- Research article
- Open Access
Constant or fluctuating hyperglycemias increases cytomembrane stiffness of human umbilical vein endothelial cells in culture: roles of cytoskeletal rearrangement and nitric oxide synthesis
© Chen et al.; licensee BioMed Central Ltd. 2013
Received: 11 November 2012
Accepted: 16 April 2013
Published: 22 April 2013
Previous studies have implicated continuous or intermittent hyperglycemia in altered endothelium-derived nitric oxide (NO) synthesis. NO can regulate both the F-actin cytoskeleton and endothelial cell membrane stiffness. Atomic force microscopy (AFM) is a powerful tool that can be used to study plasma membrane deformability at the single cell level. As membrane stiffness is partially dependent on filamentous F-actin, the interdependence of these parameters can be studied through the combined approaches of AFM and laser scanning confocal microscopy (LSCM). In the present study, we evaluated the effects of constant or fluctuating hyperglycemia on endothelial-derived NO synthesis, the cytoskeletal contribution and endothelial cell membrane stiffness.
Compared to control cells cultured in low glucose (5 mM), constant (25 mM) or fluctuating (25/5 mM) high glucose significantly decreased NO release along with stiffening of endothelial cell membranes and F-actin rearrangement. The non-selective nitric oxide synthase (NOS) inhibitor, NG-nitro-L-arginine methyl ester (L-NAME) exerted similar effects on endothelial cells. Increasing concentrations of L-NAME (from 0.1 to 1 mM) exacerbated these effects in a concentration-dependent manner.
Result from the present study suggest that stiffening endothelial cell membranes are associated with decreased NO synthesis, which was established through the F-actin cytoskeletal redistribution. The precise mechanisms of hyperglycemia-induced endothelial dysfunction require further investigation.
Vascular disease remains the major cause of increased morbidity and mortality in patients with diabetes . Endothelial cell dysfunction is a significant factor contributing to the vascular complications of diabetes . Hyperglycemia, particularly the fluctuation of glucose levels, causes a significant degree of oxidative stress, decreasing endothelial nitric oxide synthase (eNOS) expression [3, 4], reducing nitric oxide (NO) bioavailability, and impairing NO metabolism .
Decreased bioavailability of endothelial-derived NO contributes to endothelial cell contraction, which is partly dependent on reorganization of the endothelial cell cytoskeleton [6–8]. Substantial evidence exists for the involvement of endogenous NO production in the regulation of the cytoskeletal F-actin structure via mobilization of intracellular Ca2+ by either protein kinase C (PKC) or protein kinase G (PKG) [9–11].
The actin network plays a major role in determining the mechanical properties of living cells . Atomic force microscopy (AFM) can be used to study cell morphology and the micromechanical properties of both the cell surface and subsurface layers. Furthermore, recent evidence indicates that it is possible to assess the deformability of the plasma membrane at the single cell level using AFM . The combined approaches of AFM and laser scanning confocal microscopy (LSCM) extends the utility of the scanning probe approach for the evaluation of cellular mechanical properties. Using immunofluorescent dyes and LSCM to dissect the three cytoskeletal elements, elasticity of the cell membrane has been found to be related to distribution of actin and intermediate filaments, with only a minor contribution from microtubules .
Several studies have investigated the potential relationships between NO release and endothelial cell membrane stiffness using AFM [15–17]. Recently, it was reported that acute and small changes in plasma sodium concentration can lead to a significant increase in endothelial cell membrane stiffness that associated with reduced NO release , while potassium ions can soften vascular endothelial cells and increase NO production . However, it has not been definitively proven that endothelial-derived NO influences endothelial deformability via actin cytoskeletal reorganization. Interestingly, a number of studies have examined alterations in endothelial deformability by AFM in various physiological and pathological states [18–23]. However, the effects of stable or fluctuating hyperglycemia on endothelial cell membrane stiffness using AFM has received little attention.
In the present study we examined the effects of constant and fluctuating high glucose levels on the release of endothelial-derived NO, expression of eNOS, changes in endothelial cell membrane stiffness and the contribution of filamentous F-actin.
