CFTR limits F-actin formation and promotes morphological alignment with flow in human lung microvascular endothelial cells

Abstract

Micro- and macrovascular endothelial dysfunction in response to shear stress has been observed in cystic fibrosis (CF), and has been associated with inflammation and oxidative stress. We tested the hypothesis that the cystic fibrosis transmembrane conductance regulator (CFTR) regulates endothelial actin cytoskeleton dynamics and cellular alignment in response to flow. Human lung microvascular endothelial cells (HLMVEC) were cultured with either the CFTR inhibitor GlyH-101 (20 µM) or CFTRinh-172 (20 µM), tumor necrosis factor (TNF)-α (10 ng/ml) or a vehicle control (0.1% dimethyl sulfoxide) during 24 and 48 h of exposure to shear stress (11.1 dynes/cm² ) or under static control conditions. Cellular morphology and filamentous actin (F-actin) were assessed using immunocytochemistry. [Nitrite] and endothelin-1 ([ET-1]) were determined in cell culture supernatant by ozone-based chemiluminescence and ELISA, respectively. Treatment of HLMVECs with both CFTR inhibitors prevented alignment of HLMVEC in the direction of flow after 24 and 48 h of shear stress, compared to vehicle control (both p < 0.05). Treatment with TNF-α significantly increased total F-actin after 24 h versus control (p < 0.05), an effect that was independent of shear stress. GlyH-101 significantly increased F-actin after 24 h of shear stress versus control (p < 0.05), with a significant (p < 0.05) reduction in cortical F-actin under both static and flow conditions. Shear stress decreased [ET-1] after 24 h (p < 0.05) and increased [nitrite] after 48 h (p < 0.05), but neither [nitrite] nor [ET-1] was affected by GlyH-101 (p > 0.05). CFTR appears to limit cytosolic actin polymerization, while maintaining a cortical rim actin distribution that is important for maintaining barrier integrity and promoting alignment with flow, without effects on endothelial nitrite or ET-1 production.

Keywords: actin cytoskeleton; cystic fibrosis; pulmonary microvascular endothelium; shear stress.

FIGURE 1

The calculation of Feret's Angle of human lung microvascular endothelial cells. After the nucleus was stained with 0.125% crystal violet, the direction of alignment was quantified by identifying the angle formed by the longest axis of the cell (Feret's Diameter) with the abscissa axis (Feret's Angle). Each image contained >500 cells and images were collected from two randomly selected areas in each well under each incubation condition in three independent experiments

FIGURE 2

Human lung microvascular endothelial cells alignment in response to static (black bars) or shear stress (11.1 dynes/cm²; gray bars) incubation, and following treatment with vehicle control (0.1% dimethyl sulfoxide [DMSO]), GlyH‐101 (20 μM), or TNF‐α (10 ng/ml) for 24 or 48 h. Nuclei were stained with 0.125% crystal violet and images were captured at 10× magnification. Uniformity of alignment was quantified by determining the SD of the Feret's Angle for each cell within an image (>500 cells per image). Images were captured from two separate areas in a single well of three independent experiments (n = 3). HLMVECs were incubated under static conditions for 24 h (a, b, c) or 48 h (g, h, i), or under orbital flow conditions for 24h (d, e, f) or 48 h (j, k, l), in the absence (a, d, g, j) or presence of GlyH‐101 (b, e, h, k) or TNF‐α (c, f, i, l). Analysis of the variance of Feret's angle at 24h (m) and 48h (n) is shown for each of these conditions. Data were expressed as mean ± SEM. *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001

FIGURE 3

Human lung microvascular endothelial cells alignment in response to static (black bars) or shear stress (11.1 dynes/cm²; gray bars) incubation for 24 or 48 h, and following treatment with vehicle control (0.1% dimethyl sulfoxide) or CFTRinh‐172 (20 μM). Nuclei were stained with 0.125% crystal violet and images were captured at 10× magnification. Uniformity of alignment was quantified by determining the SD of the Feret's Angle for each cell within an image (>500 cells per image). Images were captured from two separate areas in a single well of three independent experiments (n = 3). Data were expressed as mean ± SEM. *p < 0.05

FIGURE 4

Human lung microvascular endothelial cells filamentous actin (F‐actin) formation in response to static (black bars) or shear stress (11.1 dynes/cm²; gray bars) incubation, and following treatment with vehicle control (0.1% dimethyl sulfoxide [DMSO]), GlyH‐101 (20 μM) or TNF‐α (10 ng/ml) for 24 h. Images included in this figure were captured at 20× magnification with 400 ms exposure to fluorescence (a–f). Images used for quantification were exposed to 200 ms of fluorescence. Images were captured from three randomly selected areas of a single well in three independent experiments (n = 3). The RawIntDen of F‐actin (Red; 100 nM of Rhodamine Phalloidin) was expressed relative to nuclei (blue; 5 μg/ml Hoechst 33342) to estimate F‐actin per cell (g). Peripheral F‐actin was analyzed using the ‘Plot Profile’ tool in ImageJ and expressed as the area under the curve for the peaks in the profile representing peripheral F‐actin (h). Data were expressed as mean ± SEM. Yellow arrows highlight paracellular gaps; white arrows highlight cortical rim F‐actin; *p < 0.05; **p < 0.01

FIGURE 5

Human lung microvascular endothelial cells culture supernatant nitrite concentrations in response to static or shear stress (11.1 dynes/cm²) incubation over time (a), and following treatment with vehicle control (0.1% dimethyl sulfoxide) or GlyH‐101 (20 μM; b). Samples were collected from duplicate wells of three independent experiments (n = 3). Data were expressed as mean ± SEM. *p < 0.05; **p < 0.01

FIGURE 6

Human lung microvascular endothelial cells culture supernatant ET‐1 concentrations in response to static (black bar) or shear stress (gray bar; 11.1 dynes/cm²) incubation over time (a), and following treatment with vehicle control (0.1% dimethyl sulfoxide) or GlyH‐101 (20 μM) for 24 h (black bar) and 48 h (gray bar; b). Samples were collected from duplicate wells of three independent experiments (n = 3). Data were expressed as mean ± SEM