Centre National de la Recherche Scientifique Unité de Recherche Associée 1283, Service de Biochimie, Centre Hospitalier Universitaire Saint-Antoine, 75012 Paris, France
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ABSTRACT |
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Oxidative
damage, which plays a major role in the early stages of
atherosclerosis, is associated with arachidonic acid (AA) release in
vascular smooth muscle cells (VSMC) as in other cell types. In this
study,
H2O2
was used to investigate mechanisms of AA release from VSMC on oxidative
stress. Cell treatment with H2O2
inhibited AA incorporation in an inverse relationship to prolonged
H2O2-induced
AA release. Identical kinetics of inhibition of AA incorporation and AA
release were observed after cell treatment with
AlF4, a process not involving
phospholipase A2
(PLA2) activation as recently
described (A. Cane, M. Breton, G. Béréziat, and O. Colard.
Biochem. Pharmacol. 53: 327-337, 1997). AA release was not specific, since oleic acid also increased in
the extracellular medium of cells treated with
H2O2
or AlF
4 as measured by gas
chromatography-mass spectrometry. In contrast, AA and oleic acid cell
content decreased after cell treatment. Oleoyl and arachidonoyl
acyl-CoA synthases and acyltransferases, assayed using a cell-free
system, were not significantly modified. In contrast, a good
correlation was observed between decreases in AA acylation and cell ATP
content. The decrease in ATP content is only partially accounted for by
mitochondrial damage as assayed by rhodamine 123 assay. We conclude
that oxidant-induced arachidonate release results from impairment of
fatty acid esterification and that ATP availability is probably
responsible for free AA accumulation on oxidative stress by preventing
its reesterification and/or transmembrane
transport.
hydrogen peroxide; aluminum fluoride; cell adenosine triphosphate content; A7r5 cells
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INTRODUCTION |
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ARACHIDONIC ACID (AA) is an important cellular mediator acting directly and after transformation to oxidized products. Metabolites of AA play important roles in regulating early growth-response genes (20) and vascular tone (22, 32) in vascular smooth muscle cells (VSMC). Recent studies have suggested additional roles for unmetabolized AA in cell signaling, such as stimulation of tyrosine-containing protein phosphorylation (5), inhibition of myosin light chain phosphatase (13), and recovery of cell growth after arrest by depletion of the Ca2+ pool (14). The cellular level of unesterified AA depends on the relative activities of enzymes that catalyze AA hydrolysis from lipids and enzymes that catalyze AA reesterification (10, 16). A major pathway for the deacylation of AA from the sn-2 position of phospholipids involves cytosolic phospholipase A2 (PLA2) activation (11). Acyl-CoA synthesis in the presence of ATP is necessary for free AA acylation by acyltransferase into phospholipids.
Oxidative damage is a major pathophysiological event in a broad range of inflammatory states, including the early stages of atherosclerosis (22). H2O2 and oxygen metabolites were shown to trigger AA release and metabolism in cultured cells (4, 8, 12, 25, 26), including smooth muscle cells (7, 21). Whereas cytosolic PLA2 activation has been observed after H2O2 treatment in some cell systems (7, 8, 21), the mechanism of AA release appears to be independent of Ca2+ in other cell systems (4, 12). Sporn et al. (25) showed that oxidative stress induced by H2O2 treatment of alveolar macrophages resulted in AA release by inhibition of AA esterification into phospholipids in association with depletion of ATP.
Recently, we characterized a
Ca2+-independent process of AA
release in
A7r5
VSMC (6). Indeed, VSMC triggering by the direct G protein activator
AlF4 induced a slow and linear
release of AA that was not accompanied by
Ca2+ mobilization or
Ca2+ entry into the cells. This AA
release did not involve any known phospholipase
A2, in contrast to the rapid and
Ca2+-dependent release of AA
induced by vasopressin that mobilizes Ca2+ and translocates cytosolic
PLA2. It was then of interest to
investigate whether the process of AA release from VSMC treated with
H2O2 was PLA2 dependent or independent.
We show that AlF
4 and
H2O2
induced an AA release that was inversely related to the acylation rate
of fatty acids and ATP content of VSMC.
