NEDD4 suppresses ferroptosis in lung ischemia-reperfusion injury by ubiquitinating and degrading SLC1A5.

NEDD4, an E3 ubiquitin ligase, has emerged as a critical regulator in ischemia-reperfusion injury across multiple organ systems. Studies demonstrate its protective roles in myocardial reperfusion injury by suppressing macrophage pyroptosis 10 and in cerebral hemorrhage by modulating ferroptosis through DMT1 ubiquitination 11 . Importantly, NEDD4’s involvement extends to ferroptosis regulation in diverse pathological contexts, including osteoarthritis 15 and coronary heart disease 16 , where it influences key ferroptosis mediators. Despite these established connections between NEDD4, IRI pathophysiology, and ferroptosis regulation, its role in pulmonary ischemia-reperfusion injury (LIRI) and associated ferroptosis remains unexplored. To address this gap, we first established an LIRI model in C57BL/6J mice via left pulmonary hilar clamping (60 min ischemia + 120 min reperfusion). To robustly confirm the induction of LIRI, we performed a comprehensive analysis encompassing histopathology, physiological function, edema formation, and inflammatory response. Histopathological analysis via H&E staining confirmed successful induction of injury, characterized by alveolar wall thickening (orange arrows), inflammatory cell infiltration (blue arrows), alveolar collapse, and eosinophilic debris in bronchioles (brown arrows) (Fig. 1 A). A semi-quantitative lung injury score confirmed a significant increase in tissue damage in the LIRI group compared to the Sham group (Fig. 1 B). Assessment of pulmonary gas exchange function by arterial blood gas analysis showed that LIRI led to severe hypoxemia, as evidenced by a marked decrease in PaO 2 and an increase in PaCO 2 (Fig. 1 C-D). The inflammatory response was evaluated by measuring pro-inflammatory cytokines. Levels of IL-1β, IL-6, and TNF-α were significantly higher in both bronchoalveolar lavage fluid (BALF) and lung tissue homogenates from LIRI mice compared to Sham mice (Fig. 1 E-F). Collectively, these multi-faceted data unequivocally demonstrate the successful establishment of the LIRI model. Given NEDD4’s protective associations in other IRI/ferroptosis models, we hypothesized it might be dysregulated in LIRI. Assessment of NEDD4 expression revealed a striking reduction. RT-qPCR showed significantly decreased NEDD4 mRNA in injured lungs (Fig.  1 G). Western blot analysis confirmed parallel downregulation at the protein level (Fig.  1 H). Immunohistochemistry further confirmed this suppression, demonstrating reduced NEDD4 level predominantly in alveolar epithelial (Fig.  1 I). Collectively, these data establish that NEDD4 expression is suppressed during LIRI, suggesting its potential role in mitigating ferroptosis-related damage in the ischemic lung. To determine the functional impact of NEDD4 in LIRI, we subjected NEDD4 −/− mice and WT mice to pulmonary ischemia-reperfusion. Histopathological analysis revealed that NEDD4 −/− mice exhibited significantly aggravated tissue damage following LIRI compared to WT-LIRI controls (Fig.  2 A), characterized by enhanced inflammatory infiltration (blue arrows), widespread alveolar wall thickening (orange arrows), eosinophilic debris in bronchioles (brown arrows), scattered lymphocyte infiltration in the interstitium (black arrows), and pronounced bronchiolar mucosal epithelial disorganization (yellow arrows). No obvious pathological alterations were observed in NEDD4 −/− -Sham lungs. Given established links between apoptosis and LIRI pathogenesis, we assessed key apoptotic regulators. Immunohistochemistry demonstrated markedly elevated expression of pro-apoptotic proteins Bax and Cleaved-Casp3 in NEDD4 −/− -LIRI lungs versus WT-LIRI (Fig.  2 B-C). Western blot analysis further confirmed dysregulated apoptosis signaling in NEDD4 −/− -LIRI compared with WT-LIRI, showing increased Bax and decreased anti-apoptotic Bcl-2 expression (Fig.  2 D). Since inflammation amplifies IRI damage, we quantified cytokine levels. ELISA revealed significantly higher concentrations of IL-1β, IL-6, and TNF-α in both BALF and lung tissues from NEDD4 −/− -LIRI mice compared to WT-LIRI controls (Fig.  2 E-F). Collectively, these data demonstrate that NEDD4 deficiency intensifies LIRI severity by promoting apoptosis and inflammatory cascades. To investigate whether NEDD4 modulates ferroptosis in LIRI, we analyzed established hallmarks of ferroptosis in WT and NEDD4 −/− lungs. ELISA assessment revealed that LIRI induced significant oxidative imbalance in WT mice, characterized by elevated lipid peroxidation (MDA) and iron accumulation alongside depleted antioxidant defenses (GSH-Px, SOD) in both BALF and lung tissues. Critically, NEDD4 −/− -LIRI mice exhibited exacerbated oxidative damage, with further increased MDA/iron levels and more pronounced reductions in GSH-Px/SOD activities compared to WT-LIRI controls (Fig.  3 A-B). We next examined key ferroptosis regulatory proteins. Western blot analysis demonstrated that LIRI suppressed expression of the ferroptosis inhibitors GPX4 and SLC7A11 in WT lungs. This suppression was markedly amplified in NEDD4 −/− -LIRI lungs, showing near-abolition of the two protective factors (Fig.  3 C). Collectively, these findings establish that NEDD4 deficiency intensifies ferroptosis during LIRI by augmenting lipid peroxidation, iron overload, and depletion of anti-ferroptotic defenses. To model LIRI in vitro, we subjected mouse (MLE-12) and human (A549) pulmonary epithelial cells to H/R [hypoxia (8 h) + reoxygenation (4, 8, or 12 h)]. Western blot analysis revealed that NEDD4 protein expression decreased progressively with prolonged reoxygenation, reaching minimal levels at 12 h (Fig.  4 A). This time point was selected for subsequent experiments. We next transfected cells with NEDD4-overexpressing plasmid, which significantly elevated NEDD4 mRNA and protein levels in both cell lines (Fig.  4 B-C). Functional assessment demonstrated that H/R injury markedly reduced cell viability and proliferation, effects that were partially reversed by NEDD4 overexpression (Fig.  4 D-E). Given the central role of oxidative stress in ferroptosis, we measured ROS accumulation. H/R injury