| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
J. Biol. Chem., Vol. 276, Issue 26, 23421-23429, June 29, 2001
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
,,,,,§
**
From the Division of Bioengineering and Environmental
Health, the § Harvard-Massachusetts Institute of Technology
Division of Health Sciences and Technology, and the Center for
Biomedical Engineering, Massachusetts Institute of Technology,
Cambridge, Massachusetts 02139 and the ¶ Department of Medicine,
St. Elizabeth's Medical Center, Tufts University School of Medicine,
Boston, Massachusetts 02135
Received for publication, November 29, 2000, and in revised form, March 28, 2001
 |
ABSTRACT |
|---|
 TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
For a number of growth factors and cytokines,
ligand dimerization is believed to be central to the formation of an
active signaling complex. In the case of fibroblast growth factor-2
(FGF2) signaling, heparin/heparan sulfate-like glycosaminoglycans
(HLGAGs) are involved through interaction with both FGF2 and its
receptors (FGFRs) in assembling a tertiary complex and modulating FGF2
activity. Biochemical data have suggested different modes of
HLGAG-induced FGF2 dimerization involving specific protein-protein
contacts. In addition, several recent x-ray crystallography studies of
FGF·FGFR and FGF·FGFR·HLGAG complexes have revealed other modes
of molecular assemblage, with no FGF-FGF contacts. All these different
biochemical and structural findings have clarified less and in
fact raised more questions as to which mode of FGF2 dimerization, if
any, is essential for signaling. In this study, we address the issue of
FGF2 dimerization in signaling using a combination of biochemical, biophysical, and site-directed mutagenesis approaches. Our findings presented here provide direct evidence of FGF2 dimerization in mediating FGF2 signaling.
Fibroblast growth factors
(FGFs)1 are involved in a
wide range of physiological processes, including morphogenesis, as well as disease processes such as tumor angiogenesis (1-3). The FGF family
consists of at least 20 members, including the well characterized acidic FGF (FGF1) and basic FGF (FGF2), both of which are potent mitogens of many cell types (4, 5). FGF signaling is mediated primarily
through high affinity interactions with cell-surface FGF
receptors (FGFRs), transmembrane polypeptides composed of immunoglobulin-like and tyrosine kinase domains (6, 7). FGF binding to
different isoforms of FGFR is believed to trigger receptor
dimerization, followed by transphosphorylation of specific tyrosine
residues (8). In turn, phosphorylated tyrosine residues activate other
signaling proteins, leading to cell proliferation, migration, and survival.
For proper presentation to FGFR, FGF2 and other members of the FGF
family interact with heparin/heparan sulfate-like glycosaminoglycans (HLGAGs). Consisting of a disaccharide repeat of glucosamine and uronic
acid, HLGAGs are heterogeneous in length (10-100 disaccharide units)
and chemical composition (including differential sulfation, acetylation, and epimerization of each disaccharide unit) (9-12). Found in the extracellular matrix and on the cell surface as part of
proteoglycans, HLGAGs modulate FGF2 activity by low affinity interactions with specific FGF2- and FGFR-binding sites (13-15), facilitating FGF2 binding to FGFR. HLGAGs promote FGF2-induced activation of FGFR through a number of mechanisms, including regulating the diffusion rate of FGF2 (16, 17) and possibly dictating the
specificity of FGF2-FGFR binding through interactions with both FGF2
and FGFR (18-20).
Another hypothesis is that FGF2 binding to HLGAGs induces ligand
oligomerization, which in turn induces dimerization and
transphosphorylation of FGFR. Biochemical studies have demonstrated
that HLGAGs facilitate FGF oligomerization (21-23); however, due to a
lack of direct evidence, it is unclear whether this biochemical
phenomenon is important for FGF2 signaling. Furthermore, different
modes of FGF-FGF interactions have been observed in various studies,
drawing into question what modes of FGF oligomerization, if any, are
biologically relevant.
Examination of apo-FGF and FGF·HLGAG crystal structures has led to
the proposal of preferential FGF2 self-association in a cis-mode, with substantial protein-protein interactions
between the adjacent molecules (24); and biochemical studies (including chemical cross-linking, analytical ultracentrifugation, and mass spectrometry) support this model (22, 25, 26). However, NMR studies
predict a different mode of FGF oligomerization, viz. a
symmetrical FGF2 dimer with possible disulfide bond formation between
two surface cysteines (27). Furthermore, the recently solved
FGF1·decasaccharide co-crystal points to an FGF
trans-dimer involving no FGF-FGF contacts (28), a mechanism
for dimerization that may or may not extend to other members of the FGF
family, viz. FGF2. More recently, several crystallography
studies of FGF·FGFR and FGF·FGFR·HLGAG complexes, including
FGF2·FGFR1 (29), FGF1·FGFR1 (30), FGF2·FGFR2 (30), and
FGF1·FGFR2 (31), have revealed assemblages of two FGFs bound to two
FGFRs, with no FGF-FGF contacts in the complex. Thus, conflicting
biochemical and biophysical evidence makes it unclear whether FGF
oligomerization is important for signaling through FGFR and, if so,
which dimerization mode of FGF, involving either protein contact or no
protein contact, mediates FGF signaling. This problem is compounded
when one considers that the two recent crystal structures of the
ternary complex involving FGF, FGFR, and HLGAG (32, 33) reveal
different stoichiometries for the complex with markedly divergent geometries.
In this study, we apply conformational studies and molecular
engineering techniques to systematically explore proposed modes of FGF2
oligomerization and to evaluate the importance of FGF-FGF interactions
in signaling. Together, our data suggest that an FGF2 dimer involving
substantial noncovalent protein-protein contact is readily formed and
mediates signaling.
Materials--
Ampicillin,
isopropyl- Site-directed Mutagenesis, Protein Expression, and Purification
of the Cysteine Mutant--
Site-directed mutagenesis was carried out
through a two-step PCR procedure as described previously (34). PCR
products were subcloned into the pCR2.1-TOPO vector (Invitrogen,
Carlsbad, CA). Inserts were subcloned into a variant of the pET14b
expression vector through the NdeI/SpeI sites. To
express recombinant protein, an overnight culture of BL21 cells was
transferred to 500 ml of LB medium supplemented with ampicillin
(400 mg/liter) and allowed to grow with shaking at 37 °C until cell
density reached A600 ~ 0.5. Isopropyl- Oxidative Cross-linking--
Purified protein was
buffer-exchanged into HEPES with 10-kDa molecular mass cutoff membranes
(Millipore Corp., Bedford, MA). Oxidative cross-linking was
performed by incubating 50 µg of protein (30 µM final
concentration) with 750 µM Cu2+
phenanthroline (made from a 1:1 mixture of 25 mM
CuSO4 and 130 mM phenanthroline) in a 100-µl
reaction volume at room temperature for 10 min. Longer incubation times
(up to 2 h) did not significantly increase the amount of oligomer
formed. For heparin treatment, the protein was incubated with 3 µM heparin for 1 h prior to cross-linking. The
protein/heparin ratio was 10:1, which was previously shown to be
optimal for FGF2 dimer formation (25). Other reaction conditions are
indicated in the legend to Fig. 3. The reaction was terminated with 0.1 M EDTA and 10 mM iodoacetic acid. Cross-linked products were analyzed by electrophoresis on 15% nonreducing
SDS-polyacrylamide gels, followed by silver staining.
