Spatial and Temporal Expression of Molecular Markers
and Cell Signals During Normal Development
of the Mouse Patellar Tendon
Chia-Feng Liu, Ph.D.,1Lindsey Aschbacher-Smith, M.S.,1Nicolas J. Barthelery, M.S.,1
Nathaniel Dyment, B.S.,2David Butler, Ph.D.,2and Christopher Wylie, Ph.D.1
Tendon injuries are common clinical problems and are difficult to treat. In particular, the tendon-to-bone in-
sertion site, once damaged, does not regenerate its complex zonal arrangement. A potential treatment for tendon
injuries is to replace injured tendons with bioengineered tendons. However, the bioengineering of tendon will
require a detailed understanding of the normal development of tendon, which is currently lacking. Here, we use
the mouse patellar tendon as a model to describe the spatial and temporal pattern of expression of molecular
markers for tendon differentiation from late fetal life to 2 weeks after birth. We found that collagen I, fi-
bromodulin, and tenomodulin were expressed throughout the tendon, whereas tenascin-C, biglycan, and car-
tilage oligomeric protein were concentrated in the insertion site during this period. We also identified signaling
pathways that are activated both throughout the developing tendon, for example, transforming growth factor
beta and bone morphogenetic protein, and specifically in the insertion site, for example, hedgehog pathway.
Using a mouse line expressing green fluorescent protein in all tenocytes, we also found that tenocyte cell
proliferation occurs at highest levels during late fetal life, and declines to very low levels by 2 weeks after birth.
These data will allow both the functional analysis of specific signaling pathways in tenocyte development and
their application to tissue-engineering studies in vitro.
tenocytes, and a high content of dense, highly polarized
extracellular matrix material. Tendons connect and transfer
force between the muscles and bones of the body. Tendon
injuries are common, and hard to repair.1–4Treatments
such as steroids reduce the pain and inflammation of ten-
don injury, but do not repair the tendon.5Surgical repair
often leads to chronic pain,6,7and tendon grafts often lead
to donor site morbidity. Recently, attention has focused on
the use of tissue-engineering approaches to generate re-
placement tendon tissue that will improve healing.8–10
However, the generation of correctly differentiated ten-
don tissue in culture from stem cell populations will re-
quire a detailed understanding of the normal gene
expression pattern of differentiating tenocytes, and how
this is controlled by intercellular signaling during normal
tendon development in vivo. Molecular markers of tendon
differentiation are needed to assay the success of tissue-
endons are a unique component of the musculo-
skeletal system. They contain differentiated cells, the
engineering approaches, and the normal spatial and tem-
poral pattern of their appearance. This information is
lacking for most tendons in the body.
Many cell signaling pathways, such as fibroblast growth
factor (FGF), bone morphogenetic protein (BMP), and
transforming growth factor beta (TGFb) signaling, have been
shown to be involved in the development of tendons. FGF
signaling is required for the specification of axial tendons11–13
as defined by the presence of Scleraxis (SCX)-expressing
cells.14In the limb tendons, the surface ectoderm is required
for the appearance of SCX-expressing cells.14,15It also has
been shown that BMP signaling inhibits the formation of
SCX-expressing cells, while at the same time stimulating the
formation of cartilage. In addition, TGFb signaling is essen-
tial for maintenance and recruitment of tendon progenitors
cells during embryonic stages.16
Many details of these processes are not clear. For example,
the signals that initiate the formation of limb tendons are
unknown. Once specified to form tenocytes, as defined by
the expression of SCX, it is not known what precise temporal
and spatial pattern of signals controls tenocyte proliferation
1Division of Developmental Biology, Cincinnati Children’s Hospital Research Foundation, Cincinnati, Ohio.
2Biomedical Engineering Program, School of Energy, Environment, and Biological and Medical Engineering, University of Cincinnati,
TISSUE ENGINEERING: Part A
Volume 18, Numbers 5 and 6, 2012
ª Mary Ann Liebert, Inc.
and differentiation. In fact, the precise temporal and spatial
pattern of appearance of tenocyte markers, and the degree to
which marker expression differs in the different specialized
regions of the tendon (e.g., mid-substance vs. insertion site),
have not been documented for most tendons. To create ten-
dons from stem cells in culture, which is one major goal of
tissue engineering, it will be necessary to understand the
spatial and temporal pattern of tenocyte differentiation
in vivo and the signaling pathways that control this process.
Although other animal models, such as rabbits and rats,
have been used to study tendon repair, these are not genetic
models and lack the reporter lines and targeted mutagenesis
lines that are readily available in the mouse. However, only
limited descriptions of tendon morphogenesis have been
published in the mouse.14,17–19Therefore, to take advantage of
genetic manipulations possible in the mouse, we have set out
to develop the murine patellar tendon as a model. The goal of
the present study is to establish a spatial and temporal map of
the appearance of some major tenocyte markers in the patellar
tendon in vivo, and to map the changes in activity of major
signaling pathways that accompany these changes in gene
expression. This will allow both the mechanistic analysis of
normal tendon development and the application of this
knowledge to the generation of tendon tissue in vitro.
