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Human spindle-shaped urine-derived stem cell exosomes alleviate severe fatty liver ischemia–reperfusion injury by inhibiting ferroptosis via GPX4

Stem Cell Research & Therapy volume 16, Article number: 81 (2025) Cite this article

Abstract

Background

Severe hepatic steatosis can exacerbate Ischemia–reperfusion injury (IRI), potentially leading to early graft dysfunction and primary non-function. In this study, we investigated the heterogeneity of different subpopulations of Urine-derived stem cells (USCs) to explore the most suitable cell subtype for treating severe steatotic liver IRI.

Methods

This study utilized scRNA-seq and Bulk RNA-seq to investigate the transcriptional heterogeneity between Spindle-shaped USCs (SS-USCs) and Rice-shaped USCs (RS-USCs). Additionally, rat fatty Liver transplantation (LT) model, mouse fatty liver IRI model, and Steatotic Hepatocyte Hypoxia-Reoxygenation (SHP-HR) model were constructed. Extracellular vesicles derived from SS-USCs and RS-USCs were isolated and subjected to mass spectrometry analysis. The therapeutic effects of Spindle-shaped USCs Exosomes (SS-USCs-Exo) and Rice-shaped USCs Exosomes (RS-USCs-Exo) were explored, elucidating their potential mechanisms in inhibiting ferroptosis and alleviating IRI.

Results

Multiple omics analyses confirmed that SS-USCs possess strong tissue repair and antioxidant capabilities, while RS-USCs have the potential to differentiate towards specific directions such as the kidney, nervous system, and skeletal system, particularly showing great application potential in renal system reconstruction. Further experiments demonstrated in vivo and in vitro models confirming that SS-USCs and SS-USCs-Exo significantly inhibit ferroptosis and alleviate severe fatty liver IRI, whereas the effects of RS-USCs/RS-USCs-Exo are less pronounced. Analysis comparing the proteomic differences between SS-USCs-Exo and RS-USCs-Exo revealed that SS-USCs-Exo primarily inhibit ferroptosis and improve cellular viability by secreting exosomes containing Glutathione Peroxidase 4 (GPX4) protein. This highlights the most suitable cell subtype for treating severe fatty liver IRI.

Conclusions

SS-USCs possess strong tissue repair and antioxidant capabilities, primarily alleviating ferroptosis in the donor liver of fatty liver through the presence of GPX4 protein in their exosomes. This highlights SS-USCs as the most appropriate cell subtype for treating severe fatty liver IRI.

Introduction

The management and outcomes for individuals suffering from terminal liver disease are critically influenced by the prevalent scarcity of suitable donor organs, representing a significant barrier in the realm of liver transplants [1]. To address this issue, the use of marginal donor livers has increased significantly in recent years. Marginal donor livers mainly refer to fatty livers, elderly donor livers, donor livers with a history of viral infection, and donor livers with prolonged ischemic times, with a significant proportion of fatty liver donors [2,3,4]. Nonetheless, more than 60% of livers with severe steatosis are often removed. IRI in the liver may be worsened by severe hepatic steatosis, resulting in early graft malfunction and primary nonfunction[5]. Therefore, reducing IRI in severely steatotic donor livers holds significant importance in expanding the donor liver pool for transplantation.

USCs, sourced from urine, are increasingly recognized for their multiple advantages including non-invasive collection, ease of access, and minimal costs [6]. These cells demonstrate a robust capacity for self-renewal and the ability to differentiate into various cell lineages, positioning them as a valuable resource for tissue and organ regeneration. Recent research has advanced substantially, with USCs now being capable of differentiating into neuron-like cells, indicating potential applications in neuroscience [7, 8]. Furthermore, their utility in the field of regenerative medicine and tissue engineering is expanding. Current applications involve the use of USCs for the reconstruction of diverse tissues such as skin, bone, and cartilage, thereby opening new pathways for medical treatments [9,10,11,12]. In tissue repair, USCs have shown promising therapeutic benefits, including the management of renal IRI [13], prevention of diabetic nephropathy [14], stimulation of angiogenesis, and wound healing[15]. However, their heterogeneity and uncertain differentiation potential limit the clinical application of USCs.

According to previous studies and our observations, USCs can be divided into two distinct subpopulations: SS-USCs and RS-USCs [16]. However, limited studies have confirmed the true functions of these USC subpopulations in subsequent experiments. In addition, the limited knowledge of USC heterogeneity poses a challenge for identifying the most suitable cell types for clinical therapeutic purposes. Thus, a thorough understanding of the diversity of USCs must be gained by analyzing a large number of samples. This will enable us to identify the most suitable cell subtypes for effective treatment of severe fatty liver steatosis. The functions of USCs primarily rely on their paracrine activity, with exosomes being a key paracrine factor of USCs [17, 18]. Exosomes mediate intercellular communication by transferring mRNA, non-coding RNA, and proteins between cells. They exhibit low immunogenicity and are convenient for application and storage. Therefore, exosome-based cell-free therapy holds significant potential for clinical applications. However, the potential roles and mechanisms of USC-derived exosomes in fatty liver IRI remain unclear.

In this study, we explored the potential roles and mechanisms of exosomes derived from two subtypes of USCs in fatty liver IRI. First, we examined the functional heterogeneity of SS-USCs and RS-USCs. Then, we compared the abilities of RS-USCs and SS-USCs to alleviate severe fatty liver IRI. Finally, we employed 4D label-free proteomics to analyze the protein expression profiles of SS-USC-Exo and RS-USC-Exo, aiming to identify the key molecular mechanisms by which SS-USC-Exo inhibits IRI.

Materials and methods

Cultivation of USCs and isolation of SS-USCs and RS-USCs

To isolate USCs, fresh urine samples were gathered from healthy adults aged 20 to 30 years. USCs were isolated from 100 to 200 ml urine samples using a single-centrifugation method: the urine sample was evenly divided into four 50-ml sterile centrifuge tubes and centrifuged at 1500 r/min for 5 min at room temperature. The supernatant was discarded, leaving 0.3–0.5 ml of liquid in each tube. To each tube, 3 ml of USC culture medium was added to resuspend the pellet, and the mixture was gently homogenized. The suspension was then seeded into 24-well plates pre-treated with 0.1% gelatin, with approximately 0.5 ml per well [16]. After 48 h of incubation, cell attachment was observed. The culture medium was replaced, and non-adherent cells were removed as part of the initial screening. The cells were further cultured with medium changes every two days for continuous selection and purification. Daily observations of cell morphology, growth, and contamination were performed using a light microscope. Any bacterial contamination was addressed immediately. Mixed subpopulations of the two USC subtypes were discarded, and wells containing only spindle-shaped or rice-shaped USCs were selected. Upon reaching a confluence of approximately 1/3 to 1/2 of the well surface area, the cells were passaged and labeled as Passage 1 (P1) SS-USCs or RS-USCs. Collected at passage levels between three and five (P3-P5), both SS-USCs and RS-USCs were utilized for subsequent experimental assessments. The expression levels of cell surface markers in SS-USCs and RS-USCs were analyzed using flow cytometry to confirm the characteristics of USCs.