induced pronounced ROS generation, which was attenuated in NEDD4-overexpressing cells (Fig.  4 F). Concordantly, ELISA revealed that NEDD4 overexpression reduced H/R-induced lipid peroxidation (MDA) and iron overload while restoring antioxidant capacity (GSH-Px, SOD) (Fig.  4 G). Finally, we examined ferroptosis regulatory proteins. H/R suppressed expression of GPX4 and SLC7A11, key inhibitors of ferroptosis. NEDD4 overexpression largely restored their expression toward baseline levels (Fig.  4 H). Collectively, these data demonstrate that NEDD4 overexpression protects pulmonary epithelial cells against H/R-induced ferroptosis by alleviating oxidative damage and restoring anti-ferroptotic defenses. Fig. 4 NEDD4 overexpression mitigates H/R-induced ferroptosis in pulmonary epithelial cells. ( A ) Western blot analysis of NEDD4 protein expression in MLE-12 and A549 cells subjected to hypoxia (8 h) followed by reoxygenation (4, 8, or 12 h). β-actin served as loading control. ( B - C ) Efficiency of NEDD4 overexpression validated by (B) RT-qPCR and (C) Western blot in MLE-12 and A549 cells transfected with NEDD4 plasmid. ( D - E ) Cell viability assessed by CCK-8 assay (D) and proliferation by EdU staining (E) in H/R-exposed cells (12 h reoxygenation) with or without NEDD4 overexpression. ( F ) Intracellular ROS levels detected by DCFH-DA fluorescence in H/R-injured cells. ( G ) ELISA quantification of oxidative stress markers (MDA, GSH-Px, SOD) and iron (Fe 2+ ) in cell lysates. ( H ) Western blot analysis of ferroptosis regulators GPX4 and SLC7A11 in H/R-injured cells. * P <0.05, ** P <0.01, *** P <0.001. NEDD4 overexpression mitigates H/R-induced ferroptosis in pulmonary epithelial cells. ( A ) Western blot analysis of NEDD4 protein expression in MLE-12 and A549 cells subjected to hypoxia (8 h) followed by reoxygenation (4, 8, or 12 h). β-actin served as loading control. ( B - C ) Efficiency of NEDD4 overexpression validated by (B) RT-qPCR and (C) Western blot in MLE-12 and A549 cells transfected with NEDD4 plasmid. ( D - E ) Cell viability assessed by CCK-8 assay (D) and proliferation by EdU staining (E) in H/R-exposed cells (12 h reoxygenation) with or without NEDD4 overexpression. ( F ) Intracellular ROS levels detected by DCFH-DA fluorescence in H/R-injured cells. ( G ) ELISA quantification of oxidative stress markers (MDA, GSH-Px, SOD) and iron (Fe 2+ ) in cell lysates. ( H ) Western blot analysis of ferroptosis regulators GPX4 and SLC7A11 in H/R-injured cells. * P <0.05, ** P <0.01, *** P <0.001. To elucidate the molecular mechanisms by which NEDD4 modulates ferroptosis, we employed TurboID-mediated proximity labeling and mass spectrometry to screen NEDD4-interacting proteins (Supplementary table 1). Ferroptosis-related proteins were annotated using ferrDb V2 ( http://www.zhounan.org/ferrdb ) (Supplementary table 2). Following the identification of five candidate interactors (Fig.  5 A), we prioritized SLC1A5 for validation based on its pathological relevance and mechanistic tractability. Western blot analysis revealed a significant upregulation of SLC1A5 protein that inversely correlated with NEDD4 downregulation in our LIRI model (Fig.  5 B), providing direct evidence of their association in disease. Furthermore, as a transmembrane transporter, SLC1A5 regulates ferroptosis through a direct, linear pathway by controlling glutamine uptake, making the hypothesis that NEDD4 targets it for degradation straightforward to test. In contrast, other candidates like KEAP1 operate within complex signaling networks, complicating the validation of a direct mechanistic link to NEDD4. We therefore selected SLC1A5 for subsequent functional validation.​ Exogenous Co-IP assays in HEK293T cells demonstrated robust reciprocal binding between Flag-NEDD4 and Myc-SLC1A5 (Fig.  5 C), while endogenous Co-IP confirmed NEDD4-SLC1A5 complex formation in both MLE-12 and A549 cells (Fig.  5 D). Immunofluorescence further revealed prominent co-localization of NEDD4 and SLC1A5 at the plasma membrane in A549 cells (Fig.  5 E). To determine the regulatory hierarchy, we assessed whether NEDD4 influences SLC1A5 at the transcriptional or post-transcriptional level. RT-qPCR analysis showed that NEDD4 overexpression had no appreciable effect on SLC1A5 mRNA in either cellular context (Fig.  5 F). In contrast, Western blot demonstrated that NEDD4 overexpression reduced SLC1A5 protein abundance, an effect abolished by proteasome inhibitor MG132, implicating the proteasome degradation pathway (Fig.  5 G). To directly test if NEDD4 promotes SLC1A5 ubiquitination, we performed ubiquitination Co-IP in HEK293T cells. NEDD4 overexpression markedly increased ubiquitin conjugation on SLC1A5 (Fig.  5 H). We further investigated the specific type of ubiquitin chain linkage involved. By co-expressing SLC1A5 with wild-type ubiquitin or mutants (K48R or K63R), we found that NEDD4-mediated SLC1A5 ubiquitination was predominantly K48-linked, as evidenced by the abolished signal with the K48R ubiquitin mutant (Fig.  5 I). This finding is consistent with the proteasomal degradation of SLC1A5, as K48-linked polyubiquitination is the canonical signal for targeting substrates to the proteasome. To determine whether ubiquitination was strictly dependent on NEDD4’s catalytic activity, we employed a catalytically inactive mutant, NEDD4-C867A. Co-IP assays revealed that while wild-type NEDD4 robustly enhanced SLC1A5 ubiquitination, the NEDD4-C867A mutant failed to do so (Fig.  5 J), indicating that the E3 ligase activity of NEDD4 is essential for this process. Consistently, cycloheximide chase assays revealed that NEDD4 overexpression significantly accelerated SLC1A5 protein turnover in both MLE-12 and A549 cells (Fig.  5 K). Collectively, these findings demonstrate that NEDD4 directly interacts with SLC1A5 and promotes its ubiquitination-dependent proteasomal degradation, thereby regulating SLC1A5 protein stability and potentially modulating ferroptosis. Fig. 5 NEDD4 regulates SLC1A5 protein stability via ubiquitination. ( A ) TurboID-biotin ligase-based proximity labeling and mass spectrometry were performed following fusion expression of NEDD4, screening for NEDD4-interacting proteins. Venn analysis revealed five