Conformational Studies--
Conformational studies were
performed with the Insight II package (Molecular Simulations,
Burlington, MA) on a Silicon Graphics workstation. The coordinates of
the FGF2 dimer in the FGF2·FGFR1 crystal structure (code 1CVS) and
that of free FGF2 (code 4FGF) were obtained from the Protein Data Bank.
The sequential dimer was constructed from code 4FGF by translating the
coordinates along the 31-Å axis.
The linker used in the experiment contained a tripeptide with the
sequence GAL. However, since the N and C termini of FGF2 in most of the
crystal structures are disordered, the modeled linker included the
tripeptide sequence and the disordered residues of FGF2. The sequence
of the linker was of the form Cterm-GAL-Nterm, where Cterm and Nterm are the disordered C and
N termini of FGF2, respectively. By deleting residues from the
disordered N terminus, linkers of different lengths could be obtained.
The most optimal structure for each of the linkers was obtained as
follows. Combinations of structures for the linker were generated from
the C terminus of one of the FGF2 monomers to the N terminus of
the other monomer in the receptor-bound and sequential dimer using the
homology modeling of Insight II. A good starting structure from the
randomly generated linker structures in each FGF2 dimer was subjected
to energy minimization with the Newton-Raphson method until
convergence. Potentials were assigned using the consistent valence
force field. Interchanging the N and C termini among monomers did not
lead to significant changes in the model of the cross-linked dimer; and
therefore, it did not affect the interpretation of the results.
Construction of Dimeric FGF2 (dFGF2)--
Based on the results
from the conformational studies, the two DNA sequences of FGF2 were
ligated and subcloned into an expression vector as outlined in Fig.
4A. NdeI/SacI sites were introduced by PCR 5' and
3' to the first sequence, whereas SacI/SpeI sites were introduced to the second. Both the first and second sequences encode FGF2 with the first nine N-terminal residues removed. To facilitate purification of dFGF2, a His6 tag and a thrombin
cleavage site were introduced by PCR 5' to the first sequence, and a T7 tag and another thrombin cleavage site were introduced similarly 3' to
the second sequence. Upon subcloning of the PCR product of the first
sequence into pCR2.1-TOPO (which carries an internal SpeI
site), a SacI/SpeI double digest was performed to
linearize the vector. The PCR product from the second sequence was
subcloned similarly, and the insert was excised by a
SacI/SpeI double digest. Ligation between the
linearized vector and the insert from the second sequence resulted in a
fused DNA of two tandemly linked FGF2 DNA sequences. DNA sequencing was
performed to confirm the identities of the fused DNA sequences. Protein
expression and purification were performed as described above, except
that a T7 affinity column (Novagen, Madison, WI) was used as described by the manufacturer after nickel chromatography. Biochemical studies were performed to ensure that dFGF2 was folded properly. Immunoblot analysis using a monoclonal antibody raised against the native form of
wild-type FGF2 showed that the eluants from the nickel and T7 affinity
chromatographies were recognized by the antibody in a
concentration-dependent fashion (data not shown).
CD Spectroscopy--
dFGF2 was concentrated to 1 µM and buffer-exchanged into 10 mM sodium
phosphate (pH 7.2). CD spectroscopy of dFGF2 was performed in a quartz
cell with a 1-mm path length (Starnz, Atascadero, CA) at room
temperature. Data were recorded in an average of 20 scans between 195 and 260 nm on an Aviv 62SD spectropolarimeter.
Protein Mass Spectrometry--
MALDI mass spectrometry was
completed by diluting a solution of FGF2, FGFR1, and a
heparinase-derived HLGAG decasaccharide (consisting of a trisulfated
disaccharide repeat unit) to 20 µM in 10 mM sodium phosphate (pH 7.0). To 1 µl of this sample was added an equimolar amount of the purified dFGF2 construct. The sample
was allowed to come to equilibrium for 30 min at 4 °C. 1 µl of the
sample was then immediately spotted on the MALDI target with 1 µl of
a saturated sinapinic acid solution in 50% acetonitrile. After drying,
the sample was washed with water, dried under a stream of nitrogen, and
subjected to mass spectral analysis. MALDI mass spectra were acquired
in the linear mode using a Voyager Elite Reflectron time-of-flight
instrument (PerSeptive Biosystems, Framingham, MA) fitted with a 337-nm
nitrogen laser. Delayed extraction was used to increase resolution (25 kV, grid at 91%, guide wire at 0.25%, pulse delay of 350 ns, and low
mass gate at 2000).
SMC Proliferation Assay--
SMCs isolated from bovine aorta
were maintained in propagation medium supplemented with 10% bovine
calf serum, 2 mM L-glutamine, and antibiotics.
The proliferation assay of SMCs, as measured by tritium incorporation,
was performed as follows. Cells were split at 95% confluence and
seeded onto 24-well plates at 1 ml/well. After 24 h, cells were
serum-starved in medium supplemented with 0.1% bovine calf serum for
another 24 h. An appropriate amount of growth factor was added to
eight wells for each protein concentration tested. 75 mM
sodium chlorate was added to half of the wells for each condition.
After 21 h, [3H]thymidine (1 µCi/ml) was applied
to each well and incubated for 3 h. Cells were washed with
phosphate-buffered saline, and 0.5 ml of 1 M NaOH was
subsequently added. The contents of each well were transferred to
scintillation vials filled with 5 ml of ScintiSafe Plus 50%
scintillation fluid (Fisher, Fair Lawn, NJ). Total
[3H]thymidine incorporation was measured by liquid
scintillation counting.
HUVEC Survival Assay--
HUVECs in passage 3 or 4 were cultured
on 1% gelatin-coated tissue culture plates in medium M199
(BioWhittaker, Inc., Walkersville, MD) supplemented with 20% fetal
bovine serum. After 24 h, HUVECs were trypsinized briefly at
37 °C, washed twice with phosphate-buffered saline, and resuspended
in medium containing 0.5% fetal bovine serum and 1% bovine serum
albumin. The cells were seeded at a density of ~1-2 × 104/well onto 96-well plates coated with fibronectin-like
polymer (Sigma). Appropriate amounts of growth factor were added to the wells using a multichannel pipette. Each experimental condition was
tested in six different wells. Cell viability was accessed after
18 h using a cell titer 96 proliferation assay (Promega) by
measuring absorbance at 490 nm.
Angiogenic Assay in the Rat Cornea--
Pellets containing
sucralfate with FGF2 or sucralfate alone were prepared as described by
Kenyon et al. (37). Briefly, suspensions of sterile FGF2
solution containing appropriate amounts of FGF2 (5 and 20 µg)
and dFGF2 (5 µg) were prepared and speed-vacuumed for 5 min. 10 µl
of 12% Hydron in ethanol was added, and the suspension was deposited
onto an autoclave-sterilized nylon mesh. The mesh was stacked between
two layers of fiber covered with a thin film of Hydron. After drying on
a sterile Petri dish for 30 min, the fibers of the mesh were pulled
apart. With the aid of a dissecting microscope, uniformly sized pellets
were selected from ~200 pellets produced. Each pellet contained
~1.5 and 6 pmol of FGF2 or 0.7 pmol of dFGF2. Control pellets
containing no FGF2 were also prepared.
