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Correspondence to Martin Alexander Schwartz: maschwartz{at}virginia.edu
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Despite the systemic nature of most atherogenic stimuli, atherosclerosis is a focal disease affecting discrete regions of the vasculature, such as vessel curvatures and bifurcations. These regions are characterized by complex flow patterns. including flow reversal, flow gradients, secondary flows with rapid changes in flow direction, and, in some regions, turbulence (VanderLaan et al., 2004). We group all of these flow patterns under the rubric of disturbed flow. Endothelial cells sense the force of flowing blood, termed shear stress, and different blood flow patterns regulate endothelial behavior. Regions of blood vessels exposed to undisturbed, unidirectional laminar flow (henceforth termed laminar flow) are protected from atherosclerosis, and in vitro prolonged laminar flow stimulates expression of athero-protective genes (Traub and Berk, 1998; Brooks et al., 2004). By contrast, disturbed flow patterns stimulate proatherosclerotic events, including increased monolayer permeability; decreased antioxidant capacity; and enhanced expression of proinflammatory genes, such as ICAM-1, VCAM-1, and monocyte chemotactic protein-1 (MCP-1; Jo et al., 1991; De Keulenaer et al., 1998; Phelps and DePaola, 2000; Brooks et al., 2004). The correlation between flow patterns and endothelial monolayer permeability has recently been demonstrated in vivo, where vascular permeability is inversely proportional to time-average shear stress and correlated with increased flow oscillation and flow gradients (Himburg et al., 2004; LaMack et al., 2005). Interestingly, onset of laminar shear stimulates many of the same responses as disturbed shear; however, in laminar shear, these events are down-regulated as cells adapt, whereas in disturbed shear, they are sustained. Thus, failure to adapt is thought to be critical for responses to disturbed shear (Orr et al., 2006).
The molecular mechanisms involved in flow-induced endothelial permeability are unknown. Although vesicular transport and transcellular channels may contribute to endothelial permeability, paracellular pore formation is most likely the major pathway for macromolecule transport across arterial endothelium (Ogunrinade et al., 2002). Paracellular permeability is limited by cell–cell interactions, especially those in tight junctions (TJs). Multiple molecular mechanisms implicated in regulation of endothelial paracellular permeability include changes in gene expression, phosphorylation of junctional components, myosin-dependent contractility, and stability of cortical actin (Ogunrinade et al., 2002). Many signaling pathways regulate permeability, most of which affect cortical actin or myosin (Yuan, 2002). Actin remodeling is regulated by the Rho family of small GTPases, including Rho, Rac, and Cdc42 (Jaffe and Hall, 2005). The p21-activated kinase (PAK) family of Ser/Thr kinases is important for Rac and Cdc42-induced cytoskeletal remodeling, affecting both actomyosin contractility and the stability of actin filaments (Bokoch, 2003). Recently, PAK was shown to stimulate paracellular pore formation and increased endothelial cell permeability in response to a wide range of cellular stimuli (Stockton et al., 2004). PAK-mediated permeability responses require the localization of active PAK to cell–cell junctions, where PAK stimulates the phosphorylation of myosin light chain to induce contractility (Stockton et al., 2004). In addition, PAK can also promote paracellular pore formation by phosphorylating VE-cadherin, which results in its arrestin-dependent internalization (Gavard and Gutkind, 2006). PAK contains multiple domains that bind scaffolding proteins, such as Nck and Grb2, capable of regulating PAK localization (Lu et al., 1997; Puto et al., 2003). Interestingly, both PAK localization to cell–cell junctions and PAK-mediated permeability were inhibited with a cell-permeable peptide corresponding to the Nck-binding sequence of PAK (Stockton et al., 2004).
