1 Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot
76100, Israel
2 ProChon Biotech Ltd, Kiryat Weizmann, Rehovot 76114, Israel
Author for correspondence (e-mail: yayon{at}prochon.co.il)
Accepted 10 October 2001
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Summary |
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Key words: Chondrocytes, FGF signaling, FGF receptor 3, Focal adhesions, G1 arrest
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Introduction |
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The expression of FGFRs is tightly regulated during embryonal development
and tissue regeneration (Basilico and
Moscatelli, 1992; Yamaguchi et
al., 1995
; Goldfarb,
1996
; Martin,
1998
; Szebnyi and Fallon, 1999;
Xu et al., 1999
). FGFR3 is
particularly highly expressed during embryonic development in the
pre-cartilaginous mesenchyme (Peters et
al., 1992
; Peters et al.,
1993
) and later on in the maturation zone of the epiphyseal
growth-plates, where it is involved in long bone development
(Naski et al., 1998
). The
discovery that specific activating mutations in FGFR3 underlie a variety of
human skeletal disorders, such as Achondroplasia, the most common form of
human genetic dwarfism, has linked FGFR3 signaling and skeletal development
(reviewed in Webster and Donoghue,
1997
; Burke et al.,
1998
; Naski and Ornitz,
1998
). Moreover, FGFR3-null mice exhibit bone overgrowth
accompanied by expansion of proliferating and hypertrophic chondrocytes within
the growth-plate (Colvin et al.,
1996
; Deng et al.,
1996
). Transgenic mice harboring FGFR-activating mutations
(Naski et al., 1998
;
Chen et al., 1999
;
Li et al., 1999
;
Wang et al., 1999
;
Segev et al., 2000
) or
overexpressing FGF2 (Coffin et al.,
1995
) or FGF9 (Garofalo et
al., 1999
) display a dwarf phenotype similar to the human
disorders where attenuated proliferation and differentiation of chondrocytes
result in retarded bone growth (Naski et
al., 1998
; Chen et al.,
1999
; Li et al.,
1999
). Overall, these studies indicate that FGFR3 acts as a potent
regulator of chondrocyte differentiation and as a negative regulator of bone
growth. However, the downstream events by which FGFR3 influences the
proliferation or terminal differentiation of chondrocytes remains poorly
understood.
Although the role of cell cycle regulating proteins in maintaining the
balance between proliferation and differentiation is well studied, there is
limited data on the expression pattern of cell-cycle-regulating genes during
chondrocyte maturation (LuValle and Beier,
2000). The expression of the
p21Waf1/Cip1 gene, a cyclin-dependent kinase
inhibitor, was found to be upregulated during chondrocyte differentiation in
vitro (Beier et al., 1999
) and
in vivo (Stewart et al., 1997
)
and to be controlled by FGFs along with activation of the transcription factor
STAT1 in RCS cells (Sahni et al.,
1999
). Stat-1-null mice, however, have not been reported to have
bone defects (Durbin et al.,
1996
; Meraz et al.,
1996
). In this study, we have utilized a chondrosarcoma model cell
system to further study the FGF-mediated control mechanisms. We show that FGF
signaling and growth arrest induces alterations in the subcellular
localization of FGFR3, and several candidate genes that may be involved in the
regulation of the cell cycle and cytoskeletal organization.
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Materials and Methods |
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Western blot analysis
Cells were lysed in lysis buffer (50 mM Tris-HCl, pH 7.5), 150 mM NaCl, 1
mM EDTA, 10 % glycerol, 1% NP40 and `Complete' (Boehringer Mannheim) protein
inhibitor mix according to the manufacturers instructions). Equal amounts of
cell lysates, as determined by a Bradford reaction, were loaded, resolved by
SDS-PAGE and followed by western blot analysis. Proteins were visualized by
using an enhanced chemiluminescence kit (ECL Amersham). Anti-FGFR3 antibodies,
anti-NF-B p65 antibodies, anti-c-Jun antibodies, anti-JunD antibodies.
Anti-FRS2 antibodies were a generous gift from Yaron Hadari, and anti-Id
antibodies were from Santa Cruz biotechnology. Anti-Ezrin antibodies were from
Transduction Laboratories. Anti-pMAPK antibodies and preimmune rabbit serum
were from Sigma. Secondary antibodies used were anti-rabbit or - mouse
immunoglobulins linked to horseradish peroxidase (Amersham Life Science).
