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INTRODUCTION |
Fibroblast growth factors
(FGFs)1 bind not only to
their cognate receptors (FGFRs) but also to heparan sulfate
proteoglycans (HSPGs). HSPGs are associated with the cell surface of
many, if not most, cell types. Most known HSPG functions are
contributed by the interactions of the heparan sulfate (HS) chains of
these molecules. Besides growth factors, these HSPGs interact with
various adhesion molecules, protease inhibitors, and enzymes, modifying the spatial distributions and activities of these ligands. Among FGFs,
the interaction with FGF2 has been studied most intensively, and it is
now generally accepted that HSPGs play important roles in FGF2
signaling. Initially, the association of FGF2 with HS has been proposed
to protect this FGF from proteolysis and thermal denaturation (1, 2)
and to serve as a reservoir of growth factor that can be released by
enzymes that degrade the proteoglycans (1). Later, HSPGs were
identified as co-receptors for FGF2, strongly promoting FGF-FGFR
binding and the subsequent activation of the receptor (3, 4). Recently,
genetic studies in Drosophila provided compelling evidence
that HSPGs are essential for FGF signaling in vivo (5).
Although the importance of HSPG in FGF-signaling is well documented,
the nature of the "co-receptor" and the precise mechanisms at work
are less well characterized. Distinctive core protein structures define
two major families of cell surface-associated HSPGs: syndecans and
glypicans (6). Prior work from our laboratory showed that syndecans and
glypican-1 stimulate FGF2-FGFR1 interaction and signaling in K562
cells, at least when co-expressed with receptor in these cells (4).
This agrees with the work of other groups, showing that cell surface
syndecan-1 from Raji cells acts as a positive regulator of FGF2 binding
and signaling (7), that syndecan-2 on human macrophages promotes
FGF2-mediated proliferation (8), and that glypican-1 can stimulate FGF2
signaling (9, 10). Meanwhile, there are also other, contradictory
reports. Syndecans and glypican-1 purified from human lung fibroblast
extracts are unable to promote high affinity binding of FGF2 to FGFR1
(11), overexpression of syndecan-1 in NIH 3T3 cells inhibits
FGF2-induced proliferation (12), and HSPGs purified from endothelial
secretions prevent FGF2 binding to vascular smooth muscle cells and
inhibit FGF2-induced mitogenesis (13). One possible explanation for this discrepancy is that the HSPGs were from different sources and
might have had different compositions. It is well known, indeed, that
the fine structure of HS is cell- and differentiation-specific, and
highly diverse (14, 15). Another possibility is that the membrane
association of HSPG might play a role in promoting FGF2 signaling,
since most of the inhibitory effects of HSPGs reported so far relate to
soluble forms. Syndecans and glypicans are constitutively shed from
cultured cells (16, 17), and shed soluble syndecan ectodomains can also
be found in inflammatory fluid (18), where they appear to act as
inhibitors of FGF2 (19). Finally, sometimes different end point
analyses were used as a measure of receptor activity. In the present
study, we attempted to define the relative importance of these
variables. We used FGF2-induced FGFR1 autophosphorylation as an
end point and tested the activities of HSPGs from different origins,
both in soluble and in membrane-associated form.
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EXPERIMENTAL PROCEDURES |
Plasmid Isolation and Construction--
Clones encoding FGFR1
were isolated from a human embryonic lung fibroblast 
ZAPII phage
library (4). The cDNA of the IgII/IIIc isoform of FGFR1 was used as
a polymerase chain reaction template. The primer
5'-CGGGATCCAGGCCGTCCCCGACCTTGCCT was designed to introduce a
BamHI restriction site at the 5'-end. The
BamHI-SpeI fragment of this FGFR1 cDNA was
blunted at the SpeI site and cloned into the eukaryotic
expression vector pDisplay (Invitrogen, San Diego, CA), using the
compatible BglII and SmaI sites, to produce an N-terminal hemagglutinin A (HA) epitope-tagged version of this receptor.
The cDNAs for syndecan-1 (20) and glypican-1 (21) were cloned into
the episomal expression vector pREP4 (Invitrogen), using the
KpnI and NheI sites and the HindIII
and NotI sites, respectively. The same cDNAs were also
cloned in pcDNA3/neo, using the KpnI and NotI
and HindIII and XbaI sites, respectively.
Chain valence mutants of glypican-1 were generated with the Transformer
site-directed mutagenesis kit (CLONTECH
Laboratories, Palo Alto, CA). The primer (and corresponding antisense
primer) 5'-GACGACGGCAGCGGCTCGGGCGCCGGTGATGGCTG was used to generate the two-chain form (changing Ser488 into Ala488).
The primer (and corresponding antisense primer)
5'-GACGACGGCAGCGGCGCGGGCGCCGGTGATGGCTG was used to generate the single
chain form (changing Ser488 into Ala488 and
Ser490 into Ala490). The
HindIII-XbaI fragments of the mutant cDNAs,
blunted at the XbaI sites, were cloned into the episomal
expression vector pREP4 (Invitrogen), using the HindIII and
blunted XhoI site.
Cell Transfections--
BaF3 cells (generously provided by Dr.
D. M. Ornitz) were routinely cultured in RPMI 1640 medium (Life
Technologies, Inc.) supplemented with 10% fetal calf serum (Hyclone,
Logan, Utah) and 10% WEHI cell-conditioned medium (as a source of
murine interleukin-3). Before transfection, cells were washed with
calcium/magnesium-free phosphate-buffered saline (PBS). For
transfection, 1 × 107 cells were incubated with 30 µg of linearized HA-FGFR1-pDisplay or pDisplay plasmid in 0.5 ml of
calcium/magnesium-free PBS at 4 °C for 10 min. Cells were then
electroporated at 350 V and 960 microfarads (Gene Pulser; Bio-Rad).
After 24-48 h of cell culture, selection was started with 600 µg/ml
G418 (Life Technologies). Two weeks later, stable clones were obtained,
and subclones were established after 1 month. Individual clones were
tested for HA-FGFR1 expression as described under "Western
Blotting." One of the highest expressors, clone B6, was selected for
further experiments.
