1Physiologisches Institut, Justus-Liebig-Universität, D-35392 Giessen; and 2Institut für Pharmakologie und Toxikologie, Westfälische Wilhelms-Universität, D-48149 Münster, Germany
Submitted 8 March 2004 ; accepted in final form 21 June 2004
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ABSTRACT |
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endothelial cell adhesion; endothelial permeability; isometric force; myosin light chain kinase; myosin light chain phosphatase
From the different responses in diverse experimental systems, the question arose whether the differences could be explained mechanistically. Changes in endothelial barrier function are usually due to one of two principal mechanisms: the first is based on activation or inactivation of the actin-myosin-based contractile machinery, which is triggered by the phosphorylation state of myosin light chains (MLC). The phosphorylation state of this regulatory subunit is balanced by a Ca2+-dependent MLC kinase (MLCK) and a MLC phosphatase (MLCP), the activity of which is regulated by various signaling pathways (4, 6, 15, 42). Activation of the contractile apparatus causes changes in cell shape, opening of intercellular gaps and, thereby, a change in barrier function. Not only in cultured endothelial monolayers but also in intact vessels such as coronary venules, activation of the contractile machinery has been identified as the dominant mechanism to increase permeability (43). The second mechanism is based on the stabilization or destabilization of cell-cell or cell-matrix junctions. In endothelial cell junctions, several adhesion molecules are clustered in multiprotein complexes in which transmembrane proteins such as VE-cadherins (17) or integrins (18) bind to corresponding adhesion proteins at adjacent cells or to the extracellular matrix, respectively. These proteins are linked via intermediate proteins, such as catenins or paxillin, to the actin-based cytoskeleton at the inner leaflet of the cell membrane. Assembly and disassembly of these cell adhesion complexes are controlled by intracellular signaling mechanisms, e.g., the cAMP/PKA pathway. It has been shown that phosphorylation of cell-cell and cell-matrix junction proteins, such as catenins, vinculin, or paxillin, can provoke disintegration of these complexes (9, 17, 18, 25, 26, 38, 39). As a result, cells detach from their immediate neighbors or from extracellular matrix. Concomitant shape changes and opening of intercellular gaps are the basis for an increase in permeability (6, 20, 21, 24, 26, 34). It is as yet unclear to which extent contractile machinery and cell junctions contribute to the divergent effects of cAMP/PKA signals in endothelial barriers.
In the present study, we used two well-defined in vitro models of endothelial barriers that exhibit opposite responses to stimulation of the cAMP/PKA signaling: first, endothelial monolayers derived from coronary microvessels [coronary endothelial cells (CEC)], a model that responds with a permeability rise on stimulation of the cAMP/PKA pathway, and, second, endothelial monolayers derived from aorta [aortic endothelial cells (AEC)], a model with a permeability-lowering response. We addressed the question on either model whether stimulation of the cAMP/PKA-signaling pathway could activate or inactivate the contractile machinery. For this purpose, the phosphorylation state of MLC, activation of MLCK and MLCP, and the generation of mechanical force were monitored. It was then studied whether the integrity of endothelial adhesion structures was affected by cAMP/PKA-dependent signaling. To elucidate this mechanism, the cellular localization of VE-cadherin, an integral protein of cell-cell junctions, and paxillin, an integral protein of cell-matrix adhesion plaques, was analyzed by cell fractionation and immunofluorescence microscopy. The cAMP/PKA-signaling pathway of the endothelial cells was activated by two different receptor agonists, the adenosine analog 5'-N-(ethylcarboxamido)adenosine (NECA) or isoproterenol (Iso), or by forskolin (FSK). The first two agents activate the adenylyl cyclase coupled with endothelial adenosine or -adrenergic receptor, respectively; the latter activates adenylyl cyclase directly.
