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Nuclear fibroblast growth factor receptor 2 regulates skeletal development and joint formation
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Nuclear fibroblast growth factor receptor 2 regulates skeletal development and joint formation
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Content
NUCLEAR FIBROBLAST GROWTH FACTOR RECEPTOR 2 REGULATES
SKELETAL DEVELOPMENT AND JOINT FORMATION
by
Joanna E. Salva
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(GENETICS, MOLECULAR and CELLULAR BIOLOGY)
December 2018
2
ACKNOWLEDGEMENTS
I would like to thank my mentor Dr. Amy Merrill for providing me with the opportunity to learn
from her and become a better scientist. She has taught me how to persevere in the face of
challenges, think outside the box and push my own boundaries. She has given me endless
opportunities to present, write and edit: skills that have all greatly improved under her guidance.
I also thank her for sharing her love of the chicken embryo system with me, which is something I
have grown to love myself.
I thank my committee members Drs. Henry Sucov (committee chair), Robert Maxson, Judd Rice
and Pedro Sanchez for their guidance on my project and support throughout the years. They have
helped to shape my project and make it something I am proud of today. Thank you also to Dr.
Francesca Mariani for her insightful commentary and feedback on my project.
I thank my former labmates Cynthia Neben and Ryan Roberts for being the best friends I could
have ever hoped for in the lab. They made California become my home away from home and are
a never-ending source of laughs, support and friendship. I am eternally grateful for having the
opportunity to get to know them. Thank you to Creighton Tuzon for letting me pester him with
questions, pick his brain and talk over experiments. I have learned so much from him and greatly
value his expertise, insight and sense of humor. It has been a pleasure to work with the other
members of the Merrill lab over the years: Brian Idoni, Xiaojing (Amber) Mao, Lauren Bobzin,
and Diana Rigueur. I would also like to thank all the students I mentored, especially Taylor
Stucky. Working with them has made me into a better teacher and mentor, and I appreciate all of
their help in pushing forward my project.
3
Thank you to my family and friends back home for all of their encouragement and prayers. A
special thanks to my parents for enduring late night phone calls, providing unwavering support,
words of encouragement, and always being there when I need them. To my brothers, thank you
for reminding me not to take myself so seriously and making me laugh when I needed it most.
Finally, thank you to my now husband Andrew Ross. He endured 6 years of a long distance
relationship with a three hour time difference so I could achieve my goals. I thank him for his
patience, understanding, encouragement, tough love when I needed it, and constant supply of
jokes and laughter. It is with his support and the help from all those listed here that I have been
able to accomplish this achievement.
4
TABLE OF CONTENTS
ACKNOWLEDGEMENTS .......................................................................................................................... 2
LIST of TABLES and FIGURES ................................................................................................................. 5
ABBREVIATIONS ...................................................................................................................................... 6
CHAPTER 1: Introduction ........................................................................................................................... 7
1.1 Fibroblast Growth Factor Signaling .................................................................................................... 7
1.2 Limb Development ........................................................................................................................... 11
1.3 Human Relevance ............................................................................................................................. 19
CHAPTER 2: Signaling networks in joint development ............................................................................ 28
CHAPTER 3: Nuclear FGFR2 regulates musculoskeletal integration within the developing limb ........... 59
CHAPTER 4: Conclusions ......................................................................................................................... 87
REFERENCES ........................................................................................................................................... 89
5
LIST of TABLES and FIGURES
Table 1 Limb defects in FGFR2 disorder 26
Table 2 Muscle patterning defects in bent bone dysplasias 27
Figure 2.1 Spatial expression patterns of the principal signaling pathways in joint
development
54
Figure 2.2 Signaling networks in joint development have unifying features 56
Figure 3.1 RCAS injections into the LPM of the hindlimb largely targets dense
connective tissues
78
Figure 3.2 Expression of the BBDS mutations in the LPM of the chick hindlimb
induces hindlimb abnormalities
79
Figure 3.3 Expression of the BBDS mutations in the hindlimb connective tissue
causes bent long bones and knee defects
80
Figure 3.4 The BBDS mutations induce defects in the autopod skeleton and pelvic
girdle
81
Figure 3.5 Musculoskeletal integration is disrupted by expression of the BBDS
mutations
82
Figure 3.6 Nuclear localization is enhanced by BBDS mutations and distinct
localization signals in ATDC5 cells
83
Figure 3.7 Increased nuclear localization of FGF phenocopies BBDS 84
Figure 3.8 Changes in skeletal morphology are induced by increased nuclear and
nucleolar localization
85
Figure 3.9 Irregular muscle patterning is detected in embryos expressing nuclear and
nucleolar FGFR2
86
6
ABBREVIATIONS
AER Apical ectodermal ridge
BBDS Bent Bone Dysplasia Sydrome
BMP Bone morphogenetic protein
CFU Colony forming unit
E Embryonic day
FGF Fibroblast Growth Factor
FGFR FGF receptor
GOF Gain of function
HH Hamburger Hamilton stage
Ig Immunoglobulin-like
LADD Lacrimo-dento-digital Syndrome
LOF Loss of function
LPM Lateral plate mesoderm
NLS Nuclear localization signal
NoLS Nucleolar localization signal
PFA Paraformaldehyde
rDNA Ribosomal DNA
RTK Receptor tyrosine kinase
SHH Sonic hedgehog
TM Transmembrane
WT Wildtype
ZPA Zone of polarizing activity
7
CHAPTER 1: Introduction
1.1 Fibroblast Growth Factor Signaling
One of the major signaling pathways used throughout development is the fibroblast growth
factor (FGF) pathway. This pathway is composed of four receptors and 22 ligands [1]. The FGF
receptors (FGFRs) are receptor tyrosine kinases (RTK), composed of an extracellular ligand
binding domain, a single pass α-helical transmembrane (TM) domain and two intracellular
tyrosine kinase domains [1, 2]. Upon ligand binding, the receptors dimerize, autophosphorylate
and activate their kinase domains. Active FGFRs then act on intracellular targets to trigger
signaling cascades including JAK/STAT, MAPK, PI3K/AKT and PLCγ. This leads to changes in
proliferation, differentiation, migration or cell survival [1, 3]. Here I will discuss the principles
behind FGFR ligand specificity as well as how nuclear localization of the receptor can lead to
distinct cellular responses.
The extracellular ligand binding domain is composed of three immunoglobulin-like (Ig)
domains. The IgI domain and the following linker to IgII are not necessary for FGFR function,
but play an inhibitory role in ligand binding [1, 4-6]. However, domains IgII and IgIII are
necessary and sufficient for ligand binding [3]. Alternative splicing of the IgIII domain confers
ligand specificity. Exon 7 encodes the N-terminal portion of the IgIII domain while either exon
8, which encodes the IIIb isoform, or exon 9, which encodes the IIIc isoform, provide the C-
terminal portion [7]. Expression of the different splice variants is tightly controlled and tissue
specific with IIIb found in the epithelium and IIIc in the mesenchyme [4, 8]. Each isoform binds
a specific subset of FGF ligands. For FGFR2, the IIIb isoform preferentially binds FGF3, 7, 10,
and 22, while the IIIc isoform binds FGF1, 2, 4, 5, 6, 8, 9, 16, 17, 18, and 20 [1]. FGFR2IIIc
8
does not bind all of these ligands with the same affinity; however, it will preferentially bind
those ligands over the ligands of FGFR2IIIb. The ligands bound by each isoform tend to
originate in the reciprocal tissue; i.e., FGFR2IIIc binds ligands from the epithelium while
FGFR2IIIb binds ligands from the mesenchyme. Tissue specific expression of receptors and
ligands allows the developing embryo to communicate between tissues and permits the same
mechanisms to be used repeatedly in different organ systems [1, 4]. Because the IgII and IgIII
domains control ligand specificity and affinity, mutations in these areas result in a range of
chondrodysplasia and craniosynostosis disorders. FGFR2 in particular is connected with
craniosynostosis disorders, the majority of which have mutations in the IgII-IgIII linker or IgIII
domain [9]. Mutations can cause ligand-independent dimerization of the receptor and increased
signaling or augment FGFR2 activity by increasing ligand affinity or lowering ligand specificity
[9, 10]. The phenotypes caused by mutations in FGFR2 will be discussed in Ch1.3 Human
Relevance.
FGFR behavior can be controlled not only by splice variants and tissue specific expression
patterns, but also by subcellular localization. Canonical FGFR signaling occurs at the plasma
membrane like most RTKs. However, FGFRs 1-3 can also localize to the nucleus under a variety
of conditions. How then does a transmembrane RTK get into the nucleus? Because FGFRs have
not been reported to have nuclear localization signals (NLS), the use of a nuclear chaperone
protein could provide the necessary signal for nuclear localization. In fibroblasts, FGFR1
translocates to the nucleus upon treatment with FGF1 or FGF2, which each contain a NLS,
suggesting a ligand-dependent transport mechanism [11-13]. Moreover, inhibition of the nuclear
transport protein importin-β blocks FGFR1 and FGF2 from entering the nucleus [14].
9
Immunoprecipitation data shows that FGFR2 also interacts with importin-β implying that
FGFR2 may use a similar mechanism for translocation [15]. Additional research has shown that
nuclear FGFR1 does not all come from the plasma membrane and that a highly mobile cytosolic
population of FGFR1 proteins can interact with nuclear chaperones in the cytosol [16-18]. The
TM domains of FGFR1and FGFR2 enhance the ability of the receptors to move to the nucleus
with their unusual arrangement of hydrophobic and hydrophilic amino acids resulting in more of
a β-sheet structure instead of a typical α-helix [13, 17]. FGFR4, which has a highly hydrophobic
α-helix transmembrane domain, does not localize to the nucleus, and replacing the FGFR1 TM
domain with that of FGFR4 inhibits nuclear localization [17]. Thus, this structural change
accounts, in part, for the ability of the receptor to translocate to the nucleus. Despite the
similarities between FGFR1 and FGFR2, their localization is differentially regulated. In Müller
cells of adult mouse retinas, Fgfr1 is located in the cytosol while Fgfr2 is found in the nucleus
[19]. This indicates that there are mechanisms to distinguish between receptors and separate
them within the cell. It also suggests that FGFR1 and FGFR2 may be used for different functions
in the nucleus and at the plasma membrane. On the other hand, FGFR3 uses a distinct nuclear
localization mechanism where the intracellular domain of the receptor is cleaved at the plasma
membrane by γ-secretase and then translocates to the nucleus [20, 21]. Full length receptor has
also been observed in the nucleus, but its transport mechanism is unknown. In breast cancer
cells, FGFR1 can be cleaved by Granzyme B and the intracellular domain trafficked to the
nucleus where it promotes migration [22]. This method of intracellular domain cleavage and
transportation has yet to be seen in FGFR2. Regardless, while some aspects of the nuclear
transport mechanism are known, further research is required to understand how each FGFR
translocates to the nucleus.
10
Observation of different cell types indicates a correlation between nuclear translocation of
FGFRs and changes in cellular behavior. In dopaminergic neuron development, FGFR1 moves
from the membrane to the nucleus as progenitor cells begin to differentiate [23]. During
Drosophila gonad development, FGFR2 in Sertoli precursor cells is located at the plasma
membrane and translocates to the nucleus as the cells differentiate [24]. Similarly, human
embryonic myofibers show a progressive increase in nuclear FGFR3 as the muscles develop
[25]. Several types of cancer, including breast, endometrial, and pancreatic, also exhibit
enhanced nuclear FGFR localization [22, 26-30]. Irregular nuclear FGFR localization leads to
augmented proliferation and migration, thus leading to a cancerous phenotype. One FGFR2
mutation found in endometrial cancer also causes the only known skeletal birth defect with
increased nuclear localization, Bent Bone Dysplasia Syndrome (BBDS) [15, 26, 29, 31]. Both
cancers and BBDS teach us how nuclear FGFR2 needs to be tightly regulated to keep cells
functioning properly. How then do the FGFRs enact these changes in cellular behavior? FGFR1
and FGFR2 have both been shown to interact with chromatin remodelers to change DNA
structure and regulate gene transcription [14, 15, 18, 32-34]. Nuclear FGFR1 is capable of
activating transcription of genes such as c-Jun and tyrosine hydroxylase, which promotes cellular
proliferation and neurotransmitter production, respectively [14, 18, 34]. Work from our
laboratory has shown that nuclear FGFR2 binds to the ribosomal DNA (rDNA) promoter with
chromatin remodelers to make the DNA more accessible for transcription [15, 32]. Increased
proliferation of osteoprogenitor cells and delayed osteoblast differentiation result from these
changes in rDNA transcription [15, 35]. Altogether, the data indicate that nuclear FGFRs have
11
the ability to modify DNA transcription, and by varying the gene targets in a cell specific
manner, alter cellular behavior.
1.2 Limb Development
Formation of the limb is a tightly regulated process involving many signaling pathways.
Communication between tissue types is critical in initial limb patterning as well as in the later
integration of multiple tissue types to form the musculoskeletal system. Fgfr2 signaling plays a
key role in the patterning and development of the limb. Here I will discuss the involvement of
FGFR2 in the establishment of the proximal-distal and anterior-posterior axes as well as bone
development and differentiation.
Early limb development
The limb begins as an outgrowth of lateral plate mesoderm (LPM) from the body wall, apparent
in the chicken embryo at Hamburger-Hamilton (HH) stage 16 or embryonic day (E) 9.5-10 in
mice [36, 37]. At this stage, proliferation of LPM cells within a localized region leads to
increased cell density and a protrusion of LPM from the flank mesoderm, thus making the limb
bud [38, 39]. Communication between the LPM and the overlying ectoderm forms the apical
ectodermal ridge (AER), the signaling center for proximal-distal patterning of the limb.
Signaling crosstalk between the LPM and AER is essential to establish the three proximal-distal
segments of the limb: the stylopod, zeugopod, and autopod, respectively. Loss of the AER by
either physical removal or genetic ablation results in limb truncations of varying severities,
depending on timing of the removal [40]. Early removal or ablation results in limbless embryos,
while experiments at later time points result in truncations along the proximal-distal axis with
12
retention of proximal structures and absence of more distal structures [40]. These experiments
indicate the necessity of the AER in controlling proximal-distal patterning of the limb and that
limb patterning is a tightly regulated, temporal process. The AER carries out its function by
secreting signaling factors to promote cell survival and proliferation and simultaneously inhibit
differentiation in the subjacent cells, thus forming a pool of limb mesenchymal progenitors. As
the limb grows in size, cells closest to the AER are maintained in a progenitor state while those
farther away from the AER signals begin to differentiate [41]. The mesenchymal progenitors
removed from AER influence will proceed to form the cartilage and bone of the skeleton, as well
as the connective tissue and tendons.
Several models have been proposed over the years regarding patterning of the limb and the role
of the AER in this process. The progress zone model suggests that the length of time spent under
the influence of the AER determines the fate of the cells, where cells that spend little time near
the AER become proximal elements and those that spend the longest amount of time become the
distal elements [42]. The early specification model, on the other hand, claims that the limb
segments along proximal-distal axis are predetermined and merely expand under the control of
signals from the AER [38, 43, 44]. However, with the emergence of genetic techniques, neither
of these models adequately explains the variety of phenotypes observed upon deletion of
different signaling molecules in the AER or LPM. The latest model in limb patterning called the
two-signal gradient model proposes that signals are needed from both the distal AER and the
proximal flank [38, 44]. Signals from the AER control the distalization of the limb bud while
proximal signals dictate the fate of proximal structures. The distribution of ligands from both
sides results in a gradient of signaling which can be used to distinguish proximal, intermediate
13
and distal fates. This model explains how changes in gene expression can result in the formation
of the stylopod and autopod with the intermediate zeugopod being reduced or absent [44, 45].
