1 Institut National de la Santé et de la Recherche Médicale (INSERM) U127, Institut Fédératif de Recherche Circulation Paris VII, Hôpital Lariboisière, Université Denis Diderot, 75475 Paris Cedex 10, France; 2 Institut de Recherches Cliniques de Montréal, Montreal, Quebec, H2W 1T8 Canada; and 3 INSERM U492, Institut de Médecine Moléculaire, Hôpital Henri-Mondor, 94010 Créteil Cedex, France
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ABSTRACT |
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To explore the vascular function of the angiotensin II (ANG II) AT2 receptor subtype (AT2R), we generated a vascular smooth muscle cell (SMC) line expressing the AT2R (SMC-vAT2). The involvement of AT2R in the motility response of SMCs was examined in SMC-vAT2 cells and their controls (SMC-v) cultured on either laminin or fibronectin matrix proteins with the agarose drop technique. All experiments were conducted in the presence of a saturating concentration of losartan to inactivate the AT1R subtype. Under basal conditions, both cell lines migrated outside drops, but on laminin only. Treatment with ANG II significantly inhibited the migration of SMC-vAT2 but not SMC-v cells, and this effect was prevented by the AT2R antagonist CGP-42112A. The decreased migration of SMC-vAT2 was not associated with changes in cell growth, cytoskeleton stiffness, or smooth muscle actin, desmin, and tenascin expression. However, it was correlated with increased synthesis and binding of fibronectin. Both responses were prevented by incubation with selective AT2R antagonists. Addition of GRGDTP peptide, which prevents cell attachment of fibronectin, reversed the AT2R inhibitory effect on SMC-vAT2 migration. These results suggest that activated ANG II AT2R inhibits SMC migration via cellular fibronectin synthesis and associated cell binding.
vascular smooth muscle cells; laminin and fibronectin substrates; cellular fibronectin
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INTRODUCTION |
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NORMAL OR PATHOLOGICAL VESSEL REMODELING is generally described as a consequence of growth and/or phenotype alterations of the vascular smooth muscle cells (SMCs) (20). Migration of medial SMCs within the vessel wall, however, is another essential aspect of the process of vascular remodeling (4, 55, 59). Identification of the environmental signals susceptible to control SMC migration has been derived largely from studies in cultured SMCs. These include growth factors, such as platelet-derived growth factor or fibroblast growth factor, cytokines, components of the extracellular matrix (ECM) (1), and nitric oxide (8) and endothelial nitric oxide synthase (25). It also is well documented that matrix metalloproteinases (MMPs), by cleaving ECM components, participate in promoting SMC migration (36, 61, 73).
Accumulating evidence suggests that angiotensin II (ANG II) is an important mediator of systemic vascular remodeling. ANG II has been shown to promote hyperplasia and/or hypertrophy of vascular SMC in vitro as well as in vivo in the normal arterial wall (31) and to participate in the myointimal proliferation response to vascular injury (13, 47, 50). ANG II also stimulates migration of SMCs both in vitro (6) and during restenosis formation after vascular injury (46, 51). Two major subtypes of ANG II receptors, designated AT1 and AT2, have been identified on the basis of their affinity for selective receptor antagonists (reviewed in Ref. 15). It is currently admitted that the stimulatory effects of ANG II on either growth or migration of SMCs are attributable to the AT1 subtype (18, 19, 29, 31, 34, 46, 51, 71). Much less is known about the physiological roles of the AT2 receptor, which is abundantly expressed in the developing vascular system (45, 60, 66) and is reexpressed in adult SMCs in response to vascular injury (45), arteriogenesis after myocardial infarction (70), or hypoxia-induced lung vessel remodeling (11). In vitro investigations of the function of AT2 receptor have proven difficult, mainly because of low or lack of expression of this receptor in cultured SMCs (3, 31, 39, 46), which contrasts with the abundant expression of the AT1 subtype. Ectopically expressed AT2 receptor in cultured SMCs has been shown to exert either an antiproliferative (45) or apoptotic (72) effect, thereby counteracting the growth-promoting action of the AT1 receptor. Whether such an effect is accompanied by a modulation in the migratory properties of SMCs has not been explored in cultured SMCs.
