1 Department of Veterinary Basic Sciences, The Royal Veterinary College, University of London, London NW1 0TU; and 2 Centre for Cardiovascular Biology and Medicine, King's College London, London SE1 1UL, United Kingdom
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ABSTRACT |
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We have examined the mechanisms regulating
prostacyclin (PGI2) synthesis after acute exposure of human
umbilical vein endothelial cells (HUVEC) to interleukin-1 (IL-1
).
IL-1
evoked an early (30 min) release of PGI2 and
[3H]arachidonate that was blocked by the cytosolic
phospholipase A2
(cPLA2
) inhibitor
arachidonyl trifluoromethyl ketone. IL-1
-mediated activation
of extracellular signal-regulated kinase 1/2 (ERK1/2; p42/p44mapk) coincided temporally with phosphorylation of
cPLA2
and with the onset of PGI2
synthesis. The mitogen-activated protein kinase (MAPK) kinase (MEK)
inhibitors, PD-98059 and U-0126, blocked IL-1
-induced ERK
activation and partially attenuated cPLA2
phosphorylation and PGI2 release, suggesting that
ERK-dependent and -independent pathways regulate cPLA2
phosphorylation. SB-203580 treatment enhanced IL-1
-induced MEK,
p42/44mapk, and cPLA2
phosphorylation but
reduced thrombin-stimulated MEK and p42/44mapk activation.
IL-1
, but not thrombin, activated Raf-1 as assessed by
immune-complex kinase assay, as did SB-203580 alone. These results show
that IL-1
causes an acute upregulation of PGI2
generation in HUVEC, establish a role for the
MEK/ERK/cPLA2
pathway in this early release, and provide
evidence for an agonist-specific cross talk between p38mapk
and p42/44mapk that may reflect receptor-specific
differences in the signaling elements proximal to MAPK activation.
human endothelium; interleukin-1; thrombin; mitogen-activated
protein kinases; cytosolic phospholipase A2; prostacyclin; extracellular signal-regulated kinase
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INTRODUCTION |
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PROSTACYCLIN
(PGI2), a potent vasodilator and inhibitor of platelet
aggregation, is synthesized by vascular endothelial cells (ECs) in
response to a variety of stimuli. G protein-coupled receptor agonists
(e.g., thrombin) generally elicit a very rapid, short-lived PGI2 release (8, 44, 46). In contrast, early
studies in ECs revealed that PGI2 generation in response to
proinflammatory cytokines only became apparent after continuous
exposure to agonist for several hours (24, 31). Three key
enzymatic steps are involved in PGI2 generation, all of
which are potentially important regulatory sites. Phospholipase
A2 (PLA2) enzymes catalyze the cleavage of
arachidonic acid (AA) from the sn-2 position of
phospholipids (e.g., Ref. 21) with subsequent conversion
of free AA into prostaglandin (PG) H2 by two distinct
isoforms of cyclooxygenase (COX; Ref. 15); newly formed
PGH2 is then converted to PGI2 by terminal PGI2 synthase (26). The protracted
PGI2 generation in response to interleukin-1 (IL-1) has
been attributed to enhanced expression of COX-2, the inducible form of
COX, and to increased cytosolic PLA2
(cPLA2
) expression (11, 15, 24, 31). More
recently it was suggested that IL-1 may elicit an acute prostanoid
release from ECs (6), but the early signaling events
responsible for this synthesis remain undefined.
The post-receptor signaling pathways regulating
cytokine-driven prostanoid synthesis in vascular endothelium
are poorly understood. However, the pleiotropic effects of IL-1
are known to result in part from the triggering of signaling cascades
involving activation of families of serine/threonine protein
kinases collectively known as the mitogen-activated protein kinases
(MAPKs). MAPKs are activated by phosphorylation of specific threonine
and tyrosine residues within the signature sequence Thr-X-Tyr catalyzed
by upstream, dual-specificity MAPK kinases (MEKs) (reviewed in Ref.
41). Of the three major MAPK subgroups, the c-Jun
NH2-terminal kinases (JNK/stress-activated protein
kinases) and the p38 MAPKs (p38mapk) are thought to
be important in mediating cellular responses to extracellular stress.
Thus ultraviolet radiation, proinflammatory cytokines, and
lipopolysaccharide strongly activate these MAPK families
(37), and p38mapk has been implicated in the
regulation of cytokine-stimulated E-selectin expression in endothelium
(38). In contrast, the extracellular signal-regulated
kinases (ERK1/2; also known as p42/p44mapk) are typically
activated in response to mitogenic stimuli and are generally poorly
stimulated by exposure to proinflammatory cytokines (reviewed in Ref.
25). We have recently shown that IL-1 and tumor
necrosis factor-
(TNF-
) transiently activate p42/p44mapk in human ECs (28). Because this
activation does not appear to be obligatory for IL-1
-induced
E-selection expression (45), and because its involvement
in other IL-1
-mediated responses has yet to be explored, the
functional significance of activation of the ERK pathway by
proinflammatory cytokines in ECs is unclear.
