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INTRODUCTION |
Fibroblast growth factor 2 (FGF-2)1 belongs to the
fibroblast growth factor family and is one of the most potent growth
factors regulating cellular functions including proliferation and
chemotaxis, tissue and organ development, and tissue repair
(1-3). FGF-2 has been suggested to play a key role in the
etiology and progression of several pathological conditions including
Parkinson's and Alzheimer's diseases (4-5), tumor growth and
progression (6-9), atherosclerosis, and restenosis after angioplasty
(10). A specific role has been indicated for FGF-2 in tumor
angiogenesis (11-12) and in neuronal differentiation (13-14).
In light of the key role FGF-2 plays in both physiologic and pathologic
conditions, structural-functional relationships studies on this factor
are of large interest. FGF-2 dimerization and its interaction with
heparan sulfates are necessary steps required for binding to high
affinity receptors and for the following signaling activation. Several
reports investigated functionally relevant regions of FGF-2 by
analyzing crystallographic structures or by mutagenesis studies. FGF-2
dimer has not been crystallized, and different studies based on
molecular simulations indicated different regions possibly lying at the
FGF-2 dimer interface (15-17), whereas crystallization (18-20) and
other studies (21-26) identified residues crucial for FGF-2 activity
and involved in the interaction with heparin and high affinity
receptors. Finally, specific residues were found to control the FGF-2
ability to induce production of urokinase-type plasminogen activator,
likely in the FGF-2 nuclear localization site (27-28).
Despite all ongoing investigations, a specific site regulating the
whole FGF-2 activity by controlling FGF-2 interaction with itself, has
not yet been identified. We recently showed that FGF-2 directly
interacts with PDGF-BB (29-30), leading to a marked inhibition of its
angiogenic properties in vitro and in vivo (31).
In the present study we report the novel identification of an FGF-2
functional domain controlling in vitro and in
vivo FGF-2 properties, likely by controlling dimer formation.
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EXPERIMENTAL PROCEDURES |
Structural Analysis of FGF-2, Active Site Identification, and
Peptide Synthesis
FGF-2 sequences were retrieved from the Web site of the National
Library of Medicine (www.ncbi.nlm.nih.gov) and aligned by using
ClustalW software (32). Accession numbers are reported in the legend
of Fig. 1. Three-dimensional analysis was carried out on two
high-resolution crystallographic structures from the Protein Data Bank
(PDB), i.e. 2FBH and 1FQ9 for FGF-2 and the
FGF-2/FGFR-1/heparin complex, respectively. Solvent accessibility analysis was performed using NACCESS software (33) to identify FGF-2
regions interacting with FGF-R1, according to a procedure previously
used (17, 29). Solvent accessibility of each amino acid was evaluated
for FGF-2 alone and for receptor-bound FGF-2; amino acids undergoing a
marked accessibility modification within the complex were assigned at
the ligand-receptor interface. FGF-2 secondary structure was evaluated
by analyzing 2FBH molecular co-ordinates with DSSP software (34).
Regions potentially falling at the FGF-2 dimerization sites were
predicted by molecular docking simulations performed with GRAMM
software (35, 36), through a procedure used for both high- (37, 38) and
low-resolution (29, 39) simulations. In this study low-resolution
docking parameters for globular proteins were used according to the
GRAMM software manual. Regions at the interface of the predicted FGF-2 dimer were identified by solvent accessibility analysis as described above. According to these analyses, the FGF-2 region including the
sequence DPHIKLQLQAE (here referred to as FREG-(48-58)) was identified. FREG-(48-58) was then custom synthesized by Research Genetics Inc. As a sequence-specificity control, two different scrambled versions of FREG-(48-58) (KHIAQLDEPLQ and KLQLDIEAHPQ), were
synthesized by Neosystem Laboratoire. They are here referred to as SCR.
Peptide purity, assessed by reverse-phase HPLC and mass spectrometry,
was higher than 95% in all cases. Different batches of FREG-(48-58)
and SCR gave similar results in all biological assays.
Cell Isolation and Cell Culture
BAEC were derived from adult bovine aorta and cultured as
previously reported (40). Cell purity was evaluated by DiI-Ac-LDL uptake and was consistently >97%. BAEC at passage 3-7 and at 80% confluence were used in all assays.
Proliferation and Migration Assays
Proliferation assays were carried out as previously
reported (31). After 24 h of starvation, BAEC were incubated with
Dulbecco's modified Eagle's medium-BSA (0.1%) alone or containing
human recombinant growth factors (10 ng/ml) as reported, in the absence
or in the presence of FREG-(48-58) or SCR, the antibody raised against
FREG-(48-58) (AbFREG-(48-58)), or preimmune control serum (AbP-I).
BSA fraction V (Sigma) was used throughout the study. Time course
experiments were carried out at 1-, 2-, and 3-day time points at 10 ng/ml FREG-(48-58) concentration. Cells were then harvested and
counted with a hemacytometer. The FREG-(48-58) dose used in most
experiments (i.e. 10 ng/ml) was identified in dose
dependence tests carried out in cell proliferation and receptor
phosphorylation experiments.