For pellet implantation, male Harlan Sprague-Dawley rats (400-450 g;
n = 5) were anesthetized with ketamine (80 mg/kg) or xylazine (10 mg/kg). Using an operating microscope, an intrastromal linear keratotomy was performed with a surgical blade
(Bard-Parker No. 15, Becton Dickinson Labware, Franklin Lakes,
NJ) parallel to and 2 mm away from the limbus. A lamellar
micropocket was dissected toward the limbus. A single pellet was placed
at the base of the pocket with jeweler's forceps. On day 6 after the
implantation, the corneal angiogenesis was photographed with a slit
lamp, and the area of angiogenesis was assessed as described (37).
Framework for This Study--
The three-dimensional structure of
FGF2 has been thoroughly elucidated by a variety of biophysical
techniques, including solution NMR and crystallography (13, 27-33,
38-40). All have pointed to roughly the same basic structure for FGF2,
whether free, bound to its HLGAG ligand, or complexed with the
receptor. An analysis of all of these structures suggests that three
orthogonal surfaces exist on FGF2 (Fig.
1). As indicated in Fig. 1, the first
surface has been implicated in binding of FGF2 to its high affinity
protein receptor. Through rigorous biochemical and site-directed
mutagenesis studies, a second orthogonal surface has been implicated in
HLGAG binding. The third surface, orthogonal to both of the first two, has been implicated in FGF2 oligomerization.
Biochemical and structural studies have suggested different modes of
FGF2 oligomerization within the third surface both in the presence and
absence of HLGAGs (24, 27, 28). As schematically represented in Fig.
2, three modes of HLGAG-induced FGF2
dimerization have been proposed. Specific protein-protein contacts are
involved in both the sequential and symmetrical FGF2 dimers (Fig.
2A and C, respectively), but not in the
HLGAG-bridged or sandwich dimer (Fig. 2B). It was earlier
demonstrated that FGF2 was capable of dimerization and oligomerization
in the absence of heparin using an amine-specific chemical cross-linker
with an 11-Å spacer (25). This observation is not consistent with the
proposed HLGAG-bridged dimer in Fig. 2B since, in this FGF
sandwich model, there are no residues on neighboring FGF2 molecules
proximal to one another and thus available for covalent cross-linking
with an 11-Å spacer (additional experiments described below also are
not consistent with this dimer mode). Therefore, we focused our initial
experiments on determining whether either of the dimer models involving
protein contacts (represented in Fig. 2, A and C)
is an accurate representation of FGF2 dimerization mediated by
HLGAGs.
Strategy to Investigate FGF Dimerization: Oxidative
Cross-linking through Surface-exposed Cysteine Residues on
FGF2--
To establish the presence of proximal contacts
between FGF2 molecules and to distinguish between different modes of
FGF2 dimerization, we performed oxidative cross-linking experiments
targeting the cysteine residues of FGF2 using copper phenanthroline, an
oxidative agent used widely for disulfide bond formation (41). This
approach is anticipated to probe for atomic distance interactions
between the FGF molecules through the introduction of a disulfide bond between two FGF2 molecules. As discussed below, by taking advantage of
the surface-exposed cysteine residues in FGF2 and through rationally engineering cysteine residues on the surface of FGF2, we systematically explored possible modes of FGF2 dimerization.
Oxidative Cross-linking of Wild-type FGF2--
There are four
cysteines in FGF2, two of which are surface-exposed (Cys69
and Cys87), and two of which are buried in the protein core
(Cys25 and Cys92). The surface positions of the
two exposed cysteines (Cys69 and Cys87) in
wild-type FGF2 are related to each other by 90°. Taking advantage of
the surface-exposed cysteine residues in the wild-type structure of
FGF2, we performed oxidative cross-linking studies to test the proposed
symmetrical mode of FGF2 dimerization in Fig. 2C, as this
model predicts facile cross-linking between two FGF2 molecules (27).
Under mild oxidative conditions, wild-type FGF2 showed very little
oligomer formation in the presence and absence of heparin (Fig.
3A, lanes 1 and
2). (Several control experiments were performed to ensure
authenticity of the data and are described below.) The absence of
significant dimers or oligomers suggests either that the FGF-FGF
interface does not involve molecular contacts or that the contacts are
such that the two surface-exposed cysteines are not at the dimer
interface. Our observation is not consistent with the proposed
symmetrical mode of FGF2 dimerization wherein dimerization is mediated
by disulfide bond formation between Cys69 of each monomer
(27).
Rational Design of the Cysteine Mutant--
In a previous study
(24), we had performed an extensive analysis of all FGF2 crystal
structures available at that time and identified protein-protein
interfaces (p-p' and q-q') that were conserved along the two unit-cell
axes. Based on our analysis, we had proposed an FGF2 dimerization model
in which FGF2 molecules were preferentially self-associated in a
sequential fashion and HLGAG binding stabilized FGF2 dimers and
oligomers that were subsequently presented to FGFR for signaling. In
this model, noncovalent FGF-FGF interactions translated along the
oligomerization direction (Fig. 2B) are expected to lead to
FGF2 oligomerization. If this model indeed describes a mode of FGF
oligomerization, then we would predict that, by substituting cysteine
residues near the protein-protein interface between adjacent FGF2
molecules, intermolecular disulfide bonds could be created under mild
oxidative conditions. The sequential dimer formed in this fashion would
be stabilized by significant protein-protein contacts. As a first step
toward testing this hypothesis, we searched for candidate pairs of
residues in the p-p' interface that, when mutated to cysteine residues,
could generate a disulfide linkage in a facile manner upon oxidative cross-linking. Through conformational studies, we found that optimal disulfide bond formation would be achieved when Arg81 and
Ser100 were mutated into cysteines, as schematically
represented in Fig. 3B. The two introduced cysteines are
located on the opposite sides of FGF2 such that intramolecular
disulfide bond formation would be disfavored. The two original
cysteines, Cys69 and Cys87, were mutated to
serines such that the total number of surface cysteines within the
primary amino acid sequence of FGF2 remained the same. This protein,
with four mutations (R81C/S100C/C69S/C87S), is hereafter referred to as
the cysteine mutant. The cysteine mutant was constructed by
site-directed mutagenesis as described under "Experimental
Procedures." The protein retained biological activity to stimulate
cell proliferation compared with the wild-type protein (data not
shown), suggesting that the introduced mutations did not grossly alter
protein folding.
Oxidative Cross-linking of the Cysteine Mutant--
Under exactly
the same oxidative conditions as applied to the wild-type protein, the
cysteine mutant yielded a markedly higher amount of oligomers. Notably,
the extent of oligomerization was elevated by preincubating the protein
with heparin (Fig. 3A, lanes 3 and 4).
In addition, cross-linking of a FGF mutant that lacked one of these
cysteines at the interface (i.e. either the R81C or S100C
mutation) resulted in significantly less oligomer formation, further
suggesting that the covalent dimer was formed through disulfide bond
formation between the designed Cys81 and
Cys100. Together, these observations strongly support the
sequential mode of FGF2 dimerization and also suggest that the extent
and stability of FGF2 oligomers are increased by binding to HLGAGs (24). Several controls were performed to ensure the authenticity of
specific cysteine-mediated FGF2 oligomerization. Addition of a reducing
agent such as dithiothreitol converted the observed dimers and
oligomers into monomers (Fig. 3C, lane 4),
indicating that the original cross-linking pattern was the result of
disulfide-linked oligomers. Also, oligomerization was abolished when
the cysteine mutant was denatured prior to cross-linking (Fig.
