OTUD1-mediated deubiquitination stabilizes SLC7A11 to suppress ferroptosis in hepatocellular carcinoma

Abstract

Background

Evasion of ferroptosis has been increasingly recognized as an important mechanism underlying lenvatinib resistance in hepatocellular carcinoma (HCC). However, the roles of deubiquitinating enzymes in ferroptosis regulation and the post-translational control of key ferroptosis regulators in HCC remain poorly understood.

Methods

Public lenvatinib resistance-related and ferroptosis-related datasets were analyzed to identify candidate deubiquitinating enzymes associated with HCC therapeutic resistance and ferroptotic stress. Loss- and gain-of-function assays were performed to evaluate the effects of OTUD1 on lenvatinib sensitivity, erastin-induced ferroptosis, and HCC cell growth. Subcutaneous xenograft and orthotopic tumor models were used to assess the role of OTUD1 in tumor growth and ferroptosis resistance in vivo. Western blotting, RT-qPCR, co-immunoprecipitation, cycloheximide chase, MG132 rescue, ubiquitination, SLC7A11 rescue, and ERK1/2 pathway modulation assays were conducted to investigate the underlying mechanism. Clinical HCC specimens and public datasets were analyzed to determine OTUD1 expression and its association with patient prognosis.

Results

OTUD1 was identified as a gene associated with lenvatinib resistance in HCC. Knockdown of OTUD1 sensitized HCC cells to lenvatinib, and this effect was abolished by ferroptosis inhibition. OTUD1 knockdown enhanced ferroptosis in HCC cells, whereas OTUD1 overexpression suppressed it. Moreover, OTUD1 promoted HCC growth and conferred ferroptosis resistance in mice. Mechanistically, OTUD1 interacted with solute carrier family 7 member 11 (SLC7A11) and removed its K48-linked polyubiquitin chains, thereby stabilizing SLC7A11 protein levels. In addition, ERK1/2-dependent phosphorylation of OTUD1 at Ser216 strengthened its interaction with SLC7A11, further enhancing SLC7A11 stability and promoting ferroptosis resistance. Analysis of clinical samples and public datasets further revealed that OTUD1 was upregulated in HCC tissues and correlated with poor prognosis.

Conclusions

Our findings identify the ERK1/2–OTUD1–SLC7A11 axis as a key mechanism mediating ferroptosis resistance in HCC. These results suggest that OTUD1 may serve as a potential therapeutic target for HCC.

Supplementary Information

The online version contains supplementary material available at https://doi.org/10.1007/s13402-026-01282-7.

Background

Hepatocellular carcinoma (HCC) accounts for the majority of primary liver cancers and remains one of the most common and lethal malignancies worldwide [1, 2]. Although systemic therapy has advanced substantially, the prognosis of patients with advanced HCC remains unsatisfactory. Frequent recurrence, intrahepatic dissemination, and therapeutic resistance are major contributors to poor outcomes [1–3]. Lenvatinib is a multi-kinase inhibitor approved as a first-line treatment for advanced HCC. However, intrinsic and acquired resistance frequently limit its long-term efficacy, despite its significant clinical benefit [4–6].

Ferroptosis is an iron-dependent form of regulated cell death driven by excessive lipid peroxidation [7, 8]. Increasing evidence shows that ferroptosis is closely involved in HCC progression and therapeutic response, and induction of ferroptosis has emerged as a promising strategy for HCC treatment [8–12]. Notably, ferroptosis contributes to the anti-tumor activity of lenvatinib, whereas ferroptosis suppression has been linked to lenvatinib resistance [13, 14]. Recent studies further showed that restoring ferroptosis can enhance lenvatinib sensitivity in HCC [15–18]. These findings suggest that molecular regulators of ferroptosis resistance may also contribute to therapeutic resistance.

The regulation of ferroptosis in HCC involves complex metabolic and signaling networks, with post-translational modification emerging as an important layer of regulation [9–12, 19, 20]. Ubiquitination is a key post-translational modification that controls protein stability and signaling output, whereas deubiquitinating enzymes (DUBs) counteract ubiquitin-mediated protein degradation by removing ubiquitin chains from substrate proteins [21–23]. Accumulating evidence suggests that DUBs contribute to tumor progression and multiple forms of regulated cell death, yet their roles in ferroptosis regulation in HCC remain incompletely understood [19–25].

Among DUB family members, OTUD1 has been implicated in oxidative stress and inflammatory signaling. Emerging evidence further suggests that OTUD1 is functionally linked to ferroptosis-related pathways. In tissue injury models, OTUD1 has been reported to suppress ferroptosis through NRF2-related [26] or AMPK/GSK3β/β-catenin-related mechanisms [27], whereas in other settings it can also enhance iron-dependent oxidative stress by stabilizing IREB2 and promoting TFRC-mediated iron transport [28]. These observations suggest that OTUD1 plays a context-dependent role in ferroptosis-associated processes. However, whether OTUD1 contributes to ferroptosis resistance in HCC and the underlying molecular mechanism remain unknown. In our preliminary analyses, OTUD1 was found to be elevated in lenvatinib-resistant HCC cell models and was subsequently identified as a candidate DUB potentially involved in ferroptosis-related processes.

In this study, we investigated the role of OTUD1 in lenvatinib resistance and ferroptosis in HCC, and explored the underlying molecular mechanisms. We found that OTUD1 promoted lenvatinib resistance in HCC cells by enhancing ferroptosis resistance. Mechanistically, ERK1/2-mediated phosphorylation of OTUD1 facilitated its binding to SLC7A11, leading to deubiquitination and stabilization of SLC7A11, thereby suppressing ferroptosis. Analysis of clinical samples revealed that OTUD1 was highly expressed in HCC and correlated with poor prognosis. These findings provide novel insights into the molecular mechanisms driving lenvatinib resistance in HCC and suggest OTUD1 as a potential therapeutic target.

Methods

Patients and clinical specimens

Sixteen pairs of fresh-frozen HCC tissues and matched adjacent non-tumor tissues, as well as 50 paraffin-embedded specimens containing both tumor and adjacent non-tumor tissues, were collected from patients with HCC who underwent hepatectomy at the First Affiliated Hospital of Xi’an Jiaotong University (Xi’an, China). The fresh-frozen tissues were used for Western blotting (WB) and RT-qPCR, whereas the paraffin-embedded specimens were used for immunohistochemistry (IHC). All patients included in this study were pathologically diagnosed with HCC and had received no preoperative anticancer therapy. Written informed consent was obtained from all patients before sample collection.

