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Am J Physiol Heart Circ Physiol 288: H1296-H1305, 2005. First published November 4, 2004; doi:10.1152/ajpheart.00687.2004
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Regulation of actin dynamics is critical for endothelial barrier functions

J. Waschke,1 F. E. Curry,2 R. H. Adamson,2 and D. Drenckhahn1

1Institute of Anatomy and Cell Biology, University of Würzburg, Würzburg, Germany; and 2Department of Human Physiology and Membrane Biology, School of Medicine, University of California, Davis, California

Submitted 12 July 2004 ; accepted in final form 27 October 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We tested the hypothesis that the equilibrium between F- and G-actin in endothelial cells modulates the integrity of the actin cytoskeleton and is important for the maintenance of endothelial barrier functions in vivo and in vitro. We used the actin-depolymerizing agent cytochalasin D and jasplakinolide, an actin filament (F-actin) stabilizing and promoting substance, to modulate the actin cytoskeleton. Low doses of jasplakinolide (0.1 µM), which we have previously shown to reduce the permeability-increasing effect of cytochalasin D, had no influence on resting permeability of single-perfused mesenteric microvessels in vivo as well as on monolayer integrity. The F-actin content of cultured endothelial cells remained unchanged. In contrast, higher doses (10 µM) of jasplakinolide increased permeability (hydraulic conductivity) to the same extent as cytochalasin D and induced formation of intercellular gaps in cultured myocardial endothelial (MyEnd) cell monolayers. This was accompanied by a 34% increase of F-actin and pronounced disorganization of the actin cytoskeleton in MyEnd cells. Furthermore, we tested whether an increase of cAMP by forskolin and rolipram would prevent the cytochalasin D-induced barrier breakdown. Conditions that increase intracellular cAMP failed to block the cytochalasin D-induced permeability increase in vivo and the reduction of vascular endothelial cadherin-mediated adhesion in vitro. Taken together, these data support the hypothesis that the state of polymerization of the actin cytoskeleton is critical for maintenance of endothelial barrier functions and that both depolymerization by cytochalasin D and hyperpolymerization of actin by jasplakinolide resulted in an increase of microvessel permeability in vivo. However, cAMP, which is known to support endothelial barrier functions, seems to work by mechanisms other than stabilizing F-actin.

permeability; actin; jasplakinolide; adenosine 3',5'-cylic monophosphate; vascular endothelia cadherin

THE AIM OF THE PRESENT EXPERIMENTS was to evaluate the contribution of the state of actin polymerization to the endothelial barrier function in individually perfused mammalian microvessels and endothelial cells in culture. The approach extends previous investigations, using the expertise in our two laboratories, to evaluate the contributions of cell-cell adhesion mechanisms, contractile mechanisms, and actin depolymerization to the regulation of endothelial permeability under different conditions in intact microvessels and in culture (48, 49). Our studies indicated that contractile forces, although present in the resting endothelium in vivo, do not significantly contribute to increased permeability in response to various stimuli in vivo (3, 49). In contrast, regulation of intercellular junctions such as the endothelial tight and adherens junction seems to be primarily important for permeability control. Both types of intercellular junctions are linked to the actin cytoskeleton. Therefore, in this study, we focused on the contribution of actin regulation in endothelial barrier control and investigated the effect of actin polymerization and depolymerization on microvessel permeability in vivo and the integrity of endothelial adherens junctions in cultured microvascular myocardial endothelial (MyEnd) cells.

We previously demonstrated that depolymerization of F-actin by cytochalasin D increases microvessel hydraulic conductivity (Lp) in vivo and reduces binding of vascular endothelia cadherin (VE-cadherin)-coated microbeads to the surface of cultured endothelial cells (48, 49). Jasplakinolide, a spongeal toxin from jaspis johnstoni, has been used to investigate the role of actin reorganization in different cellular processes. It binds to pointed ends of actin filaments (F-actin) and thereby inhibits filament depolymerization (50). Moreover, jasplakinolide favors filament nucleation and polymerization by stabilizing the binding interface between adjacent actin monomers (G-actin) (11, 20, 32). In single-perfused mesenteric microvessels, jasplakinolide reduced the permeability-increasing effects of cytochalasin D (48, 49) suggesting that jasplakinolide, under these conditions, stabilized F-actin and prevented its depolymerization by cytochalasin D. It is well known that the actin cytoskeleton of endothelial cells consists of at least two morphologically and functionally different systems: the junction-associated actin filament web and the cytoplasmatic system of stress fibers (16). These subsets of the actin cytoskeleton are highly dynamic and can be regulated in response to various extracellular stimuli (51). In this context, members of the Rho family GTPases have been identified as important regulators of actin dynamics (23, 52).

To test the hypothesis that regulation of the actin cytoskeleton is critical for maintenance of endothelial barrier functions, in this study we used jasplakinolide to promote actin polymerization and to change the equilibrium between G- and F-actin toward F-actin. Our study demonstrates that both depolymerization and hyperpolymerization of F-actin reduces endothelial barrier properties in vivo and in vitro, whereas stabilization of F-actin is protective for barrier function. Because cAMP, well known to protect endothelial barrier functions, has been shown to be involved in the regulation of the endothelial actin cytoskeleton (10, 24, 27, 33, 46), we also investigated the effects of cAMP on cytochalasin D-induced endothelial barrier breakdown. In our study, increased cAMP did not protect endothelial barrier properties against the action of cytochalasin D. These data support the hypothesis that tight regulation of actin dynamics is required for maintenance of endothelial barrier functions. This underlines the role of the actin cytoskeleton as a possible key player in the response to inflammatory stimuli.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animal preparation. Rats were kept under conditions that conformed to the National Institutes of Health Guide for the Care and Use of Laboratory Animals, approved by the Institutional Animal Care and Use Committee of the University of California (Davis, CA). Rats (male, Sprague-Dawley, 350–450 g, Hilltop Laboratory Animals) were anesthetized with pentobarbital sodium (65 mg/kg body wt) given subcutaneously. Anesthesia was maintained by giving additional pentobarbital (3 mg/dose) as needed. At the end of the procedure, rats were killed by pentobarbital overdose. Because we used male rats, our results are not necessarily representative of females.

Preparation of rats for Lp measurement. Anesthetized rats were placed on a heating pad to maintain normal body temperature. A midline surgical incision of ~1 cm was made in the abdominal wall, and the mesentery was gently taken out and spread over a pillar. The upper surface of the mesentery was continuously superfused with mammalian Ringer solution (37°C). All the experiments were carried out in straight nonbranched segments of venular microvessels, which were typically 25–35 µm in diameter. All vessels selected had brisk blood flow and were free of white blood cells.

