The extracellular matrix (ECM) confers mechanical strength to
the tissue and provides microenvironment and inductive cues
essential for proper cell proliferation, migration
differentiation. The ECM undergoes continuous remodeling
during animal development as well as during various
pathological processes including wound healing, osteoporosis,
rheumatoid arthritis and cancer. Proteases of the matrix
metalloproteinase (MMP) family (more than 20 members) are
believed to play major roles in the destructive aspects of ECM
remodeling (Egeblad and Werb, 2002; Vu and Werb, 2000).
The importance of several MMP family members in mouse
development has been documented using gene-targeting
techniques (Holmbeck et al., 1999; Itoh et al., 1997; Oh et al.,
2004; Vu et al., 1998; Zhou et al., 2000). A group of
endogenous, secreted MMP inhibitors, termed tissue inhibitors
of metalloproteinases (TIMP1-TIMP4) have been identified,
although their roles in development remain to be elucidated
(Lambert et al., 2004).
RECK (reversion-inducing cysteine-rich protein with Kazal
motifs) was first isolated as a transformation suppressor gene
by cDNA expression cloning (Takahashi et al., 1998).
glycoprotein that negatively regulates at least three members
of the MMP family (MMP2, MMP9 and MT1-MMP) in vitro
and in cultured cells (Oh et al., 2001; Takahashi et al., 1998).
Although RECK is widely expressed in normal human organs
and non-transformed cell lines, it is downregulated in various
tumor tissues and transformed cells (Takahashi et al., 1998).
Restored expression of RECK in tumor-derived cell lines
results in strong suppression of their abilities to invade,
metastasize, and induce angiogenesis (Oh et al., 2001;
Extracellular matrix (ECM) undergoes
remodeling during mammalian development. Although
involvement of matrix metalloproteinases (MMPs) in ECM
degradation has been well documented, how this process is
regulated to allow proper ECM accumulation remains
unclear. We previously showed the involvement of a
membrane-anchored MMP regulator, RECK (reversion-
inducing cysteine-rich protein with Kazal motifs), in
vascular development in mice. Here we report that Reck
mRNA can be detected in developing cartilage in
E13.5~16.5 mouse embryos
upregulated during differentiation of a chondrogenic cell
line ATDC5 in vitro. In the early phase of ATDC5
differentiation, RECK expression stays low, multiple
MMPs are upregulated, and there is ECM degradation at
the sites of cellular condensation. In the later phase, RECK
is upregulated inside the expanding cartilaginous nodules
and is progressively
where type II collagen is accumulated while active ECM
degradation persists along the rim of the nodules.
Constitutive RECK expression suppressed initial cellular
condensation, whereas RECK knockdown suppressed the
later ECM accumulation in the cartilaginous nodules.
These results suggest that RECK expression at the right
place (in the core of the nodules) and at the right time (only
in the later phase) is important for proper chondrogenesis
and that RECK, together with MMPs, plays a crucial role
in regulating dynamic processes of tissue morphogenesis.
Supplementary material available online at
Key words: RECK, MMP, ECM remodeling, Tissue morphogenesis,
Dual effects of the membrane-anchored MMP
regulator RECK on chondrogenic differentiation of
Shunya Kondo1,2, Chisa Shukunami2, Yoko Morioka1, Naoya Matsumoto1,3, Rei Takahashi4, Junseo Oh1,*,
Tadao Atsumi5, Akihiro Umezawa6, Akira Kudo7, Hitoshi Kitayama1, Yuji Hiraki2and Makoto Noda1,‡
1Department of Molecular Oncology, Kyoto University Graduate School of Medicine, Yoshida-Konoe-cho, Sakyo-ku, Kyoto 606-8501, Japan
2Department of Cellular Differentiation, Institute for Frontier Medical Sciences, Kyoto University, 53 Shogoin-Kawahara-cho, Sakyo-ku, Kyoto 606-
3Department of Anatomy and Neurobiology and 4Department of Pathology and Tumor Biology, Kyoto University Graduate School of Medicine,
Yoshida-Konoe-cho, Sakyo-ku, Kyoto 606-8501, Japan
5Antibiotics Laboratory, Discovery Research Institute, RIKEN, 2-1 Hirosawa, Wako, Saitama, 351-0198, Japan
6Department of Reproductive Biology and Pathology, National Research Institute for Child and Health Development, Okura, Setagaya, 157-8535
7Department of Biological Information, Tokyo Institute of Technology, Nagatsuda, Midori-ku, Yokohama, 226-8501, Japan
*Present address: Korea University Graduate School of Medicine, Ansan, Korea
‡Author for correspondence (e-mail: email@example.com)
Accepted 3 January 2007
Journal of Cell Science 120, 849-857 Published by The Company of Biologists 2007
Journal of Cell Science
Takahashi et al., 1998). Positive correlation between residual
RECK expression in tumor tissues and survival of the patients
have been documented in a variety of cancers (Furumoto et al.,
2001; Masui et al., 2003; Span et al., 2003; Takenaka et al.,
2004; Takeuchi et al., 2004). Mice lacking Reck die around
embryonic day 10.5 (E10.5) with defects in angiogenesis (Oh
et al., 2001). In these mutant mice, MMP activity is elevated,
and the amount of type I collagen is greatly reduced. This
phenotype is partially suppressed in mice lacking both Reck
and Mmp2. These findings support the idea that RECK is an
essential regulator of MMPs. Because of the embryonic
lethality of Reck-deficient mice, however, the roles of RECK
in mouse development after E10.5 remain to be elucidated.