Cell cycle analysis
Cells were trypsinized, centrifuged, resuspended in 300 µl PBS, fixed in
5 ml methanol and stored at -20°C for at least 24 hours. On the day of
analysis, cells were collected by centrifugation and washed with 1 ml PBS.
Cell pellets were then resuspended in PBS (between 0.5-1ml) containing 100
µg/ml RNase and 12.5 µg/ml propidium iodide and incubated in the dark
for 20 minutes. Filtered samples were then analyzed for cell cycle content by
using FACsort (Becton-Dickinson).
Screening of Atlas cDNA expression array
Total RNA was extracted from untreated RCS cells or RCS cells that were
incubated with 20 ng/ml FGF9 and 1 µg/ml heparin by using a Tri Reagent kit
(Molecular Research Center). After DNase treatment,
P32-radiolabeled cDNA was prepared from 9 µg of total RNA and
hybridized to the membranes (Clontech, 7738-1) according to the manufacturer's
instructions.
Immunohistochemistry
Isolated bones from mouse hind limbs were fixed in 4% paraformaldehyde (pH
7.4), declacified in EDTA, dehydrated in an ethanol gradient and embedded in
paraffin. 5-µm thick sections were cut, dewaxed in xylene, hydrated through
graded ethanol to water and then rinsed in PBS. Endogenous peroxidase activity
was quenched by incubation in 3% hydrogen peroxide for 10 minutes.
Immunostaining was performed using the Histostain-plus broad-spectrum
peroxidase kit (Zymed Laboratories). Sections were blocked in normal serum for
10 minutes and incubated with anti NFB p65 polyclonal antibody diluted
1:200 in PBS overnight at 4°C. Sequentially, the sections were incubated
with biotinylated secondary antibodies, followed by avidin horseradish
peroxidase conjugate and diaminobenzidine substrate as a chromogen. Finally,
the sections were counterstained with methyl green, dehydrated in graded
ethanols, cleared in xylene and mounted. Negative controls for immunostaining
were performed by substitution of the primary antibody with PBS or preimmune
serum.
Immunofluorescence
Cells were cultured on glass coverslips for 48 hours, permeabilized with
0.5% TritonX-100 and fixed with 3% paraformaldehyde in phosphate-buffered
saline. Fixed cells were incubated with the relevant primary antibodies to
FGFR3, pTYR (Upstate) and vinculin (Transduction Labratories) and detected
using Cy3 goat anti-rabbit IgG (Jackson ImmunoResearch Labratories) and Alexa
488 goat antimouse IgG (Molecular probes) as secondary antibodies. Actin was
detected by direct phallodin (Sigma) staining. Image acquisition was performed
using an Axiphot (Zeiss) microscope equipped with an AttoArc (Zeiss) camera
using a 100x objective (Zeiss) or with the Delta Vision system (Applied
Precision, Issaquah, WA, USA) equipped with a Zeiss Axiovert 100 microscope
(Oberkochen, Germany) and photometrics 300 series scientific-grade cooled CCD
camera (Tucson, AZ, USA) reading 12 bit images using a 100x1.3 NA
plan-Neoflaur objective (Zeiss, Oberkochen, Germany).
RNA preparation and northern blot analysis
Total RNA was extracted from cells using the EZ-RNA kit (Biological
Industries) according to the manufacturer's instructions. 15 µg of total
RNA were loaded on a 1% agarose MOPS/formaldehyde gel and transferred to a
nylon membrane (Sartorius). The blot was hybridized overnight to
32PdCTP-labeled FGFR3 cDNA probe (nucleotides 60-667) in
NorthernMax buffer (Ambion), washed and exposed to a Biomax film (Kodak).
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Results |
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We found, in agreement with Sahni et al.
(Sahni et al., 1999), that
FGF9, a more specific ligand for FGFR3
(Hecht et al., 1995
), not only
induces activation of the receptor but also results in rapid downregulation of
its expression. A profound decrease in the level of the FGFR3 protein was
already detected 4 hours after addition of the ligand and fell below detection
levels after 8 hours. ERK phosphorylation decreased concomitantly, with
complete loss of this downstream signal after 8 hours
(Fig. 1A). Northern blot
analysis demonstrated that the level of the transcript encoding for FGFR3
decreased with similar kinetics (Fig.
