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HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY
From the Laboratorio di Patologia Vascolare, Istituto
Dermopatico dell'Immacolata, Istituto di Ricovero e Cura a Carattere
Scientifico, Rome; Dipartimento Anatomia Umana e Istologia,
Università degli Studi, Bari; Dipartimento Istologia ed
Embriologia Medica, Università La Sapienza, Rome; Dipartimento
Biologia, Università Tor Vergata, Rome; and Dipartimento
Chirurgia Vascolare, Ospedale San Giovanni-Addolorata, Rome, Italy.
Basic fibroblast growth factor (bFGF) and platelet-derived growth
factor-BB (PDGF-BB) modulate vascular wall cell function in vitro and
angiogenesis in vivo. The aim of the current study was to determine how
bovine aorta endothelial cells (BAECs) respond to the simultaneous
exposure to PDGF-BB and bFGF. It was found that bFGF-dependent BAEC
migration, proliferation, and differentiation into tubelike structures
on reconstituted extracellular matrix (Matrigel) were inhibited by
PDGF-BB. The role played by PDGF receptor The endothelial layer represents a physical and
chemical barrier between the vessel lumen and the underlying tissues.
Endothelial cells (ECs) exert a variety of functions and modulate
underlying smooth muscle cells by releasing molecules with vasoactive
and growth-regulatory properties.1 Endothelial cells
present an active replication phenotype in vitro, but in vivo they are
quiescent.2 The different expression of membrane-bound
receptors3,4 and the in vivo action of specific
inhibitors5-7 may account, at least in part, for the
different replication pattern observed.
Basic fibroblast growth factor (bFGF) is a potent EC growth factor; it
is known to induce a proangiogenic phenotype in ECs and is released
under acidosis conditions that induce EC protection from
apoptosis.8 bFGF plays a critical role in physiologic and
pathologic angiogenesis, including tumor angiogenesis.9,10 It exerts its functions by direct action,11 by inducing
vascular endothelial growth factor (VEGF) synthesis,12 or
by potentiating VEGF activity.13,14
Platelet-derived growth factor (PDGF) is a growth factor known to be
active on ECs. Three PDGF isoforms have been identified as
disulfide-linked dimers, namely PDGF-AA, PDGF-BB, and PDGF-AB, expressed by ECs under various conditions.15-19 They
interact with different affinity with 2 tyrosine-kinase receptors,
PDGF-R We have recently shown that bFGF inhibits PDGF-BB mitogenic and
chemotactic activity on primary rat aorta smooth muscle cells in vitro
and that this inhibitory action is mediated by PDGF-R Therefore, we have hypothesized that PDGF may inhibit bFGF-induced
angiogenesis, and we have, in the current study, determined whether
PDGF modulates bFGF activity on ECs in vitro and in vivo. We found that
PDGF, through PDGF-R Cell culture
Migration assay
PDGF-BB heat inactivation was performed by repeating 3 times a cycle consisting of heating at 100°C for 10 minutes followed by fast refrigeration. A polyclonal anti-PDGF-BB neutralizing antibody (AB-220-NA; R&D Systems) was placed in the lower chamber of the Boyden apparatus, at the final concentration of 1 µg/mL, to neutralize PDGF-BB activity. The same concentration was used to block PDGF-BB-dependent receptor phosphorylation. Neutralizing goat anti-human bFGF (Ab-233-NA; R&D Systems) was used at 1 µg/mL concentration. Migration assays were carried out at 37°C in 5% CO2 for 6 hours, and filters were then removed, fixed with absolute ethanol, and stained with toluidine blue (Sigma). Cells that migrated across the filter were counted at × 400 magnification; 10 fields/filter were evaluated, and the average number of cells/field was reported. All experiments were performed at least 3 times in duplicate. Proliferation assay BAECs plated in 6-well plates (1 × 105 cells/plate) were grown for 24 hours in DMEM supplemented with 10% FCS, at 37°C in 5% CO2. Medium was then replaced with serum-free DMEM for 24 hours. Subsequently, the medium was replaced with fresh medium containing either 0.1% BSA alone or 0.1% BSA with growth factors. After treatment, cells were harvested and counted with a hemacytometer. All experiments were carried out at least 3 times in duplicate.Cell transfection with dominant-negative PDGF receptor constructs BAECs were transfected as previously reported.37 Either a dominant-negative PDGF-R vector (DN-PDGF-R) or an equal
amount of dominant-negative PDGF-R vector (DN-PDGF-R) (generous
gifts of Dr C. H. Heldin, Ludwig Institute for Cancer Research,
Uppsala, Sweden)44 or PcDNA3 (Invitrogen, Groningen, The
Netherlands) empty vector was used. Cells were cotransfected with
pEGFP-N1 (Clontech, Temecula, CA) reporter vector, with a
dominant-negative versus a reporter-vector molar ratio 4:1. BAEC
transfection (8 × 105 cells/60-mm diameter dish) was
carried out with Lipofectamine Plus reagent (Gibco) for 5.5 hours at
37°C in a 5% CO2 environment. Medium was then replaced
with DMEM-10% FCS, and migration and phosphorylation assays were
performed. Cotransfection with 2 vectors, at the reported ratio,
results in the internalization of both plasmids by the same
cell.45 Therefore, migrated cells were counted under a
fluorescence microscope to evaluate only GFP-positive cells to
overcome the potential limitations of low transfection efficiency.