Materials and Methods
Knee joints from CD-1?(Charles River), Gli1-LacZ (The
Jackson Lab Laboratory, Strain: Gli1tm2Alj), and scleraxis-green
fluorescent protein (ScxGFP) mice20were collected at four
specific stages of patellar tendon development; embryonic day
seventeen and half (E17.5), and postnatal days 1, 7, and 14 (P1,
P7, and P14). Three to five CD-1 and ScxGFP samples from
studies, and three Gli1-LacZ mice from each time point were
used for X-gal staining. Detailed experimental procedures are
were reviewed and approved by the Institutional Animal
Care and Use Committee at CCHMC and were performed in
Knee joints were freshly embedded in OCT (Tissue-Tek)
and then frozen immediately in liquid nitrogen. The em-
bedded tissues were cryostat sectioned at 10mm thickness,
then mounted onto slides. Slides were fixed in 2% parafor-
maldehyde (PFA), washed in phosphate-buffered saline
(PBS), and then stained using hematoxylin and eosin (H&E)
according to standard procedures.21Slides were dehydrated
and then mounted using Mounting Medium Xylene (Fishier
Knee joints were isolated and embedded in OCT and then
frozen immediately in liquid nitrogen. For ScxGFP samples,
tissues were fixed in 4% PFA for 2–6h and then washed with
PBS. The samples were cryostat sectioned at 10mm thickness.
For unfixed samples, sections were fixed with 2%–4% PFA
for 10min or with 2% Trichloroacetic acid for 6min and
washed in PBS and permeabilized with PBST (0.1% Trion
X-100 in PBS). Slides were then incubated in blocking solu-
tion (10% donkey or goat serum, 0.1% Triton X-100, and 4%
bovine serum albumin in PBS) for 1h at room temperature,
then incubated in primary antibodies (see below) at 4?C
overnight. The next day, sections were washed thrice for
10min each with PBST and PBS followed by incubation in
the corresponding secondary antibodies. After secondary
antibody treatment, sections were washed with PBST and
PBS thrice for 10min, counterstained with 4¢,6-diamidino-2-
phenylindole, and mounted in mounting medium. For color
development, sections were treated with diamino benzidine
(DAB) substrate using Metal Enhanced DAB Substrate
Kit (Thermo Scientific, Cat No.34065). After washing with
PBS, slides were counterstained with nuclear fast red
(VECTASHIELD?), dehydrated through a graded series of
ethanols, and mounted in Mounting Medium Xylene (Fishier
The dilutions and sources of primary antibodies used
were as follows: anti-collagen I (1:200; Novus Biological
Research), anti-Tenomodulin (TNMD) (1:50; Santa Cruz),
anti-Bigylcan (1:100; AbCam), anti-cartilage oligomeric pro-
tein (COMP) (1:50; AbCam), anti-p-FGFR1 (1:50, antibodies
fibromodulin (FMOD) (1:200; AbCam), anti-Ki67 (1:500;
Abcam), and anti-p-Smad1/5/8 (1:100; Cell Signaling). All
the secondary antibodies were purchased from Jackson Im-
munoResearch Laboratories, Inc. and used according to the
Image capture and semi-quantitative analysis
A ZEISS AxioPlan 2 microscope (Carl Zeiss MicroImaging
LLC) was used for imaging. Images were acquired at 20·
using the same exposure settings for all samples within each
experimental group. The mean intensity of fluorescence was
measured using the outline tools and measurement in the
AxioVision Rel 126.96.36.199 software (Carl Zeiss MicroImaging
LLC). A total of 12 slides from 3 animals at 4 stages were
analyzed per immunostaining within each experiment
group. Data were analyzed using one-way analysis of vari-
ance (ANOVA) followed by an LSD test for pair-wise com-
parisons using SPSS (IBM, SPSS Statistics 19).
Sections were fixed with 2% PFA for 2min at 4?C, washed
in PBS, X-Gal washing buffer (2mM MgCl2, 0.02% NP-40 in
PBS), and placed in X-gal reaction substrate containing 2mM
MgCl2, 0.02% NP-40, 3.5mM K ferrocyanide, and 3.5mM K
ferricyanide overnight at 37?C in the dark. After washing
with X-Gal washing buffer, slides were counterstained with
nuclear fast red (VECTASHIELD), dehydrated through eth-
anol gradient, and mounted with Mounting Medium Xylene
Cell proliferation counting
Knee joints were obtained from three CD-1 animals for
each time point (E17.5, P1, P7, and P14). Samples were
processed according to the immunohistochemistry proce-
dure just described. Ten random sections were selected from
NORMAL DEVELOPMENT OF THE MOUSE PATELLAR TENDON599
a total of 60 sections of each knee joint for imaging. The
pictures were taken with a ZEISS AxioPlan 2 microscope
(Carl Zeiss MicroImaging LLC). The number of cells in
the cell cycle in each section was obtained by counting Ki67-
positive cells. Data were analyzed using one-way ANOVA
followed by a Tukey test for pair-wise comparisons using
SPSS (IBM, SPSS Statistics 19).