Cell culture

The human HCC cell lines Huh7, HCCLM3, HepG2, Hep3B, MHCC97H, MHCC97L, and human embryonic kidney HEK293T cells were purchased from the National Collection of Authenticated Cell Cultures (Shanghai, China). The SNU449 cell line was kindly provided by Dr. Yilei Zhang’s laboratory (Xi’an Jiaotong University Health Science Center, Xi’an, China). The human normal hepatocyte cell line (THLE-2) was purchased from Shanghai Jinyuan Biotechnology Co., Ltd. (JY658, Shanghai, China) and cultured in THLE-2 cell culture medium (Shanghai Jinyuan Biotechnology Co., Ltd.). All cell lines were authenticated by short tandem repeat (STR) profiling every 6 months. HCC and HEK293T cells were cultured in DMEM (11965092, Gibco) supplemented with 10% fetal bovine serum (FBS; 1027 − 106, Gibco; Thermo Fisher Scientific, Waltham, MA, USA) and 1% penicillin-streptomycin (15070063, Gibco) at 37 °C in a humidified incubator with 5% CO₂.

Plasmids, antibodies and reagents

Plasmids and antibodies used in this study are listed in Supplementary Tables 1 and 2, respectively. Z-VAD-FMK (S7023), Nec-1s (S8641), Erastin (S7242), and Ferrostatin-1 (Fer-1; S7243) were purchased from Selleck Chemicals. 3-Methyladenine (3-MA; HY-19312), MG132 (HY-13259), and Cycloheximide (CHX; HY-12320) were purchased from MedChemExpress (MCE).

Cell transfection and lentivirus packaging

Plasmids were transfected into cells using Lipo8000 transfection reagent (C0533, Beyotime Biotechnology, Shanghai, China) according to the manufacturer’s instructions. For lentivirus packaging, 5 × 10⁶ HEK293T cells were seeded in a 10-cm dish. The indicated lentiviral vector (6 µg), psPAX2 (4 µg), and pMD2.G (2 µg) were co-transfected into HEK293T cells using polyethylenimine (PEI). After 6 h, the medium was replaced with 10 mL of fresh culture medium. Viral supernatants were collected 48 h after transfection, filtered through 0.45-µm filters, and then added to HCC cells in the presence of 8 µg/mL polybrene (Beyotime Biotechnology).

Immunohistochemistry (IHC)

Paraffin-embedded tissue sections were deparaffinized in xylene and rehydrated through a graded ethanol series. Antigen retrieval was performed in citrate buffer by microwave heating. IHC staining was performed according to the manufacturer’s instructions using immunohistochemistry kits (PV-90001 and PV-90002, ZSGB-BIO, Beijing, China). Target proteins were visualized using a DAB substrate kit (ZLI-9018, ZSGB-BIO). Hematoxylin was used for nuclear counterstaining. The staining intensity was scored as 0, negative; 1, weak; 2, moderate; or 3, strong. The percentage of positive cells was scored as 0, negative; 1, 1–25%; 2, 26–50%; 3, 51–75%; or 4, 76–100%. The final IHC score was calculated by multiplying the staining intensity score by the percentage score.

RNA extraction and RT-qPCR

Total RNA was extracted from cultured cells or tissues using TRIzol reagent (15596018, Invitrogen, USA) according to the manufacturer’s instructions. Complementary DNA (cDNA) was synthesized from 1 µg total RNA using a reverse transcription kit (K1622, Thermo Fisher Scientific) according to the manufacturer’s protocol. RT-qPCR was then performed using 2× Universal SYBR Green Fast qPCR Mix (RK21203, ABclonal, Woburn, MA, USA). Primers were synthesized by Tsingke Biotechnology (Beijing, China), and their sequences are listed in Supplementary Table 3.

Western blot analysis

Cells were lysed in RIPA buffer (P0013B, Beyotime, Shanghai, China) supplemented with protease and phosphatase inhibitor cocktails. Protein concentrations were determined using a BCA protein assay kit (P0012, Beyotime). Equal amounts of protein, typically 20–40 µg, were separated by SDS-PAGE and transferred onto PVDF membranes (IPVH00010, Millipore, USA). After blocking with 5% non-fat milk in TBST for 1 h at room temperature, the membranes were incubated overnight at 4 °C with primary antibodies. The membranes were then incubated with HRP-conjugated secondary antibodies for 1 h at room temperature. Protein bands were visualized using enhanced chemiluminescence (ECL) reagent (WBLUF0500, Millipore) and imaged with an Amersham Imager 680 (GE Healthcare Life Sciences, Pittsburgh, PA, USA). Band intensities were quantified using ImageJ software (National Institutes of Health, USA) and normalized to vinculin or β-actin.

Immunoprecipitation and ubiquitination assay

Cells were lysed in IP lysis buffer (P0013, Beyotime, Shanghai, China) supplemented with protease and phosphatase inhibitors. Co-immunoprecipitation was performed with Dynabeads™ Protein G Immunoprecipitation Kit (10007D, Invitrogen; Thermo Fisher Scientific) according to the manufacture’s protocols. Briefly, the indicated antibody was incubated with magnetic beads in antibody-binding buffer in 1.5-mL microcentrifuge tubes for 1 h with rotation. The supernatant was then removed. Cell lysates containing 500 µg of total protein were incubated with the antibody-bound beads at 4 ℃ overnight with rotation. The beads were washed 3 times with washing buffer and boiled in SDS loading buffer. Immunoprecipitated proteins were subjected to western blot analysis.

For ubiquitination assays, cells were transfected with the indicated plasmids and treated with the proteasome inhibitor MG132 for 6 h before harvest where indicated. Cell lysates were immunoprecipitated with anti-SLC7A11 or anti-HA antibodies, followed by immunoblotting with antibodies against ubiquitin chains.

Cell viability assay

Cell viability was determined using a Cell Counting Kit-8 (CCK-8) according to the manufacturer’s instructions. Briefly, cells were seeded into 96-well plates at a density of 5 × 10³ cells per well. Cells were then treated with the indicated drugs for the indicated times. Subsequently, 10 µL of CCK-8 reagent was added to each well and incubated for 1 h at 37 °C. Absorbance at 450 nm was measured using a microplate reader.

Colony formation assay

For the colony formation assay, 1,000 cells from each group were seeded into 6-well plates and cultured in 4 mL of complete medium. After 14 days, colonies were fixed with 4% paraformaldehyde and stained with crystal violet. Colony numbers were then counted and analyzed.

Xenograft tumor and orthotopic HCC models

Male BALB/c nude mice were housed under specific pathogen-free conditions in the Laboratory Animal Center of Xi’an Jiaotong University and randomly assigned to the indicated groups.

