1 Division of Pulmonary and Critical Care Medicine, Department of Medicine, Johns Hopkins University School of Medicine, Baltimore, Maryland 21224; and 2 Department of Pediatrics, University of Chicago School of Medicine, Chicago, Illinois 60637
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ABSTRACT |
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The treatment of endothelial cell monolayers with phorbol 12-myristate 13-acetate (PMA), a direct protein kinase C (PKC) activator, leads to disruption of endothelial cell monolayer integrity and intercellular gap formation. Selective inhibition of PKC (with bisindolylmaleimide) and extracellular signal-regulated kinases (ERKs; with PD-98059, olomoucine, or ERK antisense oligonucleotides) significantly attenuated PMA-induced reductions in transmonolayer electrical resistance consistent with PKC- and ERK-mediated endothelial cell barrier regulation. An inhibitor of the dual-specificity ERK kinase (MEK), PD-98059, completely abolished PMA-induced ERK activation. PMA also produced significant time-dependent increases in the activity of Raf-1, a Ser/Thr kinase known to activate MEK (~6-fold increase over basal level). Similarly, PMA increased the activity of Ras, which binds and activates Raf-1 (~80% increase over basal level). The Ras inhibitor farnesyltransferase inhibitor III (100 µM for 3 h) completely abolished PMA-induced Raf-1 activation. Taken together, these data suggest that the sequential activation of Ras, Raf-1, and MEK are involved in PKC-dependent endothelial cell barrier regulation.
mitogen-activated protein kinases; extracellular signal-regulated kinase; cytoskeleton
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INTRODUCTION |
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THE CONFLUENT ENDOTHELIUM serves as a selective barrier between the vascular space of blood vessels and underlying tissues. Compromise of endothelial cell barrier integrity leads to an increase in vascular permeability, which is a cardinal feature of acute inflammatory lung injury. Garcia and colleagues (13, 14, 16) and others (22, 35, 48, 60, 61) have previously shown barrier integrity to be critically dependent on the cytoskeleton that regulates actin stress fiber formation, cell shape, and cellular adherence, with involvement of actomyosin-driven contraction in specific models of permeability. For example, thrombin-induced increases in endothelial cell permeability and elevation of isometric tension are dependent on myosin light chain (MLC) kinase (MLCK)-catalyzed phosphorylation of MLCs (13, 21). However, other models of endothelial cell barrier dysfunction that occur independently of MLCK-mediated MLC phosphorylation exist. For example, phorbol 12-myristate 13-acetate (PMA), a direct protein kinase C (PKC) activator, significantly increased endothelial cell permeability without a rise in intracellular Ca2+ and MLCK activation (13, 43). In smooth muscle, PMA produces a slowly developed but sustained increase in tension, without a further increase in MLC phosphorylation (47, 53, 57), suggesting an important role for additional proteins in the regulation of the contractile apparatus. In both smooth muscle and endothelial cells, the actin-, myosin-, and Ca2+/calmodulin (CaM)-regulatory protein caldesmon can potentially regulate actomyosin interactions in the absence of MLCK activation (2, 40, 51, 52). In vitro studies indicate that in the absence of MLC phosphorylation and in the presence of a low intracellular Ca2+ concentration, caldesmon binding to actin filaments inhibits myosin ATPase activity and actin-myosin binding that can be reversed by either increased Ca2+/CaM availability or caldesmon phosphorylation (2, 18, 20, 34, 40, 44, 51).
Extracellular signal-regulated kinases (ERK1 and ERK2, also referred to as p44 and p42 kinases) are responsible for phorbol ester-stimulated phosphorylation of smooth muscle caldesmon (1-3). The ERK members of the Ser/Thr mitogen-activated protein (MAP) kinase (MAPK) family require Tyr and Thr phosphorylation for maximal activation (for a review, see Ref. 23). MAPKs are often referred to as "proline directed" because the consensus sequence for their substrate recognition includes proline residues. The diverse extracellular stimuli known to activate ERKs (including growth factors, thrombin, and phorbol esters) suggest that multiple signaling pathways are involved in their regulation (23). The major pathway involved in ERK activation appears to require the sequential activation of Ras, Raf-1, and MEK. Activation of this signaling pathway leads to cytoskeletal changes and correlates with Ca2+-independent contraction in smooth muscle (12, 19). However, the precise roles of ERKs and other signaling intermediates in agonist-stimulated cytoskeletal rearrangement of the endothelium remain unclear. In this study, we examine the role of the MAPK signaling cascade in phorbol ester-induced barrier dysfunction in bovine endothelium.
