Am J Physiol Cell Physiol Journal of Applied Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Cell Physiol 294: C324-C332, 2008. First published October 31, 2007; doi:10.1152/ajpcell.00319.2007
0363-6143/08 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
294/1/C324    most recent
00319.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Müller-Marschhausen, K.
Right arrow Articles by Drenckhahn, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Müller-Marschhausen, K.
Right arrow Articles by Drenckhahn, D.

VASCULAR BIOLOGY

Physiological hydrostatic pressure protects endothelial monolayer integrity

K. Müller-Marschhausen,* J. Waschke,* and D. Drenckhahn

Institute of Anatomy and Cell Biology, University of Würzburg, Würzburg, Germany

Submitted 23 July 2007 ; accepted in final form 26 October 2007


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Endothelial monolayer integrity is required to maintain endothelial barrier functions and has found to be impaired in several disorders like inflammatory edema, allergic shock, or artherosclerosis. Under physiologic conditions in vivo, endothelial cells are exposed to mechanical forces such as hydrostatic pressure, shear stress, and cyclic stretch. However, insight into the effects of hydrostatic pressure on endothelial cell biology is very limited at present. Therefore, in this study, we tested the hypothesis that physiological hydrostatic pressure protects endothelial monolayer integrity in vitro. We investigated the protective efficacy of hydrostatic pressure in microvascular myocardial endothelial (MyEnd) cells and macrovascular pulmonary artery endothelial cells (PAECs) by the application of selected pharmacological agents known to alter monolayer integrity in the absence or presence of hydrostatic pressure. In both endothelial cell lines, extracellular Ca2+ depletion by EGTA was followed by a loss of vascular-endothelial cadherin (VE-caherin) immunostaining at cell junctions. However, hydrostatic pressure (15 cmH2O) blocked this effect of EGTA. Similarly, cytochalasin D-induced actin depolymerization and intercellular gap formation and cell detachment in response to the Ca2+/calmodulin antagonist trifluperazine (TFP) as well as thrombin-induced cell dissociation were also reduced by hydrostatic pressure. Moreover, hydrostatic pressure significantly reduced the loss of VE-cadherin-mediated adhesion in response to EGTA, cytochalasin D, and TFP in MyEnd cells as determined by laser tweezer trapping using VE-cadherin-coated microbeads. In caveolin-1-deficient MyEnd cells, which lack caveolae, hydrostatic pressure did not protect monolayer integrity compromised by EGTA, indicating that caveolae-dependent mechanisms are involved in hydrostatic pressure sensing and signaling.

vascular-endothelial cadherin; actin

IMPAIRED BARRIER FUNCTIONS are caused by disturbed integrity of the endothelial monolayer and are involved in the pathogenesis of several disorders like inflammatory edema, allergic shock, or atherosclerosis (21, 38, 44). Under physiological conditions in vivo, endothelial cells are exposed to different mechanical forces such as hydrostatic pressure, shear stress, and cyclic stretch (22, 27). However, understanding on the role of hydrostatic pressure in the regulation of endothelial cell biology is incomplete at present. It has been reported that hydrostatic pressure can affect endothelial cell morphology, at least when applied for long incubation periods or at high arterial pressure levels (1, 24, 26, 27, 36). Moreover, we demonstrated previously that exposure of endothelial monolayers [pulmonary artery endothelial cells (PAECs)] to a hydrostatic pressure of 10 cmH2O for 2 h resulted in a significant reduction of monolayer permeability to albumin (30, 37). All these data suggest that hydrostatic pressure profoundly affects endothelial cell biology. Nevertheless, most in vitro studies on endothelial barrier functions in the literature have been carried out in the absence of significant hydrostatic pressure.

Therefore, in the present study, we tested the hypothesis that physiological hydrostatic pressure is effective to protect endothelial monolayer integrity in vitro. We investigated the effect of low hydrostatic pressure of up to 15 cmH2O (pressure typically occurring in capillaries) on the efficacy of different pharmacological mediators that are known to alter endothelial monolayer integrity in the absence (<1 cmH2O) of hydrostatic pressure. We used an immortalized microvascular endothelial cell line obtained from the mouse myocardium [myocardial endothelial (MyEnd) cells] because of the prominent role of endothelial barrier regulation in the microvascular endothelium (21, 22). Essentially, experiments were repeated in primary pulmonary artery endothelial cells (PAECs), which have been used previously to study the effect of hydrostatic pressure or albumin permeability (30, 37). We found that Ca2+ depletion by EGTA, actin depolymerization by cytochalasin D, and inhibition of Ca2+/calmodulin by trifluperazine (TFP) severely disturbed monolayer integrity and reduced vascular endothelial cadherin (VE-cadherin)-mediated adhesion in the absence of hydrostatic pressure. These effects were largely abolished by the simultaneous application of physiological hydrostatic pressure of up to 15 cmH2O. Similarly, thrombin-induced gap formation in PAECs was reduced by hydrostatic pressure. These data indicate that physiological mechanical conditions such as hydrostatic pressure profoundly modulate and protect endothelial cell functions in vivo. Moreover, the absence of these forces in most of the in vitro studies might contribute to inconsistencies compared with investigations in vivo.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. Both immortalized wild-type and caveolin-1-deficient mouse MyEnd cells were grown in DMEM (Life Technologies, Karlsruhe, Germany) supplemented with 50 U/ml penicillin G, 50 µg streptomycin, and 10% FCS (Biochrom, Berlin, Germany) in a humidified atmosphere (95% air-5% CO2) at 37°C. Immortalization and characterization of MyEnd cells were carried out as previously described (13, 42), and cells were found to be immunopositive for several endothelial markers. Expression of the junctional proteins VE-cadherin, {alpha}-catenin, β-catenin, {gamma}-catenin, zonula occludens-1, and claudin-5 as well as von Willebrand factor was verified by immunostaining and immunoblot analysis. Cultures were used for experiments when grown to confluent monolayers (day 3 up to day 7). Primary macrovascular porcine PAECs were isolated by collagenase digestion from porcine pulmonary trunks of adult Large White pigs as previously described (30) and then cultured in medium 199 (M199; Sigma-Aldrich, Taufkirchen, Germany) supplemented with 50 U/ml penicillin G, 50 µg streptomycin, and 10% FCS in a humidified atmosphere (95% air-5% CO2) at 37°C. For all experiments, PAECs were used from passages 1 to 2 when grown to confluent monolayers.