The histology of the developing patellar tendon
Four time points were used in this study: starting at E17.5,
after the patellar tendon has become clearly visible as a dis-
crete structure in the embryo but before birth, and ending at
P14 when the animal is fully motile, and, therefore, the tendon
is functional. Figure 1 shows representative sagittal sections of
patellar tendons from these four stages in their differentiation
(E17.5, P1, P7, and P14), all stained with H&E. Higher mag-
at each stage are shown in Figure 1E–L. At E17.5 (Fig. 1A, E, I),
the cells in both regions of the tendon are rounded, and sepa-
rated by relatively little extracellular matrix. There are no ob-
vious differences in cell morphology or intensity of staining
(Fig. 1B, F, J), there are clear differences in cell morphology
between the midsubstance and insertion site. In the insertion,
site cells are becoming aligned into rows of approximately
square cell profiles, and their nuclei are rounded. In the mid-
substance, cell boundaries are indistinct, but the nuclei are be-
coming elongated. The cells in both regions are becoming
clearly more separated from each other by the increasing ex-
tracellular matrix. At P7 (Fig. 1C, G, K) and P14 (Fig. 1D, H, L),
become progressively more elongated, and the extracellular
matrix/cell ratio increases. In the tibial insertion site, there is
also a continued increase in matrix. However, the cells remain
rounded in shape and arranged in linear arrays. We conclude
from H and E staining that histological differentiation of the
matrix and its spatial differentiation into midsubstance and
insertion site, starts in the neonatal period in the mouse.
Molecular tendon markers expressed
throughout the tendon
Figure 2 shows the expression pattern of several tendon
markers that start to accumulate before birth, and are syn-
insertion site. These included type I collagen (COL1) (Fig. 2A–
E), the proteoglycan FMOD (Fig. 2F–J), and the glycoprotein
TNMD (Fig. 2K–O). All these markers accumulated through-
any obvious quantitative differences in expression along the
tendon length, whereas the expression of TMOD was reduced
significantly in the insertion site starting at P7 (Fig. 2O). The
expression of these should, therefore, be controlled by cell
signals occurring along the whole length of the tendon.
Molecular markers expressed differentially
in the tendon
The expression of three tendon markers that are asym-
metrically expressed in the tendon is shown in Figure 3. In-
terestingly, they are also expressed in different temporal
patterns. The extracellular matrix glycoprotein tenascin-C
(TNC) was found to be expressed only in the insertion site at
E17.5 (Fig. 3A, E). After birth, an increasing expression was
seen in the mid-substance. However, the protein was con-
centrated at all stages in the insertion site (Fig. 3B–E). The
proteoglycan biglycan (BGN) was synthesized in all regions of
the tendon at all stages examined (Fig. 3F–J). However, its
expression became more concentrated in the insertion site at
P7 and P14 (Fig. 3H–J). A third pattern was seen for COMP,
which was expressed at low levels throughout the tendon
until P7, when it became concentrated at the insertion site.
These data show that the initial differentiation of the tendon
into mid-substance and insertion site occurs at the earliest
stages of its development, but is progressive, with new tendon
markers either appearing, or becoming more concentrated in,
the insertion site during the 2 weeks after birth. The fluores-
cence images were subjected to semi-quantitative analysis
using the Zeiss Axiovision software for pixel intensity mea-
surements. The data are shown for TNC and BGN in Figure
3E and J. High levels of staining in the adjacent tissues at early
stages confounded quantitation of COMP expression.
Signaling pathways active during patellar
The activation of several major signaling pathways was
examined, to identify any potential correlations between
these and the temporo-spatial patterns seen in tendon dif-
ferentiation. Cells responding to TGFb and BMP signaling
were identified using antibodies against phosphorylated
Smad3 (PSmad3)and phosphorylated
(PSmad1/5/8) respectively. Cells responding to FGF sig-
naling were identified using an antibody against the acti-
vated (phosphorylated) form of FGF receptor 1. Cells
responding to Wnt signaling were identified using the Top-
Gal strain of mice.22Cells responding to hedgehog (Hh)
signaling were identified using the Gli1-b-galactosidase
strain of mice (Gli1-LacZ).23
Canonical Wnt signaling was not seen at any stage (data
not shown). All cells in the tendon were found to be re-
sponding to TGFb and BMP signaling at all stages examined
(Fig. 4A–H, Q). FGF-responding cells were present in both
regions of the tendon from E17.5 to P7. However, there was
reduced antibody staining in the midsubstance at P14 and in
the insertion site at P7 and P14. (Fig. 4I–L, Q), thus sug-
gesting reduced or absent FGF signaling activity at later
postnatal stages. Most interesting was the distribution of Hh-
responding cells. At E17.5, a small number of cells at the
insertion site were seen to be responding to Hh (Fig. 4M, Q).