Migration assays were carried out in modified Boyden chambers (Costar
Scientific Corporation) as previously reported (31). Briefly, BAEC
seeded on gelatin-coated polycarbonate filters (12-µm pores) (Costar)
were exposed to FGF-2 (Invitrogen), vascular endothelial growth factor
(VEGF) (RnD Systems), epidermal growth factor (EGF) (Invitrogen), or
fibronectin (Invitrogen) human recombinant factors, dissolved at the
reported concentration in Dulbecco's modified Eagle's medium, 0.1%
BSA. FREG-(48-58) or SCR were supplemented to the growth factor
solution at the reported final concentration. Assays were carried out
at 37 °C in 5% CO2, for 6 h. Migrated cells were
counted at ×400 magnification in 15 fields for each filter, and the
average number ± S.D. of cells/field was reported. All
experiments were performed at least three times in duplicate.
Histidine Residue Chemical Modification
Chemical modification of histidine residues was achieved by
diethylpyrocarbonate (DEPC) treatment, which specifically modifies the
imidazolic ring of histidine residues (41). Briefly, an aliquot of a
freshly prepared DEPC solution (Sigma) in anhydrous ethanol was added
to FGF-2 in PBS, to reach 200 µM DEPC and 55 µM FGF-2 final concentration. After 15 min at 25 °C
the reaction was stopped with 100 mM Tris-HCl (pH 7.6).
DEPC-induced modification of FREG-(48-58) was carried out similarly.
In control experiments DEPC was first inactivated by prior incubation
with 100 mM Tris-HCl, and then added to the
sample. Histidine-modified FREG-(48-58) and FGF-2 were then tested in
SPR (see below) and migration assays.
Peptide and FGF-2 Biotinylation
Biotinylation was carried out according to the
manufacturer's instructions (Pierce). Briefly, FREG-(48-58) (2 mg/ml)
or FGF-2 (20 µg/ml) in PBS (500 µl) (pH 7.4) were incubated with 2 mM EZ-Linksulfo-NHS-biotin (Pierce) for 30 min at room
temperature to label amines. The excess reagent was quenched by
incubation with 100 mM Tris-HCl (pH 7.4) for 15 min at room
temperature. The mixture was then dialyzed against PBS; biotinylated
FREG-(48-58) was then immediately used in an overlay assay, and
biotinylated FGF-2 was immediately used in internalization assays.
Fluorescence Analysis
Fluorescence spectroscopy experiments were carried out as
previously reported with modifications (29, 42) on a Perkin Elmer LS55
fluorimeter at constant temperature (22 °C). Briefly, 10 aliquots
(0.5 µl each) of FREG-(48-58) or SCR (85 µM, 0.1 mg/ml in PBS) were subsequently added to 90 µl of FGF-2 in PBS (2 µM, 36 µg/ml). Under these conditions, the FGF-2 sample
reached only 5% dilution. Fluorescence emission spectra were collected
by excitation at 277 nm. Peak fluorescence was at 310 nm as previously
reported for similar experimental conditions (43). Data were
corrected for the buffer, FREG-(48-58), and SCR contributions and for
the dilution factor, and its value was plotted as a function of the amount of peptide supplemented. Data were analyzed according to the
single site tight binding model (44).
FGF-2 Interaction with FGF-2, with Heparin, or with FGF Receptor
1 by Surface Plasmon Resonance (SPR) Analysis
SPR experiments were carried out as previously reported (29, 45)
on the BIAcoreX instrument (Amersham Biosciences Biosensor AB).
Immobilization--
FGF-2 or recombinant human FGF-R1
(IIIC)/FC chimera (RnD Systems) were covalently coupled to the CM5
sensor chip by injecting FGF-2 or FGF-R1 (80 µl, 1.25 µg/ml and
55.5 µg/ml, respectively) dissolved in 30 mM acetate
buffer, pH 4.8. Immobilized FGF-2 and FGF-R1 achieved about 500 and
7500 Resonance Units (RU), respectively, corresponding approximately to
0.5 and 7.5 ng/mm2, respectively (46). Immobilized FGF-2
was recognized by injecting a goat anti-human FGF-2 antibody (RnD
Systems). All experiments were performed using HBS (10 mM
Hepes, 0.15 M NaCl, 3 mM EDTA, 0.005% (v/v)
surfactant P20, pH 7.4) as running buffer and as a dilution buffer for
FGF-2. In additional experiments, biotinylated low molecular weight
heparin (2.6 mg/ml in HBS containing 0.3 M NaCl, 30 µl)
was captured onto a BIAcore SA chip (carrying streptavidin onto the
surface) via biotin-streptavidin interaction at a 5 µl/min flow rate.
Under these conditions immobilized heparin reached about 200 RU.
Injection--
FGF-2 (20 µl, 1.25 µg/ml) was injected for a
2-min association phase with or without increasing concentrations of
FREG-(48-58) or SCR, followed by HBS flow for a 30-s dissociation
phase. Response, monitored at 25 °C, was expressed in RU. Sensor
chip regeneration was achieved by NaOH injections (50 mM,
10 µl each). A 10 µl/min flow rate was used throughout the experiments.
Overlay Experiments
Either FGF-2 or BSA (300 ng) were spotted onto a
nitrocellulose membrane as previously reported (29), then blocked with 5% dry milk (Bio-Rad) in TPBS (0.1% Tween 20 in PBS). In addition to
BSA, an additional control was introduced, corresponding to the mix of
molecular weight standards (Amersham Biosciences) listed in the legend
of Fig. 4. After the TPBS wash, the membrane was incubated with
biotinylated FREG-(48-58) (2 ml, 200 µg/ml) for 4 h at room
temperature and washed again. Interaction was detected by an
avidin/biotinylated horseradish peroxidase kit (Vectstain ABC, from
Vector) followed by chemiluminescent reaction and exposure to Kodak
film (Eastman Kodak).