Test reagents. EGTA, cytochalasin D, and TFP were used for 45 min in the absence or presence of hydrostatic pressure as indicated. Thrombin was used for 15 min in the absence of pressure or following 45 min of preincubation in the presence of hydrostatic pressure. EGTA (3 mM) was bought from Applichem (Darmstadt, Germany). Cytochalasin D (10 µM), TFP (100 µM), and thrombin (10 U/ml) were purchased from Sigma-Aldrich (Steinheim, Germany).

Application of hydrostatic pressure. For each of the separate experiments, different passages of MyEnd cells and different preparations of PAECs were used. If not indicated otherwise, hydrostatic pressure was applied as follows (Fig. 1, left): cells were grown on coverslips coated with gelatine cross-linked with glutaraldehyde (28). Coverslips were placed in a six-well dish where the chambers were sealed by a silicone stopper in the center of which a hole was drilled for the insertion of a standard 10-ml pipette (Greiner bio-one, Frickenhausen, Germany). The chamber and pipette were then filled with cell culture medium in the absence or presence of test reagents. For thrombin experiments, hydrostatic pressure was applied for 45 min and thrombin was added to the culture medium in the absence of pressure afterward. Previous experiments have shown that pressure application before the addition of EGTA, cytochalasin, and TFP was similarly effective to simultaneous application (not shown). For the application of a continuous pressure gradient, cells were grown on gelatine-coated glass slides. Afterward, two slides were fixed to each other by metal clips and placed in an upright position in a Perspex column filled with culture medium (Fig. 1, right). To test the effect of hydrostatic pressure on VE-cadherin-mediated adhesion, the laser tweezer setup (see below) was modified in a way that the coverslips were mounted in a metal chamber, which was filled with culture medium and afterward sealed by a Perspex/Lucite lid. A silicone tube was inserted by a side port and filled with medium to apply a hydrostatic pressure of up to 15 cmH2O. We chose low hydrostatic pressures because it is well established that the pressure in the pulmonary microcirculation is ~15 cmH2O (8, 23). Because higher pressures of ~20–50 cmH2O have been shown for tracheal, skeletal, and mesenteric capillaries (3, 11), and the pressure in coronary microvessels changes during the cardiac cycle, we tested the effect of higher levels of hydrostatic pressures in preliminary experiments. Because we found that a hydrostatic pressure of 50 cmH2O did not yield different results, the hydrostatic pressure in this study was increased only up to 15 cmH2O in both cell lines.


Figure 1
View larger version (31K):
[in this window]
[in a new window]

  		Fig. 1. Setup for application of hydrostatic pressure. For experiments using hydrostatic pressure, myocardial endothelial (MyEnd) cells and pulmonary artery endothelial cells (PAECs) were grown on coverslips, which were placed in a 6-well dish where chambers were sealed by a silicone stopper perforated centrally by a standard 10-ml pipette (left). For the application of a continuous pressure gradient, cells were grown on glass slides, which were afterward placed in an upright position fixed to each other by metal clips in a Perspex column containing culture medium (right).

 
Cytochemistry. Cells grown on gelatine-coated coverslips were incubated with various substances. Afterward, the culture medium was removed, and monolayers were fixed for 10 min at room temperature (RT) with 2% formaldehyde (freshly prepared from paraformaldehyde) in PBS. Cell membranes were then permeabilized with 0.1% Triton X-100 in PBS for 5 min. After being rinsed with PBS at RT, cells were preincubated for 30 min with 10% normal goat serum (NGS) or normal donkey serum (both from Dianova, Hamburg, Germany) containing 1% BSA at RT and incubated for 16 h at 4°C with primary antibodies. For staining of VE-cadherin in MyEnd cells, cells were incubated with rat monoclonal antibody 11D4.1 (undiluted hybridoma supernatant) directed to the ectodomain of mouse VE-cadherin (14). PAECs were treated with goat polyclonal antibody against VE-cadherin (Santa Cruz Biotechnology, Heidelberg, Germany). For the staining of claudin-5, a rabbit polyclonal antibody (Zymed Laboratories/Invitrogen, Karlsruhe, Germany, diluted 1:100 in PBS) was used.

After several rinses with PBS (3 x 5 min), monolayers were incubated for 60 min at RT with Cy3-labeled goat anti-rat IgG, Cy3-labeled goat anti-rabbit IgG, or Cy3-labeled donkey anti-goat IgG, respectively (all from Dianova, diluted 1:600 in PBS). For the visualization of F-actin, monolayers were incubated with Alexa-phalloidin (Molecular Probes, Eugene, OR, diluted 1:60 in PBS for 1 h at RT). Cells incubated with antibodies or with Alexa-phalloidin were rinsed with PBS (3 x 5 min). Coverslips were mounted on glass slides with 60% glycerol in PBS, which contained 1.5% n-propyl gallate (Serva, Heidelberg, Germany) as an antifading compound.

Recombinant VE-cadherin-Fc. As previously described, 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 (5, 14). The protein was expressed by stably transfected Chinese hamster ovary cells and purified from culture supernatants by affinity chromatography using protein A agarose (Oncogene, Cambridge, MA).

Coating of polystyrene beads. After being vortexed, 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 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 HBSS (GIBCO, Karlsruhe, Germany) containing 10 µg of either VE-cadherin-Fc or of the Fc part of human IgG (for control experiments) and allowed to react for 16 h at 4°C under permanent slow overhead rotation to avoid aggregation. After being washed for 3 x 5 min in 100 µl buffer A and 3 x 5 min in buffer B (100 mM sodium borate; pH 9.0), beads were incubated for 45 min at RT in 100 µl buffer B containing 0.54 mg dimethyl pimelimidate dihydrochloride (DMP; Pierce, Rockford, IL) to covalently cross-link protein A and bound Fc parts. After being washed 2 x 5 min in buffer C (100 µl of 0.2 M ethanolamine; pH 8.0), beads were incubated in buffer C for 2 h at RT. Finally, beads were washed 3 x 5 min in HBSS and stored in HBSS at 4°C for up to 8 days under permanent slow overhead rotation to avoid aggregation of beads. The concentration of beads in these stocks was ~1.6 x 108 beads/ml.

