1Physiology Program, Department of Environmental Health, Harvard School of Public Health, Boston; 2Pulmonary and Critical Care Division, Department of Medicine/Tupper Research Institute, Tufts-New England Medical Center and Tufts University School of Medicine, Boston, Massachusetts; 3Institute of Biochemistry, Medical School Hannover, Hannover, Germany; and 4Division of Pulmonary and Critical Care Medicine, The Johns Hopkins University School of Medicine, Baltimore, Maryland
Submitted 30 August 2004 ; accepted in final form 21 April 2005
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ABSTRACT |
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endothelial barrier; cytoskeleton; actin dynamics; stiffness; tensile stress
We have recently shown in rat pulmonary microvascular endothelial cells (RPMEC) that hypoxia causes redistribution of the actin cytoskeleton (CSK) in a fashion that is dependent on p38 mitogen-activated protein kinase (MAPK) activation and leads to the phosphorylation of the small heat shock protein HSP27 (30, 31). HSP27 is a member of the stress-inducible small HSP family that is thought to act as a microfilament capping protein in vitro (41) and to be constitutively expressed at high levels in the lung (33). During cellular stress and growth, HSP27 undergoes rapid phosphorylation, which in turn promotes actin polymerization and stress fiber formation (8, 26, 36). However, it remains unclear to what extent these structural changes contribute to functional changes in biophysical properties of the endothelium at cellular and subcellular levels.
Herein we report changes in biophysical properties of the CSK of RPMEC in response to hypoxia, as well as the modulatory effects of HSP27. To quantify cytoskeletal remodeling events, we measured spontaneous nanoscale movements of an individual microbead that was coated with a peptide containing Arg-Gly-Asp (RGD). Such beads bind avidly to cell surface integrin receptors (58) and form focal adhesions (40); they become well integrated into the CSK scaffolding (4, 20, 58) and display tight functional coupling to stress-bearing cytoskeletal structures and the contractile apparatus (3, 27, 59). We reasoned that the bead can move only if microstructures to which it is attached rearrange; accordingly, spontaneous nanoscale motions report cytoskeletal rearrangements in space and time. To quantify cell stiffness, we used magnetic twisting cytometry (20, 58), and to quantify traction forces exerted by the cell on its substrate, we used Fourier transform traction microscopy (11, 59). We present evidence indicating the existence of differentially regulated biochemical and biophysical events in response to hypoxia. On the one hand, hypoxia causes early Rho-kinase-mediated myosin phosphorylation, leading to the development of contractile stress within the endothelial cell and, on the other hand, it causes belated p38-mediated HSP27 phosphorylation, leading to stabilization of the actin CSK at or near focal adhesions. These findings help to explain how hypoxia alters endothelial cell barrier function.
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MATERIALS AND METHODS |
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Cell culture and exposure to hypoxia. Rat pulmonary microvascular endothelial cells (RPMEC) were a gift from Dr. Una Ryan (Avant Immunotherapeutics, Needham, MA) and have been well characterized by us and others (15). These cells exhibit typical endothelial "cobblestone" morphology and stain positively with antibodies against von Willebrand factor. RPMEC were grown in culture as previously described (15). Unless otherwise specified, 1 day before the experiments, cells were harvested with 0.25% trypsin-0.02% ethylenediaminetetraacetic acid (EDTA) solution, plated on tissue culture petri dishes at subconfluence (9.6 cm2 growth area; Becton Dickinson), and maintained in serum-free media at 37°C in humidified air containing 5% CO2.
For hypoxic exposure, cells were placed in humidified, airtight incubation chambers (Billups-Rothenberg, Del Mar, CA) and gassed with 3% O2, 5% CO2, and balanced N2. The hypoxic chambers were kept at 37°C in a tissue culture incubator for the duration of 0.5, 1, or 4 h. For normoxic exposure (0.5, 1, or 4 h), cells were maintained at 37°C in humidified air containing 5% CO2.
