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FGFR2 Regulates Connective Tissue Development in the Skull

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FGFR2 Regulates Connective Tissue Development in the Skull

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FGFR2 Regulates Connective Tissue Development in the Skull

by

Lauren Grace Bobzin

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

December 2023

Copyright 2023 Lauren Grace Bobzin

ii
Epigraph

“For a research worker, the unforgotten moments of their life are those rare ones which
come after years of plodding work, when the veil over nature’s secret seems suddenly
to lift and when what was dark and chaotic appears in a clear and beautiful light and
pattern.”- Gerty Cori

iii
Acknowledgements
Dissertation Committee members: Dr. Francesca Mariani, Chair; Dr. Robert Maxson,
Co-Chair; Dr. Jian Xu, Member; Dr. Amy Merrill, Mentor

I would like to thank my family and friends for supporting me through this difficult
journey. Through the rants, crying, and threatening to quit you have never wavered in
your support and confidence in me. Thank you to my parents and sister for loving me
unconditionally and always making sure I was safe and provided for. Thank you to my
best friends for the commiseration, memes, and always taking my calls. Thank you to
my therapist for giving me the strength and skills I needed to support and love myself.
Thank you to Dr. Merrill and all the members of my lab for helping me do science I am
proud of. We did it!

iv
TABLE OF CONTENTS
Epigraph ........................................................................................................................... ii
Acknowledgements ......................................................................................................... iii
List of Figures ................................................................................................................... v
Abbreviations .................................................................................................................. vii
Chapter 1: FGF signaling regulates development within the anterior fontanelle ............. 1
Chapter 2: Development and maintenance of tendons and ligaments .......................... 42
References ..................................................................................................................... 72

v
List of Figures
Chapter 1: FGF Signaling Regulates Development Within the Anterior Fontanelle
Figure 1: NCC;Fgfr2
-/-
mutant mice show a widening and patency of the
anterior fontanelle starting at E18.5. ....................................................................... 27
Figure 2: NCC;Fgfr2
-/-
mutants show defects in frontal bone approximation
and lack endocranial cartilage that prefigures the frontal suture. ........................... 28
Figure 3: Anterior fontanelle connective tissue expresses canonical
tendon/ligament marker Scx. .................................................................................. 29
Figure 4: Anterior fontanelle connective tissue is composed of heterogeneous
cell populations with distinct gene expression patterns and spatial organization.
................................................................................................................................. 30
Figure 5: Both Scx and Sox9 expressing cells contribute to the fused frontal
suture. ...................................................................................................................... 31
Figure 6: EdU pulse chase experiments show a minor decrease in proliferation
of NCC;Fgfr2
-/-
mutant mice vs littermate controls. .................................................. 32
Figure 7: NCC;Fgfr2
-/-
mice lose expression of cartilage and osteoblast specific
markers in midsuture FCT of in favor of maintaining Scx expressing connective
tissue. ..................................................................................................................... 33
Figure 8: RNA-sequencing of AF tissue reveals differential gene enrichment
between control and NCC;Fgfr2
-/-
samples. ........................................................... 34
Figure 9: Key WNT pathway members are dysregulated in NCC;Fgfr2
-/-
mice,
suggesting increased signaling ............................................................................... 35
Figure 10: Ectopic activation of Wnt/𝛽- catenin signaling in Scx expressing FCT
phenocopies NCC;Fgfr2
-/-
mutant phenotype ......................................................... 36
Figure 11: The connective tissue recruitment model of frontal suture
development ............................................................................................................ 37
Supplemental Figure 1: Mice of Scx-Cre;Fgfr2
-/-
and Sox9-Cre;Fgfr2
-/-
genotypes
are indistinguishable from littermate controls at P5 and P7. ................................... 38
Supplemental Figure 2: A subcluster of CT2 cells have high Fgfr2 but low Scx
and Sox9 expression. ............................................................................................. 39
Supplemental Figure 3: Cleaved caspase staining shows no significant levels of
apoptosis in control or NCC;Fgfr2
-/-
littermate mice. ............................................... 40
vi
Chapter 2: Development and Maintenance of Tendons and Ligaments
Figure 1: Tissue morphology of tendon, ligament, and musculoskeletal
junctions of the human elbow. ................................................................................ 67
Figure 2: Embryonic origins and molecular regional regulators of tendon and
ligament progenitors in the developing mouse. ...................................................... 68
Figure 3: Tendon progenitor induction, differentiation, maintenance, and
maturation in the mouse limb. ................................................................................. 69
Figure 4: Regulation of cell fate during development of the tendon-bone
attachment in the mouse limb. ................................................................................ 70
Figure 5: Cellular lineages contributing to tendon and enthesis repair in mice. ...... 71

vii
Abbreviations
Anterior fontanelle ........................................................................................ AF
Attachment progenitors ................................................................................ APs
Cranial neural crest cells .............................................................................. CNCCs
Collagen ...................................................................................................... COL
Extracellular matrix ....................................................................................... ECM
Early migrating mesenchyme ...................................................................... EMM
Enthesis stem and progenitor cells .............................................................. ESPCs
Fibroblast growth factor ................................................................................ FGF
Fibroblast growth factor receptor 2 .............................................................. FGFR2
Gain of function ........................................................................................... GOF
Hall-Brunt Quadruple ................................................................................... HBQ
Hedgehog ..................................................................................................... Hh
Heterotopic ossification ................................................................................ HO
Indian Hedgehog .......................................................................................... IHH
Myotendinous junction .................................................................................. MTJ
Neural crest Cre- ......................................................................................... NCC
Posterior frontal suture ................................................................................. PFS
Progressive osseous heteroplasia ............................................................... POH
Scleraxis ....................................................................................................... SCX
Sonic Hedgehog ........................................................................................... SHH
Supraorbital mesenchyme ............................................................................ SOM
Transforming growth factor beta .................................................................. TGFβ
Tenomodulin ................................................................................................. TNMD
Tendon stem and progenitor cells ................................................................ TPSCs