3C, lane 3), suggesting that oligomerization was
mediated through the native structure of the protein and that the
observed oligomers were not formed due to nonspecific protein
aggregation. In addition, since two cysteines (Cys25 and
Cys92) were buried in the protein core, they could
potentially contribute to the observed oligomerization if the protein
was unfolded during cross-linking. To exclude this possibility, the
primary amino acid sequence of the cysteine mutant was further altered
by substituting the two internal cysteines with serines
(i.e. additional C25S/C92S mutations were introduced). The
introduction of these two additional mutations did not change the
cross-linking pattern (data not shown), further indicating that only
the surface-exposed Cys81 and Cys100
contributed to disulfide-induced oligomerization. Taken together, these
oxidative cross-linking studies support a model wherein FGF2 monomers
form sequential dimers via a substantial protein-protein interface, and
this interaction is further promoted by binding to HLGAGs. These
results are consistent with other experimental studies, including
analytical ultracentrifugation of FGF2 with an octasaccharide, chemical
cross-linking, and mass spectrometry of FGF2 with or without addition
of exogenous HLGAGs (21, 22, 25, 26).
Attempts to purify the cross-linked dimers for further biochemical and
biological characterizations were unsuccessful. Therefore, we adopted
an alternative strategy of constructing an FGF2 dimer using a
combination of conformational studies and genetic engineering tools,
enabling us to investigate the biological importance of FGF2 dimers.
This latter point is of special importance since the above biochemical
studies indicate that, although a cis-FGF dimer does
preferentially form in solution, it might form only under
non-physiological conditions (i.e. high protein
concentrations, heparin/protein ratios of 1:10, etc.). However, by
constructing a defined FGF2 dimer and testing its biological activity,
we can determine whether the oligomer mode indicated by the biochemical studies, viz. a cis-dimer involving substantial
protein contact, is able to form an active signaling complex at the
cell surface.
Engineering of a Tandemly Linked FGF2 Dimer through a Linker to
Probe Contact and Non-contact FGF-FGF Interactions: Design of
dFGF2--
Conformational studies of FGF-FGFR interactions led to the
proposal that receptor clustering is facilitated by receptor binding to
an FGF dimer (42). However, the recently solved structures of 2:2
FGF·FGFR complexes, which are proposed to be active signaling complexes, revealed no contact between the two FGF molecules
(29-33).
To determine whether FGF-FGF interaction is important for FGFR binding
and concomitant signaling, we "forced" FGF2 molecules into a
cis-dimerization mode by engineering a dimeric FGF2 protein containing a tripeptide linker. By deleting residues from the N
terminus of the protein, we could control the size of the linker between the two FGFs. Since there are at least 15 N-terminal residues that are disordered in all the FGF2 crystal structures, including the
proposed active FGF2·FGFR crystal structures, we expect that these
deletions would not significantly affect the folding of the protein. To
find the optimal linker sequence length that would facilitate the
distinction between the two modes of FGF-FGF interaction, we explored
combinations of linker sequences with different lengths that could link
the FGF2 monomers in both FGF-FGF interaction modes as described under
"Experimental Procedures." Our conformational studies showed that a
linker with nine residues deleted from the N terminus would optimally
link two FGF2 molecules in the sequential dimer, but would form a
highly constrained structure when linking the two FGF2 molecules
observed in the FGF2·FGFR1 crystal structure (data not shown). A
dimeric protein (dFGF2) containing a tripeptide linker and two FGF2
molecules, linked C- to N-terminal, each with the nine N-terminal
residues removed, was constructed (Fig.
4). This engineered FGF2 dimer is an
ideal candidate to discriminate between a contacting FGF2 dimer and a
non-contacting FGF2 dimer as observed in the FGF2·FGFR1 structure.
The protein was expressed in Escherichia coli and purified
as described under "Experimental Procedures."
Prior to investigating the biological activity of dFGF2, we performed
biochemical studies to ensure that dFGF2 was folded properly. First, as
mentioned under "Experimental Procedures," we assessed the overall
folding of the protein by immunoblotting. The dFGF2 construct stained
at approximately twice the level of wild-type FGF2. In addition, when
the purified protein was heat-denatured in the presence of 1% SDS, the
intensity upon immunoblotting was drastically reduced to the background
levels (data not shown). The above results suggested that dFGF2 was
properly folded with respect to the epitope recognized by this
antibody. To assess the overall secondary structure, the banding
position from near-UV CD spectroscopy of dFGF2 was analyzed. The CD
spectrum showed a negative minimum near 200 nm (Fig.
5), which is characteristic of native
FGF2 (25). In addition, dFGF2 bound to a heparin-POROS column and was
eluted only at 1.8 M NaCl (compared with 1.2 M NaCl for FGF2; data not shown). Not only did this latter result suggest
that dFGF2 was properly folded, it also suggested that dFGF2 had a
higher affinity for HLGAGs than did FGF2, perhaps through a cooperative
binding interaction between the two linked FGF units and the heparin
column. If this is the case, then dFGF2 might have a reduced dependence
on exogenous HLGAGs for activity. We therefore explored the functional
attributes, including the effect of HLGAGs on dFGF2 activity, as
described below.
Stoichiometry of FGF2-FGFR-HLGAG Interactions--
Mass
spectrometry was used to determine whether dFGF2 could compete with
wild-type FGF2 for binding to FGFR2. MALDI mass spectrometric analysis
of dFGF2 yielded a species consistent with the expected mass for dFGF2
of 37,066 Da (data not shown). As a next step, we investigated the
nature of wild-type FGF2-FGFR interactions both in the presence and
absence of HLGAGs. These experiments indicated that, in the absence of
HLGAGs, wild-type FGF2 bound FGFR with a stoichiometry of 1:1 (Fig.
6A), consistent with
FGF·FGFR crystal structures (29, 30). However, addition of an HLGAG decasaccharide (consisting of a trisulfated disaccharide repeat unit
that is known to bind with high affinity to FGF2 and to support FGF2-mediated signaling) resulted in the formation of a detectable 2:2:1 FGF·FGFR·HLGAG complex (Fig. 6B). Addition of
dFGF2 to this complex resulted in the formation of a new 1:2 complex of
dFGF2·FGFR (Fig. 6B, inset). Notably, we could
detect no dFGF2·FGFR species with decasaccharide bound. This species
could be absent because the complex either does not form in solution or
is not ionized and detected under the conditions of this experiment. In
addition, since the ionization efficacies of the various species
undoubtedly differ from one another, with the larger species
(especially those containing the decasaccharide) being less amenable to
ionization than the smaller species, quantitative estimates of the
amount of complex formed in this case are not warranted. However,
detection of a 1:2 dFGF2·FGFR complex indicates that this species
does form at protein levels that approximate those present at the cell
surface.
Together, these results indicate that 1) one molecule of dFGF2 having
protein contact is able to support receptor dimerization; 2) one of the
roles for HLGAGs in FGF binding to FGFR is to support FGF and/or FGFR
oligomerization; and 3) biochemically, one mode of FGF oligomerization
and receptor binding involves a dimer with substantial protein-protein
contact. To determine whether the complexes observed via mass
spectrometry have a biological role, we tested the ability of dFGF2 to
signal both in vitro and in vivo.
