doi: 10.1161/01.RES.0000060201.41923.88
© 2003 American Heart Association, Inc.
Reviews |
Spatial Microstimuli in Endothelial Mechanosignaling
From the Institute for Medicine and Engineering (P.F.D., J.Z.), Department of Pathology and Laboratory Medicine (P.F.D.), University of Pennsylvania, Philadelphia, Pa; Department of Biomedical Engineering (B.P.H.), University of Virginia, Charlottesville, Va.
Correspondence to Dr P.F. Davies, Institute for Medicine and Engineering, University of Pennsylvania, 1010 Vagelos Laboratories, 3340 Smith Walk, Philadelphia, PA 19104. E-mail pfd{at}pobox.upenn.edu
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Abstract |
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 Top Abstract IntroductionLength ScalesWhere Are the Hemodynamic...Microstimuli at the Luminal...Four-Dimensional Cytoskeletal...Flow-Mediated Chemical SignalingFuture DirectionsReferences |
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Descriptive and quantitative analyses of microstimuli in living endothelial cells strongly support an integrated mechanism of mechanotransduction regulated by the spatial organization of multiple structural and signaling networks. Endothelial responses to blood flow are regulated at multiple levels of organization extending over scales from vascular beds to single cells, subcellular structures, and individual molecules. Microstimuli at the cellular and subcellular levels exhibit temporal and spatial complexities that are increasingly accessible to measurement. We address the cell and subcellular physical interface between flow-related forces and biomechanical responses of the endothelial cell. Live cell imaging and computational analyses of structural dynamics, two important approaches to microstimulation at this scale, are briefly reviewed.
Key Words: mechanotransduction • endothelial cells • shear stress • cytoskeletal dynamics
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Introduction |
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TopAbstractIntroductionLength ScalesWhere Are the Hemodynamic...Microstimuli at the Luminal...Four-Dimensional Cytoskeletal...Flow-Mediated Chemical SignalingFuture DirectionsReferences |
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The endothelia of different vascular beds express specialized morphologies and functions appropriate to their location; the local biomechanical environment is an important component of this. A feature common to all endothelial cells in a variety of vascular locations is the ability to respond to local changes in blood flow, particularly acute and sustained changes of hemodynamic shear stresses that act at the luminal cell surface.1–3 Of particular interest in mechanotransduction studies is the endothelium of the arterial side of the systemic circulation where many important vasoregulatory mechanisms are located. Arterial endothelium is generally subjected to the greatest magnitudes and variations in hemodynamic forces. Acute dilatation and constriction of arteries in response to changes of flow are controlled by the endothelium through regulatory nitrovasodilators, prostaglandins, lipoxygenases, hyperpolarizing factors, and related molecules.4–8 Chronic changes in flow characteristics stimulate structural remodeling of the artery wall through a process that is also endothelium-dependent.9 Furthermore, most vascular pathologies originate and develop their morbidity in the arterial circulation where hemodynamics plays a critical role in the focal initiation and development of vascular dysfunction and atherosclerosis.10,11
Investigations of arterial hemodynamics are complicated by flow pulsatility, wall compliance, and the geometry of arterial branches, bifurcations and curvatures through which a high-pressure, high-flow, non-Newtonian fluid is propelled. Curves and branching geometries cause flow separations, rapid vortex formation-dissolution throughout the cardiac cycle, and complex spatial and temporal gradients operating over short distances. This leads to regional heterogeneity of endothelial exposure to flow forces within the same vascular bed12 as well as between structurally diverse locations within the arterial tree, eg, elastic and muscular arteries, arterioles.13 The key findings of several previous reviews of mechanotransduction mechanisms3,14,15 are briefly restated; however, our intent in this review is to outline recent and ongoing work on vascular cell and subcellular microstimulation of cell structures of biomechanical importance.
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Length Scales |
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TopAbstractIntroductionLength ScalesWhere Are the Hemodynamic...Microstimuli at the Luminal...Four-Dimensional Cytoskeletal...Flow-Mediated Chemical SignalingFuture DirectionsReferences |
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In considering the mechanisms that regulate flow-mediated endothelial mechanotransduction, the nature of the stimuli must be carefully considered. The physical deformation of cellular elements initiates the biological responses, yet the mechanisms by which forces such as shear stresses at the luminal surface of endothelium are converted to specific cellular responses are unclear. Until fairly recently, the macro (or bulk) fluid dynamics was considered without reference to cell and subcellular length scales. In addition, studies of the role of the endothelium in arterial wall remodeling associated with basement membrane stretch have considered wall tension as a stimulus only at the length scale of vessel diameter or wall thickness. However, microstimulation of the cell monolayer observed by optical and scanning probe imaging techniques is revealing in living cells the spatial dynamics of structures directly involved in mechanotransduction.
Endothelial mechanotransduction ranges over a breadth of scales, from entire systemic and pulmonary circulatory systems, comparative vascular beds, regional differences in a single vascular bed, groups of cells within a region, discrete single cells, and spatially defined subcellular locations (Figure 1).16 The integration of microscale investigations into higher-order organizational levels is a major challenge. For example, large focal differences found from cell to cell may become less significant when averaged over several thousand cells in a particular region. In other cases, a few uniquely different endothelial phenotypes within a region may profoundly influence focal vascular physiology, eg, in the localization of atherosclerosis.17,18 In such cases, an understanding of the biomechanical microstimuli of just a few cells may be important.
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Where Are the Hemodynamic Microstimuli? |
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TopAbstractIntroductionLength ScalesWhere Are the Hemodynamic...Microstimuli at the Luminal...Four-Dimensional Cytoskeletal...Flow-Mediated Chemical SignalingFuture DirectionsReferences |
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Endothelial cells respond to shear stress by initiating a cascade of intracellular events that begins with the generation of second messengers, quickly followed by transcriptional, synthetic, and structural changes (see reviews3,19). The temporal relationships are complex, in part because the cellular structural geometry (eg, surface topography, cytoskeletal arrangement) can be quickly rearranged as a consequence of downstream signaling and this may lead to repetitive initial signaling through the same or different pathways. Furthermore, it is unclear what filtering capability the cell has for responding to changes in biomechanical forces; some selectivity would seem to be essential in a rapidly changing hemodynamic environment. Stimuli may also be "remembered," further complicating interpretation of the temporal hierarchy of responses. Nevertheless, an estimation can be obtained from the timed measurement of major responses3,19 and upstream and downstream relationships established within specific pathways.15 For temporal information, the reader is referred to those publications.
Initiation of mechanosignaling has been measured within seconds of exposure to flow. Rapid responses include potassium channel activation,20 intracellular calcium release,21 G protein activation,22,23 and stimulation of protein kinases.24,25 Shearing forces may directly deform the cell surface to generate local biochemical responses arising from undefined sensor proteins26 and/or deformation of the lipid bilayer.27 A more extended model has been proposed that does not limit signal transduction exclusively to the luminal surface. In this decentralized model,3,19,28 forces acting on the cell surface are also transmitted by the cytoskeleton to other intracellular locations where signaling can occur. Candidate sites where integral membrane proteins are connected to the cytoskeleton include focal adhesions, intercellular junctions, and the nuclear membrane3 as well as specialized lipid microdomains such as cell surface caveolae. The model predicts mechanotransduction as an integrated response of multiple signaling networks that are spatially organized in the endothelial cell (Figure 1, bottom panel). Evidence in support of decentralized endothelial mechanotransduction includes the flow-induced remodeling of focal adhesion sites29 and activation of proteins localized at adhesion sites30 as well as a well-developed literature in cytoskeleton mechanics.31,32 Cytoskeleton-dependent transfer of forces across the endothelial cell surface via transmembrane proteins31 and the demonstration of mechanical continuity between the cell plasma membrane and nuclear structures33 are consistent with force transmission through cytoskeletal tensional networks. There is now considerable evidence not only for the reorganization of the cytoskeleton as cells adapt to shear stress2,34 but also for the rapid displacement and deformation of cytoskeletal filaments that suggest direct transmission of forces by the cytoskeleton in response to a change of flow28,35–40 (and see below).
Endothelial cells exist in a state of tension associated with the maintenance of cell shape. Tension is generated through anchorage of the cytoskeleton at multiple locations throughout the cell, particularly at sites of connection to focal adhesions,41,42 nucleus,43 and neighboring cells.44,45 When external forces are loaded onto the cell during flow, the internal cellular tension changes to equalize the external force. The structural deformation associated with shear stress–induced mechanotransduction in endothelial cells is induced by (1) local displacement of sensors at the cell surface, (2) transmission via cytoskeletal elements to distribute the force throughout the cell, which (3) concentrate the strain at other sites of cytoskeletal attachment. The physical effect is to deform or change the conformation of structural elements of the cell. Overall, the transmitted mechanical stress is converted to mechanotransduction responses at sites both local to, and remote from, the initial physical stimulus, most likely as a combined integrated cellular response. Thus, an important component of the decentralized model is that hemodynamic forces acting at the cell surface are distributed to other sites in the cell through microscale deformation/displacement of surface elements and cytoskeletal filaments.
