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,21
* Harvard Medical School, Boston, Massachusetts 02115, USA;
Harvard-MIT Division of Health Sciences and Technology,
Division of Bioengineering and Environmental Health, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, USA
1Correspondence: MIT, 77 Massachusetts Ave., Building 16–561, Cambridge, MA 02139, USA. E-mail: ramnat{at}mit.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: FGF receptor • thermodynamic parameters • extracellular matrix
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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2
3)
. FGF family members
bind to two types of receptors: a high-affinity signaling receptor
present on the cell surface and a low-affinity binding receptor present
both on the cell surface and in the extracellular matrix (ECM)
(2
, 4
5
6)
. The high-affinity signaling receptors are the
fibroblast growth factor receptors (FGFR), of which there are four
(FGFR1–FGFR4) (5
, 7)
. The low-affinity receptor is the
heparin-like glycosaminoglycan (HLGAG) component of heparin sulfate
proteoglycans (HSPG) (5
, 8
9
10
11
12)
.
Understanding the nature of the interactions of FGF-2 (basic FGF) with
FGFR and the role of HLGAGs in this process is a fundamental issue that
is still poorly understood (8)
. This is complicated by the
fact that HLGAGs are complex polysaccharides present not only on the
cell surface, but also in the ECM. As a result, they interact with
FGF-2 both on the cell surface and in the ECM compartments (8
, 13
, 14)
. Thus, HLGAGs can play many roles along the trajectory
of FGF-2 signaling events. HLGAGs provide a locale for a stable
reservoir of FGF-2 in the ECM that can be released, upon appropriate
stimuli, for cell signaling (15)
. FGF-2 bound by HLGAG in
the ECM is protected from heat, acid, and protease degradation
(16)
. HLGAGs have been implicated in the induction
of FGF-2 dimerization or in the stabilization of self-associated FGF-2
molecules (11
, 17
18
19)
. This activity is thought to be
important as a prerequisite for FGFR clustering. Finally, HLGAGs are
thought to participate in interactions between FGF-2 and FGFR in a
ternary signaling complex on the cell surface that could also involve
FGFR:FGFR interactions directly induced or stabilized by
HLGAG (20
, 21)
. In physiological systems, HLGAGs
have been shown to be important in various normal and pathological
processes (3
, 22)
. However, an ongoing debate in the
literature centers around the nature of the role of HLGAGs in FGF-2
signaling and whether their presence is an absolute requirement for the
activity (23
24
25
26)
.
In this study, we propose a framework to study signaling by FGF-2
through FGFR1, in both the presence and absence of HLGAGs. Results
presented herein indicate, as has been suggested by others (23
, 24)
, that HLGAGs are not absolutely required for FGF-2 signaling
but that HLGAGs facilitate signaling at a much lower FGF-2
concentration than is possible in its absence. More important, we
demonstrate that the absence of FGF-HLGAG interactions can be
compensated by increasing the FGF-2 concentration to achieve signaling
in the absence of HLGAG. Similarly, HLGAG can be used to compensate for
the reduction in the interactions between FGF-2 and FGFR. Through
site-directed mutagenesis, we have explored the contribution of the
HLGAG and FGFR binding residues of FGF-2 in the signaling process and
how changes in the binding energy resulting from a mutation can be
compensated. The experimental data provide a basis for the development
of a thermodynamic formalism, enabling the determination of parameters
for FGF-2 signaling as a function of both FGF-2 and HLGAG
concentrations. We present a framework for systematically analyzing
these compensatory effects and discuss how nature may take advantage of
these principles in physiological and pathological processes.
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MATERIALS AND METHODS |
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Site-directed mutagenesis
FGF-2 was produced as a soluble protein in BL21(DE3)
Escherichia coli host, using the pET14b system (Novagen,
Madison, Wis.). This construct has a histidine tag (six consecutive
histidines) and a thrombin cleavage site in a 21 amino acid
amino-terminal leader sequence, which constitute a high-affinity site
for Ni2+ that can be cleaved with thrombin. The
mutations were introduced into FGF-2 by the overlap extension
polymerase chain reaction (PCR) methodology developed by Higuchi et al.