1
Chapter 1: FGF Signaling Regulates Development Within the
Anterior Fontanelle

Introduction
The calvarial bones of the infant skull are linked by transient fibrous joints called
sutures and fontanelles. Sutures, which join two adjacent calvarial bones, and fontanelles,
which occupy large unossified regions between multiple bones where sutures have yet to
form, are critical for calvarial reshaping during birth and major sites of calvarial growth.
Numerous genetic disorders present with craniofacial deformities when sutures fuse
prematurely (craniosynostosis), or fontanelles remain patent. For example, a spectrum of
gain-of-function variants in Fibroblast growth factor receptor 2 (FGFR2) cause syndromes
including Apert, Pfeiffer, Crouzon, and Bent bone dysplasia which present with multi-
suture craniosynostosis, as well as a persistently open anterior fontanelle (AF) [1-4]. The
co-occurrence of these seemingly opposite calvarial joint phenotypes suggests that the
molecular pathways through which FGF signaling regulates development of the suture vs
the anterior fontanelles are distinct.
The AF is a broad area of connective tissue that forms at the apex of the embryonic
calvaria where the two frontal and two parietal bones meet. In postnatal development, the
AF gives way to the apically advancing intramembranous bone fronts and, ultimately, is
replaced by the posterior frontal suture (PFS) that joins the paired frontal bones from the
bregma to jugum limitans. Once the PFS forms, it undergoes a normal process of fusion
through endochondral-like ossification that requires Sox9 and regional inhibition of WNT
signaling [5-8]. The ultimate fate of the AF connective tissue and the extent to which it
plays an active role in frontal suture development, as well as normal and abnormal fusion,
has remained unclear.
2
While FGFR2-related syndromes strongly suggest a role for FGF signaling in the
AF, studies to date have largely focused on the receptor’s role in the context of the
sutures, where it is expressed within the mid-suture mesenchyme and osteoprogenitors
leading the advancing bone fronts. Genetic studies in mice have shown that FGFR2
signaling, activated primarily by FGF18, is critical to maintain the balance of proliferation
and osteoblast differentiation within the sutures. In mice harboring disease-causing
variants of Fgfr2 that are associated with a gain-of-function (GOF), craniosynostosis of
the coronal suture occurs through a mechanism that either decreases or increases
proliferation depending on the variant, while also promoting ectopic osteoblast
differentiation [9-11]. Patency of the AF in an Apert mouse model, on the other hand, is
linked to increased proliferation and decreased osteoblast differentiation of the mid-suture
mesenchyme [10]. In Fgf18-deficient mice, midline ossification defects in the calvaria
arise due to decreased proliferation and delayed osteoblast differentiation [12]. While a
midline ossification defect has also been reported in mice with conditional loss of Fgfr2 in
skeletogenic mesenchyme using Twist2-Cre, the mechanism of the phenotype was not
investigated [13].
Here, we investigate the necessity of Fgfr2 in the development of the AF and PFS.
We find that Fgfr2 is necessary for closure of the AF and subsequently, PFS formation
and fusion. We show that the AF is occupied by a heterogeneous population of Scx+
connective tissue-like progenitors, and that loss of FGFR2 signaling blocks their
differentiation into bone, cartilage, and ligament-like connective tissue, preventing normal
PFS fusion that occurs through endochondral-like ossification. Gene expression analysis
indicates that Fgfr2 regulates the differentiation of Scx+ cells into osteoblasts and the
3
formation of Sox9+ chondrocytes through downregulation of WNT/b-CATENIN signaling.
Correspondingly, genetic elevation of WNT/b-CATENIN signaling in Scx+ cells in mice
recapitulates the patent AF phenotype seen in Fgfr2 mutant mice. Together, these results
suggest a key role for Fgfr2 in orchestrating divergent cell fate decisions of AF connective
tissue to promote formation and closure of the PFS.
Results
Conditional loss of Fgfr2 in the developing AF blocks PFS formation.
Genetic lineage analyses using Wnt1-Cre has shown that neural crest-derived
mesenchyme gives rise to the frontal bones, as well as the mesenchyme that occupies
the AF in mice [14, 15] (Fig. 1A). To investigate the role of Fgfr2 within the AF and PFS,
we therefore performed Wnt1-Cre-mediated deletion of Fgfr2 using a floxed allele
previously shown to induce a functional null allele. Because of the ablation of Fgfr2 in the
neural crest cells and their derivatives, we will refer to these mice as Neural Crest Cre
mice or NCC; Fgfr2
-/-
mice [13]. Calvarial bone and cartilage development was examined
in NCC;Fgfr2
-/-
and littermate controls using wholemount alizarin red and alcian blue
staining. The AF of NCC;Fgfr2
-/-
mice is wide and patent beginning at embryonic day (E)
18.5 compared to control (Fig. 1B,C). In control mice at postnatal day (P) 3, the AF begins
to progressively close, while that of NCC;Fgfr2
-/-
mice remains wide open (Fig. 1D,E). At
P5, the frontal bones of control mice have closely approximated to form the PFS, while
AF remains persistent in the mutant (Fig. 1F,G). Quantitative measurement showed that
the normalized average area between the frontal bones (total area/ length of PFS from
jugum limitans to bregma) is significantly larger in NCC;Fgfr2
-/-
at P3 (p=.007, n = 7) and
P5 (p=.005, n=9) compared to littermate controls (Fig. 1N). By P10, the PFS of control
4
mice begins to show cartilaginous fusions that follow a joint-gap pattern, while the PFS
has still failed to form in NCC;Fgfr2
-/-
mice (Fig. 1H,I). MicroCT analysis shows that the
PFS of control mice has fused by P30, with distinct endocranial and ectocranial bony
layers visible in orthoslice views (Fig. 1J,K). Meanwhile, in NCC;Fgfr2
-/-
mice, the AF
persists and regions of the frontal and sagittal sutures it prefigures do not form (Fig.
1L,M).
To identify histogenic changes in the AF and PFS of NCC;Fgfr2
-/-
mice, skeletal
and connective tissues were discriminated using Hall-Brunt Quadruple stain (HBQ), which
utilizes direct red and alcian blue to identify bone collagen and cartilage proteoglycans,
respectively [16]. In control calvaria at P5, the paired frontal bones are spanned by
connective tissue and the ectocranial and endocranial layers are discernible. The frontal
bones of NCC;Fgfr2
-/-
mice, however, are comparatively further apart than in control
animals and the endocranial bone layer is missing (Fig. 2A,B). At P7, the cartilage
template that prefigures the fused endocranial layer of the PFS begins to condense and
becomes more pronounced at P11 and P13 (Fig. 2C,E,G). Conversely, the frontal bones
of NCC;Fgfr2
-/-
mice remain separated and show no evidence of cartilage (Fig. 2D,F,H).
All together these results show that Fgfr2 is necessary for closure of the AF, as well as
formation and fusion of the PFS.
Anterior fontanelle connective tissue expresses Scx and is composed of
heterogeneous cell populations
Very little is known about the AF connective tissue and its role in frontal suture
formation. Since ligament is a connective tissue that often links bone to bones, we
examined expression of Scx, a transcription factor that promotes ligament cell fate,
5
utilizing the Scx-GFP reporter line [17]. At E18.5, Scx-GFP was expressed in all calvarial
sutures and fontanelles, as well as immediately adjacent bone (Fig. 3A). At P3, Scx-GFP
expression is more confined to the sutures themselves (Fig. 3B). Sections through the AF
at these stages reveals a bi-layered expressional pattern of Scx-GFP representing the
ecto- and endocranial layers of the suture mesenchyme. Some GFP positive cells were
also identified in the osteogenic fronts of the frontal bones, seen more clearly at P3 (Fig.
3C,D). Expression of Scx-GFP in the AF suggests that connective tissue in the AF exhibits
ligament-like characteristics.
To better understand Scx + cell populations in the AF, we analyzed a previously
published single cell RNA sequencing (scRNA-seq) dataset of the developing PFS at
E18.5 (FaceBase data set 1-4TT6) [18]. This dataset was derived from a region of AF
that will give rise to the PFS, including the paired osteogenic fronts of the frontal bones
and the intervening connective tissue, but not skin or brain. We re-analyzed this dataset
using Seurat and identified 8 mesenchymal clusters similar to those previously reported,
including those that represent the hypodermis, dura, hypodermis, preosteoblasts, and
osteoblasts (Fig. 4A). Interestingly, three of the clusters (CT1-3) show relatively enriched
expression for Scx (Fig. 4B). Gene enrichment analysis of Scx+ clusters indicate they are
uniquely marked by enriched expression of Fgf18 (CT1), Sp7 (CT2) and Sox9 (CT3). CT2
and CT3, as well as preosteoblasts have enriched expression of Fgfr2 (Fig. 4B). To
spatially map CT1, CT2, and CT3 in the AF at E18.5, we used RNAScope fluorescent in
situ hybridization. Scx is expressed throughout the AF with highest expression in the
ectocranial AF connective tissue (CT1), while Fgfr2 expression is highest in cells just
apical to the bone fronts (CT2) and endocranial AF connective tissue (CT3) (Fig. 4C).
6
Fgf18 is primarily restricted to the ectocranial region of the AF connective tissue (CT1)
that overlays the frontal bones (Fig. 4D), while Sp7 marks cells that lead the advancing
osteogenic fronts (CT2), but is excluded from the midsuture (Fig. 4D,E). Finally, Sox9
expression marks the endocranial layer of AF connective tissue (CT3) (Fig. 4E). This is
consistent with a previous study showing that Sox9 is indispensable for closure of the
PFS through endochondral ossification [8].
During normal PFS closure, cells at the osteogenic fronts undergo
intramembranous ossification, while the endocranial layer undergoes endochondral-like
ossification. Pseudotime analysis performed with Monocle predicts that the CT2 and CT3
populations followed distinct developmental trajectories that converge on an osteogenic
identity (Fig. 4F). Together, this data suggests that transcriptionally distinct populations
of Scx+ cells occupy specific domains within the AF connective tissue and have the
potential to give rise to osteogenic lineages in the PFS.
Scx-lineage cells contribute to PFS fusion and are non-autonomously regulated by
Fgfr2
To determine the fate of Scx+ cells during AF closure and PFS formation, we
traced Scx-lineage cells in Scx-Cre;Ai9 mice collected at P4 and P7. At P4, Scx -lineage
cells occupy the osteogenic fronts of the paired frontal bones and AF connective tissue,
with relative enrichment to the endocranial layer and a thin layer of cells overlaying the
entire suture (Fig. 5A). By P7, the contribution of Scx-lineage cells in both the ecto- and
endocranial layers has increased, where they occupy the mid-suture and osteogenic
fronts of the frontal bones (Fig. 5B). The scRNA-seq results suggest that a population of
Scx+ cells activate expression of Sox9 and subsequently undergo endochondral-like
7
ossification during PFS fusion. To determine the contribution of Sox9-lineage cells to
PFS, we performed lineage tracing in Sox9-CreERT2;Ai9 mice induced with tamoxifen at
P2 and P3 and then collected at either P4 or P7. Sox9-lineage cells contribute to the
leading edges of the frontal bones as well as the condensing cartilage in the endocranial
domain of the PFS (Fig. 5A,B). These result support the idea that Scx+ populations within
the AF connective tissue make direct and region-specific contributions to the PFS.
To test the extent to which Fgfr2 is required in Scx+ and Sox9+ cells during AF
closure and PFS formation, we examined Scx-Cre;Fgfr2
-/-
and Sox9-CreERT2;Fgfr2
-/-
mice. Wholemount skeletal staining of Scx-Cre;Fgfr2
-/-
mice at P5 and P7 showed that
the calvaria bones were indistinguishable from littermate controls (Supplemental Fig. 1A-
D). This suggests that loss of Fgfr2 in Scx+ cells is not sufficient to recapitulate the AF
and PFS phenotypes described in NCC;Fgfr2
-/-
mice. We next examined Sox9-CreERT;
Fgfr2
-/-
mice, inducing recombination by tamoxifen administration in timed pregnant
females at E17.5 and 18.5. Wholemount skeletal staining at P5 revealed that mutants
were indistinguishable from littermate controls (Supp. 1E,F). The same result was found
when Sox9-CreERT;Fgfr2
-/-
mice were induced at P0 and collected at P7 (Supp. 1G,H).
This suggests that loss of Fgfr2 in Sox9+ cells is not sufficient to recapitulate the AF and
PFS phenotypes described in NCC;Fgfr2
-/-
mice. Higher resolution clustering of the
scRNA-seq dataset shows that there is a subpopulation of cells within CT2 that are
enriched for expression of Fgfr2 and show little to no expression of Scx and Sox9 (Supp.
2). This indicates that these cells are unlikely to be targeted by Scx-Cre or Sox9-
CreERT2, and that Fgfr2 non-autonomously regulates Scx+ and Sox9+ cells in the
AF/PFS.
8
Differentiation of Scx cells is disrupted in the AF of NCC;Fgfr2
-/-
mice
Fgfr2 has a well-established role in regulating proliferation and differentiation in the
developing sutures of mice. We next investigated how loss of Fgfr2 impacts proliferation
during AF closure using EdU pulse-chase experiments. At E18.5, no statistically
significant differences were identified in the percentage of proliferating cells within the AF
connective tissue or osteogenic fronts of the paired frontal bones (p>.02) (Fig. 6A,B,C).
When these regions were combined, a statistically significant difference in proliferation of
less than 1% is seen between control and NCC;Fgfr2
-/-
mice (6% in total control vs. 5.1%
in total mutant, p=.02, n=7) (Fig. 6C). At P3, EdU pulse-chase shows a significant
difference in proliferation of 2.3% in NCC;Fgfr2
-/-
mice ( 9.6% in total controls vs. 7.3% in
total mutant, p=.004, n=6) (Fig. 6D,E,F). Interestingly, this difference can be accounted
for by a decrease in the proliferative rate of the NCC;Fgfr2
-/-
AF connective tissue cells
(6.8% in control vs. 4.9% in mutants, p=.001, n=6). Proliferation rates in the bone fronts
were not statistically significantly different between mutant and control (Fig. 6F). Together
this indicates that while proliferation within bone front cells appeared normal, a small
decrease in the proliferation rate of AF connective tissue could contribute to AF patency
in NCC;Fgfr2
-/-
mice.
We next examined osteogenic differentiation within the AF of NCC;Fgfr2
-/-
mice
using immunofluorescent detection of RUNX2, an early marker of osteoblasts, along with
Scx-GFP to discriminate the AF connective tissue. At P0, RUNX2+ cells are enriched
within the osteogenic fronts of the paired frontal bones, while Scx-GFP expression is
localized to cells in the ecto- and endocranial layers of the AF in both control and
NCC;Fgfr2
-/-
mice (Fig. 7A,B). In control mice at P3, RUNX2 expression is activated in
9
Scx+ cells located in the endocranial layer of the AF connective tissue, and
Scx+/RUNX2+ cells are found in the advancing bones (Fig. 7C). Scx+/RUNX2+ double
positive cells in the endocranial layer fail to form and remain Scx+ within the AF of
NCC;Fgfr2
-/-
mice (Fig. 7D). An ectocranial layer of Scx+ cells spanning and overlaying
the frontal bone remains similar between control and mutant (Fig. 7C-D). In controls at
P5, RUNX2+ cells coalesce at the approximating endocranial osteogenic fronts to form
the PFS, and Scx+/RUNX2+ cells show contribution to the approximating bone fronts.
Meanwhile, the ectocranial layer in control mice is still occupied by Scx+ connective
tissue. In NCC;Fgfr2
-/-
mice, on the other hand, the AF remains patent and very few
Scx+/RUNX2+ are detected in the osteogenic fronts and bone (Fig. 7E,F).
To detect chondrogenic differentiation, we performed immunofluorescent detection
of SOX9, an early marker for chondrocytes. While SOX9 is not detected in the PFS at P5
in control or NCC;Fgfr2
-/-
mice (Fig. 7G,H), a population of SOX9+ cells appears in the
endocranial region of the suture midline of controls by P7 (Fig. 7I). In NCC;Fgfr2
-/-
mice,
however, SOX9 expression remains undetectable, and instead the persistent AF is
occupied by Scx+ cells (Fig. 7J). We then investigated whether the loss of SOX9 could
be accounted for by increased instance of cell death in the NCC;Fgfr2
-/-
mice.
Immunofluorescence for cleaved-CASPASE showed no cell death in controls or
NCC;Fgfr2
-/-
mutants at P0, P3, or P5. (Supp. 3). All together these results strongly
indicate that the Scx+ cells of the AF connective tissue undergo differentiation into
skeletogenic cells to form the PFS and that the persistent AF in NCC;Fgfr2
-/-
mice is
largely caused by failed skeletogenic differentiation of Scx+ AF cells.

