Knockdown of PTEN Inhibits Autophagy-Dependent Ferroptosis to Alleviate LPS-Induced Sepsis-Associated Acute Kidney Injury

Abstract

Purpose: This research aims to explore the mechanism whereby PTEN modulates autophagy-dependent ferroptosis in sepsis-associated acute kidney injury (SA-AKI). Methods: Bioinformatics analysis was performed using the GSE65682 dataset to screen genes associated with autophagy-dependent ferroptosis. qRT-PCR was performed to validate the expression of key genes. Cell proliferation was determined via the Cell Counting Kit-8 and colony formation assays. Apoptosis, mitochondrial membrane potential, Fe2+ and ROS levels were measured via flow cytometry. Mitochondrial function and autophagy were observed using transmission electron microscopy. Protein analysis was conducted via Western blotting. To verify the role of the key gene PTEN in SA-AKI, an in vivo model of SA-AKI was established via lipopolysaccharide (LPS) induction. Results: PTEN was identified as a key gene, with findings showing that it is highly expressed in SA-AKI. In the LPS-induced in vitro cell model of SA-AKI, PTEN knockdown enhanced cell proliferation, inhibited apoptosis, and reduced intracellular Fe2+ and ROS levels (P < 0.05, P < 0.01). Additionally, LPS-induced mitochondrial swelling and autophagosome accumulation were alleviated by PTEN knockdown, with fewer autophagic structures observed. PTEN knockdown upregulated the expression of Nrf2, SLC7A11, and GPX4, leading to reduced expression of COX2 and 4-HNE, and regulated autophagy-dependent ferroptosis by activating the Nrf2/SLC7A11/GPX4 signaling pathway. Furthermore, PTEN knockdown mitigated LPS-induced renal injury in SA-AKI, demonstrating its protective effect against SA-AKI. Notably, Nrf2 inhibition by ML385 reversed the protective effects of PTEN knockdown. Conclusions: This study elucidates that the knockdown of PTEN activates the Nrf2/SLC7A11/GPX4 signaling pathway to modulate autophagy-dependent ferroptosis in SA-AKI.

Supplementary Information

The online version contains supplementary material available at 10.1007/s10753-026-02529-z.

Introduction

Triggered by renal ischemia-reperfusion, sepsis, nephrotoxic drugs, contrast agents, and other factors, acute kidney injury (AKI) is a renal clinical syndrome that exhibits a sudden drop in glomerular filtration rate and is correlated with high mortality [1]. Nowadays, sepsis is the most prevalent cause of AKI. AKI caused by sepsis is termed sepsis-associated acute kidney injury (SA-AKI). Sepsis involves a process in which the body’s immune response to infection is disrupted, causing the dysfunction or even failure of its own organs. In severe cases, it can be life-threatening [2, 3]. Within the intensive care unit, sepsis constitutes the primary contributor to AKI. Sepsis accounts for 45% to 70% of AKI etiologies, and the mortality rate often exceeds 50% [4]. Currently, the main treatment methods for SA-AKI include renal replacement therapy, mechanical ventilation, infection control, fluid resuscitation, and vasoactive drugs [5]. However, due to the restrictions on the treatment approaches and the deficiency in early identification and diagnostic methods, many patients not only cannot receive radical treatment but may even require lifelong renal replacement therapy, which greatly increases the economic burden on patients. Hence, studying the pathogenesis underlying SA-AKI and finding practical therapeutic targets are of great significance for its treatment and prognosis assessment.

The pathogenesis underlying SA-AKI is remarkably intricate, encompassing microcirculatory disorders, immune inflammatory responses, activation of the coagulation system, and renal tubular epithelial injury, among others. Nevertheless, the specific mechanisms remain incompletely elucidated [6]. Recent research has zeroed in on programmed cell death, including apoptosis, pyroptosis, and autophagy [7–9]. However, a growing body of research has confirmed that autophagy-dependent ferroptosis may be involved in the inflammatory response and immune regulation of SA-AKI [10, 11]. Autophagy-dependent ferroptosis, a mode of cell death, exhibits characteristics of both autophagy and ferroptosis, while autophagy-related mechanisms propel the progression of ferroptosis [12, 13]. Yang et al. validated that ferroptosis and lipophagy play potentially vital roles in sepsis-induced AKI [14]. Wang et al. aimed to explore how 4-OI exerts its protective effect in SA-AKI, finding that 4-OI was able to suppress lipopolysaccharide (LPS)-induced activation of the STING pathway. Thereby, it suppressed the autophagic degradation of GPX4 mediated by STING, reduced the accumulation of reactive oxygen species (ROS), and alleviated ferroptosis [15]. At present, the research on the role and mechanism of autophagy-dependent ferroptosis in SA-AKI is still in its infancy, and relevant exploration is extremely limited.

This study utilized bioinformatics to investigate the differentially expressed genes (DEGs) distinguishing SA-AKI patients from healthy individuals. From these genes, we selected the DEGs at the intersection of autophagy and ferroptosis, verified their expression levels in clinical samples using quantitative reverse transcription polymerase chain reaction (qRT-PCR) and enzyme-linked immunosorbent assay (ELISA), and ultimately designated PTEN as the target for subsequent research. Subsequently, we set up an LPS-induced SA-AKI in vitro model in HK-2 cells, and also established a mouse SA-AKI model by means of intraperitoneal LPS injection. We explored the biological functions and mechanisms of PTEN in autophagy-dependent ferroptosis triggered by SA-AKI, with the aim of providing new clinical treatment strategies for SA-AKI.

Materials and Methods

Dataset Collection and Processing

GSE65682 was retrieved from the Gene Expression Omnibus (GEO) database (https://www.ncbi.nlm.nih.gov/geo/), which contains 760 blood samples from individuals with sepsis and 42 healthy control blood samples. A total of 137 autophagy-related genes (ARGs) were retrieved by searching the keyword “autophagy” in the MsigDB database (https://www.gsea-msigdb.org/gsea/index.jsp) [16] and 567 ferroptosis-related genes (FRGs) were sourced from the FerrDb V2 database (http://www.zhounan.org/ferrdb/current/) [17].

DEGs Analysis

In GSE65682, the limma package (version 3.46.0) was used to compare gene expression differences between the Sepsis and Healthy control groups. The screening criteria were: padj < 0.05 and |log2FoldChange| > 0.5.

Functional Enrichment Analysis

Enrichment analysis based on the Kyoto Encyclopedia of Genes and Genomes (KEGG) and Gene Ontology (GO) databases was performed using the clusterProfiler package and org.Hs.eg.db package in R software to identify the common functions and related pathways of DEGs. A padj < 0.05 was used as the screening criterion.