For the subcutaneous xenograft model, HCC cells with the indicated genetic modifications were harvested and resuspended in PBS. A total of 5 × 10⁶ cells were subcutaneously injected into the flank of each mouse. Erastin was administered by intraperitoneal injection at a dose of 15 mg/kg every other day. Tumor size was measured every 3 days using a digital caliper, and tumor volume was calculated using the following formula: volume = (length × width²)/2. After 24 days, the mice were sacrificed, and xenograft tumors were excised, weighed, photographed, and processed for subsequent analyses, including immunohistochemistry and malondialdehyde (MDA) assay.

For the orthotopic HCC model, male BALB/c nude mice aged 6–8 weeks were anesthetized, and the liver lobe was exposed through a small upper abdominal incision. Hep3B cells suspended in PBS were injected into the liver parenchyma. The mice were monitored daily after surgery and treated as indicated. At the experimental endpoint, orthotopic tumors were harvested for gross observation, tumor burden assessment, immunohistochemistry, and MDA assay.

Determination of iron, malondialdehyde (MDA), and glutathione (GSH) levels

After the indicated treatments, HCC cells, subcutaneous xenograft tumors, or orthotopic tumor tissues were collected for biochemical analyses. Fresh tissue samples were weighed and homogenized in PBS or normal saline, whereas cell samples were prepared according to the requirements of each assay kit. Intracellular iron, MDA, and GSH levels were measured using an iron assay kit (A039-2-1, Nanjing Jiancheng Bioengineering Institute, Nanjing, China), an MDA assay kit (A003-4-1, Nanjing Jiancheng Bioengineering Institute), and a GSH assay kit (A006-2-1, Nanjing Jiancheng Bioengineering Institute), respectively, according to the manufacturers’ instructions.

Measurement of intracellular ROS and lipid peroxidation

Intracellular ROS levels were detected using a Reactive Oxygen Species Assay Kit containing the DCFH-DA probe (S0033M, Beyotime, China). Lipid peroxidation was assessed using BODIPY™ 581/591 C11 (D3861, Invitrogen, USA). Briefly, cells were treated as indicated and washed twice with PBS. The probes were diluted to working concentrations according to the manufacturers’ instructions and incubated with cells at 37 °C in the dark for 20–30 min. After incubation, the cells were harvested, resuspended in PBS, and immediately analyzed using a NovoCyte flow cytometer. Data were analyzed using FlowJo software.

Statistical analysis

All experiments were repeated at least three times unless otherwise stated. Data are presented as the mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism 9.5. The normality of quantitative data was assessed using the Shapiro–Wilk test. For normally distributed data, comparisons between two groups were performed using Student’s t-test, whereas comparisons among multiple groups were performed using one-way or two-way ANOVA, as appropriate. For non-normally distributed data, the Mann–Whitney U test or Kruskal–Wallis test was used, as appropriate. For paired clinical samples, paired Student’s t-test or Wilcoxon matched-pairs signed-rank test was used according to data distribution. Survival curves were generated using the Kaplan–Meier method and compared using the log-rank test. Correlation analyses were performed using Pearson or Spearman correlation analysis, as appropriate. P < 0.05 was considered statistically significant. ns, not significant; *P < 0.05; **P < 0.01; ***P < 0.001.

Results

OTUD1 confers ferroptosis resistance in HCC cells

First, we analyzed the expression profiles of deubiquitinating enzymes in lenvatinib resistance-related datasets GSE186191 and GSE211850. Among these candidates, OTUD1 showed consistently elevated expression in lenvatinib-resistant HCC cell lines (Fig. S1A), suggesting a potential role for OTUD1 in therapeutic resistance. We next examined OTUD1 expression in the normal liver cell line THLE-2 and a panel of HCC cell lines, including HepG2, Hep3B, SNU449, MHCC97H, HCCLM3, Huh7, and MHCC97L. OTUD1 expression was higher in most HCC cell lines, including HepG2, Hep3B, MHCC97H, Huh7, and MHCC97L, than in THLE-2 cells (Fig. S1B). We further performed colony formation assays under lenvatinib treatment. OTUD1 knockdown reduced clonogenic survival in lenvatinib-treated HCC cells, whereas OTUD1 overexpression increased clonogenic survival (Fig. S1E, F). Moreover, in a lenvatinib-treated xenograft model, tumors derived from OTUD1-knockdown cells showed reduced tumor growth compared with control tumors (Fig. S1G-I). Using Hep3B and Huh7 cells, which exhibited relatively high OTUD1 expression, we further found that OTUD1 knockdown enhanced the sensitivity of HCC cells to lenvatinib. Notably, this increased sensitivity was reversed by the ferroptosis inhibitor Ferrostatin-1 (Fer-1), but not by the apoptosis inhibitor Z-VAD-FMK, the autophagy inhibitor 3-MA, or the necroptosis inhibitor Nec-1s. These results indicate that OTUD1 contributes to lenvatinib tolerance by inhibiting ferroptosis in HCC cells (Fig. S1C, D).

To further validate the association between OTUD1 and ferroptosis, we next analyzed the ferroptosis-related dataset GSE104462. In this dataset, OTUD1 mRNA expression was reduced in HepG2 cells following erastin treatment among multiple deubiquitinating enzymes (Fig. S2A). Moreover, treatment of Hep3B and Huh7 cells with increasing concentrations of erastin led to a dose-dependent decrease in both OTUD1 mRNA and protein levels (Fig. S2B, C). Together, these findings indicate that OTUD1 is closely associated with ferroptotic stress in HCC cells.

We next investigated whether OTUD1 affects ferroptosis-related phenotypes in HCC cells. The knockdown efficiency of OTUD1 in Hep3B cells and the overexpression efficiency of OTUD1 in HCCLM3 cells were confirmed by western blotting before functional analyses (Fig. 1A, I). OTUD1 knockdown markedly increased lipid peroxidation and intracellular ROS accumulation under erastin treatment (Fig. 1B, C). Consistently, OTUD1 depletion decreased GSH levels and increased intracellular Fe²⁺ and MDA accumulation (Fig. 1D-F), accompanied by reduced cell viability and clonogenic survival under ferroptotic stress (Fig. 1G, H). In contrast, OTUD1 overexpression significantly reduced lipid peroxidation and intracellular ROS accumulation (Fig. 1J, K), increased GSH levels, and decreased MDA and intracellular Fe²⁺ levels (Fig. 1L-N), thereby enhancing cell viability and clonogenic survival (Fig. 1O, P). Importantly, these effects were largely reversed by Fer-1 treatment. Collectively, these results demonstrate that OTUD1 confers ferroptosis resistance in HCC cells.