Laser tweezer. As previously described (6), 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-numberical 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). For all experiments, the laser intensity was 42 mW in the focal plane. Coated beads (10 µl of stock solution) were suspended in 500 µl culture medium and allowed to interact with microvascular endothelial cell monolayers for 30 min at 37°C before the initiation of experiments. Beads were considered tightly bound when resisting laser displacement. For every condition, 100 beads were counted. The percentage of beads resisting laser displacement under various experimental conditions was normalized to control values.

Statistics. Values are expressed as means ± SE. Possible differences in bead binding were assessed using one-way ANOVA and Dunnett's multiple-comparison test. Statistical significance was assumed for P < 0.05.


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Hydrostatic pressure abrogates the effect of extracellular Ca2+ depletion on VE-cadherin immunolocalization. In microvascular MyEnd control monolayers, VE-cadherin was continuously distributed along cell junctions (Fig. 2A). Depletion of extracellular Ca2+ by EGTA (3 mM, 45 min) resulted in a diffuse immunostaining pattern (Fig. 2B). In parallel experiments, MyEnd cells were exposed to hydrostatic pressure, as shown in Fig. 1, left. In 41 of 50 separate experiments, the effect of EGTA on VE-cadherin immunolocalization was largely abrogated by the simultaneous application of hydrostatic pressure of 15 cmH2O (Fig. 2C). Similar results were obtained in seven of eight experiments when cells were preincubated with hydrostatic pressure for 45 min before the addition of EGTA (not shown). In contrast to VE-cadherin, distribution of the endothelial tight junction adhesion molecule claudin-5 was not affected by either EGTA or hydrostatic pressure (Fig. 2, D–F; n = 4).


Figure 2
View larger version (121K):
[in this window]
[in a new window]

  		Fig. 2. Effect of extracellular Ca2+ depletion on vascular-endothelial cadherin (VE-cadherin) localization in the presence and absence of hydrostatic pressure. MyEnd cells were immunostained for VE-cadherin (A–C) or claudin-5 (D–F). In controls, VE-cadherin was continuously localized along cell junctions (A). In contrast, extracellular Ca2+ depletion by EGTA resulted in a loss from junctions and caused a completely diffuse staining pattern (B). Hydrostatic pressure (15 cmH2O) almost completely blocked this effect in 41 of 50 experiments. In contrast, EGTA and hydrostatic pressure did not affect the localization of claudin-5 (n = 4). Scale bar = 12 µm for A–F.

 
Hydrostatic pressure inhibits actin depolymerization and intercellular gap formation. MyEnd control cells formed regular monolayers and displayed numerous stress fibers running parallel along the cellular long axis (Fig. 3, A–C). Hydrostatic pressure (15 cmH20, 45 min) alone did not affect the organization of the actin cytoskeleton (not shown). Cytochalasin D (10 µM, 45 min) induced cell dissociation, leading to intercellular gap formation (arrows in Fig. 3, D and E) and complete loss of stress fibers and fragmentation of the peripheral actin band (Fig. 3E). Intercellular gaps were located at sites where the peripheral actin was missing (arrows in Fig. 3E; n = 13). When hydrostatic pressure (15 cmH2O, 45 min) was applied simultaneously with treatment with cytochalasin D, intercellular gap formation and peripheral actin depolymerization were largely reduced in 8 of 13 separate experiments, whereas the protective effect on stress fibers was less pronounced. However, it has to be emphasized that some gaps were still detectable under these conditions (arrows in Fig. 3, G and I), indicating that the effect of thrombin was not completely abolished by hydrostatic pressure. Exposure of MyEnd cells for 45 min prior to treatment with cytochalasin D was equally effective (n = 3; not shown).


Figure 3
View larger version (159K):
[in this window]
[in a new window]

  		Fig. 3. Effect of actin depolymerization on endothelial monolayer integrity in the presence and absence of hydrostatic pressure. MyEnd cells were immunostained for VE-cadherin (A, D, and G) or stained for F-actin using Alexa-phalloidin (B, E, and H). C, F, and I: merged images. In controls, MyEnd cells formed intact monoalyers (A–C) and displayed numerous stress fibers throughout the cytoplasm (B). Cytochalasin D (CytoD) caused intercellular gap formation (arrows in D), fragmentation of the peripheral actin band (arrows in E), and loss of stress fibers (D–F). Hydrostatic pressure (15 cmH2O) reduced gap formation and interruption of VE-cadherin staining and actin fragmentation in 8 of 13 experiments (G–I). Note that some gaps were still detectable under these conditions (arrows), indicating that the effect of thrombin (Thr) was not completely abolished by hydrostatic pressure (G–I). Scale bar = 20 µm for A–I.

 
Hydrostatic pressure blocks cell detachment in response to Ca2+/calmodulin inhibition. Compared with controls (Fig. 4, A and B), TFP (100 µM, 45 min) caused disruption of VE-cadherin immunostaining and cell detachment, leading to the formation of large intercelluar gaps (arrows in Fig. 4, C and D) and a reduction of stress fibers (Fig. 4D). In some areas of the coverslip, many cells became detached following treatment with TFP (not shown). TFP-induced cell dissociation and detachment from the substratum were almost completely abolished by the simultaneous application of hydrostatic pressure (Fig. 4, E and F) in 10 of 11 separate experiments.


Figure 4
View larger version (128K):
[in this window]
[in a new window]

  		Fig. 4. Effect of Ca2+/calmodulin inhibition on endothelial monolayer integrity in the presence and absence of hydrostatic pressure. MyEnd cells were immunostained for VE-cadherin (A, C, and E) or stained for F-actin using Alexa-phalloidin (B, D, and F). In controls, MyEnd cells formed intact monoalyers and displayed numerous stress fibers throughout the cytoplasm (A and B). Trifluperazine (TFP) led to cell detachment, intercellular gap formation, and reduction of stress fibers (C and D). Hydrostatic pressure (15 cmH2O) almost completely blocked gap formation in 10 of 11 experiments. Scale bar = 12 µm for A–F.

