Statin mediated cholesterol depletion exerts coordinated effects on the alterations in rat vascular smooth muscle cell biomechanics and migration

Abstract

Not only does cholesterol induce an inflammatory response and deposits in foam cells at the atherosclerotic plaque, it also regulates cellular mechanics, proliferation, and migration in atherosclerosis progression. Statins are HMG-CoA reductase inhibitors that are known to inhibit cellular cholesterol biosynthesis and are clinically prescribed to patients with hypercholesterolemia or related cardiovascular conditions. Nonetheless, the effect of statin mediated cholesterol management on cellular biomechanics is not fully understood. In this study, we aimed to assess the effect of fluvastatin-mediated cholesterol management on primary rat vascular smooth muscle cell (VSMC) biomechanics. Real-time measurement of cell adhesion, stiffness and imaging were performed using atomic force microscopy (AFM). Cellular migration on extra cellular matrix (ECM) protein surfaces was studied by time-lapse imaging. Effect of changes in VSMC biomechanics on aortic function was assessed using an ex-vivo myograph system. Fluvastatin-mediated cholesterol depletion (−27.8%) lowered VSMC migration distance on fibronectin (FN) coated surface (−14.8%) but not on type 1 collagen (COL1) coated surface. VSMC adhesion force to FN (+33%) and integrin α5 expression were enhanced but COL1 adhesion and integrin α2 expression were unchanged upon cholesterol depletion. In addition, VSMC stiffness (−46.6%) and ex-vivo aortic ring contraction force (−40.1 %) were lowered and VSMC actin cytoskeletal orientation was reduced (−24.5%) following statin mediated cholesterol depletion. Altogether, it is concluded that statin mediated cholesterol depletion may coordinate VSMC migration and adhesion to different ECM proteins and regulate cellular stiffness and cytoskeletal orientation, thus impacting the biomechanics of the cell and aortic function.

Introduction

Atherosclerosis has been defined as a disease of aging, arterial wall remodeling, and prolonged inflammation (Libby et al., 2011; Basatemur et al., 2019) and is the most common cause of death worldwide. Clinical disease manifestations of atherosclerosis are highlighted by lipid-rich necrotic core formation, core rupture, erosion, and thrombosis. Although cholesterol is a major contributing factor in fatty atherosclerotic lesions, it also participates in regulating cellular migration (Yan et al., 2011), mechanics (Khatibzadeh et al., 2013), and spreading (Hong et al., 2013). Vascular smooth muscle cells (VSMCs) are the most prominent cellular component of blood vessels and play important roles during vascular lesion formation and progression. VSMCs residing in atheroma sites possess a multitude of receptors responsible for modified lipoprotein trafficking (Johnson, 2014; Chistiakov et al., 2016). The down-regulation of cholesterol efflux mediators and reverse cholesterol transport in tandem promote and contribute to VSMCs derived foam cell formation (Choi et al., 2009). Preceding studies validated the presence of excess cholesterol loaded into VSMCs and demonstrated that approximately 35% of total foam cells in advanced human coronary atherosclerosis were of VSMC origin (Allahverdian et al., 2012; Allahverdian et al., 2014). Furthermore, VSMCs isolated from atherosclerotic rabbits showed an enriched membrane cholesterol composition (Chen et al., 1995).

3-Hydroxy-3-methyl-glutaryl coenzyme A (HMG-CoA) reductase inhibitors, also known as statins, block cholesterol synthesis via inhibition of HMG-CoA reductase, the rate-limiting enzyme in the mevalonate pathway. While statin therapy is known to decrease the incidence of clinical cardiovascular disease through the lowering of serum cholesterol levels (Scandinavian Simvastatin Survival Study, 1994; Shepherd et al., 1995), evidence suggests statins exert additional benefits including decreased VSMC proliferation (Corsini et al., 1996; Bellosta et al., 2004). Apart from lowering cholesterol biosynthesis, statins also halt the synthesis of various metabolic intermediates associated with the mevalonate pathway, such as isoprenoid synthesis (Ward et al., 2019). These cholesterol-independent effects of statins potentially mediate pleiotropic effects on cardiovascular physiology by modifying plaque stability, signaling pathways, platelet aggregation, endothelium, and VSMC function (Oesterle et al., 2017). During atherosclerosis, VSMCs undergo a shift from a contractile to a synthetic phenotype, disengage from neighboring cells and extracellular matrix (ECM), and migrate towards the inflammatory site. Sakamoto et al. demonstrated that fluvastatin prevented VSMC hyperplasia, proliferation, and phenotype modulation in rat tail arteries (Sakamoto et al., 2005). VSMCs are also mechanoadaptive as they exhibit continuous and coordinated changes in cellular components and cytoskeletal architecture in response to extracellular mechanical cues (Martinez-Lemus et al., 2009). Moreover, VSMCs experience noteworthy cytoskeletal architecture remodeling as their phenotypic shift progresses. In turn, alterations in VSMCs cytoskeletal architecture orchestrate their contraction and adhesion to the ECM (Sehgel et al., 2013; Sehgel et al., 2015; Xie et al., 2018). Recently we demonstrated that membrane cholesterol and substrate stiffness coordinate to induce the remodeling of the cytoskeleton and alter the biomechanics of VSMCs (Sanyour et al., 2019). Significant work has been undertaken regarding the effects of statins on cholesterol and lowering cardiovascular events (Ramos et al., 2018).

In this study, we aimed to exploit the underlying effects of statin mediated cholesterol depletion on the coordination of primary cultured aortic VSMC biomechanics and cytoskeletal architecture. VSMC sub-membranous actin and total actin orientation were evaluated using both atomic force microscopy (AFM) and confocal microscopy, respectively. Migration of VSMCs on ECM proteins collagen-I (COL1) or fibronectin (FN) coated substrates were assessed. VSMC stiffness and cell-ECM adhesion forces were characterized using AFM and compared to cell adhesion molecules expression. In addition, the effect of statin mediated cholesterol depletion on native thoracic aorta cholesterol levels and its contractile response to a vasoconstrictor was evaluated.

Methods

Ethical Approval

Male Sprague-Dawley rats were used for this study and were maintained in accordance with the NIH guidelines (The Eighth Edition of the Guide for the Care and Use of Laboratory Animals (NRC 2011). Animal use in these studies was approved by the Laboratory Animal Use Committee of the University of South Dakota and Sanford Research (#13-09-15-18C) and conform to the ethical principles and regulations outlined by The Journal of Physiology (Grundy, 2015). Rats at 200-250g were euthanized using carbon dioxide (CO₂) narcosis at a fill rate of approximately 30% of the chamber volume per minute of CO₂. Following euthanasia, death was confirmed by bilateral pneumothorax. Afterwards, the thoracic rat aorta was surgically removed and collected.