Cell morphology

Cell images were captured using CKX53 imaging equipment provided by Olympus Corporation (Tokyo, Japan). Subsequently, the morphology of these cells was analyzed with ImageJ software.

Bulk RNA-Seq

Three healthy individuals provided SS-USCs and RS-USCs and bulk RNA sequencing was extracted from P3 SS-USCs and RS-USCs. Extraction of total RNA from the samples was performed to ensure that the concentration of RNA in each specimen was maintained at a minimum of 20 ng/ul, accumulating a total RNA yield of no less than 2 ug.Subsequently, sequencing libraries were prepared. After library preparation and quality assessment, sequencing was performed using the Illumina HiSeq/NextSeq platform.

scRNA-Seq

SS-USCs- and RS-USCs were extracted from three healthy individuals and P3 SS-USCs- and RS-USCs from each donor were mixed in proportion. scRNA-seq was performed using the NBelab C Series High-throughput Single-cell RNA Library Preparation Set (BGI, China). Sequencing was performed on the BGI DNBSEQ-T7RS platform.

Establishment of a rat model for fatty LT

Sprague–Dawley rats were gradually transitioned to a 100% high-fat diet starting from six weeks of age and maintained until 26–30 weeks [19]. The Animal Ethics Committee of Qingdao University granted approval for this study. All animals were euthanized using pentobarbital solution (100–200 mg/kg) in accordance with the 2020 edition of the American Veterinary Medical Association (AVMA) Guidelines for the Euthanasia of Animals.

Prior to the surgical procedure, recipient animals underwent an eight-hour fasting period, although no fasting was required for donor animals. Throughout the preoperative phase, unrestricted water access was provided to both groups. Anesthesia was induced through inhalation of isoflurane and intraperitoneal injection of Rompun (2 mg/kg). The surgical method employed for LT was a single-operator, two-cuff anastomosis technique [20].

The experimental design included seven distinct groups, each composed of five rats: sham, PBS, Fer-1, RS-USCs, SS-USCs, RS-USC-Exos, and SS-USC-Exos. The sham group was subjected to only laparotomy followed by suturing of the incision. In the postoperative phase, rats in the PBS group were administered 500 μL of PBS intravenously through the portal vein. Rats in the Fer-1 group received 10 mg/kg/day of Fer-1(MCE, USA) via intraperitoneal injection. Both the RS-USCs and SS-USCs groups received portal vein injections of 2 × 10^6 cells from their respective cell types. The groups receiving exosomes, RS-USC-Exos and SS-USC-Exos, were each administered 20 μg per rat through the portal vein. On the third day following the surgery, all rats were euthanized to collect their liver tissues and serum for analysis.

Establishment of the mouse fatty liver IRI model

From the age of six weeks, C57BL/6 mice were transitioned to an exclusive high-fat diet and this regimen was maintained over a period of 30 weeks [19]. Approval for this study was granted by the Animal Ethics Committee at Qingdao University. All animals were euthanized using pentobarbital solution (100–200 mg/kg) in accordance with the 2020 edition of the American Veterinary Medical Association (AVMA) Guidelines for the Euthanasia of Animals.

Anesthesia was induced through intraperitoneal injection of pentobarbital solution (50–60 mg/kg).Temporary occlusion of the portal vein, hepatic artery, and bile duct in the left and middle liver lobes was achieved using microvascular clamps, which resulted in about 70% ischemia of the liver. This ischemic condition was sustained for one hour prior to the restoration of blood flow. Twelve hours post-reperfusion, samples of liver tissue and serum were collected.

Seven experimental groups were established: sham, PBS, Fer-1, RS-USCs, SS-USCs, RS-USC-Exos, and SS-USC-Exos, each comprising five mice. The procedure for the sham group included only laparotomy followed by suturing. For the PBS group, a 500 μL intravenous PBS injection was administered through either the splenic or portal vein. In the Fer-1 group, Fer-1(MCE, USA) was administered intraperitoneally at a dosage of 5 mg/kg/day. RS-USCs group: Intravenous injection of 1 × 10^6 RS-USCs via the splenic or portal vein. SS-USCs group: Intravenous injection of 1 × 10^6 SS-USCs via the splenic or portal vein. RS-USCs-Exo group: Intravenous injection of RS-USCs-Exo 10 μg/mouse via splenic vein or portal vein. SS-USCs-Exo group: Intravenous injection of SS-USCs-Exo exosomes 10 μg/mouse via splenic vein or portal vein. Mice from each group were euthanized 12 h after reperfusion and the liver tissues and serum were collected.

Establishment of the SHP-HR model

HL7702 cells were cultured on a 12-well plate and then treated with oleic acid (Sigma, USA) and palmitic acid (Sigma, USA) for a duration of 24 h to induce steatosis. Subsequently, the cells were rinsed and incubated in DMEM medium devoid of fetal bovine serum and glucose (Gibco, USA), supplemented with mineral oil (Solarbio, China), for four hours at a temperature of 37 °C and an atmosphere containing 5% CO2. Following multiple washes using PBS, the cells underwent an exposure to DMEM enriched with 10% FBS and high glucose (Gibco, USA) over a period of six hours to facilitate reoxygenation [21].

Experimental groups: The control group comprised HL7702 cells with or without steatosis induction and without HR treatment. HL7702 group: HL7702 cells without steatosis induction, however, with HR treatment (HP-HR). S-HL7702 group: Steatosis-induced HL7702 cells treated with HR (SHP-HR). Erastin group: S-HL7702 cells treated with Erastin (MCE, USA). Fer-1 group: SHP-HR, supplemented with Fer-1 (MCE, USA). Z-VAD-FMK group: SHP-HR, supplemented with Z-VAD-FMK (MCE, USA). Necrostatin-1 group: SHP-HR, supplemented with Necrostatin-1 (MCE, USA). RS-USCs group: SHP-HR, co-cultured with RS-USCs using 0.4 μm Transwell chambers (Corning, USA). SS-USCs group: SHP-HR, co-cultured with SS-USCs using 0.4 μm Transwell chambers (Corning, USA). SS-USCs + GW group: SHP-HR, co-cultured with SS-USCs using 0.4μm Transwell chambers (Corning, USA) and treated with GW4869 (MCE, USA). GW group: SHP-HR, supplemented with GW4869 (MCE, USA). RS-USCs-Exo group: SHP-HR, supplemented with 100μL RS-USC-Exo solution (20 μg/mL). SS-USCs-Exo group: SHP-HR, supplemented with 100 μL SS-USC-Exo solution (20 μg/mL). SS-USCsNCshRNA-Exo group: SHP-HR, supplemented with 100 μL SS-USCsNCshRNA-Exo solution (20 μg/mL). SS-USCsshGPX4#1-Exo group: SHP-HR, supplemented with 100 μL SS-USCsshGPX4#1-Exo solution (20 μg/mL). SS-USCs-Exo + RSL3 group: SHP-HR, supplemented with 100 μL SS-USC-Exo solution (20 μg/mL) and RSL3. SS-USCs-Exo + ML210 group: SHP-HR, supplemented with 100 μL SS-USC-Exo solution (20 μg/mL) and ML210.