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METHODS |
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Reagents.
Endothelial cell cultures were maintained in medium 199 (GIBCO BRL,
Life Technologies, Chagrin Falls, OH) supplemented with 20% (vol/vol)
colostrum-free bovine serum (Irvine Scientific, Santa Ana, CA), 15 µg/ml of endothelial cell growth supplement (Collaborative Research,
Bedford, MA), 1% antibiotic-antimycotic (10,000 U/ml of penicillin, 10 mg/ml of streptomycin, and 25 µg/ml of amphotericin B; K. C. Biologicals, Lenexa, KS), and 0.1 mM nonessential amino acids (GIBCO
BRL). Unless specified, reagents were obtained from Sigma (St. Louis,
MO). PBS, Hanks' balanced salt solution without phenol red, and
LIPOFECTAMINE were purchased from GIBCO BRL (Life Technologies, Grand
Island, NY). Diperoxovanadate (DPV) was kindly provided by Dr. V. Natarajan (Johns Hopkins University, Baltimore, MD).
Bisindolylmaleimide I hydrochloride (BIM), Ro-31-8220, olomoucine,
PD-98059, farnesyltransferase (FPT) inhibitor III, and C3 exotoxin were
purchased from Calbiochem (La Jolla, CA). Anti-pan ERK monoclonal
antibody and anti-PKC- antibodies were purchased from Transduction
Laboratories (Lexington, KY), and anti-rabbit polyclonal antibody
against phospho-ERK was purchased from New England Biolabs (Beverly,
MA). Myelin basic protein (MBP) was purchased from Upstate
Biotechnology (Lake Placid, NY). Hemagglutinin (HA)-tagged ERK2 and MEK
constructs were kindly provided by Drs. R. Pestell (Albert Einstein
College of Medicine, Bronx, NY) and M. Rosner (University of
Chicago, Chicago, IL), respectively.
Bovine pulmonary artery endothelial cell culture. Bovine pulmonary artery endothelial cells (BPAECs) were obtained frozen at 16 passages from American Type Culture Collection (Manassas, VA) and were utilized at passages 19-24 (13). BPAECs were cultured in complete medium and maintained at 37°C in a humidified atmosphere of 5% CO2-95% air. The endothelial cells grew to contact-inhibited monolayers with the typical cobblestone morphology. Cells from each primary flask were detached with 0.05% trypsin, resuspended in fresh culture medium, and passaged to appropriate size flasks or dishes.
Endothelial cell monolayer electrical resistance determinations.
Electrical resistance of endothelial monolayers was measured with an
electrical cell impedance sensor technique and system (Applied
Biophysics, Troy, NY) as Garcia et al. (15) and Shi et al.
(49) have previously described. The cells were cultured on
a small gold electrode (104 cm2) in DMEM
(GIBCO BRL) supplemented with 20% (vol/vol) colostrum-free bovine
serum, antibiotics, and growth factors. Before the experiment, medium
containing serum was replaced with the same medium without serum, which
functioned as the electrolyte with the cells acting as insulating
particles. The total resistance across the monolayers was composed of
the resistance generated between the ventral cell surface and the
electrode as well as by the resistance between cells. A 4,000-Hz AC
signal with a 1-V amplitude was applied to the endothelial cell
monolayer through a 1-M
resistor, creating an approximate
constant-current source (1 µA). The lock-in amplifier attached to the
electrodes detected changes in both the magnitude and phase of the
voltage appearing across the endothelial cells and was controlled by an
IBM-compatible personal computer that was used to both run the
experiments and process the data. Electrical resistance increased
immediately after cell attachment and achieved a steady state on
confluence. Resistance data were normalized to the initial voltage and
plotted as normalized resistance.
MLC phosphorylation assay. MLC phosphorylation assay was performed as Garcia et al. (13) and Shi et al. (49) have previously described in detail.
Detergent fractionation of endothelial lysates.