In the present study, we have generated a vascular SMC line expressing the human AT2 receptor by retrovirus-mediated gene transfer to compare the effect of the AT2 receptor on SMC migration on either laminin or fibronectin substrate. To further define the molecular basis of the action of the AT2 receptor on SMC migration, we also have investigated the effect of AT2 receptor activation on SMC growth, cytoskeletal stiffness, and expression of several ECM and cytoskeleton components.
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MATERIALS AND METHODS |
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Reagents. Human ANG II was purchased from American Peptide Company. The radioligand 3-[125I]iodotyrosyl4[Sar1, Ile8]-ANG II was obtained from Amersham (specific activity 2,000 Ci/mmol) or prepared by radioiodination of [Sar1,Ile8]ANG II (sarile) using a solid-phase method. The AT1 and AT2 receptor-selective antagonists used were losartan (Merck Sharp & Dohme Research Laboratories), PD-123319 (kindly provided by Prof. B. Levy, Univ. Diderot, Paris, France), and CGP-42112A (Neosystem Laboratory, Strasbourg, France). The GRGDTP and GRGETP peptides were obtained from Sigma. Dulbecco's minimum essential medium (DMEM), fetal calf serum (FCS), L-glutamine, antibiotics, geneticin, trypsin-EDTA, phosphate-buffered saline (PBS), bovine serum albumin (BSA), and mouse laminin (Lam) were from Life Technologies, and human plasma fibronectin (FN) was from Sigma.
Infection of vascular SMC with the human AT2 receptor
gene.
A 1.3-kb genomic fragment containing the entire coding sequence of the
human AT2 receptor gene (10) was subcloned
into the retrovirus expression vector pLNCX (40) to
generate pLNCX-AT2. Empty pLNCX vector and
pLNCX-AT2 were transfected with Lipofectamine (Life
Technologies) into the amphotropic retrovirus packaging cell line
CRIP (14), and stably transfected cells were selected in complete medium containing 0.4 mg/ml G418 (Life Technologies). For
infection of vascular SMCs, viral supernatants were harvested from
helper-free retrovirus producer cell lines and applied to 30%
confluent cultures of rat aortic SMCs in the presence of 8 µg/ml
polybrene for 24 h, as previously described (24). The infected SMCs were then selected, expanded into cell lines, and screened by whole cell radioligand binding assay. The results presented
here were obtained with a representative cell line, designated
SMC-vAT2, which expresses a physiological number of AT2 receptors. A population of cells infected with empty
retroviruses, referred to as SMC-v, was used as control.
Receptor binding assays.
For whole cell binding assays, infected cells were grown to confluence
in six-well culture plates washed twice with DMEM and incubated with
~2 × 1010 M 125I-labeled sarile and
10
5 M losartan, in the absence or presence of competing
agents (PD-123319, 3 × 10
6 M; unlabeled sarile,
10
6 M), for 90 min at 25°C in a total volume of 1 ml of
DMEM, 25 mM HEPES (pH 7.4), and 0.1% heat-inactivated BSA. After
incubation, the cells were washed rapidly three times in ice-cold PBS.
Bound 125I-sarile was determined by counting the
cell-associated radioactivity in an Auto-Gamma counter after
solubilization in 0.1 M NaOH.
Cell culture and treatments. SMC-v and SMC-vAT2 cells were routinely grown in DMEM supplemented with 10% FCS, 2 mM L-glutamine, antibiotics (50 µg/ml streptomycin and 50 U/ml penicillin), and 0.4 mg/ml G418 at 37°C in a 5% CO2-95% air atmosphere. Subcultures were performed by incubating cells with 0.025% trypsin-0.02% EDTA in calcium- and magnesium-free PBS for 5 min at 37°C. Experiments were performed with cells at passages 7 and 8.
Culture plates were precoated with either Lam/H2O or FN/PBS (~1 µg/cm2). Before use, the plates were washed twice with 0.2% FCS-DMEM and then dried at room temperature for 1 h. Cells were incubated for 24 or 48 h in 0.2% FCS-DMEM supplemented with 0.4 mg/ml G418 and 10Cell migration assay.
