Phorbol esters increase MLC phosphorylation and actin remodeling in bovine lung endothelium without increased contraction

Natalia V. Bogatcheva, Alexander D. Verin, Peiyi Wang, Anna A. Birukova, Konstantin G. Birukov, Tamara Mirzopoyazova, Djanybek M. Adyshev, Eddie T. Chiang, Michael T. Crow, and Joe G. N. Garcia

Division of Pulmonary and Critical Care Medicine, Center for Translational Regulatory Medicine, Johns Hopkins University School of Medicine, Baltimore, Maryland 21224

Submitted 21 September 2001 ; accepted in final form 16 April 2003


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Direct protein kinase C (PKC) activation with phorbol myristate acetate (PMA) results in the loss of endothelial monolayer integrity in bovine lung endothelial cells (EC) but produces barrier enhancement in human lung endothelium. To extend these findings, we studied EC contractile events and observed a 40% increase in myosin light chain (MLC) phosphorylation in bovine endothelium following PMA challenge. The increase in PMA-mediated MLC phosphorylation occurred at sites distinct from Ser19/Thr18, sites catalyzed by MLC kinase (MLCK), and immunoblotting with antibodies specific to phosphorylated Ser19/Thr18 demonstrated profound time-dependent Ser19/Thr18 dephosphorylation. These events occurred in conjunction with rearrangement of stress fibers into a grid-like network, but without an increase in cellular contraction as measured by silicone membrane wrinkling assay. The PMA-induced MLC dephosphorylation was not due to kinase inhibition but, rather, correlated with rapid increases in myosin-associated phosphatase 1 (PPase 1) activity. These data suggest that PMA-mediated EC barrier regulation may involve dual mechanisms that alter MLC phosphorylation. The increase in bovine MLC phosphorylation likely occurs via direct PKC-dependent MLC phosphorylation in conjunction with decreases in Ser19/Thr18 phosphorylation catalyzed by MLCK due to PMA-induced increases in PPase 1 activity. Together, these events result in stress fiber destabilization and profound actin rearrangement in bovine endothelium, which may result in the physiological alterations observed in these models.

phorbol myristate acetate; myosin light chain kinase; myosin phosphatase; protein kinase C


ENDOTHELIAL CELL (EC) exposure to a variety of protein growth factors and edemagenic agonists initiates a coordinate series of biological responses (shape change, cell migration, or intercellular gap formation), which are dependent on the assembly and reorganization of cytoskeletal microfilaments. In nonmuscle cells, the actin cytoskeleton forms two functionally distinct structures, cytoplasmic contractile stress fiber cables that span the cell and a submembranous cortical actin ring. Although an increase in stress fiber formation spanning the cell has typically been associated with cell contraction and endothelial barrier dysfunction (7), we recently noted that increases in cortical actin may be involved in the barrier-protective effect of specific agonists on the integrity of monolayer (12).

The role of myosin light chain (MLC) phosphorylation in these cytoskeletal events has emerged; however, it is quite clear that increases in MLC phosphorylation are not at all a prerequisite for EC barrier dysfunction (29, 33). We have previously shown that the direct protein kinase C (PKC)-activating agent phorbol 12-myristate 13-acetate (PMA) induces an increase in albumin clearance occurring in bovine pulmonary artery EC (BPAEC) (33) and a rapid decrease in transendothelial electrical resistance (TER) (42), events indicative of increased vascular permeability. Human lung EC do not appear to respond similarly to PMA with barrier dysfunction (5) and, in fact, demonstrate increases in vascular integrity, although the exact mechanism responsible for this important difference is unknown. Because of the central role of the cytoskeleton in barrier regulation (7), it has been suggested that the differential effect of phorbol esters on bovine and human EC barrier properties may be related to differences in actin cytoskeleton responses (33, 42). The mechanism of PKC-mediated cytoskeletal rearrangement is not completely understood; however, given the diverse signaling targets described for PKC, actomyosin contraction could potentially be modulated by several mechanisms. As noted above, in specific in vivo and in vitro models of barrier disruption, an increase in MLC phosphorylation is a critical event (17, 23). PKC has been demonstrated to have direct and indirect effects on this process, including the capacity to phosphorylate the Ser/Thr phosphatase 1 (PPase 1) at sites that inhibit enzymatic activity toward MLC (35, 36) and to phosphorylate CPI-17, a recently described PPase 1 inhibitory protein, which potentiates PPase 1 inhibition (9), potentially increasing MLC phosphorylation. PKC has also been noted to directly phosphorylate MLC at sites Ser1, Ser8, and Thr9, sites that are distinct from the myosin light chain kinase (MLCK)-preferred phosphorylation sites (Ser19, Thr18) (19, 37). PKC-mediated MLC phosphorylation inhibits subsequent MLC phosphorylation by MLCK at Ser19/Thr18 as well as diminishing the activation of myosin prephosphorylated with MLCK (16, 19, 28). Despite these potential avenues by which PKC activation might increase EC MLC phosphorylation, our prior work in bovine pulmonary artery endothelium using a standard urea gel electrophoresis assay to detect MLC phosphorylation failed to note an increase in MLC phosphorylation following PMA challenge (10).