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Microstimuli at the Luminal Endothelial Surface |
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TopAbstractIntroductionLength ScalesWhere Are the Hemodynamic...Microstimuli at the Luminal...Four-Dimensional Cytoskeletal...Flow-Mediated Chemical SignalingFuture DirectionsReferences |
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Cell Surface Topography
Hemodynamic forces act most directly on the cell surface. Cell geometry, particularly surface topography, influences the magnitude and localization of shear stresses and consequently affects the transmission of such forces across the plasma membrane and throughout the cell. Knowledge of the bulk flow characteristics in arteries provides a macroscopic profile of flow and the predicted shear stresses acting near the endothelial surface, but it is the detailed spatial characteristics at the fluid-cell interface that more precisely influence signaling. The microscale geometry of the cell surface is therefore important. The first theoretical estimates of magnitude and spatial characteristics of endothelial cells at a subcellular level were made by Satcher et al,46 by modeling the endothelial monolayer as a wavy surface and using computational methods to estimate the influence of the waviness on local flow forces. The geometries of living endothelial cells in tissue culture were approximated by contour imaging using differential interference contrast microscopy47 and by fluorescence exclusion combined with confocal microscopy.48 However, the first high-resolution images of the continuous geometry of the luminal surface of living endothelial cells in real-time were made by atomic force microscopy (AFM).49,50 Significant differences of surface topography from cell to cell and over short distances within the same cell were revealed by AFM. Finite element analyses were then conducted to simulate flows over the endothelial surface geometries defined by AFM of living endothelial cells and thus determine the shear stress distribution. Flow near the cell surface can be considered quasi-steady, meaning that for time-varying macroscopic shear rates, the flow at the surface could be determined as if it were steady flow at any instantaneous macro shear rate.50
Cellular and Subcellular Shear Stress Distribution
Flow perturbations due to the undulating surface produced cell-scale variations of shear stress magnitude and consequently large subcellular shear stress gradients. The surface topography of cells aligned by flow was observed to change when compared with no-flow controls; there was significant streamlining of the endothelial monolayer when the cells aligned. This reorganization of the endothelial surface in response to prolonged exposure to steady flow resulted in significant reductions in the peak shear stresses and shear stress gradients compared with no-flow control cells. The studies demonstrated that there are microscopic departures from a flat boundary due to the presence of the endothelial cells that, in turn, create a localized perturbation of the macroscopic flow field. Topographical differences were noted from cell to cell that resulted in differential stress distributions associated with individual endothelial cells,50 suggesting that different endothelial phenotypes may exist even within a small area, each determined by the interplay of hemodynamic microstimulation and surface geometry.18,51,52
Since the flow perturbation due to the wavy surface propagates only a short distance into the bulk flow, the surface shear stress distribution is dictated by the prevailing macroscopic shear rate. However, from the perspective of the endothelial cell, the shear stress at the endothelial luminal surface will vary within the macroscopic flow field as a function of the microscopic surface geometry. Thus, small differences in cell topography within the monolayer result in substantial differences in average and peak shear stresses affecting neighboring cells. One cell may be well above a response threshold while a neighboring cell is unaffected by the same macro-scale flow. In aligned cells, there was an approximate 40% decrease in the average shear stress gradients compared with no-flow cells.50 An analysis of the upstream/downstream symmetry of the cell surface in vitro revealed no preferential gradient. Simulations of flow in the "reverse" direction revealed little differences in the distribution of shear stress magnitude, which was approximately proportional to cell height. Cells were often asymmetrical with regard to the upstream and downstream slopes, but without bias in the flow direction after preexposure to flow; however, the shear stress gradient was always concentrated on the leading (upstream) slope. Butler et al53 have addressed one aspect of this issue functionally by measuring the plasma membrane fluidity of endothelial cells in relation to the flow direction by using a fluorescent lipophilic probe. A strip of membrane on the upstream or downstream slopes of the cell was photobleached, and fluorescence recovery was then measured to determine the diffusion coefficient of the fluorophore. Transient (<10 seconds) upstream increases and downstream decreases of fluidity were reported followed by a secondary increase that was limited to the upstream membrane and that peaked at 7 minutes. Values returned to baseline when flow was stopped. These data demonstrate both temporal and spatial variations in shear-induced membrane lipid fluidity. The studies also support the importance of the cell surface topography to the magnitude of fluidity changes. Barbee et al50 computed symmetrical shear stresses on the upstream and downstream portions of the cell but noted the concentration of gradients of shear stress onto the upstream face. It has been proposed that membrane fluidity is altered when the front face of the cell is unequally stretched.53 The greater displacement of cytoskeletal filaments near the top of the cell than the base36 in flow is also consistent with this. The evidence cited above demonstrates that the detailed cell surface topography is a critical contributor to the stress response and, together with the resulting spatially defined microstimuli, must be considered together with the macro hemodynamics when evaluating the fluid-tissue interface. These in vitro measurements and conclusions are also likely to be valid in vivo; AFM images of endothelial cells in situ at the artery wall showed a morphology and geometry very similar to that of cultured cells.19
Surface Deformation Involves More Than the Plasma Membrane
One of the most striking consequences of endothelial realignment by flow is increased rigidity of the cell surface as measured by decreased membrane deformability on micropipette aspiration of the cell surface after exposure to shear stress.54,55 This arises less from the fluidity of the plasma membrane itself than from the cortical cytoskeleton lying immediately beneath it and to which it is attached via membrane proteins. By AFM, longitudinal ridges apparently caused by the presence of cytoskeletal structures underlying the plasma membrane were revealed by the force of the AFM stylus.50 Further experiments using lower spring-constant cantilevers (thus reducing the imaging forces) showed these features to be proportional to the stylus force applied. However, such features were largely absent in nonaligned endothelial cells, and it can be concluded that bundles of cytoskeletal filaments form realigned arrays just below the luminal plasma membrane after exposure to flow,49 resulting in decreased deformability of the surface region. Also contributing to cell surface rigidity are ankyrin-like and spectrin-like proteins that serve as anchors at the cell membrane for other cytoskeletal proteins. It is not known if these rearrange in response to flow forces, although they do so in response to direct micropipette aspiration in other cells.56 The physical continuity of the cell surface and cytoskeleton is an important subcellular component of decentralized force transmission that leads to the redistribution of intracellular tension when the exogenous forces change (see below).
Amplification of Mechanical Stimuli by Leukocyte Adhesion
An intriguing aspect of the endothelial microhemodynamic interface that is directly relevant to vascular pathophysiology is the recruitment of leukocytes from the blood. In the larger arteries, monocyte adhesion, spreading, and migration across the endothelium occur early in atherogenesis where intimal monocyte-derived macrophages are one of the most prominent characteristics of lesion development.10 Monocyte adhesion and transmigration occur at sites of complex hemodynamics where there are steep cyclical gradients of macroscopic and microscopic shear stress.57 In recent work, the adherent monocyte-endothelial topography was imaged by AFM, and perturbations of the localized shear stress distribution on the endothelium were estimated after finite element modeling and simulated flow computations (J.Z., P.F.D., unpublished data, 2002). As shown in Figure 2, the presence of an adherent monocytic cell creates a hot spot of increased shear stress at the surface of the monocyte. Whereas the peak shear stress levels over the endothelium surrounding the adhesion event were similar to those in control monolayers (no monocytes), the peak shear stresses increased by an average of 50% to 60% at the surface of adherent monocytes. As the monocytes spread, the differential declined. In addition to estimating forces on the monocyte itself, the studies show that there is a transient period of greatly increased hemodynamic strain acting focally on the endothelium to which monocytes attach; this may also be transmitted to adjacent endothelial cells creating a local zone of altered vascular mechanics during the period of leukocyte attachment and extravasation. The degree to which such transient changes influence, or are influenced by, the contemporaneous chemical signaling between leukocyte and endothelium is unknown.