Transfection of endothelial cells. Phospho-mimicking mutant human HSP27 (HSP27-PM) construct was generated as previously described (19, 48). This construct was made in the pcDNA3 vector (Invitrogen), in which the cytomegalovirus promoter drives the eukaryotic expression of the corresponding protein. The vectors used for transfection were pcDNA3 alone and pcDNA3-HSP27-PM, in which the residues S15D, S78D, S82D were mutated to mimic phosphorylated HSP27. The vectors were introduced (5 µg) into endothelial cells by electroporation. Stable cell lines were obtained by selection with geneticin, and resistant colonies were isolated, expanded, and then screened for the level of human HSP27 expression. Unless otherwise noted, cells in passages 6 and 7 were used that typically represented a pure population of stable cell lines.
Characterization of spontaneous bead motion.
The dynamics of the CSK network were measured as described previously (4). Using microscopic observation, we visualized spontaneous nanoscale movements of an individual RGD-coated microbead tightly anchored to the CSK of the endothelial cell (50100 beads/field of view) and recorded its positions every 83 ms. The trajectories of bead motions in two dimensions were then characterized by computing the mean square displacement of all beads as a function of time:
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Magnetic twisting cytometry with optical detection.
The CSK stiffness of each individual RPMEC was measured as described previously (20). Briefly, the RGD-coated ferrimagnetic microbeads bound to adherent cells were first magnetized horizontally (parallel to the surface on which cells were plated) with a brief 1,000-Gauss pulse and then twisted in a vertically aligned homogeneous magnetic field (40 Gauss) at a frequency of 0.75 Hz. This sinusoidal twisting magnetic field was too weak to remagnetize the beads and instead caused both a rotation and a pivoting displacement of the beads. As the beads move laterally to and fro, therefore, the cell resists their motion by developing internal stresses that depend on the mechanical properties of the cell (20, 27). Accordingly, lateral bead displacements in response to the resulting oscillatory torque were detected optically (in spatial resolution of 10 nm), and the ratio of specific torque to lateral bead displacements was computed and expressed as the cell stiffness in units of Pa/nm. Cell stiffness was measured both before and after each exposure (0.5, 1, or 4 h) to hypoxia or normoxia.
Traction microscopy. A detailed description of this technique was provided by Butler and colleagues (11, 59). Using traction microscopy, we measured the distribution of contractile stresses arising at the interface between each adherent cell and its substrate (traction field). In brief, RPMEC were plated sparsely on a polyacrylamide elastic gel block coated with collagen type I (0.2 mg/ml) and allowed to spread and stabilize for 6 h. Cells were then exposed for 0.5 h with either normoxia or hypoxia. After each normoxic or hypoxic exposure, images of fluorescent microbeads (0.2 µm in diameter; Molecular Probes, Eugene, OR) embedded near the gel apical surface were taken both before and after the cell was completely detached from the substrate by trypsin. The fluorescent image of the same region of the gel after trypsin was used as the reference (traction free) image. The displacement field between a pair of images was then obtained by identifying the coordinates of the peak of the cross-correlation function (11, 59).
From the displacement field and known elastic properties of the gel (Young's modulus of the gel was determined to be 1,300 Pa, and a Poisson ratio was taken to be 0.48), the traction field was calculated using both constrained and unconstrained Fourier transform traction cytometry (FTTC) as described previously (11, 59). The computed traction field was then used to obtain 1) the contractile stress (prestress), defined as the net tensile force transmitted by the actin CSK across a cross-sectional area of the cell per unit area; and 2) the net contractile moment, defined as a scalar measure of the cell's contractile "strength" (11, 59). In this study, prestress is expressed in pascals (Pa) and the net contractile moment is expressed in piconewton meters (pNm).
Statistical analysis. Data are presented as means ± SE, and n represents the number of cells. Statistical differences were determined using either Student's t-test for comparison of two sample means or ANOVA for comparison of more than two sample means, followed by Bonferroni post hoc testing for multiple comparisons between two sample means. P < 0.05 was considered statistically significant.