In this study, we have tested the hypothesis that PMA-induced bovine lung EC barrier dysfunction is mediated through the activation of MLC-dependent contractile machinery with subsequent stress fiber and cytoskeleton rearrangement. We determined that PKC produces a significant increase in bovine endothelial permeability, whereas human EC barrier responses are enhanced. Using a combination of biochemical detection assays and molecular techniques, we found that PMA challenge of bovine EC is associated with rapid increases in MLC phosphorylation at sites other than those preferred by MLCK. The PMA-mediated increase in MLC phosphorylation occurred in conjunction with transient stress fiber formation and was followed by sustained MLC dephosphorylation at Thr18 and rearrangement of the actin cytoskeleton. MLCK activity did not change significantly during PMA stimulation, whereas myosin PPase 1 activity significantly increased in a time-dependent manner. These data suggest that PMA-induced bovine lung EC barrier dysfunction involves PPase 1-dependent actin rearrangement and Ser19/Thr18 MLC dephosphorylation.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Materials. Rabbit anti-MLC antiserum was produced against purified smooth muscle MLC by Biodesign International (Saco, ME). Mouse anti-MLC antibody MY-21 was purchased from Sigma (St. Louis, MO). Anti-MLCK (D119) antibody was kindly provided by Dr. Patricia Gallagher (Indianapolis, IN). Phosphospecific MLC antisera production was performed as we have previously described (30). Antisera to phospho-Ser and phospho-Thr were purchased from Zymed Laboratories (South San Francisco, CA). MLC double-mutant T18AS19A was kindly provided by Dr. Anne Bresnick (Bronx, NY).

Cell culture. BPAEC were obtained frozen at 16 passages (culture line CCL 209) from American Type Culture Collection (Rockville, MD) and were utilized at passages 19–24 as described (33, 42). Cells were cultured in complete medium consisting of DMEM (GIBCO-BRL, Grand Island, NY) supplemented with 20% fetal calf serum (Paragon Biotech, Baltimore, MD), 1% antibiotic-antimycotic mixture (K. C. Biologicals, Lenexa, KS), and 0.1% endothelial cell growth supplement (Collaborative Research, Bedford, MA) and maintained at 37°C in a humidified atmosphere of 5% CO2-95% air. Cells from each primary flask were detached with 0.05% trypsin-EDTA, resuspended in fresh culture medium, and passaged to culture dishes or 12-well plates with coverslips. Human pulmonary artery endothelial cells (HPAEC) were purchased from Clonetics (Walkersville, MD), cultured in EBM-2 medium (Clonetics) with 10% FBS, 1% antibiotic-antimycotic mixture, and 0.1% growth supplement, and treated as mentioned above.

Measurement of TER. TER was measured as we have previously reported (13, 42). In this system, referred to as electrical cell-substrate impedance sensing (ECIS), the cells are cultured on small gold electrodes, with culture media serving as electrolyte. The electrode array (Applied Biophysics, Troy, NY) is mounted into a holder that made electrical contact through the gold contact pads and connected to the ECIS instrumentation. In all measurements reported here, 1 V and 4 kHz were applied to the sample through a 1-M{Omega} resistor. Cells' monolayer resistance data were taken every 0.5 s and processed with a Pentium 100 MHz computer that controlled the output of amplifier and relay switches to different electrodes.

Measurement of endothelial contractility. To visualize BPAEC contractility, we generated flexible rubber substrates by a modification of the protocol of Harris et al. (14) as previously described. Approximately 15 µl of silicone monomer (dimethyl diphenyl polysiloxane, Sigma) were applied onto 18-mm glass coverslips and allowed to spread for 30 min. The upper layer of silicone was polymerized by brief exposure of the coverslip to an open flame. EC were plated onto thin silicon film, and coverslips were placed onto 12-well culture plates. Contractility of subconfluent cells on silicon membrane plates on the surface of liquid silicone was assessed by formation of wrinkles seen with x20 objective on a Nikon Eclipse TE 300 microscope.

Immunofluorescence. BPAEC or HPAEC were seeded onto gelatinized glass coverslips and grown to 80% confluence and then challenged with 100 nM PMA for 5 min to 18 h. After fixation with 4% paraformaldehyde (10 min, room temperature), slides were rinsed and permeabilized with 0.25% Triton X-100 for 5 min, blocked with 2% BSA (30 min), and incubated with blocking solution containing antiphospho-MLC antibodies. After being washed with PBS, cells were incubated in darkness with Alexa 488 goat anti-rabbit conjugate and Texas red-phalloidin. Slides washed with PBS were mounted with Slow Fade kit (Molecular Probes, Eugene, OR) and observed with a x60 objective on a Nikon Eclipse TE 300 microscope.

Subcloning of T18AS19A MLC mutant. T18AS19A MLC mutant lacking Thr18 and Ser19 phosphorylation sites cloned into the pUHD10–3 expression vector was obtained from Dr. Anne Bresnick and subcloned into pcDNA 3.1/V5-His TOPO mammalian expression vector with a pcDNA3.1/V5-His TOPO TA expression kit (Invitrogen, Carlsbad, CA). Sense and antisense primers for the PCR amplification of the T18AS19A MLC were synthesized at the DNA Analysis Facility (Johns Hopkins University). This pair of primers was designed on the basis of Xenopus laevis cytoplasmic myosin II regulatory light chain mRNA sequence (4) as follows: 5'-CAAGATGTCCAGCAAAAGAGCA-3'(sense) and 5'-GTCATCCTTGTCTTTAGCTCCA-3' (antisense). The GeneAmp PCR System 9600 (Perkin Elmer, Norwalk, CT) was used for the amplification of the MLC mutant. We confirmed the DNA sequences of the PCR products before and after cloning into pcDNA3.1/V5-His TOPO vector. The PCR fragments were purified with 1% agarose gel and QIAquick gel extraction kit (Qiagen, Valencia, CA) and cloned into pcDNA3.1/V5-His TOPO expression vector.

MLC phosphorylation analysis by urea gel electrophoresis. This assay was performed as we have previously described in detail (10). In brief, confluent endothelium from 100-mm culture dishes was scraped into 10% trichloracetic acid (TCA) and 10 mM dithiothreitol. After centrifugation, pellets were washed three times with diethyl ether, suspended in 6.7 M urea, and run on a 10% polyacrylamide-40% glycerol gel to separate unphosphorylated MLC from the more rapidly migrating phosphorylated MLC. The proteins were transferred to nitrocellulose and detected by immunostaining with MLC-specific antibodies. After densitometry, MLC phosphorylation degree was compared in EC stimulated with vehicle (5 min), 100 nM thrombin (2 min), and 100 nM PMA (10 min).

