Abstract
Pulsed electromagnetic field (PEMF) therapy has been extensively investigated in clinical studies for the treatment of angiogenesis-related diseases. However, there is a lack of research on the impact of PEMFs on energy metabolism and mitochondrial dynamics during angiogenesis. The present study included tube formation and CCK-8 assays. A Seahorse assay was conducted to analyze energy metabolism, and mitochondrial membrane potential assays, mitochondrial imaging, and reactive oxygen species assays were used to measure changes in mitochondrial structure and function in human umbilical vein endothelial cells (HUVECs) exposed to PEMFs. Real-time polymerase chain reaction was used to analyze the mRNA expression levels of antioxidants, glycolytic pathway-related genes, and genes associated with mitochondrial fission and fusion. The tube formation assay demonstrated a significantly greater tube network in the PEMF group compared to the control group. The glycolysis and mitochondrial stress tests revealed that PEMFs promoted a shift in the energy metabolism pattern of HUVECs from oxidative phosphorylation to aerobic glycolysis. Mitochondrial imaging revealed a wire-like mitochondrial morphology in the control group, and treatment with PEMFs led to shorter and more granular mitochondria. Our major findings indicate that exposure to PEMFs accelerates angiogenesis in HUVECs, likely by inducing energy metabolism reprogramming and mitochondrial fission.
Keywords: Pulsed electromagnetic field, Angiogenesis, Mitochondria, Energy metabolism
Subject terms: Mechanisms of disease, Mitochondria, Energy metabolism
Introduction
Pulsed electromagnetic field (PEMF) therapy is a fascinating approach that uses electromagnetic fields to address specific health conditions1. PEMF therapy is a non-invasive technique that uses low-energy and non-ionizing electromagnetic fields to modulate biological and physiological systems2. The efficacy of PEMFs with specific parameters in modulating angiogenesis has been extensively demonstrated in numerous animal experiments and in vitro studies, and exposure resulted in an increased rate of vascular growth and capillary density3–6. PEMF therapy has been extensively studied in clinical settings for the management of angiogenesis-related diseases, such as cerebral ischemic stroke, myocardial ischemia, and diabetes-related peripheral vascular disease6–8. There has been growing research interest in the effects of PEMFs on various targets, including promotion of the proliferation, migration, and tube formation of vascular endothelial cells, increasing the expression level of angiogenic growth factors, and activation of voltage-gated calcium channels (VGCCs), as reviewed by Wei et al.6.
Energy metabolism plays a crucial role in the expression of the angiogenic phenotype in endothelial cells (ECs)9. Significant attention has been given to the role of EC metabolism in angiogenesis. Accumulating evidence suggests that during angiogenesis, ECs primarily rely on aerobic glycolysis instead of the oxidative phosphorylation pathway for ATP production. “Aerobic glycolysis”, first known as the Warburg effect, is a phenomenon that pertains to the distinctive characteristics of glucose metabolism in tumor cells10. Under typical oxygen conditions, tumor cells primarily use pyruvate to produce lactate for ATP generation, which enhances their glycolytic capacity11,12. Recent research indicated that the Warburg effect extends beyond cancer cells13, as it has been observed in other cell types involved in immunity, pluripotency, angiogenesis, and other biological processes. The Warburg effect in non-cancerous cells is referred to as the “non-cancerous Warburg effect” or “aerobic glycolysis” in non-cancerous cells14. The non-cancerous Warburg effect is observed in rapidly proliferating cells under physiological and pathological conditions and is triggered by endogenous and exogenous stimuli13,14.
Recent research revealed that the Warburg effect correlated with alterations in mitochondrial dynamics, such as increased mitochondrial fission and decreased mitochondrial fusion12,15. The intricate relationship between the Warburg effect and mitochondrial dynamics involves various components, including oncogenes, tumor suppressor genes, and metabolic enzymes. Clarification of the interplay between the Warburg effect and mitochondrial dynamics will help elucidate the potential mechanism of PEMF promotion of angiogenesis.
To the best of our knowledge, there is a limited amount of research on the impact of PEMFs on energy metabolism and mitochondrial dynamics during angiogenesis. In the present study, we examined alterations in the pattern of energy metabolism and the structure and functionality of mitochondria in ECs exposed to PEMFs to provide new insights into the underlying mechanism of PEMF facilitation of angiogenesis.
