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INTRODUCTION |
Basic fibroblast growth factor
(bFGF)1 mediates many
biological processes including cell proliferation, differentiation,
angiogenesis, and wound healing (1). It is a member of a family of at
least 12 heparin-binding proteins. The activity of bFGF is mediated by
binding to heparan sulfate proteoglycans (HSPG) and to high affinity,
cell surface receptor tyrosine kinases (FGFR).
bFGF can be regulated by HSPG, which serve as low affinity receptors
for bFGF (2). The role of HSPG in modulating bFGF activity has been
described at many levels (2-8). In their role as a co-receptor for
bFGF, HSPG participate in a ternary complex with bFGF and FGFRs
(9-12). This complex has high affinity for bFGF, with
Kd values ranging from 10
10 to
10
12 M. The high affinity binding of bFGF has
been attributed to a slow dissociation of bFGF from the ternary complex
(9, 13). The generation of stable, high affinity bFGF·FGFR complexes
is probably a major mechanism leading to HSPG-dependent
bFGF activity. In addition, bFGF has been localized to the
extracellular matrix, associated with HSPG (6, 14, 15). Release of bFGF
by matrix degradation may serve as a mechanism for mobilizing bFGF in
response to injury or tissue reorganization (16, 17).
Studies using HSPG-deficient cells have demonstrated that bFGF can bind
FGFR and induce activity in the absence of HSPG, yet HSPG appear
necessary for physiologic, bFGF-induced cellular response (3, 4, 5,
11). HSPG do not always lead to potentiation of bFGF. Endothelial
cell-derived HSPG have been demonstrated to have an inhibitory effect
on smooth muscle cell proliferation, indicating that HSPG are able to
sequester bFGF from the cell surface (6). Furthermore, HSPG have an
emerging role in the intracellular processing of bFGF in which HSPG are
thought to stabilize bFGF through endosomal and lysosomal trafficking
(8).
Since HSPG can influence bFGF binding and activity, much research has
focused on the regulation of HSPG as a major control mechanism for
bFGF. Studies on differentiating neural cells have demonstrated changes
in binding affinity of HSPG for bFGF, attributable to differential HSPG
expression (18). Further, studies in vivo have demonstrated
localized HSPG expression at the leading edge of healing corneal wounds
(19). The corneal stroma has a highly defined tissue architecture,
where cells make contact via gap junctions between collagen lamellae
(20). The stromal cells do not constitutively express HSPG. This
suggests a role for intercellular contact as a regulatory mechanism for
differential HSPG expression where cells sense the environment via
contact with adjacent cells and matrix components. Hence, intracellular
signaling mediated by cell density changes could potentially modulate
HSPG expression. Since the cornea is both avascular and devoid of
direct lymphatic supply and thus elicits the wound response without
significant, direct influence of blood and lymph, it is hypothesized
that the corneal stroma is very sensitive to localized changes in
intercellular contact and extracellular matrix organization. These
changes might signal the cells at the wound edge to express HSPG and
provide stromal fibroblasts with a mechanism for bFGF modulation.
In the present study, the regulation of bFGF binding to the cell
surface of stromal fibroblasts was analyzed and correlated to changes
in bFGF-induced proliferation. Cells were cultured at various cell
densities to model changes in intercellular contact that would be
predicted to be associated with wound healing. Binding of bFGF to the
cell surface was measured, and expression of FGFR1 and syndecan 4 was
determined. In addition, bFGF-induced cell proliferation was measured
at various cell densities. Results from these experiments showed that
bFGF-stimulated cell proliferation was cell
density-dependent in stromal fibroblasts. Furthermore, bFGF
binding was dramatically reduced as cell density increased. These
studies suggest that cell-cell contact modulates HSPG expression and
thus, controls bFGF binding and activity in corneal stromal fibroblasts. These findings might have implications for wound healing,
development, and tissue regeneration.
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EXPERIMENTAL PROCEDURES |
Reagents--
Human recombinant bFGF was from Scios/Nova
(Mountainview, CA). 125I-Bolton-Hunter reagent for bFGF
labeling was purchased from NEN Life Science Products. Human
recombinant epidermal growth factor was from Promega (Madison, WI), and
human recombinant TGF-
1 was from R & D Systems (Minneapolis, MN).
Monoclonal mouse anti-bovine FGF-R1(flg) was from Calbiochem.