Namalwa cells were grown in RPMI 1640 medium supplemented with 10%
fetal calf serum. For transfection, 1 × 107 cells
were incubated with 30 µg of linearized syndecan-1-pcDNA/neo, glypican-1-pcDNA/neo, or pcDNA/neo plasmid in 0.5 ml of
calcium/magnesium-free PBS at 4 °C for 10 min. Cells were then
electroporated at 240 V and 960 microfarads. Selection was started
24-48 h later, with 1.5 mg/ml G418. Stable transfection was achieved
after 2 weeks. The selected population was characterized by
quantitative immunofluorescence flow cytometry, using HS and core
protein-specific antibodies (see below).
K562 cells were routinely cultured in Dulbecco's modified Eagle's
medium/F-12 medium (Life Technologies) supplemented with 10% fetal
calf serum. For transfection, 5 × 106 cells were
incubated with 30 µg of syndecan-1-pREP4, glypican-1-pREP4, or pREP4
plasmid in 0.5 ml of calcium/magnesium-free PBS for 10 min at 4 °C.
Cells were then electroporated at 250 V and 960 microfarads. Selection
with hygromycin B (Roche Molecular Biochemicals) at 200 µg/ml
resulted in stable cell populations that were not further subcloned.
The population was characterized by quantitative immunofluorescence flow cytometry, using HS and core protein-specific antibodies. Transfections with pRep4 vectors encoding one-chain (SAA), two-chain (SSA), and wild type (SSS) forms of glypican-1 into Namalwa cells were
performed in similar ways.
Immunofluorescence Cytometry--
Namalwa cells or K562 cells
were incubated with 5 µg/ml 10E4 (monoclonal antibody (mAb)
recognizing HS) or the core protein-specific antibodies BB4 (mAb
recognizing syndecan-1; Serotec Ltd., Oxford, United Kingdom) and S1
(mAb recognizing glypican-1). After 1 h at 4 °C, the cells were
washed with PBS plus 2% bovine serum albumin (BSA) and incubated for
another 1 h at 4 °C with fluorescein isothiocyanate-labeled goat anti-mouse antibodies (Nordic Immunology, Tilburg, The
Netherlands). Cells were washed again with PBS plus 2% BSA and fixed
with formaldehyde. The fluorescence was measured with a FACSort (Becton
Dickinson, Mountain View, CA). The value obtained for cells that were
incubated with fluorescein isothiocyanate-labeled goat anti-mouse
antibodies only was taken as background fluorescence.
Western Blotting--
Proteins were extracted from cells with
lysis buffer (0.5% Triton X-100 in Tris-buffered saline, supplemented
with 1 mM phenylmethylsulfonyl fluoride, 1 µg/ml
pepstatin, 5 µg/ml leupeptin, and 50 mM NaF). Cell
lysates were clarified by centrifugation at 14,000 × g
for 15 min. Supernatants were subjected to 4-12% SDS-polyacrylamide gel electrophoresis (PAGE) (Bio-Rad). Proteins were then transferred for 3 h at 0.5 A to PVDF membranes (Millipore, Bedford, MA).
Membranes were blocked with Tris-buffered saline (pH 7.4) containing
0.2% I-block (Tropix, Bedford, MA) and 0.01% Tween, at 37 °C for
1 h and incubated with the anti-HA mAb 3F10 (Roche Molecular
Biochemicals). HA-FGFR1 was visualized by chemiluminescence, using
peroxidase-conjugated goat anti-rat antibody (Santa Cruz Biotechnology,
Inc., Santa Cruz, CA) and ECL Western blotting detection reagents
(PerkinElmer Life Sciences).
Characterization of HA-FGFR1--
HA-FGFR1 was extracted from
BaF3 B6 cells with lysis buffer. Total cell lysate was presorbed with
protein A-Sepharose beads (Amersham Pharmacia Biotech). The supernatant
was incubated with the anti-HA mAb 12CA5 (10 µg/ml; Roche Molecular
Biochemicals) at 4 °C for 2 h. The immune complex was collected
by further incubation with protein A beads at 4 °C overnight. After
extensive washing with lysis buffer and PBS, the beads were incubated
with 0.5 milliunits of endoglycosidase H, 1 unit of
N-glycosidase F (Roche Molecular Biochemicals), or only
assay buffer at 37 °C overnight. Then the protein A beads were
boiled in 2× SDS sample buffer in the presence of 10 mM
dithiothreitol. Samples were fractionated by SDS-PAGE and transferred
to PVDF membranes. mAb 3F10 was used to detect the HA-FGFR1 in Western blotting.
Cell Surface Biotinylation--
HA-FGFR1-transfected or
sham-transfected BaF3 cells were washed with cold PBS and incubated
with 0.5 mg/ml sulfo-N-hydroxysuccinimide-biotin (Pierce) at 4 °C for 20 min. Cells were washed and incubated with 0.5 mg/ml sulfo-N-hydroxysuccinimide-biotin for
another 20 min at 4 °C. After three washes with cold PBS, cells were
lysed with lysis buffer. Streptavidin beads were added to the cell
lysate to capture all biotinylated proteins. After 2 h of
incubation at 4 °C, the streptavidin beads were washed and boiled in
2× SDS sample buffer in the presence of dithiothreitol. Both the
biotinylated and nonbiotinylated fractions were subjected to SDS-PAGE
and then transferred to PVDF membranes. mAb 3F10 was used to detect the HA-FGFR1 in Western blotting.
FGF2-induced HA-FGFR1 Autophosphorylation--
BaF3 B6 cells
were serum-starved in RPMI medium supplemented with 1 mg/ml BSA and 2%
WEHI cell-conditioned medium. After 2 days of starvation, 2 ng/ml FGF2
(Roche Molecular Biochemicals) was added to cells in HEPES-buffered
medium (RPMI medium, 25 mM HEPES, pH 7.5, 1 mg/ml BSA, 0.1 mM orthovanadate), in the presence or absence of 100 ng/ml
heparin (Calbiochem), at 4 °C for 1.5 h. Then the cells were
warmed up at 37 °C for 5-30 min and extracted with lysis buffer in
the presence of tyrosine phosphatase inhibitor (1 mM
orthovanadate). Cell lysates were clarified by centrifugation at
14,000 × g for 15 min. The supernatants were incubated
with PY-20 (anti-phosphotyrosine antibody)-conjugated protein A-agarose beads (Santa Cruz Biotechnology) overnight at 4 °C. The beads were
washed with lysis buffer and boiled in 2× SDS sample buffer in the
presence of dithiothreitol. Samples were fractionated by SDS-PAGE and
then transferred to PVDF membranes. HA-FGFR1 was detected with mAb 3F10.