Results
PEMFs increased human umbilical vein endothelial cell (HUVEC) viability
We performed a Cell Counting Kit‐8 (CCK‐8) assay to verify the effects of PEMFs on HUVEC viability. Time-course experiments were performed to verify the effects of PEMF exposure on specific parameters. Under identical peak intensities of 20 Gs PEMF exposure, the metabolic activity of HUVECs treated with 40 or 80 Hz PEMF for 1 or 2 days was greater than the control group (Fig. 1C, 1D). Under identical frequencies of 40 Hz PEMF exposure, no significant alteration in metabolic activity was detected in the 20 Gs PEMF group compared to the control group (Fig. 1A). However, after 2 days of exposure to PEMFs, cell metabolic activity was significantly increased in both PEMF groups, with a greater increase in the 40 Gs-exposed cells compared to control cells (Fig. 1B). We chose the 40 Gs, 80 Hz PEMF exposure protocol for further study.
Figure 1.
Cell Counting Kit‐8 assay to verify the effects of PEMFs on HUVECs viability. The cell viability of HUVECs treated with various intensity PEMFs of 40 Hz for 1 (A) or 2 days (B); The cell viability of HUVECs treated with various frequency PEMFs of 20Gs Hz for 1 (C) or 2 days (D). N = 5 per group. ns: not significant, **P < 0.01, ***P < 0.001, ****P < 0.0001.
PEMFs accelerated HUVEC tube formation
There are several ways to quantify tube network formation. The most common methods of analysis involve quantification of the number of tubes, nodes, or loops/meshes or the length of tubes. The imaging program NIH ImageJ with the Angiogenesis Analyzer plugin was used for the quantification of tube networks. Compared to the control group, tube formation occurred quickly in HUVECs treated with PEMFs within 2 h, and the number of junctions and meshes, total mesh area (TMA), mean mesh size (MMS), total length (TL) and total segment length (TSL) of the HUVECs were significantly greater (Fig. 2). The tube network became more extensive over time. Although this effect of PEMF was not statistically significant within 4 h, the tendency was clearly visible. By 16 h, the tubes in the control group began to disconnect and deteriorate, but the tubes in the PEMF group remained intact.
Figure 2.
The effects of PEMFs on HUVECs tube formation assay for the in vitro study of angiogenesis. (A) Phase contrast microscopy imaging of HUVECs tube formation over time. Scale bar: 100 μm. Extremities (dark dots), Twigs (cyan), Branches (green), Segments (yellow) and Junctions (red); (B) Comparative measurement of parameters of tubes network of HUVECs in tube formation assay. N = 3 per group. TMA: total mesh area, MMS: mean mesh size, TSL: total segment length, TL: total length, t: time, ns: not significant, **P < 0.01, ***P < 0.001, ****P < 0.0001.
PEMFs transformed the energy metabolism patterns of HUVECs
Glycolysis stress tests revealed that the basic glycolysis level and glycolytic capacity were significantly greater in the HUVECs treated with PEMF than in the control cells, but the difference in the glycolytic reserve between the groups was not significant, which revealed that PEMF-exposed HUVECs were highly glycolytic for energy generation (Fig. 3). A mitochondrial stress test revealed that ATP-linked respiration, maximal respiration, and spare respiratory capacity were lower in the PEMF group than in the other groups (Fig. 4). ATP-linked respiration was significantly decreased in HUVECs treated with PEMFs. In summary, PEMFs promoted the transformation of the energy metabolism pattern of HUVECs from oxidative phosphorylation to aerobic glycolysis.
Figure 3.
The effects of PEMFs on glycolysis in HUVECs. N = 3 per group. ECAR, extracellular acidification rate, ns not significant, **P < 0.01.
Figure 4.
The effects of PEMFs on mitochondrial oxidative phosphorylation in HUVECs. N = 3 per group. OCR: oxygen consumption rate, ns: not significant, *P < 0.05.
To clarify the underlying mechanisms, changes in the gene expression of several mediators involved in glycolysis was analyzed. Compared to the control groups, the mRNA expression levels of genes (PFKL, PFKLM, PFKP, PKM2, and HK2) encoding the three key regulatory enzymes of glycolysis, hexokinase, phosphofructokinase, and pyruvate kinase, sharply increased when HUVECs were exposed to PEMFs (Fig. 8). A trend was also observed in the expression of GLUT1/2, which encodes a glucose transporter that facilitates the transport of glucose across the plasma membrane.
