neural crest cells spread well showing a
large number of focal adhesion complexes and a web-like actin filament network.
In contrast, the mutant neural crest cells displayed a very few focal adhesion
complexes and a few actin filaments, which were arranged parallel along the long
stretched cell membrane (Figure 3-4 E-L).
To identify migrating neural crest cells in vivo, E9 and E10 embryos
carrying the R26 reporter were stained for the β-galactosidase activity. Staining
patterns were carefully compared between controls and mutants; however, this
comparison did not reveal any detectable differences between these two
genotypes (Figure 3-5). Similarly, in situ hybridization for Cadherin-6 and Sox10
(markers for migrating NCCs; Figure 3-5, Figure 3-7) did not demonstrate any
significant overall differences in NCC migration at E9 and E10. To conclude,
NCCs deficient in Rac1 display defective cell-matrix interactions, abnormal
spreading and fail to migrate in vitro, but still display remarkably normal migration
in vivo, which is consistent with the histological findings that demonstrated a
large number of NC-derived mesenchymal cells in pharyngeal arches.
68
Control Control Rac1/Wnt1-Cre
Rac1/Wnt1-Cre

Fig.3-4. NCCs deficient in Rac1 failed to migrate in vitro, spread poorly and
showed a reduced amount of focal adhesion complexes.
Neural tubes from E9 embryos were cultured for 48 hours. NCCs were identified
using the R26 reporter assay (A, B). While control cultures showed a large
number of NCCs (blue cells, black arrows) that had migrated out from neural
tube explants (A), the mutant cultures failed to demonstrate any migrating NCCs
(B). Control NCCs spread well on fibronectin after 3 days in culture (C, black
arrows) and showed a web-like F-actin network (E, FITC-phalloidin staining,
green). In contrast, the mutant NCCs appeared spindle shaped, spread poorly
(D, black arrows), and showed only a few long stress fibers (J). The neural crest
cells were identified with immunostaining for β-gal (G, H; red). Immunostaining
for FAK demonstrated that NCCs deficient in Rac1 showed a reduced number of
focal adhesion complexes on fibronectin (J; green) when compared to control
cells (I). NCCs were identified with the ß-galactosidase staining (K, L; red).
Magnification: A-B, 10x; C-D and I-L, 20x; E-H, 40x.
69

Fig. 3-5. NCCs deficient in Rac1/Wnt-Cre migrate normally in vivo.
(A-D) Whole mount lacZ staining of E8.5, E9 and E10 embryos. The whole
mount lacZ staining showed that the neural crest cell migration occurs normally
in mutant embryos. Whole mount in situ hybridization for Cadherin6 (a marker for
migrating NCCs) showed no differences between the controls (E, G) and mutants
(F, H) (E, F; lateral view, G, H; dorsal view).

3-4-5. NCC survival, proliferation or rhombomere identity is not impaired
in Rac1/Wnt1-Cre mutants
Next, we analyzed whether the observed neural crest phenotypes could
be caused by deficient cell proliferation or aberrant cell death. Both control and
mutant samples displayed a similar number of BrdU and TUNEL tunnel positive
cells (Figure 3-6 A-D) suggesting that Rac1 deficiency does not cause dramatic
changes in cellular proliferation or cell death in postmigratory NCCs. We also
considered a possibility that some of the observed defects might be caused by
the loss of rhombomere identity of neural crest cells before or during their
migration. Therefore, we examined the maintenance of rhombomere identity at
Control Control
Rac1/Wnt1-Cre Rac1/Wnt1-Cre
70
E9 by analyzing expression of Krox20 and EphA4 (markers from rhombomeres 3
and 5, respectively) expression using whole-mount in situ hybridization (Ishii et
al., 2005). EphA4, which acts as an exclusive cell sorting factor, is also slightly
expressed in the rhombomeres 2 and 4. However, both Krox20 and EphA4
expression patterns in Rac1/Wnt1-Cre mutants were perfectly normal suggesting
that the Rac1 deficiency in the neural crest cells does not interfere with the
establishment of the rhombomere identity (Figure 3-6 E-H).

3-4-6. Rac1 is required for appropriate patterning and differentiation of
postmigratory NCCs in vivo
Neural crest cells contribute to development of afferent neurons and glial
cells of the cranial ganglia (Le Douarin and Kalcheim, 1999Go; Barlow, 2002).
Therefore, we analyzed patterning of cranial ganglia in Rac1/Wnt1-Cre embryos
using whole-mount immunohistochemistry at E11.0. In all analyzed samples, the
trigeminal ganglia were reduced in size, the superior cervical ganglia (III) were
misorganized, and the oculomotor nerve, superior ganglia (IX) and acoustic

71

Fig. 3-6. Normal apoptosis, cell proliferation and rhombomere identity in
Rac1/Wnt1-Cre mutants. Analysis of programmed cell death using TUNEL
staining at E10 (A-B), cell proliferation using BrdU staining at E10 and
rhombomere identity using whole mount in situ hybridization for Krox20 (specific
for the rhombomeres 3 and 5) and EphA4 (specific for the rhombomere 3) at
E9.0 showed no differences between controls (A, C, E, and G) and mutants (B,
D, F, H).

Control Rac1/Wnt1-Cre
72
ganglia (VIII) were missing. The glossopharyngeal nerve branched out from
superior ganglia (IX) as a combined structure with vagus nerve instead branching
out from jugular ganglia (X). In concordance with these findings, expression
pattern of Sox10 in Rac1/Wnt1-Cre mutants was similarly reduced in size in
trigeminal ganglia when compared to controls (Figure 3-7 A-B). To conclude,
these analyses imply that Rac1 in NCCs is required for appropriate patterning of
the cranial ganglia.
Another well-known function of NCCs is to populate PAAs and
differentiate to aSMA-positive smooth muscle cells during patterning of aortic
arch arteries. Immunostaining for aSMA showed that while positively staining
cells around PAAs could be seen both in controls and mutants, only in controls
the aSMA-positive cells formed a well-defined compact ring surrounding the
PAAs. In Rac1/Wnt1-Cre mutants the aSMA-positive cells formed diffuse clusters
suggesting that Rac1 function in NCCs is required for appropriate differentiation
to SMCs during PAA remodeling (Figure 3-7 E-F).