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RESULTS |
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Hypoxia tethers spontaneous bead motions via the activation of p38 MAPK. The formation of F-actin in response to hypoxia is thought to be mediated via the activation of p38 MAPK (30, 31). Once activated, p38 phosphorylates and activates its downstream effector MAP kinase-activated protein kinase 2/3 (MK2), which in turn phosphorylates HSP27 and, in doing so, prevents it from binding to actin monomers and causes F-actin polymerization (26, 49). To assess the role of p38 and its downstream signals on spontaneous bead motions, RPMEC were treated for 0.5 h with or without a p38 MAPK inhibitor SB-203580 before 1-h exposure to hypoxia. Under baseline conditions, cells treated with 3 µM SB-203580 exhibited an increase in bead motions as evaluated by MSD at 300 s (Fig. 3A) and, more important, displayed no further decrease in bead motions in response to hypoxia (1 h). Cells treated with a Rho-kinase inhibitor (Y-27632; 3 µM) also exhibited an increase in bead motions and showed no appreciable decrease in bead motions in response to hypoxia. Such hypoxic exposure in the case of untreated cells caused a significant decrease in bead motions (Fig. 3A).
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Hypoxia causes dynamic changes in cell stiffness. We used forced nanoscale motions of those same RGD-coated microbeads firmly attached to the CSK of the RPMEC (see Magnetic twisting cytometry with optical detection) to measure changes in cell stiffness in response to hypoxia vs. normoxia (0.5, 1, or 4 h). Exposure of RPMEC to hypoxia caused time-dependent changes in cell stiffness (Fig. 4). Cells exposed to hypoxia for 0.5 h exhibited a significant increase (P < 0.05) in cell stiffness from baseline control (from 1.09 ± 0.03 to 1.37 ± 0.05 Pa/nm; means ± SE), whereas cells exposed to normoxia for the same duration showed no appreciable increase in cell stiffness (1.08 ± 0.08 Pa/nm). By contrast, cells exposed to hypoxia for 1 or 4 h did not show appreciable changes in cell stiffness compared with their respective normoxic controls (Fig. 4). Thus hypoxia caused a rapid (0.5 h) but transient increase in endothelial cell stiffness.
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Hypoxia causes increases in cell traction, tensile stress, and contractile moment. Using traction microscopy, we next quantified the level of tensile stress within individual RPMEC (prestress), their contractile moments (contractile "strength"), and their changes in response to hypoxia. Figure 7 shows phase-contrast images of RPMEC (normoxic and hypoxic) cultured on a flexible polyacrylamide gel and their respective traction fields computed from the corresponding bead displacement fields using constrained FTTC (see MATERIALS AND METHODS). The arrows in Fig. 7 show the relative magnitudes and directions of the tractions, and the colors show the absolute magnitude of the traction vector. The greatest traction, in general, occurred at the cell periphery and was directed centripetally; the root mean square traction averaged across the entire cell-projected area increased appreciably with hypoxic exposure. Accordingly, compared with cells exposed to normoxia (1,274 ± 204 Pa), cells exposed to hypoxia (0.5 h) developed significantly greater mean prestress levels (2,077 ± 234 Pa) (Fig. 8A). Nonetheless, to avoid any systematic errors associated with the estimation of the prestress, we separately computed net contractile moment (a scalar measure of the contraction intensity) directly from the traction field without invoking assumptions concerning the cell cross-sectional area (11, 59). Compared with cells exposed to normoxia (5.49 ± 0.99 pNm), cells exposed to hypoxia (10.20 ± 2.23 pNm) generated significantly higher net contractile moment (Fig. 8B). These findings provide, for the first time, direct mechanical evidence that hypoxia causes development of contractile stresses in the endothelial cell.