FGF-2 Internalization
FGF-2 internalization was measured as described (47). BAEC
(5 × 105) were incubated with biotinylated FGF-2 (10 ng/ml) alone or in the presence of either FREG-(48-58) (20 ng/ml), SCR
(20 ng/ml), mAb 125 (monoclonal antibody neutralizing FGF-2 receptor,
Chemicon) (500 ng/ml), rabbit AbFREG48-58 (1:200 dilution), or rabbit
preimmune serum (AbP-I) (1:200) at 37 °C in a 5% CO2
atmosphere. After 1 h, cells were washed and cell surface-bound
material was extracted by washing with 2 M NaCl in 20 mM Hepes buffer (pH 7.4) and with 2 M NaCl in
20 mM acetic acid (pH 4.0) and discarded. Cells were then
lysed with lysis buffer containing 20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA,
1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM
-glycerol phosphate, 1 mM
Na3VO4, 1 µg/ml leupeptin, and 1 mM phenylmethylsulfonyl fluoride (Sigma), and the total
lysate was immobilized onto nitrocellulose and blocked with 5% dry
milk in TPBS. After washing with TPBS, biotinylated FGF-2 was then
revealed with the Vectstain ABC Kit as in overlay experiments. The film
was then subjected to densitometry analysis (GS 710, Bio-Rad).
FGF-R1 Phosphorylation
After 24 h of starvation, BAEC were incubated with FGF-2
(10 ng/ml) diluted in 0.1% BSA, in the absence or in the presence of
FREG-(48-58) or SCR (10 ng/ml) for 10 min. Cells were then washed in
ice-cold PBS and scraped with lysis buffer (as above). Cell extracts
were sonicated and centrifuged, and supernatants were collected.
Immunoprecipitation was then carried out on samples (60 µg of
protein) by incubating with anti-FGF receptor antibody (mAb 125, Chemicon) (2 µg) at 4 °C overnight on a rocking bath, followed by
incubation with 10 µg of protein G-Sepharose 4 Fast Flow (Amersham
Biosciences) for 2 h at 4 °C. After washing with lysis buffer,
complexes were dissolved in Laemmli buffer, separated by 8% SDS-PAGE,
blotted onto nitrocellulose membrane, blocked as reported above,
incubated with a monoclonal anti-phosphotyrosine antibody (clone PT66,
Sigma) (1:2000 dilution) for 1 h at room temperature, revealed by
chemiluminescence (Amersham Biosciences), and quantified by densitometry.
Production and Testing of the Rabbit Anti-FREG-(48-58) Antibody
(AbFREG48-58)
Antibody against FREG-(48-58) (AbFREG48-58) was developed
according to standard protocols. Briefly, FREG-(48-58) (1 mg)
dissolved in 2 ml of PBS and supplemented with Complete Freund's
Adjuvant, was administered by intramuscular injection in two
5-month-old New Zealand rabbits. After 22 and 44 days the antigen
injection was repeated with un-complete Freund's Adjuvant. Fifteen
days after the third injection, blood was collected, and the serum was
separated and stored at
20 °C in aliquots. Before some
experiments, rabbit serum was collected after four antigen
injections and showed activity at a higher dilution. The ability of
this antibody to recognize the whole FGF-2 was tested by spotting
increasing quantities of FGF-2 (up to 100 ng/spot) onto nitrocellulose
membranes and then blocking with 5% dry milk. After TPBS washing,
membranes were incubated with AbFREG48-58 (1:200 in PBS) for
1 h at room temperature. Membranes were then washed and incubated
with anti-rabbit peroxidase-conjugated IgG (Pierce) for 1 h at
room temperature; detection was achieved by chemiluminescence (Amersham
Biosciences). FREG-(48-58) up to 100 ng/ml was used to compete for the
observed interaction between FGF-2 and AbFREG48-58. AbFREG48-58 was
then used in internalization and proliferation assays as reported.
In Vivo Angiogenesis Assays
Procedures involving animals and their care were conducted
according to the institutional guidelines, in compliance with
international laws and policies (Guide for the Care and Use of
Laboratory Animals; United States National Research Council, 1996).
Angiogenesis in Chick Embryo Chorioallantoic Membrane (CAM
Assay)
CAM assays were performed as previously reported (31).
Fertilized White Leghorn chicken eggs (10/group) were incubated at 37 °C at constant humidity. On incubation day 3, 2 ml of albumin were removed to allow detachment of the developing CAM. The window was
sealed with a glass and the eggs returned to the incubator. On day 8, CAM was treated by applying the sponge as reported in the legend to
Fig. 8. On day 12, blood vessels entering the sponge within the focal
plane of the CAM and were counted at ×50 magnification in a
double-blind fashion under a Zeiss SR stereomicroscope (Zeiss) and
photographed in ovo with the MC63 Camera System (Zeiss).
Embryos and their membranes were fixed in ovo in Bouin's
fluid. Sponges, the underlying and immediately adjacent portions of CAM
were removed, embedded in paraffin, sectioned at 8 µm along a plane
parallel to the CAM surface, and stained with a 0.5% aqueous solution
of toluidine blue (Merck). The angiogenic response was assessed
histologically by a planimetric method of "point counting" (48).
The vascular density was indicated as the mean number of the occupied
intersection points ± S.D.