In mammals, cartilage is one of the most ECM-rich tissues
where type II collagen and aggrecan are particularly abundant.
Formation of cartilage (or chondrogenesis) is a multi-step
process involving (1) condensation of mesenchymal precursor
cells at the future sites of skeletal elements; (2) differentiation
of mesenchymal precursors in these dense cell masses along a
chondrogenic pathway; (3) deposition of specialized ECM; and
(4) cellular hypertrophy and calcification (Behonick and Werb,
2003; Cancedda et al., 1995; Hall and Miyake, 2000; Knudson
and Toole, 1985). Expression of Mmp2, Mmp9, and Mt1-mmp
in mesenchymal cells surrounding developing cartilage has
been observed (Apte et al., 1997; Reponen et al., 1992;
Reponen et al., 1994), and inhibition of cartilage formation by
a synthetic MMP inhibitor in a mandibular explant culture
system documented (Chin and Werb, 1997). Thus, MMPs seem
Journal of Cell Science 120 (5)
Fig. 1. Abundant expression of Reck in cartilage. (A) Mouse embryo
multiple tissue northern blot (Clontech) was hybridized sequentially
with cDNA probes for Reck and Gapdh (loading control). Each lane
contained 2 ?g of poly(A)+RNA from embryos at the indicated
stage (days after gestation). (B-G) Detection of Reck mRNA in
mouse embryos by in situ hybridization. Sagittal sections of an E13.5
embryo were hybridized with Reck antisense (B) or sense (C)
riboprobe. Magnified views of the boxed areas in B are shown in D
and E (*), respectively. The areas containing ribs (F) and a femur (G)
in a section of E16.5 embryo hybridized with Reck anti-sense probe
are also shown. Northern blot data (not shown) indicated that the
apparently strong signals in the liver (labeled ‘L’in panel B and F)
are non-specific. Bars, 1.28 mm in B and C; 200 ?m in D and E; 320
?m in F and G. (H-J) Detection of Reck mRNA in cultured cells by
RNA blot hybridization. Total RNA extracted from the following
cultured cells was analyzed by RNA blot hybridization using the
Reck cDNA probe. (H) Lane 1: H4-1 (a primary culture derived from
human bone marrow containing the cells capable of differentiating
into chondrocytes) (Imabayashi et al., 2003; Mori et al., 2005). Lane
2: DEC (chondrocytes obtained from human cartilage) (Imabayashi
et al., 2003). Lane 3: the human fibrosarcoma cell line HT1080-
transfected with control vector (negative control). Lane 4: HT1080
transfected with RECK expression vector (positive control).
(I) Lanes 1 and 2: longer exposure of lane 1 and 2 in H, (under the
same conditions as lanes 3~6). Lane 3: KUM9 (multipotential
progenitor cells derived from mouse bone marrow and capable of
differentiating into osteocytes, adipocytes, myocytes and neurons)
(Kohyama et al., 2001). Lane 4: KUSA-A1 (osteoblasts derived from
mouse bone marrow) (Kohyama et al., 2001). Lanes 5 and 6: MDBM
(osteoclast progenitor cells derived from mouse bone marrow) and
RANKL-treated MDBM (mature osteoclasts), respectively
(Takeshita et al., 2000). The amount of total RNA loaded was 10 ?g
for lanes 1-4 and 4 ?g for lanes 5-6. Patterns of ribosomal RNA
bands are also shown (bottom panels). (J) Relative intensity of the
bands on the blot shown in I. The band intensity determined by
densitometry and normalized against the RNA amount is presented
as a bar graph for direct comparison. (K) Temporal expression
pattern of Reck mRNA in ATDC5 cells during chondrogenic
differentiation in vitro. Total RNA (20 ?g) from ATDC5 cells that
had been allowed to differentiate in culture for the indicated times
(days) was subjected to RNA blot hybridization using a Reck cDNA
probe (top panel). The same blot was re-probed with a type II
collagen cDNA (middle panel; Col-II) to monitor chondrogenic
Journal of Cell Science
A role for RECK in chondrogenesis
to be important for ECM remodeling during chondrogenesis
(Chin and Werb, 1997).
The mouse embryonal carcinoma-derived cell line ATDC5
has properties resembling
mesenchymal cells and is able to recapitulate many aspects of
chondrogenesis including cellular condensation, cartilaginous
nodule formation, switching of collagen gene expression from
type I to types II and X, accumulation of cartilage-type
proteoglycans, etc. (Atsumi et al., 1990; Shukunami et al.,
1996). Thus, ATDC5 cells have been widely used for studying
temporal and spatial changes in morphology, gene expression,
etc. during chondrocyte differentiation and for assessing the
effects of various exogenous agents on these processes (Fujii
et al., 1999; Fujita et al., 2004; James et al., 2005; Newman et
al., 2001; Scheijen et al., 2003; Shen et al., 2002; Shukunami
et al., 1997; Shukunami et al., 1998; Woods et al., 2005).