MLC immunoprecipitation under denaturing conditions. Confluent BPAEC monolayers in 60-mm tissue culture dishes were labeled with [32P]orthophosphate (0.5 mCi/dish) for 2.5 h in phosphate-free DMEM without serum followed by stimulation with either 100 nM PMA (10 min) or 100 nM thrombin (2 min). The monolayers were rinsed with 2 ml of media and PBS and scraped into 0.1 ml of SDS/denaturing stop solution (PBS, pH 7.4, 1 mM EDTA, 1 mM EGTA, 50 mM NaF, 10 mM NaPP, 0.2 mM sodium orthovanadate, 1% SDS, and 14 mM 2-mercaptoethanol). The homogenates were passed several times through a 16-gauge needle, boiled for 5 min, diluted [1/10] with PBS, and incubated with 0.05 ml of 10% pansorbin (Calbiochem, San Diego, CA) suspension. Samples were clarified by precipitation and incubated with 0.01 ml of anti-MLC rabbit antiserum (60 min at room temperature), then with 0.05 ml of pansorbin suspension (under identical conditions). Immunocomplexes were pelleted by centrifugation for 5 min, washed three times with 1 ml of PBS, boiled for 5 min in 0.1 ml of sample buffer, separated from pansorbin by centrifugation, subjected to SDS electrophoresis, and transferred to nitrocellulose membrane. To evaluate the MLC loading, we subsequently stained membranes with MLC antibodies. The relative intensities of the 32P-labeled MLC were detected by autoradiography and were quantified by scanning densitometry.

We performed immunoprecipitation of MLC from unlabeled BPAEC and HPAEC as described above using mouse monoclonal antibodies MY-21 and protein G-Sepharose (Amersham Pharmacia Biotech, Piscataway, NJ). We assessed MLC phosphorylation using phospho-Ser/Thr antisera and MY-21 to evaluate loading.

MLC immunoprecipitation using V5 antibody from MLC mutant-transfected BPAEC. We transfected confluent BPAEC monolayers in 60-mm tissue culture dishes with the T18AS19A MLC mutant construct using FuGENE transfection reagent (Roche Diagnostic, Indianapolis, IN) according to the manufacturer's protocol. Briefly, cells were rinsed with 4 ml of Opti-MEM-0.5% fetal calf serum (Invitrogen) and then incubated for 4 h with 2 µg of DNA in transfection reagent. After transfection, the medium was changed to normal growth medium, and cells were labeled with [32P]orthophosphate, washed two times with PBS on ice, and then scraped into 0.5 ml of cold immunoprecipitation buffer [1% Triton X-100, 150 mM NaCl, 10 mM Tris, pH 7.4, 1 mM EDTA, 1 mM EGTA, pH 8.0, 0.2 mM sodium orthovanadate, 0.5% Nonidet P (NP)-40, 0.4 µM aprotinin, 25 µM bestatin, 10 µM leupeptin, and 5 µM pepstatin A]. The scraped cells were passed several times through a 26-gauge needle to disperse any large aggregates, and cell homogenates were centrifuged (16.000 g at 4°C) for 15 min. The supernatants (total cell lysates) were incubated with 10 µl/ml IgG on the rotator at 4°C for 30 min. Samples were precleared by centrifugation (8.000 g at 4°C for 2 min), and supernatants were collected and used for immunoprecipitation with either V5 antibody (7 µg per 0.5 ml of cell extract, Invitrogen) or control IgG. Samples were incubated on the rotator at 4°C for 1.5 h, then 30 µl of protein A agarose (Amersham Biosciences, Uppsala, Sweden) were added, and samples were incubated for additional 60 min. After centrifugation (8.000 g at 4°C for 2 min), the protein A-conjugated immune complexes were washed three times with cold immunoprecipitation buffer. Samples were subjected to 10% SDS-PAGE gels followed by either Western immunoblotting with V5 antibody or autoradiography.

MLC detergent fractionation. Confluent BPAEC labeled with [32P]orthophosphate were stimulated with 100 nM PMA (0–10 min), rinsed with 2 ml of PBS, and incubated with extraction buffer (1% NP-40, 150 mM NaCl, 50 mM NaF, 0.5 mM sodium orthovanadate, 50 mM Tris, pH 8.0, and 30 mM mercaptoethanol) for 30 min at 4°C. Pellet was separated by centrifugation and washed with the same buffer. Both supernatant (detergent-soluble fraction) and pellet (detergent-insoluble fraction) were subjected to immunoprecipitation under denaturing conditions with anti-MLC antibodies as described above. Immunoprecipitates of both fractions were analyzed by autoradiography-Western blotting.

MLC phosphorylation analysis by phospho-specific MLC antibodies. BPAEC and HPAEC grown on 35-mm dishes were stimulated with 100 nM PMA (5–120 min), rinsed with PBS, and scraped into 0.1 ml of SDS-PAGE sample buffer. Homogenates were boiled, subjected to SDS electrophoresis, transferred to nitrocellulose membrane, and stained with antipan-, mono-, or diphospho-specific MLC antibodies (30). The signals obtained from these phospho-specific MLC blots after scanning were normalized to the total MLC signal.