Angiogenesis in Matrigel Plugs
The angiogenesis assay in Matrigel plugs was carried out as
reported (31, 49). Briefly, Matrigel, a mixture of reconstituted basement membrane proteins (BD PharMingen, 600 µl) supplemented with
FGF-2 (150 ng/ml) alone or in the presence of FREG-(48-58) or SCR (10 µg/ml), was injected subcutaneously in CD1 mice (female, 20 g
body weight). Matrigel plugs were removed after 8 days and processed
for histology analysis. Histologic sections (7 µm) were stained with
Trichrome-Masson procedure (Bio-Optica). Vessels within the Matrigel
were recognized by morphology and by the presence of red blood cells.
Angiogenesis was evaluated blindly by two operators, by considering at
least five different sections per Matrigel plug; each section was 100 µm from the next. The total number of neo-vessels in the whole
Matrigel area was measured with an Axioplan microscope (Zeiss) and was
expressed as the number of vessels/mm2. When 2 vessels were
cut longitudinally, as indicated by a long axis greater than 3-fold of
the short axis, and were close to each other, they were counted as a
single vessel. Ten animals were treated with FGF-2 alone, 14 with FGF-2 + FREG-(48-58), 10 with FGF-2 + SCR.
Statistics
Data were expressed as mean ± S.D.
Student's two-tailed t test was performed, and
p
0.01 was considered statistically significant.
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RESULTS |
Analysis of FGF-2 Structure--
Primary sequence analysis and
three-dimensional structure evaluation were carried out on human FGF-2.
A multiple sequence alignment was performed on the 23 members of the
human fibroblast growth factor family (Fig.
1A). According to the
consensus sequence at the 60% identity level, several residues are
conserved within the family, while other regions show a high
variability. In particular, the FGF-2 region 43-60 was found to be
poorly conserved within the FGF family.


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Fig. 1.
Sequence alignment and structure analysis.
A, FGF family alignment. Sequence alignment of 23 FGFs (all
human, except FGF 15, of unknown source) was achieved by ClustalW
software available at emb1.bcc.univie.ac.at/embnet/progs/clustal/. FGFs
accession numbers are: FGF1, XP_054732; FGF2, XP_055784; FGF3,
NP_005238; FGF4, XP_053627; FGF5, P12034; FGF6, P10767; FGF7,
XP_017651; FGF8, BAA28605; FGF9, NP_002001; FGF10, NP_004456; FGF11,
AAL15439; FGF12, XP_003135; FGF13isoform 1A, NP_004105; FGF14,
NP_004106; FGF15, AR015075; FGF16, NP_003859; FGF17, NP_003858; FGF18,
NP_387498; FGF19, AAH17664; FGF20, NP_062825; FGF21, NP_061986; FGF22,
NP_065688; FGF23, NP_065689. The alignment is reported for the
FGF-2 portion whose crystallographic coordinates are available. The
consensus sequence at 60% identity level is reported (boxed
residues on yellow background). FREG-(48-58) is indicated
in red. Blue cylinders indicate regions predicted
to be involved in FGF-2 dimerization according to the predicted dimer
reported in panel B. Residues interacting with the receptor,
identified according to solvent accessibility analysis on the PDB
structure 1FQ9 (20), were: His-16, Phe-17, Lys-18, Lys-21, Tyr-24,
Lys-26, Gly-28, Gly-29, Phe-31, Arg-44, Lys-46, Gln-54, Leu-55, Gln-56,
Ala-57, Gln-58, Glu-59, Arg-60, Gly-61, Val-63, Ser-64, Tyr-73, Val-88,
Glu-96, Arg-97, Leu-98, Glu-99, Ser-100, Asn-101, Asn-102, Tyr-103,
Asn-104, Gly-131, Pro-132, Gly-133, Gln-134, Lys-135, Leu-138, Leu-140,
Pro-141. Predicted dimerization sites and observed
receptor-interacting residues partially overlap in the FREG-(48-58)
region. Gaps longer than two positions were reported as the number
indicating the gap length. B, docking simulation of two
FGF-2 monomers (PDB code: 2BFH) achieved with GRAMM software. The two
chains are colored in blue and green;
red segments correspond to residues 48-58 selected to
designate FREG-(48-58) and appear to be at the dimer
interface.
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A docking simulation was carried out on two FGF-2 monomers (PDB
code: 2BFH) leading to the prediction of a FGF-2 dimer (Fig.
1B); therefore regions at the dimer interface were predicted by solvent accessibility evaluation. Finally, on the high-resolution crystallographic model of the FGF-2/FGR1-heparin complex (PDB file:
1FQ9), regions interacting with the FGF-R1 were identified by solvent
accessibility analysis (reported in the legend of Fig. 1A).
According to all these analyses, the FGF-2 region 43-60 shows one of
the lowest intrafamily identities and, at the same time, many residues
of this segment were predicted to be involved in FGF-2 dimerization and
were found to be involved in receptor binding. All together these
observations suggested that this portion of FGF-2 may play a specific
role in FGF-2 dimerization and receptor binding. To investigate this
hypothesis, the following strategies were addressed: (i) one peptide
derived from this region was designed, synthesized, and tested in
vitro and in vivo, (ii) an antibody was developed
against this peptide and tested in FGF-2-dependent in
vitro assays, and (iii) histidine chemical modification was carried out because one of the three FGF-2 histidines falls in the
peptide region (i.e. His in position 50).
The secondary structure of residues 43-60 is characterized by a
disordered segment, a short helix, a
-strand, and an additional short disordered segment. We selected a peptide including the short
helix and the short
-strand, i.e. region 48-58
(DPHIKLQLQAE; FREG-(48-58)), to avoid excessive flexibility.