10
Expression of WNT pathway members is altered in the AF of NCC;Fgfr2
-/-
mice
To determine the molecular mechanisms through which Fgfr2 regulates
differentiation within the AF, we performed bulk RNA-seq of the control and NCC;Fgfr2
-/-
AF at E18.5, when a phenotypic difference is first detected. Heat map analysis of the
differentially expressed genes showed that RNA expression profiles of control and
NCC;Fgfr2
-/-
littermates have significant, genotype specific differences (Fig. 8A). GO
analysis of the genes indicates differential expression of members of the WNT signaling
pathway, as well as genes related to focal adhesion, extracellular matrix (ECM) -receptor
interaction, and axon guidance (Fig. 8B). Previous studies have identified a role for WNT
signaling in the developing suture, where this pathway works cooperatively with FGF
signaling to determine skeletogenic cell fate [19-21]. Close examination of WNT pathway
members showed that inhibitors Sfrp4, Notum, and Wif1, along with the WNT potentiator
Lgr6 are downregulated in the NCC;Fgfr2
-/-
AF (Fig.8C). On the other hand, the WNT/b-
CATENIN target gene and WNT signaling potentiator Lgr5 and the context dependent
WNT modulator Sfrp2 are upregulated in the NCC;Fgfr2
-/-
AF (Fig. 8C). String analysis of
the WNT pathway genes found in our analysis coupled with Fgfr2 shows interactions
between these pathways is likely facilitated by interactions between Fgfr2 and Wnt2 (Fig.
8D). RNA-seq results for several WNT pathway members were validated using in situ
hybridization with RNAScope. As suggested by RNA-seq, the NCC;Fgfr2
-/-
AF show
decreased Wif1 expression particularly within the frontal bones and increased Lgr5
expression in the AF connective tissue compared to the control AF (Fig. 9A,B). Regulation
of Wif1 expression by Fgfr2 is likely regionalized to a subset of CT2 cells which co-
express the two genes (Fig. 9C). These results suggest an overall increase in WNT
11
signaling in NCC;Fgfr2
-/-
mice precedes failed differentiation of Scx+ AF cells into bone
and cartilage.
Ectopic activation of WNT/b-CATENIN signaling in Scx cells blocks AF closure and
PFS formation
To test the impact of increased WNT/b-CATENIN signaling in Scx+ cells of the AF,
we crossed mice harboring the Scx-Cre driver to mice with a stabilizing mutation in exon
3 in 𝛽-catenin (Catnb
lox(Ex3)
) [22]. Stabilization of the β-CATENIN protein blocks its
degradation through the destruction complex and mimics conditions of constitutively
active WNT signaling. Wholemount skeletal staining of Scx-Cre;Catnb
lox(Ex3)
(Scx-
Cre;Catnb
GOF
) mice with alizarin red and alcian blue show that the AF is larger compared
to littermate controls (Fig. 10A,B). At P5, while the frontal bones of control mice
approximate to form the PFS, the AF remains patent and the PFS fails to form in Scx-
Cre;Catnb
GOF
mice (Fig. 10C,D). In addition, mineralization of the calvarial bones in Scx-
Cre;Catnb
GOF
mice was abnormal, with bony spicules forming at the frontal bone margins
(Fig. 10B,D). The observation that increased WNT/b-CATENIN signaling in Scx+ cells
recapitulates the phenotype of NCC;Fgfr2
-/-
mice supports a mechanistic connection
between FGF and WNT signaling during AF closure and PFS formation.
Discussion
The results presented in this study characterize the nature of neural crest derived,
AF connective tissue and its contribution to the formation of the posterior frontal suture.
By conditionally deleting Fgfr2 in NCC-derived tissue, we have demonstrated its necessity
in regulating the closure of the AF, as well as formation and fusion of the PFS. We show
that the AF connective tissue is made up of heterogeneous Scx+ cells which are
12
regionally organized into distinct sub-populations and have differing contributions to
skeletogenic cell types in the developing PFS. In the absence of FGFR2 signaling,
WNT/b-CATENIN signaling is increased, and Scx+ cells fail to differentiate into
osteoblasts and chondrocytes that make the PFS joint (Figure 11). Our results reveal that
paracrine FGF signaling between the approximating bones and AF connective tissue
coordinates differentiation of divergent skeletogenic cell fates during PFS formation and
fusions by modulating the WNT/b-CATENIN pathway.
This study shows for the first time that Scx+ cells within the AF connective tissue
directly contribute to joint cells within PFS joint, suggesting that sutures that form within
a fontanelle during postnatal development are distinct. We previously identified Scx+ cells
in the ectocranial region of the non-fusing murine coronal suture which go on to form
ligament-like tissue that is maintained into adulthood, [23]. Scx+ cells in the AF appear to
be distinct in from those in the coronal in several important ways. Our single cell analysis
of the AF at E18.5 identified three Scx+ populations that express varying levels of Fgfr2
and can be distinguished by differential expression of Fgf18 (CT1), Sp7 (CT2), and Sox9
(CT3) and Wnt pathway members. Through high resolution mapping and genetic lineage
tracing, we show that these Scx+ populations occupy distinct domains within the AF and
give rise to distinct lineages within the PFS including osteoblasts, chondrocytes, and
ligament-like connective tissue.
We also find that within these Scx+ cells, differential expression of FGF and WNT
pathway members establish a signaling center necessary for AF closure and PFS joint
formation. In the AF, ectocranial Scx+ cells (CT1) express Fgf18 and low levels of Fgfr2,
suggesting these cells have low autocrine FGF signaling. FGF18, a ligand for FGFR2,
13
promotes proliferation and osteogenesis, but inhibits chondrogenesis in calvarial
mesenchyme [8, 12, 24]. Low levels of FGF signaling in the ectocranial domain is
therefore consistent with our finding that ectocranial Scx+ cells contribute to ligament-like
tissue of the PFS, and likely do not make major contributions to the advancing fronts of
the frontal bone. Instead, FGF18 produced by CT1 acts as paracrine signal on Scx+ cells
located apically to the bone fronts (CT2) to promote osteogenic over chondrocyte fate
during AF closure [8, 12, 24]. This is supported by their expression of Sp7, a key regulator
of osteoblast differentiation that has been implicated in regulating cellular recruitment at
advancing bone fronts [25, 26]. In the absence of Fgfr2, CT2 cells fail to respond to Fgf18
and do not form the articulating edges of the frontal bones that establish the PFS joint.
CT2 cells also express Wif1, a secreted inhibitor of WNT signaling. Within the
fusing murine PFS, canonical WNT signaling is dynamic and sharply down-regulated
immediately preceding cartilage formation in the endocranial domain [27]. Conversely,
ectopically activated canonical WNT signaling in the PFS prevents endochondral
ossification and delayed PFS fusion [6]. We therefore predict that CT2 is a key source of
WNT/b-CATENIN inhibition in endocranial Scx+ cells (CT3) that are marked by Sox9
expression and contribute to endochondral-like fusion of the PFS. This is supported by
our finding that loss of Fgfr2 reduces Wif1 expression in CT2, increases expression of the
WNT target Lgr5 in CT3, and blocks formation of cartilage in the PFS. LGR5 is a known
stem cell marker in multiple epithelial tissues, and in this context may represent
undifferentiated connective tissue progenitors [28-30].
We propose that in the AF, moderate levels of FGF and WNT act together to
maintain multipotent skeletal cells and their combined withdrawal in the endocranial
14
domain promotes chondrocyte fate. In the ectocranial domain, continued exposure to
WNT promotes ligament-like fate, while continued exposure to FGF in the apical region
of the bone fronts promotes osteoblast fate. Loss of Fgfr2 in the AF leads to upregulation
of WNT signaling and prevents both osteoblast and chondrocyte differentiation. Canonical
WNT signaling causing inhibition of chondrogenesis in sutures is supported by the finding
that loss of WNT signaling results in ectopic endochondral ossification and
craniosynostosis in the murine coronal and sagittal sutures [27, 31]. Furthermore,
sustained activation of WNT signaling in Scx+ cells in Scx-Cre;Catnb
GOF
mice prevents
AF closure, as well as PFS formation and fusion as seen in NCC;Fgfr2
-/-
mice. This model
is analogous to the developing limb, where a spatially restricted source of FGF and WNT
signals synergistically promotes proliferation of multipotent progenitor cells but act
separately to determine cartilage versus connective tissue cell fate [32].
Studies of embryonic stages show that the calvaria develops from a layer of head
mesenchyme that is derived from mesoderm or neural crest, and regionally patterned into
osteogenic and non-osteogenic domains. Mesenchyme that resides in the supraorbital
ridge just above the eyes, called the supraorbital mesenchyme (SOM), generates the
ossification centers for the frontal and parietal bones [14, 15, 33, 34]. In contrast,
mesenchyme that lies apical to the SOM, called the early migrating mesenchyme (EMM),
remains non-osteogenic through embryonic stages of calvaria development [35, 36].
Growth of the frontal and parietal bone rudiments towards the vertex of the head in
embryogenesis is fueled by the SOM through intrinsic growth rather than recruitment of
adjacent EMM mesenchyme [15]. Instead, the EMM contributes to the sutures and
fontanelles [35]. By examining the fate of fontanelle connective tissue in the postnatal
15
period, our findings elaborate on this model. Using genetic lineage tracing, we find that
Scx+ cells in the AF become osteogenic and chondrogenic to make a significant
contribution to PFS joint tissue including the articulating bone fronts, cartilage, and
ligament. This indicates that Scx+ cells of the AF serve as joint progenitors. Our model
suggests that as the frontal bone advance, they recruit Scx+ cells of the AF to initiate
frontal suture joint formation, which is characterized by the formation of a secondary
osteogenic front within the endocranial layer which then undergoes endochondral fusion.
This may be a key factor that distinguishes the bilayered, fusing murine PFS from the
overlapping coronal suture, which does not fuse during normal mouse development.
Surprisingly, we observed that loss of Fgfr2 in Scx+ and Sox9+ cells is not
sufficient to recapitulate the AF/PFS phenotype seen in the NCC specific knockout. This
suggests that FGFR2 signaling acts non-autonomously to regulate AF closure and PFS
development, a phenomenon that has been observed in differentiating osteoprogenitors
in the long bone [37, 38]. While this study did not specifically identify which population of
Fgfr2-expressing cells are critical, there are two likely possibilities. Close examination of
the scRNA-seq dataset suggests that co-expression of Fgfr2 and Wif1 are inversely
correlated with Scx expression in CT2, where there is a subset of Fgfr2+/Wif1+ cells that
have little to no Scx or Sox9 expression. Since targeted deletion of Fgfr2 by Scx-Cre in
these cells in unlikely, they may represent the critical source of FGF signaling. Another
possibility is that Fgfr2+/Wif1+ cells in the pre-osteoblast population, which does not
express Scx and has relatively lower levels of Sox9, are a key source of FGF signaling.
In future studies, we would like to examine other changes in gene expression
which may contribute to the phenotype seen in the NCC-specific Fgfr2 loss of function
16
mice. A small number of genes implicated in HEDGEHOG (Hh) signaling were
differentially regulated in NCC;Fgfr2
-/-
mice compared to control. While Hh signaling is
known to be important in bone development, little is known about its role in suture
mesenchyme [39-41]. Two other classes of differentially regulated genes seen in our
RNA-seq, those involved in focal adhesion and ECM-receptor interaction, have been
previously implicated in controlling suture fusion, often through interactions with FGF and
WNT family genes [8, 42, 43]. Genes associated with axonal guidance were also found
to be differentially expressed between control and NCC;Fgfr2
-/-
mice. Previous work has
shown that axon guidance during neurodevelopment shares many key similarities with
normal cellular migration; both of these processes rely on cell-ECM interactions and cell-
cell adhesion for cellular pathfinding and invasion [44]. Genes originally thought to be
specific to axon guidance, including semaphorins, netrins, ephrins, and neurotrophins,
are now known to be expressed within osteoblasts and osteoclasts and facilitate migration
[45-48]. Additionally, cell adhesion molecules may act in non-adhesive roles as guidance
cues for migrating cells and axons [49]. Furthermore, β-CATENIN itself plays a role in cell
adhesion and cellular migration in addition to its role in canonical WNT signaling
[50]. Though differential gene expression in these pathways together suggests possible
changes to cell migration, the late onset of the phenotype of NCC;Fgfr2
-/-
mice as well as
the observed changes in cellular differentiation led us to conclude that deficient migration
over long-distances is not likely to cause the observed phenotype.
Overall, the results presented in this study offer further insight into regional
differences in the development of sutures that are prefigured by fontanelles. It also
suggests a possible etiology for the co-occurrence of craniosynostosis and AF patency
17
observed in FGFR2 GOF related diseases; Fgfr2 mediated repression of WNT/b-
CATENIN signaling causes premature differentiation of SOM derived osteogenic cells but
prevents differentiation of EMM derived Scx+ cells into PFS joint-forming cartilage and
ligament. Contemporary studies have indicated that an increase in WNT/b-CATENIN
signaling in Fgfr2 GOF models of Apert’s syndrome leads to coronal suture synostosis
due to increased activation of WNT co-receptors LRP5/6 [51]. We hope that by providing
further information about the interactions of WNT and FGF signaling in the developing
sutures, we contribute to the discovery of genetic tools to improve clinical treatments and
outcomes for patients suffering from suture defects.
Materials and Methods
Mice
To conditionally knockout Fgfr2, Fgfr2
flx/flx
(JAX Stock No. 007569) mice were
crossed with the Wnt1-Cre2 driver (JAX Stock No. 022137), the Sox9-CreERT2 driver
(provided by Dr. Francesca Mariani), or the ScxGFP-Cre driver (provided by Dr. Ronen
Schweitzer). The Fgfr2
flx/flx
and Wnt1-Cre2 lines were previously described [13, 52]. Scx-
GFP was used to mark dense connective tissue fibroblasts [17]. ScxGFP-Cre, and Sox9-
Cre-ERT2 lines were described previously [53, 54]. The Ai9 allele (JAX Stock No.
007909) was used as a lineage marker for those tissues targeted by Wnt1-Cre2, ScxGFP-
Cre, and Sox9-Cre-ERT2 [55]. Finally, constitutive activation of 𝛽-Catenin in NCCs was
driven by crossing Catnb
lox(Ex3)
mice [22] to mice carrying the Wnt1-Cre2 driver.
Embryonic samples were collected from timed pregnant females. Postnatal samples were
staged according to the date of birth. All experimental protocols were approved by the
USC’s Institutional Animal Care and Use Committee.
18
Skeletal preparation
Mice processed for whole mount imaging were collected as above and following
PBS rinse were fixed in 100% ethanol for a minimum of 4 days. Fixed samples were
stained for cartilage using 0.15mg/mL Alcian Blue (Sigma-Aldrich CAT. A5268) in 80%
ethanol and 20% glacial acetic acid. Samples were then de-stained in 100% ethanol for
a minimum of two days. Tissue clearing was performed using KOH between 0.5-1% w/v
for between 2-8 days depending on size of sample. Calcified tissue was then
discriminated using 0.02 mg/mL Alizarin Red S (Sigma-Aldrich CAT. A5533) in 0.5-1%
KOH. Destaining and further clearing was performed as needed in 0.5-1% KOH. Samples
were equilibrated, stored, and imaged in 75% glycerol.
Micro-computed tomography (μCT)
All μCT scans were performed by the USC Molecular Imaging Center using a
μCT50 (Scanco Medical). Samples were rotated 360° and X-ray settings were
standardized to 90 kV and 155 µA with an exposure time of 0.5 seconds per frame to
yield a nominal resolution of 20 μM. A 0.5-mm-thick aluminum filter was employed to
minimize beam-hardening artifacts. Morphometric analysis was performed using the
Amira 6.2 and VG Studio MAX 3.0 software packages. Isosurface renderings with equal
threshold were measured using the 3D measuring tool. All jaw measurements and
landmarks were measured as previously described [56]. These experiments were
performed on at least 3 biological replicates, which we defined as 3 same-sex littermate
pairs (control and mutant) derived from 3 different litters. Statistical significance was
determined using unpaired two-tailed t-tests.