NEDD4-specific interactors overlapping with the pro-ferroptosis genes set, including SLC1A5, a functionally validated ferroptosis driver. ( B ) Western blot analysis of NEDD4 and SLC1A5 protein expression in lung tissues from Sham and LIRI mice. (C-D) Validation of NEDD4-SLC1A5 interaction by Co-IP. ( C ) Exogenous Co-IP: HEK293T cells were co-transfected with Flag-tagged NEDD4 and Myc-tagged SLC1A5. Reciprocal immunoprecipitation was performed using anti-Flag or anti-Myc antibodies, followed by immunoblotting with antibodies against the reciprocal tag. ( D ) Endogenous Co-IP: MLE-12 and A549 cell lysates were immunoprecipitated using anti-NEDD4 antibody, followed by immunoblotting with anti-SLC1A5 antibody. IgG served as negative control. ( E ) Immunofluorescence detection of NEDD4 and SLC1A5 co-localization in A549 cells. ( F ) RT-qPCR analysis of SLC1A5 mRNA in MLE-12 and A549 cells overexpressing NEDD4. ( G ) Western blot analysis of SLC1A5 protein in NEDD4-overexpressing cells with or without MG132 proteasome inhibition (5 µM, 6 h). ( H ) Ubiquitination Co-IP: Myc-SLC1A5 ubiquitination assessed in HEK293T cells co-expressing HA-Ub ± Flag-NEDD4. ( I ) Ubiquitination Co-IP assay in HEK293T cells co-expressing Myc-SLC1A5, Flag-NEDD4, and wild-type HA-Ub (WT) or mutant form (K48R and K63R). ( J ) Ubiquitination Co-IP assay in HEK293T cells co-expressing Myc-SLC1A5, HA-Ub, and either wild-type Flag-NEDD4 or catalytically inactive mutant Flag-NEDD4-C867A. ( K ) Cycloheximide chase assay in MLE-12 and A549 cells transfected with NEDD4 or vector, and SLC1A5 protein stability analyzed by WB after CHX (10 µg/mL) treatment at indicated timepoints (3, 6, or 9 h). ns, not significant, ** P <0.01, *** P <0.001. NEDD4 regulates SLC1A5 protein stability via ubiquitination. ( A ) TurboID-biotin ligase-based proximity labeling and mass spectrometry were performed following fusion expression of NEDD4, screening for NEDD4-interacting proteins. Venn analysis revealed five NEDD4-specific interactors overlapping with the pro-ferroptosis genes set, including SLC1A5, a functionally validated ferroptosis driver. ( B ) Western blot analysis of NEDD4 and SLC1A5 protein expression in lung tissues from Sham and LIRI mice. (C-D) Validation of NEDD4-SLC1A5 interaction by Co-IP. ( C ) Exogenous Co-IP: HEK293T cells were co-transfected with Flag-tagged NEDD4 and Myc-tagged SLC1A5. Reciprocal immunoprecipitation was performed using anti-Flag or anti-Myc antibodies, followed by immunoblotting with antibodies against the reciprocal tag. ( D ) Endogenous Co-IP: MLE-12 and A549 cell lysates were immunoprecipitated using anti-NEDD4 antibody, followed by immunoblotting with anti-SLC1A5 antibody. IgG served as negative control. ( E ) Immunofluorescence detection of NEDD4 and SLC1A5 co-localization in A549 cells. ( F ) RT-qPCR analysis of SLC1A5 mRNA in MLE-12 and A549 cells overexpressing NEDD4. ( G ) Western blot analysis of SLC1A5 protein in NEDD4-overexpressing cells with or without MG132 proteasome inhibition (5 µM, 6 h). ( H ) Ubiquitination Co-IP: Myc-SLC1A5 ubiquitination assessed in HEK293T cells co-expressing HA-Ub ± Flag-NEDD4. ( I ) Ubiquitination Co-IP assay in HEK293T cells co-expressing Myc-SLC1A5, Flag-NEDD4, and wild-type HA-Ub (WT) or mutant form (K48R and K63R). ( J ) Ubiquitination Co-IP assay in HEK293T cells co-expressing Myc-SLC1A5, HA-Ub, and either wild-type Flag-NEDD4 or catalytically inactive mutant Flag-NEDD4-C867A. ( K ) Cycloheximide chase assay in MLE-12 and A549 cells transfected with NEDD4 or vector, and SLC1A5 protein stability analyzed by WB after CHX (10 µg/mL) treatment at indicated timepoints (3, 6, or 9 h). ns, not significant, ** P <0.01, *** P <0.001. To determine whether SLC1A5 mediates NEDD4’s anti-ferroptotic effects, we first established efficient SLC1A5 overexpression in MLE-12 and A549 cells, confirmed by RT-qPCR and Western blot (Fig.  6 A-B). We then performed rescue experiments in H/R-injured cells. NEDD4 overexpression partially restored cell viability and proliferation impaired by H/R. Critically, co-expression of SLC1A5 abolished this protective effect, reducing viability and proliferation to near-H/R levels (Fig.  6 C-D). Consistent with functional outcomes, SLC1A5 co-expression reversed NEDD4’s suppression of H/R-induced ROS accumulation (Fig.  6 E). ELISA analysis further showed that SLC1A5 overexpression counteracted NEDD4’s ability to reduce lipid peroxidation (MDA) and iron overload while blunting its restoration of antioxidant activities (GSH-Px, SOD) (Fig.  6 F). At the molecular level, Western blot revealed that SLC1A5 co-expression attenuated NEDD4-mediated upregulation of ferroptosis inhibitors GPX4 and SLC7A11 (Fig.  6 G). Collectively, these rescue experiments demonstrate that SLC1A5 is a critical downstream effector through which NEDD4 exerts its anti-ferroptotic function in pulmonary epithelial cells. Fig. 6 SLC1A5 overexpression reverses NEDD4-mediated protection against H/R-induced ferroptosis. ( A - B ) Efficiency of SLC1A5 overexpression validated by (A) RT-qPCR and (B) Western blot in MLE-12 and A549 cells transfected with SLC1A5 plasmid. ( C - D ) Functional rescue assays: Cell viability by CCK-8 (C) and proliferation by EdU staining (D) in H/R-injured cells with indicated treatments. ( E ) Intracellular ROS levels detected by DCFH-DA fluorescence. ( F ) ELISA quantification of oxidative stress markers (MDA, GSH-Px, SOD) and iron (Fe 2+ ) in cell lysates. ( G ) Western blot analysis of ferroptosis regulators GPX4 and SLC7A11. * P <0.05, ** P <0.01, *** P <0.001. SLC1A5 overexpression reverses NEDD4-mediated protection against H/R-induced ferroptosis. ( A - B ) Efficiency of SLC1A5 overexpression validated by (A) RT-qPCR and (B) Western blot in MLE-12 and A549 cells transfected with SLC1A5 plasmid. ( C - D ) Functional rescue assays: Cell viability by CCK-8 (C) and proliferation by EdU staining (D) in H/R-injured cells with indicated treatments. ( E ) Intracellular ROS levels detected by DCFH-DA fluorescence. ( F ) ELISA quantification of oxidative stress markers (MDA, GSH-Px, SOD) and iron (Fe 2+ ) in cell lysates. ( G ) Western blot analysis of ferroptosis regulators GPX4 and SLC7A11. * P <0.05, ** P <0.01, *** P <0.001.

