CTRP13 Mitigates Endothelial Cell Ferroptosis via the AMPK/KLF4 Pathway: Implications for Atherosclerosis Protection

Background

Atherosclerosis cardiovascular disease has become a global public health concern because it causes the death of more people than all cancers combined and is one of the leading causes of morbidity and mortality worldwide [1]. Atherosclerosis characteristics include lipid substance accumulation, pro-inflammatory response in macrophages, endothelial cell injury, and arterial wall sclerosis. Recent evidence demonstrates that atherosclerosis begins long before clinical manifestations, with onsets that can be traced back to adolescence or early adulthood [2]. New anti-atherosclerosis agents are required to supplement our current arsenal of anti-atherosclerosis drugs and provide therapeutic options for all patients at high risk for atherosclerosis.

The mechanism of atherosclerosis remains to be established. Some scholars believe hyperlipidemia and inflammation-induced endothelial dysfunction are crucial initiating factors and key links to atherosclerosis development. Extensive research has demonstrated that endothelial dysfunction is a common feature of atherosclerosis. Endothelial dysfunction results from increased oxidative stress and nitric oxide (NO) reduction [3–5]. Ferroptosis is a new type of regulatory cell death characterized primarily by lipid peroxidation and iron dependence but lacks the typical apoptosis and necrosis manifestations [6]. Ferroptosis participates in various pathological processes, including cancers, atherosclerosis, and cardiomyopathy. Our previous studies revealed that high glucose or oxidized low-density lipoprotein (ox-LDL) stimulation reduces endothelial NO synthase (eNOS)/NO bioavailability and inhibits the AMP-activated kinase (AMPK) pathway in cardiomyocytes and human umbilical vein endothelial cells (HUVECs) [7,8]. However, little is known about how ferroptosis impacts HUVECs, particularly under ox-LDL conditions.

In contrast to other cell death pathways, ferroptosis has shrunken mitochondria, increased membrane density, and decreased or disappeared mitochondrial ridges. Ferroptosis is a non-apoptotic cell death characterized by an imbalance of iron metabolism and lipid peroxidation [9]. Recent evidence suggests elevated extracellular glutamate levels present in atherosclerosis-inhibited system xc transportation (xCT), triggering ferroptosis [10]. Abundant experimental data support that ferroptosis inhibition can attenuate lipid peroxidation and endothelial dysfunction in endothelial cells. Lv et al indicated that treating HUVECs with ox-LDL or the ferroptosis inducer erastin and estradiol inhibited iron death in HUVECs by activating the NRF2/glutathione peroxidase 4 (GPX4) pathway [11]. Liu et al found that HUVECs phagocytized large qualities of γ-Fe2O3 and Fe3O4 nanoparticles, increasing intracellular iron levels. They further observed the disrupted cystine/glutamate reverse transporter and GPX4 signaling in γ-Fe2O3 and Fe3O4 nanoparticle-challenged HUVECs. The γ-Fe2O3 and Fe3O4 nanoparticles could also cause mitochondrial fusion and fission dysregulation and activate lipid peroxidation and iron metabolism-related genes in a P53-dependent manner [12]. Bai et al demonstrated that ferroptosis inhibition alleviates atherosclerosis by attenuating lipid peroxidation and endothelial dysfunction in mouse aortic endothelial cells [13]. Vascular endothelial cells play an essential role, forming the barrier between blood and tissues, regulating blood vessel function, and maintaining a stable internal environment [14]. Oxidative stress and mitochondrial dysfunction can cause endothelial dysfunction in the initial atherosclerosis stage. Moreover, iron overload causes endothelial dysfunction by aggravating the abnormal mitochondria structure and oxidation reaction in endothelial cells [13]. Whether ferroptosis is related to endothelial dysfunction is an interesting remaining question. Hence, in this study, we investigated the regulatory mechanism and important role of ferroptosis on endothelial dysfunction.