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METHODS |
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Cell culture. Rat aortic smooth muscle cells (A7r5) were obtained from European Collection Animal Cell Cultures. The cells were grown at 37°C in DMEM supplemented with 10% (vol/vol) FCS, 2 mM glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin (DMEM-FCS) under a 5% CO2 atmosphere. A7r5 cells were subcultured every 7 days using trypsin-EDTA.
AA release.
Confluent cells in 35-mm dishes were labeled for 1 h with 0.4 µCi of
[3H]AA per dish in 1 ml of DMEM-FCS. Alternatively, cells were labeled overnight in the same
conditions. The medium was changed, and H2O2
(200 µM) or AlF4 (5 mM NaF + 10 µM AlCl3) was added for
various times. The supernatants were collected and counted for
radioactivity by liquid scintillation.
AA incorporation.
DMEM-FCS contained ~0.5 nmol/ml as measured by gas
chromatography-mass spectrometry (GC-MS). AA incorporation was assayed in FCS-containing medium (0.5 µM AA) or in FCS plus 1.5 nmol AA/ml (2.0 µM AA). [3H]AA
was present in the incubation medium such that AA specific activity was
0.8 µCi/nmol in both cases.
H2O2
or AlF4 was added together with
[3H]AA. Supernatants
were discarded, and cells were washed and scraped in 1:0.8 (vol/vol)
methanol-H2O. Lipids were
extracted by addition of chloroform and methanol (2), the organic phase
was evaporated under N2, and the
radioactivity was counted.
ATP assay. Cellular ATP was determined by the luciferase-luciferin assay. After cell incubation, culture media were removed and cells were washed twice with ice-cold PBS. They were then scraped into 1 ml of 10 mM KH2PO4-4 mM MgSO4 buffer, pH 7.4 (buffer A), and transferred to ice. Cell suspensions were placed in a 90-95°C water bath for 4 min and then on ice until assay. Twenty microliters of sample diluted in 1 ml of buffer A were added to 2 ml of 50 mM Na2HAsO4-20 mM MgSO4 buffer, pH 7.4 (buffer B). Fifty microliters of luciferase-luciferin reconstituted in sterile water (20 mg/ml) were added to the assay mixture, and chemiluminescence was measured over 10 s in a Lumat LB 9501 apparatus (Berthold).
Rhodamine 123 uptake. The changes in mitochondrial membrane potential were evaluated by measuring the cellular retention of rhodamine 123 (17). Cells were incubated for 30 min at 37°C with 5 µM rhodamine 123, then washed and allowed to stand at 37°C for 45 min in a rhodamine-free medium. After removal of the medium, cells were treated with agonists for 2 h. The dye trapped in treated and nontreated cells was determined by fluorometric analysis after lysis in 1% Triton. The excitation wavelength was 490 nm, and the emission wavelength was 515 nm.
Thiazolyl blue assay. Cellular reductive capacity was assayed by reduction of tetrazolium salt {3-[4,5-dimethylthiazol-2-yl]-2,3-diphenyltetrazolium bromide (MTT)} (24). Cells were incubated for 4 h at 37°C with MTT (50 µg/ml) and then washed and treated with agonists for 2 h. Control and treated cells were lysed in 5% SDS, and the absorbance of the MTT reduced form was read at 540 nm.
Neutral red retention.
Cell viability was evaluated by measuring the cellular retention of
neutral red (3). Controls and cells triggered with AlF4 or
H2O2
for 2 h were loaded for 3 h at 37°C with medium containing 0.05%
neutral red. After cell lysis in 5% SDS, the absorbance of neutral red
trapped in cells was read at 535 nm.
Acyl-CoA synthase assay.