Measurement of Lp of the microvessel wall. Measurements were based on the modified Landis technique, which measures the volume flux of water across the wall of a microvessel perfused via a glass micropipette after occlusion of the vessel. The assumptions and limitations of the measurement have been evaluated in detail (36). The initial transcapillary water flow per unit area of the capillary wall [(Jv/S)0] was measured at predetermined capillary pressures of 30–60 cmH2O. The volume flux per unit surface area of vessel wall (Jv/S) was estimated during single occlusions, lasting ~10 s each, at one constant hydraulic pressure (usually 50 cmH2O) with the assumption that the net effective pressure determining fluid flow (Peff) was equal to the applied hydraulic pressure minus 3.6 cmH2O, the approximate oncotic pressure contributed by BSA in all perfusates (10 mg/ml). Lp was estimated for each occlusion as (Jv/S)/Peff.. All perfusates were mammalian Ringer solution additionally containing serum albumin at 10 mg/ml (Sigma A-4378). Measurements of (Jv/S)0 were made at ~10-min intervals for up to 90 min. Jasplakinolide and cytochalasin D as well as forskolin and rolipram were added to the perfusate and delivered via the micropipette continuously as previously described (48).

Cell culture and test reagents. The immortalized mouse MyEnd cell line was grown in DMEM (Life Technologies; Karlsruhe, Germany) supplemented with 50 U/ml penicillin G, 50 µg streptomycin, and 10% FCS (Biochrom; Berlin, Germany) in a humidified atmosphere (95% air-5% CO2) at 37°C. Generation and characterization have been described before (21, 48). The cultures were used for experiments when grown to confluent monolayers (from day 3 to day 7). Jasplakinolide (Calbiochem) was used at 10 and 0.1 µM. Cytochalasin D (Sigma) at 10 µM. Forskolin and rolipram (both from Sigma) were used at 5 and 10 µM, respectively.

Cytochemistry. MyEnd cells were grown on coverslips coated with gelatin cross-linked with glutaraldehyde (42). After incubation with jasplakinolide, culture medium was removed, and monolayers were fixed for 10 min at room temperature (RT) with 2% formaldehyde (freshly prepared from paraformaldehyde) in PBS (consisting of 137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, and 1.5 mM KH2PO4; pH 7.4). Afterward, monolayers were treated with 0.1% Triton X-100 in PBS for 5 min. After being rinsed with PBS at RT, MyEnd cells were preincubated for 30 min with 10% normal goat serum (NGS) and 1% BSA at RT and incubated for 16 h at 4°C with rat monoclonal antibody 11D4.1 (undiluted hybridoma supernatant) directed to the ectodomain of mouse VE-cadherin (22) or a mouse monoclonal IgM directed against actin (Amersham; Braunschweig, Germany). The monoclonal actin antibody selectively labeled the junction-associated actin filament web. Stress fibers were not labeled by this antibody, which may be due to some kind of steric hindrance of binding. In previous studies, we have shown that this antibody reacts with all three actin isoforms (absorption studies, immunoblotting). After several rinses with PBS (3 x 5 min), monolayers were incubated for 60 min at RT with Cy3-labeled goat anti-rat IgG or Cy3-labeled goat anti-mouse IgG, respectively (both Dianova; Hamburg, Germany; diluted 1:600 in PBS). For visualization of F-actin, some monolayers were incubated with ALEXA-phalloidin (Mobitec; Göttingen, Germany; diluted 1:60 in PBS, 1 h at RT). ALEXA-phalloidin caused bright labeling of stress fibers as well as of the junction-associated belt of actin. The latter structure was less intensively stained and partly covered by stress fibers running parallel to the cell borders. Cells incubated with antibodies or ALEXA-phalloidin were rinsed with PBS (3 x 5 min). Coverslips were mounted on glass slides with 60% glycerol in PBS containing 1.5% n-propyl gallate (Serva; Heidelberg, Germany) as antifading compound.

Transfection of MyEnd cells. Cells were transfected with pEGFP-actin vector (Clontech; Palo Alto, CA) 1 day after being plated in 6-well dishes (Greiner; Frickenhausen, Germany) using effectene transfection reagent (Quiagen; Hilden, Germany). Twenty-four hours later, cells were treated with jasplakinolide and assessed by immunostaining.

Quantification of F-actin content by Triton-soluble/insoluble extraction. MyEnd cells were incubated with jasplakinolide (0.1 and 10 µM) for 60 min. After being rinsed briefly with PBS, cells were incubated in extraction buffer [50 mM 2-(N-morpholino)ethanesulfonic acid (pH 6.8), 25 mM EGTA, 5 mM MgCl2, and 0.5% Triton X-100] containing protease inhibitor solution (leupetin, pepstatin, and aprotinin at 20 µg/ml each, Sigma) for 5 min at 37°C. The extraction buffer was then collected and defined as the Triton-soluble fraction. Cells were then removed from the culture dish using a rubber policeman and dissolved immediately in 10% SDS-containing sample buffer at 95°C for 5 min. This was defined the Triton-insoluble fraction (53). Afterward, samples were subjected to SDS-PAGE and subsequent Western blotting and immunodetection with a mouse monoclonal IgM directed against actin (Amersham; Braunschweig, Germany) using horseradish peroxidase-labeled goat anti-mouse IgG (Dianova; Hamburg, Germany) and the enhanced chemilumeniscence technique (ECL, Amersham). Densitometry was performed by transillumination scanning (Arcus II, Agfa; Mortsel, Belgium) of Western blots, followed by grayscale analysis using Adobe Photoshop 7.0 (Adobe Systems; San Jose, CA).

Quantification of F-actin by phalloidin-binding assay. MyEnd monolayers (untreated or treated with various substances) were fixed at RT with 3% formaldehyde in PBS, rinsed in PBS briefly, and permeabilized with 0.1% (vol/vol) Triton X-100 in PBS for 5 min. Each coverslip was then incubated for 1 h at 37°C with 500 µl (1 µg/ml) phalloidin covalently labeled with tetramethyl-rhodamine isothiocyanate (TRITC). These conditions have been shown to allow complete saturation of binding to cellular F-actin (18, 19). Cells were washed 3 x 1 min in PBS to remove unbound TRITC-phalloidin. TRITC-phalloidin was extracted from cells by two incubation steps for 1 h each with 1 ml methanol at 37°C. For control of complete extraction of TRITC-phalloidin, coverslips were routinely screened by fluorescence microscopy. Methanol supernatants were pooled, centrifuged at 100,000 g for 20 min, and quantified in a fluorescence spectrometer at an excitation wavelength of 540 nm and an emission wavelength of 563 nm.

Recombinant VE-cadherin-Fc. As described before, we used the VE-cadherin-Fc fusion protein consisting of the complete extracellular domain of mouse VE-cadherin (EC1-EC5) fused to the Fc portion of human IgG1, including the hinge region and Ig domains CH2 and CH3 (8, 22). The protein was expressed by stably transfected Chinese hamster ovary (CHO) cells and purified from culture supernatants by affinity chromatography using protein A agarose (Oncogene; Cambridge, MA).

