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Am J Physiol Cell Physiol 294: C178-C188, 2008. First published November 7, 2007; doi:10.1152/ajpcell.00273.2007
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VASCULAR BIOLOGY

The role of VASP in regulation of cAMP- and Rac 1-mediated endothelial barrier stabilization

N. Schlegel ,1,* S. Burger,1,* N. Golenhofen,1 U. Walter,2 D. Drenckhahn,1 and J. Waschke1

1Institute of Anatomy and Cell Biology and 2Institute for Clinical Biochemistry and Pathobiochemistry, University of Würzburg, Würzburg, Germany

Submitted 27 June 2007 ; accepted in final form 2 November 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Regulation of actin dynamics is critical for endothelial barrier functions. We provide evidence that the actin-binding protein vasodilator-stimulated phosphoprotein (VASP) is required for endothelial barrier maintenance. Baseline permeability was significantly increased in VASP-deficient (VASP–/–) microvascular myocardial endothelial cells (MyEnd) in the absence of discernible alterations of immunostaining for adherens and tight junctions. We tested whether VASP is involved in the endothelium-stabilizing effects of cAMP or Rac 1. Forskolin and rolipram (F/R) to increase cAMP and cytotoxic necrotizing factor 1 (CNF-1) to activate Rac 1 were equally efficient to stabilize barrier functions in VASP–/– and wild-type (wt) cells. In wt cells, VASP was phosphorylated in response to F/R but did not localize to intercellular junctions. In contrast, CNF-1 and expression of constitutively active Rac 1 induced translocation of VASP to cell borders in wt cells, where it colocalized with active Rac 1. In VASP–/– cells, Rac 1 activity was reduced to 0.4 of wt levels in controls and increased ~20-fold in response to CNF-1 compared with 7-fold activation in wt cells. Moreover, inactivation of Rac 1 by lethal toxin led to a greater increase of permeability compared with wt cells. All these data suggest that VASP is involved in the regulation of Rac 1 activity. Taking these findings together, our study indicates that VASP at least in part stabilizes endothelial barrier functions by control of Rho-family GTPases.

endothelial barrier functions; vasodilator-stimulated phosphoprotein; Rho guanosine 5'-triphosphatase; adenosine 3',5'-cyclic monophosphate

ENDOTHELIAL CELLS PROVIDE a selective barrier between blood vessels and the surrounding interstitial tissue. Therefore, their appropriate function is essential for the maintenance of tissue fluid and blood homeostasis (28). In response to inflammatory stimuli, maintenance of endothelial barrier functions is deranged, leading to vascular leakage, edema, and intravascular coagulation. We know that inflammatory mediators increase microvascular permeability by formation of gaps between endothelial cells (26, 27, 31, 35). Accumulating evidence points to a significant role of the actin cytoskeleton to anchor intercellular contacts such as adherens and tight junctions and, thus, for the maintenance of endothelial barrier properties in vivo and in vitro (5, 47).

It has been proposed that the actin-binding protein vasodilator-stimulated phosphoprotein (VASP) is involved in the regulation of endothelial barrier functions by cAMP (10). Furthermore, we found that stabilization of endothelial barrier functions in response to activation of the Rho GTPase Rac 1 was accompanied by translocation of VASP to intercellular junctions (50). VASP is a member of the Ena/VASP family (22, 23, 34), which besides its localization at focal adhesion sites can associate with adherens junctions via {alpha}-catenin (34, 46) and bind to endothelial tight junctions via ZO-1 (10). VASP has a polyproline-rich domain that associates with profilin, an actin monomer-binding protein (33, 38), and regulates multiple processes involved in actin polymerization such as nucleation, bundling, and capping (1, 6, 39). Recent data indicate that actin dynamics in endothelial cells are regulated via phosphorylation of VASP (7). In endothelial cells, VASP can be phosphorylated by several serine/threonine kinases, including cAMP-dependent protein kinase (PKA) and cGMP-dependent protein kinase (PKG), as well as by AMP-activated protein kinase (AMPK) (7, 44), which preferentially target specific residues of endothelial VASP. PKA and PKG primarily phosphorylate S157 and S239, respectively, whereas AMPK only phosphorylates T278 (7). Comerford et al. (10) reported that VASP, upon its phosphorylation on S157 by PKA, translocated to endothelial tight junctions and proposed that this mechanism is important for the barrier-stabilizing effects of cAMP. On the other hand, it has been shown that PKA-mediated VASP phosphorylation and membrane localization also occur in response to thrombin (32). Because cAMP and PKA are potent stabilizers of endothelial barrier functions in vivo and in vitro (10, 19, 20, 52), whereas thrombin is known to induce a barrier breakdown, these data have been interpreted as a mechanism to restore barrier functions following thrombin challenge. However, it has to be emphasized that the role of VASP phosphorylation and translocation for the barrier-stabilizing effects of cAMP and Rac 1 is unclear at present, and the importance of VASP in this respect has been hypothesized based on a spatiotemporal correlation. Thus Mehta and Malik (28) recently suggested that endothelial cells from VASP-deficient mice (VASP–/–) would be informative to address the functional role of VASP in the regulation of endothelial permeability.