Monoclonal mouse anti-bovine bFGF was purchased from Upstate
Biotechnology, Inc. (Lake Placid, NY). Mouse anti-syndecan-4 antibody
was a gift from Dr. John Couchman (University of Alabama, Birmingham,
AL). Secondary antibodies, sheep anti-mouse, and goat anti-rabbit
IgG-horseradish peroxidase were purchased from Sigma. Heparinase III
was purchased from Sigma, and heparinase I was a gift from Dr. Ram
Sasisekharan (MIT, Cambridge, MA). Chondroitinase ABC and keratanase
II/endo-
-galactosidase were from Seikagaku (Ijamsville, MD). Fetal
bovine serum and cell culture reagents were obtained from Life
Technologies, Inc. Collagenase A from Clostridium
histolyticum was purchased from Roche Molecular Biochemicals.
Disuccinimidyl suberate was purchased from Pierce.
Cell Culture--
Stromal fibroblasts were isolated from rabbit
corneas (Pel-Freez Biologicals, Rogers, AS). Epithelial and endothelial
layers were removed (21), and stromas were minced with a razor and digested for 3 h with 2 mg/ml of collagenase A (22). Cells were cultured for 3 days with 10% FBS in Dulbecco's modified Eagle's medium (DMEM), low glucose (1% amino acids, 1%
penicillin/streptomycin), and maintained in DMEM containing 4% FBS.
After cells reached confluence, they were used for experiments. For all
experiments, cells were trypsinized and plated at various cell
densities in DMEM containing 4% FBS and allowed to adhere for 4 h. Cells were incubated in DMEM containing 10% FBS and cultured for
20-24 h prior to treatment.
Binding of 125I-bFGF--
Preparation of
125I-bFGF with 125I-Bolton-Hunter reagent was
conducted as described by Nugent and Edelman (9). Equilibrium binding was measured on cultures at 4 °C in binding buffer (DMEM, low glucose, with 25 mM HEPES, 0.05% gelatin) (5, 9). Cultures were incubated for 2.5 h at 4 °C with 125I-bFGF and
rinsed with ice-cold binding buffer three times. bFGF bound to HSPG was
released with rapid washes of a high salt buffer (2 M NaCl,
20 mM HEPES, pH 7.4) and phosphate-buffered saline (PBS).
Receptor-bound bFGF was released with a 10-min wash with a high salt,
low pH buffer (2 M NaCl, 20 mM
NaCH3COOH, pH 4.0), followed by a quick rinse with PBS.
Bound 125I-bFGF was quantified by counting in an Auto-Gamma
5650
-counter (Packard Instruments, Downer's Grove, IL).
Nonspecific binding of 125I-bFGF was measured by adding
unlabeled bFGF (278 nM) prior to the addition of
125I-bFGF. To determine the effects of removing specific
glycosaminoglycans on bFGF binding, cells were treated with specific
glycosaminoglycan-degrading enzymes prior to conducting the binding
studies. Cells were treated with heparinase I (1.5 µg/ml),
chondroitinase ABC (0.1 units/ml), or keratanase/endo-
-galactosidase
(0.1 unit/ml). The enzymes and glycosaminoglycan digestion products
were removed by washing with binding buffer, and 125I-bFGF
binding was measured.
Identification of FGFR1 by Immunoblot--
To determine the
relative expression of FGFR at various cell densities, cells were
plated at various cell densities as described and incubated at 37 °C
for 24 h. Cultures were homogenized by scraping in extraction
buffer (20 mM HEPES, 1% Triton X-100, 10% glycerol, 2 mM phenylmethylsulfonyl fluoride), and nuclei and insoluble
debris were removed by centrifugation at 12,000 × g for 10 min (5). Cell extracts were subjected to SDS-PAGE (7%) under
reducing conditions, and the proteins were transferred to Immobilon-P
membrane (Millipore Corp., Bedford, MA). The membrane was blocked with
PBS (1% Tween 20), 5% nonfat milk for 1 h at room temperature,
and the antibody to FGFR1 (1:1000) was incubated with the membrane for
1 h at 37 °C. After rinsing three times with PBS (0.1% Tween
20), the membrane was incubated with anti-mouse IgG-horseradish
peroxidase (1.5:5000) secondary antibody and rinsed three times with
PBS, 0.1% Tween 20. Visualization was conducted by enhanced
chemiluminescence using Renaissance reagents from NEN Life Science Products.