FREG-(48-58) was therefore used in the following in vitro
and in vivo analyses, and for rabbit immunization to raise a
specific antibody.
Proliferation and Migration Assays--
FGF-2-induced BAEC
proliferation was evaluated at 48 h in the absence and presence of
increasing FREG-(48-58) doses up to 1000 ng/ml (Fig.
2A). FREG-(48-58) showed a
dose-dependent inhibitory activity and, at 10 ng/ml,
lowered the FGF-2 mitogenic effect to the control level, while both
scrambled peptides used as control (SCR) showed no activity. Therefore
FREG-(48-58) and SCR at 10 ng/ml were used for the following in
vitro experiments.

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Fig. 2.
BAEC proliferation. A,
FREG-(48-58) effect on FGF-2-induced BAEC proliferation. Cells were
incubated for 48 h with FGF-2 (10 ng/ml) in Dulbecco's modified
Eagle's medium, BSA (0.1%) alone or in the presence of increasing
doses of either FREG-(48-58) or SCR. Results are expressed as average
percent of cell number versus cells treated with FGF-2
only ± S.D. Absolute value corresponding to 100% is 9.5 × 105 cells. Experiments were performed four times in
duplicate. Asterisks indicate p < 0.01 versus FGF-2 alone. B, time course experiments on
BAEC proliferation. BAEC were incubated with FGF-2 (10 ng/ml) alone or
in the presence of FREG-(48-58) or SCR (10 ng/ml) up to 3 days.
FREG-(48-58) was added at time 0 and time 48 h. Results are
expressed as average cell number ± S.D. Experiments were
performed three times in duplicate. Asterisks indicate
p < 0.01 versus time 0.
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In time-course experiments FREG-(48-58) significantly inhibited
FGF-2-induced proliferation, while SCR was ineffective (Fig. 2B). In the absence of FGF-2, FREG-(48-58) and SCR did not
affect cell number, indicating that these compounds are not toxic on BAEC at the tested doses (not shown). Additional experiments showed that FREG-(48-58) markedly inhibited FGF-2-induced BAEC migration, while SCR was not effective (Fig.
3A). In contrast, EGF-,
fibronectin-, and VEGF-induced migration were not impaired by
FREG-(48-58) nor by SCR (Fig. 3, B-D, respectively),
indicating that the FREG-(48-58) effect was FGF-2-specific.

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Fig. 3.
BAEC migration. Effect of FREG-(48-58)
and SCR (10 ng/ml) on FGF-2-induced (A), EGF-induced
(B), fibronectin-induced (C), and
VEGF-induced (D) BAEC migration. FREG-(48-58) showed a
significant inhibitory effect on FGF-2 only, while SCR was inactive in
all cases. Asterisk indicates p < 0.01 versus FGF-2 alone.
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FGF-2 Interaction with FREG-(48-58)--
FREG-(48-58) was
designed from the region at the predicted interface of the FGF-2 dimer.
The hypothesis of a direct FREG-(48-58)/FGF-2 interaction was then
tested in solid phase (overlay) assays and in liquid phase
(fluorescence) experiments. Overlay assays showed that biotinylated
FREG-(48-58) (200 µg/ml) specifically interacted with FGF-2
immobilized onto nitrocellulose (300 ng/spot), while it did not
interact either with immobilized BSA or with a pool of immobilized
purified proteins (molecular weight standards) (Fig.
4A).

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Fig. 4.
FREG-(48-58) interacting properties.
A, overlay assay of biotinylated FREG-(48-58) on
immobilized FGF-2. Biotinylated FREG-(48-58) interacted with FGF-2
immobilized onto nitrocellulose (300 ng/spot), whereas no interaction
was observed with immobilized BSA (300 ng/spot) or with immobilized
molecular weight standards (Amersham Biosciences, 300 ng/spot) (Mix)
consisting of a mixture of myosin, phosphorylase B, BSA, ovalbumin,
carbonic anhydrase, trypsin inhibitor, and lysozyme. A representative
experiment is shown. This experiment was performed three times with
similar results. B, fluorescence spectra of FGF-2 (2 µM) in the presence of increasing doses of FREG-(48-58).
Spectra were obtained by exciting at 277 nm and collecting spectra
between 300 and 400 nm. Data were corrected for the dilution (which
never exceeded 5%), for the buffer, FREG-(48-58), and SCR
contribution. Peak fluorescence at 310 nm, plotted as a function of
FREG-(48-58)/FGF-2 molar ratio, shows a dose-dependent and
saturable quench. C and D, SPR analysis showed
that FGF-2 interaction with immobilized FGF-2 (C) or with
immobilized heparin (D) reduces as a function of
FREG-(48-58), while SCR shows no effect. E, FGF-2
interaction with FGF-R1 by SPR. Increasing concentrations of
FREG-(48-58) (1, 0 ng/ml; 2, 0.7 ng/ml;
3, 2.1 ng/ml; 4, 20 ng/ml; 5, 180 ng/ml; 6, 1.4 µg/ml; 7, 14 µg/ml)
dose-dependently inhibited FGF-2 interaction with
immobilized FGF-R1, up to 40%, reaching the plateau at 180 ng/ml. A
representative experiment is shown. This experiment was repeated three
times with similar results.