Transfection was carried out according to the manufacturer's instructions, and GFP transfection efficiency was higher than 70%.
PDGF receptor phosphorylation in DN-transfected cells was markedly
inhibited compared with mock-transfected cells in all experiments,
indicating that the transfection process was effective. All
transfection experiments were performed at least 3 times in duplicate.
Western blot analysis and receptor phosphorylation BAECs were grown to 70% confluence and were incubated for 2 days in DMEM containing 1% FCS and for 1 day in serum-free DMEM.46 Transfection with DN-PDGF-R was carried out as
reported above; then transfected cells were incubated overnight in
serum-free conditions. Cells were treated with growth factor or
antibodies at the indicated concentrations for 5 minutes, rinsed with
ice-cold phosphate-buffered saline (PBS), scraped, and lysed for 15 minutes with 1% triton, 10% glycerol, 100 mM NaCl, 20 mM HEPES pH
7.4, 5 mM EDTA, 1% AEBSF, 1% pepstatin A, 1% E-64, 1% bestatin, 1% leupeptin, 1% aprotinin, 1 mM phenylmethylsulfonyl fluoride (PMSF), 10 µM NaVO3, and 50 mM NaF (Sigma). Lysates were then
centrifuged at 14 000 rpm for 15 minutes. After the determination of
protein concentration, 200 µg total proteins were subjected to 7.5%
sodium dodecyl fluoride-polyacrylamide electrophoresis (SDS-PAGE).
Proteins were electrotransferred to a nitrocellulose membrane
(Amersham, Uppsala, Sweden) and were blocked with PBS containing 0.1%
Tween 20 and 5% nonfat milk (Bio-Rad, Hercules, CA). Ponceau S (Sigma) staining of the filters was also performed to verify the equal loading.
Western blot analysis was carried out by probing the nitrocellulose
membrane with the antiphosphotyrosine antibody (1 µg/mL; Upstate
Biotechnology, Lake Placid, NY), for 1 hour, followed by washing and
incubating with horseradish peroxidase-conjugated secondary antibody
(Pierce, Rockford, IL). Bands reported in PDGF receptor phosphorylation
figure migrated at the apparent molecular weight corresponding
to PDGF receptors. A neutralizing anti-PDGF antibody significantly
reduced the phosphorylation, and DN-PDGF-R-transfected cells showed
marked reduction of the band phosphorylation; therefore, the reported
bands were identified as PDGF receptors and PDGF-R, respectively.
Bands were then revealed by means of the enhanced chemiluminescence
(ECL) detection system (Amersham) and were quantified with a calibrated
imaging densitometer (GS 710; Bio-Rad). All experiments were performed
at least 3 times.
