Epithelial SLC39A1 prevents acute lung injury through zinc-mediated transcriptional activation of autophagy in male mice

Abstract

Zinc transporters regulate intracellular zinc homeostasis, but their role in acute lung injury (ALI) or acute respiratory distress syndrome (ARDS) remains underexplored. Here, we show that the zinc transporter SLC39A1 is highly upregulated in alveolar type II (AT2) cells from male murine ALI models and patients with ARDS. AT2-specific Slc39a1 deletion or zinc chelation exacerbates lung injury, whereas overexpression or zinc supplementation attenuates it. Notably, zinc supplementation fails to rescue Slc39a1-deficient mice, indicating SLC39A1 governs zinc uptake to control ALI. Zinc likely directly binds to and activates TFEB, TFE3, and MITF, inducing transcriptional activation of autophagy to eliminate damaged mitochondria and suppress apoptosis/pyroptosis in AT2 cells. Lc3b- or Tfe3-deficient mice show heightened lung injury, which remain unmitigated by zinc supplementation. Importantly, administration of AAV-shLc3b to AT2 Slc39a1-deficient mice did not further aggravate lung injury beyond that caused by either intervention alone. This epistatic relationship places SLC39A1 upstream of autophagy activation within a linear pathway. Collectively, we define an essential role for epithelial SLC39A1 in host defense against ALI/ARDS, which is mediated by a protective zinc-autophagy axis.

This study establishes the link between epithelial zinc transporter SLC39A1 and acute lung injury. The authors uncover a protective mechanism in lung epithelium mediated by the SLC39A1-zinc-autophagy axis and highlight SLC39A1 as a potential therapeutic target.

Introduction

Acute lung injury (ALI) and its severe manifestation, acute respiratory distress syndrome (ARDS), represent critical syndromes with a high mortality rate of nearly 40%^1,2. The pathogenesis of ALI/ARDS is intricate, involving the disruption of the alveolar-capillary barrier and dysregulated pulmonary inflammation as the fundamental hallmarks^3,4. This barrier consists of both epithelial and endothelial components. Within the epithelium, alveolar type I (AT1) and type II (AT2) cells collectively form a tight barrier that limits the passage of proteins and solutes. However, during ALI, this critical function is impaired, leading to compromised edema prevention. The underlying mechanism is primarily the death of alveolar epithelial cells, triggered by intrapulmonary or extrapulmonary insults^5,6. Furthermore, alveolar epithelial cells can be activated by proinflammatory cytokines secreted by resident alveolar macrophages, exacerbating and perpetuating inflammatory responses and lung injuries. Consequently, damage to the lung epithelium emerges as a crucial factor in the pathogenesis of ALI/ARDS⁷.

Zinc is the second most abundant essential element in the human body. As a crucial cofactor, zinc binds to a plethora of proteins to maintain normal cellular functions⁸. Zinc deficiency is implicated in the onset and progression of various diseases, and the significance of zinc to health is increasingly recognized and appreciated⁹. Clinical observations have revealed that lower serum zinc levels are positively correlated with disease severity in critically ill ARDS patients^10,11, hinting at the potential therapeutic benefit of zinc supplementation¹². Given that zinc cannot easily permeate through phospholipid bilayers, a complex regulatory system is necessary to control its transport and availability within different cellular compartments¹³. Intracellular zinc homeostasis is mainly hinged on two families of zinc transporters, SLC39A1-SLC39A14 and SLC30A1-SLC30A10, which either increase or decrease cytosolic zinc concentration, respectively. Specific zinc transporters assume pivotal roles in various pathophysiological processes, some of them exhibiting detrimental effects while others have protective effects^14–19. In particular, SLC39A12 was reported to regulate the pulmonary vascular response to chronic hypoxia²⁰. Nonetheless, the precise role of zinc transporters in ALI/ARDS remains largely elusive. In this study, we aim to identify the key zinc transporter involved in ALI/ARDS and to elucidate its underlying mechanism. We demonstrate that epithelial SLC39A1 facilitates zinc uptake, thereby suppressing cell death in AT2 cells through activation of the zinc-autophagy axis. Our findings indicate that SLC39A1 acts as a protective factor for the pulmonary epithelium in ALI/ARDS.

