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Department of Pharmacology, Graduate School of Medical Sciences, Kyushu University, Fukuoka 812-8582, Japan

	Abstract

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We have investigated the cellular mechanisms of mechanical stress-induced immediate responses in human umbilical vein endothelial cells (HUVECs). Hypotonic stress (HTS) induced ATP release, which evoked a Ca²⁺ transient, followed by actin reorganization within a few minutes, in HUVECs. Disruption of the actin cytoskeleton did not suppress HTS-induced ATP release, and inhibition of the ATP-mediated Ca²⁺ response did not affect actin reorganization, thereby indicating that these two responses are not interrelated. ATP release and actin reorganization were also induced by lysophosphatidic acid (LPA). HTS and LPA induced membrane translocation of RhoA, which occurs when RhoA is activated, and tyrosine phosphorylation of focal adhesion kinase (FAK) and paxillin. Tyrosine kinase inhibitors (herbimycin A or tyrphostin 46) inhibited both HTS- and LPA-induced ATP release and actin reorganization, but did not affect RhoA activation. In contrast, Rho-kinase inhibitor (Y27632) inhibited all of the HTS- and LPA-induced responses. These results indicate that the activation of the RhoA/Rho-kinase pathway followed by tyrosine phosphorylation of FAK and paxillin leads to ATP release and actin reorganization in HUVECs. Furthermore, the fact that HTS and LPA evoke exactly the same intracellular signals and responses suggests that even these immediate mechanosensitive responses are in fact not mechanical stress-specific.

(Received 25 March 2004; accepted after revision 19 May 2004; first published online 21 May 2004)
Corresponding author M. Oike: Department of Pharmacology, Graduate School of Medical Sciences, Kyushu University, Fukuoka 812-8582, Japan. Email: moike{at}pharmaco.med.kyushu-u.ac.jp

	Introduction

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It is now widely accepted that mechanical stresses regulate endothelial functions. Sustained application of shear stress or membrane deformation induces various responses in vascular endothelium over hours or days (Davies, 1995; Chien et al. 1998), including changes in cell alignment (Malek & Izumo, 1996) and gene expression (McCormick et al. 2001). However, mechanical stresses also induce immediate responses in endothelium, such as the opening of stretch-activated cation channels (Popp et al. 1992), ATP release (Oike et al. 2000), Ca²⁺ responses (Schwarz et al. 1992; Oike et al. 2000) and activation of kinases (Koyama et al. 2001). It can be speculated that mechanical stress-induced chronic changes in endothelium may be the eventual consequence of immediate responses. For instance, DNA microarray assay revealed in human umbilical cord vein endothelial cells (HUVECs) that shear stress applied for 24 h altered the expression level of 52 genes more than twofold (McCormick et al. 2001) and 12 genes more than fivefold (Dekker et al. 2002), but the latter study revealed that all of these genes except for KLF2 gene were not shear stress-specific, but were expressed in a pattern similar that observed after stimulation with cytokines (Dekker et al. 2002).

Until now, little has been known about the very first intracellular signals by which mechanical stresses evoke immediate responses. This is partly because it is difficult to evaluate cellular responses properly after applying mechanical stresses for a very short period, i.e. a few minutes. To overcome this problem, we have used hypotonic stress (HTS), which swells the cells within a few minutes (Voets et al. 1999), thereby inducing membrane deformation. We have shown in bovine aortic endothelial cells (BAECs) that HTS induces ATP release (Oike et al. 2000) and actin reorganization (Koyama et al. 2001). Released ATP binds to P₂ receptors and induces Ca²⁺ responses (Oike et al. 2000) and nitric oxide production (Kimura et al. 2000). Mechanical stress-induced ATP release can also be obtained by shear stress (Bodin et al. 1991) and membrane distortion (Moerenhout et al. 2001) in vascular endothelium. Furthermore, it has been suggested that extracellular ATP may control vascular growth (Erlinge et al. 1996) and endothelial gene expression (von Albertini et al. 1998). Thus we propose that the extracellular ATP release is one of the central immediate endothelial responses to mechanical stresses.

In this study we attempted to clarify the intracellular signalling cascades by which HTS leads to immediate responses in HUVECs. We have previously reported in BAECs that tyrosine phosphorylation and RhoA/Rho-kinase are involved in HTS-induced ATP release and actin reorganization (Koyama et al. 2001). However, we did not clarify whether the activation of these signals is sequential or independent, nor did we identify the tyrosine-phosphorylated proteins involved in HTS-induced responses. We used these intracellular signals, tyrosine phosphorylation and RhoA/Rho-kinase, as initial clues to clarify the signalling cascade of mechanotransduction in HUVECs. The results obtained demonstrate that sequential activation of RhoA/Rho-kinase and FAK/paxillin plays a central role in mechanosensitive ATP release and actin reorganization in HUVECs.