Fig. 1.

OTUD1 confers ferroptosis resistance in HCC cells. (A, I) Western blot analysis showing OTUD1 knockdown efficiency in Hep3B cells and OTUD1 overexpression efficiency in HCCLM3 cells. (B) Analysis of lipid peroxidation in OTUD1-knockdown HCC cells. Lipid peroxidation was quantified using the oxidized/total C11-BODIPY fluorescence ratio, calculated as FITC MFI/ (FITC MFI + PE MFI), and normalized to the control group. (C) Flow cytometric analysis of intracellular ROS levels in OTUD1-knockdown HCC cells. (D–F) Detection of GSH, intracellular Fe²⁺, and MDA levels in OTUD1-knockdown HCC cells. (G, H) CCK-8 and colony formation assays of OTUD1-knockdown HCC cells. (J) Analysis of lipid peroxidation in OTUD1-overexpressing HCC cells. Lipid peroxidation was quantified using the oxidized/total C11-BODIPY fluorescence ratio, calculated as FITC MFI/ (FITC MFI + PE MFI), and normalized to the control group. (K) Flow cytometric analysis of intracellular ROS levels in OTUD1-overexpressing HCC cells. (L–N) Detection of GSH, intracellular Fe²⁺, and MDA levels in OTUD1-overexpressing HCC cells. (O, P) CCK-8 and colony formation assays of OTUD1-overexpressing HCC cells. Cells were treated with DMSO, Erastin, or Erastin plus Ferrostatin-1 (Fer-1), as indicated. Data are presented as the mean ± SD. Statistical significance was determined by Student’s t-test or two-way ANOVA, as appropriate. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001

OTUD1 promotes ferroptosis resistance and tumor growth in vivo

To further validate the role of OTUD1 in ferroptosis regulation and HCC progression in vivo, we established subcutaneous xenograft models by inoculating nude mice with Hep3B cells stably expressing shOTUD1 or HCCLM3 cells stably overexpressing OTUD1, together with their corresponding control cells, followed by intraperitoneal administration of erastin. In the Hep3B xenograft model, OTUD1 knockdown markedly inhibited tumor growth, as reflected by reduced tumor volume and tumor weight compared with the control group (Fig. 2A, B). Moreover, OTUD1 depletion was accompanied by increased MDA levels in xenograft tissues, indicating enhanced lipid peroxidation (Fig. 2C). Consistently, immunohistochemical staining for 4-hydroxynonenal (4-HNE), a representative product of lipid peroxidation, showed that 4-HNE accumulation was further enhanced in tumors with OTUD1 knockdown (Fig. 2G, H). In contrast, in the HCCLM3 xenograft model, OTUD1 overexpression significantly promoted tumor growth, as evidenced by increased tumor volume and tumor weight. (Fig. 2D, E). In parallel, MDA levels were reduced in xenograft tissues with OTUD1 overexpression, suggesting decreased lipid peroxidation (Fig. 2F). Consistently, OTUD1 overexpression markedly reduced 4-HNE staining in tumor tissues (Fig. 2I, J).

Fig. 2.

OTUD1 promotes ferroptosis resistance and tumor growth in vivo. (A, B) Tumor volume and tumor weight in the Hep3B subcutaneous xenograft model with stable OTUD1 knockdown following vehicle or erastin treatment (n = 6). (C) MDA levels in xenograft tissues from the Hep3B model. (D, E) Tumor volume and tumor weight in the HCCLM3 subcutaneous xenograft model with stable OTUD1 overexpression following vehicle or erastin treatment (n = 6). (F) MDA levels in xenograft tissues from the HCCLM3 model. (G, H) Representative images and quantification of OTUD1 and 4-hydroxynonenal (4-HNE) immunohistochemical staining in tumors from the Hep3B xenograft model. (I, J) Representative images and quantification of OTUD1 and 4-HNE immunohistochemical staining in tumors from the HCCLM3 xenograft model. (K, L) Maximum tumor diameter in the orthotopic HCC model with OTUD1 knockdown following vehicle or erastin treatment (n = 5). (M) MDA levels in orthotopic tumor tissues. (K, N) Representative images and quantification of OTUD1 and 4-HNE immunohistochemical staining in orthotopic tumors. Data are presented as the mean ± SD. Statistical significance was determined by two-way ANOVA. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001

To further confirm these findings in a more physiologically relevant setting, we established an orthotopic HCC model in nude mice. Consistent with the results obtained in the subcutaneous xenograft model, OTUD1 knockdown markedly suppressed orthotopic tumor growth, as reflected by a reduction in maximum tumor diameter (Fig. 2K, L). In addition, MDA levels were increased in orthotopic tumor tissues after OTUD1 knockdown (Fig. 2M). Consistently, immunohistochemical staining showed enhanced 4-HNE accumulation in tumors with OTUD1 depletion (Fig. 2K, N). Collectively, these findings indicate that OTUD1 promotes tumor growth and suppresses ferroptosis in vivo.

OTUD1 interacts with and positively regulates SLC7A11 in HCC cells

To investigate the molecular mechanism by which OTUD1 suppresses ferroptosis in HCC cells, we first examined whether OTUD1 affects key ferroptosis-related proteins. In Hep3B and Huh7 cells, OTUD1 knockdown markedly reduced the protein level of SLC7A11 and also decreased GPX4 expression, although the change in GPX4 was less pronounced than that in SLC7A11. By contrast, the protein levels of FTH1 and ACSL4 were not obviously altered after OTUD1 depletion (Fig. 3A; Fig. S3A). Consistently, ectopic expression of OTUD1 in HCCLM3 and SNU449 cells significantly increased the protein abundance of SLC7A11 and, to a lesser extent, GPX4, whereas FTH1 and ACSL4 remained largely unchanged. Notably, the catalytically inactive mutant OTUD1-C320S failed to increase the expression of SLC7A11 or GPX4 (Fig. 3B; Fig. S3A). These findings suggest that SLC7A11 may represent the ferroptosis-related protein most consistently regulated by OTUD1.

Fig. 3.