Primary Rat VSMC Isolation and Culture

Following a previously published protocol (Wu et al., 1998; Sanyour et al., 2018), isolated cells were placed and cultured in 60 mm polystyrene tissue culture dishes (Corning Life Sciences, Tewksbury, MA) and kept in a humidified incubator (Thermo Fisher Scientific, Waltham, MA) with 5% CO₂ at 37°C. Cells were incubated with DMEM/F-12 medium supplemented with 10% FBS (Atlanta Biologicals, Flowery Branch, GA). All experiments were conducted at 60-80% cell confluency and were maintained in primary culture for 3-7 days without passaging. Before conducting AFM experiments, cells were incubated with CO₂ independent medium (Gibco, cat#18045088) for at least 30 min in a 37°C humidified incubator. All AFM experiments were conducted at room temperature in CO₂ independent medium for a 1-2 h duration (Sanyour et al., 2018).

Rat VSMC and Aortic Ring Cholesterol Manipulation and Quantification

(3R, 5S, 6E)-rel-7-[3-(4-Fluorophenyl)-1-(1-methylethyl)-1H-indol-2-yl]-3,5-dihydroxy-6-heptenoic acid monosodium salt (Fluvastatin sodium, Sigma-Aldrich, St. Louis, MO. cat# 93957-55-2) was used to manipulate primary cultured VSMC cholesterol. 30-40% confluent primary VSMCs were starved with 0.4% FBS for 48 h, treated with 1 μM fluvastatin supplemented with 10% FBS for 72 h, and kept in a humidified incubator (Thermo Fisher Scientific, Waltham, MA) with 5% CO₂ at 37°C (Corsini et al., 1993; Bellosta et al., 2004). Surgically harvested aortas were cut into approximately 2 mm long aortic ring segments and cultured for 72 h in 1 μM fluvastatin supplemented with 10% FBS followed by cholesterol quantification and contractile response assessment. Gas chromatography/mass spectrometry (GC/MS) was used to analyze VSMC cholesterol content. 3.5-7.5 × 10⁵ VSMCs were rinsed in PBS and flash frozen on dry ice. Cell pellets were reconstituted in 1 mL of water and lysed by successive freeze/thaw cycles in a 37°C bead bath. 50 μL per sample was removed for protein quantification (Micro BCA Protein Assay, Pierce Biotechnology). 1 mL saponification buffer containing 92% ethanol (Fisher Scientific), 7% KOH (Fisher Scientific), and 1% coprostan-3-ol (Abcam, ab143882) was added to 900 μL sample. Samples were saponified while shaking at 60°C for 1 h. After the addition of 1 mL water to each sample, the aqueous phase was extracted with 3 mL EtOAc (Sigma). Samples were vortexed, centrifuged at 2200 rpm for 5 min, and the organic phase was removed. A second extraction was performed with an additional 2 mL of water. The organic phase was concentrated to dryness by heating at 50°C under nitrogen flow and the residue was dissolved in 50 μL pyridine (Sigma). Extracted lipids were derivatized by the addition of 50 μL N,O-bis(trimethylsilyl) trifluoroacetamide with 1% trimethylchlorosilane (Sigma) and incubated at 60°C for 1 h. Sterol levels were determined by GC/MS analysis as previously described (Kelley, 1995). Samples were analyzed by automatic injection of 1 μL of the derivatized sterol mixture into an Agilent 7890 GC using a 1:5 split injection port (4 mm ID × 78.5 mm quartz wool liner, Restek, 23309) leading to a 0.18 mm x 20 m 1,4-bis(dimethylsiloxy)phenylene dimethyl polysiloxane column (Restek, 43602). Helium was used as a carrier gas at a linear rate of 46.9 cm/sec. After 0.5 min at 170°C, the oven temperature was raised to 250°C at 18°C/min, then to 280°C at 3°C/min, and finally to 320°C at 20°C/min and held for 7 min. An Agilent 5977B mass spectrometer was operated in the electron impact mode at 70 eV with anion source temperature of 275°C. Analysis was performed using MassHunter software. Peak identification for cholesterol (RT ~13.9) and coprostan-3-ol (internal standard, RT ~12.7) was based on retention times from known standards (Avanti Polar Lipids, Inc) as well as comparison of fragmentation patterns to the National Institute of Standards and Technologies Standard Reference Database. For peak quantitation, the total cholesterol peak area was normalized to coprostanol area, protein concentration, and expressed relative to wild-type samples using GraphPad Prism statistical software. Details of native rat aorta contractile response measurement upon cholesterol depletion and cholesterol quantification method were previously described (Sanyour et al., 2019).

Rat Aortic Ring Stiffness Testing

5 mm long segments were cut from descending rat aortas and treated with 1 μM fluvastatin in 10% FBS + DMEM for 72 h. After treatments, segments were washed twice with PBS and loaded onto the MTS Insight mechanical tester (MTS Systems, USA). Segments were stretched to 2 mm as a baseline and stretched several times to 0.2 N load to precondition the vessels. Final mechanical testing was done with a stretch rate of 10 mm/min and sampling rate of 10 Hz. From the resulting stress-strain curves a linear section was chosen and linear regression was done to get a line of best fit. The slope of the line of best fit was the elastic modulus of the aorta. Vessel wall thickness was measured on an Olympus IXplore Pro microscope system (Olympus life science, USA) for an average thickness of 175 μm and the sample thickness was set to 350 μm for our testing set up.

VSMC Migration Tracking

50 mm glass bottom dishes (MatTek Corporation, Ashland, MA) were coated with 0.15 mg/mL collagen I (COL1, Millipore-Sigma, St. Louis, MO cat# 11179179001) or fibronectin (FN, ThermoFisher Scientific, cat# 33016015) for 4 h followed by several washes with 50 mM 4-(2-hydroxyethy)-1-peperazinessthanesulfonic acid buffer (HEPES, ThermoFisher Scientific, cat# 15630080). Treated cells were trypsinized, counted, and plated 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. Migration experiments were carried out using a JuLI Stage Microscope (NanoEnTek Inc, South Korea). For each dish, five to ten regions of interest were chosen and imaged with a 10× objective every 10 minutes for 24 h. FIJI (FIJI is Just Image J) (NIH, Bethesda, MD, USA) 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 tracking was done using the built-in program MtrackJ to trace the position of the nucleus. For each experiment, 50 cells were tracked and cells with a maximum displacement of less than 10 μm were excluded in this analysis.