	Methods

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Culture of human umbilical cord vein endothelial cells (HUVECs)

HUVECs were purchased from Cambrex (East Rutherford, NJ, USA). Cells were cultured in M199 medium supplemented with 15 µg ml^–1 endothelial cell growth supplement (Sigma, St Louis, MO, USA), 5 U ml^–1 heparin and 15% fetal bovine serum.

Measurement of intracellular Ca²⁺ concentration

[Ca²⁺]_i was measured with fura-2 membrane-permeable ester of fura-2 (fura-2 AM; Dojindo, Kumamoto, Japan) by using an Attofluor digital fluorescence microscopy system (Atto Instruments, Rockville, MD, USA) as previously described (Koyama et al. 2001). Cells grown on coverslips were loaded with and mounted on a chamber of 0.5 ml volume. The chamber was continuously perfused with each solution at a rate of 0.5 ml min^–1. All experiments were performed at room temperature (20–25°C).

Luciferin–luciferase bioluminescence assay

Extracellular ATP concentration ([ATP]_o) was measured from the cells seeded on a 96-well plate at a density of 5000 cells per well by using luciferin–luciferase bioluminescence. After the culture medium had been carefully removed, 50 µl of isotonic or hypotonic Krebs solution containing 10 mg ml^–1 luciferin–luciferase (Wako, Co., Osaka, Japan) was added to each well. Emitted photons were then counted for 10 min by a luminescence detection system (Argus-50/2D luminometer, Hamamatsu Photonics, Hamamatsu, Japan). Because cationic concentration and various drugs can affect luciferin bioluminescence, standard curves for converting the photon counts into [ATP]_o were obtained with the same solution and drug contents as each experiment.

Western blot analysis of tyrosine phosphorylation and RhoA activation

Tyrosine phosphorylation of cellular proteins and RhoA activation were assessed by chemiluminescence Western blotting, by using an enhanced chemiluminescence system (SuperSignal West Dura, Pierce Co., Rockford, IL, USA). Cells were lysed after exposure to HTS or LPA for 1, 2, 5 or 10 min. Western blot analysis for phosphotyrosine was performed by using monoclonal anti-phosphotyrosine antibody (clone PY20, Exalpha Biologicals, Watertown, MA, USA), polyclonal anti-FAK antibody (NeoMarkers, Fremont, CA, USA), polyclonal anti-phosphorylated FAK antibody against pTyr³⁹⁷ (Biosource International, Camarillo, CA, USA) and monoclonal anti-paxillin antibody (clone 5H11, NeoMarkers).

For the assessment of RhoA activation, cell lysate was centrifuged for 1 h at 100 000g, and the pellet was harvested as a membrane fraction. A constant amount of membrane fraction (50 µg protein per lane) was separated with SDS-PAGE, and RhoA was detected with monoclonal anti-RhoA antibody (Cytoskeleton, Inc., Denver, CO, USA). Expression of ß-actin protein was also assessed as an internal control, using monoclonal anti-ß-actin antibody (Sigma).

In each experiment, emitted chemiluminescence was detected and analysed with a lumino image analyser (FAS-1000, Toyobo, Osaka, Japan).

Immunological staining of endothelial F-actin

Rearrangement of F-actin was examined immunologically by using rhodamine-conjugated phalloidin (Molecular Probes, Eugene, OR, USA) according to the previously reported method (Knudsen & Frangos, 1997).

Solutions and drugs

The standard extracellular solution was a modified Krebs solution (1.5 mM Ca²⁺ solution) containing (mM): 132 NaCl, 5.9 KCl, 1.2 MgCl₂, 1.5 CaCl₂, 11.5 glucose and 11.5 Hepes; pH adjusted to 7.3 with NaOH. Hypotonic solutions were made by adding appropriate amounts of distilled water to standard Krebs solution. We have previously confirmed that the reduction of the ionic concentrations did not influence the [Ca²⁺]_i responses (Oike et al. 2000).

Tyrphostin 46 was obtained from Biomol Research Laboratory Inc. (Plymouth Meeting, PA, USA), suramin from Bayer (Leverkusen, Germany) and herbimycin A from Kyowa (Tokyo, Japan). Y-27632 was kindly provided by Mitsubishi Pharma Co. (Osaka, Japan). All other drugs were purchased from Sigma.