ALT/AST

Automated biochemical analyzers utilizing enzymatic protocols quantified the serum levels of alanine aminotransferase (ALT) and aspartate aminotransferase (AST).

Oil red O staining

Deparaffinization of paraffin-embedded sections involved sequential steps starting with xylene immersion, followed by alcohol-based gradient dehydration, and culminating with a water rinse. Conversely, fixation of frozen sections employed 60% isopropanol. The process for Oil Red O staining required the sections to be immersed in the staining solution for 10–15 min, followed by a wash in 60% isopropanol and a final rinse in water.

TUNEL staining

Initial sample preparation involves deparaffinization using xylene followed by a graded series of ethanol washes (100% 5 min, 90% 2 min, 70% 2 min) and a brief rinse in water for 2 min. Subsequently, the tissue is treated with DNase-free proteinase K for a duration ranging from 15 to 30 min. Following treatment, samples are rinsed thrice using PBS. For the apoptosis detection, 50 μL of the detection solution from the One Step TUNEL Apoptosis Assay Kit (Beyotime Biotechnology) is applied to each sample, which is then incubated at 37 °C for 60 min. Post-incubation, the samples are washed and prepared for microscopic examination.

Immunohistochemical staining

For antigen retrieval, tissues undergo a heating process before being subjected to a 25-min incubation with 3% hydrogen peroxide to quench endogenous peroxidase activity. Blocking of non-specific binding sites is achieved by incubating the tissue with 3% bovine serum albumin (BSA) for 30 min. Overnight incubation with the primary antibody is conducted at 4 °C, followed by a subsequent 1-h incubation with the secondary antibody at room temperature.

CCK-8

The viability of cells was evaluated using the CCK-8 Cell Proliferation and Cytotoxicity Assay Kit (Solarbio, China). Initially, cells were plated in 96-well plates. Following this, CCK-8 reagent was administered and the cells were incubated for three hours at 37 °C. The optical density was then measured at 450 nm to ascertain cell viability.

Flow cytometry

Cell death was quantified utilizing the Annexin V-PE Apoptosis Detection Kit (Beyotime Biotechnology, China). All cells, inclusive of detached dead cells, were harvested and resuspended in 195 μL of Annexin V-PE Binding Buffer. Subsequently, 5 μL of Annexin V-PE was added and the mixture was incubated in darkness at room temperature (20–25 °C) for 10–20 min. Flow cytometry was employed to determine the proportion of PE-positive cells, providing an estimate of cell mortality.

Iron assay/malondialdehyde (MDA) assay

Intracellular and hepatic levels of Fe2 + were measured using an iron assay kit (Abcam, USA), with absorbance recorded at 593 nm on a microplate reader. Similarly, MDA levels in cells and liver tissues were assessed with an MDA assay kit (Beyotime Biotechnology, China), where absorbance was determined at 532 nm using a microplate reader.

C11-BODIPY

To assess lipid reactive oxygen species, cells were harvested and incubated in a culture medium supplemented with 5μM BODIPY 581/591 C11 at ambient temperature for a duration of 30 min. Subsequently, the cells were rinsed twice using phosphate-buffered saline before flow cytometric analysis was conducted.

Western blotting

For cellular and exosomal lysis, samples were treated with RIPA buffer and phenylmethylsulfonyl fluoride on ice. In parallel, liver tissues were processed by adding the same lysis agents, then homogenized and lysed under chilled conditions. Protein concentrations in the lysates were determined using a BCA assay. Following this, 30 µg of each protein sample underwent SDS-PAGE, after which proteins were electrotransferred to a PVDF membrane. The membrane was blocked using a 5% solution of skim milk at room temperature for one hour. Primary antibodies, after being diluted, were applied to the membrane and incubated overnight at 4 °C. Afterward, the membrane was washed thrice with TBST, each lasting 30 min. A horseradish peroxidase-conjugated secondary antibody was then applied at a dilution of 1:2000 and left at room temperature for one hour, followed by three additional 30-min washes in TBST. Proteins were developed using BeyoECL Plus (ECL like Western reagent; Beyotime Biotechnology, China). Quantification of bands was performed using Image Lab 4.1 and ImageJ software. Antibodies: GPX4 (67,763–1-Ig, 1:1000, Proteintech); CD9 (EXOAB-KIT-1, 1:1000, SBI); CD63 (EXOAB-KIT-1, 1:1000, SBI); CD81 (EXOAB-KIT-1, 1:1000, SBI); HSP70 (EXOAB-KIT-1, 1:1000, SBI); and β-actin (81,115-1-RR, 1:10,000, Proteintech).

Isolation, identification, and uptake of exosomes

Culture medium supernatant underwent a series of centrifugations: initially at low speed (300 g for 10 min), followed by medium speed (2000 g for 20 min), then high speed (10,000 g for 30 min), and finally two sequential ultracentrifugation steps (100,000 g, 70 min each) [22]. The pellet containing exosomes was resuspended in Phosphate-Buffered Saline (PBS) post these steps. The prepared exosomes were utilized immediately for experimental analysis or preserved at − 80 °C for future experiments.

The morphology and size distribution of the exosomes were documented using Transmission Electron Microscopy (TEM, Kabuskiki Kaisha, JEM 2100F, Japan). A nanoparticle analyzer (Brookhaven, OMNI, USA) was employed to evaluate the size distribution and concentration. Detection of exosome-specific proteins such as CD9, CD63, CD81, and HSP70 was conducted using an ExoAb Antibody Kit (SBI, Japan).

For visualization studies, exosomes were tagged with a specific volume of CM-DiI working solution, and cell nuclei were stained with DAPI. The internalization and localization of these marked exosomes within target cells were subsequently examined through confocal microscopy using equipment from Carl Zeiss AG (LSM 900 & Axio Imager M2, Germany).

4D label-free proteomics

Proteomic analysis without labels was executed at Jingjie PTM BioLab in China on exosome specimens.

Each sample underwent sonication for a duration of three minutes and was subsequently centrifuged for ten minutes at 12,000 g and a temperature of 4 °C. A BCA kit was employed to ascertain the protein concentration. Protein reduction was achieved using 5 mM DTT, followed by alkylation with 11 mM iodoacetamide, and digestion with trypsin. Following digestion, peptides were purified using a C18 column, dissolved in solvent A, and then introduced onto a reversed-phase analytical column. MS/MS data acquisition and processing were facilitated by Proteome Discoverer (v2.4.1.15). Bioinformatics techniques were applied to analyze proteins that exhibited differential expression between SS-USC-Exos and RS-USC-Exos.