Arachidonoyl-CoA synthase and oleoyl-CoA synthase were assayed as
described by Wilson et al. (31). Control and treated cells were
sonicated in 50 mM Tris · HCl, pH 8.0, containing
phenylmethylsulfonyl fluoride (20 µg/ml). The incubation medium
consisted of 20 mM MgCl2, 6.7 mM
ATP, 0.7 mM CoA, 1 mM -mercaptoethanol, 2 mM Triton X-100, 100 µM
fatty acid {a mixture of unlabeled AA or oleic acid (OA) and 30 nCi of [1-14C]AA (55 mCi/mmol; Amersham) or 30 nCi of
[9,10-3H]OA (10 Ci/mmol; Dupont-New England Nuclear)}, and homogenate in a total
volume of 150 µl of Tris buffer, pH 8.0. After incubation at 37°C
for 10 min, the reaction was terminated by addition of 2.25 ml of
40:10:1 (vol/vol/vol)
isopropanol-heptane-H2SO4
(2 M); 1.5 ml of heptane and 1 ml of water were then added, and the mixture was vortexed. The aqueous phase containing the radiolabeled acyl-CoA formed was washed twice with 2 ml of heptane, and
radioactivity was determined by scintillation counting.
Lysophosphatidyl acyltransferase assay. Arachidonoyl-CoA lysophosphatidylcholine (LPC) acyltransferase and oleoyl-CoA LPC acyltransferase were assayed. Control and treated cells were sonicated in 140 mM KCl and 20 mM HEPES, pH 7.4, containing 20 µg/ml phenylmethylsulfonyl fluoride. The reaction mixture contained 20 µM arachidonoyl-CoA or oleoyl-CoA, 32 µM LPC {a mixture of unlabeled LPC and 10 nCi of L-1-[palmitoyl-1-14C]LPC (56 mCi/mmol; Amersham)}, 0.2% BSA, 140 mM KCl, and homogenate (20-40 µg) in a total volume of 250 µl of HEPES buffer, pH 7.4. The incubation was performed at 37°C for 10 min, the reaction was stopped by addition of chloroform-methanol, and the lipids were extracted according to Bligh and Dyer (2). Extracted lipids were subjected to TLC on silica gel plates using 50:25:8:4 (vol/vol/vol/vol) chloroform-methanol-acetic acid-water. The LPC and phosphatidylcholine bands, visualized with I2 vapor, were scraped, and radioactivity was counted.
Fatty acid analysis by GC-MS.
Cells were incubated for 2 h in DMEM-FCS containing
AlF4 or
H2O2.
The free fatty acids, from supernatants and attached cells scraped in 1 ml of cold PBS, were extracted with 5 vol of 10:40:0.1
isopropanol-hexane-H2SO4
and methylated with diazomethane. They were then separated by GC on a
capillary column containing Supelcowax 10 bonded phase (0.32 mm
diameter, 30 m long) on a Hewlett-Packard 5890 series II gas
chromatograph. Fatty acids were detected by MS (model R10-10C, Nermag)
in the chemical ionization mode with ammonia (0.1 bar) as the reagent gas. The positive quasi-molecular ions were monitored and time integrated. Quantification was referred to heptadecanoic methyl ester
as an internal standard, and the response factors of the fatty methyl
esters were calibrated for each experiment.
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RESULTS |
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We first compared kinetic curves of AA release, induced by 200 µM
H2O2
or 10 µM AlF4, from VSMC
prelabeled with [3H]AA
for 60 min (Fig. 1). The
release of AA from cells exposed to 10 µM
AlF
4 for 2 h represented ~20%
of lipid labeling (6). The release of AA induced by 200 µM
H2O2
was in the same range. This noncytolytic concentration was then chosen to compare kinetic curves. Both compounds induced, after a 30-min latency period, a linear AA release up to 120 min representing 21.6 ± 2.8 and 16.8 ± 3.2% after treatment with
AlF
4 and
H2O2,
respectively. Linear release after the latency period was also observed
after overnight prelabeling. AA release after overnight prelabeling was
somewhat lower than after 1 h of labeling in control and stimulated
cells, such that the ratio of stimulated to control cells remained
constant (Table 1). This finding favors involvement of the same lipid pools in AA release observed in the two
prelabeling conditions. The kinetics of AA release after AlF
4 and
H2O2
treatment were then similar and completely different from
vasopressin-mediated AA release, which resulted in
Ca2+ mobilization and cytosolic
PLA2 translocation (6).