Live VSMC Stiffness and Cell-ECM Adhesion Measurement Using AFM

Real-time adhesion and stiffness properties of VSMCs were performed 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 the tip of an AFM probe (MLCT-O10-D, Santa Barbara, CA; Bruker Corp.) and used for Young’s modulus (E-modulus) measurement. For estimating the E-modulus of the VSMC cortex, a parabolic Hertz equation (eq. 1) was used to fit the 150-300 nm length of approaching AFM force curve immediately after the VSMCs cortex contact point.

$$E=\frac{3}{4}\cdot \frac{1-v^2}{\sqrt{r_b}}\cdot \frac{F}{{\delta }^{\frac{3}{2}}}$$ (1)

Where r_b is the radius of the bead at 2500 nm, F is the force exerted on the cell, and v is the Poisson ratio assumed to be 0.5 for a cell.

An AFM probe (MLCT-C, Santa Barbara, CA; Bruker Corp.) was coated with COL1 or FN for adhesion force measurement using the protocol described previously (Hong et al., 2015). Briefly, polyethylene glycol (PEG, Sigma, St. Louis, MO) was used as a linker molecule between ECM proteins and the AFM probe. The probe was submerged in 10 mM of PEG for 5 minutes, rinsed with Dulbecco’s phosphate buffered saline (DPBS) three times, and then incubated with 50 μl of COL1 (0.1 mg/ml) or FN (0.1 mg/ml) solution for 2 min at room temperature followed by three times PBS rinse. Prior to the experiment, the AFM cantilever was submerged in CO₂ independent medium for approximately 30 min to equilibrate the AFM system and minimize drifting. Primary VSMCs attached to at least two other cells were selected for AFM experiment and the site between their nucleus and cell edge were indented. For cell stiffness measurements, a 6×6 position of a 40×40 μm cell surface area were automatically scanned and indented with a spherical AFM probe at 0.5 Hz indentation frequency with a 1 μm/s approach/retraction velocity. For cell adhesion measurements, cells were indented with the ECM coated stylus AFM probe at 0.05 Hz indentation frequency and 0.1 μm/s approach/retraction velocity at a single spot. Thermal noise amplitude method was used to calibrate each AFM probe after each adhesion experiment and before each stiffness experiment (Hutter & Bechhoefer, 1993; Butt & Jaschke, 1995). A proprietary software written in MATLAB (R2016a, Mathworks) was used to analyze AFM force curves. Adhesion forces for COL1 and FN to integrins were determined by the product of cantilever spring constant and the height of ruptures on a retraction force curve. The adhesion force loading rate was determined by the product of retraction speed and the slope of a force curve right before a rupture (Hong et al., 2012b; Hong et al., 2014; Sanyour et al., 2019). The measured spring constant values fell between 18-19 pN/nm for adhesion and 37-52 pN/nm for stiffness experiments, respectively.

Cell Adhesion Molecule Expression Analysis

Western blotting assay to detect level of protein expression was performed according to the modified procedure described previously (Fang et al., 2018). Briefly, cells were lysed by addition of 1× Laemmli Sample Buffer (Bio-rad, #1610737) and sonicated with a Branson Sonifier-250. Equal amounts of protein from each treatment group were separated by 4-20% Mini-PROTEAN® TGX™ Precast Protein Gels (Bio-rad, #4561094). Proteins were transferred onto a Polyvinylidene fluoride (PVDF) membrane (Bio-Rad, # 1704272) using a semi-dry Bio-Rad Trans-blot apparatus. The membrane was then probed with one of the following primary antibodies: anti-β-actin (Santa Cruz, sc-47778) (1:1,000 dilution), anti-Integrin alpha 2 (Abcam, ab181548) (1:5,000 dilution) and anti-Integrin alpha 5 (ab150361) (1:2,500 dilution). The following secondary antibodies were used: horseradish peroxidase-conjugated goat anti-rabbit (Fisher Scientific, PI31460) and goat anti-mouse (Fisher Scientific, PI31436). Proteins were detected using a GE Healthcare Amersham™ ECL™ Prime Western blotting chemiluminescent detection reagent kit (Fisher Scientific, 45-002-401) and protein expression was quantified with the gel analysis function of ImageJ software (NIH, Bethesda, MD).

Live VSMC AFM Imaging

Contact mode AFM imaging was used to study live VSMCs cytoskeleton. Using a stylus AFM probe (model MLCT-C, k=15 pN/nm, Bruker, Santa Barbara, CA, USA), a 40×40 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 code written in MATLAB (R2016a, Mathworks). All experiments were conducted in previously warmed CO₂ independent medium at room temperature for a 1-2 h duration (Sanyour et al., 2019).

AFM Image Analysis for Stress Fiber Orientation and Density

The ratio of the cell surface area covered by actin stress fiber compared to the whole cell surface area was defined as the stress fiber area fraction. To evaluate the stress fiber area fraction, obtained AFM height images were first flattened to eliminate background noise resulted from the AFM stylus scanning lines. This improves the contrast between the background (non-stress fiber area) and the foreground (stress fiber). Cell surface roughness average (RA) was determined using the built-in function of the AFM software (Asylum Research). AFM deflection images were used to compute submembranous stress fibers orientation of VSMCs using a custom developed software (Sanyour et al., 2019).

F-actin Fluorescent Staining and Confocal Image Processing

Freshly isolated VSMCs were cultured on 50 mm glass bottom tissue culture dishes (MatTek Corporation, Ashland, MA) and followed by fluvastatin-treatment. VSMCs were fixed using 500 μL of 4% paraformaldehyde solution in PBS (Affymetrix, CA) for 20 min at room temperature followed by 2 washes with PBS. 0.1 % Triton X-100 (500 μL) was added to the cell bath for 5 min and washed with PBS twice. A working solution (1/1000 dilution) of phalloidin (Phalloidin-iFluor 488 - Abcam, Cambridge, UK) dissolved in 1% bovine serum albumin (BSA)/PBS was used to stain F-actin cytoskeleton (20 min). Following 2 quick washes with PBS, nuclear counter staining was performed using a 1:1000 dilution working solution of Hoechst 33342 (BD Biosciences, San Jose, CA) dissolved in PBS (10 min). VSMCs were imaged using a laser scanning confocal microscope (Olympus IX83 FV1200, Olympus life science, USA) at a 1024×1024 pixels resolution with an optimum z-step interval (0.2-0.4 μm). VSMCs z-stack images were flattened and manually traced for subsequent segmentation and analysis. As described previously in our publication, a series of elongated Laplacian-of-Gaussian (eLoG) filters were used to convolve z-stack confocal images to detect total cellular cytoskeletal fibers orientation (Hong et al., 2014; Sanyour et al., 2019).