1B), possibly indicating the induction of a transcriptional
inhibitory loop mechanism.
|
Long-term exposure of RCS cells to FGF9 resulted in a dramatic inhibition
of their proliferation (Fig.
1C); a similar effect had previously been observed for FGF1
(Sahni et al., 1999). The
density of the cell cultures incubated with FGF9 for various periods (8, 16,
24 and 72 hours) was essentially unchanged from the initial seeding density,
as estimated by microscopic examination, although some dead cells were
observed following incubations of more than 16 hours (data not shown). This
qualitative observation indicated that FGF9 caused cell growth arrest.
Following removal of FGF9 and continuation of growth for a further 72 hours,
the numbers of cells in the cultures were found to be
7.2x105, 6.8x 105, 3.5x105,
2.9x105 and 1.1x105 for FGF exposures of 0,
8, 16, 24 and 72 hours, respectively. Since the original number of cells
seeded was 0.6x105 in each case, we conclude that a
significant proportion of the cells exposed to FGF9 for 8-24 hours was capable
of restarting growth after removal of FGF9. However, cell death and apoptosis
can not be ruled out for the cells that failed to regain proliferative
capacity.
In order to analyze the growth inhibition in greater detail, RCS cells were incubated with or without FGF9 for 16 hours and subjected to FACS analysis. Cells incubated with FGF9 either alone or together with heparin exhibited a significant increase in the percentage of cells in the G1 phase (86% compared with 62% in untreated cells; Fig. 2A) and a concomitant decrease in the percentage of cells in S phase (5% compared with 22% in the untreated cells; Fig. 2A), strongly suggesting that FGF9 induces growth arrest at the G1 phase of the cell cycle. Addition of heparin alone had no effect on the cell cycle (Fig. 2A). In a detailed time course analysis, a transient accumulation of the surviving cells at the G2 phase of the cell cycle was noted after 8 hours of treatment (28% compared with 13% in the untreated cells) (Fig. 2B), preceding the G1 growth arrest detected after 10 hours of incubation with FGF9.
|
FGF signaling modulates multiple genes in RCS cells
In an attempt to identify the genes that are modulated by FGF and that
might be involved in FGF-induced growth arrest, we utilized DNA array
technology (Fig. 3A). Total RNA
extracted from RCS cells before (0 hours) and after incubation with FGF9 and
heparin for 3 hours was used for screening an Atlas membrane containing 588
known rat cDNAs (7738-1, Clontech). This array is composed mainly of genes
reported to play key roles in processes such as signal transduction,
apoptosis, tumor suppression and oncogenesis. The screen identified 11
distinct genes whose expression level changed more than two-fold upon FGF9
stimulation. Five of these were known FGF target genes, including c-Jun
(Pertovaara et al., 1993;
Cao et al., 1998
), the
urokinase receptor (Mignatti et al.,
1991
), the LDL receptor (Hsu
et al., 1994
), cyclin D1 (Rao
et al., 1995
) and p21 (Sahni
et al., 1999
). Among the genes that were significantly upregulated
but had not been previously described as direct FGF targets were JunD, FRA 2,
NF-
B1 (p50/p105), STAT3 and Ezrin. The expression of Id1 was markedly
decreased (Fig. 3A).
Computerized analysis of the data using AtlasImage 1.01 software (Clontech)
demonstrated that the induction of c-Jun, JunD, FRA 2, cyclin D1,
NF-
B1(p50/p105), STAT3 and Ezrin was 2.45-, 15.13-, 5.63-, 2.27-,
3.55-, 3.05- and 3.98-fold higher, respectively. There was a threefold
reduction in the expression of the Id1 gene.
|
We confirmed the above results by determining the levels of the protein products of some of the genes in immunoblots (Fig. 3B). In agreement with the gene array results, the levels of c-Jun, JunD and p21 proteins increased at 2 hours after FGF addition and peaked at 4 hours. The increase in the levels of Ezrin was detectable only 6 hours after stimulation, and it remained high for at least 24 hours. The level of Id1 protein fell to below detection limit within 2 hours of treatment (Fig. 3B).
In a search for a possible link between the expression of some of the
induced genes and FGFR3 activation in vivo, we have analyzed epiphyseal
growth-plates from normal and from transgenic mice carrying the Achondroplasia
G380R hFGFR3 gene. Immunostaining of paraffin-embedded sections with
anti Rel A (p65) antibodies (Rel A together with NF-B1(p50/p105) forms
the active NF-
B dimer) revealed a significantly denser distribution of
Rel A in the growth-plates of the transgenic mice than in their normal
littermates (Fig. 3C).