The two-signal model combines elements from the previous models in that the AER is required
for distalization of the limb and is necessary for continued limb outgrowth, while adding new
insight into the specification of each segment. Sheeba et al. contribute a modification to the two-
signal model by incorporating a temporal element [46]. Evidence implies that the proximal and
distal gradients work in conjunction with a time-dependent factor to convey positional
information in a sequential fashion thus allowing limb segments to be specified even after they
have moved away from the signaling sources. As new information comes to light, the current
models may be further revised and modified to best explain the complexity of developmental
limb patterning.
Working along with the AER, the zone of polarizing activity (ZPA) controls patterning along the
anterior-posterior axis. The ZPA is found in the mesenchyme of the posterior margin of the limb
bud and uses a signaling gradient to establish posterior and anterior zones. This role was
uncovered by observing that transplantation of the ZPA to the anterior portion of the chick wing
creates a mirror image duplicate set of digits [47, 48]. Initiation of the ZPA is dependent on the
AER; however, both regions require crosstalk in order to maintain their signaling gradients and
establish proper patterning of the limb [49, 50].
Fgfr2 signaling in limb patterning
Fgf signaling is the primary signaling pathway used by the AER to pattern the limb. Experiments
have shown that Fgfs are needed for limb development with loss of Fgf4, 8, 9, 10 and 17, alone
14
or in combination, resulting in limb defects of varying severities [45, 51-55]. Ligands are
secreted by both the mesenchyme and the overlying epithelium, and it is through differential
expression of Fgfrs with their varying ligand affinities that crosstalk between the tissues occurs.
Fgfr2, specifically, is critical in creating this communication loop between the mesenchyme of
the LPM and the epithelium of the AER.
To study Fgfr2 in the context of limb development, a variety of techniques need to be applied, as
universal deletion of Fgfr2 results in embryonic lethality in mice [56]. By deleting the IgIII
domain of Fgfr2, which is responsible for ligand specificity, the resulting embryos are viable but
limbless [57]. As mentioned in Ch1.1 Fibroblast Growth Factor Signaling, the IgIII domain of
Fgfr2 can be alternatively spliced into two differentially expressed isoforms: Fgfr2IIIb in the
epithelium and Fgfr2IIIc in the mesenchyme [8, 58]. In the absence of Fgfr2IIIb, embryos fail to
form the AER and are limbless [50, 59]. On the other hand, Fgfr2IIIc deletion results in a full
limb; however, development of the bones is affected [60]. This indicates that Fgfr2IIIb is the
critical isoform needed for establishing the limb and that Fgfr2IIIc plays a role in the later
development of the skeleton. Use of the Cre-LoxP system to conditionally delete Fgfr2 in a
spatiotemporal manner reveals further insight into the role the receptor plays in patterning of the
limb [61]. In Msx2-Cre; Fgfr2
flx/flx
mice, the forelimb AER has been established prior to Fgfr2
being knocked out, thus the proximal components of the limb are formed, but the distal autopod
is missing. In the hindlimb, however, Fgfr2 is removed prior to AER establishment resulting in
complete absence of the hindlimbs [62, 63]. This shows that Fgfr2 plays an important temporal
role in limb patterning and that its continued expression is required for all limb segments to
form.
15
Fgf signaling is also used by the ZPA in a feedback loop with Shh and Bmp to establish a
morphogen gradient along the anterior-posterior axis [64]. Bmps are expressed in the
mesenchyme adjacent to the AER and inhibit Fgf signaling. Fgf from the AER stimulates
production of Shh in the posterior mesenchyme to form the ZPA. Shh in turn promotes Gremlin1
that then inhibits Bmp thus allowing for more Fgf to be produced [65]. This feedback loop helps
to maintain signals from both the AER and ZPA. If one of the components in the loop is
disrupted, irregular patterning of the digits results. Creation of chimeric embryos with Fgfr2-
deleted cells leads to embryos with pre-axial polydactyly, oligodactyly and missing zeugopod
skeletal elements [66]. Given that the chimeric cells did not contribute to the AER and they
cause changes in digit number, these results suggest that mesenchymal Fgfr2 plays a role in
establishing normal patterning along anterior-posterior axis. Knockdown of Fgfr2 expression in
the distal mesenchyme using AP2-Cre causes absence of digit 3 and hypoplasia of digits 1 and 2
[67]. Tamoxifen induced deletion of Shh at E9.5-10 results in the absence of digit 3 with varying
hypoplasia or loss of digits 4 and 5, mirroring the phenotype in the AP2-Cre Fgfr2 knockdown
[68]. This implies that Fgfr2 works opposite of Shh to help establish the anterior portion of the
digital plate. The receptor involved in the Fgf-Shh-Bmp signaling loop has previously been
unknown, but this work suggests that Fgfr2 is the receptor involved.
Endochondral bone development
As the limb bud grows, the LPM derived mesenchyme in the center of the limb bud condenses.
Once a certain density is attained, the mesenchymal cells begin to differentiate into chondrocytes
[9, 69, 70]. The perichondrium, which is derived from the surrounding mesenchyme,
16
encapsulates the cartilage template, produces signaling molecules to regulate chondrocyte
growth and differentiation, and is a source of osteoprogenitors that will make the bone [71-73].
The phases of cartilage differentiation resting, proliferating, pre-hypertrophic and hypertrophic
chondrocytes are visible simultaneously within the stratified structure of the growth plate. Within
the growth plate the rates of proliferation and hypertrophy and the size of the hypertrophic
chondrocytes and mineralized matrix formed dictate the final size of the bone [74, 75]. After an
extracellular matrix scaffold is deposited by the hypertrophic chondrocytes, signals from the
perichondrium and periosteum induce the cells to apoptose. Recent work shows that not all
hypertrophic chondrocytes die and that some cells at the chondro-osseous junction will
transdifferentiate into osteoblasts [76]. Osteoblasts from the perichondrium and periosteum
migrate into the cartilage template to replace the hypertrophic chondrocytes and continue to
mineralize the tissue to form trabecular bone. The osteoblasts become embedded in the matrix
where they terminally differentiate into osteocytes that will maintain the mineralized bone
matrix. This process of ossification proceeds until the entire cartilage template, except the
epiphyseal surface, is replaced with bone.
Fgfr2 in bone development
Expression patterns reveal that Fgfr2 is expressed throughout the stages of bone development.
Fgfr2, specifically the IIIc isoform, is initially expressed within the condensing mesenchyme in
the limb buds of human, mouse and chicken embryos [8, 58, 60, 77, 78]. Little is known about
the function of Fgfr2 in the condensing mesenchyme; however, in vitro experiments suggest that
Fgfr2 in the pre-chondrogenic mesenchyme inhibits the adjacent mesenchyme from becoming
cartilage. In combination with differential ligand expression patterns, it provides a potential
17
mechanism by which spacing of skeletal elements can be controlled [70]. As the condensation
begins to differentiate, Fgfr2 becomes restricted to the perichondrium and regions of future
joints. Fgfr2 expression is further restricted as chondrocyte differentiation continues to the
epiphyseal chondrocytes, periarticular cartilage, periosteum and perichondrium [58, 77].
To uncover Fgfr2’s function in these tissues, mouse models have proved vital. Both complete
and conditional knockouts of mesenchyme-specific Fgfr2 result in viable offspring with small,
proportional skeletal elements. On further examination, ossification is delayed with reduced
mineralization and decreased expression of bone specific genes. BrdU staining reveals that
without Fgfr2 osteoprogenitors have a decreased proliferative capacity, leading to a thin,
disorganized perichondrium and periosteum with fewer osteoprogenitors and osteoblasts and
subsequent decreased bone mineral density [79, 80]. Additionally, growth plate architecture is
altered with smaller zones of proliferating and hypertrophic chondrocytes. Expression of gain of
function (GOF) mutations found in disorders like Crouzon, Pfeiffer and Apert syndromes also
induce bone phenotypes in mice. Despite different causal mutations, there are several
phenotypic similarities. Long bones in all mutants are smaller in size but overall proportional.
Augmented numbers of osteoblasts and osteoprogenitors, particularly at the chondro-osseous
junction, are observed in Crouzon and Apert mouse models [79, 81]. Elevated expression of
osteoblast-specific genes indicates increased differentiation of osteoprogenitors. Mild changes in
the growth plate are also observed with smaller zones of proliferating and hypertrophic
chondrocytes, but the primary defects lie in the osteoprogenitor/osteoblast lineage. The effects of
Fgfr2 in preosteoblasts have been thoroughly characterized in the calvarium, which is formed by
intramembranous ossification, and similarly show that Fgfr2 promotes proliferation and
18
differentiation of osteoblasts [79, 81-83]. Work from our laboratory has uncovered how Fgfr2
can regulate these two distinct processes in preosteoblasts. In using the mutations from BBDS,
which enhance nuclear localization of FGFR2, we find that nuclear FGFR2 promotes
proliferation of osteoprogenitor cells while membrane bound FGFR2 promotes differentiation
[15]. Altogether, Fgfr2 plays a critical role in the proper proliferation and differentiation of
osteoprogenitors and osteoblasts during skeletal formation.
Interestingly, the phenotypes between GOF and loss of function (LOF) models overlap. The
skeletons of both are smaller overall compared to littermate controls. Precocious proliferation
and differentiation of the perichondrial/periosteal preosteoblasts with GOF mutations results in
stunted growth of the bones as the progenitor pool is rapidly exhausted. Loss of Fgfr2, on the
other hand, causes smaller bones due to a small pool of osteoprogenitors that proliferate and
differentiate at a slower rate. Despite being smaller in size, the bones remain proportional. The
disproportionally short long bones observed in achondroplasia and thantatophoric dysplasia are
due to activated FGFR3 signaling inhibiting chondrocyte proliferation and hypertrophy [84].
While slightly smaller zones of proliferating and hypertrophic chondrocytes are observed in
Fgfr2 mutants, these changes are insufficient to change the overall morphology of the bone [80,
82]. As Fgfr2 is expressed in the resting, periarticular chondrocytes which become proliferating
chondrocytes, and Fgfr2 regulates proliferation and differentiation of osteoblasts that replace the
hypertrophic chondrocytes, it is unsurprising that the proliferating and hypertrophic
chondrocytes are mildly affected.
19
Fgfr2 in joint development
Often overlooked, the joint is a distinct organ composed of bone, cartilage, tendon and ligaments
and is a critical component in movement of an organism. Little is known about Fgfr2 in synovial
joint development, but it has been well-studied in the fibrous joints of the cranial sutures. Within
the cranial suture Fgfr2 controls both proliferation and differentiation of the osteoprogenitor cells
at the bone fronts and is necessary to maintain patent sutures. Disruption of the pathway leads to
craniosynostosis, the premature fusion of the cranial sutures. Deletion of the cranial joint marker
Gdf6 results in fused sutures that fail to express Fgfr2, suggesting that joint specification is
required for Fgfr2 expression in the suture [85]. As there is considerable overlap in the signaling
pathways used between synovial and fibrous joints, Fgfr2 likely influences synovial joint
development as well [86]. It is expressed at the proper spatiotemporal interval during limb
development with Fgfr2 found in the developing joint space and the periarticular and epiphyseal
cartilage, which compose the interface between bones and the joint space [58, 77]. Various
mouse models removing Fgfr2 from the limb result in metatarsal and tarsal joint fusions, as well
as irregular articulations or fusions of the ankle [67, 70, 80]. Expression of activating Fgfr2
mutations induces small or fused elbow and knee joints and ectopic ossification within the joint
[79, 87]. In light of these experiments, more research is needed to fully uncover Fgfr2’s role in
synovial joint development.
1.3 Human Relevance
Mutations in FGFR2 are responsible for at least 10 different skeletal birth defects [9, 10, 31].
Despite the different mutations, these FGFR2 disorders exhibit a number of skeletal
abnormalities with phenotypic similarities. While the role of FGFR2 in craniofacial development
20
has been more broadly studied, much remains unknown about how aberrant FGFR2 signaling
influences limb development. Here I will discuss the limb abnormalities found in FGFR2
disorders and how they lend insight into normal limb development. Additionally, I will discuss
what we can learn from other bent bone dysplasias to better understand phenotypic abnormalities
in FGFR2 disorders and how this information can impact treatment for patients with these
conditions.
Most FGFR2 skeletal disorders exhibit some degree of digit abnormalities ranging from
clinodactyly and brachydactyly to split-hand malformation and syndactyly indicating abnormal
proximal-distal and anterior-posterior patterning (Table 1). FGF signaling is well-known for
regulating proximal-distal patterning of the limb through the AER; however, it is also important
in anterior-posterior limb patterning. FGF works together with SHH and BMP to establish and
maintain the anterior-posterior axis, but little is known about which FGF receptor is employed in
this process [64, 88, 89]. Polydactyly, seen in GOF Apert and Pfeiffer syndromes, and
oligodactyly, in LOF LADD syndrome, suggests FGFR2 influences anterior-posterior patterning
of the digits [90-93]. Apert syndrome patients typically have pre-axial polydactyly, where the
anterior digits are duplicated [93]. While polydactyly tends to be post-axial in Pfeiffer syndrome
patients, broadened first digits are observed in the hands and feet suggesting that the anterior
digit field has widened but not completely split [88, 90]. The big toes of patients with Jackson-
Weiss syndrome are similarly broad and deviated [94]. Defects in LADD syndrome also tend to
impact the anterior digits with absent, hypoplastic or bifid thumbs [91, 92]. Additionally, LADD
syndrome patients exhibit split-hand malformations. Split-hand malformations have been
connected more with deficiencies of the AER than anterior-posterior patterning; nevertheless,
21
this phenotype correlates with irregular digit patterning and could be caused by a combination of
defects in proximal-distal and anterior-posterior signaling [95]. The prevalence of patterning
defects along the anterior-posterior axis in the FGFR2 skeletal disorders indicates a novel role
for FGFR2 in anterior-posterior patterning that has been previously unstudied.
Synovial joint fusions are commonly found throughout the limb in FGFR2 skeletal dysplasias,
yet little is known about the role of FGFR2 in synovial joint development. Examination of
normal human embryonic limbs shows that FGFR2 is highly expressed within synovial joints
throughout development and is continued to be expressed within the epiphyseal and articular
chondrocytes as joint cavitation proceeds [77, 96]. This finding shows that FGFR2 is
spatiotemporally present during formation of the joint, indicating a possible role in joint
development. Studying the joint defects present in FGFR2 disorders provides insight into this
role. A hallmark feature of Apert syndrome is bony and cutaneous syndactyly of the hands and
feet [96-98]. While syndactyly is not a joint defect, it is an irregular fusion of bones between
fingers accompanied by fusions within each digit. It has been suggested that abnormal FGFR2
signaling maintains the interdigital mesenchyme in Apert syndrome hands, preventing its
apoptosis and thus providing a bridge between digits that later ossifies [96]. Along with
syndactyly, the digits exhibit symphalangism, the absence of interphalangeal joints. Histological
examination reveals that the interphalangeal joint does not cavitate and maintains a cartilage
intermediate that becomes ossified over time [97]. Symphalangism is also observed in Pfeiffer
syndrome, but not in Jackson-Weiss syndrome despite the similar broad, deviated big toes [90].
Patients with Jackson-Weiss display more proximal defects of tarsal and metatarsal fusion
instead [94]. Joint defects are not limited to the hands and feet with elbow ankylosis being
22
observed in patients with Apert, Pfeiffer, Antley-Bixler, and LADD syndromes [92, 97, 99, 100].
Varying combinations of radioulnar, radiohumeral, and radioulnahumeral fusions are found in
these disorders and all result in restricted mobility at the elbow. Pfeiffer and Antley-Bixler
syndromes have also exhibited knee ankylosis. While joint ankylosis is not a standard diagnostic
criterion for most FGFR2 disorders, bilateral radiohumeral synostosis is a diagnostic requirement
for Antley-Bixler syndrome [99, 101, 102]. Restricted elbow movement without bony fusion in
BBDS and radial dislocation in Crouzon and Pfeiffer syndromes suggests that FGFR2 influences
the soft tissue components of the joint as well as the articular surface of the bones [90, 103, 104].