After PMA treatment (100 nM for 10 min), BPAECs were fractionated into
cytosolic, membrane, and nuclear or cytoskeletal fractions as
previously described (45), with some modifications. The
cells were rinsed with PBS and incubated in ice-cold cytosolic buffer [0.01% digitonin, 10 mM PIPES, pH 6.8, 300 mM sucrose, 100 mM NaCl, 3 mM MgCl2, 5 mM EDTA, 5 µM phallicidin, and protease
inhibitory cocktail (1:500 dilution; Calbiochem)] with agitation for
10 min at 4°C. The soluble (cytosolic) fraction was collected, the
dishes were rinsed with cytosolic buffer without protease inhibitors, and the residual material was extracted with membrane buffer (0.5% Triton X-100, 10 mM PIPES, pH 7.4, 300 mM sucrose, 100 mM NaCl, 3 mM
MgCl2, 3 mM EDTA, 5 µM phallicidin, and protease
inhibitory cocktail) with agitation for 20 min at 4°C. The soluble
(membrane) fraction was collected, and the protein material remaining
on the dishes was scraped into SDS buffer (0.5% Triton X-100, 0.5% SDS, 10 mM Tris · HCl, pH 6.8, and protease inhibitory
cocktail), sonicated, heated at 100°C for 5 min, and centrifuged, and
the supernatant (cytoskeletal fraction) was collected. Each fraction was used for SDS-PAGE and subsequent Western blotting analysis with
PKC--specific antibodies (Transduction Laboratories).
Endothelial cell permeabilization. Because the Rho inhibitor toxin generated by Clostridium botulinum, C3 exoenzyme, does not readily pass through the cell membrane under native conditions, we developed a method based on the detergent LIPOFECTAMINE for cell permeabilization. Briefly, BPAEC monolayers (80-100% confluence) grown on 60-mm culture dishes were rinsed with OPTI-MEM I medium, and LIPOFECTAMINE reagent (GIBCO BRL) was added at final concentration of 20 µg/ml. The cells were incubated for 1 h, and C3 exoenzyme (2.5 µg/ml) was added and remained present for 11 h.
Western immunoblotting. Proteins were extracted from BPAEC preparations with SDS sample buffer as previously described (32). The extracts were separated by SDS-PAGE, transferred to nitrocellulose (30 V for 18 h or 90 V for 2 h), and reacted with the antibodies of interest. Immunoreactive proteins were detected with an enhanced chemiluminescence (ECL) detection system according to the manufacturer's directions (Amersham, Little Chalfont, UK). The relative intensities of the protein in the bands were quantified by scanning densitometry.
Raf-1 activity assay.
Raf-1 kinase activity was assessed by using a commercially available
kit (c-Raf immunoprecipitation kinase cascade assay kit, Upstate
Biotechnology, Lake Placid, NY) according to the manufacturer's recommendation, with minor modification. Confluent BPAECs grown in
60-mm dishes were treated with 100 nM PMA or 0.1% DMSO as a vehicle
control for different time periods after 18 h of serum starvation
in medium 199. The cells were lysed in 500 µl of lysis buffer
A (50 mM Tris, pH 7.5, 1 mM EDTA, 1 mM EGTA, 0.5 mM
Na3VO4, 50 mM NaF, 5 mM sodium pyrophosphate,
10 mM sodium glycerophosphate, 0.1% -mercaptoethanol, and 0.1%
Triton X-100) including a 1:500 dilution of a protease inhibitory
cocktail [200 µM 4-(2-aminoethyl)benzenesulfonyl fluoride, 160 nM aprotinin, 10 µM bestatin, 3 µM E-64, 4 µM leupeptin, and 2 µM pepstatin A; Calbiochem] for 30 min. Cell debris was removed by a
10-min centrifugation at 16,000 g, and the supernatant was
incubated with 4 µg of anti-human c-Raf kinase COOH-terminal antibodies at 4°C for 2 h followed by incubation with 100 µl
of a PBS-prewashed protein G Sepharose slurry (containing 30% Protein G Sepharose 4 Fast Flow, Amersham Pharmacia Biotech, Piscataway, NJ)
for 2 h at 4°C with gentle agitation. Immunoprecipitated active Raf-1 was used to phosphorylate and activate glutathione
S-transferase (GST)-MEK, which, in turn, phosphorylates and
activates p42 GST-ERK2. Active GST-ERK2 was then used to phosphorylate
MBP with [
-32P]ATP. The radiolabeled substrates were
allowed to bind to P81 phosphocellulose paper (Whatman, Clifton, NJ),
and the radioactivity per minute was measured in a scintillation counter.