Therefore, in the present study, we established microvascular myocardial endothelial cells (MyEnd) from VASP wild-type (wt) and VASP–/– mice to further characterize the role of VASP in endothelial barrier regulation as well as the requirement of VASP for the barrier-stabilizing effects of cAMP and Rac 1. We found that VASP is required for the maintenance of endothelial barrier functions under resting conditions but is not essential for cAMP and Rac 1 to enhance barrier functions. Rather, because Rac 1 activity was significantly reduced in VASP–/– cells, it is possible that VASP is involved in the mechanisms regulating activity of Rac 1 in microvascular endothelium.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. The immortalized mouse microvascular endothelial cell lines from myocardium (MyEnd wt and MyEnd VASP–/–) were grown in Dulbecco's modified Eagle's medium (DMEM; Life Technologies, Karlsruhe, Germany) supplemented with 50 U/ml penicillin-G, 50 µg streptomycin, and 10% fetal calf serum (FCS) (Biochrom, Berlin, Germany) in a humidified atmosphere (95% air–5% CO2) at 37°C. MyEnd cells have been prepared from mouse myocardial tissue of wt and VASP–/– mice (18). Immortalization was carried out as described previously for MyEnd cells (14). In brief, myocardial tissue of newborn mice was minced, digested with 0.05% trypsin (Biochrom) and 0.02% collagenase (Boehringer; Mannheim, Germany), and seeded onto gelatin-coated culture dishes. One day after plating, adherent cells were transfected with Polyoma virus middle T antigen. Polyoma virus middle T antigen transfection causes growth advantage of endothelial over nonendothelial cells, leading to a homogenous monolayer of cells with endothelial morphology after 4–6 wk of culture. MyEnd cells were immunopositive for several endothelial markers. For the present study, expression of von Willebrand factor and VASP, as well as junctional proteins VE-cadherin, β-catenin, ZO-1, claudin-5, and platelet endothelial cell adhesion molecule-1 (PECAM-1), were verified by immunoblotting. Cultures were used for experiments when grown to confluent monolayers (day 3 up to day 7).

Test reagents. Cytotoxic necrotizing factor (CNF-1) from Escherichia coli was supplied by Gundula Schmidt (Department of Pharmacology and Toxicology, University of Freiburg, Freiburg, Germany) and has been prepared and described previously (40, 41). According to our previous studies, we used CNF-1 at 300 ng/ml for 120 min for all experiments (50). Previously, we demonstrated that CNF-1 specifically activates Rac 1 and Cdc42, but not RhoA, in the MyEnd cell line (50). Forskolin and rolipram (F/R; both from Sigma-Aldrich, Taufkirchen, Germany) were used for 60 min at 5 and 10 µM, respectively. Lethal toxin (LT), which we confirmed to inactivate Rac 1 in MyEnd cells (49), was a gift from Torsten Giesemann (Department of Pharmacology and Toxicology, University of Freiburg) and was used at 50 ng/ml for 180 min.

Rac activation assay. For measurement of Rac 1 activation, the Rac G-Lisa Activation Assay Biochem kit (Cytoskeleton, Denver, CO) was used according to the manufacturer's recommendations. In brief, normal and VASP–/– MyEnd cells were grown to ~70% confluency in 35-mm dishes. Cells were incubated in the presence or absence of CNF-1 (300 ng/ml) in DMEM for 120 min. Afterward, monolayers were washed with ice-cold PBS (consisting of 137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, and 1.5 mM KH2PO4; pH 7.4) to remove serum proteins. Ice-cold cell lysis buffer was added, and cell lysates were harvested by centrifugation at 14,000 (~15,000 g) rpm at 4°C for 2 min. An aliquot was kept on ice for protein concentration measurement, and samples were snap frozen at –70°C. After adjustment of protein concentration, cell lysates were thawed and 50 µl of lysate were added to the wells of the RAC-GTPase binding plate, which is coated with a Rac-GTP binding domain (p21 binding domain, PBD). Additional wells were filled with lysis buffer or nonhydrolyzable Rac as a negative or positive control, respectively. The plate was placed immediately on a cold orbital shaker at 400 rpm at 4°C for 30 min and then washed twice with wash buffer at room temperature (RT), and 200 µl of antigen presenting buffer were added to each well for 2 min at RT. After the plate was washed three times, 50 µl of Rac primary antibody (diluted 1:200 in antibody dilution buffer) were added and incubated for 45 min on an orbital microplate shaker (400 rpm, RT). After the plate was washed three times, 50 µl of secondary anti-horseradish peroxidase (HRP)-labeled antibody (diluted 1:100) were added to each well on the orbital microplate shaker (RT, 45 min). The wells were then washed three times, and 50 µl of horseradish peroxidase (HRP) detection reagent were added and incubated at 37°C for 15 min. HRP stop buffer (50 µl) was added, and the signal was measured immediately at 490 nm using a microplate spectrophotometer (Sunrise; Tecan, Crailsheim, Germany). The results were analyzed using Microsoft Excel software.

Cytochemistry. MyEnd cells were grown on coverslips coated with gelatin cross-linked with glutaraldehyde (42). After incubation with F/R or CNF-1, culture medium was removed and monolayers were fixed for 10 min at RT with 2% formaldehyde (freshly prepared from paraformaldehyde) in PBS. Afterward, monolayers were treated with 0.1% Triton X-100 in PBS for 5 min. After being rinsed with PBS at RT, MyEnd were preincubated for 30 min with 10% normal goat serum (NGS) and 1% bovine serum albumin (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 (15), a mouse monoclonal IgG Alexa Fluor 555 conjugate (Upstate, Schwalbach, Germany) directed against cortactin (1:100 in PBS), a mouse monoclonal antibody against ZO-1 (Chemicon-Millipore, Schwalbach, Germany), or a polyclonal IgG raised against vasodilator-stimulated phosphoprotein (1:100 in PBS) (16). We used the following phospho-VASP-specific antibodies: a rabbit polyclonal IgG directed against phospho-S157-VASP (44), two mouse monoclonal IgG directed against phospho-S239-VASP (44), or a phospho-T278-VASP (7), respectively. After several rinses with PBS (3 times for 5 min), monolayers were incubated for 60 min at RT with Cy3-labeled goat anti-rat IgG, Cy3-labeled goat anti-mouse IgG, or Cy3-labeled goat anti-rabbit IgG, respectively (all from Dianova, Hamburg, Germany; diluted 1:600 in PBS). For visualization of filamentous actin (F-actin), some monolayers were incubated with Alexa phalloidin (Mobitec, Goettingen, Germany; diluted 1:60 in PBS, 1 h at RT). Cells incubated with antibodies or Alexa phalloidin were rinsed with PBS (3 times for 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 wt cells. Cells were transfected with a pEGFP-Rac 1V12 constitutive active (pEGFP-Rac 1 CA) mutant or a p-EGFP-Rac 1N17 dominant negative (pEGFP-Rac 1 DN) mutant 1 day after plating using effectene transfection reagent (Quiagen, Hilden, Germany) as described previously (51). The plasmids were a kind gift of Kozo Kaibuchi (University of Nagoya, Nagoya, Japan) (30). Twenty-four hours after transfection, cells were fixed for 10 min at RT with 2% formaldehyde (freshly prepared from paraformaldehyde) in PBS, and immunostaining for VASP or cortactin was carried out as described above.