Determination of FGFR Identity by Cross-linking--
To further
identify FGFR, bFGF·FGFR complexes were covalently cross-linked by
disuccinimidyl suberate (5). FGFR and bFGF were identified by
immunoblots. Cells were plated at high density and cultured for 24 h as described. The bFGF binding assay was performed as described
above, using unlabeled bFGF. Then cell layers were washed three times
with PBS. Fresh PBS (5 ml) was then added to cultures, and
disuccinimidyl suberate (300 µM) was added to cross-link
bFGF·FGFR complexes. Culture dishes were gently shaken for 20 min at
room temperature, and the reaction was quenched by adding 500 µl of
buffer (500 mM Tris-HCl, pH 8.0, 1.0 M
glycine). Cells were rinsed three times with PBS and homogenized by
scraping in extraction buffer (20 mM HEPES, 1% Triton
X-100, 10% glycerol, 2 mM phenylmethylsulfonyl fluoride).
Nuclei and insoluble debris were removed by centrifugation at
12,000 × g for 10 min. Cell extracts were subjected to
SDS-PAGE (7%), proteins were transferred to Immobilon-P membrane, and
the membrane was blocked with PBS (1% Tween 20), 5% nonfat milk for
1 h at room temperature. For FGFR analysis, immunoblots were
performed as described. For bFGF visualization, blots were hybridized
with anti-bFGF (1:1000) using the procedure described for FGFR.
Density Dependence of Mitogen-stimulated
Proliferation--
Mitogen activity was assessed by determining the
relative amount of acid phosphatase activity present in each culture
after incubation with various mitogens (23). The level of acid
phosphatase has previously been demonstrated to relate directly to cell
number in a variety of systems. We performed controls in our studies to
ensure that acid phosphatase levels showed a linear relationship with
corneal fibroblast cell number under our culture conditions. Cells were
plated at several cell densities and incubated for 24 h, as
described. Cultures (t = 24) were treated with PBS,
bFGF (0.56 nM), TGF-
(0.4 nM), epidermal
growth factor (1 nM), or FBS (to a total FBS concentration
of 20%) and incubated for 24 h (t = 48). Cell
number in replicate cultures was determined by a Coulter counter. One
plate was rinsed with PBS and placed at
20 °C to serve as the
initial cell density condition for acid phosphatase activity
determination at t = 24. At t = 48, cells were rinsed with PBS and placed at
20 °C for 2 h. The
amount of cellular acid phosphatase activity was quantified in each
culture to determine relative cell numbers. Cells were lysed with
buffer (0.1% Triton X-100, 100 mM NaCH3COO, pH
5.5) and assayed for acid phosphatase activity upon the addition of
para-nitrophenol phosphate. The role of HSPG in regulating
proliferation was determined by treating cultures at cell density of
14,000 cells/cm2 with 1.0 units/ml heparinase III for
1 h prior to the addition of bFGF.
Determination of Syndecan-4 by Immunoblot--
Cells were
cultured for 24 h at various cell densities as described above.
Cells were digested with heparinase III, extracted, and subjected to
SDS-PAGE (8.5%). After transfer of proteins to polyvinylidene
difluoride, the membrane was blocked with PBS (0.1% Tween 20), 5%
nonfat milk overnight and hybridized with anti-syndecan-4 antibody
(1:500) for 1 h at 37 °C. The membrane was then washed three
times with PBS (0.1% Tween 20) and hybridized with an anti-mouse IgG-horseradish peroxidase secondary antibody (1.5:10,000) for 1 h
at 37 °C. After three washes with PBS (0.1% Tween 20), syndecan-4 was visualized as described above.