19
Histology
Samples used for histology were processed as follows; animals were humanely
sacrificed and decapitated. Heads were then fully or partially skinned, rinsed in PBS, and
fixed in 10% neutral buffered formalin (NBF) for between 24-36 hours depending on
stage. Following fixation, samples were decalcified in 10% EDTA for 3-7 days, until
calvariae began to concave. Samples were then dehydrated in serial ethanol washes of
increasing concentration, equilibrated in Citrus Clearing Solvent (VWR CAT.72060-044),
and embedded following 3 washes in pure paraffin wax undergone at 65˚C with vacuum
pressure. Embedded samples were then cut into 8 µM sections. Sections were stained
using the Hall-Brunt quadruple stain [16].
Immunofluorescent analysis
Samples for immunofluorescent staining were harvested and fixed in 4% PFA for
between 30-60 mins depending on stage. Decalcification of bone was performed via
incubation in 10% EDTA (pH 7.4) for between 3-7 days at 4°C until calvaria began to
concave, Subsequently, samples were equilibrated sequentially in 15% and 30%
sucrose/PBS at 4°C until they sank, then embedded in optimal cutting temperature
(O.C.T.) compound (Electron Microscopy Sciences). Cryosectioning was performed at a
section thickness of 8μm onto Superfrost+ slides. Sections were washed three times with
PBS, permeabilized with 0.1% TritonX-100 in PBS, then blocked for one hour at room
temperature in 10% serum (either donkey or goat from Sigma). Slides were incubated in
primary antibody overnight at 4°C (Supplemental Table 1), then washed in PBS and
incubated in Alexa Fluor secondary antibodies at a concentration of 1:500 for one hour at
room temperature. Finally, slides were washed in PBS and mounted in Vectashield with
20
DAPI (Vector Labs). Slides used for lineage tracing were washed three times in PBS and
counterstained with DAPI as described above. Slides were imaged using either a Leica
TCS SP5/8 or Stellaris 5 confocal system.
Single cell sequencing
Data for scRNA-seq analysis was obtained from FaceBase (Accession #1-4TT6)
[18]. Seurat 3 R-Package was used to explore QC metrics and filter for high-quality cells.
The filtered dataset representing 3366 cells (median of 3750 genes per cell) was
normalized (log normalized), scaled (linear transformation), and clustered into 14
communities using unsupervised graph-based clustering based on principal component
analysis scores. The identities of the clusters were resolved using previously reported
markers. Pseudotime analysis was conducted using Monocle.
In situ hybridization
Samples for in situ hybridization were collected and processed in the same manner
as histological samples, described above. Transcripts were detected using RNAscope
Fluorescent Multiplex Assay (ACD) as per manufacturer’s instructions. Briefly, slides with
paraffin sections were baked on a hot plate for 60 mins. at 65°C. Slides were then
deparaffinized using xylenes, dehydrated in 100% EtOH, and allowed to completely air
dry. Endogenous peroxidase activity was quenched using hydrogen peroxide provided
within the kit, slides were washed in deionized water. Antigen retrieval was performed
using the provided Target Retrieval Reagent for 15 mins. in an Oster steamer heated to
99°C, then dehydrated in 100% EtOH and allowed to completely air dry. The slides were
treated with ACD Protease plus in a humidified slide chamber for 15 mins. at 40°C. Probe
hybridization was also performed in a 40°C humidified slide chamber for 2 hours
21
(Supplemental Table 1). Slides were then stored in 5x SSC solution overnight.
Amplification steps were performed as prescribed by manufacturer and signal
development for channels one and two were done using TSA Plus fluorophores Cyanine
3 (Perkin and Elmer NEL744E001KT) and Cyanine 5 (Perkin and Elmer NEL705A001KT)
diluted 1:1000 in the ACD provided TSA buffer. Slides were then counterstained using
Vectashield mounting medium with DAPI and imaged using the Leica TCS SP5/8 or
Stellaris 5 confocal system. These experiments were performed on 3 biological replicates,
which we defined as 3 littermate pairs (control and mutant) derived from 2 different litters.
EdU Proliferation Assay
Proliferation of cells was assayed using an in vivo EdU Click kit according to
manufacturer instructions (Sigma BCK647-IV-IM-M). Briefly, E18.5 samples were
obtained by intraperitoneally injecting timed pregnant females with 1.5 mg EdU dissolved
in PBS, while P3 mice received subcutaneous injections of .1mg EdU dissolved in PBS.
In both cases, a chase period of four hours was observed before sample collection.
Samples were cryo-embedded and sectioned at 8μm as described above. Detection of
EdU took place in the dark, at room temperature for 30 mins. Slides were then
counterstained and mounted with Vectashield with DAPI. For analysis, littermate control
and NCC;Fgfr2
-/-
pairs were selected, and a minimum of 6 different sections from each
sample were imaged and uploaded to ImageJ. Each image was quantified for total AF
cells via individual nuclei, and the number of proliferating cells was counted. Statistical
analysis was performed on total proliferation in mutant vs control, proliferation of AF
fibrous connective tissue, and proliferation in bone front cells. Statistical significance was
determined using unpaired two-tailed t-tests.
22
RNA Isolation and Gene Expression Analysis
Mice of desired age were sacrificed by decapitation and the region of the anterior
fontanelle was excised (including both suture connective tissue and bone but excluding
skin and brain). Samples were placed in 300µL of DNA/RNA Shield (Zymo) on ice until
ready for further processing. RNA isolation from tissue was carried out using the Quick-
RNA Miniprep Plus Kit (Zymo) according to manufacturer’s instructions with the addition
of the optional DNase digestion and a 1 min dounce homogenization step following the
proteinase-K digestion. Samples were eluted from the column using 80 µL of
DNase/RNase- free water and then analyzed via nano-drop and to check concentration
and purity. Library preparation and sequencing was performed at the UCLA TCBG facility
using the Kapa Stranded Kit (Roche). Sequencing was performed using Hiseq3000 at
1x50 read length and 30 million reads per sample. Differential gene expression analysis
was performed using Partek Flow.