Statistical Analysis

One-way ANOVA with Bonferroni’s post hoc test was used to infer statistical significance of cholesterol content, migration, stiffness, adhesion, stress fiber density, surface roughness, stress fiber orientation, and aortic contraction force between fluvastatin-treated and control VSMCs. Circular statistics was employed in analyzing actin orientation. The T-test for circular uniformity (Rayleigh test) was employed in determining significant actin orientation distribution. Values ranging from 0 to 1 indicate the levels of actin fiber organization, i.e., 0 indicates a complete random orientation of actin fibers, and 1 indicates a complete anisotropic orientation of actin fibers. A value of P ≤ 0.05 was considered significantly different. All data were reported as the mean ± standard deviation (SD).

Results

Fluvastatin Modified VSMC Migration and Cholesterol Content.

In order to study the effect of fluvastatin on VSMC migration on different ECM protein substrates, VSMCs were treated with 1 μM fluvastatin and seeded on COL1 or FN coated glass surfaces and allowed to migrate for 24 h in serum free media. Fig. 1A and 1B represent the position plots of all VSMC migration on COL1 and Fig. 1E and 1F on FN coated surfaces for 24 h. Moreover, total distance covered by all VSMCs on both protein substrates are represented in Fig. 1C, 1D, 1G and 1H. Fluvastatin-treatment resulted in a significant drop in total distance covered by VSMCs on FN but not on COL1 coated substrates (Fig. 1I-1L). It can be noted that the treatment did not affect the displacement (the change in start and end position) of VSMCs on either substrate indicating a relatively unidirectional migration (Fig. 1L). Cholesterol quantification of VSMCs post fluvastatin-treatment showed an approximately 30 % reduction in total cellular cholesterol (Fig. 1M).

Figure 1.

VSMC cholesterol content and migration on collagen 1or fibronectin coated glass substrates. A, B, E, F. Position plots of VSMC migration on collagen 1 or fibronectin surface. C, D, G, H. Migration distance covered by VSMCs on collagen 1 or fibronectin surface. I, J. Average migration distance versus time for all cells. K. Average migration distance covered by each VSMCs experimental group. L. Displacement of each VSMCs experimental group. M. GC/MS cholesterol quantification. (n= 8 for CTRL, n=9 for Fluvastatin). Displacement is defined as the difference between the starting and ending position of a cell. For each dish, five to ten regions of interest were chosen and imaged with a 10× objective every 10 min for 24 h. For each experiment, 50 cells were tracked. All data were presented as the mean ± SD (n=150 cells from 3 independent experiments).

Fluvastatin-mediated Cholesterol Depletion Increased the Expression of Integrin α5 in VSMCs, Enhanced VSMC Adhesion to FN, and Concurrently Reduced Cellular Stiffness.

Primary VSMCs attached to at least two other cells were selected for AFM experiment and indented at a site between their nucleus and cell edge. Cell-ECM adhesions were carried out using ECM protein functionalized AFM stylus probes (Sun et al., 2017). Fig. 2A is a schematic illustration representing a stylus AFM probe coated with COL1 or FN probing VSMCs surface. A typical representative AFM force curve generated upon surface indentation provides quantitative information about adhesion events (ruptures), adhesion force loading rate, and adhesion force of each rupture (Fig. 2B). Our results demonstrated that fluvastatin-treatment significantly increased VSMC adhesion probability and average adhesion force to FN but not to COL1 (Fig. 2C, 2D). In addition, the average adhesion force loading rates to FN significantly increased upon fluvastatin-treatment (Fig. 2E). Observing changes in cell-ECM adhesion properties, we set forth to explore adhesion protein expression. Our results suggested a significant increase in integrin α5 (Itg α5) protein expression. However, there was no significant change in integrin α2 (Itg α2) protein expression (Fig. 2F-2I).

Figure 2.

Effect of fluvastatin-mediated cholesterol depletion on VSMC biomechanics and adhesion proteins expression. A. Schematic illustration of adhesion force measurement using AFM. ECM-coated stylus AFM probe set to contact cell surface, establish binding with receptors, and retract to dissociate the ligand-receptor binding. B. A representative AFM force curve (approach-blue, retraction-red). Black numbers indicate adhesion force of each rupture; Green numbers indicate adhesion force loading rate of each rupture; Blue numbers indicate adhesion event in this individual sampling cycle. C. Adhesion probability represented by the number of ruptures per curve. D. Average adhesion force obtained by multiplying individual rupture heights by the cantilever spring constant. E. Average adhesion force loading rate, which was defined by the product of retraction speed and slope of the force curve right before the rupture. F, H. Full cell lysate protein expression for integrins α5 and α2. G, I. Average relative protein expression for integrins α5 and α2 to β-actin average of 3 independent experiments. All AFM data are reported as the mean ± SD. (n=100 cells from 7 independent experiments). All protein expression data are presented as the mean ± SD (n=3 from 3 independent experiments conducted in triplicates).

Given changes in VSMC cholesterol content, migration, adhesion properties, and protein expression, we aimed to assess the effect of fluvastatin-mediated cholesterol depletion on live VSMC stiffness. Primary VSMCs were scanned with a microbead-attached probe for real time stiffness-map measurement. Fig. 3A-3D describe the process by which a 40×40 μm VSMC surface area was scanned. Fig. 3E and 3F respectively represent the 6×6 position stiffness color maps of a control and a fluvastatin-treated live VSMC. The results demonstrated that VSMCs E-modulus significantly decreased in response to fluvastatin-mediated cholesterol depletion (Fig. 3G).

Figure 3.