Human umbilical vein endothelial cells (HVUECs-12, CRL-2480) were obtained from the American Type Culture Collection, plated in 25 cm2 flasks (Costar, Japan) and cultured in low glucose (5 mM) Dulbecco’s Modified Eagles Medium (DMEM; GIBCO, Invitrogen, USA) supplemented with 10% fetal bovine serum (TBD, China). The flasks were incubated at 37°C in a humidified atmosphere containing 5% CO2. After reaching confluence, the cells were treated with 0.01% trypsin-EDTA (Amresco, USA). One ml cell suspensions containing 105 cells/ml were re-seeded in 60-mm dishes (Corning, USA). The endothelial cells were allowed to attach overnight, and were exposed to the appropriate experimental conditions for seven days. In brief, eight groups of cells were prepared, each receiving the following fresh media every 24 h, respectively: 1)control group exposed to continuous low glucose (5 mM) DMEM medium; 2)continuous high glucose (25 mM) DMEM media (GIBCO, Invitrogen, USA); 3) low (5 mM) and high (25 mM) glucose media alternating every 24 h; 4) continuous low glucose (5 mM) DMEM media containing 0.1 mM non-selective NOS inhibitor NG-nitro-L-arginine methyl ester (L-NAME, Biyotime Co., China); 5) continuous low glucose (5 mM) DMEM media containing 0.5 mM L-NAME; 6) continuous low glucose (5 mM) DMEM media containing 1 mM L-NAME; 7) continuous osmotic control, low glucose (5 mM) DMEM media containing 20 mM mannitol (Sigma, USA); or 8) intermittent osmotic control, low glucose (5 mM) DMEM media containing mannitol (0/20 mM, alternating every 24 h).
Nitric oxide measurement
Where ODT was the absorbance of the test solution; ODB was the absorbance of a regent blank; ODS was the absorbance of the 100 μmol/L standard. All experiments were performed in triplicate.
Expression of endothelial nitric oxide synthase (eNOS)
Assays were performed using western immunoblot analyses using specific antibodies against eNOS (Genscrip, USA) and β-actin (Newmark, USA). β-actin expression was used as a loading control. One ml cell suspension was plated at a density of 105/ml in 60-mm dishes (Corning, USA), allowed to attach and cultured under various conditions. Cells were subsequently washed with chilled PBS and homogenized in modified RIPA buffer (1 M Tris–HCl PH 8, 1 M NaCl, 1% Triton X-100, 0.5M EDTA PH 8, 1 M MgCl2, 1 mM PMSF, Boster Co., China) for 5 min. Cell lysates containing equal amounts of protein were dissolved by adding 5×SDS-PAGE sample buffer (Boster Co., China) followed by heating for 10 min. 100 μg of total protein was separated on polyacrylamide gels (the gel percentage was chosen depending on the protein being observed.) Following electrophoresis, samples were transferred to a PVDF membrane using a 25 mM Tris buffer containing 192 mM glycine and 20% methanol using a Bio-Rad mini-Blot transfer apparatus. The PVDF membranes were blocked with 5% (w/v) non-fat milk in 0.01M PBS containing 0.1% Tween 20 for 2 h at room temperature, and then incubated in the above antibodies against eNOS (1:1000 dilution) and β-actin (1: 400) overnight at 4°C. After washing with PBS/Tween (3 × 15 min), the membranes were incubated in secondary antibody (goat anti-rabbit IgG, 1:5,000 dilution, Keygen Co., China) for 2 h and washed. Protein bands were detected using chemiluminescent substrate (Pierce Biotechnology, USA) and developed on film (Kodak, USA) following a 5 min exposure. Relative band densities were quantified using densitometry with Image J software (NIH, Bethesda, MD). Data were then normalized to β-actin levels. All western immunoblot experiments were repeated at least in triplicate with separate cell preparations.