(27)
. Briefly, two primary PCR reactions (12 cycles) were
set up using the original FGF-pET14b construct as template, followed by
secondary PCR reactions using the primary PCR products as the template.
The amplified secondary PCR product was cloned into pET-14b expression
vector (Novagen) using NdeI and SpeI restriction
enzymes. After amplification and cloning in pET-14b vector, the
sequence of the mutated genes was verified using Sequanase (U.S.
Biochemicals Inc., Cleveland, Ohio). The plasmid containing the gene in
pET-14b was isolated, purified, and used to transform the host
BL21(DE3) cells.
Expression, isolation, and purification of mutant FGF-2 in
E. coli
Overnight cultures (100 ml) of BL21(DE3) (Novagen) containing
the FGF-2 genes in pET-14b were grown to an OD600
of 0.5, induced with 1 mM iso-phenyl-thio-galactoside (IPTG) for 2 h and harvested. The cell pellet was resuspended in
1/20th volume binding buffer (20 mM Tris, 500 mM
NaCl, 5 mM imidazole). The resuspended culture was placed in an ice
bath, sonicated for 2 min using a Branson sonicator (model no. 450,
power 3, 50% pulse; CT) and centrifuged at 4°C and 15,000 x
g for 30 min. The supernatant was purified by
Ni2+ affinity chromatography using Sepharose 6B
Fast Flow resin covalently linked to nitrilotriacetic acid (Novagen).
Briefly, the resin was charged with five column volumes of 200 mM
NiSO4 and equilibrated with five column volumes
of binding buffer. A 6–10 ml sample was then applied, followed by 12
ml binding buffer, 9 ml 15% elution buffer (20 mM Tris, 500 mM NaCl,
200 mM imidazole), and 10 ml 100% elution buffer. Typically, 1 mg of
purified FGF was obtained per liter of culture. Purity of the eluted
protein was determined by sodium dodecyl sulfate-polyacrylamide gel
electrophoresis analysis with a Mini Protean II electrophoresis
apparatus (Bio-Rad, Hercules, Calif.). Visualization of proteins in
gels (12% gels) was accomplished with 0.1% Coomassie blue stain or
silver stain (Bio-Rad). Heparin-POROS chromatography was used to
confirm the loss of heparin binding for the mutants K26A, K119A/R120A,
and K125A. Circular dichroism was performed on each of the mutant
proteins and shown to be identical with wild-type FGF-2 protein (data
not shown).
FGF-2 assay (ELISA)
Samples of standard (Scios, FGF-2) and test (recombinant and
mutant) proteins were prepared by serial dilution in phosphate-buffered
saline in the concentration range of 1–200 ng/ml. They were added to a
96-well microtiter plate and incubated at 37°C for 2 h to coat
the wells. The wells were washed with water and incubated with blocking
buffer (borate-buffered saline containing 0.05% Tween 20, 1 mM EDTA,
0.25% BSA, 0.05% NaN3). Fifty microliters of
murine mAb 1 (mAb 11.1, which binds to linear sequence residues 21–27,
concentration 1000 ng/ml) was added to the FGF-2-coated wells and
incubated at 37°C for 2 h. After washing and blocking, 50 µl
of HRP-conjugated anti-mouse immunoglobulin G 1 antibody (Pierce
Chemicals) was added at a concentration of 1000 ng/ml and incubated at
37°C for 2 h. After washing and blocking the wells, they were
incubated with 150 µl of a substrate for HRP (3, 3', 5, 5'
tetramethylbenzidine; Pierce) for 30 min at room temperature. One
hundred microliters of 1 M
H2SO4 was added to stop the
reaction. The plate was read in a microtiter plate reader at 450 nm and
the readings were plotted vs. the standard FGF concentration to
determine the concentrations of the test samples.
