Abstract
N-methyl-D-aspartate receptors (NMDARs) are ligand-gated, voltage-dependent channels of the ionotropic glutamate receptor family. The present study explored whether NMDAR activation induced ferroptosis in vascular endothelial cells and its complicated mechanisms in vivo and in vitro. Various detection approaches were used to determine the ferroptosis-related cellular iron content, lipid reactive oxygen species (LOS), siRNA molecules, RNA-sequence, MDA, GSH, and western blotting. The AMPK activator Acadesine (AICAR), HMGB1 inhibitor glycyrrhizin (GLY), PP2A inhibitor LB-100, and NMDAR inhibitor MK801 were used to investigate the involved in vivo and in vitro pathways. The activation of NMDAR with L-glutamic acid (GLU) or NMDA significantly promoted cellular ferroptosis, iron content, MDA, and the PTGS2 expression, while decreasing GPX4 expression and GSH concentration in human umbilical vein endothelial cells (HUVECs), which was reversed by ferroptosis inhibitors Ferrostatin-1(Fer-1), Liproxstatin-1 (Lip-1), or Deferoxamine (DFO). RNA-seq revealed that ferroptosis and SLC7A11 participate in NMDA or GLU-mediated NMDAR activation. The PP2A-AMPK-HMGB1 pathway was majorly associated with NMDAR activation-induced ferroptosis, validated using the PP2A inhibitor LB-100, AMPK activator AICAR, or HMGB1 siRNA. The role of NMDAR in ferroptosis was validated in HUVECs induced with the ferroptosis activator errasin or RSL3 and counteracted by the NMDAR inhibitor MK-801. The in vivo results showed that NMDA- or GLU-induced ferroptosis and LOS production was reversed by MK-801, LB-100, AICAR, MK-801, and GLY, confirming that the PP2A-AMPK-HMGB1 pathway is involved in NMDAR activation-induced vascular endothelium ferroptosis. In conclusion, the present study demonstrated a novel role of NMDAR in endothelial cell injury by regulating ferroptosis via the PP2A-AMPK-HMGB1 pathway.
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Introduction
Vascular endothelial cells (VECs), located in an oxygen-rich vascular environment, are vital for supporting the structure and biological function of blood vessels and maintaining vascular tension by mediating the balance of vasodilation, contraction, growth inhibition, and growth promotion [1] as well as anti-inflammatory or pro-inflammatory effects [2, 3]. VEC dysfunction may result in arteriosclerosis [4, 5] and cardiovascular diseases such as thromboangiitis obliterans [6, 7], which are closely related to cardiovascular risk factors. In contrast to atherosclerosis, endothelial dysfunction can also be reversed. Therefore, early prevention and treatment should be performed to prevent vascular disease [8,9,10,11]. Ferroptosis is a new type of regulated cell death [12, Human umbilical vein endothelial cells (HUVECs) were obtained from ScienCell Research Laboratories (San Diego, CA, USA) and cultured in plates pre-coated with 0.2% gelatin in endothelial cell medium supplemented with 5% fetal bovine serum, 1% penicillin/streptomycin, and 1% endothelial cell growth supplement (ScienCell) at 37 °C with 5% CO2. For each experiment using cultured HUVECs, cells were seeded in the incubator and cultured to 70–80% confluence, then exposed to L-glutamic acid (GLU, 20 mM, Cat#: G8415, Sigma-Aldrich, St. Louis, MO, USA) or N-Methyl-D-aspartic acid (NMDA, 1 mM, Cat#: HY-17551, MedChemExpress, Shanghai, China) for 24 h in the absence or presence of Ferrostatin-1 (Fer-1, 10 μM, Cat#: HY-100579, MedChemExpress), Liproxstatin-1 (Lip-1, 2 μM, Cat#: HY-12726, MedChemExpress), LB-100 (5 μM, Cat#: S7537, Sellechchem, Houston, TX, USA), Acadesine (AUCAR, 2 μM, Cat#: HY-13417, MedChemExpress), Deferoxamine (DFO, 10 μM, Cat#: HY-B0988, MedChemExpress), and Erastin (5 μM, Cat#: HY-15763, MedChemExpress) for 24 h. These plating conditions were used in all experiments, unless mentioned otherwise. Cell viability was assessed using the 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) assay (Solarbio Technology, Bei**g, China), as described previously. Briefly, HUVECs were seeded in 96-well plates and cultured for 24 h. Cells were pretreated with the corresponding inhibitors or small interfering RNA (siRNA), and then exposed to a designated concentration of NMDA and GLU for the indicated times. After treatment, the cells were incubated with 0.5 mg/mL MTT for 4 h and resuspended in 150 μL DMSO (Sigma-Aldrich, CA, USA). The absorbance was measured at 495 nm using an Infinite M200 Pro NanoQuant (TECAN, Switzerland). FerroOrange (Do**do Laboratories, Kumamoto, Japan) was used to determine Fe2+ levels by immunofluorescence staining. Briefly, HUVECs were plated on confocal culture dishes and treated with GLU or NMDA in the absence or presence of Fer-1, Lip-1, and DFO for 24 h. Cells were washed three times with phosphate-buffered saline (PBS), and images were captured using a scanning confocal microscope (EVOS™ FL Auto 2 Imaging System, Thermo Fisher Scientific, MA, USA). The treated HUVECs were harvested and lysed by sonication at 0 °C for 20 min, followed by centrifugation at 15,000 × g at 4 °C for 10 min. The cleared supernatant was prepared to detect the total protein concentration using a Protein Assay Kit (Thermo Fisher Scientific) and to measure the amount of GSH using the GSSG/GSH Quantification Kit. To remove proteins from samples, 1/6 volume of 5% 5-sulfosalicylic acid dihydrate (Wako Pure Chemical Corporation, Japan) in distilled water was used. Samples were centrifuged at 15,000 × g at 4 °C for 10 min, and the supernatant was used for the GSH assay. The protein and GSH levels in the samples were detected according to the manufacturer’s instructions by measuring the absorbance at 405 nm using a plate reader. Values for total GSH levels were calculated, corrected for protein concentration in the same sample, and normalized to the control. MDA (A003-1; Nan**g Jiancheng Bioengineering Institute, Nan**g, China) and LPO (A106-1, Nan**g Jiancheng Bioengineering Institute) in vascular tissues or cells were measured using commercial kits, according to the manufacturer’s instructions. Briefly, cells (1 × 106) were collected in 200 μL of lysis buffer and homogenized on ice. Subsequently, an MDA test solution (1000 μL) was added to each experimental sample or vial containing the standard sample to form an MDA-TBA adduct, followed by incubation for 40 min at 95 °C. For LPO analysis, 800 μL of the LPO test solution was added to each experimental sample or vial containing the standard sample and then incubated for 60 min at 45 °C. The mixture was cooled to 25 °C in an ice bath and centrifuged at 4000 rpm for 10 min. The insoluble material was removed, and 250 μL of each reaction mixture was pipetted into 96-well plates for colorimetric assays to measure the absorbance of MDA and LPO at 530 nm and 586 nm, respectively, with an Infinite M200 Pro NanoQuant (TECAN). siRNA was used to silence the expression of specific genes in HUVECs. The cells at 60–70% confluence were transfected with specific siRNA duplexes (60 nM, Santa Cruz Biotechnology, CA, USA) using Lipofectamine RNAiMAX Reagent (Thermo Fisher Scientific) following the manufacturer’s instructions. After 48 h of transfection with control siRNA, HMGB1 siRNA, PP2A siRNA, and AMPK siRNA (Santa Cruz Biotechnology), the cells were incubated with GLU or NMDA for the indicated times and then collected for western blot analysis and cell viability. The animal study protocol was approved by the Animal Care and Ethics Committee of ** according to the complete random number method; experimental operation and data analysis were according to animal No. and not grouped information.): (1) control (n = 8); (2) GLU (1 g/kg, n = 8); (3) GLU + LB-100 (n = 8, 2 mg/kg); (4) GLU + AICAR (n = 8, 100 mg/kg); (5) GLU + MK-801 (0.5 mg/kg, n = 8); (6) GLU + Glycyrrhizin acid (GLY, n = 8, 20 mg/kg, Cat#: HY-N0184, MedChem Express); (7) NMDA (75 mg/kg, n = 8); (8) NMDA + LB-100 (2 mg/kg, n = 8); (9) NMDA + AICAR (100 mg/kg, n = 8); (10) NMDA + MK-801(0.5 mg/kg, n = 8); and (11) NMDA + GLY (20 mg/kg, n = 8). Animals were intraperitoneally injected with different compounds twice daily for ten consecutive days, and control animals received equal volume of vehicle (0.9% saline) or DMSO subcutaneously. All animals were maintained at room temperature (23 ± 2 °C) with a 12 h light/dark cycle and free access to a basic diet and water. When the animals were sacrificed at the end of the experiment, vascular tissues were collected for further data analysis. Cell fluorescence levels of total intracellular ferrous ion and LOS were detected using FerroOrange (10 μM) and Liperfluo (1 μM), respectively. When cells were treated with the test compounds at the indicated times, cells in 6-well dishes were resuspended in 500 ml of fresh Hank’s balanced salt solution (HBSS) for Liperfluo evaluation for 30 min at 37 °C in a culture incubator, and cells in 24-well plates were used for FerroOrange staining in HBSS for 30 min at 37 °C. After washing three times, intracellular FerroOrange fluorescence and LOS were determined in the single-cell suspension using a flow cytometer (Beckman Coulter, USA) with an excitation wavelength of 488 nm and an emission wavelength of 525 nm. Increased percentages of LOS levels were determined by quantifying and calculating the ratio of 488 nm channel intensity in treated groups compared to that in control groups. Vascular tissues isolated from mice or cultured HUVECs were homogenized in saline or PBS. The protein concentration was measured using the BCA Protein Assay Kit. The levels of iron in the blank (ddH2O), iron standard solution, and test samples (supernatant) were examined using an Iron Assay Kit (TC1015; Leagene Biotechnology, Bei**g, China) according to the