The 85-kDa cPLA2 is one of a growing family of
PLA2 enzymes (35, 40, 42) implicated in
stimulus-evoked AA mobilization and is regulated by at least two major
posttranslational mechanisms, both of which are thought to be essential
for the initiation of AA release: 1) calcium-stimulated
membrane association via its calcium-binding domain (39),
and 2) phosphorylation of Ser505 within the
sequence Pro-Leu-Ser-Pro typically recognized by proline-directed protein kinases, such as members of the MAPK families.
p42mapk-mediated phosphorylation of Ser505
results in an increase in the specific activity of cPLA2
(13) and a characteristic shift in the electrophoretic
mobility of the phosphorylated, active form of the enzyme
(33). Evidence from studies carried out in platelets and
neutrophils also implicates the p38mapk pathway in
regulating the phosphorylation state of cPLA2
(14, 18, 43). Our recent studies have shown that p42mapk
is likely to be an important regulator of PGI2 generation
in thrombin- and vascular endothelial growth factor (VEGF)-stimulated human umbilical vein endothelial cells (HUVEC) via its effects on
cPLA2
phosphorylation (44, 45), but the
roles of the ERK and p38mapk pathways in controlling
cytokine-stimulated prostanoid production are unknown. In the present
study we establish that IL-1
acutely releases
PGI2 from HUVEC and examine the involvement of
cPLA2
and the p42/44mapk and
p38mapk pathways in this early synthesis. Our results
demonstrate the importance of early ERK-mediated activation of
cPLA2
as a regulator of acute cytokine-stimulated
PGI2 release and identify a novel, receptor-specific cross
talk between the p38mapk and ERK pathways that may regulate
the extent of prostanoid synthesis in agonist-stimulated human
endothelial cells.
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METHODS |
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Materials.
AA, human -thrombin, bovine serum albumin (BSA; fraction V),
pepstatin, aprotinin, leupeptin, sodium fluoride, sodium pyrophosphate, sodium orthovanadate, and sodium glycerophosphate were all purchased from Sigma (Poole, Dorset, UK). 4-(2-Aminoethyl)benzenesulfonyl fluoride (AEBSF), myelin basic protein, and protein G agarose were
obtained from Calbiochem (Nottingham, UK). Human recombinant IL-1
was from R&D Systems (Oxford, UK). PD-98059, SB-203580, and U-0126 were
generously provided by Dr. A. Saltiel (Parke Davis), Dr. J. Lee (Smith
Kline Beecham), and Dr. J. M. Trzaskos (DuPont Merck),
respectively. Anti-ACTIVE MAPK antibody recognizing the active, dually
phosphorylated forms of p42/p44mapk, was from Promega
(Southampton, UK); anti-p42/p44mapk antibody was from
Affiniti Research Products (Nottingham, UK). Phosphospecific
anti-p38mapk and anti-phospho-MEK1/2 antibodies were
purchased from New England Biolabs (Beverly, MA).
Anti-p38mapk antibody, protein A/G PLUS agarose, and
glutathione-S-transferase (GST)-activating transcription
factor (ATF)-21-96 were from Santa Cruz (Santa
Cruz, CA). The polyclonal anti-cPLA2
antibody was a kind
gift from Dr. R. Kramer (Eli Lilly, Indianapolis, IN). Horseradish
peroxidase-conjugated goat anti-mouse/rabbit immunoglobulins were from
Pierce and Warriner (Cheater, Cheshire, UK). Reagents for SDS-PAGE were
from Bio-Rad (Hemel Hempstead, Hertfordshire, UK) and National
Diagnostics (Hessle, Hull, UK). Polyvinylidene difluoride (PVDF)
membranes (Immobilon-P) and I-Block were from Sigma and Tropix
(Warrington, UK), respectively. Enhanced chemiluminescence (ECL)
Western blotting detection reagents, Hyperfilm-ECL, and
[5,6,8,9,11,12,14,15-3H]AA were all obtained from
Amersham International (Amersham, UK). 125I-labeled
6-ketoprostaglandin F1
(6-keto-PGF1
) was
purchased from Metachem Diagnostics (Piddington, Northampton, UK).
[
-32P]ATP and Renaissance 4CN Plus were from DuPont
(Dreieich, Germany). Raf-1 immunoprecipitation kinase cascade assay
kits were from Upstate Biotechnology (Lake Placid, NY). Culture media
were purchased from Sigma or Life Technologies (Paisley, UK). All other
reagents were obtained from Sigma or BDH (Poole, Dorset, UK) at the
equivalent of analytical reagent grade.