Results

SLC39A1 is elevated in ALI murine models and ARDS patients

To investigate alterations in zinc transporters expression in ALI, we analyzed the mRNA expression levels of 24 zinc transporters (including SLC39A1-SLC39A14 and SLC30A1-SLC30A10) in lung tissues of mice subjected to LPS instillation. Our results showed that Slc39a1 mRNA had a relatively high basal expression level and underwent the most significantly altered upon LPS treatment among the 24 zinc transporters (Fig. S1A, B). The upregulation of SLC39A1 was evident as early as 6 h and lasts for 24 h after LPS treatment (Fig. 1A, D). This upregulation was consistent across various ALI murine models established by cecal ligation and puncture (CLP) operation (Fig. 1B, E) and acid aspiration (Fig. 1C, F). Immunofluorescence and immunohistochemical staining further confirmed the increase of SLC39A1 levels in the lung tissues of these ALI models (Fig. 1G–I). Of particular interest, the elevation of SLC39A1 appeared notably prominent in AT2 cells, as evidenced by immunofluorescence staining (Fig. 1G, H) and RNA-seq data (GSE109913) (Fig. S1C, D). This elevation of SLC39A1 was also observed in primary rat AT2 cells, A549 and BEAS-2B cell lines exposed to LPS in a time-dependent manner (Fig. 1J, K; and Fig. S1E). SLC39A1 upregulation was found to depend on TLR4 signaling, a conclusion supported by the dose-dependent attenuation of SLC39A1 induction upon treatment with the TLR4-specific inhibitor TAK-242 (Fig. S1F, G). Furthermore, cycloheximide inhibition assays revealed that LPS stimulation significantly slowed the degradation rate of SLC39A1 protein compared to untreated cells (Fig. S1H, I). Together, these results indicate that the upregulation of SLC39A1 is mediated through both TLR4-dependent transcriptional activation and post-translational stabilization of the protein. Importantly, both immunohistochemical staining and qPCR analyses revealed elevated expression of SLC39A1 in lung tissues from ARDS patients compared to non-ARDS controls (Fig.1L, M; Table S1, S2), suggesting a potential role for SLC39A1 in ARDS pathogenesis.

Fig. 1. Increased SLC39A1 expression in AT2 cells in acute lung injury.

A–C The mRNA expression level of Slc39a1 in mouse lung tissues was determined by qPCR analysis following (A) intratracheal LPS instillation (2 mg/kg) (n = 3 per group), B CLP surgery (n = 5 per group), or (C) acid aspiration (n = 5 per group). D–F Western blot analysis determined SLC39A1 protein expression in lung tissues from mouse models of ALI induced by (D) LPS, E CLP surgery, or (F) acid aspiration (n = 3 or 4 per group). G, H Representative immunofluorescence images showing colocalization of SPC (AT2 cell marker, red) and SLC39A1 (green) in lung tissues from mice treated with (G) LPS or (H) CLP surgery. Yellow signal indicates colocalization (white arrows). Nuclei were counterstained with DAPI (blue). Scale bar, 10 μm. I Representative immunohistochemical (IHC) staining of SLC39A1 in lung tissues from mice treated with LPS (left panel) or CLP surgery (right panel). Red arrows indicate SLC39A1 protein localization. Scale bars, 100 or 10 μm. J, K SLC39A1 protein expression in (J) primary rat alveolar type II cells (pAT2) and (K) A549 cells treated with LPS (1 μg/mL) at indicated time points was assessed by western blot analysis. L Representative immunochemical staining of SLC39A1 in lung tissues of patients with or without ARDS. Scale bars, 50 μm. M The mRNA expression levels of SLC39A1 in the lungs of patients with or without ARDS were determined by qPCR (non-ARDS patients, n = 10; ARDS patients, n = 6). Data are presented as mean ± SD. A One-way ANOVA, followed by Tukey’s multiple comparisons test. B, C Unpaired two-tailed Student’s t-test. M Mann-Whitney U test (two-tailed, independent samples). Source data are provided as a Source Data file.

SLC39A1 controls the uptake of zinc in AT2 cells

To assess potential alterations in zinc homeostasis within AT2 cells during ALI, we employed Fluozin 3, AM, a specific fluorescence probe for zinc ions, to label intracellular zinc. Our fluorescence staining revealed an elevation in zinc levels specifically within AT2 cells, rather than endothelial cells, in the lung tissues of mice subjected to either LPS or CLP operations (Fig. 2A, B). A similar trend of increased zinc ions was also observed in live fluorescent staining and flow cytometry analysis of zinc ions in rat primary AT2 cells (pAT2) (Fig. 2C, D) or A549 cells (Fig. 2E, F) treated with LPS. To further elucidate the role of SLC39A1 in zinc uptake by AT2 cells, we employed specific siRNA targeting SLC39A1 and engineered lentiviruses containing SLC39A1 gene stable expression plasmids to knockdown or overexpress SLC39A1 (Fig. 2G, I). The live fluorescent staining results revealed that SLC39A1 knockdown diminished zinc uptake in A549 cells in response to LPS (Fig. 2H), whereas SLC39A1 overexpression had the opposite effect (Fig. 2J). Notably, the signal increase in zinc ions was significantly diminished upon addition of the zinc chelator N,N,N′,N′-Tetrakis (2-pyridylmethyl) ethylenediamine (TPEN), confirming the specificity of Fluozin 3,AM for zinc ions (Fig. 2J). Collectively, these findings suggest that SLC39A1 mediates zinc uptake in AT2 cells.

Fig. 2. SLC39A1 regulates zinc uptake in AT2 cells.

