Endothelial Barrier Strengthening by Activation of Focal Adhesion Kinase*

Sadiqa K. QuadriDagger , Mrinal BhattacharjeeDagger , Kaushik ParthasarathiDagger , Tatsuo Tanita§, and Jahar BhattacharyaDagger

From the Dagger  Lung Biology Laboratory, College of Physicians and Surgeons, Columbia University, St. Luke's-Roosevelt Hospital Center, New York, New York 10019, and the § Department of Thoracic Surgery, Iwate Medical University, School of Medicine, Iwate, 020-8505, Japan

Received for publication, September 27, 2002, and in revised form, January 17, 2003

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Endothelial cell barrier (EC) properties regulate blood tissue fluid flux. To determine the role of endothelial-matrix interactions in barrier regulation, we induced cell shrinkage by exposing confluent endothelial monolayers to hyperosmolarity. The dominant effect of a 15-min hyperosmolar exposure was an increase in the trans-endothelial electrical resistance, indicating the induction of barrier strengthening. Hyperosmolar exposure also increased activity of focal adhesion kinase and E-cadherin accumulation at the cell periphery. Concomitantly, the density of actin filaments increased markedly. In EC monolayers stably expressing constitutively active or dominant negative isoforms of Rac1, the actin response to hyperosmolar exposure was enhanced or blocked, respectively, although the response in trans-endothelial resistance was unaffected, indicating that the endothelial barrier enhancement occurred independently of actin. However, in monolayers expressing a kinase-deficient mutant of focal adhesion kinase, the hyperosmolarity-induced increases in activity of focal adhesion and peripheral E-cadherin enhancement were blocked and the induced increase of electrical resistance was markedly blunted. These findings indicate that in EC exposed to hyperosmolar challenge, the involvement of focal adhesion kinase was critical in establishing barrier strengthening.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Endothelial cells (EC)1 lining blood vessels form the principal barrier to fluid flux from blood to tissue. A decrease in barrier properties increases fluid flux and promotes tissue edema that in vital organs such as lung or brain could potentially be life threatening. Permeability of the EC barrier depends largely on the restriction to fluid transport across the paracellular pathway that contains tight junctions and adherens junctions (1). Tight junctions are the primary determinants of barrier function (2). Barrier-deteriorating agents cause EC contraction by activating Ca2+-dependent myosin light chain kinase, thereby widening the junctions and causing hyperpermeability of the EC barrier (3). In a previous study, we reported that exposing lung capillaries to a 1-min hyperosmolar stimulus caused an immediate hyperpermeability response followed by a gradual return of barrier properties to normal (4). These findings suggest that even while challenged by barrier-deteriorating stimuli, EC institute repair mechanisms that reestablish adequacy of the barrier. However, these repair processes remain inadequately understood.

In this regard, the role of focal adhesions requires consideration. EC exposed to barrier deteriorating stimuli develop focal adhesion complexes at points of cell-matrix contact (5, 6). Although the barrier regulatory role of this response remains unclear, evidence from other cell types indicates that focal adhesion formation causes activation of focal adhesion kinase (FAK) (1, 6). It is proposed that in EC exposed to the barrier-deteriorating agent, thrombin, actin-induced translocation of FAK to focal adhesions reduces barrier deterioration (7). The actin cytoskeleton may be particularly relevant in this regard, because receptor-mediated enhancement of actin (8) or actin stabilization by phallicidin (9) strengthens, whereas actin depolymerization by cytochalasins deteriorates the barrier (10). However, the specific mechanisms induced by FAK leading to barrier strengthening in EC remain unclear.

Here we considered the possibility that FAK may be responsible for cross-talk between focal adhesions and cadherins. Dynamic regulation of the EC barrier is attributable to E-cadherin (1, 11, 12) or VE-cadherin (13). At intercellular junctions, the cadherins form homophilic interactions between their extracellular domains on adjacent cell membranes, whereas their cytoplasmic domains bind beta -catenin or plakoglobin (gamma -catenin) that in turn associates with the actin-binding protein, alpha -catenin, thereby establishing a linkage between the cadherin-catenin complex and the actin cytoskeleton (1, 14). Evidence that this linkage is important for barrier regulation comes from findings that barrier-deteriorating stimuli deplete both the cadherin-catenin complex (15) as well as actin (16) from the cell periphery. The FAK substrate alpha -actinin also binds beta -catenin (17), thus providing a link between focal adhesions and the cadherin-catenin complex and thereby raising the possibility that focal adhesion assembly stabilizes the cadherin complex.

We considered these issues in the context of hyperosmolar exposure that causes a cell shrinkage-induced activation of focal adhesion proteins as indicated by increased tyrosine phosphorylation of FAK (18). This response provided an opportunity to test whether in EC FAK is involved in the stabilization of the cadherin complex and possibly of barrier properties. Accordingly, we generated EC expressing a truncated form of deleted FAK (del-FAK) in which cadherin expression in the plasma membrane was markedly diminished. Our findings indicate that although hyperosmolarity increased barrier properties and peripheral cadherin recruitment in wild type EC, both effects were markedly blunted in EC-expressing del-FAK, indicating that EC employ FAK-dependent signaling mechanisms as a means to barrier strengthening.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Materials-- Chemicals were obtained from Sigma unless otherwise stated. Cell culture media and growth supplements, M199 medium, Lipofectin, G418, and Opti-MEM were obtained from Invitrogen. All reagents for immunofluorescence studies were obtained from Molecular Probes Inc. (Eugene, OR). Anti-phosphotyrosine mAb PY99 (mouse and monoclonal) and protein A/G-agarose beads were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-paxillin mAb was purchased from Zymed Laboratories Inc. (South San Francisco, CA). Anti-FAK and E-cadherin mAbs were obtained from Transduction Laboratories, Inc. (Lexington, KY). Anti-FAK polyclonal antibody (BC3) was obtained from Upstate Biotechnology (Lake Placid, NY). Anti-actin rabbit polyclonal antibody and alpha -tubulin mAb were obtained from Sigma.