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Four-Dimensional Cytoskeletal Motion in Living Cells Under External Fluid Mechanical Forces |
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GFP-Intermediate Filament Imaging
By using conventional antibody staining in fixed, permeabilized cells, cytoskeleton remodeling has been observed after exposure to shear stress.2,34,58 However, to measure the dynamics of filament deformation with high temporal resolution necessitates the use of live cells. Transfection of cells with gene constructs of green fluorescent protein fused to cytoskeletal proteins (GFP and other fluorophores) has made it possible to observe spatiotemporal dynamics directly in living cells.35,39,59,60 We have used a GFP-vimentin fusion protein expressed in endothelial cells to evaluate the dynamics of the intermediate filament (IF) cytoskeleton during a step change in hemodynamic shear stress. Time-lapse optical sectioning and deconvolution microscopy produced 3-dimensional analyses of fluorescent IF movement in 90-second intervals.35 After transfection of confluent endothelial cells in vitro, maximum-intensity volume projections were constructed from stacks of deconvolved optical sections acquired at 0.1- to 0.5-µm intervals (Figure 3). The images were consistent with immunocytochemically defined IF distribution in endothelium and other mammalian cells.61 A prominent perinuclear ring was visible, with IFs extending above and below the nucleus. A mesh network of IFs radiated throughout the cytoplasm from the perinuclear ring, and thick circumferential IF bundles were sometimes present near the cell edge or reaching into regions containing lamellipodia. The GFP-vimentin images colocalized with a monoclonal antibody directed against vimentin, indicating that GFP-vimentin was incorporated into the preexisting endogenous vimentin network.
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Subcellular Microstimuli: Heterogeneous Spatial Displacement of Filaments by Shear Stress
Images from the beginning and end of timed intervals were false-colored red and green, respectively, so that yellow represented no change in GFP-vimentin position in merged color images, and green and red identified filament displacement. Time-lapse images of deconvolved optical sections revealed the dynamics of the filaments even in the absence of externally imposed forces (Figure 4A). This constitutive "wiggling" motion did not change the number of filaments nor their inter-IF connections and appeared to be random fluctuations. However, in response to a step change of flow in which the monolayer was subjected to a shear stress of 13 dyn/cm2, significant directional IF displacements of up to 1 µm occurred (Figure 4B) during the earliest image acquisition times after flow stimulation (90 seconds). The displacements were not attributable to cell migration or change in the position of each cell within the confluent monolayer. The spatial distribution of the responses was heterogeneous; filaments nearer the top of the cell were more greatly displaced than those near the basal surface36 but throughout each optical plane, irrespective of the height of the plane in the cell, displacement was heterogeneous. These data demonstrate that significant initial displacement of IFs occurs in response to shear stress and that the distribution of flow-induced IF movement is heterogeneous within the cell, reflecting inhomogeneity of the mechanical responses of the cytoskeleton to externally loaded stresses. Subcellular differences in the responses of IFs to shear suggest that IF dynamics plays a role in the cytoskeletal coordination of multiple signaling networks. It has been proposed that shear stresses are transmitted over the cell surface to junctions and basal adhesion sites, largely bypassing the cytoskeleton.62 Although such a mechanism undoubtedly contributes to force redistribution, our studies support a broader interpretation of force transmission through transcellular cytoskeletal displacement and consistent with mechanotransduction activated through the complex, organelle-rich body of the cell rather than the cell periphery alone. It is also increasingly clear from our studies and those of others that exogenous force transmission via filamentous elements linked to membrane surfaces and organelles provide exquisite sensitivity to allow appropriate cellular responses and that treating the cell as a solid body or gel-filled membrane is inappropriate.
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w=12 dyn cm-2); yellow represents zero displacement during the interval. Flow direction in all images was left to right. A, A small amount of filament displacement ("wiggling") in the absence of flow represents constitutive filament dynamics. B, Extensive displacement of filaments after the onset of flow at 13 dyn/cm2 shear stress. Reprinted from Helmke BP, Goldman RD, Davies PF. Rapid displacement of vimentin intermediate filaments in living endothelial cells exposed to flow. Circ Res. 2000;86:745–752, by permission of the American Heart Association ©2000.
Quantitation of Cytoskeletal Displacement in Living Cells
Quantitation of the displacement of filaments revealed important spatial insights into subcellular force transmission and its distribution. A displacement index (DI) was computed based on the spatial product moment cross correlation between 2 images to measure the degree of overlap or separation between red and green images.36 DI revealed both spatial and temporal patterns of flow-induced IF displacement. During no-flow intervals, small values of DI were computed in most spatial regions of cells ("wiggling"). However, DI was significantly increased in regions of the cell during the 90-second interval after the onset of shear stress (Figure 5). After the initial period of exposure to shear stress, DI was decreased and reached a new steady-state average rate of displacement, demonstrating that displacement of IF position was increased coincident with and after the onset of flow. On average, DI increased with height in the cell and in spatial regions downstream from the nucleus; however, DI was unchanged in subregions below the nucleus. Larger IF displacement may be expected near the luminal surface where shear forces act directly, and smaller displacement below the nucleus may represent relative structural stability in this area of the cell. Increased IF displacement with distance along the flow axis reveals a dependence on filament connections throughout the cytoplasm, suggesting that cytoskeletal network morphology plays a role in transmitting force through the cell away from the upstream luminal surface. It is important to note that heterogeneous spatial distribution of DI at all heights in the cell demonstrates that cellular deformation is not simple shear but a more complex displacement pattern that depends on mechanical interactions among cytoskeletal elements.
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Computation of Intracellular Strain From Cytoskeletal Displacement
The coordinated displacement of a group of interconnected IFs can occur as a local displacement without mechanical strain of the cytoskeletal network, for example, the positional movement of a group of interconnected IFs (translation). To separate translation and deformation, a method was developed to compute the intracellular strain field by detecting the relative deformation of locally connected IFs from images of optical sections that were skeletonized to represent in 2 dimensions the positions of IFs.37 Coordinates of vertices, or connection points, among three or more IF segments were extracted at each time point, and a particle-tracking algorithm determined the path of each vertex as a function of time. Thus, the projected displacement field could be computed using image features extracted directly from the cytoskeleton morphology. The smallest possible set of triangles connecting all vertices tracked in the data set was computed, and the strain tensor was solved to describe the average strain field in the spatial regions among every set of 3 vertices. A strain map for an optical plane through the basal region of an endothelial cell is shown in Figure 6. Strain was heterogeneous with small regions of high-stretch concentrations located at the cell periphery and at several locations in the cell interior. The former corresponds to the border with adjacent (untransfected, unmapped) cells in the confluent monolayer, whereas the latter may represent adhesion sites. Spatially localized peaks in IF strain were repositioned after the onset of shear stress. This approach also computed not only the magnitudes but also the directions of principal strain at each spatial location. We noted that stretch orientation was often changed from parallel to perpendicular by the imposition of flow, an effect that was most notable at the upstream boundary of the cell. These studies are the first high-resolution strain maps to be constructed using an endogenous reporter molecule, GFP-vimentin, in living cells. This redistribution of flow-induced strain indicated by the intermediate-filament cytoskeleton is consistent with the displacement of microtubules, microfilaments, and associated structures such as mitochondria that has been measured previously in response to force application at the cell surface.40 The measurements indicate that extracellular forces are transmitted by the cytoskeleton throughout the cytoplasm to sites such as cell junctions and adhesion sites where mechanochemical signals have been detected.45,42
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I during a 3-minute interval immediately after onset of shear stress (12 dyn/cm2, left to right). C, Filamentous skeleton image superimposed on the magnitude of principal stretch as a spatial reference. Note strain focusing at dense interconnections among IF segments in panel B. Also see Reference
Microstimuli Initiate Molecular Mechanisms of Mechanotransduction
Although the exact mechanisms of mechanotransduction remain unclear, there is evidence for multiple molecular interactions. As discussed above, fluid forces act directly on cell surface transmembrane proteins, the lipid bilayer itself, the cytoskeleton, and, through the cytoskeleton, to connected proteins at multiple locations throughout the cell. The plasma membrane contains structures that are capable of both transmitting force to the cytoskeleton and transducing force into biochemical signals. Signal transduction may be localized to membrane regions such as caveolae that are both rich in signaling molecules and exhibit structure that is capable of transmitting tension.63 Spatial localization of signaling molecules that associate with actin stress fibers also implicates caveolae in directly transmitting force to the cytoskeleton,64 although the structural and signaling links between caveolae and the cytoskeleton are poorly understood. At the endothelial cell luminal surface membrane, domains composed of 
5ß1 integrin, vinculin, talin, and paxillin are structurally linked to the cytoskeleton65 analogous to focal adhesion sites. Shear stress may deform or otherwise stimulate these complexes in a manner similar to that described for basal adhesion signaling.41 The biophysical properties of the endothelial surface glycocalyx, a layer enriched with glycoproteins, are poorly understood in relation to mechanosignaling. Branching carbohydrate structures extend into the fluid layer at the luminal surface and may be deformed by flow, possibly resulting in conformational changes in the protein core. This in turn may modulate the ability of transmembrane proteins or ion channels to sense the fluid mechanical environment. The physical properties of the plasma membrane itself may be altered by shear stress, perhaps through membrane fluidity changes,53 leading to activation of adjacent G proteins within seconds.27
The direct transmission of extracellular forces to the submembranous cytoskeleton is strongly suggested by experiments that bound small microspheres coated with ligands to integrin receptors of the cell surface.31 Resistance to optical66 or magnetic31 forces applied to the beads demonstrated direct connection to an intracellular mechanical structure, most likely the cortical cytoskeleton and its connecting cytoplasmic cytoskeletal network. The cytoskeleton quickly remodels to reinforce the strength of attachment66 through a mechanism that involves activation of signaling molecules such as the tyrosine kinase src.67 Such investigations highlight the close relationship that exists between locally applied shear stress and the detailed geometry of the endothelial luminal surface. Physical connections to the surface via integrins and scaffolding proteins65 transmit the deformation to microtubules and microfilaments almost simultaneously,39 and cross-linking to IFs68 also implicates network force transmission. Although IFs in endothelial cells are mechanically connected to intercellular junctions by desmoplakin,69 it is not yet clear whether they interact directly with protein scaffolds at the luminal cell surface or indirectly through crosslinks to other cytoskeletal structures. Although it appears reasonable to conclude that locations of greatest shear stress concentration at the cell surface will correspondingly influence the 2-dimensional and 3-dimensional subcellular distribution of cytoskeletal strain, the complex and dynamic interconnections of microfilaments, IFs, and microtubules represent a formidable quantitative challenge for detailed analyses. It will be illuminating to extend the imaging of deformation to multiple cytoskeletal components in living cells and map the resulting strain as described above for IFs. The roles of accessory or regulatory proteins and organelles, currently poorly defined, will need to be better understood.