Cell culture. HUVEC were isolated by collagenase digestion using a modification of a procedure originally described by Jaffe et al. (16). Primary HUVEC cultures were grown in medium 199 (M199) supplemented with 20% (vol/vol) fetal calf serum, L-glutamine (5 mM), NaHCO3 (25 mM), penicillin (50 U/ml), and streptomycin (50 U/ml) and maintained at 37°C in 5% CO2-95% air. All tissue culture plastic was precoated with gelatin (1%) before the cells were plated. On reaching confluence (~1 × 106 cells/25-mm2 flask), primary cultures were trypsinized by brief exposure to a trypsin/EDTA solution (0.1%/0.025% in phosphate-buffered saline; PBS), plated onto 75-mm2 flasks, and cultured in M199 additionally supplemented with heparin (90 µg/ml) and endothelial cell growth supplement (ECGS; 20 µg/ml). ECGS was prepared from porcine brain as described by Maciag et al. (23). When confluent (~3.5 × 106 cells/75-mm2 flask), cells were plated onto either 24-well tissue culture trays (~1 × 105 cells/well) or 60-mm-diameter tissue culture dishes (~1 × 106 cells/dish). Confluent passage 2 cells were subsequently used for experimentation 4-5 days after plating; all experiments were routinely conducted at 37°C.
Measurement of PGI2 release.
Confluent cultures of HUVEC in 24-well tissue culture trays were washed
with serum-free M199 supplemented with HEPES (25 mM) and glutamine (5 mM) (H-M199, pH 7.4). Cell monolayers were treated as described in
legends to Figs. 1A and 4, and the PGI2
levels in the cell supernatants were quantified using a specific
radioimmunoassay for 6-keto-PGF1, the stable hydrolysis
product of PGI2, as previously described (46).
Supernatants from HUVEC monolayers used in immunoblotting studies were
also routinely assayed for 6-keto-PGF1
content.
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Measurement of [3H]AA release. Release of AA from endothelial cells was quantified as previously described (44). Briefly, confluent HUVEC monolayers in 24-well trays were incubated for 24 h with [5,6,8,9,11,12,14,15-3H]AA (1 µCi/ml) in serum-containing M199. After incubation, cells were washed twice in serum-free H-M199 and subsequently exposed to the same medium supplemented with 0.3% fatty acid-free BSA and other additions as indicated. At the end of the incubations the medium above the monolayers was collected and centrifuged for 5 min at 13,000 g, and the radioactivity in aliquots of supernatant was determined by beta-scintillation counting.
Immunoblotting procedures.
Immunoblotting studies were performed essentially as previously
described (45, 46). Briefly, confluent HUVEC in
60-mm-diameter dishes (~1 × 106 cells/dish) were
serum deprived for 16 h in serum- and ECGS-free M199 supplemented
with glutamine (5 mM). Those cultures that exhibited significant cell
detachment on serum starvation were discarded. Monolayers were
subsequently washed twice in H-M199 and challenged as detailed in
legends to Figs. 2, 3, and 5-8. Incubations were terminated by washing with ice-cold PBS containing
Na3VO4 (0.4 mM) and whole cell lysates prepared
in lysis buffer [63.5 mM Tris · HCl (pH 6.8), 10% glycerol,
2% SDS, 1 mM Na3VO4, 1 mM AEBSF, 50 µg/ml
leupeptin, 5% -mercaptoethanol, and 0.02% bromphenol blue]. The
protein content of cell lysates was measured using a bicinchoninic acid
(BCA) protein assay (Pierce and Warriner), and equal quantities of
protein (100 µg/lane) were resolved by SDS-PAGE (10%). Gels were
cast using a Protean II XI (20 cm) electrophoresis system (Bio-Rad).
Samples were subjected to prolonged electrophoresis overnight and then
transferred onto PVDF (Immobilon-P) membrane. Membranes were blocked
for 3 h in TBST [50 mM Tris, 150 mM NaCl, and 0.02% (vol/vol)
Tween 20, pH 7.4] containing 3% (wt/vol) BSA. For immunodetection of
cPLA2
, blots were blocked for 2 h in 0.2% I-Block.
Membranes were incubated overnight in TBST/0.2% BSA containing anti-p42/44mapk (1:25,000), anti-ACTIVE p42/p44
(1:20,000), anti-phospho-MEK1/2 (1:1,000), anti-cPLA2
(1:5,000), anti-p38mapk (1:1,000), or
phosphospecific anti-p38mapk (1:500) antibody. Blots were
then washed in TBST (8 × 15 min) and incubated with horseradish
peroxidase-conjugated goat anti-rabbit/mouse IgG as appropriate
(1:10,000) for 1 h. After further washing (8 × 15 min),
immunoreactive bands were visualized either colorimetrically with
Renaissance 4CN Plus or by enhanced chemiluminescence (ECL) according
to the manufacturer's instructions. In experiments employing phosphospecific antisera, equal loading was verified by reprobing with
antibody recognizing total protein. Enhanced phosphorylation/activation of either p42mapk or cPLA2
is reflected in a
decreased electrophoretic mobility such that the active, phosphorylated
forms of the enzymes are retarded in the gel. Where indicated,
densitometric analysis of cPLA2
or p42mapk
gel shifts was performed using a Bio-Rad Gel Doc 1000 system and
Molecular Analyst software.
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Immunoprecipitation and immune-complex kinase assay of
p38mapk.