Cell Culture-- Rat lung microvascular endothelial cells (RLMEC) were cultured as described previously (4, 5) under 5% CO2 in M199 medium supplemented with 5% fetal bovine serum and 5% bovine calf serum. Cells were plated at a density of 1 × 105/cm2. EC phenotype was confirmed by cell uptake of fluorescent labeled-acetylated low density lipoprotein in imaged monolayers.

Transendothelial Electrical Resistance (TER)-- For EC barrier quantification, we determined TER in RLMEC monolayers grown on sterile polycarbonate inserts held at 37 °C (Endohm, World Precision Instruments, Sarasota, FL). After a 30-min base-line period, experimental solutions were added and TER was determined every 15 s for the first 10 min and then at 1-min intervals for the subsequent 20 min. The data were corrected for the resistance of the insert alone.

Plasmid Construction Del-FAK-- The plasmid pBS-FAK (a gift of Dr. James Parsons, Department of Microbiology, School of Medicine, University of Virginia, VA) was prepared by cloning the full-length FAK cDNA into the pBluescript, K5 vector (Stratagene, La Jolla, CA) (19). This plasmid was used for the generation of del-FAK DNA by deleting sequences among the EaeI sites at 1176 and 2793 bp (amino acids 392-931) in the FAK gene (Fig. 1A). This deletion includes a segment containing tyrosine residues that are critical for FAK activation. These residues include Tyr-397 at which FAK autophosphorylates (20), Tyr-576 and Tyr-577 at which phosphorylation determines the kinase activity of FAK (21), and Tyr-925 at which phosphorylation leads to activation of Src (22). Del-FAK DNA (1.6 kb) was subcloned into the plasmid vector pBK-CMV (7.76 kb) at BamH1-XhoI sites and then introduced into the bacterial strain MV10. We selected the clone (kanamycin, 50 µg/ml) containing the 1.6-kb segment (pSLRCU33) corresponding to del-FAK variant (Qiagen, Valencia, CA) using restriction enzymes (BamH1-XhoI). Each 2 µg of plasmid pSLRCU33 or the empty vector was stably transfected in RLMEC using nominal procedures (Lipofectin, Invitrogen).


View larger version (49K):
[in this window]
[in a new window]
 
Fig. 1.   Transfections of del-FAK construct. A, a map of FAK gene shows a segment deleted in the del-FAK construct. The tyrosine (Y) residues, amino acids, and the sites of the sense (S) and antisense (As) primers are indicated. B, G418-resistant EC clones of stably transfected RLMEC were subjected to RT-PCR. Gel shows RT-PCR products for PCR buffer (PCR mix), wild type EC (wt), and cells containing empty vector (vec) or del-FAK.

To confirm transfection of the del-FAK construct in RLMEC, we prepared primers based on the full-length FAK cDNA (GenBankTM accession number M86656) (19): sense primer (873-895 nucleotides), 5'-CCC AGA GGA AGG AAT CAG CTA C-3', and antisense primer (3085-3065 nucleotides), 5'-GCT GGT CAT GAC GTA CTG CTG-3' (Fig. 1A). For PCR, the parameters were: denaturing at 95 °C for 15 min followed by 35 cycles of denaturing, annealing, and extension at 95, 59, and 72 °C for 1, 1, and 2 min, respectively, in 3 mM MgCl2. In the presence of these primers, amplification by RT-PCR is expected to yield a 600-bp product only in cells containing the del-FAK construct (Fig. 1B), confirming successful transfection of the del-FAK construct. In wild type and empty vector-transfected cells, the same primers yielded only the expected RT-PCR product of 2.2 kb (Fig. 1B).

V12Rac1GFP and N17Rac1GFP-- V12Rac1GFP and N17Rac1GFP constructs were generously provided by Dr. P. Jurdic (Laboratoire de Biologie Moléculaire et Cellulaire, Ecole Normale Supérieure de Lyon, Lyon, France) (23). Chimeras between enhanced GFP and GTPases were derived by insertion of the mutated GTPase open reading frames between EcoR1/SalI into the pEGFP-Cl expression vector downstream to the enhanced green fluorescent protein (GFP) coding sequence (Clontech, Palo Alto, CA). The resulting cDNA encoded chimeric GTPase-GFP expression vector. Plasmid DNA, V12Rac1GFP, N17Rac1GFP, and vector pEGFP-Cl were amplified by the standard protocol and adjusted at a final concentration of 1 mg/ml in water. All of the plasmids were expressed by stable transfection using Lipofectin reagent and following the manufacturer's instructions.

Lysate Preparation-- These procedures are routine in our laboratory (4, 5). Cells were exposed to isosmolar medium, or medium was made hyperosmolar by the addition of sucrose (except where stated) at 37 °C under 5% CO2 in M199 for indicated periods. Subsequently, cells were washed twice with ice-cold PBS and lysed in radioimmune precipitation assay buffer (50 mM Tris-HCl, pH 7.6, 150 mM NaCl, 2 mM EDTA, 50 mM NaF, 0.5% SDS, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 1 mM phenylmethylsulfonyl fluoride) containing 1 mM sodium orthovanadate, 10 mM sodium pyrophosphate, 25 mM beta -glycerophosphate, 0.1% SDS, and 1% Triton X-100 at 4 °C. Total cell lysate was clarified by centrifugation at 10,000 × g for 10 min. Protein concentrations were determined using a protein analysis kit (BCA, Pierce).

Cell Fractionation-- Cells were rinsed 2× PBS and solubilized in Triton X-100 buffer (50 mM NaCl, 10 mM Pipes, pH 6.8, 3 mM MgCl2, 0.5% Triton X-100, 300 mM sucrose, 1.2 mM phenylmethylsulfonyl fluoride, 10 µg/ml leupeptin) for 20 min at 4 °C on a rocking platform. The cells were scraped from the plate and centrifuged (10 min). The supernatant was collected. The cell pellet was suspended in 100 µl of SDS immunoprecipitation buffer (15 mM Tris, pH 7.5, 5 mM EDTA, 2.5 mM EGTA, 1% SDS) and then boiled for 10 min and diluted to 300 µl with Triton X-100 buffer. Equal amounts of extracted proteins were immunoprecipitated and loaded for SDS-PAGE gel electrophoresis.

