1Physiology Program, Harvard School of Public Health, Boston 02115; 2Department of Chemistry and Chemical Biology, Harvard University, Cambridge, Massachusetts 02138; and 3EOL Eberhard, CH-4104 Oberwil, Switzerland
Submitted 6 May 2004 ; accepted in final form 16 June 2004
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ABSTRACT |
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cytoskeleton; prestress; stress fibers; mechanotransduction; mechanical deformation
Various tools have been used to deform living cells, including microrods and cell poking, micropipette aspiration, elastomeric substrates, microchannels and fluid shear, scanning probe microscopes, and optical traps (1, 4, 6, 8, 9, 13, 27, 32). Magnetic particles attached to the cell surface can be manipulated using magnetic field gradients (2, 5), and the manipulations can include linear or oscillating forces or torques involving rotation of up to 90° (34, 35). The magnetic and optical approaches are among the most versatile because the probes can be coated with specific ligands and a wide range of force amplitudes and frequencies can be applied; they have been useful in characterizing the rheological properties of cells and the pathways by which cells transduce forces (2, 3, 10, 11, 17, 20, 21, 3437). All of these approaches are limited, however, in the range of motions that can be used to apply force; therefore, they are not ideally suited to characterize the mechanical anisotropy of the cell.
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MATERIALS AND METHODS |
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Three-dimensional magnetic device. Our three-dimensional (3-D) magnetic twisting cytometer contains the following seven components: 1) a high-voltage generator for generating the current in the coils used to magnetize the magnetic particles, 2) three separate bipolar current sources to twist the particles, 3) a personal computer to control the twisting, 4) an inverted microscope to observe the sample, 5) a charge-coupled device camera using self-written software to synchronize image capture with oscillatory magnetic fields, 6) a temperature controller for living cells, and 7) a microscope insert (see Fig. 6 in the APPENDIX).
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To estimate noise associated with the microscope stage, we determined the rigid body displacement by quantifying movements of a magnetic bead embedded in epoxy in the absence of an external load: at 125 ms/frame, the standard deviation of x displacement was 1.3 ± 0.02 nm and that of y displacement was 1.1 ± 0.1 nm (means ± SE). Thermal noise was 10 nm in living cells for YFP mitochondria or the nucleolus. Taking advantage of the periodicity of the sinusoidal inputs, we averaged 510 cycles of images of data to improve the signal-to-noise ratio for the YFP mitochondria and the nucleolus. The noise floor was reduced to
4 nm after averaging nine cycles of images and all load-induced displacements were synchronized with the frequency of the applied load at 0.3125 Hz (see Fig. 8 in the APPENDIX). The signal-to-noise ratio <2:1 for the YFP mitochondria or <1:1 for the nucleolus and 4 nm were chosen as the cutoff values (see Fig. 5).
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Micropatterning technique.
Polydimethylsiloxane (PDMS) membranes were made using photolithography according to procedures described previously (25). Briefly, PDMS membranes with the same area (2,500 µm2) holes but different shapes were fabricated: either 56-µm diameter circles, or 125 x 20-µm and 156 x 16-µm rectangles (length-to-width ratios of 6.25 and 9.75, respectively; normal airway smooth muscle cell length-to-width ratio is >6:1). We coated the plastic dishes with extracellular matrix (ECM) molecules (type 1 collagen, 40 µg/ml) or poly-L-lysine (40 µg/ml) through the holes of the membranes sealed onto the dishes overnight and then lifted off the membrane. After blocking the rest of the dish area with 1% BSA, we plated cells sparsely overnight. Single cells that assumed circular or rectangular shapes of the same area were used for obtaining mechanical measurements.
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RESULTS AND DISCUSSION |
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Mechanical anisotropy depends on stress fiber orientation and cytoskeletal tension.