Based on the defects found in these FGFR2 disorders, a failure for the joint to cavitate and
improper soft tissue connections surrounding the joint are suggested. In addition to synovial joint
fusions and malformations, nearly all FGFR2 disorders have craniosynostosis, and many also
have fusions of the cervical vertebrae [99, 100]. That fibrous, cartilaginous and synovial joints
are all affected in FGFR2 disorders suggests that the same mechanism is repeatedly used in each
type of joint and warrants further study.
It is well known that FGFR2 influences bone development. Altered bone growth is observed in
FGFR2 disorders through changes in digit outgrowth as well as mineralization and ectopic
ossification. Crouzon syndrome patients do not typically exhibit overt changes in the digits of
hands or feet; however, upon assessment of the metacarpophalangeal pattern profile, which
measures bone proportions in the hand, Crouzon patients have shorter phalanges than normal and
have generally smaller hands [103, 105, 106]. The observation of shorter phalanges indicates a
change in bone growth. In BBDS, the digits simultaneously demonstrate brachydactyly and
trident hand malformation, where the digits are equal in length giving the hand a fork-like
23
appearance, also signifying changes in bone outgrowth [31, 100, 104]. Additionally, patients
exhibit reduced mineralization of the calvarium and osteopenia [15, 31, 104]. Although BBDS
patients seem to show a decrease in bone growth, they also display irregular bony nodules on the
phalanges, metacarpals and metatarsals giving them an “angel-shape” [31, 100, 104]. These extra
nodules imply ectopic ossification. Further support for excess bone growth is exemplified by the
radiohumeral fusions present in Antley-Bixler syndrome and other FGFR2 disorders as well as
ectopic ossification between digits that causes syndactyly in Apert syndrome [96, 99, 101].
Taken together it seems that FGFR2 plays distinct roles in ossification and bone growth
depending on its location. At tendinous junctions or synovial joints abnormal FGFR2 signaling
induces ectopic bone formation, while intramembranous and endochondral ossification may be
reduced in the bone. Despite the variation, it is clear that FGFR2 regulates bone development.
FGFR2 is connected with two bent bone disorders, Antley-Bixler syndrome and Bent Bone
Dysplasia Syndrome, both which present with bent femoral long bones and bilateral femoral
fractures, with additional bowed tibias in BBDS [31, 99, 101, 104, 107]. While little is known
about the cause of bent femurs in either of these disorders, there are over 40 other bent bone
disorders that can lend insight [108]. Many of the bent bone disorders are attributed to defects in
the structural integrity of the bone. However, the genes responsible for these disorders often also
impact muscle and tendon. For example, defects in musculoskeletal integration are observed in
Campomelic Dysplasia, Osteogenesis imperfecta, Schwartz-Jampel Syndrome Type 1, and
Stüve-Wiedemann syndrome (Table 2). Contribution of muscle to the bowed limb phenotypes in
Schwartz-Jampel Syndrome and Stüve-Wiedemann syndrome is apparent with progressive
worsening of the bending with age due to myotonia [109-111]. Campomelic dysplasia and
24
osteogenesis imperfecta, on the other hand, have small, weak muscles with irregular tendon and
ligament attachments evidenced by changes in joint laxity and bone dislocation [112-115]. Bone
development requires mechanical input from the muscle. Absent muscle forces in animal models
induce changes in bone mineral density, structural integrity, epiphyseal morphology, and in some
models, bowed long bones [116-123]. Reduced muscle activity in the human disorders could be
further exacerbating the already weakened bones, making bowing even more likely. Several
studies have proposed this very idea where changes in the musculature in combination with poor
bone integrity lead to bowed long bones [111, 112, 124-126]. Supporting this concept is the
disparity between bowing in the upper and lower limbs with the long bones of the legs being
more susceptible to bowing. Analysis of the musculoskeletal system between upper and lower
extremities reveals that while the skeletal structure is rather homologous, the musculature is
vastly different with more points of contact muscle-to-muscle and muscle-to-bone, mediated by
tendons, in the lower limb than in the upper limb [127]. Therefore, changes in muscle patterning
and connectivity are likely to have more severe consequences in the lower limb. Treating muscle
defects through physical therapy and regular exercise routines improved muscle function and
strength in patients with osteogenesis imperfecta leading to better daily functioning [114, 115].
Continued exercise is required though with lapse in treatment causing a return to the weakened
muscle state. Whether other patients with bent bone disorders could benefit from physical
therapy warrants further study. Altogether the data suggest that there may be muscle defects in
patients with BBDS and Antley-Bixler syndrome. While the musculature has yet to be examined
in either condition, ligamentous laxity in BBDS implies a defect in ligaments surrounding the
joints [104]. Additionally, work from our laboratory indicates a role for FGFR2 in enthesis
patterning in the jaw, implying that FGFR2 could be affecting the entheses of the limbs in BBDS
25
and Antley-Bixler syndrome [128]. Further examination of the muscles and tendons should be
conducted in both of these disorders and other bent bone dysplasias to gain a better
understanding of the bowed limb phenotype and find possible treatments for surviving patients.
Moreover, muscles in the other FGFR2 disorders should be examined as they could enhance our
understanding of the role of FGFR2 in musculoskeletal integration.
26
Table 1. Limb defects in FGFR2 disorders
Disorder
(MIM)
Mutation
Activity
Digit Joint Long Bone Reference
Antley-
Bixler
Syndrome
(207410)
GOF Arachnodactyly
Radiohumeral
synostosis,
radioulnar
synostosis, elbow
and knee ankylosis
Femoral bowing,
femoral fracture
[99, 101,
102, 129]
Apert
Syndrome
(101200)
GOF
Bony and soft
tissue syndactyly
of hands and
feet, polydactyly
Elbow ankylosis,
symphalangism
N/O
[93, 97,
98, 130]
Bent Bone
Dysplasia
Syndrome
(614592)
?
Angel-shaped
phalanges,
brachydactyly,
trident hand
configuration
Limited elbow
extension
Femoral and
tibial bowing,
femoral fracture
[31, 104,
107]
Crouzon
Syndrome
(123500)
GOF
N/O
Short phalanges
Radial dislocation,
small joint
between humerus
and ulna
N/O
[103, 105,
106]
Jackson
Weiss
Syndrome
(123150)
GOF
Broad medially
deviated big
toes, normal
hands
Tarsal and/or
metatarsal fusion
N/O [94]
Lacrimo-
auriculo-
dento-
digital
(LADD)
Syndrome
(149730)
LOF
Clinodactyly,
brachydactyly,
syndactyly,
hypoplastic or
bifid thumbs,
ectrodactyly
Radioulnar
synostosis, elbow
ankylosis
N/O
[91, 92,
131]
Pfeiffer
Syndrome
(101600)
GOF
Broad, deviated
thumbs and
big toes,
clinodactyly,
polydactyly
Elbow and knee
ankylosis, Radial
dislocation,
symphalangism
N/O
[90, 94,
100, 101]
N/O Not observed
27
Table 2. Muscle patterning defects in bent bone dysplasias
Disorder
(MIM)
Gene Long Bone Muscle/Tendon Joint Reference
Antley-Bixler
Syndrome
(207410)
FGFR2
Femoral bowing,
femoral fracture
?
Radiohumeral
synostosis,
radioulnar
synostosis, Joint
ankylosis
[99, 101,
102]
Bent Bone
Dysplasia
Syndrome
(614592)
FGFR2
Femoral and
tibial bowing,
femoral fracture
Ligament laxity
Limited elbow
mobility
[31, 104,
107]
Campomelic
Dysplasia
(114290)
SOX9
Femoral and
tibial bowing
Small, shortened
muscles,
hypotonia,
irregular tendon
attachments,
clubfoot, split
tendons
Knee contracture,
Hip dislocation
[112, 132]
Osteogenesis
Imperfecta
(166200)
COL1
Femoral bowing,
bone fractures
Small muscles,
hypotonia,
reduced muscle
force, ligament
laxity
Joint laxity,
radial dislocation
[108, 113-
115, 133]
Schwartz-
Jampel
Syndrome
Type 1
(255800)
HSPG2
Femoral bowing,
humeral fracture,
short limbs,
slender diaphysis
and wide
metaphysis
Myotonia,
progressive
muscle stiffness
Multiple joint
contractures
[108, 109,
111, 134]
Stüve-
Wiedemann
Syndrome
(601599)
LIFR
Progressive
femoral bowing,
tibial bowing,
osteopenia, wide
metaphysis
Myotonia,
hypotonia
Joint contracture,
limited knee and
elbow mobility
[110, 126]
28
CHAPTER 2: Signaling networks in joint development
#
Joanna E. Salva
1, 2
and Amy E. Merrill
1, 2*
1
Center for Craniofacial Molecular Biology, Ostrow School of Dentistry, University of Southern
California, Los Angeles, CA, 90033
2
Department of Biochemistry and Molecular Biology, Keck School of Medicine, University of
Southern California, Los Angeles, CA, 90033
*
Correspondence: amerrill@usc.edu
Keywords: synovial joint, cartilaginous joint, fibrous joint, IVD, suture, FGF, hedgehog, Wnt,
and Bmp
#This chapter has previously been published in Developmental Dynamics. Salva JE, Merrill AE. 2017.
Signaling networks in joint development. Dev Dyn 246:262-274.
29
Abstract
Here we review studies identifying regulatory networks responsible for synovial, cartilaginous,
and fibrous joint development. Synovial joints, characterized by the fluid-filled synovial space
between the bones, are found in high-mobility regions and are the most common type of joint.
Cartilaginous joints unite adjacent bones through either a hyaline cartilage or fibrocartilage
intermediate. Fibrous joints, which include the cranial sutures, form a direct union between
bones through fibrous connective tissue. We describe how the distinct morphologic and
histogenic characteristics of these joint classes are established during embryonic development.
Collectively, these studies reveal that despite the heterogeneity of joint strength and mobility,
joint development throughout the skeleton utilizes common signaling networks via long-range
morphogen gradients and direct cell-cell contact. This suggests that different joint types
represent specialized variants of homologous developmental modules. Identifying the unifying
aspects of the signaling networks between joint classes allows a more complete understanding of
the signaling code for joint formation, which is critical to improving strategies for joint
regeneration and repair.
30
Introduction
Joints connect articulating elements of the vertebrate skeleton. While all joints share this role,
their morphologic diversity produces a broad range of mechanical possibilities. The degree of
strength and mobility is controlled by the composition of joint connective tissues, whereas the
type and range of motion is conferred by joint shape. There are three major classes of joints:
freely movable synovial joints, slightly movable cartilaginous joints, and immovable fibrous
joints. The distinct morphologic and histogenic characteristics of each joint class are established
during embryonic development. Despite this heterogeneity, joint development collectively
utilizes common signaling mechanisms via long-range morphogen gradients and direct cell-cell
contact. These signaling processes are critical to establish and maintain a transcriptional profile
unique to the joint-forming compartment.
Here we review studies that have identified the principal signaling pathways within the
regulatory networks for synovial, cartilaginous, and fibrous joint development. In doing so, we
reveal that these signaling pathways, as well as their hierarchical relationships, are reiteratively
used across joint types. We discuss overlapping signaling architecture between distinct joint
classes and discuss evidence that different joint types are homologous developmental modules
that have undergone specialization. By identifying the unifying aspects of the signaling networks
between joint classes, we hope to gain a more complete understanding of the signaling code for
joint formation, which is a critical first step in unlocking the potential for joint regeneration and
repair.
31
Signaling networks in joint development
1. Synovial joints
1.1 Synovial joint structure
Synovial joints, found in high-mobility regions such as the limbs and their girdles, are the most
common and structurally variable of all the joint types. A distinguishing characteristic of a
synovial joint is the fluid-filled cavity that separates the articulating surfaces of the bones. The
jointed bones, which come in a variety of configurations, are composed of hyaline cartilage, a
permanent avascular tissue that reduces joint friction through its secretion of hyaluronate and
lubricin [135]. The joint cavity is enclosed by the stratified structure of the joint capsule. Lining
the inner surface is a thin synovial membrane that secretes synovial fluid to lubricate the joint.
The capsule’s outer layer of dense fibrous connective tissue is fastened into the boney epiphyses
to structurally support the articulation. In most joints, the fibrous layer is locally thickened into
the capsular ligament, which in some cases may be replaced by tendon. Frequently, the capsular
tissue invades the synovial joint cavity, dividing it completely or incompletely, as a
fibrocartilage articular disc or meniscus, respectively. Further structural support for the joint
capsule is provided by accessory ligaments, which can lie inside or outside the capsule and
prevent damage from overextension.
1.2 Signals in synovial joint development
The first morphological sign of synovial joint development is the emergence of the interzone, a
dense population of pre-chondrogenic mesenchyme that lies between adjacent cartilaginous
anlagen [136]. Early studies demonstrated that removal of the interzone leads to fusion of
articulating skeletal elements, indicating its necessity for joint formation [137]. The interzone is
32
composed of three layers: two dense outer layers of round cells separated by an inner layer of
flattened cells. Light and electron microscopic analyses suggest that the outer layers contribute to
the growing long bone epiphyses while the inner layer forms the articular surfaces [138]. Taken
together, the evidence indicates that the interzone cells are a population of joint progenitor cells
critical for forming multiple tissues of the mature joint.
1.2.1 Bmp
Gdf5, a member of the BMP family of secreted factors, is one of the earliest known markers for
the presumptive interzone and is an autonomous regulator of synovial joint development. Gdf5
expression is initially limited to the site of the future joint prior to the emergence of the interzone
[139, 140]. Lineage tracing of Gdf5+ cells demonstrates their contribution to the epiphysis,
articular cartilage, joint capsule, and intra-articular ligaments [141-143]. Mice null for Gdf5
exhibit abnormalities in the synovial joints of the limb, with partial or complete fusion of the
jointed bones [140, 144]. While Gdf5 is expressed in nearly all synovial joints of the
appendicular skeleton, some joints remain unaffected in the limbs of Gdf5 null mice. This is
likely due to functional redundancy with Gdf6 and Gdf7, fellow Bmp family members with
joint-specific expression and a high degree of homology to Gdf5 [85, 145]. Mice null for both
Gdf5 and Gdf6 exhibit more extensive joint fusions not seen in the individual mutants,
suggesting functional redundancy of these genes in joint development [85].
While Gdf5 is required for synovial joint development, it is not sufficient to induce formation of
an ectopic joint. Instead, treatment of developing limbs with recombinant Gdf5-soaked beads
promotes cartilage growth [144]. Additionally, Gdf5 overexpression in chicks and mice expands
33
the epiphyses, lengthens the bone, and ablates the joint [139, 146]. It is interesting that both too
much and too little Gdf5 signaling result in joint loss. That the developing synovial joint is
exquisitely sensitive to gradients of Bmp signaling is also supported by the dynamic expression
patterns of Bmp pathway components. Bmp2 is co-expressed with Gdf5 after the interzone is
established [147, 148]. Bmp inhibitors Noggin and Chordin are expressed in the early interzone,
with Noggin later being regionally restricted to the epiphysis several layers from the interzone
[135, 147, 148]. Cells between the Chordin-expressing interzone and the Noggin-expressing
region of the epiphysis are responsive to the pro-chondrogenic signal from Bmp and form the
articular cartilage [135]. Increasing the zone of Bmp signaling through Noggin inactivation or
ectopic activation of Bmp receptors Bmpr1b and Acrv1/Alk2 leads to mis-differentiation of the
interzone and articular cartilage into growth plate-like cartilage [135, 149-151].
While it is largely accepted that Gdf5 expression marks joint progenitor cells, the source of these
progenitor cells has been debated. It was initially thought that all joint progenitor cells were
specified in a single early event and originated in the interzone. However, evidence suggests that
cells from outside the interzone also contribute to the developing joint. Lineage tracing in avian
embryos with DiI shows mesenchymal cells adjacent to the interzone migrating into the
developing synovial joint [152]. Genetic lineage tracing in mice shows that Col2a1+ cells within
the pre-chondrogenic anlagen give rise to the interzone and subsequently the articular cartilage,
ligaments, and medial meniscus. However, Col2a1- cells from outside the anlagen later
contribute to the lateral meniscus [153].