Recombinant VE-cadherin-Fc. As described previously, 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 (3, 4, 15). The protein was expressed by stably transfected Chinese hamster ovary cells (CHO) and purified from culture supernatants by affinity chromatography using protein A-agarose (Oncogene, Cambridge, MA).

Coating of polystyrene beads. After vortexing, 10 µl of protein A-coated superparamagnetic polystyrene microbeads (Dynabeads; diameter 2.8 µm; Dynal, Oslo, Norway) 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 VE-cadherin-Fc, and allowed to react for 16 h at 4°C under permanent slow overhead rotation to avoid aggregation. After being washed three times for 5 min in 100 µl of buffer A and three times for 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 of dimethyl pimelimidate dihydrochloride (Pierce, Rockford, IL) to covalently cross-link protein A and bound Fc parts. After being washed twice for 5 min in buffer C (100 µl of 0.2 methanolamine, pH 8.0), beads were incubated in buffer C for 2 h at RT. Finally, beads were washed three times for 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 tweezers. As described previously (4), 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 Oberkochen, Germany), coupled through the epi-illumination port of an Axiovert 135 microscope (Zeiss) and reflected to the objective by a dichroic mirror (FT 510; Zeiss). Throughout all experiments, the laser intensity was 42 mW in the focal plane. Coated beads (10 µl of stock solution) were suspended in 200 µl of culture medium and allowed to interact with microvascular endothelial cell monolayers for 30 min at 37°C before initiation of experiments. Beads were considered tightly bound when resisting laser displacement at 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.

Measurement of FITC-dextran flux across monolayers of cultured endothelial cells. As described previously (50), endothelial cells were seeded on top of gelatin-coated transwell chambers for six-well plates (0.4-µm pore size; Falcon, Heidelberg, Germany) and grown to confluence. After being rinsed with PBS, cells were incubated with fresh DMEM without phenol red (Sigma) containing 1 mg/ml FITC-dextran (4 or 70 kDa; Sigma) in the presence or absence of CNF-1 and F/R. Paracellular flux was assessed by taking 100-µl aliquots from the outer chamber over 2 h of incubation. Fluorescence was measured using a Wallac Victor2 fluorescence spectrophotometer (Perkin-Elmer, Überlingen, Germany) with excitation and emission at 485 and 535 nm, respectively. For transendothelial flux under baseline conditions, the permeability coefficient (PE) was calculated using the following formula as described in detail previously (8, 9): PE = [({Delta}CA/{Delta}t)·VA]/(S·{Delta}CL), where PE is the diffusive permeability (cm/s), {Delta}CA is the change of FITC-dextran concentration, {Delta}t is the change of time, VA is the volume of the abluminal medium, S is the surface area, and {Delta}CL is the constant luminal concentration. Changes of permeability after F/R, CNF-1, or LT treatment were expressed as FITC-dextran clearance through the monolayer compared with the clearance of cell monolayers under control conditions.

Statistics. Values are means ± SE. Possible differences in bead binding and FITC-dextran flux (and Rac activation) between groups were assessed using unpaired Student's t-test and the nonparametric Mann-Whitney statistic. Statistical significance is assumed for P < 0.05.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Permeability was increased in VASP–/– cells in the absence of discernible morphological alterations. To investigate the role of VASP for baseline endothelial barrier functions, we compared the permeability coefficient PE of FITC-dextran flux across VASP–/– and wt endothelial monolayers under resting conditions. Over 2 h of measurements, the mean PE was 0.4 ± 0.1 x 10–6 cm/s for 70-kDa FITC-dextran and 1.4 ± 0.2 x 10–6 cm/s for 4-kDa FITC-dextran (Fig. 1). This was comparable to the PE of other endothelial cell lines (8). In VASP–/– monolayers, the PE was significantly increased threefold (4-kDa FITC-dextran) and ninefold (70-kDa FITC-dextran) relative to wt cells. Interestingly, fluxes of 4- and 70-kDa FITC-dextran were significantly different in wt but not in VASP–/– cells, indicating that the size selectivity of the endothelial barrier was compromised in VASP–/– cells. To investigate whether reduced barrier functions were caused by alterations of intercellular junctions or the actin cytoskeleton, we performed immunostaining of wt and VASP–/– monolayers. VASP–/– cells showed shape and growth pattern comparable to primary cultures of microvascular endothelial cells (21, 29, 37), and both cell lines displayed intact monolayers with a whirl-like growth pattern (Fig. 2). Moreover, immunostaining revealed VE-cadherin (Fig. 2, a and b), β-catenin (Fig. 2, c and d), the tight junction-associated protein ZO-1 (Fig. 2, e and f), and claudin-5 (not shown) regularly distributed along cell borders in both cell lines. Multiple stress fibers were running across the cytoplasm in both cell lines as shown by staining of F-actin using Alexa phalloidin (Fig. 2, g and h). Taken together, these data demonstrate that VASP-deficiency led to decreased endothelial barrier functions in the absence of obvious morphological alterations.