Cell suspensions (total 105 cells) were plated on glass coverslips and positioned in 60 mm culture dishes. Following seven days of incubation in various media, cells were washed with chilled PBS, fixed in 3.75% formaldehyde solution, permeabilized with Triton X-100, and blocked with 1% BSA. To label F-actin, one unit of Rhodamine Phalloidin (Biotum, USA) was diluted in 200ul PBS with 1% BSA, and then added to each cover slip. The samples were stained for 20 min, and subsequently washed in triplicate with PBS. Stained samples were imaged using a Zeiss LSCM (510 META Duo Scan; Zeiss, Germany) at excitation (540 nm) and emission (565 nm) wavelengths.
AFM imaging and measurement
Cells were planted on coverslips and cultured under various conditions for seven days. The coverslips, with seeded cells, were washed in triplicate to clear the culture media. Following preparation of the samples, an Autoprobe Cp AFM (Autoprobe CP Research, Veeco, USA) was operated in contact mode in air to obtain AFM images and perform force spectroscopy. Stiffness of the endothelial cells was determined using AFM techniques as described previously and measured with soft cantilevers, since the area of interaction between the tip and cell was larger and thus mechanically less noisy [12, 15, 16, 25]. Silicon nitride tips (UL20B, Park Scientific Instruments) were used for all AFM measurements. The curvature radius of the tips was less than 10 nm, and the length, width and thickness of the cantilevers were 115, 30, and 3.5 μm, respectively, with an oscillation frequency of 255 kHz and a force constant of 0.01 N/m (manufacture offered). The same approach velocity was maintained to detect the endothelial cells.
Topographical morphology and deflection images of the surface of HUVECs were acquired over an area of 5-80 μm2. According to these images, parameters of the endothelial cells were obtained, such as Rp-v and Ra. The valley-to-peak value (Rp-v) defines the difference between the maximum and minimum values of the z coordinate on the surface of the analytical area. Ra denotes the average roughness in the analytical area. All parameters were directly generated by the software IP2.1. Subsequently, endothelial cell membrane stiffness of each chosen point was evaluated using the indentation of the local force-distance curve .
Five randomly selected endothelial cells were studied for each experimental treatment. Membrane stiffness of each cell was sampled at approximately 30 randomly chosen points on the cell center (including perinuclear) or peripheral membrane. The total average stiffness was acquired from 300 force curves.
Results are expressed as mean ± SD. When the experimental treatments were compared to either the control group, or the group under continuous high glucose exposure, a one-way variance analyses was performed. A value of P<0.05 was considered statistically significant. The statistical package SPSS13.0 was used for all analyses.
Release of endothelial-derived NO
Effects of high glucose and NOS inhibitor L -NAME on the viability, NO release, eNOS expression and F-actin fluorescence intensity of human umbilical vein endothelial cells incubated in different media (as described in Figure 2 ) for seven days
Cell viability (Optical density)
NO concentration (μmol/L)
eNOS expression (%)
F-actin florescence intensity
Expression of eNOS
AFM imaging and morphologic parameters
Data of different parameters obtained by AFM in human umbilical vein endothelial cells incubated in various media (as described in Figure 2 ) for seven days
Endothelial cell membrane stiffness was estimated through the detection of the point-by-point force-distance curves (Figure 4E). Half of the 300 force curves for each group were acquired by measuring the surface of the cellular periphery and the other half in the perinuclear region (Figure 4B b and c).
F-actin localization and quantification of the fluorescence intensity
Cellular function is largely determined by its structure. Thus, cellular deformability, a mechanical property of the cellular structural organization, varies in a number of physiological processes (including cell differentiation, growth and adhesion) and in pathological states (including oxidative stress, viral infection and, parasitic infection) . Recent studies have shown that alterations in endothelial cell membrane stiffness are accompanied with changes in NO production [15–17]. For example, a small physiological increase in extracellular sodium directly increases the stiffness of vascular endothelium and decreases NO release . An acute increase in potassium, within a physiological range, swells and softens endothelial cells and increases the release of NO . Nebivolol, a β1-receptor blocker, decreases membrane stiffness of endothelial cells, which is dependent on the increased content of NO outcome and is abrogated by L-NAME .