Wild-type C57BL/6J mice (male, 8 weeks, 22–25 g) and NEDD4 −/− mice were acquired from Cyagen Biosciences (Suzhou, China). All animals were housed under specific pathogen-free conditions at 22 ± 1 °C with 55% humidity and a 12 h light/dark cycle, with ad libitum access to irradiated chow and autoclaved water. Animal procedures were approved by the Animal Care and Use Committee of Fujian Medical University Union Hospital and complied with ARRIVE 2.0 guidelines. The LIRI model was induced by left lung ischemia as previously described 12 with modifications. Mice were anesthetized with pentobarbital sodium (50 mg/kg, i.p.) and received preoperative buprenorphine analgesia (0.1 mg/kg, s.c.). Orotracheal intubation was performed using a 22G catheter connected to a rodent ventilator (Harvard Apparatus), with tidal volume set at 10 mL/kg, respiratory rate at 120 breaths/min, and inspiratory-to-expiratory ratio of 1:1. A left thoracotomy at the fourth intercostal space exposed the pulmonary hilum, which was occluded with a non-traumatic microclip for 60 min to induce ischemia. SpO 2 typically decreased from a baseline of ~ 98% to a stabilized plateau ranging from 91% to 94%. Reperfusion was initiated by clip removal and maintained for 120 min. Sham-operated mice underwent identical procedures without hilar clamping. For Fig.  1 analyses, mice were divided into two groups ( n  = 5/group): WT-Sham (sham surgery + 180 min observation) and WT-LIRI (60 min ischemia + 120 min reperfusion). One mouse in WT-LIRI group was excluded from the study due to SpO 2  < 90% during ischemia. At the experimental endpoint, mice were deeply anesthetized with pentobarbital sodium (100 mg/kg, i.p.) and euthanized by cervical dislocation after confirming the absence of reflex response. The left lungs were fixed in 4% paraformaldehyde for histology, while right lungs were snap-frozen in liquid nitrogen for molecular analyses. Bronchoalveolar lavage fluid (BALF) was collected by tracheal cannulation with three instillations of 0.8 mL sterile PBS, followed by centrifugation (500 ×g, 10 min, 4 °C); supernatants were stored at −80 °C. Fig. 1 NEDD4 expression is downregulated in murine LIRI model. ( A ) Histopathological assessment of lung tissues from sham-operated (Sham) and LIRI mice ( n  = 5/group) by H&E staining. Scale bar: 50 μm. ( B ) Semi-quantitative assessment of lung injury score. ( C and D ) Arterial blood gas analysis showing PaO 2 and PaCO 2 levels. ( E - F ) ELISA quantification of pro-inflammatory cytokines (IL-1β, IL-6, TNF-α) in bronchoalveolar lavage fluid (BALF) (E) and lung tissue homogenates (F). ( G ) RT-qPCR analysis of NEDD4 mRNA in lung tissues. Data normalized to β-actin. ( H ) Western blot analysis of NEDD4 protein in lung homogenates. β-actin served as loading control. ( I ) Immunohistochemical staining of NEDD4 in lung sections. Scale bar: 20 μm. * P <0.05, *** P <0.001. NEDD4 expression is downregulated in murine LIRI model. ( A ) Histopathological assessment of lung tissues from sham-operated (Sham) and LIRI mice ( n  = 5/group) by H&E staining. Scale bar: 50 μm. ( B ) Semi-quantitative assessment of lung injury score. ( C and D ) Arterial blood gas analysis showing PaO 2 and PaCO 2 levels. ( E - F ) ELISA quantification of pro-inflammatory cytokines (IL-1β, IL-6, TNF-α) in bronchoalveolar lavage fluid (BALF) (E) and lung tissue homogenates (F). ( G ) RT-qPCR analysis of NEDD4 mRNA in lung tissues. Data normalized to β-actin. ( H ) Western blot analysis of NEDD4 protein in lung homogenates. β-actin served as loading control. ( I ) Immunohistochemical staining of NEDD4 in lung sections. Scale bar: 20 μm. * P <0.05, *** P <0.001. For Figs.  2 and 3 , four experimental groups ( n  = 5/group) were utilized: WT-Sham, NEDD4 −/− -Sham, WT-LIRI, and NEDD4 −/− -LIRI. Surgical protocols and tissue processing mirrored Fig.  1 procedures. Two mice in NEDD4 −/− -LIRI group were excluded due to SpO 2  < 90% during ischemia to ensure model consistency.​. Fig. 2 NEDD4 deficiency exacerbates LIRI-induced lung injury, apoptosis, and inflammation. ( A ) H&E-stained lung sections from WT and NEDD4 −/− mice under sham or LIRI conditions ( n  = 5/group). Scale bars: 50 μm. ( B - C ) Immunohistochemical analysis of pro-apoptotic markers (B) Bax and (C) Cleaved-Casp3 in lung tissues. Scale bars: 20 μm. ( D ) Western blot analysis of apoptosis regulators Bax and Bcl-2 in lung homogenates. ( E - F ) ELISA quantification of inflammatory cytokines in (E) bronchoalveolar lavage fluid (BALF) and (F) lung tissue homogenates. * P <0.05, ** P <0.01, *** P <0.001. NEDD4 deficiency exacerbates LIRI-induced lung injury, apoptosis, and inflammation. ( A ) H&E-stained lung sections from WT and NEDD4 −/− mice under sham or LIRI conditions ( n  = 5/group). Scale bars: 50 μm. ( B - C ) Immunohistochemical analysis of pro-apoptotic markers (B) Bax and (C) Cleaved-Casp3 in lung tissues. Scale bars: 20 μm. ( D ) Western blot analysis of apoptosis regulators Bax and Bcl-2 in lung homogenates. ( E - F ) ELISA quantification of inflammatory cytokines in (E) bronchoalveolar lavage fluid (BALF) and (F) lung tissue homogenates. * P <0.05, ** P <0.01, *** P <0.001. Fig. 3 NEDD4 deficiency exacerbates LIRI-induced ferroptosis. ( A - B ) ELISA quantification of oxidative stress markers (MDA, GSH-Px, SOD) and iron (Fe 2+ ) levels in BALF (A) and lung tissue homogenates (B) from WT and NEDD4 −/− mice under sham or LIRI conditions. ( C ) Western blot analysis of ferroptosis regulators GPX4 and SLC7A11 in lung homogenates. * P <0.05, ** P <0.01, *** P <0.001. NEDD4 deficiency exacerbates LIRI-induced ferroptosis. ( A - B ) ELISA quantification of oxidative stress markers (MDA, GSH-Px, SOD) and iron (Fe 2+ ) levels in BALF (A) and lung tissue homogenates (B) from WT and NEDD4 −/− mice under sham or LIRI conditions. ( C ) Western blot analysis of ferroptosis regulators GPX4 and SLC7A11 in lung homogenates. * P <0.05, ** P <0.01, *** P <0.001. Lung tissues designated for histopathological analysis were fixed in 4% paraformaldehyde at 4 °C for 24 h. Following fixation, tissues underwent dehydration through a graded ethanol series (70%, 80%, 95%, and 100% ethanol, 1 h per concentration), cleared in xylene (two changes, 1 h each), and embedded in paraffin blocks. Sections were cut at 4 μm thickness using a rotary microtome and mounted on poly-L-lysine-coated glass slides. For