23
Figure Legends:
Figure 1: NCC;Fgfr2
-/-
mutant mice show a widening and patency of the anterior
fontanelle starting at E18.5.
A Schematic representing the tissue origins of the calvarial vault; neural crest
derived tissue is depicted in blue while mesoderm is labeled in pink (A). Whole mount
skeletal prep staining using alizarin red and alcian blue shows morphological differences
between control and NCC;Fgfr2
-/-
littermate AF development. At E18.5 NCC;Fgfr2
-/-
mice
show a slight widening of the anterior fontanelle (B,C, n=3 pairs). By P3 and P5 the
widening becomes more pronounced, and the frontal bones fail to approximate (D,E,F,G,
n=3 pairs each). In P10 controls, regions of cartilaginous fusion in the frontal suture
become evident (H) and are not observed in NCC;Fgfr2
-/-
mutants (I). (J,L) µCT scans of
NCC;Fgfr2
-/-
mutant and littermate control at P30 show AF dysmorphia in mutant including
patency and bone mineralization defects (asterisk n=2 pairs). Lines in panel J & L
represent the plane of orthoslice shown to the right (K,M). (N) Graph representing the
differences in normalized average AF area between control and NCC;Fgfr2
-/-
mutants at
P3 (n= 7 pairs) and P5 (n= 9 pairs). Normalized area was calculated individually by
sample by dividing the total area of the AF by the sample’s frontal suture length from the
jugum limitans to the top of the bregma. AF, anterior fontanelle; Ctrl, control.
Figure 2: NCC;Fgfr2
-/-
mutants show defects in frontal bone approximation and lack
endocranial cartilage that prefigures the frontal suture.
HBQ stained histological sections of control samples (A, C, E, G) show progressive
stratification of the ectocranial and endocranial layers of bone (red) that form the mature
frontal suture. By P7, the cartilage that prefigures frontal suture fusion begins to condense
(C) and by P11 these cells take on a chondrocyte morphology and stain positively for
proteoglycans via alcian blue (E). By P13, the cartilage has begun to undergo
endochondral ossification, forming the bony endocranial fusion of the frontal suture. In
NCC;Fgfr2
-/-
mutant samples (B, D, F, H) approximation of the frontal bones lags that of
control and mineralized tissue shows no evidence of stratification into ecto- and
endocranial layers. By P13 mutants still show no sign of forming cartilage at the suture
midline, which is still occupied by CT (H). (n=3 pairs for each timepoint)
Figure 3: Anterior fontanelle connective tissue expresses canonical
tendon/ligament marker Scx
As seen in both wholemount (A,B) and in section (C, D) the fibrous connective
tissue of the control AF expresses Scx-GFP. At E18.5, Scx-GFP is present in all calvarial
sutures and fontanelles as well as immediately adjacent bone (A, n=2). By P3, Scx
expression is more confined to the sutures themselves (B, n=5). Sections through the
anterior fontanelle at these stages revealed a bi-layered expressional pattern of Scx, with
cells at both the ecto- and endo-cranial layers of the suture mesenchyme. Some GFP
positive cells were also identified in the frontal bones at E18.5 (n=5), seen more clearly
at P3 (n=5) (C,D). AF, anterior fontanelle; CS, coronal suture; FB, frontal bone; FS, frontal
suture; PB, parietal bone; SS, sagittal suture; LS lambdoid suture.
24
Fig. 4: Anterior fontanelle connective tissue is composed of heterogeneous cell
populations with distinct gene expression patterns and spatial organization.
UMAP plot showing cluster analysis of sequenced frontal suture cells (A) reveals
multiple cell populations. Feature maps reveal that three of these clusters (CT1, CT2, and
CT3) show high expression of Scx and variable expression of Fgfr2 (B). In situ
hybridization confirms that collectively these three clusters represent cells of both the
advancing frontal bones as well as the intervening FCT (C). Expressional analysis of the
three CT clusters revealed differential gene expression and identified Fgf18, Sp7, and
Sox9 as being enriched in each population, respectively (B). Finally, further in situ
analysis shows that each of the three CT populations is spatially distinct within the AF;
CT1 represents the ectocranial connective tissue of the AF (D, n=3), CT2 is localized to
the frontal bones and the region immediately preceding their growth (D, E, n=6), and CT3
represents the endocranial layer of connective tissue (E, n=3). Predicted lineage
trajectory shows two paths of differentiation, which suggests CT1/CT2 and CT3 converge
on an osteoblast identity (F). CT, connective tissue; DM, dura mater; EC, endothelial cells;
HC, hematopoietic cells; HD, hypodermis; OB, osteoblasts; PO, proliferating osteoblasts;
PreO, pre-osteoblasts.
Figure 5: Both Scx and Sox9 expressing cells contribute to the fused frontal suture
ScxGFP-Cre;TdTom mice have a constitutively active Cre and were collected at
P4 (n=3) and P7 (n=3). At P4, labeled cells appear throughout the bone and CT and are
most highly concentrated within the endocranial layer and a thin layer of cells overlaying
the entire suture (A). By P7, the number of labeled cells in both the ecto- and endocranial
layers has increased, and the cells contribute both the midsuture CT and bone (B). Sox9-
CreERT2;TdTom were given tamoxifen at stages P2 and P3 to induce recombination,
then collected at P4 (C, n=3) and P7 (D, n=4). Labeled cells are present at the bone fronts
and within the midsuture at P4 (C). By P7, the Sox9 cells at the midsuture have
condensed in preparation for cartilage formation, while bone front labeling is comparable
to P4 (D).
Figure 6: EdU pulse chase experiments show a minor decrease in proliferation of
NCC;Fgfr2
-/-
mutant mice vs littermate controls.
Representative sections of EdU labeling at E18.5 (A, B, n=7 pairs) and P3 (D,E,
n=6 pairs) show proliferating cells within the bone fronts (white dotted lines) and AF
connective tissue (yellow dotted lines). Percentages of proliferating cells were calculated
by dividing number of labeled cells by total cells in a region as determined by counting
nuclei (C, F). Graphs show calculated averages of connective tissue (CT), bone fronts
(BF) and of both areas combined (total). At E18.5 significant differences in proliferation
were only seen when counting both the bone front and CT regions (6% in control vs. 5.1%
in cKO, p=.02) (C). By P3, total proliferation was overall higher in control samples (9.6%
in control vs. 7.3% in cKO, p= .004) with the bulk of this difference occurring in the suture
CT (6.8% in control vs. 4.9% in cKO, p= .001) (F).
Fig. 7: NCC;Fgfr2
-/-
mice lose expression of cartilage and osteoblast specific
markers in midsuture FCT of in favor of maintaining Scx expressing connective
tissue.
25
Immunofluorescence staining of control samples (A, C, E, n=3 each) show
progressive increase of Runx2 expression within Scx expressing midsuture tissue
beginning at P3 (C) and contributing to the endocranial bone layer by P5 (E). In littermate
NCC;Fgfr2
-/-
mice (B,D,F, n=3 each), these cells do not upregulate Runx2, but instead
maintain Scx expression and a more connective tissue-like morphology. At P5, control
samples show little to no expression of the cartilage marker Sox9 (G, n=3), but by P7
expression of Sox9 is visible within the endocranial midsuture as endochondral fusion is
initiated (I, n=3). In the mutant samples, these Sox9 expressing cells are not seen (H, J,
n=3 each).
Figure 8: RNA-sequencing of AF tissue reveals differential gene enrichment
between control and NCC;Fgfr2
-/-
samples.
Samples of the anterior fontanelle, including both bone fronts and intervening
connective tissue (dotted region in A schematic) were collected and analyzed for RNA
content. Heatmap analysis reveals substantial differential gene enrichment between
NCC;Fgfr2
-/-
(top rows) and littermate control samples (bottom rows) (A, n=5 pairs). GO
analysis reveals significant differential regulation of genes associated with focal adhesion,
axon guidance, and Wnt signaling (B). Differentially regulated genes associated with the
Wnt pathway included LGR6, SFRP4, Notum, and Wif1 (upregulated in control vs. cKO)
as well as LGR5 and SFRP2 (downregulated in control vs. cKO) (C). String analysis
shows the relationship of these genes to members of the FGF signaling pathways as well
as WNT pathway ligands Wnt2 and Wnt11 (D).
Figure 9: Key WNT pathway members are dysregulated in NCC;Fgfr2
-/-
mice,
suggesting increased signaling
ISH performed with RNAscope on E18.5 control mice revealed high levels of Wif1
expression in and immediately surrounding the frontal bones (dotted outlines), and low
levels of Lgr5 expression in the midsuture and a thin layer of cells overlying the suture (A,
n=3 pairs). Littermate NCC;Fgfr2
-/-
mice show overall lower expression of Wif1 and have
greatly increased Lgr5 expression in the midsuture and overlying the suture. A subset of
the CT2 population (dotted ellipse) shows high co-expression of Fgfr2 and Wif1 (C).
Figure 10: Ectopic activation of Wnt/𝛽- catenin signaling in Scx expressing FCT
phenocopies NCC;Fgfr2
-/-
mutant phenotype
At E18.5, Scx-Catnb
GOF
mice have much wider AF regions than littermate controls
(A,B, n=4 pairs). In addition, bone quality in the leading edges of the frontal bones is of
poor quality. At P5, frontal bones of P5 control mice have approximated to form the
presumptive PFS (C, n=5). Meanwhile, Scx-Catnb
GOF
mice maintain a wide-open AF, and
still demonstrate decreased bone mineralization (D, n=1).
Figure 11: The connective tissue recruitment model of frontal suture development
In control mice, signaling through Fgfr2 down regulates Wnt signaling within the
presumptive PFS. This allows conditions permissive to the differentiation of mesenchymal
suture progenitors into Scx+/Sox9+ or Scx+/Runx2+ skeletogenic cells which are
recruited to contribute to the fusion of the PFS. Reciprocal interactions between Wnt and
Fgf18 facilitate a feedback loop to regulate levels of Fgf and Wnt during this process. In
26
NCC;Fgfr2
-/-
mice, loss of Fgfr2 expression prevents the downregulation of Wnt signaling
during suture establishment and fusion. Scx+ progenitor cells in the midsuture fail to
differentiate leading to developmental arrest and AF patency, the cells instead maintain
Scx expression and a more fibrous connective tissue-like identity.
Supplemental Figure 1: Mice of Scx- Fgfr2
-/-
and Sox9-Fgfr2
-/-
genotypes are
indistinguishable from littermate controls at P5 and P7.
Scx- Fgfr2
-/-
mice and littermate controls were collected at P5 (A, B, n=5 pairs)
and P7 (C,D, n=2 pairs) but had no noticeable phenotype. Sox9-Fgfr2
-/-
were generated
either via tamoxifen induction at E17.5/ 18.5 and collection at P5 (E,F, n=5 pairs), or
induction at P0 and collection at P7 (G, H, n=3 pairs) and were indistinguishable from
control littermates.
Supplemental Figure 2: A subcluster of CT2 cells have high Fgfr2 but low
Scx and Sox9 expression.
High resolution clustering of the scRNA-seq dataset shows that there is a
subpopulation of cells within the CT2 population (A). Feature maps reveal that this
subset has high relative expression of Fgfr2 (B,C, green) but low expression of Scx (B,
pink) and Sox9 (C, pink).
Supplemental Figure 3: Cleaved caspase staining shows no significant levels of
apoptosis in control or NCC;Fgfr2
-/-
littermate mice.
Immunofluorescence staining of cleaved caspase at P0 (A,B), P3 (C,D), and P5
(E,F) shows no significant levels of apoptosis in either NCC;Fgfr2
-/- or
littermate control
mice (n=3 pairs for each stage). Antibody viability was confirmed with etoposide treated
and untreated control slides (CST 8104S) (G,H).

27
Figure 1:

28
Figure 2:

29
Figure 3:

30
Figure 4:

31
Figure 5:

32
Figure 6:

33

Figure 7:

34
Figure 8:

35
Figure 9:

36
Figure 10:

37
Figure 11:

38
Supplemental Figure 1:

39
Supplemental Figure 2:

40

Supplemental Figure 3:

41
Supplemental Table 1:

Target Company- Cat. # Application Notes
Scx ACD- 439981 RNA-scope Channel 1
Fgfr2 ACD- 443501 RNA-scope Channel 2
Fgf18 ACD- 495421 RNA-scope Channel 1
Sp7 ACD- 403401 RNA-scope Channel 3
Sox9 ACD- 401051 RNA-scope Channel 2/3
Lgr5 ACD-312171 RNA-scope Channel 1
Wif1 ACD- 412361 RNA-scope Channel 3
Cleaved Caspase CST- 9661S Immunofluorescence 1:400 conc.
Runx2 CST- 12556 Immunofluorescence 1:400 conc.
Sox9 Novus- NBP1-
85551
Immunofluorescence 1:400 conc.