There are many interactions between the 3 principal cytoskeleton networks that influence intracellular force transmission. These include the interaction of vimentin with F-actin and phospholipids to regulate polymerization,70 vimentin transport to sites of IF network formation that requires intact microtubule tracks dependent on the motor protein kinesin,71 and IF cross-linking to microfilaments and microtubules by plectin68 and nestin72 in stable structures. Multiple signaling networks involving small GTPases can regulate this balance of cytoskeletal interactions, emphasizing the relationship between force transmission and cytoplasmic biochemical pathways. Shear stress–induced redistribution of IFs near the basal side of the cell affects the dynamics of focal adhesion sites.73 Because the perinuclear ring of vimentin IFs may be directly or indirectly linked to the nuclear lamina,33 force redistribution under flow may also affect the karyoskeleton, consistent with other mechanical perturbations. Through interactions between nuclear IF proteins, the nuclear lamins, DNA, and histones,70,71,74 changes in gene expression may be directly mediated by flow.
These molecular interactions proposed for force transmission and transduction suggest that flow induces biochemical signal initiation at discrete sites in the cell in response to an altered intracellular force distribution. The overall cellular response is integrated, as suggested by the decentralized mechanotransduction hypothesis, but the key molecular events that transduce mechanical force into a biochemical reaction remain unknown. To emphasize the integrated nature of mechanotransduction, Ingber,42 while outlining the subcellular architecture that constitutes an appropriate response, has succinctly noted that "the whole cell is the mechanosensor" (page 882). However, it behooves us to define the many sets of molecules that contribute to mechanotransduction and to obtain a detailed understanding of their regulation in physiological and pathological states.
Microstimulation of Adhesion Sites
Using tandem scanning confocal microscopy, a xenon-wavelength confocal version of interference reflection microscopy, we imaged the endothelial basal cell surface involvement in adhesion in living confluent endothelial cells.75 Time-lapse image subtraction revealed the dynamic nature of the adhesion sites, their locations relative to each other, and permitted measurement of the area of membrane associated with close and focal adhesion sites. Cell adhesion was also estimated by integration of the distance between the cell surface and the substrate with the area of contact. Significant rearrangements of attachment areas during periods of several minutes were recorded.76 Adhesion site remodeling was random in control no-flow cells. In contrast, in confluent endothelial cells subjected to 10 dyn/cm2, focal adhesions showed directional remodeling during a similar interval. Thus, these abluminal membrane regions are mechanically responsive to luminal shear stress. Over extended periods of flow, separate adhesion sites frequently joined together and became aligned with the flow. However, although there were fewer sites, they were larger in area, an observation recently also noted in migrating cells.77 The total area of membrane contact remained relatively constant, and when membrane-substrate distance was factored in, a nominal measure of cell adhesion in real time indicated little change in adhesion, despite substantial cellular remodeling over periods as long as 24 hours. When we substituted collagen for vitronectin as the substrate, the rate of remodeling of focal adhesions decreased by half. These results probably reflect the fundamental importance of cell adhesion (and membrane integrin-adhesion protein interactions) in the context of normal differentiation and survival42 or the epigenetic regulation of migration, proliferation, and apoptosis in transformed cells.78 The unexpected shear stress responsiveness of a subpopulation of adhesion sites in confluent endothelium demonstrates that the topography of the abluminal surface may be of equal importance as that of the luminal endothelium. The adhesion sites are not only foci of resistance to the torsional forces that shear stress imposes on the cell body but also, as shown above, are sites of strain concentration resulting from the transcellular transmission of forces by the cytoskeleton. Biochemical evidence of adhesion site-specific responses to flow have been reported by several groups and includes phosphorylation of focal adhesion kinase,79 paxillin dephosphorylation/rephosphorylation,19 and movement of kinases such as PYK-2 from the cytosol to the adhesions.80 Paxillin phosphorylation and integrin clustering occur within minutes after onset of shear stress, and focal adhesion–associated molecules serve as a scaffold for recruitment of signaling molecules such as Grb2 and Shc,81 which play a role in activating mitogen-activated protein kinase signaling.82
Microstimuli at Cell-Cell Junctions and Associated Proteins
Several molecular interactions suggest a role for intercellular junctions in mediating mechanotransduction. Adherens junctions, composed of VE-cadherin, -catenin and ß-catenin, and plakoglobin, adapt their structure concurrently with the actin cytoskeleton. As actin-dense peripheral bands dissociate during the first several hours after onset of unidirectional steady laminar shear stress, the distributions of VE-cadherin, -catenin, and ß-catenin change from continuous structures along cell edges to punctate complexes that are located only in areas of contact between adjacent cells. With continued exposure to shear stress, junctional complexes elongate into small dashes and associate with the ends of F-actin stress fibers as they assemble.83 In some endothelial cell types, vimentin IFs are also inserted into adherens junctions through association with desmoplakin,69 a molecule that associates with plakoglobin in epithelial junctions. Because IFs maintain their network structure and are displaced by similar direction and magnitude in adjacent cells after onset of shear stress,35,36 this cytoskeletal component probably plays a role in the structural integrity of the monolayer by maintaining the associations among VE-cadherin, -catenin, and ß-catenin as F-actin dissociates and reassembles in response to the extracellular force. Temporary translocation of plakoglobin away from the junctions to the nucleus during adaptation to shear stress may serve to regulate gene expression by interacting with TCF-type transcription factors.83 Interestingly, ß-catenin also signals to these factors downstream of Wnt-1 activation in cardiomyocytes.84 In these cells, ß-catenin is the effector molecule for Wnt-mediated regulation of connexin43 expression, thereby modulating cell-cell communication via gap junctions. Both connexin43 and cadherins are associated with the Triton-insoluble cytoskeletal cell fractions,85 and shear stress induces redistribution of connexin43 that affects cell-cell communication.86 Fujiwara and colleagues45 have proposed that PECAM-1, a cell adhesion molecule localized to the interendothelial adhesion site at the cell periphery in confluent endothelium, may play a role in endothelial mechanosensing. PECAM-1 is rapidly phosphorylated by mechanical forces, including hyperosmotic stretch and flow-induced shear stress. When magnetic beads coated with antibodies directed against the extracellular domain of PECAM-1 were placed at the endothelial surface, PECAM accumulated at the bead interface. An attractive magnetic field exerted a pull on the molecule for a 10-minute period, after which the isolated PECAM was found to be phosphorylated. Neither binding alone nor irrelevant control antibodies resulted in PECAM phosphorylation. Coincident with PECAM phosphorylation was phosphorylation of the extracellular signal-regulated kinase (ERK), a result that was dependent on SHP-2 binding to phospho-PECAM and SHP-2 phosphatase activity.45 SHP-2 was also observed to transfer to the interendothelial junction upon flow exposure.
These findings suggest that shear stress transmitted from the luminal surface to cell-cell junctions via the cytoskeleton may regulate a complex molecular signaling network that modulates endothelial barrier permeability and intercellular communication.