Immunoprecipitation and Immunoblotting-- Immunoprecipitation and immunoblotting were performed as described previously (4). Cell lysates containing equal amounts of protein were precleared for 30 min with 20 µl of protein A/G-agarose beads followed by incubation with primary antibodies (4 µg for 2 h). Antibody-antigen complexes were precipitated with 30 µl of protein A/G-agarose beads overnight at 4 °C. Nonspecific bound proteins were removed by washing the agarose beads three times with radioimmune precipitation assay buffer and one time with PBS. Bound proteins were eluted in 40 µl of 4× Laemmli's loading buffer. The proteins were resolved by SDS-polyacrylamide gel electrophoresis, blotted onto nitrocellulose membrane, and analyzed by immunoblotting.

FAK Activity-- For the analysis of FAK autophosphorylation, we used the reported immune complex kinase assay (21). We immunoprecipitated p125FAK from wild type or del-FAK-transfected RLMEC. The immunoprecipitated complexes were washed three times with radioimmune precipitation assay buffer and once with kinase buffer (pH 7.4, 20 mM Hepes, 50 mM NaCl, 5 mM MgCl2, 5 mM MnCl2). The pellet was resuspended in 30 µl of the kinase buffer containing 10 µCi of [gamma -32P]ATP (6000 Ci/mmol), and the sample was incubated for 30 min at 37 °C with frequent agitation. The reaction was terminated by the addition of 10 µl of 4× SDS-PAGE gel sample buffer followed by boiling for 5 min, and all of the products were resolved by SDS-polyacrylamide gel electrophoresis and transferred to a nitrocellulose membrane (Bio-Rad). Direct exposures were used to visualize FAK autophosphorylation. The amount of FAK present in the reactions was visualized by immunoblotting of the same membrane with anti-FAK monoclonal antibody.

Rac Activation Assay-- Using glutathione S-transferase-tagged p21- activated kinase-p21 binding domain (PAK-PBD) protein beads that specifically bind active Rac1, we obtained immunoprecipitates from EC lysates as per the methods reported previously (24) using the assay kit from Cytoskeleton Inc. (Denver, CO). For positive and negative controls for the active and inactive small GTPases, the non-hydrolyzable GTP analog, GTPgamma S, and GDP were used, respectively. Cell lysates obtained from untreated cells were incubated with 100 µM GTPgamma S or 1 mM GDP in the presence of 10 mM EDTA for 15 min at 30 °C to ensure efficient loading with the added nucleotide. To terminate the reaction, the lysates were placed on ice and supplemented with 60 mM MgCl2. These control samples were then incubated with the glutathione S-transferase-tagged PAK-PBD protein beads and washed in the same manner as the other samples. Captured proteins were removed from the beads by boiling the samples in Laemmli buffer, and the samples were subjected to SDS-PAGE and Western blotting.

Immunofluorescence and Confocal Microscopy-- RLMEC monolayers grown on glass coverslips were fixed (4% formaldehyde in PBS, pH 7.4, 20 min, 22 °C), rinsed (3× PBS), permeabilized (0.1% Triton X-100), and stained using rhodamine-phalloidin. For immunofluorescence, cells were incubated with diluted primary antibodies (1:50) in blocking solution, 4% goat serum in PBS (1 h at 22 °C), and washed with 3× PBS. Fluorescence-conjugated antibodies then were added (1:500, 1 h at 22 °C) and washed with 3× PBS. The glass coverslips were mounted upside down on object slides using fluorescent-mounting medium (Dako Corporation Carpinteria, CA). Confocal images were obtained by means of a laser-scanning microscope (Pascal LSM, Carl Zeiss) and subjected to image analysis as described below (MCID 5, Imaging Research, St. Catherine, Canada).

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Barrier Response-- TER of RLMEC monolayers was stable at 37 ± 3 ohm/cm2 for up to 40 min (mean ± S.E., n = 6). Exposure of monolayers to isosmolar medium (300 mosM) did not change TER from base line (Fig. 2A). However, in monolayers exposed to medium-made hyperosmolar by the addition of sucrose (350 mosM), TER decreased in the first minute and then increased in the subsequent ~10 min (Fig. 2A). The increase was sustained at a steady level for 5-7 min after which TER gradually returned to base line in 20-30 min. The increase of TER was abrogated by the addition of isosmolar medium (Fig. 2B), and it could be repeated subsequently in the same monolayer (data not shown). These findings indicated that subsequent to an initial decrease of barrier properties, the dominant effect of hyperosmolar exposure was to enhance the EC barrier.


View larger version (21K):
[in this window]
[in a new window]
 
Fig. 2.   TER responses to hyperosmolarity in RLMEC. A and B, single tracings exemplify responses to sucrose medium (added at arrows) at indicated levels of hyperosmolarity and replicated six times each. C, group responses to 350 mosM sucrose medium in monolayers treated for 30 min with each of the indicated treatments. Each bar is mean ± S.E. for four experiments. D, maximum TER increase is in relation to base-line value. Each point is mean ± S.E. for seven experiments.

Because hyperosmolar exposure increases protein tyrosine phosphorylation in EC (18) and because actin polymerization is proposed to cause barrier strengthening (8, 9), we exposed monolayers to inhibitors of tyrosine kinase and actin polymerization. The TER increase was inhibited by the tyrosine kinase inhibitor, genistein, and by the inhibitors of actin polymerization, latrinculin B (Fig. 2C) and cytochalasin D (data not shown). Although hyperosmolar exposure is reported to increase the cytosolic Ca2+ (25), the intracellular calcium chelator, BAPTA-AM had no effect on the present TER response to hyperosmolarity (Fig. 2C). Not shown are results from experiments in which we incubated monolayers separately with the p38 mitogen-activated protein kinase blocker SB203580 (25 µM), the protein kinase C blocker calphostin C (500 nM), or the phosphatidylinositol 3-kinase blocker wortmannin (50 nM) (n = 3 for each inhibitor). A single monolayer was pretreated with the nitric-oxide synthase inhibitor L-NAME (30 µM). None of these treatments affected the TER response to hyperosmolarity. The maximum increase in TER correlated non-linearly with an osmolar concentration (Fig. 2D) with 80% of the response being established at 350 mosM. Hence, for the studies described below, we exposed EC to this hyperosmolar concentration for 15 min.