3-4-7. The blistering cyst of the mutant embryos is caused by the absence
of the anchoring fibrils in the basement membrane
Our histological analyses and immunostaining for the basement
membrane protein laminin suggested that epithelial blistering was caused by
detachment of the basement membrane from the underlying mesenchyme
73

Fig. 3-7. Abnormal patterning of cranial ganglia and smooth muscle cells in
Rac1/Wnt1-Cre. Embryos lacking Rac1 in NCCs demonstrate patterning defects
in NC-derived cranial nerve ganglia (A-D). Projections of the trigeminal ganglia
were significantly shorter in mutants (B, D) than in controls (A, C). The
oculomotor nerve was missing and glossopharyngeal (IX) and vagus nerves (X)
were fused (B; asterisk). A-B, Whole mount immunohistochemistry for
neurofilaments (NF) at E11; C-D, wholemount in situ hybridization for Sox10
(E10). Patterning defects in vascular smooth muscle cells surrounding the
common arterial trunk (E-H). In controls the endothelial lining of the aortic sac
and pharyngeal arch arteries was surrounded by a relatively compact layer of
aSMA-positive smooth muscle cells (E and G; arrowheads). In contrast, the
aSMA positive cells were diffusely distributed around the common arterial trunk
in Rac1/Wnt1-Cre mutants (F and H; arrowheads).

74
(Figure 3-8 A-B). To further explore this phenomenon, we used transmission
electron microscopy (TEM). Already at E11, the anchoring fibrils connected the
basement membrane to the NC-derived pharyngeal arch mesenchyme in
controls. In contrast, the mutants failed to demonstrate well-formed anchoring
fibrils (Figure 3-8 C-D). Moreover, mesenchymal cells adjacent to the blister
appeared more organized than those in other locations of the pharyngeal arch
mesenchyme (either in controls or in mutants), and displayed prototypical tight
junctions and visualized by TEM or ZO1 immunostaining (Figure 3-8 E-F).

75

Fig. 3-8. Abnormal connection between the basal lamina and the underlying NC-
derived mesenchyme in Rac1/Wnt1-Cre mutants.
In control mandibular arches at E11, the laminin-positive (red) basal lamina was
normally located underneath the epithelial cells layer and was in tight contact
with the underlying NC-derived mesenchyme (A). In the mutant mandibular arch,
the basal lamina has detached from mesenchyme (B; white arrow).Magnification,
20 x. TEM analysis of the basement membrane in mandibular arches at E11 (C,
D). In controls, the epithelium (E) and mesenchyme (M) were connected with
easily identifiable anchoring fibrils (arrow in C), while in mutants, there were only
a few rudimentary appearing fibrils visible (arrowhead in D). M, mesenchymal
cell; E, epithelial cells; B, epithelial cyst. Analysis of the formation of tight junction
in the mesenchyme under the blistering cyst with ZO-1 antibody (E, F). The
epithelium of nasal process is connected with the tight junction (E) and the
mesenchyme in the blistering cyst also form the prototypical tight junction (F,
white arrow)
Control Rac1/Wnt1-
Cre
76
3-5. Discussion

3-5-1. Epithelial blisters and mid-facial clefting
Deletion of Rac1 in the NC led to embryonic lethality around E12.5 - E13
with defects in craniofacial structures and the cardiac outflow tract. A consistent
early phenotype of mutant embryos is epithelial blistering that bears some
similarity to a rare human genetic disease epidermolysis bullosa (Ross et al.,
2008). According to our TEM results, the epidermal blistering in Rac1/Wnt1-Cre
mutant embryos may be caused by abnormalities in formation of appropriate
anchoring fibrils, which are mainly composed of collagen type VII produced by
the underlying NC-derived mesenchyme. In mutant embryos, the blistering
always starts to form from lateral aspects of the mandibular arch around E11, but
occurs later also in other sites in which the underlying mesenchyme is populated
by NC-derived cells, e.g., maxillary processes. Nevertheless, the lower jaw
developed relatively normally, while the upper jaw is severely malformed with a
characteristic mid-facial cleft. It has been previously reported that exposure of
mouse embryos at E11 to a terotogenic amino acid analogue Diazo-Oxo-
Norleucine (DON), causes mid-facial clefts (Burk and Sadler, 1983). This
phenotype was reported to result from a low mesenchymal cell density in the
midline of the nasal process. Our experiments showed that Rac1/Wnt1-Cre
embryos demonstrated a remarkably low cell density in this apparently critical
location, while there were large mesenchymal clusters lateral to the midline
region. This implies that the mid-face clefts seen in Rac1 mutants result from an
77
abnormal patterning of the NC-derived craniofacial mesenchyme during facial
development.

