Toxic effect of silica nanoparticles on endothelial cells through DNA damage response via Chk1-dependent G2/M checkpoint

Abstract

Silica nanoparticles have become promising carriers for drug delivery or gene therapy. Endothelial cells could be directly exposed to silica nanoparticles by intravenous administration. However, the underlying toxic effect mechanisms of silica nanoparticles on endothelial cells are still poorly understood. In order to clarify the cytotoxicity of endothelial cells induced by silica nanoparticles and its mechanisms, cellular morphology, cell viability and lactate dehydrogenase (LDH) release were observed in human umbilical vein endothelial cells (HUVECs) as assessing cytotoxicity, resulted in a dose- and time- dependent manner. Silica nanoparticles-induced reactive oxygen species (ROS) generation caused oxidative damage followed by the production of malondialdehyde (MDA) as well as the inhibition of superoxide dismutase (SOD) and glutathione peroxidase (GSH-Px). Both necrosis and apoptosis were increased significantly after 24 h exposure. The mitochondrial membrane potential (MMP) decreased obviously in a dose-dependent manner. The degree of DNA damage including the percentage of tail DNA, tail length and Olive tail moment (OTM) were markedly aggravated. Silica nanoparticles also induced G2/M arrest through the upregulation of Chk1 and the downregulation of Cdc25C, cyclin B1/Cdc2. In summary, our data indicated that the toxic effect mechanisms of silica nanoparticles on endothelial cells was through DNA damage response (DDR) via Chk1-dependent G2/M checkpoint signaling pathway, suggesting that exposure to silica nanoparticles could be a potential hazards for the development of cardiovascular diseases.

Figure 1. Characterization of silica nanoparticles.

(A) Transmission electron microscopy image: TEM images of silica nanoparticles had a spherical shape with the average diameter of 62 nm. (B) Size distribution: The size distribution measured by ImageJ software showed approximately normal distribution.

Figure 2. Subcellular localization of silica nanoparticles.

(A) LSCM images of HUVECs after incubation for 24 h with Ruthenium (II) hydrate labeled silica nanoparticles (50 µg/mL, red) of size 62 nm. The cell skeleton was stained with Phalloidin-FITC (green), and the cell nucleus with 4,6-diamidino-2-phenylindole (DAPI; blue). (B) TEM images of HUVECs exposed to silica nanoparticles for 24 h. Both TEM and LSCM results showed that the silica nanoparticles were internalized into cells compared to control group.

Figure 3. Cytotoxicity of HUVECs induced by silica nanoparticles.

(A) Morphological changes of HUVECs after exposure to silica nanoparticles for 24 h. Cell density reduction, irregular shape and cellular shrinkage were observed by optical microscope. (B) Cell viability of HUVECs treated with silica nanoparticles was measured by MTT assay after 6 h, 12 h, 24 h exposure. (C) LDH leakage of HUVECs exposed to viarous concentrations of silica nanoparticles for 24 h. The results indicated that silica nanoparticles induced cytotoxicity in a dose- and time-dependent manner. Data are expressed as means ± S.D. from three independent experiments (*p<0.05).

Figure 4. Apoptosis of HUVECs after exposure to silica nanoparticles for 24 h.

(A) Apoptotic and necrotic populations of cells double-stained with PI- and FITC-labled Annexin V were depicted by flow cytometry. FITC negative and PI negative were designated as live cells in the lower left quadrant; FITC positive and PI negative as apoptotic cells in the upper left quadrant; FITC positive and PI positive as necrotic cells in the upper right quadrant; and FITC negative and PI positive as large nuclear fragments in the lower right quadrant. (B) HUVECs exposure to silica nanoparticles caused increase of both necrosis and apoptosis rate. The apoptosis rate was much lower than the necrosis rate. Data are expressed as means ± S.D. from three independent experiments (*p<0.05).

Figure 5. Oxidative stress and oxidative damage induced by silica nanoparticles on HUVECs.

The intracellular levels of ROS and MDA were obviously increased (A, B). While SOD and GSH-Px levels were decreased significantly with a dose-dependent way (C, D). Silica nanoparticles-induced ROS generation caused oxidative damage followed by the production of MDA as well as the inhibition of SOD and GSH-Px. Data are expressed as means ± S.D. from three independent experiments (*p<0.05).

Figure 6. Mitochondrial membrane potential changes after silica nanoparticles exposure for 24 h detected with JC-1 probe by flow cytometry.

The green/red fluorescence intensity ratio was used to express the changes of MMP and the increased ratio indicates decrease of MMP. Data are expressed as means ± S.D. from three independent experiments (*p<0.05).

Figure 7. DNA damage of HUVECs after exposed to silica nanoparticles for 24 h determined by comet assay.

(A) Control group, (B) 25 µg/mL treated group, (C) 50 µg/mL treated group, (D) 75 µg/mL treated group, (E) 100 µg/mL treated group. More severe DNA injury was reflected by larger area of the comet tail. The DNA damage caused by silica nanoparticles was getting more serious with the dosages increasing. The magnification was 200× by fluorescence microscope.

Figure 8. Cell cycle arrest of HUVECs induced by silica nanoparticles.

After exposure to various concentrations of silica nanoparticles for 24 h, flow cytometry were used to determine the cell cycle distribution of HUVECs. The images showed that cell cycle was arrested in G2/M phase. The percentage of cells in G2/M phase increased progressively in a dose-dependent manner, while in G0/G1 and S phase the percentage of cells declined irregular.

Figure 9. Effects of silica nanoparticles on G2/M DNA damage checkpoint signaling pathway

. (A) Effect of silica nanoparticles on the expression of Chk1, Cdc25C, cyclin B1, Cdc2. GAPDH was used as an internal control to monitor for equal loading. (B) Relative densitometric analysis of the proteins bands was performed and presented. Silica nanoparticles induced G2/M arrest through the upregulation of Chk1 and the downregulation of Cdc25C, cyclin B1/Cdc2. Data are expressed as means ± S.D. from three independent experiments (*p<0.05).