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MATERIALS AND METHODS |
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Cell cultures. Macrovascular AEC from porcine aortas were isolated and cultured as previously described (36). Microvascular CEC were isolated from 250-g male Wistar rats and grown in culture as previously described (32). Confluent cultures of primary endothelial cells were trypsinized in PBS [composed of (in mM) 137 NaCl, 2.7 KCl, 1.5 KH2PO4, and 8.0 Na2HPO4, at pH 7.4, supplemented with 0.05% (wt/vol) trypsin, and 0.02% (wt/vol) EDTA] and seeded at a density of 7 x 104 cells/cm2 on either 24-mm round polycarbonate filters (pore size 0.4 µm), glass coverslips, or 60-mm plastic culture dishes for determination of albumin permeability, immunohistochemistry, or protein analysis, respectively. Experiments were performed with confluent monolayers 4 days after seeding.
Experimental protocols. The basal medium used in incubations was modified Tyrode's solution (composition in mM: 150 NaCl, 2.7 KCl, 1.2 KH2PO4, 1.2 MgSO4, 1.0 CaCl2, and 30.0 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid; pH 7.4, 37°C) supplemented with 5% (vol/vol) heat-inactivated newborn calf serum (10 min, 60°C). Agents were added as indicated in the figure legends. Stock solutions of FSK, ML-7, NECA, and Rp-8-CPT-cAMPS were prepared with DMSO. Where indicated, aliquots of the stock solutions were added to the cells. Final solvent concentrations were <0.1% (vol/vol). The same final concentrations of DMSO were included in all respective controls. All other agents were dissolved in basal medium and were added where indicated.
Macromolecule permeability. Endothelial cells were cultured on polycarbonate filter membranes. Permeability of the endothelial cell monolayer was determined by the flux of Trypan blue-labeled albumin across the monolayer as previously described (29).
Force measurement. Isometric force measurements were performed as previously described by Kessler et al. (14) with minor modifications. Briefly, endothelial cells were seeded at 300,000 cells/cm2 on the collagen lattices that were precast in 1 x 1.5-cm molds. On these lattices, the cells were grown to confluence within 2 days. Afterward, the molds were transferred to specially designed force apparatus, the lattices were connected to force transducers (KG 7A with bridge-amplifier DUBAM 7C; Scientific Instruments, Heidelberg, Germany) and were maintained in a humidified 5% CO2 atmosphere at 37°C. After an equilibration period of 4 h, isometric force of CEC or AEC reached a plateau of 180 ± 19 or 126 ± 15 µN, respectively. These values are referred to as stable baseline forces. Only those cell-loaded lattices with a stable baseline were used for force experiments. Integrity of the endothelial monolayers was routinely checked by phase-contrast microscopy or immunofluorescence staining of the cell-cell adhesion protein VE-cadherin.
Determination of MLC phosphorylation.
The phosphorylation of MLC was determined by glycerol-urea polyacrylamide gel electrophoresis and Western blot analysis as described previously (7, 31). This procedure allows separation of nonphosphorylated from phosphorylated MLC protein, the latter of which migrates more rapidly. Briefly, electrophoretically separated proteins were transblotted on PVDF membranes and incubated with an anti-MLC antibody followed by an alkaline phosphatase-conjugated anti-IgM antibody. Blots were scanned densitometrically, and the stoichiometry of MLC phosphorylation (expressed as mol PO4/mol MLC) was calculated from the densitometric values of non-(MLC), mono-(MLCP), and diphosphorylated MLC (MLC
PP) as follows
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As all MLC can become diphosphorylated, MLC phosphorylation varies between 0 and 2 mol PO4/mol MLC.
MLCK activity assay.