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Flow-Mediated Chemical Signaling |
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The microspatial relationships discussed throughout this review are most relevant to shear stress forces that result in displacement of specific localized membrane sensors and/or multiple connected components extending throughout the cell. However, labile chemicals at the cell surface may also activate endothelial flow responses independently of physical displacement of the cell.3,87,88 Thus, when high local concentrations of labile agonists are released close to the endothelial surface, increased flow improves convective delivery to endothelial receptors in an autocrine loop, whereas decreased flow slows convection. The endothelium possesses modifying (activating and degradative) ectoenzymes for important blood-borne agents such as bradykinin, adenonucleotides, angiotensin, and proproteins. When the removal rate at the endothelial surface exceeds convective and diffusive delivery rates from the bulk fluid, a steep concentration gradient exists between the fluid and the cell surface and this in turn is influenced by the flow characteristics. Flow-mediated chemical responses and shear stress displacement responses can be separated experimentally, although it is unlikely they actually occur independently of each other. When shear stress was greatly increased by changing the fluid viscosity, with only small changes in mass transport, flow-mediated relaxation of intact arteries was enhanced indicating that shear stress was the principal effector.89 Ando et al90 have also provided evidence for a direct mechanism of force transduction in evoking [Ca2+]i responses that is additional to the effects of flow on mass transport. Thus, the potency of the local chemical environment should be considered whenever possible, and the spatial relationships within chemical gradients near the cell surface can become a complex factor in flow-mediated cellular responses. Such measurements are difficult to obtain. It is likely that both physical displacement of the cell and local chemical concentrations interact to evoke a mechanochemical transduction response.
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TopAbstractIntroductionLength ScalesWhere Are the Hemodynamic...Microstimuli at the Luminal...Four-Dimensional Cytoskeletal...Flow-Mediated Chemical SignalingFuture DirectionsReferences |
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The endothelium is a multifunctional effector of vessel wall biology, and a sensitive responder to the local mechanical environment, roles that are closely intertwined and of great physiological and pathological importance. A decentralized model of endothelial mechanotransduction readily accommodates a variety of mechanisms including stress sensors at the luminal endothelial surface, force transmission throughout the cell via the cytoskeleton to create hot spots of strain, and sensors of tension change at basal, junctional, and nuclear sites. The interface between fluid forces and the cell, however, remains the endothelial luminal cell surface, the geometry of which influences the magnitude and distribution of mechanical force transmission. Therefore, variations in endothelial surface topography from region to region and cell to cell, and the dynamics of these microstimulatory changes, remain important determinants of flow responses in the endothelial monolayer. Modern optical techniques are revealing the astonishing temporal changes of membranes, organelles, and tagged molecules that occur both constitutively and after cell stimulation. These will provide unpredicted insights into the dynamics of both cell structures and metabolic function associated with biomechanical responses.
Continuing advances in understanding the material properties of intermediate filaments, microfilaments, and microtubules is essential for accurate intracellular force mapping. Furthermore, interactions between IFs and other cytoskeletal elements, including cross-linking proteins such as plectin, will play a role in determining the dynamic response to shear stress. Measurements at the functional level in vimentin-null mice have revealed impaired wound healing91 and loss of flow-induced arterial remodeling.92 These observations of compromised vascular tissue function implicate IFs in the maintenance of contractility and mechanical integrity at the cellular level.93 However, the contribution of IFs to mechanisms regulating cell mechanics and initiation of mechanotransduction has only recently been investigated.28,35–40
Recent genomics investigations of the links between hemodynamic macrostimuli and cultured endothelial cells have revealed the identities of novel shear stress–responsive genes94,95 and proteins. RNA amplification techniques now permit small numbers of cells to be transcriptionally profiled with reliable fidelity,96 and fingerprints of gene expression at the level of single cells18 may soon be routine, an approach that permits spatial (and thus hemodynamic) precision of gene expression.16 Considerable insights will be gained by the convergence, in a single cell, of mechanotransduction imaging and the expression levels of many genes (and eventually proteins).16 Once subsets of genes/proteins activated or suppressed in mechanotransduction are identified, it should be possible to take such an analysis to the subcellular level of protein localization and dynamics in living cells.
New techniques for the 3-dimensional visualization of organelles and molecular complexes at high resolution in living or flash-frozen cells are emerging into the mainstream and will soon become more widely available. A recent example is the development of cryoelectron tomography that enables 3-dimensional projections to be made of volumes extending 300 to 600 nm into the cell.97 Two-dimensional 60-nm-thick slices displayed as tomograms can be projected as volume images to reveal 3-dimensional structures. Although a noninvasive technique, the cells must be quickly frozen before imaging, limiting its usefulness for studies of cellular dynamics associated with biomechanical stimuli. Nevertheless, this and other approaches to near-field imaging will provide unique insights into the structural elements that are central to many aspects of cellular biomechanics.
The availability of fluorescent cytoskeleton fusion proteins to visualize filament location in living cells is a powerful tool for spatiotemporal studies of intracellular mechanics. GFP-actin, GFP-tubulin, and GFP-vimentin are readily available and, together with increasingly sophisticated optical systems and advances in fluorescence resonance energy transfer (FRET) and associated techniques, are leading to ever more detailed studies of the cytoskeleton and its relationships to other cellular components. Although quantitative spatial studies of filament movement can be readily performed, as illustrated here, interpretation of the biological significance is limited by the lack of detailed knowledge of the material properties of the filaments themselves. This is attributable in part to the complex organization of filamentous bundles and the dynamic reorganization that some components undergo over short intervals. Perhaps more difficult to determine, however, are the properties of critical linking molecules between the cytoskeleton and its anchoring points near cell membranes. From an engineering perspective, little is known about these molecules. Furthermore, the interplay between different cytoskeletal elements that connect at nodal locations through such linkers is an added complication. As some of these gaps in knowledge are filled, an accurate mechanochemical model of the cell that reflects its complexity will become clearer.
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Acknowledgments |
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This work was supported by NIH grants HL36049 (MERIT Award), HL64388, HL62250, and HL10058 (to B.P.H.). We thank Dr Robert Goldman of Northwestern University who kindly provided the original GFP-vimentin gene construct.
Received December 13, 2002; revision received January 22, 2003; accepted January 23, 2003.
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References |
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|
TopAbstractIntroductionLength ScalesWhere Are the Hemodynamic...Microstimuli at the Luminal...Four-Dimensional Cytoskeletal...Flow-Mediated Chemical SignalingFuture DirectionsReferences |
|---|
1. Flaherty JT, Pierce JE, Ferrans VJ, Patel DJ, Tucker WK, Fry DL. Endothelial nuclear patterns in the canine arterial tree with particular reference to hemodynamic events. Circ Res. 1972; 30: 23–33.
2. Dewey CF Jr, Bussolari SR, Gimbrone MA Jr, Davies PF. The dynamic response of vascular endothelial cells to fluid shear stress. J Biomech Eng. 1981; 103: 177–185.[Medline] [Order article via Infotrieve]
3. Davies PF. Flow-mediated endothelial mechanotransduction. Physiol Rev. 1995; 75: 519–560.
4. Frangos JA, Eskin SG, McIntire LV, Ives CL. Flow effects on prostacyclin production by cultured human endothelial cells. Science. 1985; 227: 1477–1479.
5. Pohl U, Holtz J, Busse R, Bassenge E. Crucial role of endothelium in the vasodilator response to increased flow in vivo. Hypertension. 1986; 8: 37–47.
6. Ayajiki K, Hindermann M, Hecker M, Fleming I, Busse R. Intracellular pH and tyrosine phosphorylation but not calcium determine shear stress–induced nitric oxide production in native endothelial cells. Circ Res. 1996; 78: 750–758.
7. Huang A, Sun D, Carroll MA, Jiang H, Smith CJ, Connetta JA, Falck JR, Shesely EG, Koller A, Kaley G. EDHF mediates flow-induced dilation in skeletal muscle arterioles of female eNOS-KO mice. Am J Physiol. 2001; 280: H2462–H2469.
8. Griffiths TM. Endothelial control of vascular tone by nitric oxide and gap junctions: a haemodynamic perspective. Biorheology. 2002; 39: 307–318.[Medline] [Order article via Infotrieve]
9. Langille BL, O’Donnell F. Reductions in arterial diameter produced by chronic decreases in blood flow are endothelium-dependent. Science. 1986; 231: 405–407.
10. Ross R. Cell biology of atherosclerosis. Annu Rev Physiol. 1995; 57: 791–804.[CrossRef][Medline] [Order article via Infotrieve]
11. Cornhill JF, Henderick EE, Stary HC. Topography of human aortic sudanophilic lesions. In: Liepsh DW, ed. Monographs on Atherosclerosis. Basel, Switzerland: Karger; 1990: 15: 13–19.
12. Glagov S, Zarins CK, Giddens DP, Ku DN. Hemodynamics in atherosclerosis: insights and perspectives gained from studies of human arteries. Arch Pathol Lab Med. 1988; 316: 1371–1375.