staining, sections were deparaffinized in xylene (2 × 10 min) and rehydrated through descending ethanol concentrations (100%, 95%, 80%, 70%, 2 min each) to distilled water. Nuclear staining was performed with Mayer’s hematoxylin solution for 5 min, followed by rinsing under running tap water for 5 min. Differentiation was achieved by brief immersion in 1% acid ethanol (1% HCl in 70% ethanol) for 5 s, and sections were subsequently blued in 0.2% ammonia water for 1 min. Cytoplasmic counterstaining was conducted with eosin Y for 2 min. After staining, sections were dehydrated through ascending ethanol series (70%, 80%, 95%, 100% ethanol, 30 s each), cleared in xylene (2 × 3 min), and permanently mounted with neutral balsam. Lung injury degree was observed, including alveolar wall thickening, inflammatory cell infiltration, hemorrhage, and edema. The degree of lung injury was assessed using a well-established semi-quantitative scoring system based on the following criteria: alveolar congestion, hemorrhage, neutrophil infiltration in the airspace or vessel wall, and alveolar wall thickening. Each criterion was scored on a 0 to 4 point scale: 0 = minimal damage, 1 = mild damage, 2 = moderate damage, 3 = severe damage, and 4 = maximal damage. The total lung injury score for each sample was calculated as the sum of the scores for all criteria, yielding a maximum possible score of 16. At the end of the reperfusion period, arterial blood was collected by direct puncture of the left ventricle. Blood gas analysis was immediately performed on a handheld blood gas analyzer according to the manufacturer’s instructions. The partial pressure of oxygen (PaO 2 ) and carbon dioxide (PaCO 2 ) were directly measured and recorded for each sample. Total RNA was extracted from lung tissues or cultured cells (MLE-12 and A549) using TRIzol ® reagent (Invitrogen) according to the manufacturer’s protocol. RNA concentration and purity were determined spectrophotometrically, with A260/A280 ratios between 1.8 and 2.0 deemed acceptable. First-strand cDNA synthesis was performed using 1 µg RNA with the RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific) and oligo(dT) primers. Quantitative PCR amplification was carried out in triplicate using SYBR Green Master Mix (Bio-Rad) on a QuantStudio 5 Real-Time PCR System (Applied Biosystems). Each 20 µL reaction contained 10 µL SYBR Green mix, 1 µL cDNA template, 0.8 µL each of forward/reverse primers (10 µM), and 7.4 µL nuclease-free water. The thermal cycling protocol consisted of initial denaturation at 95 °C for 10 min, followed by 40 cycles of 95 °C for 15 s and 60 °C for 1 min. Relative mRNA expression was calculated using the 2 −ΔΔCt method, normalized to β-actin, and expressed as fold-change relative to controls. Protein lysates were extracted from lung tissues or cultured cells (MLE-12, A549, HEK293T) using RIPA lysis buffer (Beyotime) supplemented with protease inhibitors (Roche). Lysates were centrifuged at 12,000 ×g for 15 min at 4 °C, and supernatants were collected. Protein concentrations were determined via BCA assay according to the manufacturer’s instructions. Equal amounts of protein (30 µg per lane) were resolved on 10% SDS-polyacrylamide gels and electrophoretically transferred to PVDF membranes (Millipore). Membranes were blocked with 5% non-fat milk in TBST for 1 h at room temperature and subsequently incubated overnight at 4 °C with primary antibodies diluted in blocking buffer. After three washes with TBST, membranes were incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies for 1 h at room temperature. Protein bands were visualized using enhanced chemiluminescence (ECL) substrate and imaged. Band intensities were quantified using ImageJ software and normalized to β-actin. Key primary antibodies used were listed as follows: NEDD4 (Proteintech, #21698-1-AP, 1:8,000), Bax (Proteintech, #50599-2-Ig, 1:8,000), Bcl-2 (Proteintech, #26593-1-AP, 1:3,000), GPX4 (Proteintech, #67763-1-Ig, 1:3,000), SLC7A11 (Proteintech, #26864-1-AP, 1:3,000), Flag (Proteintech, #20543-1-AP, 1:8,000), Myc (Proteintech, #60003-2-Ig, 1:8,000), SLC1A5 (Proteintech, #20350-1-AP, 1:8,000), HA (Proteintech, #51064-2-AP, 1:8,000), and β-actin (Proteintech, #66009-1-Ig, 1:20,000). Tissue Sect. (4 µm thickness) from paraffin-embedded lungs were deparaffinized in xylene (2 × 10 min) and rehydrated through a graded ethanol series (100%, 95%, 80%, and 70%). Antigen retrieval was performed by heating slides in citrate buffer (pH 6.0) at 95 °C for 20 min using a microwave oven. Endogenous peroxidase activity was quenched with 3% hydrogen peroxide in methanol for 15 min at room temperature. Non-specific binding was blocked with 5% bovine serum albumin (BSA) in PBS for 1 h. Sections were incubated overnight at 4 °C with primary antibodies diluted in blocking buffer. After washing with PBS, slides were incubated with HRP-conjugated secondary antibodies for 1 h at room temperature. Signal development used 3,3’-diaminobenzidine (DAB) substrate for 3 min, followed by hematoxylin counterstaining for 1 min. Sections were dehydrated, cleared in xylene, and mounted with neutral balsam. Key primary antibodies used were listed as follows: NEDD4 (Proteintech, #21698-1-AP, 1:300), Bax (Proteintech, #50599-2-Ig, 1:800), Cleaved-Casp3 (Proteintech, #19677-1-AP, 1:300). ELISA quantified inflammatory cytokines (IL-1β, IL-6, TNF-α), oxidative markers (MDA, GSH-Px, SOD), and iron (Fe 2+ ) in bronchoalveolar lavage fluid (BALF) and lung tissue homogenates using commercial kits (Beyotime: #PI301, #PI326, #PT512, #S0131S, #S0056, #S0101S; Mlbio: #ml095093). BALF supernatants (centrifuged at 500 ×g, 10 min, 4 °C) and lung homogenates (PBS-homogenized, 12,000 ×g, 15 min) were analyzed per manufacturer protocols. Samples/standards (100 µL) underwent sequential incubations with detection antibodies and HRP-streptavidin, followed by TMB substrate development. Absorbance (450 nm) was measured, with concentrations calculated from standard curves. Mouse lung epithelial cells (MLE-12, Sunncell Biotech, #SNL-414), human alveolar adenocarcinoma cells (A549, Procell, #CL-0016), and human embryonic kidney cells (HEK293T, Procell, #CL-0005) were cultured in DMEM supplemented with 10% fetal bovine serum (Gibco) and 1% penicillin/streptomycin at 37 °C under 5% CO 2 . For H/R modeling, cells were serum-starved for 12 h, transferred to an anaerobic