MLCK activity was determined according to Verin et al. (40) with some modifications. Briefly, endothelial cells in 60-mm culture dishes were incubated in the presence of ML-7 (1100 µM) and FSK (510 µM) or respective volumes of basal medium (controls) for 10 min at 37°C. Afterward, the dishes were rinsed twice with PBS and cells were lysed with 0.3 ml of lysis buffer (20 mM MOPS, pH 7.4, 25 mM -glycerol phosphate, 5 mM EGTA, 10% glycerol, 10 mM NaF, 1.5 mM Na3VO4, 2 mM DTT, 1 mM PMSF, and 0.1 µg/ml complete proteinase inhibitor cocktail) containing 1% NP-40 for 10 min on ice. The lysates were scraped, passed five times through a 30-gauge needle, and centrifuged for 5 min at 4°C. Kinase activity was measured with MLC from bovine muscle as substrate in NP-40-free lysis buffer containing 0.1 µCi of [
-32P]ATP in a final volume of 50 µl for 20 min at 30°C. The reaction was terminated by transferring aliquots onto P-81 Whatman filter membranes. The membranes were immediately rinsed with ice-cold 10% TCA, washed five times with 0.75% phosphoric acid (wt/vol), and were subsequently rinsed with acetone and dried. The filter squares were counted in a scintillation counter. Specific MLCK activity was estimated by subtracting kinase activity, which was insensitive to specific inhibition with MLCK pseudosubstrate (342352) amide or ML-7. For both inhibitors, maximum effective doses were determined to be 100 and 50 µM, respectively.
Protein phosphatase assay.
Protein phosphatase activity was determined according to Neumann et al. (27). To determine protein phosphatase activity, [32P]-labeled phosphorylase a was prepared as substrate by phosphorylation of phosphorylase b with phosphorylase kinase. Briefly, to obtain [32P]-labeled phosphorylase a, phosphorylase b (5 mg/ml) was incubated with phosphorylase kinase (200 U/ml) in a 2-ml incubation cocktail containing 50 mM Tris, pH 7.4, 20 mM MgCl2, 31 mM -mercaptoethanol, 0.5 mg/ml BSA, 1 mM CaCl2, 1 mM ATP, and 1 mCi
-[32P]-ATP for 2 h at 30°C. The [32P]-labeled phosphorylase a was precipitated by addition of 2 vol of ice-cold saturated ammonium sulfate solution. The cocktail was incubated for 20 min on ice and was centrifuged for 30 min at 12,000 g at 4°C. The precipitate was solubilized in 2 ml dialysis buffer (10 mM Tris·HCl, pH 7.4, 1 mM EDTA) dialyzed at room temperature against 2x 2-liter dialysis buffer and finally stored at 4°C. [32P]-labeling was estimated by measuring aliquots of the substrate in a liquid scintillation counter.
For determination of protein phosphatase activity (2), diluted protein fractions from endothelial cells were preincubated in a total volume of 30 µl for 10 min at 30°C in the presence or absence of okadaic acid, the protein phosphatase 2A inhibitor. The reaction was started by addition of 20 µl of [32P]-labeled phosphorylase a in an incubation cocktail containing 50 mM Tris·HCl, pH 7.4, 12.5 mM caffeine, 0.25 mM EDTA, 0.25% (vol/vol) -mercaptoethanol. After 20-min incubation at 30°C, the reaction was terminated by addition of 20 µl of 50% (wt/vol) ice-cold TCA and 30 µl of 2% (wt/vol) BSA. After 15 min on ice, the suspension was centrifuged for 5 min at 12,000 g and 4°C. Aliquots of the supernatant were measured in a liquid scintillation counter. Reactions were carried out in duplicate. To ensure linear rates of phosphorylase a dephosphorylation, dephosphorylation was restricted to >25%.