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If regular and long-lasting appearance of AA in the extracellular
medium does not depend on phospholipid hydrolysis, it can be the
consequence of decreased reacylation mechanisms. Therefore, we measured
the effect of both agonists on the rate of AA incorporation (Fig.
2). In control cells, incorporation of
[3H]AA into lipids was
nearly linear up to 120 min, where it represented 25% of initial
radioactivity. Addition of AlF4 or
H2O2
together with [3H]AA
resulted in a decrease in AA incorporation at 30-60 min followed by a complete inhibition of the incorporation.
H2O2
had slightly less effect than AlF
4
on AA incorporation as on AA release (Figs. 1 and 2). Triacylglycerols
and phospholipids were separated on TLC in two experiments. The AA
incorporation into triacylglycerols remained very low (2.6 ± 0.6%)
in control and treated cells. Thus the decrease in AA acylation of
treated cells relative to control cells was essentially due to
decreased AA incorporation into phospholipids.
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It remained a possibility that a decrease in labeled AA incorporation
might be observed if the treatments led to a decrease in AA specific
activity. Such would be the case if the treatments led to an increase
in fatty acid deacylation, thus enhancing AA concentration. Therefore,
we incorporated AA at two different concentrations. As shown in Fig.
3, the amount of AA incorporated into
lipids of control and stimulated cells was increased by 3.4 ± 0.2-fold in the presence of 2 µM AA compared with 0.5 µM in the
control medium. The decrease in AA incorporation when cells were
incubated with AlF4 or
H2O2
was in the same range whether or not 1.5 µM AA was added to
FCS-containing medium. This experiment demonstrates that an increase in
AA deacylation, because it enhanced AA concentration, would not
decrease AA incorporation and rules out the possibility that decreased
AA incorporation was due to increased AA deacylation.
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The accumulation of free fatty acids in the extracellular medium was
then measured by GC-MS (Table 2). In two
separate experiments, AA increased from 529 ± 41 pmol in
supernatants from control cells to 730 ± 106 and 681 ± 98 pmol
when 5 cells were treated with
AlF4 and
H2O2,
respectively. To investigate whether this increase was specific for AA,
OA content in the medium was also examined. The medium had a much
higher content of OA (4,079 ± 453 pmol/ml) than of AA. Cell
treatment also increased the amount of OA in the extracellular medium.
Surprisingly, the ratio of the amount of free fatty acids in cells and
medium (1:20) was in the same range for OA and AA. There was no
accumulation, rather there was a decrease, of the intracellular content
of either fatty acid after exposure to agonists.
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Fatty acid incorporation into phospholipids requires two enzymatic
activities: an acyl-CoA synthase and an acyl-CoA acyltransferase. If
one of these steps is altered, free fatty acids should accumulate in
the medium. We assayed the enzyme activities with OA and AA in
sonicated cells (Table 3). The oleoyl
transferase activity was considerably higher, ~10-fold, than the
arachidonoyl transferase activity, whereas oleoyl-CoA synthesis was
only twice as high as arachidonoyl-CoA synthesis. In our incubation
conditions, no significant change in these different enzyme activities
was observed after cells were treated with
AlF4 or
H2O2.
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It then appears that decreased AA incorporation is not accounted for by
inhibition of enzyme activities. Because ATP is necessary for acyl-CoA
synthase (18) and
H2O2
decreased ATP content in alveolar macrophages (27), we measured cell
ATP content. As shown in Fig. 4, the ATP
content declined by one-half in cells treated with
AlF4 for 2 h. Treatment with
H2O2 diminished the ATP content slightly less than treatment with
AlF
4. These decreases in ATP
content were well correlated with the inhibition of AA acylation (Fig.
5).
