Exploring the difference in the mechanics of vascular smooth muscle cells from wild-type and apolipoprotein-E knockout mice

Abstract

Atherosclerosis-related cardiovascular diseases are a leading cause of mortality worldwide. Vascular smooth muscle cells (VSMCs) comprise the medial layer of the arterial wall and undergo phenotypic switching during atherosclerosis to a synthetic phenotype capable of proliferation and migration. The surrounding environment undergoes alterations in extracellular matrix (ECM) stiffness and composition and an increase in cholesterol content. Using an atherosclerotic murine model, we analyzed how the mechanics of VSMCs isolated from Western diet-fed apolipoprotein-E knockout (ApoE^−/−) and wild-type (WT) mice were altered during atherosclerosis. Increased stiffness of ApoE^−/− VSMCs correlated with a greater degree of stress fiber alignment, as evidenced by atomic force microscopy (AFM)-generated force maps and stress fiber topography images. On type-1 collagen (COL1)-coated polyacrylamide (PA) gels (referred to as substrate) of varying stiffness, ApoE^−/− VSMCs had lower adhesion forces to COL1 and N-cadherin (N-Cad) compared with WT cells. ApoE^−/− VSMC stiffness was significantly greater than that of WT cells. Cell stiffness increased with increasing substrate stiffness for both ApoE^−/− and WT VSMCs. In addition, ApoE^−/− VSMCs showed an enhanced migration capability on COL1-coated substrates and a general decreasing trend in migration capacity with increasing substrate stiffness, correlating with lowered adhesion forces as compared with WT VSMCs. Altogether, these results demonstrate the potential contribution of the alteration in VSMC mechanics in the development of atherosclerosis.

INTRODUCTION

Despite strides of progress, cardiovascular disease has become the world’s leading cause of mortality, and atherosclerosis is a major contributor to this global epidemic (1). Atherosclerosis is a complex disease described as a chronic state of inflammation resulting in the formation of arterial plaques filled with lipid and cellular debris. Atherosclerotic plaque instability, fibrous cap thinning, and rupture cause a plethora of cardiovascular diseases such as heart attack, stroke, and peripheral vascular disease (2, 3).

Vascular smooth muscle cells (VSMCs) are present in the medial layer of the vasculature providing arterial contraction and extracellular matrix (ECM) production to maintain optimal hemodynamic conditions (4). During the progression of atherosclerosis, VSMCs “respond to injury” and shift from a contractile to a synthetic phenotype to stabilize the plaque and form a fibrous cap. The VSMC phenotypic shift entails the reduction of contractile protein expression and the upregulation of cell proliferation, migration, and secretion of ECM proteins (5, 6). VSMCs detach from neighboring cells and the surrounding ECM to migrate toward the intima during the development of atherosclerosis. Migrating VSMCs experience a wide range of microenvironments, as the ECM within the plaque varies in composition and stiffness (7). Type-1 collagen (COL1) was shown to be abundant near the stiffer fibrous cap but nearly absent within the softer, lipid-rich necrotic core (8, 9). In addition, migrating VSMCs actively modify ECM composition and stiffness through COL1 and fibronectin (FN) deposition (10, 11).

Mechanotransduction at the cell-matrix interface plays a critical role in regulating cell adhesion to the ECM and cell migration (12–18). ECM stiffness has emerged as a prominent mechanical cue that precedes disease and drives its progression, by altering cellular behaviors such as phenotypic shifting (19) and aberrant cell migration in response to disease development (13, 20). Cell-matrix mechanotransduction is regulated by factors including integrin expression and activity (21) and ECM composition (18); the latter has been demonstrated to be a critical determinant of cell behaviors (18, 22, 23). In our recent study, ECM proteins and substrate stiffness (where substrate refers to the material the cells were growing on) were found to synergistically regulate VSMC migration and cortical cytoskeleton organization (24). Moreover, VSMCs from apolipoprotein-E knockout (ApoE^−/−) mice were shown to constitute 30–70% of macrophage marker-positive (CD68) cells (25) and foam cells (26). In humans, ∼30–40% of CD68-positive cells and 50% of foam cells are of VSMC origin (25, 27). VSMCs from ApoE^−/− were shown to have altered structural and functional properties (28–31). Moreover, phenotypically altered VSMCs were shown to metabolize lipids differently to contractile VSMCs, partly due to the reduced cholesterol efflux mediated by ATP-binding cassette transporters and the decreased expression of cholesterol esterase, facilitating foam cell formation (27, 32). Collectively, membrane cholesterol and substrate stiffness were shown to coordinate and induce VSMC cytoskeleton remodeling and alteration of cell mechanics (33).