Figure 1
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  		Fig. 1. Endothelial barrier functions are reduced in vasodilator-stimulated phosphoprotein-deficient (VASP–/–) cells. FITC-dextran flux across untreated endothelial monolayers was used to assess barrier functions in myocardial endothelial cells (MyEnd) wild-type (wt) and VASP–/– cells under resting conditions. Permeability coefficient (PE) was determined for 4- and 70-kDa FITC-dextran. Measurements over 2 h demonstrated that 4-kDa FITC-dextran flux was significantly increased 3.2-fold across VASP–/– monolayers (n = 20) compared with MyEnd wt cells (n = 24). Similarly, the flux of 70-kDa FITC-dextran was increased 9.0-fold in VASP–/– cells (n = 12) compared with wt cells (n = 6). Values are means ± SE.

 

Figure 2
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  		Fig. 2. VASP–/– cells showed no morphological alterations compared with wt cells. Immunostaining of endothelial adherens and tight junction protein was performed in MyEnd wt (a, c, e, g) and VASP–/– monolayers (b, d, f, h), and F-actin was stained using Alexa phalloidin (g and h). VE-cadherin (a and b), β-catenin (c and d) and tight junction-associated protein ZO-1 (e and f) were regularly distributed along cell borders in both MyEnd wt and VASP–/– cells. Multiple stress fibers were distributed throughout the cytoplasm in both cell lines (g and h). Results shown are representative of n ≥ 5 experiments. Scale bar, 20 µm (a–h).

 
VASP was not required for the mechanisms involved in endothelial barrier stabilization by cAMP and Rac 1. The cAMP-dependent phosphorylation of VASP has been suggested to play an important role in regulating endothelial barrier functions (10). To sensitively detect changes in permeability, we measured the flux of 4-kDa FITC-dextran across VASP wt and VASP–/– monolayers following incubation with F/R to increase intracellular cAMP. FITC-dextran flux was significantly reduced to a comparable extent in both VASP wt (37 ± 13% of controls) and VASP–/– monolayers (31 ± 8% of controls; Fig. 3). To activate Rac 1 in MyEnd cells, we used CNF-1, because we previously showed that it effectively activates Rac 1 in virtually all endothelial cells of the monolayer within 120 min (50). Similar to treatment with F/R, after incubation with CNF-1 to activate Rac 1, permeability was also significantly reduced in wt (74 ± 13% of controls) and VASP–/– monolayers (72 ± 9% of controls). These effects were not significantly different in both cell lines. Together, these experiments indicate that VASP is not required for barrier-stabilizing effects of cAMP and Rac 1.


Figure 3
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  		Fig. 3. Increased cAMP levels and activation of Rac 1 stabilized endothelial barrier functions in MyEnd wt and VASP–/– cells. Measurements of FITC-dextran flux across endothelial monolayers after treatment with either forskolin and rolipram (F/R) or cytotoxic necrotizing factor 1 (CNF-1) resulted in a significant stabilization of endothelial barrier functions compared with controls in both cell lines. F/R reduced FITC-dextran flux to 37 ± 13% in wt and to 31 ± 8% in VASP–/– cell monolayers (n = 8). Activation of Rac 1 by CNF-1 resulted in a reduction of FITC-dextran flux to 74 ± 13% in wt and to 72 ± 9% in VASP–/– cells (n = 12) compared with controls. Values are means ± SE. *P ≤ 0.05 compared with controls.

 
Treatment with F/R to increase intracellular cAMP did not affect VE-cadherin distribution in wt and VASP–/– cells (Fig. 4, a–d). In contrast to previous reports (10), VASP remained diffusely distributed in the cytoplasm in a punctuated staining pattern reminiscent of focal adhesions of wt cells and did not localize to cell borders after F/R treatment (Fig. 4, e–h). However, Western blotting revealed that in contrast to CNF-1 treatment, incubation with F/R induced a shift of the VASP-specific band from ~46 to ~50 kDa (Fig. 5A), indicating that F/R treatment was effective to induce phosphorylation of VASP under these conditions (10, 17). The shift of the VASP-specific band was previously demonstrated to be due to a mobility change of phosphorylated VASP (16). In VASP–/– cells, no VASP-specific band was detected. To rule out the possibility that only a subset of phosphorylated VASP molecules was recruited to cell borders that was too small to be detected by the VASP antibody, we tested the recruitment of phosphorylated VASP to cell borders in MyEnd wt cells after F/R treatment, using specific antibodies directed against VASP phosphorylated at S157, S239, or T278. Antibodies against phospho-S239-VASP and phospho-T278-VASP yielded an unspecific staining pattern in controls and in F/R- and CNF-1 treated cells, indicating that VASP was phosphorylated at neither S239 nor T278 under these conditions (not shown). Compared with controls (Fig. 5Ba), staining of phospho-S157-VASP was increased in F/R-treated cells (Fig. 5Bc) but not in CNF-1-treated cells (Fig. 5Bb), indicating that VASP was phosphorylated at S157 after F/R treatment. However, translocation of phospho-S157-VASP to cell borders was not observed in any experiment.


Figure 4
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  		Fig. 4. F/R treatment did not affect distribution of VE-cadherin and VASP. MyEnd wt (a, e, c, g) and VASP–/– cell monolayers (b, f, d, h) were immunostained with VE-cadherin and VASP. VE-cadherin was regularly distributed at the cell borders in controls (a and b). This was not altered after incubation with F/R in both cell lines (c and d). VASP was diffusely distributed in the cytoplasm in a punctuated staining pattern reminiscent of focal adhesions in MyEnd wt cells under control conditions (e) as well as after F/R treatment (g). No specific VASP immunostaining was observed in VASP–/– cells (f and h). Results shown are representative of n ≥ 5 experiments. Scale bar, 20 µm (a–h).