Focal Adhesion Proteins-- Because genistein blocked the TER response to hyperosmolarity, we considered that hyperosmolar cell shrinkage might increase cell-matrix interactions, leading to activation of focal adhesion proteins. By confocal microscopy of wild type cells under base-line conditions, immunofluorescence of the focal adhesion marker protein paxillin was largely localized to the nuclear and perinuclear regions (Fig. 3A, left image). Following hyperosmolar exposure, the fluorescence became pronounced as aggregates localized to the cell periphery in a pattern characteristic of focal adhesion formation (Fig. 3A, right image) (6). This focal adhesion response was absent in del-FAK-expressing monolayers (Fig. 3B). Exposure of plated EC to hyperosmolar sucrose also increased tyrosine phosphorylation of FAK and paxillin (Fig. 3C). FAK activity, which was less in del-FAK-transfected monolayers than in wild type monolayers under base-line conditions (Fig. 3, D and E), increased 3-fold in wild type monolayers exposed to hyperosmolar medium (Fig. 3, D and E). By contrast, hyperosmolar exposure caused no enhancement of FAK activity in del-FAK-transfected cells (Fig. 3, D and E). Under non-stimulated conditions, TER was 30 ± 5% less in del-FAK-expressing monolayers than in wild type monolayers or in monolayers expressing vector alone (p < 0.05; n = 4). Following hyperosmolar exposure, the increase of TER was blunted in del-FAK-expressing monolayers to 60 ± 5% of that of wild type monolayers (Fig. 3F). Taking these findings together, we interpret that hyperosmolar exposure increased focal adhesion formation and that inhibition of this response inhibited the hyperosmolarity-induced barrier enhancement.


View larger version (34K):
[in this window]
[in a new window]
 
Fig. 3.   Responses to hyperosmolarity in del-FAK-transfected and wild type RLMEC. EC were exposed to isosmolar (300 mosM) or hyperosmolar (350 mosM) conditions for 15 min each as indicated. A and B, immunofluorescence of paxillin (arrows) shown in images obtained by confocal microscopy in wild type (A) and del-FAK-expressing (B) cells. Monolayers were fixed and permeabilized and then stained with mouse anti-paxillin mAb followed by Alexa Red-linked anti-mouse IgG with each set replicated four times. C, plated EC were subjected to immunoprecipitation (IP) using indicated antibodies. Immunoblots were obtained using anti-phosphotyrosine antibody. Immunoblots using the indicated protein-specific antibodies confirm loading of equal protein amounts in each lane with each set replicated four times. D, gel shows enzymatic activity of FAK autophosphorylation as determined by immune complex kinase assay of FAK immunoprecipitates from EC lysates. Upper panel shows FAK activity. Lower panel shows the amount of FAK present in the reactions as indicated by immunoblotting the same membrane with anti-FAK mAb. E, bar diagram shows FAK activity as optical densities of bands relative to protein content (n = 3 for each bar). F, TER responses to hyperosmolar (350 mosM) exposure in wild type (wt) monolayers and in monolayers transfected with vector alone (vec) or FAK mutant (del-FAK). n = 4 for each bar; mean ± S.E., *, p < 0.01 against bar at left.

To further consider the role of the cytosolic Ca2+, we carried out immunoprecipitations from monolayers treated with the intracellular Ca2+ chelator BAPTA-AM. Predictably, BAPTA-AM blocked the tyrosine phosphorylation of the Ca2+-dependent focal adhesion protein Pyk2 (Fig. 4A) (26), but it did not modify phosphorylation responses for FAK or paxillin (data not shown). These results together with the above lack of an inhibitory effect on TER by BAPTA-AM indicate that the hyperosmolarity-induced TER increase was Ca2+-independent. In EC lysates, bands were evident at 125 and 68 kDa (Fig. 4, B and C), corresponding to FAK and paxillin, respectively (Fig. 3A). However, as opposed to the responses in plated EC, no increase of tyrosine phosphorylation was evident in lysates prepared either from suspended EC exposed to hyperosmolar sucrose (Fig. 4B, lanes 3 and 4) or from plated EC exposed to hyperosmolar urea (Fig. 4C, lane 3). The implications of these findings are discussed below.


View larger version (19K):
[in this window]
[in a new window]
 
Fig. 4.   Effects of cell suspension, urea, and intracellular Ca2+ chelation in RLMEC. A, plated EC exposed for 15 min to conditions shown were subjected to immunoprecipitation (IP) using anti-Pyk2 mAb antibodies. Immunoblots are for phosphotyrosine (upper panels) and protein (lower panels) contents. BAPTA-AM (100 µM) with each set replicated three times. B and C, data shown for plated EC (monolayer) or EC held in suspension in medium (suspended). Medium was made hyperosmolar by adding sucrose except where stated (urea). Cell lysates were subjected to SDS-PAGE, transfer, and immunoblotting (IB) using anti-phosphotyrosine antibody (Tyrp), and each set replicated three times each.

E-cadherin-- In considering the barrier regulatory function of FAK, we addressed the role of cadherins. Immunoprecipitation studies using mAbs against either VE- or E-cadherin indicated that although E-cadherin was well expressed in RLMEC (Fig. 5A), the expression of VE-cadherin was weak (Fig. 5B), indicating that E-cadherin was the dominant cadherin type expressed in these EC. We confirmed that the mAb against VE-cadherin was capable of recognizing rat antigens (Fig. 5B) and that as reported by others (27) it recognized a band in human umbilical vein endothelial cells (Fig. 5B).