Confluent, quiescent HUVEC in 60-mm dishes were washed and treated as
described for the immunoblotting studies. The medium was aspirated, and
incubations were terminated by washing in ice-cold PBS supplemented
with 0.4 mM Na3VO4; all subsequent procedures were carried out at 4°C. Monolayers were lysed in a buffer comprising 20 mM Tris · HCl, 137 mM NaCl, 2 mM EDTA, 25 mM
-glycerolphosphate, 2 mM sodium pyrophosphate, 10% glycerol, 1%
Triton X-100, 1 mM Na3VO4, 10 µg/ml
leupeptin, 1 mM phenylmethylsulfonyl fluoride (PMSF), and 40 µg/ml
aprotinin (pH 7.4) and allowed to stand on ice for 30 min with gentle
agitation. Whole cell lysates were centrifuged (4°C, 10 min, 13,000 g), and the supernatants were retrieved and transferred to
fresh 1.5-ml microcentrifuge tubes. Equal amounts of lysate protein
were subsequently incubated with anti-p38mapk antibody (10 µl/incubation) for 3 h at 4°C with constant rotation. Immune
complexes were captured with protein A/G plus agarose (4°C, overnight). The bead suspensions were pelleted, and immunoprecipitates were washed twice in lysis buffer and twice in kinase buffer (25 mM
HEPES, 25 mM
-glycerolphosphate, 25 mM MgCl2, 2 mM
dithiothreitol (DTT), 100 µM Na3VO4; pH 7.4).
Immune-complex kinase activity was assayed by incubation for 30 min at
30°C in kinase buffer (25-µl final volume) containing
[
-32P]ATP (5 µCi, 50 µM) and GST-ATF-2 (3 µg).
Reactions were terminated by the addition of 2× concentrated Laemmli
sample buffer. Proteins were resolved by SDS-PAGE (12%) and
transferred to PVDF membrane, and GST-ATF-2 phosphorylation was
assessed by autoradiography (
70°C).
Raf-1 immunoprecipitation and kinase activity assay.
Quiescent HUVEC monolayers in 60-mm dishes were washed and treated as
described in the legend to Table 1.
Incubations were terminated by washing in ice-cold PBS supplemented
with 0.4 mM Na3VO4; all subsequent procedures
were carried out at 4°C. Raf-1 activity was assessed using a Raf-1
immunoprecipitation kinase cascade assay kit, according to the
manufacturer's instructions. Briefly, 200 µl of buffer A
[50 mM Tris (pH 7.5), 1 mM EDTA, 1 mM EGTA, 0.5 mM
Na3VO4, 0.1% -mercaptoethanol, 1% Triton
X-100, 50 mM NaF, 5 mM Na pyrophosphate, 10 mM Na glycerophosphate, 0.1 mM PMSF, 1 µg/ml aprotinin, 1 µg/ml leupeptin, 1 µg/ml pepstatin, and 1 µM okadaic acid] was applied to the monolayers, which were then placed on ice for 1 h. Whole cell lysates were collected and
centrifuged (15 min, 16,000 g, 4°C). The resulting
supernatants were precleared with protein G agarose and incubated (2 h
with rotation; 4°C) with anti-Raf-1 antibody previously conjugated to
protein G agarose. The bead suspensions were pelleted, and immunoprecipitates were washed twice in buffer A and once in
assay dilution buffer (ADB: 20 mM MOPS, pH 7.2, 25 mM
-glycerol
phosphate, 5 mM EGTA, 1 mM Na3VO4, 1 mM DTT).
The protein G agarose/enzyme immune complexes were subsequently
incubated with inactive MEK1 and inactive ERK2 for 30 min at 30°C.
Samples were then briefly centrifuged to pellet the beads, and 4 µl
of the supernatant were removed and mixed with myelin basic protein
(final concn 0.6 mg/ml) and [
-32P]ATP (10 µCi/tube),
made up to a final volume of 34 µl with ADB. Samples were incubated
for 10 min at 30°C and then slowly spotted onto P81 phosphocellulose
squares. Membranes were washed in 0.75% phosphoric acid (3 × 5 min) and acetone (1 × 5 min) and transferred to scintillation
vials containing 5 ml of scintillation cocktail, and radioactivity was
quantified on a Beckman beta counter.
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Statistical analysis. Data were analyzed for statistically significant differences between experimental conditions using ordinary ANOVA (Bonferroni multiple comparison test). Data are expressed as means ± SE; P < 0.05 was considered statistically significant.
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RESULTS |
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IL-1 stimulates arachidonate release and PGI2
synthesis in HUVEC monolayers.