For testing the effects of membrane-associated HSPGs on FGF2-induced
HA-FGFR1 phosphorylation, 3 × 106 B6 cells were
incubated with 1 × 106 HSPG-presenting cells (fetal
human lung fibroblasts; syndecan-1-transfected, glypican-1-transfected,
or sham-transfected K562 cells; or transfected Namalwa cells) in the
presence of FGF2 at 4 °C for 1.5 h. To remove the cell surface
HS, cells were treated with 0.01 units/ml heparinase (Seikagaku Corp.,
Tokyo, Japan) at 37 °C for 40 min. Treated cells were incubated with
B6 cells and FGF2 and analyzed as described above.
For analyzing the time course of the effect of a heparitinase
digestion, cells were incubated with 0.006 units/ml heparitinase (Seikagaku) for 2, 5, 10, or 40 min at 37 °C. Cells were washed twice with assay buffer (HEPES-buffered medium) and then mixed with B6
cells in the presence of FGF2. The phosphorylation of HA-FGFR1 was
assayed as described above.
To test the soluble forms of these HSPGs, HSPG-presenting cells were
treated with 100 µg/ml trypsin (Sigma) at 4 °C for 10 min. Trypsin
inhibitor (Sigma) was then added to quench the effect of trypsin.
Further treatment with 0.006 units/ml heparitinase was performed at
37 °C for 40 min. Soluble HS chains or chain clusters prepared from
immunopurified syndecan-1 or glypican-1 (see below) were incubated with
B6 cells in the presence of FGF2, and tested as described above.
Extraction and Purification of Cell Surface HSPG--
Cell
surface proteoglycans were extracted from fetal human lung fibroblasts,
syndecan-1- or glypican-1-transfected K562 cells, and transfected
Namalwa cells as described previously (4, 22). Briefly, cells were
labeled with [35S]sulfate and lysed with a Triton X-100
buffer in the presence of protease inhibitors. The cell extract was
then centrifuged and concentrated on a DEAE-Trisacryl M column (Life
Technologies). Proteoglycans were immunopurified with specific mAb,
immobilized on CNBr-activated Sepharose 4B (Amersham Pharmacia
Biotech). After further purification by ion exchange chromatography on
a RESOURCE Q column (Amersham Pharmacia Biotech) in Triton-urea-Tris
buffer, these HSPGs were treated with chondroitinase ABC (Seikagaku)
(20 milliunits/ml, 3 h at 37 °C). Soluble HS fractions were
obtained from these HSPGs by trypsin digestion (60 µg/ml, 30 min at
37 °C) or alkaline treatment (0.5 M KOH, overnight at
4 °C). All treated proteoglycan fractions were repurified on DEAE
beads. Heparitinase digestion (0.015 units/ml) was at 37 °C for
3 h.
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RESULTS |
Heparin- and FGF2-dependent FGFR1 Phosphorylation in
BaF3 Cells--
To test the effects of heparin or HSPGs on receptor
activation, a low to zero background of endogenous HSPG expression is required. BaF3 cells express no detectable levels of endogenous HSPG.
These cells also cannot be induced to express any heparan sulfate upon transfection with relevant core protein cDNAs, which excludes their use for the design of receptor and HSPG co-transfection experiments. However, several reports suggest that cell surface HSPGs
can mediate FGF2 binding to an FGFR that is expressed on neighboring
cells and induce the signaling of this receptor (7, 23). We hence
developed tools for measuring such "transactivation" of the FGFR,
using receptor-autophosphorylation as assay. BaF3 cells transfected
with an HA epitope-tagged human FGFR1 IgII/IIIc isoform were
constructed to act as FGFR1-presenting cells. Namalwa and K562 cells,
transfected with syndecan-1 or glypican-1 cDNAs, and fetal human
lung fibroblasts, which express high levels of endogenous syndecans and
glypican-1, were chosen to act as HSPG-presenting cells.
The expression of HA-FGFR1 in the BaF3 cells was confirmed by Western
blotting. HA-FGFR1 appeared as two bands migrating around 100 and 120 kDa. The predicted molecular mass of this HA-FGFR1 is 81.5 kDa, and
glycosidase susceptibility tests indicated that the 120-kDa
(endoglycosidase H-resistant) and 100-kDa (endoglycosidase H-susceptible) forms represented two different isoforms of glycosylated receptor (both N-glycosidase F-susceptible) (Fig.
1A). Cell surface biotinylation revealed that the majority of the cell surface-exposed HA-FGFR consisted of the 120-kDa isoform (Fig. 1B).
HA-FGFR1-transfected BaF3 cells were cloned, and one of the highest
expressors, clone B6, was chosen for further experiments. Clone B6 and
sham-transfected BaF3 cells were then incubated with or without FGF2 in
the presence or absence of heparin, and the phosphorylation of HA-FGFR1
was analyzed as described under "Experimental Procedures"
(anti-Tyr(P) pull-down; blotting with anti-HA). Clearly, the
phosphorylation (pull-down) of HA-FGFR1 in BaF3 cells was strictly
dependent upon the addition of both FGF-2 and heparin (Fig.
2A). A time course experiment
showed that FGFR1 autophosphorylation reached peak levels around 15 min
of exposure to ligand at 37 °C (Fig. 2B).
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Fig. 1.
Characterization of HA-FGFR1 in BaF3
cells. A, glycosylation of FGFR1. HA-FGFR1 was
immunoprecipitated from BaF3 (clone B6) cell extracts with the murine
anti-HA mAb 12CA5 and incubated with endoglycosidase H
(EndoH), N-glycosidase F (PNGaseF), or
assay buffer only (control). The digested samples were fractionated by
SDS-PAGE. Rat anti-HA mAb 3F10 was used to detect HA-FGFR1 by Western
blotting. HA-FGFR1 appears as two bands migrating around 100 and 120 kDa, which represent two different glycoforms of the receptor.
B, cell surface expression of FGFR1. Receptor-transfected
(B6) and sham-transfected (pDisplay) BaF3 cells were cell
surface-biotinylated. Biotinylated proteins present in the lysates of
these cells were captured on streptavidin beads. Both the bound
materials and the materials remaining in the supernatants of the
streptavidin beads were subjected to SDS-PAGE. HA-FGFR1 was detected
with mAb 3F10 in Western blotting. The 120-kDa, endoglycosidase
H-resistant isoform represents the major from of cell surface-exposed
receptor. B, bound, biotinylated protein; S,
supernatant, nonbiotinylated protein.
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Fig. 2.