Mitogen-activated protein kinase activation assay BAECs were seeded onto 150-mm culture plates (1 × 106 cells) incubated in DMEM containing 0.5% FCS for 2 days and in serum-free DMEM for the following day. Medium was then replaced with fresh medium containing either 0.1% BSA alone or 0.1% BSA with growth factors (10 ng/mL bFGF, 10 ng/mL PDGF-BB, or both) for 10 minutes. Cells were washed in ice-cold PBS and scraped in lysis buffer containing 20 mM Tris pH 7.5, 150 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1% triton, 2.5 mM sodium pyrophosphate, 1 mM-glycerol-phosphate, 1 mM Na3VO4, 1 µg/mL leupeptin, and 1 mM PMSF. Cell extracts were sonicated and
centrifuged, and supernatants were collected. After protein content
determination with the Bio-Rad protein assay system (Bio-Rad), samples
(30 µg total protein each), suspended in Laemmli sample buffer and
boiled for 5 minutes, were resolved on 8% SDS-PAGE and transferred to
nitrocellulose membrane (Amersham). Ponceau S (Sigma) staining of the
filters was also performed to verify the equal loading. Filters were
then probed with primary antibody antiphosphorylated ERK p44/42 (New
England Biolabs, Hitchin, United Kingdom), which specifically
recognizes activated forms of ERK1/2, and with horseradish
peroxidase-conjugated secondary antibody (Pierce). Detection was
carried out using the ECL detection system (Amersham) and was
quantified with a calibrated imaging densitometer (GS 710; Bio-Rad).
Six experiments were performed.
Differentiation assay Endothelial cell differentiation into tubular structures (TS) was assessed as previously reported.42 Briefly, 200 µL reconstituted extracellular matrix protein (Matrigel), growth factor-reduced and without phenol red (Collaborative Research, Bedford MA), was applied to 24-well culture plates and was incubated at 37°C for 60 minutes. BAEC (8 × 104 in 1 mL medium) were seeded on Matrigel-coated wells in the presence of 0.1% BSA, 1% FCS, or bFGF (10 ng/mL), or bFGF/PDGF-BB (10 ng/mL each), or PDGF-BB alone (10 ng/mL), and were kept at 37°C. Morphology changes and TS appearance were monitored at 0, 2, 3, 4, 6, 8, and 24 hours after plating by phase-contrast microscopy. Quantification of TS formation was expressed by the mean number of branching points in 10 fields. This experiment was carried out 3 times in duplicate, and similar results were collected.Angiogenesis on gelatin sponge chick chorioallantoic membrane assay Chick embryo chorioallantoic membrane (CAM) assay was performed as previously reported.47 Briefly, fertilized White Leghorn chicken eggs (30/group) were incubated at 37°C at constant humidity. On incubation day 3, a square window was opened in the shell, and 2 to 3 mL albumen was removed to allow detachment of the developing CAM. The window was sealed with glass, and the eggs were returned to the incubator. At day 8, 1 µL sterilized gelatin sponges (Gelfoam, Upjohn, Kalamazoo, MI) adsorbed with bFGF alone (500 µg), PDGF-AA alone (500 µg), PDGF-BB alone (500 µg), bFGF/PDGF-AA, or bFGF/PDGF-BB (500 µg each) dissolved in 2 mL PBS were implanted on the top of growing CAMs under sterile conditions. Sponges containing vehicle alone were used as negative control. At day 12, blood vessels entering the sponge within the focal plane of the CAM were recognized macroscopically (at 50×), counted by 2 observers in a double-blind fashion48 under a Zeiss SR stereomicroscope (Zeiss, Oberkochen, Germany), and photographed in vivo with an MC63 Camera System (Zeiss). Percentage inhibition reported in the text was computed after subtracting spontaneous angiogenesis ie, vessels in the presence
of PBS only.