Exposure of confluent HUVEC monolayers to IL-1
at a maximally
effective concentration (100 U/ml) resulted in significant accumulation
of 6-keto-PGF1
after a 30-min incubation, which continued to increase for the duration of the experiment (Fig. 1A). In
seven independent experiments the mean increase in PGI2 synthesis evoked by a 30-min exposure to IL-1
was 197 ± 27 (% over basal; ± SE). Parallel studies showed that neither COX-2 mRNA nor protein levels were enhanced in cells exposed to IL-1
for
30 min (data not shown); thus these early changes in prostanoid synthesis occurred in the absence of enhanced COX-2 expression. To
examine the effects of IL-1
on AA mobilization, HUVEC monolayers were prelabeled with [3H]AA and exposed to IL-1
at
various concentrations for different time periods. Consistent with
PGI2 synthesis, early [3H]AA
mobilization (30 min) in IL-1
-stimulated HUVEC was
concentration dependent with maximal release achieved at 100 U/ml
IL-1
(Fig. 1B) and occurred in a time-dependent manner
(Fig. 1C). All subsequent experiments employed 100 U/ml IL-1
. TNF-
promoted similar early changes in AA release and
PGI2 synthesis in HUVEC (data not shown). In accordance
with our previous findings (44), thrombin at 1 U/ml
elicited both a greater and more rapid release of AA compared with a
maximal dose of IL-1
(15 min thrombin: 380 ± 48; 30 min thrombin 605 ± 82% increase over basal; mean ± SE from 3 independent experiments).
Effects of a cPLA2 inhibitor on AA release and
PGI2 synthesis.
Arachidonyl trifluoromethyl ketone (AACOCF3),
a trimethyl ketone analog of AA, has been shown to inhibit
cell-associated 85-kDa cPLA2
, as well as human
recombinant 85-kDa cPLA2
(1, 27). To
evaluate further the potential role of cPLA2
in
cytokine-driven PGI2 generation, we examined the effects of
AACOCF3 on [3H]AA release and
PGI2 synthesis. Pretreatment (30 min) with
AACOCF3 (10 µM) completely suppressed AA release in
response to a 30-min exposure to IL-1
(IL-1, 118 ± 4%; IL-1
plus AACOCF3, 103 ± 5% increase over
basal; mean ± SE, n = 3). AA release under
basal conditions was also partially, but not significantly, inhibited after AACOCF3 treatment. In parallel experiments
AACOCF3 (10 µM) also attenuated IL-1
-stimulated
PGI2 formation (IL-1, 182 ± 24%; IL-1 plus
AACOCF3, 100 ± 13% increase over basal; mean ± SE, n = 3) without significantly modifying basal
PGI2 synthesis (AACOCF3 alone, 105 ± 20%). These results demonstrate that IL-1
promotes early AA release
in a cPLA2
-dependent manner.
Phosphorylation of MEK, p42mapk, and
cPLA2 in HUVEC exposed to IL-1
.
We have previously reported that inhibition of MEK, the upstream
activator of p42mapk, attenuates thrombin- or VEGF-induced
PGI2 release and that this results from inhibition of
p42mapk-mediated cPLA2
phosphorylation
(44, 45). To determine whether these mechanisms contribute
to the acute release of PGI2 from IL-1
-stimulated HUVEC,
we examined the effects of IL-1
on activation of MEK and
p42mapk and on the phosphorylation state of
cPLA2
. Phosphorylated MEK was detected in whole cell
HUVEC lysates under basal conditions and increased in a time-dependent
manner in response to IL-1
with maximal phosphorylation observed
after 20-30 min (Fig.
2A). Consistent with this time
course, and confirming our recent observations (28, 45),
IL-1
promoted a pronounced, time-dependent increase in the upper,
electrophoretically slower band of p42mapk and a
concomitant decrease in the lower band (Fig. 2B). Increased phosphorylation was detected by 15-20 min, reached a maximum at 30 min, and decreased after 60 min of incubation with IL-1
. As previously described (44), phosphorylated and
nonphosphorylated forms of cPLA2
are present in
nonstimulated HUVEC (Fig. 2C). Enhancement of
cPLA2
phosphorylation by IL-1
followed kinetics similar to those for p42mapk, with maximally elevated
phospho-cPLA2
evident at 20-30 min (Fig.
2C); in contrast, cPLA2
phosphorylation was
maintained for up to 2 h after IL-1
stimulation (Fig.
2C and data not shown). Thus enhanced activation of MEK,
p42mapk, and cPLA2
coincides temporally with
the onset of AA mobilization and PGI2 synthesis in
IL-1
-stimulated HUVEC.
Inhibition of MEK modulates IL-1-induced p42mapk
activation, cPLA2
phosphorylation, and PGI2
synthesis.
The close correlation between IL-1
-induced
p42mapk/cPLA2
phosphorylation and
PGI2 generation prompted us to investigate further the
potential role of the MEK/ERK/cPLA2
pathway in acute,
cytokine-driven PGI2 formation. Prior treatment of HUVEC
(30 min) with the cell-permeant MEK inhibitors PD-98059
(9) or U-0126 (10) dose-dependently attenuated IL-1
-induced ERK activation, with complete inhibition observed at 5 or 1 µM, respectively (Fig.