FGFR1-phosphorylation in BaF3 cells.
A, FGF2 and heparin dependence. Receptor-transfected (B6)
and sham-transfected (pDisplay) BaF3 cells were serum-starved. Aliquots
of 3 × 106 cells were incubated with or without 2 ng/ml FGF2, in the absence or presence of 100 ng/ml heparin at 4 °C.
Cells were warmed up at 37 °C and then lysed. The
anti-phosphotyrosine mAb PY-20 was used to precipitate all of the
tyrosine-phosphorylated proteins present in the lysates. The
immunoprecipitates and supernatants were then subjected to SDS-PAGE and
transferred to PVDF membranes. HA-FGFR1 was detected with mAb 3F10. The
autophosphorylation of HA-FGFR1 is clearly dependent on the addition of
both FGF2 and heparin. C, control; F, FGF2;
H, heparin; FH, FGF2 plus heparin; IP,
PY20 immunoprecipitate, represents phosphorylated FGFR1; SN,
supernatant, represents unphosphorylated FGFR1. B, kinetics
of FGF2-induced FGFR1 phosphorylation. After incubation with FGF2 and
heparin at 4 °C, the cells were warmed up at 37 °C for 5-30 min,
as indicated, and analyzed as above. FGFR1 autophosphorylation reaches
the highest level around 15 min.
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Effect of Membrane-associated HSPG on FGF2-induced HA-FGFR1
Phosphorylation--
Cultured human lung fibroblasts accumulate large
amounts of syndecans, glypican-1, and perlecan on their cell surfaces
(22). Namalwa and K562 cells, in contrast, express very low to low
levels of endogenous cell surface HSPG, mainly glypican-1 (in Namalwa cells) and syndecans (in K562 cells) (4). Phorbol diester-stimulated K562 cells secrete significant amounts of perlecan into their culture
media (24), but under basic conditions only very little of this
proteoglycan can be detected on the surfaces of these cells (result not
shown). To construct cells that express only or primarily a single
major form of proteoglycan, both the Namalwa and K562 cell lines were
transfected with syndecan-1 or glypican-1 cDNA. The expression
levels of cell surface syndecan-1 and glypican-1 proteoglycan were
monitored by quantitative immunofluorescence flow cytometry, using the
HS-specific mAb 10E4 and protein-specific antibodies. In all
HSPG-transfected cells, the expression of cell surface HS was markedly
increased, by at least 1 order of magnitude. The protein-specific
antibodies BB4 and S1 confirmed the expression of cell surface
syndecan-1 and glypican-1, respectively (Fig. 3).
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Fig. 3.
Cell surface expression of HSPG in Namalwa
and K562 cells. Cell surface expression was measured by
quantitative immunofluorescence cytometry. The curves illustrate the
background fluorescent signal, the heparan sulfate expressions, and the
core protein expressions in empty vector-transfected Namalwa cells
(A), glypican-1-transfected Namalwa cells (B),
syndecan-1-transfected Namalwa cells (C), empty
vector-transfected K562 cells (D), glypican-1-transfected
K562 cells (E), and syndecan-1-transfected K562 cells
(F). The results indicate very low levels of endogenous HSPG
on the cell surfaces of sham-transfected cells and marked increases in
cell surface HSPG expression in transfected cells. 2nd
Ab, background signal obtained with goat anti-mouse
secondary antibody only; 10E4, HS-specific mAb;
BB4, syndecan-1-specific mAb; S1,
glypican-1-specific mAb.
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HA-FGFR1-transfected BaF3 cells (clone B6) were then incubated with
HSPG-presenting cells in the presence of FGF2 and assessed for
autophosphorylation of HA-FGFR1. Fig. 4
shows that, independently of their origin, all membrane-associated
HSPGs (transfectant syndecan-1 and glypican-1 in the case of Namalwa or
K562 cells and total cell surface HSPGs in the case of fetal human lung
fibroblasts) markedly enhanced the FGF2-dependent
phosphorylation of HA-FGFR1. In contrast, K562 or Namalwa cells that
had been treated with heparinase or transfected with empty vectors had
no significant effects on FGFR1 autophosphorylation. This confirmed
that the effects of these HSPG-presenting cells on FGFR1
phosphorylation were HSPG-dependent. Interestingly,
treating the HSPG- presenting cells with heparitinase instead of
heparinase, for increasing lengths of time, first increased and then
decreased the capacity of these cells to support FGF2-induced receptor
autophosphorylation (Fig. 5).
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Fig. 4.
Activation of FGF2-FGFR1 signaling by cell
surface HSPG. Effects of the HSPGs from Namalwa cells
(A), K562 cells (B), and fetal human lung
fibroblasts (C) on FGF2-induced receptor phosphorylation in
BaF3 cells. Aliquots of 3 × 106 HA-FGFR1-transfected
BaF3 cells (clone B6) were incubated with 1 × 106
HSPG-presenting cells (nontreated or treated with heparinase) or
trypsin-solublized HSPGs from these cells (without or with an
additional heparitinase digestion of these soluble fractions) in the
presence of 2 ng/ml FGF2. Receptor phosphorylation was assayed as
described under "Experimental Procedures." Membrane-associated
HSPGs of all three cell lines enhance FGF2-induced HA-FGFR1
phosphorylation. The effect can be abolished by a heparinase
treatment of the cells. Trypsin-released soluble forms of these HSPGs
fail to promote receptor autophosphorylation. Heparitinase converts
these soluble forms into activators of receptor signaling.
Na-syn1, syndecan-1-transfected Namalwa cells;
Na-GPC1, glypican-1-transfected Namalwa cells;
Na-pcDNA, sham-transfected Namalwa cells;
K562-syn1, syndecan-1-transfected K562 cells;
K562-GPC1, glypican-1-transfected K562 cells;
K562-pRep4, sham-transfected K562 cells; HLF,
fetal human lung fibroblasts; Cell, untreated presenting
cells; Hepn, heparinase-treated cells; Tsn,
trypsin-released soluble HSPG; Hept, heparitinase-digested
trypsin-released soluble HSPG. Activation by FGF only (F)
and by the combination of FGF plus heparin (FH) is given as
a reference.
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Fig. 5.
Effect of heparitinase on HSPG-presenting
cells. Syndecan-1-transfected Namalwa cells were treated with
heparitinase at 37 °C for different lengths of time, as indicated.