View larger version (33K):
[in this window]
[in a new window]
 
Fig. 5.   E-cadherin expression in RLMEC. A and B, gels show immunoprecipitation (IP) and immunoblot (IB) with anti E-cadherin (A) or anti-VE-cadherin (B) mAb. Wild type (wt), del-FAK-transfected (del-FAK), and vector-transfected (vec) cells were exposed to isosmolar (300 mosM) or hyperosmolar (350 mosM) sucrose medium for 15 min. Monolayer lysates were detergent-fractionated into cytosolic (C) and membrane (M) fractions, and each set replicated three times. C, bar diagram shows E-cadherin as optical densities of bands (n = 3 for each bar). p < 0.05 compared with corresponding value for 300 mosM (*) or the corresponding cytosolic value (#). D and E, E-cadherin distribution (arrows) shown in images obtained by confocal microscopy in wild type (D) and del-FAK-expressing (E) cells. Monolayers were fixed and permeabilized and then stained with mouse anti-E-cadherin mAb followed by Alexa Red-linked anti-mouse IgG. Each set replicated four times. HUVEC, human umbilical vein endothelial cells.

Detergent-based fractionation of EC lysates into cytosolic and membrane fractions followed by quantitative immunoprecipitation and immunoblotting revealed that in wild type EC, E-cadherin content was greater in the membrane than the cytosolic fraction (Fig. 5A, lanes 1 and 2). Hyperosmolar exposure increased the membrane content (Fig. 5A, lanes 2 and 4) while decreasing the cytosolic content (Fig. 5A, lanes 1 and 3). These compartmental changes were approximately equal as quantified by band densitometry (Fig. 5C). Accordingly, hyperosmolarity increased the membrane-cytosol ratio for E-cadherin (Fig. 5C). In del-FAK-expressing cells, E-cadherin content was less under control conditions than in wild type cells (p < 0.01) (Fig. 5, A and C). Moreover, the membrane content was less than the cytosolic (Fig. 5A, lanes 5 and 6, and C). Furthermore, in contrast to the wild type response, in del-FAK-expressing cells, E-cadherin failed to increase following hyperosmolar exposure (Fig. 5A, lanes 6 and 8), resulting in similar membrane-cytosol ratios under control and hyperosmolar conditions (Fig. 5C). Responses in monolayers expressing the empty vector were similar to those of wild type monolayers (Fig. 5A, lanes 9-12, and C).

In wild type cells under base-line conditions, confocal microscopy revealed the distribution of E-cadherin as a discontinuous line of fluorescence that marked the cell periphery (Fig. 5D, left image). Following hyperosmolar exposure, this peripheral fluorescence became pronounced and was now evident as a continuous line (Fig. 5D, right image), indicating increased E-cadherin expression. In del-FAK-expressing monolayers, the peripheral fluorescence was poorly developed under both base-line and hyperosmolar conditions (Fig. 5E), indicating abrogation of E-cadherin expression and pointing to FAK as critical factor in the hyperosmolarity-induced enhancement of E-cadherin.

Actin-- Because latrinculin B inhibited the hyperosmolarity-induced TER increase (Fig. 2C), we considered the involvement of the actin cytoskeleton in the present barrier response. Confocal microscopy of untreated wild type cells revealed actin distribution as reflected in rhodamine-phalloidin fluorescence as a thin band at the cell periphery and a perinuclear condensation (Fig. 6A, left image). A 15-min hyperosmolar exposure markedly enhanced the density of filamentous actin in wild type cells but not in del-FAK-expressing cells (Fig. 6, A-C). Immunoblotting with an actin-recognizing mAb indicated that, as expected, actin content was higher in the membrane than in the cytosolic fraction under control conditions (Fig. 7, A and B). In wild type cells, hyperosmolar exposure increased membrane actin content almost 2-fold (Fig. 7B, compare second and fourth bars) while decreasing the cytosolic content (Fig. 7B, compare first and third bars). However, in del-FAK-expressing cells, these responses were completely blocked (Fig. 7, A and B).


View larger version (34K):
[in this window]
[in a new window]
 
Fig. 6.   Immunofluorescence of actin in RLMEC. A and B, images obtained by confocal microscopy images show fluorescence of rhodamine-phalloidin depicting distribution of actin in wild type (A) and del-FAK-transfected (B) cells. As indicated, monolayers were exposed to isosmolar (300 mosM) or hyperosmolar (350 mosM) sucrose medium for 15 min. Increase of filamentous actin is indicated by double-headed arrow. Each set replicated four times. C, determinations of fluorescence intensity by image analysis. Linear fluorescence was determined along a 5-µm-wide band that followed the cell contour midway between the periphery and the center (white line in inset). Data are for wild type (wt) and del-FAK-expressing (del-FAK) cells exposed to isosmolar (300 mosM) or hyperosmolar (350 mosM) conditions. Mean ± S.E.; n = 4 for each bar. *, p < 0.05 compared with control (300 mosM).


View larger version (29K):
[in this window]
[in a new window]
 
Fig. 7.   Compartmental actin distribution in RLMEC. A, monolayer lysates were detergent-treated to obtain cytosolic (C) and membrane (M) fractions. Gels show immunoblots (IB) conducted sequentially on same membrane with anti-actin (actin) and anti-alpha -tubulin (tubulin) antibodies. Immunoblot with anti-alpha -tubulin mAb confirms equal protein loading in each lane. B, bars show densitometric analyses of bands in A expressed as optical density normalized to alpha -tubulin density. For each paired determination, density was higher for membrane than cytosolic fractions (p < 0.05). *, p < 0.05 compared with 300 mosM. Mean ± S.E.; n = 4 for each bar.