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Further studies indicated that FGF-2 fluorescence intensity at 310 nm
was progressively quenched by increasing FREG-(48-58) concentration
(Fig. 4B). The fluorescence quench reached a plateau around
a FREG-(48-58)/FGF-2 molar ratio = 1, whereas no
significant shift of the maximum fluorescence was observed. The quench
reached 50% of the maximum effect (IC50) at 1.33 ± 0.076 µM FREG-(48-58) concentration (versus
FGF-2, 2 µM). The equilibrium dissociation constant
(KD), according to a non-linear least-square fitting, was equal to 579 ± 272 nM. Under similar
experimental conditions SCR was ineffective.
Taken together these data indicate that a FREG-(48-58)/FGF-2
heterocomplex formation is observed in both solid phase and liquid phase conditions, with a KD (in fluorescence
studies) falling in the nanomolar range.
FGF-2 Interaction with FGF-2, with Heparin and with FGF-R1, in the
Presence of FREG-(48-58)--
FGF-2 dimerization is considered to be
necessary for receptor binding and activation; therefore, a potential
FREG-(48-58) interference with FGF-2/FGF-2 interaction was tested. SPR
analyses showed that interaction of soluble FGF-2 with immobilized
FGF-2 is inhibited by increasing doses of FREG-(48-58) up to about
50% (Fig. 4C). Further SPR experiments carried out on
immobilized heparin showed that FREG-(48-58) markedly and
dose-dependently reduced FGF-2 interaction with heparin
(Fig. 4D). FREG-(48-58) IC50 values in FGF-2
self-interaction and heparin-binding experiments were 820 ng/ml and 980 ng/ml, respectively. Additional SPR experiments showed that increasing
FREG-(48-58) doses (from 0 to 14 µg/ml) inhibited up to 40% FGF-2
interaction with immobilized FGF-R1, reaching the plateau at 180 ng/ml
(Fig. 4E).
FREG-(48-58) also inhibited FGF-2/FGF-2 interaction in overlay
experiments, while the interaction of immobilized FGF-2 with a
polyclonal anti-FGF-2 antibody was unaffected (data not shown). In all
cases SCR was ineffective. Taken together, these data show that
FREG-(48-58) strongly inhibits FGF-2 interaction with itself, with
heparin, and with FGF-R1.
FGF-2 Receptor Phosphorylation and FGF-2 Internalization in the
Presence of FREG-(48-58)--
It was then tested whether the observed
action of FREG-(48-58) would affect FGF-2-dependent
receptor phosphorylation and FGF-2 internalization.
FGF-2-dependent phosphorylation of FGF-R1 in BAEC was
markedly diminished by FREG-(48-58) (10 ng/ml), while SCR did not show any effect (Fig. 5A). Further,
biotinylated FGF-2 internalization was measured in BAEC. A large excess
of unlabeled FGF-2, as well as mAb 125, a FGF-2 neutralizing antibody
(Chemicon), abolished FGF-2 internalization, as expected. Under such
conditions, FREG-(48-58) significantly reduced FGF-2 internalization
by about 50% while SCR was inactive. Quantification of this effect is
reported in Fig. 5B, while Fig. 5C shows one
representative experiment.

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Fig. 5.
FREG-(48-58) effect on FGF-R1
phosphorylation and FGF-2 internalization. A, FGF-R1
phosphorylation. FGF-R1 was immunoprecipitated and samples were
subjected to electrophoresis and detected with an anti-phosphotyrosine
antibody. FGF-2-induced FGF-R1 phosphorylation was
dose-dependently inhibited in the presence of FREG-(48-58)
(0.1-1000 ng/ml), while SCR was ineffective. Where not specified,
peptides were used at a 10 ng/ml dose. Results represent the average of
three experiments ± S.D. Asterisk indicates
p < 0.01 versus FGF-2 alone. B,
FGF-2 internalization. Membrane-bound biotinylated-FGF was discarded;
internalized biotinylated-FGF-2 was spotted onto nitrocellulose and
quantified. Internalization was significantly inhibited in the presence
of an excess of unlabeled FGF-2, in the presence of the FGF-2
neutralizing monoclonal antibody mAb 125, and in the presence of
FREG-(48-58) and of AbFREG48-58, while preimmune rabbit serum (AbP-I)
and SCR were ineffective. Results represent the average of three
experiments ± S.D. C, representative experiment of
FGF-2 internalization.
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These results indicate that, in the presence of FREG-(48-58), FGF-2
internalization, which depends upon its binding to membrane-bound receptors, and FGF-R1 phosphorylation are markedly inhibited.
Developing the Anti-FREG-(48-58) Antibody (AbFREG48-58)--
To
further investigate the role of FGF-2 region 48-58, a rabbit
polyclonal antibody was raised against FREG-(48-58) (AbFREG48-58) and
was tested in vitro. It almost abolished FGF-2
internalization at a 1:200 dilution, and this effect was comparable to
that of the commercially available neutralizing antibody mAb 125 used at a 1:500 dilution (Fig. 5, B and C). This
finding supported the hypothesis that region 48-58 of FGF-2
may be functionally relevant. Furthermore, AbFREG48-58 recognized
increasing doses of FGF-2 immobilized onto nitrocellulose (Fig.
6A), and FREG-(48-58) competed dose-dependently with this specific interaction
(Fig. 6B). These results indicated that FREG-(48-58)
represents an exposed epitope and may be a functionally relevant region
of the whole FGF-2. This hypothesis was further confirmed in
proliferation experiments in which AbFREG48-58 acted as neutralizing
antibody by significantly reducing the FGF-2 mitogenic effect at 1:4000 dilution (Fig. 6C). These data indicate that AbFREG48-58
blocks FGF-2 internalization and the FGF-2 mitogenic effect, supporting the hypothesis that the region encompassing FREG-(48-58) is a crucial
functional domain.