Aliquots of 1 × 106 digested cells were washed with
assay buffer and mixed with aliquots of 3 × 106 B6
cells in the presence of FGF2. Phosphorylated HA-FGFR1 was detected as
described under "Experimental Procedures." Heparitinase transiently
enhances activity. Symbols are as in Fig. 4.
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Effect of Soluble HSPG on FGF2-induced HA-FGFR1
Phosphorylation--
Previous reports had demonstrated that purified
syndecans and glypican-1 originating from cultured fetal human lung
fibroblasts could not assist FGF2 in high affinity receptor binding
(11). Nevertheless, in the present assay, cell-associated forms of
these HSPGs were able to promote FGF2-induced receptor phosphorylation. To test whether this membrane association played a significant role,
the HSPG-presenting cells were treated with trypsin. Trypsin would be
predicted to cleave the core proteins of these proteoglycans and to
release the ectodomain and ectodomain fragments that bear the HS chains
into medium. Analysis of the residual HSPG expression by quantitative
immunofluorescence flow cytometry confirmed that after trypsinization,
barely any HSPG remained on the cell surface (data not shown). The
supernatants of these trypsin digestions (containing the released HSPG)
were supplemented with trypsin inhibitor and then added to the
HA-FGFR1-transfected B6 cells in the presence of FGF2. The soluble
forms of these HSPGs all failed to enhance FGF2-induced HA-FGFR1
phosphorylation (Fig. 4). To ensure that no HS had been lost during the
collection of the supernatants, trypsinized HSPG-presenting cells and
supernatants were also left together, supplemented with trypsin
inhibitor, and added to the B6 cells. None of these recombinations
stimulated FGF2-induced receptor phosphorylation (data not shown).
Furthermore, when non-trypsin-treated HSPG-presenting cells were mixed
with soluble HS, obtained by trypsinizing the equivalent of 10 times more cells, no significant phosphorylation of HA-FGFR1 could be detected in the presence of FGF2 (Fig.
6). Thus, soluble forms of HSPG inhibited
the stimulatory effects of cell surface-associated HSPGs.
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Fig. 6.
Inhibitory effects of soluble HSPGs.
Aliquots of 1 × 107 syndecan-1-transfected Namalwa or
K562 cells were treated with trypsin at 4 °C. The supernatant
(containing soluble HSPG) was collected and supplemented with trypsin
inhibitor. The soluble HSPG was mixed with either 1 × 106 non-trypsin-treated syndecan-1-transfected cells or
with the heparitinase digests from 1 × 106
syndecan-1-transfected cells (obtained as mentioned under
"Experimental Procedures"), and the mixtures were added to 3 × 106 B6 cells in the presence of FGF2.
Tyrosine-phosphorylated HA-FGFR1 was detected as described under
"Experimental Procedures." Soluble forms of HSPG clearly inhibit
the stimulatory effects of membrane-associated HSPGs and
heparitinase-generated soluble fragments on FGF2-induced
FGFR1-phosphorylation. Similar results were obtained for fetal human
lung fibroblasts, and for glypican-1-transfected Namalwa and K562 cells
(data not shown). Symbols are as in Fig. 4.
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When cells are treated with trypsin, the core proteins of the
proteoglycans (PGs) are at least partially destroyed. To test whether
the protein part of the PGs might play a role in assisting FGF2-induced
FGFR1 autophosphorylation, glypican-1-transfected Namalwa cells were
treated with phosphatidylinositol-specific phospholipase C. This
treatment cleaves glycosylphosphatidylinositol tails, leaving the
ectodomain of this PG intact. Nevertheless, like trypsin-released HSPG
fragments, phosphatidylinositol-specific phospholipase C-released
ectodomains of glypican-1 failed to promote FGF2-induced HA-FGFR1
phosphorylation (data not shown).
Since heparitinase can convert inactive HS fractions into activators of
FGF2-induced mitogenesis
(25),2 and because a
brief heparitinase treatment of the presenting cells stimulated
the activity of these cells (see above; Fig. 5), we further treated the
trypsin-released HSPGs with heparitinase. Consistently,
heparitinase-treated HSPGs stimulated FGF2-induced HA-FGFR1
phosphorylation in all the cases (Fig. 4). In these cases, however,
extending the heparitinase-treatment never suppressed the activity of
the digest (results not shown). Importantly, adding increasing
concentrations of soluble, undigested HS fractions to these
heparitinase digests inhibited the stimulatory effect of these digests
on HA-FGFR1 phosphorylation (Fig. 6).
In summary, all membrane-associated forms of HSPG tested here activated
FGF2-induced FGFR1 phosphorylation, but corresponding soluble forms,
obtained by trypsinization of the cells, were ineffective, even
inhibitory. Heparitinase, however, converted these soluble fragments
into activators.
Structural Characteristics of HSPGs Purified from Fetal Human Lung
Fibroblasts, K562 Cells, and Namalwa Cells--
Thus, HSPGs from three
different sources were acting similarly in terms of FGF2-FGFR1
signaling. To assess whether this also reflected similarity in HS
structure, syndecan-1 and glypican-1 were isolated from detergent
extracts of 35SO4-labeled fetal human lung
fibroblasts, K562 cells, and Namalwa cells.
After prepurification on DEAE and affinity purification on
corresponding protein-specific antibody columns, the purified PGs were
fractionated by ion exchange chromatography over RESOURCE Q. HSPGs
derived from human lung fibroblasts and Namalwa cells had fairly
homogeneous compositions. Syndecan-1 and glypican-1 from human lung
fibroblasts were eluted from RESOURCE Q as single sharp peaks
(syndecan-1 at 0.58-0.70 M NaCl; glypican-1 at 0.56-0.67 M NaCl). Syndecan-1 and glypican-1 from Namalwa cells were
also eluted as single but slightly broader peaks (at 0.60-0.82
M and 0.48-0.74 M NaCl, respectively). The
immunopurified HSPGs from K562 cells, in contrast, were more
heterogeneous. Both syndecan-1 and glypican-1 eluted as three
incompletely separated peaks, with increasing charge densities from
peak 1 to peak 3 (Fig. 7) (4). Accordingly, these materials were collected separately, as three distinct subfractions.
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Fig. 7.
Structural heterogeneity of the HSPGs.
35S04-labeled HSPGs were extracted from cells
with detergent. After affinity purification on corresponding core
protein-specific antibody columns, the PGs were subjected to ion
exchange chromatography over RESOURCE Q. Shown are elution profiles
obtained for glypican-1 from fetal human lung fibroblasts
(A) and K562 cells (B). Bars indicate
the fractions collected as pools 1, 2, and 3.