The staining increased in both intensity and in the number of
positive cells at P1 (Fig. 4N, Q). At P7 and P14, the number of
positive cells decreased to very low levels (Fig. 4O–Q). These
data show that cells of the insertion site are responding to Hh
signals at precisely the time at which we see the first mo-
lecular marker of the insertion site (TNC) appearing, and
suggest a role for Hh signaling in this process.
The timing of cell proliferation during patellar
Tendon growth during late fetal and early postnatal life
could be due to cell proliferation, the accumulation of
600 LIU ET AL.
and eosin stained mid-
sagittal sections of the mouse
patellar tendon at E17.5 (A, E,
I), P1 (B, F, J), P7 (C, G, K),
and P14 (D, H, L). Higher
magnification images of the
midsubstance are shown in
E–H and of the tibial inser-
tion sites in I–L. P, patella; T,
tibial. Scale bar=100mm.
E17.5, embryonic day
seventeen and half; P1, P7,
P14, postnatal days 1, 7, 14.
NORMAL DEVELOPMENT OF THE MOUSE PATELLAR TENDON601
extracellular matrix during differentiation, or both. To assay
the spatiotemporal pattern of cell proliferation, we used the
Ki67 antibody, which stains the nuclei of cells still in the cell
cycle.22To distinguish between tendon progenitor cells and
connective tissue cells of the endo-, peri-, and epitenons, we
used the ScxGFP reporter mouse, which expresses GFP in all
cells that express the molecular marker of tendon precursor
cells SCX.14Ki67 positive cells were found in the mouse
patellar tendon at all four time points. However, their
number and distribution was different at each stage. There
were more Ki67 positive cells in the tendon at E17.5 and P1
than at P7 and P14 (Fig. 5E). In addition, more of these cells
were double positive (Ki67 and ScxGFP) (Fig.5A, B). By P7
and P14, most of the Ki67-positive, proliferating cells were
not positive for ScxGFP, thus indicating that the cycling
cells by P7 were not tenocytes (Fig. 5C, D). Most of the Ki67
cells at P7 and P14 were located in the epitenon. These data
show that by P7 there are very few tenocytes (as defined
by the expression of SCX) that are still in the cell cycle.
This suggests either that they can re-enter the cycle after
tendon injury, or that cells that repair tendons after injury
arise from the connective tissue cells of the tendon, such as
bromodulin, and K–N show tenomodulin expression using specific antibody staining. Specific staining in each case is red,
and blue shows nuclear staining using DAPI. E, J, and O show semi-quantitation of the antibody staining by measurement of
pixel intensity. Scale bar=100mm. Error bar represents standard error, and the asterisk represents statistical significance
(p<0.05). DAPI, 4¢,6-diamidino-2-phenylindole.
Shows tenocyte markers expressed throughout the patellar tendon. A–D show type I collagen, F–I show fi-
602 LIU ET AL.
the epitenon, rather than the tenocytes themselves. The data
also show that most, if not all, tenocytes, exit from the cell
cycle during the time of major deposition of ECM, and, thus,
A fundamental understanding of normal tendon devel-
opment is needed for any future biologically based treat-
ments of tendon injuries. The current studies are aimed at
producing information that will be useful in the identifica-
tion of novel strategies for potential biological therapies. In
particular, we have identified spatial and temporal land-
marks of mouse patellar tendon proliferation and differen-
tiation, which can be used to assess the degree of tendon
healing, and some of the signaling pathways that may con-
Tendons contain highly abundant ECM glycoproteins and
proteoglycans, which are essential for both organization of
the tendon and its healing. The timing and position of ap-
pearance of these in the tendon can be used as molecular
markers for healing, or for attempts to develop tendons from
biglycan, and K–N show COMP staining using specific antibodies. In each panel, red is specific stain, and blue shows cell
nuclei stained using DAPI. Scale bar=100mm. E and J show semi-quantitation of the antibody staining by measurement of
pixel intensity. Scale bar=100mm. Error bar represents standard error, and the asterisk represents statistical significance
(p<0.05). COMP, cartilage oligomeric protein.