Measurement of MLCK activity in nondenaturing immunoprecipitates. MLCK activity was assessed in immunoprecipitated samples as we have previously described (11). Confluent EC in 100-mm culture dishes were challenged with 100 nM PMA for 5 min–24 h, rinsed with PBS, and lysed with 0.3 ml of lysis buffer (20 mM MOPS, pH 7.0, 50 mM MgCl2, 10% glycerol, 0.5 mM EGTA, and 30 mM mercaptoethanol) containing 1% NP-40 for 5 min on ice. The lysates were scraped and centrifuged for 5 min at 4°C, supernatants were diluted up to 1 ml with the same buffer containing 0.1% NP-40, incubated with anti-MLCK antibodies, and then with 0.05 ml of 10% pansorbin. The immunoprecipitation complexes were harvested by centrifugation, washed with PBS, and resuspended in 0.1 ml of 50 mM MOPS, pH 7.4, 10 mM magnesium acetate, 1 mg/ml BSA, and 8 mM mercaptoethanol with or without 10 µM KT-5926. Kinase activity in immunoprecipitates was measured with MLC from bovine muscle (1 mg/ml, Sigma) as a substrate in the above mentioned buffer, containing 1 µM CaM, 0.1 mM [32P]ATP (1 Ci/mM), and 0.3 mM CaCl2 for 30 min at 25°C. We stopped this reaction by pipetting aliquots onto Whatman 3-mm filters and immediately rinsing filters with ice-cold 10% TCA and 2% sodium pyrophosphate. Filters were washed with the same solution, rinsed with ethanol, dried, and counted by liquid scintillation counting. Specific MLCK activity was estimated by subtracting kinase activity, which was insensitive to specific MLCK inhibition with KT-5926 (BioMol Research Laboratories, Plymouth Meeting, PA).

PPase 1 assay. Total and myosin-specific PPase 1 activity was determined as we have previously described (43) with minor modification. Briefly, BPAEC and HPAEC from 100-mm dishes were treated with 100 nM PMA for 0–60 min and rinsed twice with buffer A (50 mM Tris ·HCl, pH 7.0, 0.1 mM EDTA, and 28 mM mercaptoethanol). Buffer A was then added to the dishes (500 µl), and we quickly froze the cells at -70°C and scraped and homogenized them by passing the cell suspension several times through a 1-ml tuberculin syringe to prepare total cell lysates. To prepare myosin-enriched fractions, we treated 400 µl of total cell lysates with 0.6 M NaCl and 0.1% Tween 20 at 4°C for 1 h with constant agitation followed by low-speed centrifugation (5,000 rpm) at 4°C for 30 min. The myosin-containing supernatant was diluted 10 times with buffer A, and myosin was pelleted by centrifugation at 10,000 rpm for 30 min at 4°C. The supernatant was carefully removed, and the pellet was dissolved in buffer A containing 0.6 M NaCl and 0.1% Tween 20 (~20 µl). Phosphatase activity in total lysate or myosin-enriched fractions was determined by Malachite green microtiter-plate assay, where the enzyme reaction was carried out in a final volume of 20 µl in the wells of 96-well plates. One microliter of total cell lysate or myosin-enriched fraction was incubated with 0.25 mM of phosphopeptide KRpTIRR (Upstate Biotechnology, Lake Placid, NY) in assay buffer (50 mM Tris ·HCl, pH 7.0, 0.1 mM EDTA, 28 mM mercaptoethanol, and 0.1 mg/ml BSA) at 30°C for 20 min. As a control, the cell suspension and phosphopeptide were incubated in assay buffer containing Ser/Thr phosphate inhibitors (1 mM EGTA and 5 µM okadaic acid). The reactions were terminated by addition of 100 µl of Malachite green solution (Upstate Biotechnology), and the plates were incubated at room temperature for 15 min before measurement of absorbance at 620 nm. We determined the phosphate released in this enzyme reaction by comparing the absorbance over control to the phosphate standard curve. The amount of proteins contained in cell lysate or myosin-enriched fraction was assessed by TCA precipitation followed by bicinchoninic acid (BCA) protein assay. In brief, 10 µl of sample were added to 1 ml of H2O, mixed with 100 µl of 0.15% sodium deoxycholate, and incubated at room temperature for 10 min. The mixture was blended with 100 µl of 72% TCA and subjected to centrifugation (3,000 g, 15 min, at room temperature). The protein assay was conducted with the BCA protein assay kit (Pierce, Rockford, IL).


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
PKC-induced alterations in electrical resistance across EC monolayers. PMA, a potent activator of PKC, induced the rapid decline in TER across BPAEC grown on gold electrodes over a wide range of PMA concentrations (10–1,000 nM) with maximal decline within 30–60 min after challenge (Fig. 1A). PMA-induced decreases in TER values gradually resolved in a time-dependent manner, returning to near baseline values by 15 h (Fig. 1B). These effects on TER were species specific, as PMA failed to reduce TER values across human pulmonary artery endothelium (HPAEC) (Fig. 1C). This species-specific divergence was observed despite comparable translocation of PKC-{alpha} from the cytosolic to the membrane compartment in both human and bovine pulmonary endothelium (data not shown). In human EC, PMA produced reproducible marginal increases in TER lasting <30 min (Fig. 1C).



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Fig. 1. Effect of phorbol myristate acetate (PMA) on transendothelial electrical resistance (TER) across bovine (BPAEC) and human pulmonary artery endothelial cell (HPAEC) monolayers. Endothelial cell (EC) monolayer resistance was monitored for 45 min, then cells were treated with either 0.1% DMSO (1) or PMA at varying concentrations: 10 (2), 100 (3), or 1,000 nM PMA (4). Shown are results from 2–3 representative experiments. Arrows depict the time of PMA addition. Bovine lung EC responses to PMA at short (A) and prolonged (B) time points. C: the response of human lung EC to PMA.

 

Involvement of MLC phosphorylation in PMA-mediated actin cytoskeleton reorganization. Vascular barrier regulation is known to involve the endothelial cytoskeleton and spatially defined alteration in MLC phosphorylation (7, 10, 12). To assess the level and distribution of phosphorylated MLC during PMA stimulation, we performed immunofluorescent studies utilizing mono- or diphosphorylated MLC-specific antibodies, which recognize sites of MLCK-dependent phosphorylation (Ser19 and Thr18). Double immunofluorescent staining of subconfluent bovine EC challenged for 5 min with PMA (Fig. 2) revealed that F-actin undergoes complex redistribution with colocalization of monophospho- and diphospho-MLC within newly formed stress fibers. Prolonged exposure to PMA (15–120 min) resulted in a complex array of polymerized actin, including formation of a grid-like filamentous network. Despite dramatic declines in diphospho-MLC immunoreactivity (Ser19 and Thr18), MLC remained colocalized with F-actin (Fig. 2). In contrast, immunofluorescent staining with monophospho-specific (Ser19) MLC antiserum remained apparently unchanged in PMA-stimulated cells (5–120 min), although modest increases were observed in areas that did not involve F-actin colocalization. Restoration of stress fibers began after 4 h of PMA stimulation (data not shown), correlating well with the restoration of barrier function (Fig. 1B). Pretreatment of BPAEC monolayers with the specific PKC inhibitor bisindolylmaleimide (BIM) completely abolished PMA-induced alterations in both BPAEC cytoskeletal structure (Fig. 2C) and TER (42), indicating that PMA-induced BPAEC cytoskeletal remodeling, as predicted, is entirely PKC dependent.