Measurement of Raf-1 phosphorylation. Bovine endothelium grown to confluence in 60-mm dishes was serum deprived for 18 h in medium 199 followed by incubation with 100 nM PMA for the indicated times. Cells were lysed in 500 µl of lysis buffer A, and the total Raf-1 protein was immunoprecipitated with 4 µg of anti-human c-Raf kinase COOH-terminal antibody (Upstate Biotechnology). After electrophoresis on a 10% SDS-PAGE and Western transfer, the proteins were probed with 1.25 µg/ml of rabbit c-Raf p-Ser621 phospho-specific antiserum (Quality Controlled Biochemicals, Hopkinton, MA) and were visualized by ECL.
Ras GTPase activity assay. To measure Ras activity, we used two complementary approaches. In the first method, endothelial cells were cultured in 35-mm dishes until 100% confluent. The cells were serum starved for 18 h in 1 ml of phosphate-free DMEM and loaded with 200 µCi of [32P]orthophosphate for 4 h. The cells were treated with 100 nM PMA or DMSO for 2, 5, 10, and 30 min and then lysed in lysis buffer containing 25 mM Tris, pH 7.5, 150 mM NaCl, 16 mM MgCl2, 1% Nonidet P-40 (NP-40), a 1:1,000 dilution of proteinase inhibitor cocktail (Calbiochem), and 10 µg/ml of anti-v-H-Ras (Calbiochem). The cell lysates were scraped into Eppendorf tubes and centrifuged for 10 min at 16,000 g at 4°C. An additional 2 µg of anti-v-H-Ras were added to the supernatants, and the mixtures were incubated at 4°C for 1 h. Ras protein was then precipitated by protein G Sepharose; the Ras immunoprecipitates were eluted with buffer containing 2 mM EDTA, 2 mM dithiothreitol, and 0.2% SDS; and the remaining GTP and GDP were separated by TLC. The amount of GTP and GDP was quantitated with Molecular Dynamics PhosphorImager 445 SI.
In the second approach to assess Ras activity, BPAECs were grown to confluence in 60-mm dishes and serum starved for 18 h in medium 199 followed by incubation with stimuli for specified times. At the end of the treatment, the cells were washed with ice-cold PBS once and lysed in 500 µl of lysis buffer B containing 25 mM HEPES, pH 7.5, 150 mM NaCl, 10 mM MgCl2, 1 mM EDTA, 1 mM Na3VO4, 25 mM NaF, 0.25% sodium deoxycholate, 1% NP-40, 10% glycerol, and a 1:500 dilution of proteinase inhibitor cocktail (Calbiochem). The lysates were homogenized by pipetting up and down, and cell debris was removed by centrifuging at 16,000 g for 10 min at 4°C. The supernatants were incubated for 30 min with 8 µl of agarose-conjugated Raf-1-GST corresponding to the human Ras binding domain (residues 1-149), which specifically binds to activated Ras (RasGAP; Upstate Biotechnology). After being washed with 500 µl of the lysis buffer three times, the precipitated agarose complex was resuspended in 30 µl of 2× Laemmli sample buffer (32) and boiled for 5 min. The supernatants were collected, and 15 µl were loaded on a 15% SDS-PAGE (32). The gel was transferred to nitrocellulose membrane and then probed with 0.5 mg/ml of anti-human Ha-Ras monoclonal antibody (Transduction Laboratories) for 1 h. A dilution of horseradish peroxidase-conjugated goat anti-mouse antibody (1:10,000; Bio-Rad Laboratories, Richmond, CA) was used as the secondary antibody, and the ECL reagents were used for the final protein detection.Cotransfection of MEK- and ERK2-expressing constructs.