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DISCUSSION |
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Current thinking on endothelial cell barrier function holds that vascular permeability occurs mainly through paracellular pathways and is tightly regulated by a balance of physical forces (18, 23). On the one hand, myosin-based contractile forces are developed within the endothelial cell, and, on the other, those internal forces must be taken up at the cell boundary by adhesive tethering forces at cell-cell contacts and cellular-extracellular matrix (ECM) interactions (9, 14, 38, 43, 55, 62). In the endothelial cell, myosin exerts its mechanical effects by interacting with actin within an integrated scaffolding comprising scores of cytoskeletal and signaling molecules (18, 24, 25, 58). It is now known that this scaffolding is dynamic and in a continuous state of remodeling; the actin lattice, the myosin filament, and the focal adhesion complex are all considered to be evanescent structures that can be virtually demolished in some circumstances and then reconfigured and stabilized in others (4, 7). Accordingly, disruption of these molecular events may modulate the balance of physical forces that regulate endothelial cell barrier function and thereby contribute to the formation of intercellular gaps and altered endothelial cell permeability. We begin the DISCUSSION by addressing methodological issues and then elaborate on the findings of this report and their implications.
Spontaneous bead motions, CSK remodeling dynamics, and cellular-ECM tethering interactions. The anomalous bead motions that we observed are inconsistent with simple Brownian motion. We suggest instead that they track rearrangements of underlying structures to which they are firmly attached. This is in keeping with the reports of others, but with some notable differences. A microbead coated with fibronectin, when placed briefly on the surface of lamellipodia of migrating fibroblasts, moves a large distance (>4 µm) and preferentially rearward toward the nucleus (13, 51). Such directed motions are thought to be driven by the rearward flow of cortical actin CSK to which the bead is attached (16, 17, 51). In our experiments with RGD-coated microbeads on the surface of the endothelial cell, we observed no directed motions. Unlike motions of beads on locomoting fibroblasts, spontaneous motions of beads on the RPMEC were random and were smaller by 100- to 1,000-fold; they were nanoscalar, spanned only a small fraction of the bead diameter, and were certainly much less than the lateral extent of the cell boundaries. A number of factors could account for these differences.
First, the RPMEC was not visibly motile; cells seemed to be firmly adherent to the substrate and displayed no characteristic polar (asymmetric) morphology that is commonly observed in locomoting cells (13, 51). Experiments conducted on micropatterned substrates on which the cell could adhere but not crawl (45) showed similar bead motions (data not shown). Second, whereas actin filaments in motile cells are mainly distributed as a network in the cellular cortex (17), actin filaments of RPMEC are bundled with other proteins into contractile stress fibers (18, 23, 31, 55). The strong attachment of the bead to these stress-bearing cytoskeletal structures by RGD-integrin binding is well established (3, 27, 59), and the development of focal adhesions and complex interconnections to the actin CSK are readily observed at the bead locations (20, 40, 58). Taken together, therefore, the differences in the nature of bead behavior on migrating fibroblasts vs. rather stationary RPMECs are likely attributable to differences in cell biophysics as well as in the mode of bead anchorage to underlying cytoskeletal structures.
Exposure of endothelial cells to hypoxia caused time-dependent changes in spontaneous bead motions, whereas exposure to normoxia caused no appreciable changes in bead motions for the indicated times (Fig. 2). Interestingly, changes in bead motions in response to hypoxia recapitulated remarkably in time structural changes in the actin CSK that were reported previously by us (31). In that study, we observed the greatest increase in F-actin polymerization after a 1-h exposure to hypoxia, which corresponds closely to the greatest decrease in spontaneous bead motions (Fig. 2B). Conversely, spontaneous bead motions increase dramatically when the actin CSK is disrupted by cytochalasin D or latrunculin A (4). Consistent with these observations, cells treated with Rho-kinase inhibitor Y-27632 exhibited an increase in spontaneous bead motions that persisted even after a 1-h exposure to hypoxia (Fig. 3A): Rho-kinase inhibition has been reported to disrupt the actin CSK (1, 39). Taken together, these findings imply that the more polymerized the network of F-actin, the more tethered are the beads to the cytoskeletal scaffolding and the less they move. We suggest that the decrease in spontaneous bead motions in response to hypoxia is probably due to stabilization of the actin CSK, perhaps reflecting an increase in the cellular-ECM tethering forces at or near focal adhesions.