3A). Treatment with IL-1
caused a nearly complete gel shift of cPLA2
that was
partially inhibited by PD-98059 concentrations of 5 µM and above and
by 0.1-3 µM U-0126 (Fig. 3B). These findings were
confirmed by densitometric analysis of the immunoblots, with a maximal
inhibition of ~50% with either inhibitor (Fig. 3C). In
accordance with this partial inhibitory effect, IL-1
-stimulated
[3H]AA release was inhibited by 12 ± 2% (mean ± SE; n = 3 independent experiments) after treatment
with 5 µM PD-98059. PD-98059 also caused a dose-dependent reduction
of basal and IL-1
-induced PGI2 release (Fig.
4A). Interestingly,
PGI2 synthesis was totally blocked at 1 µM PD-98059, a
concentration that partially inhibited IL-1
-stimulated p42mapk activation (Fig. 3A) and did not
detectably reduce cPLA2
phosphorylation (Fig.
3B). In contrast, U-0126, at a concentration (1 µM) that abolished IL-1
-stimulated ERK activation and maximally affected cPLA2
phosphorylation (Fig. 3), reduced IL-1
-evoked
PGI2 synthesis by only 59 ± 5% (mean ± SE,
n = 3 independent experiments). These results show that
ERK-dependent and -independent mechanisms regulated cPLA2
phosphorylation in response to IL-1
and that
inhibition of cPLA2
activation correlates with
PGI2 synthesis.
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Activation of p38mapk by IL-1 and thrombin.
To address the potential role of p38mapk in cytokine-driven
prostanoid synthesis, we initially measured p38mapk
activation by immune-complex kinase assay. Treatment with IL-1
or
thrombin enhanced p38mapk activity, which was partially
attenuated by the p38mapk inhibitor SB-203580 (Fig.
5C). The incomplete inhibition
of p38mapk reflects the rapid reversibility of
SB-203580:p38mapk interaction during immunoprecipitation
(43). SB-203580 also dose-dependently suppressed
agonist-stimulated p38mapk activation when added directly
to immunoprecipitates (data not shown). IL-1
and thrombin enhanced
phosphorylation of p38mapk (Fig. 5, A and
B), and this was unaffected by SB-203580 treatment (data not
shown). The kinetics of the responses differed, with transient or
sustained activation observed in response to IL-1
or thrombin,
respectively. In addition, IL-1
-stimulated p38mapk
activation (5 min) occurred more rapidly than p42mapk
phosphorylation (20-30 min; Fig. 2), whereas p42mapk
activation by thrombin (1 min; Refs. 44 and 45) was
earlier than p38mapk activation (Fig. 5B).
Neither PD-98059 nor U-0126 affected the phosphorylation state or
activity of p38mapk as assessed by in-gel kinase assay
(data not shown). Thus both IL-1
and thrombin activate
p38mapk but with different kinetics.
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Effects of SB-203580 on agonist-stimulated PGI2
synthesis, p42mapk activation, and
cPLA2 phosphorylation.
In contrast to the inhibitory effects of MEK inhibition on
IL-1
-induced p42mapk activation and cPLA2
phosphorylation, blockade of p38mapk enhanced the
phosphorylation of p42mapk (
1 µM SB-203580; Fig.
6B) and cPLA2
(Fig. 6C). Consistent with these effects SB-203580 strongly
potentiated IL-1
-stimulated MEK phosphorylation (Fig.
6A). However, despite potentiation of the
MEK/ERK/cPLA2
pathway, SB-203580 markedly
inhibited basal and IL-1
(100 U/ml)-stimulated PGI2
release (Fig. 4B). In contrast to our findings in
IL-1-stimulated HUVEC, SB-203580 dose-dependently reduced activation of
MEK, p42mapk, and cPLA2
in response to
thrombin (Fig. 7).
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Effects of IL-1 and thrombin on Raf-1 activation.
To determine whether the differential effects of SB-203580 on IL-1
-
vs. thrombin-mediated ERK activation reflect differences in the
signaling components upstream of ERK, we measured Raf-1 activation in
immunoprecipitates from thrombin- and IL-1-stimulated cells (Table 1).
At the time points examined, IL-1
, but not thrombin, significantly
elevated Raf-1 activity, suggesting that thrombin uses alternative
pathway(s) to activate MEK. Because Raf-1 activity was enhanced in the
presence of SB-203580 alone, SB-203580 treatment resulted in strong
potentiation of Raf-1 activity in both IL-1
- and thrombin-stimulated
HUVEC. Thus the opposing effects of p38mapk blockade on
IL-1
- (Fig. 6) vs. thrombin-induced p42/44mapk
activation (Fig. 7) cannot be explained by differential effects of the
inhibitor on Raf-1 activation.
MEK inhibitors block COX activity in HUVEC.
To examine whether inhibition of IL-1-stimulated PGI2
release by SB-203580 (despite enhancement of
MEK/ERK/cPLA2
phosphorylation) reflects an independent
action of the p38mapk inhibitor on COX activity, we
measured PGI2 synthesis after application of exogenous AA
(Table 2). SB-203580 attenuated
AA-induced PGI2 formation. Parallel studies showed
that PD-98059 also inhibited AA-stimulated PGI2
synthesis, whereas U-0126 (1 µM) had no significant effect (Table 2),
confirming that SB-203580 and PD-98059 block COX activity at
concentrations that maximally effect ERK/p38mapk
activation, whereas U-0126 does not. Because indomethacin did not
modulate IL-1
-induced p42/44mapk activation or
cPLA2
phosphorylation (Fig.