We examined the relationship between the direction of mechanical loads applied to an elongated cell and the structure of the CSK. We applied an oscillating torque (45 Pa at 0.83 Hz, a physiological load that produced an average cellular surface deformation 0.15 ± 0.03 µm) to a single RGD-coated spherical bead attached to the apical surface of human airway smooth muscle cells via integrin receptors in two different ways: 1) the bead was magnetized (M) in the x direction, and a twisting field B was applied in the z direction: this combination of fields induced a torque T about the y-axis (Ty) (Fig. 2A, left bead); and 2) the bead was magnetized in the y direction, and a twisting field was applied in the z direction: this combination induced a torque T about the x-axis (Tx) (Fig. 2A, right bead).
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We hypothesized that the structure of the CSK might influence mechanical anisotropy. To test this hypothesis, we disrupted the F-actin in the CSK of the cells using two different drugs: latrunculin A (Fig. 2C) and cytochalasin D (Fig. 2C). Within 3 min of adding the drug to the medium, and in the absence of changes in cell shape, exposure to either drug resulted in greatly increased displacements of the bead (at constant magnetic torque) and almost completely abolished mechanical anisotropy (Fig. 2C). These data suggest that F-actin is involved in determining the mechanical anisotropy.
To control the projected area and shape of the cell and to explore further the underlying origin of the observed mechanical anisotropy, we plated cells on micropatterned islands having the same area (2,500 µm2, an area chosen on the basis of normal projected areas of smooth muscle cells), but with different shapes and different length-to-width aspect ratios. We applied a constant magnetic torque to a magnetic bead and measured the displacement both parallel and perpendicular to the long axis of these patterned cells. We define the "anisotropy index" as the ratio of stiffness, that is, the specific torque (torque per bead volume or apparent stress) divided by displacement along the long axis of the cell to the stiffness transverse to the long axis of the cell. An index of 1.0 indicates that the mechanical response is isotropic. The cells plated on ECM-coated rectangular islands (Fig. 3B; aspect ratio 9:1) had anisotropic mechanical behavior, because the deformation was twice as large when the bead rotated across (i.e., transverse to) the long axis of the cell than when the bead rotated along (i.e., parallel to) the long axis (Fig. 3A). The anisotropy index was 2.1 ± 0.3 (P < 0.007 vs. isotropy or index of 1.0) and similar to that of unpatterned, natural cells, which was 2.0 ± 0.2 (P < 0.00003 vs. isotropy) (Fig. 4). In contrast, the stiffness of the cells plated on circular islands of ECM (Fig. 3C; aspect ratio 1:1) did not depend on the direction of the applied torque and thus exhibited no mechanical anisotropy (Fig. 3A). These results are consistent with recent results at our laboratory demonstrating deep cytoskeletal anisotropy detected by differential phase lags in x vs. y directions (17), with a recent report showing that cyclically strained adherent cells are stiffer along the long axis (33), and with the finding of an anisotropic strain in the erythrocyte cortical CSK in response to a large deformation (22). It remains to be seen, however, to what degree these smooth muscle cells in a 3-D culture environment and other elongated nonmuscle cells (e.g., endothelial cells and epithelial cells) exhibit mechanical anisotropy.
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Recently, researchers at our laboratory have shown that endogenous cytoskeletal tension (i.e., prestress) dictates cell stiffness (36, 37). To test further the hypothesis that the tension in the stress fibers and/or actin bundles along the long axis of the cell contributes to the mechanical anisotropy of the cell, we transfected caldesmon into the cell. Caldesmon decreases cytoskeletal tension by inhibiting the interaction between actin and myosin (24). Transfecting a low level of caldesmon into the cell decreased cytoskeletal tension by 50% without altering the shape of the cell, the distribution of focal adhesions, or stress fibers (17, 24). Interestingly, these cells also exhibited lower mechanical anisotropy than the wild type; the anisotropy index decreased from 2.1 to 1.6 ± 0.1 (P < 0.0005 vs. isotropy). This difference represents a 45% decrease in anisotropy (a 100% decrease in anisotropy would be equivalent to an anisotropy index of 1.0) (Fig. 3A, open bars; Fig. 4).