 

Figure 5
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  		Fig. 5. A: Western blot analyses demonstrated phosphorylation of VASP after F/R treatment. Western blots show that incubation of MyEnd wt cells with CNF-1 did not alter the VASP-specific band at ~46 kDa detected in controls, whereas treatment with F/R induced a shift of the VASP-specific band from ~46 to ~50 kDa, indicating a phosphorylation of VASP. No VASP-specific bands were detected in VASP–/– cells. The experiment shown is representative of n = 3 individual experiments. B: immunostaining showed an increase of phospho-S157-VASP in MyEnd wt cells after F/R treatment. MyEnd wt cells were immunostained for phospho-S157-VASP using a specific antibody. We observed a diffuse staining pattern in controls (a) and in CNF-1-treated cells (b). In MyEnd wt cells treated with F/R, phospho-S157-VASP was augmented, indicating that VASP had been phosphorylated at S157 after F/R treatment (c). However, no recruitment of phospho-S157-VASP to cell borders was obvious in all F/R-treated cells. Results shown are representative of n = 4 experiments. Scale bar, 20 µm.

 
Compared with controls (Fig. 6A, a, b, e, and f), incubation of endothelial monolayers with CNF-1 resulted in increased staining of the junction-associated actin belt in both wt and VASP–/– cells and reduction of stress fibers (Fig. 6A, g and h) without any effect on the localization of VE-cadherin at cell borders (Fig. 6A, c, d, g, and h). Cortactin, a downstream effector of Rac 1 in the regulation of the junction-associated actin cytoskeleton, redistributed to cell borders after incubation with CNF-1 in both wt and VASP–/– cells (Fig. 6A, k and l). Similarly, VASP translocated to cell borders after CNF-1 treatment in wt cells compared with a punctuated staining pattern reminiscent of focal adhesions in untreated monolayers (Fig. 6A, m and o). In contrast, no specific VASP immunostaining was found in VASP–/– monolayers (Fig. 6A, n and p).


Figure 6
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  		Fig. 6. A: activation of Rac 1 strengthened the junction-associated actin belt and caused distribution of cortactin and VASP to cell borders. MyEnd wt (a, c, e, g, i, k, m, o) and VASP–/– (b, d, f, h, j, l, n, p) cell monolayers were immunostained for VE-cadherin (a–d), cortactin (i–l), and VASP (m–p) and stained for F-actin with Alexa phalloidin (e–h). Cortactin and VASP immunostaining are shown at higher magnification (i–p). VE-cadherin distribution was regular at cell borders in MyEnd wt (a) and VASP–/– controls (b). No alterations were discernible upon incubation with CNF-1 in both cell lines (c and d). Staining of F-actin showed multiple stress fibers in controls (e and f). Activation of Rac 1 resulted in an increase of the junction-associated actin belt and reduction of stress fibers (g and h). Cortactin was diffusely distributed over the cytoplasm in controls (i and j) and redistributed to the cell borders after CNF-1 treatment in both cell lines (k and l). Diffusely distributed VASP in MyEnd wt cells (m) also translocated to the cell membrane upon Rac 1 activation. Specific VASP immunostaining in VASP–/– cells (n and p) was absent. Results shown are representative of n ≥ 5 experiments. Scale bars, 20 µm (a–p). B: after Rac 1 activation, VASP colocalized with VE-cadherin and ZO-1 at cell borders. MyEnd wt cells treated with CNF-1 were stained for the adherens junction marker VE-cadherin (a) and the tight junction marker ZO-1 (d). VASP (b and e) was recruited to cell borders upon Rac 1 activation and colocalized largely with either VE-cadherin (c) or ZO-1 (f). However, VASP staining was missing in some limited areas (arrows in f) where ZO-1 staining was observed. Results shown are representative of n = 4 experiments. Scale bar, 20 µm (a–f).

 
Colocalization studies showed that following CNF-1 treatment VASP largely colocalized with the adherens junction marker VE-cadherin (Fig. 6B, a–c) and the tight junction marker ZO-1 (Fig. 6B, d–f). However, in some limited areas of the cell border, ZO-1 was observed where VASP was missing (arrow in Fig. 6Bf).

Effects of F/R and CNF-1 on VE-cadherin-mediated adhesion in VASP wt and VASP–/– cells. To study the role of VASP in the regulation of VE-cadherin-mediated adhesion, we performed laser tweezer experiments using VE-cadherin-coated microbeads as described previously (49). Microbeads were allowed to settle on the cell surface for 30 min, which resulted in formation of cell-to-bead-contacts immunopositive for VE-cadherin, β-catenin, and F-actin (2, 50). Afterward, beads were subjected to a laser beam to displace unbound beads. Under control conditions, the percentage of bound beads in MyEnd wt was not significantly different compared with VASP–/– cells. On the surface of wt cells, 79 ± 2% beads were bound after 30 min, whereas VASP–/– cells had a binding rate of 74 ± 3% under the same conditions. These control values were set to 100% (Fig. 7). Incubation of both wt and VASP–/– monolayers with EGTA for 30 min led to a significant decrease in the number of bound beads and served as a negative control (42 ± 2% of controls for wt cells; 47 ± 3% of controls for VASP–/– cells). F/R did not alter the number of bound beads in wt (102 ± 2% of controls) as well as VASP–/– cells (100 ± 2% of controls), indicating that cAMP does not regulate VE-cadherin-mediated adhesion. Similarly, in wt cells, incubation with CNF-1 did not cause alterations in the number of bound beads (98 ± 3% of controls). However, in VASP–/– cells, the number of bound beads was increased to 14% above control levels (114 ± 4% of controls) following treatment with CNF-1. It has to be emphasized that the laser tweezer assays only allow quantification of bead binding to isolated VE-cadherin molecules on the cell surface and therefore might not completely reflect the situation within the intercellular cleft.