Cell culture
BaF3 and F32 cells were a generous gift of Dr. David Ornitz
(Washington University, St. Louis, Mo.). These cells were maintained in
suspension culture in RPMI 1640 with 10% calf serum, 10%
WEHI-3-conditioned media, 2 mM L-glutamine, 100 U/ml penicillin, and
100 µg/ml streptomycin. Cultures were grown in 25
cm2 flasks in a 37°C/5%
CO2 incubator and were passaged 1:10 by dilution
thrice weekly. WEHI-3 cells (ATCC, TIB 68) were grown in suspension
culture in RPMI 1640 medium supplemented with 10% calf serum and 0.05
mM 2-mercaptoethanol in a humidified 37°C/5%
CO2 incubator. Supernatants from the WEHI-3 cells
[which produce interleukin 3 (IL-3), an antiapoptotic cytokine for F32
cells] were collected by incubating 3 x
106 cells/ml in their media for 3 days.
Supernatants were recovered by centrifugation at 800 x
g, filtered through an 0.45 µM filter and stored at
-70°C. RPMI 1640, calf serum, 100x
penicillin-streptomycin-glutamine, and 2-mercaptoethanol were from
Sigma Chemical (St. Louis, Mo.).
Proliferation studies
F32 cells were collected from the propagating culture, washed
three times with IL-3-deficient media (i.e., without WEHI-3-conditioned
media), and counted with an electronic cell counter (Coulter, Inc.,
Hialeah, Fla.). The cells were resuspended in IL-3-deficient media to a
density of 1 x 105 cells/ml. This cell
suspension (1 ml) was added to each well of a 24-well tissue culture
plate along with the appropriate concentration of FGF-2 and heparin.
The negative control wells contained the cell suspension without growth
factor or heparin; positive control wells contained the cell suspension
with 2.9 nM (50 ng/ml) recombinant FGF-2 and 29.4 nM (500 ng/ml)
heparin. The cells were incubated for 72 h in the 37°C/5%
CO2 humidified incubator and counted with a
Coulter counter at the experimental end point. These experiments
benefit from the use of the F32 cell line, which lacks cell surface
HLGAGs and has been genetically engineered to express FGFR1
(9)
. The concentration of HLGAG is dependent only on the
amount added in the media; no pretreatment with chlorate or heparinase
was needed to rid the cells of cell surface HLGAGs, allowing an
accurate measurement of concentration effects.
Data analysis
For simplicity, the data have been normalized as follows. The
proliferative index was defined as the increase in cell number for the
experimental case divided by the increase in cell number for the
positive control. The positive control were F32 cells incubated with 50
ng/ml wild-type FGF-2 and 500 ng/ml heparin. The positive control
conditions were found to yield the maximal proliferative response for
FGF-2, using the F32 cells. The proliferative index was independently
determined for each experiment in order to eliminate potentially
confounding factors, such as receptor number per cell, [R] in the
equations that follow.
The model described below predicts that at very high FGF-2
concentrations in the absence of HLGAG (Ho=0), the signaling tends
toward k3/k4. In the
experiments shown (as well as in data that are not shown), the maximal
signaling for FGF-2 in the absence of HLGAG (HLGAG-independent
signaling in Fig. 2
) is ~65% of the maximal signaling for FGF-2 in
the presence of HLGAG (HLGAG-dependent signaling in Fig. 2
). Therefore,
we have set k3 = 0.65 and
k4 = 1 to account for this information.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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shows a FGF-2 cell-mediated response in a concentration-dependent
fashion in the absence of HLGAGs, as described in earlier reports
(23
, 24)
. These experiments were carried out in the F32
cell line, which is devoid of endogenous HLGAGs. Increasing the FGF-2
concentration in the absence of any HLGAGs increases the proliferation
index (cellular response or signal) of the F32 cells. The data suggest
that FGF-2 can signal even in the absence of HLGAGs.
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We propose a simple and unified framework for addressing
HLGAG-dependent and HLGAG-independent signaling by FGF-2, shown in
Fig. 2
A. In this framework, efficient and optimal signaling can be
achieved by a ternary interaction between FGF-2:FGFR:HLGAG. The
signaling in the absence of HLGAGs need not be as efficient, but can
nevertheless be assumed to be a parallel signaling mechanism. Such a
framework can account for the observed HLGAG-mediated FGF-2
dimerization or oligomerization and consequent receptor dimerization
and clustering observed (3)
. Furthermore, no assumptions
regarding the stoichiometry of the interactions were made, enabling us
to develop a set of thermodynamic parameters for the different
interactions based on experimental data.