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Fig. 2. Effect of PMA on actin and phosphorylated myosin light chain (MLC) localization in BPAEC. Shown are immunofluorescent images of BPAEC challenged with 100 nM PMA for the times indicated, fixed, and double-stained with Texas red-phalloidin and anti-monophospho MLC (p-MLC, A) or anti-diphospho MLC (pp-MLC, B) antibodies as described in MATERIALS AND METHODS (x60 magnification). C: BPAEC were pretreated with either vehicle (0.1% DMSO) or bisindolylmaleimide (Bis, 1 µM) for 60 min, then challenged with 100 nM PMA for 30 min, fixed, and stained for F-actin or diphospho-MLC as described above. Bovine EC demonstrated biphasic actin rearrangement after exposure to phorbol ester with rapid stress fiber formation at 5 min followed by reduction in stress fibers after 15 min of stimulation and complete reorganization into grid-like structures. Mono- and diphospho-MLC staining showed preferential colocalization of phospho-MLC with stress fibers and grid-like structures. Pharmacological inhibition of PKC completely abolished the effect of PMA on EC cytoskeletal structure, indicating that PMA-induced EC cytoskeletal changes are entirely PKC dependent.

 

Subconfluent human EC also demonstrated actin rearrangement in response to PMA, albeit with the distinct differences from the alterations observed in bovine endothelium. Stimulation of human EC with PMA led to initial stress fiber formation accompanied by marginal increases in MLC diphosphorylation (Fig. 3). Prolonged PMA treatment (15–120 min) resulted in decreased levels of stress fibers without formation of the actin-containing grid-like structures. After 1 h of exposure to PMA, human EC appeared comparable with control monolayers. It is important to note that postconfluent EC did not show an induction of stress fiber formation or alterations in MLC phosphorylation during the initial phase of response to PMA (data not shown). The cell behavior during the long-term phase remained very similar to that of subconfluent cells.



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Fig. 3. Effect of PMA on actin and phosphorylated MLC localization in HPAEC. Shown are immunofluorescent images of HPAEC challenged with 100 nM PMA for the time indicated, fixed, and double-stained with Texas red-phalloidin (A) and anti-diphospho-MLC (B) antibodies as described in MATERIALS AND METHODS (x60 magnification). Human EC demonstrated transient stress fiber formation followed by the reversion to the control state with slight increases in cortical actin content. Diphospho-MLC immunoreactivity was colocalized with stress fibers and cortical actin.

 

MLC phosphorylation during PMA stimulation. To directly measure the level of MLC phosphorylation after PMA treatment, we employed two complementary methods. First, MLC phosphorylation induced by PMA treatment was detected by MLC immunoprecipitation from bovine EC prelabeled with 32P followed by SDS electrophoresis. A second method was based on the separation of diphospho-, monophospho-, and unphosphorylated MLC by urea gel electrophoresis followed by Western blotting with MLC antibody. Autoradiography of MLC immunoprecipitates revealed an ~40% increase in MLC phosphorylation after 15 min of PMA stimulation (Fig. 4A), a modest increase compared with the 145% increase evoked by thrombin. This increase in MLC phosphorylation was observed only in bovine EC, as PMA challenge of human EC revealed a slight decrease in the phospho-Ser/Thr content (Table 1). Phospho-MLC detection via urea gel electrophoresis detected a remarkably similar level of thrombin-induced MLC phosphorylation; however, unlike studies of immunoprecipitated MLC, urea gel electrophoresis failed to detect a significant PMA-mediated change in bovine MLC phosphorylation (Fig. 4B). The discrepancy between these two biochemical methods of phospho-MLC detection, as well as with diphospho-MLC immunolocalization, prompted us to analyze MLC phosphorylation in both detergent-soluble and detergent-insoluble fractions. Figure 5 depicts the significant increase in MLC phosphorylation (65%) within the detergent-insoluble (cytoskeletal) fraction of bovine lung EC following 15 min of PMA stimulation, whereas these values were similar to vehicle controls in the detergent-soluble fraction. Experiments conducted with antisera specific for the sites of MLC phosphorylation catalyzed by MLCK (Ser19, Thr18), however, revealed significant and time-dependent decreases in mono- (data not shown) and diphosphorylation of MLC, beginning at 5 min of PMA challenge, reaching maximum reduction at 30 min (~60% reduction), and sustained after 2 h of stimulation (Fig. 6A). Dephosphorylation of Ser19 revealed by monophospho-MLC antibody was less dramatic than Thr18 dephosphorylation in PMA-treated cells (data not shown). Ser19 phosphorylation decreased slightly within 5–15 min after PMA stimulation and returned to the previous level of basal Ser19 phosphorylation after 60 min of stimulation. In contrast to bovine EC, human cells did not demonstrate significant changes in MLC Ser19/Thr18 diphosphorylation level (Fig. 6B). To examine the contribution of phosphorylation sites at other than Ser19 in PMA-stimulated EC, we transfected bovine lung EC with the T18AS19A MLC double-mutant construct in which Ser19/Thr18 were substituted for Ala. Cells were next labeled with [32P]orthophosphate and treated with PMA for 30 min, and the phosphorylation status of the V5 fusion MLC was determined by autoradiography of V5 immunoprecipitates subjected to SDS-PAGE followed by transfer to nitrocellulose. Figure 6C demonstrates significant increases in 32P incorporation in immunoprecipitate MLC after PMA treatment, indicating that PMA stimulation increases MLC phosphorylation at site(s) distinct from Ser19/Thr18. This MLCK-independent MLC phosphorylation suggests a major contribution to the overall increase in MLC phosphorylation after PMA stimulation.