Figure 7
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  		Fig. 7. Role of VASP in VE-cadherin-mediated adhesion was assayed using laser tweezers. VE-cadherin-coated microbeads were allowed to settle on the cell surface for 30 min. The number of bound beads was evaluated (controls) and was not significantly different in MyEnd wt and VASP–/– cells. Ca2+ depletion by EGTA resulted in a significant decrease in the number of bound beads in both cell lines (n = 6). Treatment with F/R did not alter the number of bound beads in wt and VASP–/– cells (n = 6). In contrast, CNF-1 significantly increased the number of bound beads compared with controls in VASP–/– cells (n = 9), whereas no such increase could be observed in wt cells (n = 20). Values are means ± SE. *P ≤ 0.05 compared with controls.

 
Rac 1 activity and activation by CNF-1 were significantly altered in VASP–/– cells. To study whether compromised Rac 1 activity in VASP–/– cells accounted for the differences in VE-cadherin binding in response to CNF-1, and because previous studies showed that Rac 1 activity was altered in VASP–/– fibroblasts (12), GTP-bound activated Rac 1 was assayed as a measure of Rac 1 activation. Interestingly, baseline levels of active Rac 1 were significantly reduced to ~ 60% (0.4 ± 0.3-fold) of wt levels (Fig. 8). Incubation with CNF-1 resulted in a 6.8 ± 0.1-fold activation of Rac 1 in wt cells and a 19.5 ± 0.1-fold activation in VASP–/– cells. Thus activation of Rac 1 was significantly stronger in VASP–/– cells. However, compared with that of wt controls, activity of Rac 1 in VASP–/– cells was augmented 7.9 ± 0.1-fold, i.e., to an absolute level similar to that in wt cells.


Figure 8
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  		Fig. 8. VASP–/– cells show significantly altered Rac 1 activity and activation by CNF-1. Rac activation analyses demonstrated that Rac 1 activity was significantly reduced 0.4 ± 0.3-fold in VASP–/– cells (n = 7) to ~60% of wt cell levels (n = 6) under resting conditions. Incubation with CNF-1 resulted in strong activation of Rac 1 in both cell lines. Compared with wt controls, activation was increased 6.8 ± 0.1-fold in wt cells (n = 6) and 7.9 ± 0.1-fold in VASP–/– cells (n = 8). *P ≤ 0.05 compared with MyEnd wt controls. #P ≤ 0.05 compared with CNF-1-treated MyEnd wt.

 
Active Rac 1 colocalized with VASP and cortactin in transfected MyEnd cells. From our experiments, it became obvious VASP was not necessary to mediate the barrier-stabilizing effect of Rac 1. However, the decreased baseline activity of Rac 1 under control conditions in VASP–/– and the observation that VASP was recruited to cell borders in MyEnd wt cells after Rac 1 activation indicated a functional association of VASP and activated Rac 1. We transfected MyEnd wt cells either with a constitutive active Rac 1 (pEGFP-Rac 1 CA) mutant or a dominant negative Rac 1 (pEGFP-Rac 1 DN) mutant to test whether activated Rac 1 and VASP colocalize. In transfected cells, pEGFP-Rac 1 CA was located at cell borders (arrows in Fig. 9, a and g). Immunostaining demonstrated that VASP was recruited to cell borders (Fig. 9b) and exactly colocalized with active Rac 1 (Fig. 9c). This was not observed in cells transfected with pEGFP-Rac 1 DN, in which VASP remained diffusely distributed in the cytoplasm (Fig. 9e). Staining of cortactin in the pEGFP-Rac 1 CA-transfected cells confirmed that activated Rac 1 led to recruitment of cortactin to cell borders (Fig. 9h), where it colocalized with active Rac 1 (Fig. 9i), similar to our experiments where CNF-1 was used to activate Rac 1 (Fig. 6A, i and k). Again, this was not observed in cells transfected with pEGFP-Rac 1 DN (Fig. 9, f and l).


Figure 9
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  		Fig. 9. Colocalization of activated Rac 1 with VASP and cortactin in MyEnd wt cells transfected with pEGFP-Rac 1V12 constitutive active (pEGFP-Rac 1 CA) mutant. MyEnd wt cells were transfected with either pEGFP-Rac 1 CA (a–c, g–i) or p-EGFP-Rac 1N17 dominant negative (pEGFP-Rac 1 DN) mutants (d–f; j–l). Cells transfected with Rac 1 CA showed active Rac 1 located at the cell borders (arrows in a and g), which was not observed in Rac 1 DN-transfected cells (d and j). Costaining with VASP (a–f) showed that VASP was recruited to the cell borders in cells transfected with pEGFP-Rac 1 CA (b) but not in pEGFP-Rac 1 DN-transfected cells (e). VASP and activated Rac 1 exactly colocalized at cell borders (arrows in c). Cortactin was also recruited to cell borders in pEGFP-Rac 1 CA-transfected cells (h) and colocalized with active Rac 1 (arrows in i). This was not observed in pEGFP-Rac 1 DN-transfected cells (k and l). Results are representative of n = 4 experiments. Scale bar, 50 µm (a–l).

 
The endothelial barrier of VASP–/– cells is more vulnerable to Rac 1 inactivation. Because Rac 1 activity was reduced in VASP–/– cells and permeability was increased under control conditions, we tested whether this cell line was more sensitive to an additional inactivation of Rac 1 than wt cells. Therefore, we performed FITC-dextran (4 kDa) flux measurements after treatment of both cell lines with LT, which is known to inhibit Rac 1 activity in MyEnd cells (49). FITC-dextran flux across endothelial monolayers was significantly increased in both cell lines, i.e., to 366 ± 28% in MyEnd wt and to 481 ± 29% in VASP–/– cells compared with controls (Fig. 10). Thus VASP–/– monolayers displayed a significantly higher (1.3-fold) increase of FITC-dextran flux than wt cells. This indicates that VASP–/– cells were more sensitive to Rac 1 inhibition.