chamber (95% N 2 /5% CO 2 ) for 8 h hypoxia, then reoxygenated in normoxic conditions (21% O 2 ) for 4, 8, or 12 h. Cells (MLE-12, A549, HEK293T) were seeded at 60–80% confluency 24 h prior to transfection. Lipofectamine 3000 (Thermo Fisher Scientific) was used for all plasmid transfections according to manufacturer’s protocol. Full-length human NEDD4 cDNA was cloned into pcDNA3.1 with a Flag tag, named Flag-NEDD4. Human SLC1A5 cDNA was subcloned into pcDNA3.1 with a Myc tag, named Myc-SLC1A5. The culture supernatant was replaced after 6 h. Cells were harvested 48 h post-transfection for downstream analyses. Transfection efficiency was validated by RT-qPCR and WB. Cell viability was assessed using the Cell Counting Kit-8 (Beyotime, #C0041). Cells (MLE-12/A549) were seeded in 96-well plates (5 × 10 3 cells/well) and subjected to experimental treatments (H/R, transfection). After interventions, 10 µL CCK-8 reagent was added per well and incubated for 2 h at 37 °C. Absorbance at 450 nm was measured using a microplate reader. Cell proliferation was evaluated using the Click-iT™ EdU-488 cell proliferation assay kit (Servicebio, #G1601) following the manufacturer’s instructions. Cells were seeded onto glass coverslips in 24-well plates and subjected to experimental treatments. EdU (10 µM final concentration) was added to the culture medium and cells were incubated at 37 °C for 2 h. After incubation, cells were fixed with 4% paraformaldehyde at room temperature for 15 min, permeabilized with 0.5% Triton X-100 for 10 min, and incubated with the Click-iT reaction cocktail to detect incorporated EdU. Nuclei were counterstained with DAPI. EdU-positive cells were visualized using a fluorescence microscope and five random fields were imaged per group. The proportion of EdU positive cells was calculated as the percentage of EdU positive cells relative to total DAPI-stained nuclei. Intracellular ROS levels were measured using the DCFH-DA fluorescent probe (Beyotime, #S0033S). After the indicated treatments, cells were incubated with 10 µM DCFH-DA diluted in serum-free medium at 37 °C for 30 min in the dark. Following incubation, cells were washed three times with PBS to remove excess probe and immediately imaged under a fluorescence microscope. ROS production was quantified by measuring mean fluorescence intensity in at least five random fields using ImageJ software. The NEDD4-TurboID fusion construct was generated by cloning the human NEDD4 coding sequence into the pLVX-EnCMV-V5-TurboID-NES-PGK-Puro vector. A549 cells were transfected with NEDD4-TurboID (NEDD4) or empty TurboID vector (Vec). After 48 h, proximity-dependent biotinylation was initiated with 100 µM biotin for 2 h at 37 °C 13 , 14 . Cells were immediately cooled on ice, washed with PBS, and lysed in RIPA buffer. Lysates were incubated with streptavidin magnetic beads overnight at 4 °C. Beads were stringently washed (20 mM HEPES, 150 mM NaCl, 5 mM MgCl 2 , 1% NP-40). Eluted proteins were subjected to Liquid Chromatography-Tandem Mass Spectrometry (LC-MS/MS). NEDD4-specific interactors were distinguished from background through comparative profiling against empty TurboID vector controls. Ferroptosis-related proteins were annotated using ferrDb V2 ( http://www.zhounan.org/ferrdb ). Protein samples enriched via streptavidin pull-down were processed using the MagicOmics-MMB8X kit (QLBIO). Briefly, 20 µL protein lysate was mixed with MMB beads, incubated with binding buffer for 15 min at room temperature, and washed three times. Proteins were digested with trypsin in digestion buffer at 37 °C for 4 h, followed by reaction quenching. Peptides were lyophilized and reconstituted in mobile phase A (0.1% formic acid in H 2 O). LC-MS/MS analysis was performed on a Q Exactive HF-X mass spectrometer coupled to a RIGOL L-3000 nanoflow HPLC system. Peptides (1 µg) were separated over a 30-min gradient: 8% to 12% mobile phase B (0.1% formic acid in 80% acetonitrile) in 2 min, 12% to 30% in 15 min, 30% to 40% in 3 min, and 95% B for 9 min. MS parameters included NSI ion source (2.4 kV), 275 °C ion transfer tube, and data-dependent acquisition (top 40 precursors; 120,000 resolution for MS1, 15,000 for MS2). Raw files were processed using Proteome Discoverer 2.4 against the Homo sapiens UniProt database (20240307 release). For exogenous Co-IP, HEK293T cells were co-transfected with Flag-tagged NEDD4 and Myc-tagged SLC1A5 using Lipofectamine 3000. At 48 h post-transfection, cells were lysed in NP-40 buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1% NP-40) supplemented with protease inhibitors. Lysates (500 µg) were precleared with Protein A/G beads, then incubated with anti-Flag M2 magnetic beads or anti-Myc magnetic beads overnight at 4 °C with rotation. Beads were washed four times with lysis buffer, and bound proteins were eluted at 95 °C for 10 min. Immunoprecipitates were analyzed by Western blot using anti-Myc or anti-Flag antibodies. For endogenous Co-IP, MLE-12 and A549 cells were lysed in RIPA buffer. Cell lysates (1 mg) were incubated with 4 µg anti-NEDD4 antibody (Proteintech, #21698-1-AP) or control IgG overnight at 4 °C, followed by Protein A/G bead capture for 4 h. Beads were washed stringently (three times with RIPA, once with PBS), and complexes were eluted for Western blot analysis with anti-SLC1A5 antibody (Proteintech, #20350-1-AP) and anti-NEDD4 antibody (Proteintech, #21698-1-AP). To investigate the spatial interaction between NEDD4 and SLC1A5, cells were fixed with 4% paraformaldehyde (15 min, RT), permeabilized with 0.1% Triton X-100 (10 min), and blocked with 5% BSA in PBS (1 h). Dual staining was performed using Rabbit anti-NEDD4 (Proteintech, #21698-1-AP, 1:200) detected with Alexa Fluor 488 (green) and Mouse anti-SLC1A5 (Proteintech, #68540-1-Ig, 1:200) detected with Cy5 (far-red). Cells were incubated with primary antibodies overnight at 4 °C. The next day, cells were incubated with species-matched secondary antibodies for 1 h at RT in the dark. Nuclei were counterstained with DAPI (1 µg/mL, 5 min). Images were acquired using a confocal microscope with appropriate filter sets (488 nm excitation for Alexa Fluor 488, 647 nm excitation for Cy5). To assess NEDD4-mediated ubiquitination of SLC1A5, HEK293T cells were co-transfected with Myc-tagged SLC1A5, HA-tagged ubiquitin, and either Flag-tagged NEDD4 or empty vector. At 36 h post-transfection, cells