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(Circulation Research. 1996;79:1000-1006.)
© 1996 American Heart Association, Inc.

Articles

Macromolecular Composition of Stress Fiber–Plasma Membrane Attachment Sites in Endothelial Cells In Situ

Yumiko Kano, Kazuo Katoh, Michitaka Masuda, Keigi Fujiwara

the Department of Structural Analysis, National Cardiovascular Center Research Institute, Osaka, Japan.

Correspondence to Yumiko Kano, Department of Structural Analysis, National Cardiovascular Center Research Institute, 5 Fujishiro-dai, Suita, Osaka 565, Japan.

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Stress fibers (SFs) are present along the apical (apical SF) and the basal (basal SF) portions of cultured cells. We have recently shown that apical SFs are anchored to the apical plasma membrane (PM) in a manner similar to how basal SFs are attached to focal adhesion sites. We propose calling such apical SF–membrane attachment sites "apical plaques." To study the macromolecular composition of the apical plaque and the focal adhesion in endothelial cells (ECs) in situ, we examined by confocal laser scanning and fluorescence microscopy guinea pig aortae stained with various antibodies against focal adhesion–associated proteins. Basal SFs oriented parallel to the blood flow direction were mainly located in the upstream half of the cell. Thin apical SFs were also observed. Spotty staining patterns were observed in the basal and the apical portions of cells stained with anti-vinculin, anti-talin, anti-paxillin, or anti-fibronectin receptor, indicating the presence of focal adhesions and apical plaques in ECs in situ. Although fibronectin receptors were present in the apical plaque, fibronectin was not detected on the apical cell surface. Our data suggest that the molecules responsible for the SF-PM association are the same between in vitro and in situ cells. Our results appear to support a hypothesis that the SF system is involved in sensing and/or signal transduction of fluid mechanical forces.

Key Words: endothelium • focal adhesion • mechanotransduction • vascular system • shear stress

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Endothelial cells lining the inner surface of blood vessels are constantly exposed to blood flow, and both in vivo and in vitro studies indicate that aspects of EC physiology, morphology, and biosynthetic activity are influenced by hemodynamic factors.^{1 2 3} In addition, the areas within the artery where atherosclerotic legions are likely to develop often correspond to the so-called "low shear stress" region.⁴ Thus, it has been phenomenologically established that ECs are able to sense fluid flow and to respond to it in many ways. In addition, our laboratory has recently demonstrated that PECAM-1 is rapidly tyrosine phosphorylated in ECs exposed to fluid flow.^{5 6} Using ECs exposed to fluid flow in vitro, investigators have demonstrated that flow of culture medium rapidly increases K⁺ conductance,^{7 8} activates trimeric G proteins,^{9 10} and induces Ca²⁺ mobilization.^{11 12 13 14} These are some of the early events occurring in ECs under fluid flow that may be related to flow sensing or cellular signal transduction and are thought to cause various later responses.

Of various types of EC responses to flow, the morphological responses, such as changes in cell shape,^{15 16 17} cytoskeletal orientation,^{18 19 20} and cell migration,^{21 22} reflect the direction of fluid flow. Whether or not other types of responses express directionality is not known, but it is difficult to think that responses such as flow-activated gene expression and flow-induced production of vasoactive substances occur in a polarized manner with respect to the flow direction. How ECs transduce directional information is an important question to ask. This is important from the standpoint of not only basic cell biology but also the study of the pathogenesis of atherosclerosis, since it is known that the endothelium at the atherosclerotic region shows much reduced polarized morphology.⁴ The most straightforward way to transmit directional information between two points within a cell is to have a physical link between them. Recently, the importance of the cytoskeleton in mechanotransduction has been proposed,^{2 23 24 25} and the SF system may be a good candidate for this role.