Figure 10
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  		Fig. 10. Inactivation of Rac 1 by lethal toxin (LT) led to a higher increase of permeability in VASP–/– cells than in MyEnd wt cells. Measurements of 4-kDa FITC-dextran flux across endothelial monolayers after treatment with LT led to a significant increase of permeability across VASP–/– monolayers (481 ± 29%) as well as across MyEnd wt cells (366 ± 28%) compared with controls (n = 4). This increase of permeability was significantly higher in VASP–/– monolayers than in wt monolayers. Values are means ± SE. *P ≤ 0.05 compared with controls. #P ≤ 0.05 compared with MyEnd wt.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
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 DISCUSSION
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 REFERENCES
 
The present studies extend our investigations on the role of the actin cytoskeleton in the regulation of endothelial barrier functions as well as on the mechanisms involved in the barrier-stabilizing effects of cAMP and Rac 1. We provide the first evidence that the actin-binding protein VASP is required for the maintenance of endothelial barrier functions in vitro. In a VASP-deficient microvascular myocardial endothelial cell line, baseline permeability was significantly increased compared with wt cells. However, increase of cellular cAMP by F/R as well as activation of Rac 1 by CNF-1 were equally efficient to enhance barrier functions in both wt and VASP–/– cells, indicating that VASP is not an essential downstream regulator in cAMP- and Rac 1-dependent barrier stabilization. In contrast, because baseline activity of Rac 1 was significantly reduced in VASP–/– cells, Rac 1 activation by CNF-1 was 20-fold increased in these cells, and VASP exactly colocalized with active Rac 1 in wt cells, our data indicate that VASP might stabilize endothelial barrier functions by regulating Rac 1 activity.

The role of VASP in the regulation of endothelial barrier functions under resting conditions as well as in response to increased cAMP. The anchorage of endothelial adherens and tight junctions to the actin cytoskeleton is generally thought to be essential for strong intercellular adhesion (5). Accordingly, disruption of F-actin leads to a breakdown of the endothelial barrier in vivo and in vitro (11, 43, 45, 50). VASP has been demonstrated to promote actin polymerization by multiple mechanisms, including nucleation, bundling, and capping (1, 39). Moreover, it was recently reported that VASP is important for F-actin assembly in endothelial cells (7). In the present study, we found that baseline permeability of VASP–/– MyEnd monolayers was about threefold higher for 4-kDa FITC-dextran and ninefold higher for 70-kDa FITC-dextran compared with wt cells. This is the first direct evidence that VASP is required for the maintenance of microvascular endothelial barrier functions.

In VASP–/– monolayers, baseline fluxes of 4- and 70-kDa FITC-dextran were not significantly different in contrast to wt monolayers. These data indicate that in addition to a reduced overall barrier function the size selectivity of the endothelial barrier was compromised in VASP–/– cells. On the other hand, we did not detect alterations in the distribution of adherens junction or tight junction components. Although we were also unable to detect ultrastructural alterations of cell junctions using electron microscopy (Schlegel N and Waschke J, unpublished observations) more detailed studies are necessary to address this important issue.

Our results support a recent study that showed that downregulation of VASP by small interference RNA increased FITC-dextran flux, whereas VASP overexpression reduced permeability (36). In contrast, previous studies concluded from experiments using bovine macrovascular arterial endothelial cells transiently transfected with VASP mutants defective for actin bundling that VASP may negatively regulate barrier functions in the resting state (10). Comerford et al. (10) proposed that VASP is required to maintain an open paracellular pathway in endothelial cells and that VASP, upon S157 phosphorylation by PKA, translocates to cell junctions and binds to ZO-1 to decrease permeability by reducing the cytoskeletal tension exerted on tight junctions. According to this hypothesis, VASP deficiency would lead to reduced baseline permeability, and the response to cAMP-mediated activation of PKA would be blunted. We cannot rule out the possibility that cAMP was still effective to reduce permeability in VASP–/– cells, because other members of the Ena/VASP protein family such as Mena or EVL, which share a conserved domain structure, are able to compensate for VASP deficiency. It has indeed been demonstrated that other members of the Ena/VASP family are substrates of PKA and PKG (13, 24, 48). Nevertheless, in our study, cAMP-mediated barrier stabilization in wt MyEnd monolayers was not accompanied by redistribution of phosphorylated or unphosphorylated VASP to cell junctions, although cAMP induced phosphorylation of VASP at S157. Taking these findings together, we found no indication for a prominent role of VASP in endothelial barrier regulation by cAMP.

VASP is not involved in the barrier-stabilizing effects downstream of Rac 1. Rac 1, similarly to cAMP, is required for the maintenance of endothelial barrier functions and is capable to enhance barrier properties in vivo and in vitro (49, 50, 54–56). We recently found that barrier stabilization in response to activation of Rac 1 was paralleled by translocation of VASP to intercellular junctions (50), although VASP is not known to be a downstream effector of Rac 1. Therefore, we studied whether VASP was involved in the mechanisms by which Rac 1 regulates barrier functions. Activation of Rac 1 by CNF-1 was equally effective to recruit the Rac 1 effector cortactin to cell borders, to strengthen the peripheral actin belt, and to reduce endothelial permeability in both VASP–/– and wt cells. These data demonstrate that VASP is not essential for barrier stabilization by Rac 1. In wt cells, barrier stabilization was paralleled by translocation of VASP to cell borders. From the fact that none of the phospho-specific antibodies detected CNF-1-induced relocalization, we conclude that Rac 1 activation caused translocation of unphosphorylated VASP, and therefore VASP phosphorylation seems not to be required for VASP membrane localization. Moreover, the results indicate that VASP relocalization to cell borders, which has been described in response to VASP phosphorylation by PKA (10, 32), may not necessarily be involved in the barrier-stabilizing effects downstream of cAMP and Rac 1.