Furthermore, the protein was predominantly expressed, like FGFR3, in the
maturation/upper hypertrophic zones of the growth-plate. Immunostaining of
growth-plate sections with antibodies against NF-
B1(p50/p105) and c-Jun
also showed similar qualitative differences (data not shown), although these
were significantly less pronounced than for Rel A (p65). These results
indicate that the constitutively active, mutant FGFR3 induces in vivo gene
expression, which parallels that observed in FGF9-stimulated RCS cells.
FGF activates FGFR3 localized to focal adhesions and disrupts the
cytoskeletal organization
It is well known that proliferation and cell cycle progression are tightly
associated with cell shape and the organization of the cytoskeleton
(Assoian and Zhu, 1997). This,
together with the fact that FGF signaling induces the expression of Ezrin, a
prominent cytoskeletal protein, led us to examine more carefully the
morphological changes induced in these cells by FGF. In general, the majority
of RCS cells in culture have a polygonal shape, which is typical of mature
chondrocytes. A small percentage of RCS cells in the culture exhibit round
morphology, which might represent a different differentiation stage.
Incubation of RCS cells with FGF9 dramatically changed their morphology, with
complete rounding of the cells apparent 6 hours after stimulation, whereas
untreated cells or cells treated with heparin alone retained their polygonal
shape (Fig. 4A).
|
The fact that the cells seem to at least partially detach from the plate
prompted us to examine the effect of FGF9 on the organization of their focal
adhesions, the major anchor sites of cells to their substrate. RCS cells were
therefore subjected to double immunofluoresence staining with an anti-FGFR3
polyclonal antibody and an anti-vinculin monoclonal antibody
(Fig. 4B). Unexpectedly, we
found that a significant portion of the FGFR3 protein was localized in
arrowhead-shaped structures typical of focal adhesions and colocalized with
vinculin. Notably, not all focal adhesions contain the receptor. Next, we
examined whether activation of FGF signaling enhances phosphotyrosine (pTyr)
activity at the focal adhesions by immunofluorescent staining of
FGF9-stimulated cells with anti-phosphotyrosine antibodies
(Fig. 5A). It is well
documented that pTyr activity is abundant in the focal adhesions of
unstimulated cells (reviewed in Vuori et al., 1998;
Cary and Guan, 1999), as can
clearly be seen in untreated RCS cells. Detailed time-course analysis showed
that pTyr activity in the focal adhesions increases 10 minutes after FGF9
addition, which correlated with activation of FGFR3 and its downstream targets
in the focal adhesions. This activity decreased after 1 hour and almost
completely disappeared after 6 hours (Fig.
5A).
|
Most dramatic, however, was the disruption of the focal adhesions and its correlation with the kinetics of FGFR3- and vinculin-associated focal adhesions following FGF stimulation (Fig. 5B). Although 10 minutes after stimulation FGFR3 was still associated with the focal adhesions, an hour later it was almost undetectable in these sites. This was in marked contrast with vinculin which was retained in these adhesion sites, and its distribution was apparently unchanged up to 6 hours after stimulation as the cells became more rounded and the focal adhesions disintegrated (Fig. 5B). Staining with phalloidin, which labels actin filaments, demonstrated a similar pattern whereby exposure to FGF9 for several hours induced a major breakdown of the organized actin network in these cells, as well as in ruffling and lamelipodia extensions, which were observed already one hour after FGF stimulation (Fig. 5C).
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Discussion |
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Cell cycle regulation and alterations in transcriptional pattern
induced by FGF
We have chosen a rat chondrosarcoma cell line (RCS) as an in vitro model
system because it expresses a high level of FGFR3 protein as well as other
chondrocyte-specific markers (Sahni et
al., 1999). The validity of this model is confirmed by the
inhibition of RCS cell proliferation in response to FGF9, a considerably
specific ligand for FGFR3 (Hecht et al.,
1995
). We show that the mitotic arrest occurs at the G1 phase of
the cell cycle (Fig. 2).
Interestingly, a careful analysis of the cell cycle data suggests that just
before the G1 arrest, the cells transiently accumulate at the G2 phase
(Fig. 2B), a mechanism shown
previously to be directly associated with cell differentiation
(Aloni-Grinstein et al., 1995
;
Schwartz and Rotter, 1998
).