manufacturer’s instructions. The reaction mixture was then incubated at room temperature for 15 min. The absorbance was measured at 562 nm using a microplate reader (Infinite M200 Pro Nano Quant, TECAN). Vascular tissues isolated from mouse or cultured HUVECs were homogenized in saline or PBS, followed by centrifugation. PP2A activity was measured according to the manufacturer’s protocol (R&D Systems, DYC3309-2). Western blot analysis was used to determine the expression of specific proteins in the vascular tissues and cultured HUVECs. Proteins of tissue homogenate lysates or HUVECs lysates prepared in sodium dodecyl sulfate (SDS) lysis buffer were extracted with RIPA buffer supplemented with protease and phosphatase inhibitors on ice, and protein concentration was determined using the BCA Protein Assay Kit (Solarbio), as described previously. SDS-PAGE and polyvinylidene difluoride membranes (Bio-Rad, Hercules, CA, USA) were used to separate proteins. The membranes were blocked with 5% fat-free dry milk in 0.1% Tris-buffered saline with Tween for 2 h and then probed with different primary antibodies at 4 °C overnight, including anti-PTGS2 (abcom, ab283574, 1:1000), Anti-Glutathione Peroxidase 4 antibody (abcom, ab125066, 1:1000), Anti-Transferrin Receptor antibody (abcom, ab214039, 1:1000), Anti-xCT antibody (abcom, ab175186, 1:1000), Anti-PP2A antibody (abcom, ab32104, 1:1000), Anti-Phospho-PP2A antibody (santacruz, sc-271903, 1:1000), Anti-AMPKα antibody (CST, #2532, 1:1000), Anti-Phospho-AMPKα antibody (CST, #2535, 1:1000), Anti-HMGB1 antibody (abcom, ab79823, 1:10000) and Anti-β-Actin antibody (santacruz, sc-47778, 1:1000). After washing, the membranes were incubated with the secondary antibody (1:10000) for 1 h at room temperature. Blots were visualized using ECL ™ reagents (Advansta, Menlo Park, CA, USA). Protein signals were captured using the FluorChem E chemiluminescence detection system (ProteinSimple, San Jose, CA, USA). All western blots were performed at least five times. The signal intensity of the immunoreactive bands was quantified using ImageJ software (NIH, Bethesda, MD, USA) and normalized to that of β-actin in each sample. VECs cultured with NMDA and GLU for 24 h were collected and prefixed in 2.5% glutaraldehyde phosphate (0.1 M, pH 7.4) overnight at 4 °C, post-fixed in 2% buffered osmium tetraoxide, and then embedded in Epon812 (Merck, NJ, USA), followed by dehydration. Ultrathin sections (60 nm thick) were cut and stained with uranyl acetate as well as lead citrate. Images were examined using a Hitachi HT7800 TEM instrument (Tokyo, Japan). Statistical analyses were performed using GraphPad Prism 9.0 (GraphPad Software Inc., San Diego, CA, USA). All results are presented as the mean ± SEM from at least three separate experiments unless otherwise described. One-way analysis of variance followed by the Bonferroni post-hoc test was used to compare the groups. The variance was similar among the groups that were being statistically compared. Statistical significance was set at P < 0.05. The exact sample size was provided in the figure legend.Materials and methods
Cell culture and exposure
Cell viability
Immunofluorescence for Fe2+ analysis
Glutathione (GSH) measurement
Measurement of malondialdehyde (MDA) and lipid peroxidation (LPO) levels
siRNA
Animal experiments
Detection of intracellular ferrous ions and lipid reactive oxygen species (LOS)
Iron content assay
PP2A activity assay
Western blot analysis
AMPK activity assay
Statistical analysis
Data availability
All the data and experimental details in this article may be obtained from the corresponding author upon reasonable request.
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Funding
This work was supported by Fujian provincial health technology project (Grant No. 2021QNB022, Rui-Ying Wang) and the National Natural Science Foundation of China (Grant No. 82270417, Gang Li; Grant No. 82200373, Rui-Ying Wang).
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LG designed experiments and guided the whole project. WRY wrote and modified manuscript. XGS analyzed the data and supervised the work. HWM and HYX conducted the experiments in vitro and in vivo. LG and WRY provided funding for all experiments. All authors read and approved the final manuscript.
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The animal study protocol was reviewed approved by the Animal Care and Ethics Committee of **amen University (**amen, Fujian, China).
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Han, WM., Hong, YX., **ao, GS. et al. NMDARs activation regulates endothelial ferroptosis via the PP2A-AMPK-HMGB1 axis. Cell Death Discov. 10, 34 (2024). https://doi.org/10.1038/s41420-023-01794-3
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DOI: https://doi.org/10.1038/s41420-023-01794-3
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