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The purified and fractionated HSPGs were further digested with
chondroitinase ABC and then treated with trypsin, to retain the
clustering of the HS chains as in the intact proteoglycan, or with
alkali to generate single protein-free HS chains. The fractionation of
the single chains from different HSPGs by SDS-PAGE and autoradiography
revealed that human lung fibroblasts produced the longest HS chains
among the three cell lines (mass of 45-100 kDa for syndecan-1-HS and
40-80 kDa for glypican-1-HS). Namalwa (syndecan-1-HS, 18-30 kDa;
glypican-1-HS, 18-40 kDa) and K562 cells (14-35 kDa) produced shorter
chains. Further analysis indicated that the heterogeneity of the K562
materials was due to the differences in both HS chain clustering (one-,
two-, and three-chain forms of HSPG) and charge density (data not
shown). Altogether, we could deduce from these data that, on average,
K562 HS chains derived from peak 3 were more highly sulfated than the
HS chains from peak 1 or HS chains from Namalwa or human lung
fibroblast HSPGs.
Effects of Purified HSPGs on FGF2-induced HA-FGFR1
Phosphorylation--
When trypsin is used to release HS from cells in
culture, a mixture of HS chains, HS chain clusters, and degraded cell
surface proteins is released into the conditioned medium. To avoid
possible interference by these other degraded proteins, single HS
chains and chain clusters derived from affinity-purified HSPGs,
isolated from fetal human lung fibroblasts, K562 cells, or Namalwa
cells, were also tested in the FGF2-induced HA-FGFR1 phosphorylation assay.
HS fractions derived from fetal human lung fibroblasts or Namalwa cells
similarly failed to promote FGF2-induced FGFR1 phosphorylation. Again,
after heparitinase treatment all these fractions promoted FGFR1
signaling (Fig. 8A). The more
highly sulfated HS derived from the peak 2 and 3 materials, only
present in K562 cells, represented an exception, and significantly
enhanced FGF2-induced FGFR1 phosphorylation (Fig. 8B). This
was somewhat unexpected, since trypsin-released K562 HSPGs were not
able to promote FGF2-induced receptor phosphorylation (see above).
Therefore, the three subfractions from K562 cells were also tested
after recombination in one fraction. From this it appeared that the
lesser sulfated subfractions of HS inhibited the stimulatory effect of
the more highly sulfated subfractions (Fig. 8B).
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Fig. 8.
FGFR1-activation by purified HS chains and
chain clusters of differing compositions. Syndecan-1 and
glypican-1 were isolated from Namalwa cells and fetal human lung
fibroblasts (A) and from K562 cells (B), by
affinity purification on corresponding protein-specific antibody
columns and ion exchange chromatography. Then either alkali
(A) or trypsin (T) was used to generate single HS
chains or chain clusters respectively, from these HSPGs or subfractions
of these HSPGs. Aliquots of 3 × 106
HA-FGFR1-transfected BaF3 cells were incubated with 100 ng/ml of these
soluble HS fractions, in the presence of 2 ng/ml FGF2. Phosphorylated
HA-FGFR1 was detected as described under "Experimental Procedures."
Except for HS subfractions of high charge density, derived from K562
cells, purified soluble native single HS chains and chain clusters are
unable to enhance FGF2-induced FGFR1 phosphorylation, while the
heparitinase digests of all of these materials are activators of
receptor signaling. Hept, heparitinase. A1,
A2, and A3, single HS chains, from peaks 1, 2, and 3; A1 + A2 + A3, pooled peak
fractions; T1 and T2, HS chain clusters, from
peaks 1 and 2. See Fig. 8 for peak definitions. Other
symbols are as in Fig. 4.
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Chain Valence Mutants of Glypican-1 in FGF2-induced FGFR1
Phosphorylation--
All syndecans and glypicans carry multiple HS
chains, which are clustered in one small domain of the protein. Since
both single chains and chain clusters derived from highly sulfated
proteoglycan fractions (in K562 cells) promoted FGF2-induced FGFR1
phosphorylation, we tended to conclude that the multivalence of the
cell surface HSPGs was not essential for the formation of active
FGF2·HS·FGFR1 complexes. To confirm this, chain valence
mutants of glypican-1 were constructed and transfected into Namalwa
cells. The cell surface expression of glypican-1 in these cells was
analyzed by quantitative immunofluorescence flow cytometry (data not
shown), and the effects of cells expressing one-chain (SAA), two-chain (SSA), or wild-type (SSS) forms of glypican-1 on FGF2-induced FGFR1
autophosphorylation were compared. Should a single chain glypican be
unable to assist FGFR1 signaling, overexpression of this form would be
expected to act as a dominant negative mutant, inhibiting any FGFR1
phosphorylation supported by endogenous multichain forms. Clearly,
cells expressing the single chain mutant strongly promoted FGF2-induced
HA-FGFR1 phosphorylation in B6 cells (Fig. 9), indicating that multivalence is not a
requirement for the activity of membrane-associated forms of HSPG.
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Fig. 9.
Effect of chain valence mutants of glypican-1
on FGF2-induced FGFR1 phosphorylation. Namalwa cells were
transfected with chain valence mutants of glypican-1. Aliquots of
1 × 106 cells were incubated with 3 × 106 B6 cells in the presence of FGF2. The phosphorylation
of HA-FGFR1 was analyzed as described under "Experimental
Procedures." Wild type, the two-chain mutant, and the single chain
mutant of this glypican all promote FGF2-induced HA-FGFR1
phosphorylation. SSS, wild type; SSA, two-chain
mutant; SAA, one-chain mutant. Other symbols are
as in Fig. 4.
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DISCUSSION |
Since Yayon et al. (3) first demonstrated the
importance of HSPG for high affinity FGF2-FGFR1 binding, a large amount
of biochemical and biological data has been produced, indicating that
HSPG is essential for FGF/FGFR signaling (4, 5, 26, 27). Although
several models have been proposed from binding studies, functional
analyses, and crystal structures (see below), the mechanism through
which HSPGs/heparin assist FGF/FGFR signaling remains less well
understood. Moreover, the contribution of individual HSPGs to FGFR
signaling is also a matter of debate. Two possible reasons for the
inconsistency of the published data were addressed here: the cell
specificity of the HSPG structure and the importance of membrane association.