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Inasmuch as mitochondria represent a major source of ATP and were
proposed to be the primary target of
H2O2
action in neuronal cells (30), we investigated the effect of
AlF4 or
H2O2
on the integrity of mitochondria using rhodamine 123 assay (Table
4). Rhodamine 123 is selectively taken up
by mitochondria and then reflects maintenance of the mitochondrial
potential (
) (17). Exposure of cells to
AlF
4 or
H2O2
for 2 h slightly reduced the retention of rhodamine 123 by
mitochondria. We also used the MTT assay to evaluate the reductive
capacity of cells (24). The assay is based on the ability of the cell to produce formazan (reduced form of MTT). No inhibitory effect of
AlF
4 or
H2O2
was observed on this enzyme activity. Finally, neutral red was used to
evaluate the cytotoxic effect of treatments on membrane structures (3)
(Table 4). Control or treated cells were incubated for 2 h with the
dye, and the neutral red trapped in the cells was measured. Again, there was no significant difference between treated and nontreated cells. The two last assays demonstrated that treatment of cells with
H2O2
or AlF
4 for 2 h did not alter cell viability.
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DISCUSSION |
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H2O2
is a major oxygen metabolite produced by activated inflammatory cells,
including macrophages and neutrophils (23). These cells, together with
T lymphocytes, are present in the intima at the early stages of
atherosclerosis (22).
H2O2
produced by these inflammatory cells can thus be in contact with VSMC.
In this study we investigated the mechanisms involved in
H2O2-induced AA release from VSMC. Our data strongly suggest that oxidant-induced AA
release from VSMC does not depend on
PLA2 activation but results from
impairment of fatty acid reesterification. First, incorporation of
[3H]AA into cell
phospholipids was inhibited in an inverse way to AA release from
prelabeled cells on exposure to
H2O2.
Inhibition of AA esterification cannot be accounted for by a decrease
in AA specific activity due to increased deacylation, since inhibition rates were similar in the presence of 0.5 and 2.0 µM AA. Moreover, increasing AA concentration considerably enhances its incorporation. Second, inhibition of AA incorporation together with AA release was
also observed after treatment of cells with
AlF4, a process not involving
PLA2 activation (6). Third, AA
release was not specific, since OA also increased in the extracellular medium of cells treated with
H2O2
or AlF
4, as demonstrated by GC-MS.
These data are not consistent with the liberation of AA by an
arachidonate-specific PLA2 (11).
Oxygen species have been shown to trigger AA release in a number of cell types (4, 8, 12, 25), including VSMC (7, 21). A role for cytosolic PLA2 has often been suggested but not clearly demonstrated. Increased synthesis of cytosolic PLA2 is indeed involved in AA release after ultraviolet B injury of keratinocytes (8), among many other long-term effects. In growth-arrested VSMC, H2O2 induced very slight phosphorylation of cytosolic PLA2 and mitogen-activating protein kinase (21), which could not completely account for the large AA release observed. In endothelial cells, involvement of a PLA2 specific for arachidonate was deduced from the absence of [3H]OA release from prelabeled cells treated with H2O2 (4). However, we observed an accumulation of OA as well as AA in the medium of VSMC treated with H2O2.
On the other hand, Sporn et al. (25) demonstrated that, in alveolar
macrophages,
H2O2
increased the availability of AA by inhibiting its acylation into
phospholipids. Our data are in agreement with their results. AA
esterification is inhibited in our VSMC cultured in 10% FCS and
treated with
H2O2
or AlF4. Moreover, not only AA but
also OA accumulates in the medium. It is then likely that
esterification of all fatty acids is prevented. In addition, cell
content in free fatty acids decreased on oxidative stress, despite
their accumulation in the medium. This impairment of fatty acid
esterification could be a general process by which H2O2
resulted in free AA accumulation. Continual fatty acid remodeling between cell phospholipids (10) implicates basal
PLA2 activity together with
acyltransferase and transacylase activities. Part of the cytosolic
PLA2 appears to be constitutively
bound to membranes of resting cells, as observed in our VSMC line (6),
as in other cells (11), and could account for fatty acid remodeling and basal release. This basal release would be responsible for fatty acid
accumulation when fatty acid reesterification is prevented.
In a second set of experiments we investigated the mechanism of the
impairment of fatty acid esterification. We first assayed, in a
cell-free system, the two enzyme activities necessary for free fatty
acid incorporation into phospholipids: the acyl-CoA synthase, which
forms acyl-CoA complex from free fatty acid, CoA, and ATP, and the
acyl-CoA acyltransferase, which incorporates the fatty acyl moiety of
acyl-CoA into a lysophospholipid (18). When saturating concentrations
of the various substrates and cofactors were used, the oleoyl
transferase was considerably higher than the arachidonoyl transferase.