Shear stress activates the integrin family of ECM receptors, and new integrin ligation mediates effects of flow on Rac, Cdc42, and Rho activity (Jalali et al., 2001; Tzima et al., 2001, 2002, 2003). Flow-induced GTPase regulation mediates cell alignment in the direction of flow and stimulates the transcription factor NF-
B, which is important for expression of inflammatory genes in the endothelium (Tzima et al., 2002). The idea that integrin ligation mediates these effects suggested that alterations in the subendothelial matrix composition would affect which integrins become ligated, resulting in differential signaling in response to flow. Indeed, shear stress activates NF-B when endothelial cells are plated on a fibronectin (FN) or fibrinogen matrix, but not when cells are plated on collagen or laminin. Furthermore, FN and fibrinogen were deposited at sites of disturbed flow in vivo, which correlated with expression of NF-B target genes (Orr et al., 2005). These results suggest that matrix remodeling plays a causal role in atherogenesis. In this work, we investigate the role of flow and ECM in endothelial permeability in atherogenesis.
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Matrix-dependent PAK signaling regulates flow-induced endothelial permeability
Because PAK regulates permeability of endothelial monolayers (Stockton et al., 2004; Gavard and Gutkind, 2006), we tested whether matrix-specific PAK activation correlates with permeability. To assay flow-mediated endothelial cell permeability, we developed a novel transwell assay that used a modified cone and plate device adapted to 75-cm transwell chambers (Fig. 2 A).
Using this system, we applied shear to endothelial cell monolayers and assessed the movement of a tracer across the filter. Membranes were then fixed and stained to ensure that no cell loss occurred during the assay. Consistent with previous results, we found that laminar flow transiently increased endothelial cell permeability, which returned to baseline by 4 h (Fig. 2 B).
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80%, whereas an inactive control peptide containing mutations in key proline residues involved in Nck binding (Kiosses et al., 1999) had no effect (Fig. 3 C). Though transfection efficiency with the PAK AID was 50%, the decrease in flow-induced permeability approached 50%, indicating that it is also highly effective (Fig. 3 D).
In addition to HRP, Alexa 488–labeled BSA was also used to determine flow-induced permeability. Absolute permeability to both BSA and HRP were similar (Fig. 3, A and E), and both showed sensitivity to PAK inhibition (Fig. 3 E).
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Effects of cytokines and oxidized LDL (oxLDL)
Although flow patterns regulate susceptibility to atherosclerosis, a number of soluble factors also promote atherosclerotic plaque development and likely contribute to endothelial permeability in atherosclerosis. OxLDL stimulates endothelial cell permeability through a Rho-dependent pathway (Essler et al., 1999; Siess et al., 1999). In early atherogenesis, activated endothelial cells and macrophages produce MCP-1, which also stimulates endothelial permeability (Stamatovic et al., 2003), as do the macrophage-derived cytokines TNF
and IL-1ß (Martin et al., 1988; Brett et al., 1989). Furthermore, mice deficient in either MCP-1 or TNF show reduced atherosclerosis (Gu et al., 1998; Ohta et al., 2005). We previously showed that TNF-induced endothelial permeability was reduced by the PAK-Nck inhibitory peptide (Stockton et al., 2004). To analyze the matrix dependence of these factors, PAK phosphorylation was assessed in endothelial cells plated on FN or MG. Though the time courses were distinct, MCP-1, TNF, and oxLDL stimulated PAK phosphorylation in cells on FN but not on MG (Fig. 5). In all cases, phosphorylated PAK localized to cell–cell junctions, and this localization was inhibited by the Pak-Nck peptide (Fig. S3, available at http://www.jcb.org/cgi/content/full/jcb.200609008/DC1).
We next examined monolayer permeability. All of these factors triggered matrix-dependent increases in permeability (Fig. 6 A) that were inhibited by the PAK-Nck blocking peptide (Fig. 6 B) and by expression of the PAK AID (Fig. 6 C). Thus, effects of a number of atherogenic soluble factors on PAK- dependent permeability are strongly modulated by the ECM.