The cell migration assay was similar to those previously described by
Varani and Ward (65) and Kiernan and ffrench-Constant (30), with a few modifications (49). Cells
were resuspended at 40 × 106 cells/ml in a 0.3%
agarose-DMEM solution containing 0.2% FCS and 105 M
losartan. Drops (2.5 µl) of the cellular agarose suspension were
plated on 24-well plates precoated with either Lam/H2O or FN/PBS and kept at 4°C for 20 min to allow gelation of the agarose. The drops were then incubated at 37°C for 24 or 48 h in a 0.5-ml volume of medium as described in Cell culture and
treatments. Afterward, the medium was removed and the samples were
fixed and stained with the Diff Quick kit (Dade). To assess the degree
of radial migration of cells from drop side to external environment, we
recorded light microscopy images of drops onto a computer after digitization using image analysis software (Perfect Image, Paris, France) that enables measurement of surface areas by enclosing them in
a cell-by-cell freehand curve. For each sample, both the area enclosed
by the drop side (i.e., the drop surface) and the area enclosed by the
cell front line (i.e., the drop surface plus the area between the drop
side and the cell front line) were measured. The subtraction of the
former from the latter gave a relative value representing cell
population migration. Because of the possibility of slight variations
in the surface area of the original drops, values were normalized to
the surface area of each corresponding original drop to obtain cell
migration indexes that were comparable among drops according to the
following formula: cell migration index = [(drop area + cell
migration area)
drop area]/drop area, expressed in arbitrary
units (AU).
Staining of cellular fibronectin on migrating cells by immunocytochemistry. Cellular drops were plated in the center of eight-well cover slides precoated with Lam/H2O and incubated as described above. At the end of incubation, cells were fixed with 4% paraformaldehyde and rinsed twice with ice-cold PBS. Immunocytochemistry was performed by using the biotin-avidin-peroxidase technique with 3-amino-9-ethylcarbazol (AEC) chromogen as a substrate (Sigma). Briefly, samples were preincubated for 30 min at 25°C in PBS supplemented with 5% BSA and then incubated at 4°C overnight with anti-cellular fibronectin (c-Fn) monoclonal antibody (MAb) (clone FN-3E2, Sigma) diluted 1:100 in PBS-2% BSA. After being washed in PBS, they were incubated for 30 min at 25°C with mouse biotinylated Ig (Dako) diluted 1:100 in 5% serum-PBS, washed with PBS, and treated with 3% H2O2 for 10 min. After being washed in PBS, the samples were incubated for 30 min at 25°C with avidin-peroxidase and then for 10 min with AEC substrate to allow adequate chromogen development. The cover slides were mounted and observed by microscopy.
Immunoblot analysis. The cellular contents in desmin, tenascin, smooth muscle actin, and fibronectin (c-Fn) were determined by Western blot analysis. Lysate proteins (4-10 µg) were resolved by electrophoresis on 7.5% SDS-acrylamide gel and electrophoretically transferred to Hybond-C nitrocellulose membranes (Amersham). The membranes were incubated for 1 h 45 min at 25°C with either anti-smooth muscle actin MAb (1:7,000; clone 1A4, Dako), anti-c-Fn MAb (1:7,000; clone FN-3E2), anti-tenascin C MAb (1:3,000; clone BC-24, Sigma), or anti-desmin polyclonal antibody (1:4,000) (52) in Tris-buffered saline (pH 7.4) containing 0.1% Tween 20. After being washed, the membranes were incubated with anti-mouse IgG or anti-rabbit IgG conjugated to horseradish peroxidase (1:5,000) (Amersham). Immunoreactive bands were visualized by enhanced chemiluminescence (Amersham) and quantified by densitometry using a computer-based imaging system (Gel Doc 1000; Bio-Rad).
Analysis of c-Fn secretion activity. Cells were stimulated with ANG II as described. After 12 h, [35S]methionine was added at a final concentration of 100 µCi/ml, and the cells were incubated for another 12 h. The medium was diluted with 3 vols of immunoprecipitation (IP) buffer [50 mM Tris · HCl (pH 7.4), 150 mM NaCl, 1 mM sodium orthovanadate, 5 mM EGTA, and 0.1% SDS], and c-Fn was immunoprecipitated at 4°C overnight with 5 µl of rabbit anti-plasmatic Fn polyclonal antibody (Chemicon). Immune complexes were collected by incubation with protein A/G-Sepharose beads for 2 h at 4°C. The beads were washed four times with IP buffer, and immunoprecipitates were analyzed by SDS-gel polyacrylamide on 7.5% acrylamide gels and visualized with a PhosphorImager.