Sample Collection

A total of 6 pairs of blood samples from patients with SA-AKI and healthy control subjects were collected at the China-Japan Union Hospital of Jilin University. This study adhered to the Declaration of Helsinki (revised in 2013) and was granted approval by the Ethics Committee of the China-Japan Union Hospital of Jilin University. Informed consent to use their samples and data was given by all patients.

qRT–PCR

Total RNA was isolated from SA-AKI clinical samples and Human Kidney-2 (HK-2) cells with TRIzol™ reagent (15596018CN, Thermo Fisher Scientific, Waltham, MA, USA), followed by reverse transcription. The Real-time PCR reaction was executed with a Q-PCR instrument (CFX96 TOUCH, Bio-Rad, California, USA). Primer sequences were listed in Table 1. With GAPDH as the internal control, the 2^−ΔΔCT method was applied for quantification.

Table 1.

Primer sequences information.

Gene name	Sequence
PTEN	Forward	GCAGAAAGATTGAAGGCGTAT
	Reverse	GCTGTGGTGGGTTATGGT
GAPDH	Forward	GATTGTTGCCATCAACGACC
	Reverse	GTGCAGGATGCATTGCTGAC

Cell Culture and Transfection

HK-2 cells were provided by Wuhan Procell Life Science & Technology Co., Ltd. (CL-0109, Wuhan, China). The culture medium for HK-2 cells was prepared by supplementing Minimal Essential Medium (MEM; C11095500BT, Gibco, Grand Island, USA) with 10% fetal bovine serum (FBS; 40130ES76, Yeasen, Shanghai, China) and 1% penicillin-streptomycin (BL505A, Biosharp, Beijing, China). Cells were cultured in an incubator at 37℃ with 5% CO₂.

To generate the stable PTEN knockdown HK-2 cell line, HK-2 cells were first seeded into 6-well plates at a density of 2 × 10⁵ cells per well and cultured overnight in the above-described complete MEM medium until reaching 70–80% confluence. Cells were then stably transduced with PTEN-targeted short hairpin RNA (shRNA) lentivirus (Lv-shPTEN) and negative control lentivirus (Lv-shNC) provided by Jintusi Biotechnology Co., Ltd. (Wuhan, China), at a multiplicity of infection (MOI) of 5. The corresponding sequences of Lv-shPTEN and Lv-shNC were presented in Table 2. Lipofectamine 2000 reagent (11668-019, Invitrogen, Carlsbad, CA) was employed for cell transduction according to the manufacturer’s protocol. After 8 h of transduction, the medium was replaced with fresh complete MEM medium, and the cells were continuously cultured for 72 h. Stable cell lines were selected using 800 µg/mL neomycin (G418, Sigma-Aldrich, USA), and single-cell clones were picked and expanded in complete MEM medium containing 400 µg/mL neomycin for maintenance to ensure stable PTEN knockdown. The knockdown efficiency of the stable PTEN-knockdown HK-2 cell line was verified by qRT-PCR at the mRNA level. The SA-AKI HK-2 cell model was established by treating the cells with 100 ng/mL LPS (BS904, Biosharp, Beijing, China) for 0, 24, 48, and 72 h.

Table 2.

Lentivirus sequences information.

Gene name	Sequence
sh-PTEN−1	CTAGAACTTATCAAACCCTTT
sh-PTEN−2	CGTGCAGATAATGACAAGGAA
sh-PTEN−3	AGGCGCTATGTGTATTATTAT.
sh-NC	TTCTCCGAACGTGTCACGT

Cell Counting Kit–8 Assay (CCK8)

To assess the proliferation of HK-2 cells, the CCK8 assay (KC0301-500, Zhengzhou Keremei Biotechnology Co., Ltd., Zhengzhou, China) was performed. First, morphologically healthy HK-2 cells were seeded into a 96-well plate with three replicate wells. Second, 10 µL of CCK8 solution was added to each well, and the plate was then incubated in a cell incubator under standard conditions for 1 h. Finally, the absorbance readings at 450 nm were measured using a microplate reader.

Colony Formation Assay

A total of 10³ well-shaped HK-2 cells were carefully seeded into a 6-well plate, with 3 replicates per group. Cells were cultured in an incubator at 37℃ with 5% CO₂. The culture medium for the cells was refreshed every 2–3 days. The cells were fixed using 4% paraformaldehyde (BL539A, Biosharp, Beijing, China). Subsequently, they were stained with crystal violet staining solution (BL802A, Biosharp, Beijing, China), and images were captured. For quantitative assessment, colonies were clearly distinguishable by the naked eye, and manual counting was performed using the Multi-point Tool in ImageJ software across the entire well. Data were presented as the absolute number of colonies to facilitate comparison between groups.

Cell Apoptosis

Upon completion of cell culture, the cells were harvested and rinsed with pre-chilled PBS. The cell number was adjusted to 5 × 10⁶ cells per tube before treatment, with 3 replicates per group. According to the operation instructions, cell death was evaluated using the Annexin V-FITC Apoptosis Detection Kit (C1062S, Beyotime, Shanghai, China). Briefly, cells were resuspended by adding 300 µL of 1×Binding Buffer. Next, 5 µL of Annexin V-FITC was added, mixed thoroughly, and incubated at room temperature in the dark for 15 min. Subsequently, 5 µL of PI was added for staining. Finally, analysis was performed using an Agilent NovoCyte Advanteon flow cytometer (Agilent Technologies, Beijing, China). Notably, fluorescence intensity was quantified using the FITC-height (FITC-H) parameter rather than the conventional FITC-area (FITC-A). In standard flow cytometry, FITC-A is widely adopted for fluorescence quantification; however, FITC-H records the peak fluorescence signal height of individual events, which minimizes signal overlap and reduces interference from cell aggregation and doublets in the present cell model. Therefore, we specifically employed the FITC-H parameter for Annexin V-FITC signal acquisition and analysis. For quantitative assessment, live cells were gated first, and compensation was adjusted with single-stain control tubes. Apoptotic cells were quantified as the sum of the cell percentages in Q2 and Q3 quadrants, and data were presented as the percentage of apoptotic cells for comparative analysis.

Detection of Mitochondrial Membrane Potential in Cells

Mitochondrial membrane potential was measured using the JC-1 mitochondrial membrane potential assay kit (C2006, Beyotime, Shanghai, China). The JC-1 staining working solution was prepared according to the manufacturer’s instructions. Then, 1 mL of cell culture medium and 1 mL of JC-1 staining working solution were added to each well of a 6-well plate and mixed thoroughly. After that, the mixture was incubated at 37 °C for 20 min. Finally, the mitochondrial membrane potential was analyzed using a flow cytometer (excitation: 490 nm, emission: 530 nm).