Biological Activity of dFGF2--
To test if FGF-FGF contacts are
involved in signaling, dFGF2 was first assayed for its biological
activity in vitro. Mitogenicity of dFGF2 was tested on SMCs
treated with or without chlorate. Because chlorate treatment inhibits
the biosynthesis of HLGAGs and thereby depletes cell-surface HLGAGs,
the dependence of HLGAG binding on the activity of dFGF2 for signaling
can be evaluated. With intact cell-surface HLGAGs (no chlorate
treatment), both wild-type FGF2 and dFGF2 were active in mediating a
proliferative response on SMCs (Fig.
7A). Importantly, the molar
concentrations required to achieve half-maximal proliferation by
wild-type FGF2 and dFGF2 were 270 and 60 pM, respectively.
Hence, dFGF2 exhibited 4.5-fold more activity compared with wild-type
FGF2 in promoting cell proliferation under these in vitro
conditions. In chlorate-treated SMCs, whereas wild-type FGF2 produced
only a moderate response in proliferation, a marked increase in the
proliferative response was exhibited by dFGF2, achieving ~80% of
full proliferation in HLGAG-depleted cells (Fig. 7B). The
results from the SMC proliferation assay suggest a higher potency in
stimulating proliferation and a lower dependence on HLGAG for signaling
by dFGF2.
In addition to SMCs, FGF2 is a potent angiogenic factor well known for
its ability to induce cell survival in endothelial cells. Therefore, we
determined the ability of dFGF2 to promote cell viability in HUVECs.
Using the colorimetric dye, which reflects the mitochondrial integrity
of viable cells, the HUVEC survival assay provides a sensitive way to
measure endothelial cell viability mediated by the growth factors
added. In serum-deprived HUVECs, cell viability was ~50% of that of
HUVECs grown in 10% serum (Fig. 8).
Addition of various concentrations of wild-type FGF2 and dFGF2 partially recovered cell viability in a dose-dependent
manner. Again, dFGF2 was more active than wild-type FGF2 in stimulating survival in HUVECs on a molar basis, consistent with its elevated potency observed in SMCs. Taken together, the biological activity of
dFGF2 from two independent cell types demonstrates that the dimeric
construct binds to and activates FGFR to elicit various downstream
signals as measured by the biological assays.
In Vivo Potency of dFGF2--
To extend the above in
vitro findings, the ability of dFGF2 to induce angiogenesis in an
experimental in vivo model was investigated. The activities
of FGF2 and dFGF2 were compared, side by side, using the rat corneal
pocket assay, the results of which are shown in Fig.
9. As anticipated, control pellets
containing no FGF2 (i.e. no angiogenic stimuli) failed to
induce appreciable angiogenesis (Fig. 9A). FGF2 induced an
angiogenic response in a dose-dependent manner, with little
angiogenesis induced at a protein level of 1.5 pmol (Fig.
9B) and more extensive angiogenesis induced at 6 pmol (Fig.
9C). Thus, the extent of angiogenesis induced by FGF2 is
accurately reflected both by the length of induced vessels and the
circumference of those vessels. Compared with FGF2, dFGF2 induced more
extensive angiogenesis in the corneas of rats at a lower concentration
of 0.7 pmol (Fig. 9D). With dFGF2, induced blood vessels
were longer, of larger circumference, and more plentiful, as measured
by "clock hours" or the extent of angiogenesis around the limbus.
In fact, 0.7 pmol of dFGF2 was a better angiogenic stimulus than FGF2
at an 8-fold higher level, viz. 6 pmol. Thus, the biological
potency of dFGF2, as measured in in vitro cell culture
experiments, was retained in an in vivo animal model, suggesting that the dFGF2 construct is a potent biological
mediator.
FGF·FGF Dimer Involves Protein Contact--
Several signaling
pathways mediated by growth factors and cytokines involve binding of
dimeric or oligomeric ligands to their cell-surface receptors to
facilitate receptor dimerization (43), a key step leading toward
activation of intracellular signaling cascade. For the FGF family of
growth factors, many biochemical studies have pointed to different
modes of dimerization and oligomerization in the presence and absence
of HLGAGs that are essential for signaling. In this study, we examined
three possible modes of HLGAG-mediated FGF2 association (Fig. 2,
A-C) by evaluating earlier and current biochemical
findings. The observation that FGF2 can oligomerize in the absence of
HLGAG (25) is inconsistent with the non-contacting HLGAG-bridged
dimerization mode (Fig. 1B). Our results from the oxidative
cross-linking studies of wild-type FGF2 do not support the proposed
symmetrical dimerization, which is potentially mediated by
intermolecular disulfide bonds (27). Through rational design of a
disulfide-mediated sequential dimer (cysteine mutant) based on earlier
extensive analysis of FGF2 crystal structures (24), we demonstrated
(a) a marked increase in the amount of oligomers formed
compared with wild-type FGF2, which has the same number of surface
cysteines, but at different positions; (b) a higher extent
of oligomerization by preincubating this cysteine mutant with heparin;
and (c) that the observed oligomers involve specific protein
contacts and are disulfide-mediated. The above findings strongly
support a model in which FGF2 molecules self-associate through specific
FGF-FGF interactions in a sequential fashion and that HLGAG serves to
provide a "platform" to stabilize the intermolecular interactions
between FGF2 molecules (24).
dFGF Is Biologically Active--
To test our hypothesis of an
active FGF2 dimer involving protein-protein contact and to distinguish
it from the FGF2 dimer observed in the FGF-FGFR co-crystal structures
that lack protein-protein contact, we constructed a tandemly linked
dFGF2 molecule using conformational studies and genetic engineering
tools. dFGF2 was designed such that the short distance between the two
FGF2 molecules within the dimeric protein would allow for substantial
FGF-FGF interactions while making the non-contacting dimer mode less
favorable and therefore enable us to determine whether a contacting
FGF2 dimer can elicit biological activity. Though mass spectrometry, we
show that dFGF2 interacts with FGFR at a ratio of 1:2, suggesting that
dFGF2 can bind to a dimer of FGFR. Furthermore, these results indicate
that one mode, involving substantial protein contact, by which FGF2 and
its receptor can interact is through the binding of FGFR to an FGF2
dimer. These biochemical findings are supported by the biological
activity of the dFGF2 molecule.
To test whether a contacting FGF2 dimer can elicit biological activity,
dFGF2 was subjected to two independent in vitro assays. As
determined by both the SMC proliferation and HUVEC survival assays,
dFGF2 exhibited elevated biological activity compared with wild-type
FGF2. This effect was especially pronounced in the SMC assays, in which
dFGF2 was severalfold more active than wild-type FGF2 and only 30%
less active in the absence of HLGAGs as in their presence (as opposed
to wild-type FGF2, whose activity was significantly reduced in
the absence of cell-surface HLGAGs). These findings suggest that dFGF2,
in which FGF-FGF interactions are predicted to be substantial, forms an
active signaling complex with the receptor. In addition, proliferation
of chlorate-treated SMCs demonstrated that dFGF2 was less
HLGAG-dependent for signaling. This observation can be
rationalized if one considers that one primary mechanism by which
HLGAGs modulate FGF2 activity is by stabilizing two FGF2 molecules in a
dimer mode to facilitate receptor dimerization. Because dFGF2 is
already dimeric, its dependence on HLGAGs for proper presentation to
the receptor is lower compared with wild-type FGF2. We have also
extended these studies to show that the dFGF2 construct is a potent
pro-angiogenic agent in vivo, much more so than wild-type
FGF, thus providing compelling evidence that the dFGF2 construct,
involving substantial protein-protein contact, forms an active
signaling complex at the cell surface.