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PAK inhibition reduces permeability in atherosclerosis in vivo
To determine whether PAK is responsible for the increased permeability during development of atherosclerosis, 32-wk-old ApoE–/– mice (chow diet) were given intraperitoneal injections of the PAK-Nck blocking peptide or a control peptide. Mice under these conditions are reported to develop moderate atherosclerotic lesions, though plaque development is slower than in animals on a high-fat Western diet (Reddick et al., 1994). Vascular permeability within the aorta was then assessed by measuring leakage of Evans blue dye into the vascular wall. Aortas from C57Bl/6 mice were used as a source for healthy, atherosclerosis-free vessels. Each mouse received 1 mg of peptide at 24 h and 1 h before Evans blue injection via the tail vein. After 30 min, leakage of dye into the aorta was assessed. Although little Evans blue accumulated in the aorta of C57Bl/6 mice, in ApoE–/– mice, dye was apparent at the lesser curvature of the arch and at branch points for major arteries in both the nontreated and control peptide–treated animals (Fig. 8), consistent with known athero-prone regions.
The Pak-Nck peptide inhibited 67% of the increase in permeability, relative to healthy vessels. These data suggest that PAK makes an important contribution to permeability in atherogenesis.
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The mechanisms regulating the matrix specificity of PAK activation are presently unclear. Flow-induced Rac activation is equivalent on all matrices (unpublished data), suggesting that there may be matrix-specific signals that inhibit PAK activation. Known mechanisms limiting PAK activation include binding of PAK to Nischarin or hPIP1 and dephosphorylation by the phosphatases PP2A and POPX1/2 (Xia et al., 2001; Koh et al., 2002; Kumar and Vadlamudi, 2002; Alahari et al., 2004). Phosphorylation of PAK by protein kinase A also inhibits PAK activation (Howe and Juliano, 2000). Further examination of matrix-specific PAK activation will be an interesting avenue for future work.
The current data suggest that reducing either PAK activation or localization to cell–cell junctions should reduce the permeability of the endothelial cell layer. Recently, the Ser/Thr kinases Akt and protein kinase G (PKG) were found to phosphorylate PAK at Ser21 within the Nck-binding sequence, inhibiting the interaction between PAK and Nck (Zhou et al., 2003; Fryer et al., 2006). Because blocking the PAK–Nck interaction inhibits localization of PAK to cell–cell borders and decreases endothelial permeability, these kinases might decrease permeability in a similar manner. Indeed, both Akt and cyclic GMP/PKG can decrease vascular permeability (Pearse et al., 2003; Chen et al., 2005; Moldobaeva et al., 2006). Whether PAK is the relevant target for these effects remains to be explored.
The mechanisms by which permeability is elevated in the plaque endothelium are not well understood. Dissolution of intercellular interactions during endothelial cell division and apoptosis, both of which are elevated at athero-prone sites in vivo (Weinbaum et al., 1985; Lin et al., 1988), has been suggested as a possible mechanism. However, the correlation between endothelial cell turnover and enhanced permeability in vivo is weak (Penn and Chisolm, 1991; Malinauskas et al., 1995). A more likely mechanism involves TJs in athero-prone regions, which are discontinuous compared with athero-resistant regions (Okano and Yoshida, 1994). Changes in TJ protein expression, phosphorylation, and reorganization could all contribute to decreased barrier function (Ogunrinade et al., 2002). Both flow and cytokines induce permeability too rapidly for changes in gene expression to be an attractive mechanism. Shear stress stimulates occludin phosphorylation on Ser/Thr residues, which could alter occludin localization to TJs or function (Sakakibara et al., 1997; DeMaio et al., 2001). VEGF stimulates PAK-dependent VE-cadherin phosphorylation, resulting in its arrestin- dependent internalization and the formation of paracellular pores (Gavard and Gutkind, 2006). Myosin light chain phosphorylation triggers cell contraction and the formation of paracellular pores (Stockton et al., 2004), and contractility appears to be a common pathway for endothelial cell permeability by multiple atherogenic stimuli (Takeya et al., 1993; Essler et al., 1999; Siess et al., 1999; Ogunrinade et al., 2002; Stamatovic et al., 2003). PAK inhibition decreases myosin phosphorylation and contractility in endothelial cells (Kiosses et al., 1999; Stockton et al., 2004). Thus, effects of PAK on the cytoskeleton appear to be involved in regulation of permeability, though other events, such as VE-cadherin and occludin phosphorylation, are likely to contribute.