ELISA Assay

Human GTPase KRas (KRAS) ELISA Kit (CSB-EL012493HU) was obtained from CUSBIO (Wuhan, China), while Human PTEN ELISA kit (JLC-Y3331) and Human SQSTM1 ELISA kit (JLC6162) were purchased from Jingkang Bioengineering Co., Ltd. (Shanghai, China). The mouse interleukin-6 (IL-6) ELISA kit (PI326), mouse tumor necrosis factor-alpha (TNF-α) ELISA kit (PT512), total superoxide dismutase (SOD) assay kit with WST-8 (S0101S) and malondialdehyde (MDA) assay kit (S0131S) were from Beyotime (Shanghai, China); the creatinine (Cr) assay kit (D799853), and urea nitrogen assay kit (D799850) were from Sangon Biotech (Shanghai, China). These kits were employed to determine KRAS, PTEN and SQSTM1 in SA-AKI clinical samples, as well as the levels of IL-6, TNF-α, SOD, MDA, serum creatinine, and blood urea nitrogen in cells or mice, respectively. During the measurement process, the corresponding reagents were added carefully, strictly following the manufacturer’s instructions for the respective kits. Subsequently, each well’s optical density (OD) values were measured using a microplate reader. The wavelengths for measuring KRAS, PTEN, SQSTM1, IL-6, TNF-α and SOD were all 450 nm, that for MDA was 532 nm, that for serum creatinine was 546 nm, and that for blood urea nitrogen was 630 nm.

Fe²⁺ Level Detection

The intracellular Fe²⁺ levels were determined using the Cell Ferrous Iron (Fe²⁺) Fluorometric Assay Kit (E-BC-F101, Elabscience Biotechnology Co., Ltd., Wuhan, China). Briefly, the working solution was prepared strictly according to the manufacturer’s instructions, and then incubated at 37℃ in the dark for 30–60 min. Subsequently, intracellular Fe²⁺ levels were measured using a flow cytometer. Upon interaction between the probe and Fe²⁺, an irreversible orange (red) fluorescent product was formed (excitation: 542 nm, emission: 575 nm).

ROS Level Detection

The ROS Assay Kit (50101ES01, Yeasen, Shanghai, China) and the fluorescent probe DCFH-DA were used to detect ROS. HK-2 cells were inoculated into 6-well plates and treated with 100 ng/mL LPS. Subsequently, 2 mL of DCFH-DA, diluted to a final concentration of 10 µmol/L, was added to each well, and the cells were incubated at 37℃ for 20 min. Cells were washed three times with serum-free medium. After collecting the cells, they were analyzed using a flow cytometer (excitation: 488 nm, emission: 525 nm).

Transmission Electron Microscopy (TEM) for Cell Morphology Observation

The morphology of mitochondria and autophagy in HK-2 cells was observed by TEM. Cells were collected and fixed in 2.5% glutaraldehyde at 4℃ overnight. Next, samples were fixed in 1% osmium tetroxide at 4℃ for 2 h. Thereafter, the samples were carefully dehydrated using graded ethanol, then replaced with acetone, and embedded. Then, ultrathin sections were prepared, stained with uranyl acetate and lead citrate, and finally examined under a JEM1400 electron microscope (JEOL, Japan).

Establishment of the SA-AKI Animal Model

A total of thirty male C57BL/6 mice, aged 6–8 weeks and weighing 20–25 g, were purchased from Beijing Vital River Laboratory Animal Technology Co., Ltd. (Beijing, China). The mice were housed in a specific pathogen-free (SPF) environment, with free access to food and water. Approval for this study was obtained from the BaiRen Medical Laboratory Animal Welfare and Ethics Committee (BR.No20240713001). A total of 6 groups were formed by randomly assigning the mice (5 mice in each group): Control, LPS, LPS + sh-NC, LPS + sh-PTEN, LPS + sh-NC+ML385, and LPS + sh-PTEN+ML385 groups. Mice in the LPS + sh-NC, LPS + sh-PTEN, LPS + sh-NC+ML385, and LPS + sh-PTEN+ML385 groups were intravenously injected with 0.1 mL of adenovirus vector (control adenovirus for LPS + sh-NC and LPS + sh-NC+ML385 groups, sh-PTEN adenovirus for LPS + sh-PTEN and LPS + sh-PTEN+ML385 groups) via the tail vein at a dose of 1 × 10¹⁰ plaque-forming units (PFU) per mouse. The adenoviral vector injection was performed 24 h prior to LPS administration, which ensured sufficient time for the adenovirus to infect renal tubular epithelial cells and achieve stable expression of the target gene (PTEN knockdown), avoiding the impairment of adenovirus infection efficiency caused by early LPS-induced systemic inflammation and renal injury. After 24 h, the LPS, LPS + sh-NC, LPS + sh-PTEN, LPS + sh-NC+ML385, and LPS + sh-PTEN+ML385 groups each received an intraperitoneal injection of 10 mg/kg LPS, whereas the Control group was intraperitoneally administered 0.9% normal saline. For the LPS + sh-NC+ML385 and LPS + sh-PTEN+ML385 groups, mice were intraperitoneally injected with ML385 (30 mg/kg, MedChemExpress, New Jersey, USA) 2 h prior to LPS administration to specifically inhibit Nrf2 activity, while mice in the other groups received an equal volume of 0.9% normal saline. After 24 h, all the mice were moved to metabolic cages, fasted, and allowed free access to water. Twenty-four-hour urine output was collected. At 48 h post-injection, each group of mice was humanely euthanized, with kidney tissues and blood collected for further analysis.

Hematoxylin-Eosin (H&E) Staining

Mouse kidney tissues were placed in 4% paraformaldehyde for 6 h of fixation. Subsequently, they underwent dehydration with ethanol of varying concentrations, followed by embedding and sectioning. Each tissue section was stained using an Hematoxylin and Eosin Staining Kit (C0105S, Beyotime, Shanghai, China). After mounting the slides, the sections were observed under a microscope and photographed.

Renal Tubular Injury Score Assessment

Renal tubular injury was evaluated in H&E-stained renal tissue sections, as previously described methods [18]. Briefly, two independent pathologists, who were blinded to the experimental groups, assessed the severity of tubular injury by examining 10 randomly selected high-power fields per tissue sample. The injury score was graded on a 0–5 scale based on the extent of brush border loss, tubular dilation, cast formation, tubular necrosis, and neutrophil infiltration: 0: normal (no injury); 1: mild injury (0%-10% involvement); 2: moderate injury (11%–25% involvement); 3: severe injury (26%-49% involvement); 4: very severe injury (50%-75% involvement); 5: extensive injury (> 75% involvement). The final score for each sample was determined as the average score from the two independent assessors to minimize inter-observer variability.

Immunohistochemical Examination

The tissue sections were deparaffinization with xylene, after which they were hydrated using alcohols of various grades. After that, they were placed in freshly boiled citrate antigen retrieval solution for antigen retrieval. Subsequently, the endogenous peroxidase was blocked. The sections were successively treated with the primary antibody and the HRP-labeled goat anti-rabbit IgG polymer through incubation. Then, DAB (P0203, Beyotime, Shanghai, China) was used for color development, and hematoxylin was used for counterstaining. Subsequently, the sections were viewed under a microscope and images were taken.