13. Shepherd JT, Vanhoutte PM. The Human Cardiovascular System: Facts and Concepts. New York, NY: Raven Press; 1979.
14. Resnick N, Gimbrone MA. Hemodynamic forces are complex regulators of endothelial gene expression. FASEB J. 1995; 9: 874–882.[Abstract]
15. Shyy JY, Chien S. Role of integrins in endothelial mechanosensing of shear stress. Circ Res. 2002; 91: 769–775.
16. Davies PF, Polacek DC, Shi C, Helmke BP. The convergence of haemodynamics, genomics, and endothelial structure in studies of the focal origin of atherosclerosis. Biorheology. 2002; 39: 299–306.[Medline] [Order article via Infotrieve]
17. Davies PF, Reidy MA, Goode TB, Bowyer DE. Scanning electron microscopy in the evaluation of endothelial integrity of the fatty streak lesion of atherosclerosis. Atherosclerosis. 1976; 25: 125–130.[CrossRef][Medline] [Order article via Infotrieve]
18. Davies PF, Shi C, DePaola N, Helmke BP, Polacek DC. Hemodynamics and the focal origin of atherosclerosis: a spatial structural, gene expression, and functional approach. Ann N Y Acad Sci. 2001; 947: 7–18.[Medline] [Order article via Infotrieve]
19. Davies PF, Barbee KA, Volin MV, Robotewskyj A, Chen J, Joseph L, Griem ML, Wernick M, Jacobs E, Polacek DC, DePaola N, Barakat A. Spatial relationships in early signaling events of flow-mediated endothelial mechanotransduction. Annu Rev Physiol. 1997; 59: 527–549.[CrossRef][Medline] [Order article via Infotrieve]
20. Olesen SP, Clapham DE, Davies PF. Haemodynamic shear stress activates a K+ current in vascular endothelial cells. Nature. 1988; 331: 168–170.[CrossRef][Medline] [Order article via Infotrieve]
21. Shen J, Luscinskas FW, Connolly A, Dewey CF Jr, Gimbrone MA Jr. Fluid shear stress modulates cytosolic free calcium in vascular endothelial cells. Am J Physiol. 1992; 262: C384–C390.[Medline] [Order article via Infotrieve]
22. Gudi SRP, Clark CB, Frangos JA. Fluid flow rapidly activates G proteins in human endothelial cells: involvement of G proteins in mechanochemical signal transduction. Circ Res. 1996; 79: 834–839.
23. Jo H, Sipos K, Go YM, Law R, Rong J, McDonald JM. Differential effect of shear stress on extracellular signal–regulated kinase and N-terminal Jun kinase in endothelial cells: Gi2- and Gß-dependent signaling pathways. J Biol Chem. 1997; 272: 1395–1401.
24. Ishida T, Peterson TE, Kovach NL, Berk BC. MAP kinase activation by flow in endothelial cells: role of ß1 integrins and tyrosine kinases. Circ Res. 1996; 79: 310–316.
25. Jalali S, Li YS, Sotoudeh M, Yuan S, Li S, Chien S, Shyy JY-J. Shear stress activates p60src-Ras-MAPK signaling pathways in vascular endothelial cells. Arterioscler Thromb Vasc Biol. 1998; 18: 227–234.
26. Davies PF, Tripathi SC. Mechanical stress mechanisms and the cell: an endothelial paradigm. Circ Res. 1993; 72: 239–245.
27. Gudi SRP, Nolan JP, Frangos JA. Modulation of GTPase activity of G proteins by fluid shear stress and phospholipid composition. Proc Natl Acad Sci U S A. 1998; 95: 2515–2519.
28. Helmke BP, Davies PF. The cytoskeleton under external fluid mechanical forces: hemodynamic forces acting on the endothelium. Ann Biomed Eng. 2002; 30: 284–296.[CrossRef][Medline] [Order article via Infotrieve]
29. Davies PF, Robotewskyj A, Griem ML. Quantitative studies of endothelial cell adhesion: directional remodeling of focal adhesion sites in response to flow forces. J Clin Invest. 1994; 93: 2031–2038.[Medline] [Order article via Infotrieve]
30. Li S, Kim M, Hu YL, Jalali S, Schlaepfer DD, Hunter T, Chien S, Shyy JY-J. Fluid shear stress activation of focal adhesion kinase: linking to mitogen-activated protein kinases. J Biol Chem. 1997; 272: 30455–30462.
31. Wang N, Butler JP, Ingber DE. Mechanotransduction across the cell surface and through the cytoskeleton. Science. 1993; 260: 1124–1127.
32. Janmey PA. The cytoskeleton and cell signaling: component localization and mechanical coupling. Physiol Rev. 1998; 78: 763–781.
33. Maniotis AJ, Chen CS, Ingber DE. Demonstration of mechanical connections between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear structure. Proc Natl Acad Sci U S A. 1997; 94: 849–854.
34. Galbraith CG, Skalak R, Chien S. Shear stress induces spatial reorganization of the endothelial cell cytoskeleton. Cell Motil Cytoskel. 1998; 40: 317–330.[CrossRef][Medline] [Order article via Infotrieve]
35. Helmke BP, Goldman RD, Davies PF. Rapid displacement of vimentin intermediate filaments in living endothelial cells exposed to flow. Circ Res. 2000; 86: 745–752.
36. Helmke BP, Thakker DB, Goldman RD, Davies PF. Spatiotemporal analysis of flow-induced intermediate filament displacement in living endothelial cells. Biophys J. 2001; 80: 184–194.[Medline] [Order article via Infotrieve]
37. Helmke BP, Rosen AB, Davies PF. Mapping mechanical strain of an endogenous cytoskeletal network in living endothelial cells. Biophys J. In press.
38. Stamenovic D, Mijailovich SM, Tolic-Norrelykke IM, Chen J, Wang N. Cell prestress, II: contribution of microtubules. Am J Physiol. 2002; 282: C617–C624.
39. Wang N, Naruse K, Stamenovic D, Fredberg JJ, Mijailovich SM, Tolic-Norrelykke IM, Polte T, Mannix R, Ingber DE. Mechanical behavior in living cells consistent with the tensegrity model. Proc Natl Acad Sci U S A. 2001; 98: 7765–7770.
40. Wang N, Stamenovic D. Contribution of intermediate filaments to cell stiffness, stiffening, and growth. Am J Physiol. 2000; 279: C188–C194.
41. Burridge K, Fath K, Kelly T, Nuckolls G, Turner C. Focal adhesions. Annu Rev Cell Biol. 1988; 4: 487–525.[CrossRef]
42. Ingber DE. Mechanical signaling and the cellular response to extracellular matrix in angiogenesis and cardiovascular physiology. Circ Res. 2002; 91: 877–887.
43. Pienta KJ, Coffey DS. Nuclear-cytoskeletal interactions: evidence for physical connections between the nucleus and cell periphery and their alteration by transformation. J Cell Biochem. 1992; 49: 357–365.[CrossRef][Medline] [Order article via Infotrieve]
44. Takeichi M. The cadherins: cell-cell adhesion molecules controlling animal morphogenesis. Development. 1988; 102: 639–655.
45. Osawa M, Masuda M, Kusano K, Fujiwara K. Evidence for a role of PECAM-1 in endothelial cell mechanosignal transduction: is it a mechanoresponsive molecule? J Cell Biol. 2002; 158: 773–785.
46. Satcher RL, Bussolari SR, Gimbrone MA Jr, Dewey CF Jr. The distribution of fluid forces on model arterial endothelium using computational fluid dynamics. J Biomech Eng. 1992; 114: 309–316.[Medline] [Order article via Infotrieve]
47. Sakurai A, Nakano A, Yamaguchi T, Masuda M, Fujiwara K. A computational fluid mechanical study of flow over cultured endothelial cells. Adv Bioeng. 1991; 20: 299–302.
48. Yamaguchi T, Hoshiai K, Okino H, Kakurai A, Hanai S, Masuda M, Fujiwara K. Shear stress distribution over confluently cultured endothelial cells studied by computational fluid mechanics. Adv Bioeng. 1993; 20: 167–170.
49. Barbee KA, Davies PF, Lal R. Shear stress-induced reorganization of the surface topography of living endothelial cells imaged by atomic force microscopy. Circ Res. 1994; 74: 163–171.