Coating of polystyrene beads. After being vortexed, a 10-µl solution of protein A-coated superparamagnetic polystyrene microbeads (diameter 2.8 µm, Dynabeads; Dynal, Oslo) containing 2 x 109 beads/ml were washed three times using 100 µl of buffer A (100 mM sodium phosphate buffer; pH 8.1). Washing was performed by immobilization of beads for 1 min in a magnetic tube holder (MPC-E-1, Dynal) and reuptake in the corresponding buffer. Washed beads were suspended in 100 µl of 100 mM sodium phosphate buffer (pH 8.1) in Hanks' balanced salt solution (HBSS; GIBCO; Karlsruhe, Germany) containing 10 µg of either VE-cadherin-Fc or the Fc part of human IgG (for control experiments) and allowed to react for 16 h at 4°C under permanent slow overhead rotation to avoid aggregation. After being washed 3 x 5 min in 100 µl of buffer A and 3 x 5 min in buffer B (100 mM sodium borate; pH 9.0), beads were incubated for 45 min at RT in 100 µl of buffer B containing 0.54 mg dimethylpimelimidate dihydrochloride (DMP; Pierce; Rockford, IL) to covalently crosslink protein A and bound Fc parts. After being washed 2 x 5 min in buffer C (100 µl of 0.2 M ethanolamine; pH 8.0), beads were incubated in buffer C for 2 h at RT. Finally, beads were washed 3 x 5 min in HBSS and stored in HBSS at 4°C for up to 8 days under permanent slow overhead rotation to avoid aggregation of beads. The concentration of beads in these stocks was ~1.6 x 108 beads/ ml.

Laser tweezer. As described previously (9), the home-built laser tweezer setup consisted of a Nd:Yag laser (1,064 nm), the beam of which was expanded to fill the back aperture of a high-numerical aperture objective (x100, 1.3 oil, Zeiss), coupled through the epi-illumination port of an Axiovert 135 microscope (Zeiss; Oberkochen, Germany) and reflected to the objective by a dicroic mirror (FT 510, Zeiss). Through all experiments, the laser intensity was 42 mW in the focal plane. Coated beads (10 µl of stock solution) were suspended in 500 µl of culture medium and allowed to interact with MyEnd monolayers for 30 min at 37°C before the initiation of experiments (addition of lethal toxin alone or together with forskolin and rolipram). Beads were considered tightly bound when they resisted laser displacement at the 42-mW setting. For every condition, 100 beads were counted. The percentage of beads resisting laser displacement under various experimental conditions was normalized to control values.

Statistics. Values are expressed as means ± SE. Baseline Lp distributions are non-Gaussian in both frog mesentery capillaries and rat mesentery venules (28, 37). Therefore, we used the nonparametric Mann-Whitney statistic to test for differences in Lp between groups. Possible differences in bead binding and actin content between groups were assessed using the unpaired Student's t-test. Statistical significance was assumed for P < 0.05.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Effects of jasplakinolide on microvessel Lp are dose dependent. When individual rat venular microvessels were perfused with the F-actin-promoting drug jasplakinolide at a low dose (0.1 µM), Lp did not significantly change during 80 min of perfusion and was 3.3 ± 2.3 x 10–7 cm·s–1·cmH2O–1 after 30 min compared with a control value of 2.1 ± 1.0 x 10–7 cm·s–1·cmH2O–1 (n = 5). In two experiments, measurements were carried out for 110 min without any change of Lp (not shown). In contrast, when jasplakinolide was applied at 10 µM, Lp rose from 1.06 ± 0.2 x 10–7 to 99.6 ± 29.9 x 10–7 cm·s–1·cmH2O–1 after 30 min (n = 5; Fig. 1A). This effect was comparable to the effect of the actin-depolymerizing agent cytochalasin D (Fig. 1B), which increased Lp from 1.7 ± 0.3 x 10–7 to 86.1 ± 16.5 x 10–7 cm·s–1·cmH2O–1 after 30 min of perfusion (n = 5). Moreover, the onsets of the Lp increase induced by jasplakinolide (10 µM) and cytochalasin D were not statistically different. After 10 min of perfusion with jasplakinolide, Lp was 26.8 ± 10.3 x 10–7 compared with 22.2 ± 4.7 x 10–7 cm·s–1·cmH2O–1 in the experiments using cytochalasin D.



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  		Fig. 1. Effect of jasplakinolide (Jasp) and cyotochalasin D (Cyto D) on microvessel hydraulic permeability (Lp). A: data from a representative vessel perfused with 10 µM jasplakinolide demonstrating the increase in Lp during 30 min of perfusion. B: venule perfused with cytochalasin D (10 µM). Both cytochalasin D and high-dose jasplakinolide drastically increased microvessel Lp (n = 5 for each condition).

 
Jasplakinolide induced intercellular gap formation in cultured endothelial monolayers in dose-dependent manner. To test the dose-dependent effects of jasplakinolide on endothelial monolayer integrity, cultured mouse microvascular MyEnd cells were treated with jasplakinolide and immunostained for VE-cadherin and actin (n = 9). Endothelial cells in control monolayers displayed a continuous beltlike immunolabeling of VE-cadherin (Fig. 2A). Immunostaining for actin resulted in a similar beltlike pattern (Fig. 2C). In addition (see MATERIALS AND METHODS), ALEXA-phalloidin visualized numerous stress fibers throughout the cytoplasm (Fig. 2B). After treatment with jasplakinolide (0.1 µM, 60 min), the immunostaining for VE-cadherin and the junction-associated belt of actin were not affected (Fig. 2, D and F). However, ALEXA-phalloidin staining of both the junctional belt and stress fibers was strongly reduced due to competitive binding of jasplakinolide to F-actin, which blocks the binding sites for phalloidin (Fig. 2E) (11). When jasplakinolide was applied at 10 µM, ALEXA-phalloidin staining was completely abolished, suggesting almost total saturation of jasplakinolide binding sites on F-actin (Fig. 2H). Although the total contents of F-actin had increased under these conditions (see below), the junction-associated VE-cadherin and actin became locally interrupted. Additionally, intercellular gaps were found at sites where either immunostaining for actin or VE-cadherin were interrupted (Fig. 2, G and I, arrows). These experiments indicate that hyperpolymerization of actin by treatment with 10 µM jasplakinolide results in local loss of actin from the junctional belt and redistribution of actin to other cellular sites (see below).