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Fig. 4. Biochemical determination of MLC phosphorylation in PMA-stimulated BPAEC. A: 32P-labeled BPAEC were challenged with thrombin (2 min, 100 nM) or PMA (10 min, 100 nM), immunoprecipitated under denaturing conditions, followed by SDS-PAGE electrophoresis, Western blotting, and autoradiography. The densitometric intensities of bands on the autoradiogram are presented as the percentage of control intensity (n = 3). B: proteins from EC challenged with thrombin and PMA as described in A were precipitated with TCA, dissolved in urea-containing buffer, and separated by urea-gel electrophoresis followed by Western blotting with anti-MLC antibodies. Intensities of bands characteristic for mono- (Mono-P) and diphospho-MLC (Di-P) were analyzed and presented as percentage of control (n = 3). The 2 biochemical methods utilized provide similar results after thrombin stimulation (high level of MLC phosphorylation) but conflicting results with urea gel analysis, which failed to detect the significant increase in MLC phosphorylation after PMA (low level of phosphorylation).

 

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Table 1. Myosin light chain phosphorylation on Ser/Thr residues in PMA-treated BPAEC and HPAEC

 


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Fig. 5. Level of MLC phosphorylation in detergent-soluble and detergent-insoluble fractions. 32P-labeled bovine cells were challenged with 100 nM PMA for 10 min and extracted with Triton X-100-containing buffer. MLC from the supernatant and pellet were immunoprecipitated under denaturing conditions, separated by SDS-PAGE electrophoresis, followed by Western blotting and autoradiography. The intensity of the radioactive bands was analyzed and expressed as the percentage of control (n = 3). Phosphorylated MLCs were only detectable in the detergent-insoluble fraction after PMA stimulation.

 


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Fig. 6. Analysis of site-specific MLC phosphorylation in PMA-treated cells. A: bovine EC were challenged with PMA for the times indicated and lysed with SDS-containing buffer, and the diphospho-MLC content was analyzed with antisera specific to phospho-MLC (Ser19, Thr18). The values provided below each lane represents the mean densitometric values (means ± SE) for each point from 2–3 experiments compared with control. MLC dephosphorylation at Ser19 and Thr18 occurs promptly after PMA treatment of BPAEC with maximal dephosphorylation at 30 min. B: similar experiments were performed in human EC as described in A. The level of diphospho-MLC in HPAEC did not change significantly during PMA treatment. C: BPAEC were transfected with T18AS19A MLC mutant construct in pcDNA 3.1/V5-His TOPO expression vector. After transfection, cells were labeled with [32P]orthophosphate and challenged with either vehicle (0.1% DMSO) or PMA (100 nM) for 30 min, and cell lysates were subjected to immunoprecipitation with either control IgG or V5 antibody. After SDS-PAGE and transfer to nitrocellulose, the level of 32P incorporation in V5-MLC mutant fusion protein was visualized by autoradiography (32P autorad) and immunoblotting with V5-specific antibody. PMA caused significant increase in MLC phosphorylation at sites distinct from Ser19 and Thr18.

 

Effect of PMA on bovine endothelial contractility. To assess the contribution of PMA-induced, site-specific alterations in MLC phosphorylation to EC contractile activity, we monitored the number of wrinkles produced by a population of subconfluent BPAEC grown on a silicone membrane. Resting bovine endothelium demonstrated moderate wrinkling (Fig. 7A), which was completely unaffected by 100 nM PMA challenge (Fig. 7B). In contrast, 3 µM nocodazole, known to stimulate isometric contraction and MLC phosphorylation in fibroblasts and EC (24, 43), immediately induced robust wrinkling (Fig. 7C). These data indicate that PMA-mediated decreases in TER and actin rearrangement in bovine endothelium occur without increased cellular contraction, which is consistent with the declining levels of Ser19 and Thr18 phosphorylation after PMA treatment.



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Fig. 7. Effect of PMA and nocodazole on contractility of bovine pulmonary artery endothelium by membrane wrinkling. A: the sparse level of basal contractility in BPAEC before agonist treatment. B: effect of 10-min PMA (100 nM) challenge that fails to induce significant wrinkle formation compared with basal contractility. C: the robust activation of the contractile apparatus in bovine lung EC produced by 10-min exposure to nocodazole (3 µM) treatment.

 

Role of MLCK and PPase 1 in PMA-induced MLC phosphorylation. To elucidate the mechanism underlying the observed reduction in Ser19/Thr18 MLC phosphorylation after PMA, we next analyzed MLCK activity in MLCK immunoprecipitates. These studies failed to detect any significant changes in MLCK activity in cells treated with PMA regardless of the duration (5 min–4 h), whereas thrombin induced a significant (90%) increase in MLCK activity (Table 2), as we previously reported (11). We next analyzed the total and myosin-specific activity of PPase 1 responsible for dephosphorylating MLC in bovine EC and observed a rapid (1 min) and time-dependent increase in both the overall activity of bovine PPase 1 and myosin-targeted phosphatase activity measured in myosin-enriched fraction reaching maximum at 30 min (Fig. 8, A and B). Human EC challenged with PMA also demonstrated slight increases in total PPase 1 activity in a time-dependent fashion but failed to achieve >50% of the total PPase 1 activity observed in bovine lung EC after PMA challenge (Fig. 8C). Furthermore, the duration of PMA-induced PPase 1 activity was much less substantial than bovine EC returning to normal values by 60 min.