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RESULTS |
Cell Density Dependence of bFGF-stimulated Proliferation--
The
corneal stroma has a defined architecture of cells, collagen lamellae,
and associated proteoglycans. The cells comprising this tissue
communicate via gap junctions (20, 24). To investigate the role that
cell-cell communication plays in modulating the mitogenic response of
corneal fibroblasts, growth factor-induced cell proliferation was
analyzed in cells at different densities. To determine if
responsiveness was dependent on initial plating density or on cell
density in general, we examined mitogen-induced cell proliferation in
cultures plated at specific cell densities and in cultures plated at a
single low density and then grown to various higher densities. To
conduct the experiments, cells were cultured at a particular density
for 24 h, thereby establishing density-dependent
levels of intercellular contact. Then the cultures were treated with
epidermal growth factor, TGF-
, bFGF, or serum. From replicate
cultures, cell number was determined with a Coulter counter to yield
the exact cell density at the time of growth factor inducement. Cell
proliferation was assayed by determining relative levels of acid
phosphatase activity. Specifically, the percentage of proliferation was
determined by measuring acid phosphatase activity at the time of
mitogen administration and again after 24 h and comparing the
changes to those observed for untreated controls. bFGF stimulated cell
proliferation at all cell densities tested (Fig.
1). However, at cell densities between
12,000 and 16,000 cells/cm2, bFGF-stimulated cell
proliferation was dramatically increased (190-280%) compared with
that observed at other cell densities (40-100%). This effect was
experimentally determined five times. Epidermal growth factor
stimulated cell proliferation at every density tested, resulting in
cell numbers 5-25% higher than untreated controls. In contrast,
TGF-
only slightly stimulated cell proliferation (20%) at the
highest cell density (30,000 cells/cm2) and had no effect
at the other cell densities evaluated. Cultures treated with fetal
bovine serum exhibited an increased stimulation of proliferation of
30-60% compared with untreated controls at all densities.

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Fig. 1.
Cell density dependence of bFGF-stimulated
cell proliferation. Corneal stromal fibroblasts were plated at an
array of cell densities. After 24 h of incubation
(t = 24), cultures were treated with bFGF (0.56 nM) ( ), epidermal growth factor (1 nM)
( ), TGF- (0.4 nM) ( ), or FBS (20%) ( ). Cell
numbers were determined in replicate cultures to determine the cell
density at the time of mitogen administration. Additional cultures at
t = 24 were rinsed with PBS and frozen overnight at
20 °C for direct comparison of acid phosphatase activity levels
with those of treated cultures. Treated cells were then cultured an
additional 24 h (t = 48), rinsed with PBS, and
frozen at 20 °C for 2 h. All cells were thawed, and relative
acid phosphatase activity was determined. Individual experiments for
each mitogen are summarized graphically. Percentage of stimulation is
given by changes in absorbance at 440 nm as follows: % stimulation = 100 × ((t = 48treated t = 24) (t = 48untreated t = 24))/(t = 48untreated t = 24). The results
presented represent the average of duplicate or triplicate
determinations. S.E. values are given and are not visible if smaller
than the symbol. Similar results were observed in eight
additional experiments.
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For cultures plated at a single low density, grown to various higher
densities, and then assayed, the results demonstrated a similar profile
to Fig. 1, with bFGF stimulating proliferation at all cell densities
and with marked increased stimulation at intermediate cell densities
(not shown).
bFGF Binding Decreases as Cells Proliferate--
Since mitogenic
responsiveness to bFGF varied with cell density, it was possible that
the expression of cell surface binding sites also varied. Thus, we
investigated the role of cell density as a mechanism by which bFGF
binding to the cell surface can be modulated. As cells grew from low
density (2000 cells/cm2) to high density (19,000 cells/cm2) over 96 h (Fig.
2A), we observed a 3-5-fold
decrease in binding of bFGF per cell (Fig. 2B). Cells in low
density cultures (500-3000 cells/cm2) bound 0.5-1.4 × 106 molecules of bFGF/cell, whereas cells at higher
densities (20,000-35,000 cells/cm2) bound 150,000-250,000
molecules/cell. bFGF binding to cells plated at various cell densities
and cultured for 24 h exhibited a similar 3-5-fold increase in
bFGF binding per cell at low cell density compared with high cell
density (Fig. 2C).

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Fig. 2.
bFGF binding per cell decreases as cell
density increases. A, cells were plated at low cell
density (1000 cells/cm2) and cultured for 4 days.
B, bFGF binding (250 pM) at equilibrium
conditions (4 °C) was measured at each indicated time point. Total
binding was determined by extracting bound 125I-bFGF using
a high salt, low pH buffer. bFGF bound was normalized to cell number,
as determined by a Coulter counter. C, cells were plated at an array of
cell densities and cultured for 24 h. bFGF binding per cell was
measured as described for A. Results represent the average
of quadruplicate determinations. S.E. values are given and are not
visible if smaller than the symbol. Similar results were
observed in six additional experiments.