Downregulation of FGFR3 in response to FGF was also confirmed for RCS cells by
the dramatic reduction in the levels of FGFR3 mRNA and protein following the
addition of FGF9 (Fig. 1).
Furthermore, a consequence of the failure to turn off FGFR3 signaling, such as
occurs with the constitutively activated G380R mutant of FGFR3
(Monsonego-Ornan et al.,
2000
), is shown in Fig.
3C.
Stimulation by FGF, as is the case with all growth factors, has a profound
effect on the cellular gene transcription profile. In the exploratory screen
carried out in the present work, FGF was found to activate multiple genes and
to repress one. Since FGF arrests chondrocyte proliferation and acts as a
differentiation trigger, it is not unexpected that seven of the ten induced
proteins (c-jun, Jun D, Fra2, NFB, STAT 3, Cyclin D1 and p21) as well
as the repressed Id1 are involved in cell cycle regulation. c-Jun, Jun D and
Fra1 are members of the AP-1 family of transcription factors (reviewed in
Karin et al., 1997
;
Leppa and Bohmann, 1999
),
which are upregulated by FGF in RCS cells. Several studies have indicated that
c-Jun and JunD inhibit the differentiation of chondrocytes in vitro
(Kameda et al., 1997
). In
addition, Jun D plays a major role in osteoblast maturation
(McCabe et al., 1996
) and has
been implicated as a negative regulator of cell proliferation in several other
cell types (Wang et al.,
1996
). The Fra2-related transcription factor, Fral, is associated
with enhanced osteoblast differentiation, resulting in increased bone
formation (Jochum et al.,
2000
). Although the observed elevation in cyclin D1 mRNA levels
does not typically correlate with a G1 arrest, FGF has been found to suppress
MCF-7 human breast cancer cell proliferation concomitantly with an increase in
cyclin D1 (Wang et al., 1997
).
Also, cyclin D1 is directly activated by the transcription factor ATF-2, which
inhibits chondrocyte proliferation in mice
(Beier et al., 1999
).
Therefore, this array of genes may participate in regulating cell cycle
progression in order to establish the differentiation phenotype of mature
chondrocytes.
Induction of the NF-B transcription factor and the cyclin inhibitor
p21Waf1/Cip1 is usually associated with stress or injury. In
chondrocytes, as other cells, NF
B has been shown to attenuate
Fas-mediated apoptosis (Beg et al.,
1995
). Therefore, in the epiphyseal growth plate it may play a
permissive role, by allowing the cells to reach maturity while inhibiting
entry into the apoptosis pathway. The p21Waf1/Cip1 protein blocks
the cell cycle and therefore may have a role in the FGF-induced G1 arrest. In
this respect, its rise may serve a similar function to the fall in the levels
of the Id1 protein, whose downregulation has been associated with decreased
mitotic activity in chondrocytes (Asp et
al., 1998
). The levels of NF
B in chondrocytes of transgenic
mice expressing this mutation are dramatically elevated throughout the
epiphyseal growth plates compared to those of control littermates. The
phenotypic consequence of the G380R mutation is restrained chondrocyte
proliferation and maturation, leading to inhibition of bone growth and
dwarfism (Segev et al., 2000
;
Deng, 1996
). It remains to be
determined to what extent the excessive expression of the pleiotropic
activator NF
B is responsible for this phenotype.
Members of the Id family of transcription factors function as general
inhibitors of terminal differentiation by directly inactivating basic
helix-loop-helix proteins (bHLH), thus controlling cell growth
(Massari and Murre, 2000). Id1
and Id3 were already suggested to be involved in the control of proliferation
and differentiation of cartilage (Asp et
al., 1998
). Most recently, Id1 and Twist, a b-HLH transcription
factor family member, and a known downstream effector of NF-
B
(Tickle, 1998
), were suggested
to regulate the expression of FGFR2 during osteoblast maturation. In this
system, FGF stimulation was proposed to induce Twist, which in turn is thought
to inhibit FGFR2 and is counteracted by the action of Id
(Rice et al., 2000
). Moreover,
loss-of-function mutations in the Twist gene in humans result in the
Saethre-Chotzen syndrome, which is characterized by craniofacial and limb
anomalies (Dixon et al., 1997). Although most patients with this syndrome have
Twist-related mutations, some patients with an overlapping phenotype have a
mutation either in FGFR3 or FGFR2
(Paznekas et al., 1998
).