Organized actin filaments in cells forming a monolayer (including ECs) are generally found as the circumferential actin bundle and the SF. The majority of SFs are located within the basal cell cortex, hence called the basal SFs, and their ends are associated with focal adhesion sites.^{26 27} However, some SFs are located above the nucleus and we have proposed to call them apical SFs (see Reference 28 for review). In a recent study, we have shown that apical SFs in cultured human fibroblasts are associated with the apical PM via a specialized structure that we call the apical plaque.²⁸ The basic macromolecular composition of the apical plaque is essentially the same as that of the focal adhesion, indicating that the apical SF in cultured cells is firmly anchored to the apical PM. The other end of the apical SF is often anchored to the basal PM via the focal adhesion. On the basis of these morphological data, we have proposed that the apical plaque and apical SFs might work as mechanotransducers, directly transmitting mechanical forces exerted on the apical surface to the interior and the base of the cell.²⁸

Shear stress has been shown experimentally to induce F-actin redistribution in ECs.^{19 29 30} Under high shear stress conditions, ECs contain long, thick SFs while the expression of circumferential actin bundle is diminished. The SFs described in these studies are the basal type. There are a few reports that deal with fluid flow and the focal adhesion,^{29 31} but no data are available at present on the apical SFs and plaques in ECs in situ.

If one were to suggest a role for apical SFs in flow sensing by ECs, it would be first necessary to establish the presence of the apical SFs firmly attached to the luminal PM of ECs in situ. In this study, blood vessels were immunofluorescently stained with various antibodies against focal adhesion–associated proteins, and the staining patterns were analyzed by CLSM. Semithin cryosections were also used. Our results have indicated that the apical SFs and apical plaques are indeed present in ECs in situ and that the basic molecular organization of the apical and the basal SF-PM connections in ECs in situ is essentially identical to that of the focal adhesion of cultured cells.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Preparation of Blood Vessels for Fluorescence Microscopy
Aortae were obtained from normal adult guinea pigs weighing 450 to 650 g. Animals were anesthetized by an intraperitoneal injection of pentobarbital sodium (50 mg/mL). Before they were killed, 200 U of heparin sodium in 200 mL of 0.85% NaCl or HBSS (pH 7.0 to 7.2) with 0.2 mmol/L MgCl₂ was infused into the animals via the left ventricle. After perfusion, the descending thoracic and abdominal aortae were excised. The vessel was cut open along the dorsal wall and pinned onto a dental wax plate, exposing the luminal surface.

The aortae were fixed on a dental wax plate using a microwave irradiation method.³² Each aorta was put into a 100-mL beaker containing 50 mL of ice-cold 2% paraformaldehyde in PBS. The beaker was placed in a plastic box containing 600 mL water with ice and irradiated in a 500 W microwave oven for 30 seconds. After microwave irradiation, they were further treated with the same fixative for 30 minutes at room temperature. After they were rinsed with PBS, vessels were cut crosswise into small segments (4 to 8 mm in length) and processed for immunofluorescence microscopy.

Antibodies and Fluorescent Reagents
Monoclonal anti-vinculin (Sigma Chemical Co), anti-talin (Sigma), anti-paxillin (Zymed), and anti-phosphotyrosine (clone PY-20³³ ; ICN) were purchased. Polyclonal anti-fibronectin (Yagai), anti-fibronectin receptor (Chemicon), and fluorescein-labeled goat anti-rabbit IgG (Cappel) and anti-mouse IgG (Cappel or Sigma) were also purchased. PI and fluorescein-labeled phalloidin were obtained from Sigma.

Immunofluorescence Procedures
For the whole mount preparation, aorta segments were permeabilized with 0.01% to 0.5% Triton X-100 in PBS for 5 minutes and then washed with PBS. They were incubated with 10% normal goat serum for 20 minutes and then treated with one of the following primary antibodies for 60 to 90 minutes at room temperature: anti-vinculin (dilution, 1:400), anti-talin (1:50), anti-paxillin (1:100), anti-phosphotyrosine (1:200), anti-fibronectin (1:50), and anti-fibronectin receptor (1:100). After they were washed in PBS, they were incubated with fluorescein-labeled goat anti-rabbit IgG (1:100 to 1:200) or anti-mouse IgG (1:100 to 1:200) for 60 to 90 minutes and washed again in PBS. Actin filaments were stained with fluorescein-labeled phalloidin. Stained aorta pieces were mounted in 90% glycerol in PBS containing 2.5% 1,4-diazabicyclo [2.2.2]-octane (DABCO, Aldrich). Some specimens were also stained with 10 µg/mL PI in PBS to identify nuclei.