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Fig. 6.
Investigating AbFREG48-58. A,
AbFREG48-58 interaction with immobilized FGF-2. Increasing quantities
of FGF-2 were spotted onto nitrocellulose membrane (0, 6, 12.5, 25, 50, and 100 ng/spot, respectively); membrane was then incubated with
AbFREG48-58 (1:200 in PBS), and bound antibody was revealed with
anti-rabbit peroxidase-conjugated IgG and chemiluminescence.
AbFREG48-58 recognized the immobilized FGF-2. This experiment was
performed three times with similar results. B, FREG-(48-58)
competition. Increasing quantities of FREG-(48-58) (up to 100 ng/ml)
dose-dependently competed for the interaction between FGF-2
and AbFREG48-58, carried out as reported in panel A. C, FGF-2 induced BAEC proliferation at 48 h, in the
presence of AbFREG48-58 (1:2000; 1:4000) or in the presence of
preimmune rabbit serum (same dilutions). AbFREG48-58 (1:4000)
significantly inhibited FGF-2-induced proliferation, while the control
antibody was ineffective. The two antibodies did not modify cell number
under basal conditions.
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Histidine Residue Chemical Inactivation--
Histidines are very
reactive residues and often occupy functionally relevant sites in
proteins. Human FGF-2 contains three histidines, His-16, His-35, and
His-50, the latter falling within the FREG-(48-58) region. In order to
test whether histidine residues play a role in FGF-2 activity, their
direct and specific chemical inactivation was achieved by DEPC
treatment. Interestingly, DEPC-treated FGF-2 was unable to interact
with itself (Fig. 7A) and
failed to induce BAEC chemotaxis (Fig. 7B). Further,
inactivating FREG-(48-58) histidine by DEPC incubation, abolished the
FREG-(48-58) ability to inhibit FGF-2-induced chemotaxis (Fig.
7B). Taken together these results show that histidines play
a crucial functional role for FGF-2 activity in vitro, and
indicate that the histidine present in FREG-(48-58) is essential for
its biological effects.

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Fig. 7.
Histidines inactivation. Histidines were
chemically modified by incubating FGF-2 and FREG-(48-58) with
DEPC. As a control, inactivated DEPC (i.e. DEPC in 100 mM Tris-HCl) was added. A, FGF-2 dimerization by
SPR. Native FGF-2 interacted with immobilized FGF-2 reaching about 50 RU. In contrast, FGF-2 previously incubated with DEPC, entirely lost
the ability to interact with immobilized FGF-2. Inactive DEPC did not
modulate FGF-FGF interaction. A representative experiment is reported.
This experiment was carried out three times, with similar results.
B, BAEC migration induced by FGF-2, in the presence or in
the absence of FREG-(48-58), previously treated or not treated with
DEPC. FGF-2 entirely lost its ability to induce BAEC migration when
histidines were modified; FREG-(48-58) (0.1-1000 ng/ml)
dose-dependently inhibited BAEC migration, and at all
tested doses the inhibitory effect was abolished after DEPC treatment.
Where not specified, FREG-(48-58) was tested at 10 ng/ml. FGF-2
chemotactic activity was unaffected by incubation with inactive DEPC.
Data are expressed as percent of the effect of FGF-2 alone ± S.D.
In response to FGF-2, 31 ± 6 cells/field migrated.
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In Vivo Angiogenesis Assays--
In order to evaluate whether
FREG-(48-58) modulates FGF-2 effects in vivo, two different
assays were performed, i.e. the CAM assay and the Matrigel assay.
FREG-(48-58) Effect in the CAM Assay--
Ten CAM per group were
treated with PBS, FGF-2 alone, FREG-(48-58) alone, or FGF-2 + FREG-(48-58). Experiments were stopped at day 12 and examined
macroscopically and histologically. FREG-(48-58) abolished the new
vessel formation induced by FGF-2, either inside the sponge trabeculae
or at the boundary between the sponge and the CAM mesenchyme.
Quantification of the effect achieved by histologic examination and
planimetric vessel counting is reported in Table I, and a representative experiment is
reported in Fig. 8A.
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Table I
Chick embryo CAM-sponge assay: microvessel density
For histologic assessment, every third section of 30 serial slides from
each specimen was analyzed under a 144-point mesh inserted into the
eyepiece of the photomicroscope. The total number of intersection
points occupied by transversally cut vessels (3-10 µm in diameter)
were counted at ×250 magnification inside the sponge and at the
boundary between the sponge and surrounding CAM mesenchyme in 6 randomly chosen microscopic fields per section and reported as
mean ± S.D. Asterisks indicate statistically significant
difference from FGF-2 (p < 0.01).
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Fig. 8.
Angiogenesis assays in CAM and Matrigel.
A, on 8-day-old chick embryos, 1 mm3 gelatin
sponges (Gelfoam, Upjohn Company) loaded with 3 µl of PBS alone
(d), or FREG-(48-58) alone (50 µg/ml) (c), or
FGF-2 alone (500 ng) (a), or FGF-2 + FREG-(48-58) (500 ng
and 50 µg, respectively) (b), were implanted; CAM were
then analyzed at day 12. In panel a small blood vessels
converge like spokes toward the sponge, while in b,
c, and d there are very few small blood vessels
around the sponge or converging toward it. Original magnification:
×50. B, quantification of angiogenesis in Matrigel plugs
injected subcutaneously in CD1 mice, in the presence of FGF-2 alone
(150 ng/ml), FGF-2 + FREG-(48-58) (150 ng/ml and 10 µg/ml,
respectively), or FGF-2 + SCR (150 ng/ml and 10 µg/ml, respectively).