Under the experimental conditions, different groups of
interactions can be unified into single thermodynamic parameters by
simplifications of the equations in Fig. 2B
. In Fig. 2C
, the thermodynamic parameters k1
and k3 describe FGF-2 signaling in the absence of
heparin. Within k1 are embedded FGF-2:FGFR
interactions, FGFR:FGFR interactions, and FGF-2:FGF-2 interactions.
Similarly, the parameters k2 and
k4 relate to the possible interactions of HLGAG,
FGF-2, and FGFR and the signaling resulting from them. The parameters
k3 and k4 are also a
measure of the efficiency of signaling of the FR and FHR complexes,
respectively. Therefore, the following two equations will be used in
conjunction with experimental data to determine these parameters.
Assuming that the observed saturation results from limiting
receptor concentrations, we can derive from first principles the
relationship between normalized signaling (y) and the initial
concentrations of FGF (Fo) and HLGAGs
(Ho).3Assuming these two parallel reactions are additive, the signaling for
this unified model can be written as:
 |
Wild-type FGF-2
The thermodynamic parameters that govern the interaction between
FGF-2 and FGFR in the absence of HLGAG can be determined from the data
presented in Fig. 1
. Setting Ho = 0 in Equation 3
and using the
values of k3 and k4
described in Materials and Methods, the value of
k1 calculated from the data is 1.08 nM. The
calculated trend for the signaling using this k1
value in Equation 3
is also shown in Fig. 1
. It can be seen that this
model fits the data very well.
Figure 3
shows FGF-2 signaling as a function of FGF-2 concentration in the
presence of 500 ng/ml heparin. Maximal proliferation occurs at an FGF-2
concentration of 0.6 nM (10 ng/ml), corresponding to a 50x reduction
in the amount of FGF-2 needed to reach plateau than when the experiment
is run in the absence of heparin (Fig. 1)
. Furthermore, increasing
either the FGF-2 concentration or the heparin concentration beyond
these points does not alter signaling, confirming that the receptor
concentration on the cell surface is the rate-limiting reactant
(22)
. Using the above determined value for
k1, the value of k2 can be
determined from Equation 3
. The value of k2 for
wild-type FGF-2 is calculated to be 4.0 nM. The signaling calculated
from Equation 3
is plotted on the same graph, and again shows that the
fit between the experimental data and the model is quite good. Using
these parameters for wild-type FGF-2, signaling can be written as:
 |
shows the predicted signaling at this FGF-2 concentration as a function
of heparin concentration.
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|
The experimental data corresponding to an FGF-2 concentration of 50
ng/ml with varying heparin concentration are also plotted in Fig. 4
. At
low heparin concentrations (<0.06 nM), the experimental data indeed
show that the signaling is equivalent to that of signaling in the
absence of heparin at the same FGF-2 concentration, as predicted by the
model. The proliferation index reaches a plateau at a heparin
concentration of 5.8 nM, as predicted by the model.
As mentioned above, Equation 4
predicts a 2-dimensional surface for the
dependence of signaling on FGF-2 and heparin concentration.
Figure 5
shows the dependence of signaling (z axis) on the
concentration of FGF-2 and heparin (x and y axis,
respectively). This surface shown in Fig. 5
allows us to predict the
proliferation response as a function of heparin concentration at a
given FGF-2 concentration, and vice versa. Furthermore, the
experimental data shown in Figs. 1
, 3
, and 4
can be superimposed on
this 3-dimensional plot. Such a figure combines the results of the
modeling and the experimental data into one plot; this format will be
used to present data on the FGF-2 mutants. It is important to note that
in this plot, two sets of data (varying FGF concentrations at Ho=0 and
Ho=29 nM) are used to regress the model parameters, and this is used to
predict the third data set (FGF constant, varying Ho). For this data
set, the predicted data are compared to the experimental data for
varying Ho, keeping Fo constant.
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FGF-2 mutants
Site-directed mutagenesis studies have identified specific sites
on FGF-2 as being involved in HLGAG binding (28
, 29)
or
FGFR binding (30)
. Specific mutants that alter the binding
interaction of FGF-2 to HLGAG or FGFR can therefore be used to further
investigate and corroborate the unified model presented above. The
FGF-2 heparin binding mutants K26A, K119A/R120A, and K125A and the
FGF-2 receptor binding mutants (Y103A and Y111A/W114A) were chosen for
study; the results are described below.