For cryosectioning, fixed aortae were immersed in 2.3 mol/L sucrose solution in PBS or 1/15 mol/L PBS, pH 7.4, and frozen in liquid N₂. Semithin cryosections (0.4 to 1.0 µm in thickness) were cut on a Reichert ultramicrotome equipped with cryokits and stained with various antibodies as described above. As a control, fixed aortae were stained with the secondary antibodies alone or treated with normal rabbit serum and then the secondary antibodies. These samples did not exhibit the spotty staining pattern described in the "Results." The internal elastic lamina stained nonspecifically with some secondary antibodies.

Cryosections were observed using an Axiophoto (Carl Zeiss) epifluorescence microscope with an apochromat x63 (N.A. 1.4, oil) objective lens. Whole mount specimens were examined using a confocal laser scanning microscope (GB-200, Olympus) with a plan-apochromat x60 (N.A. 1.4, oil) objective lens.

Electron Microscopy
After a brief perfusion with 0.85% NaCl containing heparin sodium (1 U/mL), the descending aorta was perfusion-fixed with 2.5% glutaraldehyde and 2% paraformaldehyde in 0.1 mol/L sodium cacodylate buffer, pH 7.4. The aorta was cut into small pieces and further fixed in fresh fixative for 3 hours at room temperature and then overnight at 4°C. After they were washed in 0.1 mol/L sodium cacodylate buffer, the vessel pieces were post-fixed with 1% OsO₄ for 2 hours on ice, stained en bloc with 0.5% uranyl acetate for 2 hours, and then dehydrated and embedded in Epon 812. Thin sections were cut and examined using a JEOL JEM 2000FX–type electron microscope (JEOL) operated at 80 kV.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
SFs in the Apical Portion of ECs In Situ
Whole mount aorta specimens stained simultaneously with fluorescein-labeled phalloidin and PI were analyzed by CLSM (Fig 1). When the basal portions of the ECs were brought into focus, basal SFs oriented parallel to the direction of blood flow were observed. The strongly stained and prominent SFs were usually located in the proximal half of a cell in relation to blood flow (Fig 1a). When the microscope focus was brought up to cells' summit, apical SFs were observed over the PI-stained nucleus (Fig 1b). Although the actin bundles in the apical portion also run parallel to the direction of blood flow, they were considerably thinner than the basal SFs. In situ ECs stained with anti-myosin or anti–-actinin revealed dotty, linear staining along both the apical and the basal SFs (data not shown, see Reference 18). Our results suggest that the apical SFs have the same basic macromolecular composition and organization as the basal SFs in ECs in situ.

View larger version (95K): [in this window] [in a new window]   Figure 1. In situ ECs stained doubly with fluorescein-labeled phalloidin (green) and PI (red). Two optical sections obtained by CLSM are illustrated. Because the specimen was not perfectly flat, each optical section captured a portion of each cell. a, Basal portion of ECs showing basal stress fibers. b, A confocal section near the top of ECs. Arrowheads indicate apical stress fibers. Blood flow was from left to right.

By electron microscopy, in addition to basal SFs,³⁴ we observed microfilament bundles in the apical portion (Fig 2). These apical SFs were smaller in size but also appeared to attach to the PM. Increased electron density was noted at the SF-PM attachment site. Most of the apical SFs were over the nucleus and closely associated with the PM near the top of the cell. Cryosectioned specimens stained with fluorescein-labeled phalloidin exhibited, in addition to basal staining, dotty intense staining along the apical PM (Fig 3). The latter staining represents sectioned apical SFs.

View larger version (140K): [in this window] [in a new window]   Figure 2. Electron micrographs showing apical stress fibers in ECs in situ. Arrowheads indicate portion of apical microfilament bundles. These apical stress fibers attach to the apical PM with electron dense apical plaques (a, b). N indicates nucleus; L, vessel lumen.

View larger version (85K): [in this window] [in a new window]   Figure 3. A fluorescence micrograph of a semithin cryosection stained with fluorescein-labeled phalloidin. In the apical portion of the endothelium, dotty phalloidin-staining representing apical stress fibers is seen (arrowheads). At the base of the endothelium, another set of dots representing basal stress fibers is seen. Smooth muscle cells in the media are intensely stained. EC indicates endothelial cells; IE, internal elastic lamina; and SM, smooth muscle cells.

