Actomyosin Cortical Mechanical Properties in Nonadherent Cells Determined by Atomic Force Microscopy

Abstract

The organization of filamentous actin and myosin II molecular motor contractility is known to modify the mechanical properties of the cell cortical actomyosin cytoskeleton. Here we describe a novel method, to our knowledge, for using force spectroscopy approach curves with tipless cantilevers to determine the actomyosin cortical tension, elastic modulus, and intracellular pressure of nonadherent cells. We validated the method by measuring the surface tension of water in oil microdrops deposited on a glass surface. We extracted an average tension of T ∼ 20.25 nN/μm, which agrees with macroscopic experimental methods. We then measured cortical mechanical properties in nonadherent human foreskin fibroblasts and THP-1 human monocytes before and after pharmacological perturbations of actomyosin activity. Our results show that myosin II activity and actin polymerization increase cortex tension and intracellular pressure, whereas branched actin networks decreased them. Interestingly, myosin II activity stiffens the cortex and branched actin networks soften it, but actin polymerization has no effect on cortex stiffness. Our method is capable of detecting changes in cell mechanical properties in response to perturbations of the cytoskeleton, allowing characterization with physically relevant parameters. Altogether, this simple method should be of broad application for deciphering the molecular regulation of cell cortical mechanical properties.

Figure 1

Free body diagram of the top section of a nonadherent cell. The applied cantilever normal force (F) is balanced by: 1) the net contractile force generated from the cortical tension (T) and 2) the hydrostatic pressure excess (P) of the incompressible cytosolic fluid. The gap between the cytosolic fluid and the actomyosin cortex is not real, but just illustrates that hydrostatic pressure acts in all directions. Note that the substrate does not contribute to the force balance if tension and pressure balance are taken in the upper-half of the cell. In the actomyosin cortex zoom, the red filaments illustrate the myosin II minifilaments and the green filaments are the actin filaments. To see this figure in color, go online.

Figure 2

Estimation of surface tension and hydrostatic pressure of water-in-oil microdrops. (A) Schematic representation of a water-in-oil microdrop deposited on glass being slightly deformed by a tipless AFM cantilever. When deforming the microdrop, changes in laser (red line) location in the photodetector are acquired and transformed to deflection in length units. The depicted parameters represent: d is the cantilever deflection, Z is the piezo movement, k_c is the cantilever spring constant, and R is the microdrop radius. (B) Acquired force-distance curve on a water microdrop. (Inset) Bright field image of the microdrops to record location as well as determine the radius. The red slope in the plot shows the linear region that is fitted to determine the surface tension and the hydrostatic pressure of each microdrop. (C) Determined surface tension for nontilting and 10° tilting conditions. No significant statistical differences were found between both conditions (p > 0.05). The red dashed line represents the previous reported macroscale measurement for surface tension of T = 20 nN/μm, which is in excellent agreement with our measurements. To see this figure in color, go online.

Figure 3

Nonadherent HFF cells shape is approximately spherical. (A) A representative midplane cross-section view showing the circular shape of a primary HFF cell stained with fluorescently conjugated wheat germ agglutinin for marking the glycocalyx on the plasma membrane imaged by confocal microscopy. (B) Z-stacks were collected to show the spherical shape of the nonadherent cells, satisfying an important assumption for the method to be valid on nonadherent cells.

Figure 4

Applicability of the method to nonadherent HFF cells. (A) Schematic representation of a nonadherent cell being slightly deformed by a tipless AFM cantilever. When deforming the spherical cell, changes in laser location in the photodetector are acquired and transformed to deflection in length units. F is the applied normal force, d is the cantilever deflection, Z is the piezo movement, k_c is the cantilever spring constant, R is the initial cell radius, and h is the cortical actin thickness. (B) Velocity-dependent compression force curves performed on the same HFF cell. Successive curves show negligible viscous losses with negligible deviation from each other for deformations <∼400 nm. (C) Typical stress-relaxation curve performed on a nonadherent untreated HFF cell. The tipless cantilever approaches and is pushed against the spherical cell until a deformation of ∼500 nm is reached, then the cantilever is held at a constant height for 10 s. The marked area shows the comparison of rapid compression 4 μm/s (equal to 1 s compression of cell) to the region of fast decay in force immediately after holding the cantilever height constant. The decay is moderately small, ≤20% the maximum force. (D) Acquired force-distance curve on a HFF cell. (Inset) Bright field image of the HFF cells to identify the location and viability as well as to determine the cell radius. The red slope in the plot shows the linear region that is fitted to determine the actin cortex tension and the intracellular pressure of each cell. (E) Cortical actomyosin tension of untreated HFF cells extracted at different Z-piezo distances from 0 to 600 nm. It can be observed that from 100 to 400 nm these are not statistically different (p > 0.05) in extracted cortex tension, confirming that our method is robust for quantitatively estimating the mechanical properties for small deformations. For Z-distance of 500 nm or greater, the estimated tension increases, demonstrating significant differences using a one-way analysis-of-variance test (p < 0.05). (F) Cortical actomyosin tension of untreated HFF cells measured at different spreading stages. The progressive cell spreading and adhesion is observed by phase contrast (inset). At early stages (time 0–20 min) there are no statistically significant differences (p > 0.05) in extracted cortical tension, confirming that weak adhesion contributions are negligible. However, when cells exhibit lamellipodia and filopodia projections, the measured tension significantly increases—demonstrating that adhesion contribution cannot be ignored at advanced spreading stages (time >30 min). To see this figure in color, go online.

Figure 5

Determination of nonadherent HFF cells’ cortical actomyosin tension and intracellular pressure after pharmacological drug treatments. (A) Cortical actomyosin tension after drug treatments perturbing the cortex. (B) Intracellular pressure after treatments. All treated cases were found to have statistically significant differences versus untreated cases (p < 0.05). To see this figure in color, go online.

Figure 6

Determination of actin cortex thickness and elastic moduli after pharmacological drug treatments. (A) A representative fixed nonadherent HFF cell, transiently transfected with the plasma membrane marker FusionRed-CAAX and labeled for F-actin with Alexa-Fluor 488 phalloidin, were imaged by confocal microscopy. (B) Normalized fluorescence intensity peaks of actin and membrane extracted by line scan after background corrections. (C) Calculated actin cortex thickness, h, between cells after drug treatments using the method of Clark et al. (22). All cases were found to have no statistically significant differences (p > 0.05), except for LatA treatments (p < 0.05). (D) Cortical actomyosin elastic modulus after treatments perturbing the cortex. All cases were found to have statistically significant differences (p < 0.05), except for LatA treatments (p > 0.05). To see this figure in color, go online.