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Table 2. MLCK activity in PMA- and thrombin-treated BPAEC

 


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Fig. 8. Analysis of total and myosin-associated phosphatase (PPase) 1 activities in PMA-challenged bovine endothelium. Bovine (A, B) and human EC (C) were challenged with 100 nM PMA for the times indicated. We determined the activity of PPase 1 in cell homogenates (A, C) and myosin-enriched fractions (B) using a synthetic phosphopeptide as substrate (see MATERIALS AND METHODS). Summarized are data from 3 experiments; *set of results that are significantly different from control (P < 0.05). Both total and myosin-associated PPase 1 activities in bovine EC significantly increase after PMA, whereas total PPase 1 activity in human lung EC fail to achieve this level of activation observed in bovine EC.

 


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
We have demonstrated that EC barrier properties are affected by phorbol ester in a species-specific manner. Bovine pulmonary EC respond to PMA by significant decreases in TER, whereas TER values across human pulmonary EC are briefly but significantly increased without any decline in TER (Fig. 1). These results are consistent with a prior observation that PMA causes a time-dependent increase in hydraulic conductivity of bovine aortic EC monolayer, whereas water flux across human umbilical vein EC monolayer is not significantly affected (5). According to published data, cytoskeletal rearrangements that occur in response to phorbol esters are highly cell specific. For example, PMA leads to the appearance of myosin-containing stress fibers, accompanied by an increase in MLC phosphorylation in Chinese hamster ovary cells (34) but depolymerizes stress fibers in the immortalized Madin-Darby canine kidney renal epithelial cell line (39).

To determine the type of cytoskeleton reorganization that occurs in PMA-treated EC, we utilized a combination of immunofluorescent and biochemical strategies. Our data (Fig. 2) indicate that phorbol ester-treated bovine EC exhibit biphasic cytoskeletal responses. The initial step includes refinement of the actin cortical ring in association with transient induction of cytoplasmic stress fiber formation. The second phase is accompanied by the replacement of polymerized actin-containing stress fibers with actin grid-like structures (Fig. 2). PMA-induced EC cytoskeletal changes are completely abolished by the specific PKC inhibitor BIM (Fig. 2C), indicating that these changes are PKC dependent. These results are consistent with the observation that treatment of bovine EC with 1 µM PMA for 30 min leads to the loss of dense peripheral bands of actin (44), unlike the thick cytoplasmic stress fiber cables produced by thrombin (10). The induced response of human EC to PMA is similar to those of bovine EC, as brief (0–5 min) exposure to phorbol ester also induces stress fiber formation (Fig. 3). However, in human EC, most newly formed stress fibers dissolve completely, and cells revert to the cytoskeletal organization observed in the control state without formation of grid-like structures. As PMA does not decrease but rather augments transendothelial resistance in human EC, we speculate that rearrangement of actin filaments into grid-like structures observed only in bovine lung EC may be important to the subsequent induction of EC barrier dysfunction. The transient stress fiber formation common to both bovine and human EC is observed only in subconfluent cells but not in postconfluent monolayers (data not shown), suggesting a limited role in barrier regulation. This initial stress fiber formation is not involved in PMA-induced barrier dysfunction but could potentially explain why decreases in electrical resistance develop slowly during the initial 30-min period and reach maximum decreases only after 45 min.

Our data clearly indicate that PMA does not increase MLC Ser19 and Thr18 phosphorylation, biochemical events necessary for activation of actomyosin contractile interaction in smooth muscle and nonmuscle cells such as EC (7). Consistent with these data, we failed to observe enhanced PMA-induced bovine EC contraction (Fig. 7), even at time points when transient stress fiber formation occur (Fig. 2). These results differ from reports of phorbol ester-induced isometric tension in porcine pulmonary artery EC (3), again indicating the cell-specific nature of physiological responses to PMA.

As demonstrated in Figs. 4, 5, 6, bovine lung EC, but not human lung EC, respond to PMA with increased MLC phosphorylation. Experiments utilizing MLC antibodies specific for the MLCK-specific sites of MLC phosphorylation (Ser19/Thr18) (Fig. 6) do not support a role for MLCK in this increased MLC phosphorylation (Fig. 7). In contrast to smooth muscle, where PMA produces a slowly developing contractile response (18), our data utilizing a silicone wrinkling assay suggest the increase in MLC phosphorylation after PMA does not increase bovine EC contractility. MLC can be directly phosphorylated in vivo by PKC at Ser1, Ser8, and Thr9 (6, 26), sites distinct from the preferred MLCK-mediated phosphorylation sites (Ser19 and Thr18) (13, 32). Our data (Fig. 6C) indicate that an MLC mutant lacking MLCK-mediated phosphorylation sites is heavily phosphorylated after PMA stimulation, suggesting an importance of sites distinct from Ser19 and Thr18 in the overall increase in MLC phosphorylation after PMA stimulation. Phosphorylation of these MLC sites by PKC may inhibit subsequent MLCK-catalyzed MLC phosphorylation (28, 37) or inhibit the ATPase activity of myosin prephosphorylated by MLCK, thereby decreasing the affinity of myosin to actin (16, 28). Thus the MLC phosphorylation catalyzed by PKC may directly contribute to the decrease in Ser19/Thr18 phosphorylation observed in PMA-treated bovine cells. Although direct MLC phosphorylation by PKC has no effect on transition between extended (filamentous) and folded (monomeric) conformations of myosin (16), this event could have a profound effect on stress fiber stability, as prevention of MLCK-mediated MLC phosphorylation shifts the equilibrium toward the folded, nonfilamentous form of myosin (22). Levels of phosphorylated at Ser19/Thr18 MLC are restored after2hof PMA stimulation and precede the restoration of the normal actin network in EC that occurs at 4 h. In contrast, HPAEC do not exhibit PMA-induced increases in total MLC phosphorylation, MLC dephosphorylation at Ser19/Thr18, or actin stress fibers rearrangement into grid-like structures. Our unpublished data indicate that there are two MLC isoforms in BPAEC and human umbilical vein EC (MLCa and MLCb), with the relative abundance of each isoform distinct for each cell type. We speculate that the differential ability of the two MLC isoforms to undergo PKC-dependent phosphorylation could contribute to the apparent differences in the stability of stress fibers in PMA-treated BPAEC and HPAEC.