Drosophila Twist is also thought to inhibit DFR1, the fly FGF
receptor homologue (Shishido et al.,
1993
). In addition, NF-
B inhibits signaling by BMP4
(Tickle, 1998
), a factor that
directly regulates the expression of Id1
(Hollnagel et al., 1999
).
BMP4, on the other hand, is downregulated by FGFR3 in the growthplates of
transgenic mice harboring the Achondroplasia mutant FGFR3
(Naski et al., 1998
), which is
consistent with their excessive NF
B levels
(Fig. 3C). We hypothesize that
the FGF-induced upregulation of NF-
B and reduction in Id1 in RCS cells
mark their entry into the differentiation pathway. This signaling network is
schematized in the partial working model presented in
Fig. 6, where NF-
B
together with Id1 and a b-HLH protein, such as Twist, interact to turn off
FGFR3.
|
FGFR3 localization, the cytoskeleton and growth control
Chondrocytes produce a thick cartilage matrix containing collagen and
sulfated proteoglycans (Cancedda et al.,
1995), which can serve as a substrate for focal adhesions.
Although the functional importance of the clustering of FGFR3 in focal
adhesions is unclear, it may be analogous to the aggregation of FGFR1 in focal
adhesions isolated from endothelial cells using immobilized beads coated with
a synthetic RGD tripeptide or with fibronectin
(Plopper et al., 1995
). FGFR2
is expressed, although not in this location in RCS cells (not shown),
suggesting selectivity for FGFR3. The FGFR3-FGF complexes are most likely
active in the focal adhesions, as evidenced by the increased
tyrosine-phosphorylation in their immediate vicinity and the subsequent
alterations in their structure. Among the most prominent
focal-adhesion-resident signaling complexes are tyrosine-specific kinases
including focal adhesion kinase (FAK) and members of the Src family of
cytoplasmic tyrosine kinases, as well as several other proteins including
tensin, paxillin and Cas, which can be phosphorylated on tyrosine residues
(Vuori, 1998
;
Cary and Guan, 1999
). Basal
tyrosine phosphorylation of focal adhesion proteins is essential for their
formation and maintenance. However, increased tyrosine phosphorylation of
focal adhesions, such as that which occurs in RCS cells upon FGF signaling,
can disrupt these structures, as was previously shown for cells expressing the
oncogenic form of Src kinase
(Rohrschneider, 1980
;
Volberg et al., 1991
).
Interestingly, the Src family of kinases is also a potential substrate for FGF
receptors (Yayon et al.,
1997
), and activation of FAK was observed in response to FGF
stimulation (Klint and Claesson-Welsh,
1999
).
Upon activation of FGF signaling, FGFR3 selectively disappears from the
focal adhesions, leaving vinculin behind. This ligand-receptor complex is
internalized and targeted for degradation with a half-life of 30 minutes
(Monsonego-Ornan et al.,
2000), which corresponds with the observed disappearance of FGFR3
from the focal adhesions within 1 hour and the decrease in the
tyrosine-phosphorylation level within a similar time frame. These events are
subsequently (> 1 hour later) followed by dissolution of the focal adhesion
microstructure that together with changes in cytoskeleton-associated proteins
such as Ezrin (Tsutita and Yonemura, 1997;
Bretscher, 1999
) may lead to
the observed reorganization of the actin network and cell rounding. We
speculate that activation of FGFR3 at the substrate adhesion sites is the
trigger for its removal, which may contribute to the cells' detachment from
the substrate.
Multiple studies have shown that cell cycle events require signals provided
by both soluble factors and the cytoskeleton and that these effects are
usually restricted to the G1 phase of the cell cycle
(Assoian and Zhu, 1997). In an
early study, Folkman and Moscona (Folkman
and Moscona, 1978
) demonstrated that cell shape is tightly coupled
to DNA synthesis and growth. In RCS cells, changes in FGF receptor
localization were already observed 1 hour after induction with FGF9, whereas
changes in cell shape and cell cycle distribution were detected later, after 6
and 10 hours, respectively. It is tempting to speculate that the cell-surface
localization of the activated FGFR may determine the nature of its signal.
FGFR in focal adhesions may activate growth arrest whereas extrajunctional
FGFR may lead to mitogenesis. It also remains a challenge to elucidate whether
growth arrest and cell shape are coupled or represent independent consequences
of FGF induction.
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Acknowledgments |
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