This study aims to investigate the difference in the mechanics of VSMCs isolated from ApoE^−/− and wild-type (WT) mice. Atomic force microscope (AFM) was used to study N-cadherin (N-Cad)-mediated cell-cell adhesion, integrin-mediated cell-ECM adhesion forces, and stiffness of VSMCs cultured on elastically tunable substrates. AFM was also used to examine live VSMC submembranous cytoskeleton architecture. In addition, we inspected VSMC migration dynamics and global cytoskeleton organization on elastically tunable COL1-coated substrates, mimicking the variation in environmental stiffness VSMCs experience in atherosclerosis. Our results demonstrated a significant difference in cell mechanics, cytoskeletal organization, and migratory behavior of VSMCs isolated from WT and ApoE^−/− mice.

MATERIALS AND METHODS

Mouse Vascular Smooth Muscle Isolation

The mice used in this study were kept in accordance with the National Institutes of Health (NIH) guidelines (8th edition of the Guide for the Care and Use of Laboratory Animals), and the animal use protocol was approved by the Laboratory Animal Use Committee of the University of South Dakota (No. 13-09-15-18 C) and Sanford Institutional Care and Use Committee (No. 153-03-21 C). For this study, male ApoE^−/− (B6.129P2-Apoe^tm1Unc/J, Jackson Laboratory) and male WT (C57BL/6J, Jackson Laboratory) mice were subjected to 10 wk of Western diet after reaching 8 wk of age. To discern the influence of the Western diet on cell mechanics, additional ApoE^−/− and WT mice were fed normal chow for 18 wk for stiffness measurements on a plastic substrate only. Mice were euthanized using carbon dioxide (CO₂) asphyxiation, and VSMCs were enzymatically isolated from the descending thoracic aorta and seeded onto 60-mm plastic dishes (Corning, Corning, NY). Cells were maintained in a DMEM-F12 (Invitrogen) medium supplemented with 10% fetal bovine serum (FBS, ATLANTA Biologicals, Lawrenceville, GA) in a humidified incubator with 5% CO₂ at 37°C (34).

Elastically Tunable ECM Protein-Coated Polyacrylamide Gel Preparation

Elastically tunable COL1-coated PA gel preparation was described in detail in our previous studies (24, 33, 35). Glass-bottom 50-mm dishes (MatTek, Ashland, MA) or glass-bottom six-well plates (Cellvis, Mountain View, CA) were activated using 0.1 N sodium hydroxide at 37°C and grafted with one layer of (3-aminopropyl) triethoxysilane (APTS; Sigma, St. Louis, MO). The aminated glass surface was then grafted with a 25-mm-diameter PA gel using 0.5% glutaraldehyde (Sigma, St. Louis, MO) as a crosslinker between the PA gel and the aminated glass surface. The PA substrate was washed with 50 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid buffer (Thermo Fisher Scientific, Waltham, MA) to remove the unreacted monomer. The PA gel surface was coated with 300 µL of 1 mM sulfosuccinimidyl 6-(4′-azido-2′-nitrophenylamion) hexanoate (sulfo-SANPAH; Thermo Fisher Scientific, Waltham, MA), activated for 10 min under UV light, and quickly washed with phosphate-buffered saline (PBS). This step was repeated to ensure sufficient coating of the crosslinker. After quick washes with PBS, one layer of COL1 (Sigma, St. Louis, MO; 0.15 mg/mL) was grafted to the PA surface. The different elasticities of the gels were previously determined using AFM indentation (35).