RT-qPCR

In the RT-qPCR procedure, total RNA from hepatocytes was isolated utilizing TRIzol reagent from Invitrogen, adhering to the supplied instructions. This RNA was then converted into cDNA using the GoScript Reverse Transcription Mix from Promega, USA. Detection of this cDNA was carried out employing the GoTaq qPCR Master Mix from Promega, USA. The PCR regimen consisted of an initial denaturation at 95 °C for two minutes, followed by forty cycles each comprising denaturation at 95 °C for twenty seconds, annealing at 60 °C for thirty seconds, and elongation at 72 °C for thirty seconds. The primers for GPX4 (human) were as follows: GPX4 (human) forward, GCCCCACTATTTCTAGCTCCAC; reverse, TGTCTGTTTATTCCCACAAGGTAGC. GAPDH (human): forward, GGTATGACAACGAATTTGGC; reverse, GAGCACAGGGTACTTTATTG.

Inhibition of GPX4

The GPX4shRNA lentivirus was provided by GeneChem (Shanghai, China). Cell transfection was performed according to manufacturer’s instructions. Briefly, the required amount of virus was calculated based on the virus titer and Multiplicity of Infection (MOI), which represents the average number of virus particles infecting each cell. Then, 40 μL of HitransG A transfection enhancer was added to the culture medium. The medium was changed 12–16 h after viral infection. After 72 h of infection, 2.5 μg/ml puromycin (Sigma, USA) was added for selection. The sequences were as follows: sh-GPX4#1, GGATGAAGATCCAACCCAA; sh-GPX4#2, GAGGCAAGACCGAAGTAAA; sh-GPX4#3, GGGAGTAACGAAGAGATCA; and negative control (NC) shRNA, TTCTCCGAACGTGTCACGTAA. The GPX4 overexpression vector is GV341, with the element sequence: Ubi-MCS-3FLAG-SV40-puromycin. The restriction enzyme sites are NheI and AgeI. This plasmid carries a 3xFLAG fusion protein. The expression of the target gene is verified using Western blot analysis.

Statistical analysis

Data are presented as mean plus or minus the standard deviation (SD). To evaluate differences between two distinct groups, an unpaired Student’s t test was applied. Conversely, a one-way analysis of variance (ANOVA) was employed for assessing variations among three or more groups. These statistical analyses were conducted using GraphPad Prism software. Statistically significant differences were acknowledged when the P value was less than 0.05.

Results

Comparison of the morphology between SS-USCs and RS-USCs

From sixty urine samples collected from ten healthy individuals, two morphologically distinct types of USCs were identified: SS-USCs and RS-USCs. Flow cytometry analysis revealed that both SS-USCs and RS-USCs expressed common MSC surface markers, including CD29, CD44, and CD73 (Fig. S1A, B). Despite repeated culturing, the morphological traits of each subtype persisted unaltered, as illustrated in Fig. S1C. A comparative analysis of dimensions between these subtypes demonstrated that the major axis of SS-USCs was considerably longer at 84.9 ± 18.6 μm compared to 41.6 ± 8.9 μm for RS-USCs, with a statistically significant difference (P < 0.05). Conversely, the minor axes of both subtypes were statistically indistinguishable, measuring 13.4 ± 4.6 μm for SS-USCs and 14.9 ± 3.2 μm for RS-USCs (P > 0.05), as shown in Fig. S1D. The statistical significance of these disparities was confirmed (P < 0.01).

Characterization of heterogeneity in SS-USCs and RS-USCs populations

Bulk RNA-Seq of P3 generation SS-USCs and RS-USCs extracted from three healthy individuals was performed (Fig. 1A). The results revealed that SS-USCs were enriched in key biological pathways related to mesenchymal stem cell development, stem cell differentiation, developmental growth involved in morphogenesis, regulation of wound healing, and apoptotic cell clearance, indicating similarities to mesenchymal stem cells and strong tissue repair capabilities (Fig. 1B, C, and Table 1). RS-USCs were primarily enriched in pathways associated with renal and urogenital system development, proximal/distal pattern formation, loop of Henle development, embryonic skeletal system development, and axonogenesis, suggesting their potential for differentiation and development in the kidney, nervous system, and skeletal system (Fig. 1B, C and Table 1).

Fig. 1
figure 1

Characterization of Heterogeneity in SS-USCs and RS-USCs. A Experimental design and analysis of extracting USCs from human urine samples. B Heatmap displaying the differentially expressed genes (DEGs) between SS-USCs and RS-USCs groups (n = 3) based on Bulk RNA-seq data. C GO analysis of DEGs from Bulk RNA-seq, showing enrichment in 11 biological processes. D Score analysis based on characteristic gene sets of SS-USCs/RS-USCs, dividing them into three major clusters (C1, C2, and C3). E scRNA-seq heatmap: Top 10 DEGs in subgroups C1/C3. F scRNA-seq Bar plots performing GO and KEGG analysis on DEGs, showing enrichment in 10 biological processes for SS-USCs and RS-USCs respectively

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SS-USCs and RS-USCs were isolated from three healthy individuals and subjected to scRNA-Seq analysis after mixing them (Fig. 1A). A total of 33,559 cells passed the quality control standards (Fig. S2A, B, D), and after filtering, the USCs were classified into 14 distinct clusters (Fig. S2C).

By scoring these 14 clusters based on the characteristic gene sets of SS-USCs/RS-USCs, we classified them into three main clusters: high expression (C1), moderate expression (C2), and low expression (C3) (Fig. 1D and Table 2). The C1 cluster corresponds to SS-USCs, whereas the C3 cluster corresponds to RS-USCs. Differentially expressed genes (DEGs) were detected among the three clusters. Analysis of DEGs revealed that the ten most significant DEGs in clusters C1 and C3 effectively discriminated these subgroups as shown in Fig. 1E and Table 3. In contrast, cluster C2 exhibited no notable overexpression of genes.

Subsequent exploration into the surface markers of USC identified variable expression levels of CD24, CD29, CD44, CD73, CD90, CD105, CD133, Vimentin (VIM), and Collagen I across the three clusters. The results indicated no significant differences in the presence of any marker among the three clusters. However, from C1 to C3, a slight increase in the expression difference of CD24 was observed (Fig. S2E).

GSEA revealed that both clusters C1 and C3 exhibited tendencies toward osteogenic, adipogenic, and chondrogenic differentiation (Fig. S3A). Subsequent predictions of developmental fate were based on the levels of mRNA for osteogenesis markers IGFBP3 and MCAM, adipogenesis markers FABP5 and MEST, and chondrogenesis markers DCN and LUM. The results showed that both clusters C1 and C3 exhibited tendencies toward osteogenic, adipogenic, and chondrogenic differentiation, with no significant differences in differentiation ability (Fig. S3B). Regarding cell stemness, compared to the C1 subgroup, the C3 subgroup was enriched in the expression of stemness markers (such as BIRC5, CCNA2, TUBA1B, CDK1, CCNB1, and E2F1), indicating a stronger capability for self-renewal and maintenance of stem cell status in the C3 cluster (Fig. S3C).