were treated with 10 µM MG132 (proteasome inhibitor) for 6 h to stabilize ubiquitinated proteins. Cells were lysed in NP-40 buffer supplemented with 10 mM N-ethylmaleimide (ubiquitin chain preservation) and protease inhibitors. Lysates (800 µg) were incubated with anti-Myc magnetic beads overnight at 4 °C with rotation. Beads were washed four times with high-stringency buffer (50 mM Tris pH 7.5, 150 mM NaCl, 1% NP-40, 0.5% deoxycholate) to remove nonspecific binders. Immunoprecipitated complexes were eluted at 95 °C for 10 min and resolved by SDS-PAGE. Ubiquitinated SLC1A5 was detected by Western blot using anti-HA antibody (Proteintech, #51064-2-AP), with total Myc-SLC1A5 as loading control. Protein degradation kinetics were assessed in MLE-12 and A549 cells transfected with NEDD4 overexpression plasmid or empty vector. At 48 h post-transfection, cells were treated with 10 µg/mL CHX to inhibit de novo protein synthesis. Cells were harvested at specified timepoints (0, 3, 6, or 9 h) post-CHX addition and lysed in RIPA buffer containing protease inhibitors. Total protein concentrations were determined by BCA assay, with equal amounts (30 µg per lane) resolved by SDS-PAGE. SLC1A5 protein levels were detected by Western blot using anti-SLC1A5 antibody (Proteintech, #20350-1-AP, 1:8,000), while β-actin served as loading control. All quantitative data are presented as mean ± standard deviation (SD). Statistical analyses were performed using GraphPad Prism 9.0 software. Normality of data distribution was confirmed by Shapiro-Wilk testing. For comparisons between two groups, unpaired Student’s t -tests were applied. Multiple group comparisons utilized one-way analysis of variance (ANOVA) with Tukey’s post hoc test for normally distributed data. Significance thresholds were defined as * P  < 0.05, ** P  < 0.01, *** P  < 0.001.

LIRI poses a significant clinical challenge in thoracic surgery and transplantation, contributing substantially to postoperative morbidity and mortality 17 , 18 . Dysregulated cell death contributes centrally to LIRI pathogenesis 17 . Ferroptosis, an iron-dependent cell death mechanism characterized by uncontrolled lipid peroxidation, represents a critical driver of LIRI injury 2 . While therapeutic strategies targeting ferroptosis show promise, endogenous regulators of this pathway in pulmonary contexts remain incompletely defined. This study identifies the E3 ubiquitin ligase NEDD4 as a novel suppressor of ferroptosis in LIRI, operating through targeted degradation of the pro-ferroptotic transporter SLC1A5. NEDD4 and its homolog NEDD4L are key members of the HECT family of E3 ubiquitin ligases, regulating diverse disease processes through substrate ubiquitination 19 – 21 . NEDD4 is critically involved in ferroptosis regulation. For instance, PUM2 promotes PTEN-mediated chondrocyte ferroptosis and osteoarthritis progression by degrading NEDD4 mRNA 15 . SGK1 modulates ferroptosis in coronary heart disease via the NEDD4L/NF-κB pathway 16 . NEDD4 also exerts protective functions in ischemia-reperfusion injury. It mitigates myocardial reperfusion injury by suppressing macrophage pyroptosis 10 , while celastrol targeting Nedd4 reduces Nrf2-mediated oxidative stress in astrocytes after ischemic stroke 22 . Crucially, NEDD4-mediated DMT1 ubiquitination protects against ferroptosis in cerebral hemorrhage 11 . Despite these established roles, NEDD4’s function in pulmonary ferroptosis during LIRI remains unexplored. Our study aims to addresses this gap to provide a foundation for NEDD4-targeted therapies. We first demonstrated significant downregulation of NEDD4 in murine LIRI. Critically, NEDD4 −/− mice exhibited exacerbated lung injury, apoptosis, inflammation, and ferroptosis markers. Enhanced lipid peroxidation (MDA), iron overload, and depletion of GPX4/SLC7A11 in knockout animals established ferroptosis as a primary pathological driver. These findings extend NEDD4’s known anti-ferroptotic functions in osteoarthritis 15 , coronary disease 16 , melanoma 23 , and endometriosis 24 to the pulmonary system. To elucidate the molecular mechanism underlying NEDD4-mediated protection, we employed TurboID proximity labeling coupled with mass spectrometry. This approach screened for proteins interacting with NEDD4 and closely related to ferroptosis. TurboID utilizes an engineered biotin ligase to label proximal proteins in living cells, capturing transient or weak interactions inaccessible to conventional methods 25 , 26 . Screening identified SLC1A5 as a key NEDD4 interactor. Emerging evidence identifies glutamine metabolism as a critical regulator of ferroptosis 27 . Glutamine provides carbon for TCA cycle replenishment via α-ketoglutarate conversion, fueling lipid peroxidation through oxidizable PUFA generation. Central to this process is SLC1A5, the primary transporter mediating Na + -dependent glutamine uptake. Genetic ablation of SLC1A5 or key glutamine-metabolizing enzymes (e.g., glutaminase GLS2, transaminase GOT1) may attenuate ferroptotic injury in LIRI 28 , 29 , positioning SLC1A5 as a key regulator in ferroptotic cascades. Wang et al.. reported that SLC1A5 acts as a ferroptosis-inducing gene in myocardial infarction, aggravating myocardial injury by mediating cardiomyocyte ferroptosis 30 . Ma et al.. demonstrated that SLC1A5 acts as a pro-ferroptotic mediator in endometriosis, amplifying ferroptosis sensitivity through a ROS/c-Myc/SLC1A5 positive feedback loop that drives glutamine-dependent oxidative cell death 31 . Subsequent validation confirmed the physical interaction between NEDD4 and SLC1A5. We further determined that NEDD4 ubiquitinates SLC1A5 and regulates its stability, leading to proteasomal degradation. Functionally, SLC1A5 overexpression reversed NEDD4-mediated protection against H/R-induced ferroptosis in vitro, establishing SLC1A5 as the pivotal downstream effector. This mechanism contrasts with NEDD4-mediated ferroptosis regulation via VDAC2/3 in melanoma 23 or DMT1 in intracerebral hemorrhage 11 , underscoring its context-dependent substrate specificity. These findings highlight promising therapeutic avenues. First, restoring NEDD4 expression (e.g., via adeno-associated virus delivery or small-molecule NEDD4 activators) attenuated ferroptosis in pulmonary epithelia, suggesting clinical potential. Second, direct inhibition of SLC1A5 (e.g., via pharmacological agents like