3-5-2. Rac1 deficiency and neural crest cell migration
Neural crest cells are very motile capable of migrating long distances in
vivo. Several genes have been associated with this fundamental property of
NCCs. Loss of Snail/Slug gene function, the earliest neural crest cell marker
related to neural crest induction causes defects in the NCC delamination from the
neural plate border and results in abnormal NCC migration (LaBonne and
Bronner-Fraser , 2000). Moreover, the ErbB4 and Krox20 genes have been
suggested to play an important role in establishment of the rhombomere identity.
The disruption of those two genes leads to an abnormal NCC migration (Golding
et al., 2000; Swiatek and Gridley , 1993), which differs from that caused by a
deficiency of Snail/Slug. Our present studies demonstrate that the Rac1/Wnt1-
Cre mutant embryos have a normal establishment of the rhombomere identity.
Moreover, the R26R lineage tracing and in situ hybridization for Cadherin6 and
Sox10 at E9 and E10 show that in the Rac1/Wnt1-Cre mutant embryos do not
have severe defects in NCC delamination or migration. However, some of the
Rac1/Wnt1-Cre mutant embryos have abnormal fusion of the superior (IX) and
jugular (X) ganglia. This may suggest the Rac1/Wnt1-Cre mutant embryos suffer
from defects in communication between migrating NCCs and the surrounding
matrix or other cell types similar to those seen in ErbB4 mutant embryos.
78
Perhaps, the most surprising finding of our study was the apparently
normal migration of Rac1-deficient NCCs in vivo, while the mutant cells appeared
non-migratory on fibronectin in vitro. Although the Wnt1-Cre driverline has been
used in numerous studies, and has been proven to be a consistent and robust
inducer of recombination, it is still possible that the presence of some of the
migratory Rac1-/- cells could be explained either by the partial recombination
efficiency or by exceptional protein stability. However, our neural tube culture
assay showed that at E9.0 the recombination efficiency is close to 100% and the
Rac1 function is completely lost, since, unlike the control cells, practically all
Rac1-deficient NCCs were unable to migrate in vitro. Therefore it is highly
unlikely that the apparently normal NCC migration in Rac1/Wnt1-Cre mutants in
vivo, particularly after E9.0, could result from a failure of Wnt1-Cre to efficiently
ablate the functional Rac1 from NCCs.
So far, most of the Rac1 function on a cellular level related to the cell
migration has been derived from migration studies in vitro. Cell migration in vivo
is a result of a tightly-regulated coordination of autonomous cell behavior and
dynamic changes in the extracellular matrix (Zamir EA et al. 2008). These
dynamic changes will be ignored in standard migration assays in vitro, which will
likely explain our findings that Rac1 plays a non-redundant role in NCC migration
in vitro, but not in vivo.

79
3-6. Chaper III References
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Cornell RA, Eisen JS., 2005. Notch in the pathway: the roles of Notch signaling in
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Creuzet S, Couly G, Le Douarin NM., 2005. Patterning the neural crest
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Golding JP, Trainor P, Krumlauf R, Gassmann M., 2000. Defects in pathfinding
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Guillou H, Depraz-Depland A, Planus E, Vianay B, Chaussy J, Grichine A,
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Hogan B., Beddington R., Constantini F and Lacy E., 1994 Manipulating the
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Huang X, Saint-Jeannet JP., 2004. Induction of the neural crest and the
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Ishii M, Han J, Yen HY, Sucov HM, Chai Y, Maxson RE Jr. 2005. Combined
deficiencies of Msx1 and Msx2 cause impaired patterning and survival of the
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Jiang X, Rowitch DH, Soriano P, McMahon AP, Sucov HM., 2000. Fate of the
mammalian cardiac neural crest. Development. 127, 1607-16.

LaBonne C, Bronner-Fraser M., 2000. Snail-related transcriptional repressors are
required in Xenopus for both the induction of the neural crest and its subsequent
migration. Dev. Biol. 221, 195-205.

Le Douarin NM, Creuzet S, Couly G, Dupin E., 2004. Neural crest cell plasticity
and its limits. Development. 131, 4637-50.

McMahon JA, Takada S, Zimmerman LB, Fan CM, Harland RM, McMahon AP.,
1998. Noggin-mediated antagonism of BMP signaling is required for growth and
patterning of the neural tube and somite. Genes Dev. 12, 1438-52.

Monsoro-Burq AH, Fletcher RB, Harland RM., 2003. Neural crest induction by
paraxial mesoderm in Xenopus embryos requires FGF signals. Development.
130, 3111-24.

Nichols DH., 1986. Formation and distribution of neural crest mesenchyme to the
first pharyngeal arch region of the mouse embryo. Am. J. Anat. 176, 221-31.

Nie X, Luukko K, Kettunen P., 2006. BMP signalling in craniofacial development.
Int. J. Dev. Biol. 50, 511-21.

Noden DM, Trainor PA., 2005. Relations and interactions between cranial
mesoderm and neural crest populations. J. Anat. 207, 575-601.

Pankov R, Endo Y, Even-Ram S, Araki M, Clark K, Cukierman E, Matsumoto K,
Yamada KM., 2005. A Rac switch regulates random versus directionally
persistent cell migration. J. Cell. Biol. 170, 793-802.

Ross R, DiGiovanna JJ, Capaldi L, Argenyi Z, Fleckman P, Robinson-Bostom L.,
2008. Histopathologic characterization of epidermolytic hyperkeratosis: a
systematic review of histology from the National Registry for Ichthyosis and
Related Skin Disorders. J. Am. Acad. Dermatol. 59, 86-90.

Richman JM, Lee SH., 2003. About face: signals and genes controlling jaw
patterning and identity in vertebrates. Bioessays. 25, 554-68.

81
Ridley AJ, Schwartz MA, Burridge K, Firtel RA, Ginsberg MH, Borisy G, Parsons
JT, Horwitz AR., 2003. Cell migration: integrating signals from front to back.
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Sasai Y, De Robertis EM., 1997. Ectodermal patterning in vertebrate embryos.
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Sugihara K, Nakatsuji N, Nakamura K, Nakao K, Hashimoto R, Otani H,
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82
Chapter IV

Conditional and conventional mutants for Trim33 - the Trim33 conventional
knockout mice are embryonic lethal

4-1. Summary
Trim33 (Tif1γ, ectodermin, moonshine), a member of the TIF1 family of
transcriptional coactivators and corepressors, is a large nuclear protein that
contains an N-terminal tripartite (Trim) domain composed of a RING domain, two
B-box domains, and a coiled coil domain. It has been suggested that Trim33
(Ectodermin) mediates ectodermal induction in the Xenopus by functioning as a
Smad4 ubiquitin ligase, while in the zebrafish Trim33 (moonshine) has been
reported to act as a R-Smad binding protein in induction of erythroid
differentiation. Since the developmental role of Trim33 in mammals is currently
unknown, we generated mice carrying the conditional Trim33 (Trim33FX) allele
by flanking exons 2–4 encoding most of the functionally critical N-terminal
tripartite domain by loxP sites. We confirmed the null genotype by using the EIIa-
Cre transgenic approach to create mice that lack exons 2–4. Embryos deficient
in Trim33 die during early somitogenesis, demonstrating that Trim33 plays an
important nonredundant role in mammalian embryonic development.