8), the effects of the
MEK/p38mapk inhibitors on IL-1
-stimulated
ERK/cPLA2
phosphorylation did not result from their
ability to block COX activity.
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The cytosolic, AA-selective 85-kDa cPLA2 is
important in mediating agonist-driven AA mobilization in several cell
types (21). This release initiates the biosynthesis of
PGs, leukotrienes, and platelet-activating factor, all of which are
implicated in the modulation of diverse physiological and
pathophysiological processes, including vascular homeostasis and
inflammation. The proinflammatory cytokines IL-1 and TNF-
have been
shown to promote PGI2 generation by ECs, but their effects
are generally delayed in onset and require de novo protein synthesis
(5, 15). In the present study we have focused on the
post-receptor transduction pathways regulating early changes in
cytokine-driven prostanoid synthesis and have demonstrated that IL-1
stimulates acute PGI2 synthesis in part via a
MEK/p42mapk/cPLA2
pathway. Thus
1) MEK, p42mapk, and cPLA2
were
phosphorylated by IL-1
with kinetics coinciding with the onset of AA
mobilization and PGI2 synthesis, 2)
AACOCF3, at concentrations previously reported to modify
stimulus-induced AA generation, markedly suppressed IL-1
-driven AA
release and PGI2 synthesis, consistent with a key role for
the 85-kDa cPLA2
, and 3) inhibition of MEK
attenuated cytokine-stimulated p42mapk activation,
cPLA2
phosphorylation, and PGI2 formation.
We (44, 45) and others (7) have
previously shown that rapid PGI2 synthesis in response
to Ca2+-mobilizing agonists that promote
cPLA2
translocation also requires p42mapk-mediated cPLA2
phosphorylation, and
several studies have suggested that phosphorylated cPLA2
fails to mobilize AA in the absence of an increase in intracellular
Ca2+ concentration ([Ca2+]i)
(e.g., Ref. 48). Because acute exposure to IL-1
neither raises [Ca2+]i nor causes translocation of
cPLA2
(Ref. 11; Houliston and Wheeler-Jones, unpublished observations), these studies provide evidence that enhanced cPLA2
phosphorylation alone may
be sufficient to promote AA mobilization in IL-1
-stimulated HUVEC.
It is possible that cPLA2
activation by IL-1
may
result from phosphorylation of discrete pools of the enzyme that are
already localized within the membrane at resting
[Ca2+]i (34). Although there are
likely to be several alternative cell-specific mechanisms for
cPLA2
regulation (12, 30, 32, 38, 47), our
findings are in keeping with recent studies suggesting that
cPLA2
phosphorylation becomes most important for
activation when the rise in [Ca2+]i is
insufficient to fully translocate the enzyme (14). The contribution, if any, of the novel, Ca2+-independent
cPLA2
and -
isoforms to IL-1
-stimulated AA release in HUVEC remains to be defined (35, 40, 42).
Our findings implicating ERK activation in IL-1-induced
AA/PGI2 release are based on pharmacological inhibition
with PD-98059 or U-0126, two structurally and mechanistically distinct
inhibitors of MEK. Under conditions where p42/44mapk
activation by IL-1
was abolished, IL-1
-evoked
cPLA2
phosphorylation was only partially inhibited by
either of these agents, indicating that MEK-dependent and
MEK-independent pathway(s) both regulate early changes in
cPLA2
phosphorylation in response to cytokine. The
partial reductions in IL-1
-stimulated AA mobilization and PGI2 synthesis after MEK inhibition are also consistent
with the effects on cPLA2
phosphorylation.
In contrast to our present findings in human ECs,
p42/p44mapk does not mediate phosphorylation of
cPLA2 in thrombin- or collagen-stimulated platelets
(3), and current evidence suggests that
p38mapk plays a central role in these cells
(2, 14, 43). Our results suggest that p38mapk
activation is unlikely to account for the MEK-independent
phosphorylation since SB-203580, a pyridinyl imidazole inhibitor of
p38mapk activity (49), did not inhibit
IL-1
-induced cPLA2
phosphorylation in HUVEC. It is
possible that p38mapk directly or indirectly
(14) phosphorylates cPLA2
on
Ser727, which does not alter the mobility of
cPLA2
on SDS-PAGE (36), or alternatively,
that an SB-203580-insensitive p38mapk isoform contributes
to Ser505 phosphorylation in IL-1
-stimulated endothelium
(19).