Further analysis for enrichment of GO and KEGG pathways indicated enrichment of the C1 cluster in pathways including glutathione metabolism, oxidative stress response, zinc ion response, pathways regulating longevity across multiple species, cell growth, cellular senescence, epidermal growth factor stimulus response, wound healing, and apoptosis as shown in Fig. 1F. Conversely, the C3 cluster showed enrichment in pathways associated with regeneration, organ regeneration in animals, positive cell cycle regulation, chromosome segregation, mitotic nuclear division, endoplasmic reticulum protein processing, pyrimidine metabolism, DNA replication, mineral absorption, and cell cycle processes, also illustrated in Fig. 1F.

SS-USCs alleviate severe hepatic IRI and SHP-HR by inhibiting ferroptosis

In steatotic HL7702 cells, exposure to HR treatment markedly elevated Lipid ROS and MDA levels, as depicted in Fig. S4A, B. Subsequent investigations demonstrated that both HR treatment and Erastin, a ferroptosis inducer, considerably diminished the viability of these cells. Conversely, Fer-1, a ferroptosis inhibitor, notably enhanced cell viability, as shown in Fig. S4C. The influence of inhibitors for pyroptosis (Z-VAD-FMK) and apoptosis (Necrostatin-1) on cell viability was minimal. Additionally, a significant escalation in cell mortality was observed following treatments with HR and erastin, which was substantially mitigated by Fer-1 administration (Fig. S4E). Similarly, HR treatment induced a notable increase in the intracellular concentrations of Lipid ROS, MDA, and Fe2 + , all of which were significantly reduced following Fer-1 treatment, suggesting a decline in these ferroptosis indicators (Fig. S4D, F, G).

Steatotic livers exhibited extensive necrosis following transplantation or IRI, accompanied by an increase in Suzuki score (Fig. S5A, E). Markers associated with ferroptosis, such as elevated levels of Fe2 + and MDA, were significantly increased (Fig. S5B, C and S5F, G), whereas GPX4 levels were notably decreased (Fig. S5D, H). Application of Fer-1 significantly reduced widespread necrosis in liver tissue, decreased the levels of ferroptosis markers, such as Fe2 + and MDA, and increased GPX4 content.

S-HL7702 cells were subjected to HR treatment and intervention was carried out by co-culturing with SS-USCs or RS-USCs (Fig. 2A, B). The addition of SS-USCs effectively reduced cell death (Fig. 2C); restored cell viability (Fig. 2D); decreased Lipid ROS, MDA, and Fe2 + levels; and increased GPX4 expression (Fig. 2E–H). In contrast, co-culture with RS-USCs did not suppress ferroptosis or alleviate SHP-HR.

Fig. 2
figure 2

SS-USCs inhibit ferroptosis to alleviate SHP-HR. A Diagram illustrating the SHP-HR experimental design. B Oil Red O staining showing lipid degeneration in HL7702 cells subjected to the S-HL7702 treatment. C Flow cytometry analysis comparing cell death rates among different groups (Control, HR, RS-USCs, and SS-USCs), with quantification (n = 3). D Cell viability assessed by the CCK-8 assay (n = 3). E Lipid-ROS levels measured using C11-BODIPY staining (n = 3). F GPX4 levels in each group, normalized to β-actin (n = 3). G, H Measurements of Fe2 + and MDA levels in cells across all groups (n = 3), *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

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Subsequently, we established a severe fatty liver transplantation (LT) rat model (Fig. 3A, C) and a severe fatty liver IRI mouse model (Fig. 3B, D). Liver damage was compared between the sham, PBS, RS-USCs, and SS-USCs groups. The SS-USCs treatment group showed significantly alleviated hepatic tissue necrosis (Fig. 3E, F) and reduced serum transaminase levels (Fig. 3G, H). Additionally, the application of SS-USCs markedly decreased the Fe2 + and MDA levels (Fig. 3I–L). However, RS-USCs did not exhibit similar inhibitory effects.

Fig. 3
figure 3

SS-USCs reduce ferroptosis, alleviating IRI in rat and mouse livers with severe steatosis. A Experimental protocol for creating a rat model of severe fatty liver transplantation. B Experimental protocol for the mouse model of severe fatty liver IRI. C, D Oil Red O staining of severe fatty liver tissues in both rats and mice. E, F H&E and TUNEL staining of liver tissues from different treatment groups (Sham, PBS, RS-USCs, and SS-USCs), with necrotic regions highlighted (n = 5). G, H Measurement of serum ALT and AST levels in both rat and mouse groups (n = 5). (I, J) Quantification of Fe2⁺ and MDA levels in rat liver tissues (n = 5). (K, L) Quantification of Fe2 + and MDA levels in mouse liver tissues (n = 5). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

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SS-USCs alleviate severe fatty liver IRI and SHP-HR by inhibiting ferroptosis through exosomes

SS-USCs were treated with the exosome inhibitor GW4869 (GW). After co-culture, the experimental results revealed that, compared to using SS-USCs alone or GW alone, treatment with GW + SS-USCs significantly reduced cell viability (Fig. 4A) and protective capacity against cell damage (Fig. 4F). This treatment also increased Lipid ROS levels (Fig. 4B) and Fe2 + content (Fig. 4J). This confirmed that exosomes serve as the primary pathway through which SS-USCs exert their biological functions.

Fig. 4
figure 4

SS-USCs inhibit ferroptosis to alleviate SHP-HR through the utilization of Exosomes. A Flow cytometry analysis of cell death rates; the exosome inhibitor GW4869 significantly attenuates the protective effect of SS-USCs on cell damage (n = 3). B Elevated levels of Lipid ROS after treatment of SS-USCs with GW4869. C Transmission electron microscopy image showing exosomes isolated from USCs conditioned medium (scale bar = 200 nm).D Analysis of size distribution of USCs-Exo, revealing an average diameter of 102.4 ± 9.8 nm. E Western blot validation of exosome markers CD9, CD63, CD81, and HSP70, with conditioned medium (CM) as the control group. F Cell viability measured by CCK-8 assay (n = 3). G Fluorescence images showing S-HL7702 cells uptake CM-DiI-labeled (red) exosomes. H Flow cytometry analysis of cell death rates across different treatment groups (HR, RS-USCs-Exo, SS-USCs-Exo) (n = 3). I CCK-8 assay measuring cell viability (n = 3). J Elevated Fe2⁺ levels after treating SS-USCs with GW4869. K Lipid ROS levels (C11-BODIPY) detection in various treatment groups (HR, RS-USCs-Exo, SS-USCs-Exo) (n = 3). L, M Measurement of MDA and Fe2 + levels in cells from each group (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

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Extracted exosome secreted by USCs: Through transmission electron microscopy observation, we confirmed that these exosomes exhibited typical double-layered lipid bilayer membrane structures (Fig. 4C). Particle size analysis revealed that the average diameter of these exosomes was 102.4 ± 9.8 nm (Fig. 4D). Furthermore, Western blotting detected exosome-associated protein markers, such as CD9, CD63, CD81, and HSP70 (Fig. 4E). Using confocal microscopy, we observed the distribution of exosomes labeled with CM-DiI (red) within HL7702 cells (Fig. 4G). These exosomes were primarily localized in the cytoplasm.