C1q/tumor necrosis factor-related protein 13 (CTRP13), an adipokine secreted by adipocytes, has many systemic effects and important roles in developing atherosclerosis and cardiovascular diseases. CTRP13 has a restored mitochondrial calcium ion balance, lowering reactive oxygen species (ROS) production [15]. Recent studies have discovered that CTRP13 significantly recovered NO production and eNOS coupling in the aortae of diabetic mice and high glucose-treated HUVECs. These findings showed that CTRP13 treatment reversed the downregulation of GCH1 and the ratio of eNOS dimers to monomers in the renal arteries of patients with diabetes [4]. Soudabeh et al demonstrated that lower circulating levels of CTRP13 increased the risk of coronary artery disease and type 2 diabetes mellitus [16]. To the best of our knowledge, the molecular mechanism and pharmacological action of CTRP13 in endothelial cell ferroptosis remains uninvestigated.

Krüppel-like factor 4 (KLF4) is a zinc finger-containing transcription factor, the expression of which is connected with growth arrest. KLF4 participates in the regulation of processes including cell proliferation, differentiation, invasion, and migration [17,18]. Studies have shown that the knockdown of KLF4 by siRNA enhances the proliferation inhibitory effect of Polyphyllin III by promoting ferroptosis in breast cancer [19]. However, it remains unknown whether KLF4 mediates endothelial cell ferroptosis. AMPK is a serine/threonine kinase [20,21]. Accumulating evidence indicates that AMPK acts as an important regulator in the pathogenesis of cardiovascular diseases [22,23]. Kurabayashiet et al indicated that dysregulation of Elovl6-driven long-chain fatty acid metabolism induces phenotypic switching of vascular smooth muscle cells via ROS production and AMPK/KLF4 signaling, which leads to growth arrest and downregulation of vascular smooth muscle cell marker expression [24].

This study aimed to investigate the effects of CTRP13 on HUVECs ferroptosis induced by ox-LDL and to explore the underlying mechanisms of CTRP13 against ferroptosis via activating the AMPK/KLF4 pathway.

Material and Methods

CELL CULTURE AND TREATMENT:

HUVECs were obtained from Science Cell Research Laboratories (Carlsbad, CA, USA), and the purity of HUVECs was more than 89.67%, which was confirmed by immunofluorescence identification (iCell Bioscience Inc, Shanghai, China). Cells were cultured in Dulbecco’s modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (Gibco, Life Technologies, Waltham, MA, USA), 100 U/mL penicillin, and 0.1 mg/mL streptomycin (Gibco, Grand Island, NY, USA). The cells were cultured in an incubator under 5% CO2 at 37°C, and the culture medium was exchanged every 2 days. The cells within 7 passages were used for in vitro study. The cells were treated with trypsin (Beyotime Biotechnology, Shanghai, China) and seeded in plates for reagent intervention. Ox-LDL (YB-002) was obtained from Yiyuan Biotechnology, Guangzhou, China. HUVECs were cultured in DMEM containing ox-LDL (0–200 μg/mL) for ox-LDL treatment for 24 or 48 h [25].

CHEMICALS AND REAGENTS:

CTPR13 (purity >99.37%) was purchased from R&D Systems (9115-TN) and reconstituted at 500 μg/mL in water. Ferrostatin-1 (Fer-1) was purchased from Sigma (SML0583), and a 5-mmol/L stock solution was prepared in dimethyl sulfoxide (Sigma, St. Louis, MO). APTO (HY-16291, MedChem Express) was used as an inducer of KLF4. Primary antibodies against GPX4, recombinant solute carrier family 7, member 11 (SLC7A11), acyl-CoA synthetase long-chain family member 4 (ACSL4), heme oxygenase 1 (HO-1), and KLF4 were obtained from Abcam (Cambridge, MA, USA). Primary antibodies against phosphor-specific anti-AMPK (Thr172), AMPK, superoxide dismutase 1 (SOD-1), and SOD-2 were obtained from Cell Signaling Technology (Danvers, MA, USA; 1: 1000 dilution). β-actin, Goat anti-Rabbit IgG, and HRP Goat anti-Mouse IgG were provided by ZSGB-BIO (Beijing, China). Biological reagents, such as JC-1, Mito-Tracker Red, and 2′,7′-dichlorofluorescein di-acetate (DCFH-DA), were obtained from Beyotime Biotechnology, China. Nucleic acid sequences were obtained from Integrated DNA Technologies (Genewiz, IA, USA).