Thus, the biochemical, cell culture, and in vivo assays are
consistent with the proposal that a contacting FGF2 dimer is involved in the active signaling complex. These findings appear to be
inconsistent with the different FGF2·FGFR crystal structures, which
show no FGF-FGF interactions. Such an inconsistency may reflect the
inherent complexity and multifaceted nature of the FGF system. One
possible explanation is that the different structural configurations of FGF·FGFR may reflect the different states, viz. "on"
or "off" states, of the signaling complex. A similar observation
has been made in other systems (44, 45). For instance, in the case of
erythropoietin, it has been noted that certain mimetic peptides can
dimerize the receptor, but fail to induce signaling (46) due to the
formation of an inactive complex at the cell surface. Furthermore, the
ultimate biological end point of erythropoietin signaling,
i.e. whether erythropoietin signaling results in
proliferation or differentiation, is sensitive to how the receptors are
brought together (44). Another system in which the mode of dimerization plays a critical role in determining activity is tumor necrosis factor
binding to tumor necrosis factor receptor-1. Unliganded tumor necrosis
factor receptor-1 exists as an inactive dimer, with its catalytic
domains over 100 Å apart (47). Binding of tumor necrosis factor to its
receptor brings the catalytic domains of the receptors proximal to one
another, initiating intracellular signaling cascades (48). Thus,
similar to these cases, it is conceivable that a mode of FGF2
dimerization involving protein-protein interactions could lead to a
cooperative FGF2-FGFR interaction by promoting subsequent
oligomerization and signaling, whereas non-contacting FGF2 dimerization
may lead to an inactive complex.
In addition, it must be noted that other studies have suggested that
monomeric forms of FGF2 may form active signaling complexes (49, 50).
For instance, in a recent study, it was found that covalently linked
complexes of monomeric FGF with a pool of heparin dodecasaccharides
were able to promote cell proliferation in vitro (50).
However, as observed in that study, the covalent FGF·HLGAG complex
was less active than uncomplexed FGF in promoting
[3H]thymidine incorporation. In contrast, the dFGF2
construct presented in this study is several times more
potent in biological assays than wild-type FGF, with reduced dependence
on exogenous HLGAGs for activity. Nevertheless, it is possible that
monomeric complexes of FGF do signal, albeit with less apparent
activity compared with oligomeric forms of FGF2.
In summary, we report here that FGF2 does have a preference to
oligomerize, and the studies contained herein point to the fact that
this oligomerization interface involves protein-protein contact. In
addition, a dFGF2 construct based on these biochemical findings has
potent biological activity, consistent with the hypothesis that FGF
oligomers are potent mediators of FGFR dimerization and concomitant signaling.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-D-thiogalactopyranoside, 1,10-phenanthroline,
sodium chlorate, and dithiothreitol were from Sigma. Human recombinant
wild-type FGF and anti-FGF2 antibody were gifts from Scios Nova
(Mountain View, CA). The pET14b expression vector variant and FGFR were
a generous gift from D. M. Ornitz (Washington University). Heparin
sodium from porcine intestinal mucosa was from Kabi Pharmacia
(Franklin, OH). Ready-Gel (15% polyacrylamide gel) and Bradford assay,
immunoblot assay, and silver staining kits were from Bio-Rad.
-D-thiogalactopyranoside (1 mM)
was added to induce protein expression for 2 h. Protein
purification by nickel chromatography was performed as previously
described (35, 36). Purity of the protein was assessed by
SDS-polyacrylamide gel electrophoresis under nonreducing conditions,
and concentration was determined by Bradford assay using recombinant
wild-type FGF2 as a control.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (32K):
[in a new window]
Fig. 1.
Analysis of various binding sites on
FGF2. The surface of an FGF2 molecule can be approximated as the
faces of a parallelepiped. Of the six faces, two opposite faces
represent the receptor-binding sites (pointing into and out of the
plane of the paper), whereas the other four (denoted as
oligomerizing and heparin binding) represent
directions about which FGF can associate. Note that two of the three
oligomerizing directions are aligned along the same plane. Translation
of FGF2 molecules along these two directions forms the basis of FGF2
oligomerization.
View larger version (14K):
[in a new window]
Fig. 2.
Proposed modes of FGF dimerization.
Either a closed or an opened triangle is drawn
inside each FGF molecule to distinguish different orientations. A
round indentation within FGF represents the heparin-binding
domain. HLGAG is depicted as a chain of beads. A,
two FGF molecules, oriented asymmetrically in cis, bind to
the same side of HLGAG in a "side-by-side" fashion (22, 24, 26).
B, two FGF molecules are oriented in trans to the
axis of HLGAG in a "head-to-head" fashion (28). C, four
FGF molecules interact both in cis and in trans
with HLGAG (27). Note that, for the cis-interaction, the two
FGF molecules are symmetrically related, as opposed to the dimer in
A.
View larger version (21K):
[in a new window]
Fig. 3.
Oxidative cross-linking studies.
A, oxidative cross-linking of wild-type FGF2 and the
cysteine mutant. Wild-type FGF2 was oxidized with (lane 1)
or without (lane 2) heparin as described under
"Experimental Procedures." A minor amount of dimer was detected,
likely resulting from the cross-linking reaction of the unfolded
protein. The cysteine mutant, which was designed based on the model of
FGF2 dimerization (24), was oxidized with (lane 3) or
without (lane 4) heparin under the same conditions as the
wild-type protein. All reaction products were separated by nonreducing
SDS-polyacrylamide gel electrophoresis (15%), followed by silver
staining. Note the extent of oligomerization achieved by the cysteine
mutant compared with the wild-type protein. B, schematic
representation of the protein-protein and protein-HLGAG interactions in
cysteine mutants. Two cysteine mutant molecules are shown, each with
two dimer interfaces as represented by striped (site p) and
white (site p') rectangles. Two solvent-exposed
cysteines (Cys81 and Cys100 as shown near sites
p' and p, respectively) were engineered so they would position in close
proximity with each other at the interface. C, dimerization
and oligomerization of the cysteine mutant were mediated by the native
structure of the protein. Lane 1, cysteine mutant alone;
lane 2, cysteine mutant oxidized without heparin; lane
3, same as lane 2, but heat/SDS-denatured prior to
oxidative cross-linking; lane 4, same as lane 2,
but treated with 1 mM dithiothreitol (DTT). Note
that oxidative cross-linking of the cysteine mutant was abolished by
either denaturing or reducing treatments.
View larger version (35K):
[in a new window]
Fig. 4.
Engineering, cloning, and purification of
dFGF2. A, a scheme is shown for linking two FGF2 genes
and subcloning them into an expression vector for protein expression.
Restriction sites (NdeI, SacI, and
SpeI) were introduced 5' and 3' to FGF2 cDNA by PCR.
B, shown are the results from restriction digestion of the
expression vector with two tandemly linked FGF2 cDNAs. Lane
1, NdeI/SpeI digest of the expression
vector; lane 2, NdeI/SacI digest;
lane 3, SacI/SpeI digest.
kb, kilobase pairs. C, shown is a
schematic of the protein product obtained upon expression of the
genetic construct of A. An N-terminal His tag, a C-terminal
T7 tag, and two thrombin cleavage sequences (shaded
rectangles) are present to facilitate protein purification. The
arrows indicate the positions of thrombin cleavage.