Figure 8.
The mRNA expression of antioxidants gene, glycolytic pathway and mitochondrial dynamics related genes in HUVECs exposed to the PEMFs compared to control group cells. n = 3 per group. ns: not significant, *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.
PEMFs maintained low levels of intracellular ROS in HUVECs
Mitochondrial respiration is widely accepted as the primary sources of ROS, and the results of the mitochondrial stress test showed decreased mitochondrial respiration. Therefore, ROS detection was performed to examine the effect of PEMFs on intracellular ROS. As shown in Fig. 5C, flow cytometry showed that PEMFs dramatically reduced ROS levels in HUVECs treated with PEMFs compared to the control group. Taken together, these results clearly demonstrate that PEMFs improved the health status of HUVECs. Furthermore, a significant increase in the expression of superoxide dismutase 1/2 (SOD1/2) and glutathione peroxidase 1/4 (GPX1/4) was observed in HUVECs in the PEMF group (Fig. 8).
Figure 5.
The effects of PEMFs on mitochondrial membrane potential and ROS in HUVECs. (A) JC-1 microscopy assay for mitochondrial membrane potential in HUVECs. Scale bar: 20 μm; (B) JC-1 Cytometry assay for mitochondrial membrane potential in HUVECs; (C) ROS detection by flow cytometer in HUVECs. n = 3 per group. ROS: reactive oxygen species, ns: not significant, *P < 0.05.
PEMFs did not damage mitochondrial membrane potential in HUVECs
The mitochondrial membrane potential provides a valuable indicator of cell health and functional status. Cytometry- and microscopy-based analyses are widely used to study mitochondrial behavior. When the mitochondrial membrane potential is high, JC‐1 aggregates and emits red fluorescence after staining. When the mitochondrial membrane potential is low, JC‐1 is present as a monomer and emits green fluorescence after staining. Microscopy revealed that the red fluorescence intensity in the PEMF group seemed higher compared to the control group, but there was no significant difference in the ratio of red-to-green fluorescence intensity between the two groups according to flow cytometry, which suggested that PEMF stimulation did not damage the mitochondrial membrane potential (Fig. 5A, 5B).
PEMFs regulated the mitochondrial dynamics of HUVECs
Mitochondria are dynamic organelles that behave as a dynamic network in most cells and change their biogenesis and structure depending on cell needs or in response to different conditions. Mitochondrial-specific fluorescence staining revealed that the shape of mitochondria in HUVECs in the control group were wirelike, but the mitochondria became shorter and more granular after treatment with PEMFs. We used mitochondrial network analysis (MiNA) software to analyze the structure of the mitochondria. The results showed that the morphologies of the mitochondria in the cells treated with PEMF exhibited increased numbers of “individual” structures with no branches and decreased numbers of networks and mean length of branches, which indicated that PEMF induced mitochondrial fission (Fig. 6). These tendencies were clearly visible on holo-tomographic microscopy (HTM) and transmission electron microscopy (TEM) (Fig. 7).
Figure 6.
Representative images for MitoTracker™ Deep Red FM staining of HUVECs (A) and results of MiNA between the two groups (B). Scale bar: 100 um (white); 20 um (red). MiNA: Mitochondrial Network Analysis, **P < 0.01, ****P < 0.0001.
Figure 7.
Representative images for Holo-tomographic microscopy (A) (Scale bar: 20 um) and TEM (B) (Scale bar: 1um) of HUVECs. Red arrow: mitochondria. TEM: transmission electron microscopy.
However, the effects of PEMF on the mRNA expression of mitochondrial fission- and fusion-related genes are complex. PCR revealed that the expression of mitochondrial fission-related genes, including DRP1 and FS1, increased, but the expression of mitochondrial fission-related genes, including MFN1/2 and OPA1, decreased slightly in PEMF-exposed HUVECs compared to the control group (Fig. 8).