Cell Migration Studies

Cells were trypsinized, counted, and plated on elastically tunable COL1-coated PA gels at a density of 2,500 cells/cm². Cells were allowed to attach to the plate in serum-free medium in a humidified incubator with 5% CO₂ at 37°C for 2 h. Afterward, migration experiments were performed using a JuLI stage microscope (NanoEnTek Inc, South Korea). For each PA gel, five to ten regions of interest were chosen and imaged with a ×10 objective every 10 min for 24 h. FIJI (FIJI is Just Image J, NIH, Bethesda, MD) was used to analyze image stacks. Time-lapse image stacks were aligned using the plugin StackReg (Biomedical Imaging Group, Swiss Federal Institute of Technology Lausanne). Manual cell tracking was completed by tracing the position of the cell nucleus using the MTrackJ plugin. For each experiment, 90 cells were tracked from at least three replicates (24, 36).

VSMC Biomechanical Characterization Using AFM

VSMC stiffness, cell-cell adhesion, and cell-ECM adhesion were assessed in real time using an Asylum AFM system (Model MFP-3D-BIO, Asylum Research, Santa Barbara, CA) mounted on an inverted microscope (Model IX81, Olympus America Inc.). A 5-μm-diameter glass microbead was glued to an AFM probe (MLCT-O10-D, Santa Barbara, CA; Bruker Corp.) and used for Young’s modulus (E-modulus) measurement. Cell surface areas of 30 × 30 μm were automatically scanned and indented at 6 × 6 positions with a 0.5 Hz indentation frequency and 1 µm/s approach/retraction velocity. A parabolic Hertz equation was used to estimate VSMC stiffness (35, 37). For adhesion force measurement, cell surfaces were probed at a 0.05 Hz indentation frequency and 0.1 µm/s approach/retraction velocity using an AFM probe (MLCT, Santa Barbara, CA; Bruker Corp.) functionalized with N-cadherin (Human N-Cad, R&D Systems, Minneapolis, MN; 10 µg/mL) or COL1 (1 mg/mL). The AFM force curves were analyzed using a proprietary MATLAB program (R2016a, MathWorks). The product of the AFM probe spring constant and the height of ruptures (adhesion events) on a retraction force curve were identified and used to compute adhesion forces. The total average adhesion force is the product of average adhesion force and number of ruptures. Adhesion and stiffness testing of VSMCs cultured on elastically tunable PA gels were limited to measuring a single position. Primary VSMCs attached to at least two other cells were selected and indented at a site between the cell edge and the nucleus. The thermal noise amplitude method was used to calibrate each AFM probe after each adhesion measurement and before each stiffness measurement experiment (38, 39).

Live VSMC Cytoskeletal Imaging Using AFM and Image Processing

Contact mode AFM imaging was used to assess VSMC cytoskeleton architecture in real time. Using an AFM stylus probe (model MLCT-C, k = 15 pN/nm; Bruker, Santa Barbara, CA, a 30 × 30-μm cell surface area was imaged with the digital density of 512 × 512 pixels. The scanning frequency was 0.3 Hz. The obtained height and deflection images were analyzed using a proprietary MATLAB program to analyze cell fiber orientation and density as described in our previous work (24, 33, 36). Stress fiber area fraction is defined as the ratio of the whole cell surface area compared with that covered by stress fibers. The area fraction was computed from AFM height images post flattening and background noise elimination to improve the contrast between the background (nonstress fiber area) and the foreground (stress fibers). AFM deflection images were used to determine cytoskeletal stress fiber orientation. Finally, cell surface roughness was determined from height images using the built-in function of the AFM Asylum Research software.