Shows tenocyte markers expressed predominantly in the tibial insertion site. A–D show tenascin-C, F–J show
NORMAL DEVELOPMENT OF THE MOUSE PATELLAR TENDON603
stem cells in culture. In this work, we found that COL1,
FMOD, and TNMD were present throughout the developing
mouse patella tendon; whereas COMP, TNC, and BGN were
concentrated at the insertion site. COL1 is the most abundant
protein in the tendon and is important for its strength.24
Therefore, it is logical to see the dramatic increase in COL1
during the first 2 weeks after birth, when the patellar tendon
becomes functional. FMOD and TNMD are thought to con-
tribute to the distribution and patterning of the tendon
collagen.25,26Null mutations in Fmod mice have shown
irregular fiber bundles and abnormal morphology in Achilles
and tail tendons.27Mice in which Tnmd had been targeted
also developed abnormally organized collagen fibrils.25
These findings indicated that FMOD and TNMD regulate the
composition of fibril bundles in tendons, and correlate well
with the increase in both of these proteins at the same period
as the increase in COL1. Although several studies showed
that type II collagen (COL2) is present at the insertion site,
we did not observe COL2 expression by immunocytochem-
istry during the period studied (data not shown). COL2 was
expressed at high levels by adjacent articular cartilage cells,
thus indicating that the anti-COL2 antibody used in the
current study was able to detect COL2 protein. It may be that
COL2 appears later in the differentiation of the patellar
The localization of BGN at the insertion site of the patella
tendon was consistent with the previous findings using the
Achilles tendon.28COMP expression has been observed in
the tendons/ligaments of both humans and different animal
models29, whereas TNC is known to be expressed in many
tissues during normal embryogenesis as well as in some
cases after tendon rupture.30,31However, to our knowledge,
the current study is the first to show that the COMP and
TNC are concentrated in the insertion site during normal
tendon development at early postnatal stages. Interestingly,
it has been shown that the expression of COMP and TNC are
elevated during the healing process in the tendon-to-bone
insertion site.30,32This correlation between normal tendon
morphogenetic protein signaling (brown stain) using an antibody against pSmad1/5/8. Cell nuclei are stained using fast red.
E–H show cells responding to transforming growth factor beta signaling using an antibody against pSmad2/3. Cell nuclei are
stained with fast red. I–L show cells responding to FGF signaling (brown stain) using an antibody against pFGFr1. Nuclei are
stained with DAPI (blue). M–P show cells responding to hedgehog signaling (blue stain), using the Gli1-LacZ reporter line.
Cell nuclei are stained with fast red. Q show the summary of intensity levels of pSmad1/5/8, pSmad2/3, pFGFr-1, and Gli1-
LacZ. Scale bar=100mm. FGF, fibroblast growth factor.
Shows activated cell signaling pathways in the developing patellar tendon. A–D show cells responding to bone
604 LIU ET AL.
development and healing suggests that COMP and TNC
might play roles during the formation of tendon-to-bone
insertion sites. In addition, ECM proteins such as FMOD,
bigylcan, and tenasin-c have been suggested to be involved
in the maintenance of stem cell niches.33,34Therefore, COMP,
BGN, and TNC might also play roles in regulating niches of
tendon progenitor cells at insertion sites. However, how
these might function in tendon cell generation is still unclear.
It will be important to further investigate their functional
roles in the formation of the insertion site. Further, it will be
important to identify ECM components that have different
expression patterns in the normal developing patellar tendon
in culture models of tendon differentiation, because they
may serve as markers for differentiation of mid-substance vs.
In the current study, we have identified the spatial and
temporal expression patterns of activated cell signaling
pathways during mouse patellar tendon development.
Among these, Hh signaling is particular interesting. We ob-
served that Gli1-LacZ -positive cells appeared in the tibial
NORMAL DEVELOPMENT OF THE MOUSE PATELLAR TENDON605
insertion site, thus indicating that the Hh signaling pathway
was active in the insertion site. We also observed that the
Gli1-LacZ positive cells were present at the cruciate ligament
insertion sites (data not shown), thus suggesting the general
importance of Hh signaling at tendon/ligament-to-cartilage
insertion sites. The maturation of tendon-to-cartilage inser-
tion sites requires contributions from both the tenocyte and
chondrocyte.35In addition, it is known that the differentia-
tion and proliferation of chondrocytes is regulated by Hh
signaling.36,37Therefore, the localization of Hh-responding
cells in the insertion site suggests that the Hh signaling
pathway might control the chondrification of the tenocytes at
the insertion site. There are also spatial and temporal corre-
lations between the expression of BGN and TNC and Hh
responding cells at the insertion site, thus suggesting that the
Hh signaling pathway might regulate the expression of these
ECM proteins in the developing insertion site of mouse pa-
tella tendon. Alternatively, the ECM glycoproteins and pro-
teoglycans may actually control the function of the Hh
signaling pathway in the tendon. It has been shown that
proteoglycans such as BGN can bind to signaling ligands and
control their range of action.38It will be important to learn
the relationship between these ECM components and the Hh
signaling pathway in the insertion site. In addition, we also
found that there were fewer Gli1-LacZ positive cells at P7
and P14 than there were at P1, thus suggesting that the ac-
tivity of Hh signaling was decreased at the insertion site after
P7. It is well known that the tendon-to-bone insertion is a
antibody against Ki67 (red stain). Nuclei are stained blue with DAPI, tenocytes are green, using the ScxGFP reporter mouse line.
Cycling cells in the connective tissue of the epitenon are GFP-negative (arrowheads), whereas tenocytes in the cell cycle are GFP-
positive (arrows). Scale bar=100mm. (E) shows counts of cycling cells in the patellar tendon at each stage (mean–SEM) gained
by counting Ki67-positive cells from 10 random sections from 3 knee joints at each stage. The asterisk represents statistical
significance (p=0.3387 at E17.5, p=0.338 at P1, p=0.001 at P7, and p=0.0001 at P14). ScxGFP, scleraxis-green fluorescent protein.