Detection of protein phosphatases in the myosin-enriched and -depleted cell fraction. The content of protein phosphatases 1 and 2A (PP1 and PP2A) in the myosin-enriched cell fraction was analyzed as previously described (4). Briefly, confluent endothelial monolayers on 10-cm dishes were stimulated as indicated in the text. Afterward, the monolayers were rinsed twice with PBS to remove the incubation medium, 200 µl of homogenization buffer (0.1 mM EDTA, 28 mM mercaptoethanol, 1 µg/ml Pefabloc, Tris·HCl, pH 7.4) were added, and dishes were cooled immediately to 80°C. Afterward, the cells were scraped and homogenized. Homogenates were incubated with a high-salt buffer [0.6 M NaCl, 0.1% (vol/vol) Tween 20, 1 µg/ml Pefabloc] for 1 h at 4°C and centrifuged at 4,500 g for 30 min at 4°C. Supernatants were diluted 10-fold with assay buffer (0.1 mM EDTA, 28 mM mercaptoethanol, Tris·HCl, pH 7.0) and centrifuged again at 8,200 g for 40 min at 4°C. The pellet (myosin-enriched fraction) and the supernatant (myosin-depleted fraction) were eluted in Laemmli sample buffer (16) and analyzed by Western blot using antibodies against the catalytic subunits of PP1 and PP2A. In accordance with previous reports (7), the myosin-enriched fractions contained PP1 but not PP2A.
Immunofluorescence microscopy. Confluent endothelial monolayers were rinsed three times with PBS (10 mM phosphate buffer, 2.7 mM KCl, and 137 mM NaCl, pH 7.4), then fixed with 100% methanol for 10 min at 20°C, and washed again three times with PBS. The cells were covered with 100 µl of an anti-pan-cadherin (diluted 1:100 in PBS) or a FITC-conjugated anti-paxillin antibody (diluted 1:100 in PBS) and incubated for 12 h at 37°C. Afterward, the coverslips were washed three times with PBS. In the case of VE-cadherin staining, specimens were covered by 100 µl of a FITC-conjugated anti-mouse IgG (diluted 1:200 in PBS) and incubated for 3 h at 37°C. The coverslips were finally mounted on glass slides with a drop of a HEPES-buffered glycerol solution [1:2 (vol/vol), pH 8.5]. Cell monolayers were visualized using an Olympus IX 70 microscope (Hamburg, Germany).
Detergent fractionation and Western blot. Paxillin was determined in detergent-soluble and -insoluble fractions of cell proteins to estimate its translocation between the cytoskeleton and cytosolic compartment. Endothelial monolayers were rinsed three times with PBS and incubated with 0.3 ml of NP-40 solubilization buffer [1% Nonidet P-40 (vol/vol), 150 mM NaCl, 20 mM dithiothreitol, 10 mM PMSF, 50 mM Tris·HCl, pH 8] for 10 min at 4°C. The supernatant (detergent-soluble, cytosolic fraction) was removed, the culture dish was rinsed three times with ice-cold PBS, and the remaining detergent-insoluble proteins (detergent-insoluble, cytoskeleton-rich fraction) were solubilized into 0.3 ml of Laemmli SDS buffer (16). To determine the total paxillin concentration, endothelial monolayers were rinsed three times with PBS and solubilized into 0.3 ml Laemmli SDS buffer (total cell lysate). Equal volumes of either detergent-soluble, detergent-insoluble fractions, or total protein lysate were applied for SDS-PAGE [12% (wt/vol) total acrylamide concentration] and transferred onto PVDF membranes. The membranes were blocked with 3% (wt/vol) bovine serum albumin and probed with an anti-paxillin antibody (0.2 µg/ml). The signal was visualized with an alkaline phosphatase-conjugated anti-rabbit IgG antibody (0.1 µg/ml).
Statistical analysis. Data are given as means ± SD of n experiments of independent cell preparations. The comparison of means between groups was performed by one-way ANOVA followed by a Bonferroni post hoc test. Changes of parameters within the same group were assessed by multiple ANOVA analysis. Probability (P) values of <0.05 were considered significant.
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RESULTS |
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In the next step, we analyzed whether stimulation of this pathway also influenced the contractile apparatus of AEC. On addition of NECA (100 nM), Iso (10 µM), or FSK (5 µM), isometric force significantly declined within 10 min from a control level of 126 ± 15 to 25 ± 16, 32 ± 17, or 24 ± 19 µN (P < 0.05 vs. time 0 min, n = 5), respectively. As shown for FSK in Fig. 4B, the FSK-induced contractile response was identical to that observed in microvascular coronary monolayers. The FSK-stimulated decline in isometric force was paralleled by a reduction of MLC phosphorylation (Fig. 5B). This effect was inhibited by the PKA inhibitor Rp-8-CPT-cAMPS. The response of MLC phosphorylation in AEC was also comparable to that of CEC.