Macromolecular Composition of the Focal Adhesion and the Apical Plaque in ECs In Situ
Vinculin
A piece of aorta was cryosectioned along the long axis and stained with anti-vinculin. Smooth muscle cells in the media were heavily stained mostly along the PM. Dotty staining was observed along the base of ECs. The stained spots are presumably focal adhesions and other basal SF-PM attachment sites.^{28 35} Smaller and less bright staining spots were also observed in the apical portion of the endothelium (Fig 4). We also studied whole mount samples by CLSM and observed anti-vinculin staining along both the apical and the basal regions of ECs. Fig 5 is a CLSM image of the apical part of cells, showing small anti-vinculin staining spots. Some spots are yellow, not green, due to superimposition of green anti-vinculin with red PI staining. Note that the cell border is also stained. This staining is presumably associated with the adherens junction between neighboring cells. At the basal portion of the same sample, short linear staining patterns running parallel to the blood flow direction were detected. Simultaneous staining of aortae with rhodamine-labeled phalloidin and anti-vinculin revealed that the basal anti-vinculin staining patterns were superimposable with the basal SFs (data not shown, see Reference 34).

View larger version (92K): [in this window] [in a new window]   Figure 4. A semithin cryosection stained with anti-vinculin. Vinculin staining is observed in both the apical and the basal portions of ECs in dotty configurations. Arrowheads indicate apical anti-vinculin staining. Note that the membrane portion of smooth muscle cells is heavily stained outlining individual smooth muscle cells. The internal elastic lamina stained nonspecifically.

View larger version (163K): [in this window] [in a new window]   Figure 5. An en face confocal image of anti-vinculin staining near the apical surface of the endothelium. A whole mount preparation doubly stained with anti-vinculin (green) and PI (red). Small but distinct dots stained with anti-vinculin (arrowheads) are present against the red nucleus.

Paxillin
Paxillin is a vinculin³⁶ and integrin³⁷ binding protein. Immunofluorescent localization of paxillin by CLSM is shown in Fig 6. In the apical portion, anti-paxillin staining was detected in a faint dotty pattern over the nucleus (Fig 6a). Here again, due to superimposition with red PI staining, many of the anti-paxillin staining spots appear yellow. The overall distribution of the anti-paxillin staining spots over the nucleus was similar to that of the anti-vinculin spots. Sometimes, whole mount preparations have small folds. Such a folded area presents a side, not en face, view of the endothelium (Fig 6b). A whole mount preparation viewed from its side exhibited anti-paxillin staining as dots in both the apical and the basal portions. Anti-paxillin staining was associated with the smooth muscle cell periphery.

View larger version (127K): [in this window] [in a new window]   Figure 6. Confocal immunofluorescence micrographs of whole mount preparations doubly stained with anti-paxillin (green) and PI (red). a, An en face view of the top portion of ECs. Arrowheads show apical anti-paxillin staining spots. b, A side view of the endothelium, showing the apical (arrowheads) and the basal (arrows) anti-paxillin staining spots. The lumen of the vessel is the dark area toward the top of the figure.

Talin
Anti-talin staining patterns are shown in Fig 7. Dotlike staining also occurred in the apical and the basal portions of cells. In some places, the basal dots aligned linearly, presumably along basal SFs (Fig 7a). Apical staining is best appreciated in cryosections (Fig 7b). Heavy staining was associated with the smooth muscle cell periphery. Although simultaneous staining with anti-vinculin, anti-paxillin, or anti-talin has not been done due to the fact that all of the probes are mouse monoclonal antibodies, we found that anti-vinculin, anti-paxillin, and anti-talin staining patterns over the nucleus were strikingly similar.

View larger version (108K): [in this window] [in a new window]   Figure 7. Immunofluorescent localization of talin. a, Linear dotty staining presumably associated with the basal SFs is seen at the base of ECs. b, A semithin cryosection showing dotty staining patterns at the basal (arrows) and the apical (arrowheads) portions of the endothelium. The internal elastic lamina stained nonspecifically.

Fibronectin
Since some specific extracellular matrix proteins are known to be associated with both focal adhesions and apical plaques in cultured cells,²⁸ we stained whole mount preparations and semithin cryosections of aorta with anti-fibronectin. Underneath the endothelium, anti-fibronectin showed a linear pattern parallel to the blood flow direction (Fig 8a), as we reported previously.³⁴ However, when the focus was brought up to the apical portion of these cells, no fluorescent signal could be detected. Cryosectioned aortae stained with anti-fibronectin are shown in Fig 8b. Although heavy spotty staining patterns along the basal cell surface and general staining of the internal elastic lamina were observed, no staining was associated with the luminal surface.