Real time VSMC stiffness measurement. A. A spherical glass bead (5 μm in diameter) attached to an AFM probe. B. Schematic illustration of overall cell E-modulus (E-map) measurement using spherical AFM probe by scanning over a 40×40 cell surface with approximately 500 pN indentation force. C. 40×40 μm (cyan square) cell surface area was scanned for E-mapping. D. 40×40 μm cell surface area was divided to 6×6 sub-regions. E, F. Representative 6×6 E-map of a control VSMC and fluvastatin-treated VSMC. G. Effect of statin on overall VSMC E-modulus. Fluvastatin-treatment significanly decreased VSMC E-modulus. All data are presented as the mean ± SD (n=58 for control and n=59 for fluvastatin-treated cells from 4 independent experiments).

Live AFM Imaging Revealed Fluvastatin-mediated VSMC Submembranous Cytoskeletal Remodeling.

We speculated that the decrease in stiffness and alteration in adhesion force were partly due to cholesterol depletion induced cytoskeletal remodeling. Therefore, we evaluated VSMC submembranous cytoskeletal architecture by live cell AFM imaging. To compute the variation in submembranous stress fiber area fraction, AFM height images were acquired and analyzed. Fig. 4A and 4B present the three-dimensional stress fiber height topography of a control and fluvastatin-treated VSMC, respectively. Fig. 4C and 4D respectively display representative flattened AFM height images of submembranous stress fibers for a control and fluvastatin-treated VSMCs. Using the corresponding flattened height images, the stress fibers (red) were distinguished from the analogous background (blue) and stress fiber area fraction was computed (Fig 4E, 4F). The results indicated that there was no significant change in actin stress fiber density (Fig. 4G). Alternatively, the cell surface RA, the average absolute value of a set of measured microscopic surface peaks and valleys of VSMCs height images demonstrated a significant decrease in stress fiber roughness after fluvastatin-treatment (Fig 4H). Next, the AFM deflection images were used to compute and quantitatively analyze the VSMC stress fiber orientation. The representative AFM images of VSMC stress fiber were shown in Fig. 5A and 5B. The corresponding circular histograms of the submembranous stress fiber orientations for a control (blue) and fluvastatin-treated (red) VSMC are presented in Fig. 5C and 5D, respectively. Normalized percentage circular histograms along the dominant orientation (Fig. 5E, 5F), show a clear difference in stress fiber orientation between statin treated and control VSMCs. Consistent with the representative VSMCs images, the summarized results of the two groups of VSMCs demonstrated a significant difference in stress fiber orientation (Fig. 5G, 5H). The circular variance was used to find a significant difference in total stress fiber orientation between control and fluvastatin-treated VSMCs (Fig. 5I). These results indicated significant alterations in stress fiber orientation after fluvastatin-mediated cholesterol depletion without notable changes in stress fiber density.

Figure 4.

AFM image for live VSMC submembranous stress fibers. A, B. Representative three-dimensional stress fibers height topography of a control and fluvastatin-treated VSMCs. Alteration in stress fiber height from low to high is presented as color changes from blue to red. C, D. Representative 40×40 μm flattened AFM height images of control and fluvastatin-treated VSMCs. E, F. Representative submembranous stress fibers surface area fraction images were obtained from their respective AFM height images. The red represents the surface area of stress fiber, while the blue represents the empty background. G. The summarized area fraction of VSMC submembranous actin stress fibers. H. The summarized stress fiber surface roughness average. All data are presented as the mean ± SD (n=20 cells from 3 independent experiments).

Figure 5.

Live VSMCs submembranous stress fiber organization. A, B. Representative AFM deflection images of control and fluvastatin-treated VSMCs submembranous stress fibers. C, D. Circular histograms of stress fiber orientation frequency of control (blue) and fluvastatin (red) treated VSMCs. The histogram data were computed from the representative images in panel A and B, respectively. E, F. Normalized percentage circular histograms along the dominant orientation. G, H. Summarized percentage circular histograms of VSMCs stress fiber orientation for each experimental group. I. Circular variance of stress fiber orientation for each individual VSMC. All data are reported as the mean ± SD (n=20 cells from 3 independent experiments).

Global VSMC Cytoskeletal Architecture Remodeling in Response to Fluvastatin-mediated Cholesterol Depletion was Observed by Confocal Microscopy.

Following live cell imaging, we studied the impact of fluvastatin on the overall cytoskeletal architecture in the entire cell. Primary VSMCs were cultured on glass bottom dishes and treated with fluvastatin. VSMCs were fixed, permeabilized, and stained with F-actin and DNA binding fluorescent dyes and visualized using confocal microscopy. Fig. 6A and 6C show representative control and fluvastatin-treated VSMCs confocal images. Using a proprietary software developed in MATLAB, VSMCs were segmented and stress fiber orientation color maps were computed for control (example cells 1-4 in Fig. 6B) and fluvastatin-treated VSMCs (example cells 1-4 in Fig. 6D). A pseudo color map representing different stress fibers orientation angles were calculated from corresponding VSMCs stained images. The colors of actin stress fiber orientation for each control VSMC were almost uniform, whereas the color maps for fluvastatin-treated VSMCs displayed diverse colors, demonstrating a greater dispersion of actin stress fiber orientation. Moreover, the normalized percentage circular histograms of each control cell’s stress fiber angles exhibited a well-organized stress fiber orientation (Fig. 6E-6H), while fluvastatin-treated cells (Fig. 6I-6L) depicted a more disorientated characteristics of actin stress fibers. The summarized results displayed a significant difference between the stress fiber orientations of control (Fig. 7A) and fluvastatin-treated (Fig. 7B) VSMCs groups. The average stress fiber orientation for both control and fluvastatin-treated VSMCs were plotted in a percentage frequency histogram, in which the fiber angles were normalized along the dominant orientation and fitted with a first order Gaussian function (Fig. 7C). Significance of the difference in the fiber orientation was determined using the comparision of the cirular variance of fibers in the two experimental groups.

Figure 6.

The global effect of fluvastatin on VSMC actin stress fiber architecture. A, C. Representative confocal images of control and fluvastatin-treated VSMCs. Actin was stained in green and the nucleus was in blue. B, D. A pseudo color map of stress fibers orientation computed from the corresponding confocal images with various colors representing different fibers orientation angles. Cells with a clear boundary without overlay with neighboring cells were analyzed for their fiber orientation. E-H. Representative percentage circular histograms of detected stress fiber orientation frequency of 4 control cells. I-L. Representative percentage circular histograms of detected stress fiber orientation frequency of 4 fluvastatin-treated cells.

Figure 7.