83
4-2. Introduction
Trim33 was identified in 1999 as a new member of the TIF1
(transcriptional intermediary factor) family by screening a cDNA library using a
Tif1α probe (Venturini et al., 1999). So far, 37 Trim genes have been discovered
in mammals, which can, as a result of alternative spicing, generate more than 70
different Trim proteins (Table 4-1) (Reymond et al., 2001). All Trim proteins share
an evolutionary conserved N-terminal tripartite (RBCC) motif (Peng et al., 2002),
which is composed of the Ring, B-box type 1, B-box type 2 and Coiled-coil
domains. While the N-terminal RBCC domain offers a common structural
character for the Trim protein family, the other motifs, such as RFP-like, NHL,
TSS, PHD/TTC or Bromodomain determine functional features of each Trim
protein. According to their functional activities, the Trim proteins can be
subdivided into several subfamilies. Among the more than 70 Trim proteins,
Trim24 (Tif1a), Trim28 (Tif1β) and Trim33 (Tif1g) proteins are sub-categorized
into the TIF1 (transcriptional intermediary factor) family due to the presence of
the highly conserved TSS-PHD/TTC-Bromodomains in the C-terminus (Nielsen
et al., 1999).

84

Table. 4-1 Trim family proteins.

85
The TSS-PHD/TTC-Bromodomain has been found only in nuclear proteins
having a role in chromatin remodeling. The Bromodomain recognizes acetylated
lysine residues in other transcription factors, while the PHD/TTC domain is
conserved in transcriptional mediators. By functional interaction with CBP/p300
histone acetyltransferase, PHD/TTC domain functions as a transcriptional co-
activator. TSS, as a TIF1 signature sequence, is composed of 25 amino acids
and has an essential role in the transcriptional inhibitory activity of the TIF1 family
proteins. Unlike Trim33 (Tif1g), Trim24 (TIF1α) and Trim28 (TIF1β) proteins have
a HP1 interaction domain that is involved in the heterochromatin-mediated
silencing through interaction of heterochoromatin-associated factors, such as
HP1α, HP1β and HP1γ. They have a binding site to KRAB domain of vertebrate
Krüppel-type zinc finger proteins that repress transcription (Abrink et al., 2001).
Moreover, Trim24 has a NRID domain (nuclear receptor interaction domain)
consistent with its function as a regulated nuclear receptor (Heery et al., 1997;
Yan et al., 2004). This domain is not present in Trim28 and Trim33 (Fig. 4-1). To
conclude, while TIF1 family proteins share similar structural motifs and may
share comparable transcriptional inhibitory functions, the detailed regulatory
mechanism may be quite different.
So far, functions of Trim24 and Trim28 have been analyzed in vivo by
using the conventional gene targeting technique. Trim24 shows the liver-specific
tumor suppressor function (Khetchoumian et al., 2007). However, it does not
seem to have any major nonredundant developmental functions. In contrast, a
86

Fig. 4-1 Nucleic and amino acid sequences of hTIF1α (a) and hTIF1γ (b).
Sequence positions are indicated on the left; characteristic domains and motifs
are outlined and named on the right. The putative nuclear localization signals
(KRK/KKK or KK/KKK, respectively) and an LxxLL motif in hTIF1g, as well as
stop codons, are underlined [Reproduced from (Venturini et al., 1999)].

87
Trim28 knockout study shows that this gene is essential in embryogenesis,
especially during the early postimplantation period (Cammas et al., 2000). So far,
several groups have reported Trim33 functions in several organisms including
zebrafish, frog and mouse. Depending on the organism, the Trim33 has several
synonyms; moonshine in the zebrafish, ectodermin in the frog, TIF1γ in the
mouse and human. Zebrafish having a mutation in the moonshine gene show
severe defects in the primitive hematopoiesis affecting a function of the key
hematopoietic transcription factors, e.g., gata1, scl, gata2 and ikaros (Figure 4-2
A) (Ransom et al., 2004). The rare surviving adult mutant zebrafish show a
dramatic reduction in a number of differentiated erythroid cells. Another study
with the human umbilical cord blood also identified a function for TIF1γ in
erythoropoiesis (He et al., 2006). Attenuation of TIF1γ using shRNA in CD34+
hematopoietic stem/progenitor cells resulted in reduction of a number of fully
differentiated erythrocytes (Figure 4-2 B). In addition, these investigators
discovered that TIF1γ bound specifically to activated Smad2/3, and thus
functioned as an inhibitor of Smad4. They also suggested that, when TIF1γ
interrupts the canonical Tgf-β signaling by competing with Smad4, the
hematopoietic stem/progenitor cells escape from their undifferentiated stage and
differentiate to erythrocytes (Figure 4-3). At the same time, another group
reported that in the Xenopus, Trim33 (Ectodermin) is responsible for the
ectoderm formation (Dupont et al., 2005). The function of Ectodermin was

88
A
B

Fig. 4-2. Defective Trim33 function cause abnormal hematopoiesis in zebrafish
(A) and human (B) [Reproduced from(Ransom et al., 2004; He et al., 2006)].

Fig. 4-3 TIF1γ as a branch in the TGFb-activated Smad pathway [Reproduced
from (He et al., 2006)].
89
discovered by screening a blastula-stage cDNA library. Each mRNAs expressed
in the cDNA library was injected into frog embryos and cDNAs which could
induce the ectoderm specific marker genes were identified. One of these cDNAs
encoded a novel protein that was named Ectodermin, which turned out to be a
frog homolog of Trim33. The function of Ectodermin as an ectoderm determinant
was confirmed with the knock-down of Ectodermin using morpholinos (Figure 4-4
A). These investigators discovered that the function of Ectodermin is to maintain
the ectoderm identity by inhibiting Tgf-β signaling coming from the mesoderm.
They suggested that when ectodermal cells are stimulated with Tgf-βs,
Ectodermin recognizes the R-Smad/Smad4 complex. After binding to the Smad4
of the complex, Ectodermin recruits the ubiquitin proteinase and blocks the
canonical Tgf-β signaling by degradation of R-Smad/Smad4 complexes (Figure
4-4 B). In conjunction with the suggested function of TIF1γ in the hematopoiesis,
it sems that Trim33 acts as a regulator of Tgf-β signaling. However, the detail
functional mechanisms are still controversial. In this study, the Trim33 function is
investigated by a generation mice that carry both conventional and conditional
alleles of Trim33 (Kim and Kaartinen, 2008). While Trim28 conventional knockout
embryos die at the postimplantation stage, the Trim33 conventional knockout
embryos die during early somatogenesis. This shows that Trim33 has an
essential nonredundant function in the embryogenesis. Moreover, an induced
deletion of the Trim33 gene after early somatogenesis shows that the Trim33
90
may be responsible for the embryonic survival after neurulation and body axis
formation.