Confocal Imaging and Image Processing

VSMCs were passaged and seeded onto COL1-coated PA gels at a density of 10,000 cells/cm². Cells were fixed at 60–80% confluency with 4% paraformaldehyde in phosphate-buffered saline (PBS; Affymetrix, CA) for 20 min at room temperature followed by several rinses with PBS. VSMCs were permeabilized with 0.1% Triton X-100 in PBS for 5 min and rinsed with PBS. F-actin cytoskeleton was stained with a 1:1,000 dilution of phalloidin (Phalloidin-iFluor 488, Abcam, Cambridge, UK) in 1% bovine serum albumin/PBS for 20 min and rinsed with PBS. The nuclei were counterstained with a 1:1,000 dilution of Hoechst 33342 (BD Biosciences, San Jose, CA) dissolved in PBS for 10 min followed by a final rinse with PBS. VSMCs were imaged using a laser scanning confocal microscope (Olympus IX83 FV1200, Olympus Life Science) at a 1024 × 1024-pixel resolution and z-height of 0.38 μm. Z-stacks were flattened and manually segmented by tracing VSMCs in contact with at least one other cell. As previously described, a series of elongated Laplacian-of-Gaussian (eLoG) filters were used to convolve flattened z-stacks to detect total cellular cytoskeletal fiber orientation (24, 40).

Statistical Testing

One-way ANOVA with Tukey’s post hoc test was used to infer statistical significance for all experiments. A value of *P ≤ 0.05, **P ≤ 0.01, and ***P ≤ 0.001 was considered statistically significant. All data are reported as means ± standard error of the mean.

RESULTS

WT and ApoE^−/− VSMC Stiffness and Submembranous Stress Fiber Orientation Measurement Using AFM

Using a 5-μm-diameter glass microbead glued to an AFM probe, 30 × 30-μm cell surface areas were automatically scanned and indented at 6 × 6 positions (Fig. 1A). Normal diet fed ApoE^−/− and WT mice had no significant difference in cell stiffness (Fig. 1B). Stiffness maps for WT and ApoE^−/− VSMCs are presented in Fig. 1C and 1D, respectively, with Western diet-fed ApoE^−/− VSMCs cultured on a plastic plate significantly stiffer than WT VSMCs (Fig. 1E).

Figure 1.

Live wild-type (WT) and apolipoprotein-E knockout (ApoE^−/−) vascular smooth muscle cell (VSMC) stiffness maps. A: 30 × 30-μm cell surface areas were automatically scanned and indented at 6 × 6 positions with a glass bead. B: average stiffness for normal diet-fed ApoE^−/− and WT VSMCs. C, D: stiffness force maps for WT and ApoE^−/− VSMCs, respectively. E: average stiffness for Western diet-fed WT and ApoE^−/− VSMCs. No significant difference in E-modulus was observed between normal diet-fed ApoE^−/− and WT VSMCs. Western diet-fed ApoE^−/− VSMCs had a significantly higher E-modulus compared with WT VSMCs. All data are presented as means ± SE (n ≥ 60 cells across six different mice). Scale bar in lower right corner represents 10 μm. A was created with BioRender.com.

Submembranous stress fiber topography was acquired by scanning a 30 × 30-μm area in contact mode (Fig. 2A). Figure 2B and Fig. 2C are representative three-dimensional 30 × 30-μm cell surface area stress fiber topography of a WT and ApoE^−/− VSMC, respectively. The stress fiber area fraction was unchanged for both cell types, suggesting similar stress fiber area coverage within the cell body (Fig. 2D). The normalized percentage circular histograms along the dominant orientation (Fig. 2, E and F) demonstrate a clear difference in stress fiber orientation between WT and ApoE^−/− VSMCs. For WT and ApoE^−/− VSMCs, Fig. 2G illustrates the percent frequency of average stress fiber orientation, normalized along the dominant angle, fitted with a first-order Gaussian function. The summarized percent frequency of stress fibers oriented around the dominant angle (±20°) indicates that ApoE^−/− VSMCs have significantly greater stress fiber alignment as compared with the WT VSMCs (Fig. 2H). The surface roughness average (the absolute difference in the peak and valley value of measured microscopic surface peaks and valleys of VSMC topographical images) of ApoE^−/− VSMCs was significantly greater than that of WT VSMCs (Fig. 2I).