RS-USC-Exo and SS-USC-Exo were separately added to S-HL7702 cells after HR treatment. The results showed that compared to RS-USC-Exo treatment, SS-USC-Exo treatment not only significantly improved cell viability (Fig. 4I), but also markedly reduced the rate of cell death (Fig. 4H). In the studies, SS-USC-Exo demonstrated a notable reduction in the increase of lipid ROS, MDA, and Fe2 + concentrations when contrasted with RS-USC-Exo (Fig. 4K–M). The inhibitory response by RS-USC-Exos was comparatively less potent.

Investigating the impact on severe fatty liver models in rats (LT) and mice (IRI), treatment with SS-USC-Exo proved more efficacious than RS-USC-Exo in mitigating liver tissue necrosis (Fig. 5A, B) and lowering serum ALT and AST levels (Fig. 5C, D). Moreover, this treatment notably decreased Fe2 + and MDA concentrations within the liver (refer to Fig. 5E–H). The levels of GPX4 protein in the liver were also significantly elevated following SS-USC-Exo administration (Fig. 5I–L).

Fig. 5
figure 5

SS-USCs inhibit ferroptosis by utilizing exosomes in rat donor liver with severe steatosis and mouse liver with severe steatosis. A, B H&E and TUNEL staining of liver tissues from various treatment groups (Sham, PBS, RS-USCs-Exo, SS-USCs-Exo) in both rat and mouse models, highlighting necrotic areas (n = 5). C, D Measurement of serum ALT and AST levels in rat and mouse subjects (n = 5). E, F Quantification of Fe2 + and MDA levels in liver tissues of rat groups (n = 5). G, H Analysis of Fe2 + and MDA levels in mouse liver tissues (n = 5). I, J GPX4 levels in liver tissues from rats and mice, normalized to β-actin (n = 3). K, L Immunohistochemical staining to display GPX4 levels across treatment groups (n = 5). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

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Protein expression profiles of SS-USCs-exo and RS-USCs-exo

Through 4D label-free quantitative proteomic analysis, the proteome landscapes of SS-USC-Exos and RS-USC-Exos were delineated. Collectively, proteomic profiling resulted in the identification of 1751 protein groups across both exosome types (Fig. 6A). The comparative analysis highlighted that 317 proteins were significantly elevated in SS-USC-Exos, while 631 showed reduced levels when contrasted with RS-USC-Exos (Figs. 6B, S6A, and Table 3). Subsequent biological examination of these differentially expressed proteins revealed a notable association with pathways like apoptosis, intracellular sequestration, and the regulation of iron ions within cells (Fig. 6C). Among the proteins analyzed, GPX4 stood out with the most significant differential in expression between SS-USC-Exos and RS-USC-Exos (Fig. 6D). Public datasets were used to compute the expression ratios of iron death-related proteins between SS-USC-Exo and SS-USC, underscoring a substantial prevalence of GPX4 in SS-USC-Exo (Fig. S6B) [15]. This was corroborated by Western blot analysis, aligning with the findings from the proteomic evaluation (Fig. 6E).

Fig. 6
figure 6

SS-USCs-Exo alleviates SHP-HR by inhibiting ferroptosis through GPX4. A Venn diagram of protein expression profiles of SS-USCs-Exo and RS-USCs-Exo. B Volcano plot illustrating the proteins with increased and decreased expression in SS-USCs-Exo and RS-USCs-Exo. C Bar plots showing KEGG and GO enrichment analyses of differentially expressed proteins. D Top 10 proteins with elevated expression in SS-USCs-Exo and RS-USCs-Exo. E Western blot analysis validating GPX4 protein levels in SS-USCs-Exo and RS-USCs-Exo (n = 3). F qRT-PCR analysis confirming the inhibitory effect of shRNA targeting GPX4 (n = 3). G Western blot detecting the impact of shGPX4#1 silencing on GPX4 protein in SS-USC and SS-USCs-Exo (n = 3). H Flow cytometry analysis of cell death rates in various treatment groups (HR, SS-USCsNCshRNA-Exo, SS-USCsshGPX4#1-Exo) (n = 3). I, J Measurement of Lipid ROS levels (C11-BODIPY) in each group (n = 3). K MDA levels in cells of each group (n = 3). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001

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SS-USC-exos alleviate SHP-HR by delivering GPX4 protein

We used shRNAs, namely shGPX4#1, shGPX4#2, and shGPX4#3, to target and downregulate GPX4 expression in SS-USCs. Among these, shGPX4#1 exhibited the strongest inhibitory effect (Fig. 6F). Western blotting confirmed that GPX4 expression was significantly downregulated in GPX4-silenced USCs (SS-USCsshGPX4#1) and in their secreted Exos (SS-USCsshGPX4#1-Exo) (Fig. 6G). After silencing GPX4 in SS-USCs, the ability of SS-USCsshGPX4#1-Exo to inhibit cell death significantly decreased (Fig. 6H). Elevations in lipid ROS and MDA concentrations were noted, demonstrating that the reduction in GPX4 diminishes the suppressive impacts of SS-USC-Exo on ferroptosis (Fig. 6I–K). We employed two inhibitors specific to GPX4, namely RSL3 and ML210. When SS-USCs-Exo was administered in conjunction with either RSL3 or ML210, a marked increase in the rate of cell mortality was observed (Fig. S6C). Furthermore, there was a significant rise in the levels of lipid ROS and MDA (Fig. S6D–E). Overexpression of GPX4 in RS-USCs was found to enhance the ability of RS-USC-Exo to inhibit ferroptosis; however, its ferroptosis-inhibitory effect remained weaker compared to that of SS-USC-Exo (Fig. S6F–I).

Discussion

Stem cells have been widely applied in tissue engineering and regenerative medicine and have shown promising therapeutic effects in various diseases[23,24,25,26]. However, despite the favorable outcomes observed in preclinical animal models, most clinical trials have not achieved the anticipated therapeutic results. Thus, stem cell heterogeneity has become a crucial issue.

USCs are among the most recently discovered stem cells[6]. Previous studies have reported morphologically distinct USC types, indicating that they are not homogeneous cell populations. In our culture experiments, we observed two major morphological types of USCs: spindle- and rice-shaped. Importantly, even after several culture passages, the morphological characteristics of the two USCs subpopulations remained stable and showed no significant changes, as visualized in Fig. S1B. The stability of these morphological features provides novel perspectives for exploring the diversity and complexity of USCs.

RNA-Seq analysis of SS-USCs and RS-USCs revealed the enrichment of pathways involved in mesenchymal cell differentiation, regulation of wound healing, and apoptotic cell clearance. Conversely, RS-USCs were predominantly enriched in biological pathways, such as renal system development, embryonic skeletal system development, and axonogenesis. Previous studies have confirmed the angiogenic and wound healing effects of USCs[15]. Bharadwaj et al.[27] illustrated that under controlled in vitro environments, USCs have the capacity to transform into various bladder-specific cell types. These include cells from both the urothelial and the smooth muscle lineages, exhibiting functional characteristics. This suggests that SS-USCs play an important role in tissue repair. However, RS-USCs exhibit inherent potential for differentiation toward specific directions, such as the kidney, nervous system, and skeletal system, with significant promise for renal system reconstruction.

