Role of Ras-dependent ERK activation in phorbol ester-induced endothelial cell barrier dysfunction

Alexander D. Verin1, Feng Liu1, Natalia Bogatcheva1, Talaibek Borbiev1, Marc B. Hershenson2, Peiyi Wang1, and Joe G. N. Garcia1

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


    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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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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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-alpha 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 (10-4 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-MOmega 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-alpha -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% beta -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 [gamma -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 [gamma -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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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-alpha 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-alpha 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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Fig. 1.   Effect of phorbol 12-myristate 13-acetate (PMA) on electrical resistance of bovine pulmonary artery endothelial cell (BPAEC) monolayers. Endothelial cell monolayer resistance was monitored for 3 h. At time 0, cells were rinsed with medium 199 to remove serum, incubated in the same medium for 60 min to stabilize basal electrical resistance, and then treated with either vehicle (DMSO) or PMA. Shown are results from a representative experiment (n = 3). Inset: protein kinase (PK) C-alpha immunoblots of PMA-treated (100 nM for indicated time periods) endothelial cell subcellular fractions obtained by differential detergent fractionation as described in METHODS. PKC-alpha immunoreactivity was completely translocated from the cytosolic to the membrane fraction, reflecting PKC activation.



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Fig. 2.   Effect of PKC inhibition on PMA-induced endothelial cell barrier dysfunction. Endothelial cell monolayer resistance was monitored for 3.5 h. At time 0, cells were rinsed with medium 199 to remove serum, incubated in the same medium for ~90 min to stabilize basal electrical resistance, and then pretreated with either vehicle (0.1% DMSO; control) or specific PKC inhibitor bisindolylmaleimide I (BIM; 1 µM) for 1 h after challenge with either vehicle or 100 nM PMA. Shown are results from a representative experiment (n = 3). Inhibition of PKC by BIM almost completely blocked the PMA-induced decrease in resistance of endothelial cell monolayers, consistent with the primary role for PKC activation in PMA-induced endothelial cell barrier dysfunction. Inset: dose-dependent effect of BIM on PMA-induced endothelial cell barrier dysfunction. Cells were pretreated with indicated concentrations of BIM for 1 h, and electrical resistance was monitored after addition of PMA (100 nM) for 90 min.

PMA-induced recruitment of PKC to the cell membrane may be dependent on Rho GTPase (26) and the activity of the small monomeric G protein involved in endothelial cell contractile regulation (17, 55). Our results indicate that PMA-induced PKC-alpha translocation from the cytosolic to the membrane compartment was not affected by C3 exotoxin, a specific Rho inhibitor (Fig. 3A), although under identical conditions, C3 exotoxin treatment completely attenuated DPV (Fig. 3B)- and thrombin-induced MLC phosphorylation (17). Taken together, these data indicate the lack of Rho involvement in PMA-induced PKC activation in bovine endothelium.


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Fig. 3.   A: effect of Rho inhibition on PMA-induced PKC-alpha translocation. BPAECs were pretreated with C3 exoenzyme (2.5 µg/ml for 10 h) in the presence of LIPOFECTAMINE, then stimulated for 15 min with PMA (100 nM) followed by differential detergent fractionation as described in METHODS. Obtained fractions were analyzed by Western immunoblotting with specific anti-PKC-alpha antibodies. All subcellular fractions were quantified with bicinchoninic acid assay, and equal amounts of proteins were loaded onto SDS-PAGE. +, Presence; -, absence. B: effect of C3 exotoxin on diperoxovanadate (DPV)-stimulated myosin light chain (MLC) phosphorylation. BPAECs were pretreated with C3 exotoxin in the presence of LIPOFECTAMINE followed by replacement with LIPOFECTAMINE-free medium containing C3 exotoxin or vehicle as described in METHODS. Then DPV (5 µM for 10 min) or vehicle (0.1% DMSO) was added, and MLC phosphorylation was monitored by urea gel electrophoresis followed by immunoblotting with anti-MLC antibodies. These results indicate the successful access of the C3 exotoxin into the endothelial cell intracellular compartment, validating that pretreatment of BPAECs with C3 exotoxin has no effect on PMA-induced PKC translocation. un-P, unphosphorylated; mono-P, monophosphorylated; di-P, diphosphorylated.

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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Fig. 4.   Effect of PMA stimulation on extracellular signal-regulated kinase (ERK) activation in BPAECs. BPAEC monolayers were stimulated with either 100 nM PMA for indicated periods of time (A) or different doses of PMA for 10 min (B). The level of ERK activation was analyzed by Western immunoblotting of endothelial cell homogenates with specific anti-phospho-ERK (p-ERK) antibodies (ab). The level of ERK protein expression was analyzed by pan-ERK antibody, which preferentially recognizes ERK2 in the cell homogenates.