Summarized overall VSMCs actin stress fiber orientation. A, B. Summarized percentage circular histograms of control (blue) and fluvastatin-treated (red) VSMCs stress fiber. C. The average normalized stress fiber orientation percentage histogram for all VSMCs, in which the dominant stress fiber orientation angle was set as zero degree for each cell. D. Circular variance of stress fiber orientation for each experimental group of VSMCs. All data for panel D is presented as the mean ± SD (n = 120 cells from 6 independent experiments)

Fluvastatin Diminished Native Aorta Cholesterol Content and Modified Contractile Activity.

To evaluate the effect of fluvastatin-mediated cholesterol depletion on native aorta cholesterol levels and contractile activity, rat aortic rings were incubated with fluvastatin for 72 h followed by contractile activity examination using a myograph. Native vessel cholesterol loading significantly reduced after fluvastatin-treatment by 40% (Fig. 8A). Moreover, the contractile activity of fluvastatin-treated vessel ring in response to stimulation with phenylephrine (PE) (100 μM) was diminished (Fig. 8C and 8D). However, there was no change in the aortic vessel stiffness after fluvastatin-mediated cholesterol depletion (Fig. 8B). The summarized results showed that fluvastatin-treatment lowered aortic tissue cholesterol level and altered its contractility in response to PE stimulation (Fig. 8D).

Figure 8.

Fluvastatin-mediated native aorta cholesterol depletion and its effect on the vessel ring contractile activity. A. Cholesterol loading in control and fluvastatin-treated aortic rings (n=11 aortic rings from 4 independent experiments). B. Stiffness of control and fluvastatin-treated rat aortic rings (n= at least 6 aortic rings from 6 independent experiments). C. Average real-time contractile response of aortic rings to phenylephrine (PE) stimulation. D. Summarized aortic ring maximum constriction force (n=8 aortic rings from 8 independent experiments). All data are presented as the mean ± SD.

Discussion

A number of studies have elucidated some of the key roles that cholesterol plays in cell spreading, migration, and mechanics of different cell types (Yan et al., 2011; Hong et al., 2013; Khatibzadeh et al., 2013). In addition, linage studies have demonstrated that a large number of foam cells retain VSMC markers suggesting cholesterol to be a possible regulator of phenotypic shifting (Allahverdian et al., 2012; Allahverdian et al., 2014). A significant number of publications have described statins’ ability to lower cholesterol levels and reduce cardiovascular events (Corsini et al., 1993; Scandinavian Simvastatin Survival Study, 1994; Shepherd et al., 1995). In particular, fluvastatin-treatment significantly reduced the possibility of adverse cardiac events in patients with average cholesterol levels undergoing percutaneous coronary intervention (Serruys et al., 2002). However, clinical use of statins as hypercholesterolemia lowering drugs are still debated due to their pleiotropic effects (Oesterle et al., 2017). A critical step during atherosclerosis progression is the phenotypic shift of VSMCs accompanied by detachment from neighboring cells and migration towards the intima. Hayashi et al. reported that VSMC phenotype modulation from contractile to synthetic is governed by a signaling pathway through mitogen-activated protein kinase (MAPK) and extracellular signal-regulated kinase 1 and 2 (ERK1/2) or p38MAPK (Hayashi et al., 1999). Moreover, Sakamoto et al. demonstrated that fluvastatin prevented VSMCs hyperplasia, proliferation, and phenotype modulation by ERK1/2 and p38MAPK inactivation (Sakamoto et al., 2005). Furthermore, the same study suggested that the physicochemical properties of the ECM are important and implicated in maintaining VSMCs contractile state.

To understand the mechanosensory nature of VSMCs and statin mediated cholesterol depletion, we aimed to examine and unravel the effect of fluvastatin-treatment on VSMC cholesterol content, biomechanics, and cytoskeletal structure in freshly isolated rat aortic VSMCs. In order to minimize VSMCs phenotypic changes in our study, primary VSCMs were cultured for a short period without passaging to allow their recovery from the enzymatic isolation process, then treated and used in each experiment. Adhesion of cells to the ECM is mainly governed by integrins. A key event during the development of atherosclerosis is the migration of VSMCs within the blood vessel wall. Therefore, we initially assessed the effect of fluvastatin on VSMC migration on COL1 and FN coated substrates. Fluvastatin-treatment significantly reduced VSMC migration distance on FN substrates, but did not change VSMC migration distance on COL1 substrates (Fig. 1). This result indicates that fluvastatin distinctly regulates the migration behavior of VSMCs on different ECM protein substrates. It is well established that the migration of VSMCs from the media to the intima is a critical event in atherosclerosis (Wang et al., 2015), thus our result could suggest that fluvastatin may reduce VSMC migration and impede the atherosclerosis process. We then studied the effect of fluvastatin on VSMC cholesterol content. Our results showed an approximately 30% decrease in total cholesterol per VSMC upon fluvastatin-treatment (Fig. 1). Cotsini et al. first demonstrated 1 μM of fluvastatin (3R, 5S) reduced rat VSMC cholesterol synthesis by approximately 70% in vitro (Corsini et al., 1993). Moreover, patients diagnosed with type IIa hypercholesterolemia supplemented with therapeutic doses of fluvastatin (0.1-1 μM) had a 28% and 16% decrease in their VSMCs growth and total serum cholesterol respectively (Tse et al., 1992; Corsini et al., 1996). A study conducted by Schaefer et al. confirmed a dose of 40 mg/day fluvastatin was able to reduce low density lipoprotein (LDL) cholesterol levels in serum of patients with coronary heart disease by 20% (Schaefer et al., 2004). During phenotypic switching, detachment, and migration towards the inflammatory lipid rich site in the progression of atherosclerosis, VSMCs undergo continuous mechanoadaptations partly due to their ability to sense and react to mechanical loading and chemical stimuli (Leung et al., 1976; Hong et al., 2016).