D, wild-type FGF2 (lane 1) and dFGF2 (lane
2) were separated by SDS-polyacrylamide gel electrophoresis under
reducing conditions. The molecular mass sizes are shown on the
left.
View larger version (15K):
[in a new window]
Fig. 5.
Structural properties of dFGF2. The
near-UV CD spectrum of dFGF2 is shown. dFGF2 was concentrated to 1 µM and buffer-exchanged into 10 mM sodium
phosphate (pH 7.2) as described under "Experimental Procedures."
Data were recorded in an average of 20 scans between 195 and 260 nm. Of
note is the characteristic intense negative CD signal observed near 200 nm that is indicative of properly folded FGF2. deg,
degrees.
View larger version (32K):
[in a new window]
Fig. 6.
Competitive binding of dFGF2 to
FGFR2. A, MALDI mass spectrometric profile of a mixture
of wild-type FGF2 and the ectodomain of FGFR2. Observed in the mass
spectrum are the (M + H)+ ions for an FGF2 dimer
(m/z 30,214) and trimer (m/z 45,132), an FGFR2
monomer (m/z 24,888) and dimer (m/z 49,572), and
a 1:1 FGF2·FGFR2 complex (m/z 39,896). The theoretical
molecular masses for FGF2 and FGFR2 are 15,114 and 24,864 Da,
respectively. B, mass spectrum of the FGF2/FGFR2 mixture in
the presence of a homogeneous HLGAG decasaccharide. Addition of
a decasaccharide (Deca) to FGF2/FGFR2 promoted the formation
of a 2:2 FGF2·FGFR2 complex with an observed (M + H)+ ion
at m/z 82,750 (with the decasaccharide) or m/z
79,872 (without the decasaccharide). The (M + H)+ ions for
two dimeric FGFR2 species were also observed. The first (at
m/z 49,592) represents the apo complex, and the second (at
m/z 52,474) is a 2:1 FGFR2·decasaccharide complex.
Inset, mass spectrum of dFGF2 added to the
decasaccharide/FGF2/FGFR2 mixture shown above. Three high molecular
mass complexes were observed: a 2:2 FGF2·FGFR2 complex with or
without the decasaccharide and a 1:2 dFGF2·FGFR2 complex without the
decasaccharide.
View larger version (13K):
[in a new window]
Fig. 7.
SMC proliferation assay. Serum-starved
SMCs were stimulated with the indicated molar concentrations of
wild-type FGF2 ( 
) and dFGF2 (
). SMCs were grown in the absence
(A) or presence (B) of 75 mM
chlorate. After 21 h at 37 °C, [3H]thymidine was
added for 3 h. Cells were harvested and washed, and
[3H]thymidine incorporation was measured. Maximal
counts/min for wild-type FGF2 and dFGF2 were about 6000 and 5000, respectively. Note that the proliferation curve of dFGF2 is shifted
toward the left of wild-type FGF2. The molar concentrations for
half-maximal proliferation by wild-type FGF2 and dFGF2 were 270 and 60 pM, respectively.
View larger version (20K):
[in a new window]
Fig. 8.
HUVEC survival assay. Serum-starved
HUVECs were stimulated with the indicated concentrations of wild-type
FGF2 and dFGF2 or without any growth factor. Cells supplemented with
10% fetal calf serum (FCS) served as a positive control.
After 18 h, cell viability was determined colorimetrically. Note
that both wild-type FGF2 and dFGF2 restored HUVEC viability following
serum starvation and that dFGF2 achieved the same levels of cell
viability at a lower molar concentration compared with wild-type
FGF2.
View larger version (93K):
[in a new window]
Fig. 9.
In vivo potency of dFGF2.
Shown are slit lamp photographs of rat corneas on day 6 after
implantation with Hydron pellets containing no FGF2 as a control
(A), 1.5 pmol of FGF2 (B), 6.0 pmol of FGF2
(C), or 0.7 pmol of dFGF2 (D). The area of pellet
implantation is designated with arrows. The control pellet
did not induce a significant angiogenic response, whereas pellets
containing dFGF2 induced an intense neovascular response originating
from the limbal vessels and reaching the pellet on day 6 after the
implantation. Pellets containing FGF2 (B and C)
induced a less vigorous, but still detectable, angiogenic response on
day 6 after implantation. The extent of corneal angiogenic response is
expressed as linear length and circumferential clock hours in the
table. *, standard error.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
| |
ACKNOWLEDGEMENTS |
|---|
We thank David Berry and Cindy Ku for technical assistance and Isabel de Aos Scherpenseel for advice. We would also like to thank Scios Nova and Dr. David Omitz for making available reagents used in this study.
| |
FOOTNOTES |
|---|
* This work was supported in part by the National Institutes of Health Grant RO1HL59966 (to R. S.), the Burroughs Wellcome Foundation (to R. S.), the CapCure Foundation (to R. S.), a Merck fellowship (to Z. S.), and a Whitaker Health Sciences Fund fellowship (to Z. S.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
** To whom correspondence should be addressed: Center for Biomedical Engineering, MIT, 77 Massachusetts Ave., Bldg. 16-561, Cambridge, MA 02139. Tel.: 617-258-9494; Fax: 617-258-9409; E-mail: rams@mit.edu.