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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Fig. 5.   Effect of ERK inhibition on PMA-induced endothelial cell barrier dysfunction. A: endothelial cell monolayer resistance was monitored for 3 h. At time 0, cells were rinsed with medium 199 to remove serum, incubated in the same medium for ~20 min to stabilize basal electrical resistance, and then pretreated with either vehicle (0.1% DMSO) or the specific ERK kinase (MEK) inhibitor PD-98059 (PD; 25 µM) for 1 h after challenge with either vehicle or 100 nM PMA. Shown are results from a representative experiment (n = 3). B: endothelial cell monolayer resistance was monitored for 4 h. At time 0, cells were rinsed with medium 199 to remove serum. After 1 h, olomoucine (Olo; 625 µM) or vehicle was added, and cells were incubated for an additional hour after challenge with either vehicle or PMA. Shown are results from a representative experiment (n = 3). C: BPAEC monolayers were pretreated with antisense oligonucleotides against ERK2 for indicated times as described in METHODS and then challenged with 100 nM PMA, and electrical resistance was monitored for 1.5 h. Inset: Western immunoblotting of ERK2 with pan-ERK antibodies shows time-dependent ERK2 depletion after antisense treatment. Each lane represents equal protein loading. No. at left, molecular marker. Decrease in ERK activity by either specific inhibitors or antisense depletion significantly attenuated but did not completely abolish PMA-induced decrease in resistance of endothelial cell monolayer, suggesting involvement of ERKs in PMA-induced endothelial cell barrier dysfunction.

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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Fig. 6.   Effect of MEK inhibition on PMA-induced ERK activation in bovine endothelium. A: BPAEC monolayers were pretreated with specific MEK inhibitor PD-98059 for 15 min and then stimulated with PMA for 10 min. The level of ERK activation was analyzed by Western immunoblotting of endothelial cell homogenates with specific anti-phospho-ERK antibodies. No. at left, molecular marker. B: BPAECs from 35-mm dish with 50-80% confluence, seeded a day before experiment, were transiently transfected with plasmids encoding a hemagglutinin (HA)-tagged ERK2 and either a constitutively active (CA) or dominant negative (DN) MEK1, followed by treatment with vehicle (0.1% DMSO) or PMA (100 nM) for 10 min. HA-ERK2 kinase activity was assessed by immunoprecipitation with specific anti-HA antibody followed by in vitro kinase activity assay with phospho myelin basic protein (pMBP; top) as a substrate as described in METHODS. The cell lysates after PMA challenge were used for Western immunoblotting with anti-HA (middle) or anti-MEK1 (bottom) antibody. Inhibition of MEK by PD-98059 or dominant negative MEK1 significantly attenuated PMA-induced ERK activation.

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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Fig. 7.   Effect of PMA on Raf-1 activity. Confluent BPAEC monolayers were serum deprived for 18 h and then incubated with vehicle (0.1% DMSO) or PMA (100 nM) for indicated periods of time. Cells were lysed, and cell lysates were immunoprecipitated by anti-human c-Raf kinase COOH-terminal antibodies. Raf-1 activity was measured with Raf-1 kinase cascade assay kit as described in METHODS. Data are means ± SE; n = 3 experiments. * Significant difference compared with basal activity, P < 0.05. Inset: PMA-induced Raf-1 phosphorylation. Immunoprecipitated Raf-1 after PMA challenge was subjected to 15% SDS-PAGE followed by transfer to nitrocellulose membrane and then immunoblotted with either c-Raf p-Ser621 phospho-specific antibody (pRaf; 1.25 µg/ml; top) or rabbit polyclonal Raf-1 antibody (1 µg/ml; bottom). Signals were detected by enhanced chemiluminescence (ECL). PMA treatment significantly increased Raf-1 activity, which was correlated with PMA-induced Raf-1 phosphorylation.



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Fig. 8.   Effect of forskolin (FSK) on PMA-induced endothelial cell barrier dysfunction. Endothelial cell monolayer resistance was monitored for 6 h. At time 0, cells were rinsed with medium 199 to remove serum, incubated in the same medium for ~60 min to stabilize basal electrical resistance, and then pretreated with either vehicle (0.1% DMSO) or FSK (50 µM) for 20 min after challenge with either vehicle (control) or 100 nM PMA. Shown are results from a representative experiment (n = 3). FSK significantly attenuated the PMA-induced decrease in resistance of endothelial cell monolayers. Inset: p-ERK immunoblots of BPAECs pretreated with either vehicle or FSK for 20 min after challenge with either vehicle or PMA for 10 min. FSK treatment significantly attenuated PMA-induced ERK activation.