Integrins are a large family of cytoskeleton connected mechanosensors. Of particular note, the integrin β₁ family mediates VSMC adhesion to the ECM. α5β1 integrin binds to FN and allows focal adhesion development. In addition, previous evidence showed that membrane cholesterol and phospholipid composition changed integrin α5β1 adhesion to FN (Gopalakrishna et al., 2000; Gopalakrishna et al., 2004). Graf et al. demonstrated that statins were able to regulate COL1 specific integrin α2β1 expression in human VSMCs (Graf et al., 2003). In this scenario, we aimed to study the integrin mediated cell-ECM adhesion, particularly integrins binding to FN and/or COL1 following fluvastatin-mediated cholesterol depletion. As shown in Figure 2, the adhesion probability significantly increased for VSMC-FN adhesion without any change for VSMC-COL1 adhesion. Moreover, the average VSMC-FN adhesion force was significantly enhanced upon fluvastatin-treatment (Fig. 2D). In addition, fluvastatin-treatment significantly increased the adhesion force loading rate for VSMC-FN adhesion (Fig. 2E). We speculate that integrins are recruited and aggregated upon statin treatment and AFM tip probing stimulation, which leads to a compounded increase in adhesion force and loading rates. Moreover, the increase in adhesion probability to FN is likely attributed to the ability of several different integrin subunits to bind to FN, and possibly the over expression of Itg α5. Interestingly, there was no change in VSMC-COL1 adhesion forces, which was consistent with no change in migration on the COL1 coated substrate. Furthermore, the significant drop in migration on the FN coated substrate and significant increase in adhesion force and loading rate may indicate a stronger integrin-FN adherence rendering VSMCs to lock down in position and migrate less. Overall, a significant difference in VSMCs migration was observed on FN coated substrates. In healthy human arteries, VSMCs reside within ECM that is mainly composed of collagen and laminin. However, the ECM of atherosclerotic arteries contain abundant deposits of FN in both humans and murine models, suggesting a functional role for FN in the pathophysiology of atherosclerosis (Doddapattar et al., 2015). Therefore, fluvastatin-mediated cholesterol depletion exhibited a significant role in modifying VSMC-ECM interactions through affecting α5β1-FN binding. Our findings are consistent with what was reported by Ramprasad et al. that membrane cholesterol content has a specific effect on certain signaling pathways involving FAK and Erk1/2 MAPKs that regulate cell motility on FN and actin cytoskeleton organization, thus regulating the shape, adhesion, and migration of cells plated on FN (Ramprasad et al., 2007). Several previous studies elucidated the vital role cholesterol played in modifying E-moduli of different cell types (Hong et al., 2012a; Hong et al., 2013; Ayee & Levitan, 2016). To follow up, we assessed real time VSMCs E-modulus after fluvastatin-treatment. A 40×40 μm area was scanned and indented 150-250 nm into VSMCs cortex using an AFM tip attached with a 5 μm spherical glass bead (Hong et al., 2012b; Sehgel et al., 2013; Hong et al., 2015). We observed a significant drop in VSMCs E-modulus in response to fluvastatin-mediated cholesterol depletion (Fig. 3), thus the decrease in VSMCs stiffness by fluvastatin could serve to offset the increasing arterial stiffness seen during arteriosclerosis progression.

Recently we demonstrated that VSMCs undergo cytoskeletal remodeling and changes in their biomechanical adhesion due to membrane cholesterol and substrate stiffness coordination (Sanyour et al., 2019). Similarly, a number of researchers have revealed the direct and indirect roles played by membrane cholesterol in reorganizing cellular cytoskeleton (Chubinskiy-Nadezhdin et al., 2011; Efremova et al., 2012; Chubinskiy-Nadezhdin et al., 2013; Khatibzadeh et al., 2013; Dason et al., 2014). In this study, live VSMC imaging was performed using AFM to interpret the reduction in VSMC stiffness upon fluvastatin cholesterol depletion. A custom developed novel computational approach enabled us to quantitatively analyze stress fiber organization (Fig. 4 and 5). Our results showed that fluvastatin-treatment did not change VSMC actin stress fiber density (Fig. 4) but significantly dispersed the stress fiber orientation (Fig. 5 and 6). These results suggest that stress fiber orientation contribute to VSMCs stiffness alteration. Recognizing the pleiotropic effects of statins, especially on the non-steroid isoprenoid pathway as well as the ability of cholesterol to directly modulate signaling pathways involved in cytoskeletal organization, we examined total actin orientation in VSMCs using confocal microscopy (Papanikolaou et al., 2005; Kang et al., 2014; Kang et al., 2016; Wang et al., 2017; Zhang et al., 2018). Our results suggest that fluvastatin significantly alters overall stress fiber orientation throughout VSMCs, another indicative observation explaining the drop in VSMC stiffness upon fluvastatin cholesterol depletion (Figs. 6 and 7). Similar findings have been reported that generation of stress fibers together with their alignment, are associated with stiffening of the cell body (Rabbani et al., 2017), and it is also recognized that cell elongation and stiffening in response to cyclic stretch is caused by actin alignment and elevation of cellular protein content (Krishnan et al., 2009). Regarding the underlying mechanisms, Krishnan et al. found that cells can fluidize their cytoskeleton to reduce their structural stiffness and remodel in response to a change in their mechanical environment. In addition, Gavara and Chadwick elegantly modeled and assessed the relationship between cell stiffness, stress fiber amount, and orientation using both simultaneous AFM and live-cell fluorescence imaging. The study demonstrated that the amount, thickness, and actin stress fiber orientation directly reinforce cell stiffness. In addition, stress fiber orientation was shown as a second order modulator of cell stiffness (Gavara & Chadwick, 2016). It is established that VSMCs not only undergo quantitative changes in contractile and cytoskeletal proteins, but also undergo reorganization and significant remodeling of their cytoskeletal structure during atherosclerosis development. Therefore, fluvastatin induced cytoskeletal remodeling might be an alternative means by which fluvastatin prevents atherosclerosis progression while decreasing cellular cholesterol. A previous report suggesting membrane cholesterol depletion results in Src kinase-mediated Rho activation and caveolin phosphorylation which collectively participate to form stress fibers, might provide a mechanistic explanation underlying the cytoskeleton remodeling following cholesterol depletion by fluvastatin (Qi et al., 2009).

It is worth mentioning that all the results in our present study are derived from the observations of cells cultured on 2 dimensional (2D) substrates. Not only does cellular cytoskeletal dynamics in 3D differ from those observed in 2D, but also differences in F-actin organization are observed. For example, cellular F-actin depolymerization in 3D following both sustained stretch and stress fiber compression, was different to the reinforcement and fluidization effects observed upon stretch and compression in 2D (Wu & Feng, 2015; Mierke, 2018). Cells in 3D were observed to exhibit a single natural axis defined by stretch, in contrast to cells in 2D with an additional axis defined by the normal to the culture dish (Lee et al., 2012). Additionally, different from that in a 2D culture, where cells exhibited branched and multiple-directed F-actin stress bundles at the cell edge and strengthened stress fibers traversing the cell body, cells cultured in spheroids showed compact cell body, relaxed cytoskeleton tension with very thin cortical actin filament outlining the cell (Zhou et al., 2017). Therefore, these observations of cells cultured on 2D substrates are not completely representative of behaviors that these same cells would exhibit in a natural tissue environment in vivo.