Fig. 4-4. Ectodermin is required for the formation of ectoderm (A) by preventing
Tgf-β and Bmp signaling in the ectoderm area (B) [Reproduced from (Dupont et
al., 2005)].

A
B
91
4-3. Materials and Methods
4-3-1. Construction of a targeting vector
RP2468C16 Bac DNA was used as a template to PCR amplify both short
and long arms as ClaI-XhoI and KpnI-SalI fragments, respectively. High-fidelity
Supermix (Qiagen, Valencia, CA) polymerase was used for amplification, and the
generated arms were sequenced to verify that no PCR-generated mutations
were introduced. A single loxP site was inserted into intron 4 by replacing a small
553-bp SacI-PstI fragment with a loxP sequence flanked by SacI and PstI
restriction sites. Subsequently, a loxPNeoloxP cassette was inserted as a XhoI-
NotI fragment into the pKODT plasmid. A long arm containing the single loxP
sequence in intron 4 was inserted as a KpnI-SalI fragment into
loxPNeoloxP/pKODT, and finally a short arm was added into the construct as a
ClaI-XhoI fragment. The targeting vector was electroporated into TVB2 mouse
ES cells, and recombinant ES cell clones were selected with G418 as described
(Kaartinen et al., 2004, 1995).

4-3-2. Screening of correctly targeted embryonic stem cells
ES cell DNAs were first screened for correct targeting by PCR using a
forward primer (a’) 5’ CACGACACAAAGAACTGTAG 3’ and a reverse primer (b’)
5’ CAAGCAAAACCAAATTAAGG 3’. Subsequently, the presence of the single
loxP site in intron 4 was verified by PCR using a forward primer (c) 5’
CATTGTGCTTCACCTCCTCCTCTTCG 3’ and a reverse primer (d) 5’
92
GGGAGGGAAAATCTGGCTGAA 3’.

4-3-3. Generation of mutant mice
Mouse chimera were generated by injecting correctly targeted ES clones
into the C57BL/6J mouse blastocysts. Trim33
fxneo/+
mice were generated by
crossing Trim33 chimeric male mice (which were potent germ-line transmitters)
with wild type female, B6xCBA (F1). To remove the neo selection marker, the
Trim33
fxneo/+
female mice were crossed with EIIa-Cre male mice and
Trim33
mosaic
/EIIa-Cre mice were generated. By crossing again with wild type
mice, B6CBAF1, the partial and complete Trim33 recombination alleles were
segregated into Trim33
fx/+
and Trim33
ko/+
alleles. Trim33
ko/+
/Rosa;rtTA/tet-O-Cre
mice were obtained by introducing the Rosa;rtTA and tet-O-Cre alleles into the
Trim33
ko/+
background. Fig. 4-5 shows the overall breeding scheme to get Trim33
flox and Trim33 null mice.
Trim33FX mice were genotyped using forward and reverse primers (a) 5’
GCACCTTGATGAGATCTTCCTCCTCC 3’ and (b) 5’
GACGACATACTGGACACCGTA 3’, while Trim33KO mice were genotyped using
forward and reverse primers (a) 5’ GCACCTTGATGAGATCTTCCT CCTCC 3’
and (d) 5’ GGAGGGAAAATCTGGCTGAA 3’.

93

Fig. 4-5 Breeding scheme to get the Trim flox and Trim null knockout mice.

4-3-4. Reverse transcription – PCR
Total RNA was isolated from E8 embryos using the RNeasy mini kit
(Qiagen), and cDNAs were synthesized by the Omniscript reverse transcription
kit (Qiagen) according to the manufacturers’ protocols. Subsequently, the cDNAs
were analyzed by PCR for Trim33 expression using the following primer pairs. β-
actin was used as a quality and loading control.
Trim33-ex4-S 5’ GAGTCTGTTGGAACATCTGGTCAGCG3’
Trim33-ex7-AS 5’ GGCCTGTGATATCATTCTGCTGCTGT3’
94
Trim33-ex1-S 5’ GGTGTGTCAGCAGAGCTTGCA3’
Trim33-ex6-AS 5’ GATAAGGGTGAAGATGGCCACT3’
Trim33-ex15-S 5’ ACCTCATGCACAGGTCGGCAAGGAT3’
Trim33-ex20-AS 5’ GCTCAAACTCTGGCAAAGGAGTGAAG3’
b-actin-S 5’ GTGGGCCGGTCTAGGCACCAA3’
b-actin-AS 5’ CGGTTGCCTTAGGGTTCAGG3’

4-3-5 Immunohistochemistry
This method has been described in the chapter III

4-4. Results and Discussion
4-4-1. Trim33 expression pattern during mouse embryogenesis
The Trim33 protein is produced ubiquitously in many tissues including the
nasal process, palatal shelves and the oral ectoderm (at E14) and the spinal
cord, heart lung and muscle (E17) (Figure 4-6). Even though Trim33 is also
expressed in the mesenchyme, it is highly expressed in the epithelium, especially
the palatal shelves’ epithelium, oral ectoderm, nasal process and lung epithelium.

95

Fig. 4-6 Trim33 protein expression in the craniofacial area at E14 (A) and thorax
at E17 (B). Trim33 protein is expressed ubiquitously, however, it is strongly
expressed in the epithelium.