Measurements of DNA and protein contents.
Cells were plated (40 × 103 cells/cm2) on
six-well plates and allowed to attach in DMEM-0.2% FCS for 24 h
at 37°C. They were preincubated for 15 min with losartan in the
absence or presence of PD-123319 and then treated for 24 or 48 h
as described in Cell culture and treatments. The DNA total
content was measured by fluorometry using the bisbenzimide dye Hoechst
33342 as described previously (53). After measurement, the
medium containing the Hoechst 33342 dye was removed, and the plates
were rinsed with PBS, frozen with liquid nitrogen, and stored at
80°C. Protein measurement was performed as already described.
Statistical analysis. Results are expressed as means ± SE. The statistical significance of differences between the various cell treatments was determined by one-way analysis of variance, and group-to-group comparison was made by Scheffé's F test. The accepted level of significance was P < 0.05.
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RESULTS |
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Expression of AT2 receptor in rat vascular SMCs.
Vascular SMCs infected with the human AT2 receptor gene
display a high increase in binding activity in contrast to cells
infected with empty vector (Fig.
1A). The AT2
receptor binds the radioligand in a saturable manner with a binding
capacity of 770 fmol/mg protein (data not shown). Binding assays were
conducted in the presence of a saturating concentration of losartan to
prevent binding of the radioligand to the AT1 receptor
subtype (Fig. 1B). The potency order of the ligands in
competing for 125I-sarile binding was sarile
(Kd = 0.7 nM) > CGP-42112A
(Kd = 1.2 nM) > ANG II
(Kd = 4.6 nM) > PD-123319
(Kd = 10.0 nM), similar to that reported
previously for the endogenous or cloned AT2 receptor (44). These results indicate that the stably expressed
recombinant receptor exhibits the typical pharmacological properties of
the AT2 receptor.
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AT2 receptor activation inhibits SMC migration.
The migratory properties of SMC-v and SMC-vAT2 cells on FN
and Lam substrates were analyzed in the presence of a saturating concentration of losartan by using the agarose drop technique. As shown
in Fig. 1, incubation with 105 M losartan completely
blocked the binding of ANG II to the AT1 receptor subtype,
thereby allowing study of the action of the AT2 receptor.
Figure 2 shows representative pictures of
cell drops incubated for 24 h under diverse conditions on either
FN (A) or Lam (B) substrate. In the presence of
serum, SMC-v and SMC-vAT2 cells migrated out from their
drops on either substrate, whereas under control conditions both cell
lines displayed migration on Lam only, indicating that under our
conditions Lam, but not FN, substrate is able per se to promote cell
migration.
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Inhibition of cell migration by the AT2 receptor is
concomitant with increased c-Fn secretion and fiber formation.
To define the molecular basis of the inhibitory action of the
AT2 receptor on SMC migration, we examined the impact of
AT2 receptor activation on both SMCv-AT2 cell
growth and expression of cytoskeletal (actin, desmin) and ECM
(tenascin, c-Fn) proteins. Total DNA and protein contents of
SMCv-AT2 cells plated on Lam substrate were unchanged after
either 24 or 48 h of ANG II treatment compared with controls
(Table 1), indicating that inhibition of
SMC migration in response to AT2 activation was not
associated with changes in cell growth properties. Thus the lower
number of cells observed around ANG II-treated drops results from a
decrease in the motility, not the proliferation, of
SMCv-AT2 cells.
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AT2-mediated inhibition of SMC migration is due to the
attachment of newly synthesized c-Fn.
To determine whether the increased binding of c-Fn is responsible for
the inhibition of cell migration, we examined the effect of the GRGDTP
peptide on the expression of c-Fn and the migration properties of
SMC-vAT2 cells. GRGDTP, as well as GRGETP control peptide,
had no effect on either basal or ANG II-stimulated c-Fn synthesis and
secretion in SMC-vAT2 cells (Fig. 6, A and
B). However, GRGDTP markedly
reduced the ANG II-dependent increase in c-Fn staining at the cell
surface (Fig. 6C vs. 5C), indicating that this
peptide specifically prevents cell attachment of c-Fn.