Figure 2.

Live wild-type (WT) and apolipoprotein-E knockout (ApoE^−/−) vascular smooth muscle cell (VSMC) submembranous stress fiber orientation measurement. A: 30 × 30-μm areas were scanned for overall VSMC stiffness measurement. B, C: representative 30 × 30-μm three-dimensional stress fiber topography of WT and ApoE^−/− VSMCs used for live VSMC stress fiber orientation and area fraction analysis. D: average area fraction of VSMC submembranous actin stress fibers. E, F: circular histogram showing the normalized stress fiber orientation of WT and ApoE^−/− deflection images, respectively. G: the average stress fiber orientation percentage histogram for WT and ApoE^−/− VSMCs, in which the dominant stress fiber orientation angle was set as zero degrees for each cell. H: the summarized percentage frequency of dominant fiber orientation (−20° ∼ +20°). I: average stress fiber surface roughness average (RA). ApoE^−/− VSMCs have significantly greater stress fiber alignment and surface roughness compared with WT VSMCs. All data are presented as means ± SE (n= 16 cells across six different mice). A was created with BioRender.com.

Stiffness, Cell-Cell Adhesion, and Cell-ECM Adhesion of WT and ApoE^−/− VSMCs on COL1-Coated Gel Substrates

WT and ApoE^−/− VSMCs were cultured on elastically tunable COL1-coated gels and probed with the AFM to measure stiffness (Fig. 3A). The stiffness of these substrates was previously measured with AFM and determined to be 28 and 103 kPa (35). ApoE^−/− VSMCs cultured on 28 and 103 kPa substrates had a significantly higher E-modulus compared with WT VSMCs, and those cultured on the stiffer substrate had a higher E-modulus for both ApoE^−/− and WT VSMCs (Fig. 3B).

Figure 3.

Measuring stiffness of wild-type (WT) and apolipoprotein-E knockout (ApoE^−/−) vascular smooth muscle cells (VSMCs) on type-1 collagen (COL1)-coated gel substrates. A: WT or ApoE^−/− VSMCs were cultured on elastically tunable COL1-coated polyacrylamide (PA) gels and indented with a glass bead at a single point. B: average E-modulus of VSMCs on 28 and 103 kPa substrates. ApoE^−/− VSMCs cultured on 28 and 103 kPa substrates had a significantly higher E-modulus compared with WT VSMCs and demonstrated increased E-modulus when cultured on stiffer substrates. All data are presented as means ± SE (n ≥ 60 cells across six different mice for each group). AFM, atomic force microscope. A was created with BioRender.com.

Cell-ECM adhesion and cell-cell adhesion were investigated with COL1- or N-Cad-coated stylus probes (Fig. 4A). Cell-ECM average adhesion force of ApoE^−/− VSMCs on the 28 and 103 kPa substrates was significantly lower compared with that of WT VSMCs, with both WT and ApoE^−/− VSMCs exhibiting slightly reduced adhesion force on the 103 kPa substrate (Fig. 4B). Cell-cell adhesion was measured using an AFM stylus probe coated with N-Cad (Fig. 4A). ApoE^−/− VSMCs had a significantly lower adhesion force to N-Cad compared with WT VSMCs on the 28 and 103 kPa substrates. The average adhesion force increased slightly with increased substrate stiffness for both WT and ApoE^−/− VSMCs (Fig. 4C). Cell counting of VSMC nuclei showed a slight decreasing trend in the adhesion rate of ApoE^−/− compared with WT VSMCs after allowing to attach to substrate for 30 min (Supplemental Fig. S1; all supplemental material is available at https://doi.org/10.6084/m9.figshare.20682595). However, the difference was not statistically significant (P > 0.05).