OTUD1 interacts with and positively regulates SLC7A11 in HCC cells. (A) Western blot analysis of SLC7A11 expression in Hep3B and Huh7 cells with OTUD1 knockdown. (B) Western blot analysis of SLC7A11 expression in HCCLM3 and SNU449 cells expressing OTUD1 or the catalytically inactive mutant OTUD1-C320S. (C, D) Endogenous co-immunoprecipitation assays of OTUD1 and SLC7A11 in Hep3B and Huh7 cells. (E, F) Exogenous co-immunoprecipitation assays of FLAG-OTUD1 and HA-SLC7A11 in HEK293T cells. Data are presented as the mean ± SD. Statistical significance was determined by one-way ANOVA. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001

Given the central role of SLC7A11 in ferroptosis regulation, we next examined whether OTUD1 regulates SLC7A11 expression at the post-translational level. As shown above, OTUD1 modulation markedly affected the protein abundance of SLC7A11, but had little effect on its mRNA level (Fig. S3B). Endogenous co-immunoprecipitation assays confirmed the interaction between OTUD1 and SLC7A11 in Hep3B and Huh7 cells (Fig. 3C, D). This interaction was further validated by exogenous co-IP in HEK293T cells co-transfected with FLAG-OTUD1 and HA-SLC7A11 (Fig. 3E, F). Together, these results indicate that OTUD1 interacts with and positively regulates SLC7A11 at the post-translational level in HCC cells.

OTUD1 stabilizes SLC7A11 by removing K48-linked polyubiquitin chains

Given that OTUD1 is a deubiquitinating enzyme, we next investigated whether it regulates SLC7A11 stability through the ubiquitin–proteasome pathway. In Hep3B and Huh7 cells, MG132 largely rescued the reduction in SLC7A11 protein levels induced by OTUD1 knockdown (Fig. 4A, B; Fig. S4A, B), indicating that OTUD1 protects SLC7A11 from proteasomal degradation. CHX chase assays further showed that OTUD1 depletion accelerated SLC7A11 degradation and shortened its half-life in Hep3B and Huh7 cells (Fig. 4C; Fig. S4C), whereas OTUD1 overexpression prolonged SLC7A11 stability in HCCLM3 and SNU449 cells. In contrast, the catalytically inactive mutant OTUD1-C320S failed to stabilize SLC7A11 (Fig. 4D; Fig. S4D). Ubiquitination assays in HEK293T cells showed that OTUD1 knockdown increased, whereas OTUD1 overexpression decreased, SLC7A11 ubiquitination; OTUD1-C320S had no obvious effect (Fig. 4E, F). Endogenous ubiquitination assays further confirmed these findings (Fig. S4E, F). Moreover, OTUD1 selectively removed K48-linked, but not K63-linked or K33-linked, polyubiquitin chains from SLC7A11 (Fig. 4G, Fig. S4G). To further identify the lysine residues involved in OTUD1-regulated K48-linked ubiquitination of SLC7A11, we selected six potential ubiquitination-related lysine residues of SLC7A11, including K4, K12, K30, K37, K222, and K483, based on annotations from the PhosphoSitePlus database. Site-specific lysine mutants of SLC7A11 were then generated and subjected to ubiquitination assays. Among these mutants, mutation of K37 most markedly attenuated the ability of OTUD1 to reduce K48-linked polyubiquitination of SLC7A11, whereas mutation of K30 also partially affected this process (Fig. S4H). These results suggest that K37 is a major lysine residue involved in OTUD1-regulated K48-linked ubiquitination of SLC7A11, with K30 playing a contributing role. Together, these results demonstrate that OTUD1 stabilizes SLC7A11 by removing K48-linked polyubiquitin chains and thereby limiting its proteasomal degradation.

Fig. 4.

OTUD1 stabilizes SLC7A11 by removing K48-linked polyubiquitin chains. (A, B) Western blot analysis of SLC7A11 expression in OTUD1-knockdown Huh7 and Hep3B cells treated with or without MG132. (C) Cycloheximide (CHX) chase assay of SLC7A11 protein stability in OTUD1-knockdown Hep3B cells. (D) CHX chase assay of SLC7A11 protein stability in HCCLM3 cells expressing OTUD1 or the catalytically inactive mutant OTUD1-C320S. (E, F) Ubiquitination assays of SLC7A11 in HEK293T cells with OTUD1 knockdown, OTUD1 overexpression, or OTUD1-C320S expression. (G) Ubiquitination assays of K48-linked and K63-linked polyubiquitin chains on SLC7A11 in the presence of OTUD1. Data are presented as the mean ± SD. Statistical significance was determined by two-way ANOVA. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001

SLC7A11 mediates the ferroptosis-suppressive effect of OTUD1

To determine whether SLC7A11 is functionally required for OTUD1-mediated ferroptosis resistance, we performed rescue experiments in HCC cells under erastin treatment. In OTUD1-overexpressing cells, silencing SLC7A11 significantly restored ferroptosis-associated phenotypes, as evidenced by increased lipid peroxidation (Fig. 5A), elevated intracellular ROS, Fe²⁺, and MDA levels, and reduced GSH content (Fig. 5B–E). In parallel, knockdown of SLC7A11 largely abolished the pro-survival effect of OTUD1 overexpression under ferroptotic stress, as shown by decreased cell viability and clonogenic survival in CCK-8 and colony formation assays (Fig. 5F; Fig. S5A). These results indicate that SLC7A11 is required for the ferroptosis-suppressive effect of OTUD1.

Fig. 5.

SLC7A11 mediates the ferroptosis-suppressive effect of OTUD1 in HCC cells and xenografts. (A) Analysis of lipid peroxidation in HCC cells from the control, OTUD1-overexpression, SLC7A11-knockdown, and OTUD1-overexpression plus SLC7A11-knockdown groups following erastin treatment. Lipid peroxidation was quantified using the oxidized/total C11-BODIPY fluorescence ratio, calculated as FITC MFI/ (FITC MFI + PE MFI), and normalized to the control group. (B–E) Detection of intracellular ROS, Fe²⁺, MDA, and GSH levels in HCC cells from the control, OTUD1-overexpression, SLC7A11-knockdown, and OTUD1-overexpression plus SLC7A11-knockdown groups following erastin treatment. (F) CCK-8 assay of HCC cells from the control, OTUD1-overexpression, SLC7A11-knockdown, and OTUD1-overexpression plus SLC7A11-knockdown groups following erastin treatment. (G–K) Detection of lipid peroxidation, intracellular ROS, Fe²⁺, MDA, and GSH levels in HCC cells from the control, OTUD1-knockdown, SLC7A11-overexpression, and OTUD1-knockdown plus SLC7A11-overexpression groups following erastin treatment. (L) CCK-8 assay of HCC cells from the control, OTUD1-knockdown, SLC7A11-overexpression, and OTUD1-knockdown plus SLC7A11-overexpression groups following erastin treatment. (M, N) Tumor volume and tumor weight in erastin-treated xenografts from the control, OTUD1-overexpression, and OTUD1-overexpression plus SLC7A11-knockdown groups. (O) MDA levels in xenograft tissues following erastin treatment. (P, Q) Representative images and quantification of OTUD1, SLC7A11 and 4-hydroxynonenal (4-HNE) immunohistochemical staining in xenograft tumors following erastin treatment. Data are presented as the mean ± SD. Statistical significance was determined by one-way/two-way ANOVA, as appropriate. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001