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DISCUSSION |
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In this study, we have demonstrated that AT2 receptor activation inhibits vascular SMC migration through the secretion and subsequent cell attachment of c-Fn. These results provide the first direct evidence for a negative influence of the AT2 receptor on SMC migrating capacity through a process involving modifications of proximal matrix environment.
Previous studies have documented the promoting effect of ANG II on migration of vascular SMCs in culture (6, 18, 19, 29, 34, 46, 71) or in vivo during neointimal thickening after balloon injury (46, 51). The blockade of ANG II-stimulated SMC migration by various AT1 but not AT2 receptor-selective antagonists (18, 19, 29, 34, 46, 71) has led to the conclusion that migration-promoting effects of ANG II are mediated through AT1 receptors. The role of the AT2 receptor had remained elusive in this setting.
Although AT2 receptors have been found to be expressed in
the vasculature of adult rats (37), their expression is
barely detectable in cultured SMCs (3, 39, 46, 64),
presumably because AT2 receptors are easily lost after
subculturing as postulated by Moriguchi et al. (42). In
agreement with these observations, we noted in this study the scarcity
of endogenous AT2 receptors in both the empty
virus-infected SMC-v (Fig. 1A) or the untransfected parental
cells (not shown). To circumvent this problem, we ectopically expressed
the AT2 receptor in aortic SMCs by retrovirus-mediated gene
transfer and conducted all experiments in the presence of a saturating
concentration of losartan to inactivate AT1 receptors. This
"gain-of-function" strategy enabled us to demonstrate that activation of AT2 receptor by ANG II increases the
synthesis, secretion, and cell binding of c-Fn. To our knowledge, this
is the first direct evidence for a stimulatory effect of
AT2 on c-Fn synthesis, which thereby extends previous
results that blockade of AT1 receptors in adult rat
increases aortic levels of c-Fn, presumably through the increased
stimulation of the "accessible" AT2 vascular receptors
(58). Further support for our finding is provided by
reports that 1) in SMCs, AT2 activates the
nuclear transcription factor NF-B (56), which enhances
c-Fn expression in gliastoma multiform-derived cell lines
(54) and 2) after arterial injury, both
AT2 receptors and c-Fn increase at the site of injury
(5, 45). It is noteworthy that AT2 receptor
also stimulates collagen synthesis in cultured SMCs (39)
and in vasculature in vivo (9, 33). It is noteworthy that
CGP-42112A, which per se had no effect on c-Fn synthesis (data not
shown), completely inhibited the stimulatory effect of ANG II on c-Fn
production (Fig. 5), indicating that this compound acts as a full
antagonist in our cell system. There have been conflicting reports on
the pharmacological properties of CGP-42112A, with some studies
describing it as an antagonist (7, 35, 41, 56, 62) and
others as a partial or full agonist (26, 63).
Because MMPs regulate the cell matrix environment by cleaving ECM components (36, 61, 73), we also examined the effect of AT2 receptor activation on MMP cell activity. We found that SMCv-AT2 cells produce mainly MMP-2, in agreement with previous reports that MMP-2 is the prevalent isozyme expressed in cultured (21) as well as in situ SMCs (67). Activation of the AT2 receptor did not change the levels of activated MMP-2 (data not shown). Thus the AT2-dependent increase in c-Fn deposition is not related to a decrease in cell MMP activity.
Newly synthesized c-Fn is secreted as a soluble monomer and polymerizes
in a insoluble, multimeric complex that modifies cell properties
(57). Changes in the strength of linkages between cell and
ECM components can be inferred from the measurement of cytoskeletal
stiffness by either magnetocytometry (32, 68, 69) or by
optical trapping (12). To determine whether the increased
c-Fn binding modifies cell-substrate interactions, we measured the
cytoskeletal stiffness of SMC-vAT2 cells by a
magnetocytometry approach, which is based on the application of
controlled mechanical stresses directly to cell surface integrins using
RGD-coated microbeads. Activation of AT2 receptor did not
change cytoskeletal stiffness of SMC-vAT2 cells (data not
shown), suggesting that cellular binding of newly polymerized c-Fn is
not enough to alter the strength of linkages between integrins and the
cytoskeletal apparatus (68). However, the
observation that GRGDTP peptide prevented both
AT2-dependent cell binding of c-Fn (Fig. 6) and inhibition
of cell migration (Fig. 7) indicates that the inhibitory action of
AT2 is due to cell attachment of c-Fn. This is in agreement
with the lack of migration of SMCs on FN substrate (Fig.