To determine mechanisms underlying actin assembly, we considered the role of the small GTPase, Rac1 that regulates actin polymerization (8, 28). We immunoprecipitated increased amounts of active Rac1 from cells exposed to hyperosmolar sucrose (Fig. 8A), which indicated that hyperosmolar exposure increased Rac1 activity in these cells. To modify Rac1 activity, we expressed GFP-tagged constitutively active (V12Rac1) or dominant negative (N17Rac1) Rac1 mutants in EC monolayers. These isoforms enhance and reduce actin filament formation, respectively (8, 28). Responses in RLMEC are exemplified in the images shown in Fig. 8B. Under isosmolar conditions, actin density was higher in V12Rac1-expressing monolayers than in monolayers expressing N17Rac1 or vector alone. Following hyperosmolar exposure, actin density increased in cells expressing empty vector, which is consistent with the above immunoprecipitation data in which hyperosmolar exposure increased membrane actin. Hyperosmolarity caused greater increases in actin filaments in V12Rac1-expressing cells than in vector controls. The overlay of rhodamine on GFP resulted in yellow discoloration along the cell margin in V12Rac1- and N17Rac1-transfected cells, indicating that transfected Rac1 was targeted to the cell periphery. In contrast, hyperosmolarity failed to increase actin density in cells expressing N17Rac1, indicating that Rac1 activation was critical for the actin response to hyperosmolar exposure (28). However, despite these differences, TER was unaffected by the expression of either V12Rac1 or N17Rac1 in both base-line and hyperosmolar conditions (Fig. 8C). We interpret from these findings that actin assembly was irrelevant in the barrier-enhancing response to hyperosmolarity.


View larger version (54K):
[in this window]
[in a new window]
 
Fig. 8.   Rac activity in RLMEC. Wild type (wt), constitutively active Rac1 (V12Rac1GFP) or dominant negative Rac1 (N17Rac1GFP) or empty vector pEGFP (vec) transfected cells were exposed to isosmolar (300 mosM) or hyperosmolar (350 mosM) sucrose medium for 15 min. A, upper panel shows bands for active Rac1 immunoprecipitated from EC lysates using glutathione S-transferase-tagged PAK-PBD protein beads. Bands for GDP and GTPgamma S denote negative and positive controls obtained, respectively, in untreated cells. Immunoblots using anti-Rac1 polyclonal antibody in the lower panel indicate that lysate samples contained equal amounts of Rac1 protein prior to immunoprecipitation. Each set replicated three times. B, images show fluorescence in single cells of monolayers for rhodamine-phalloidin (rhod) and GFP (GFP). Arrows point to actin filaments. Double-headed arrows indicate site of increased actin density. Yellow pseudocolor in the overlay panel indicates co-localization of filamentous actin with the transfected vector (double arrows). Each set replicated three times. C, TER responses to hyperosmolar (350 mosM) exposure in wild type-, vec-, V12Rac1-, and N17Rac1-transfected cells. n = 4 for each bar; mean ± S.E.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Our findings may be summarized as follows. A 15-min exposure of EC to hyperosmolarity resulted in (i) an immediate decrease and then a relatively sustained increase of TER, (ii) increases in focal adhesion formation and FAK activity, (iii) increased membrane accumulation of E-cadherin, (iv) increased Rac1 activity, and (v) increased density of filamentous actin.

We conclude from these findings that the dominant effect of hyperosmolarity was to enhance EC barrier properties. This counterintuitive interpretation opposes the expectation that in the presence of external hyperosmolarity, cell water extraction leading to cell shrinkage should widen intercellular junctions and consequently decrease the barrier (4). Although such an effect did in fact occur as indicated by the initial decrease in TER, the effect was short-lived and lasted for only the first minute of the response. However, this initial cell shrinkage was critical for the subsequent barrier enhancement as was evident when we replaced sucrose with urea, the membrane-permeable osmolyte that does not extract cell water. Hyperosmolar urea failed to alter EC barrier properties, indicating that cell shrinkage initiated the complex barrier response to hyperosmolar sucrose. An important consequence of cell shrinkage was the induction of tyrosine phosphorylation in plated EC. Although this result is similar to previous findings that we (4) as well as others (18) have reported in lung microvascular and aortic EC, no hyperosmolarity-induced increase of tyrosine phosphorylation occurred in EC suspended in buffer. These results indicate that cell shrinkage alone was not sufficient to initiate signaling but that cell-matrix interactions were a key element in this response.

The Role of FAK-- In blood vessels, mechanical stresses such as flow-induced shear and mechanical stretch cause EC-matrix interactions, leading to activation of focal adhesion proteins and enhanced protein tyrosine phosphorylation (29). Here, hyperosmolar exposure enhanced focal adhesion formation, tyrosine phosphorylation of FAK and paxillin, and FAK activity. These responses suggest that focal adhesions were induced following hyperosmolarity-induced cell shrinkage, possibly as a result of displacement of the cell membrane on the underlying matrix. In an analogous mechanism, cell contraction induces focal adhesion formation in tracheal smooth muscle cells (30). In del-FAK-expressing EC, the hyperosmolarity-induced enhancement of the barrier was markedly blunted, indicating that FAK was critical in this barrier-strengthening response.

The presence of cross-talk between FAK and the cadherin complex was indicated in that hyperosmolarity also induced the increased expression of E-cadherin at the cell periphery. This increase occurred within the duration of the TER increase, suggesting that the peripheral E-cadherin enrichment was responsible for the strengthening of adherens junctions (11, 12). The link to focal adhesions was indicated in that in del-FAK-expressing cells, the hyperosmolarity-induced peripheral E-cadherin enrichment was markedly diminished. Taking this result together with the diminished barrier enhancement response in del-FAK-expressing cells, we interpret that FAK-dependent junctional E-cadherin enrichment induced barrier enhancement.

According to the cell contraction hypothesis of vascular permeability, deterioration of the EC barrier is determined importantly by myosin light chain kinase activation that is itself induced by an increase in the cytosolic Ca2+ (3). The intracellular Ca2+ chelator, BAPTA-AM, blocks increases of cytosolic Ca2+ and inhibits Ca2+-induced barrier deterioration (3). However, in the present experiments, BAPTA-AM modified neither the hyperosmolarity-induced TER response nor the associated enhancements of tyrosine phosphorylation on FAK and paxillin. Nevertheless, the Ca2+-blocking effect of this agent was evident in that BAPTA-AM blocked the hyperosmolarity-induced enhancement of tyrosine phosphorylation of the Ca2+-sensitive kinase Pyk2. These findings indicate that the present barrier enhancement occurred through Ca2+-independent mechanisms.