View larger version (76K): [in this window] [in a new window]   Figure 8. Immunofluorescent localization of fibronectin. a, A confocal micrograph of a whole mount preparation stained with anti-fibronectin showing fibrous staining under ECs. Fibronectin fibrils are lined parallel to the blood flow direction. The entire basement membrane is also stained. b, A semithin cryosection stained with anti-fibronectin. A heavy spotty staining pattern is detected in the basal portion of the endothelium (arrows), but no labeling is present along the apical portion of the cell. Arrowheads delineate the luminal surface of the endothelium.

Fibronectin Receptor (₅ß₁)
Spotty anti-fibronectin receptor staining was observed along the base of ECs. In addition, despite the lack of anti-fibronectin staining on the apical surface of in situ ECs, spotty staining was observed with anti-fibronectin receptor in the apical portion. Fig 9 is a confocal image of a whole mount preparation focused at the apical part of the endothelium. Many brightly stained dots are present over the nucleus. This staining pattern is similar to those of anti-vinculin, anti-talin, and anti-paxillin.

View larger version (119K): [in this window] [in a new window]   Figure 9. A confocal microscope image of the apical portion of EC stained with anti-fibronectin receptor (green) and PI (red). Anti-fibronectin receptor (5ß1) staining (arrowheads) is dotty against red nuclei.

Tyrosine-Phosphorylated Protein(s)
Phosphorylation of tyrosine residues of various proteins plays an important role in cellular signaling. It is known that there are several proteins with phosphorylated tyrosines in the focal adhesion of cultured cells.^{23 33 38 39} To examine whether or not tyrosine-phosphorylated proteins are also present at the cell-substrate contact site in situ, aorta specimens were stained with anti-phosphotyrosine (PY-20). This monoclonal antibody has been shown to stain focal adhesion sites in cultured cells.³³ In ECs in situ, PY-20 staining was detected in both the apical and the basal portions of the cell as well as the cell-to-cell junction. Semithin longitudinal cryosections also clearly revealed PY-20 staining in the apical and the basal portions of cells and the cell-cell attachment site (Fig 10).

View larger version (231K): [in this window] [in a new window]   Figure 10. Immunofluorescent localization of tyrosine phosphorylated proteins in ECs in situ. Cryosection (a) and whole mount (b) preparations stained with anti-phosphotyrosine (PY-20). a, PY-20 staining is detected at the cell-cell junction as well as at both the apical (arrowhead) and the basal (arrow) portions of the cell. Smooth muscle cells hardly stained with PY-20. b, Double staining with PY-20 and PI. A confocal optical section is cut in the middle portion of the cell. At this level, staining is seen primarily at the cell-cell junction.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
For cultured cells, a considerable amount of data is available on the molecular composition and organization of focal adhesions,^{26 27 40} and we have recently reported on the partial characterization of apical plaques.²⁸ However, little is known about the similar structures in cells in situ. In this study, we investigated the distribution of representative focal adhesion–associated proteins at the SF-PM attachment sites in ECs in situ. We identified specialized structures analogous to the focal adhesion and the apical plaque that were described for tissue-cultured cells. Our immunofluorescence data indicate that the molecular makeup of these SF-PM attachment sites is essentially identical between in vitro and in situ cells.

Hemodynamic forces are known to influence actin filament organization in ECs both in vitro^{19 29 30 41} and in situ.^{20 34 42 43 44 45 46} SFs in the basal portion of in situ endothelium ran parallel to the direction of blood flow, and prominent SFs were located within the proximal (in relation to blood flow) half of the cell^{18 19 46} and within cultured ECs exposed to fluid flow.²⁹ Since the ends of the basal SF are associated with focal adhesions, the distribution of focal adhesions may also be influenced by fluid flow. Indeed, vinculin localization in cultured endothelial cells is affected by flow.²⁹ Davies et al³¹ showed flow-dependent directional remodeling of focal adhesions in live cultured ECs. While these in vitro studies suggest that fluid flow affects the distribution pattern of focal adhesions, this study and our earlier³⁴ studies on the in situ endothelium failed to provide evidence for flow-dependent uneven distribution of focal adhesions. The in vitro data are from cells exposed to fluid flow for only a short period of time. Thus, although it is clear that focal adhesions are remodeled and change their distribution when fluid flow is first applied to ECs, it is not known whether such alterations perpetually occur in cells exposed continuously to flow for an extended period of time, such as months and years. It is conceivable that the observed in vitro events are of a temporary nature and that in situ ECs have more stable and evenly distributed focal adhesions.