2A). These results also are consistent with the
demonstration that "superfibronectin," which resembles matrix
fibers, greatly enhances cell adhesive properties and suppresses cell
migration (43) and that accumulation of c-Fn within ECM
suppresses the motility and growth potential of fibrosarcoma tumor
cells, most likely through 5
1-integrins (2). Thus we propose that the twofold increase in c-Fn
synthesis in response to AT2 activation results in an
increase in the adhesive properties of SMC-vAT2 cells, with
this effect leading to inhibition of SMC-vAT2 cell
migration. The observation that PD-123319 is able to inhibit SMC
migration in an AT2-independent manner (Fig. 4) provides a
unifying explanation for the paradoxical results that PD-123319 fails
to antagonize the inhibitory action of AT2 receptor on cell
migration despite its inhibitory effect on c-Fn binding.
The amplitudes of the inhibitory effects of AT2 receptor on cell migration are comparable to the range of stimulated migration responses by AT1 receptor activation reported in the literature (6, 18, 29, 34), indicating that our results with this "artificial" expression system may be physiologically relevant. However, the observations that engagement of both AT1 and AT2 receptor stimulates c-Fn production and yet that the two receptors have apparently opposite effects on SMC migration, raise questions. Interestingly, DiMilla et al. (17) reported that the migration speed of human vascular SMCs on FN depends in a biphasic manner on both ECM surface density and attachment strength. It is maximal at an intermediate level of cell-substratum adhesiveness and minimal when the attachment strength is either weak or high. Thus it is conceivable that the increase in c-Fn synthesis that occurs in response to AT1 receptor activation results in an intermediate level of cell adhesiveness that allows cell migration, whereas that observed upon AT2 receptor activation is incompatible with migration stimulation.
In conclusion, we have shown that AT2 receptor activation
enhances both the synthesis and secretion of Fn and its binding to
SMCs, resulting in the inhibition of SMC migration. This novel insight
into AT2 receptor function may provide a new basis for understanding the exact role of AT2 in the structural
changes occurring during normal and pathological growth of blood
vessels, especially during postangioplasty restenosis. Indeed,
AT2 receptors have been implicated in the reduction of
neointimal formation after vascular injury (28, 45).
Interestingly, Pickering et al. (48) reported that a
subpopulation of 5
1-integrin-bearing SMCs
orchestrates integrin-mediated Fn assembly in the repairing artery
wall. It is therefore conceivable that AT2 receptors might, through the mechanism described in this study, participate in inhibiting vascular SMC migration from media to neointima, thus resulting in the reduction of arterial thickening.
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ACKNOWLEDGEMENTS |
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We thank Edith Giasson and Stéphane Pelletier (Institut de Recherches Cliniques de Montréal), Dominique Rideau and Antoine Mary (INSERM U492), and Françoise Marotte (INSERM U127) for technical help and advice.
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FOOTNOTES |
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* C. Chassagne and C. Adamy contributed equally to this work.
This work was supported by INSERM and Centre National de la Recherche Scientifique, by grants from the Canadian Institutes for Health Research (FRN 14168) and Fondation de France, and by a medical school grant from Merck Frosst Canada. S. Meloche is an Investigator of the Canadian Institutes for Health Research.
Previously presented in part at the 74th Scientific Sessions of the American Heart Association, Anaheim, Ca., November 11-14, 2001.
Address for reprint requests and other correspondence: C. Chassagne, INSERM U127, Hôpital Lariboisière, 41 bvd de la Chapelle, 75475 Paris Cedex 10, France (E-mail: catherine.chassagne{at}inserm.lrb.ap-hop-paris.fr).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajpcell.00318.2001
Received 12 July 2001; accepted in final form 20 November 2001.
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