Endothelial cells grown to 50-80% confluence in 35-mm dishes were
transiently transfected with plasmids encoding either HA-tagged ERK2
(HA-ERK2), dominant negative MEK1 (EE-MEK-2E), or a constitutively active MEK1 (EE-MEK-2A). Briefly, the cells were incubated with 1 µg
of the total amount of DNA (1:1 ratio for two plasmids) and 10 µl of
LIPOFECTAMINE (GIBCO BRL) in 1 ml of OPTI-MEM for 6 h. The
solution was replaced by 1 ml of normal growth medium, and the cells
were incubated for 24 h and then serum-starved in DMEM for 20 h. The transfected endothelial cell monolayers were then treated with
either vehicle (0.1% DMSO) or PMA (100 nM) for 10 min. HA-ERK2 kinase
activity was assessed by immunoprecipitation with anti-HA antibody,
followed by an in vitro kinase assay with MBP as a substrate. Briefly,
the cells were quickly rinsed with PBS after treatment with agonists
and lysed with 150 µl of immunoprecipitation buffer containing 10 mM
Tris · HCl, pH 7.4, 1% Triton X-100, 0.5% NP-40, 150 mM NaCl,
20 mM NaF, 0.2 mM sodium orthovanadate, 1 mM EDTA, 1 mM EGTA, and 1%
inhibitor cocktail for 30 min at 4°C. The cells were scraped,
homogenized by passage through a 26-gauge syringe three times, and
centrifuged for 10 min at 4°C. The soluble cell lysate (100 µl)
containing ~100 µg of total protein was incubated with mouse
anti-HA antibody overnight at 4°C and then with 15 µl of protein G
Sepharose at 4°C for 1 h. The immune complexes were washed three
times with immunoprecipitation buffer and three times with kinase
buffer containing 10 mM Tris · HCl, pH 7.4, 150 mM NaCl, 10 mM
MgCl2, and 0.5 mM dithiothreitol. The immune complexes were
resuspended in 40 µl of kinase buffer supplemented with 0.5 mg/ml of
MBP, 25 µM ATP, and 2.5 µCi of [-32P]ATP and
incubated at 30°C for 30 min. The reaction was stopped by adding 14 µl of boiling 4× Laemmli sample buffer (32). Then the
sample was heated to boiling for 5 min and centrifuged for 5 min, and
15 µl of the supernatant were loaded on a 10% SDS-PAGE (32). After electrophoresis, the gel was stained with
Coomassie blue R250, destained, dried, and exposed to X-OMAT film (Kodak).
ERK2 depletion by antisense oligonucleotides. Endothelial cells were grown to confluence in 96-well plates (for Western immunoblotting) or on the electrical cell impedance sensor gold microelectrode wells (for resistance measurement). The complete medium was then replaced with OPTI-MEM (GIBCO BRL) containing the antisense phosphorothioate oligonucleotides (final concentration 5.6 µg/ml of bovine ERK2; Chemicon, Temecula, CA) for 30 min with LIPOFECTAMINE (final concentration 40 µg/ml; GIBCO BRL). After 3 h of incubation at 37°C in 5% CO2, the cells were washed with medium 199 and further incubated for different time periods with 2.9 µg/ml of antisense oligonucleotide followed by PMA treatment. The predicted decrease in ERK2 production by antisense treatment was monitored by Western blotting analysis with anti-pan-ERK antibodies.
Protein concentrations. Protein concentrations were determined by using either the Bradford method (8) or the bicinchoninic acid protocol (Pierce, Rockford, IL) with BSA as a standard.
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RESULTS |
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PKC-induced decreases in electrical resistance across endothelial
cell monolayers.
Treatment with PMA, a recognized potent activator of PKC, increased
bovine endothelial cell intercellular gap formation and permeability to
Evans blue-bound albumin, processes that correlate with increased
activity and translocation of PKC- from the cytosolic to the
membrane compartment (43, 52). Consistent
with these findings, Fig. 1 demonstrates
that PMA challenge induced a rapid decline in transendothelial
electrical resistance across endothelium grown on gold microelectrodes,
with ~90% of the response complete in 30 min after agonist
stimulation. Relatively low concentrations of PMA (5 nM) produced
nearly maximal declines in electrical resistance, indicating the
sensitivity of this method over conventional measurements of Evans
blue-bound albumin or 125I-BSA flux where the maximal
increase in permeability was achieved at 1 µM PMA (36,
43, 52). The PMA-induced decrease in
electrical resistance strongly correlated with the translocation of
PKC-
from the cytosolic to the membrane compartment, reflecting PKC activation (Fig. 1, inset). To further establish the
relationship between PMA-induced decreases in endothelial cell
electrical resistance and PKC activity, we pretreated bovine
endothelium with the specific PKC inhibitor BIM and measured electrical
resistance after PMA challenge. Figure 2
demonstrates that BIM significantly dose dependently attenuates the
PMA-induced decreases in endothelial cell electrical resistance.
Similar results were obtained with another PKC-specific inhibitor,
Ro-31-8220 (data not shown).
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Effect of PMA on ERK activity in bovine endothelium.