34
During early joint development there is a significant increase in the cellularity of the interzone;
however, the Gdf5+ cells in the interzone proliferate at a very slow rate [135]. Lineage tracing of
Col2a1+ cells showed a zone of proliferative cells at the distal ends of the cartilage anlagen
contributing to the growth of the interzone and articular cartilage. The notion that joint
progenitor cells are indeed derived from multiple origins is further substantiated in another
recent study. Transient labeling of Gdf5+ cells at distinct time points during joint development
identifies a continuous influx of new Gdf5+ cells into the interzone to sustain joint development
[154]. The source of the recruited cells is proposed to be a population of Sox9+/Gdf5- cells
flanking the developing joint, which is consistent with previous studies showing that all joint
structures are derived from Sox9+ cells [155]. Timing of Gdf5+ cell recruitment influences the
tissue type to which the cells contribute [154]. That there is a continuous influx of Gdf5+ cells
into the developing joint seems to conflict with earlier studies showing that the interzone is
necessary for joint development [137]. However, this could be explained by an instructive role
for the interzone in new Gdf5+ cell recruitment.
1.2.2 Wnt
Wnt signaling is also an early regulator of synovial joint formation and maintenance. Wnt4,
Wnt9a (previously known as Wnt14) and Wnt16 are expressed in overlapping and
complementary patterns in and around the presumptive joint. While all are expressed in the joint
interzone, Wnt4 expression is higher in the mesenchyme flanking the joints that gives rise to the
joint capsule, and Wnt9a is enriched in the mesenchyme surrounding the cartilage primordium in
the tissue that will become tendon [156, 157]. Wnt16, on the other hand, is joint-specific with
high expression levels restricted to the joints of the digits [156]. Mis-expression of Wnt9a in
35
chicks produces gaps in the cartilage matrix that have the morphological and molecular
hallmarks of an interzone [157]. That Wnt signaling is sufficient for joint specification has also
been shown in mice expressing Wnt9a or a constitutively active form of the canonical Wnt
effector β-catenin in Col 2+ cells [156].
However, knockout studies demonstrate that Wnt signaling is not necessary for joint induction
but rather for maintenance. Wnt9a knockout mice initiate joint formation; however, shortly after
specification, the interzone cells ectopically differentiate into cartilage, subsequently causing
joint fusions [158, 159]. Mice null for Wnt9a and Wnt4 have a more severe phenotype with
ectopic cartilage and fusion in additional joints [158, 159]. Furthermore, in β-catenin knockout
mice the joint interzone initially forms but later fails to maintain joint identity and undergoes
chondrogenesis [156, 159]. These studies suggest that canonical Wnt signaling blocks
chondrogenesis in the interzone during joint development. While Wnt exhibits anti-chondrogenic
activity in the interzone, it supports formation of the articular cartilage. A recent study shows
that Wnt and Bmp signaling oppose one another to control the zone of chondrogenic activity.
Noggin expression in the epiphysis blocks Bmp signaling in a region of interzone-adjacent cells,
allowing their differentiation into articular cartilage under the influence of Wnt from the
interzone [135]. Thus, Bmp signaling activates programs for transient cartilage of the growth
plate, whereas Wnt signaling promotes programs for articular cartilage.
While Wnt9a is critical for joint development, it is only necessary after joint specification,
suggesting the presence of an upstream regulator. In a study aimed to identify direct
transcriptional activators of Wnt9a in the developing joint, c-Jun was identified as a critical
36
regulator of cell fate in the interzone [160]. Conditional deletion of c-Jun decreases canonical
Wnt signaling, prevents formation of the characteristic flat interzone cells, and leads to a range
of abnormalities from ectopic cartilage between the articular surfaces to irregular articular
surfaces and ligament hypoplasia. The chondrogenic switch in the cell fate of the interzone cells
can be explained by enhanced Bmp signaling due to down-regulation of Chordin.
1.2.3 Hedgehog
There is strong evidence to suggest that the developing bones influence the establishment and
maintenance of the interzone as signaling center. Signals from the adjacent cartilage anlagen that
regulate growth and maturation of chondrocytes within the growth plate also influence joint
formation. Indian hedgehog (Ihh), expressed by prehypertrophic chondrocytes in the growth
plate, controls the distance between the hypertrophic zone and the articular surface of the joint
through a negative feedback loop with Pthrp. Ihh indirectly promotes Pthrp expression in the
periarticular joint region and in turn, the range of Pthrp signaling determines the length of the
proliferative zone [161-163]. Mice lacking Ihh exhibit long bone defects, as well as joint fusions
[141, 164, 165]. While Gdf5+ joint progenitors are specified in these mice, the cells mislocalize
to the perichondrium at the periphery of the future joint [141, 164]. This suggests that Ihh is
critical for the influx of joint progenitors flanking the prospective joint.
Although Pthrp and Ihh work together in the growth plate, this regulatory relationship is not
entirely conserved in joint development. Constitutively active Pth1r is not sufficient to rescue
joint fusions in Ihh knockout mice [166]. In fact, the developing joint can respond directly to
hedgehog by expressing pathway effectors such as Gli1, Gli3, Hip1, and Patched1 [141, 167].
37
Activation of Ihh signaling in Col2-expressing cells directly stimulates mis-differentiation of the
periarticular cells into columnar chondrocytes by increasing Bmp expression [167, 168].
Correspondingly, joint fusions in these mice are rescued by treatment with Noggin [167].
Furthermore, increased Ihh signaling in the joint interzone induced by loss of Patched1 or
expression of SmoM2 blocks formation of articular cartilage and menisci and induces ectopic
cartilage formation by inhibiting Wnt/β-catenin target genes such as Fgf18 [169]. Fgf18
treatment blocks ectopic cartilage formation induced by hedgehog activation in these mice.
Together these results show that opposing Wnt/β-catenin-Fgf and Ihh-Bmp signals regulate cell
fate in the interzone.
1.2.4 Mechanotransduction
During joint cavitation, the articulating cartilage surfaces of the jointed skeletal elements become
separated by a fluid-filled space. Early studies proposed that the cavity between the bones was
generated through apoptosis of the cells located in the center of the interzone [170]. However,
cell death is quite restricted and thus unlikely to be the driving force in cavitation [138, 171]. An
alternative view is that separation relies on mechanically induced changes in the extracellular
matrix (ECM), particularly the production of hyaluronan, which contributes to the loss of cell-
cell integrity at the plane of cleavage [172, 173]. Since synovial joints are specifically adapted
for motion, it is not surprising that their formation requires input from extrinsic mechanical
forces. Indeed, skeletal muscle paralysis prevents joint cavitation despite normal formation of the
interzone; moreover, mechanical stimulation from the skeletal muscle is necessary for
maintenance of already cavitated joints [174-177].
38
Less is known about how mechanical cues are translated into cellular signals to mediate a
transcriptional response in the interzone cells. Candidate pathways for mechanotransduction in
the joint have been suggested by the identification of genes with mechanoresponsive expression.
Integrin-linked kinases work in an ERK1/2-dependent manner to promote proliferation and
stimulation of SOX9, VEGF and c-Myc in articular chondrocytes under mechanical stress [178].
Mechanical stimuli activate two key effectors downstream of Fgf2 signaling, mitogen-activated
protein kinases P38 and Erk1/2, in the presumptive joint line and promote production of
hyaluronan-rich matrices [179, 180]. On the other hand, immobilization of avian embryos
diminishes expression of Fgf2 in the presumptive joint line prior to cartilaginous fusion [181].
More localized joint immobilization in mice leads to a decrease in Fgf2 at the articular surface,
loss of Bmp2 within the interzone, and expansion of Pthrp expression in the periarticular
cartilage into the joint line region [120]. Changes in ion concentration, specifically Ca
+2
, within a
cell are a common response to mechanical stress. Expressed in the articular and proliferating
chondrocytes, TRPV4 is a membrane protein that induces cartilage specific gene expression
through an influx of Ca
+2
upon mechanical stimulation, while simultaneously suppressing pro-
inflammatory genes [182, 183]. Co-expression of PIEZO1 and 2, mechanically activated Ca
+2
ion channels, in articular chondrocytes is needed for induction of Ca
+2
ion uptake in response to
high levels of stress [184]. Elevated levels of PTHrP, PGE2, ATP and intracellular Ca
+2
are
required for mechanical induction of Prg4 expression [185]. A recent genome-wide survey to
identify mechanosensitive genes differentially expressed in the limb skeleton of the Pax3
Spd
muscle-less mouse model uncovered changes in membrane-associated proteins including
members of the Wnt, Fgf, Notch, and Eph/ephrin pathways [186].
39
1.3 Future directions
Despite advancements in our understanding of interzone specification, maintenance, and
eventual cavitation, many questions still remain. While a variety of signals that regulate synovial
joint development have been identified (Figure 2.1A), the signal or combination of signals that
are both necessary and sufficient for specification of joint progenitor cells is unknown. Wnt
signaling is the only pathway shown to be sufficient for joint establishment, and yet it appears
not to be necessary. One possibility is that there is more Wnt pathway redundancy during
specification than has been explored with the β-catenin knockout or Wnt9a; Wnt4 double
knockout mouse lines. After the interzone is specified, Bmp, Wnt, Ihh, and Pthrp signaling
coordinate histogenesis of the joint. It is unclear how the combinatorial effects of these pathways
on synovial joint progenitor cells lead to the multiple cell types within the mature joint.
Certainly, signaling mechanisms that facilitate cell-cell communication and cellular boundaries
to coordinate cell fate decisions are likely at play, but little is known about the role of such
regulators including Eph/ephrin and Notch. These very same pathways are also candidate
regulators for the process of cavitation. As cavitation advances, the articular surfaces of the
opposing bones undergo the process of morphogenesis, in which their three-dimensional
structure takes shape. Synovial joints are the most structurally diverse of all joint types, including
ball-and-socket joints in the hip and shoulder, saddle joints in the digits, hinge joints in the
elbows and knees, and gliding joints in the wrist. The signaling mechanisms that produce this
variation and also coordinate the shapes of the interlocking articular ends are largely unknown.
Synovial joints show variable expression of and different sensitivity to signaling pathway
components according to their anatomical location. It will be important to explore correlations
between expression differences, signaling intensity and duration, and joint patterning.
40
2. Cartilaginous joints
2.1 Cartilaginous joint structure
Cartilaginous joints unite adjacent bones through either a hyaline cartilage or fibrocartilage
intermediate. There are two types of cartilaginous joints, synchondroses and symphyses.
Synchondroses are immovable and usually temporary joints made of hyaline cartilage that allow
interstitial growth between ossification centers before their eventual fusion. Temporary
synchondroses are located between the primary ossification centers of the growing cranial base
and the developing hip, as well as within the growth plates of the long bones. Permanent
synchondroses that remain unossified are located in the thoracic cage between the ribs and the
sternum. Symphyses, on the other hand, are joints that allow for limited movement and are
composed of fibrocartilage. A symphysis can span a narrow or wide joint space, as in the narrow
strip of fibrocartilage that fills the pubic symphysis, or the thick pad of fibrocartilage that fills the
space between adjacent vertebrae.
Here we will discuss the signaling mechanisms involved in the most studied example of a
cartilaginous joint apart from the growth plate: the intervertebral symphysis, commonly known
as the intervertebral disc (IVD). Along the spinal column, adjacent vertebrae are linked together
by IVD, which provide mechanical stabilization, flexibility, and the ability to bear a significant
load. In the mature IVD, concentric connective tissue rings of the annulus fibrosus (AF) enclose
a gel-like center known as the nucleus pulposus (NP). The outer layer of the AF is tendon-like in
its morphology, while the inner layer resembles hyaline cartilage [187]. The AF anchors the IVD
between two articulating vertebral bodies through the hyaline cartilage endplates. Specifically,
41
the outer AF is anchored to the vertebral body and endplate while the inner AF inserts directly
into the endplate [188].
2.2 Signals in IVD development
The AF, along with the vertebral bodies and their endplates, is embryonically derived from the
sclerotome [189, 190]. While the sclerotome arises from cells within the ventral medial somites
and somitocoele, it is the sclerotome cells of somitocoele lineage that give rise to the AF [191,
192]. Interestingly, the somitocoele is known as the joint-forming compartment of the somites.
The NP component of the IVD, on the other hand, is derived from the notochord [193, 194].
Perturbations to the processes leading up to the formation of the sclerotome and/or notochord can
therefore result in hypoplastic or dysmorphic IVD. Since the relationship between these early
processes and IVD development have been recently reviewed, we will focus on signaling events
after specification of the sclerotome and notochord [195, 196].
Following specification, the ventral sclerotome cells expressing Pax1 and Pax9 migrate and
condense around the notochord to form a continuous perichordal tube [197, 198]. The
perichordal tube acquires a metameric pattern of high and low condensed regions that correspond
to the AF and vertebral body, respectively. Pax1, which inhibits chondrogenesis, remains up-
regulated in the AF anlagen and is down-regulated in the presumptive vertebral bodies through
signals that are yet unknown [198-200]. However, it is clear that migration and segmentation of
the perichordal tube is dependent on signals that emanate from the adjacent notochord. That the
notochord regulates development of the perichordal tube was first shown by notochord excision
experiments, which resulted in the perichordal tube forming an unsegmented cartilage rod [201].
42
Later it was shown that Sonic Hedgehog (Shh) derived from the notochord is required for
multiple steps in IVD development including maintenance of Pax1 expression in the ventral
sclerotome, formation of the perichordal tube, and eventual patterning of the IVDs [202, 203].
As the presumptive vertebral bodies undergo chondrogenesis, the notochord regresses in the
vertebral regions and expands in the center of the IVD to form the NP. This regression is a result
of notochord cell migration towards the presumptive IVD in response to mechanical cues and/or
growth factor gradients originating from the segmenting perichordal tube [204-206]. That signals
from the perichordal tube are critical for NP development is supported by studies showing that
changes in perichordal structure or ECM composition disrupt NP formation. In the absence of
hedgehog signaling, the perichordal tube does not form and notochord cells disperse throughout
the vertebral column instead of coalescing in the presumptive NP [205]. Furthermore, loss of
Pax1 in cells of the ventral sclerotome leads to abnormalities in the metameric arrangement of
the perichordal tube as well as deficiencies in notochord regression and NP formation [200, 207].
2.2.1 Notch
Only a few candidate factors are known to mediate the reciprocal interactions between the
notochord and perichordal tube during vertebral morphogenesis. Deciphering the precise
functions of factors identified thus far has been complicated by their initial roles in
somitogenesis and sclerotome determination. The Notch pathway, which first regulates
periodicity of somite formation and somite patterning, also regulates the patterning of the
perichordal tube. Mice null for Lunatic fringe, a glycosyltransferase that enhances Notch
activation though the Delta1 ligand, fail to form the metameric pattern of high and low density
43
mesenchymal condensations around the notochord [207]. While Notch signaling through Delta1
in the caudal region of the somites is critical for rostral-caudal polarity, overexpression of Delta1
throughout the somites does not affect somite polarity but instead results in vertebral column
defects including loss of the IVD and incomplete extrusion of the notochord [208, 209]. This
suggests that negative feedback mechanisms that regulate somite polarity do not function later
during IVD morphogenesis.
2.2.2 Hedgehog
Hedgehog signaling is another pathway redeployed throughout axial skeletal development.
During somite development, Sonic hedgehog (Shh) from the notochord is required for pacing the
somitogenesis clock, inducing formation and survival of the sclerotome, as well as stimulating
the sclerotome to be competent for subsequent differentiation into chondrocytes [203, 210-213].
Consistent with their origins in the notochord, NP cells continue to express Shh through the early
postnatal period to activate growth and differentiation of the NP, AF, and endplate [214].
Another hedgehog ligand, Indian hedgehog (Ihh) is released by the hypertrophic zone of the
vertebral body growth plate and influences cartilage maturation in the IVD. In mice with
postnatal conditional deletion of Ihh in Col2-expressing cells, the IVDlose the AF and hyaline
cartilage endplate [215]. As seen in synovial joints, Ihh is critical for maintaining distinct regions
of cartilage maturation at the articulating surface of the IVD.