A
B
96
4-4-2. Conventional and conditional knockout alleles for Trim33.
To address the role of Trim33 in mammalian embryogenesis, we generated
mice harboring the conditional knockout allele for Trim33. Our strategy was to
flank exons 2–4 encoding the functionally critical RING and B1 and B2 Box
domains by loxP sites. The short (1.6 kb) and long (8.2 kb) arms of the targeting
vector were PCR amplified from a Bac genomic DNA. The loxPNeoloxP cassette
was inserted into intron 1 and a single loxP site was inserted into intron 4 (Figure
4-7). As a negative selection marker we used the diphtheria toxin (DT) gene. The
linearized targeting vector was electroporated into TVB2 embryonic stem cells as
described (Yang and Kaartinen, 2007). Fifteen out of 261 G418-resistant
colonies were targeted to the correct locus; however, only three of them
contained the distant 3’ loxP site. All three correctly targeted ES clones were able
to produce highly chimeric male mice, which in turn were potent germ line
transmitters. To remove the Neo selection marker and to generate a presumed
knockout allele for Trim33, we crossed the Trim33
FXNeo/FXNeo
mice (homozygotes
for the targeting vector) with EIIa-Cre transgenic mice (Xu et al., 2001). In these
mice, Cre is expressed under the control of the adenoviral EIIa promoter that
targets expression of the Cre recombinase to the early mouse embryo (Lakso et
al., 1996). The EIIa-Cre transgene creates both partial and complete
recombinations and therefore EIIa-Cre transgenic mice can be used both as a
deleter mouse to generate knockout alleles and to remove selection markers

97

Fig. 4-7 Trim33 targeting vector and screening of ES colonies. A: A schematic
presentation of the Trim33 protein domains, Trim33 genomic locus depicting a
segment from exon1 to intron 4, the Trim33 targeting vector and the Trim33
targeted allele. B: An example of a PCR screen (upper panel); positive clones
#102 and #113 can be easily identified. PCR analysis using c/d primers and
template DNA from clones that suggested the correct targeting demonstrated
that only clones number #40, #119, and #259 (not shown) had retained the 30
loxP site (lower panel). The wild type allele produces a 713-bp amplification
product. Since the strategy to insert the 30 loxP site into intron 4 involved the
replacement of a 553-bp SacI-PstI fragment with a loxP site, the mutant allele
gives rise only to a 265-bp amplification product.

98
(e.g., loxP-pGKNeo-loxP) when a triple loxP-strategy is used (Holzenberger et
al., 2000; Xu et al., 2001). The obtained mosaic male mice progeny were
subsequently crossed with wild-type female mice to obtain the floxed (Trim33
FX
)
and null (Trim33
KO
) alleles (Figure 4-8). Homozygote Trim33
FXFX
mice were
viable and fertile, and they did not display any recognizable phenotypes.

Fig. 4-8 Generation of mice carrying the floxed (Trim33
FX
) and knockout
(Trim33
KO
) Trim33 alleles. Transgenic EIIa-Cre mice were crossed with mice
homozygous for the targeting vector. The obtained mosaic males were further
crossed with wild-type females to obtain Trim33
FX/WT
mice (type I recombination;
red arrow) and Trim33
KO/WT
mice (type II recombination; blue arrows).

99
4-4-3. The Trim33 null knockout cause embryonic lethality around E9.5.
To confirm that the Trim33
KO
allele encoded the true null allele and to
provide initial information about the biological role of Trim33 during
embryogenesis, we intercrossed the heterozygote Trim33
KO/WT
mice to obtain
homozygote Trim33
KO/KO
mice (Figure 4-9). Genotype analyses of newborn
offspring revealed that all the homozygote mutant pups died during gestation
(Table 4-2). To examine the time frame during which embryonic lethality
occurred, we harvested embryos at different time points and discovered that at
E9 the mutant embryos displayed a dramatic developmental delay when
compared with controls (Table 4-2 and Figure 4-9 H–J). Nevertheless, they had
formed a body axis, displayed head folds and the neural tube, and showed 5–6
somite pairs. We were unable to discover any living mutant embryos after E9.5.
At E8.0–8.5 (3–6 somite pairs in controls), the Trim33 mutant embryos were
aligned at the base of the yolk sac, and while they had formed the anterior–
posterior body axis and identifiable anterior structures, for example, head folds
(arrows in Figure 4-9 D, E, and G), it was difficult to identify any other embryonal
structures. RT-PCR using primers with target sequences in exons 4 and 7 did not
produce any detectable amplification product, while the heterozygote and wild-
type samples showed the expected 423-bp fragment (Figure 4-9 B). This is
concordant with the lack of sequences encoded by exon 4. To further validate the
Trim33
KO/KO
allele, we used exon 1- and exon 6-specific primers. The wild type
and homozygote Trim33
KO/KO
alleles produced the expected 733-bp and 366-bp
100
products, respectively, while the heterozygote samples gave rise to both PCR
fragments (Figure 4-9 B). RT-PCR analysis using primers specific for 30 exons
15 and 20 produced the expected 684-bp fragment from wildtype, heterozygote
(Trim33
KO/WT
) and homozygote (Trim33
KO/KO
) samples suggesting that the
mutated allele lacking exons 2–4 is able to produce a stable mRNA (Figure 4-9
B). To conclude, our RT-PCR analyses demonstrated that the mRNA encoded by
Trim33
KO/KO
allele lacks sequences encoded by exons 2–4, and that it is highly
likely that the phenotype observed in homozygote samples results from a loss of
Trim33 function, particularly since most of the possible splicings, for example,
exon 1 to 4, will lead to a frameshift and premature translational stop. These
mutant mRNAs would produce only the very N terminal peptide encoded by
Trim33 exon 1. Only the splicing from exon 1 to 9 or exon 1 to 13 would maintain
the open reading frame. In these unlikely hypothetical cases the protein product
would lack the tripartite motif, but would contain the C-terminal PHD and Bromo
domains. Based on the current knowledge, it is impossible to say whether these
aberrant proteins lacking the functionally important tripartite motif would posses
any biological activity.