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bFGF Binding to FGFR1 and HSPG--
Both FGFR and HSPG can
independently bind bFGF and, can act together to form a high affinity,
ternary complex (7, 9, 10). We sought to determine the role of each in
binding bFGF by measuring the relative amount of bFGF bound to each
class of sites. The amount of bFGF bound to these two classes of
binding sites can be measured by using selective extraction conditions to remove HSPG-bound bFGF from FGFR-bound bFGF (2). Binding of bFGF was
measured at bFGF concentrations of 27.8 pM to 1.11 nM in cell cultures at densities of 4000, 15,800, and
25,900 cells/cm2. At the highest concentration of bFGF
tested (1.11 nM), cells, at densities of 4000, 15,800, and
25,900 cells/cm2, bound a total of 13.3 × 105, 7.19 × 105, and 5.11 × 105 molecules of bFGF/cell, respectively (Fig.
3A). bFGF bound to HSPG sites
at the same densities was measured to be 13.0 × 105,
6.70 × 105, and 4.46 × 105
molecules/cell, respectively (Fig. 3B).
Kd values for HSPG interactions were calculated to
be 7.8 ± 6.7 and 7.8 ± 7.8 nM for cell
densities of 4000 and 15,800 cells/cm2, respectively. The
receptor-bFGF interactions were calculated to have
Kd values of 23 ± 15, 200 ± 77, and
384 ± 104 pM for cell densities of 4000, 15,800, and
25,900 cells/cm2, respectively. Receptor number/cell was
calculated to be (6.9 ± 0.7) × 104, (6.3 ± 0.8) × 104, and (10.0 ± 1.2) × 104
molecules/cell.

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Fig. 3.
Analysis of bFGF binding to stromal
fibroblasts. Cells were plated at various densities and cultured
for 24 h, and then cell number was determined by a Coulter counter to yield the cell
density at the time of binding analysis. Cell densities were determined
to be 4000 ( ), 15,900 ( ), and 31,600 ( ) cells/cm2.
A, total, specific binding of bFGF was determined by
measuring 125I-bFGF binding at the specified concentrations
under equilibrium conditions (4 °C). Specific, HSPG-bound bFGF
(B) was determined as the high salt releasable fraction, and
specific, FGFR-bound bFGF (C) was determined as the
remaining high salt, low pH releasable fraction. Calculation of
dissociation constants and number of total binding sites was performed
using Kaleidograph (version 3.0.1) by fitting the data to a generalized
single site binding model as follows: [FGFb] = [FGFf] × [Rtot]/(Kd + [FGFf]), where [FGFb] is the amount of bFGF
bound, [FGFf] is the amount of bFGF free,
[Rtot] is the total number of receptors per cell, and
Kd is the equilibrium dissociation constant (5).
Results represent the average of triplicate determinations. S.E. values
are given and are not visible if smaller than the
symbol.
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Receptor Expression Does Not Vary with Cell Density--
The
binding data suggested that the levels of FGFR1 expression did not vary
significantly. Further, differences in the Kd for
bFGF·FGFR1 binding at different cell densities appeared to result
from HSPG. To further evaluate the changes in bFGF binding to receptor,
immunoblots were performed to compare relative levels of FGFR1
expression at different cell densities using an antibody raised against
the extracellular domain of FGFR1. One prominent 73-kDa species was
detected, and the expression level did not appear to vary significantly
with cell density (Fig. 4A).
This band was also detected using a second monoclonal antibody (Upstate Biotechnology; data not shown).

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Fig. 4.
Identification of FGFR by immunoblot and
cross-linking analysis. A, Cells were plated at various
densities and cultured for 24 h, and cell number was determined
with a Coulter counter to yield cell density at time of extraction of
replicate cultures. Whole-cell lysates were subjected to SDS-PAGE,
transferred to a polyvinylidene difluoride membrane, hybridized with a
monoclonal antibody to FGFR, and visualized by ECL. Cell densities were
determined to be 1200, 2500, 7900, 12,100, and 24,900 (cells/cm2) for VL, L, ML, M, and H, respectively.