Data analysis

Pooled data are given as means ±S.E.M. Statistical significance was determined using Student's unpaired t test. Probabilities less than 5% (P < 0.05) were regarded as significant.

	Results

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Hypotonic stress (HTS)-induced responses in human umbilical vein endothelial cells (HUVECs)

First we examined the effects of HTS (–30%) on [Ca²⁺]_i in HUVECs. HTS induced a gradual increase in [Ca²⁺]_i (Fig. 1Aa), which was significantly inhibited by phospholipase C inhibitor (10 µM U73122) and P₂ receptor antagonist (30 µM suramin, Fig. 1B). Ca²⁺ responses were also inhibited by tyrosine kinase inhibitors (1 µM herbimycin A or 100 µM tyrphostin 46) and Rho-kinase inhibitor Y27632 (10 µM, Fig. 1A and B). Simultaneous treatment of the cells with herbimycin A and phospholipase A2 inhibitor p-bromophenacyl bromide (pBPB, 10 µM) almost completely inhibited HTS-induced Ca²⁺ transients (Fig. 1Ad and B). An exogenously applied low concentration of ATP (0.1 µM) induced a gradual increase in [Ca²⁺]_i, as in case of HTS (Fig. 1Ae). These results suggest that at lease two signals, i.e. extracellular ATP and arachidonic acid, are involved in the HTS-induced Ca²⁺ responses in HUVECs.

View larger version (28K): [in this window] [in a new window]   Figure 1.  HTS-induced increase in [Ca2+]i and ATP release in HUVECs A, HTS (–30%) induced a gradual increase in [Ca2+]i (a), which was partly inhibited by pretreatment with herbimycin A for 24 h (b) or Y27632 for 30 min (c) and almost completely abolished by pretreatment with herbimycin for 12 h and subsequent 30 min with pBPB and herbimycin A. A low concentration of ATP also induced a gradual increase in [Ca2+]i (e). B, statistical analysis of the net maximal elevation of HTS-induced increase in [Ca2+]i ([Ca2+]i,peak). HUVECs were pretreated either with U73122 (30 min), herbimycin A, Y27632 or herbimycin A with pBPB, and HTS was applied. In other experiments HTS was applied to untreated cells in the presence of suramin or tyrphostin 46. *P < 0.05 versus untreated control cells. P < 0.05 versus herbimycin A alone. C, the extracellular ATP concentration ([ATP]o) was measured 10 min after exchanging solutions to 10 mg ml–1 luciferin–luciferase-containing isotonic or –30% hypotonic solution. **P < 0.01 versus control. n.s., P > 0.05 versus control.

HTS also induced a transient reorganization of the actin cytoskeleton. Dense actin stress fibres were observed in the cytosolic area 2 min after starting HTS application, and redistributed into the peripheral focal adhesion complex in 10 min (Fig. 2). Transient formation of dense actin stress fibres was inhibited by herbimycin A and Y27632 (Fig. 2, lower panels), thereby suggesting that the same intracellular signals, i.e. tyrosine phosphorylation and Rho-kinase activation, are involved in actin reorganization and ATP release in HUVECs.

View larger version (104K): [in this window] [in a new window]   Figure 2.  HTS-induced actin reorganization in HUVECs In control cells, HTS (–30%) induced dense actin bundles by 2 min, which were converged into peripheral adhesion complex by 10 min. In contrast, actin bundles were not observed in herbimycin A- or Y27632-treated cells. The P2 receptor antagonist, suramin, did not inhibit HTS-induced actin reorganization. Each scale bar indicates 30 µm.

Tyrosine phosphorylation and RhoA activation with HTS in HUVECs

We then tried to identify the tyrosine-phosphorylated proteins which are involved in the HTS-induced responses in HUVECs. Western blotting revealed that HTS induced tyrosine phosphorylation of at least two proteins, 68 and 125 kDa, which were identified as paxillin and focal adhesion kinase (FAK), respectively, by using selective antibodies against these molecules (Fig. 3A). Furthermore, time-dependent tyrosine phosphorylation of FAK was confirmed with monoclonal anti-phosphorylated FAK antibody (Fig. 3A). Densitometric analysis of the phosphotyrosine bands revealed that both proteins were maximally phosphorylated between 2 and 5 min after starting HTS, and declined by 10 min (Fig. 3A).