101

Fig. 4-9 Development of embryos deficient in Trim33 is arrested at the early
somitogenic stage. A: PCR genotyping of embryos from the crossing between
Trim33KO/WT males and females at E8.5. HE, heterozyggotes; WT, wildtype;
HO, homozygotes. B: RT-PCR analysis of embryos harvested at E8.5 using
primers for exons 4 and 7 demonstrates that Trim33KO/KO embryos do not
display any detectable mRNA product, while wild-type and heterozygote controls
show an expected 423-bp PCR product (B, upper panel). Corresponding analysis
using primers for exons 1 and 6 show that the wild-type sample produces the
expected 733-bp PCR product, the homozygote mutant sample produces the
expected 336-bp product, while the heterozygote samples gives rise to both 733-
bp and 336-bp products (second panel from the top). Corresponding analysis
using primers for exons 15 and 20 shows that both the wild-type and mutant
alleles produce a 684-bp amplification product (second panel from bottom). b-
actin specific primers produced a 245-bp amplification product with comparable
intensity from all samples (bottom panel). At E8.2–8.5, Trim33 null mutants (D, E,
G) demonstrate retarded development when compared to controls (C, F). D and
E depict both lateral and frontal images of the same embryo, respectively. I:
Development of the Trim33 2/2 embryos is arrested at the early somitogenic
stage (E9.0). J: A high power picture of a Trim33 2/2 embryo shown in (I). H:
Control littermate. Arrows in C, D, E, G, and J point to head folds, white
arrowhead in E points to the primitive streak, and black arrowhead in J points to
the somites.

102
4-4-4. The Trim33 conditional knockout during E9 to E15 cause severe
embryonic axis formation defect and failure of neural tube closure.
By using the inducible conditional knockout strategy, the Trim33 function
after early neurlation were analyzed. As a Cre driver mouse, the Rosa rtTA;tet-O-
Cre mouse was crossed with Trim33
fxfx
mice (Figure 4-10 A). The cre expression
was induced by doxycycline food from E9 to E15 stage. The mutant embryo died
with severe embryonic axis formation defect and the failure of neural tube closure
(Figure 4-10 B).

Fig. 4-10 The deletion of Trim33 function after neurulation causes the twisted
body axis and neural tube closure failure.
A
B
103
4-4-5. Male mosaic EIIa-cre mice can be efficiently used to segregate the
floxed alleles.
During the generation of conditional knockout mice, loxPneoloxP cassette
is introduced into one of the intron. This cassette contains a positive selection
marker (the neomycin resistance gene) flanked by loxP sites. The neo gene is
usually driven by a strong mammalian promoter, such as the PGK promoter.
Even though the neo gene is introduced into an intron, due to its robust
expression capacity, the neo gene can alter the endogeneous gene expression
pattern. Therefore, it is important to remove the neo gene from a conditional
allele. Several methods have been reported to remove the neo gene as well as to
segregate the floxed allele from other alleles. While Kaartinen V. et al used
adenoviral Cre recombinase to remove the neo cassette, Martin H. et al and Xu
X et al used the transgenic EIIa-Cre mouse line, which was generated in 1996 by
Lakso et al. In an early undifferentiated embryo, such as one-cell stage zygote,
there is specific protein function which has “E1A-like activity.” E1A is an
adenoviral transactivator. It was discovered that this E1A also could activate the
adenoviral EIIa promoter. By introducing the adenoviral EIIa promoter into the
mouse genome, EIIa-Cre mouse was generated. In this mouse Cre is expressed
only for a short period of time simultaneously with the “E1A-like” protein. To
remove the neo cassette and to segregate the floxed allele from the mosaic EIIa-
Cre mice, we used both female and male mosaic EIIa-Cre mice. We obtained 7
pups with the floxed allele by crossing of the male mosaic EIIa-Cre mouse and
104
wild type females. However, the female mosaic mouse produced efficiently
knockout alleles, but failed to produce any floxed alleles. Perhaps, in oocytes
there are pre-existing “E1A-like activity” proteins, and therefore a high level of
Cre would be expressed. If the “E1A-like activity” protein is introduced later, a
level of Cre expression is lower and the three different kinds of recombination
events can occur. To conclude, it is recommendable to use male mosaic EIIa-cre
mice to segregate the floxed alleles.

105
4-6. Chaper IV References
Abrink M, Ortiz JA, Mark C, Sanchez C, Looman C, Hellman L, Chambon P,
Losson R., 2001. Conserved interaction between distinct Krüppel-associated box
domains and the transcriptional intermediary factor 1 beta. Proc Natl Acad Sci U
S A. 98, 1422-6.

Cammas F, Mark M, Dollé P, Dierich A, Chambon P, Losson R., 2000. Mice
lacking the transcriptional corepressor TIF1beta are defective in early
postimplantation development. Development. 127, 2955-63.

Dupont S, Zacchigna L, Cordenonsi M, Soligo S, Adorno M, Rugge M, Piccolo
S., 2005. Germ-layer specification and control of cell growth by Ectodermin, a
Smad4 ubiquitin ligase. Cell. 121, 87-99.

He W, Dorn DC, Erdjument-Bromage H, Tempst P, Moore MA, Massagué J.,
2006. Hematopoiesis controlled by distinct TIF1gamma and Smad4 branches of
the TGFbeta pathway. Cell. 125, 929-41.

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Mol Biol. 320, 629-44.

106
Ransom DG, Bahary N, Niss K, Traver D, Burns C, Trede NS, Paffett-Lugassy N,
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107
Chapter V
Conclusions

Tgf-ß signaling plays a critical role in craniofacial development. So far,
many studies have elucidated detailed features of the canonical signaling
pathway and its essential role in the craniofacial development. However, the non-
canonical Tgf-ß signaling pathway, particularly in vivo, is still very poorly
understood. In this study, I have used three different strategies to investigate the
function and mechanisms of non-canonical Tgf-ß signaling pathways, especially,
in the craniofacial development.