PAK regulates cytoskeletal organization, proliferation, and movement in many cell types, making PAK activity by itself an unlikely target for long term therapy. For example, PAK inhibition in mice with a cell-permeable peptide was recently shown to mimic Alzheimer's disease (Zhao et al., 2006). However, specific interactions, such as Nck, may offer more attractive therapeutic targets. The ECM dependence of PAK activity may provide an especially attractive means for therapeutic intervention that would be less perturbing than global inhibition of kinase activity.
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Immunoblotting and immunocytochemistry
Cell lysis and immunoblotting were performed as previously described (Orr et al., 2002). Antibodies used include rabbit anti–phospho-PAK (Ser141; 1:5,000; Biosource International) and rabbit anti-PAK (1:1,000; Cell Signaling Technologies). For immunocytochemistry, cells were fixed with PBS containing 2% formaldehyde, permeabilized with 0.2% Triton X-100, and blocked for 1 h in PBS containing 1% BSA and 10% goat serum. Primary antibodies were incubated with cells in blocking buffer as follows: rabbit anti–phospho-PAK (Ser141; 1:500 overnight), rabbit anti–ß-catenin (1:200 overnight; Santa Cruz Biotechnology, Inc.), and mouse anti–ZO-1 (1:500 overnight). Cells were then incubated in 1 µg/ml Alexa 488– conjugated goat anti–rabbit IgG or goat anti–mouse IgG (Invitrogen). Slides were mounted with Fluoromount G, and images were taken using the 60x oil-immersion objective on a microscope (DiaPhot; Nikon) equipped with a video camera (CoolSnap; Photometrics) using the Inovision ISEE software program.
Permeability assays
A novel transwell well-flow device was developed to assay macromolecule permeability across an intact endothelial monolayer using previously established methods (Stockton et al., 2004). In brief, a previously developed cone-and-plate flow device was adapted to accept a 75-mm chamber transwell insert (Blackman et al., 2002). Custom flanges mounted on the lip of the Petri dish hold inlet and outlet tubing for the top and lower chambers, respectively, to inject and remove HRP without interrupting flow. Transwell chambers (3.0-µm pore size; Costar) were coated with either MG or FN, and BAE cells were allowed to attach for 4–24 h. Some transwells were coated with a fixed concentration of MG followed by increasing concentrations of FN. For flow experiments, cells on 75-mm chambers were serum deprived for 4 h in phenol red-free DME containing 0.5% CS and 2% dextran (wt/vol) and loaded onto the flow device stage, and shear stress was applied using the modified cone-and-plate device. At desired times, the medium was replaced with fresh medium containing 60 µg/ml HRP (Sigma-Aldrich) or Alexa 488–conjugated BSA (Invitrogen). After 1 h, medium was removed from the lower chamber, and cells were fixed in 2% formaldehyde and stained with Coomassie blue to detect cell loss or examined by immunocytochemistry for Ser141 phosphorylated PAK. For cytokine and LDL-induced permeability assays, cells grown on 6.5-mm filters were serum deprived for 4 h in phenol red–free DME containing 0.5% CS and transferred to fresh medium containing soluble factors for 90 min. HRP was then added to the top well to give a final concentration of 60 µg/ml. After 30 min, medium from the bottom well was removed, incubated with 0.5 mM guaiacol, 50 mM Na2HPO4, and 0.6 mM H2O2, and formation of O-phenylenediamine was determined by measure of absorbance at 470 nm. Alexa 488–conjugated BSA was measured using a spectrofluorometer (FluoroLog; Jobin Yvon). Results are shown as a fold increase in HRP activity or in absolute solute permeability. Solute permeability coefficients for the endothelial monolayer were calculated as Ps = 
CaVa/CtS, where Ca is the final concentration in the lower well, Va is the volume of the bottom well (ml), C is the concentration in the top well, t is the sampling interval (s), and S is the surface area of the transwell (cm2; Kajimura et al., 1997).