2.2.3 Wnt
The effect of Shh on growth and differentiation of the NP relies on Wnt signaling. Conditional
knockout of Wntless, which is necessary for Wnt secretion, in Shh-expressing cells results in
44
downregulation of Shh target genes [216]. Since canonical Wnt signaling is up-regulated in the
NP upon loss of Shh, a negative feedback loop has been suggested: Wnt signaling activates Shh
signaling, which in turn represses Wnt signaling. The effects of Wnt signaling on IVD
development are more extensive as its activity and localization dynamically change in the IVD
between embryonic and postnatal stages [217]. Outside the NP, Wnt signaling is a known
inhibitor of chondrogenesis and is active in the end plate and AF. Conditional deletion of β-
catenin in Col2-expressing cells leads to increased endochondral bone formation in the endplate
[217]. Furthermore, transient over-activation of Wnt signaling in the postnatal IVD caused
deterioration of the AF [217]. This suggests that Wnt signaling regulates cell-fate determination
and maintenance of the AF.
2.2.4 Bmp and Tgf β
In addition to Wnt, Gdf is also involved in AF cell-fate determination and maintenance. Some
Gdf5+ cells, which give rise to the AF, were found to reside adjacent to the cartilage endplates
and migrate along the lamellae into the disc [218, 219]. Loss of Gdf5 leads to histogenic
abnormalities in the AF including replacement of the normal lamellar architecture with
chondroid tissue that invades the NP [220]. Although primary formation of the NP is normal in
the absence of Gdf5, the AF abnormalities secondarily result in NP deformity and degeneration
[221].
As seen in the synovial joint, inhibitors of the BMP pathway control responsiveness to BMP
signals from the adjacent bones. A Noggin-expressing layer in the IVD blocks pro-chondrogenic
45
Bmp signals from the adjacent vertebral bodies (DiPaola et al., 2005). Noggin null mice exhibit
severe vertebral column defects that include vertebral fusions [150, 222].
AF histogenesis is also regulated by Tgfβ receptors, which are among the earliest known markers
of the presumptive IVD [223, 224]. Conditional inactivation of Tgfβ type II receptor ( T gfβ r2) in
Col2-expressing cells of the AF leads to inappropriate chondrogenesis, fusion of the AF and
vertebral body, and subsequently, loss of the IVD [225]. The anti-chondrogenic role of Tgfβr2 is
in part mediated through the induction of AF specific transcription factors such as Scx,
fibromodulin, Erg1, and Mohawk in the sclerotome [226, 227]. Correspondingly, loss of
Mohawk, which promotes responsiveness to Tgfβ and reduces responsiveness to Bmp in the
outer AF layer, leads to down-regulation of tendon/ligament markers and up regulation of the
chondrogenic factor Sox9 [228]. That the balance between TGFβ and BMP signaling is critical
for AF maintenance is supported by loss of Filamin B, a scaffold protein that regulates signaling
attenuation of the Tgfβ family of receptors. In Filamin B knockout mice, which exhibit enhanced
Tgfβ and Bmp signaling, AF cells acquire the molecular signature of hypertrophic chondrocytes
undergoing endochondral-like ossification [229]. The latent ability of cells in the outer layer of
the AF to differentiate into chondrocytes can be explained by the finding that the AF is derived
from Scx+/Sox9+ bipotent progenitor cells that differentiate into either chondrocytes or
tenocytes [230]. Correspondingly, conditional deletion of Sox9 in Scx+/Sox9+ cells causes
hypoplasia of the inner AF and expansion of the outer AF layer [230].
46
2.3 Future directions
What is known and reviewed here is likely only part of a larger signaling network controlling
IVD development and maintenance (Figure 2.2B). Expression analyses have identified multiple
components of the Wnt, Hedgehog, Fgf, Bmp, and Tgfβ signaling pathways in the IVD [222,
231]. However, functional studies have yet to reveal the full extent to which these pathways
regulate IVD formation and maintenance. Basic knowledge of the signaling pathways regulating
the IVD, as well as a better understanding of their hierarchy and interconnectedness, will be
critical to identifying therapeutic targets in IVD degeneration, for which there are few effective
treatments.
3. Fibrous joints
3.1 Fibrous joint structure
Fibrous joints form a direct union between bones through fibrous connective tissue. There are
three types of fibrous joints: syndesmoses, gomphoses, and sutures. Syndesmoses traverse
widely spaced parallel bones through bands of ligaments or sheets of connective tissue called
interosseous membranes. These joints provide strength, stability, and limited movement between
the shafts of the radius and ulna in the forearm and the tibia and fibula of the leg. Gomphoses are
specialized fibrous joints that anchor the roots of teeth to the bony sockets in the upper and lower
jaw via a thin fibrous membrane called the periodontal ligament. Sutures bind the contiguous
margins of closely opposed bones in the skull through short connective tissue fibers that restrict
most movement and yet provide compliance and elasticity during parturition and postnatal brain
growth. By adulthood in humans, many sutures ossify to permanently fuse adjacent bones. The
signaling mechanisms that regulate suture development are the best described of all fibrous joints
and therefore will be the example we discuss here.
47
3.2 Signals in suture development
A suture is established when the osteogenic fronts of opposing calvarial bones meet following
their apical growth. Once formed, the osteogenic fronts and intervening mid-suture mesenchyme
organize subsequent calvarial bone growth by coordinating intramembranous ossification in
response to mechanical forces from the enlarging brain until the eventual fusion of the sutures in
late adolescence. Intramembranous ossification is spatially organized within the sutures.
Preosteoblasts located within the osteogenic front terminally differentiate and withdraw from the
growing front to incorporate into the bone. Proliferating osteoprogenitor cells, on the other hand,
are maintained at the tip of the osteogenic fronts and advance with the developing bone [232].
The mid-suture mesenchyme maintains a non-osteogenic fate during the embryonic period, and
as the suture matures, the mid-suture zone becomes occupied by fibrous connective tissue [233].
Calvarial bones and their intervening sutures are embryonically derived from neural crest and
paraxial mesoderm. Frontal bones in the rostral calvaria arise from neural crest mesenchyme,
while occipital and parietal bones of the caudal calvaria develop from mesoderm [234-237].
Interestingly, the coronal suture, situated between the frontal and parietal bones, coincides with a
physical boundary between neural crest and mesoderm in mammals [236, 237]. While neural
crest cells and mesoderm each contribute an osteogenic front to the coronal suture, the mid-
suture mesenchyme here is of mesodermal origin. The coronal mid-suture mesenchyme is pre-
specified as a population of Gli1+ cells within the head mesoderm prior to the formation of the
calvarial bone rudiments [238]. These Gli1+ cells, activated by Shh from the notochord and
prechordal plate, transiently express En1 and migrate first to the supraorbital ridge and then
apically to their position at the neural crest-mesoderm boundary. Lineage tracing of a similar
48
population of Gli+ cells in the adult suture shows they can contribute to the osteogenic front,
dura, periosteum, and calvarial bones [239].
3.2.1 Eph/Ephrin
Suture development depends on the establishment and maintenance of cellular boundaries
between osteogenic and non-osteogenic territories via direct cell-cell signaling mechanisms.
Loss of boundary integrity between osteogenic and non-osteogenic compartments induces
ectopic ossification of the mid-suture mesenchyme and subsequently, joint fusion. Normally, as
the calvarial bones grow, osteogenic precursor cells from the bone rudiments migrate apically
along the developing bone, guided by EphA/ephrinA signaling, to join the leading edge of the
bones [237, 240, 241]. EphrinA-expressing osteogenic precursors migrate along an EphA-
expressing cell layer located on the apical surface of the bone. Activation of the EphA by
ephrinA initiates bidirectional signaling that promotes cell-cell repulsion. When expression
levels of EphA or ephrinA are reduced, unrestrained osteogenic cells migrate off their path into
the non-osteogenic compartment of the mid-suture mesenchyme and cause suture fusion [241,
242].
3.2.2 Notch
Notch/Jagged signaling is also critical for delineating boundaries between osteogenic and non-
osteogenic compartments. Active Notch2 signaling is restricted to the osteogenic fronts of the
coronal suture by a layer of Jagged1-expressing cells in the mid-suture mesenchyme [243]. Upon
conditional loss of Jagged1 in the mesoderm, but not the neural crest, Notch2 signaling expands
into the mid-suture mesenchyme, triggering misspecification to an osteogenic fate, and
49
ultimately leading to joint fusion [243]. While the roles of Notch/Jagged and EphA/ephrinA
signaling in suture boundaries are not directly linked, both pathways lie downstream of Twist1, a
transcriptional regulator critical for suture morphogenesis [242, 243].
3.2.3 Fgf
Fgf signaling is a critical regulator of suture development and maintenance. Osteoprogenitor
cells at the osteogenic front express Fgfr2, which is down-regulated upon differentiation. The
onset of osteoblast differentiation in the osteogenic front is concomitant with a transient increase
in Fgfr1 expression that is later reduced in the mature osteoblasts of the bone [244-246]. Human
mutations that increase the activity of FGFR1 or FGFR2 are a major cause of premature suture
fusion, known as craniosynostosis [247]. Similarly, mice carrying orthologous mutations in
Fgfr1 and Fgfr2 develop craniosynostosis before birth due to enhanced osteoprogenitor cell
proliferation and differentiation [79, 81-83, 248, 249].
Fgfr2 functions cell-autonomously within the sutures. Conditional expression of a disease-
causing Fgfr2 mutation in En1-expressing coronal suture progenitor cells is sufficient to induce
craniosynostosis [238, 250]. Interestingly, loss of Fgfr2 simultaneously leads to craniosynostosis
of the coronal suture and open gaps where frontal and sagittal sutures should form [60, 80]. This
apparent paradox is later reconciled through growth. The osteogenic fronts of paired frontal and
paired parietal bones are slow to approximate, due to decreased osteoprogenitor cell proliferation
and differentiation, and eventually fuse during postnatal development [60, 248].
50
A precise spatial gradient of Fgf signaling is critical to establish and maintain the sutures.
Multiple Fgfs are expressed in and around the sutures. Fgf2 and Fgf9 are expressed in the mid-
suture mesenchyme, while Fgf18 is expressed in the osteogenic mesenchyme of the bone [246,
251, 252]. These Fgfs act on Fgfr1- and Fgfr2-expressing cells in the osteogenic fronts and Fgfr3
in the bone. Thus, Fgf signaling is thought to be highest in the osteogenic fronts [244, 245].
Fgf18 knockout mice exhibit suture widening, while no phenotypes have been identified in Fgf2
and Fgf9 knockout mice [251, 253]. Ectopic treatment of the developing sutures with Fgf2-
soaked beads show that Fgf2 can promote mid-suture cell identity by inducing Twist1, a negative
regulator of osteogenesis that directly inhibits Runx2 [252, 254]. The spontaneous mouse mutant
Elbow knee synostosis (Eks) harbors a point mutation in Fgf9 that increases ligand diffusion.
More diffusible Fgf9 activates Fgfr3 in the bone, a region that normally receives low levels of
Fgf, leading to fusion of the coronal and sagittal sutures [255, 256]. That spatial localization of
the Fgf gradient determines the cellular response is further supported by the finding that Fgf4
bead placement at the osteogenic fronts accelerates proliferation and differentiation, whereas
placement on the mid-suture mesenchyme strictly enhances proliferation [246]. Additionally, the
spatial responsiveness of receptors to the Fgf gradient is critical for suture development:
inappropriate localization of Fgfr1 or changes in ligand-binding specificity of Fgfr2 cause
coronal suture fusion [257, 258].
3.2.4 Bmp
Fgf signaling in the suture is tightly linked with Bmp signaling. Bmp4 in the osteogenic fronts
and mid-suture mesenchyme is necessary to induce transcription of Msx1 and Msx2, which are
critical regulators of osteoprogenitor cell proliferation and differentiation [246, 259]. Noggin, a
51
secreted antagonist of Bmp expressed in the suture mesenchyme and the underlying dura,
restricts Bmp activity to the osteogenic fronts of patent sutures [260]. While Bmp induces
Noggin expression, Fgf signaling inhibits it. The fact that craniosynostosis can result from
inappropriate Fgf-mediated repression of Noggin is supported by experiments showing that
ectopic application of Noggin can block suture fusion in a chimeric nude rat model [261].
The Bmp family member Gdf6 plays a critical role early in establishing the coronal suture. In the
developing skull, Gdf6 is first expressed in a supraorbital domain that coincides with the frontal
bone rudiment and shortly thereafter is localized to the mid-suture mesenchyme with Fgfr2 [85,
262]. Interestingly, this Gdf6+ population appears to spatially overlap with the Gli1+/En1+ cells
identified by Deckelbaum et al. that give rise to the coronal suture. Gdf6 mutant mice do not
show morphologic or molecular evidence of suture formation [85, 262]. In particular, Fgfr2
expression is lost in the coronal suture of Gdf6-null mice in a pattern quite similar to what is seen
in En1-null mice, suggesting that these may be the same group of cells [85, 238].
3.2.5 Hedgehog
The hedgehog pathway controls both early and late events in suture formation. Early in
development, Shh from the notochord induces formation of the Gli1+ cells in the mesoderm that
eventually give rise to coronal suture progenitors [238]. Functional studies suggest that Ihh in the
osteogenic fronts integrates with Bmp and Fgf to regulate later events in suture development.
Ihh-null mice have delayed calvarial osteogenesis and wide sutures due to a decrease in Bmp-
mediated recruitment of osteoprogenitor cells to the osteogenic fronts [263]. Loss of negative
regulators in the hedgehog pathway also demonstrates that Ihh promotes osteogenesis in the
52
suture. Excessive Ihh signaling in mice caused by reduced levels of Gli3 and Ptch1 leads to
increased proliferation, enhanced osteogenesis, and premature fusion of the lambdoid suture
[264-266]. In the mid-suture mesenchyme of Gli3 knockout mice, Twist1 expression is reduced,
while BMP signaling and Runx2 expression are expanded [265, 266]. Allelic reduction of Runx2
in the Gli3 mutant background rescues suture fusion [266]. Interestingly, treatment of the Gli3
mutant with Fgf2 reinstates Twist1 expression and also rescues suture fusion [265]. Together
these studies suggest that Ihh signaling promotes osteogenesis through Bmp-mediated activation
of Runx2 and that this activity is counterbalanced by Fgf2-mediated maintenance of Twist1.
3.2.6 Wnt
Acting upstream of Fgf and Bmp signaling, Wnt is critical for maintaining suture patency.
Wnt/β-catenin promotes the commitment of skeletal progenitor cells into osteoprogenitors, and
suppresses their chondrogenic potential [267, 268]. Axin2, a negative regulator of canonical
Wnt signaling that promotes β-catenin degradation, is expressed in the mid-suture mesenchyme
as well as the osteogenic fronts and periosteum of the calvarial bones. Axin2 knockout mice have
fusion of the metopic and coronal sutures due to enhanced osteoprogenitor cell proliferation and
differentiation [269, 270]. This phenotype is concomitant with increased levels of Fgf18, Fgf4
and Fgfr1 expression, as well as elevated BMP signaling [270, 271]. Similarly, mice with
constitutive activation of β-catenin in Axin2-expressing cells also exhibit excessive
intramembranous ossification that results from increased Fgf and Bmp signaling [272]. Thus,
these studies strongly suggest that Wnt signaling promotes the expansion of osteoprogenitor cells
and the commitment of these cells into the osteoblast lineage through Fgf and Bmp signaling.
Reducing elevated Fgf signaling in Axin2 knockout mice through allele reduction of Fgfr1 does
53
not rescue suture fusion. Instead, these double mutant mice develop craniosynostosis due to
ectopic endochondral ossification [273]. This change in cell fate within the mid-suture
mesenchyme is blocked by treatment with the Bmp inhibitor Noggin [273]. Together these
findings show that Wnt regulates lineage specification within the suture by balancing levels of
Bmp and Fgf signaling.