 
Protective effects of hydrostatic pressure on monolayer integrity were also present in primary artery endothelial cells. Our results demonstrate that physiological hydrostatic pressure protects microvascular endothelial monolayer integrity against various different stimuli in vitro. To rule out the possibility that these effects were due to immortalization artifacts in MyEnd cells, experiments were essentially repeated under the same conditions using PAECs. As shown in Fig. 5, the effects of hydrostatic pressure in PAECs were similar to the results obtained from MyEnd cells. Hydrostatic pressure (15 cmH2O, 45 min) almost completely blocked the effect of EGTA (3 mM) on VE-cadherin immunolocalization (Fig. 5, A–C) in 12 of 24 separate experiments. Cell dissociation in response to cytochalasin D (10 µM, 45 min) was blocked in 9 of 12 experiments (Fig. 5, D–F), and TFP-induced (100 µM, 45 min) gap formation was found to be abolished in all 5 experiments performed in the presence of hydrostatic pressure (Fig. 5, G–I). Exposure of PAECs to a lower pressure of 5 cmH2O was nearly similarly effective to block the effects of cytochalasin D (Fig. 5E) and EGTA (not shown) but not TFP (Fig. 5H). Because PAECs in contrast to MyEnd cells respond to thrombin, which is widely used as a physiological mediator to alter endothelial barrier functions (21), we tested whether hydrostatic pressure would affect the response of PAECs to thrombin. In the absence of hydrostatic pressure, thrombin (10 U/ml, 15 min) induced profound cell dissociation, leading to the formation of multiple intercellular gaps (arrows in Fig. 5J). Pretreatment of monolayers with a hydrostatic pressure of 5 or 15 cmH2O for 45 min significantly reduced intercellular gap formation (Fig. 5, K and L) in response to thrombin. However, in some areas, gaps were still observed (arrows in Fig. 5, K and L), indicating that the effect of thrombin was not completely abolished.


Figure 5
View larger version (141K):
[in this window]
[in a new window]

  		Fig. 5. Effect of hydrostatic pressure on monolayer integrity in the macrovascular endothelium. PAECs were immunostained for VE-cadherin. In controls, PAECs formed intact monoalyers with continuous localization of VE-cadherin along cell junctions (A). Extracellular Ca2+ depletion by EGTA resulted in a completely diffuse staining pattern (B), which was almost completely blocked by hydrostatic pressure (15 cmH2O) in 12 of 24 experiments (C). CytoD-induced intercellular gap formation (D) was inhibited by hydrostatic pressures of 5 and 15 cmH2O in 9 of 12 experiments (E and F). TFP caused intercellular gap formation (G), which was also inhibited in part by a hydrostatic pressure of 5 cmH2O (H) and almost completely by 15 cmH2O (I). Thr-induced gap formation (J) was reduced but not completely abolished by prereatment with hydrostatic pressures of 5 and 15 cmH2O in 8 of 13 experiments (arrows in K and L). Scale bar = 20 µm for A–L.

 
The protective effect of hydrostatic pressure increases up to 15 cmH2O and is dependent on caveolin-1. To study exactly how much pressure was required to block the effect of EGTA on VE-cadherin immunolocalization, a continuous pressure gradient was applied, as described in METHODS. Because glass slides coated with endothelial cells were placed upright in the center of a culture medium-filled column, cells were equally exposed to hydrostatic pressure from all sides. This setup allowed us to completely rule out a possible contribution of strain that might be exerted on the cell layer by pressure-induced minuscule deformations of the culture well and its adhering coverslips. In MyEnd cells and PAECs, gradual protective effects of hydrostatic pressure on VE-cadherin immunolocalization were detected already at ~1 cmH2O, and the protective effect increased gradually up to 15 cmH2O, indicating that no threshold of hydrostatic pressure exists that is required to trigger protective mechanisms (shown for MyEnd cells in Fig. 6). However, 5 cmH2O was almost as effective as 15 cmH2O to block the effect of EGTA in MyEnd cells. These effects were present in seven of nine experiments.


Figure 6
View larger version (183K):
[in this window]
[in a new window]

  		Fig. 6. Effects of continuous hydrostatic pressures ranging from 0 to 15 cmH2O. MyEnd cells were immunostained for VE-cadherin following exposure to EGTA in the absence (A) or presence of hydrostatic pressure (B–I). Extracellular Ca2+ depletion by EGTA resulted in a completely diffuse staining pattern (A). Hydrostatic pressure blocked this effect already at 1 cmH2O (B) and was more effective with increased pressure up to 15 cmH2O (C–I). This effect was observed in 7 of 9 experiments. Scale bar = 12 µm for A–I.

 
Finally, because caveolae have been reported to be involved in sensing of cyclic stretch in vascular smooth muscle cells (31), we used the caveolin-1-deficient (caveolin-1–/–) MyEnd cell line we have recently established (42). In caveolin-1–/– cells, EGTA (3 mM, 45 min) led to a completely diffuse VE-cadherin staining pattern compared with controls, which was similar to the effects seen in normal caveolin-1-containing MyEnd cells (Fig. 7, A and B). However, in contrast to normal MyEnd cells, the protective effect of hydrostatic pressure was completely abrogated (Fig. 7C; n = 6). These results are a first indication that caveolin-1 and/or caveolae might be involved in the mechanisms underlying pressure-induced mechanosensation.


Figure 7
View larger version (61K):
[in this window]
[in a new window]

  		Fig. 7. Effect of hydrostatic pressure required the expression of caveolin-1. MyEnd cells were immunostained for VE-cadherin. In controls, VE-cadherin was continuously localized along cell junctions (A). Extracellular Ca2+ depletion by EGTA resulted in a completely diffuse staining pattern (B). In contrast to wild-type cells, hydrostatic pressure did not block the effect of EGTA in caveolin-1-deficient (caveolin-1–/–) cells (C; n = 6). Scale bar = 20 µm for A–C.