We note that decreasing the cytoskeletal tension by overexpressing caldesmon and abolishing stress fibers by plating the cells on poly-L-lysine decreased the anisotropy index from 2 toward 1 (i.e., isotropy) while decreasing stiffness by a factor of 3 in micropatterned cells (Fig. 4). Similarly, when the CSK was disrupted with drugs, there was a shift from anisotropy toward isotropy, a threefold decrease in stiffness after treating the cells with latrunculin A, and a >10-fold decrease in stiffness after treatment with cytochalasin D in unpatterned cells (Fig. 4). These observations suggest that the common structural origin of mechanical anisotropy in these living cells is the preferential orientation of stress fibers and/or actin bundles along the long axis of the cell.
Anisotropic mechanical signaling to the CSK and to the nucleolus. To explore further the utility of 3-D MTC in intracellular mechanical signaling, using a recently developed synchronous detection method (17), we mapped the cytoskeletal displacement field of an elongated cell (Fig. 5A) with transfected YFP-mitochondria as fiducial markers (17, 36) (Fig. 5B). The load-induced displacement patterns of the CSK were quite different in response to the applied torque of the same magnitude and frequency, but along different directions (compare Fig. 5D in response to Ty with Fig. 5E in response to Tx). Interestingly, when a more complex load was applied with all three oscillatory twisting fields switched on simultaneously with different phase lags (x = 0°, y = 90°, z = 0° in Fig. 5F, or x = 0°, y = 90°, z = 90° in Fig. 5G, respectively), distinctly different displacement patterns emerged (compare Fig. 5F with Fig. 5G). All of these results demonstrate that spatial distributions of the cytoskeletal deformation depend on not only on the magnitude and frequency but also the direction of loading.
We next mapped the displacements of the nucleolus, an intranuclear organelle crucial for ribosomal RNA synthesis (31), in response to the load applied at the cell surface. The nucleolus of an interphase cell is visible under phase-contrast microscopy without staining or GFP (Fig. 5C). The displacement map within the nucleolus varied with the loading direction: changing the torque direction by 90° (from Ty in Fig. 5H to Tx in Fig. 5I) drastically altered displacement patterns of the nucleolus. Applying a complex load, with all three twisting fields switched on simultaneously with different phase lags, led to dramatically different nucleolar displacement maps (compare Fig. 5J with Fig. 5K): concentrated displacements occurred at different sites and pointed in various directions within the nucleolus, demonstrating different intranucleolar deformation patterns. These results suggest that physiological loads applied at the cell surface might be able to regulate and control nuclear functions directly (e.g., synthesis of ribosomal RNA) via the structural pathways of the CSK and the nuclear matrix. The specific biological responses of the nucleus may therefore depend on the direction of mechanical loading. This result, together with the result of cytoskeletal deformation, extends the previous finding of Maniotis et al. (23) and demonstrates that small but physiologically relevant surface deformation applied via integrin receptors along different directions can elicit different mechanical deformation patterns deep in the cytoplasm and in the nucleus.
In summary, we demonstrate that elongated adherent cells exhibit intrinsic mechanical anisotropy. This anisotropy may originate in the prestress residing in stress fibers and/or actin bundles oriented preferentially along the long axis of the cell. We also show that intracellular deformation patterns of the CSK and the nucleolus are sensitive to directions of loading. Much of the current research on the mechanics of the cell focuses on elucidating mechanotransduction pathways at the cell surface and especially in elucidating the role of focal adhesions (12, 29). Our results regarding intracellular anisotropic mechanical signaling suggest that load direction-dependent direct mechanotransduction may also occur deep in the cytoplasm and inside the nucleus via the structural pathways of the CSK and within the nucleus (17, 23). We think that this new mechanical loading device will be a valuable tool for probing mechanisms of mechanotransduction when used in combination with single live cell biochemical assays (30, 38).
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APPENDIX |
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GRANTS |
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ACKNOWLEDGMENTS |
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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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