3.3 Future directions
While the structure of the suture may be less complex then the other joints reviewed here, the
signaling regulating its development is no less complicated (Figure 2.1C). Enhanced or
diminished activation of a single signaling pathway, such as Fgf, yields the same phenotype –
suture fusion. It should be noted that the underlying pathologies of “suture fusion” in these
situations are likely distinct. The term “suture fusion” implies that the suture was established but
not maintained. However, a subset of these phenotypes are expected to be the result of defective
joint specification, or failure to form a suture altogether. In future studies it will be important to
distinguish between lack of suture specification and failed maintenance by looking at the
development of En1+ suture progenitor cells.
54
Figure 2.1. Spatial expression patterns of the principal signaling pathways in joint development. The
expression domains of critical signaling pathway components are regionally restricted during
development of the (A) synovial, (B) cartilaginous, and (C) fibrous joints. Articular cartilage (AC);
interzone (IZ); joint capsule (JC); intervertebral disc (IVD); endplate (EP); annulus fibrosus (AF); nucleus
pulposus (NP); osteogenic front (OF); and mid-suture mesenchyme (SM).
Unifying features of joint development
Review of the mechanisms that regulate development of synovial, cartilaginous, and fibrous
joints reveals that there are unifying features in signaling architecture between distinct joint
classes (Figure 2.2). First, the joint progenitor cells of different joint types express genes that
belong to the same principal signaling pathways. For example, the Gdfs, Wnts, and Bmp
inhibitors are all expressed in the presumptive joint compartments of synovial, cartilaginous, and
fibrous joints. Second, the regulatory relationships between the principal signaling pathways are
reiteratively used during the development of distinct joint types. A signaling axis of Wnt-Fgf
from within the joint-forming compartment opposes chondrogenic signals from Ihh-Bmp in
order to promote specification and differentiation of specialized tissues at the joint-bone
interface. In synovial and cartilaginous joints, the balance between Wnt and Ihh signaling
promotes formation of intermediate tissues such as hyaline cartilage and fibrous connective
55
tissue by limiting chondrogenic potential. In the fibrous joint of the suture, Wnt and Ihh
signaling maintain the osteogenic front by balancing osteoprogenitor cell specification with
terminal osteoblast differentiation. Third, the same signal pathway is utilized for the same
function in distinct joint types. Cell-cell signaling through the Notch pathway is employed in the
IVD and suture to define osteogenic and non-osteogenic compartments. Similarities in the
molecular regulatory network between joint types are coincident with similarities in joint
morphogenesis. Joint progenitor cells that make synovial, cartilaginous, and fibrous joints share
a common feature: they are sourced from the outside and move into the joint space.
One possibility raised by previous studies is that diverse types of joints have a common
evolutionary origin. For example, the structure of some cranial joints is taxonomically variable in
lizards [274]. While the quadrate-pterygoid joint of the jaw is fibrous in gekkotans, it is a mixed-
phenotype joint—part fibrous and part synovial—in iguanidae. Similarly, joints within the axial
and appendicular skeleton also vary between tetrapods. The costal joint is fibrous in most
mammals and synovial in birds [275, 276]. The knee joint, which is synovial in mammals, is
fibrous and lacks a synovial cavity in amphibians such as the salamander and frog [277].
Additionally, joints within the amphibian ankle and wrist show a mixed phenotype of fibrous and
synovial. These studies suggest that the signaling network responsible for specifying the joint is
competent to generate more than one joint class.
56
Figure 2.2 Signaling networks in joint development have unifying features. There is overlap in the
principal signaling pathways and their regulatory interactions between (A) synovial joints, (B)
cartilaginous joints, and (C) fibrous joints.
Conclusions
Overlap in the signaling networks that regulate distinct joint types is certainly more extensive
than is appreciated here. The role of Fgf signaling in joint development has been largely defined
by studies in the sutures. However, craniosynostosis syndromes caused by mutations in FGFR2
are also associated with synovial joint fusions within the limb as well as intervertebral fusions
[278, 279]. To understand this regulatory connection, it will be important to more closely
examine the role of Fgf signaling in synovial and cartilaginous joint development.
Despite overlapping signaling mechanisms in the development of the synovial, cartilaginous, and
fibrous joints, the final structures of these joints are quite specialized, and there is significant
morphological variation within each category. How can use of similar signaling networks across
57
joint types produce the unique histogenic structure of each joint? Signaling pathways can
produce varied biological responses depending on the intensity and duration of the signal.
Differences in the spatiotemporal expression patterns of signaling pathway components or
inhibitors can account for the modulations in dose- and time-dependent response. In the joints,
this is evidenced by varied involvement of select Wnt, Gdf, and Fgf ligands, as well as Bmp
inhibitors.
Conservation of the regulatory network across joint types may suggest that synovial,
cartilaginous, and fibrous joints are homologous developmental modules that have undergone
specialization. If they are indeed homologous modules, then the specialized transcription factors
that define the joint compartment should be shared across joint types. These joint-determining
transcription factors should be expressed within the cells of the presumptive joint compartment,
required for initial specification of joint identity, and sufficient for joint specification under the
correct conditions [280]. A transcription factor that fits this description has yet to be identified in
any of the joint types.
Here we have illustrated the similarities and differences between distinct joint types in an effort
to better understand joint development. By comparing the different joint classes, an overlap in
the signaling networks becomes clear. Discoveries in one joint type can therefore provide
mechanistic insights into the development of the other joint classes. Since all joint classes are
commonly affected by disease and injury, there is a clinical need to develop molecular-based
strategies to repair and/or rebuild joint tissues. Identifying unifying features in the mechanisms
that instruct joint development enhance our potential to reach this goal.
58
Acknowledgements
This work was supported by the National Institutes of Health [R01DE025222 to A.E.M.] and
March of Dimes [#6-FY15-233 to A.E.M.]. We thank Bridget Samuels for her help in editing
this manuscript.
59
CHAPTER 3: Nuclear FGFR2 regulates musculoskeletal integration within the developing
limb
#
Joanna E. Salva
1, 2
, Ryan R. Roberts
1,2
,
Taylor S. Stucky
1, 2
, Amy E. Merrill
1, 2,*
1
Center for Craniofacial Molecular Biology, Ostrow School of Dentistry, University of Southern
California, Los Angeles, CA, 90033
2
Department of Biochemistry and Molecular Medicine, Keck School of Medicine, University of
Southern California, Los Angeles, CA, 90033
*
Correspondence: amerrill@usc.edu
Key words: limb development, Bent Bone Dysplasia Syndrome, connective tissue
Grant Sponsor and Number: NIH/NIDCR R01DE025222, NIH/NIDCR R01DE025222-01S1
NIH/NIDCR T90DE021982, NIH/NICHD 5T32HD060549-02, and March of Dimes #6-FY15-
233
# This work has been submitted for publication in Developmental Dynamics, August 2018.
60
ABSTRACT
Background: Bent Bone Dysplasia Syndrome (BBDS), a congenital skeletal disorder caused by
dominant mutations in Fibroblast growth factor receptor 2 (FGFR2), is characterized by bowed
long bones within the limbs. We previously showed that the FGFR2 mutations in BBDS enhance
nuclear and nucleolar localization of the receptor; however, exactly how shifts in subcellular
distribution of FGFR2 affect limb development remained unknown.
Results: Targeted expression of the BBDS mutations in the lateral plate mesoderm of the
developing chick induced angulated limbs, a hallmark feature of the disease. Whole mount
analysis of the underlying skeleton revealed bent long bones with shortened bone collars and, in
severe cases, dysmorphic epiphyses. Epiphyseal changes were also correlated with joint
dislocations and contractures. Histological analysis revealed that bent long bones and joint
defects were closely associated with irregularities in skeletal muscle patterning and muscle-to-
bone attachment. The spectrum of limb phenotypes induced by the BBDS mutations were
recapitulated by targeted expression of wild type FGFR2 appended with nuclear and nucleolar
localization signals.
Conclusions: Our results indicate that the bent long bones in BBDS arise from disruptions in
musculoskeletal integration and that increased nuclear and nucleolar localization of FGFR2 plays
a mechanistic role in the disease phenotype.
61
INTRODUCTION
Fibroblast growth factor receptors (FGFRs) are a central branch point in the FGF signaling
network that allows for context-specific regulation of cell growth, proliferation, differentiation,
and survival during development [1]. In response to extracellular FGF ligands, membrane bound
FGFRs dimerize, autophosphorylate, and launch downstream signaling cascades including
MAPK, JAK/STAT, and PI3K [1]. While the role of FGFRs as transmembrane signal
transducers are well-defined, it is unclear how these receptors produce a transcriptional signature
distinct from other receptor tyrosine kinases that utilize the same downstream signaling cascades.
Our lab and others have shown that FGFRs and their ligands act in the nucleus to regulate gene
expression [13, 15, 27, 32, 33, 281, 282]. Although nuclear FGFR signaling could provide
insight into FGF signaling specificity in development and disease, little is known about the
significance of this activity in vivo.
FGFR signaling is fundamental for development, acting in the earliest stages of embryogenesis
and then reiteratively in organogenesis of diverse tissue types. One of the best-described roles of
the FGF pathway is in the developing limb and its underlying skeleton. During limb bud
initiation, FGFR signaling mediates a positive feedback loop between the epithelium and
mesenchyme to induce formation of the apical ectodermal ridge (AER). Subsequently, FGF
signaling promotes outgrowth, proximal-distal patterning, and anterior-posterior patterning of the
appendicular skeleton [45, 63-65, 283-288]. Fgfr2, which is expressed in both the epithelium and
mesenchyme, plays a critical role in these processes. The epithelial-specific Fgfr2 isoform, Fgfr2
IIIb, is expressed in the limb ectoderm and AER, where it is required for limb formation [50, 59].
The mesenchyme-specific Fgfr2 isoform, Fgfr2 IIIc, is expressed in the skeletal anlagen and
62
later in the perichondrium and periosteum, and is not required for limb formation [9, 77].
Instead, loss of Fgfr2 IIIc results in short limbs, reduced long bone growth, and delayed bone
mineralization [60]. A key role for mesenchymal Fgfr2 in limb skeletogenesis is further
supported by conditional deletion of Fgfr2 in the skeletal anlagen, which reduces thickness of the
perichondrium/periosteum and size of the hypertrophic chondrocyte zone [80].
The significance of Fgfr2 in limb formation is underscored by limb malformations in congenital
human disorders with FGFR2 mutations. Loss-of-function FGFR2 mutations in Lacrimo-
auriculo-dento-digital (LADD) syndrome lead to shortened, curved, and missing digits, as well
as duplicated thumbs, and radioulnar fusions [91, 92, 131]. Gain-of-function mutations in
FGFR2 result in shortened digits with soft or osseous fusions as well as radio-humeral joint
fusion in Pfeiffer, Crouzon, Apert, and Antley-Bixler syndromes [99, 103, 130, 289-291]. Mouse
models for these syndromes phenocopy many of the limb abnormalities and reveal that increased
Fgfr2 activity elevates osteoprogenitor cell proliferation and differentiation [79, 81-83, 87].
While human disorders that result from gain or loss of FGFR2 activity primarily exhibit defects
in distal bones and joints of the limb, Bent Bone Dysplasia Syndrome (BBDS) [OMIM
#614592], the most recent addition to the FGFR2 spectrum of disorders, affects bones along the
entire proximal-distal axis, including the girdle. BBDS patients have brachydactyly and trident
configuration of the digits: bending of the tibia, fibula, and femur: and hypoplasia of clavicles,
scapulae and pubis [31, 104]. The distinct limb phenotype in BBDS suggests that these mutations
are functionally distinct from other disease-causing FGFR2 mutations that lead to gain or loss of
receptor function.
63
We showed previously that the BBDS mutations FGFR2
M391R
and FGFR2
Y381D
are located in the
transmembrane domain of FGFR2 and enhance receptor localization to the nucleus in skeletal
progenitor cells [15, 31, 32]. While we are beginning to understand the molecular consequences
of the BBDS mutations on nuclear FGFR2 signaling, it remains unclear how altered localization
of FGFR2 affects skeletal development in vivo. Here we employ the chick model to better
understand the etiology of the limb abnormalities in BBDS. We find that in ovo expression of the
BBDS mutations in the developing hindlimb mesenchyme induces bent long bones, a phenotype
that can be recapitulated with enhanced nuclear localization of wild type FGFR2. Furthermore,
we uncover strong evidence to suggest that long bone bending results primarily from defects in
musculoskeletal integration.
RESULTS
The BBDS mutations alter limb development in ovo
To target expression in the skeletogenic mesenchyme of the developing chick hindlimb,
concentrated RCAS virus was unilaterally injected into the coelom between the two layers of the
lateral plate mesoderm (LPM) at Hamburger Hamilton (HH) stage 15 [36, 292] (Fig3.1 A).
Injection of RCAS-GFP in the LPM at HH15 showed GFP expression in all three limb segments
by HH29 (Fig3.1 B). Immunofluorescent staining of frozen sections at HH29 showed that in the
developing flexor muscles along the posterior of the bone GFP
+
cells were colocalized with
Tcf4, a transcription factor that is highly expressed in connective tissue fibroblasts of tendons,
ligaments, and muscle connective tissue (Fig3.1 C-E; arrows) [293]. While GFP
+
cells were
associated with Tcf4
+
muscle connective tissue, immunofluorescent detection of the muscle
marker myosin heavy chain (MF20) identified only a few muscle cells that were GFP
+
64
confirming that the skeletal muscle was not readily targeted (Fig3.1 F,G). LPM injections at
HH15 were not expected to target skeletal muscle cells, as migration of myogenic precursors
from the somites into the hindlimb begins at HH17 [294, 295]. Tcf4 is also expressed in pre-
hypertrophic and hypertrophic chondrocytes of the limb [296]; however, our infections resulted
in only sporadic GFP
+
cells in the cartilage (Fig3.1 C,G; asterisks). Clusters of GFP
+
cells were
also identified within the Tcf4
-
cells located between the developing muscle and bone, where
tendons form (Fig3.1 C,D; arrowheads). Together this data suggests that our injection strategy
for RCAS expression largely targets cells associated with the Tcf4
+
and Tcf4
-
cells that give rise
to muscle connective tissue and tendon.
To determine the effects of the BBDS mutations on limb development, RCAS virus expressing
GFP or the IIIc isoforms of FGFR2
WT
, FGFR2
M391R
, and FGFR2
Y381D
were injected into the
LPM at HH15. Embryos were collected for analysis at HH29 after the cartilage anlagen of the
long bones are established [297-300]. Whole mount analysis of the embryos showed that targeted
expression of GFP (n=16) and FGFR2
WT
(n=11) resulted in phenotypically normal hindlimbs
(Fig3.2 A-F). This suggests that expression of the RCAS virus or expression of FGFR2
WT
alone
do not interfere with normal limb development. Targeted expression of FGFR2
M391R
(n=15) and
FGFR2
Y381D
(n=17), on the other hand, resulted in angulated and splayed hindlimbs, which are a
hallmark of BBDS (Fig3.2 G-L) [31, 104]. Embryos expressing FGFR2
M391R
and FGFR2
Y381D
also exhibited autopod defects, including an anterior-posterior spreading of the foot plate with
pronounced deflection of digit 1 (n=11) and, in some cases, a small footplate with split-foot
malformation (n=15) (Fig3.2 I, L). Together these data show that expression of FGFR2
M391R
and
65
FGFR2
Y381D
in a subpopulation of the LPM that gives rise to muscle connective tissue and
tendon in ovo induces hindlimb abnormalities that are consistent with BBDS.