 
Hydrostatic pressure blocks the loss of VE-cadherin-mediated adhesion caused by both extra- and intracellular stimuli. The experiments described above show that hydrostatic pressure blocked cell dissociation and loss of continuous VE-cadherin localization along cell junctions. To investigate whether hydrostatic pressure directly affects VE-cadherin-mediated homophilic binding, which is a known prerequisite for tight intercellular adhesion in endothelial cells, we tested the binding of VE-cadherin-coated microbeads by laser tweezers. Therefore, microbeads were coated with recombinant VE-cadherin-Fc and allowed to settle on the cell surface to establish adherens-like cell-to-bead contacts (6, 40). Afterward, beads were subjected to laser displacement. Beads that could not be displaced by the laser beam were considered as tightly bound. Experiments were performed in the presence or absence of 15 cmH2O hydrostatic pressure (Fig. 8). Experiments were carried out in MyEnd cells but not in PAECs because recombinant mouse VE-cadherin does not bind to its porcine cell homolog. In accordance with our previous studies, in control MyEnd monolayers, ~70–80% of coated beads tightly adhered to the cell surface and could not be displaced by the laser beam (13, 42). These values were normalized to 100%. Extracellular Ca2+ depletion by EGTA (3 mM, 45 min) reduced the number of bound beads to 54 ± 2% of control levels. This effect was significantly diminished by a hydrostatic pressure of 15 cmH2O (91 ± 5% beads remained bound, n = 12) in 12 of 15 experiments. Actin depolymerization (10 µM. 45 min) led to a significant loss of bead binding to 62 ± 4% of controls, as previously described (6, 40). This effect was significantly reduced in the presence of hydrostatic pressure (5 of 7 experiments), where 84 ± 4% of beads resisted laser displacement. TFP-induced reduction of bead binding (76 ± 7%) was completely blocked by hydrostatic pressure (97 ± 2%) in 8 of 11 experiments. Taken together, these results demonstrate that hydrostatic pressure protects VE-cadherin-mediated adhesion against both extra- and intracellular destabilizing stimuli.


Figure 8
View larger version (11K):
[in this window]
[in a new window]

  		Fig. 8. Effect of hydrostatic pressure on VE-cadherin-mediated adhesion as revealed by laser tweezers. In MyEnd controls, ~70–80% of microbeads coated with recombinant VE-cadherin tightly adhered to the cell surface and could not be displaced by the laser beam. EGTA, CytoD, and TFP resulted in a significant loss of bead adhesion. These effects were significantly reduced in the presence of hydrostatic pressure (15 cmH2O) in 12 of 15 experiments (EGTA) and in 5 of 7 experiments (CytoD), respectively. Hydrostatic pressure completely blocked the TFP-induced loss of bead binding in 8 of 11 experiments. Values are means ± SE. *Significant changes compared with controls (P < 0.05); #significant change compared with experiments using the same pharmacological mediator in the absence of hydrostatic pressure (P < 0.05).

 

   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The aim of the present study was to investigate the role of hydrostatic pressure in the maintenance of monolayer integrity in microvascular (MyEnd cells) and macrovascular endothelial cells (PAECs). We used EGTA-mediated Ca2+ depletion, cytochalasin D-induced actin depolymerization, and TFP-mediated inhibition of Ca2+/calmodulin function because these conditions have been shown in our previous studies to disrupt monolayer integrity in the absence of hydrostatic pressure in vitro (6, 35, 41, 43). We found that a hydrostatic pressure of 15 cmH2O was sufficient to largely prevent the junctional loss of VE-cadherin in response to EGTA in both microvascular (MyEnd cells) and macrovascular (PAECs) endothelial cells. Similarly, intercellular gap formation in response to actin depolymerization as well as cell detachment induced by TFP were reduced in the presence of hydrostatic pressure. Moreover, our laser tweezer experiments showed that hydrostatic pressure completely blocked the loss of VE-cadherin-mediated adhesion in response to EGTA as well as to TFP and partially rescued the cytochalasin D-induced decrease of VE-cadherin binding.

One possibility to explain these phenomena could be that hydrostatic pressure prevents the uptake of TFP and cytochalasin D. However, this is unlikely because has been shown previously that TFP prevents the A23187 [GenBank] -induced rise in permeability in PAEC monolayers exposed (conditioned) to a hydrostatic pressure of 10 cmH2O (30). Likewise, cytochalasin D caused a delayed increase of permeability under same hydrostatic pressure conditions in PAECs, clearly showing that both TFP and cytochalasin D were taken up and exerted cellular responses. It has to be noted that despite the fact that hydrostatic pressure in our present study was effective to reduce cytochalasin D-induced gap formation, gaps were still detectable under these conditions (Fig. 6), indicating that cytochalasin D in the presence of hydrostatic pressure is still effective to compromise endothelial barrier properties. This is in line with our previous study (41) showing that cytochalasin D increased the permeability of single-perfused mesenteric microvessels in vivo. Since cytochalasin D directly compromises actin filament polymerization and TFP modulates the organization of the actin filament system by interference with myosin light chain phosphorylation (16, 18, 25, 33, 34), we assume that hydrostatic pressure triggers signaling pathways that stabilize the actin filament system. Possible candidates are members of the Rho protein family, which are known to be key regulators of both endothelial adhesion and cytoskeleton. Further studies are in progress to address this aspect of hydrostatic pressure signaling.