Stasek et al. (52) have previously shown that a
PMA-induced increase in endothelial cell permeability is correlated
with the phosphorylation of caldesmon, a key regulatory cytoskeletal protein potentially involved in actin-dependent cytoskeletal
regulation. ERK members of the MAPK family are responsible for phorbol
ester-stimulated in vivo phosphorylation of caldesmon in smooth muscle
(1-3). To further elucidate relevant PKC targets and
biochemical pathways involved in PMA-induced endothelial cell barrier
dysfunction, we next studied the effect of PMA stimulation on ERK
activity in BPAECs. Figure 4 demonstrates
that PMA stimulation significantly increased ERK activity in a time-
and dose-dependent manner as evidenced by Western immunoblotting of
PMA-treated cell lysates with specific anti-phospho-ERK antibodies.
Activation started as early as 10 min after PMA treatment and persisted
for at least 60 min (Fig. 4A), followed by a slow decline
and a return to the basal level at 3 h of treatment (data not
shown), and was observed at concentrations of PMA as low as 5 nM (Fig.
4B).
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Effect of ERK activity on PMA-induced endothelial cell barrier
dysfunction.
Figure 5, A and B,
demonstrates that ERK inhibition with either PD-98059 (a specific
inhibitor of MEK) (4) or olomoucine (a general inhibitor
of MAPK) significantly attenuated (~30-50% inhibition) but did
not completely abolish the PMA-induced decline in transendothelial
electrical resistance. Consistent with these data, depletion of ERK2 by
treatment with specific anti-ERK2 antisense oligonucleotide
significantly attenuated (~30%) the PMA-induced drop in BPAEC
electrical resistance (Fig. 5C), suggesting the potential
existence of ERK-dependent and -independent mechanisms of PMA-induced
endothelial cell barrier dysfunction. It is interesting to note that
the ERK inhibitors themselves caused transient but significant
reductions in electrical resistance, suggesting that basal ERK activity
may be necessary for maintaining the integrity of the endothelial
barrier.
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Role of MEKs in PMA-induced ERK activation.
To characterize the signaling sequence involved in PMA-induced ERK
activation, we pretreated cells with PD-98059, a specific inhibitor of
the dual-specific MEK (4). PD-98059 significantly decreased PMA-induced ERK activation, with complete inhibition at 50 µM (Fig. 6A), consistent
with the recognized role of MEK in ERK activation. To further examine
the requirement of MEK for PMA-induced ERK activation, we cotransfected
BPAECs with MEK1 and HA-ERK2 constructs, then immunoprecipitated ERK2
with HA antibody and measured the activity of ERK in the
immunoprecipitates with MBP as a substrate. Figure
6B demonstrates that unstimulated cells that
overexpress HA-ERK2 have limited basal enzymatic activity, which is
significantly increased by PMA or by cotransfection with a construct
encoding constitutively active MEK1. Coexpression of a dominant
negative MEK1 construct with HA-ERK2 significantly attenuated but,
interestingly, did not completely abolish the effect of PMA on HA-ERK2
activity, suggesting that besides MEK1, additional MEK isoforms such as
MEK2 may contribute to PMA-induced ERK activation.
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Role of Raf-1 and Ras in PMA-stimulated ERK activation in bovine
endothelium.
To examine the requirement of Raf-1 for PMA-induced ERK activation, we
measured Raf-1 kinase activity by immunoprecipitation (with an antibody
generated against the COOH terminus of Raf-1), followed by sequential
phosphorylation of MEK, ERK, and finally MBP (see METHODS
for details). Figure 7
demonstrates that PMA significantly increased Raf-1 kinase activity in
a time-dependent manner, with maximal activation (~6-fold increase
over basal values) at 5 min, which correlated with an increase in Raf-1
autophosphorylation (Fig. 7, inset). Previous studies
(11, 56) have shown that forskolin, a
well-known activator of cAMP-dependent PKA, decreases growth
factor-induced Raf-1 activity in several cell types including fibroblasts and endothelial cells. To evaluate the role of Raf-1 in
PMA-induced ERK activation and endothelial cell barrier dysfunction, we
next examined the effect of forskolin on PMA-induced decreases in
transendothelial electrical resistance across endothelial cell monolayers. Figure 8
demonstrates that forskolin alone (50 µM) transiently increased
electrical resistance and significantly attenuated but did not abolish
PMA-induced decreases in electrical resistance. The effect of forskolin
on electrical resistance correlates with significant attenuation of
PMA-induced ERK activation by forskolin (Fig. 8, inset).