Conversely, we examined whether reintroduction of SLC7A11 could rescue the enhanced ferroptosis sensitivity caused by OTUD1 depletion under erastin treatment. As expected, ectopic expression of SLC7A11 markedly attenuated the ferroptotic phenotypes induced by OTUD1 knockdown, including lipid peroxidation, ROS accumulation, Fe²⁺ overload, MDA elevation, and GSH depletion (Fig. 5G–K). Consistently, SLC7A11 overexpression also partially restored cell viability and clonogenic capacity in OTUD1-silenced cells under ferroptotic stress (Fig. 5L; Fig. S5B). Together, these findings demonstrate that SLC7A11 is a critical downstream effector mediating OTUD1-induced ferroptosis resistance in HCC cells.

We next validated this functional relationship in vivo using a xenograft tumor model under erastin treatment. OTUD1 overexpression significantly promoted tumor growth, whereas simultaneous knockdown of SLC7A11 markedly attenuated this effect, as reflected by reduced tumor volume and tumor weight (Fig. 5M, N). Consistent with the in vitro results, biochemical analysis of xenograft tissues showed that SLC7A11 knockdown reversed the decrease in MDA levels caused by OTUD1 overexpression (Fig. 5O). Moreover, immunohistochemical staining for 4-HNE revealed that OTUD1 overexpression reduced lipid peroxidation in tumor tissues, whereas this effect was largely reversed by SLC7A11 depletion (Fig. 5P, Q). Collectively, these results indicate that SLC7A11 is functionally required for OTUD1-mediated ferroptosis resistance and tumor-promoting effects in HCC. However, SLC7A11 knockdown did not completely abolish the ferroptosis-suppressive effect of OTUD1 overexpression, indicating that additional OTUD1-regulated ferroptosis-related factors may also contribute to this phenotype.

In addition, to determine whether SLC7A11 mediates the effect of OTUD1 on lenvatinib response, we performed rescue experiments under lenvatinib treatment. In Hep3B and Huh7 cells, OTUD1 knockdown reduced clonogenic survival under lenvatinib treatment, whereas re-expression of SLC7A11 markedly restored colony formation (Fig. S6A). Similarly, CCK-8 assays showed that SLC7A11 restoration attenuated the reduction in cell viability caused by OTUD1 knockdown following lenvatinib treatment (Fig. S6C). Conversely, in HCCLM3 and SNU449 cells, OTUD1 overexpression increased clonogenic survival and cell viability under lenvatinib treatment, whereas SLC7A11 knockdown weakened these effects (Fig. S6B, D). Together, these results indicate that SLC7A11 is an important downstream effector involved in OTUD1-associated lenvatinib tolerance in HCC cells.

ERK1/2-mediated phosphorylation of OTUD1 enhances SLC7A11 stabilization and ferroptosis resistance

To explore the upstream mechanism regulating OTUD1 activity, we used the PhosphoSitePlus (https://www.phosphosite.org/) database and identified Ser216 as a potential ERK1/2-mediated phosphorylation site in OTUD1. To verify whether ERK1/2 regulates OTUD1 phosphorylation, HCC cells were treated with the ERK1/2 inhibitor SCH772984 or the ERK1/2 activator EGF. Immunoprecipitation assays showed that inhibition of ERK1/2 markedly reduced the serine phosphorylation level of OTUD1, whereas EGF treatment significantly increased OTUD1 serine phosphorylation (Fig. 6A, B). To further validate this phosphorylation site, we generated phosphorylation-deficient (OTUD1-S216A) and phosphomimetic (OTUD1-S216D) OTUD1 mutants. Under ERK1/2 activation conditions, the serine phosphorylation signal associated with OTUD1 was maintained in wild-type OTUD1 and was further enhanced in the S216D mutant, whereas it was markedly diminished in the S216A mutant (Fig. 6C), indicating that Ser216 is a critical site for ERK1/2-mediated phosphorylation of OTUD1. To further determine whether ERK1/2 directly phosphorylates OTUD1, we performed an in vitro kinase assay using recombinant active ERK1/2 and purified OTUD1 proteins. Immunoblotting with an anti-phosphoserine antibody showed that active ERK1/2 induced a strong serine phosphorylation signal on OTUD1-WT in the presence of ATP. In contrast, this phosphorylation signal was markedly reduced in the absence of ATP and was also decreased in the OTUD1-S216A mutant group (Fig. S7A). These results provide direct in vitro evidence that ERK1/2 phosphorylates OTUD1 and further support Ser216 as an important ERK1/2-responsive phosphorylation site.

Fig. 6.

ERK1/2-mediated phosphorylation of OTUD1 enhances SLC7A11 stabilization. (A, B) Immunoprecipitation analysis of OTUD1 serine phosphorylation in HCC cells treated with the ERK1/2 inhibitor SCH772984 or the ERK1/2 activator epidermal growth factor (EGF). (C) Immunoprecipitation analysis of serine phosphorylation in wild-type OTUD1 and the S216A/S216D mutants under ERK1/2 activation conditions. (D–F) Co-immunoprecipitation assays of OTUD1 and SLC7A11 following ERK1/2 inhibition or activation, and in cells expressing wild-type OTUD1 or the S216A/S216D mutants. (G) Western blot analysis of SLC7A11 expression in cells expressing wild-type OTUD1 or the S216A/S216D mutants. (H) Cycloheximide (CHX) chase assay of SLC7A11 protein stability in cells expressing wild-type OTUD1 or the S216A/S216D mutants. (I) Ubiquitination assays of SLC7A11 in cells expressing wild-type OTUD1 or the S216A/S216D mutants under control or ERK1/2 inhibition conditions

We next investigated whether ERK1/2-mediated phosphorylation of OTUD1 affects its interaction with SLC7A11. Co-immunoprecipitation assays showed that ERK1/2 inhibition weakened the interaction between OTUD1 and SLC7A11, whereas ERK1/2 activation enhanced their binding. Consistently, the S216A mutant exhibited reduced binding to SLC7A11, while the S216D mutant showed enhanced interaction with SLC7A11 compared with wild-type OTUD1 (Fig. 6D–F). These findings suggest that ERK1/2-mediated phosphorylation of OTUD1 at Ser216 promotes the OTUD1–SLC7A11 interaction.