In contrast, arachidonoyl and oleoyl acyl-CoA synthase activities were
in the same range. Neither of these activities was decreased when cells
were pretreated with H2O2
or AlF4 for 2 h. Because no change
in these enzyme activities could explain the impairment of fatty acid
esterification and because ATP is involved in acyl-CoA synthesis, we
then measured cell ATP content.
H2O2
and AlF
4 treatment decreased cell
ATP content by 36% and 50%, respectively. These decreases correlated
well with the inhibition of AA esterification. Such a correlation
between accumulation of free AA and depletion of ATP has been observed
in cultured myocardial cells (9, 15) and in alveolar macrophages
triggered with
H2O2
(25). Muscle cells are known to contain a large amount of ATP, required
essentially for muscle contraction-relaxation. The ATP content of our
cell line, even after
H2O2
or AlF
4 treatment and a 50%
decrease in ATP, is indeed ~100-fold higher than the ATP content of
alveolar macrophages (27). We could wonder whether such a decrease in
ATP induced by agonists would be sufficient to prevent fatty acid
reesterification. However, despite the relatively high amount of ATP
remaining in cells after treatment with the agonists, a good
correlation was observed between ATP content and AA esterification.
Then it is likely that different ATP pools exist in muscle cells and
that the pool used for acyl-CoA synthesis is sufficiently decreased to
prevent acyl-CoA formation. Alternatively, membrane proteins that might
be involved in the transmembrane transport of fatty acids were recently
discovered (29), discrediting the hypothesis that fatty acids crossed
the membrane by simple diffusion. Because free fatty acid cell content
slightly decreased after
H2O2
or AlF
4 treatment, the
transmembrane transport of fatty acids appears somewhat altered by
oxidative stress and might also be involved in the decrease of
reesterification.
Because activities of mitochondria have been suggested in various
models of oxidative stress to represent a major target for prooxidant
molecules (28, 30) and mitochondria represent a source of ATP, we
investigated the action of AlF4 and
H2O2
on cell viability and integrity of mitochondria. Cell viability
evaluated by neutral red uptake or the MTT assay was not significantly
modified by 2 h of treatment with agonists, whereas mitochondrial
potential, as assayed by rhodamine 123, was slightly affected. However,
it is likely that these low reductions, which appear not to be
comparable to the marked decrease in ATP content, account only
partially for the decrease in ATP content.
AA esterification in pancreatic islets was impaired by an inhibitor of
ATP synthase (19). Changes in enzyme activities involved in ATP
synthesis might then account for the decrease in cell ATP content
observed after treatment with
H2O2
or AlF4. Moreover, Balsinde et al.
(1) found that cholera and pertussis toxins had no effect on AA
incorporation into phospholipids in mouse peritoneal macrophages,
implying that G proteins do not regulate acylation in these cells. It
is then likely that AlF
4 depletes
ATP and inhibits fatty acid acylation by a non-G protein-dependent mechanism.
In summary, our data demonstrate that oxidant-induced AA release from VSMC results from impairment of fatty acid reesterification and that this prevention in esterification may be due to a decrease in cell ATP content, which appears to be provoked only partially by damage to the mitochondria. The transmembrane transport of fatty acids might also be involved in the diminution of fatty acid esterification. The level of ATP, free AA, and other fatty acids, depending on oxidative stress, could then play an important role in atherosclerosis.
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ACKNOWLEDGEMENTS |
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This work was supported in part by Ministère de l'Enseignement et de la Recherche Grant ASV 9 and Institut National de la Santé et de la Recherche Médicale Grant CRE 930502. A. Cane is a recipient of a fellowship from the Ministère de l'Enseignement et de la Recherche.
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FOOTNOTES |
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Address for reprint requests: O. Colard, CNRS URA-1283, CHU Saint-Antoine, 27 Rue Chaligny, 75012 Paris, France.
Received 7 April 1997; accepted in final form 23 December 1997.
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