REVERSE TRANSCRIPTION-QUANTITATIVE POLYMERASE CHAIN REACTION:

Total cellular RNA was extracted using TRIzol reagent (Invitrogen, CA, USA). Real-time reverse transcription-polymerase chain reaction (RT-PCR) was performed using the SYBR Premix reagent (cat# 1725125, BIO-RAD) and ABI Fast 7500 Real-time PCR System (Applied Biosystems, South San Francisco, CA). The 2(−ΔΔCt) method was used to calculate relative fold changes with GAPDH as the internal control. Primers for real-time RT-PCR are listed in Table 1.

CELL COUNTING KIT-8 ASSAY:

Cell Counting Kit-8 (CCK-8) assay was used per the manufacturer’s instructions. Briefly, 5×10³ cells/well were cultured in a 96-well plate for specific time points (24 or 48 h) at different concentrations. Then, 10 μL of CCK-8 (Dojindo, Kumamoto, Japan) reagent was added to each cell. The cells were further incubated for 2 h at 37°C. The absorbance was measured at 450 nm using a microplate reader (Bio-Rad Laboratories, Inc.).

TRANSMISSION ELECTRON MICROSCOPY:

HUVECs were seeded in 6-well plates. After treatment, the cells were digested with trypsin and re-suspended in phosphate-buffered saline (PBS) twice. The samples were obtained by centrifuging at 3000 rpm. The supernatant was removed and fixed with 2.5% glutaraldehyde overnight at 4°C. Finally, the mitochondria ultrastructure was observed using transmission electron microscopy.

HOECHST/PROPIDIUM IODIDE STAINING:

Hoechst/propidium iodide (PI) staining was performed according to the manufacturer’s instructions. HUVECs (1×10⁵ cells/well) were cultured in a 12-well plate to detect cell membrane integrity. The cells were washed twice with PBS and stained with Hoechst staining (5 μg/mL; staining for all cell nuclei) and PI (2 μg/mL; staining for membrane-damaged cells) at 4°C for 30 min. Finally, cells were washed with ice-cold PBS once and observed under a fluorescent microscope (Olympus, Japan).

MEASUREMENT OF LACTATE DEHYDROGENASE ACTIVITY:

The culture supernatant was obtained after centrifugation and assayed for lactate dehydrogenase (LDH) activity (Beyotime, Shanghai, China, C0017). Then, 120 μL of supernatant from each well was added to the corresponding well of a new 96-well plate. The absorbance was measured at 490 nm.

IRON CONTENT ASSAY:

Intracellular iron ion levels were examined using an iron assay kit (TC1015, Leagene, Beijing, China) according to the manufacturer’s instructions. Briefly, the supernatant was centrifuged at a low speed, mixed, and incubated at 37°C for 10 min. Then, the absorbance was measured at 562 nm.

MEASUREMENT OF INTRACELLULAR OXIDATIVE STRESS LEVELS:

Total ROS production was detected using the fluorescence probe DCFH-DA (S0033S, Beyotime, Shanghai, China) according to the manufacturer’s instructions. DCFH-DA was diluted in serum-free medium at 1: 1000 to a final concentration of 10 mM and incubated in the cells at 37°C for 20 min. Then, the absorbance was measured at 488 and 525 nm. A commercially available malondialdehyde (MDA) assay kit (Beyotime Institute of Biotechnology) was used to measure lipid peroxidation parameters by detecting MDA content, following the instructions for when cells undergo oxidative stress. The γ-glutamyl-cysteinyl-glycine (GSH) level was measured according to the manufacturer’s instructions (Beyotime, China).

IMMUNOFLUORESCENCE:

The HUVECs were fixed with 4% paraformaldehyde for 15 min and blocked with 5% bovine serum albumin for 30 min. Then, the cells were incubated overnight at 4°C with anti-GPX4 antibody (1: 200; Abcam; Cambridge, MA, USA), followed by incubation with horseradish peroxidase (HRP)-conjugated secondary antibody (Amresco) for 1 h at room temperature in the dark. The nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI) for 5 min. Subsequently, the stained cells were observed and photographed using a fluorescence microscope (Olympus, Japan).

MITOCHONDRIAL MEMBRANE POTENTIAL ASSAY:

Mito-Tracker staining (C1049B, Beyotime Institute of Biotechnology) specifically labeled mitochondria within live cells utilizing the mitochondrial membrane potential. JC-1 (C2003S, Beyotime Institute of Biotechnology), whose emission of red and green fluorescence relies on mitochondrial membrane potential, was an ideal fluorescence probe to detect mitochondrial membrane potential. After cell washing, JC-1 staining solution was added, thoroughly mixed, and incubated at 37°C for 20 min, and then stained cells were observed and photographed using a fluorescence microscope.