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  		Fig. 2. Effects of jasplakinolide on endothelial monolayer integrity and junction-associated actin filament web were dose dependent. Myocardial endothelial (MyEnd) cells were stained for vascular endothelial (VE-)cadherin (A, D, and G) and for F-actin using either ALEXA-phalloidin (B, E, and H) or an antibody directed to actin (C, F, and I) (n = 5 for each condition). Control cells display continuous distribution of VE-cadherin along cell borders (A) accompanied by a continuous junction-associated actin filament web (C) as well as a dense cytoplasmatic system of stress fibers (B). Treatment with 0.1 µM jasplakinolide did not affect the distribution of VE-cadherin and junction-associated actin (D and F) but reduced phalloidin staining, indicating competitive binding of jasplakinolide to phalloidin binding sites on actin filaments (E). High doses of jasplakinolide (10 µM) saturated all phalloidin binding sites (H) and induced intercellular gap formation coinciding with fragmentations of VE-cadherin and actin immunostaining (arrows in G and I). Scale bars = 20 µm for all images.

 
Jasplakinolide-induced intercellular gaps appear at sites where the junction-associated actin system is fragmented. As outlined above, visualization of changes in the morphology of the actin cytoskeleton induced by jasplakinolide was hampered by the fact that jasplakinolide competitively binds to F-actin at the same binding sites as ALEXA-phalloidin and thereby ALEXA-phalloidin staining was abolished by treatment with high-dose jasplakinolide (Fig. 2H). Moreover, the monoclonal actin antibody did not label stress fibers and thus did not allow staining of all subsets of actin filaments. To use a different approach that was independent of labeling efficiency to visualize the effects of jasplakinolide on the actin cytoskeleton, we transfected MyEnd cells with enhanced green fluorescent protein (EGFP)-actin. However, transfection efficiency of endothelial cells is not as good as known from other cell types, and thus only some cell groups can be evaluated. These groups are indicated by the green color of EGFP-actin and are surrounded by nontransfected cells that are either not visible (Fig. 3, A–D) or only revealed by VE-cadherin staining (for example, a large cell group in the bottom left corner of Fig. 3F). However, only confluent monolayers were used for the jasplakinolide challenge. Controls displayed diffuse fluorescence of the cytoplasm, most probably reflecting the distribution of G-actin (Fig. 3, A and B). Low-dose jasplakinolide (0.1 µM) did not change this pattern of EGFP-actin distribution (not shown). However, treatment with high-dose jasplakinolide (10 µM) resulted in the formation of strongly fluorescent aggregates of actin throughout the cell body. The junction-associated actin filament system became locally interrupted (arrows in Fig. 3, C and D), whereas it was well preserved in other localizations (arrowheads in Fig. 3D). In addition, the diffuse EGFP-actin fluorescence of the cytoplasm was strongly reduced, possibly reflecting the decrease of G-actin induced by jasplakinolide (see below). Intercellular gaps mostly were found at sites where the VE-cadherin immunostaining and the junction-associated actin belt were fragmented (compare localization of arrows in Fig. 3, C and E as well as D and F), whereas distribution of VE-cadherin was continuous where the junctional actin belt was well preserved (compare arrowheads in Fig. 3, D and F). It has to be emphasized that because of transfection efficiency, no quantitative but only qualitative information can be obtained from the presented data.



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  		Fig. 3. Jasplakinolide-induced intercellular gap formation coincided with sites where the junction-associated actin web is fragmented. MyEnd cells were transfected with enhanced green fluorescent protein (EGFP)-actin (A–F) and additionally immunostained for VE-cadherin (E and F) (n = 3 for each condition). Controls (A and B) displayed thin stress fibers throughout the cytoplasm and a brightly fluorescent background, most likely reflecting the distribution of G-actin. Treatment with 10 µM jasplakinolide induced profound disorganization of the actin cytoskeleton including fragmentations of the junction-associated actin filament web (C and D). These fragmentations typically coincided with intercellular gap formation as well as with disruptions of VE-cadherin immunostaining (E and F). Scale bars = 20 µm for all images.

 
Effect of jasplakinolide on F-actin content in MyEnd cells was dose dependent. To test whether the jasplakinolide-induced disorganization of the actin filament system correlated to changes of the ratio of F-actin and G-actin, we performed Triton X-100 extractions of MyEnd cells after treatment with jasplakinolide. The phalloidin-binding assay that we used to measure F-actin content in response to cytochalasin D in this study (see below) and previous studies (48) could not be applied after treatment with jasplakinolide due to competitive binding of phalloidin and jasplakinolide for same binding sites on actin filaments. Therefore, relative amounts of actin in Triton-resistant (F-actin) and Triton-soluble (G-actin) fractions were assayed by immunoblotting (29). Figure 4 shows a representative experiment of three total experiments. Low-dose jasplakinolide (0.1 µM) had no effect on the Triton solubility in both fractions (Fig. 4A). In controls (lanes 1 and 3) as well as in cells treated with the low dose of jasplakinolide (lanes 2 and 4), actin was found in the Triton-soluble and Triton-insoluble fractions, indicating that actin was present as both monomeric (G-actin) and F-actin, respectively. In contrast, high-dose jasplakinolide (10 µM) strongly reduced actin in the Triton-soluble fraction by ~96% compared with control levels and increased actin in the Triton-resistant fraction by 34%, suggesting that under these conditions almost all actin is present as F-actin (Fig. 4B).



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  		Fig. 4. High-dose jasplakinolide increased actin in the Triton-resistant fraction in MyEnd cells. Western blots show Triton-soluble (sol., lanes 1 and 2) and Triton-resistant (res., lanes 3 and 4) fractions of actin from MyEnd cells after treatment with low-dose (0.1 µM; A) or high-dose (10 µM; B) jasplakinolide. Low doses of jasplakinolide had no effect on the relative amount of actin in the Triton-soluble (G-actin) and Triton-resistant (F-actin) fractions. High-dose jasplakinolide reduced actin in the soluble fraction and increased actin in the resistant fraction, indicating increased actin polimerization (n = 3).