FGF-2-induced angiogenesis was significantly inhibited by FREG-(48-58)
(p < 0.01), while SCR was ineffective. Matrigel
without FGF-2 presented 25 ± 8 vessels/mm2. This
number was subtracted from the other groups as background. Data are
expressed as % versus control ± S.D. (100% = 250 ± 28 vessels/mm2).
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FREG-(48-58) Effect in the Matrigel Assay--
FGF-2-induced
angiogenesis in Matrigel plugs injected subcutaneously in CD1 mice was
also evaluated. Eight days after implant, Matrigel plugs were removed,
histologically processed, and new blood vessel formation was measured.
As shown in Fig. 8B, FREG-(48-58) significantly inhibited
(p < 0.01) FGF-2-induced vessel formation by 49 ± 5%, while SCR was ineffective. Taken together, in vivo studies indicate that FREG-(48-58) markedly inhibits FGF-2-induced angiogenesis in two different in vivo assays.
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DISCUSSION |
FGF-2 dimerization is reported to be a necessary event for
receptor dimerization and signaling activation; therefore, region(s) directly involved in FGF-2 dimerization may represent crucial site(s)
controlling the whole FGF-2 activity. The dimer in the absence of
heparin has not been crystallized for FGF-2 although it is known for
FGF-9 (50). Conversely, complexes in the presence of heparin (but
lacking a direct FGF-FGF interaction) have been crystallized for FGF-1
(51) and FGF-2 (20). These studies revealed different
dimerization/oligomerization features for different members of the FGF
family, likely due to the differences in the
1,
2,
9, and
10 loops, outside the
trefoil core (50). Different
three-dimensional simulation-based studies predicted different
organizations of the FGF-2 dimer in the absence of heparin (15-17) and
indicated the theoretical possibility that FGF-2 monomers may interact
and associate with different orientations. The dimer model predicted in
this study shows a high content of hydrophobic residues at the dimer
interface (i.e. 41%), which might contribute stability to
the predicted dimer. FREG-(48-58) was selected because its sequence is
poorly conserved within the FGF family (Fig. 1A) (being
specific for FGF-2, whereas other portions are conserved throughout the
entire family), and because it falls at the predicted interface of the
FGF-2 dimer. Further, it overlaps, or is close to, receptor- and
heparin-binding residues. FREG-(48-58) strongly and specifically
inhibited FGF-2 activity in vitro and markedly reduced
FGF-2-induced new vessel formation in vivo. An
FGF-2/FREG-(48-58) complex formation was observed and, likely as
consequence of this, FGF-2 interaction with itself, with heparin, and
with FGF-R1 was strongly reduced, as well as FGF-2-induced FGF-R1
phosphorylation and FGF-2 internalization. It is known that FGF-2
internalization follows its binding to either high affinity receptors
or to heparan sulfate proteoglycans (51). Therefore, the observed
FREG-(48-58)-induced inhibition of FGF-2 internalization (Fig. 5,
B and C) may be due either to reduced interaction
with high affinity receptors (Fig. 4E), to the inhibition of
FGF-2 self-interaction (Fig. 4C), or to reduced interaction
with heparan sulfate proteoglycans (Fig. 4D). In either
case, the extensive inhibitory effect indicates that FREG-(48-58) may
interfere with an early event essential for the receptor activation.
FGF-2 region 48-58 may control an early event, likely dimerization,
because the region 48-58 stays at the predicted FGF-2 dimerization
interface, essential to activate FGF-2 signaling. In this case, the
observed reduced heparin-binding, receptor binding, and receptor
activation may be consequences of dimer formation inhibition. However,
a direct effect of FREG-(48-58) on heparin binding and receptor
binding cannot be ruled out at this stage. FREG-(48-58) may affect
FGF-2 dimer formation by directly interacting with FGF-2 and masking
the dimerization site, although presently we do not have a definitive
indication of the site of interaction with FGF-2.
Alternative approaches confirmed that region 48-58 may be an FGF-2
crucial functional domain. In fact, the antibody raised against
FREG-(48-58) recognized the whole FGF-2 and blocked FGF-2 internalization and its mitogenic effect, indicating that region 48-58
is a relevant epitope and that it may have a crucial functional role.
Further, histidine modification abolished FGF-2 and FREG-(48-58) in vitro activities, indicating that histidines, and
specifically His-50, play a crucial role for FGF-2 and FREG-(48-58)
activity, respectively. Site-directed mutagenesis studies will further
characterize active residues in this region.
Several endogenous factors control angiogenesis (31, 53-54) and a
number of molecules inhibiting FGF-2 are currently being investigated
(47, 55-62). Specifically inhibiting FGF-2-dimerization with
FREG-(48-58) may represent a novel effective approach to modulate
FGF-2 activity with potential therapeutic applications. To this
respect, a biocomputing tool potentially helpful to investigate protein
active sites has been recently made available (63) at crisceb.unina2.it/ASC/. In conclusion, results of the present study
indicate a novel FGF-2 regulatory domain and identify a short peptide,
which strongly inhibits FGF-2 activity both in vitro and
in vivo. Further studies will be aimed at characterizing the
potential therapeutic role of this peptide.