WESTERN BLOTTING:

After different treatments, the HUVECs were washed twice with cold PBS. Total cellular proteins were extracted using radioimmunoprecipitation assay buffer (RIPA) buffer. Protein concentrations were determined using a bicinchoninic acid protein assay kit (Beyotime Institute of Biotechnology). Equal amounts of proteins were separated using 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). The separated proteins were transferred onto polyvinylidene difluoride membranes (Millipore, Billerica, MA, USA). Then, the membranes were blocked with 5% skimmed milk in TBST at room temperature for 1 h. The blots were incubated at 4°C overnight with the following primary antibodies from Abcam: anti-GPX4 (1: 5000), anti-SLC7A11 (1: 1000), anti-ACSL4 (1: 10000), anti-HO-1 (1: 2000), anti-KLF4 (1: 1000), and anti-β-actin (1: 1000), and with the following primary antibodies from Cell Signaling Technology: anti-p-AMPK (1: 1000), anti-AMPK (1: 1000), anti-SOD-1(1: 1000), and anti-SOD-2 (1: 1000). Then, the blots were incubated with HRP-conjugated secondary antibodies (1: 5000) for 1 h at room temperature. Subsequently, blots were developed using enhanced chemiluminescence (Model 6600; Tanon, Shanghai, China).

STATISTICAL ANALYSIS:

The data were analyzed using GraphPad Prism version 7.0 (GraphPad Software Inc. San Diego, CA, USA). Differences between 2 groups were analyzed using the t test, whereas differences between multiple groups were analyzed using one-way analysis of variance, followed by Tukey’s test. All experiments were performed at least thrice. All data are presented as means±SEM. A P value of 0.05 or less was statistically significant.

Results

OX-LDL INDUCED HUVECS FERROPTOSIS IN A DOSE-DEPENDENT MANNER:

We used ox-LDL (0–200 μg/mL) for 24 or 48 h to induce endothelial cell ferroptosis and to explore the effect of ox-LDL on HUVECs. The CCK8 results showed that treatment with ox-LDL in a dose- and time-dependent manner markedly reduced the cell viability in HUVECs (Figure 1A, 1B). Hoechst/PI staining was used to evaluate the integrity of the cellular membrane. The results indicated that the number of PI-positive cells was higher in the ox-LDL group (100 μg/mL) for 48 h than in the control group (Figure 1C). HUVEC mitochondria of the ox-LDL-induced group (100 μg/mL) for 48 h showed morphological characteristics of ferroptosis, such as shrinkage, membrane density increasing, and mitochondrial ridges decreasing or disappearing (Figure 1D). These results indicated that the cellular viability of HUVECs was significantly decreased following treatment with 100 μg/mL ox-LDL at 48 h. Our results revealed that the ferroptosis protein ACSL4 expression level was increased after ox-LDL stimulation, compared with the control (Figure 1E, 1F). Meanwhile, GPX4 and SLC7A11 expressions were reduced in the ox-LDL group (Figure 1G, 1H). These results suggested that ox-LDL induced HUVECs ferroptosis.

CTRP13 SUPPRESSED OX-LDL-INDUCED HUVEC FERROPTOSIS:

We treated HUVECs with different CTRP13 concentrations for 48 h to further confirm whether CTRP13 was involved in ferroptosis responses. CTRP13 had no significant toxic effect on HUVECs at concentrations below 1200 ng/mL, compared with the control group (Figure 2A). HUVECs had approximately 45% viability at 1200 ng/mL after being cultured with 100 μg/mL ox-LDL (Figure 2B). Viability at 150, 300, and 450 ng/mL CTRP13 was consistent with a previous report [26]. Then, we treated HUVECs with 100 μg/mL ox-LDL at indicated time points. The results revealed that 450 ng/mL CTRP13 unregulated GPX4/SLC7A11 protein and mRNA levels, compared with an ox-LDL group (Figure 2C–F, 2H, 2I). Conversely, CTRP13 treatment of 450 ng/mL significantly blocked ACSL4 signal activity in the ox-LDL group (Figure 2G). The results indicated that 450 ng/mL CTRP13 significantly inhibited HUVEC ferroptosis.