 
cAMP did not block the effect of cytochalsin D on microvessel Lp in vivo and gap formation in vitro. Our study demonstrates that maintenance of endothelial barrier functions depends on the integrity of the actin filament system. Because previous studies have reported that cAMP, which is well known to protect endothelial barrier functions, also is involved in the regulation of the endothelial actin cytoskeleton (10, 24, 27, 33, 46), we investigated the effect of cAMP on cytochalasin D-induced endothelial barrier breakdown. As shown above in mesenteric microvessels perfused with cytochalasin D (10 µM), Lp drastically increased during the first 30 min of perfusion (Fig. 1B). Increased cAMP by forskolin (5 µM) and rolipram (10 µM) did not significantly reduce the cytochalasin D-induced Lp increase. Lp increased from 0.6 ± 0.2 x 10–7 to 98.4 ± 6.7 x 10–7 cm·s–1·cmH2O–1 after 40 min of perfusion (Fig. 5, compare with Fig. 1B). Thus cAMP did not seem to stabilize F-actin in vivo. To test the effect of increased cAMP on endothelial F-actin content, we measured the F-actin content of MyEnd cells using the phalloidin-binding assay. In contrast to cytochalasin D, which significantly reduced F-actin by 22 ± 0.7% (n = 3), increased cAMP had no significant effect on F-actin content (108 ± 1.4%, n = 4). We next investigated whether increased cAMP would modulate the cytochalsin D-induced breakdown of adherens junctions in cultured endothelial cells. Cytochalasin D (10 µM) induced formation of large intercellular gaps in MyEnd monolayers, which coincided with fragmentations of VE-cadherin immunostaining (Fig. 6C). Also, profound alterations of the actin filament system became visible. Incubation with cytochalasin D resulted in complete loss of stress fibers as well as fragmentations (arrows) of the junction-associated actin filament system (Fig. 6D) compared with control MyEnd cells (Fig. 6B). Forskolin (5 µM) and rolipram (10 µM) neither blocked cytochalasin D-induced gap formation nor reorganization and displacement of VE-cadherin and F-actin (arrows in Fig. 6, E and F). Similarly, forskolin and rolipram did not block gap formation induced by jasplakinolide (not shown). These experiments also indicate that in vitro cAMP is not protecting endothelial barrier properties by stabilizing F-actin against cytochalasin D-induced depolymerization.



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  		Fig. 5. cAMP did not block the cytochalasin D-induced Lp increase. Shown is a graph from a single experiment showing perfusion with cytochalasin D (10 µM) in presence of forskolin (5 µM) and rolipram (10 µM) (F/R). Increased cAMP had no significant effect on the cytochalasin D-induced permeability increase (n = 5).

 


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  		Fig. 6. In contrast to jasplakinolide, cAMP did not block cytochalasin D-induced effects on endothelial cell morphology. MyEnd cells were immunostained for VE-cadherin (A, C, and E) and stained for F-actin using ALEXA-phalloidin (B, D, and F) (n = 5 for each condition). In contrast to controls (A), VE-cadherin staining (C) and the junction-associated actin cytoskeleton (D) were fragmented after treatment with cytochalasin D (10 µm, 60 min). The effects of cytochalasin D were not blocked by simultaneous treatment with forskolin (5 µM) and rolipram (10 µM) (E and F). Scale bars = 20 µm for all images.

 
In contrast to cAMP, jasplakinolide increased VE-cadherin-mediated adhesion by stabilizing F-actin. Our previous studies demonstrated that jasplakinolide, when applied at low doses (0.1 µM), stabilized F-actin because it blocked the effect of the actin-depolymerizing drug cytochalasin D on microvessel Lp as well as on adhesion of VE-cadherin-coated microbeads. To see whether increased cAMP enhances endothelial barrier function by regulating VE-cadherin-mediated adhesion via comparable mechanisms as low doses (0.1 µM) of jasplakinolide, we carried out laser tweezer studies using VE-cadherin-coated microbeads (Fig. 7). Beads were allowed to settle on the surface of MyEnd cells for 30 min to induce formation of cell-to-bead contacts (9, 49). Beads resisting displacement by laser beam were counted as bound (control value). As shown previously, depolymerization of F-actin by cytochalasin D (10 µM, 60 min) resulted in significant reduction of bead binding to 48 ± 1% (n = 5). As demonstrated previously, low-dose jasplakinolide (0.1 µM) prevented the cytochalasin D-induced Lp increase (49) by stabilization of F-actin without increasing the overall cellular F-actin content (Fig. 4A). Under the same conditions, VE-cadherin-mediated adhesion was increased to 139 ± 6% of control levels after treatment with jasplakinolide (0.1 µM, 60 min, n = 5). When cytochalasin D was applied in the presence of jasplakinolide, the cytochalasin D-induced loss of bead binding was completely abolished (130 ± 4% bound beads, n = 5). These results again indicate that stabilization of F-actin is sufficient to enhance VE-cadherin-mediated adhesion.



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  		Fig. 7. Jasplakinolide blocked the cytochalasin D-induced reduction of VE-cadherin-coated bead binding, whereas cAMP had no effect. VE-cadherin-coated beads were allowed to settle on the surface of MyEnd cells for 30 min. Beads resisting displacement by laser beam were counted as bound (control). Jasplakinolide completely blocked the cytochalasin D-induced reduction of bead adhesion, whereas forskolin (5 µM) and rolipram (10 µM) had no effect.

 
Increasing intracellular cAMP by forskolin (5 µM) and rolipram (10 µM) had no effect on number of bound beads and did not block the cytochalasin D-induced reduction of bead binding because the number of bound beads dropped to 41 ± 5% of controls (n = 5 in both groups). These data suggest that cAMP does enhance endothelial barrier functions by neither stabilizing F-actin nor direct regulation of VE-cadherin-mediated adhesion.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
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 REFERENCES
 
Organization of the actin cytoskeleton is critical for maintenance of endothelial barrier properties. Our study demonstrates that maintenance of endothelial barrier properties depends critically on an undisturbed equilibrium between G-actin and F-actin. Both depolymerization of actin by cytochalasin D and hyperpolymerization of actin by high-dose jasplakinolide increased microvascular permeability in vivo. In contrast, low-dose jasplakinolide under conditions where no change in F-actin content was detectable increased VE-cadherin-mediated adhesion and antagonized the effect of actin depolymerization on vascular permeability. On the other hand, elevated cAMP did not prevent barrier breakdown by cytochalasin D and thus was found not to enhance barrier functions by stabilization of F-actin. In the following, we will first explain the rationale to study the actin cytoskeleton as an important actuator of endothelial barrier function and then focus on the role of the actin filament system in endothelial barrier regulation in vivo and in vitro.

As early as 1961, the permeability increase in response to inflammatory mediators was shown to be caused by the formation of paracellular gaps between endothelial cells (34, 35). However, the mechanisms that actually cause formation of intercellular gaps and increase permeability in vivo are not completely understood. This is in part due to the problem that large intercellular gaps, as seen between cultured endothelium after treatment with various stimuli, are not necessarily representative of the gaps that may be responsible for measurable permeability increases in vivo (49). An important role of both myosin-dependent contraction (44) and proteins that regulate contraction, such as the Rho family (4, 52), in the regulation of cultured endothelial barrier functions has been favored in the past. However, recent studies using rat mesenteric microvessels indicate that myosin-dependent contraction is not needed to increase permeability in response to various stimuli (3, 48) and favor the hypothesis that contractile forces might be increased under cell culture conditions (15). Indeed, it has been demonstrated that passive recoil without additional contractile forces are sufficient for endothelial gap formation (6). However, it is reasonable that reorganizations of the cortical actin system would affect the integrity of intercellular junctions because both adherens and tight junctions are linked to actin filaments (45). Although the tight junction is the main permeability limiting structure, the adherens junction provides the mechanical strength to intercellular adhesion (14). Molecular characterization of binding properties of VE-cadherin revealed an extremely low affinity and lifetime of single cadherin bonds (8) necessitating tight anchorage of cadherins to the cytoskeleton to reduce lateral mobility and increase local concentration in the plane of the lipid bilayer thereby increasing number of bonds and improving binding strength (7).