Western Blot

Total proteins were isolated from HK-2 cells and kidney tissues using RIPA lysis buffer (P0013C, Beyotime, Shanghai, China) supplemented with a protease inhibitor. Then, we quantitatively detected the protein concentration with the BCA Protein Assay Kit (P0011, Beyotime, Shanghai, China). Samples were loaded, subjected to electrophoresis and transferred onto a membrane. Afterward, the PVDF membrane (IEVH85R, Merck KGaA, Darmstadt, Germany) was incubated overnight at 4℃ with primary antibodies diluted according to the manufacturer’s instructions. HRP-conjugated secondary antibodies (AS092, ABclonal, Wuhan, China, 1:5000 dilution) were used to incubate the membranes at 37℃ for 1 h. After the incubation was completed, development of the bands was performed using BeyoECL Plus (P0018S, Beyotime, Shanghai, China). The detailed information of the antibodies used in this study is shown in Table S1.

Statistical Analysis

Using the limma package, differences in gene expression among groups were compared. The clusterProfiler package and the org.Hs.eg.db package in R software were employed to analyze the enrichment results. All bioinformatics statistical analyses were performed using R software (version 4.2.1). GraphPad Prism 8.0 software was employed for statistical analysis, and group differences were analyzed using the t-test and one-way analysis of variance (ANOVA). For all experimental data (n ≥ 3), data were expressed as the mean ± standard deviation (SD), and P < 0.05 was considered statistically significant.

Results

DEGs Analysis in SA-AKI

By conducting differential expression analysis on the GSE65682 microarray, 1053 DEGs were found in total between Sepsis and Healthy samples. Among them, 1296 DEGs were upregulated in the Sepsis samples, and 2782 DEGs were downregulated in the Sepsis samples (Fig. 1A). We drew an expression heatmap that included the top 10 upregulated and downregulated DEGs (Fig. 1B). Based on the enrichment analysis of the GO and KEGG databases, we searched for the common functions and related pathways of a large number of genes within the set of DEGs (Fig. 1C-D). By drawing a Venn diagram to visualize the intersection of DEGs, FRGs, and ARGs, we obtained 15 differentially expressed autophagy-ferroptosis-related genes (A-FRGs) (Fig. 1E). Among them, the top 5 genes ranked by Degree were SQSTM1, HIF1A, PTEN, KRAS, and MAPK1, which were regarded as hub genes for subsequent analysis (Fig. 1F). Some studies have reported that HIF1A and MAPK1 are highly expressed in the serum of SA-AKI patients, and regulating their expressions can achieve the goal of treating SA-AKI [19, 20]. Therefore, subsequent studies were conducted on SQSTM1, PTEN, and KRAS.

Fig. 1.

DEGs analysis in SA-AKI. (A) A volcano plot of DEGs between Sepsis and Healthy. (B) A heatmap of the top 10 up-regulated and down-regulated DEGs between Sepsis and Healthy. (C) A treemap showing GO enrichment of DEGs. (D) KEGG pathways enriched analysis by DEGs. (E) A Venn diagram of overlapping genes among DEGs, ARGs, and FRGs. (F) A PPI network was constructed with 15 overlapping genes. (G) qRT-PCR verification of SQSTM1, PTEN, and KRAS expression in the blood of patients with SA-AKI. (H) ELISA verification of SQSTM1, PTEN, and KRAS expression in clinical samples. **p < 0.01. Abbreviations: DEGs: differentially expressed genes; ARGs: autophagy-related genes; FRGs: ferroptosis-related genes; GO: Gene ontology; qRT-PCR: quantitative reverse transcription polymerase chain reaction; SA-AKI: sepsis-associated acute kidney injury

The results of PCR verification on clinical samples revealed that SQSTM1, PTEN and KRAS expressions were significantly upregulated in the blood of patients with SA-AKI (P < 0.01, Fig. 1G). Consistently, ELISA validation results further confirmed that the levels of these three genes were also significantly elevated in the blood of SA-AKI patients (P < 0.01, Fig. 1H). Among them, PTEN exhibited the highest absolute expression level. Therefore, the PTEN gene was selected for experimental study.

The Influence of the Knockdown of PTEN on the Biological Functions of HK-2 Cells After Being Induced by LPS

To examine the function of PTEN in SA-AKI, HK-2 cells were induced using LPS to construct a SA-AKI cell model. Through qRT-PCR detection of PTEN expression, it was revealed that PTEN was overexpressed in HK-2 cells treated with LPS (P < 0.01, Fig. 2A). Subsequently, the knockdown efficiency was detected via qRT-PCR (P < 0.01, Fig. 2B), and sh-PTEN-1 was selected for subsequent research. Notably, when PTEN was knocked down, its expression level decreased markedly (P < 0.01), indicating that the lentivirus could potently inhibit PTEN expression in HK-2 cells (Fig. 2C).

Fig. 2.

The influence of the knockdown of PTEN on the biological functions of HK-2 cells after being induced by LPS (A) qRT-PCR detection of PTEN expression in LPS-induced HK-2 cells. (B) qRT-PCR analysis of PTEN expression in LPS-induced HK-2 cells after stable transfection with sh-PTEN lentiviruses targeting PTEN. (C) PTEN expression in LPS-induced HK-2 cells following stable PTEN knockdown. (D) CCK8 assay for cell viability. (E) Colony formation assay of cell proliferation in each group. (F) Flow cytometry analysis of cell apoptosis in each group. n = 3 replicates per group; Gating: live cells, single-stain compensation, scatter plot clusters; quantification: apoptotic cells = Q2 + Q3. (G) Detection of TNF-α and IL-6 levels in each group. (H) Western blot analysis of Beclin 1 and LC3 protein expression in LPS-induced HK-2 cells. *p < 0.05, **p < 0.01. Abbreviations: qRT-PCR: quantitative reverse transcription polymerase chain reaction; CCK8: Cell Counting Kit-8

We further explored and verified the function of PTEN in SA-AKI through in vitro cell behaviors. The CCK8 assay revealed that, compared with the sh-NC group, cell proliferation viability was significantly reduced in the LPS + sh-NC group (P < 0.01), confirming LPS-induced inhibition of cell viability. Furthermore, compared with the LPS + sh-NC group, the cell proliferation viability in the LPS + sh-PTEN group was significantly increased (P < 0.01), indicating that the knockdown of PTEN could improve the LPS-induced reduction in cell viability. Notably, the sh-PTEN group showed no significant difference in viability compared with the sh-NC group, suggesting that PTEN knockdown alone does not affect basal cell viability. The viability of cells at 72 h exceeded that at 48 h, so 72 h was selected for subsequent experiments (Fig. 2D). In addition, compared with the sh-NC group, the number of cell clones decreased and cell proliferation was inhibited in the LPS + sh-NC group. When PTEN was knocked down in LPS-treated cells, cell proliferation was restored (P < 0.01, Fig. 2E). According to flow cytometry findings, compared with the sh-NC group, cell apoptosis was significantly promoted in the LPS + sh-NC group, while the knockdown of PTEN reversed cell apoptosis (P < 0.01, Fig. 2F). Notably, PTEN knockdown alone did not alter basal apoptosis levels in the sh-PTEN group compared with the sh-NC group.