The angiogenic response was also assessed histologically by a planimetric method of point counting.47 Briefly, in every third section within 30 serial slides, an individual specimen was analyzed by a 144-point mesh inserted in the eyepiece of a Leitz-Dialux 20 photomicroscope. Six randomly chosen microscopic fields were evaluated for each section at × 250 magnification. To this purpose, the total number of the intersection points occupied by vessels transversally cut (diameter, 3 to 10 µm) inside the sponge and at the boundary between the sponge and the surrounding CAM mesenchyme were counted. Mean values ± SD were determined for each analysis. Vascular density was indicated by the final mean number of occupied intersection points. Angiogenesis in Matrigel plugs Angiogenesis assays in reconstituted basement membrane (Matrigel) plugs were carried out according to a previously reported procedure.49 Matrigel containing bFGF (150 ng/mL) alone or bFGF/PDGF-BB (150 ng/mL each) was injected subcutaneously in CD1 mice (female, 19 weeks of age). Under these conditions bFGF induces a vascular network formation in the Matrigel plug within 8 days; Matrigel plugs were excised 8 days after injection and included in paraffin. Slides were stained with the Trichrome-Masson staining procedure (Bio-Optica, Milan, Italy), and vessel quantification of the portion of histologic sections occupied by Matrigel was obtained by an integrated image analysis system (Quantimet 970; Cambridge Instruments, United Kingdom), according to a published procedure.50 The number of newly formed vessels within the whole Matrigel area was expressed as number of vessels/mm2.Statistics Data are expressed as the mean ± SD. Student 2-tailed paired t test was performed, and P .01 was
considered statistically significant.
Effect of PDGF-BB on bFGF-induced BAEC migration, proliferation, and differentiation BAEC migration was investigated in response to bFGF alone, PDGF-BB alone, and bFGF/PDGF-BB, in concentration-dependence experiments, with increasing concentrations of one factor and a fixed concentration of the other. PDGF-BB alone (1 to 50 ng/mL) showed a weak chemotactic action, but it inhibited, in a concentration-dependent manner, migration induced by bFGF (10 ng/mL), and at 10 ng/mL it inhibited bFGF-induced migration by approximately 50% (Figure 1A). Basic FGF-induced migration was inhibited at all tested concentrations by 10 ng/mL PDGF-BB (Figure 1B). PDGF-BB did not show any inhibitory effect when it was heat-denatured (see inset in Figure 1A) nor when its action was neutralized with a specific antibody (1 µg/mL), whereas a control antibody was ineffective (not shown). Further, EGF did not show any inhibitory activity on bFGF-induced BAEC migration (not shown), indicating that the bFGF chemotactic effect on BAEC is specifically inhibited by PDGF-BB.
Additional experiments were aimed at evaluating the effect of PDGF-BB
on bFGF mitogenic action. PDGF-BB (10 ng/mL) abolished the mitogenic
effect of bFGF on BAEC at all time points in a 3-day proliferation
assay (Figure 2). As observed in
migration experiments, heat-denatured PDGF-BB did not inhibit bFGF
mitogenic effect (not shown), PDGF-BB neutralization with a specific
antibody abolished its inhibitory effect, and a control antibody was
ineffective (not shown). These results show that the native and active
forms of PDGF-BB are required to inhibit chemotactic and mitogenic
actions of bFGF.
BAECs plated on Matrigel formed TS in a bFGF-dependent way; therefore,
we examined whether PDGF-BB modulated bFGF's ability to induce TS
formation. Basic FGF alone and PDGF-BB alone (10 ng/mL) induced TS
formation, and this effect was less pronounced than in response to 1%
FCS (Figure 3). When bFGF was mixed with PDGF-BB (10 ng/mL each), BAECs differentiated significantly less than
in response to either factor alone, and the magnitude of the response
was comparable to the control treated with BSA alone. PDGF-BB
inhibitory effect on bFGF was evident at 3 hours (Figure 3) and at 6 and 24 hours (not shown). These experiments indicated that PDGF-BB,
though a weak TS inducer, substantially inhibits BAEC differentiation
into TS induced by bFGF.
In additional experiments, PDGF-BB inhibited FCS-induced BAEC migration (not shown), indicating that PDGF-BB may exert this inhibitory effect in the presence of FCS. Taken together, these results show that PDGF-BB has a marked and specific inhibitory action on bFGF's proangiogenic effect on BAECs in vitro. Role of PDGF- R and PDGF-R can initiate
positive or negative signals, depending on the cell
type,51 and that PDGF-R stimulation inhibits smooth
muscle cell migration and proliferation.37-40 According to
the literature,4,20-22 we found that PDGF and bFGF
receptors are expressed on BAECs (data not shown); therefore; we
hypothesized that PDGF-BB-dependent inhibition of bFGF-induced BAEC
migration, proliferation, and differentiation may be mediated by PDGF
receptors. The role of PDGF-R was then investigated (1) by testing
PDGF-AA, which is a selective PDGF-R agonist, (2) by inhibiting
PDGF-BB binding to PDGF-R with neomycin, which has been shown to
specifically inhibit PDGF-BB binding to PDGF-R without affecting its
binding to PDGF-R,43,52 (3) by determining the effect
of PDGF-BB on BAECs transfected with dominant-negative PDGF-R or
dominant-negative PDGF-R, and (4) by evaluating PDGF-R
phosphorylation in the presence of PDGF and bFGF.