Our results demonstrate that actin hyperpolymerization by jasplakinolide increases microvascular permeability to extents comparable to those due to depolymerization of F-actin induced by cytochalasin D. On the other hand, low-dose jasplakinolide at least partially blocked the cytochalasin D-induced Lp increase, suggesting that jasplakinolide stabilized actin filaments under these conditions and protected them against depolymerization (49). However, treatment with low-dose jasplakinolide alone did not reduce baseline Lp, indicating that, in the resting microvessel endothelium, depolymerization and repolymerization of actin are in an equilibrated state. Taken together, these results indicate that both increased actin polymerization and depolymerization interfere with endothelial barrier functions in vivo, whereas stabilization of F-actin without favoring polymerization seems to be protective for endothelial barrier functions, at least over the short period of time covered by our experiments. These data are consitent with previous studies using drugs that at least in part seemed to increase microvascular permeability by affecting the actin cytoskeleton. A-23187, a calcium ionophore that increases cytoplasmic calcium levels, induced interendothelial gaps and increased microvessel permeability in rats and frogs (39). Recent studies pointed to the possibility that A-23187 reduces endothelial barrier functions by calcium-induced actin depolymerization (9, 30). Moreover, Clostridium botulinum C2 toxin has been shown to induce lung edema in rabbit lungs by ADP-ribosylation of G-actin (17). These studies suggest an important role of actin filament regulation in endothelial barrier functions in vivo.

In cultured endothelial cells, our results demonstrate that both actin depolymerization and hyperpolymerization lead to formation of large intercellular gaps. This is indicated by the formation of intercellular gaps (Figs. 2, 3, and 6) under conditions where the amount of F-actin was either decreased by cytochalasin D or increased after treatment with jasplakinolide (10 µM). Additionally, these images suggest that gaps coincided with sites where the junction-associated actin system was disrupted, further conforming to the concept that actin filaments are important for stabilizing intercellular junctions. VE-cadherin was also lacking at these sites suggesting that an intact actin filament system might be important for stabilizing VE-cadherin at intercellular junctions. However, no quantitative results can be obtained from these observations. This view that actin is required for stabilizing VE-cadherin-containing contacts was further supported by our previous studies using the laser tweezer, which showed a significant reduction of VE-cadherin-mediated adhesion in response to cytochalasin D- or A-23187-induced depolymerization of actin (9). In contrast, low-dose jasplakinolide, which had no effect on monolayer morphology or F-actin content in cultured endothelial cells, significantly increased VE-cadherin-mediated adhesion. These results conform to earlier studies that reported that stabilizing F-actin by phalloidin and related toxins enhanced endothelial barrier functions in vitro (5, 40). Taken together, these studies indicate that actin filaments are required for maintenance of endothelial barrier functions possibly by forming a scaffold for anchorage of intercellular junctional proteins and support the hypothesis that control of VE-cadherin-mediated adhesion is involved in endothelial barrier regulation. However, it has to be emphasized that the mechanisms contributing to the formation of large intercellular gaps in vitro do not necessarily reflect the underlying mechanisms of increased permeability in vivo. Therefore, further parallel studies investigating microvascular endothelium in vivo and in vitro are required to determine the mechanisms that are involved in endothelial barrier regulation in both systems.

cAMP does not enhance endothelial barrier functions by stabilizing F-actin. Our study strengthened the view that a well-balanced regulation of the actin filament system is required for endothelial barrier function and emphasizes the important role of the actin cytoskeleton as one possible actuator in the permeability response to inflammatory mediators. It is well established that cAMP enhances endothelial barrier functions in vivo and in vitro (1–3, 12, 13, 25, 26, 31, 38, 43, 44). Several mechanisms such as reduction of myosin light chain phosphorylation (54), regulation of vasodilator-stimulated phosphoprotein (13), and inhibition of Rho A (41) have been reported to be involved in endothelial barrier regulation by cAMP. Moreover, it has been reported that cAMP participates in the regulation of the actin cytoskeleton, suggesting that these mechanisms might play a role in the cAMP-mediated maintenance of endothelial barrier function (10, 24, 27, 33, 46). However, the studies have all been performed using cultured endothelial cells mainly derived from macrovascular vessels. Thus the relevant mechanisms involved in cAMP-dependent endothelial barrier regulation in microvessels remain unclear. Previous studies from our lab demonstrated that myosin-dependent contraction is not primarily important for endothelial barrier regulation in vivo and indicated that cAMP is involved in tight junction regulation (1–3, 48). In the present study, we focused on the actin cytoskeleton to find out whether cAMP would stabilize F-actin by comparable mechanisms as jasplakinolide. In contrast to jasplakinolide, cAMP neither prevented the cytochalasin D-induced permeability increase in vivo nor prevented intercellular gap formation and the reduction of VE-cadherin-mediated adhesion in vitro. Moreover, increased cAMP did not significantly affect the content of F-actin in cultured endothelial cells. These results do not support the hypothesis that cAMP stabilizes F-actin in microvascular endothelium in vivo and in vitro. However, we cannot exclude the possibility that cytochalasin D does not act via the same mechanisms as intracellular actin regulating proteins even though it binds to the barbed ends like is typical for other capping proteins (47). Moreover, it is possible that cAMP would modulate the effect of a less powerful mediator than cytochalasin D. In a parallel study, we found that cAMP regulates the GTPase Rac-1 and thus more likely mediates its barrier-promoting effects at the level of signaling events that control the linkage of adhesion molecules to the actin cytoskeleton. Future studies will have to elucidate mechanisms involved in the regulation of actin dynamics and in the regulation of endothelial adhesion proteins in response to inflammatory mediators.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This study was supported in part by National Heart, Lung, and Blood Institute Grants HL-44485 and HL-28607 and by Deutsche Forschungsgemeinschaft Grant SFB 487, TP B5.