View larger version (42K): [in this window] [in a new window]   Figure 3.  HTS-induced tyrosine phosphorylation and RhoA activation in HUVECs A, HTS-induced tyrosine phosphorylation of FAK and paxillin. HTS (–30%) induced a transient increase in tyrosine phosphorylation of at least two proteins, 125 and 68 kDa. A representative result is shown. These bands could be stained with antibodies against focal adhesion kinase (FAK) and paxillin, respectively. By using anti-phosphorylated FAK antibody, the transient phosphorylation of FAK was confirmed (a). Densitometric analysis of phosphorylated tyrosine bands revealed that both 125 kDa FAK and 68 kDa paxillin were maximally phosphorylated between 2 and 5 min after starting HTS (b). Data were obtained from four measurements. B, HTS (–30%) induced membrane translocation of the small G protein RhoA in HUVECs. Band densities of RhoA and ß-actin were measured, and RhoA/ß-actin values were expressed as relative to that of the isotonic control. RhoA in the membrane fraction was increased by HTS, and maximal expression was observed between 2 and 5 min. Data were obtained from 4 repeated measurements.

We then investigated the interrelation between tyrosine phosphorylation and the RhoA/Rho-kinase cascade. As shown in Fig. 4A, Y27632 significantly inhibited the HTS-induced tyrosine phosphorylation of 125 kDa FAK and 68 kDa paxillin. In contrast, herbimycin A did not affect the HTS-induced membrane translocation of RhoA (Fig. 4B). These findings suggest that the RhoA/Rho-kinase cascade is essential for the HTS-induced tyrosine phosphorylation, whereas tyrosine phosphorylation of FAK/paxillin is not required for the activation of the RhoA/Rho-kinase pathway.

View larger version (25K): [in this window] [in a new window]   Figure 4.  Interrelation between tyrosine phosphorylation and RhoA activation in HUVECs A, pretreatment with 10 µM Y27632 for 30 min abolished the –30% HTS-induced tyrosine phosphorylation of 125 kDa FAK and 68 kDa paxillin in HUVECs. **P < 0.01 versus control. Data were obtained from 4 measurements. Control data (open symbols) are the same as those shown in Fig. 3Ab. B, pretreatment with 1 µM herbimycin A for 12 h did not affect the –30% HTS-induced membrane translocation of RhoA in HUVECs. Data were obtained from 4 measurements. No significant difference was observed in the RhoA band density between control () and herbimycin A-treated cells (•). n.s., P > 0.05 versus control.

Lysophosphatidic acid (LPA) binds to its specific receptors and activates RhoA and Rho-kinase (Moolenaar, 1995). We observed that LPA (1 µM) induced a membrane translocation of RhoA both in control and in herbimycin A-treated HUVECs (Fig. 5A). As in the case of HTS, LPA also induced tyrosine phosphorylation of 125 and 68 kDa proteins, and Y27632 significantly inhibited the phosphorylation of both proteins (Fig. 5B).

View larger version (24K): [in this window] [in a new window]   Figure 5.  Lysophosphatidic acid (LPA)-induced intracellular signals in HUVECs A, LPA induced a membrane translocation of RhoA. Control or herbimycin A-treated cells were stimulated with 1 µM LPA or vehicle for 2 min, and a membrane fraction was prepared. Note that LPA induced an increase in RhoA in the membrane fraction, and herbimycin A did not affect it. n.s., P > 0.05 versus control. B, LPA (1 µM) induced tyrosine phosphorylation of 125 and 68 kDa proteins, and the tyrosine phosphorylation of those proteins was inhibited by 10 µM Y27632. Data were obtained from 4 measurements. **P < 0.01 versus untreated control.

View larger version (57K): [in this window] [in a new window]   Figure 6.  LPA induces ATP release, an increase in [Ca2+]i and actin reorganization in HUVECs A, LPA (1 µM) increased [ATP]o and this was significantly inhibited by Y27632, herbimycin A and tyrphostin 46. n.s., P > 0.05 versus control; **P < 0.01 versus control. B, LPA (1 µM) induced a gradual increase in [Ca2+]i, which was blocked by the P2 receptor antagonist suramin (30 µM). The net maximal increment of [Ca2+]i ([Ca2+]i,peak) was measured. **P < 0.01. C, LPA (1 µM) induced a transient formation of dense actin bundles, which appeared by 2 min and converged by 10 min. Y27632 and herbimycin A inhibited the LPA-induced formation of dense actin fibers. Each scale bar indicates 30 µm.