Discussion
Angiogenesis is crucial for tissue repair and has been extensively studied, especially in the context of therapeutic angiogenesis for ischemic diseases and tissue regeneration1,16. Although the impact of PEMFs on angiogenesis has been investigated in numerous experimental and clinical studies6,7, the effect of PEMFs on angiogenesis is controversial. These controversies may arise from inconsistencies in experimental settings, such as variations in the parameters for electromagnetic field intervention, the cell type and disease models used, and other related factors2,17. Therefore, the presence of multiple variables is challenging for performing quantitative analyses and reaching definitive conclusions. The potential impact of extremely low-frequency pulsed electromagnetic fields (ELF-EMFs) on human health and its therapeutic applications have garnered significant attention. ELF-EMFs are low-energy, non-ionizing electromagnetic fields that induce various biological effects, including angiogenesis and bone healing2. The PEMFs used in the present study are also classified as ELF-EMFs. The specific parameters for PEMFs in our study were selected based on commonly used clinical treatment parameters for bone healing. The final protocol for PEMF exposure was determined by considering the impact of magnetic field frequency and peak intensity on the viability of HUVECs. Our CCK-8 assay results demonstrated that the effect of PEMFs was not directly proportional to its frequency or intensity. Therefore, it is more appropriate to evaluate the biological response to PEMFs using the "biological window" hypothesis rather than relying solely on a dose–effect relationship18. Although PEMFs modulate an extensive array of biological effects, including cell migration, proliferation, and differentiation, cytokine and growth factor expression, and alterations of nitric oxide signaling during angiogenesis, few studies have focused on the impact of PEMFs on energy metabolism in endothelial cells during angiogenesis.
The role of endothelial cell metabolism in angiogenesis has received significant attention recently. Numerous novel metabolic characteristics of angiogenic ECs have been revealed, particularly their glycolytic features. Unique glucose metabolism traits have recently been identified in ECs. During vessel sprouting in HUVECs, the glycolysis pathway accounts for approximately 85% of ATP production19, whereas cultured quiescent aortic ECs convert approximately 99% of glucose into lactate20. This phenomenon is similar to the Warburg effect in tumor cells, known as aerobic glycolysis in non-cancerous cells13. The distinctive aerobic glycolytic characteristics of ECs provide several beneficial effects9,21,22. First, glycolysis enables faster ATP production kinetics. Second, glycolysis prevents the generation of reactive oxygen species (ROS), and third, glycolysis supplies metabolic intermediates for the synthesis of lipids, amino acids, and nucleotides. Our findings from the Seahorse assay demonstrated that PEMF-treated HUVECs showed higher glycolytic flux compared to the control group, with obviously upregulated gene expression of glucose transporters (GLUT1/4) and three key regulatory enzymes (PFK, PKM, HK) of the glycolytic pathway, which are essential for efficient glucose uptake and the promotion of enhanced glycolytic flux. We also found that glycolytic flux was accompanied by increased antioxidant gene expression and decreased intracellular ROS, which resulted in a healthier microenvironment for cell proliferation and function. The process of mitochondrial respiration involves the consumption of oxygen and the generation of reactive oxygen species (ROS), and mitochondria are widely recognized as the primary sources of ROS23. Therefore, the PEMF-induced decrease in ROS may be related to the downregulation of oxidative phosphorylation (OXPHOS) pathways and enhanced ROS scavenging.
The maintenance of mitochondrial function and adaptation to diverse environmental conditions require the complex coordination of mitochondrial dynamics, including the processes of mitochondrial fission and fusion. These processes collectively contribute to cellular homeostasis. In the present study, we observed visible mitochondrial fission in HUVECs treated with PEMFs, which was closely associated with an increase in aerobic glycolysis. Several studies suggest that PEMFs modulate mitochondrial dynamics and play a crucial role in regulating mitochondrial quality control and cellular metabolism. Ren et al. reported that the activation of PKM2 enhanced angiogenesis in endothelial cells (ECs) by modulating glycolysis, mitochondrial fission, and fusion24. Based on these findings, we speculate that PEMF-induced mitochondrial fission is associated with the upregulation of PKM2 expression under electromagnetic field exposure.
Our study has several limitations. First, we report the effects of PEMFs on the energy metabolism and mitochondrial dynamics of endothelial cells at the phenotypic and genetic levels. Further studies, such as protein expression and genomic or proteomic analyses, may help identify new therapeutic targets for regulating angiogenesis. Second, we did not perform in vivo animal experiments to verify the effects of PEMFs on angiogenesis, energy metabolism and mitochondrial dynamics.