Published, JBC Papers in Press, April 5, 2001, DOI 10.1074/jbc.M010786200
| |
ABBREVIATIONS |
|---|
The abbreviations used are: FGFs, fibroblast growth factors; FGFR, fibroblast growth factor receptor; dFGF2, dimeric fibroblast growth factor-2; HLGAG, heparin/heparan sulfate-like glycosaminoglycan; PCR, polymerase chain reaction; MALDI, matrix-assisted laser desorption/ionization; SMC, smooth muscle cell; HUVEC, human umbilical vein endothelial cell.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
| 1. | Ornitz, D. M. (2000) Bioessays 22, 108-112 |
| 2. | Taipale, J., and Keski-Oja, J. (1997) FASEB J. 11, 51-59 |
| 3. | Hanahan, D., and Folkman, J. (1996) Cell 86, 353-364 |
| 4. | Gospodarowicz, D., Ferrara, N., Schweigerer, L., and Neufeld, G. (1987) Endocr. Rev. 8, 95-114 |
| 5. | Nugent, M. A., and Iozzo, R. V. (2000) Int. J. Biochem. Cell Biol. 32, 115-120 |
| 6. | Jaye, M., Schlessinger, J., and Dionne, C. A. (1992) Biochim. Biophys. Acta 1135, 185-199 |
| 7. | Wilkie, A. O., Morriss-Kay, G. M., Jones, E. Y., and Heath, J. K. (1995) Curr. Biol. 5, 500-507 |
| 8. | Schlessinger, J., Lax, I., and Lemmon, M. (1995) Cell 83, 357-360 |
| 9. | Gallagher, J. T. (1997) Biochem. Soc. Trans. 25, 1206-1209 |
| 10. | Salmivirta, M., Lidholt, K., and Lindahl, U. (1996) FASEB J. 10, 1270-1279 |
| 11. | Turnbull, J., Powell, A., and Guimond, S. (2001) Trends Cell Biol. 11, 75-82 |
| 12. | Sasisekharan, R., and Venkataraman, G. (2000) Curr. Opin. Chem. Biol. 4, 626-631 |
| 13. | Faham, S., Hileman, R. E., Fromm, J. R., Linhardt, R. J., and Rees, D. C. (1996) Science 271, 1116-1120 |
| 14. | Ornitz, D. M., Herr, A. B., Nilsson, M., Westman, J., Svahn, C. M., and Waksman, G. (1995) Science 268, 432-436 |
| 15. | Kan, M., Wang, F., Xu, J., Crabb, J. W., Hou, J., and McKeehan, W. L. (1993) Science 259, 1918-1921 |
| 16. | Dowd, C. J., Cooney, C. L., and Nugent, M. A. (1999) J. Biol. Chem. 274, 5236-5244 |
| 17. | Flaumenhaft, R., Moscatelli, D., and Rifkin, D. B. (1990) J. Cell Biol. 111, 1651-1659 |
| 18. | Guimond, S. E., and Turnbull, J. E. (1999) Curr. Biol. 9, 1343-1346 |
| 19. | Kan, M., Wu, X., Wang, F., and McKeehan, W. L. (1999) J. Biol. Chem. 274, 15947-15952 |
| 20. | Pye, D. A., Vives, R. R., Turnbull, J. E., Hyde, P., and Gallagher, J. T. (1998) J. Biol. Chem. 273, 22936-22942 |
| 21. | Ornitz, D. M., Yayon, A., Flanagan, J. G., Svahn, C. M., Levi, E., and Leder, P. (1992) Mol. Cell. Biol. 12, 240-247 |
| 22. | Herr, A. B., Ornitz, D. M., Sasisekharan, R., Venkataraman, G., and Waksman, G. (1997) J. Biol. Chem. 272, 16382-16389 |
| 23. | Spivak-Kroizman, T., Lemmon, M. A., Dikic, I., Ladbury, J. E., Pinchasi, D., Huang, J., Jaye, M., Crumley, G., Schlessinger, J., and Lax, I. (1994) Cell 79, 1015-1024 |
| 24. | Venkataraman, G., Sasisekharan, V., Herr, A. B., Ornitz, D. M., Waksman, G., Cooney, C. L., Langer, R., and Sasisekharan, R. (1996) Proc. Natl. Acad. Sci. U. S. A. 93, 845-850 |
| 25. | Davis, J. C., Venkataraman, G., Shriver, Z., Raj, P. A., and Sasisekharan, R. (1999) Biochem. J. 341, 613-620 |
| 26. | Venkataraman, G., Shriver, Z., Davis, J. C., and Sasisekharan, R. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 1892-1897 |
| 27. | Moy, F. J., Safran, M., Seddon, A. P., Kitchen, D., Bohlen, P., Aviezer, D., Yayon, A., and Powers, R. (1997) Biochemistry 36, 4782-4791 |
| 28. | DiGabriele, A. D., Lax, I., Chen, D. I., Svahn, C. M., Jaye, M., Schlessinger, J., and Hendrickson, W. A. (1998) Nature 393, 812-817 |
| 29. | Plotnikov, A. N., Schlessinger, J., Hubbard, S. R., and Mohammadi, M. (1999) Cell 98, 641-650 |
| 30. | Plotnikov, A. N., Hubbard, S. R., Schlessinger, J., and Mohammadi, M. (2000) Cell 101, 413-424 |
| 31. | Stauber, D. J., DiGabriele, A. D., and Hendrickson, W. A. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 49-54 |
| 32. | Schlessinger, J., Plotnikov, A. N., Ibrahimi, O. A., Eliseenkova, A. V., Yeh, B. K., Yayon, A., Linhardt, R. J., and Mohammadi, M. (2000) Mol. Cell 6, 743-750 |
| 33. | Pellegrini, L., Burke, D. F., von Delft, F., Mulloy, B., and Blundell, T. L. (2000) Nature 407, 1029-1034 |
| 34. | Higuchi, R. (1990) in PCR Protocols: A Guide to Methods and Applications (Innis, M. A. , Gelfand, D. H. , Sinsky, J. J. , and White, T. J., eds) , pp. 177-185, Academic Press, Inc., San Diego, CA, 177-183 |
| 35. | Ernst, S., Venkataraman, G., Winkler, S., Godavarti, R., Langer, R., Cooney, C. L., and Sasisekharan, R. (1996) Biochem. J. 315, 589-597 |
| 36. | Padera, R., Venkataraman, G., Berry, D., Godavarti, R., and Sasisekharan, R. (1999) FASEB J. 13, 1677-1687 |
| 37. | Kenyon, B. M., Voest, E. E., Chen, C. C., Flynn, E., Folkman, J., and D'Amato, R. J. (1996) Investig. Ophthalmol. Visual Sci. 37, 1625-1632 |
| 38. | Moy, F. J., Seddon, A. P., Bohlen, P., and Powers, R. (1996) Biochemistry 35, 13552-13561 |
| 39. | Zhang, J. D., Cousens, L. S., Barr, P. J., and Sprang, S. R. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 3446-3450 |
| 40. | Eriksson, A. E., Cousens, L. S., and Matthews, B. W. (1993) Protein Sci. 2, 1274-1284 |
| 41. | Bisaccia, F., Zara, V., Capobianco, L., Iacobazzi, V., Mazzeo, M., and Palmieri, F. (1996) Biochim. Biophys. Acta 1292, 281-288 |
| 42. | Venkataraman, G., Raman, R., Sasisekharan, V., and Sasisekharan, R. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 3658-3663 |
| 43. | Heldin, C. H. (1995) Cell 80, 213-223 |
| 44. | Ballinger, M. D., and Wells, J. A. (1998) Nat. Struct. Biol. 5, 938-940 |
| 45. | Jiang, G., and Hunter, T. (1999) Curr. Biol. 9, R568-R571 |
| 46. | Livnah, O., Johnson, D. L., Stura, E. A., Farrell, F. X., Barbone, F. P., You, Y., Liu, K. D., Goldsmith, M. A., He, W., Krause, C. D., Pestka, S., Jolliffe, L. K., and Wilson, I. A. (1998) Nat. Struct. Biol. 5, 993-1004 |
| 47. | Naismith, J. H., Devine, T. Q., Brandhuber, B. J., and Sprang, S. R. (1995) J. Biol. Chem. 270, 13303-13307 |
| 48. | Naismith, J. H., Devine, T. Q., Kohno, T., and Sprang, S. R. (1996) Structure 4, 1251-1262 |
| 49. | Pantoliano, M. W., Horlick, R. A., Springer, B. A., Van Dyk, D. E., Tobery, T., Wetmore, D. R., Lear, J. D., Nahapetian, A. T., Bradley, J. D., and Sisk, W. P. (1994) Biochemistry 33, 10229-10248 |
| 50. | Pye, D. A., and Gallagher, J. T. (1999) J. Biol. Chem. 274, 13456-13461 |
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati What's this?
This article has been cited by other articles:
|
|
 |
|
  M. A. Dupree, S. R. Pollack, E. M. Levine, and C. T. Laurencin Fibroblast Growth Factor 2 Induced Proliferation in Osteoblasts and Bone Marrow Stromal Cells: A Whole Cell Model Biophys. J., October 15, 2006; 91(8): 3097 - 3112. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Dityatev, G. Dityateva, V. Sytnyk, M. Delling, N. Toni, I. Nikonenko, D. Muller, and M. Schachner Polysialylated Neural Cell Adhesion Molecule Promotes Remodeling and Formation of Hippocampal Synapses J. Neurosci., October& |