Furthermore, we constructed a comprehensive single-cell transcriptomic atlas of human USCs. Based on scoring analysis using the characteristic gene sets of SS-USCs/RS-USCs, they were classified into three major clusters: C1, C2, and C3. C1 cluster corresponds to SS-USCs, whereas C3 cluster corresponds to RS-USCs. Analysis of traditional CD markers revealed a limited ability to distinguish between C1–C3 clusters. However, in contrast to previous studies[16], we observed a slight increase in CD24 expression from C1 to C3. To further explore the differentiation potential differences between the C1 and C3 clusters, we performed GSEA. The results revealed that both clusters C1 and C3 exhibited osteogenic, adipogenic, and chondrogenic differentiation capabilities. The prediction of developmental outcomes was conducted by analyzing mRNA levels for markers indicative of osteogenesis (IGFBP3, MCAM), adipogenesis (FABP5, MEST), and chondrogenesis (DCN, LUM). The findings indicated that both clusters C1 and C3 showed tendencies toward osteogenic, adipogenic, and chondrogenic differentiation, with no significant differences in differentiation capacity, which is consistent with previous research. Chen et al.have indicated that SS-USCs exhibit significantly enhanced potential for osteogenic and adipogenic differentiation. In contrast, RS-USCs are more proficient in chondrogenic differentiation, a variance that may stem from differences in the culture conditions employed. In terms of stemness, compared to the C1 cluster, the C3 cluster showed enrichment of stemness markers (such as BIRC5, CCNA2, TUBA1B, CDK1, CCNB1, and E2F1), indicating a stronger ability of the C3 cluster to self-renew and maintain stem cell status.

Rao et al.[28] confirmed that USCs exert aging effects through PLAU and TIMP1 proteins. Chen et al.[15] demonstrated that USCs enhance angiogenesis and aid in healing diabetic wounds. Our investigations reveal that Cluster 1 displays an enrichment in several oxidative stress response pathways, such as glutathione metabolism, reactions to oxidative stress, and cellular responses to zinc ions. Activation of these pathways may be associated with the ability of C1 cluster cells to resist oxidative stress and maintain cellular stability. Additionally, C1 cluster is enriched in pathways, such as longevity regulating pathway-multiple species, cell growth, cellular senescence, cellular response to epidermal growth factor stimulus, wound healing, and apoptosis, indicating that C1 cluster may play important roles in anti-aging and tissue regeneration. In contrast, the C3 cluster was mainly enriched in pathways related to the positive regulation of cell cycle processes, chromosome segregation, mitotic nuclear division, DNA replication, and cell cycle. The enrichment of these pathways suggests that the C3 cluster is in a relatively quiescent state of stemness with strong proliferation and self-renewal capabilities. These analyses revealed the similarity between SS-USCs and MSCs, and their robust tissue repair functions. In contrast, RS-USCs possess the potential to differentiate and develop into the kidney, nervous, and skeletal systems.

In both in vitro and in vivo models, the use of SS-USCs markedly mitigated the effects of reperfusion injury when compared with RS-USCs. This improvement can likely be linked to the various biological roles of USCs, such as their anti-inflammatory and antioxidant properties[29, 30]. Non-alcoholic fatty liver disease (NAFLD) is widely recognized as a chronic liver ailment marked by an excessive fat buildup within liver cells, frequently coinciding with disruptions in iron homeostasis[31]. Research indicates that blocking ferroptosis can curtail the advancement of NAFLD and hepatic ischemia–reperfusion injury[21, 32, 33]. Characterized as a type of regulated cell death dependent on iron and driven by lipid peroxidation, ferroptosis plays a pivotal role in IRI[34, 35]. Studies conducted in vitro and in vivo reveal that treatment with the ferroptosis inhibitor Fer-1 notably enhances liver functionality, diminishes liver pathology, augments cell survival, reduces the rate of cell death related to injury, and decreases markers associated with ferroptosis. Meanwhile, agents that inhibit pyroptosis, like Z-VAD-FMK, and apoptosis, such as Necrostatin-1, exhibited minimal impact on cellular survival. These findings underscore the significant impact of ferroptosis on cell damage induced by hepatic reperfusion. Subsequent studies revealed that SS-USCs significantly suppressed ferroptosis.

To determine whether exosomes are the key mechanisms by which SS-USCs exert protective effects in the SHP-HR model, SS-USCs were treated with the exosome inhibitor GW4869. The experimental results showed that GW weakened the inhibition of ferroptosis and alleviated cell damage induced by SS-USCs. These findings emphasize that exosomes are the main pathway through which SS-USCs exert their biological functions in the SHP-HR model.

Exosome-based cell-free therapies serve as a promising avenue in tissue engineering due to their non-immunogenic and non-tumorigenic nature, high stability, effective delivery to wound areas, and their ability to avoid vascular blockages[36,37,38,39]. USCs also secrete exosomes that have potential therapeutic effects in various aspects. Previous studies confirmed that USC-Exos can be used to treat renal ischemia–reperfusion injury, promote vascularization and wound healing in diabetic wounds, and exert anti-aging effects. Experimental results from both in vitro and in vivo studies revealed a crucial role for exosomes derived from SS-USCs in mitigating ferroptosis, in contrast to the comparatively weaker effects observed with RS-USC-Exos.

Advanced 4D label-free quantitative proteomic analysis identified distinct protein expression profiles in SS- and RS-USC-Exos, with the former showing higher levels of proteins related to apoptosis, intracellular iron ion trapping, and the maintenance of iron ion equilibrium. Additionally, this analysis pointed to a marked increase in GPX4 in SS-USC-Exo when compared to both RS-USC-Exo and SS-USCs themselves. Validation through Western blot techniques affirmed the GPX4 enrichment in SS-USC-Exos.

GPX4 is a unique antioxidant enzyme belonging to the glutathione peroxidase (GPX) family, which utilizes glutathione (GSH) as a substrate to reduce peroxides, thereby protecting cells from oxidative damage[40]. Furthermore, GPX4 exists in several forms within the cell, including the cytoplasmic, mitochondrial, and nuclear forms, allowing it to function in multiple cellular compartments. GPX4 inhibits ferroptosis by directly reducing lipid peroxides, making it a key antioxidant enzyme in preventing ferroptotic cell death[41, 42]. Decreased activity or inhibition of GPX4 expression increases the cellular sensitivity to ferroptosis. The role of GPX4 extends beyond ROS scavenging to maintaining cell membrane integrity and stability[35, 43, 44]. In vitro experiments have demonstrated that reducing the expression of GPX4 in SS-USC-Exos markedly lessens its protective capabilities. To further validate the role of GPX4 in preventing ferroptosis, the GPX4 inhibitors RSL3 and ML210 were applied to selectively inhibit GPX4 function in cells. The protective efficacy of SS-USC-Exos was notably diminished with the addition of RSL3 or ML210[45, 46].