In the present study, we attempted to explore the functions
of RECK after E10.5 using tissue slices prepared from wild-
type mice at later developmental stages and cultured ATDC5
those of undifferentiated
cells. Our data indicate that RECK is critically involved in the
dynamic regulation of ECM remodeling and tissue
morphogenesis during chondrogenesis.
RECK is abundant in developing cartilage
RNA blot hybridization with poly(A)+RNA from whole mouse
embryos at different stages indicated that although Reck
mRNA is barely detectable at embryonic day 7 (E7), it is
clearly detectable at E11 and thereafter (Fig. 1A). The
observed onset of Reck expression is consistent with our
previous finding that Reck-deficient mice died around E10.5
with prominent defects in vascular development (Oh et al.,
2001). In an attempt to address the question of whether RECK
plays any roles in events other than vascular development, we
analyzed the expression of Reck in wild-type embryos at E13.5
and E16.5 by in situ hybridization. At E13.5, Reck-specific
hybridization signals (blue signals in Fig. 1B) were found
ubiquitously in the entire embryo and were especially intense
in cartilaginous tissues, such as Meckel’s cartilage
(Fig. 1D), vertebrae and ribs (Fig. 1E). At E16.5,
intense Reck signals were found in rib cartilage (Fig.
1F) and terminal regions of long bones (Fig. 1G,
To identify the cells expressing RECK, we
analyzed the relative abundance of Reck mRNA in
cultured cells of various origins by RNA blot
hybridization (Fig. 1H-J).Strong Reckexpression was
detectable in the cells of the chondrocytic lineage
Fig. 2. Expression patterns of MMP, collagens and
RECK during cartilaginous nodule formation by ATDC5
cells. (A) Temporal expression patterns of Mmps and
Reck in differentiating ATDC5 cells. Total RNA (20 ?g)
extracted from ATDC5 cells that had been incubated for
the indicated times was analyzed by RNA blot
hybridization using indicated probes. (B) Collagenase
activity expressed by differentiating ATDC5 cells.
ATDC5 cells were incubated for the indicated times and
then overlaid with semi-solid medium containing
reconstituted type I collagen and DQ collagen. After an
additional 24 hours incubation, morphology (DIC) and
collagenolytic activity (green DQ-fluorescence) were
recorded with a confocal microscope. Bars, 100 ?m in
the day-5 and day-10 panels; 200 ?m in the day-24
panel. Arrowheads indicate the rim of a large nodule.
(C) Sensitivity of the DQ-fluorescent signals to an MMP
inhibitor GM6001. The experimental conditions were the
same as in B, day 24 except that the cells were exposed
to GM6001 (100 ?M; bottom panels) or an inactive
analogue (100 ?M; control) for the last 36 hours (i.e. 12
hours before overlay plus 24 hours after overlay). Bars,
100 ?m. (D) Temporal expression pattern of type I
collagen mRNA in differentiating ATDC5 cells. The
same blot used in A was re-probed with a type I collagen
cDNA. (E and F) Localization of type I and type II
collagen at cartilaginous nodules formed by ATDC5
cells. ATDC5 cells incubated for 15 days were fixed and
stained with anti-RECK (red) plus anti-type I collagen
(green; E) or anti-RECK (red) plus anti-type II collagen
(green; F). Reconstituted z-axis images along two cutting
lines (a,b) are shown in the top and right panels,
respectively. Bars, 100 ?m in E,F.
Journal of Cell Science
(Fig. 1H-J, lanes 1 and 2). The level of Reck expression in these
cells was much higher than that in the cells of the osteocytic (Fig.
1I-J, lanes 3 and 4) or osteoclastic lineage (Fig. 1I-J, lanes 5 and
6). Thus, RECK is abundantly expressed in chondrocytes.
To confirm and extend these findings, we examined the
temporal expression patterns of Reck and type II collagen (a
chondrocyte differentiation marker) mRNA in ATDC5 cells.
Interestingly, the amount of Reck mRNA was relatively low in
undifferentiated ATDC5 cells and was progressively increased
as the chondrogenic differentiation proceeded (Fig. 1K).
Cartilaginous nodule formation is accompanied by
differentially regulated MMP activity and RECK
We also examined the temporal expression patterns of three
MMPs, Mmp2, Mmp9, and Mt1-mmp, in differentiating
ATDC5 cells by RNA blot hybridization, since these MMPs
are known to be regulated by RECK. Interestingly, all the genes
were upregulated around day 7 and gradually declined
thereafter, whereas Reck was progressively upregulated (Fig.
2A). Spatial distribution of MMP activity in differentiating
ATDC5 cells was also monitored by in situ zymography using
a cleavage-dependent fluorigenic substrate, DQ-collagen (Fig.