Our results demonstrated that SB-203580 and PD-98059, but not U-0126,
reduced PGI2 generation in response to exogenous AA, suggesting additional inhibitory effects on COX (4, 18). Inhibition of COX activity at low concentrations may explain why PD-98059 had a more potent effect on PGI2 synthesis
compared with its effects on p42mapk/cPLA2
activation. However, inhibition of COX activity with indomethacin did
not mimic the effects of either SB-203580 or PD-98059 on cytokine- or
thrombin-stimulated p42mapk/cPLA2
activation, suggesting that the effects of these inhibitors did not
result from COX inhibition and are mediated by blockade of
p38mapk and ERK, respectively. More importantly, U-0126, an
inhibitor of MEK with no capacity to block COX in HUVEC, mimicked the
partial inhibitory effects of PD-98059 on ERK/cPLA2
phosphorylation and also partially inhibited PGI2
synthesis. These results confirm that PGI2 generation
is regulated in part via a MEK/ERK/cPLA2
-dependent pathway.
The present studies also confirmed that SB-203580 treatment inhibited
agonist-stimulated p38mapk activity. As discussed above,
blocking p38mapk activity had no inhibitory effect on
IL-1-stimulated cPLA2
phosphorylation but, in
contrast, potentiated cytokine-driven cPLA2
phosphorylation and p42mapk activation. These results imply
that early triggering of the p38mapk pathway by IL-1
limits the extent of subsequent activation of p42/p44mapk
and thus of the downstream events regulated by these kinases. The
earlier (5 min) activation of p38mapk in
cytokine-stimulated HUVEC compared with that of p42/44mapk
(20-30 min; Ref. 28) is also consistent with this
hypothesis. The modulatory effects of p38mapk on
p42/44mapk activation are unlikely to reflect direct
association of p38 with ERK (51) because 1) MEK
activation in SB-203580-treated cells is modulated in parallel with
ERK, and 2) immunoprecipitation of p38mapk does
not coprecipitate p42/p44mapk (Houliston and Wheeler-Jones,
unpublished observations); communication between the p38 and
p42/p44mapk pathways is therefore likely to occur at the
level of upstream regulatory kinases. Furthermore, because
IL-1
-induced p38mapk activation was not affected by
inhibiting the MEK/ERK pathway, the interaction between ERK and
p38mapk pathways occurs in a unidirectional manner. In
marked contrast to IL-1
, blocking p38mapk activation had
a differential effect on thrombin-mediated signaling, causing
inhibition of MEK, p42/44mapk, and cPLA2
phosphorylation. These data suggest that p38mapk activation
positively regulates the ERK pathway in thrombin-stimulated cells and
may therefore contribute to both the early (45) and sustained phases of p42/44mapk activation observed in
response to thrombin (Houliston and Wheeler-Jones, unpublished
observations). Because MEK1/2 and p42/44mapk were modified
in parallel by SB-203580 treatment, it is likely that the level of
regulation by p38mapk is proximal to MEK1/2 activation.
However, measurement of Raf-1 activation by immune-complex kinase assay
showed that SB-203580 in the absence of agonist strongly activated
Raf-1 (17). In addition, IL-1
but not thrombin elevated
Raf-1 activity in HUVEC, demonstrating that alternative
Raf-1-independent pathways leading to MEK activation are recruited in
thrombin-stimulated cells. The ability of SB-203580 alone to activate
Raf-1 caused potentiation of activity in cells subsequently exposed to
either IL-1
or thrombin, suggesting that the different effects of
SB-203580 on signaling by these two agonists cannot be explained by
effects at the level of Raf-1. Different types of cross talk between
MAPK pathways are emerging, and the functional outcomes of such
interactions are likely to be cell-type specific (20, 22, 29,
50). Our results suggest that differences in the onset and
duration of p38mapk vs. p42/44mapk activation
may determine how these pathways are coordinated to produce the
physiological prostanoid responses of IL-1
- or thrombin-challenged ECs. A putative signaling pathway that could account for our
observations is shown in Fig. 9.
|
In summary, we have shown that IL-1 stimulates early
cPLA2
phosphorylation, AA mobilization, and
PGI2 synthesis in HUVEC through activation of MEK-dependent
and -independent pathways. SB-203580-sensitive p38mapk(s)
negatively regulate ERK signaling after IL-1
treatment but
facilitate thrombin signaling via the MEK/ERK pathway. These effects
may reflect differential usage by IL-1
and thrombin of signaling elements upstream of p42/44mapk activation. These findings
underscore the importance of MAPK cascades in regulating acute events
in human endothelium.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. A. Saltiel, Dr. J. Lee, Dr. J. M. Trzaskos, and
Dr. R. Kramer for gifts of PD-98059, SB-203580, U-0126, and
anti-cPLA2 antibody, respectively. We also thank the
midwives and delivery staff at St. Mary's and the Portland Hospitals,
London, UK, for assistance in obtaining umbilical cords.
![]() |
FOOTNOTES |
---|
This work was supported by the British Heart Foundation.
Address for reprint requests and other correspondence: C. P. D. Wheeler-Jones, Dept. of Veterinary Basic Sciences, The Royal Veterinary College, Univ. of London, Royal College St., London NW1 0TU, UK (E-mail: cwheeler{at}rvc.ac.uk).
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.
Received 22 February 2001; accepted in final form 12 June 2001.
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