	Discussion

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We have previously reported the involvement of protein tyrosine phosphorylation and Rho/Rho-kinase in HTS-induced ATP release in BAECs (Koyama et al. 2001). We observed in the present study that the same signals were also involved in the HTS-induced responses in HUVECs. Inhibitors of tyrosine kinase and Rho-kinase inhibited the HTS-induced ATP release (Fig. 1), Ca²⁺ transient (Fig. 1) and actin reorganization (Fig. 2). Furthermore, we have identified the tyrosine-phosphorylated proteins involved in ATP release for the first time as FAK and paxillin (Fig. 3A), and demonstrated the HTS-induced membrane translocation of RhoA (Fig. 3B), a hallmark of its activation (Kranenburg et al. 1997). We have previously shown in BAECs that ATP release is also induced by LPA (Koyama et al. 2001), and this study further clarified that LPA mimicked all of the HTS-induced responses in HUVECs, i.e. RhoA activation (Fig. 5A), tyrosine phosphorylation of FAK and paxillin (Fig. 5B), ATP release (Fig. 6A) and actin reorganization (Fig. 6B). The pharmacological profiles of these LPA-induced responses were also identical to those of HTS-induced ones. LPA is a bioactive lipid that is generated in extracellular fluid by secretory lysophospholipase D (Tokumura, 2002). LPA binds to its specific receptor(s) and, of three known LPA receptor subtypes, LPA₁ and LPA₂ receptors have been reported to be involved in the activation of RhoA (Ishii et al. 2000). Furthermore, activation of FAK by LPA was reported in Swiss 3T3 fibroblasts (Hunger-Glaser et al. 2003). Therefore, here we propose that simultaneous activation of RhoA/Rho-kinase and FAK/paxillin, either by mechanical stresses or by receptor activation, leads to ATP release and actin reorganization in HUVECs.

We have found that RhoA/Rho-kinase activation and tyrosine phosphorylation are not independent phenomena but are interrelated. Y27632 inhibited HTS- and LPA-induced tyrosine phosphorylation of FAK and paxillin (Figs 4A and 5B). Furthermore, tyrosine kinase inhibitors did not affect HTS- and LPA-induced RhoA activation (Figs 4B and 5A), but suppressed LPA-induced ATP release (Fig. 6A). Therefore, these findings indicate that HTS and LPA consecutively activate FAK/paxillin and RhoA/Rho-kinase, and the phosphorylation of FAK/paxillin is located downstream of the RhoA/Rho-kinase cascade in HUVECs. Previous studies have also suggested the sequential activation of these intracellular signals. For instance, microinjection of RhoA or the activation of endogenous RhoA with LPA or bombesin induced the activation of FAK/paxillin, and this lead to the formation of actin stress fibres in Swiss 3T3 fibroblasts (Flinn & Ridley, 1996). Furthermore, inhibition of RhoA with botulinum toxin C3 inhibited cyclic strain-induced tyrosine phosphorylation of FAK/paxillin in BAECs (Yano et al. 1996).

ATP release has been considered as one of the main mechanical stress-induced immediate responses not only in endothelium (Burnstock, 1999) but also in other cell types, such as astrocytes (Joseph et al. 2003) and epithelium (Burnstock, 1999; Jans et al. 2002). Released extracellular ATP binds to P₂ receptors and induces Ca²⁺ responses (Oike et al. 2000; Joseph et al. 2003) and various subsequent cellular responses including NO production (Kimura et al. 2000). Furthermore, a recent report suggested that vasorelaxation induced by third-generation ß-adrenoceptor antagonists is due to ATP release from endothelium (Kalinowski et al. 2003). However, it should be noted that ATP release is not the only mechanism for the HTS-induced Ca²⁺ mobilization in HUVECs, since the inhibition of ATP release did not abolish the HTS-induced Ca²⁺ transients (Fig. 1). Reorganization of endothelial actin cytoskeleton leads to the alterations in cell shapes and plays a central role in vascular permeability (Amerongen et al. 2000) and adaptation of cell alignment to a new haemodynamic environment (Birukov et al. 2002). We have shown that these mechanosensitive responses, i.e. ATP release and actin reorganization, are not related each other, but are activated by the same series of intracellular signals. This may also suggest a possibility that a variety of mechanosensitive responses in endothelium in fact originate from a few common intracellular signals. Furthermore, since LPA mimicked all of the HTS-induced responses, this study also indicates that even these initial signals are not mechanical stress-specific, thereby supporting the previous report using DNA microarray assay that most of the genes mobilized by shear stress are in fact not shear-specific (Dekker et al. 2002).

In conclusion, HTS-induced immediate endothelial responses are mediated by the sequential activation of RhoA/Rho-kinase and FAK/paxillin in HUVECs. LPA, which activates these intracellular signalling cascades, induces the same responses.

	References

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	Acknowledgements

 
This study was carried out as a part of ‘Ground Research Announcement for the Space Utilization’ promoted by the National Space Development Agency of Japan and Japan Space Forum.

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