Our data indicate that cell surface syndecans and glypican-1 from three
different cell lines (fetal human lung fibroblasts, K562, and Namalwa
cells) do differ in fine structure. These structural differences do not
appear to play a major role in the activation of FGF2/FGFR1 signaling
by membrane-associated HSPGs but correlate with differential activities
observed among soluble HSPGs, derived from these membrane forms. Among
the soluble HS forms, activity was restricted to highly sulfated
subfractions (from K562 cells), the fractions with the highest
resemblance to heparin, whose effects on FGF/FGFR signaling are well
established. Irrespective of this, the markedly differential effects of
soluble and membrane-associated syndecans and glypicans, derived from
identical sources and thus with similar glycosylations, suggest that
the membrane association of these HSPGs plays an important role for
their activity in FGF2/FGFR1 signaling.
It is generally accepted that growth factors activate their cognate
receptor tyrosine kinases by inducing receptor dimerization or
oligomerization. Various models for FGFR dimerization have been
proposed, including a cooperative bridging model (28), a beads on a
string model (29, 30), and a preexisting FGFR·HS duplex model
(31), to name only some. Recently published crystal structures of a
complex between FGF2 and FGFR1 provide a rather convincing receptor
dimerization model (32). The crystal contains two 1:1 FGF2·FGFR1
complexes, which are assembled in a "back-to-back" fashion, without
direct ligand-ligand contact. The two D2 domains of the FGFRs form a
positively charged canyon, and a positive potential extends onto the
topside of both ligands on either side of the canyon. A heparin
oligosaccharide could fit in this canyon and provide a cross-link,
connecting all four components in the assembly. However, unless one
accepts degeneracy of the binding sequences, it would seem logical to
propose that symmetric dimerizations of FGF2·FGFR1 complexes would be
facilitated by HS structures with antiparallel orientations, supported
by two different HS chains, belonging to one PG molecule or to two
different PG molecules with restrained diffusions. Indeed, accepting a
single mode of FGF2-HS interaction, FGF2 molecules docked to the same
HS chain would always adopt "cis" or side-by-side orientations.
Mckeehan's model proposes that a 2 FGF2·2 FGFR1 complex is formed by
the assistance of different HS chains belonging to one PG molecule (31,
33). Nonetheless, our data on the chain valence mutants demonstrate
that a one-chain mutant of glypican-1 is able to augment the
FGF2-dependent phosphorylation of FGFR1 just as well as
forms that carry two or three chains. Highly sulfated single HS chains from K562 cells have similar activities as chain clusters on FGFR1 phosphorylation, further evidence that the cluster structure of HS
chains is not required for their activity in FGF2/FGFR1 signaling. Then
how would noncontiguous HS structures with antiparallel orientations result in the symmetric dimerization of two FGF2·FGR1
complexes? Here, the "two-end" model proposed by Schlessinger
et al. provides an interesting solution (34). In
this crystal structure, obtained by soaking Plotnikov's preformed
FGF2-FGFR1 crystals with HS-oligosaccharide, a dimer consisting of two
1:1:1 FGF·FGFR·heparin ternary complexes is stabilized via direct
FGFR-FGFR contact, by a secondary interaction between FGF2 in one
ternary complex and FGFR1 in the other complex, and by indirect
heparin-mediated FGFR-FGFR contacts. This structure is similar to
Plotnikov's model (32), except that two HS structures are
occupying the canyon while adopting antiparallel end-to-end orientations. The heparin structures bind with their nonreducing ends
in the center of the canyon and run out onto the high affinity heparin-binding sites of the ligands. In addition to promoting FGF-FGFR
interaction within a 1:1 FGF·FGFR complex, the nonreducing ends of
the heparin structures also interact with the adjoining receptor across
the 2-fold dimer. In this way, the model provides a molecular basis for
the dual role of HS in augmenting 1:1 FGF-FGFR affinity and promoting
dimerization of two FGF·FGFR complexes. In the case of intact PGs,
this structure would imply the contribution of two heparan sulfate
chains, each belonging to a different PG, the protein cores of these
PGs (linked to the reducing ends) adopting a peripheral position in the
complex. At the moment, it is not clear whether the "two-end" model
can be extended to other FGF-FGFR systems. Different FGFs show
different HS requirements for the activation of their cognate
receptors, and pentameric structures have been reported for
FGF1·FGFR2·heparin complexes (35).
Unlike heparin, heparan sulfate chains have typical block structures
and are composed of highly sulfated, heparin-like "S-domains," alternating with low or nonsulfated domains (36). In the context of the
"two-end" model, this would imply that formation of the receptor
complex involves two heparan sulfate chains, with relevant S domains as
end structures, or very flexible HS chains, with irrelevant end
structures that protrude from the canyon. The stimulation resulting
from a brief treatment of the cells with heparitinase, interpreted as
an increase in the number of heparin-like end structures on cell
surfaces, and the activities of the heparitinase-generated fragments of
the various soluble HS chains fit well in this "two-end" model.
Heparitinase cuts the low sulfated regions of the HS chain but respects
the S domains. When heparitinase cleaves cell surface-bound heparan
sulfate on the nonreducing side of these S domains, it will convert
these domains in end structures. These will remain associated with the
cell surface until further cleavage of the chain in the nonsulfated and
less sulfated regions, which invariably include the protein-proximal
parts of the chains, releases the S domain. The lack of activity of
most of the soluble native HS chains and the marked activity of (what
we propose to be) the heparitinase-resistant (heparin-like) segments of
these HS chains would indicate that most of the relevant S-domains (in
terms of receptor dimerization) are embedded in the chains and have
only virtual or latent activities. HS chains with appropriate S domains at their nonreducing ends might be sufficiently preponderant among the
highly sulfated HS subfractions from K562 cells but would seem to occur
less often in other HS fractions tested in this assay. Competition of
the internally positioned S domains of these chains for FGF2 binding
would easily explain their inhibitory effects on the relatively rare
active end structures. In this sense, the effect of heparitinase is to
be considered as dually reinforcing, since it both generates active end
structures and destroys, by number, the internal inhibitory sequences.
Why then are membrane-associated HSPGs active where their soluble forms
fail to activate FGFR signaling? Both forms should have similar ratios
of active "end" and inhibitory "internal" structures.
Conceivably, direct core protein interactions with growth factor and/or
receptor could also account for or contribute to receptor activation.