Taken together, our data demonstrate that hydrostatic pressure is capable of maintaining endothelial monolayer integrity in cells exposed to several pharmacological agents known to alter monolayer integrity. These findings suggest that the absence of hydrostatic pressure in most studies may account for some of the apparent inconsistencies with the in vivo situation. For instance, for mesenteric and muscular microvessels in vivo, it has been demonstrated that endothelial barrier function is insensitive to extracellular Ca2+ depletion (10). Although differences in the method to deplete Ca2+ do not allow direct comparison, these results cannot be easily reconciled with the observation from our group and others that, in cultured endothelium in vitro, permeability was found to be dependent on both hydrostatic pressure and extracellular Ca2+ (2, 12, 20, 29, 30, 32, 37). The requirement of extracellular Ca2+ for the maintenance of endothelial monolayer integrity can at least in part be explained by the fact that millimolar extracellular Ca2+ is needed for both the stabilization of the VE-cadherin ectodomain and homophilic binding across the intercellular space (5, 39). VE-cadherin mediates homophilic intercellular binding in endothelial adherens junctions and is thought to be required for the maintenance of monolayer integrity and thus for endothelial barrier function in vivo and in vitro (9, 14, 17, 19). Accordingly, in MyEnd cells, we have shown that, in the absence of hydrostatic pressure, extracellular Ca2+ depletion by EGTA resulted in a profound loss of VE-cadherin immunostaining along cell junctions paralleled by impaired VE-cadherin-mediated adhesion probed by laser tweezers (4, 6).

Our present study now suprisingly suggests that in the absence of extracellular Ca2+, hydrostatic pressure prevents junctional loss of VE-cadherin and, in addition, promotes adhesion of VE-cadherin-coated microbeads. We have no explanation for this phenomenon since drop of extracellular Ca2+ below 0.5 mM inhibits transinteraction of VE-cadherin measured by atomic force microscopy and laser tweezers (5). One possibility would be that hydrostatic pressure signaling stabilizes cytoskeletal tethering of VE-cadherin and prevents its disappearance from junctions (including bead-to-cell junctions) by lateral diffusion. Since homophilic binding between opposing VE-cadherin molecules is not possible at low Ca2+, one has to consider the possibility that other Ca2+-independent adhesion molecules associated with cadherin junctions such as PECAM-1, nectin, or juction-associated adhesion molecules (JAMs) (7, 21) may form heterophilic bonds to VE-cadherin even or preferentially at low extracelluar Ca2+ to compensate for the loss of homophilic VE-cadherin adhesion. Further studies need to be done to test whether PECAM, nectin, and JAMs may bind to VE-cadherin at low Ca2+.

However, it has to be emphasized that the experimental design of our study on the cultured endothelium was not identical to the conditions used for in vivo investigations from the literature in which either Ca2+-free solutions or very low EGTA concentrations (µM) were used (10, 15) and, therefore, these studies cannot be directly compared. Nevertheless, our results indicate that the maintenance of endothelial barrier properties in vitro is dependent on the presence of physiological mechanical conditions such as hydrostatic pressure.

In contrast to previous studies using cultured endothelium in which either high pressures of 80–100 mmHg were used or endothelial cells were exposed to hydrostatic pressure for several days (1, 26, 27, 36), we did not observe any significant changes in cell morphology like elongation or reorganization of filamentous actin under the short-term conditions used in our study. It has been reported that exposure of PAECs to 8–12 mmHg for 7 days resulted in cell elongation and multilayer formation (1, 26). When applied for 9 days, high pressures of 80–100 mmHg were reported to cause cell elongation but not multilayer formation in bovine aortic cells (36). In a recent study, application of high hydrostatic pressure of 100 mmHg for 24 h to bovine aortic cells induced stress fiber formation and loss of VE-cadherin immunstaining along cell junctions (24, 27). Especially, in contrast to Ohashi et al. (18), we did not observe fragmentation of VE-cadherin staining when physiological pressure values were used (24, 27). This observation has lead to the speculation that adherens junctions might be disassembled in response to high pressure. Nevertheless, our data show that physiological pressure in the microvascular endothelium protects immunolocalization of VE-cadherin and also stabilizes VE-cadherin-mediated adhesion against several agents that reduced VE-cadherin binding in the absence of pressure.

Because the mechanisms underlying mechanosensation of hydrostatic pressure in endothelial cells are completely unclear at present, we first performed experiments to characterize the mechanisms involved in the protective effects of hydrostatic pressure. Based on the observation that caveolin-1 is essential for the senation of cyclic stretch in vascular smooth muscle cells (31), we used the caveolin-1–/– MyEnd cell line established recently by our laboratory (42). We found that hydrostatic pressure was ineffective in the caveolin-1-deficient endothelium, indicating that caveolin-1 and caveolae indeed are part of the structures participating in the sensation of hydrostatic pressure. However, the mechanisms by which caveolin-1 or /and caveolae are important for the protective effects of hydrostatic pressure are unknown. Several possibilities could be envisaged: although inhibition of endothelial nitric oxide (NO) synthase by N-nitro-L-arginine methyl ester had no effect on the efficacy of hydrostatic pressure to maintain (unpublished observations), it is tempting to speculate that NO might be involved in the mechanisms underlying the sensation of hydrostatic pressure. Based on the experiments in which endothelial cells were exposed to a continuous pressure gradient, we believe that cell or substrate deformation is not required for pressure sensing. Therefore, we raised the hypothesis that small intracellular gas bubbles (e.g., NO bubbles) that might be located inside caveolae might provide a sensor for hydrostatic pressure. Unlike fluid, gas bubbles would undergo pressure-dependent volume changes (Boyle-Mariotte's law) and could function as a sensor by causing mechanical strain to the caveolar membrane, if located there. Since our experiments do not entirely exclude this possibility, further, more-refined studies are planned to address this aspect of pressure sensing. Nevertheless, it cannot be completely ruled out that caveolae may provide a store of plasma membrane that may allow pressure-induced cell deformation, which, in turn, could contribute to the initiation of cell signaling mechanisms.

Taken together, our study supports the hypothesis that physiological hydrostatic pressure protects endothelial monolayer integrity and thus profoundly regulates endothelial cell biology. This has been neglected by most cell culture studies in the literature and may at least in part explain some inconsistencies with previous in vivo studies.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by Deutsche Forschungsgemeinschaft Grants SFB 487, TP B5 and SFB 688, TP A4.


   ACKNOWLEDGMENTS
 
We are grateful to Stefanie Imhof, Nadja Niedermeier, and Jens Günther for skillful technical assistance. Prof. Axel Wilke (Department of Orthopaedics, Elisabeth Hospital, Olsberg, Germany) and Dr. Jochen Steinbrenner (Emergency Unit, Spital Grabs, Grabs, Switzerland) kindly provided unpublished supplementary material of thesis and diploma work conducted in our laboratory several years ago.