Conclusion
Our study demonstrated that exposure to PEMFs accelerated angiogenesis in HUVECs by inducing the reprogramming of energy metabolism from oxidative phosphorylation to aerobic glycolysis and mitochondrial fission, which reduced intracellular ROS levels. These findings are promising and provide novel insights into the underlying mechanism of PEMF-promoted angiogenesis. Our findings support the application of PEMFs as a new non-pharmacological strategy for regulating cellular energy and promoting tissue repair. However, further investigations are required to elucidate the molecular mechanisms underlying the enhanced glycolytic flux induced by PEMFs and to determine the optimal parameters for promoting angiogenesis.
Methods
PEMF exposure system
The custom-designed PEMF exposure system consisted of a signal generator (DG1022, RIGOL, China) (Fig. 9B), a power amplifier (FN-3080, Nanjing, China) (Fig. 9C), and a Helmholtz coil (CHY20-60 J, CH-Magnetoelectricity Technology, China) (Fig. 9D) with a uniform magnetic field in a Φ130-mm sphere. The peak intensity of the output magnetic fields was approximately 60 Gs using a Gaussmeter (Lake Shore Cryotronics, Westerville, OH). The signal generator's peak magnetic field fluctuated less than 2% throughout a one-hour PEMF treatment. The output waveform was comprised of a burst of pulses with a pulse width of 2 ms.
Figure 9.
The design schematic diagram and physical diagram of PEMFs exposure system.
Cell culture
HUVECs were purchased from American Type Culture Collection (ATCC, USA). The cells were cultured in DMEM (GIBCO, USA) supplemented with 10% fetal bovine serum (FBS) (GIBCO, USA). The cells were maintained in an incubator at 37 °C with 5% CO2 and 95% O2.
Cell Counting Kit‐8 assay
CCK‐8 (Dojindo, Japan) assays were performed to detect the impact of PEMFs on HUVEC survival according to the manufacturer’s instructions. HUVECs were seeded in 96‐well plates (0.5 × 104 cells/well) and cultured for 24 h. Adherent cells were cultured under PEMF exposure (20 Gs with different frequencies for 30 min or 40 Hz with different intensities for 30 min daily) for 2 days. The original medium was replaced with fresh medium containing 10% CCK‐8 on days 1 and 2, and the cells were incubated for 1 h. The absorbance was measured at a wavelength of 450 nm using a microplate reader (Synergy Mx, BioTek, USA).
Tube formation assay
Tube formation assays were performed as described previously25. ABW Matrigel (#082,704, Nova, Shanghai, China) was dispensed into 96‐well plates (50 μl/well) and incubated at 37 ℃ for 1 h for solidification. HUVECs were seeded on the Matrigel (3 × 104 cells/well) and cultured under PEMF exposure of 40 Gs, 80 Hz for 30 min. Within 16 h after plating, images were acquired with a Zeiss imaging system (Axio Observer D1, Carl Zeiss, Germany) and analyzed using the image analysis software ImageJ with the Angiogenesis Analyzer plugin for the quantification of tube networks26.
Seahorse assay
The extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) were detected using a Seahorse XF Cell Mito Stress Test Kit and an XF Glycolysis Stress Test Kit (#103,015-100, 103,020-100; Seahorse Bioscience, USA), respectively, in a Seahorse XFe24 Extracellular Flux Analyzer (Seahorse Bioscience) according to the manufacturer’s instructions. Briefly, HUVECs cultured in XFe24 cell culture microplates were treated as described for the PEMF exposure protocol (40 Gs, 80 Hz) for 30 min daily for 2 consecutive days. Before measurement, a sensor cartridge was hydrated in a Seahorse calibrant in a non-CO2 incubator at 37 °C for 4 h before the experiments. The microplates were loaded into the XFe24 Analyzer, and 5 μM oligomycin (Oligo), 1 μM carbonylcyanide-4-(trifluoromethoxy) phenylhydrazone (FCCP), and a mixture of rotenone and antimycin A (1 μM) were sequentially added to reveal several parameters of metabolic function, including basal respiration, ATP production, maximal respiration and spare capacity. The ECAR was measured after the sequential injection of glucose (10 mM), oligomycin (1.0 μM), and 2-deoxyglucose (2-DG, 50 mM) as the basal ECAR, maximal ECAR, and reserve capacity, respectively. After measurement, the cells were stained with DAPI and counted using a Celigo Image Cytometer (Celigo 200-BFFL-5C, Nexcelom Bioscience, USA). The OCR and ECAR of the cells were normalized to the cell number. The datasets were analyzed using Wave software (Agilent Technologies).