The core proteins of some proteoglycans (e.g. NG2 and
phosphacan) bind to FGF2 (37, 38). Direct core protein interactions
with signaling components might be lost upon tryptic release of the
proteoglycan from the cell surface, explaining the failure of these
soluble forms. However, at least in the case of glypican-1 this would
appear to be insufficient as an explanation, since
phosphatidylinositol-specific phospholipase C-released glypican-1 (with
an intact ectodomain) also fails to promote FGF2-induced FGFR1
phosphorylation. Possible direct core protein interactions certainly do
not suffice for explaining the activity of the membrane-associated forms of syndecan-1 or glypican-1, since this HSPG-assisted FGF2/FGFR1 signaling is strictly HS-dependent. The binding of
syndecans and glypican to FGF2 is solely HS-mediated (4), but
interactions with receptor are not excluded. Such interactions would
appear to be endowed with little specificity, since syndecan-2 (8) and
the membrane-associated HS-substituted splice variant of CD44 (39) also
promote FGF2 signaling (mitogenesis) in BaF3 cells. We therefore tend
to conclude that membrane association and proteoglycan assemblies
rather than a specific direct core protein contribution are needed for
efficient HSPG-supported FGF signaling.
It is likely that the concentration of the reactants and the need for
threshold concentrations to be reached play a significant role. Cell
surface HSPGs binding of FGF will reduce the dimensionality of ligand
diffusion from three to two dimensions. Thus, the local concentration
of the bound ligands at the plasma membrane and the probability of
their interaction with the high affinity receptors might be greatly
enhanced. In the present experimental setup, cell binding and cell-cell
contacts between HS-presenting cells and receptor-bearing cells are
likely to play an important role, which is suggested by the inhibitory
effects of adding soluble HSPG to the presenting cells (see above) and
adding increasing numbers of presenting cells for fixed amounts of
receptor-bearing cells and added FGF (results not shown). Possibly,
additional aspects have to be considered. Membrane association of the
HSPG might further stabilize the FGF·FGFR·HSPG complex. This might not be essential, since soluble heparitinase-resistant fragments are
active in assisting FGF2/FGFR1 signaling, but might occur, since in
further studies we were unable to demonstrate "high affinity" binding of FGF2 to cell surfaces supported by such
fragments.2 In contrast, high affinity binding of FGF2
could be measured when FGF2 was presented by cell-bound HSPGs or by
heparin, which itself may tether to cell surfaces (40).
Further possible aspects of the membrane association of HSPG, which are
not exclusive with the above, include facilitated diffusion and
cooperativity. Due to their highly negative charges, HS chains would be
expected to repel each other. On most cells, the abundance and
molecular dimensions of cell surface HSPGs are such that, assuming a
random distribution, they should cover the entire cell surface and even
invade each other's domains. Typical concentrations of HSPGs on the
cell surface are in the range of 105 to 106
molecules/cell (41). Fetal human lung fibroblasts and the
HSPG-transfected K562 and Namalwa cells studied here express around
5 × 105 to 2 × 106 HSPG
molecules/cell. According to their molecular masses, the average
lengths of the HS chains would vary from 30 to 60 nm in Namalwa and
K562 cells, and from 70 to 140 nm in fetal human lung fibroblasts.
Around 5 × 105 HSPGs uniformly placed on fetal human
lung fibroblasts (diameter of 16 µm), K562 cells (14 µm), or
Namalwa cells (12 µm), assuming spherical cell shapes, would
encompass the entire cell surface. High densities of HS may facilitate
FGF diffusion from chain to chain and along chains (42). High density
and cell surface association may impose particular chain orientations
(all reducing ends oriented toward cells), so that highly
sulfated domains at the nonreducing ends of HS chains have better
chances to approximate each other than when free in solution. Moreover,
when FGFs are added, they will neutralize the negative charges on some
chains, reducing the repellent forces between chains. In a simplified
model, consisting of two PGs, each with three chains but only one
loaded with FGF, fixed in position but with rotational freedom, one
would easily conceive how the neutralized chains will be driven
together assisted by the repulsion of the nonneutralized chains. This
would be more plausible if the neutralized domains were end structures
and would result in the FGFs forming a "trans-dimer," supported by
two antiparallel oriented chains.
Glypicans can associate with rafts, resulting in their oligomerization
(43). Syndecan core proteins exhibit a propensity to form noncovalently
linked dimers and higher order oligomers (44, 45), and adding ligands
such as FGF2 to syndecan-1-expressing cells induces clustering of this
syndecan (46). Syndecans bind to a variety of intracellular molecules,
and the cytoplasmic domain of syndecan-4 seems to play an important
role in cellular responses to FGF2 stimulation (47). Thus, syndecans
and glypican may preexist as clusters on the cell surface or be
dimerized/oligomerized in response to extracellular ligand binding or
inside out signaling, implying that the local concentrations of
reactants may even be underestimated in the above reasonings.
In summary, we showed that several HSPG-presenting cells can stimulate
FGF2/FGFR1 signaling in neighboring HS-negative/receptor-positive cells
but that "proteolytic" shedding of these HSPGs severely impairs
their activity in this sense. This indicates HSPG shedding could have a
major role in negatively regulating growth factor activity. The
physiological relevance of this experimental paradigm remains to be
proven but is plausible, since in vivo the HS expressions are often restricted to particular cells and since inhibitory shed
ectodomains accumulate in inflammatory fluid (19). Bacterial heparitinase converted these inhibitory shed HS species into activators of FGF2-dependent FGFR1 autophosphorylation, interpreted as
the generation of "end structures" that can cooperate in receptor dimerization (34). Mammalian heparanase is an endoglucuronidase, cleaving heparan sulfate at specific intrachain sites (48). Whether
mammalian heparanases produce "end-structures" of the types that
were used in modeling and crystal structures and that we suggest to be
generated here remains to be proven. Nevertheless, mammalian
heparanases can also convert inhibitory soluble HSPG ectodomains,
recovered from inflammatory fluid, into potent mitogenic activators of
FGF2 (19). All cells tested here express mammalian heparanase.2 In the extension of this model, the form of
HSPG that activates FGF2 signaling might be a heparanase-generated product.
TOP
ABSTRACT
INTRODUCTION
To whom correspondence should be addressed: Center for Human
Genetics, Campus Gasthuisberg, O & N, Herestraat 49, B-3000 Leuven, Belgium. Tel.: 32-16-34-58-63; Fax: 32-16-34-71-66; E-mail:
guido.david@med.kuleuven.ac.be.