We then analyzed whether the activation of cAMP/PKA signaling by FSK caused a decline of MLC phosphorylation by inactivation of MLC kinase. Endothelial cells were exposed to FSK and MLCK activity was determined in cell homogenates of treated cells. Exposure of endothelial cells to 5 µM FSK caused a reduction of MLCK activity by 25 ± 12% compared with control (P < 0.05; n = 5), indicating that MLCK activity is modulated by the cAMP pathway in endothelial cells. However, this reduction of MLCK activity is too moderate to account for the cAMP-induced MLC dephosphorylation determined in intact cells. Therefore, in the first step, the role of MLC phosphatase was analyzed in intact endothelial cells using a pharmacological approach. For that reason, FSK-stimulated MLC phosphorylation was determined on one hand under inhibition of MLC kinase by ML-7 and on the other hand under inhibition of MLC phosphatase by calyculin A. If endothelial cells were exposed to increasing concentrations of the inhibitor of MLC kinase, ML-7, MLC phosphorylation declined concentration dependently (Fig. 7A), indicating that the basal activity of MLC kinase contributes to the basal state of MLC phosphorylation. FSK (5 µM) in copresence of 50 µM ML-7, a dose that completely blocked MLCK activity in endothelial cells, caused a further drop of the MLC phosphorylation level. These data indicate that a cAMP-dependent pathway can increase MLC phosphatase activity.
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As recently shown (4, 7), dephosphorylation of MLC in endothelial cells requires translocation and binding of the catalytic subunit of protein phosphatase to myosin. Therefore, in a second approach, translocation of the protein phosphatase 1 subunit into the myosin-enriched cell fraction and the phosphatase activity were analyzed in FSK-stimulated endothelial cells. As shown in Fig. 8A, FSK increased phosphatase content in the myosin fraction. The increase of the phosphatase content in the myosin-enriched cell fraction went along with a reduction of the phosphatase content in the corresponding myosin-depleted cell fraction. The analysis of the phosphatase activity (Fig. 8B) showed that the FSK-stimulated signaling mechanism not only shifted protein phosphatase 1 into myosin-enriched cell fraction but also caused a distinct increase in phosphatase activity in that fraction.
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DISCUSSION |
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In the past, it was documented in a number of studies that stimulation of the cAMP/PKA pathway causes opposing effects on the control of endothelial barrier function. However, the divergent responses in endothelial cells of different origins have remained unexplained. In the present study, we observed that stimulation of cAMP production provokes an increase in macromolecule permeability not only in CEC from rat but also in CEC derived from guinea pig and swine. This suggests that the increase in permeability is a unique cAMP effect in microvascular CEC and rules out the possibility that this cAMP effect on barrier function is due to heterogeneity of species.
We used agonists for receptors coupled in a stimulating manner to adenylyl cyclase in endothelial cells as well as FSK, which activates adenylyl cyclase directly. We showed that the investigated effects on endothelial barrier function and contractile activation were comparable at similar elevations of cellular cAMP contents. These effects were blocked by the highly specific inhibitor of PKA, Rp-8-CPT-cAMPS. With these maneuvers, we analyzed the response of endothelial monolayers to activation of cAMP/PKA signaling. Although effects of cAMP/PKA signaling on endothelial barrier function have been studied before, it has remained unclear which mechanisms are responsible for the divergent effects of cAMP on endothelial barrier function in different experimental models. Patterson et al. (30) demonstrated that stimulation of the cAMP/PKA pathway reduces phosphorylation of MLC in endothelial cells. The effect was attributed to inhibition of MLCK, which was shown to be directly phosphorylated by PKA. These data and findings from other groups (3, 10, 22) have led to the assumption that stabilization of endothelial barriers via cAMP/PKA signaling is due to an inactivation of the contractile machinery based on inactivation of MLCK.