Measurement of mitochondrial membrane potential
A JC-1 kit (#C2006, Beyotime, Shanghai, China) was used to determine the mitochondrial membrane potential according to the manufacturer’s instructions. HUVECs were seeded in 6‐well plates (2 × 105 cells/well) and incubated for 24 h. Adherent cells were cultured under PEMF exposure (40 Gs, 80 Hz) for 30 min daily for 2 consecutive days. After treatment as described, the cells were gently washed once with warm PBS. For the flow cytometry assay, cells were trypsin-digested and suspended in PBS. The cells were incubated in JC-1 working solution for 20 min at 37 ℃, then gently washed three times with JC-1 dyeing buffer. The fluorescence intensity was measured using a flow cytometer (Cytoflex, Beckman, China) and analyzed using FlowJo™ Software (v10.8, USA). Data were obtained from three independent experiments, and 10,000 cells were examined per group in each experiment. For microscopy, cells stained with JC-1 were immersed in culture medium and imaged as quickly as possible. Images were acquired using a fully automatic fluorescence microscope (IX83, Olympus, Japan).
Mitochondrial-specific fluorescence staining
HUVECs were seeded in CellCarrier Ultra microplates (#6,005,530, PerkinElmer, USA) (3000 cells/well) and cultured for 24 h. Adherent cells were cultured under PEMF exposure (40 Gs, 80 Hz) for 30 min daily for 2 consecutive days then stained with MitoTracker™ Deep Red FM (#M22426, Invitrogen, USA) according to the manufacturer's protocol. Images were acquired using a high-content screening system (PE/Opera Phenix Plus, PerkinElmer, USA). Mitochondrial network morphology was analyzed using the MiNA toolset, which is a relatively simple pair of macros in existing ImageJ plug-ins, as previously reported27.
Intracellular ROS assay
HUVECs were seeded in 6‐well plates (2 × 105 cells/well) and treated as described for the PEMF exposure protocol (40 Gs, 80 Hz) for 30 min daily for 2 consecutive days. Intracellular ROS were detected using a Reactive Oxygen Species Assay Kit (#BL714A, Biosharp, China) according to the manufacturer's protocol. The fluorescence intensity was measured by a flow cytometer (Cytoflex, Beckman, China) and analyzed using FlowJo™ Software (v10.8, USA). Data were obtained from three independent experiments, and 10,000 cells were examined per group in each experiment. The geometric average of the fluorescence intensity or percentage of positive cells was compared between groups.
HTM
HTM is a label-free non-phototoxic microscopy method that reports fine changes in cell refractive indices (RIs) in 3D. Mitochondria show a specific RI signature that distinguishes these structures with high resolution and contrast. HUVECs were seeded in BeyoGold™ 35-mm cell culture dishes (#FCFC020, Beyotime, Shanghai, China) (3 × 104 cells/dish) and treated as described for the PEMF exposure protocol (40 Gs, 80 Hz) for 30 min daily for 2 consecutive days. The cells were analyzed using 3D Cell Explorer-fluo (NANOLIVE, Ecublens, Switzerland).
TEM
HUVECs were seeded in 6‐well plates (2 × 105 cells/well) and treated as described for the PEMF exposure protocol (40 Gs, 80 Hz) for 30 min daily. Cell suspensions were sequentially fixed in 2.5% glutaraldehyde and 1% osmium tetroxide, dehydrated in acetone, embedded in epoxy resin, and polymerized at 60 °C for 3 days. Sections were created and collected on copper grids. The sections were observed for mitochondrial ultrastructure using TEM (JEM1400, JEOL, Japan).