2B). The collagenolytic activity was detectable at sites of
cellular condensation on day 5 until approx. day 10 and
persisted in the rim of cartilaginous nodules at later stages (e.g.
day 24, arrowheads). These fluorescent signals were barely
detectable in the presence of a broad-spectrum MMP inhibitor,
GM6001, suggesting that these signals most likely represent
MMP activity (Fig. 2C).
Type I collagen is a major collagen species expressed in
chondrogenic mesenchymal precursors. Expression of the type
I collagen (?2 chain) gene was also upregulated at day 7 and
declined thereafter (Fig. 2D). Immunofluorescent staining
indicated that on day 15, type I collagen was prominent near
the surface of the nodules (Fig. 2E, green signals;
supplementary material Fig. S1B), whereas chondrocyte-
specific type II collagen could be detected inside the nodules
where RECK was also abundant (Fig. 2F, green and red signals,
respectively; supplementary material Fig. S1, C and A).
These observations suggest that initial cellular condensation
is accompanied by MMP upregulation and low RECK
expression, i.e. a condition favorable for active ECM
remodeling.Bycontrast, the later phase of ATDC5
differentiation is accompanied by increasing RECK expression
inside the nodules and persistent collagenolysis along the rim
of the nodules, a condition favorable for ECM accumulation
inside the cartilaginous nodules and continuous enlargement of
Forced expression of RECK affects cellular
To address the question of whether progressive increase in Reck
expression has any significance in chondrogenesis, we tried to
manipulate RECK expression in ATDC5 cells. First, to study
Journal of Cell Science 120 (5)
Fig. 3. Effects of forced RECK expression on nodule formation by
ATDC5 cells. (A) ATDC5 cells were stably transfected with a
mammalian expression vector pCXN2 (V) (Niwa et al., 1991) or the
vector expressing human RECK (R). Expression of RECK protein
(top panel) and RECK mRNA (middle panel) in the pooled
transfectants harvested under the proliferative conditions was
analyzed by immunoblot assay (30 ?g protein per lane; top panel)
and RNA blot hybridization (10 ?g total RNA per lane; middle and
bottom panels), respectively. Relative band intensity is shown under
each lane. (B) Growth curves of the transfectants. The plating density
at day 0 was 3?104cells per 35-mm dish. The data are average
values from duplicate dishes. (C) Phase-contrast micrographs of the
vector-transfected cells (left panels) and RECK-transfected cells
(right panels) incubated for the indicated times. Bars, 300 ?m.
Arrows indicate compact nodules. (D) Number of foci (areas of cell
condensation) formed by the vector-transfected cells (hatched bars)
and RECK-transfected cells (white bars) after incubation for the
indicated times. The data are given as mean ± s.e.m. of quadrate
dishes. Consistent results were obtained in two separate experiments
using the same set of transfectants and in another experiment using
retroviral vectors for gene transfer.
Journal of Cell Science
A role for RECK in chondrogenesis
the effects of constitutive RECK expression, we generated
ATDC5 cells stably transfected with a vacant expression vector
or the vector containing human RECK cDNA and compared
their behaviors (Fig. 3). Significant increase in RECK
expression was detectable in the RECK-transfected cells (R) as
compared to the control cells (V) under growing conditions
(Fig. 3A). These pooled transfectants showed similar growth
rates before reaching confluence (Fig. 3B, up to day 3). After
this time point, however, RECK-transfected cells began to
show lower cell density than the control (Fig. 3B, day 5 and
thereafter). Between days 5 and 8, the control cells formed
numerous foci, representing active cellular condensation (Fig.
3C, Vector, Day 8), and these foci continued to grow into
cartilaginous nodules (Fig. 3C, Vector, Day 15~36). By
contrast, the RECK-transfected cells formed markedly fewer
foci than the control cells (Fig. 3C, RECK, day 8; Fig. 3D),
and these foci grew into compact nodules (Fig. 3C, RECK, Day
At day 14, the number of spots of Alcian Blue staining,
representing cartilaginous ECM accumulation, was far fewer
in RECK-transfected culture than the control (Fig. 4A, Day
14), although by day 24, smaller spots did appeared in the
RECK-transfected culture (Fig. 4A, Day 24). Expression of
two marker genes, type II collagen and type X collagen, was
also significantly retarded in RECK-transfected cells around
day 14 but was markedly increased by day 24 (Fig. 4B, Col-II
and Col-X). Elevated baseline RECK expression in the RECK-
transfected cells was evident up to day 14, but it was masked
by the elevated endogenous Reck expression on day 24 (Fig.
4B, RECK). Thus, constitutive RECK expression lowers the
efficiency of cellular condensation and delays the expression
of cartilage-specific ECM components.
To understand how the over-expressed RECK suppressed
cellular condensation, we analyzed the level of gelatinases (i.e.
MMP2 and MMP9) present in the culture supernatant of the
transfectants by gelatin zymography. Consistent with our
previous findings (Takahashi et al., 1998), the level of secreted
pro-MMP9 was significantly reduced in RECK-transfected
cells (Fig. 4C, lanes R). We also found that the ATDC5 cells
stably transfected with an Mmp9- or MT1-MMP-expression
vector showed more rapid and extensive cellular condensation
than the cells transfected with the empty vector or an Mmp2-
expression vector (Fig. 4D). These findings suggest that some
MMPs have a potential to promote cellular condensation of
ATDC5 cells and that RECK may suppress cellular
condensation by regulating such MMPs.