In some endothelial models, it has also been observed that stimulation of the cAMP/PKA signaling can cause a reorganization of the F-actin-based cytoskeleton and stabilizes cell adhesion structures (19). Stimulation of cAMP synthesis can increase junctional strands between endothelial cells of intact microvessels (1) and reduces gap formation in venules induced by inflammatory stimuli (21). An interesting observation was also made by Huxley and colleagues (12, 13), who showed that in intact microvessels the glycocalyx of endothelial cells can substantially contribute to the cAMP-mediated modulation of endothelial barrier function. None of these studies could explain, however, why cAMP/PKA signaling leads to a loss of barrier function in other endothelial models and microvessel preparations.
In the present study, the effects of cAMP/PKA-signaling mechanisms on MLCK and MLCP were studied using two different approaches. Direct determination of the MLCK activity showed that these signals cause only a moderate inhibition of MLCK activity, too small to account for the pronounced dephosphorylation of MLC found in cAMP-stimulated cells. Analysis of the MLC phosphorylation level in endothelial cells in which MLCK was completely inhibited revealed that stimulation of cAMP/PKA signaling causes a drop in MLC phosphorylation even under these conditions. This observation indicated that dephosphorylation of MLC is largely due to a cAMP-induced increase in MLCP activity. Indeed, we found that translocation of the catalytic subunit of protein phosphatase 1 into the myosin-enriched fraction, indicative of site-specific phosphatase activation, is increased under these conditions.
In the case of AEC, MLC dephosphorylation and decline in force were closely correlated with the decrease in barrier permeability when the cAMP/PKA-signaling pathway was activated. Comparable elevations of the cellular cAMP levels by the different stimuli were accompanied by similar effects on force, MLC phosphorylation, and monolayer permeability. This indicates a causal relationship between isometric tension and permeability and a tight coupling of the cAMP-signaling mechanism to these cellular responses, as has been suggested by several studies before (8, 15, 30, 31, 34).
In monolayers of CEC, however, permeability was increased despite contractile inactivation when cAMP/PKA signaling was activated. This occurred even though the two experimental models were very similar in terms of culture conditions. In contrast to AEC, cAMP/PKA signaling caused a rapid disintegration of cellular adhesion complexes in CEC. The changes in VE-cadherin and paxillin delocalization occurred almost simultaneously with the changes in permeability. Previous studies have shown that endothelial barrier function is disturbed when adhesion complexes disintegrate (8, 17, 18, 25). One possible explanation is that if the tethering forces keeping cells spread on the extracellular matrix are lost and intercellular adhesion is weakened, cellular shape changes lead to the opening of gaps between cells. Because delocalization of paxillin proceeded faster than that of VE-cadherins in cAMP/PKA-stimulated CEC, one may speculate that the loss of cell adhesion starts at the cell-matrix adhesion sites rather than at cell-cell junctions. The molecular mechanisms by which cAMP/PKA signaling can cause destabilization of cell adhesion structures in endothelial cells are largely unknown. It has been shown previously in adrenal cortex-derived cells (9, 33) that PKA can stimulate dephosphorylation of paxillin due to activation of a protein tyrosine phosphatase possessing a Src homology (SH)2 domain (SHP-2). Dephosphorylation of paxillin went along with translocation of the phosphorylated protein to cytoplasm, reorganization of focal adhesion plaques, and detachment of the cells from the extracellular matrix. Recently, it has been shown that SHP-2 is a component of cell adhesion also in endothelial cells (38, 39).
In conclusion, the cAMP/PKA-mediated control of contractile elements in macro- and microvascular endothelial cells is identical. Tension is reduced, which, per se, stabilizes barrier function. In coronary microvascular cells, there is an additional cAMP-mediated effect, i.e., cell detachment, that predominates in the control of barrier function.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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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