We next studied the effect of PMA on Ras, a small monomeric G protein and recognized Raf-1 upstream effector when in the active GTP-bound Ras conformation (37, 54, 58). Figure 9 demonstrates that PMA produces a significant time-dependent increase in Ras activity (maximal at 5 min, 80% increase over basal level) as shown by TLC that separates GTP from GDP content in Ras immunoprecipitates. PMA treatment also significantly increased Ras content in Raf-1 immunoprecipitates (Fig. 9, inset), consistent with prior reports that Raf-1 only stably associates with active, GTP-bound Ras (41, 58). The PKC inhibitor BIM completely abolished PMA-induced Ras activation (Fig. 10), indicating that PMA activates Ras in a PKC-dependent manner. To estimate the contribution of Ras-dependent and -independent pathways in Raf-1 activation after PMA, we measured Raf-1 activity after pretreatment with the Ras inhibitor FPT inhibitor III (26). Pretreatment of cells with FPT inhibitor III nearly completely abolished PMA-induced Raf-1 (Fig. 11) and ERK activation (data not shown), suggesting that PMA-induced activation of Raf-1 and ERK is Ras dependent.


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Fig. 9.   Effect of PMA on Ras activity. Confluent endothelial cell monolayers were serum starved for 18 h. After being labeled with [32P]orthophosphate for 4 h, monolayers were treated with PMA (100 nM) or vehicle (0.1% DMSO) for indicated periods of time. Cell lysates were immunoprecipitated with 14 µg/ml of anti-v-H-Ras (Ab-1), and then GTP and GDP were eluted from the immunoprecipitates and separated by TLC. Radioactivity was quantitated with a phosphorimager, and the ratio of GTP to (GTP+GDP) was calculated. Data are means ± SE; n = 3 experiments. * P < 0.05 compared with basal activity. Inset: Western blotting analysis of Ras activation. Serum-starved confluent endothelial cell monolayers were treated with PMA or DMSO for 5 min. Cells were lysed with magnesium-containing lysis buffer and incubated with 10 ml of agarose-conjugated Raf-1-glutathione S-transferase containing human Ras binding domain (residues 1-149) for 30 min. After incubation, the agarose complex was resuspended in 2× Laemmli sample buffer (31), boiled for 5 min, and microcentrifuged for 5 min, and the supernatant was subjected to 15% SDS-PAGE. Proteins were transferred to nitrocellulose membrane and immunoblotted with anti-Ras monoclonal antibody. ECL was used for the detection of activated Ras protein. No. at left, molecular marker. PMA stimulation significantly increased RAS activity in bovine endothelium.



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Fig. 10.   Effect of PKC inhibition on PMA-induced Ras activation. Photograph of TLC plate shows intensity of GDP and GTP spots after pretreatment with either DMSO (0.1%) or BIM (1 µM for 60 min) followed by treatment with either 0.1% DMSO or 100 nM PMA for 10 min. PKC inhibition by BIM completely abolished PMA-induced Ras activation. Orig, original spots.



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Fig. 11.   Effect of Ras inhibition on PMA-induced Raf-1 activation. Confluent BPAEC monolayers were serum deprived from the culture medium for 18 h. Cells were incubated with 100 µM farnesyltransferase (FPT) inhibitor III for 3 h followed by incubation with 100 nM PMA for 5 min; then Raf-1 was immunoprecipitated from the cell lysates, and Raf-1 kinase activity was measured with Raf-1 kinase cascade assay kit as described in METHODS. PMA-induced Raf-1 activity with and without FPT inhibitor III pretreatment was calculated. Data are means ± SE of multiple of increase over control value; n = 3 experiments. * P < 0.05. Ras inhibition almost completely abolished PMA-induced Raf-1 activation, indicating that this activation occurred primarily via Ras-dependent pathway.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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-alpha 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-alpha activation in bovine pulmonary artery endothelial cells. Involvement of PKC-alpha but not PKC-varepsilon 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-alpha and PKC-varepsilon but not of PKC-zeta . In contrast, PKC-beta 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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Fig. 12.   PMA-induced endothelial cell barrier dysfunction as shown by schematic representation of the mechanism by which phorbol esters activate mitogen-activated protein kinase cascade and can cause barrier dysfunction in bovine endothelium. See text for further explanation.


    ACKNOWLEDGEMENTS

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.


    FOOTNOTES

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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