Animals and vessel harvest
Nine male ApoE-deficient mice on a C57Bl/6 background from The Jackson Laboratory, 8–12 wk of age and weighing 18–20 g, were used in these experiments. Four mice were fed a Western-type atherogenic diet (TD 88137 [Harlan-Teklad]; containing 21% fat by weight, 0.15% by weight cholesterol, and 19.5% by weight casein without sodium cholate) for 10 wk before sacrifice. Control mice were fed a chow diet during this time. At 20 wk of age (10 wk on diet), mice were perfused with 4% paraformaldehyde, and the aortic arch, left carotid sinus, and right carotid sinus were processed for paraffin embedding. For Evans blue assays, six male C57Bl/6 and nine male ApoE-deficient mice (The Jackson Laboratory) were maintained on chow diets for 8 or 32 wk, respectively.
Immunohistochemistry
5-µm paraffin sections were obtained for immunohistochemistry. Immunohistochemistry for adhesion molecules VCAM-1 (Santa Cruz Biotechnology, Inc.) was performed as previously described (McPherson et al., 2001). After microwave antigen retrieval with antigen unmasking solution (Vector Laboratories), rabbit anti-FN (1:400; Sigma-Aldrich) and rabbit anti-Ser141 phosphorylated PAK (1:250) were applied. Detection of antibodies was with Vetastain Elite kit (Vector Laboratories). Visualization was with diaminobenzidine (DakoCytomation). For en face staining, the aortic arch was cut into rings and stained for either PECAM-1 or Ser141 phosphorylated PAK using Alexa 488–conjugated goat anti–rabbit secondary antibodies to detect localization. Rings were then cut, opened, and mounted between two coverslips for en face viewing by fluorescence microscopy. Images were acquired using the 10x or 40x objective on a microscope (BX51; Olympus) equipped with a digital camera (DP70; Olympus) using ImagePro Plus software (Media Cybernetics).
Permeability to Evans blue in vivo
Mice were injected intraperitoneally with 0.1 ml of either control peptide or the PAK-Nck inhibitory peptide (10 mg/ml) at 24 h and at 1 h before Evans blue injection. Evans blue (0.1 ml of 1% dye in PBS) was injected into the tail vein. After 30 min, mice were killed with ketamine/xylazine and perfused through the left ventricle with 10 ml of 4% formaldehyde in PBS, and the aorta was excised from the cusp to the renal artery branches. Bright field microscopy of excised aortas was performed using the 0.5 and 1.2x objectives on a microscope (SZX12; Olympus) equipped with a DP70 digital camera using ImagePro Plus. Aortas were dried and weighed, Evans blue was extracted by incubation in formamide for 24 h at 60°C, and absorbance at 620 nm was determined. Concentration curves for pure Evans blue were used to calculate the total amount of dye extracted, and this value was normalized to the weight of the isolated aortas.
Online supplemental material
Independent of matrix composition, the 4-h plating time is sufficient to allow both adherens and TJ formation, as assessed by staining cells for ß-catenin and ZO-1, respectively (Fig. S1). Matrix-specific effects on monolayer permeability are not due to differences in matrix permeability, which shows no difference between MG and FN (Fig. S2). Localization to cell–cell junctions is required for PAK-dependent permeability (Stockton et al., 2004), and TNF, MCP-1, and oxLDL all stimulate active PAK localization to cell–cell junctions, which was abrogated by the addition of the PAK-Nck inhibitory peptide (Fig. S3). Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200609008/DC1.
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Acknowledgments |
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This work was supported by U.S. Public Health Service grant RO1 HL75092 to M.A. Schwartz, American Heart Association Mid-Atlantic Affiliate fellowship 0525589U to A.W. Orr, National Institutes of Health grant 1RO1HL66264 to I.J. Sarembock, and The Whitaker Foundation Biomedical Research grant RG-02-0853 to B.R. Blackman.
Submitted: 1 September 2006
Accepted: 23 January 2007
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