We further examined whether Ser216 phosphorylation affects OTUD1-mediated regulation of SLC7A11 stability. Compared with wild-type OTUD1, the S216A mutant reduced SLC7A11 protein abundance, whereas the S216D mutant further increased SLC7A11 expression (Fig. 6G). CHX chase assays showed that the S216A mutant shortened the half-life of SLC7A11, whereas the S216D mutant prolonged SLC7A11 stability relative to wild-type OTUD1 (Fig. 6H). In addition, ubiquitination assays demonstrated that the S216A mutant markedly impaired the ability of OTUD1 to remove ubiquitin chains from SLC7A11, whereas the S216D mutant further enhanced OTUD1-mediated deubiquitination of SLC7A11. Notably, this effect was dependent on ERK1/2 kinase activity (Fig. 6I). Together, these results indicate that ERK1/2-mediated phosphorylation at Ser216 enhances OTUD1-mediated deubiquitination of SLC7A11 and promotes SLC7A11 stabilization.

Finally, we investigated whether Ser216 phosphorylation of OTUD1 affects ferroptosis resistance. Compared with wild-type OTUD1, expression of the S216A mutant increased lipid ROS and reduced GSH content, indicating enhanced ferroptosis sensitivity. In contrast, the S216D mutant produced the opposite effects, with reduced lipid ROS and increased GSH levels (Fig. S7B, C). Consistently, CCK-8 assays showed that the S216A mutant weakened the pro-survival effect of OTUD1 under ferroptotic stress, whereas the S216D mutant further enhanced cell viability (Fig. S7D). Collectively, these findings indicate that ERK1/2-mediated phosphorylation of OTUD1 at Ser216 strengthens the OTUD1–SLC7A11 axis and promotes ferroptosis resistance in HCC cells.

OTUD1 is upregulated in HCC and associated with poor prognosis

To evaluate the expression pattern and clinical relevance of OTUD1 in HCC, we first examined its expression in our clinical samples. In 16 paired HCC and adjacent non-tumorous tissues, both the protein and mRNA levels of OTUD1 were significantly elevated in tumor tissues compared with matched non-tumorous tissues (P < 0.01; Fig. 7A–C). Consistently, immunohistochemical staining of 50 paired HCC and adjacent non-tumorous tissues further confirmed that OTUD1 was markedly upregulated in HCC tissues (P < 0.001; Fig. 7D, E). We next analyzed public datasets from the GEO database and found that OTUD1 was consistently upregulated in HCC tissues compared with non-tumorous liver tissues in GSE214846 (P < 0.001; Fig. 7F), GSE17856 (P < 0.05; Fig. 7G), and GSE31370 (P < 0.01; Fig. 7H). Analysis of TCGA data further showed that high OTUD1 expression was associated with shorter overall survival and progression-free survival in patients with HCC (Fig. 7I, J). Collectively, these results indicate that OTUD1 is aberrantly upregulated in HCC and is associated with unfavorable prognosis.

Fig. 7.

OTUD1 is upregulated in HCC and associated with poor prognosis. (A, B) Representative western blot images (A) and quantitative analysis (B) of OTUD1 protein expression in 16 paired HCC and adjacent non-tumorous tissues. (C) RT-qPCR analysis of OTUD1 mRNA expression in 16 paired HCC and adjacent non-tumorous tissues. (D, E) Representative immunohistochemical staining images (D) and quantitative analysis (E) of OTUD1 expression in 50 paired HCC and adjacent non-tumorous tissues. (F–H) OTUD1 expression analysis in HCC and non-tumorous liver tissues from the GEO datasets GSE214846 (F), GSE17856 (G), and GSE31370 (H). The datasets were analyzed using HepCAT (https://www.hccdatasph.cn/app/hepcat). (I, J) Kaplan–Meier survival analysis of overall survival (I) and progression-free survival (J) in patients with high or low OTUD1 expression based on TCGA-LIHC data. The data were analyzed using GEPIA2. (K) Schematic model of the ERK1/2-OTUD1-SLC7A11 axis in HCC. In panels B, C, and E, green markers indicate adjacent non-tumorous tissues, and red markers indicate HCC tissues. In panel E, overlapping data points and connecting lines may be present because IHC scores were recorded as discrete values. Data are presented as the mean ± SD. Paired Student’s t-test or Wilcoxon matched-pairs signed-rank test was used for paired clinical samples according to data distribution. OTUD1 IHC scores in paired HCC and adjacent non-tumorous tissues were analyzed using the Wilcoxon matched-pairs signed-rank test. Comparisons of OTUD1 expression between tumor and normal tissues in public datasets were performed using the Wilcoxon rank-sum test. Survival curves were generated using the Kaplan–Meier method and compared using the log-rank test. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001

To further evaluate the clinical relevance of the ERK1/2–OTUD1–SLC7A11 axis in HCC, we performed additional western blotting, immunohistochemical staining, correlation analysis, and clinicopathological analysis. Using the same set of tissue protein lysates, we examined SLC7A11 protein expression in 16 paired HCC and adjacent non-tumorous tissues by western blotting (Fig. S8A). The results showed that SLC7A11 expression was increased in HCC tissues compared with matched adjacent non-tumorous tissues (P < 0.01; Fig. S8B). Correlation analysis showed that OTUD1 protein levels were positively associated with SLC7A11 protein levels in HCC tissues (r = 0.5059, P = 0.0479; Fig. S8C). We next performed immunohistochemical staining for SLC7A11 in 50 paired HCC and adjacent non-tumorous tissue samples. SLC7A11 expression was higher in HCC tissues than in adjacent non-tumorous tissues (Fig. S8D, E). Moreover, SLC7A11 IHC scores were positively correlated with OTUD1 IHC scores in HCC tissues (r = 0.4258, P = 0.002; Fig. S8F). We also examined p-ERK1/2 expression by immunohistochemistry. p-ERK1/2 expression was increased in HCC tissues and was positively correlated with SLC7A11 expression in HCC tissues (r = 0.4444, P = 0.001; Fig. S8D, G, H). Kaplan–Meier analysis of these 50 patients showed that patients with high OTUD1 expression had shorter overall survival than those with low OTUD1 expression (P = 0.0219; Fig. S8I). Clinicopathological correlation analysis further showed that high OTUD1 expression was significantly associated with larger tumor size, lymph node metastasis, and advanced TNM stage (Supplementary Table 4). Together, these clinical findings further support the association of OTUD1 with SLC7A11 expression, ERK1/2 activation, and unfavorable clinical features in HCC.