CTRP13 INHIBITED OX-LDL-INDUCED HUVEC LIPID PEROXIDATION AND MITOCHONDRIAL DYSFUNCTION:

We detected LDH release in the supernatant to explore whether CTRP13 pre-treatment could ameliorate cellular membrane integrality. The results indicated that ox-LDL significantly increased LDH release, and CTRP13 attenuated this effect (Figure 3A). Moreover, CTRP13 treatment inhibited MDA and GSH (Figure 3B, 3C). The iron content was lower in the CTRP13 treatment group (Figure 3D) than in the ox-LDL group. ROS-mediated lipid peroxidation was the key step that induced ferroptosis. The CTRP13 and Fer-1 group significantly decreased ROS levels, compared with the ox-LDL group (Figure 3E, 3F). Also, we detected a dramatic improvement in mitochondrial membrane potential. The mito-tracker results reduced many mitochondrial membranes and fissions, compared with the control, while CTRP13 and Fer-1 alleviated this effect (Figure 3G, 3H).

CTRP13 REVERSED THE DOWNREGULATION OF MULTIFARIOUS ANTIOXIDANT PROTEINS:

CTRP13 accelerated antioxidant enzyme expression levels, such as HO-1, SOD-1, and SOD-2, compared with the ox-LDL group (Figure 4A–4E). Moreover, JC-1 staining revealed that adding CTRP13 restored mitochondrial membrane potential (Figure 4F, 4G). The immunofluorescence images indicated an extensive increase of GPX4 mean intensity in aggregates with CTRP13 and Fer-1, compared with the ox-LDL group (Figure 4H, 4I).

FERROPTOSIS PROTECTIVE EFFECT OF CTRP13 WAS ATTENUATED BY INHIBITING AMPK/KLF4 PATHWAY:

We examined the AMPK phosphorylation levels and KLF4 expression to elucidate the molecular ferroptosis mechanism induced by CTRP13. The p-AMPK/AMPK expression ratio decreased, and KLF4 expression increased in the HUVECs treated with ox-LDL (Figure 5A–5C). However, these changes were more pronounced in the HUVECs treated with ox-LDL or compound C (CC, 5 μmol/L), a potent and selective AMPK inhibitor. Ferroptosis biomarker proteins GPX4 and SLC7A11 expressions were decreased in the compound C group. In contrast, compound C groups had significantly increased ACSL4 and KLF4 levels, compared with the ox-LDL group. APTO (5 μmol/L), a KLF4 activator, did not affect AMPK phosphorylation, while APTO attenuated the anti-ferroptosis effect of CTRP13 (Figure 5D–5F).

CTRP13 RESTORED MITOCHONDRIAL FUNCTIONING BY COUNTERACTING AGAINST OX-LDL EFFECTS ON THE AMPK/KLF4 PATHWAY:

Next, we examined whether the AMPK/KLF4 pathway was responsible for the mitochondrial functioning of CTRP13 by inhibiting AMPK, the upstream activator of KLF4. DCFH and JC-1 staining indicated that compound C and APTO increased ROS and mitochondrial dysfunction (Figure 6A–6D). Expectedly, immunofluorescence staining revealed compound C and APTO diminished GPX4 expression (Figure 6E, 6F).

Hence, these results suggested that CTRP13 most likely protected ox-LDL-induced HUVEC ferroptosis by activating the AMPK/KLF4 pathway and upregulating various antioxidant enzymes (Figure 7).

Discussion

STUDY LIMITATIONS:

This study had potential limitations. First, using HUVECs instead of human aortic endothelial cells may be less reflective of actual aortic endothelial cell activity. Second, in vitro and in vivo experiments with endothelial cells are required for future studies to confirm the vascular protective effect of CTRP13 on ox-LDL-induced ferroptosis.

Conclusions

Our study showed that CTRP13 inhibited HUVEC ferroptosis induced by ox-LDL. CTRP13 depressed ROS production and decreased cell ferroptosis. We elucidated the molecular basis of the AMPK/KLF4-mediated induction of CTRP13 expression that inhibits ferroptosis. From a therapeutic perspective, these results provide a preliminary understanding of the pharmacological action of CTRP13 and can have important translational value in the treatment of endothelial dysfunction-accelerated atherosclerosis.