PDGF-AA alone showed no significant chemotactic activity; however, it
significantly inhibited migration induced by bFGF. The PDGF-AA effect
at 10 ng/mL was comparable to that of PDGF-BB, and it lowered BAEC
migration to control levels (Figure 4).
Further, PDGF-AA and PDGF-BB, when simultaneously added to bFGF, did
not exhibit a synergistic or an additive action. Therefore, PDGF-AA, by
selectively binding PDGF-R
In other experiments, PDGF-BB binding to PDGF-R The role of PDGF receptors in PDGF-BB-induced inhibition of bFGF
effects was examined further in migration experiments on BAECs
transfected either with the dominant-negative form of PDGF-R
PDGF receptor phosphorylation and mitogen-activated protein kinase activation In additional experiments carried out on BAECs, PDGF receptor phosphorylation was found to be increased on bFGF/PDGF-BB treatment (5 minutes) in comparison with PDGF-BB alone (Figure 6A). The average increase was 53% ± 15% (n = 3) by densitometric analysis. Receptor phosphorylation was markedly inhibited by anti-PDGF-BB neutralizing antibody (1 µg/mL). Receptor was then investigated by treating
cells with PDGF-AA, which binds only PDGF-R in the presence or in
the absence of bFGF. Under these conditions, PDGF-R phosphorylation
was increased in the presence of bFGF/PDGF-AA compared with PDGF-AA
alone; phosphorylation of this receptor in DN-PDGF-R-transfected
cells was almost abolished, as expected. (Figure 6B). No other bands
were found to be modulated under these conditions. These results
indicate that receptor -phosphorylation is increased in the presence
of bFGF/PDGF compared with PDGF alone.
Additional investigation was carried out on mitogen-activated protein (MAP) kinases, which play a central role in PDGF-BB and bFGF signaling. Therefore, we examined the effect of simultaneous exposure to bFGF and PDGF-BB on the ERK1/2 activation. Western blot analysis indicated that ERK1/2 phosphorylation was markedly inhibited (41% ± 14% and 46% ± 6%, respectively) on treatment with bFGF/PDGF-BB compared with bFGF alone (Figure 6C). This finding suggests that PDGF-BB-mediated inhibition of bFGF activity may involve, at least in part, the inhibition of MAP kinase-dependent signaling. Inhibitory effect of PDGF-BB on bFGF-induced angiogenesis in vivo The angiogenic effect of bFGF in the presence of PDGF-BB was tested in 2 angiogenesis assays in vivo.Chick embryo chorioallantoic membrane assay Gelatin sponges adsorbed with bFGF induced an angiogenic response, and allantoic vessels developed radially toward the implant in a spoked-wheel pattern (Figure 7A). Allantoic vessels were less numerous in the specimens treated with bFGF/PDGF-AA and bFGF/PDGF-BB (Figure 7D,F). Almost no vascular reaction was detectable around the sponges treated with PBS only or PDGF-AA (Figure 7B-C), whereas PDGF-BB induced a moderate angiogenic response (Figure 7E). Quantification of vascularization at day 12, in response to bFGF, PDGF-AA, PDGF-BB, bFGF/PDGF-AA, bFGF/PDGF-BB, and PBS, is reported in Table 1. Under these conditions, PDGF-BB and PDGF-AA induced 47% and 61% inhibition of bFGF angiogenic activity, respectively.