The BBDS mutations induce bent long bones in ovo
To identify the cause of the angulated long bone in the hindlimb, we examined the underlying
bone and cartilage in whole mount at HH36 using Alizarin red and Alcian blue, respectively. The
skeletal abnormalities in limbs expressing FGFR2
M391R
and FGFR2
Y381D
were classified as mild
or severe, depending on the degree of bending and number of missing skeletal elements. In
mildly affected limbs expressing FGFR2
M391R
(n=15) and FGFR2
Y381D
(n=11), there was mild
bowing of the femur and tibiotarsus, as well as shortening of the femoral bone collar compared
to GFP controls (n=8) (Fig3.3 A-C,E; arrows). Mildly affected limbs also exhibited knee
abnormalities including fibular dislocation, decreased joint angle, as well as malformed
epiphyses compared to controls (Fig3.3 A’,B,C’,E’). In severely affected limbs, FGFR2
M391R
and FGFR2
Y381D
expression caused more pronounced bowing of the femurs and loss of the
tibiotarsus (Fig3.3 D,F). When the tibiotarsus was missing, the fibula and the femur formed the
primary articulation of the knee, and the femur exhibited a malformed, cup-shaped epiphysis
(Fig3.3 D’,F’). Interestingly, the fibulare, which initially develops as part of the fibula and later
detaches to fuse with the epiphysis of the tibiotarsus [301], remained at the distal end of the
fibula where it connects with the metatarsals (Fig3.3 D’,F’; arrowheads). Without the tibiotarsus,
the fibulare was posteriorly oriented resulting in displacement of the metatarsals. In both mild
and severe embryos, missing digits were observed mostly affecting digit 1 and to a lesser extent
digit 2 (n=8) (Fig3.4 A-C). FGFR2
M391R
and FGFR2
Y381D
expression also caused defects in the
bones of the pelvic girdle, including a hypoplastic ilium and ischium, as well as a hypoplastic
66
and bowed pubis (Fig3.4 D-E). Altogether, these results demonstrate that expression of the
FGFR2
M391R
and FGFR2
Y381D
in the connective tissue progenitors of the hindlimb produce
skeletal defects in that are consistent with phenotypes seen in BBDS.
Musculoskeletal integration in the hindlimb is altered by BBDS mutations
RCAS-GFP injections primarily targeted cells within the muscle connective tissue and tendon,
and not those in the cartilage (Fig3.1). To better understand how FGFR2
M391R
and FGFR2
Y381D
expression in these tissues could result in long bone defects, we examined the bone and its
surrounding tissues histologically at HH36. Paraffin embedded samples were sectioned and
stained with Hall and Brunt Quadruple (HBQ) stain, which differentially stains cartilage (blue),
bone (pink), and muscle (purple) [302]. Several abnormalities were identified within the long
bones of FGFR2
M391R
and FGFR2
Y381D
expressing limbs. The epiphysis of the tibiotarsus at the
knee was morphologically abnormal with loss of its typical V-shape (Fig3.5 A-D; asterisk). In
the diaphysis of the bowed tibiotarsus and femur, mineralized bone matrix was asymmetrically
deposited at the bend (Fig3.5 E-L; arrowheads).
The most notable abnormalities in limbs expressing FGFR2
M391R
and FGFR2
Y381D
were in the
muscles and connective tissues surrounding the bone. At the knee, muscle integrity was
disrupted, the positions of the muscles relative to the bones were displaced, and the tendinous
connections between muscle and bone were irregular (Fig3.5 A-D; brackets). The muscle along
the anterior of the tibiotarsus had a broader area of attachment on the patella of mutant embryos
than controls (Fig3.5 A-D; arrows). Flexor muscles, which are responsible for knee flexion and
hip extension and insert along the posterior surface of the femur and tibiotarsus, were poorly
67
attached and disorganized, especially at the site of long bone bending (Fig3.5 E-L; brackets).
Changes in muscle connective tissue patterning result in similar muscular defects [293, 303].
Therefore, these results suggest that expression of the BBDS mutations alter development of the
connective tissues by disrupting their relative patterning and connections to the bone.
Enhanced nuclear localization of FGFR2 in the hindlimb induces BBDS-like abnormalities
Previously, we showed that the BBDS mutations enhance nuclear and nucleolar localization of
FGFR2 in patient-derived chondrocytes and osteoprogenitor cells [15, 31, 32]. In ATDC5
prechondrocytes, immunofluorescent localization of FLAG-tagged FGFR2
M391R
and FGFR2
Y381D
and the nucleolar marker Polr1D confirmed that these mutations enhanced receptor localization
to the nucleus and nucleolus of chondrogenic cells (Fig3.6 A-C). To test the idea that the
hindlimb phenotypes induced by FGFR2
M391R
and FGFR2
Y381D
result from enhanced nuclear
localization as opposed to neomorphic activity, we appended the IIIC isoform of the FGFR2
WT
cDNA with the classical nuclear localization signal (NLS) from the SV40 Large T antigen or a
synthetic nucleolar localization signal (NoLS) [15, 304, 305]. Immunofluorescent localization of
FLAG-tagged NLS-FGFR2
and NoLS-FGFR2 in ATDC5 cells along with Polr1D showed that
these modifications enhanced nuclear and nucleolar localization of the receptor, respectively,
compared to FGFR2
WT
(Fig3.6 D, E).
To determine the extent to which the enhanced nuclear FGFR2 activity induced the hindlimb
phenotypes, injections of RCAS virus expressing NLS-FGFR2
and NoLS-FGFR2 were targeted
to the hindlimb LPM at HH15. At HH29, hindlimbs expressing NLS-FGFR2 (n=16) and NoLS-
FGFR2
(n=15) exhibited whole mount phenotypes similar to those of the BBDS mutations,
68
including limb angulation and splaying, as well as pronounced deflection of digit 1 (Fig3.7 A-I).
Whole mount skeletal preps stained with Alizarin red and Alcian blue at HH36 revealed that
hindlimbs expressing NLS-FGFR2 and NoLS-FGFR2 had bent long bones (Fig3.8 A, C, E),
short bone collars (Fig3.8 B), and abnormal angulation of the knee joints (Fig3.8 A’, B, C’, E’).
In severely affected limbs, the tibiotarsus was absent and the articulation between the femur and
fibula was abnormal (Fig3.8 D, F).
Histological sections stained with HBQ at HH36 showed that NLS-FGFR2 and NoLS-FGFR2
expressing limbs had morphological changes in the tibiotarsus epiphysis with decreased
definition of the condyles (Fig3.9 A-D; asterisks). Both the femur and tibiotarsus revealed
asymmetric mineralization (arrowhead) at the point of the bend in NLS-FGFR2
hindlimbs, while
the NoLS-FGFR2 embryos were milder in phenotype with small bone collars (Fig3.9 E-L;
arrowhead). NLS-FGFR2 and NoLS-FGFR2 expressing hindlimbs also showed alterations in
muscle patterning. The muscles surrounding the knee were mispatterned and fragmented with
irregular tendon attachments (Fig3.9 A-D; brackets), and the flexor muscles along the posterior
of the femur and tibiotarsus were poorly and irregularly attached (Fig3.9 E-L; brackets).
Altogether, the hindlimb abnormalities induced by enhanced nuclear and nucleolar localization
of FGFR2 IIIc phenocopy those caused by expression of the BBDS mutations. These results
suggest that abnormal limb development in this BBDS chick model largely results from
increased nuclear FGFR2 IIIc.
69
DISCUSSION
In this study, we demonstrate that expression of BBDS mutations in the mesenchymal precursors
of the chick hindlimb induces bowed long bones, a hallmark skeletal finding of the human
disease. Our analysis suggests that bowed long bones are secondary to the mutations’ effect on
development of the soft connective tissues surrounding the bone, including muscle connective
tissue and tendon. We showed previously that the BBDS mutations enhance nuclear and
nucleolar localization of the receptor. Here we find that enhanced nuclear and nucleolar
localization of wildtype FGFR2 in the hindlimb mesenchyme recapitulates the effects of the
BBDS mutations. Altogether, these results suggest that enhanced nuclear FGFR2 in the
developing hindlimb triggers bowing of the long bones by disrupting musculoskeletal
integration.
Hallmark skeletal features of BBDS are recapitulated in the chick hindlimb
RCAS-mediated expression of FGFR2
M391R
and FGFR2
Y381D
in chick lateral plate mesoderm
causes skeletal malformations in the hindlimb that phenocopy BBDS. Bowing of the femur, tibia,
and fibula along with hypoplasia of pubis are key diagnostic features of BBDS [31, 104]. In the
chick, we showed that FGFR2
M391R
and FGFR2
Y381D
induce bowing of the femur and tibiotarsus,
as well as hypoplasia of the pubis. Additional skeletal abnormalities in the chick include
epiphysis hypoplasia, decreased angulation of the knee joint, and lateral leg splaying at the hip
joint. Likewise, abnormal joint angulation and limited joint extension occur in BBDS [104].
BBDS also presents with brachydactyly, hypoplasia of the distal phalanges, and trident-hand
deformity in which deflection between digits 1 and 2 as well as digits 3 and 4 gives the hand a
fork-like appearance [31, 104]. In the chick, FGFR2
M391R
and FGFR2
Y381D
result in pronounced
70
deflection of digit 1 in the hindlimb and, in some cases, missing anterior skeletal elements and
split-foot malformation. It is not completely clear how missing anterior skeletal elements is
related to BBDS; however, split-hand malformation is a type of brachydactyly [31, 104, 306].
Nevertheless, it was not expected that all aspects of the BBDS skeletal phenotype would be
recapitulated in the chick because our RCAS injection strategy largely targets soft connective
tissues and therefore is not highly expressed within the bony elements of the limb.
The BBDS mutations alter musculoskeletal integration in the chick hindlimb
Clinical findings, including abnormal joint angulation, bent long bones, limited joint extension,
and ligamentous laxity, suggest that musculoskeletal integration is altered in BBDS [31, 104].
The idea that the BBDS mutations interfere with musculoskeletal integration is here supported
by the chick model. Our data suggest that bowed long bones in the chick result from
abnormalities in skeletal muscle patterning and irregularities in muscle-to-bone attachment.
Proper development of the skeleton relies on feedback from the surrounding skeletal muscle. A
chick model for disorganized muscle patterning has a bowed tibiotarsus with dysmorphic,
flattened epiphyses [116, 117, 122, 123]. In immobilized chicks or ‘muscleless limb’ mice, the
absence of muscle contractions during development leads to joint abnormalities, as well as shape
changes and reduced bone formation in the skeletal elements [118-121]. During fetal
development, the flexor muscles functionally predominate over the extensor muscles in the legs,
and irregular development of the flexor muscles can lead to outward bowing of the long bones
[112]. Abnormalities in flexor muscle patterning and flexor muscle-to-bone attachment in the
chick model for BBDS could disrupt the mechanical forces experienced by the growing bones
71
and subsequently lead to the skeletal defects we observe including outward bowing of the long
bones.
While muscle cells are not targeted by our RCAS injections, the altered skeletal muscle
patterning and muscle-to-bone attachment observed in the BBDS chick model can be explained
by our targeting the LPM-derived Tcf4
+
cells that give rise to the fibroblasts of muscle
connective tissue and tendons. Muscle connective tissue fibroblasts regulate muscle
morphogenesis by establishing a pre-pattern for the muscle that the migrating myoblasts follow
[293, 295, 307-309]. In conjunction, connective tissue fibroblasts contribute to the tendons,
which mediate muscle-to-bone attachment at their origin and insertion and provide boundaries
for muscle formation [308]. Correspondingly, loss of Tcf4
+
connective tissue cells results in
misshaped muscles, delayed myoblast differentiation, altered origin and insertion sites of the
muscles and joint contractures [293, 303]. Our data suggest that increased nuclear FGFR2 in the
Tcf4
+
cells alters musculoskeletal integration by disrupting development of the muscle
connective tissue and tendons.
The pathophysiology of bowed long bone beyond BBDS
Bent long bones in the legs are a relatively common occurrence in skeletal birth defects with
more than 40 distinct disorders being associated with bent femurs [108]. Despite genetic
heterogeneity in bent bone disorders, clinical evidence suggests that long bone bending results
from a combination of poor bone material properties and altered muscle forces [108-110, 310].
Osteogenesis imperfecta caused by COL1A1 and COL1A2 mutations has weakened bone
strength with reduced muscle size and function [114, 311]. Similarly, Stüve-Wiedemann
syndrome exhibits osteopenia and myotonia due to mutations in LIFR [109, 110, 126, 310].
72
There is also evidence to suggest that imbalanced muscle forces caused by abnormal pattering of
muscle and muscle-to-bone attachment leads to long bone bending in Campomelic dysplasia, a
disorder caused by haploinsufficiency of SOX9 [112]. This idea is consistent with the known role
for Sox9 in the Sox9
+
/Scx
+
progenitor cells that give rise to the tendon-bone connection [230,
312]. Interestingly, multiple aspects of the skeletal phenotype in Campomelic dysplasia overlap
with BBDS. Our observations here in the chick model for BBDS indicate that defects in the
tissues responsible for musculoskeletal integration, including muscle connective tissue and
tendon, are sufficient to induce bowed long bones.
EXPERIMENTAL PROCEDURES
Cloning
The cDNA coding for FGFR2, FGFR2
M391R
, FGFR2
Y381D
, and NoLS-FGFR2 were subcloned
from pCMV Tag4a vector (previously described in Neben et al., 2014) into the ClaI restriction
site in the RCAS BP(A) vector using Sense and Antisense ClaI FGFR2 primers. To generate the
cDNA coding for NLS-FGFR2, Phusion Site Directed Mutagenesis (Thermo Fisher Scientific)
was used to insert the SV40 Large T antigen NLS between the signal peptide and the first Ig
domain of FGFR2 using the NLS Insertion Forward and NLS Vector Reverse primers. NLS-
FGFR2 was then subcloned from pCMV Tag4a vector into the ClaI site of RCAS BP(A) vector.
Sequence of clones was verified via Sanger sequencing. All primers are found in Supplemental
Table 1.
73
RCAS Virus Production
RCAS virus was produced according to Logan and Tabin, 1998. Briefly, DF-1 chicken
fibroblasts were plated at 7.5x10
6
cells per 100 mm dish with DMEM, 10% Fetal Bovine Serum
(FBS), Penicillin/Streptomycin/Glutamine, and after 24 hours, transfected with 17µg RCAS
BP(A) vectors using Fugene HD (Promega). The cells were expanded to six 150mm dishes over
4-5 days. Once cells reached superconfluency, media was collected and flash frozen. To
concentrate the virus, frozen viral media was thawed, filtered with a pre-filter (Nalgene) and
45µm filter (Corning), and concentrated by ultracentrifugation at 21,000 rpm for 3hrs using a
SW28 rotor. Supernatant was decanted and the viral pellet resuspended, aliquoted, and stored at -
80°C.
Viral Titration
A 24-well plate was seeded with DF-1 cells at 1.75x10
5
cells/well. Concentrated virus was
serially diluted in DMEM to concentrations ranging from 10
-3
to 10
-10
. 500µl of each dilution
was added per well and incubated for 48 hrs. Viral media was removed, cells were washed with
PBS and fixed with 4% paraformaldehyde (PFA) for 10 min. Cells were permeabilized using
PBST (1xPBS 0.1% Triton X-100) for 2 min, blocked using 1% goat serum and incubated
overnight with primary antibody against the viral coat protein Gag-Pro (AMV-3C2 supernatant,
1:10, DSHB deposited by D. Boettiger). The next day, cells were washed with PBS and
incubated with goat anti-mouse Alexa 488 (Invitrogen) secondary antibody at 1:400 in 1% goat
serum. Colony forming units (CFUs) were counted in each well until only 1 CFU was found per
well. The titer was calculated by the number of CFUs and the inverse of the dilution factor
giving the infection units (IU) per ml.
74
Targeted RCAS infection in chick embryos
Pathogen free SPF eggs (Charles River) incubated to HH15 were windowed and the LPM was
targeted for RCAS infection. The virus was injected into the coelom formed by the LPM at the
level of somites 26-30 in the hindlimb field to infect the connective tissue of the limb. Virus was
diluted to 1x10
8
IU/ml in 0.02% Fast Green. Needles were pulled from capillary tubes (0.58 mm
World Precision Instruments) using a laser-based micropipette puller (Sutter Instruments Co). A
Picospritzer III microinjector (Parker Hannifin) was utilized to inject a total of 13.2nl of virus.