Formation of ECM-rich nodules is suppressed by RECK
To assess the importance of Reck upregulation in the later
Fig. 4. Effects of forced expression of RECK or MMP on
chondrogenic differentiation of ATDC5 cells. (A) The cells
transfected with the control vector (left panels) or the vector
expressing human RECK (right panels) were incubated for the
indicated times and then stained with Alcian Blue. Bars, 2 mm.
(B) Expression of chondrogenic differentiation markers in the
transfectants. Total RNA (20 ?g) extracted from the transfectants at
the indicated times (bottom labels) was analyzed by RNA blot
hybridization using the indicated probes (top labels). (C) Effects of
forced RECK expression on the level of secreted gelatinases.
Conditioned media (7 hours conditioned) prepared from vector- (V)
or RECK-transfected ATDC5 cells (R) at the indicated times were
analyzed by gelatin zymography. The positions of the pro-MMP9,
pro-MMP2, and intermediate-MMP2 bands are indicated. The
amounts of samples applied were adjusted between V and R so as to
represent equal cell number. Day 7 samples were five times more
concentrated than day 3 samples on a per-cell basis. (D) Effects of
forced expression of MMPs on cellular condensation of ATDC5 cells.
ATDC5 cells were stably transfected with a mammalian expression
vector (pcDNA3.1/Hygro(+); Invitrogen) or the vector containing
mouse Mmp2, mouse Mmp9, or human MT1-MMP cDNA. Phase-
contrast micrographs of vector- or MMP-transfected ATDC5 cells
incubated for 5 days are shown. Expression of MMP in each
transfectant was confirmed by northern blot hybridization and gelatin
zymography. Note the accelerated cellular condensation in Mmp9-
transfected cells and MT1-MMP-transfected cells. Bars, 100 ?m.
Journal of Cell Science
stages of ATDC5 differentiation, we generated ATDC5 cells
stably transfected with a vacant expression vector or the vector
containing small hairpin RNA (shRNA) designed to
knockdown Reck mRNA (Fig. 5). Significant decrease in
RECK expression was detectable in the shRNA-expressing
cells (S) as compared to the control cells (V) (Fig. 5A). Under
differentiating conditions, the shRNA-expressing cells did give
rise to foci of cellular condensation (Fig. 5B, Days 12 and 15)
as numerous as the control cells (Fig. 5C, open bars). These
foci, however, were morphologically distinct from those of
control cells: they were more flat and less refractile (Fig. 5B,
Day 15; Fig. 5D, DIC; Fig. 5C, Day 15, hatched bars). These
abnormal foci, which showed markedly reduced RECK
immunoreactivity (Fig. 5D, RECK), were negative for Alcian
Blue staining (Fig. 5B, Day 24) and type II collagen
immunoreactivity (Fig. 5D, Col-II), indicating reduced ECM
components in these structures. These results suggest that
RECK is essential for the formation of mature, ECM-rich
Our data indicate that Reck is abundantly expressed in
developing cartilage in mouse embryos as well as in cultured
cells of the chondrocytic lineage. We also found, however, that
during the early phase of ATDC5 differentiation, Reck
expression is low and multiple MMP family genes (Mmp2,
Mmp9 and Mt1-Mmp) are upregulated (Fig. 6A). The areas of
elevated collagenolytic activity coincided with the foci of
cellular condensation (Fig. 2B), suggesting that MMPs may
play roles in cellular condensation. Our finding that forced
expression of RECK, a membrane-anchored MMP regulator,
reduced the efficiency of cellular condensation (Fig. 3) is
consistent with the model that MMPs (e.g. MMP9, MT1-
MMP) are required for this process (left part of Fig. 6B).
As differentiation proceeds, ATDC5 cultures become
heterogeneous, and RECK becomes upregulated mainly inside
the cartilaginous nodules where type II collagen is accumulated
(Fig. 2F), whereas collagenolytic activity persists around the
rim of the nodules (Fig. 2B, Day 24). This spatial arrangement
agrees with the hypothesis that elevated RECK inside the
nodule is important for ECM accumulation whereas persistent
ECM degradation along the rim is required for nodule
expansion (right panel of Fig. 6B). Our findings that RECK
knockdown reduced the number of ECM-rich nodules (Fig. 5)
Journal of Cell Science 120 (5)
Fig. 5. Effects of Reck gene knockdown on
cartilaginous nodule formation by ATDC5
cells. (A) ATDC5 cells were stably transfected
with the control vector (V) or the vector
expressing shRNA against Reck (S).
Expression of RECK in these cells were
analyzed by immunoblot assay (top panel) and
RNA blot hybridization (middle panel). Total
proteins (30 ?g) harvested on day 7 and total
RNA (20 ?g) harvested on day 4 were used.