At the microscopic level, highly vascularized tissue was present among
the trabeculae of the bFGF-treated sponges (not shown). The tissue
consisted of newly formed blood vessels, mainly capillaries, with 3- to
10-µm diameters growing perpendicularly to the plane of the CAM and
of infiltrating fibroblasts within an abundant network of collagen
fibers. Vessels were less numerous in PDGF-BB-, bFGF/PDGF-AA-, and
bFGF/PDGF-BB-treated sponges and were absent among the trabeculae of
implants treated with PBS and PDGF-AA. These observations were
confirmed by quantification of the angiogenic response by a
morphometric method of point counting (Table
2). Under these conditions, PDGF-BB and
PDGF-AA induced 50% and 42% inhibition of bFGF angiogenic activity,
respectively. These experiments show that PDGF-BB and PDGF-AA reduce in
vivo bFGF-dependent blood vessel formation.
Subcutaneous Matrigel assay New vessel formation was induced in Matrigel plugs containing bFGF (Figure 8). Under these conditions PDGF-BB alone did not induce the formation of new blood vessels, whereas it inhibited angiogenesis induced by bFGF. Quantification of new vessel formation, carried out by the Quantimet image analyzer, revealed a 30% inhibition of new blood vessel formation in the presence of bFGF/PDGF-BB versus bFGF alone (P < .01). These experiments showed that PDGF-BB reduces in vivo bFGF-dependent blood vessel formation.
The current study shows that bFGF actions on BAECs are significantly inhibited by PDGF-BB. The native form of the PDGF-BB molecule was required to inhibit bFGF-dependent migration and proliferation; in fact, inhibition was not observed when heat-denatured PDGF-BB was mixed with bFGF. In addition, blocking PDGF-BB with a specific neutralizing antibody abolished the inhibition in migration and proliferation assays, but an aspecific antibody was ineffective. Finally, combining bFGF with EGF did not show any inhibition, confirming that the observed inhibition was a specific event. We have recently shown37 that PDGF-BB-directed migration
and proliferation of smooth muscle cells are inhibited in the presence of bFGF. The inhibition observed was somehow unexpected because PDGF-BB
and, to a lesser extent, bFGF are known to be positive regulators of
migration and proliferation of these cells. However, there is evidence
for opposite effects of PDGF, depending on the receptors expressed on
the target cells and, therefore, depending on receptor dimerization
events.38,40,53,54 Results presented in other studies show
that PDGF-R The PDGF-R PDGF-BB and bFGF are detected in the serum and within the vascular wall, in normal and pathologic conditions; it has been proposed that altering their balance may underlie, at least in part, vascular wall abnormalities.59 This may help in interpreting the PDGF-BB inhibitory effect on bFGF-induced migration, and it supports the hypothesis that PDGF family members may act as angiogenesis-modulating factors. The PDGF-BB inhibitory effect was also observed in bFGF-induced
angiogenesis in vivo The imbalance of positive and negative regulators of vascular cells may
underlie many vascular abnormalities.61,62 Modulating factors include the expression of known angiogenesis
inhibitors5,63 and the altered expression of PDGF
receptors64-66 or bFGF-receptors.67 As
reviewed by Battegay et al,18 PDGF is considered a
pro-angiogenic factor through an indirect effect, according to in vitro
and in vivo experiments. Although PDGF-R Recent evidence shows that PDGF-BB and bFGF directly interact with high affinity,70 and this may, at least in part, explain the observed inhibitory effect. In conclusion, the results of the current study show that activating
PDGF-R
We thank Francesco Facchiano for useful discussions. We also thank Gabriella Ricci and Cinzia Carloni for secretarial assistance.
Submitted January 29, 2001; accepted November 2, 2001.
Supported in part by project grants from the European Union (contracts BMH4-CT95-1160 and BMH4-CT97-2270) (F.D.M., M.S., M.C.C., A.F.); from FIRC (Italian Foundation for Cancer Research) and ASI grant ASI I/R/31/00 (A.F.); and from Associazione Italiana per la lotta al Neuroblastoma, Genoa, and Ministero dell' Università e della Ricerca Scientifica, Rome (D.R.).
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Antonio Facchiano, Laboratorio di Patologia Vascolare, Istituto Dermopatico dell'Immacolata, Via dei Monti di Creta 104, 00167 Rome, Italy; e-mail: a.facchiano{at}idi.it.
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Cines DB, Pollak ES, Buck CA, et al.
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Abstract
Introduction