Eggs were resealed and incubated at 39°C until desired stage.
Whole mount embryo analysis
HH29 embryos were collected, washed in 1xPBS and fixed in 4%PFA overnight at 4°C.
Embryos were dehydrated stepwise in ethanol to 70% ethanol and imaged using a Leica M125
stereoscope with IC80 HD camera.
Whole mount skeletal staining
Chicken embryos were collected at HH35-36, dehydrated in 95% EtOH, and stained overnight in
1% Alcian blue/30% acetic acid in ethanol. Tissues were cleared in 0.25% KOH and stained
overnight with Alizarin red in 0.25% KOH. Samples were treated with ascending volumes of
glycerol in 0.25% KOH and stored in 75% glycerol/0.25% KOH. Imaging was performed using
a Leica M125 stereoscope with IC80 HD camera. Knee angle and femoral bone collar length
were measured using US National Institutes of Health software program ImageJ. To measure the
knee angle, lines were drawn from the neck of the femur to the femur patellar surface to the
groove in the distal end of the tibiotarsus, and the angle formed was measured. The length of the
75
femur was measured using the landmarks just described, and the bone collar was measured from
the proximal end of the mineralized collar to the distal end of the mineralized collar. Bone collar
size was calculated as a ratio of the length of the bone collar to the full length of the femur. Each
measurement was performed in triplicate and the average was calculated. Data is represented as
the mean ± standard error of the mean. Two-tailed Student’s t-test was used to calculate
significance. P-value of <0.05 was considered statistically significant (n=8).
Histology
Chicken embryos were harvested at HH36, fixed in 4% PFA, dehydrated stepwise in ethanol and
processed for paraffin embedding. Samples were cut in 8µm sections and placed on superfrost
slides (VWR). Slides were de-paraffinized, rehydrated, and stained using Hall and Brunt
Quadruple stain (HBQ) [302]. Samples were imaged at 10x using a Keyence BZ-X710
microscope.
Cryosectioning
RCAS-GFP embryos collected at HH29 were fixed in cold 4% PFA for 30min, washed in cold
PBS for 30min, equilibrated in 10% sucrose for 1.5hrs at 4°C, and then incubated in 30% sucrose
overnight at 4°C. Samples were transferred to O.C.T compound (Tissue Tek) to equilibrate for
1hr at 4°C prior to embedding in O.C.T compound. Cryoembedded samples were cut into 10µm
frozen sections, placed on Superfrost Plus slides (VWR) and stored at -80°C.
76
Immunofluorescence
ATDC5 cells were plated at 8x10
4
cells/well in chamber slides (LabTek) with DMEM Ham’s
F12 media (Corning) with 5% FBS and incubated at 37°C. Next day cells were transfected with
500ng pCMV Tag4a FGFR2, FGFR2
M391R
, FGFR2
Y381D
, NoLS-FGFR2, and NLS-FGFR2
vectors using Lipofectamine 2000 (Thermo Fisher Scientific). Cells were grown for 48 hrs prior
to fixation with 2% paraformaldehyde (PFA) for 10min. Samples were washed with PBS,
permeabilized with PBST for 10sec, washed again with PBS, and then blocked with 10% goat
serum (Invitrogen) for 1hr. The primary antibodies mouse anti-FLAG (Sigma, F3165; 1:400) and
rabbit anti-Polr1d (Abcam, ab104115; 1:400) were diluted in 1% goat serum/PBS and incubated
on the slides overnight at 4°C. The next day, slides were washed with PBS and incubated for 1hr
at room temperature with goat anti-mouse Alexa568 and goat anti-rabbit Alexa488 (Invitrogen)
secondary antibodies diluted to 1:400 in 1% goat serum/PBS. Slides were again washed with
PBS and coverslipped using Vectashield with DAPI (Vector Labs). Cells were imaged at 90x
using Leica SP8 confocal microscope.
Frozen sections were washed three times using PBST and blocked with 10% goat serum.
Primary antibodies were diluted in 10% goat serum/0.1% Triton-X 100 and incubated at 4°C in a
humidified chamber overnight. After washing with PBST, samples were incubated in secondary
antibody diluted 1:400 in 10% goat serum/0.1% Triton-X 100 for 1hr at room temperature.
Antibodies used include: anti-GFP (Abcam, ab6556, 1:1000), anti-myosin (DSHB deposited by
D.A. Fischman, MF20-c; 1:200), and anti-Tcf4 (Cell Signaling, cs2569, 1:50). Secondary
antibodies used were goat anti-rabbit Alexa 488 and goat anti-mouse Alexa 568 (Invitrogen).
77
Slides were coverslipped with Vectashield mounting media with DAPI and imaged at 10x using
a Keyence BZ-X710 microscope.
ACKNOWLEDGEMENTS
This work was supported by the National Institutes of Health [R01DE025222 to A.E.M,
5T32HD060549-02 to J.E.S. and R01DE025222-01S1 and T90DE021982 to R.R.R] and March
of Dimes [#6-FY15-233 to A.E.M.]. We thank Cynthia Neben, Creighton Tuzon, Diana Rigueur,
Lauren Bobzin and Francesca Mariani for their helpful discussions.
78
Figure 3.1. RCAS injections into the LPM of the hindlimb largely targets dense connective
tissues. (A) In chick embryos at HH15, the LPM was injected with RCAS-GFP at the level of the
future hindlimb (dotted line). (B) RCAS-GFP injections collected at HH29 show widespread
expression throughout the proximal-distal axis of the limb. Black dashed lines denote region of
sections for panels C-G. Sections through the autopod (C), zeugopod (D) and stylopod (E) of the
hindlimb showed GFP
+
cells primarily found within Tcf4
+
(red) connective tissue in the muscle
(arrows) and Tcf4
-
connective tissues (tendon) between the developing muscle and (arrowhead).
A few GFP
+
cells were also detected in the cartilage (C,G; asterisk). In the zeugopod (F) and
stylopod (G) of the hindlimb, GFP
+
cells were largely interstitial to myosin
+
muscle cells (red).
79
Figure 3.2. Expression of the BBDS mutations in the LPM of the chick hindlimb induces
hindlimb abnormalities. Whole mount analysis of HH29 embryos following HH15 LPM
injections of RCAS-GFP (n=16) (A-C) and RCAS-FGFR2
WT
(n=11) (D-F) resulted in normal
hindlimbs. RCAS expression of BBDS mutations FGFR2
M391R
(n=15) (G-I) and FGFR2
Y381D
(n=17) (J-L) induced angulated hindlimbs (G,J), splayed limbs (H,K) anterior-posterior
spreading of the foot plate with pronounced deflection of digit 1 (I,L). Asterisks denote region of
abnormal angulation.
80
Figure 3.3. Expression of the BBDS mutations in the hindlimb connective tissue causes bent
long bones and knee defects. Chick embryos collected at HH36 and stained with Alcian blue
(cartilage) and Alizarin red (bone) showed that RCAS-GFP expression (A-A’) resulted in normal
long bone development. Expression of FGFR2
M391R
(C,C’) and FGFR2
Y381D
(E,E’) caused mild
bending of the femur and tibiotarsus (arrows), irregular epiphyseal morphology at the knee, and
dislocation of the fibula. (B) The femoral bone collar length and knee joint angle were
significantly smaller in FGFR2
M391R
and FGFR2
Y381D
embryos compared to GFP controls. In
more severely affected limbs expressing FGFR2
M391R
(D,D’) and FGFR2
Y381D
(F,F’), the
tibiotarsus was missing and the fibulare, a portion of the fibula that contributes to the epiphysis
of the tibiotarsus, was retained and irregularly, posteriorly oriented (arrowhead). Morphology of
the knee in the severe cases (D’,F’) showed irregular articulation of the fibula to the femur with
changes in epiphyseal morphology of both bones. The foot was removed for clarity. F=femur,
Fb=fibula, T=tibiotarsus. Error bars represent standard error of the mean, n=8 per condition,
*p<0.05, ** p<0.01, *** p<0.001.
81
Figure 3.4. The BBDS mutations induce defects in the autopod skeleton and pelvic girdle.
Whole mount skeletal preps stained with Alizarin red and Alcian blue at HH36. (A) RCAS-GFP
expressing limb displays normal patterning of the autopod with four digits. (B) In a FGFR2
M391R
expressing limb, digits 1 and 2 are missing in the autopod. (C) In a FGFR2
Y381D
expressing limb,
digit 1 is missing. Numbers indicate digit identity. (D) RCAS-GFP expressing limb exhibits
normal pelvic girdle. (E,F) The ilia of FGFR2
M391R
and FGFR2
Y381D
expressing embryos are
small and have irregular morphology (brackets). The pubis is bowed in FGFR2
M391R
embryos
and hypoplastic in FGFR2
Y381D
embryos (arrowheads). Representative images were selected
(n≥4).
82
Figure 3.5. Musculoskeletal integration is disrupted by expression of the BBDS mutations.
Histological sections of HH36 embryos were stained using HBQ stain to mark cartilage (blue),
bone (pink) and muscle (purple). (A-D) Knee morphology was altered in RCAS-FGFR2
M391R
and
RCAS-FGFR2
Y381D
embryos compared to controls. The tibiotarsus epiphysis exhibited loss of
typical V-shaped condyles, resulting in a flatter articular surface (B,D; asterisk). The anterior
muscle of the tibiotarsus had a broader attachment on the patella in the mutants compared to the
controls (B,D; arrow). The femur and tibiotarsus were bent in FGFR2
M391R
and
FGFR2
Y381D
expressing embryos with asymmetric mineralization at the point of the bend with
more posterior mineralization than anterior (F,H,J,L; arrowhead). Disorganized flexor muscle
patterning with poor attachment was visible throughout the hindlimb (B,D,F,H,J,L; brackets).
F=femur, P=patella, T=tibiotarsus
83
Figure 3.6. Nuclear localization is enhanced by BBDS mutations and distinct localization
signals in ATDC5 cells. ATDC5 prechondrocytes stained with FLAG (red), DAPI (blue) and
Polr1D (green) showed localization of FLAG-tagged FGFR2 constructs. (A-A”) FGFR2
WT
exhibited membrane and nuclear localization in transfected ATDC5 cells. (B-C”) BBDS mutant
FGFR2 receptors FGFR2
M391R
and FGFR2
Y381D
exhibited increased nuclear localization
compared to wildtype receptor. (D-D”) -FGFR2 distinctly enhanced nuclear localization of the
receptor. (E-E”) NoLS-FGFR2 increased localization to the nucleolus and perinucleolar space.
Magnification 90x.
84
Figure 3.7. Increased nuclear localization of FGF phenocopies BBDS. (A-C) Control RCAS-
GFP embryos (n=15) exhibited straight hindlimbs, proper integration at the hip and regular digit
patterning at HH29. (D-I) NLS-FGFR2 (n=16) and NoLS-FGFR2
(n=15) exhibited bending of
the femur, splayed limbs, anterior-posterior spreading of the foot plate with pronounced
deflection of digit 1, and in some cases split-hand malformation. Asterisk denotes bending.
85
Figure 3.8. Changes in skeletal morphology are induced by increased nuclear and nucleolar
localization. Skeletal staining of bone and cartilage with Alizarin red and Alcian blue,
respectively, was performed at HH36. Expression of NLS-FGFR2 and NoLS-FGFR2 induced
bending of the femur and tibiotarsus (C,E; arrows), reduced bone collar length and decreased
angle of the knee (B) and irregular joint morphology (C’,E’) compared to control (A,A’).
Severely affected NLS-FGFR2 and NoLS-FGFR2 embryos exhibited bent femurs and absent
tibotarsi with retention of the fibulare (D,F; arrowhead). Irregular epiphyseal and joint
morphology were visible at the articulation of the femur with the fibula (D’,F’). The foot has
been removed for clarity. F=femur, Fb=fibula, T=tibiotarsus. Error bars represent standard error
of the mean, n≥6 per condition, * p<0.05, ** p<0.01.
86
Figure 3.9. Irregular muscle patterning is detected in embryos expressing nuclear and
nucleolar FGFR2. Histological sections of HH36 chicken embryos were stained using HBQ
stain for cartilage (blue), bone (pink) and muscle (purple). (A-D) The knee joint showed altered
morphology in NLS-FGFR2 and NoLS-FGFR2
embryos with broadened and flattened epiphyses
of the tibiotarsus (asterisk). Muscles displayed disorganization in the posterior of the knee (B,D;
brackets). Nuclear FGFR2 signaling induced bending of the femur and tibiotarsus (E-L; black
arrowhead). Asymmetric mineralization was observed in NLS-FGFR2 expressing embryos (F,J),
while NoLS-FGFR2 embryos were milder with small bone collars (H,L). Flexor muscles along
the posterior of the femur and tibiotarsus were poorly attached and disorganized, particularly in
regions adjacent to bent bones (B,D,F,H, brackets). F=femur, P=patella, T=tibiotarsus
87
CHAPTER 4: Conclusions
In conclusion, FGFR2 plays a critical role in skeletal development by influencing connective
tissues as well as the bone. Little is known about the normal function of FGFR2 in the
connective tissue, but what has been uncovered supports a role for FGFR2 in connective tissue
development. FGFR2 is expressed within intramuscular connective tissue fibroblasts and in
differentiating fibro/adipocyte progenitors, indicating that it is expressed within connective tissue
cells and is involved in the differentiation process of these cells [313-315]. Loss of Fgfr2 in
mouse bladder mesenchyme results in a thickened layer of bladder connective tissue, increased
fibrosis and smaller, weakened muscles [316]. In studying the FGFR2
S252W
Apert syndrome
mutation, researchers found that fibroblasts exhibited greater increases in proliferation,
differentiation and migration in response to the mutation than both control cells and
mesenchymal stem cells expressing FGFR2
S252W
[317]. Both activation and loss of FGFR2 result
in augmented connective tissue, indicating that FGFR2 may tightly regulate connective tissue
behavior with too much or too little FGFR2 disrupting the process. Additionally, fibroblasts
exhibited higher sensitivity to changes in FGFR2 activity than mesenchymal stem cells, implying
a normal role for FGFR2 in the connective tissue that becomes augmented by the Apert
mutation. Overexpression of FGF5 in the chick limb increased connective tissue proliferation
and subsequently altered muscle patterning and development [71]. These chick limbs exhibited
thickened periostea and bowed bones like patients with BBDS and our chick model, suggesting
that FGFR2 in the connective tissue may be the receptor activated by the ectopic FGF5 [31, 71].
Our work is novel because we demonstrate that expressing nuclear FGFR2 in the limb
connective tissue mesenchyme is sufficient to induce bowed long bones. We show in our system
88
that connective tissue influences the structure and patterning of muscle, tendon, bone, and joint
during limb development. Using the chick embryonic system, we created a model for studying
BBDS that allows us to examine the role of connective tissue defects in bent bone disorders.
Additionally, we reveal that BBDS is caused by increased nuclear localization of FGFR2. This
work provides insight into bent bone and FGFR2 disorders as well as furthers our understanding
of the role of FGFR2 in limb development.
89
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Abstract (if available)
Abstract
Background: Bent Bone Dysplasia Syndrome (BBDS), a congenital skeletal disorder caused by dominant mutations in Fibroblast growth factor receptor 2 (FGFR2), is characterized by bowed long bones within the limbs. We previously showed that the FGFR2 mutations in BBDS enhance nuclear and nucleolar localization of the receptor
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Salva, Joanna Elizabeth
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Nuclear fibroblast growth factor receptor 2 regulates skeletal development and joint formation
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Keck School of Medicine
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Doctor of Philosophy
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Genetic, Molecular and Cellular Biology
Publication Date
10/03/2019
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08/14/2018
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bent bone dysplasia syndrome,bone,Development,FGFR2,fibroblast growth factor signaling,joint,muscle,OAI-PMH Harvest
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bent bone dysplasia syndrome
FGFR2
fibroblast growth factor signaling
joint
muscle