   FOOTNOTES
 

Address for reprint requests and other correspondence: D. Drenckhahn, Institute of Anatomy and Cell Biology, Julius-Maximilians-Univ., Koellikerstrasse 6, Würzburg D-97070, Germany (e-mail: anat015{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.

* K. Müller-Marschhausen and J. Waschke contributed equally to this work. Back


   REFERENCES
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Acevedo AD, Bowser SS, Gerritsen ME, Bizios R. Morphological and proliferative responses of endothelial cells to hydrostatic pressure: role of fibroblast growth factor. J Cell Physiol 157: 603–614, 1993.[CrossRef][Web of Science][Medline]

2. Baetscher M, Brune K. An in vitro system for measuring endothelial permeability under hydrostatic pressure. Exp Cell Res 148: 541–547, 1983.[CrossRef][Web of Science][Medline]

3. Ballard ST, Nations RH, Taylor AE. Microvascular pressure profile of serosal vessels of rat trachea. Am J Physiol Heart Circ Physiol 262: H1303–H1304, 1992.[Abstract/Free Full Text]

4. Baumgartner W, Golenhofen N, Weth A, Hiiragi T, Saint R, Griffin M, Drenckhahn D. Role of transglutaminase 1 in stabilisation of intercellular junctions of the vascular endothelium. Histochem Cell Biol 122: 17–25, 2004.[Web of Science][Medline]

5. Baumgartner W, Hinterdorfer P, Ness W, Raab A, Vestweber D, Schindler H, Drenckhahn D. Cadherin interaction probed by atomic force microscopy. Proc Natl Acad Sci USA 97: 4005–4010, 2000.[Abstract/Free Full Text]

6. Baumgartner W, Schutz GJ, Wiegand J, Golenhofen N, Drenckhahn D. Cadherin function probed by laser tweezer and single molecule fluorescence in vascular endothelial cells. J Cell Sci 116: 1001–1011, 2003.[Abstract/Free Full Text]

7. Bazzoni G, Dejana E. Endothelial cell-to-cell junctions: molecular organization and role in vascular homeostasis. Physiol Rev 84: 869–901, 2004.[Abstract/Free Full Text]

8. Cope DK, Parker JC, Taylor MD, Houston M, Taylor AE. Pulmonary capillary pressures during hypoxia and hypoxemia: experimental and clinical studies. Crit Care Med 17: 853–857, 1989.[Web of Science][Medline]

9. Corada M, Mariotti M, Thurston G, Smith K, Kunkel R, Brockhaus M, Lampugnani MG, Martin-Padura I, Stoppacciaro A, Ruco L, McDonald DM, Ward PA, Dejana E. Vascular endothelial-cadherin is an important determinant of microvascular integrity in vivo. Proc Natl Acad Sci USA 96: 9815–9820, 1999.[Abstract/Free Full Text]

10. Curry FE, Mason JC, Michel CC. Proceedings: the effect of bovine gamma-globulin and Ca2+ on the filtration coefficient of individual frog mesenteric capillaries. J Physiol 234: 43P–44P, 1973.[Medline]

11. Fronek K, Zweifach BW. Microvascular pressure distribution in skeletal muscle and the effect of vasodilation. Am J Physiol 228: 791–796, 1975.[Abstract/Free Full Text]

12. Gao X, Kouklis P, Xu N, Minshall RD, Sandoval R, Vogel SM, Malik AB. Reversibility of increased microvessel permeability in response to VE-cadherin disassembly. Am J Physiol Lung Cell Mol Physiol 279: L1218–L1225, 2000.[Abstract/Free Full Text]

13. Golenhofen N, Ness W, Wawrousek EF, Drenckhahn D. Expression and induction of the stress protein alpha-B-crystallin in vascular endothelial cells. Histochem Cell Biol 117: 203–209, 2002.[CrossRef][Web of Science][Medline]

14. Gotsch U, Borges E, Bosse R, Boggemeyer E, Simon M, Mossmann H, Vestweber D. VE-cadherin antibody accelerates neutrophil recruitment in vivo. J Cell Sci 110: 583–588, 1997.[Abstract]

15. He P, Curry FE. Albumin modulation of capillary permeability: role of endothelial cell [Ca2+]i. Am J Physiol Heart Circ Physiol 265: H74–H82, 1993.[Abstract/Free Full Text]

16. Hidaka H, Asano M, Tanaka T. Activity-structure relationship of calmodulin antagonists, naphthalenesulfonamide derivatives. Mol Pharmacol 20: 571–578, 1981.[Abstract/Free Full Text]

17. Hordijk PL, Anthony E, Mul FP, Rientsma R, Oomen LC, Roos D. Vascular-endothelial-cadherin modulates endothelial monolayer permeability. J Cell Sci 112: 1915–1923, 1999.[Abstract]

18. Keller TC 3rd, Mooseker MS. Ca++-calmodulin-dependent phosphorylation of myosin, and its role in brush border contraction in vitro. J Cell Biol 95: 943–959, 1982.[Abstract/Free Full Text]

19. Lampugnani MG, Resnati M, Raiteri M, Pigott R, Pisacane A, Houen G, Ruco LP, Dejana E. A novel endothelial-specific membrane protein is a marker of cell-cell contacts. J Cell Biol 118: 1511–1522, 1992.[Abstract/Free Full Text]

20. Langeler EG, van Hinsbergh VW. Characterization of an in vitro model to study the permeability of human arterial endothelial cell monolayers. Thromb Haemost 60: 240–246, 1988.[Web of Science][Medline]

21. Mehta D, Malik AB. Signaling mechanisms regulating endothelial permeability. Physiol Rev 86: 279–367, 2006.[Abstract/Free Full Text]

22. Michel CC, Curry FE. Microvascular permeability. Physiol Rev 79: 703–761, 1999.[Abstract/Free Full Text]

23. Negrini D. Pulmonary microvascular pressure profile during development of hydrostatic edema. Microcirculation 2: 173–180, 1995.[Medline]