42
Chapter 2: Development and Maintenance of Tendons and
Ligaments
(This was previously published in Bobzin et al., Development 2021)
Tendons and ligaments are fibrous connective tissues vital to the transmission of
force and stabilization of the musculoskeletal system. Arising in precise regions of the
embryo, tendons and ligaments share many properties and little is known about the
molecular differences that differentiate them. Recent studies have revealed heterogeneity
and plasticity within tendon and ligament cells, raising questions regarding the
developmental mechanisms regulating tendon and ligament identity. Here, we discuss
recent findings that contribute to our understanding of the mechanisms that establish and
maintain tendon progenitors and their differentiated progeny in the head, trunk, and limb.
We also review the extent to which these findings are specific to certain anatomical
regions and model organisms, and indicate which findings similarly apply to ligaments.
Finally, we address current research regarding the cellular lineages that contribute to
tendon and ligament repair, and to what extent their regulation is conserved within tendon
and ligament development.
Introduction
Tendons and ligaments are closely related dense fibrous connective tissues that
play vital roles in musculoskeletal mobility and stability. Tendons connect muscle to bone
and facilitate body movement by transmitting tensile forces and storing elastic energy,
whereas ligaments join bone to bone, stabilizing joints and guiding movement through the
normal range of motion [57-59]. The functions of tendons and ligaments rely on their
strong yet flexible structure of collagen fibrils, which are hierarchically organized and
bundled by connective tissue sheaths. Though tendon and ligament are similarly
43
composed of collagen 1 (COL1) fibrils, proteoglycans, elastin and glycoproteins, a few
differences do exist. COL1 fibrils, which are aligned parallel to one another along the
longitudinal axis in tendons, are multidirectional and less densely packed in ligaments to
better support tensile load [60-63] (Fig. 1).
Individual tendons and ligaments have unique morphological properties adapted
for specific roles. Remarkable variability exists in tendon and ligament shape, which can
form cords, strap-like bands, flat ribbons, discs, and fan-like structures to accommodate
distinct mechanical and anatomical environments [57, 58]. Morphological diversity among
tendons and ligaments is further enhanced through their variable association with
synovial sheaths, bursae, fibrous retinacula and fat pads that function to lubricate, anchor,
support and provide mechanosensory input [57]. In addition, there is morphological
variability along tendons, with fibrocartilage forming at bone insertions, sesamoid bones
forming within regions that cross over joint surfaces, and myotendinous junctions where
tendon unites with muscle (Fig. 1). Genome-wide expression analysis of porcine adult
tendon and ligament tissue from distinct regions and subtypes suggests that
morphological and functional differences largely correlate with variation in gene
expression levels rather than activation of subtype-specific genes [64]. The extent to
which embryonic factors are sufficient to establish location-specific morphology has not
been directly investigated. Undoubtedly, morphological variation among tendon and
ligament subtypes will be key to interpreting region-specific differences in development
and repair.
Tendons and ligaments arise from distinct embryonic origins depending on their
position along the rostro-caudal axis (Fig. 2). Tendons and ligaments in the head arise
44
from cranial neural crest cells (CNCCs), whereas those in the neck and shoulder girdle
are derived from lateral plate mesoderm, CNCCs and paraxial mesoderm [65-71].
Lineage tracing of CNCCs and mesoderm in the murine neck suggests that the embryonic
origin of the tendon matches that of the bone to which it attaches [72]. In the limb, tendons
and ligaments originate from lateral plate mesoderm, whereas those in the trunk are
derived from somites [73]. The extent to which the developmental origin of tendons and
ligaments influences their mechanisms of formation and later properties is unclear.
Fibroblastic-like cells, often called tenocytes and ligamentocytes, are the principal
mature cell types within tendons and ligaments, respectively. The fundamental nature of
tendon and ligament cell identity and their heterogeneity have yet to be determined. A
major challenge in defining these cell types is in their seemingly strong potential to revert
to a progenitor-like state or transdifferentiate into osteoblasts or chondrocytes [74-77]. As
both tendon and ligament pathologies are commonly associated with aberrant bone
formation, there is a need to understand the factors that maintain tendon and ligament
cell fate [78].
In this review, we focus primarily on the better-understood mechanism of tendon
development, indicating where key findings are also applicable to ligaments. We discuss
region-specific developmental mechanisms in the head, trunk and limb that induce and
maintain tendon progenitors and regulate tendon cell differentiation and maturation. We
also discuss current evidence in the field that points to heterogeneity and regional
specificity within tendon cell populations. Finally, we compare tendon progenitors in
development to adult stem and progenitor cells that contribute to tendon and tendon
attachment repair.
45
Tendon and ligament development
The discovery of Scleraxis (SCX), a basic helix-loop-helix transcription factor, as a
marker of both progenitors and differentiated cells within tendon and ligament has led to
the creation of powerful genetic tools for understanding their development [17, 79, 80].
The Scx-GFP reporter and Scx-Cre alleles in mice are routinely employed in mechanistic
studies that address tendon progenitor patterning, differentiation, and maintenance [17,
81, 82]. Similarly, the Scx knockout mouse has underscored its necessity in tendon
morphogenesis [59, 83]. In zebrafish, nonsense mutations in scxa and scxb lead to
defects in cranial tendon and ligament morphogenesis, showing a conserved role for Scx
in differentiation and maturation of tendons across vertebrates [84]. Molecular studies
indicate that Scx promotes tendon cell differentiation and maturation by transcriptionally
activating a suite of tendon cell genes, including the major building blocks of the tendon
extracellular matrix (ECM), such as Col1a1 and tenomodulin (Tnmd) [85, 86].
Scx is expressed in all tendon and ligament progenitors, yet its requirement during
development varies depending on the subtype and location of the tendons. Loss of Scx in
mice preferentially disrupts formation of long-range, but not short-range, muscle-
anchoring of tendons and ligaments in the limb and trunk [59]. This confounding aspect
of the Scx knockout phenotype has been addressed in a recent study that more closely
examined long-range tendon development. Development of both long-range and short-
range tendons begins with formation of a tendon anlage that anchors to muscle and bone.
In long-range tendons, this anlage subsequently undergoes rapid elongation that is fueled
by recruitment of surrounding Scx
+
progenitors into the tenogenic lineage. Scx, although
not required for formation of the initial tendon anlagen, is necessary to recruit tendon
46
progenitors for substantial extension during growth [87]. Nevertheless, Scx remains the
earliest known marker of tendon and ligament progenitors, it does not appear to be
necessary for tendon cell fate determination.
Establishment of tendon progenitors
A central pathway key to establishment of tendon progenitors is transforming
growth factor β (TGFβ) signaling. TGFβ2/3 ligands emanating from neighboring
cartilage/muscle recruit undifferentiated mesenchymal cells to form tendon progenitors,
and then maintain them by activating Scx expression in both mouse and chick [88, 89]
(Fig. 3). Loss of Tgfb2/3 in mice leads to diminished Scx expression, considerable
reduction in tendon progenitor numbers, and eventual widespread loss of cranial, limb
and trunk tendons [88, 90]. In addition, conditional loss of Tgfbr2 in mouse limb
mesenchyme (via Prx1-Cre) results in depletion of limb tendons [88]. The dramatic
reduction of tendon and ligament progenitors in mice with mutations in the TGFβ signaling
pathway is the result of lost cellular identity, either through de-differentiation or acquisition
of different cell fates [88].
Region-specific regulation of tendon progenitors
The distinct embryonic origins of cranial, limb and trunk tendons suggest the
possibility that the establishment and differentiation of each population is uniquely
regulated (Fig. 2). Although this has been difficult to address given the paucity of
information on cranial tendon progenitors relative to those of the limb and trunk, growing
evidence supports this idea. For example, fibroblast growth factor (FGF) signaling from
the adjacent myotome promotes the induction of axial tendon progenitors by
driving Scx expression in mice and chicks [91-93] (Fig. 2). Conversely, induction of
47
cranial and distal limb (autopod) tendon progenitors is muscle independent, with muscle
interaction only required later for maintenance of cellular identity [94-98]. In addition,
interactions with cartilage are vital to the organization (and in some cases establishment)
of tendon and ligament progenitors of the craniofacial and axial skeleton [95].
Current research suggests that the relationship between FGF signaling
and Scx expression varies based on species and anatomical region. We have recently
found that Fgfr2 promotes establishment of SCX
+
tendon progenitor cells at tendon-bone
insertion sites in the jaw [83] (Fig. 2). Exogenous treatments that activate or block FGF
ligands, or its downstream effector ERK (MAPK1), in limb explants suggest that FGF
signaling promotes Scx expression in chick and inhibits Scx expression in mouse [89, 99,
100]. Conversely in zebrafish embryos, inhibition of FGF signaling leads to loss of all
Scx
+
progenitors contributing to both tendon and ligament [95]. The reason for this
species-specific difference is unclear, but the outcome of FGF signaling
on Scx expression may be related to the intensity and duration of the FGF-ERK signal
[93] or, alternatively, differences in experimental approaches between species. It is also
not yet clear what the specific role of FGF signaling is in ligament, although the common
embryonic origins and function of tendon and ligament would suggest that findings
specific to tendons in various anatomical regions could be extrapolated to nearby
ligaments. The limited available literature suggests that FGF signaling enhances
proliferation and differentiation in periodontal ligament cells in vitro by upregulating Scx
and Tnmd while suppressing osteogenic differentiation through inhibition
of Runx2 expression [101, 102]. Defining a clearer role for FGF signaling in tendon and
ligament induction will require further analysis of FGF-related mutant models.
48
Studies in the limb provide evidence that tendon progenitors of the same
embryonic origin can have distinct regulatory programs depending on their location along
the dorsal-ventral and proximal-distal axes. Whole-transcriptome expression profiling of
limb-derived Scx-GFP cells during the early stages of development has
identified Foxf2 and Six2 as tendon-enriched transcription factors with region-specific
expression [103]. Foxf2 expression is restricted to the dorsal tendons within the distal limb
(autopod), whereas Six2 expression is enriched in ventral tendons of both the proximal
(zeugopod) and distal (autopod) limb (Fig. 2). However, it is not clear whether this
difference in expression is specific to Scx-expressing tendon and ligament populations
[103]. In the proximal limb (zeugopod), differentiation and growth of tendons is dependent
on the muscle [80, 94, 104, 105] (Fig. 2). In the autopod, however, the induction,
differentiation and maintenance of tendons, progenitors of which are distinctly derived
from a Six2 lineage, is instead dependent on the cartilage that prefigures the bone [104]
(Fig. 2). The conclusion that development of autopod tendons is dependent on skeletal
cues is underscored by the finding that autopod tendons are induced in supernumerary
digits and fail to form in the absence of cartilage [104]. Whether cartilage promotes tendon
formation by secreting tenogenic factors or providing a source of progenitors with
tenogenic potential is unknown; however, candidate signaling pathways for regulation of
this process include TGFβ, FGF and WNT [83, 88, 106].
Tendon cell maturation
Once tendon cells are specified, they function primarily in producing the tendon's
ECM and establishing the patterning of collagen fibrils and their bundles [107] (Fig. 3).
The most abundant structural protein in tendon is COL1, which comprises about 95% of
49
the total collagen content. Expression of Col1a1 in tendon cells is regulated by multiple
factors, including SCX, which directly binds to E-box sites within the Col1a1 promoter
[85]. Other transcription factors that are known to have functional roles in tendon
development, such as mohawk (Mkx) and Egr1/2, are primarily required for tendon
maturation, including matrix deposition and organization [108-110] (Fig. 3). For example,
loss of Mkx decreases COL1A1 expression and impairs collagen fibril formation, reducing
tendon mass, but has no discernible effect on ligament development [108, 110]. Though
expression of Mkx and Egr1/2 has been documented in ligament fibroblasts, it is not yet
clear what their roles in ligament cell differentiation and maturation may be [110, 111].
As tendons mature, their respective fibroblasts arrange in longitudinal rows along
the collagen fibrils, flatten and become less dense. Maturing tendon cells develop a
complex network of cellular projections, including nanotubes that facilitate intercellular
communications via gap junctions [57, 112, 113]. In zebrafish, the force of muscle
contraction promotes release of TGFβ, which is required for tendon progenitors to extend
microtubule-rich projections [114]. These projections, in turn, promote tendon cell
maturation by regulating force-dependent expression of tendon- and ligament-specific
ECM components, such as Thrombospondin 4b [114-116].
Tendon cell fate maintenance
In the mature tendon, tenocytes require continuous feedback to maintain their
differentiated state and repress metaplasia into the osteochondral lineage. Feedback
mechanisms that maintain tendon cell fate (and have been loosely implicated in ligament
fibroblast maintenance) are discussed below, including growth factors, ECM components
and mechanical force (Fig. 3). Tendon cell projections are also essential for cell fate
50
maintenance, as their loss leads to the upregulation of scxa in zebrafish, suggesting a
reversion to a de-differentiated state [114].
Growth Factors
TGFβ signaling is the primary growth factor pathway known to be involved in
maintenance of tendon cell fate (Fig. 3). Targeted deletion of Tgfbr2 in tendon and
ligament cells using Scx-Cre does not appear to affect cell fate during embryonic
development. However, tendon and ligament cells in early postnatal conditional knockout
mice lose expression of differentiation markers and reactivate genes associated with
tendon progenitors at the earliest stages of embryonic tendon induction [76, 89]. The role
of TGFβR2 signaling is cell-autonomous in tendon cells, as targeted re-expression
of Tgfbr2 in de-differentiated tendon cells is sufficient to reactivate differentiation.
Together, this suggests that tendon cell fate is reversible, and that tendon cell identity
requires extrinsic growth factor signals for continuous maintenance.
ECM components
Proteoglycans within the tendon ECM control the bioavailability of growth factors,
such as TGFβ/BMP, and play important roles in maintaining tendon cell fate. The
metalloproteinases ADAMTS7 and ADAMTS12 are co-expressed in mouse hindlimb
tendons starting in the late embryonic period and persist through adulthood to maintain
tendon cell fate [117]. Combined inactivation of Adamts7 and Adamts12 alters collagen
fibrillogenesis and leads to progressive heterotopic ossification (HO) in juvenile mouse
tendons [117]. Prior to HO, tendon cells exhibit enhanced BMP signaling and upregulate
cartilage and bone markers. This suggests that ADAMTS7 and ADAMTS12 maintain the
tendon cell fate by suppressing osteochondral lineage differentiation. Similarly, the small
51
leucine-rich proteoglycans biglycan (BGN) and fibromodulin (FMOD) promote tendon cell
differentiation by repressing BMP signaling, and their combined loss results in HO within
the hindlimb tendons [118, 119]. In a model for tendon HO caused by hyperactive BMP
signaling, ectopic bone is derived from Scx-lineage cells that aberrantly activate bone and
cartilage markers [120]. These studies show that tendon ECM preserves the tendon cell
fate, in part by providing factors that inhibit bone formation, and strongly supports a role
for ECM proteins in the maintenance of tendon cell fate.
Mechanical feedback
Mechanical signals are also necessary to maintain tendon cell fate during
development and homeostasis [121]. Tendon cells respond to physical input in part
through activation of the mechanoresponsive transcription factors EGR1/2 and MKX.
Increased load activates Egr1 and Mkx expression, and decreased load reduces their
expression [122-124]. Downstream of mechanical signals, EGR1/2 and MKX promote
tendon cell fate through distinct mechanisms. EGR1 is sufficient to activate gene
expression of Scx and Col1a1 during development, and to promote tendon cell fate
during postnatal growth [109, 125]. In postnatal tendons, EGR1 occupancy is enriched at
the Tgfb2 promoter and the absence of Egr1r educes Tgfb2 expression [125]. This
suggests that, in response to mechanical signals, EGR1 acts upstream of TGFβ signaling
to maintain tendon cell identity [126]. Although all of the aforementioned factors have also
been found to be expressed in ligament, their roles in ligament cell maintenance remain
unclear. However, a wealth of evidence supports the importance of mechanical signaling
in controlling ligament cell fate, in part through regulation of RHO/ROCK signaling in the
ECM microenvironment [127, 128].
52
In Mkx knockout mice, mechanical stimuli fail to maintain tenogenic gene
expression and hindlimb tendons exhibit HO [123, 129]. These ossified sites are derived
from aberrant differentiation of Scx-lineage cells into chondrocytes that then undergo
endochondral-like bone formation like that seen in Adamts7, Adamts12, Bgn and Fmod
mutant models. Selective inactivation of Mkx in postnatal Scx
+
tendon cells recapitulates
the HO phenotype [129]. Although tendon HO is associated with increased BMP signaling
in the mouse models listed above, Hedgehog (Hh) signaling plays a crucial role in tendon
ossification in Mkx mutant mice. Interestingly, many cells at the periphery of ossifying
tendon nodules express both Scx-GFP and SOX9, like developing sesamoid bones and
bone eminences [81, 130, 131] (Fig. 4). A chromatin immunoprecipitation assay for MKX
in rat tendon-derived cells identified enrichment of both tendon- and cartilage-related
genes, suggesting a dual role for MKX in transcriptional promotion of tendon cell
differentiation, while preventing chondrogenic/osteogenic differentiation [132]. Although
these studies did not directly address ligaments, the mechanisms and signals are
assumed to be recapitulated, as many human diseases featuring HO affect both types of
tissue [133].
Tendon cell fate at the tendon-bone interface
Tendons and ligaments attach to bone through a specialized tissue called the
enthesis, which functions to dissipate mechanical stress across the hard-soft tissue
interface (Fig. 1). There are two major types of entheses: fibrous entheses, whereby
tendons and ligaments attach or insert directly to the bone or periosteum, and
fibrocartilaginous entheses, which are zonally organized into tendon or ligament,
fibrocartilage, mineralized fibrocartilage, and bone [134] (Fig. 4). In fibrocartilaginous
53
entheses, this gradient in cell types, ECM and minerals is essential for the transmission
of force during musculoskeletal movement [135]. The intermediate properties of this
tissue arise in development from attachment progenitors (APs), which are established at
the tendon-bone interface through a little-understood mechanism requiring TGFβ
signaling [81] (Fig. 4). Tgfbr2 is expressed in APs and its loss in the jaw (condyle and
angular process) or limb (deltoid tuberosity) results in loss of these key tendon-bone
attachment sites in mice [81, 136, 137]. Though the structure and function of tendon and
ligament fibrocartilaginous entheses appear to be the same, it is unclear whether
developmental mechanisms for enthesis establishment are conserved between these two
types. However, it has been shown that TGFβ signaling plays a crucial role in regulating