Shows cells that are still in the cell cycle in the patellar tendon at E17.5 (A), P1 (B), P7 (C), and P14 (D), using an
606 LIU ET AL.
common region for overuse injuries and is hard to repair.
However, why exactly it is so difficult to reconstruct an in-
jured insertion site is still unclear. We suggest two reasons:
first, the loss of Hh signaling at the insertion site at later
postnatal stages, and second, the ossification of the target
cartilage to bone at later postnatal stages. It will be important
to further explore the functional roles of Hh signaling path-
way in the differentiation of the insertion site as well as its
role in tissue repair. Two Hh ligands; sonic Hh and Indian
Hh, have been shown to regulate the development of the
musculoskeletal system.39It will be important to learn which
ligand is functional at the insertion site.
Many studies have shown that BMP, FGF, and TGFb
signaling are important during tendon development, in-
cluding the initiation of tendon cell fate.40However, how
these signals regulate tendon differentiation is not fully un-
derstood. Here, we show that TGFb and BMP signaling were
active throughout the period studied and throughout the
whole tendon. FGF signaling is responsible for the initiation
of tendon development during embryogenesis.13,15,41In the
current study, we found that FGF signaling was decreased at
P14 (Fig. 4E–H), thus suggesting that it may not be essential
after the initial period of tendon differentiation is over.
Several studies have shown that the healing process of ten-
dons and ligaments can be enhanced by FGF,42–44which
might therefore re-awaken a developmental pathway in the
tendon. Although the chief mechanism among these is not
clear, it will be important to compare activation of FGF sig-
naling during tendon healing with its timing of activation
during the normal development of tendon.
Although we did not find any evidence to support the
presence of canonical Wnt signaling in the developing mouse
patellar tendon, our results could not exclude the involve-
ment of noncanonical WNT signaling in regulation of tendon
growth. Further analysis of noncanonical WNT signaling in
tendon development is needed.
This study shows that Scx-expressing cells are still in the
cell cycle at E17.5 and P1. However, by P7 and P14, most of
the cycling cells were not ScxGFP positive, thus suggesting
that most tenocytes have exited the cell cycle by this time.
Occasional Scx-positive cells still in the cycle can be identi-
fied at the two later time points. It is not yet clear how this
data might fit with the finding that cycling tenocytes are
generated from tendon cell cultures, or with the suggestion
that cells from the epitenon and endotenon might contribute
to the healing process of tendon.45It is not yet known
whether cells in the connective tissues of the tendon can
activate Scx expression and become tenocytes, It is also not
clear whether the repair process of an injured tendon gen-
erates a fibrotic scar,46,47or a tissue containing genuine te-
nocytes. It is most likely that healing cells in the tendon
require signals that are present during normal development
of the tendon for the generation of genuine tendon at the
wound site. The markers identified in this study and the
signaling pathways identified as active during normal de-
velopment should be useful in gaining the answers to these
The authors are grateful to Dr. Ronen Shweitzer for the
ScxGFP mice. They thank all the members of the Wylie-
Heasman lab and the Butler lab. This work is supported by
NIH grants AR46574-10, AR56943-02, and an IGERT training
grant from NSF (#0333377).
No competing financial interests exist.
1. Klepps, S., Bishop, J., Lin, J., Cahlon, O., Strauss, A., Hayes,
P., et al. Prospective evaluation of the effect of rotator cuff
integrity on the outcome of open rotator cuff repairs. Am J
Sports Med 32, 1716, 2004.
2. Jost, B., Pfirrmann, C.W., Gerber, C., and Switzerland, Z.
Clinical outcome after structural failure of rotator cuff re-
pairs. J Bone Joint Surg Am 82, 304, 2000.
3. Praemer, A., Furner, S., and Rice, D. Musculoskeletal Con-
dition in the United States. Parke Ridge, IL: American
Academy of Orthopaedic Surgeons, 1999.
4. Sharma, P., and Maffulli, N. Tendon injury and tendinopathy:
healing and repair. J Bone Joint Surg Am 87, 187, 2005.
5. Riley, G. Tendinopathy—from basic science to treatment.
Nat Clin Pract Rheumatol 4, 82, 2008.
6. Austin, J.C., Phornphutkul, C., and Wojtys, E.M. Loss of
knee extension after anterior cruciate ligament reconstruc-
tion: effects of knee position and graft tensioning. J Bone
Joint Surg Am 89, 1565, 2007.
7. Petsche, T.S., and Hutchinson, M.R. Loss of extension after
reconstruction of the anterior cruciate ligament. J Am Acad
Orthop Surg 7, 119, 1999.
8. Juncosa, N., West, J.R., Galloway, M.T., Boivin, G.P., and
Butler, D.L. In vivo forces used to develop design parameters
for tissue engineered implants for rabbit patellar tendon
repair. J Biomech 36, 483, 2003.