In that connection, numerous investigators have inferred that the small heat shock protein HSP27 is likely to play an important role in stabilization of the actin CSK (26, 37), but these findings have been based exclusively on structural evidence. In rat pulmonary microvascular endothelial cells, we recently demonstrated that hypoxia reversibly triggers p38-mediated activation of MK2 that in turn results in HSP27 phosphorylation and F-actin formation (30, 31). Indeed, compared with cells stimulated with hypoxia, endothelial cells overexpressing constitutively active MK2 or HSP27-PM exhibit similar increases in F-actin formation (31). By contrast, cells overexpressing dominant-negative MK2 not only show reduced F-actin content but also exhibit no further increases in F-actin formation in response to hypoxia (31). Consistent with this structural evidence, the decrease in spontaneous bead motions in response to hypoxia (i.e., beads tethering to, and perhaps being incorporated into, the newly formed cytoskeletal scaffolding at the site of focal adhesions) was attributable largely to the activation of p38 MAPK, leading to the phosphorylation of HSP27 (Fig. 3). Under baseline conditions, cells treated with a p38 MAPK inhibitor SB-203580 exhibited the expected increase in spontaneous bead motions and, most important, displayed no decrease in bead motions in response to hypoxia (Fig. 3A). By contrast, cells overexpressing HSP27-PM demonstrated a significant decrease in bead motions compared with cells transfected with the empty vector alone (Fig. 3B). These findings provide direct functional evidence that HSP27 phosphorylation indeed stabilizes the actin CSK, thus confirming inferences of others that were based solely on structural findings (26, 31, 37).
At this time, however, we are not certain how these changes in spontaneous bead motions relate to cadherin-mediated cell-cell adhesions. We speculate that local remodeling and stabilization at sites of cellular-ECM adhesions may be accompanied by local destabilization at the cell periphery that tethers cell-cell adhesions (9, 14). In support of this idea, studies have shown that hypoxia causes disassembly of the peripheral actin bands delineating the margins of the cells and, at the same time, causes assembly of stress fibers within the cell body and subsequently the appearance of intercellular gaps (31, 46). These findings have a particular importance to the understanding of endothelial cell barrier function, because the adhesive tethering forces of cell-cell and cellular-ECM adhesions are intimately linked to the actin-based CSK (18, 23, 55), which defines not only endothelial cell morphology (58) but also its ability to generate contractile force (18, 25, 63).
Cell stiffness, tensile stress, and the brisk biomechanical response of the endothelial cell. Direct quantitative indices of the physical forces within the endothelial cell have been difficult to measure (10, 25, 63). As a result, studies have often relied on the structural changes in the F-actin polymerization and stress fiber formation as the surrogates for changes in the contractile behavior. In the airway smooth muscle, we have demonstrated that agonist-evoked cell stiffening is attributable mostly to myosin activation, whereas polymerization of the actin lattice is necessary but not sufficient to account for the observed increase in cell stiffness (3). Similarly, in the RPMEC, the time course of hypoxia-induced changes in cell stiffness (Fig. 4), as well as tensile stresses developed within the CSK of individual endothelial cells (Fig. 8), did not correspond to the temporal dynamics of spontaneous bead motions (Fig. 2) or to the reported time course of structural changes in the actin CSK (31). Indeed, both cell stiffness and tensile stress quickly increased after a 0.5-h exposure to hypoxia, at which time we found no qualitative changes in spontaneous bead motions or quantitative changes in F-actin content (31). More interestingly, the increase in cell stiffness in response to hypoxia was not blocked by SB-203580 or observed in cells overexpressing HSP27-PM (Fig. 5). These findings provide, for the first time, direct, quantitative, mechanical evidence demonstrating the contractile status, the amount of physical force within the endothelial cell, and their changes in response to hypoxia. Moreover, they draw a consistent picture in which hypoxia-activated biochemical events leading to F-actin polymerization (i.e., p38-mediated HSP27 phosphorylation) are not sufficient to account for the increase in the contractile state of RPMEC.