bone resorption following traumatic injury to both tendon and ligament entheses, pointing
to at least some conservation of enthesis development between the two tissue types [138-
140].
Comparative transcriptomic analysis together with mouse genetics has identified
that, in the appendicular skeleton, there are global and regional regulators that determine
the correct location of APs along the bone's length. Spatial organization of APs throughout
the limb requires Gli3, whereas proximal versus distal patterning of APs is regulated by
Pbx1/Pbx2 and Hoxa11/Hoxd11, respectively [131] (Fig. 2). Disruption in these global
and regional regulators of AP patterning leads to mislocalization of the bone eminence
attachment sites of both tendon and ligament. Global and regional regulators working
together to fine-tune AP patterning demonstrates modularity in enthesis development.
APs co-express lineage commitment factors for tendon (Scx) and cartilage (Sox9)
and have the potential to adopt either cell fate depending on their location along the
54
tendon-bone axis (Fig. 4). During enthesis formation, APs near the tendon form
fibroblasts of the tendon terminus, whereas those near the bone form chondrocytes, some
of which then ossify into the bone eminence [53] (Fig. 4). Loss of Scx or Sox9 disrupts
tendon-bone attachment development throughout the skeleton [53, 81, 83, 141].
Genome-wide transcriptomic and chromatin accessibility assays of the limb attachment
cells early in their differentiation have shown that APs, as a population, are marked by a
hybrid fibroblast-chondrocyte transcriptional profile with active enhancers shared
between tenocytes or ligamentocytes and chondrocytes [142]. This suggests that the
graded structure of the enthesis is derived from attachment cells with a hybrid tendon-
cartilage or ligament-cartilage cell fate (Fig. 4); however, transcriptional analysis at the
single-cell level will be required to rule out the possibility that the graded structure results
from intermingling of attachment cells of discrete types.
In recent years, several regulators of AP differentiation have been identified. The
Krüppel-like transcription factors KLF2 and KLF4, which bind the proximal regulatory
regions of genes expressed in attachment cells, such as Gli1 and Col5a1, are required
for tendon AP differentiation in the limb [142]. In periodontal ligaments, KLF5 has been
implicated as a key regulator of progenitor cell proliferation and osteogenic differentiation
via activation of both FGF and WNT signaling pathways [143]. Whereas KLF family genes
regulate the overall capacity of APs to differentiate, BMP and FGF signaling regulate the
cell fate choice of APs (Fig. 4). In the limb, BMP4 derived from the tendon tip induces
APs to differentiate into chondrocytes, some of which then undergo endochondral-like
ossification to form the bone eminence [144]. Conditional inactivation of Bmp4 using Scx-
Cre blocks formation of the cartilage anlage prefiguring the bone eminence, despite
55
normal tendon differentiation. Bmp2 and Bmp4 are also necessary for the endochondral
ossification of APs into tendon-embedded sesamoid bones [130]. Although it is
understood that BMP promotes AP differentiation into chondrocytes, it is not clear what
blocks BMP-induced chondrogenesis in APs that differentiate into tendon cells.
During a similar time frame, APs also express the BMP family member Gdf5. Gdf5
lineage tracing marks APs that contribute to linear growth of the enthesis and give rise to
fibrocartilage [145, 146]. The relationship between this Gdf5 lineage and APs that co-
express Scx and Sox9 remains unresolved. However, in the developing joint Sox9
+
cells
give rise to Gdf5
+
joint interzone progenitors that contribute to various joint tissues,
including the menisci, cruciate ligaments, articular cartilage, and intra-articular ligaments
[147, 148]. It has been suggested that the onset and duration of GDF5 signaling may
instruct lineage divergence in the joint, and a similar mechanism may also occur in the
enthesis. The one pathway currently known to promote tendon cell fate in APs is FGF
signaling (Fig. 4). We recently showed that perichondral FGFR2 signaling, activated by
FGF2 from the tendon, promotes tendon cell differentiation in cranial APs by regulating
NOTCH signaling [83]. Upon CNCC-specific deletion of Fgfr2, cranial APs lose DLL1-
NOTCH2 signaling and undergo biased differentiation into chondrocytes over tenocytes.
This indicates that FGF signaling plays a key role in promoting the differentiation of tendon
at the expense of cartilage in the face.
Hh signaling also plays an important role in the cellular complexity and maturation
of tendon-bone and ligament-bone attachment cells. Sonic hedgehog (SHH)
induces Gli1 expression in a subpopulation of newly differentiated attachment cells,
with Gli1 expression maintained postnatally by Indian hedgehog (IHH). These Gli1-
56
expressing cells give rise to the mineralized fibrocartilage in the mature enthesis [146,
149-151] (Fig. 4). Mouse genetics show that Hh signaling is necessary for fibrocartilage
mineralization: loss of the Hh transmembrane effector SMOOTHENED using Scx-
Cre greatly reduces the number of Gli1
+
cells and inhibits their subsequent mineralization
[146, 150, 151]. In addition, ablation of Gli1
+
cells early in postnatal development results
in loss of mineralized fibrocartilage [151]. The lineage relationship between postnatal
Gli1
+
cells and embryonic APs that precede them differs depending on whether the
enthesis remains stationary or migrates during bone growth. In stationary entheses, which
are found at bone ends, Gli1
+
cells are derived from APs; however, in migrating entheses
found along the bone shaft, a separate population of Gli1
+
cells replace APs [149]. The
origin of Gli1
+
cells in migrating entheses remains unclear, but the periosteum seems a
likely source.
Tendon cell fate at the tendon-muscle interface
The structure and mechanical properties of the myotendinous junction (MTJ) rely
on interaction between tendon and muscle via the ECM (Fig. 1). MTJ connections are
unique to tendons because ligament, by definition, facilitates the joining of bones
exclusively. During MTJ formation in Drosophila, myoblasts migrate to sites of integration
and extend actin-based, filopodia-like structures to recognize tendon cells via
thrombospondin-mediated cell adhesion [152]. Interactions between Drosophila tendon
and muscle are mediated by cell-cell contact through integrin heterodimers, as well as
various ECM proteins that can be classified as force transmitting and structural collagens,
scaffolding proteins, or crosslinking proteins. Key ECM proteins that make up the MTJ
include LAMININ, FIBRONECTIN, thrombospondins, and collagens [153, 154]. Recent
57
RNAi screens have expanded the predicted list of genes contributing to MTJ formation
and highlight Tango1 and Fascin as key regulators of collagen deposition and actin
bundling in filopodia, respectively [155, 156].
It remains unclear to what extent insights made in Drosophila can be applied to
vertebrate MTJ formation. In vertebrates, the migratory nature of cells contributing to the
MTJ is known to differ based on both species and anatomical region. Avian hindlimb
myoblasts elongate and adhere to tenocytes already present at attachment sites, yet in
the avian cranium it is the tenocytes that must travel to meet the established muscle [97,
105]. In the zebrafish, cranial tenocytes appear to prefigure regions of future muscle
attachment, with altered tenocyte migration in cyp26b1 mutants resulting in secondary
mislocalization of muscle attachments [157]. It is unknown whether differences in MTJ
patterning across regions and species are due to differing embryonic origins of tendon
and muscle or other, potentially species-specific, factors.
During MTJ development in vertebrates, a careful balance between ECM
deposition and remodeling is maintained in response to mechanical signals. This balance
is achieved through reciprocal interactions between structural proteins, such as LAMININ,
collagens, and matrix metalloproteinases (MMPS) [158-160]. As previously discussed,
both molecular and mechanical factors originating from the muscle play a vital role in
inducing tendon progenitors in the axial skeleton and promoting tendon cell maturation in
the limb and craniofacial complex. Mechanical signals also affect tendon cell
morphogenesis at the MTJ through a TGFβ-driven feedback loop that regulates the
growth and branching of tenocyte projections [114]. Studies in the zebrafish axial skeleton
suggest that interactions of tendon and muscle are mediated by long, microtubule-rich
58
cell projections originating from tenocytes that extend into the intersomitic space during
early development of the MTJ [114, 161]. These projections likely increase the adhesion
of tendon and muscle at the MTJ and provide support and structure to the muscle, a
hypothesis supported by the phenotypes observed in dystroglycan 1 (dag1) mutants
and col22a1 knockdown zebrafish, which model muscular dystrophy and feature
increased muscle detachment [161, 162]. Loss of function of transmembrane protein 2
(tmem2; also known as cemip2) in zebrafish also results in muscle fiber detachment,
coupled with disorganization of key MTJ ECM proteins, such as LAMININ and
FIBRONECTIN, indicating its important role in the regulation of muscle cell-ECM
interactions that facilitate muscle-tendon attachment [163]. Transcription
factors TBX4 and TBX5 originating in muscle connective tissue have also been
implicated in the patterning of tendon and muscle in the mouse limb through modulation
of N-CADHERIN (CADHERIN 2) and β-CATENIN expression [164].
Tendon cell fate in repair
Tendons, ligaments and their entheses are commonly injured and, in adults, heal
through the formation of a persistent fibrovascular scar that does not recapitulate the
developmental process or mechanical properties of the native tissue [165]. Injured
tendons and entheses heal through phases of inflammation, proliferation and remodeling
[166]. During inflammation, the release of growth factors and cytokines initiates
vascularization and recruits immune cells and activates resident stem and progenitor
cells. In the proliferative stage, resident stem and progenitor cells expand and synthesize
a granulation tissue rich in fibroblasts, capillaries, and an ECM network of COL3 [167,
168]. The granulation tissue then undergoes remodeling to form scar tissue that has
59
reduced cellularity and vascularity as well as increased COL1 [167, 168]. As reparative
strategies for tendon, ligament and enthesis, which are largely limited to surgical repair,
fail to recreate the structural and functional characteristics of the native tissue that forms
during development, there is a need to identify and harness the regenerative potential of
endogenous stem and progenitor cells.
Tendon and ligament healing
As discussed, tendon and ligament development appear to share key
morphological and functional characteristics. In fact, a frequently used surgical
intervention to repair torn or injured ligaments is tendon auto- or allografting [169, 170].
For purposes of simplicity, we focus here on the larger body of current research pertaining
to tendon healing with the hope that findings in this field can be extrapolated to improve
understanding of ligament healing. Tendon stem and progenitor cells (TSPCs) were first
identified through in vitro characterization of cells disassociated from the tendon proper
[119]. These cells were identified as expressing surface markers commonly associated
with other stem cells, such as SCA1 (also known as LY6A) and CD44, as well as
tenogenic factors such as SCX. Tissue-isolated TSPCs also exhibit characteristic traits
of stem cells, such as clonal serial passaging and multilineage potential in vitro, and give
rise to tendon-like tissue upon engraftment. More recently, genetic lineage tracing after
tendon injury has begun to characterize potential TSPCs in vivo. Emerging studies reveal
that putative in vivo TSPCs that contribute to tendon repair following injury can arise from
both the tendon proper and sheath, and that age is a major factor for whether healing is
regenerative or fibrotic (Fig. 5).
60
In neonatal mice, transection injury sustained to the Achilles tendon results in the
recruitment of resident Scx-lineage tenocytes, which promote regenerative healing
through new tendon formation [74]. In adult mice, however, the same injury induces
fibrotic healing whereby Scx-lineage cells remain quiescent and an extrinsic population
of αSMA
+
cells contribute to a permanent scar (Fig. 5). Similarly, following full thickness
transection injury in the mouse patellar and supraspinatus tendons, αSMA-lineage cells
are recruited, activate expression of Scx-GFP and become the main contributors to the
healing response [145, 171]. The αSMA
+
cells that contribute to adult tendon repair are
heterogeneous, arising from a subpopulation of resident tenocytes marked by S100a4,
as well as from extrinsic progenitors originating from the tendon sheath [145, 172-175].
Since the αSMA-lineage contributes to multiple cell types within the tendon, only some of
which turn on tenogenic genes during repair, αSMA is not a specific marker for TSPCs
[145, 176] (Fig. 5). Inducible Cre lines that more specifically trace lineages of the tendon
sheath support the idea that the sheath may be a source of TSPCs during repair. An
inducible Cre line driven by tubulin polymerization-promoting protein family member 3
(Tppp3), a developmental marker of tendon sheath cells, marks a population of
SCA1
+
/PDGFRA
+
cells that are recruited to the site of biopsy punch-injury of the patellar
tendon where they activate Scx expression and give rise to tenocytes in a PDGF-
dependent manner [176, 177] (Fig. 5). Interestingly, Tppp3 expression overlaps with that
of Bglap (osteocalcin), a marker typical of osteoblasts, in the sheath of some tendons. In
these tendons, Bglap-lineage cells from the tendon sheath differentiate and contribute to
the population of tenocytes activated during tendon repair [178]. As Bglap and Tppp3 do
not overlap in all tendons, there are likely regional differences in TSPC populations
61
(Fig. 5). Alternatively, heterogeneity in TSPCs might also indicate different states of
commitment along the differentiation pathway.
Evidence that TSPC populations differ in their capacity for repair comes from in
vitro studies comparing the relative healing potential of putative TSPCs from the tendon
proper versus the sheath. Tendon proper-derived TSPCs express higher levels of
tenogenic markers and have greater potential to form tendon-like tissue, whereas sheath-
derived TSPCs express pericyte-like markers, proliferate faster and have increased
potential for myofibroblast differentiation [179-181]. Single-cell RNA-seq analysis is
beginning to reveal the heterogeneity of cells within the tendon proper and tendon sheath;
however, whether this is an output of diversity in TSPCs is unclear [182, 183]. Taken
together, these studies suggest that TSPCs are heterogeneous depending on their
anatomical location, tissue source (tendon proper versus sheath) and degree of lineage
commitment.
Enthesis healing
Like tendons and ligaments, entheses heal via the contribution of multiple cell
lineages. The putative enthesis stem and progenitor cells (ESPCs), primarily
characterized by lineage tracing in repair models, appear to be distinct from TSPCs. For
example, αSMA-lineage cells do not contribute to enthesis healing [145, 184] (Fig. 5).
Evidence suggests that the reparative potential of intrinsic enthesis cells is age
dependent. In adult mice, Scx-, Sox9- and Gli1-lineage cells, which mark tendon,
ligament, and fibrocartilage of the enthesis, show limited involvement during mature
enthesis healing [184, 185]. In contrast, in juvenile mice following needle-punch injury,
Gli1-lineage cells proliferate and contribute to immature enthesis healing through a
62
regenerative process that more closely mimics normal development [185] (Fig. 5). This
suggests that the adult enthesis healing process could potentially be enhanced by
identifying methods to mimic the developmental process.
Although cells intrinsic to the adult enthesis have poor capacity for repair, recent
studies in mice suggest that extrinsic cells from the adjacent bone and tendon are a
source of ESPCs during enthesis healing. Following enthesis reconstruction surgery, in
which tendon grafts are inserted into bone tunnels, Gdf5-lineage cells from the underlying
bone proliferate, infiltrate the tendon graft, and give rise to a zonal tendon-bone
attachment [186] (Fig. 5). Following surgical repair of detached tendons, Axin2-lineage
cells from the tendon proper also contribute to healing, although they promote formation
of a fibrovascular scar rather than recapitulating the graded structure of the native
attachment [184].
Conclusions and future directions
In this review, we have discussed a number of mechanisms that regulate tendon
and ligament cell fate in the context of both development and repair. Recent advances in
our understanding of these mechanisms have opened several interesting questions for
future investigation. For example, mechanisms regulating tendon and ligament cell fate
in development feature regional differences in the necessity of transcription factors and
dependence on inductive signals from muscle versus cartilage. However, it is not yet
known how regional and universal developmental programs are integrated to organize
the musculoskeletal system, or the extent to which developmental insights learned in one
anatomical region can be applied to others. Additionally, tendon cell identity shows
evidence of plasticity and reversibility, but it is not yet fully understood how or what
63
transcriptional factors can function to actively induce tendon or ligament cell fate while
inhibiting osteoblast and chondrocyte cell fate. There is also a high likelihood that key
members of these transcriptional networks have not yet been identified. Additional
information is needed to determine the similarities and differences between tendon and
ligament cells in a developmental, functional, and reparative capacity. For example, can
a close study of joint development reveal regulators that are unique to ligament
development [187, 188]? To what extent are the factors regulating TSPCs conserved
between anatomical regions and between organisms with differing regenerative
potential? Can TSPCs harvested in one location be relied upon for tendon repair in
another location? Answers to these questions will help determine whether repair and
tissue engineering approaches need to be designed with regional specificity in mind.
Understanding tendon cell identity using single-cell sequencing technologies that
allow for RNA expression profiling and chromatin accessibility will be a vital first step to
begin to address many of the questions proposed here about tendon cell fate. For
example, single-cell technologies will be able to determine whether the graded connective
tissue within tendon-bone attachments arises from intermingling of canonical cell types
(tenocyte, osteoblast, chondrocyte), hybrid cell types that feature mixed properties, or a
combination of both. Single-cell technologies will allow for a better resolution of the
similarities and differences between tendon and ligament cells, which will in turn affect
our clinical approaches to injury and repair. A better understanding of the molecular
relationship between these cell types would allow for in vivo studies that advance our
knowledge of the mechanisms regulating development and repair of ligament and
ligament-bone attachments.
64
Studies in model organisms such as mouse and zebrafish have significantly
advanced our understanding of tendon cell differentiation and maintenance, yet we lack
a complete picture of how tendon cell fate is regulated. By focusing research on human
disorders that feature tendon and ligament defects, such as joint laxity, dislocations, and
contractures, it may be possible to identify key genes and mechanisms regulating the
developmental process. Examples of such disorders include geleophysic dysplasia,
which features joint contractures and shortened limbs due to tendon defects. Recent work
has identified Adamtsl2 as the likely affected gene, which had not traditionally been
studied as a regulator of tendon and ligament cell identity [189, 190]. Additionally,
disorders such as progressive osseous heteroplasia (POH) [191] and McCune–Albright
syndrome [192] feature metaplasia of bone and fibrous connective tissue caused by loss-
and gain-of-function mutations in Gnas, respectively. The opposing phenotypes of these
disorders (ossification of fibrous tissue in POH and fibrous metaplasia of bone in
McCune–Albright syndrome) indicate a key role of GNAS in tendon cell identity and
maintenance that is not yet fully understood [193, 194]. Mechanistic studies of congenital
disorders with tendon and ligament defects such as these are likely to reveal new insights
into tendon and ligament cell identity and development.