K26A
K26 is situated in the primary heparin binding site and is thought
to form salt bridges with the anionic sulfates of HLGAG (28
, 30
, 31)
. A mutation of K26 to Ala significantly reduces FGF-2
binding to HLGAGs (28
, 30)
. The values for
k1 and k2 were calculated
to be 2.53 and 18.03 nM for K26A in the same way as described for
wild-type FGF-2. Using these values, signaling as a function of K26A
and heparin concentration can be described by Equation 5
.
 |
shows the 3-dimensional graph of the signaling for this mutant as
predicted by Equation 5
. Note that although a significant amount of
binding energy has been disrupted by this mutation, complete activity
can be recovered by increasing the K26A or the HLGAG concentration. The
predicted profile for the dependence of signaling on the K26A and HLGAG
concentration match almost exactly with the experimental data plotted
on the same axes. Comparison of Fig. 6
with Fig. 5
shows that signaling
in the absence of HLGAGs is almost identical, whereas in the presence
of HLGAG, the signaling behavior of this mutant is shifted toward
higher concentrations compared with wild-type FGF-2.
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K119A/R120A
The positively charged cluster of amino acids close to the
carboxyl-terminal region of FGF-2 has also been implicated in HLGAG
binding (28
, 29)
. For instance, the amino acid residues
K119, R120, and K125 are situated in the primary heparin binding site
and are responsible for hydrogen bonding and salt bridge interactions
with HLGAG (28
, 29)
. A double mutation of K119 and R120 to
Ala significantly reduces FGF-2 binding to HLGAGs (28
, 29)
. Experiments with varying K119A/R120A concentrations in the
absence of HLGAG resulted in a k1 of 5.63 nM; in
the presence of HLGAG, k2 was found to be 7.29 nM
(2)
. Using these values, signaling as a function of
K119A/R120A and heparin concentration can be described by Equation 6
.
 |
shows the 3-dimensional graph of the signaling for K119A/R120A as
predicted by the Equation 6
. As with K26A, complete activity can be
recovered by increasing the concentration of K119A/R120A or HLGAG.
Figure 7
also shows that the experimental data fall exactly on the
contours predicted by Equation 6
. A comparison of Figs. 5
and 7
reveals
that the surface profiles are almost identical between the wild-type
FGF-2 and K119A/R120A except for a marginal shift in both the Fo and Ho
concentrations.
|
K125A
A mutation of K125A also is reported to reduce interactions
between FGF-2 and HLGAG (28
, 29)
. However, the values for
k1 and k2 from our
experiments were found to be much higher than wild-type FGF-2 (157 nM
and 84 nM (2)
for K125A. These values for
k1 and k2 indicate that the
proliferative response will be less than that of wild-type FGF-2 at a
given set of concentrations. Using these values, signaling as a
function of K125A and heparin concentration can be described by
Equation 7
.
 |
. As with the previous mutants, the experimental data fit almost exactly
with the model predictions. A comparison of Fig. 8
with Fig. 5
shows
that the signaling behavior of this mutant is shifted toward
significantly higher concentrations of K125A and heparin as compared to
wild-type FGF-2, while the overall trends remain the same.
|
Y103A
Y103A is a well-studied mutant of the primary receptor binding
site on FGF–2. The residue Y103 has been reported to be critical for
the binding of FGF-2 to FGFR, and mutation of this amino acid leads to
loss of activity (30)
. In contrast to native FGF–2, Y103A
only minimally stimulates cell proliferation in the absence of heparin
over the concentration range studied, giving a k1
value of 110 nM, a full two orders of magnitude higher than that of
wild-type FGF-2. However, Y103A is able to signal to the same extent as
wild-type FGF-2 in the presence of 29.4 nM heparin at 29.4 nM Y103A. In
essence, the combination of 29.4 nM heparin and increased Y103A
concentration is able to rescue the dramatic loss of activity that is
seen in the absence of heparin for this mutant. This corresponds to a
k2 value of 16.15 nM (2)
, as
compared to k2 of 1.08 nM (2)
for
wild-type FGF-2. Using these values, signaling as a function of FGF-2
and heparin concentration can be described by Equation 8
.