Actin-- A striking result in these EC was that hyperosmolar exposure increased the density of filamentous actin as indicated in the immunoprecipitation data for membrane actin as well as by confocal microscopy of actin immunofluorescence. Increases in filamentous actin have been reported following exposure of Swiss T3 cells to platelet-derived growth factor (31), neutrophils to hyperosmolarity (24), and pulmonary artery EC to sphingosine 1-phosphate (8). The polymerization of actin is attributable to Rac1, a member of the Rho family of small GTPases that is activated by phosphatidylinositol 3-kinase (32). Cell expression of a constitutively active form of Rac (V12Rac1) increases filamentous actin by activating LIM kinase that phosphorylates cofilin, thereby inhibiting actin depolymerization (33). The findings in mouse fibroblasts indicate that Rac1 also stabilizes the cadherin-catenin complex by inhibiting the interaction of IQGAP1 with beta -catenin (34, 35) and by activating the p21-activated protein kinase (36). However, the understanding of these interactions in the context of EC barrier regulation is complicated by the proposal that the expression of either constitutively active or inactive Rac deteriorates barrier properties in aortic EC (37) and that overexpression of Rac destabilizes actin filaments in transformed epithelial cells (38).

Here, in agreement with reported findings in neutrophils (24), hyperosmolar exposure increased Rac1 activity in EC monolayers. Furthermore, the hyperosmolarity-induced increase of actin density was inhibited in EC-expressing N17Rac1 but augmented in EC-expressing V12Rac1, consistent with the interpretation that Rac activation was responsible for the actin enhancement (8, 28). However, the phosphatidylinositol 3-kinase inhibitor, wortmannin, failed to modify the hyperosmolarity-induced TER enhancement, thereby suggesting that this kinase played no role in the present barrier response. Moreover, in V12Rac1- and N17Rac1-expressing cells, TER was not affected either under unstimulated conditions or during hyperosmolar exposure. We conclude from these findings that despite the present evidence for Rac1 activation and actin enhancement, these factors played no role in the hyperosmolarity-induced barrier strengthening.

The bulk of the understanding of actin involvement in barrier regulation comes from the application of the actin inhibitors, the latrinculins and the cytochalasins (7, 10, 27). Several groups have reported that the reduction of polymerized actin in EC caused by these agents associates with the deterioration of EC barrier properties (8, 10). Our findings are consistent with these reports, because both agents not only reduced TER at base-line (data not shown) but they also abrogated the TER increase during hyperosmolar exposure. However, these pharmacological data were not supported by TER responses in V12Rac1- and N17Rac1-expressing cells, suggesting that pharmacological inhibitors of actin polymerization may induce barrier effects through nonspecific mechanisms.

In conclusion, our results provide the first evidence that focal adhesion formation and FAK involvement are critical events leading to barrier-strengthening processes in EC. Although the role of the associated increase in actin filaments remains unclear, we believe that this response did not determine the barrier strengthening. It is possible that these EC developed the actin response to increase cell rigidity and thereby oppose imposed shape changes forced by hyperosmolar cell shrinkage (39). The ability of FAK to regulate E-cadherin accumulation at the cell periphery suggests that FAK serves as a determining factor in the induction of barrier integrity. Although a further understanding of this signaling pathway is required, we propose that the present evidence for EC barrier enhancement may provide a basis for the consideration of hyperosmolar therapy in the treatment of vascular leak syndromes.

    ACKNOWLEDGEMENTS

We thank R. Kashyap and R. Patel for assistance with the experiments and Dr. Sunita Bhattacharya for reading the paper.

    FOOTNOTES

* This work was supported by National Institutes of Health Grants HL36024, HL57556, and HL64896.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.

To whom correspondence should be addressed: St. Luke's-Roosevelt Hospital Center, 1000 10th Ave., New York, NY 10019. Tel.: 212-523-7310; Fax: 212-523-8005; E-mail: jb39@columbia.edu.

Published, JBC Papers in Press, January 28, 2003, DOI 10.1074/jbc.M209922200

    ABBREVIATIONS

The abbreviations used are: EC, endothelial cells; FAK, focal adhesion kinase; mAb, monoclonal antibody; RLMEC, rat lung microvascular endothelial cells; TER, trans-endothelial electrical resistance; del-FAK, deleted FAK; GFP, green fluorescent protein; RT, reverse transcriptase; PBS, phosphate-buffered saline; Pipes, 1,4-piperazinediethanesulfonic acid; GTPgamma S, guanosine 5'-O-(thiotriphosphate); BAPTA-AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N',-tetraacetic acid tetrakis(acetoxymethyl ester).