In addition to its localization underneath cells, fibronectin has been immunocytochemically localized on the apical surface of cultured fibroblasts^{28 47} and ECs (authors' unpublished observations, 1996). Consistent with these observations, a biochemical study has demonstrated that ECs express several types of integrins on both the basal and the apical cell surfaces in vitro and in vivo and that some of the types, such as ₃ß₁, ₅ß₁, and _vß₃, have fibronectin-binding capacity.⁴⁸ Fibronectin in the subendothelial space forms a fibrous meshwork, and its pattern sometimes coincides with the SF pattern within the cell over it.^{34 49 50} We have recently shown that the streaky fibronectin pattern on the apical surface of human fibroblasts corresponds well with the cluster of apical plaques, consisting of a fibronectin receptor, talin, vinculin, and paxillin, to which apical SFs are tightly anchored.²⁸ In our present study, however, we did not observe anti-fibronectin staining on the luminal surface of ECs in situ, although clusters of fibronectin receptors were localized within the apical surface. This result appears to indicate that the fibronectin receptor on the luminal surface of ECs in the guinea pig aorta is either unoccupied or occupied by some unknown ligand(s). If the receptors are unoccupied, an intriguing question is why they are not available to plasma fibronectin. One possibility is that these receptors are in an inactive form. The present study has identified an interesting topic for future studies.

We have previously proposed that the apical SF has an important role in maintaining the structural integrity of ECs in situ that are constantly exposed to hemodynamic forces of flowing blood.⁴⁴ In addition to their structural role, apical SFs may be able to transmit fluid dynamic forces from the apical surface to the other parts of the cell. Indeed, attractive concepts have recently been advanced that the cytoskeleton of the cell may transmit tension and be able to function as a mechanosensor in ECs^{2 51} and that integrin, and the structure that is linked to this cell adhesion molecule, is a mechanotransducer.^{2 23 24} The present work together with our earlier study on cultured fibroblasts suggest that the apical SF is a good candidate for the proposed cytoskeletal structure responsible for fluid-flow sensing and/or signal transduction. In fact, depolymerization of actin filaments with cytochalasin B inhibits EC response to fluid flow.^{52 53}

Anti-phosphotyrosine stained both the apical and the basal sides of cells as well as the lateral cell-cell adhesion site. Since the focal contact and the apical plaque of cultured fibroblasts are labeled with anti-phosphotyrosine,²⁸ it is plausible that some, if not most, of the staining in the endothelium are the SF-PM attachment sites. As we have demonstrated, there are many proteins associated with the in situ focal adhesions and apical plaques whose tyrosine residues can be phosphorylated. Such proteins include talin, vinculin, and paxillin. Although the identity of the tyrosine-phosphorylated protein(s) localized in this study is not known, it is possible that some of these focal adhesion proteins contain phosphotyrosine(s). Heavy staining at the cell-cell adhesion site is interesting because this is another area where the force from fluid flow could become concentrated and where it is known that vinculin, -actinin, paxillin, and other cytoskeletal and adhesion molecules are localized. In addition, we have recently found that PECAM-1, which is highly concentrated at the inter-EC adhesion site,⁵ is tyrosine phosphorylated especially when ECs in a monolayer are exposed to flow.⁶ Since tyrosine phosphorylation is involved in transducing various chemical signals, it may also play a role in mechanosensing by ECs.

	Selected Abbreviations and Acronyms

 

CLSM = confocal laser scanning microscopy EC = endothelial cell PECAM-1 = platelet endothelial cell adhesion molecule-1 PI = propidium iodide PM = plasma membrane SF = stress fiber

	Acknowledgments

 
This work was supported in part by grants-in-aid for Scientific Research from the Ministry of Education, Science, and Culture of Japan, grants from the Ministry of Health and Welfare of Japan, and Special Coordination Funds for Promoting Science and Technology from the Science and Technology Agency of Japan.

Received June 3, 1996; accepted July 31, 1996.

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