 |
. As with the previous mutants, the experimental data fit almost exactly
with the model predictions. In contrast to the previous mutants, a
comparison of Fig. 9
with Fig. 5
shows that the signaling behavior of
this mutant is shifted from the wild-type FGF-2 in an asymmetric
manner. The shift is much more dramatic along the Y103A axis and is
shifted to a lesser extent along the heparin axis.
|
Y111A/W114A
Y111A/W114A is a mutation of FGF-2 in the secondary receptor
binding site (30)
. Just as Y103A was an ideal mutant to
investigate the primary receptor binding site, Y111A/W114A is used to
determine whether changes in FGF-2 FGFR interaction at the secondary
site of FGF-2 can be compensated for by an increase in FGF or heparin
concentration. This mutant was inactive at all protein concentrations
both in the presence and absence of heparin (see Discussion).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Compensation of HLGAG and FGF-2 components
Mutations that affect the binding interactions between FGF-2 and
HLGAG (K26A, K119A/K120A) are able to achieve the same magnitude of
cellular response as wild-type FGF-2 in the presence of HLGAG, but at
marginally higher FGF-2 or HLGAG concentrations. This is reflected in
the much higher k2 values compared to wild-type,
especially for K26A. On the other hand, the cellular response in the
absence of HLGAG for these mutations is almost identical, as reflected
in the similar k1 values for these mutants and
wild-type FGF-2. Decoupling the HLGAG-independent and the
HLGAG-dependent cellular responses in our framework has enabled us to
identify the specific interactions that are affected by a mutation. The
K125A mutant, which was expected to affect only the HLGAG binding, was
found to influence the cellular response in both the presence and
absence of HLGAG (k1 and k2
are over 50-fold higher compared to wild-type FGF-2). It is possible
that this mutation affects additional interactions between FGF-2 and
FGFR or influences other interactions, such as FGF–FGF dimerization.
The interaction of FGF with FGFR has been mapped to two different
sites, termed the primary and secondary receptor binding sites on
FGF-2. Residues at the primary binding site (Y24, N101, Y103, L140,
M142) contribute a significant amount of binding energy to the
interaction with the receptor (24)
. Using Y103A as a model
mutant, we show here that the contribution from this residue can be
compensated for by increasing the protein concentrations. On the other
hand, the secondary binding site residues of FGF-2 (amino acids
106–115) (24)
are not conserved in the FGF-2 family
(unpublished observation). These residues (e.g., Y111, W114), although
not contributing significant binding energy, seem to be critical for
FGF-2 activity. A Y111A/W114A double mutant cannot be rescued by any of
the changes in protein or HLGAG concentration in the range tested here
(data not shown). It is possible that a mutation in the secondary
receptor binding site affects k3 and
k4, which govern the progression of the FR or FHR
complex to the measured proliferative response.
Physiological implications
A common requirement shared by all tyrosine kinase receptors is
the formation of a threshold number of appropriately phosphorylated
cytoplasmic domains to stimulate the signaling cascade. This implies
that a threshold number of active receptor-ligand complexes need to be
present on the cell surface for an adequate period of time to result in
effective signaling. Although a thermodynamic treatment of the
signaling events is able to explain and predict several of the
important trends in FGF-2 and FGF-2 mutant signaling, it is important
to point out that the kinetic parameters of the interactions between
FGF-2:FGFR:HLGAG are also very important. The off-rates of FGF-2
binding to FGFR have been shown to be dramatically reduced in the
presence of HLGAGs (32)
, suggesting that the ternary
complex is in the associated state for a much longer period than is the
binary complex between FGF and FGFR. This `duration of complex' might
also be responsible for the differences in the efficiency of signaling
in the absence of HLGAG (k3 vs.
k4) and will be affected by the changes in
concentrations. Recent studies have revealed that the intracellular
transport of FGF-2 from the cell surface to the nucleus is dependent on
HLGAGs (33)
, and this may contribute to differences
observed in cell proliferation. In addition, the role of HLGAG binding
to FGFR and how this affects FGF-2-mediated cellular response remains
to be understood (20)
.