However, the range of forskolin-induced attenuation of the PMA response
(~70%) was higher than the range of the effect of MAPK inhibition on
the PMA-induced decrease in endothelial resistance (30-50%; Fig.
5). These results suggest that forskolin attenuates PMA-induced
endothelial cell barrier dysfunction via several pathways, including
inhibition of the Raf-1-MEK-ERK pathway.
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DISCUSSION |
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We have clarified critical biochemical pathways that participate
in a model of PKC-dependent endothelial cell and barrier regulation.
Although PMA directly occupies diacylglycerol binding sites on PKC to
initiate activation (7), it was recently suggested in
porcine pulmonary artery endothelial cells and human epithelial cells
that Rho activity is required for PMA-induced recruitment of PKC- to
the cell membrane (26). Our data (Fig. 3) did not confirm
this observation, reflecting differences in either the cell types
utilized or the specificity of the Rho inhibitor used. We utilized C3
exotoxin, which specifically ADP-ribosylates, and therefore inactivates
RhoA, RhoB, and RhoC (5, 6), whereas Hippenstiel et al. (26) used the cell-permeable toxin B,
which, via UDP glucosylation, inactivates other Rho family members,
including Cdc 42 and Rac, as well (10, 28,
29).
Information is limited regarding the role of the different PKC isoforms
in phorbol ester-induced permeability response. Our data suggested a
correlation between an increase in permeability and PKC-
activation in bovine pulmonary artery endothelial cells. Involvement of
PKC-
but not PKC-
in the permeability increase after phorbol
ester treatment was demonstrated in porcine aortic endothelial cells
(25). In human umbilical vein endothelial cells
(46), PMA treatment caused translocation and activation of
PKC-
and PKC-
but not of PKC-
. In contrast, PKC-
overexpression significantly reduced PMA-induced permeability in human
dermal microvascular cells (56). We speculate that the
variety of expression patterns of PKC family members in different types
of endothelium may potentially contribute to the cell type-specific
mechanisms of PKC-mediated barrier regulation.
The biochemical events linking PMA-induced PKC activation and barrier dysfunction are incompletely understood. Thrombin- and PMA-mediated PKC activation and endothelial cell barrier dysfunction have been previously noted to be linked with redistribution and phosphorylation of two cytoskeletal proteins, caldesmon and vimentin, in bovine endothelium (52). Smooth muscle caldesmon contains distinct binding sites for actin, myosin, and Ca2+/CaM and potentially regulates actomyosin interactions and actin filament formation in the absence of MLCK activation (2, 34, 40, 44, 51). Smooth muscle caldesmon has multiple phosphorylation sites for kinases such as PKC, ERK, and Ca2+/CaM-dependent kinase II. However, only ERK appears to phosphorylate smooth muscle caldesmon in vivo (1, 3). Caldesmon phosphorylation may reverse the inhibitory effects of caldesmon on cross-bridge cycling and allow actomyosin contraction (12, 18-20, 42). Our data utilizing specific pharmacological inhibitors of the MAPK pathway or antisense strategies suggest the involvement of ERK in the signaling cascade, which ultimately results in PMA-induced endothelial cell barrier dysfunction. Although not specifically addressed in our studies, caldesmon phosphorylation catalyzed by another proline-directed kinase, p34cdc2 kinase, leads to a profound change in the cytoskeleton of intact fibroblasts preceding mitosis (33, 38, 62, 63). In human umbilical vein endothelial cells, ERK participates in the rearrangement of junctional proteins, leading to an increase in endothelial permeability after vascular endothelial growth factor stimulation (30). Taken together, these data indicate that ERK activation may be an essential element in the activation and rearrangement of the endothelial cytoskeleton that can lead to increased endothelial permeability.
Our data indicated that pretreatment of PMA-stimulated cells with PD-98059 completely abolished ERK activation, strongly suggesting that MEKs are required for PMA-induced ERK activation. Cotransfection of dominant negative MEK1 and ERK2 constructs significantly decreased but did not completely abolish PMA-induced ERK 2 activation, suggesting that MEK1 is required but is not sufficient for ERK activation induced by PMA. PD-98059 is able to effectively inhibit MEK1 and at higher concentrations inhibit MEK2 as well but does not affect other MEK homologs (4). Our data suggest that both MEK isoforms, MEK1 and MEK2, may participate in PMA-induced ERK activation in bovine endothelium.