The number under each lane indicates relative
band intensity. (B) Phase-contrast micrographs
(top 2 rows) and Alcian blue-stained foci (third
row) of the vector-transfected cells (left panels)
or RECK-siRNA-transfected cells (right
panels) incubated for the indicated times. Bars,
100 ?m (Day 12); 250 ?m (Day 15) and 400
?m (Day 24). (C) The number and
morphology of foci. After incubation for the
indicated times, the number of all visible foci
(Total foci) and the number of highly refractile
foci (e.g. in B, Vector Day 15, arrow), which
represent ECM-rich cartilaginous nodules
(Shukunami et al., 1998), were scored under a
microscope. Values are mean ± s.e.m. from
quadruple dishes. (D) Immunofluorescent
staining of foci at day 15. The cells were
doubly stained with anti-type II collagen
(green) and anti-RECK (red) antibodies.
Morphology (DIC), type II collagen signals
(Col-II), and RECK signals (RECK) around
typical foci were recorded under the same
microscopic conditions. Bars, 100 ?m. The
findings were consistent between two separate
experiments using independently derived
Journal of Cell Science
A role for RECK in chondrogenesis
and that the forced expression of RECK gave rise to compact
nodules (Fig. 3C) are consistent with this model.
Although RECK overexpression suppressed cellular
condensation and delayed the onset of marker gene expression,
it did not strongly suppress the upregulation of these markers
in the later phase (Fig. 4A,B, Day 24). In fact, at day 24, the
level of type II collagen in RECK-transfected cells exceeded
the level in the control cells (Fig. 4B). This raises the
possibility that RECK promotes differentiation in the later
phase of chondrogenesis. Alternatively, RECK may activate
the type II collagen gene specifically in certain cellular
contexts. Whatever the molecular mechanism, this result seems
to indicate the involvement of a positive feedback loop in this
process where the inhibition of matrix proteases results in
upregulation of their substrate gene, thereby accelerating ECM
Parallels to our findings can be found in some previous
reports describing studies using intact animals or tissue
explants. Abundant expressions of Mmp2, Mmp9, and Mt1-
mmp in mesenchymal cells surrounding cartilage have been
reported (Apte et al., 1997; Reponen et al., 1992; Reponen et
al., 1994). Non-overlapping expression patterns of type II
collagen (cartilage) and type I collagen (perichondrium) have
also been reported (Mizoguchi et al., 1990; Sandberg and
Vuorio, 1987). The smaller cartilage formed after treatment of
mandibular explants in culture with a synthetic MMP inhibitor
(Chin and Werb, 1997) is reminiscent of the compact nodules
formed by the ATDC5 cells constitutively expressing RECK
(Fig. 3C). Such consistency strengthens our notion that ATDC5
is a good model system for studying the dynamic processes of
chondrogenesis and that many findings in this system may have
relevance to chondrogenesis in vivo.
There may be some other roles of RECK in chondrogenesis,
however, which would be hard to explore in the ATDC5
system. For instance, blood vessel invasion into avascular
cartilage is a critical step in endochondral bone formation, and
active roles for MMP9 and MT1-MMP in this process have
been documented (Holmbeck et al., 1999; Vu et al., 1998; Zhou
et al., 2000). Given its activity in regulating angiogenesis in
vivo (Oh et al., 2001) and its abundance in cartilage, RECK is
likely to be involved in the regulation of endochondral bone
formation. Perhaps, more elaborate experimental systems in
vivo, such as conditional knockout or conditional transgenic
mice, need to be employed to test such a hypothesis.
In our previous study using an immunohistochemical
approach, little RECK expression could be detected in
developing cartilage (Echizenya et al., 2005). In this study,
however, in situ hybridization as well as experiments with
cultured cells revealed abundant expression of Reck in cartilage
and/or chondrocytes. Cartilage is notoriously difficult to analyze
in immunohistochemical studies because of its abundant ECM.
In fact, pre-treatment with hyaluronidase was necessary to stain
RECK protein in ATDC5 cartilaginous nodules in the present
study (Fig. 2E,F, Fig. 5D). Moreover, our in situ hybridization
data did indicate moderate levels of Reck expression in muscles
(e.g. intercostals regions in Fig. 1F and the areas surrounding
developing femur in Fig. 1G). It is, therefore, likely that we
failed to detect the RECK protein in cartilage for some technical
reasons in the previous immunohistochemical studies where we
focused on developing muscles.
Our previous study using Reck-deficient mice indicated that
RECK is essential for vascular development around E10.5 and
that this is probably due to the activity of RECK in regulating
MMPs and to protect fibrillar collagen (Oh et al., 2001). More
recent studies (Echizenya et al., 2005) (also this study) have
revealed that RECK is expressed in mouse embryos beyond
E10.5. This, together with our previous observation that RECK
is expressed in various human adult tissues (Takahashi et al.,
1998), suggest that RECK probably continues to play important
roles in the later stages of development and in adult life.