Data presented here clearly show that though FGF-2 can signal in the
absence of HLGAGs, the type and amount of HLGAG can significantly
modulate FGF-2 signaling. HLGAG stabilizes the intermolecular
interactions and provides a method for amplification of the system so
that signaling can occur at lower FGF-2 concentrations than would be
possible in the absence of HLGAG. This provides cells a manner whereby
they can become either more sensitive or less sensitive to the presence
of a given FGF-2 concentration in the extracellular environment.
Physiologically, the cells can up- or down-regulate the expression of
HLGAGs on their surface, and thus modulate the nature and strength of
signaling from the FGF-2 stored in the matrix (34)
.
The data shown here indicate that HLGAG levels can modulate the
activity of FGF-2. This raises an interesting issue in view of the
physiological regulation of HLGAGs by cells themselves. Proliferation
and binding data for FGF-2 have been obtained in a variety of different
cell types under various conditions, sometimes yielding perplexing or
contradictory results (35)
. Could some of these
disparities be a result of differential expression of surface HLGAGs
from cell type to cell type, or even from week to week, in a given
culture? For example, could a primary capillary endothelial cell
culture used at passage 9 have a different complement of cell surface
HLGAG than one used at passage 13, yielding different results as a
consequence? This underscores the utility of the F32 cells to
investigate the role of HLGAG in FGF signaling, where the only HLGAG in
the system is exogenously added.
Although it is clear that HLGAGs can regulate FGF-2 signaling and biological activity, the mechanistic and molecular features of regulation of FGFR activity by HLGAGs are still under investigation. This study provides a framework for the design of experiments investigating the contributions of HLGAGs in the different steps of receptor complex formation leading to cellular activation.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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3 Abbreviations: ECM, extracellular matrix; FGF, fibroblast growth factor; FGF-2, basic FGF; FGFR, FGF receptor; HLGAG,
heparin-like glycosaminoglycan; HRP, horseradish peroxidase; HSPG,
heparin sulfate proteoglycans; IL, interleukin; PCR, polymerase chain reaction.
 |
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Fo, and H Ho
(see text), and simplifying
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Received for publication December 10, 1998. Accepted for publication May 14, 1999.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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 A. Zamfir, D. G. Seidler, H. Kresse, and J. Peter-Katalinic Structural investigation of chondroitin/dermatan sulfate oligosaccharides from human skin fibroblast decorin Glycobiology, November 1, 2003; 13(11): 733 - 742. [Abstract] [Full Text] [PDF]
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 D. Qiao, K. Meyer, C. Mundhenke, S. A. Drew, and A. Friedl Heparan Sulfate Proteoglycans as Regulators of Fibroblast Growth Factor-2 Signaling in Brain Endothelial Cells. SPECIFIC ROLE FOR GLYPICAN-1 IN GLIOMA ANGIOGENESIS J. Biol. Chem., April 25, 2003; 278(18): 16045 - 16053. [Abstract] [Full Text] [PDF]
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A. Facchiano, K. Russo, A. M. Facchiano, F. De Marchis, F. Facchiano, D. Ribatti, M. S. Aguzzi, and M. C. Capogrossi Identification of a Novel Domain of Fibroblast Growth Factor 2 Controlling Its Angiogenic Properties J. Biol. Chem., February 28, 2003; 278(10): 8751 - 8760. [Abstract] [Full Text] [PDF]
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 O. Maltseva, P. Folger, D. Zekaria, S. Petridou, and S. K. Masur Fibroblast Growth Factor Reversal of the Corneal Myofibroblast Phenotype Invest. Ophthalmol. Vis. Sci., October 1, 2001; 42(11): 2490 - 2495. [Abstract] [Full Text] [PDF]
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C.-P. Kwan, G. Venkataraman, Z. Shriver, R. Raman, D. Liu, Y. Qi, L. Varticovski, and R. Sasisekharan Probing Fibroblast Growth Factor Dimerization and Role of Heparin-like Glycosaminoglycans in Modulating Dimerization and Signaling J. Biol. Chem., June 22, 2001; 276(26): 23421 - 23429. [Abstract] [Full Text] [PDF]
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