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

1. Gumbiner, B. M. (1996) Cell 84, 345-357[Medline] [Order article via Infotrieve]
2. Tsukita, S., Furuse, M., and Itoh, M. (2001) Nat. Rev. Mol. Cell. Biol. 2, 285-293[CrossRef][Medline] [Order article via Infotrieve]
3. Garcia, J. G., and Schaphorst, K. L. (1995) J. Invest. Med. 43, 117-126[Medline] [Order article via Infotrieve]
4. Ragette, R., Fu, C., and Bhattacharya, J. (1997) J. Clin. Invest. 100, 685-692[Abstract/Free Full Text]
5. Bhattacharya, S., Fu, C., Bhattacharya, J., and Greenberg, S. (1995) J. Biol. Chem. 270, 16781-16787[Abstract/Free Full Text]
6. Abedi, H., and Zachary, I. (1997) J. Biol. Chem. 272, 15442-15451[Abstract/Free Full Text]
7. Mehta, D., Tiruppathi, C., Sandoval, R., Minshall, R. D., Holinstat, M., and Malik, A. B. (2002) J. Physiol. (Lond.) 539, 779-789[Abstract/Free Full Text]
8. Garcia, J. G., Liu, F., Verin, A. D., Birukova, A., Dechert, M. A., Gerthoffer, W. T., Bamberg, J. R., and English, D. (2001) J. Clin. Invest. 108, 689-701[Abstract/Free Full Text]
9. Phillips, P. G. (1994) Semin. Thromb. Hemostasis 20, 417-425[Medline] [Order article via Infotrieve]
10. Shasby, D. M., Shasby, S. S., Sullivan, J. M., and Peach, M. J. (1982) Circ. Res. 51, 657-661[Abstract]
11. Rubin, L. L., Hall, D. E., Porter, S., Barbu, K., Cannon, C., Horner, H. C., Janatpour, M., Liaw, C. W., Manning, K., Morales, J., Tanner, L. I., Tomaselli, K. J., and Bard, F. (1991) J. Cell Biol. 115, 1725-1735[Abstract]
12. Moy, A. B., Winter, M., Kamath, A., Blackwell, K., Reyes, G., Giaever, I., Keese, C., and Shasby, D. M. (2000) Am. J. Physiol. 278, L888-L898
13. Lampugnani, M. G., Resnati, M., Raiteri, M., Pigott, R., Pisacane, A., Houen, G., Ruco, L. P., and Dejana, E. (1992) J. Cell Biol. 118, 1511-1522[Abstract]
14. Conacci-Sorrell, M., Zhurinsky, J., and Ben Ze'ev, A. (2002) J. Clin. Invest. 109, 987-991[Free Full Text]
15. Rabiet, M. J., Plantier, J. L., Rival, Y., Genoux, Y., Lampugnani, M. G., and Dejana, E. (1996) Arterioscler. Thromb. Vasc. Biol. 16, 488-496[Abstract/Free Full Text]
16. Ehringer, W. D., Yamany, S., Steier, K., Farag, A., Roisen, F. J., Dozier, A., and Miller, F. N. (1999) Microcirculation 6, 291-303[CrossRef][Medline] [Order article via Infotrieve]
17. Tsukatani, Y., Suzuki, K., and Takahashi, K. (1997) J. Cell. Physiol. 173, 54-63[CrossRef][Medline] [Order article via Infotrieve]
18. Malek, A. M., Goss, G. G., Jiang, L., Izumo, S., and Alper, S. L. (1998) Stroke 29, 2631-2640[Abstract/Free Full Text]
19. Schaller, M. D., Borgman, C. A., Cobb, B. S., Vines, R. R., Reynolds, A. B., and Parsons, J. T. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 5192-5196[Abstract]
20. Schaller, M. D., Hildebrand, J. D., Shannon, J. D., Fox, J. W., Vines, R. R., and Parsons, J. T. (1994) Mol. Cell. Biol. 14, 1680-1688[Abstract]
21. Calalb, M. B., Polte, T. R., and Hanks, S. K. (1995) Mol. Cell. Biol. 15, 954-963[Abstract]
22. Schlaepfer, D. D., Hanks, S. K., Hunter, T., and van der, G. P. (1994) Nature 372, 786-791[Medline] [Order article via Infotrieve]
23. Ory, S., Munari-Silem, Y., Fort, P., and Jurdic, P. (2000) J. Cell Sci. 113, 1177-1188[Abstract/Free Full Text]
24. Lewis, A., Di Ciano, C., Rotstein, O. D., and Kapus, A. (2002) Am. J. Physiol. 282, C271-C279[Medline] [Order article via Infotrieve]
25. Paemeleire, K., de Hemptinne, A., and Leybaert, L. (1999) Exp. Brain Res. 126, 473-481[CrossRef][Medline] [Order article via Infotrieve]
26. Tokiwa, G., Dikic, I., Lev, S., and Schlessinger, J. (1996) Science 273, 792-794[Abstract]
27. Hinck, L., Nathke, I. S., Papkoff, J., and Nelson, W. J. (1994) J. Cell Biol. 125, 1327-1340[Abstract]
28. Takaishi, K., Sasaki, T., Kotani, H., Nishioka, H., and Takai, Y. (1997) J. Cell Biol. 139, 1047-1059[Abstract/Free Full Text]
29. Chien, S., Li, S., and Shyy, Y. J. (1998) Hypertension 31, 162-169[Abstract/Free Full Text]
30. Gerthoffer, W. T., and Gunst, S. J. (2001) J. Appl. Physiol. 91, 963-972[Abstract/Free Full Text]
31. Machesky, L. M., and Hall, A. (1997) J. Cell Biol. 138, 913-926[Abstract/Free Full Text]
32. Kovacs, E. M., Ali, R. G., McCormack, A. J., and Yap, A. S. (2002) J. Biol. Chem. 277, 6708-6718[Abstract/Free Full Text]
33. Yang, N., Higuchi, O., Ohashi, K., Nagata, K., Wada, A., Kangawa, K., Nishida, E., and Mizuno, K. (1998) Nature 393, 809-812[CrossRef][Medline] [Order article via Infotrieve]
34. Kuroda, S., Fukata, M., Nakagawa, M., Fujii, K., Nakamura, T., Ookubo, T., Izawa, I., Nagase, T., Nomura, N., Tani, H., Shoji, I., Matsuura, Y., Yonehara, S., and Kaibuchi, K. (1998) Science 281, 832-883[Abstract/Free Full Text]
35. Fukata, M., Kuroda, S., Nakagawa, M., Kawajiri, A., Itoh, N., Shoji, I., Matsuura, Y., Yonehara, S., Fujisawa, H., Kikuchi, A., and Kaibuchi, K. (1999) J. Biol. Chem. 274, 26044-26050[Abstract/Free Full Text]
36. Zhao, Z. S., Manser, E., Chen, X. Q., Chong, C., Leung, T., and Lim, L. (1998) Mol. Cell. Biol. 18, 2153-2163[Abstract/Free Full Text]
37. Wojciak-Stothard, B., Potempa, S., Eichholtz, T., and Ridley, A. J. (2001) J. Cell Sci. 114, 1343-1355[Abstract/Free Full Text]
38. Quinlan, M. P., and Hyatt, J. L. (1999) Cell Growth Differ. 10, 839-854[Abstract/Free Full Text]
39. Glogauer, M., Arora, P., Yao, G., Sokholov, I., Ferrier, J., and McCulloch, C. A. (1997) J. Cell Sci. 110, 11-21[Abstract/Free Full Text]


Copyright © 2003 by The American Society for Biochemistry and Molecular Biology, Inc.