B, cells were plated at an intermediate density, cultured
24 h, rinsed with PBS, and treated with disuccinimidyl suberate in
Me2SO/PBS to induce cross-linking. Cell layers were
extracted and subjected to SDS-PAGE and transferred to a polyvinylidene
difluoride membrane. Blots were then hybridized with a monoclonal
antibody to bFGF or monoclonal antibody to FGFR and then visualized
with ECL.
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Determination of 73-kDa FGFR1--
Although the immunoreactive
73-kDa band does not directly correspond to any described isoforms of
FGFR1, FGFR isoforms have previously been shown to be subject to
differential expression and processing yielding a range of molecular
masses of 55-145 kDa (25-28). To further identify the 73-kDa species
as an FGFR, we performed immunoblots for bFGF cross-linked to its
receptor and visualized both bFGF and FGFR1 bands by enhanced
chemiluminescence. To one half of the membrane, an anti-bFGF monoclonal
antibody was hybridized and visualized. This detected a band at 91 kDa, corresponding to the combined molecular masses of bFGF (18 kDa) and the
73-kDa species (Fig. 4B). Also detected were bands at 18 and
36 kDa, corresponding to bFGF and bFGF dimers, respectively. Cross-linked bFGF to FGFR1 was also detected in samples not treated with bFGF, indicating the presence of endogenous bFGF. To the other
half of the membrane, an anti-FGFR1 monoclonal antibody was hybridized
and visualized. This detected a 73-kDa species and a 91-kDa species
only when cells were treated with disuccinimidyl suberate (Fig.
4B). Control blots hybridized only to secondary antibody did
not show any bands (not shown).
Heparinase Treatment Abrogates bFGF Binding and bFGF-stimulated
Proliferation--
The changes in bFGF binding versus cell
density were not mediated by different isoforms of FGFR nor altered
levels of FGFR1 expression. From the binding data, HSPG appeared to be
the major regulatory mechanism by which these cells modulate bFGF
binding. To test this hypothesis, cell cultures at various cell
densities were digested with enzymes to degrade specific classes of
glycosaminoglycans. Binding was measured and compared with untreated
cell cultures. At a cell density of 15,300 cells/cm2,
digestion with chondroitinase ABC and keratanase
II/endo-
-galactosidase resulted in bFGF binding levels relative to
control of 94 and 99%, respectively (Fig.
5). Heparinase I digestion resulted in complete loss of specifically bound bFGF. Similar results were obtained
for the cell density of 30,600 cells/cm2. Interestingly, at
low cell density (3000 cells/cm2), chondroitinase ABC and
keratanase II/endo-
-galactosidase digestion resulted in levels of
bFGF binding relative to control of 59 and 80%, respectively.
Heparinase I completely abrogated specific binding of bFGF to these low
density cultures, as in higher density conditions.

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Fig. 5.
Measurement of bFGF binding after enzymatic
digestion of glycosaminoglycans. Cells were plated at various
densities and cultured for 24 h, and cell number was determined
with a Coulter counter to yield cell density at the time of enzyme
treatment. Replicate cultures were treated for 1 h at 37 °C as
follows: control, untreated (filled bar);
heparinase I (1.5 mg/ml) (negligible); chondroitinase ABC (0.1 mg/ml)
(cross-hatched bar); or keratanase II/endo- -galactosidase
(0.1 mg/ml) (open bar). Binding of 125I-bFGF
(0.22 nM) was conducted as described above, and
specifically bound bFGF at the various conditions is represented as
molecules bound per cell. Results represent the average of triplicate
determinations. S.E. values are given and are not visible if smaller
than the symbol.
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To correlate bFGF binding interactions to the
density-dependent proliferation profile induced by bFGF
(Fig. 1), we treated cultures within the critical density range of
10,000-20,000 cells/cm2 with heparinase III prior to
stimulation with bFGF. The percentage of proliferation was determined
by measuring the changes in acid phosphatase activity at the time of
enzyme digestion to those 24 h later and comparing the changes to
those observed for undigested controls. Treatment with bFGF resulted in
a 139% stimulation of proliferation. Cultures digested with heparinase
III and then treated with bFGF resulted in only 8% stimulation of
proliferation. Digestion with heparinase III alone resulted in 22%
less proliferation than untreated controls, further indicating the
presence of endogenous bFGF (Fig. 6).

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Fig. 6.