The above studies, however, have only focused on how experimental conditions altered endothelial cell membrane stiffness while simultaneously affecting NO synthesis. Thus, attention has not been paid to the exact effects of NO release and eNOS expression on endothelial cell membrane stiffness. Therefore, to examine the effect of NO on cell membrane stiffness, L-NAME was used to inhibit endogenous NO generation. Expression of eNOS (Figure 3), together with NO generation (as assessed by nitrite) (Figure 2), was quantified to clarify the relationship between cell membrane stiffness and NO metabolism. The data showed that L-NAME increased endothelial cell membrane stiffness (Figure 6), inhibited NO release and suppressed eNOS protein expression in a concentration-dependent manner suggesting a direct relationship between production of NO and the mechanical properties of the cell.
Endothelial cells subjected to a diabetic environment, both in vivo and in vitro, exhibit a diminished capacity for NOS-induced generation of NO . Consistent with this we observed that both high glucose and L-NAME similarly impaired eNOS expression and reduced NO production. Exposure to high glucose has been reported to impair NO metabolism through production of oxidative stress [5, 27, 28]. Superoxide may interfere with the generation of NO by several mechanisms including a decrease in endothelial eNOS expression mediated by activator protein AP-1, a change in the electrophysiological state of endothelial cells and the availability of tetrahydrobiopterin, an essential cofactor of eNOS. Interestingly hyperglycemia contributes to a switch in eNOS expression in a time-dependent manner. It was found that eNOS protein expression was significantly up-regulated 12 h following exposure to high glucose concentrations (30 mM), reaching a peak at 48h (two fold increase over baseline levels) . Pricci et al. also demonstrated that under high glucose conditions (20 mM), eNOS expression and nitrite/nitrate levels increase the first day, returning to normal levels at day three and diminishing thereafter . Consequently, under high glucose conditions, a compensatory increase in eNOS expression appears at this early stage, thereafter declining gradually as long-term oxidative stress develops. A deleterious effect of intermittent high glucose is also mediated by free radical over-production [3, 4, 29].
In agreement with previous studies, incubation of endothelial cells in constant or intermittent high glucose for seven days decreased not only eNOS expression (Figure 3), but also NO concentration in the culture media (Figure 2). Interestingly, there was no significant difference in NO production between the cells exposed to constant hyperglycemia and those incubated in fluctuating high glucose conditions (Figure 2). However, intermittent high glucose reduced eNOS expression more significantly than a constant level of hyperglycemia (Figure 3). This apparent discrepancy may be related to the observation that endothelial cells exposed to intermittent high glucose are more seriously impaired than those exposed to continuous high glucose due to enhanced oxidative stress [3, 4, 29]. Other reports show that high glucose increases eNOS protein expression, but ultimately leads to decreased NO release .
Hyperglycemia plays an important role in the etiology of endothelial dysfunction [26, 31]. Current data suggest that the deformability of endothelial cells detected by AFM will be affected in the presence of endothelial dysfunction [15–18, 21–23, 32, 33]. Despite this, few studies have used AFM to explore the effects of constant and intermittent high glucose on endothelial cell membrane elasticity. In this study, analysis of 300 force-distance curves obtained using AFM (Figure 5), showed that increased average membrane stiffness of endothelial cells (in response to stable or intermittent high glucose) accompanied decreased NO release and eNOS expression (Figures 2 and 3). Simultaneously, no significant difference existed in membrane stiffness and NO release between continuous and intermittent high glucose groups (Figures 2 and 6). Thus, it is suggested that the increased membrane stiffness of endothelial cells may be affected by the change of NO release more than that of eNOS expression.