9. Butler, D.L., Juncosa-Melvin, N., Boivin, G.P., Galloway,
M.T., Shearn, J.T., Gooch, C., et al. Functional tissue engi-
neering for tendon repair: a multidisciplinary strategy using
mesenchymal stem cells, bioscaffolds, and mechanical
stimulation. J Orthop Res 26, 1, 2008.
10. Juncosa-Melvin, N., Matlin, K.S., Holdcraft, R.W., Nirmala-
nandhan, V.S., and Butler, D.L. Mechanical stimulation in-
creases collagen type I and collagen type III gene expression
of stem cell-collagen sponge constructs for patellar tendon
repair. Tissue Eng 13, 1219, 2007.
11. Kahane, N., Cinnamon, Y., Bachelet, I., and Kalcheim, C. The
third wave of myotome colonization by mitotically compe-
tent progenitors: regulating the balance between differenti-
Development 128, 2187, 2001.
12. Brent, A.E., Schweitzer, R., and Tabin, C.J. A somitic com-
partment of tendon progenitors. Cell 113, 235, 2003.
13. Brent, A.E., and Tabin, C.J. FGF acts directly on the somitic
tendon progenitors through the Ets transcription factors
Pea3 and Erm to regulate scleraxis expression. Development
131, 3885, 2004.
14. Schweitzer, R., Chyung, J.H., Murtaugh, L.C., Brent, A.E.,
Rosen, V., Olson, E.N., et al. Analysis of the tendon cell fate
using Scleraxis, a specific marker for tendons and ligaments.
Development 128, 3855, 2001.
15. Edom-Vovard, F., Bonnin, M., and Duprez, D. Fgf8 tran-
scripts are located in tendons during embryonic chick limb
development. Mech Dev 108, 203, 2001.
16. Pryce, B.A., Watson, S.S., Murchison, N.D., Staverosky, J.A.,
Du ¨nker, N., and Schweitzer, R. Recruitment and maintenance
during muscle development.
NORMAL DEVELOPMENT OF THE MOUSE PATELLAR TENDON607
of tendon progenitors by TGFbeta signaling are essential for Download full-text
tendon formation. Development 136, 1351, 2009.
17. Brown, D., Wagner, D., Li, X., Richardson, J.A., and Olson,
E.N. Dual role of the basic helix-loop-helix transcription
factor scleraxis in mesoderm formation and chondrogenesis
during mouse embryogenesis. Development 126, 4317, 1999.
18. Ito, Y., Toriuchi, N., Yoshitaka, T., Ueno-Kudoh, H., Sato, T.,
Yokoyama, S., et al. The Mohawk homeobox gene is a critical
regulator of tendon differentiation. Proc Natl Acad Sci USA
107, 10538, 2010.
19. Blitz, E., Viukov, S., Sharir, A., Shwartz, Y., Galloway, J.L.,
Pryce, B.A., et al. Bone ridge patterning during musculoskel-
etal assembly is mediated through SCX regulation of Bmp4 at
the tendon-skeleton junction. Dev Cell 17, 861, 2009.
20. Pryce, B.A., Brent, A.E., Murchison, N.D., Tabin, C.J., and
Schweitzer, R. Generation of transgenic tendon reporters,
ScxGFP and ScxAP, using regulatory elements of the scler-
axis gene. Dev Dyn 236, 1677, 2007.
21. Lillie, R.D. Histopathologic Technic and Practical Histochem-
istry, 3rd edition. New York: McGraw-Hill Book Co, 1965.
22. DasGupta, R., and Fuchs, E. Multiple roles for activated
LEF/TCF transcription complexes during hair follicle de-
velopment and differentiation. Development 126, 4557, 1999.
23. Bai, C.B., Auerbach, W., Lee, J.S., Stephen, D., and Joyner,
A.L. Gli2, but not Gli1, is required for initial Shh signaling
and ectopic activation of the Shh pathway. Development
129, 4753, 2002.
24. Liu, S.H., Yang, R.S., al-Shaikh, R., and Lane, J.M. Collagen
in tendon, ligament, and bone healing. A current review.
Clin Orthop Relat Res 318, 265, 1995.
25. Docheva, D., Hunziker, E.B., Fassler, R., and Brandau, O.
Tenomodulin is necessary for tenocyte proliferation and
tendon maturation. Mol Cell Biol 25, 699, 2005.
26. Hedlund, H., Mengarelli-Widholm, S., Heinegard, D., Re-
inholt, F.P., and Svensson, O. Fibromodulin distribution and
association with collagen. Matrix Biol 14, 227, 1994.
27. Svensson, L., Aszodi, A., Reinholt, F.P., Fassler, R., Heine-
gard, D., and Oldberg, A. Fibromodulin-null mice have ab-
normal collagen fibrils, tissue organization, and altered
lumican deposition in tendon. J Biol Chem 274, 9636, 1999.
28. Riley, G. The pathogenesis of tendinopathy. A molecular
perspective. Rheumatology (Oxford) 43, 131, 2004.