Interestingly, RECK expression is low during the early phase
of muscle development where myoblast proliferation, migration
and fusion are actively taking place, but RECK expression is
upregulated in the later phase where individual myofibers
become ensheathed by basement membranes (Echizenya et al.,
2005). Hence, progressive upregulation of RECK occurs in two
different tissues (skeletal muscle and cartilage) during
development; in both cases, low RECK expression is associated
with early morphogenetic steps whereas elevated RECK
expression is associated with later, consolidation steps or ECM
accumulation. The GPI-mediated membrane anchoring makes
RECK unique among endogenous MMP regulators and
particularly suitable for a task that requires delicate and
dynamic regulation of pericellular ECM remodeling.
Type II collagen
(type I collagen+) Cartilaginous nodule
Fig. 6. Possible roles of RECK and MMPs during chondrogenic
differentiation. (A) Schematic representation of the temporal changes
in the amounts of mRNA encoding MMP (representing MMP2,
MMP9 and MT1-MMP; red line), RECK (green line), and type II
collagen (blue line) during ATDC5 differentiation. (B) A working
hypothesis: elevated MMP activity is required for cellular
condensation in the early phase and nodule expansion in the later
phase in ATDC5 differentiation. Elevated RECK expression is
required for ECM accumulation inside the nodules in the later phase.
Constitutive RECK expression suppresses cellular condensation.
RECK knockdown suppresses ECM accumulation and hence nodule
Journal of Cell Science
Cartilage is known to have very limited potential of self-
repair (Buckwalter, 2002). Cell transplantation experiments to
repair injured cartilage has been unsuccessful, a major obstacle
being the difficulty in inducing proper morphogenesis
(Buckwalter, 2002). Destruction of joint ECM is one of the
major symptoms of rheumatoid arthritis (RA); involvement of
MMPs in this process has been documented in a number of
studies (Jackson et al., 2001; Martel-Pelletier et al., 2001).
Interestingly, reduced expression of RECK in RA synovial
membrane has recently been reported (van Lent et al., 2005).
Thus, further studies on RECK in the context of cartilage
development and maintenance may also yield information
useful for clinical application.
In conclusion, our data points to the importance of regulated
expression of RECK during cartilage morphogenesis. Its
ubiquitous expression in embryonic tissues and the previously
demonstrated roles in developing blood vessels (Noda et al.,
2003; Oh et al., 2001) and skeletal muscles (Echizenya et al.,
2005) support the idea that RECK serves as an essential and
versatile regulator of tissue morphogenesis.
Materials and Methods
RNA blot hybridization
Total RNA extracted from cultured cells was separated by electrophoresis in 1%
agarose gels, transferred to Hybond N+membranes (Amersham Pharmacia
Biotech), and hybridized with a 32P-labeled probe. The following cDNA fragments
were used as probes: 4.1 kb mouse Reck (Takahashi et al., 1998), 4.4 kb human
RECK (Takahashi et al., 1998), 2.58 kb human MT1-MMP (Sato et al., 1994), 2.3
kb mouse Mmp2 (Reponen et al., 1992), 3.2 kb mouse Mmp9 (Tanaka et al., 1993),
0.85 kb mouse type I collagen (?2) (pAZ1002; kindly provided by Benoir de
Crombrugghe, M. D. Anderson Cancer Center, Houston, TX), 1.4 kb rat type II
collagen (?1) (Kimura et al., 1989), 0.65 kb mouse type X collagen (?1) (Apte et
al., 1992), and mouse glyceraldehyde-3-phosphate dehydrogenase (GAPDH
Control Amplifier Set; Clontech). The quantity and quality of RNA applied to each
lane were assessed by the patterns of ribosomal RNAs visualized by ethidium
bromide or SYBR Green (Biowhittaker Molecular Applications). Densitometric
analyses of X-ray film images were performed using the NIH Image software.
In situ hybridization
The 0.68 kb 3?-UTR fragment of mouse RECK cDNA (GenBank AB006960,
nucleotides 3412-4094) was amplified by PCR and used as a template for generating
riboprobes. Digoxigenin-labeled antisense and sense (control) riboprobes were
prepared using the DIG RNA Labeling Kit (Roche Diagnostics). Mouse embryos
were embedded in OCT (Tissue-Tek), frozen on liquid nitrogen, sectioned at 12 ?m
thickness with a cryostat, and fixed briefly in 4% paraformaldehyde at room
temperature. Hybridization was performed at 58°C for 16 hours as described by
Braissant et al. (Braissant et al., 1996), and after washing under high stringency
conditions, signals were visualized using DIG Nucleic Acid Detection Kit (Roche
ATDC5 cells (Atsumi et al., 1990; Shukunami et al., 1996) were cultured in growth
medium consisting of 1:1 mixture of Dulbecco’s modified Eagle’s medium and
Ham’s F-12 (Invitrogen) supplemented with 5% fetal bovine serum (Equitech-Bio
Inc., Ingram, TX, USA), 10 ?g/ml bovine insulin, 10 ?g/ml human transferrin and
3?10–8M sodium selenite. Inoculum density for induction of chondrogenesis was
9?104cells/35-mm dish, 2.3?105cells/60-mm dish or 6.5?105cells/100-mm dish,
unless noted otherwise. Transfection was performed by the calcium-phosphate
method using 2 ?g plasmid DNA mixed with 1 ?g calf thymus DNA (carrier). After
selection in growth medium containing 1 mg/ml G418 or 208 U/ml hygromycin-B,
transfectant colonies were pooled and expanded. Preparation of conditioned media
and gelatin zymography were performed as described previously (Takahashi et al.,
1998) except that plating density, medium and pre-incubation time were adapted to
the cell line (see above).