To evaluate events that are further upstream from ERK activation, we next examined the effect of PMA on Raf-1 and Ras activities. Our data demonstrate that PMA treatment significantly increased the activity of Raf-1, a Ser/Thr kinase, which specifically phosphorylates and activates MEK after growth factor- or phorbol ester-induced cell stimulation (37, 54, 58). Recent data indicate that PKA stimulation leads to Raf-1 phosphorylation and enzymatic inhibition in several cell types (24, 50, 59) including endothelial cells (11). Consistent with prior reports by Garcia and colleagues (13, 15) and Patterson et al. (43) that cAMP is barrier protective, forskolin (50 µM for 15 min), a direct activator of adenylate cyclase leading to cAMP generation and PKA stimulation, indirectly attenuated PMA-induced decreases in endothelial cell electrical resistance, suggesting the involvement of Raf-1 in PMA-induced endothelial cell barrier dysfunction. It has been previously shown that the small GTP-binding protein Ras recruits Raf-1 to the membrane and results in its activation after growth factor receptor stimulation (39). Our data demonstrate significantly increased Ras activity after PMA and indicate the involvement of activation of Ras/Raf-1 and MEK signaling pathways in PMA-induced ERK activation and barrier dysfunction. Importantly, Ras and Raf-1 activation (maximal at 5 min; Figs. 7 and 9) precedes ERK activation (maximal at 10 min; Fig. 4) and cytoskeletal rearrangement (started at 10 min; data not shown), suggesting the sequential character of these events. After 1 h of PMA treatment, ERK activity gradually declines, but permeability persists for at least 2 more hours, suggesting that ERK activation is more important for the initiation of cytoskeletal rearrangement and increase in permeability rather than for sustained barrier dysfunction.
PMA may activate ERK via Ras-independent mechanisms (9, 41) such as direct PKC-dependent phosphorylation of Raf-1 (31, 41). However, although this event may stimulate Raf-1 autokinase activity, it does not appear to enhance MEK phosphorylation (37). Our data argue against Ras-independent Raf-1 stimulation after PMA because FPT inhibitor III, a farnesyltransferase inhibitor that prevents Ras processing and Ras-mediated transformation (27), dramatically attenuated PMA-induced Raf-1 and ERK activation. These data strongly suggest that PMA stimulates the MAPK pathway in bovine endothelium via Ras activation. Specific inhibition of PKC with BIM leads to complete inhibition of both PMA-induced Ras (Fig. 10) and ERK activation (data not shown), consistent with a key role for PKC in the activation of the Ras-ERK signaling cascade in bovine endothelium.
In summary, biochemical and physiological data provided in this report
characterize a MAPK pathway that is involved in endothelial cell
permeability in bovine endothelium after PMA stimulation. This pathway
includes sequential activation of PKC, Ras, Raf-1, MEK1, and MEK2 and
leads to ERK1 and ERK2 activation, which potentially participates in
endothelial cell barrier dysfunction via phosphorylation of specific
cytoskeletal targets like caldesmon (Fig.
12) (52).
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ACKNOWLEDGEMENTS |
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We gratefully acknowledge Lakshmi Natarajan and Anila Ricks-Cord for superb technical assistance. Special appreciation is extended to Dr. V. Natarajan (Johns Hopkins University, Baltimore, MD) for providing diperoxovanadate, Dr. K. L. Schaphorst (Johns Hopkins University, Baltimore, MD) for help in conducting the detergent fractionation experiments, and Drs. R. Pestell (Albert Einstein College of Medicine, Bronx, NY) and M. Rosner (University of Chicago, Chicago, IL) for providing the extracellular signal-regulated kinase (ERK) and ERK kinase constructs, respectively.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute Grants HL-44746, HL-50533, and HL-58064 and grants from the American Heart Association.
Present address of N. Bogatcheva: Department of Biochemistry, School of Biology, Moscow State University, Moscow, Russian Federation.
Address for reprint requests and other correspondence: A. D. Verin, Division of Pulmonary and Critical Care Medicine, 5501 Hopkins Bayview Circle, Baltimore, MD 21224 (E-mail: averin{at}welch.jhu.edu).
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. §1734 solely to indicate this fact.
Received 15 September 1999; accepted in final form 21 March 2000.
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