Quantitative real-time PCR
HUVECs were seeded in 6‐well plates (2 × 105 cells/well) and incubated for 24 h. Adherent cells were cultured under PEMF exposure (40 Gs, 80 Hz) for 30 min daily for 2 consecutive days. Total RNA from cell samples was extracted using an Eastep® Super Total RNA Extraction Kit (#LS1040, Promega, Shanghai, China) according to the manufacturer’s protocol. RNA was reverse transcribed into cDNA using an iScript cDNA Synthesis Kit (#1,708,891, Bio-Rad, USA). iTaq Universal SYBR® Green SuperMix (#1,725,121, Bio-Rad, USA) was used to perform quantitative PCR according to the manufacturer’s instructions. The signals were detected and analyzed in a CFX Connect System (Bio-Rad, USA). RNA18SN1 was used as an internal reference, and sequences were designed using an online tool at the National Institutes of Health (https://www.ncbi.nlm.nih.gov/tools/primer-blast/index.cgi). Sequences are listed in Table 1.
Table 1.
Primer design of HUVECs.
| Gene | Forward primer | Reverse primer |
|---|---|---|
| RNA18SN1 | CGGCGACGACCCATTCGAAC | GAATCGAACCCTGATTCCCCGTC |
| GPX1 | GTGCTCGGCTTCCCGTGCAAC | CTCGAAGAGCATGAAGTTGGGC |
| GPX4 | ACAAGAACGGCTGCGTGGTGAA | GCCACACACTTGTGGAGCTAGA |
| SOD1 | CTCACTCTCAGGAGACCATTGC | CCACAAGCCAAACGACTTCCAG |
| SOD2 | CTGGACAAACCTCAGCCCTAAC | AACCTGAGCCTTGGACACCAAC |
| PFKM | GCTTCTAGCTCATGTCAGACCC | CCAATCCTCACAGTGGAGCGAA |
| PFKL | AAGAAGTAGGCTGGCACGACGT | GCGGATGTTCTCCACAATGGAC |
| PFKP | AGGCAGTCATCGCCTTGCTAGA | ATCGCCTTCTGCACATCCTGAG |
| PKM2 | ATGGCTGACACATTCCTGGAGC | CCTTCAACGTCTCCACTGATCG |
| HK2 | GAGTTTGACCTGGATGTGGTTGC | CCTCCATGTAGCAGGCATTGCT |
| SLC2A1 | TTGCAGGCTTCTCCAACTGGAC | CAGAACCAGGAGCACAGTGAAG |
| SLC2A4 | GTTCTTTCATCTTCGCCGCC | TTCCCCATCTTCGGAGCCTA |
| MFN1 | GGTGAATGAGCGGCTTTCCAAG | TCCTCCACCAAGAAATGCAGGC |
| MFN2 | ATTGCAGAGGCGGTTCGACTCA | TTCAGTCGGTCTTGCCGCTCTT |
| OPA1 | GTGGTTGGAGATCAGAGTGCTG | GAGGACCTTCACTCAGAGTCAC |
| FIS1 | CAAGGAACTGGAGCGGCTCATT | GGACACAGCAAGTCCGATGAGT |
| DNM1L | GATGCCATAGTTGAAGTGGTGAC | CCACAAGCATCAGCAAAGTCTGG |
Statistical analysis
Statistical analyses were performed using the unpaired two-tailed t test or ANOVA with Sidak’s multiple comparisons test in GraphPad Prism 9 software (GraphPad Software, USA). All data are expressed as means ± standard deviation (SD). Significance is indicated by asterisks: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001; NS, not significant.
Acknowledgements
We are grateful to all the colleague who have offered us invaluable help in experiments and evaluation of this study, especially Hongying Chen, Jinkui Pi, Mengli Zhu, and Jian Yang in the Core Facilities of West China Hospital, Sichuan University.
Author contributions
Conceptualization: Q.Y., B.P., and H.L.; Methodology: L.X. and F.L.; Formal analysis: C.L., Y.Z., and Y.C.; Writing – review & editing: C.Y. All authors reviewed the manuscript.
Funding
This research was supported by the National Natural Science Foundation of China (82002395), the Department of Science and Technology of Sichuan Province (2021YJ0432) and the Talent Youth Fund of Sichuan Provincial People's Hospital (2016QN10).
Data availability
The raw data is available from the corresponding author on reasonable request.
Competing interests
The authors declare no competing interests.
Footnotes
Publisher's note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
These authors contributed equally: Chengyi Yang and Li Xu.
Contributor Information
Bo Peng, Email: lucky63@163.com.
Huifang Liu, Email: liuhuifang@med.uestc.edu.cn.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Data Availability Statement
The raw data is available from the corresponding author on reasonable request.