Regarding the regulation of contractile force in the endothelium, phosphorylation of the 20-kDa MLC plays an important role (22, 28). When phosphorylated by Ca2+/calmodulin-dependent MLCK (52), MLC promotes actomyosin motor activity and tension development within the endothelial cell (25, 34, 56). We have observed that hypoxia increases phosphorylation of MLC in pulmonary endothelial cells (data not shown), an effect that recently has been described in pulmonary arterial smooth muscle cells (60, 61). Indeed, several models of agonist-induced barrier dysfunction, such as the effects of thrombin or histamine, have implicated MLC phosphorylation for generation of contractile force and endothelial cell retraction (10, 25, 52, 56, 63). In human umbilical vein endothelial cells, Sheldon et al. (52) demonstrated that an inhibition of MLCK with ML-9 not only reduced basal MLC phosphorylation but also prevented histamine-induced increase in MLC phosphorylation and subsequently endothelial cell retraction. Consistent with their findings, we found that ML-7 decreased baseline stiffness of RPMEC; ML-7 is a derivative of ML-9 that is thought to be an equally selective and potent inhibitor of MLCK (6, 50). Nevertheless, treatments of cells with ML-7 were not sufficient to prevent the increase in cell stiffness in response to hypoxia (Fig. 6).
More recently, studies have emphasized the increasing importance of Rho/Rho-kinase signaling pathways for the regulation of endothelial cell barrier function (12, 24). In particular, activation of Rho and its downstream effector ROCK have been shown to phosphorylate and inhibit MLC phosphatase, which subsequently leads to increases in MLC phosphorylation, actomyosin interactions, focal adhesion, and stress fiber formation (2, 32, 47, 57). In this connection, Katoh et al. (29) reported two different classes of stress fibers in cultured FS-133 cells, which are differentially regulated by MLCK and ROCK. Whereas peripheral stress fibers are dependent on the activity of MLCK, central stress fibers are dependent on the activity of ROCK (29). In contrast to the effects of ML-7, we found that cells treated with a ROCK inhibitor Y-27632 exhibited not only a decrease in baseline cell stiffness but also a marked reduction in hypoxia-induced increase in cell stiffness. These findings are in agreement with the notion that MLC phosphorylation may be differentially regulated by hypoxia (29, 42, 57, 60, 61) and also extend that idea by providing direct mechanical evidence. Currently, we are in the midst of localizing the temporospatial evolution of myosin phosphorylation, physical forces exerted by the endothelial cell, and their contributions to endothelial cell permeability in response to hypoxia. Taken together, our findings demonstrate that the major part of hypoxia-induced contractile response seems to be attributable largely to actomyosin motor activation through the Rho/Rho-kinase signaling pathway, whereas hypoxia-induced stabilization of the actin CSK at or near focal adhesions requires p38-mediated HSP27 phosphorylation.
Regulation of pulmonary vascular permeability is a complex process involving several mechanisms. The current model of endothelial cell barrier function holds that vascular permeability is tightly regulated through a balance between contractile forces within the endothelial cell and adhesive forces that connect endothelial cells to each other or to the ECM. Using a variety of novel tools in combination with various pharmacological and genetic manipulations, we have shown that hypoxia differentially regulates early contractile events within the endothelial cell that are followed in time by local changes in cellular-ECM adhesive interactions. The appreciable increases in cell stiffness and tensile stress in response to hypoxia were largely attributable to Rho-mediated activation of actomyosin motors. Spontaneous nanoscale motions of microbeads anchored to the CSK were superdiffusive, and activation of p38 leading to the phosphorylation of HSP27 was found to tether bead motions, thereby providing direct functional evidence that activation of HSP27 indeed stabilizes the actin CSK at or near focal adhesions. Taken together, these findings provide direct mechanical evidence that hypoxia alters a balance of physical forces that regulate endothelial cell barrier function.
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FOOTNOTES |
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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.
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