Figure 4.

Measuring cell-extracellular matrix (ECM) adhesion and cell-cell adhesion of wild-type (WT) and apolipoprotein-E knockout (ApoE^−/−) vascular smooth muscle cells (VSMCs) on type-1 collagen (COL1)-coated gel substrates. A: atomic force microscopic (AFM) tips were coated with ECM proteins or cadherin and used to measure adhesion forces. B: total average adhesion force to cell-ECM adhesion protein COL1. C: average adhesion force to N-cadherin (N-Cad). ApoE^−/− VSMCs demonstrated a significantly lower adhesion force to COL1 and N-Cad compared with WT cells. All data are presented as means ± SE (n ≥ 60 cells across six different mice for each group). A was created with BioRender.com.

WT and ApoE^−/− VSMC Migration Dynamics on COL1-coated Gel Substrates

WT and ApoE^−/− VSMC migrations were measured on 28 and 103 kPa COL1-coated PA gels (Fig. 5A). The average migration distance over time was significantly greater for ApoE^−/− VSMCs compared with WT VSMCs on both 28 and 103 kPa substrates (Fig. 5, B and C). However, substrate stiffness did not have a statistically significant effect on migration for both WT and ApoE^−/− VSMCs (Fig. 5D).

Figure 5.

Wild-type (WT) and apolipoprotein-E knockout (ApoE^−/−) vascular smooth muscle cell (VSMC) migration dynamics on type-1 collagen (COL1)-coated gel substrates. A: VSMCs isolated from WT and ApoE^−/− mice were seeded onto COL1-coated polyacrylamide (PA) gels to track cell migration. B, C: average distance vs. time for WT and ApoE^−/− VSMCs on the 28 and 103 kPa COL1-coated substrates, respectively. D: migration distance of VSMCs on different substrate stiffnesses. ApoE^−/− VSMCs demonstrated a higher migration capacity compared with WT cells. All data are presented as means ± SE (n = 90 cells across three different mice for each group). A was created with BioRender.com.

WT and ApoE^−/− VSMC Global Cytoskeleton Architecture

Global cytoskeletal architecture was characterized using confocal microscopy through the study of stress fiber orientation in WT and ApoE^−/− VSMCs on 28 and 103 kPa COL1-coated gel substrates. Fluorescent actin cytoskeleton z-stack images, represented in Fig. 6 for WT and ApoE^−/− VSMCs for 28 and 103 kPa COL1 substrates (Fig. 6, A, B, C, and D, respectively), were used to generate the corresponding segmented stress fiber orientation color maps (Fig. 6, E, F, G, and H) using a proprietary MATLAB program. On both 28 and 103 kPa COL1-coated substrates, ApoE^−/− VSMCs exhibited a high degree of intracellular color map uniformity, indicating close alignment of actin filaments, whereas more varied coloration within WT VSMCs indicates greater dispersion of actin filaments (Fig. 6, E–H). This observation was consistent with qualitative analysis of F-actin orientation, as illustrated by circular histograms with a tighter grouping around the dominant orientation angle for ApoE^−/− VSMCs on both substrates compared with WT VSMCs, especially on the 103 kPa substrate (Fig. 6, I–L). Normalized fiber orientation histograms confirmed increased frequency of fiber orientation about the dominant angle for ApoE^−/− VSMCs compared with WT VSMCs, with a slightly higher peak on the 28 kPa substrate and a more pronounced peak on the 103 kPa substrate (Fig. 6, M and O). ApoE^−/− VSMC global stress fiber alignment around the dominant orientation angle was significantly greater than that of WT VSMCs on the 103 kPa substrate, with the ApoE^−/− VSMCs showing a more orientated structure and the WT VSMCs showing a more dispersed structure (Fig. 6P). No significant difference in stress fiber orientation was seen on the 28 kPa substrate, although ApoE^−/− orientation had a slightly higher average (Fig. 6N).