V9302) may disrupt its pro-ferroptotic activity. Third, combining ferroptosis inhibitors (e.g., Fer-1 or Liproxstatin-1) with NEDD4 pathway activators may yield synergistic benefits. Several limitations warrant consideration. While we used systemic NEDD4 −/− mice, cell type-specific knockouts would clarify compartmentalized effects, particularly in macrophages which critically regulate LIRI and where NEDD4’s function remains uncharacterized. Furthermore, while our study defines the key ubiquitin linkage type (K48) and establishes the necessity of NEDD4’s catalytic activity, the precise molecular determinants of the NEDD4-SLC1A5 interaction remain to be elucidated, including the specific protein domains and the exact ubiquitination acceptor sites on SLC1A5. Human validation of NEDD4/SLC1A5 expression in clinical LIRI samples would strengthen the translational relevance of our findings. Finally, the upstream triggers of NEDD4 downregulation require elucidation. Although our in vitro rescue experiments strongly support the model, definitive in vivo functional rescue through SLC1A5 knockdown in NEDD4-deficient mice would provide the most compelling validation of the axis’s physiological role. Despite these limitations, our findings establish NEDD4 as a molecular brake on ferroptosis in LIRI. By ubiquitinating SLC1A5 and facilitating its degradation, NEDD4 disrupts iron accumulation and lipid peroxidation cascades (Graphical Abstract). In conclusion, this research proved that reduced NEDD4 expression fails to ubiquitinate the glutamine transporter SLC1A5, leading to SLC1A5 accumulation, and then drives fuels lipid peroxidation and ferroptosis(Fig.  7 ). Here, we delineate the NEDD4-SLC1A5 axis as a central regulator of ferroptotic lung injury. Pharmacological modulation of this pathway holds significant promise for improving outcomes in LIRI. Fig. 7 Normally, NEDD4-mediated ubiquitination targets SLC1A5 for proteasomal degradation, thereby suppressing ferroptotic cell death and protecting pulmonary tissue. In lung ischemia-reperfusion injury (LIRI), reduced NEDD4 expression impairs its E3 ligase function, failing to ubiquitinate the glutamine transporter SLC1A5. This allows SLC1A5 accumulation, driving excessive glutamine uptake that fuels lipid peroxidation and ferroptosis. Normally, NEDD4-mediated ubiquitination targets SLC1A5 for proteasomal degradation, thereby suppressing ferroptotic cell death and protecting pulmonary tissue. In lung ischemia-reperfusion injury (LIRI), reduced NEDD4 expression impairs its E3 ligase function, failing to ubiquitinate the glutamine transporter SLC1A5. This allows SLC1A5 accumulation, driving excessive glutamine uptake that fuels lipid peroxidation and ferroptosis.

Lung ischemia-reperfusion injury (LIRI) remains a devastating complication of thoracic surgeries and lung transplantation, driving significant morbidity and mortality through oxidative tissue damage and dysregulated cell death 1 . Ferroptosis, an iron-dependent form of regulated cell death characterized by uncontrolled lipid peroxidation, which has been established as a critical driver of LIRI 2 . Mounting evidence underscores the therapeutic potential of targeting ferroptosis, demonstrating that specific interventions can effectively mitigate lung injury. For instance, pro-resolving lipid mediators such as Lipoxin A4 and Maresin1 have been shown to attenuate LIRI by suppressing ferroptosis in alveolar epithelial and endothelial cells through the NRF2 and PKA-Hippo-YAP signaling pathways, respectively 3 , 4 . While these pharmacologic strategies highlight the centrality of ferroptosis, the role of endogenous regulatory remains less explored. The ubiquitin-proteasome system (UPS) serves as a critical stress-responsive mechanism in ischemic pathologies 5 , 6 . It is increasingly recognized as a pivotal regulator of sterile inflammation and cell death in LIRI. For example, damage-associated molecular patterns (DAMPs) released during lung I/R promote inflammation by enhancing the K63-linked ubiquitination of TRAF6 7 . Similarly, systemic inhibition of the upstream ubiquitin-activating enzyme E1 attenuates NF-κB activation and subsequent multi-organ injury following intestinal I/R 8 . Most recently, the E3 ligase UBE3B was shown to protect against acute lung injury by ubiquitinating the ferritinophagy receptor NCOA4 to suppress ferroptosis 9 , directly linking UPS activity to the regulation of this cell death pathway. Despite these advances connecting the UPS to IRI, the role of specific E3 ubiquitin ligases in modulating ferroptosis within the context of pulmonary I/R remains largely unexplored. The E3 ubiquitin ligase neural precursor cell expressed, developmentally down-regulated 4 (NEDD4) has emerged as a key regulator of cell survival in extra-pulmonary organs. It mitigates myocardial reperfusion injury by suppressing pyroptosis 10 and protects against cerebral hemorrhage by modulating ferroptosis through DMT1 ubiquitination 11 . However, despite these established connections between NEDD4, IRI pathophysiology, and ferroptosis regulation, its function and specific substrates in pulmonary ferroptosis during LIRI are unknown. Identifying the key substrates through which NEDD4 exerts its effects is crucial, yet challenging due to the transient nature of E3 ligase-substrate interactions. To overcome this limitation and directly capture NEDD4-specific proximal proteins, we employed TurboID proximity labeling technology.​. Here, we bridge this knowledge gap through integrated in vivo and in vitro approaches. Using TurboID proximity proteomics to resolve transient interactions, we identify the ferroptosis-promoting transporter solute carrier family 1 member 5 (SLC1A5) as a novel NEDD4 substrate. We demonstrate that NEDD4 is downregulated in LIRI, and its deficiency exacerbates ferroptotic injury. Mechanistically, NEDD4 ubiquitinates SLC1A5, targeting it for proteasomal degradation, thereby disrupting glutamine metabolism and lipid peroxidation. Finally, SLC1A5 overexpression reverses the protective effects of NEDD4. Our work establishes the NEDD4-SLC1A5 axis as a crucial endogenous pathway limiting ferroptosis in LIRI, revealing a novel therapeutic target for intervention.

Below is the link to the electronic supplementary material. Supplementary Material 1 Supplementary Material 1 Supplementary Material 2 Supplementary Material 2