To investigate the effects of PTEN knockdown on SA-AKI, we detected inflammatory factors (TNF-α and IL-6) as well as autophagy-related proteins (Beclin 1 and LC3). LPS treatment alone significantly elevated TNF-α and IL-6 levels relative to the sh-NC group, and the knockdown of PTEN in LPS-treated cells reversed these LPS-induced increases in cytokine levels (P < 0.01, Fig. 2G). Furthermore, PTEN knockdown alone had no significant effect on basal TNF-α and IL-6 levels. In addition, compared with the sh-NC group, LPS treatment significantly upregulated the expression of Beclin 1 and LC3 (P < 0.01). Meanwhile, the knockdown of PTEN could reduce the increase in Beclin 1 and LC3 levels in LPS-treated HK-2 cells (P < 0.01, Fig. 2H). The above results all indicated that the knockdown of PTEN had a protective effect on LPS-treated HK-2 cells, while PTEN knockdown alone had no significant effect on basal cell function, inflammation, or autophagy.

Knockdown of PTEN Inhibited Ferroptosis in HK-2 Cells Subjected to LPS Treatment

For defining the role of ferroptosis in SA-AKI, we evaluated how PTEN knockdown affects the SA-AKI cell model. As shown in the results of the CCK8 assay, ferroptosis could reduce cell viability. The cell viability decreased significantly after LPS treatment, while the knockdown of PTEN improved the cell death after modeling. After the application of the ferroptosis inducer (Erastin), the cell viability declined, but PTEN knockdown was able to rescue ferroptosis in HK-2 cells (Fig. 3A).

Fig. 3.

Knockdown of PTEN inhibited the ferroptosis of HK-2 cells induced by LPS. (A) The cell viability was detected by CCK8. (B) Flow cytometry detection of mitochondrial membrane potential in each group. n = 3 replicates per group. Gating: Negative control validation, distinct cell clustering; quantification: Green/Red fluorescence ratio. (C) Flow cytometry detection of intracellular Fe²⁺ levels in each group. n = 3 replicates per group. Gating: Live cells, kit positive control reference; quantification: Positive cell percentage and MFI. (D) Flow cytometry detection of intracellular ROS levels in each group. n = 3 replicates per group. Gating: Live cells, negative control reference; quantification: Positive cell percentage and MFI. (E) Detection of SOD and MDA levels. (F) TEM observation of mitochondrial morphological changes. *p < 0.05, **p < 0.01. Abbreviations: CCK8: Cell Counting Kit-8; ROS: Reactive Oxygen Species; SOD: Superoxide dismutase; MDA: Malondialdehyde; MFI: Mean fluorescence intensity; TEM: Transmission Electron Microscopy

Mitochondrial membrane potential and Fe²⁺ levels in HK-2 cells were measured via flow cytometry. Compared with the LPS + sh-NC group, the Green/Red ratio was significantly decreased (P < 0.05) and the Fe²⁺ level was significantly reduced (P < 0.01) in the LPS + sh-PTEN group. In addition, the addition of Erastin promoted cellular ferroptosis, and the Fe²⁺ level in the LPS + sh-PTEN+Erastin group was significantly higher than that in the LPS + sh-PTEN group (P < 0.01, Fig. 3B-C, Figure S1 and S2). These results suggested that knockdown of PTEN could rescue cellular injury and effectively reverse cellular ferroptosis. Similarly, we confirmed that PTEN possesses antioxidant capacity in vitro. When stimulated by LPS and with the addition of Erastin, the fluorescence intensity increased significantly. When ferroptosis occurred, the level of ROS increased, but the knockdown of PTEN could significantly reduce it (P < 0.01, Fig. 3D).

To gain insight into the oxidative stress status and the degree of cell damage during SA-AKI, different kits were employed to determine the SOD and MDA levels in the cells. The knockdown of PTEN exerted an effect in raising SOD levels and lowering MDA levels (P < 0.01, Fig. 3E). TEM showed that when HK-2 cells underwent ferroptosis, their mitochondria were swollen, the membrane was blurred, the matrix was sparse, and the cristae were broken and reduced. The knockdown of PTEN improved the situation of ferroptosis and reduced cell damage (Fig. 3F).

Knockdown of PTEN Inhibited Ferroptosis by the Nrf2/SLC7A11/GPX4 Signaling Pathway

To confirm that SA-AKI could cause cellular ferroptosis, we verified ferroptosis-related proteins (COX2 and 4-HNE) by Western blot. When ferroptosis occurred, the protein levels of COX2 and 4-HNE increased, while the knockdown of PTEN decreased their levels (P < 0.01, Fig. 4A-B). Interestingly, given that the Nrf2/SLC7A11/GPX4 pathway has a vital role in ferroptosis regulation, we proposed a novel mechanism whereby this pathway modulates LPS-induced SA-AKI. Western blot analysis identified the expression levels of proteins involved in the Nrf2/SLC7A11/GPX4 signaling pathway (Nrf2, SLC7A11, and GPX4) in the cells, indicating that PTEN knockdown could activate this pathway, with related proteins showing significantly elevated expression levels (P < 0.01, Fig. 4C-D). These findings revealed that PTEN knockdown could significantly inhibit ferroptosis caused by SA-AKI and effectively alleviate cell death.

Fig. 4.

Knockdown of PTEN inhibited ferroptosis by the Nrf2/SLC7A11/GPX4 signaling pathway. The protein expressions of (A)-(B) COX2, 4-HNE and (C)-(D) Nrf2, SLC7A11, GPX4 in LPS-induced HK-2 cells were detected by Western blot. *p < 0.05, **p < 0.01.

Knockdown of PTEN Inhibited the Autophagy-Dependent Ferroptosis of HK-2 Cells Induced by LPS

For determining the association between autophagy and ferroptosis, as well as investigating how PTEN affects SA-AKI, the cell models of SA-AKI were first induced with Erastin, and then further induced with the autophagy inducer (Rapa) or autophagy inhibitor (3-MA) according to different experimental groups. The CCK8 assay demonstrated that cell viability in the LPS + sh-NC+Rapa group was lower than that in the LPS + sh-NC group, while cell viability in the LPS + sh-NC + 3-MA group was higher than that in the LPS + sh-NC group, indicating that autophagy aggravates LPS-induced ferroptosis, whereas blocking autophagy can alleviate this pathological process. In addition, cell viability in the LPS + sh-PTEN group was significantly higher than that in the LPS + sh-NC group (P < 0.01). After treatment with 3-MA, cell viability in this group was further enhanced, while the addition of Rapa reversed the protective effect induced by PTEN knockdown (Fig. 5A). Mitochondrial membrane potential, Fe²⁺, and ROS levels in the cells were detected via flow cytometry. The detection of mitochondrial membrane potential showed that the proportion of damaged cells in the LPS + sh-PTEN group was significantly lower than that in the LPS + sh-NC group. 3-MA treatment could significantly reduce the number of damaged cells, while Rapa treatment promoted cell damage. Meanwhile, the changing trends of Fe²⁺ and ROS levels were consistent with those of mitochondrial membrane potential (P < 0.01, Fig. 5B-D).