29. DiCesare, P., Hauser, N., Lehman, D., Pasumarti, S., and
Paulsson, M. Cartilage oligomeric matrix protein (COMP) is
an abundant component of tendon. FEBS Lett 354, 237, 1994.
30. Pajala, A., Melkko, J., Leppilahti, J., Ohtonen, P., Soini, Y.,
and Risteli, J. Tenascin-C and type I and III collagen ex-
pression in total Achilles tendon rupture. An immunohis-
tochemical study. Histol Histopathol 24, 1207, 2009.
31. Riley, G.P., Harrall, R.L., Cawston, T.E., Hazleman, B.L., and
Mackie, E.J. Tenascin-C and human tendon degeneration.
Am J Pathol 149, 933, 1996.
32. Wurgler-Hauri, C.C., Dourte, L.M., Baradet, T.C., Williams,
G.R., and Soslowsky, L.J. Temporal expression of 8 growth
factors in tendon-to-bone healing in a rat supraspinatus
model. J Shoulder Elbow Surg 16, S198, 2007.
33. Garcion, E., Halilagic, A., Faissner, A., and ffrench-Constant,
C. Generation of an environmental niche for neural stem cell
development by the extracellular matrix molecule tenascin
C. Development 131, 3423, 2004.
34. Bi, Y., Ehirchiou, D., Kilts, T.M., Inkson, C.A., Embree, M.C.,
Sonoyama, W., F. Identification of tendon stem/progenitor
cells and the role of the extracellular matrix in their niche.
Nat Med 13, 1219, 2007.
35. Thomopoulos, S., Genin, G.M., and Galatz, L.M. The de-
velopment and morphogenesis of the tendon-to-bone inser-
tion - what development can teach us about healing. J
Musculoskelet Neuronal Interact 10, 35, 2010.
36. Mak, K.K., Kronenberg, H.M., Chuang, P.T., Mackem, S.,
and Yang, Y. Indian hedgehog signals independently of
PTHrP to promote chondrocyte hypertrophy. Development
135, 1947, 2008.
37. Nie, X., Luukko, K., Kvinnsland, I.H., and Kettunen, P.
Developmentally regulated expression of Shh and Ihh in the
developing mouse cranial base: comparison with Sox9 ex-
pression. Anat Rec A Discov Mol Cell Evol Biol 286, 891, 2005.
38. Yoon, J.H., and Halper, J. Tendon proteoglycans: biochem-
istry and function. J o Musculoskelet Neuronal Interact 5,
39. Mullor, J.L., Sanchez, P., and Ruiz I Altaba, A. Pathways and
consequences: Hedgehog signaling in human disease.
Trends Cell Biol 12, 562, 2002.
40. Liu, C.F., Aschbacher-Smith, L., Barthelery, N.J., Dyment,
N., Butler, D., and Wylie, C. What we should know before
using tissue engineering techniques to repair injured ten-
dons: a developmental biology perspective. Tissue Eng Part
B Rev 17, 165, 2011.
41. Edom-Vovard, F., Schuler, B., Bonnin, M.A., Teillet, M.A.,
and Duprez, D. Fgf4 positively regulates scleraxis and
tenascin expression in chick limb tendons. Dev Biol 247,
42. Chan, K.M., Fu, S.C., Wong, Y.P., Hui, W.C., Cheuk, Y.C.,
and Wong, M.W. Expression of transforming growth factor
beta isoforms and their roles in tendon healing. Wound
Repair Regen 16, 399, 2008.
43. Fukui, N., Katsuragawa, Y., Sakai, H., Oda, H., and Naka-
mura, K. Effect of local application of basic fibroblast growth
factor on ligament healing in rabbits. Rev Rhum Engl Ed 65,
44. Kobayashi, D., Kurosaka, M., Yoshiya, S., and Mizuno, K.
Effect of basic fibroblast growth factor on the healing of
defects in the canine anterior cruciate ligament. Knee Surg
Sports Traumatol Arthrosc 5, 189, 1997.
45. Jones, M.E., Mudera, V., Brown, R.A., Cambrey, A.D.,
Grobbelaar, A.O., and McGrouther, D.A. The early surface
cell response to flexor tendon injury. J Hand Surg Am 28,
46. Gelberman, R.H., Manske, P.R., Vande Berg, J.S., Lesker,
P.A., and Akeson, W.H. Flexor tendon repair in vitro: a
comparative histologic study of the rabbit, chicken, dog, and
monkey. J Orthop Res 2, 39, 1984.
47. Hatano, I., Suga, T., Diao, E., Peimer, C.A., and Howard, C.
Adhesions from flexor tendon surgery: an animal study
comparing surgical techniques. J Hand Surg Am 25, 252,
Address correspondence to:
Christopher Wylie, Ph.D.
Division of Developmental Biology
Cincinnati Children’s Hospital Research Foundation
3333 Burnet Ave.
Cincinnati, OH 45229
Received: June 15, 2011
Accepted: September 21, 2011
Online Publication Date: November 9, 2011
608LIU ET AL.