Figure 6.

Confocal imaging of wild-type (WT) and apolipoprotein-E knockout (ApoE^−/−) vascular smooth muscle cell (VSMC) cytoskeleton orientation. A–D: representative fluorescent z-stack images of the actin cytoskeleton for WT and ApoE^−/− VSMCs on the 28 and 103 kPa type-1 collagen (COL1)-coated substrates. E–H: the corresponding segmented color maps showing cytoskeleton orientation computed from the confocal images. I–L: corresponding circular histograms showing the normalized F-actin fiber orientation. M, O: average normalized fiber orientation where the dominant angle is set as zero degrees on the 28 and 103 kPa substrates, respectively. N, P: summarized percentage frequency of fibers orientated around the dominant angle. ApoE^−/− VSMCs on the 103 kPa substrate demonstrated a greater cytoskeletal alignment compared with WT cells. All data are presented as means ± SE (n > 60 cells from at least 25 images across three different mice for each group). Scale bars in lower right corner represent 50 μm.

DISCUSSION

We have previously demonstrated the effects of cholesterol and substrate stiffness on VSMC biomechanics, including changes in cytoskeletal organization, cellular stiffness, adhesion forces, migration, and vasoactivity through in vitro manipulation of cholesterol using methyl-β-cyclodextrin and statins in rat VSMCs (24, 33, 36). We sought to expand these studies and use a murine atherosclerosis model. ApoE^−/− mice exhibit hypercholesterolemia and spontaneous development of atherosclerosis with a Western diet (41). Hypercholesterolemia from the Western diet significantly altered VSMC biomechanics, as normal diet ApoE^−/− and WT VSMCs had no significant difference in elastic modulus, whereas VSMCs isolated from ApoE^−/− mice fed a Western diet exhibited a higher elastic modulus compared with WT VSMCs, an effect that was also seen with increased substrate stiffness (33). ApoE^−/− VSMCs had a more aligned arrangement of cortical actin as well as lowered adhesion for both N-Cad and COL1 compared with WT. In addition, ApoE^−/− VSMCs migrated further than the WT on both 28 and 103 kPa substrates and exhibited greater cytoskeletal alignment, particularly on the stiffer substrate.

These results largely corroborate our previously reported results, providing further insight into VSMC behavior during the progression of atherosclerosis. ApoE^−/− VSMCs were stiffer than WT VSMCs which is consistent with the enhanced alignment of cytoskeletal stress fiber orientation. The cytoskeleton has been shown to be a major contributor to cell stiffness, and actin disruption resulted in decreased stiffness (42). Furthermore, several studies in erythrocytes have shown that increased cholesterol promotes tighter membrane cytoskeleton linkage and increased resistance to cell lysis (43–45). Other studies have implicated ezrin, radixin, and moesin (ERM), which link the membrane to the actin cytoskeleton, to influence cytoskeletal architecture and stiffness, though their exact function has yet to be elucidated in VSMCs as ERM function appears to differ between cell types (46). However, cholesterol has been shown to regulate ERM proteins. During adipogenic differentiation, cholesterol enrichment upregulated phosphorylated moesin and downregulated phosphorylated ezrin, decreasing stiffness while cholesterol depletion had the opposite affect (47). We assessed total ERM, total phospho-ERM, ezrin, moesin, myosin light chain 2, and myosin light chain kinase (MLCK) expression by immunoblotting and found no significant difference in expression between ApoE^−/− and WT VSMCs (Supplemental Fig. S2). However, some interesting trends were observed, particularly for ezrin and MLCK that are worth further investigation, particularly with nonpassaged cells (Supplemental Fig. S2A). In addition, the cytoskeleton is connected to the ECM through focal adhesions that physically couple cells to the matrix (34). In our previous work, we showed that both membrane cholesterol and substrate stiffness coordinate to induce the remodeling of the cytoskeleton and alter VSMC integrin-mediated biomechanics (33). Thus, increased VSMC stiffness with increased substrate stiffness could be attributed in part to enhanced transduction of tension from substrates to the cell surface and VSMC cholesterol content.