In addition to the deleterious effects on endothelial NO-dependent function, high glucose concomitantly increases extracellular osmolality, which may itself impact cellular membrane stiffness. Thus, maintained and intermittent hyperosmolality (as caused by mannitol) was used as an osmotic control for continuous and intermittent high glucose. In the presence of mannitol the release of NO (Figure 2) was unaffected while the expression of eNOS (Figure 3) was slightly decreased. Although abnormal levels of osmolality are likely harmful to endothelial cells, the absence of a NO releasing effect presumably explains why hypertonicity was not shown to increase cell membrane stiffness (Figure 5). In fact, consistent with previous observations, hypertonicity decreased membrane stiffness .
Actin is not only an essential component necessary for maintenance of cellular integrity and function (e.g., membrane polarity, tight junctions, cellular adhesions, and signal transduction), but also undergoes dynamic changes in response to physiological and pathological stresses, including shear stress, vascular pressure and harmful mediators . Several studies have shown how decreases in NO synthesis and eNOS expression induce the redistribution of actin, especially F-actin [9–11, 35, 36]. NO-induced changes in F-actin filaments are proposed to be associated with mobilization of intracellular Ca2+ mediated by the cGMP-dependent pathway, which is activated by cGMP-dependent protein kinase G (PKG) . The stronger F-actin polymerization is also observed when endothelial cells are exposed to D-glucose pre-treated aortic smooth muscle cells, while D-manntinol has no effect on endothelial Ca2+ signaling . TGF-β, a growth factor closely linked to diabetic microvascular complications can stimulate F-actin assembly via activation of NADPH oxidase, which is a mechanism implicated in hyperglycemia . Overwhelming evidence demonstrates that actin redistribution can also regulate NO synthesis and eNOS activity not only through pre-translational mechanisms , but also through posttranslational mechanisms .
Consistent with the aforementioned studies, changes in F-actin relative fluorescence intensity in response to different media (Figures 6 and 7) were associated with decreased NO production (Figure 2) and decreased expression of eNOS (Figure 3). Treatment of endothelial cells with differing concentrations of L-NAME, stable and intermittent high glucose enhanced the apparent thickness of the longitudinal F-actin filaments as demonstrated by an increase in the relative fluorescence intensity of F-actin. Exposure to maintained and intermittent hyperosmolality did not alter NO production nor was, the fluorescence intensity of F-actin significantly different from the control group.
F-actin, as one of the major cytoskeletal components, clusters to form actin filaments, which are bundled and crosslinked by several actin-binding proteins into a network. The actin network plays a major role in determining the mechanical properties of living cells . Furthermore, evidence suggests that depolymerization/polymerization of F-actin filaments results in a dramatic decline/enhancement in endothelial cell membrane stiffness. For example, studies by Cuerrier have demonstrated that laturculin A, an F-actin filament depolymerizing agent, causes a dramatic decline in endothelial cell membrane stiffness . The variation in membrane elasticity in the various regions of endothelial cells (perinuclear or cytoplasmic membrane) is related to the distribution of cytoskeletal elements within these regions . In this study, it was demonstrated that the increased average membrane stiffness of endothelial cells treated with stable/intermittently high glucose and L-NAME was consistent with the change in the corresponding F-actin cytoskeleton (Figures 5, 6 and 7). Therefore, it is speculated that constant and intermittent high glucose, as well as L-NAME treatment, may stiffen endothelial cell membranes by an alteration in F-actin expression and arrangement through the described dysfunction of NO synthesis.
The present study demonstrates that constant and fluctuating high glucose levels, as well as the treatment with L-NAME, enhances average membrane stiffness of intact endothelial cells. Alterations in membrane stiffness and structure may be mediated by the increased expression and redistribution of the F-actin cytoskeleton, which are initiated by high glucose-induced or L-NAME-induced inhibition of eNOS expression and NO synthesis. Importantly, cell membrane stiffness, which is associated with the reorganization of F-actin, can be evaluated with AFM. Thus, AFM, a novel scanning probe for the study of cellular mechanical properties, can be applied to the evaluation of endothelial dysfunction under differing pathological conditions.
The work was partially supported by the Department of Chemistry, Jinan University, Guangdong, China. This research received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.
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