24. Ohashi T, Sugaya Y, Sakamoto N, Sato M. Hydrostatic pressure influences morphology and expression of VE-cadherin of vascular endothelial cells. J Biomech 40: 2399–2405, 2007.[CrossRef][Web of Science][Medline]

25. Patel H, Margossian SS, Chantler PD. Locking regulatory myosin in the off-state with trifluoperazine. J Biol Chem 275: 4880–4888, 2000.[Abstract/Free Full Text]

26. Salwen SA, Szarowski DH, Turner JN, Bizios R. Three-dimensional changes of the cytoskeleton of vascular endothelial cells exposed to sustained hydrostatic pressure. Med Biol Eng Comput 36: 520–527, 1998.[CrossRef][Web of Science][Medline]

27. Sato M, Ohashi T. Biorheological views of endothelial cell responses to mechanical stimuli. Biorheology 42: 421–441, 2005.[Web of Science][Medline]

28. Schnittler HJ, Franke RP, Akbay U, Mrowietz C, Drenckhahn D. Improved in vitro rheological system for studying the effect of fluid shear stress on cultured cells. Am J Physiol Cell Physiol 265: C289–C298, 1993.[Abstract/Free Full Text]

29. Schnittler HJ, Puschel B, Drenckhahn D. Role of cadherins and plakoglobin in interendothelial adhesion under resting conditions and shear stress. Am J Physiol Heart Circ Physiol 273: H2396–H2405, 1997.[Abstract/Free Full Text]

30. Schnittler HJ, Wilke A, Gress T, Suttorp N, Drenckhahn D. Role of actin and myosin in the control of paracellular permeability in pig, rat and human vascular endothelium. J Physiol 431: 379–401, 1990.[Abstract/Free Full Text]

31. Sedding DG, Hermsen J, Seay U, Eickelberg O, Kummer W, Schwencke C, Strasser RH, Tillmanns H, Braun-Dullaeus RC. Caveolin-1 facilitates mechanosensitive protein kinase B (Akt) signaling in vitro and in vivo. Circ Res 96: 635–642, 2005.[Abstract/Free Full Text]

32. Shasby DM, Shasby SS. Effects of calcium on transendothelial albumin transfer and electrical resistance. J Appl Physiol 60: 71–79, 1986.[Abstract/Free Full Text]

33. Sheterline P. Trifluoperazine can distinguish between myosin light chain kinase-linked and troponin C-linked control of actomyosin interaction by Ca++. Biochem Biophys Res Commun 93: 194–200, 1980.[CrossRef][Web of Science][Medline]

34. Sobieszek A. Calmodulin antagonist action in smooth-muscle myosin phosphorylation. Different mechanisms for trifluoperazine and calmidazolium inhibition. Biochem J 262: 215–223, 1989.[Web of Science][Medline]

35. Steinbrenner J. Untersuchungen zur Permeabilität von Endothelzellen und der Wirkung von hydrostatischem Druck auf das Zytoskelett (Medical Thesis). Würzburg, Germany: University of Würzburg, 2001.

36. Sumpio BE, Widmann MD, Ricotta J, Awolesi MA, Watase M. Increased ambient pressure stimulates proliferation and morphologic changes in cultured endothelial cells. J Cell Physiol 158: 133–139, 1994.[CrossRef][Web of Science][Medline]

37. Suttorp N, Hessz T, Seeger W, Wilke A, Koob R, Lutz F, Drenckhahn D. Bacterial exotoxins and endothelial permeability for water and albumin in vitro. Am J Physiol Cell Physiol 255: C368–C376, 1988.[Abstract/Free Full Text]

38. van Nieuw Amerongen GP, van Hinsbergh VW. Targets for pharmacological intervention of endothelial hyperpermeability and barrier function. Vasc Pharmacol 39: 257–272, 2002.[CrossRef]

39. Vincent PA, Xiao K, Buckley KM, Kowalczyk AP. VE-cadherin: adhesion at arm's length. Am J Physiol Cell Physiol 286: C987–C997, 2004.[Abstract/Free Full Text]

40. Waschke J, Baumgartner W, Adamson RH, Zeng M, Aktories K, Barth H, Wilde C, Curry FE, Drenckhahn D. Requirement of Rac activity for maintenance of capillary endothelial barrier properties. Am J Physiol Heart Circ Physiol 286: H394–H401, 2004.[Abstract/Free Full Text]

41. Waschke J, Curry FE, Adamson RH, Drenckhahn D. Regulation of actin dynamics is critical for endothelial barrier functions. Am J Physiol Heart Circ Physiol 288: H1296–H1305, 2005.[Abstract/Free Full Text]

42. Waschke J, Golenhofen N, Kurzchalia TV, Drenckhahn D. Protein kinase C-mediated endothelial barrier regulation is caveolin-1-dependent. Histochem Cell Biol 126: 17–26, 2006.[CrossRef][Web of Science][Medline]

43. Wilke A. Morphologische und immunzytochemische Charakterisierung eines in vitro-Modelles zur Untersuchung der Permeabilität von Endothelzellmonolayern (Master Thesis). Marburg, Germany: University of Marburg, 1987.

44. Yuan SY. Protein kinase signaling in the modulation of microvascular permeability. Vasc Pharmacol 39: 213–223, 2002.[CrossRef]




This article has been cited by other articles:


Home page
 Am. J. Physiol. Cell Physiol.Home page
G. P. van Nieuw Amerongen, R. J. P. Musters, E. C. Eringa, P. Sipkema, and V. W. M. van Hinsbergh
Thrombin-induced endothelial barrier disruption in intact microvessels: role of RhoA/Rho kinase-myosin phosphatase axis
Am J Physiol Cell Physiol, May 1, 2008; 294(5): C1234 - C1241.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
294/1/C324    most recent
00319.2007v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via ISI Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Müller-Marschhausen, K.
Right arrow Articles by Drenckhahn, D.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Müller-Marschhausen, K.
Right arrow Articles by Drenckhahn, D.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2008 by the American Physiological Society.