Fig. 5.

Knockdown of PTEN inhibited autophagy-dependent ferroptosis in HK-2 cells induced by LPS. (A) Cell viability was detected by CCK8 assay. (B) Mitochondrial membrane potential were detected by flow cytometry. n = 3 replicates per group. Gating: Negative control validation, distinct cell clustering; quantification: Green/Red fluorescence ratio. (C) Fe²⁺ levels were detected by flow cytometry. n = 3 replicates per group. Gating: Live cells, kit positive control reference; quantification: Positive cell percentage. (D) ROS levels were detected by flow cytometry. n = 3 replicates per group. Gating: Live cells, negative control reference; quantification: Positive cell percentage. (E) Detection of SOD and MDA levels. (F) The cell morphology during autophagy was observed by TEM. **p < 0.01. Abbreviations: CCK8: Cell Counting Kit-8; ROS: Reactive Oxygen Species; SOD: Superoxide dismutase; MDA: Malondialdehyde; MFI: Mean fluorescence intensity; TEM: Transmission Electron Microscopy

Different kits were used to detect SOD and MDA levels in the cells. The results showed that, compared with the LPS + sh-PTEN group, the LPS + sh-PTEN+Rapa group had significantly decreased SOD levels and significantly increased MDA levels (P < 0.01), while the LPS + sh-PTEN + 3-MA group showed an increasing trend in SOD levels and a decreasing trend in MDA levels (P < 0.01, Fig. 5E). As observed under the transmission electron microscope, when autophagy occurred in the cells, various damaged organelles would be enclosed by double-membrane vacuoles to form autophagosomes. The quantity of autophagosomes showed a positive correlation with the degree of cell damage. LPS treatment could induce cells to produce autophagosomes. PTEN knockdown could reduce the formation of autophagosomes, and this effect was further enhanced after 3-MA treatment, confirming that PTEN knockdown can effectively inhibit excessive autophagy in cells (Fig. 5F).

Knockdown of PTEN Inhibited Autophagy-Dependent Ferroptosis by the Nrf2/SLC7A11/GPX4 Pathway

To further determine autophagy-dependent ferroptosis in cells induced by SA-AKI and its molecular mechanisms, western blot was employed to detect the expression levels of ferroptosis-related proteins (COX2 and 4-HNE) and Nrf2/SLC7A11/GPX4 pathway-related proteins (Nrf2, SLC7A11, and GPX4). The results showed that, compared with the LPS + sh-NC group, the expression levels of COX2 and 4-HNE were significantly decreased in the LPS + sh-PTEN group (P < 0.05, P < 0.01). Treatment with Rapa further upregulated these ferroptosis-related proteins, whereas treatment with 3-MA downregulated their expression (P < 0.05, P < 0.01, Fig. 6A-B). In contrast, the expression trends of Nrf2, SLC7A11, and GPX4 were the opposite. Their levels were significantly higher in the LPS + sh-PTEN group than in the LPS + sh-NC group (P < 0.05, P < 0.01). Rapa treatment reduced the expression of these pathway proteins, while 3-MA treatment further increased their expression (P < 0.05, P < 0.01, Fig. 6C-D). These results indicate that autophagy is positively correlated with ferroptosis, as enhancing autophagy aggravates LPS-induced ferroptosis, while blocking autophagy alleviates this pathological process. It is confirmed that PTEN knockdown inhibits autophagy-dependent ferroptosis by activating the Nrf2/SLC7A11/GPX4 signaling pathway.

Fig. 6.

Knockdown of PTEN inhibited autophagy-dependent ferroptosis by the Nrf2/SLC7A11/GPX4 signaling pathway. The protein expressions of (A)-(B) COX2, 4-HNE and (C)-(D) Nrf2, SLC7A11, GPX4 in HK-2 cells induced by LPS were detected by Western blot. *p < 0.05, **p < 0.01

Rescue Experiments Verified that PTEN Knockdown Inhibited Autophagy-Dependent Ferroptosis in SA-AKI via the Nrf2/SLC7A11/GPX4 Pathway

To investigate the regulatory effect of PTEN on autophagy-dependent ferroptosis via the Nrf2/SLC7A11/GPX4 pathway, rescue experiments were performed using LPS-induced SA-AKI cells that were treated with Erastin prior to the Nrf2 inhibitor ML385. CCK8 assay results showed that cell viability was significantly increased in the LPS + sh-PTEN group compared with the LPS + sh-NC group (P < 0.01), while cell viability was significantly decreased in the LPS + sh-PTEN+ML385 group after treatment with ML385 (P < 0.01, Fig. 7A). Mitochondrial membrane potential, Fe²⁺, and ROS levels were detected by flow cytometry. Mitochondrial membrane potential analysis indicated that the Green/Red ratio was markedly reduced in the LPS + sh-PTEN group relative to the LPS + sh-NC group (P < 0.01), whereas this ratio was significantly upregulated following ML385 treatment (P < 0.01), accompanied by aggravated cellular injury (Fig. 7B). The variation trends of intracellular Fe²⁺ and ROS levels were consistent with those of mitochondrial membrane potential (Fig. 7C-D). In addition, detection of oxidative stress markers revealed that compared with the LPS + sh-NC group, the LPS + sh-PTEN group exhibited a significant increase in SOD level and a remarkable decrease in MDA level (P < 0.01). Following ML385 administration, SOD level was significantly decreased and MDA level was sharply elevated (P < 0.01, Fig. 7E). Western blot was applied to examine the expression levels of ferroptosis-related proteins (COX2 and 4-HNE) and key proteins in the Nrf2/SLC7A11/GPX4 signaling pathway. The results demonstrated that the protein expression levels of COX2 and 4-HNE were significantly lower in the LPS + sh-PTEN group than in the LPS + sh-NC group (P < 0.01), and the addition of ML385 significantly reversed the downregulation of ferroptosis-related proteins mediated by PTEN knockdown (Fig. 7F). The expression changes of the pathway-related proteins Nrf2, SLC7A11, and GPX4 showed an entirely opposite trend to those of ferroptosis-related proteins (Fig. 7G). These findings confirmed that Nrf2 plays a pivotal role in the regulation of autophagy-dependent ferroptosis in SA-AKI cells by PTEN, and PTEN mediates autophagy-dependent ferroptosis during the progression of SA-AKI by modulating the Nrf2/SLC7A11/GPX4 signaling pathway.