Abrogation of bFGF-induced cell proliferation
by heparinase pretreatment. Cells were plated and cultured 24 h. Cell number was determined with a Coulter counter to yield cell
density at time of heparinase III/bFGF treatment. Replicate cultures at
a cell density of 14,000 cells/cm2 were treated with
heparinase III (1.0 unit/ml) for 1 h at 37 °C. To heparinase
III-treated or -untreated control cultures, bFGF (0.56 nM)
was added, and cultures were incubated for 24 h. Stimulation of
proliferation is represented as described above. Results represent the
average of triplicate determinations. S.E. values are given and are not
visible if smaller than the symbol.
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Syndecan-4 Expression Decreases at Higher Cell
Densities--
Differential HSPG expression was predicted to be the
regulatory mechanism by which these cells change their binding capacity for bFGF. Since syndecan-4 has been demonstrated to be a major cell
surface HSPG (29), its expression was measured at different cell
densities. Cell extracts were pretreated with heparinase III to release
the syndecan-4 core protein and then subjected to SDS-PAGE and
immunoblot analysis. The results demonstrate a sharp decrease in
syndecan-4 expression at higher cell density as indicated by the
decrease of the immunoreactive band with an apparent molecular mass of
48 kDa (Fig. 7). The results shown are
from cell densities of 2, 10, and 25 × 103
cells/cm2.

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Fig. 7.
Expression of syndecan-4 decreases at higher
cell density. Cells were plated and cultured 24 h. Cell
number was determined with a Coulter counter to yield cell density at
the time of heparinase digestion. Cells were extracted and digested
with heparinase III as described under "Experimental Procedures."
Samples were subjected to SDS-PAGE, transferred to a polyvinylidene
difluoride membrane, hybridized with a monoclonal antibody to
syndecan-4, and visualized by ECL. Cell densities were determined to be
2, 10, and 25 × 103 cells/cm2 in
lanes 2, 3, and 4,
respectively. As a control, heparinase III alone was loaded in
lane 1.
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DISCUSSION |
The importance of co-receptors, like HSPG for bFGF binding, has
been widely studied (2, 3, 9, 30-32), yielding significant insight
into the mechanism and regulation of growth factor activity. Interestingly, many tissues contain high levels of several growth factors yet do not appear to be constitutively responding to their stimuli. For example, the aqueous humor has been reported to contain high concentrations of bFGF, yet the corneal endothelial cells and
keratocytes appear quiescent in situ (33). These
observations suggest that cellular response might be regulated by the
competency of the cells and not solely by the presence of growth
factors and their effectors. In this study, we investigated the action of bFGF in primary corneal stromal fibroblast cells cultured at different cell densities. We hypothesized that cell density serves as a
mechanism by which cells sense their extracellular environment and
control their response to growth factors. We found that bFGF binding
was inversely proportional to cell density. Furthermore, we observed
that bFGF-stimulated cell proliferation was also
density-dependent. Thus, cell density might coordinate
extracellular stimuli with cellular context to provide a concerted
cellular response.
As cell density increased with proliferation, binding of bFGF per cell
decreased 3-5-fold. This effect was not simply a result of ligand
depletion at high densities, since analysis of the binding data
revealed ligand depletion of less than 20%, except at very low
concentrations of bFGF (27.8 pM). The binding data further indicated that cells plated at various cell densities and cultured for
24 h, as compared with those proliferating to specific densities, exhibited the same density-dependent changes in bFGF
binding. These data indicate that cell density, whether achieved by
plating conditions or by culturing to specific densities, provides the cells a mechanism by which bFGF binding may be modulated. Our data
emphasize a role for HSPG as the primary means by which these cells
modulate bFGF binding. However, it is important to note that the
changes we observed might not be a direct response to cell density
per se but instead could reflect the action of additional factors present in our cell culture system.
Experiments measuring bFGF·FGFR binding suggested that receptor
number did not change with cell density. Indeed, immunoblot analysis
showed no density-dependent changes in FGFR1. In contrast, receptor affinity was increased at lower cell densities. Determination of FGFR dissociation constants revealed a 10-20-fold higher affinity at the lowest cell density compared with the highest cell density (23 pM versus 200-384 pM). Thus,
receptor affinity appeared to increase with increased expression of
HSPG sites, consistent with a cooperative role for HSPG in stabilizing
bFGF·FGFR complexes. We also have investigated HSPG core protein
expression and identified syndecan-4 as an HSPG species regulated by
cell density.