65
Figure legends
Figure 1: Tissue morphology of tendon, ligament, and musculoskeletal junctions
of the human elbow.
(Clockwise from top right) Tendons (which connect muscle to bone) and ligaments
(which connect and stabilize bones) share similar properties but have slightly different
orientation of collagen fibrils; whereas tendons have parallel arrangements of fibrils,
ligament fibrils can overlap and cross. The tendon enthesis features graded, intermediate
tissue types that facilitate the transmission of force from tendon to bone, whereas the
myotendinous junction facilitates the joining of tendon and muscle through interactions of
ECM proteins.

Figure 2: Embryonic origins and molecular regional regulators of tendon and
ligament progenitors in the developing mouse.
Tendons and ligaments are derived from mesenchyme of multiple embryonic
origins with cranial tendons and ligaments originating from cranial neural crest cells
(CNCCs; blue), limb tendons and ligaments from lateral plate mesoderm (yellow), axial
tendons from paraxial mesoderm (pink), and neck and shoulder tendons and ligaments
from a mixture of all three (green). In addition to their multiple origins, tendon and ligament
progenitors require regional mechanisms for their establishment.

Figure 3: Tendon progenitor induction, differentiation, maintenance, and
maturation in the mouse limb.
Induction of Scx in tendon progenitors relies on TGFβ signaling during embryonic
development. Later, tendon cells differentiate and multiply in the presence of mechanical
stimulation from the muscle, as well as undetermined secreted factors from surrounding
tissue, including cartilage. Various inputs are required for both the establishment and
maintenance of the tendon cell fate. E, embryonic day; P, postnatal day.

Figure 4: Regulation of cell fate during development of the tendon-bone attachment
in the mouse limb.
The enthesis is derived from attachment progenitors (APs), which have the
potential to form endochondral bone, fibrocartilage, and tendon. (A) Tgfβ is necessary to
establish APs, which co-express Scx and Sox9. Once established, APs give rise to either
Scx
+
tenocytes, which make tendon, or Sox9
+
chondrocytes, which make fibrocartilage
and bone. Some APs may maintain co-expression of Scx and Sox9 and acquire a hybrid
tenochondral phenotype. (B) The APs form as a secondary condensation atop the primary
cartilage anlagen at the site of tendon insertion at E12.5. (C) By E13.5, the graded nature
of the enthesis begins to emerge; however, it is not known if the intermediate tissue
forming at the tendon-bone interface is made from hybrid tenochondral cell type (cells
that co-express Scx and Sox9) and/or from discrete cell types (tenocytes and
chondrocytes) that intermingle and/or are separated by a cellular boundary. (D) By E14.5-
15.5, the graded enthesis has formed, connecting tendon to bone. E, embryonic day.

66
Figure 5: Cellular lineages contributing to tendon and enthesis repair in mice.
Injuries to the tendon and enthesis heal through the formation of a persistent
fibrovascular scar that does not recapitulate the native morphology or mechanical
properties of the original tissue. Cells contributing to tendon repair originate from the
tendon proper (green cell) and the tendon sheath (orange cell), whereas cells contributing
to enthesis healing can be sourced from either tendon or bone (beige cell). A few cell-
specific markers that characterize putative tendon stem/progenitor cells (TSPCs) or
enthesis stem/progenitor cells (ESPCs) have been identified. Asterisks indicate markers
found in tendon sheath-derived cells.

67
Figure 1:

68
Figure 2:

69
Figure 3:

70
Figure 4:

71
Figure 5:

72
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86

Abstract (if available)

Abstract The calvarial bones of the infant skull are linked by transient fibrous joints called sutures and fontanelles. Numerous genetic disorders present with craniofacial deformities when sutures fuse prematurely (craniosynostosis), or fontanelles remain patent. The anterior fontanelle is a broad area of connective tissue that forms at the apex of the embryonic calvaria and is ultimately replaced by the posterior frontal suture. However, the ultimate fate of the AF connective tissue and the extent to which it plays an active role in frontal suture development has remained unclear. We find that Fgfr2 is necessary for closure of the AF and subsequently, PFS formation and fusion. We show that the AF is occupied by a heterogeneous population of Scx+ connective tissue-like progenitors, and that loss of FGFR2 signaling blocks their differentiation into bone, cartilage, and ligament-like connective tissue, preventing normal PFS fusion. Analysis indicates that Fgfr2 regulates the differentiation of Scx+ cells into osteoblasts and the formation of Sox9+ chondrocytes through downregulation of WNT signaling. Correspondingly, genetic elevation of WNT signaling in Scx+ cells in mice recapitulates the patent AF phenotype seen in Fgfr2 mutant mice. Together, these results suggest a key role for Fgfr2 in orchestrating divergent cell fate decisions of AF connective tissue to promote formation and closure of the PFS. Overall, the results presented in this study offer further insight into regional differences in the development of sutures that are prefigured by fontanelles.

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Creator Bobzin, Lauren Grace (author)

Core Title FGFR2 Regulates Connective Tissue Development in the Skull

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Development, Stem Cells and Regenerative Medicine

Degree Conferral Date 2023-12

Publication Date 09/01/2023

Defense Date 08/30/2023

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

Tag FGFR2,fontanelle,OAI-PMH Harvest,suture,Wnt

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Advisor Mariani, Francesca (committee chair), Maxson, Robert (committee member), Merrill, Amy (committee member), Xu, Jian (committee member)

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