   ACKNOWLEDGMENTS
 
We are grateful to Gabriele Lang and Joyce Lenz for skillful technical assistance.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Waschke, Institute of Anatomy and Cell Biology, Julius-Maximilians-Univ., Koellikerstrasse 6, D-97070 Würzburg, Germany (E-mail: jens.waschke{at}mail.uni-wuerzburg.de)

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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
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 REFERENCES
 

  1. Adamson RHJY, Lenz JF, and Curry FE. Basal hydraulic conductivity regulation in single perfused rat venules by cAMP (Abstract). FASEB J 15: A44, 2001.
  2. Adamson RH, Liu B, Fry GN, Rubin LL, and Curry FE. Microvascular permeability and number of tight junctions are modulated by cAMP. Am J Physiol Heart Circ Physiol 274: H1885–H1894, 1998.[Abstract/Free Full Text]
  3. Adamson RH, Zeng M, Adamson GN, Lenz JF, and Curry FE. PAF- and bradykinin-induced hyperpermeability of rat venules is independent of actin-myosin contraction. Am J Physiol Heart Circ Physiol 285: H406–H417, 2003.[Abstract/Free Full Text]
  4. Aepfelbacher M and Essler M. Disturbance of endothelial barrier function by bacterial toxins and atherogenic mediators: a role for Rho/Rho kinase. Cell Microbiol 3: 649–658, 2001.[CrossRef][ISI][Medline]
  5. Alexander JS, Hechtman HB, and Shepro D. Phalloidin enhances endothelial barrier function and reduces inflammatory permeability in vitro. Microvasc Res 35: 308–315, 1988.[CrossRef][ISI][Medline]
  6. Baldwin AL and Thurston G. Mechanics of endothelial cell architecture and vascular permeability. Crit Rev Biomed Eng 29: 247–278, 2001.[ISI][Medline]
  7. Baumgartner W and Drenckhahn D. Plasmalemmal concentration and affinity of mouse vascular endothelial cadherin, VE-cadherin. Eur Biophys J 31: 532–538, 2002.[CrossRef][ISI][Medline]
  8. Baumgartner W, Hinterdorfer P, Ness W, Raab A, Vestweber D, Schindler H, and Drenckhahn D. Cadherin interaction probed by atomic force microscopy. Proc Natl Acad Sci USA 97: 4005–4010, 2000.[Abstract/Free Full Text]
  9. Baumgartner W, Schutz GJ, Wiegand J, Golenhofen N, and Drenckhahn D. Cadherin function probed by laser tweezer and single molecule fluorescence in vascular endothelial cells. J Cell Sci 116: 1001–1011, 2003.[Abstract/Free Full Text]
  10. Birukova AA, Liu F, Garcia JG, and Verin AD. Protein kinase A attenuates endothelial cell barrier dysfunction induced by microtubule disassembly. Am J Physiol Lung Cell Mol Physiol 287: L86–L93, 2004.[Abstract/Free Full Text]
  11. Bubb MR, Senderowicz AM, Sausville EA, Duncan KL, and Korn ED. Jasplakinolide, a cytotoxic natural product, induces actin polymerization and competitively inhibits the binding of phalloidin to F-actin. J Biol Chem 269: 14869–14871, 1994.[Abstract/Free Full Text]
  12. Carson MR, Shasby SS, and Shasby DM. Histamine and inositol phosphate accumulation in endothelium: cAMP and a G protein. Am J Physiol Lung Cell Mol Physiol 257: L259–L264, 1989.[Abstract/Free Full Text]
  13. Comerford KM, Lawrence DW, Synnestvedt K, Levi BP, and Colgan SP. Role of vasodilator-stimulated phosphoprotein in PKA-induced changes in endothelial junctional permeability. FASEB J 16: 583–585, 2002.[Free Full Text]
  14. Corada M, Mariotti M, Thurston G, Smith K, Kunkel R, Brockhaus M, Lampugnani MG, Martin-Padura I, Stoppacciaro A, Ruco L, McDonald DM, Ward PA, and Dejana E. Vascular endothelial-cadherin is an important determinant of microvascular integrity in vivo. Proc Natl Acad Sci USA 96: 9815–9820, 1999.[Abstract/Free Full Text]
  15. Curry FE. Microvascular injury: mechanisms and modulation. Int J Angiol 11: 1–6, 2002.[CrossRef]
  16. Drenckhahn D and Ness W. The Endothelial Contractile Cytoskeleton. Stuttgart, Germany: Schauttauer, 1997.
  17. Ermert L, Bruckner H, Walmrath D, Grimminger F, Aktories K, Suttorp N, Duncker HR, and Seeger W. Role of endothelial cytoskeleton in high-permeability edema due to botulinum C2 toxin in perfused rabbit lungs. Am J Physiol Lung Cell Mol Physiol 268: L753–L761, 1995.[Abstract/Free Full Text]
  18. Faulstich H, Trischmann H, and Mayer D. Preparation of tetramethylrhodaminyl-phalloidin and uptake of the toxin into short-term cultured hepatocytes by endocytosis. Exp Cell Res 144: 73–82, 1983.[CrossRef][ISI][Medline]
  19. Franke RP, Grafe M, Schnittler H, Seiffge D, Mittermayer C, and Drenckhahn D. Induction of human vascular endothelial stress fibres by fluid shear stress. Nature 307: 648–649, 1984.[CrossRef][Medline]
  20. Gallo G, Yee HF Jr, and Letourneau PC. Actin turnover is required to prevent axon retraction driven by endogenous actomyosin contractility. J Cell Biol 158: 1219–1228, 2002.[Abstract/Free Full Text]
  21. Golenhofen N, Ness W, Wawrousek EF, and Drenckhahn D. Expression and induction of the stress protein alpha-B-crystallin in vascular endothelial cells. Histochem Cell Biol 117: 203–209, 2002.[CrossRef][ISI][Medline]
  22. Gotsch U, Borges E, Bosse R, Boggemeyer E, Simon M, Mossmann H, and Vestweber D. VE-cadherin antibody accelerates neutrophil recruitment in vivo. J Cell Sci 110: 583–588, 1997.[Abstract]
  23. Hall A and Nobes CD. Rho GTPases: molecular switches that control the organization and dynamics of the actin cytoskeleton. Philos Trans R Soc Lond B Biol Sci 355: 965–970, 2000.[CrossRef][ISI][Medline]
  24. Hastie LE, Patton WF, Hechtman HB, and Shepro D. H2O2-induced filamin redistribution in endothelial cells is modulated by the cyclic AMP-dependent protein kinase pathway. J Cell Physiol 172: 373–381, 1997.[CrossRef][ISI][Medline]
  25. He P and Curry FE. Differential actions of cAMP on endothelial [Ca2+]i and permeability in microvessels exposed to ATP. Am J Physiol Heart Circ Physiol 265: H1019–H1023, 1993.[Abstract/Free Full Text]
  26. He P, Zeng M, and Curry FE. Dominant role of cAMP in regulation of microvessel permeability. Am J Physiol Heart Circ Physiol 278: H1124–H1133, 2000.[Abstract/Free Full Text]