Since VSMCs are the major cellular component of the aorta, we tested the effects of fluvastatin on native aorta cholesterol content and contractile activity. The total cholesterol loading and contractility of rat aortic rings in response to PE (100 μM) stimulation were significantly reduced after a 3 day culture with fluvastatin (Fig. 8A). This result was consistent with a previous publication by Perings et al. that chronic fluvastatin-treatment improved endothelial and VSMC relaxation function in hyperlipoproteinemia rabbit carotid arteries (Perings et al., 2004). In addition, other statins have been also shown to inhibit conduit vessel constrictor responses (Ghaffari et al., 2011). For example, atorvastatin induced concentration-dependent relaxation of young and adult phenylephrine-contracted rat aortas, acute pravastatin pretreatment inhibited vasoconstrictor responsiveness of isolated rat mesenteric small vessels, and simvastatin was able to produce vascular relaxation in rat small arteries (de Sotomayor et al., 2000; Ghaffari et al., 2011; Nurullahoglu-Atalik et al., 2017). With respect to the underlying mechanism, it has been demonstrated that statins increase nitric oxide (NO) and carbon monoxide (CO) production in the cardiovascular system; both these “gasotransmitters” have significant vasodilatory and antiatherogenic activities (Wojcicka et al., 2011). Furthermore, it was elucidated that some statins can inhibit the vasocontraction induced by extracellular Ca²⁺ entry via the receptor-operated Ca²⁺ channel pathway, which is the primary mode of action that phenylephrine contracts VSMCs (Nurullahoglu-Atalik et al., 2017). One possible interpretation for why the aortic tissue sitffness was unchanged could be due to the overwhelming aorta ECM protein composition that governs vessel stiffness rendering the impact of cellular stiffness on the overall tissue stiffness insignificant. Collectively, we believe that fluvastatin-mediated cholesterol depletion and endothelial/VSMCs relaxation modulate and reduce PE induced vasoconstriction.

In summary, fluvastatin-treatment depleted cholesterol in primary cultured VSMCs by approximately 30% and in native aorta rings by 40%. Fluvastatin-treatment resulted in significantly less migration distance of VSMCs on FN substrate compared to control VSMCs, while having no significant effect on the migration behavior of VSMCs on the COL1 substrate. In addition, fluvastatin significantly increased Itg α5 expression in VSMCs and enhanced the VSMC-FN adhesion, while having no significant effect on Itg α2 protein expression and VSMC-COL1 adhesion. AFM and confocal microscopy images revealed a significant loss in cytoskeletal architecture orientation upon fluvastatin-treatment. Moreover, physiological functional testing of the native blood vessel ring demonstrated that fluvastatin significantly reduced vasoconstriction capability in response to the PE stimulation. Collectively, this work contributes to a better understanding of the effects of statin-mediated cholesterol management on VSMC mechanics and migration, in addition to its cholesterol lowering capability.

Translational perspective.

Atherosclerosis is a leading cause of death worldwide. Phenotypic shifting, disengagement of vascular smooth muscle cells (VSMCs) from neighboring cells/extracellular matrix (ECM), and migration toward inflammatory site of blood vessel walls are all critical contributions of VSMCs to atherosclerosis progression. Cholesterol is not only a major factor involved in fatty deposition in the atherosclerotic lesion, but growing evidence has suggested cholesterol’s role in modulating cellular biomechanics. Therefore, we investigated the impact of clinically relevant cholesterol lowering drugs, statins, on regulating VSMCs biomechanics. When statin mediated VSMC cholesterol depletion was achieved, migration on different ECM proteins was altered. This was corroborated by quantifying the adhesion forces to the corresponding ECM specific cell surface receptors. In addition, statins were able to reduce VSMC stiffness that was associated with stress fiber disorientation. When rat aortic tissue was treated with statins, tissue cholesterol content and vasoconstriction were reduced. These findings demonstrate the impact of cholesterol lowering drugs on both the cellular biomechanical function and extrapolated to tissue biomechanical function and possibly having a direct biomechanical impact on a patient’s cardiovascular disease development. In summary, our results suggest that statins lower VSMC and aortic tissue cholesterol content and modulate their biomechanical functions, which will ultimately affect patient atherosclerosis disease progression. Deciphering the extent and exact mechanism by which cholesterol modulates cellular biomechanics could offer patients and clinicians a better approach to manage and control atherosclerosis pathogenesis.

Key points.

• This study demonstrates and evaluates the changes in rat vascular smooth muscle cell biomechanics following statin mediated cholesterol depletion.

• Evidence is presented to show correlated changes in migration and adhesion of vascular smooth muscle cells to extracellular matrix proteins fibronectin and collagen. Concurrently, integrin α5 expression was enhanced but not integrin α2.

• Atomic force microscopy analysis provides compelling evidence of coordinated reduction in vascular smooth muscle cell stiffness and actin cytoskeletal orientation in response to statin mediated cholesterol depletion.

• Proof is provided that statin mediated cholesterol depletion remodels total vascular smooth muscle cell cytoskeletal orientation that may additionally participate in altering ex-vivo aortic vessel function.

• It is concluded that statin mediated cholesterol depletion may coordinate vascular smooth muscle cell migration and adhesion to different extracellular matrix proteins and regulate cellular stiffness and cytoskeletal orientation, thus impacting the biomechanics of the cell.

Biography

Hanna J. Sanyour earned his MS in Biotechnology from Ghent University in 2015 and conducted his graduate training at the Flanders Institute for Biotechnology. He also earned his Ph.D. in Biomedical Engineering from the University of South Dakota in 2019. His research interest is focused on the lipid driven cellular biomechanics and vascular tissue engineering. Na Li obtained her M.S from Jilin University in 2009 and her Ph.D. from Dalian Institute of Chemical Physics, Chinese Academy of Sciences in 2016. She worked as a postdoctoral researcher at the University of South Dakota from 2017 to 2019. Then she has worked as a postdoctoral fellow at the University of Chicago. Her research interest is focused on vascular tissue engineering and vascular mechanics.