Fig. 7.

ML385 verified PTEN regulated autophagy-dependent ferroptosis via Nrf2/SLC7A11/GPX4 pathway in LPS-induced SA-AKI cells. (A) Cell viability was detected by CCK8 assay. (B) Mitochondrial membrane potential was detected by flow cytometry. n = 3 replicates per group. Gating: Negative control validation, distinct cell clustering; quantification: Green/Red fluorescence ratio. (C) Intracellular Fe²⁺ levels were detected by flow cytometry. n = 3 replicates per group. Gating: Live cells, kit positive control reference; quantification: Positive cell percentage. (D) Intracellular ROS levels were detected by flow cytometry. n = 3 replicates per group. Gating: Live cells, negative control reference; quantification: Positive cell percentage. (E) SOD and MDA levels were detected by corresponding detection kits. (F) Protein expressions of 4‑HNE and COX2 were detected by Western blot. (G) Protein expressions of key proteins in the Nrf2/SLC7A11/GPX4 signaling pathway were detected by Western blot. *p < 0.05, **p < 0.01. Abbreviations: SA-AKI: sepsis-associated acute kidney injury; CCK8: Cell Counting Kit-8; ROS: Reactive Oxygen Species; MFI: Mean fluorescence intensity; SOD: Superoxide dismutase; MDA: Malondialdehyde

Effects of PTEN in Mice with Endotoxin-induced SA-AKI

An LPS-induced SA-AKI model in mice was established. PTEN’s effects on mice with LPS-induced SA-AKI were evaluated by mouse urine output, serum biochemical indices, and kidney-related detection indices. The results of monitoring mice’s 24 h urine output, as shown in Fig. 8A, indicated that after LPS induction, in contrast to the Control group, urine output showed a significant reduction (P < 0.05). Meanwhile, in the LPS group, serum creatinine, blood urea nitrogen, IL-6, TNF-α, and MDA levels were significantly elevated (P < 0.01), with a marked decrease in SOD level (P < 0.01). We injected the adenovirus vector sh-PTEN via the caudal vein to achieve the knockdown of PTEN expression, to investigate the functional role of PTEN in SA-AKI. Compared with the LPS group, there was a significant inhibition in the serum levels of creatinine, blood urea nitrogen, TNF-α, IL-6, and MDA (P < 0.01), and the SOD level increased significantly (P < 0.01, Fig. 8B-D). To further verify whether the protective effect of PTEN knockdown against SA-AKI was specifically mediated by the Nrf2 pathway, in vivo rescue experiments were performed using ML385. Compared with the LPS + sh-PTEN group, the LPS + sh-PTEN+ML385 group exhibited a significant reduction in 24 h urine output (P < 0.05), significantly increased serum levels of creatinine, blood urea nitrogen, IL-6, TNF-α, and MDA (P < 0.01), and a markedly decreased SOD level (P < 0.01, Fig. 8A-D). In addition, H&E staining results revealed that the structural integrity of the kidneys in the LPS group, the LPS + sh-NC group and the LPS + sh-NC+ML385 group was damaged, with diffuse erosion of tubular cells and tubular dilation. The glomeruli showed atrophy and widened interspaces, and the renal tubular epithelial cells were shed. Vacuolar degeneration, epithelial cell necrosis, and inflammatory cell infiltration were observed. The LPS + sh-PTEN group showed alleviated pathological changes, whereas the LPS + sh-PTEN+ML385 group exhibited aggravated renal pathological damage (Fig. 8E). Correspondingly, after LPS induction, the renal injury score of mice in the LPS group was significantly higher than that in the Control group (P < 0.01). Following PTEN knockdown, the renal injury score in the LPS + sh-PTEN group was significantly reduced compared with both the LPS group and the LPS + sh-NC group (P < 0.01). The renal injury score in the LPS + sh-PTEN+ML385 group was significantly higher than that in the LPS + sh-PTEN group (P < 0.01), reversing the protective effect of PTEN knockdown on LPS-induced SA-AKI (Fig. 8F). These data demonstrated that the model of SA-AKI in mice was successfully established, and the knockdown of PTEN could effectively ameliorate LPS-induced SA-AKI in mice, and this protective effect was specifically mediated by the Nrf2 pathway.

Fig. 8.

The effects of knockdown of PTEN on SA-AKI mice. (A) 24 h urine output monitoring. (B) Detection of blood creatinine and blood urea nitrogen levels. (C) The TNF-α and IL-6 levels were quantified by ELISA. (D) Detection of SOD and MDA levels. (E) The renal tissue structure of the mice was observed by H&E staining. (F) Renal injury score. *p < 0.05, **p < 0.01. Abbreviations: SA-AKI: Sepsis-associated acute kidney injury; SOD: Superoxide dismutase; MDA: Malondialdehyde

Knockdown of PTEN Inhibited Autophagy-Dependent Ferroptosis in a Model of SA-AKI

To explore the therapeutic mechanism underlying PTEN in SA-AKI in vivo, we further conducted analyses using immunohistochemistry and western blot. Beclin 1, LC3, COX2, and 4-HNE expression was increased in the SA-AKI mouse model. The condition of SA-AKI mice was rescued by the knockdown of PTEN, as the LPS + sh-PTEN group showed significantly decreased expression of Beclin 1, LC3, COX2, and 4-HNE compared with the LPS and LPS + sh-NC groups (P < 0.05). Notably, the expression levels of Beclin 1, LC3, COX2, and 4-HNE were significantly increased in the LPS + sh-PTEN+ML385 group compared with the LPS + sh-PTEN group (P < 0.05), reversing the inhibitory effect of PTEN knockdown on autophagy and ferroptosis (Fig. 9A-D). Additionally, the LPS + sh-PTEN group exhibited higher protein expressions of Nrf2, SLC7A11, and GPX4, which suggests that the knockdown of PTEN is capable of activating the Nrf2/SLC7A11/GPX4 pathway. In contrast, the expression levels of Nrf2, SLC7A11, and GPX4 were significantly decreased in the LPS + sh-PTEN+ML385 group compared with the LPS + sh-PTEN group (P < 0.05), indicating that ML385 effectively inhibited the activation of the Nrf2/SLC7A11/GPX4 pathway mediated by PTEN knockdown (Fig. 9E). These results confirmed that the knockdown of PTEN could activate the Nrf2/SLC7A11/GPX4 pathway, thus suppressing autophagy-dependent ferroptosis in SA-AKI.

Fig. 9.