Indirect immunofluorescent staining
Cells cultured on Lab-Tek Chamber Slides (Nunc) were fixed with 4%
paraformaldehyde in phosphate-buffered saline containing divalent cations [PBS
(+)], treated with 2.5 % hyaluronidase in PBS (+), and permeabilized with 0.1%
Triton X-100 in PBS (+), followed by pre-treatment to block non-specific reactions
with 5% skim milk and 1.5% non-immune goat serum in PBS (+). For collagen and
RECK double staining, the primary immunoreaction was carried out using rabbit
antiserum against either type I collagen (Rockland, #600-401-103-0.5; 1:100
dilution) or type II collagen (Rockland, #600-401-104-0.5; 1:400 dilution) mixed
with mouse monoclonal antibodies against RECK (5B11D12; a gift from Amgen;
4.5 ?g/ml). To avoid cross reaction, the secondary immunoreaction was carried out
in two steps: (1) incubation with FITC-conjugated goat anti-rabbit IgG (Jackson
ImmunoReserch Laboratories, #211-095-109; 1:500 dilution) followed by rinsing
and blocking with PBS (+) containing skim milk and 1.5% non-immune rabbit
serum, and (2) incubation with TRITC-conjugated rabbit anti-mouse IgG (DAKO,
#R0270; 1:30 dilution) followed by rinsing. The fluorescence images were recorded
with a confocal laser microscope (FV300, Olympus) using the sequential scan mode
to avoid leakage from different wavelengths. We also performed two types of control
experiments: (1) single staining, using antibodies against either RECK, type I
collagen or type II collagen, and (2) omission of primary antibodies (see
supplementary material Fig. S1). The results indicate that the signals detected in the
double-labeling experiments shown in Fig. 2E,F reflect the specific
immunoreactivity of respective primary antibodies. Specificity of the anti-RECK
antibodies (5B) has been confirmed in a number of previous experiments, which
include comparison between the following sample pairs: (1) vector-transfected and
RECK-transfected HT1080 cells (low endogenous RECK) in immunoblot assay
(Takahashi et al., 1998), (2) embryo fibroblasts derived from wild-type and Reck-
null mice in immunoblot assay (Oh et al., 2001), and (3) wild-type and Reck-null
mouse embryos after immunohistochemical staining (Oh et al., 2001).
To extract proteins, cultured cells were scraped in cell extraction buffer [10 mM
Tris-HCl (pH 8.0), 150 mM NaCl, 10% glycerol, 1% deoxycholate, 0.1% SDS,
protease inhibitors (CompleteTM; Roche Diagnostics)] at 4°C. After sonication, the
lysates were cleared by centrifugation, and the concentration of soluble proteins was
determined. The proteins (30 ?g) were separated by SDS-PAGE (10% acrylamide),
blotted onto PVDF membranes (Millipore), and detected using the anti-RECK
antibodies (5B11D12) and ECL System (Amersham Pharmacia Biotech).
Densitometric analyses were performed using NIH Image.
Alcian Blue staining
To visualize accumulation of sulfated glycosaminoglycans (a marker for
chondrogenic differentiation), cells were rinsed with PBS, fixed with 95% methanol
for 20 minutes, and stained with 0.1% Alcian Blue 8GS (Fluka, Buchs, Switzerland)
in 0.1 M HCl overnight at room temperature.
In situ zymography
Cells grown on 35 mm glass-bottomed dishes (Iwaki, Chiba, Japan) were washed
with PBS and overlaid with 1 ml medium containing freshly neutralized Cellmatrix
type I collagen (Nitta Gelatin, Osaka, Japan) and 10 ?g/ml FITC-labeled DQTM-
type I collagen (Molecular Probes). After solidification at 37°C for 30 minutes, 1
ml growth medium was gently added, and the dishes were incubated for an
additional 24 hours. FITC signals, representing digested DQTM-type I collagen,
were recorded using the confocal laser microscope.
Three small hairpin RNA (shRNA) sequences (19 mers) against the mouse Reck
mRNA were designed and inserted into the mammalian expression vector pRNA-
U6.1./Neo (GeneScript). The vector yielding highest activity of gene silencing
(target sequence: ACGCCTGCAAGAGAATTCT) was selected and used in this
We are grateful to Benoit de Crombrugghe, Karl Tryggvason and
Hidekazu Tanaka for providing cDNA clones and to Kazuki Kuroda
for technical advice on in situ hybridization. We also thank Emi
Nishimoto and Takashi Kawai for technical assistance, Aki Miyazaki
for secretarial assistance, and all the members of our laboratories for
valuable discussions. This work was supported by grants from MEXT,
Japan and the Mitsubishi Foundation. S.K. was a recipient of a JSPS
Research Fellowship for Young Scientists.
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