ApoE^−/− VSMCs had a significantly greater migration distance compared with WT VSMCs on each of the different substrate stiffnesses. This behavior aligns with VSMCs demonstrating enhanced proliferation and migration with the progression of atherosclerosis following the switch from a contractile to synthetic phenotype. Enhanced cell migration distance is likely the result of a combination of changing adhesion forces and cytoskeleton dynamics. Increased plasma membrane cholesterol content has been associated with reduction in maximum protrusion force and subsequent increase in protrusion length in human embryonic kidney cells during optical tweezer manipulation (42). As the formation of filopodia and lamellipodia drives cell migration, the ability to easily form longer protrusions could partially explain ApoE^−/− VSMC behavior. Ezrin has also been demonstrated to influence migration and cytoskeleton stiffness with constitutively activated ezrin resulting in greater migration and cytoskeleton stiffness (48). Adhesion to the ECM through focal adhesion complexes is another critical component to cell migration. A significant reduction in adhesion forces to N-Cad and COL1 was observed in ApoE^−/− VSMCs for each substrate stiffness compared with WT VSMCs. Although not significant, we observed an increase in cell adhesion force to N-Cad with increased substrate stiffness in both ApoE^−/− and WT VSMCs. ApoE^−/− VSMCs exhibited reduced N-Cad adhesion force and greater migration compared with WT VSMCs, consistent with chemotaxis migration assays that suggest N-Cad has an anti-migratory effect and that downregulation of N-Cad promotes cell migration (49, 50). Alternatively, ApoE^−/− VSMCs exhibited lower COL1 adhesion force and increased migration compared with WT VSMCs. With increased substrate stiffness from 28 kPa to 103 kPa, ApoE^−/− VSMC migration increased while COL1 adhesion was reduced. Bangasser et al. proposed a cell migration model wherein substrate stiffness modulates cell migration dependent upon the number of motors and clutches, for which an optimal range promotes fast migration by nonlinear trends (20).

Conclusion

In summary, hypercholesterolemia from the Western diet likely had a significant causal effect on the observed difference in cell biomechanics. ApoE^−/− VSMCs had a higher stiffness compared with WT VSMCs as a result of greater cytoskeleton stress fiber alignment. Increasing substrate stiffness had a synergistic affect, increasing cell stiffness for both WT and ApoE^−/− VSMCs, though to a greater degree for ApoE^−/− VSMCs, and increasing cytoskeleton stress fiber alignment in ApoE^−/− VSMCs. Adhesion forces to N-Cad and COL1 were lower for ApoE^−/− VSMCs compared with WT VSMCs, associated with the increased migration of ApoE^−/− VSMCs compared with WT VSMCs. These results support our hypothesis that atherosclerosis alters the mechanical properties of VSMCs and provide an insight into underlying mechanisms that may lead to future novel therapeutic approaches.

SUPPLEMENTAL DATA

Supplemental Figs. S1 and S2: https://doi.org/10.6084/m9.figshare.20682595.

GRANTS

This work was supported, in part, by the National Science Foundation Grant 2127031 (to Z.H.) and the National Institutes of Health Grant R15HL147214 (to Z.H.).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

A.P.R., H.J.S., C.K., and Z.H. conceived and designed research; A.P.R., H.J.S., C.K., N.K., H.V., and Z.H. performed experiments; A.P.R., H.J.S., C.K., N.K., H.V., and Z.H. analyzed data; A.P.R., H.J.S., C.K., and Z.H. interpreted results of experiments; A.P.R., H.J.S., C.K., and Z.H. prepared figures; A.P.R., H.J.S., C.K., and Z.H. drafted manuscript; A.P.R., H.J.S., C.K., N.K., H.V., and Z.H. edited and revised manuscript; A.P.R., H.J.S., C.K., N.K., H.V., and Z.H. approved final version of manuscript.