Knockdown of PTEN inhibited autophagy-dependent ferroptosis in a model of SA-AKI. (A)-(C) The expressions of LC3 and Beclin 1 in renal tissues were detected by immunohistochemical staining. The protein expressions of (D) COX2, 4-HNE and (E) Nrf2, SLC7A11, GPX4 in mouse renal tissues were detected by Western blot. *p < 0.05, **p < 0.01

Discussion

At present, the pathophysiological mechanism of SA-AKI remains unclear. Its occurrence mechanism is generally considered to be related to inflammatory response, oxidative stress, metabolic reprogramming, microvascular dysfunction, and so on [21]. Existing studies have pointed out that ferroptosis plays an essential role in damaging renal tubular epithelial cells [22–24]. Specifically, ferroptosis can be categorized as a type of autophagy-dependent cell death, referred to as autophagy-dependent ferroptosis [25]. Nevertheless, existing knowledge regarding the mechanisms that underlie autophagy-dependent ferroptosis in SA-AKI is still limited. This study involved screening SA-AKI-related DEGs from clinical samples through bioinformatics analysis, followed by verification using qRT-PCR and ELISA. Subsequently, the role of PTEN in SA-AKI was confirmed through in vitro and in vivo assays. These results showed that PTEN could effectively ameliorate autophagy-dependent ferroptosis in SA-AKI via the Nrf2/SLC7A11/GPX4 signaling pathway.

This research revealed that PTEN is a gene associated with autophagy-dependent ferroptosis in SA-AKI. Through verification using clinical samples, PTEN expression levels were noted to be raised in the blood of SA-AKI patients. The PTEN gene is the only tumor suppressor gene discovered so far that has both protein-lipid phosphatase and phosphatase activities. It is involved in diverse biological processes, including genomic stability, cell viability, proliferation, survival, and energy metabolism [26, 27]. Over the past few years, the function of PTEN in acute kidney injury has gradually emerged [28]. According to a study by Wang et al., the knockdown of PTEN was found to enhance how miR-22-3p inhibits the inflammatory response in SA-AKI [29]. However, to date, no study has explored the relationship between PTEN and autophagy-dependent ferroptosis in SA-AKI.

LPS, an endotoxin produced by Gram-negative bacteria, can promote the secretion of diverse pro-inflammatory cytokines within renal tissues. Such a process then sparks ROS generation and induces renal tubular cell demise [30, 31]. Experiments conducted in vitro and in vivo have confirmed that LPS induces autophagy-dependent ferroptosis in SA-AKI, during which the levels of inflammatory factors, Fe²⁺, and ROS increase. When SA-AKI occurs, the body’s immune system detects the invasion of pathogens and immediately triggers a systemic inflammatory response. Considerable amounts of inflammation-eliciting cytokines are released, directly impairing mitochondrial function. At the same time, the inflammatory state leads to the excessive activation of the intracellular autophagy process. A large number of autophagosomes encapsulate and degrade ferritin (an iron storage protein), and large amounts of iron ions are released into the cytoplasm, leading to a sharp rise in intracellular free iron ion concentration [32, 33]. The damage to mitochondria and the imbalance of iron ions together lead to a further increase in ROS. The Fenton reaction occurs between excessive ROS and Fe²⁺, resulting in damage to the cell membrane, oxidation of proteins and nucleic acids, and further aggravates renal cell damage [34–36]. We further discussed the mechanism of action of PTEN in autophagy-dependent ferroptosis induced by LPS in SA-AKI. Knockdown of PTEN could effectively reduce the level of the inflammatory response, alleviate mitochondrial damage, and inhibit excessive autophagic activity. These changes subsequently significantly decreased both Fe²⁺ concentration and ROS levels within the cytoplasm, thereby alleviating cell damage. The above research results fully revealed the inherent association between PTEN and autophagy-dependent ferroptosis, indicating that PTEN could regulate autophagy-dependent ferroptosis in SA-AKI.

In addition, the Nrf2/SLC7A11/GPX4 pathway serves as an important pathway in regulating ferroptosis, primarily inhibiting ferroptosis through antioxidant effects. After Nrf2 is activated, it upregulates the expression of SLC7A11, stimulates cystine uptake for GSH synthesis, and then enhances GPX4 activity to reduce lipid peroxides, preventing ferroptosis from occurring [37–39]. Guo et al. reported that suppressing ferroptosis via activating the Nrf2/SLC7A11/GPX4 pathway contributed to alleviating septic liver injury [40]. This study explored how PTEN inhibited autophagy-dependent ferroptosis through the Nrf2/SLC7A11/GPX4 signaling pathway. PTEN knockdown activated the Nrf2/SLC7A11/GPX4 signaling pathway, with significant increases in the expression levels of its target proteins Nrf2, SLC7A11, and GPX4. Meanwhile, treatment with the Nrf2 inhibitor ML385 significantly reversed the regulatory effects mediated by PTEN knockdown in both in vitro and in vivo models. In vitro changes were manifested as decreased cell viability, aggravated mitochondrial damage, elevated levels of Fe²⁺, ROS and MDA, reduced SOD levels, upregulated expression of ferroptosis-related proteins, and downregulated expression of the Nrf2/SLC7A11/GPX4 pathway-related proteins. Consistently, in vivo intervention with ML385 abrogated the renal protective effects of PTEN knockdown, leading to deteriorated renal function, intensified inflammatory infiltration, aggravated renal pathological injury, and blocked Nrf2 pathway activation in mice. In vitro and in vivo experiments confirmed that PTEN can restrain autophagy-dependent ferroptosis via the Nrf2/SLC7A11/GPX4 signaling pathway, thus ameliorating SA-AKI.

Nevertheless, several limitations must be recognized. While autophagy-dependent ferroptosis is identified as a critical pathway modulated by PTEN, the study exclusively focuses on this particular type of programmed cell death. The potential crosstalk between PTEN and other cell death modalities, such as apoptosis, necroptosis, pyroptosis, or even classical necrosis, remains entirely uncharacterized in the context of SA-AKI. Future investigations should provide a more comprehensive mechanistic framework for PTEN-targeted therapies in SA-AKI.

Conclusions

This study clarified a previously unreported mechanism, namely, that PTEN knockdown activates the Nrf2/SLC7A11/GPX4 signaling pathway to regulate autophagy-dependent ferroptosis in SA-AKI, which may serve as a potential therapeutic target for SA-AKI.

Supplementary Information

Below is the link to the electronic supplementary material.

Author Contributions

Dede Lian: Conception and design of the research, Acquisition of data, Analysis and interpretation of data, Statistical analysis, Drafting the manuscript; Qingling Zhang: Revision of manuscript for important intellectual content; Hongjun Li: Conception and design of the research, Revision of manuscript for important intellectual content.

Funding

None.

Data Availability

The datasets used and/or analyzed during the current study are available from the corresponding author via email request.

Declarations

Ethics Approval and Consent to Participate

Approved by the Ethics Committee of the China-Japan Union Hospital of Jilin University. Informed consent to use their samples and data was given by all patients. This study was obtained from the BaiRen Medical Laboratory Animal Welfare and Ethics Committee (BR.No20240713001).

Consent for Publication

Not applicable.

Competing Interests

The authors declare no competing interests.