5-1. Alk5 forms functional signaling complexes with unconventional type II
receptors.
By comparing the craniofacial phenotype of mice lacking Alk5 in neural
crest cells to comparable mutants lacking the prototypical binding partner of Alk5,
i.e., the Tgf-β type II receptor, we noticed that the phenotypes of mice deficient in
Alk5 were consistently more severe than those of mice lacking Tgfbr2
(Choudhary et al., 2006; Dudas et al., 2006; Ito et al., 2003; Wang et al., 2006).
Therefore, we decided to study, whether other type II receptors could also induce
activation of Alk5. To address this question, we used the Ariad
heterodimerization system that allows us to pull receptors together in a ligand
independent manner (Stockwell et al., 1998).
108
The cytoplasmic domain of Alk5 and TgfRII proteins were over-expressed
in the CHO cells and the formation of conventional Tgf-ß receptor complex was
induced by an artificial small molecule heterodimerizer. Phosphorylation of
Smad2 proteins was confirmed. With an identical strategy Alk5 and Acvr2B were
coexpressed to investigate the catalytical activity of this non-conventional Tgf-ß
receptor combination. We confirmed the Smad2 proteins were also
phosphorylated by the non-conventional Tgf-ß receptor complex, even though
the level of phosphorylation of Smad2 protein was less than that induced by the
conventional Tgf-ß receptor complex. This result suggests that the non-canonical
Tgf-ß receptor complex may act as a functional signaling mediator and that not
only the Alk5 and Acvr2B combinations, but also other non-canonical receptor
combinations may be possible.

5-2. Rac1 is indispensable in the craniofacial and cardiac neural crest, but
is not required for NCC migration in vivo
We discovered that a function of the small Rho-related GTPase Rac1 is
essential in the craniofacial and in cardiac development. Rac1 was deleted
specifically in the neural crest by crossing mice carrying the floxed Rac1 allele
with transgenic Wnt1-Cre mice. The mutant embryos displayed a prominent
midfacial cleft, small mandible, aortico-pulmonary septation defects and
abnormally large outflow tract vessels, and died around E13. In vitro studies
suggest that the mutant neural crest cells have defects in appropriate cell
109
attachment, and in formation of sub-cellular structures that are required for cell
migration. However, in vivo studies show that neural crest cells deficient in Rac1
can migrate relatively normally. Instead, neural crest cells lacking Rac1 function
display a defective differentiation capacity leading to an aberrant patterning of the
cranial ganglia and abnormal differentiation of smooth muscle cells surrounding
the pharyngeal arch arteries. So far, most of the knowledge on Rac1 functions in
cell migration has been derived from migration studies in vitro. Cell migration in
vivo is a result of a tightly-regulated coordination of autonomous cell behavior
and dynamic changes in the extracellular matrix (Zamir EA et al. 2008). These
dynamic changes will be ignored in standard migration assays in vitro, which will
likely explain our findings that Rac1 plays a non-redundant role in NCC migration
in vitro, but not in vivo.

5-3. Trim33 conditional and conventional knockout mice were generated -
the Trim33 conventional knockout mice are embryonic lethal
As one of the transcriptional co-regulators, the Trim33 has been identified
as a new regulator related to the Tgf-ß signaling pathway. Even though several
functional mechanisms of Trim33 have been suggested, the mechanisms are still
arguable, and the detail functional mechanism is not clear. In this study, we
generated mice carrying conditional and knockout alleles for Trim33. The null
knockout of Trim33 caused embryonic lethality around E9. The mutant embryos
started to show delayed development around E8.5 and died around E9. These
110
results demonstrate that Trim33 is an essential factor for the normal early
embryonic development. With the Trime33
fx
generated in this study, we can
identify more diverse functions of Trim33 depending on the embryonic stage and
tissue type.

111
5-4. Chaper V References
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Dudas M, Kim J, Li WY, Nagy A, Larsson J, Karlsson S, Chai Y, Kaartinen V
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Ito Y, Yeo JY, Chytil A, Han J, Bringas P Jr, Nakajima A, Shuler CF, Moses HL,
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Stockwell BR, Schreiber SL. 1998. Probing the role of homomeric and
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Wang J, Nagy A, Larsson J, Dudas M, Sucov HM, Kaartinen V 2006. Defective
ALK5 signaling in the neural crest leads to increased postmigratory neural crest
cell apoptosis and severe outflow tract defects. BMC Dev Biol. 6, 51.

112
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Abstract (if available)

Abstract Craniofacial malformations including cleft lip, cleft palate and craniosynostosis are among the most common birth defects in humans. A Tgf-β signaling pathway has been shown to be important during craniofacial development. Detailed mechanisms of the canonical Tgf-β signaling pathway in vitro have been well established, and a significant amount of information has been accumulated about a role of the canonical Tgf-β signaling pathway in craniofacial development. However, the canonical Tgf-β signaling pathway alone is not enough to explain all of the published findings of different craniofacial phenotypes in mice harboring mutations in genes encoding Tgf-β signal transduction components in vivo. Therefore, we hypothesized that a non-canonical Tgf-β signaling pathway also acts as an essential player in craniofacial development. To this end, we studied a function of the non-canonical Tgf-β signaling pathway in craniofacial development using three different approaches. First, we discovered that non-conventional Tgf-β receptor combinations can also act as functional receptor complexes. Second, we studied a role of Rac1, as a possible downstream mediator of Tgf-βs, in craniofacial development using the tissue-specific knockout mouse model system. We found out that Rac1 is an essential factor in the craniofacial development, and an important regulator of neural crest cell behavior. Third, to find out a function of Trim33, which was recently identified as a Smad4-independent regulator of Tgf-β signaling, we generated the Trim33 conditional knockout mouse line. We discovered that Trim33 is required for the early embryonic development. This study shows that the non-canonical Tgf-β signaling pathway is also essential factor in craniofacial development.

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Asset Metadata

Creator Kim, Ji Eun (author)

Core Title Non-canonical Tgf-beta signaling in craniofacial development

School School of Dentistry

Degree Doctor of Philosophy

Degree Program Craniofacial Biology

Publication Date 04/28/2009

Defense Date 03/11/2009

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

Tag conditional knockout mouse,Craniofacial Biology,OAI-PMH Harvest,Rac1,TGF-beta,Trim33

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Kaartinen, Vesa (committee chair), Bellusci, Saverio (committee member), Chai, Yang (committee member), Shi, Wei (committee member), Snead, Malcolm L. (committee member)

Creator Email jiek@usc.edu,jieunbucks@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m2133

Unique identifier UC1140233

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Document Type Dissertation

Rights Kim, Ji Eun

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

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Repository Location Los Angeles, California

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