Discussion

Ferroptosis is increasingly recognized as a critical determinant of therapeutic response in HCC [9, 10, 12, 29–33]. Recent studies have shown that ferroptosis is closely linked to the anti-tumor activity of lenvatinib, whereas impaired ferroptotic cell death may contribute to lenvatinib resistance in HCC [13–18, 34]. These findings highlight the importance of identifying intrinsic mechanisms that restrain ferroptosis in tumor cells. Ubiquitination has emerged as an important post-translational mechanism involved in ferroptosis regulation in HCC [35–38]. Our data add OTUD1 to this regulatory network and support a ferroptosis-suppressive role of OTUD1 in HCC cells.

SLC7A11 is a central anti-ferroptotic regulator that maintains cystine uptake, glutathione synthesis, and GPX4-dependent detoxification of lipid peroxides [39–42]. SLC7A11 has been widely implicated in tumor survival, therapeutic resistance, and malignant progression [43–57]. In recent years, deubiquitination has emerged as an important mechanism controlling SLC7A11 protein stability. Among the reported SLC7A11-regulating deubiquitinases, OTUB1 is one of the best-characterized examples. Similar to OTUD1, OTUB1 stabilizes SLC7A11 and suppresses ferroptosis by limiting SLC7A11 ubiquitination [58]. However, OTUB1 has been reported to regulate SLC7A11 through a non-canonical mechanism that is not strictly dependent on its catalytic deubiquitinase activity [58]. In contrast, our data indicate that OTUD1-mediated stabilization of SLC7A11 depends on its catalytic activity, as the catalytically inactive OTUD1-C320S mutant failed to effectively deubiquitinate and stabilize SLC7A11.

Other deubiquitinases, including OTUD5 and USP52, have also been reported to stabilize SLC7A11 and suppress ferroptosis in specific tumor contexts [59, 60]. These studies, together with ours, support the concept that SLC7A11 is tightly controlled by deubiquitination-mediated post-translational regulation. Compared with these previously reported SLC7A11-regulating deubiquitinases, an important distinction of our study is the identification of an upstream kinase-dependent mechanism that regulates OTUD1 function in HCC. The ERK signaling pathway is frequently activated in HCC and contributes to tumor growth, survival, and therapeutic resistance [61–65]. Our findings indicate that ERK1/2-dependent phosphorylation of OTUD1, with Ser216 serving as an important regulatory site, strengthens the OTUD1–SLC7A11 interaction and enhances OTUD1-mediated SLC7A11 stabilization. Thus, our findings do not simply add another deubiquitinase to the list of SLC7A11 regulators. Rather, they reveal a kinase-dependent regulatory mechanism that links oncogenic ERK1/2 signaling to OTUD1-mediated SLC7A11 stabilization, ferroptosis resistance, and lenvatinib tolerance in HCC.

Our findings may have translational implications. Lenvatinib resistance remains a major challenge in the treatment of advanced HCC. Recent studies have suggested that targeting oncogenic signaling or ferroptosis-related pathways may enhance lenvatinib efficacy in HCC [16–18, 34, 66] Consistent with this concept, our data showed that OTUD1 contributes to lenvatinib tolerance, at least partly through SLC7A11. Clinical analyses further supported the relevance of this axis, as OTUD1 expression was positively associated with SLC7A11 expression, p-ERK1/2 was positively associated with SLC7A11 expression, and high OTUD1 expression was associated with shorter overall survival. Nevertheless, this study has limitations. The clinical cohort was relatively small, and an independent tissue microarray cohort or a clinically annotated lenvatinib-treated HCC cohort would further strengthen the clinical relevance of our findings. In addition, although SLC7A11 is an important downstream effector of OTUD1, OTUD1 may regulate additional substrates involved in ferroptosis or redox homeostasis. Future studies are needed to systematically identify additional OTUD1 substrates involved in ferroptosis regulation.

In conclusion, our study reveals an ERK1/2–OTUD1–SLC7A11 regulatory axis that promotes ferroptosis resistance in HCC. OTUD1 stabilizes SLC7A11 by removing K48-linked polyubiquitin chains, and ERK1/2-dependent phosphorylation of OTUD1 further enhances this process. These findings link oncogenic kinase signaling to deubiquitination-mediated control of SLC7A11 stability and suggest that targeting this axis may provide a potential strategy to improve ferroptosis-based therapeutic responses in HCC.

Conclusions

In conclusion, our study identifies OTUD1 as an important regulator of ferroptosis resistance in HCC. Functional analyses in HCC cells, subcutaneous xenograft models, and orthotopic tumor models showed that OTUD1 promotes tumor growth and protects HCC cells from ferroptotic stress. Mechanistically, OTUD1 stabilizes SLC7A11 by removing K48-linked polyubiquitin chains, thereby limiting ferroptotic lipid peroxidation and supporting HCC cell survival. ERK1/2-dependent phosphorylation of OTUD1 at Ser216 further enhances the OTUD1–SLC7A11 interaction and promotes SLC7A11 stabilization. These findings reveal an ERK1/2–OTUD1–SLC7A11 regulatory axis that contributes to ferroptosis resistance in HCC and suggest that OTUD1 may represent a potential therapeutic target for this disease.

Supplementary Information

Below is the link to the electronic supplementary material.

Abbreviations

OTUD1

OTU domain-containing deubiquitinase 1

SLC7A11

Solute carrier family 7 member 11

ERK1/2

Extracellular signal-regulated kinase 1/2

ACSL4

Acyl-CoA synthetase long-chain family member 4

GPX4

Glutathione peroxidase 4

FTH1

Ferritin heavy chain 1

4-HNE

4-Hydroxynonenal

MDA

Malondialdehyde

CHX

Cycloheximide

EGF

Epidermal growth factor

Author contributions

Kangsheng Tu and Qingguang Liu conceived and designed the experiments; Runtian Li and Huanye Mo performed the experiments and analyzed the data; Lei Zhang contributed reagents/materials/analysis tools; Runtian Li wrote the paper; Kangsheng Tu reviewed and revised the paper. All authors read and approved the final manuscript.

Funding

This work was supported by the Huadong Medicine Joint Funds of the Zhejiang Provincial Natural Science Foundation of China (Grant No. LHDMD23H300002) and Key Research and Development Projects of Shaanxi Province (2025SF-YBXM-356, 2026SF-YBXM-276).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

All participants in this study signed informed consent. All procedures involving human participants were in accordance with the ethical standards of the Research Ethics Committee of The First Affiliated Hospital of Xian Jiaotong University. All animal experiments were approved by the Biomedical Ethics Committee of Health Science Center of Xi’an Jiaotong University.

Competing interests

The authors declare no competing interests.