We have provided additional mechanistic information on the PMA-induced Ser19/Thr18 MLC dephosphorylation, i.e., the absence of an effect of PKC on EC MLCK activity and the time-dependent activation of the PPase 1. Direct phosphorylation of the low-molecular-weight smooth muscle MLCK isoform by PKC results in a reduced affinity of MLCK for calmodulin (20). In PMA-stimulated intestinal epithelial cells, MLC dephosphorylation correlated with MLCK phosphorylation, although decreased enzymatic activity of phosphorylated MLCK was not confirmed (38). Our results (Table 2) indicate that the activity of the high molecular MLCK isoform first cloned by our laboratory (11) is not altered during the first phase of PMA stimulation and remains at control levels up to 4 h after stimulation. In contrast, our results indicate that the activities of both total PPase 1 and myosin-associated phosphatase increase significantly after 5 min of PMA stimulation with maximum values noted after 30 min of bovine EC challenge. The PMA-mediated increase in human PPase 1 activity was delayed and less dramatic. We speculate that early and significant PMA-induced activation of PPase 1 is responsible for the time-dependent Ser19/Thr18 MLC dephosphorylation we observed in the bovine cells. We do not yet know precisely the characteristic of the phosphatase involved in MLC dephosphorylation at PKC-catalyzed sites. PPase 1 was shown to effectively dephosphorylate all sites in isolated MLC, but with a strong preference for sites of MLCK-catalyzed phosphorylation (8), suggesting the possibility of involvement of phosphatase distinct from PPase 1 in this process.

The exact mechanism by which PPase 1 could be activated in PMA-treated cells is unknown; however, phorbol esters were found to inhibit myosin PPase activity in smooth muscle tissues (21), an event suggested to be responsible for PKC-induced Ca2+ sensitization (27) and possibly Ca2+-independent muscle contraction (18, 21). One candidate for the regulation of PPase 1 activity in a PKC-dependent manner is CPI-17, a novel PPase 1 inhibitory protein and PKC target we have noted to be expressed in human EC (unpublished data). However, phosphorylation of CPI-17 in porcine aorta smooth muscle enhances its inhibitory potency (9), and therefore, this mechanism is unlikely to be involved in the PMA-induced activation of PPase 1 in EC. Direct in vitro phosphorylation of PPase 1 by PKC has been noted with the incorporation of 2 mol of phosphate per mole of myosin phosphatase target subunit (MYPT) 1, the 110-kDa PPase 1 regulatory subunit (36). However, similar to CPI-17, this phosphorylation has been reported to diminish the stimulatory effect of MYPT1 on the activity of PPase 1 catalytic subunit (36). PKC phosphorylates the catalytic subunit of PPase 1 (CS1{alpha}) in vitro (35); however, our prior work indicates the myosin-associated PPase 1 catalytic subunit in endothelium to be the {delta}-isoform (CS1{delta}) (41). Furthermore, the corresponding region of CS1{delta} does not contain the PKC phosphorylation site (35) present in CS1{delta}, and phosphorylation of CS1{alpha} but not CS1{delta} in response to B cell receptor stimulation resulted in enzymatic inhibition of PPase 1 (35). PPase 1 activation was also described in phorbol ester-stimulated pigmented ciliary epithelial cells (25) and skeletal muscle L6 cells (32). Insulin-induced activation of PPase 1 in L6 cells is mediated by the phosphorylation of PP-1G regulatory subunit via a PI 3-kinase/PKC/PKB and/or the ras/MAP kinase/ribosomal S6 kinase kinase cascade (31). Insulin-induced myosin-bound PPase 1 activation in vascular smooth muscle cells was accompanied by dephosphorylation of the regulatory subunit MYPT produced by inhibition of Rho kinase activity (1). This observation is of potential interest as the ras/MAP kinase cascade is known to be involved in EC responses to PMA (42). We speculate that PPase 1 activation by phorbol esters and by insulin could share common pathways.

In summary, we have characterized the progressive actin rearrangement that follows PKC activation in BPAEC but not HPAEC and have linked these events to alterations in the regulation of MLC phosphorylation. PMA rapidly stimulated stress fiber formation followed by assembly of grid-like structures in bovine endothelium, which was accompanied by progressive decreases in Ser19/Thr18 MLC phosphorylation but an overall increase in total MLC phosphorylation, presumably via direct PKC-mediated MLC phosphorylation. These events precede reductions in MLC Ser19/Thr18 phosphorylation and actin reorganization and are not associated with increased cellular contraction. Activation of myosin-bound PPase 1 maintains a low level of Ser19 and Thr18 phosphorylation during prolonged PMA exposure. Human pulmonary EC exhibit brief stress fiber formation, sustained decreases in the level of MLC phosphorylation, and significant augmentation of the endothelium barrier. The results represent the potential mechanism by which phorbol ester may alter stress fiber organization and physiological properties in EC in a species-specific manner.


    ACKNOWLEDGMENTS
 
The authors gratefully acknowledge Lakshmi Natarajan for superb technical assistance, Drs. P. Gallagher (Indianapolis, IN) and A. Bresnick (Bronx, NY) for providing important reagents for this work, and Ellen Reather for expert assistance in manuscript preparation.

DISCLOSURES

This work was supported by National Heart, Lung, and Blood Institute Grants HL-0533, HL-8064, HL-7307, and HL-68062; the American Heart Association; and the Dr. Davis Marine Professorship Endowment.


    FOOTNOTES
 

Address for reprint requests and other correspondence: Joe G. N. Garcia, Johns Hopkins Univ., 1830 E. Monument St., 5th Fl., Baltimore, MD 21218 (E-mail: drgarcia{at}jhmi.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. Section 1734 solely to indicate this fact.


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