50. Barbee KA, Mundel T, Lal R, Davies PF. Subcellular distribution of shear stress at the surface of flow aligned and non-aligned endothelial monolayers. Am J Physiol. 1995; 268: H1765–H1772.[Medline] [Order article via Infotrieve]
51. Davies PF, Mundel T, Barbee KA. A mechanism for heterogeneous endothelial responses to flow in vivo and in vitro. J Biomech. 1995; 28: 1553–1560.[CrossRef][Medline] [Order article via Infotrieve]
52. Davies PF, Polacek DC, Handen JS, Helmke BP, DePaola N. A spatial approach to gene expression profiling: mechanotransduction and the focal origin of atherosclerosis. Trends Biotechnol. 1999; 17: 347–351.[CrossRef][Medline] [Order article via Infotrieve]
53. Butler PJ, Norwich G, Weinbaum S, Chien S. Shear stress induces a time- and position-dependent increase in endothelial cell membrane fluidity. Am J Physiol. 2001; 280: C962–C969.
54. Sato M, Levesque MJ, Nerem RM. Micropipette aspiration of cultured bovine aortic endothelial cells exposed to shear stress. Arteriosclerosis. 1987; 7: 276–286.
55. Sato M, Ohshima N, Nerem RM. Viscoelastic properties of cultured porcine aortic endothelial cells exposed to shear stress. J Biomech. 1996; 29: 461–467.[CrossRef][Medline] [Order article via Infotrieve]
56. Lee JC, Discher DE. Deformation-enhanced fluctuations in the red cell skeleton with theoretical relations to elasticity, connectivity, and spectrin unfolding. Biophys J. 2001; 81: 3178–3192.[Medline] [Order article via Infotrieve]
57. Malinauskas R, Herrmann R, Truskey G. The distribution of intimal white blood cells in the normal rabbit aorta. Atherosclerosis. 1995; 115: 147–163.[CrossRef][Medline] [Order article via Infotrieve]
58. Malek AM, Izumo S. Mechanism of endothelial cell shape change and cytoskeletal remodeling in response to fluid shear stress. J Cell Sci. 1996; 109: 713–726.[Abstract]
59. Ludin B, Matus A. GFP illuminates the cytoskeleton. Trends Cell Biol. 1998; 8: 72–77.[CrossRef][Medline] [Order article via Infotrieve]
60. Heidemann SR, Kaech S, Buxbaum RE, Matus A. Direct observations of the mechanical behaviors of the cytoskeleton in living fibroblasts. J Cell Biol. 1999; 145: 109–122.
61. Yoon M, Moir RD, Prahlad V, Goldman RD. Motile properties of vimentin intermediate filament networks in living cells. J Cell Biol. 1998; 143: 147–157.
62. Fung YC, Liu SQ. Elementary mechanics of the endothelium of blood vessels. J Biomech Eng. 1993; 115: 1–12.[Medline] [Order article via Infotrieve]
63. Rizzo V, Sung A, Oh P, Schnitzer JE. Rapid mechanotransduction in situ at the luminal cell surface of vascular endothelium and its caveolae. J Biol Chem. 1998; 273: 26323–26329.
64. Fujimoto T, Miyawaki A, Mikoshiba K. Inositol 1,4,5-trisphosphate receptor-like protein in plasmalemmal caveolae is linked to actin filaments. J Cell Sci. 1995; 108: 7–15.[Abstract]
65. Kano Y, Katoh K, Masuda M, Fujiwara K. Macromolecular composition of stress fiber-plasma membrane attachment sites in endothelial cells in situ. Circ Res. 1996; 79: 1000–1006.
66. Choquet D, Felsenfeld DP, Sheetz MP. Extracellular matrix rigidity causes strengthening of integrin-cytoskeleton linkages. Cell. 1997; 88: 39–48.[CrossRef][Medline] [Order article via Infotrieve]
67. Felsenfeld DP, Schwartzberg PL, Venegas A, Tse R, Sheetz MP. Selective regulation of integrin-cytoskeleton interactions by the tyrosine kinase Src. Nat Cell Biol. 1999; 1: 200–206.[CrossRef][Medline] [Order article via Infotrieve]
68. Svitkina TM, Verkhovsky AB, Borisy GG. Plectin sidearms mediate interaction of intermediate filaments with microtubules and other components of the cytoskeleton. J Cell Biol. 1996; 135: 991–1007.
69. Kowalczyk AP, Navarro P, Dejana E, Bornslaeger EA, Green KJ, Kopp DS, Boegwardt JE. VE-cadherin and desmoplakin are assembled into dermal microvascular endothelial intercellular junctions: a pivotal role for plakoglobin in the recruitment of desmoplakin to intercellular junctions. J Cell Sci. 1998; 111: 3045–3057.[Abstract]
70. Shah JV, Wang LZ, Traub P, Janmey PA. Interaction of vimentin with actin and phospholipids. Biol Bull. 1998; 194: 402–405.[Abstract]
71. Prahlad V, Yoon M, Moir RD, Vale RD, Goldman RD. Rapid movements of vimentin on microtubule tracks: kinesin-dependent assembly of intermediate filament networks. J Cell Biol. 1998; 143: 159–170.
72. Steinert PM, Chou YH, Prahlad V, Parry DA, Marekov LN, Wu KC, Jang SI, Goldman RD. A high molecular weight intermediate filament-associated protein in BHK-21 cells is nestin, a type VI intermediate filament protein. J Biol Chem. 1999; 274: 9881–9890.
73. Li S, Kim M, Hu YL, Jalali S, Schlaepfer DD, Hunter T, Chien S, Shyy JY-J. Fluid shear stress activation of focal adhesion kinase: linking to mitogen-activated protein kinases. J Biol Chem. 1997; 272: 30455–30462.
74. Guilak F. Compression-induced changes in the shape and volume of the chondrocyte nucleus. J Biomech. 1995; 28: 1529–1541.[CrossRef][Medline] [Order article via Infotrieve]
75. Davies PF, Robotewskyj A, Dull RO, Griem ML. Endothelial cell adhesion in real-time: measurements by tandem scanning confocal image analysis. J Clin Invest. 1993; 91: 2640–2652.[Medline] [Order article via Infotrieve]
76. Davies PF, Robotewskyj A, Griem ML. Quantitative studies of endothelial cell adhesion: directional remodeling of focal adhesion sites in response to flow forces. J Clin Invest. 1994; 93: 2031–2038.[Medline] [Order article via Infotrieve]
77. Li S, Butler P, Wang Y, Hu Y, Han DC, Usami S, Guan JL, Chien S. The role of the dynamics of focal adhesion kinase in the mechanotaxis of endothelial cells. Proc Natl Acad Sci U S A. 2002; 99: 3546–3551.
78. Weaver VM, Lelievre S, Lakins JN, Chrenek MA, Jones JC, Giancotti F, Werb Z, Bissell MJ. ß4 integrin-dependent formation of polarized three-dimensional architecture confers resistance to apoptosis in normal and malignant mammary epithelium. Cancer Cell. 2002; 2: 205–216.[CrossRef][Medline] [Order article via Infotrieve]
79. Takahashi M, Berk BC. Mitogen-activated protein kinase (ERK1/2) activation by shear stress and adhesion in endothelial cells: essential role for a herbimycin-sensitive kinase. J Clin Invest. 1996; 98: 2623–2631.[Medline] [Order article via Infotrieve]
80. Davies PF. Multiple signaling pathways in flow-mediated endothelial mechanotransduction: PYK-ing the right location. Arterioscler Thromb Vasc Biol. 2002; 22: 1755–1757.
81. Li S, Kim M, Hu YL, Jalali S, Schlaepfer DD, Schlaepfer T, Hunter, Chien S, Shyy JY-J. Fluid shear stress activation of focal adhesion kinase: linking to mitogen-activated protein kinases. J Biol Chem. 1997; 272: 30455–30462.
82. Chen K-D, Li Y-S, Kim M, Li S, Li Yuan, Chien S, Shyy JY-J. Mechanotransduction in response to shear stress: roles of receptor tyrosine kinases, integrins, and Shc. J Biol Chem. 1999; 274: 18393–18400.
83. Noria S, Cowan DB, Gotlieb AI, Langille BL. Transient and steady-state effects of shear stress on endothelial cell adherens junctions. Circ Res. 1999; 85: 504–514.
84. Ai Z, Fischer A, Spray DC, Brown AM, Fishman GI. Wnt-1 regulation of connexin43 in cardiac myocytes. J Clin Invest. 2000; 105: 161–171.[Medline] [Order article via Infotrieve]
85. Ko K, Arora P, Lee W, McCulloch C. Biochemical and functional characterization of intercellular adhesion and gap junctions in fibroblasts. Am J Physiol. 2000; 279: C147–C157.
86. DePaola N, Davies PF, Pritchard WP, Polacek DC. Spatial regulation of gap junction connexin 43 in endothelial cells exposed to disturbed flows in vitro. Proc Natl Acad Sci U S A. 1999; 96: 3154–3160.