  27. Hippenstiel S, Witzenrath M, Schmeck B, Hocke A, Krisp M, Krull M, Seybold J, Seeger W, Rascher W, Schutte H, and Suttorp N. Adrenomedullin reduces endothelial hyperpermeability. Circ Res 91: 618–625, 2002.[Abstract/Free Full Text]
  28. Huxley VH and Rumbaut RE. The microvasculature as a dynamic regulator of volume and solute exchange. Clin Exp Pharmacol Physiol 27: 847–854, 2000.[CrossRef][ISI][Medline]
  29. Kouloukoussa M, Aleporou-Marinou V, Angelopoulou B, Trougakos IP, Panagopoulou E, Kittas C, and Marinos E. Phorbol myristate acetate induces changes on F-actin and vinculin content in immature rat Sertoli cells. Tissue Cell 36: 149–155, 2004.[CrossRef][ISI][Medline]
  30. Kuhne W, Besselmann M, Noll T, Muhs A, Watanabe H, and Piper HM. Disintegration of cytoskeletal structure of actin filaments in energy-depleted endothelial cells. Am J Physiol Heart Circ Physiol 264: H1599–H1608, 1993.[Abstract/Free Full Text]
  31. Langeler EG and van Hinsbergh VW. Norepinephrine and iloprost improve barrier function of human endothelial cell monolayers: role of cAMP. Am J Physiol Cell Physiol 260: C1052–C1059, 1991.[Abstract/Free Full Text]
  32. Lee E, Shelden EA, and Knecht DA. Formation of F-actin aggregates in cells treated with actin stabilizing drugs. Cell Motil Cytoskeleton 39: 122–133, 1998.[CrossRef][ISI][Medline]
  33. Liu F, Verin AD, Borbiev T, and Garcia JG. Role of cAMP-dependent protein kinase A activity in endothelial cell cytoskeleton rearrangement. Am J Physiol Lung Cell Mol Physiol 280: L1309–L1317, 2001.[Abstract/Free Full Text]
  34. Majno G and Palade GE. Studies on inflammation. I. The effect of histamine and serotonin on vascular permeability: an electron microscopic study. J Biophys Biochem Cytol 11: 571–605, 1961.[Abstract/Free Full Text]
  35. Majno G, Palade GE, and Schoefl GI. Studies on inflammation. II. The site of action of histamine and serotonin along the vascular tree: a topographic study. J Biophys Biochem Cytol 11: 607–626, 1961.[Abstract/Free Full Text]
  36. Michel CC and Curry FE. Microvascular permeability. Physiol Rev 79: 703–761, 1999.[Abstract/Free Full Text]
  37. Michel CC, Mason JC, Curry FE, Tooke JE, and Hunter PJ. A development of the Landis technique for measuring the filtration coefficient of individual capillaries in the frog mesentery. Q J Exp Physiol Cogn Med Sci 59: 283–309, 1974.[Abstract/Free Full Text]
  38. Moy AB, Bodmer JE, Blackwell K, Shasby S, Kamath A, and Shasby DM. Cyclic adenosine monophosphate protects endothelial barrier function independent of inhibiting 20-kd myosin light chain-dependent tension development. Chest 116: 33S, 1999.[Free Full Text]
  39. Neal CR and Michel CC. Transcellular gaps in microvascular walls of frog and rat when permeability is increased by perfusion with the ionophore A23187. J Physiol 488: 427–437, 1995.[ISI][Medline]
  40. Phillips PG, Lum H, Malik AB, and Tsan MF. Phallacidin prevents thrombin-induced increases in endothelial permeability to albumin. Am J Physiol Cell Physiol 257: C562–C567, 1989.[Abstract/Free Full Text]
  41. Qiao J, Huang F, and Lum H. PKA inhibits RhoA activation: a protection mechanism against endothelial barrier dysfunction. Am J Physiol Lung Cell Mol Physiol 284: L972–L980, 2003.[Abstract/Free Full Text]
  42. Schnittler HJ, Franke RP, Akbay U, Mrowietz C, and Drenckhahn D. Improved in vitro rheological system for studying the effect of fluid shear stress on cultured cells. Am J Physiol Cell Physiol 265: C289–C298, 1993.[Abstract/Free Full Text]
  43. Stelzner TJ, Weil JV, and O'Brien RF. Role of cyclic adenosine monophosphate in the induction of endothelial barrier properties. J Cell Physiol 139: 157–166, 1989.[CrossRef][ISI][Medline]
  44. Suttorp N, Weber U, Welsch T, and Schudt C. Role of phosphodiesterases in the regulation of endothelial permeability in vitro. J Clin Invest 91: 1421–1428, 1993.[ISI][Medline]
  45. Vestweber D. Molecular mechanisms that control endothelial cell contacts. J Pathol 190: 281–291, 2000.[CrossRef][ISI][Medline]
  46. Vischer UM, Barth H, and Wollheim CB. Regulated von Willebrand factor secretion is associated with agonist-specific patterns of cytoskeletal remodeling in cultured endothelial cells. Arterioscler Thromb Vasc Biol 20: 883–891, 2000.[Abstract/Free Full Text]
  47. Wakatsuki T, Schwab B, Thompson NC, and Elson EL. Effects of cytochalasin D and latrunculin B on mechanical properties of cells. J Cell Sci 114: 1025–1036, 2001.[Abstract]
  48. Waschke J, Baumgartner W, Adamson RH, Zeng M, Aktories K, Barth H, Wilde C, Curry FE, and Drenckhahn D. Requirement of Rac activity for maintenance of capillary endothelial barrier properties. Am J Physiol Heart Circ Physiol 286: H394–H401, 2004.[Abstract/Free Full Text]
  49. Waschke J, Drenckhahn D, Adamson RH, and Curry FE. The role of adhesion and contraction in Rac 1-regulated endothelial barrier function in vivo and in vitro. Am J Physiol Heart Circ Physiol 287: H704–H711, 2004.[Abstract/Free Full Text]
  50. Watanabe N and Mitchison TJ. Single-molecule speckle analysis of actin filament turnover in lamellipodia. Science 295: 1083–1086, 2002.[Abstract/Free Full Text]
  51. Welch MD and Mullins RD. Cellular control of actin nucleation. Annu Rev Cell Dev Biol 18: 247–288, 2002.[CrossRef][ISI][Medline]
  52. Wojciak-Stothard B and Ridley AJ. Rho GTPases and the regulation of endothelial permeability. Vasc Pharmacol 39: 187–199, 2002.
  53. Wright TJ, Leach L, Shaw PE, and Jones P. Dynamics of vascular endothelial-cadherin and beta-catenin localization by vascular endothelial growth factor-induced angiogenesis in human umbilical vein cells. Exp Cell Res 280: 159–168, 2002.[CrossRef][ISI][Medline]
  54. Wysolmerski RB and Lagunoff D. Regulation of permeabilized endothelial cell retraction by myosin phosphorylation. Am J Physiol Cell Physiol 261: C32–C40, 1991.[Abstract/Free Full Text]



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