From this, it can be inferred that SS-USC-Exos primarily protect cells from ferroptosis by delivering GPX4 to directly reduce lipid peroxides. Inhibition of GPX4 did not completely eliminate the effects of SS-USC-Exos on SHP-HR. Additionally, overexpression of GPX4 in RS-USCs enhanced the ferroptosis-inhibitory effect of RS-USC-Exo; however, this effect remained weaker than that of SS-USC-Exo. These findings suggest that other signaling molecules are also involved in these mechanisms. This observation aligns with our proteomic findings, which revealed that SS-USC-Exos are involved in pathways related to wound healing, apoptosis, cell growth, and cellular senescence. Furthermore, SS-USC-Exos are abundant in other ferroptosis-associated proteins, including LPCAT3[47], GSS[48], FTL[49], and PCBP2[50]. Consequently, the suppression of ferroptosis and the reduction of ischemia–reperfusion injury-induced fatty liver by SS-USC-Exos are facilitated by the delivery of multiple functional molecules, rather than a single protein entity.

Conclusion

SS-USCs exhibit robust tissue repair and antioxidant properties. In contrast, RS-USCs are capable of differentiating into specific lineages, including the kidney, nervous system, and skeletal system, indicating significant potential for renal system reconstruction. Furthermore, both in vivo and in vitro experiments verified that SS-USCs and their exosomes substantially inhibited ferroptosis and mitigated severe fatty liver ischemia–reperfusion injury, while RS-USCs and their exosomes exhibited comparatively less effectiveness. Analysis comparing the proteomic differences between SS-USC-Exos and RS-USC-Exos revealed that SS-USC-Exos primarily inhibited ferroptosis and improved cellular viability by secreting exosomes containing GPX4. This highlights SS-USCs as the most suitable cell subtype for treating severe fatty liver ischemia–reperfusion injury. The role of SS-USC-Exos in suppressing ferroptosis in fatty livers has been elucidated. This discovery not only provides a novel perspective on the role of urine-derived stem cells in tissue repair and regeneration but also identifies critical molecular targets for developing stem cell exosome-based therapeutic strategies in the future.

Data availability

The raw sequence data reported in this paper have been deposited in the Genome Sequence Archive (Genomics, Proteomics and Bioinformatics 2021) in National Genomics Data Center (Nucleic Acids Res 2022), China National Center for Bioinformation/Beijing Institute of Genomics, Chinese Academy of Sciences (GSA-Human: HRA008742) that are publicly accessible at https://ngdc.cncb.ac.cn/gsa-human.

Abbreviations

IRI:

Ischemia-reperfusion injury

USCs:

Urine-derived stem cells

BMSCs:

Bone marrow mesenchymal stem cells

UCBSCs:

Umbilical cord blood stem cells

AFSCs:

Amniotic fluid-derived stem cells

SS-USCs:

Spindle-shaped USCs

RS-USCs:

Rise-shaped USCs

SHP-HR:

Steatotic hepatocyte hypoxia-reoxygenation

SHP-HR:

Hepatocyte hypoxia-reoxygenation

ALT:

Serum alanine aminotransferase

AST:

Aspartate aminotransferase

MDA:

Malondialdehyde

DEGs:

Differentially expressed genes

VIM:

Vimentin

GO:

Gene ontology

KEGG:

Kyoto encyclopedia of genes and genomes

GPX4:

Glutathione peroxidase 4

ROS:

Reactive oxygen species

GSH:

Glutathione

LT:

Liver transplantation

Exo:

Exosome

NAFLD:

Non-alcoholic fatty liver disease

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Acknowledgements

The authors declare that they have not use AI-generated work in this manuscript.

Funding

This research was funded by the National Natural Science Foundation of China, grant number 82370667 and 82272973.

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Author notes

  1. Shangheng Shi, Cunle Zhu and Shangxuan Shi contributed equally to this work.

Authors and Affiliations

  1. Organ Transplantation Center, Affiliated Hospital of Qingdao University, Qingdao, China

    Shangheng Shi, Xinqiang Li, Imran Muhammad, Qingguo Xu, Xinwei Li, Ziyin Zhao, Huan Liu, Guangming Fu, Xijian Huang, Feng Wang & Jinzhen Cai

  2. Institute of Transplantation Science, Affiliated Hospital of Qingdao University, Qingdao, China

    Shangheng Shi, Xinqiang Li, Imran Muhammad, Qingguo Xu, Xinwei Li, Ziyin Zhao, Huan Liu, Xijian Huang, Feng Wang & Jinzhen Cai

  3. Organ Transplant Center, Fujian Medical University Union Hospital, Fuzhou, China

    Cunle Zhu & Jinzhen Cai

  4. Shanghai Institute for Advanced Immunochemical Studies, Shanghai Tech University, Shanghai, China

    Shangxuan Shi

  5. Department of Immunology, Medical College of Qingdao University, Qingdao, China

    Meiying Song

Authors
  1. Shangheng Shi
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Contributions

Conceptualization, S-H.S., F.W. and J-Z.C.; methodology, S-H.S.; software, S-X.S. and X-Q.L.; validation, S-H.S., C-L.Z., and X-W.L.; formal analysis, Q-G.X.; investigation, Z-Y.Z.; resources, H.L.; data curation, G-M.F.; writing—original draft preparation, S-H.S.; writing—review and editing, S-H.S. and M.I.; visualization, X-J.H., and M-Y.S; supervision, F.W. and J-Z.C.; project administration, F.W. and J-Z.C.; funding acquisition, J-Z.C. All authors have read and agreed to the published version of the manuscript.

Corresponding authors

Correspondence to Feng Wang or Jinzhen Cai.

Ethics declarations

Ethics approval and consent to participate

All human urine samples were obtained with informed donor consent and under the approval of IRB of Affiliated Hospital of Qingdao University, Qingdao, China (Title of an ethical approved project, “Human Spindle-shaped Urine-derived Stem Cell Exosomes Alleviate Severe Fatty Liver Ischemia–Reperfusion Injury by Inhibiting Ferroptosis via GPX”, Approval number, KY2018-036-02 and date of approval, July 5, 2021). All experimental protocols were approved and carried out in accordance with the relevant guidelines and regulations of the Ethics Committee of Qingdao University, China (Title of an ethical approved project, “Exosomes Derived from Spindle-Shaped USCs Inhibit Fatty Liver Ischemia–Reperfusion Injury by Enhancing GPX4”, Approval number, 20210310C57BL4220211010061; 20210310SD4220211010062, and date of approval, March 1, 2021). All investigations were conducted according to the principles expressed in the Declaration of Helsinki.

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Shi, S., Zhu, C., Shi, S. et al. Human spindle-shaped urine-derived stem cell exosomes alleviate severe fatty liver ischemia–reperfusion injury by inhibiting ferroptosis via GPX4. Stem Cell Res Ther 16, 81 (2025). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s13287-025-04202-y

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Keywords

Stem Cell Research & Therapy

ISSN: 1757-6512