87. Dull RO, Davies PF. Flow modulation of agonist (ATP)-response (Ca++) coupling in vascular endothelial cells. Am J Physiol. 1991; 261: H149–H156.[Medline] [Order article via Infotrieve]
88. Dull RO, Tarbell JM, Davies PF. Mechanisms of flow-mediated signal transduction in endothelial cells: kinetics of ATP surface concentrations. J Vasc Res. 1992; 29: 410–419.[Medline] [Order article via Infotrieve]
89. Koller A, Sun D, Kaley G. Role of shear stress and endothelial prostaglandins in flow- and viscosity-induced dilation of arterioles in vitro. Circ Res. 1993; 72: 1276–1284.
90. Ando J, Ohtsuka A, Korenaga R, Kawamura T, Kamiya A. Wall shear stress rather than shear rate regulates cytoplasmic Ca++ responses to flow in vascular endothelial cells. Biochem Biophys Res Commun. 1993; 190: 716–723.[CrossRef][Medline] [Order article via Infotrieve]
91. Eckes B, Colucci-Guyon E, Smola H, Nodder S, Babinet C, Krieg T, Martin P. Impaired wound healing in embryonic and adult mice lacking vimentin. J Cell Sci. 2000; 113: 2455–2462.[Abstract]
92. Schiffers PM, Henrion D, Boulanger CM, Colucci-Guyon E, Langa-Vuves F, van Essen H, Fazzi GE, Levy BI, De Mey JG. Altered flow-induced arterial remodeling in vimentin-deficient mice. Arterioscler Thromb Vasc Biol. 2000; 20: 611–616.
93. Eckes B, Dogic D, Colucci-Guyon E, Wang N, Maniotis A, Ingber D, Merckling A, Langa F, Aumailley M, Delouvee A, Koteliansky V, Babinet C, Krieg T. Impaired mechanical stability, migration and contractile capacity in vimentin-deficient fibroblasts. J Cell Sci. 1998; 111: 1879–1907.
94. Topper JN, Gimbrone MA. Blood flow and vascular gene expression: fluid shear stress as a modulator of endothelial phenotype. Mol Med Today. 1999; 5: 40–46.[CrossRef][Medline] [Order article via Infotrieve]
95. Garcia-Cardena G, Comander J, Anderson KR, Blackman BR, Gimbrone MA. Biomechanical activation of vascular endothelium as a detriment of its functional phenotype. Proc Natl Acad Sci U S A. 2001; 98: 4478–4485.
96. Polacek DC, Passerini A, Manduchi E, Grant G, Shi C, Stoeckert C, Davies PF. Fidelity and enhanced sensitivity of differential transcription profiles following linear amplification of nanogram amounts of endothelial mRNA. Physiol Genomics. In press.
97. Medalia O, Weber I, Frangakis AS, Nicastro D, Gerisch G, Baumeister W. Macromolecular architecture in eukaryotic cells visualized by cryoelectron tomography. Science. 2002; 298: 1209–1213.
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 S. A. Wright, F. M. O'Prey, D. J. Rea, R. D. Plumb, A. J. Gamble, W. J. Leahey, A. B. Devine, R. C. McGivern, D. G. Johnston, M. B. Finch, et al. Microcirculatory Hemodynamics and Endothelial Dysfunction in Systemic Lupus Erythematosus Arterioscler Thromb Vasc Biol, October 1, 2006; 26(10): 2281 - 2287. [Abstract] [Full Text] [PDF]
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 C. Friedrich, N. Endlich, W. Kriz, and K. Endlich Podocytes are sensitive to fluid shear stress in vitro Am J Physiol Renal Physiol, October 1, 2006; 291(4): F856 - F865. [Abstract] [Full Text] [PDF]
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D. D'Arcangelo, V. Ambrosino, M. Giannuzzo, C. Gaetano, and M. C. Capogrossi Axl receptor activation mediates laminar shear stress anti-apoptotic effects in human endothelial cells Cardiovasc Res, September 1, 2006; 71(4): 754 - 763. [Abstract] [Full Text] [PDF]
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K. N. Dahl, P. Scaffidi, M. F. Islam, A. G. Yodh, K. L. Wilson, and T. Misteli Distinct structural and mechanical properties of the nuclear lamina in Hutchinson-Gilford progeria syndrome PNAS, July 5, 2006; 103(27): 10271 - 10276. [Abstract] [Full Text] [PDF]
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 H. Oberleithner, C. Riethmuller, T. Ludwig, V. Shahin, C. Stock, A. Schwab, M. Hausberg, K. Kusche, and H. Schillers Differential action of steroid hormones on human endothelium. J. Cell Sci., May 1, 2006; 119(Pt 9): 1926 - 1932. [Abstract] [Full Text] [PDF]
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 R. L. Armentano, J. G. Barra, D. B. Santana, F. M. Pessana, S. Graf, D. Craiem, L. M. Brandani, H. P. Baglivo, and R. A. Sanchez Smart Damping Modulation of Carotid Wall Energetics in Human Hypertension: Effects of Angiotensin-Converting Enzyme Inhibition Hypertension, March 1, 2006; 47(3): 384 - 390. [Abstract] [Full Text] [PDF]
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 E. A. Osborn, A. Rabodzey, C. F. Dewey Jr., and J. H. Hartwig Endothelial actin cytoskeleton remodeling during mechanostimulation with fluid shear stress Am J Physiol Cell Physiol, February 1, 2006; 290(2): C444 - C452. [Abstract] [Full Text] [PDF]
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J. D. Pearson Tightening the Barrier: Mechanical Forces and the Control of Endothelial Permeability Arterioscler Thromb Vasc Biol, January 1, 2006; 26(1): 10 - 11. [Full Text] [PDF]
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M. Dixit, A. E. Loot, A. Mohamed, B. Fisslthaler, C. M. Boulanger, B. Ceacareanu, A. Hassid, R. Busse, and I. Fleming Gab1, SHP2, and Protein Kinase A Are Crucial for the Activation of the Endothelial NO Synthase by Fluid Shear Stress Circ. Res., December 9, 2005; 97(12): 1236 - 1244. [Abstract] [Full Text] [PDF]
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 S. L. Cuvelier, S. Paul, N. Shariat, P. Colarusso, and K. D. Patel Eosinophil adhesion under flow conditions activates mechanosensitive signaling pathways in human endothelial cells J. Exp. Med., September 19, 2005; 202(6): 865 - 876. [Abstract] [Full Text] [PDF]
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I. Fleming, B. Fisslthaler, M. Dixit, and R. Busse Role of PECAM-1 in the shear-stress-induced activation of Akt and the endothelial nitric oxide synthase (eNOS) in endothelial cells J. Cell Sci., September 15, 2005; 118(18): 4103 - 4111. [Abstract] [Full Text] [PDF]
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R. Zaidel-Bar, Z. Kam, and B. Geiger Polarized downregulation of the paxillin-p130CAS-Rac1 pathway induced by shear flow J. Cell Sci., September 1, 2005; 118(17): 3997 - 4007. [Abstract] [Full Text] [PDF]
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 N. W. Blackstone and D. M. Bridge Model Systems for Environmental Signaling Integr. Comp. Biol., August 1, 2005; 45(4): 605 - 614. [Abstract] [Full Text] [PDF]
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J. C. R. Jones, K. Lane, S. B. Hopkinson, E. Lecuona, R. C. Geiger, D. A. Dean, E. Correa-Meyer, M. Gonzales, K. Campbell, J. I. Sznajder, et al. Laminin-6 assembles into multimolecular fibrillar complexes with perlecan and participates in mechanical-signal transduction via a dystroglycan-dependent, integrin-independent mechanism J. Cell Sci., June 15, 2005; 118(12): 2557 - 2566. [Abstract] [Full Text] [PDF]
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 D. Lominadze, N. Tsakadze, U. Sen, J. C. Falcone, and S. E. D'Souza Fibrinogen and fragment D-induced vascular constriction Am J Physiol Heart Circ Physiol, March 1, 2005; 288(3): H1257 - H1264. [Abstract] [Full Text] [PDF]
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H. A. Silber, P. Ouyang, D. A. Bluemke, S. N. Gupta, T. K. Foo, and J. A. C. Lima Why is flow-mediated dilation dependent on arterial size? Assessment of the shear stimulus using phase-contrast magnetic resonance imaging Am J Physiol Heart Circ Physiol, February 1, 2005; 288(2): H822 - H828. [Abstract] [Full Text] [PDF]
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P. A. VanderLaan, C. A. Reardon, and G. S. Getz Site Specificity of Atherosclerosis: Site-Selective Responses to Atherosclerotic Modulators Arterioscler Thromb Vasc Biol, January 1, 2004; 24(1): 12 - 22. [Abstract] [Full Text] [PDF]
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R. T. Lee Lessons From Lymph: Flow-Guided Vessel Formation Circ. Res., April 18, 2003; 92(7): 701 - 703. [Full Text] [PDF]
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