VASP is not essential for VE-cadherin-mediated adhesion. Because we found that the actin cytoskeleton is important for VE-cadherin-mediated adhesion in vitro (4, 53) as well as for the maintenance of endothelial barrier functions in vivo (51), we investigated whether VASP was required for VE-cadherin-mediated adhesion. The number of VE-cadherin-coated beads bound to the cell surface was not significantly different in wt and VASP–/– cells in resting monolayers as well as in response to increased cAMP, indicating that either VASP is not involved in the regulation of VE-cadherin-mediated adhesion or its deficiency can be compensated by other members of the Ena/VASP family. Previously, we found that inhibition of Rac 1 by LT resulted in intercellular gap formation, fragmentation of VE-cadherin immunostaining along cell junctions, and a significant loss of adhesion (49). Presently, we have shown that Rac 1 was reduced in VASP–/– cells to 40% of wt levels in the absence of gap formation or detectable alterations of adherens junctions as well as of VE-mediated-adhesion. These data suggest that reduced Rac 1 activity as observed in VASP–/– cells was still sufficient to maintain VE-cadherin binding. This is supported by the fact that activation of Rac 1 in wt cells did not further strengthen VE-cadherin-binding above control levels, likely because the effect of Rac 1 on VE-cadherin-mediated adhesion was maximal in these cells under control conditions. In contrast, in VASP–/– cells in which baseline activity of Rac 1 was reduced, activation of Rac 1 by CNF-1 significantly increased VE-cadherin binding. These data indicate that other types of cell contacts such as tight junctions or focal adhesions may be more sensitive to altered Rac 1 activity.

VASP may act as a regulator of Rac 1 activity in endothelial cells. We found that baseline activity of Rac 1 was reduced to 40% of wt levels. Because Rac 1 is required for maintenance of endothelial barrier functions (49, 55), it is possible that reduced Rac 1 activity was responsible for the increased permeability in VASP–/– cells. This view is supported by the fact that activation of Rac 1 by CNF-1 was increased in VASP–/– cells to levels of CNF-1-treated wt cells. Rac 1 was activated by CNF-1 about 7-fold in wt cells compared with 20-fold in VASP–/–, i.e., activity was increased to a similar absolute level when the different baseline activity levels are taken into account. This may explain why permeability was reduced by CNF-1 to a comparable extend in both wt and VASP–/– cells. These data support previous results from VASP–/– fibroblasts showing that Rac 1 activation by PDGF was augmented (12). The hypothesis that reduced activity of Rac 1 accounted for impaired barrier function in VASP–/– cells is also supported by our finding that VASP–/– cells were more sensitive to Rac 1 inactivation by LT.

As a second line of evidence, we demonstrated that activation of Rac 1 by CNF-1 or by transfection of cells with constitutive active Rac 1 recruited VASP to the cell junctions, where it colocalized with active Rac 1. On the other hand, we found that VASP is unlikely to be a downstream effector of Rac 1, at least in the regulation of endothelial barrier functions, because CNF-1 was effective to reduce permeability in VASP–/– cells. Therefore, from our present findings we suggest that VASP may act as a regulator of Rac 1 activity, although the exact mechanisms are still unclear. It is possible that VASP in wt cells interferes with intrinsic GTP hydrolysis, and thus the inactivation of Rac 1, because VASP can interact with p120RasGAP. This mechanism could account for the reduced activity of Rac 1 in VASP–/– cells under resting conditions (12). In addition, activation of Rac 1 by CNF-1 may be increased in VASP–/– cells, because VASP in wt cells interfered with Rac 1 activation by controlling not only GTP hydrolysis but also GTP exchange. In favor of this hypothesis, it has been shown that VASP can have opposing functions to Trio, a GTP exchange factor (GEF) for Rac 1 (25). These data suggest that VASP may be involved in negatively regulating Rac 1 activity. In support of this hypothesis, we previously found that increased cAMP under conditions where VASP is phosphorylated abolished LT-induced Rac 1 inactivation (53). Moreover, our observation that VASP relocated to cell junctions upon Rac 1 activation but was not required for the barrier-stabilizing effects downstream of Rac-1 can be explained in the way that VASP translocated together with active Rac 1 to regulate its function.

Our hypothesis of an important role of VASP as a regulator of Rac 1 in the maintenance of endothelial barrier functions might also be relevant for increased permeability under pathological conditions such as hypoxia. It was recently demonstrated that VASP expression can be downregulated by hypoxia-inducible factor 1. This has been suggested to be responsible for the increased permeability in endothelial cells under hypoxia (36). Because it is known that hypoxia can lead to an inactivation of Rac 1 (56), it can be speculated that HIF-1-induced downregulation of VASP might lead to an inactivation of Rac 1 and thereby to increased endothelial permeability.

Taking all the findings together, our study supports the hypothesis that VASP is required for the maintenance of endothelial barrier functions. In addition to its well-known regulatory functions on the level of the actin cytoskeleton, we propose that VASP also takes place in the regulation of small GTPase Rac 1, which is also a key player in the regulation of endothelial barrier properties. Future studies are needed to address the precise mechanism by which VASP interferes with Rac 1 activity.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
These studies were supported by a grant from the Deutsche Forschungsgemeinschaft (SFB 688, TP A2 and A4).


   ACKNOWLEDGMENTS
 
We are grateful to Stefanie Imhof, Nadja Niedermeier, and Lisa Bergauer for skillful technical assistance. The phospho-T278-VASP antibody was supplied by Thomas Renné (Institute for Clinical Biochemistry and Pathobiochemistry, University of Würzburg).

Present address of N. Golenhofen: Institute of Anatomy and Cell Biology, University of Ulm, Albert-Einstein Allee 11, D-89081 Ulm, Germany.


   FOOTNOTES
 

Address for reprint requests and other correspondence: J. Waschke, Institute of Anatomy and Cell Biology, Julius-Maximilians-Univ., Koellikerstr. 6, 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.

* N. Schlegel and S. Burger contributed equally to this work. Back


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