Our data indicated a 73-kDa species as a bFGF binding protein in our
culture system. This molecular mass does not correspond to any known
isoform of FGFR1, although several low molecular mass receptor species
have been described (25-27). Due to the size, we speculate that this
protein represents a previously unknown or processed form of FGFR1 or
perhaps a proteolytically cleaved ectodomain of FGFR1 liberated by our
extraction methods. The epitope of one of the monoclonal antibodies
used to detect FGFR was generated using the ectodomain of FGFR1 and
that of the other has not been determined. We therefore cannot formally
conclude whether this 73-kDa protein is a fragment or a full-length
receptor species. We also detected the presence of endogenous bFGF.
While not directly measured, evidence that endogenous bFGF exists is
2-fold. First, cells treated for cross-linking in the absence of
exogenously added bFGF exhibited a 91-kDa band corresponding to the sum
of the 73-kDa FGFR and the 18-kDa bFGF. Second, pretreatment of cell cultures with heparinase I resulted in lower levels of basal cell proliferation compared with untreated controls, indicating a loss of
sensitivity to endogenous bFGF-induced proliferation.
Changes in cell phenotype resulting from different levels of cell-cell
contact have been shown to modulate HSPG expression (18, 34-37). In
studies on proliferating smooth muscle cells, HSPG expression was
determined to depend on cell phenotype (38). Specifically, HSPG from
cells at high density had longer glycosaminoglycan chains than those at
lower cell density. HSPG derived from nondividing smooth muscle cell
cultures had 10-fold higher antiproliferative potency than HSPG
from proliferating cells. Other investigators have measured changes in
affinity and binding of bFGF in differentiating neural cells and have
determined that the changes described were due to modulation of HSPG
expression (18). Together, these studies strongly correlate cell
phenotype with differential HSPG expression.
Biochemical analyses of corneal wounds in vivo have
demonstrated changes in proteoglycan synthesis, in which heparan
sulfate and chondroitin sulfate/dermatan sulfate expression increase
and keratan sulfate decreases (19, 39, 40). Alternatively, in situ hybridization has been successfully employed to detect HSPG at the leading edge of wounds. In vitro studies of wound
healing have largely utilized linear wounds of confluent cell layers. These studies have shown somewhat similar data as in vivo,
although they have been limited by the sensitivity of the assays used
to describe the biochemistry of the wound healing process. It was our
aim to model intercellular communication levels that might be
associated with wound healing and development. It has been previously
demonstrated that functional gap junctions and focal adhesions form in
these cells (20, 24), indicating a putative sensing mechanism of cell
density. Further, in corneal fibroblasts cultured at very low cell
density, a phenotypic change has been demonstrated, whereby a
population of fibroblasts become myofibroblasts, exhibited by detection
of
-smooth muscle actin (41). This phenotypic change would
presumably allow for motility and, eventually, wound contraction.
Indeed, changes in motility induced by fibroblast growth factor have
been demonstrated in cells at low cell density, while the same growth
factor induces mitogenesis at higher cell density (42). The mechanism
of this conversion in cellular response was not resolved. Our data show
markedly increased bFGF binding in cells at low cell density,
potentially indicating a sequestration and protection of bFGF. The
implications of this are not yet clear. It is possible that bFGF could
induce cell migration until a critical cell density is achieved,
whereby the mitogenic effects of bFGF would be dominant and cell
proliferation would be stimulated. Thus, the maximal binding observed
at low cell density did not correlate to maximal mitogenic potential.
Only at intermediate densities were the mitogenic effects of bFGF
maximal. Both the mitogenic effects and the binding effects were
HSPG-mediated as demonstrated by the abrogation of bFGF-induced
proliferation and of bFGF binding after heparinase treatment on the
cell cultures.
Our data indicate that changes in bFGF binding and activity at
different cell densities are HSPG-mediated. HSPG, by serving as low
affinity, high capacity binding sites for bFGF, are required for the
unique functions of this and possibly other heparin-binding growth
factors (7). By protecting and sequestering bFGF, HSPG may serve to
provide a developing or healing tissue with a constant source of bFGF,
with the cellular response being dictated by numerous stimuli. HSPG
expression could function to provide spatial and temporal regulation of
bFGF and other heparin-binding growth factors to coordinate biological processes.