A, B Representative immunofluorescence images of lung tissues from mice treated with (A) LPS or (B) CLP surgery, showing zinc ions (FluoZin-3 AM probe, green), AT2 cells (SPC, red), and endothelial cells (CD31, purple). White arrows indicate colocalization of SPC and zinc signals in merged images (yellow). Nuclei were counterstained with DAPI (blue). Scale bar, 20 μm. C, E Representative live-cell fluorescent staining of intracellular zinc ions (probed by zinc-specific fluorescent probe FluoZin-3 AM, green) in (C) rat pAT2 or (E) A549 cells treated with LPS (1 μg/mL) for 9 h. Scale bars, 50 μm or 10 μm. D, F Representative flow cytometry analysis of intracellular zinc ions labeled with FluoZin-3 AM in (D) rat pAT2 or (F) A549 cells after 9-h LPS treatment. MFI, Mean fluorescence intensity. G Two SLC39A1-specific siRNAs were transfected into A549 cells for 48 h, followed by knockdown efficiency assessment via western blot analysis. siNC, negative control siRNA. H Representative live-cell fluorescence images showing intracellular zinc ions (stained with the zinc-specific fluorescent probe Fluozin-3 AM, green) in SLC39A1-knockdown A549 cells treated with LPS for 9 h. Scale bars, 50 μm or 10 μm (as indicated). I SLC39A1 protein expression in SLC39A1-overexpressing A549 cells was determined by western blot. J Representative fluorescence images of intracellular zinc ions (labeled with the zinc-specific probe Fluozin-3 AM, green) in SLC39A1-overexpressing A549 cells treated with LPS, in the presence or absence of the zinc chelator TPEN (1 μM). Scale bars, 50 μm or 10 μm (as indicated).

AT2-specific depletion of Slc39a1 promotes ALI

To probe for the relevance of AT2 SLC39A1 in ALI, we generated tamoxifen-inducible, AT2-specfic Slc39a1 deficient mice (referred to as Slc39a1 icKO) by crossing mice homozygous for the floxed Slc39a1 allele with Sftpc-CreERT2 mice (Fig. 3A). The expression of SLC39A1 in AT2 cells was markedly reduced in lung tissues of Slc39a1 icKO mice compared to Control mice (Slc39a1^flox/flox mice without Sftpc-CreERT2), as confirmed by immunofluorescence staining and immunochemical staining (Fig. 3B, C). In contrast, SLC39A1 expression in AT1 cells (labeled with the marker HOPX) or endothelial cells (labeled with CD31) was significantly lower than that in AT2 cells under control conditions (Fig. S2A, B). In line with the role of SLC39A1, zinc uptake in AT2 cells was significantly decreased in Slc39a1 icKO mice compared to Control mice following LPS exposure, evidenced by decreased fluorescent signal of FluzoZin, 3AM (Fig. 3D), and mRNA expression of intracellular zinc reservoir metallothionein-2 (Mt2) (Fig. 3E). Slc39a1 icKO mice displayed more severe lung injury than Control mice, manifested by increased cell count and protein concentration in BALF (Fig. 3F, G) and pathological deterioration (Fig. 3H, I). Furthermore, elevated IL-6 levels in BALF were observed in Slc39a1 icKO mice following LPS exposure (Fig. S2C). In an in vitro lung epithelial cell model, it was confirmed that LPS-induced elevation of IL-8 secretion in BEAS-2B cell supernatant was exacerbated by siRNA-mediated knockdown of SLC39A1 (Fig. S2D–G). Moreover, intraperitoneal administration of the zinc chelator TPEN resulted in increased mortality (Fig. S3A, B) and exacerbated pulmonary damage in LPS-challenged mice (Fig. S3C–I). Similarly, TPEN promoted IL-8 secretion and enhanced oxidative stress in LPS-treated BEAS-2B cells (Fig. S4A–C). These results indicate that disruption of SLC39A1 in AT2 cells impairs zinc uptake and exacerbates lung injury.

Fig. 3. AT2-specific Slc39a1 deficiency aggravates LPS-induced ALI.

A Inducible conditional knockout (icKO) mice (Slc39a1^flox/flox; Sftpc-CreERT2^+/−, denoted Slc39a1 icKO) and littermate controls (Slc39a1^flox/flox; Sftpc-CreERT2^−/−, denoted Control) received intraperitoneal tamoxifen injections for 5 consecutive days to induce AT2 cell-specific deletion of Slc39a1. After a 14-day washout period to eliminate tamoxifen effects, mice underwent intratracheal instillation of LPS (2 mg/kg) or vehicle. Lung tissues and bronchoalveolar lavage fluid (BALF) were subsequently collected for lung injury assessment. B Representative immunofluorescence staining of AT2 cells (labeled with AT2 marker SPC, green) and SLC39A1 (red) in lung tissues of Slc39a1 icKO or Control mice following tamoxifen-induced recombination. White arrows denote colocalization of SPC and SLC39A1. Nuclei were counterstained with DAPI (blue). Scale bar, 10 μm. C SLC39A1 protein expression in lung tissues of Slc39a1 icKO and Control mice following tamoxifen-induced recombination, assessed by immunohistochemical staining. Scale bars, 100 μm or 10 μm (as indicated). D Representative immunofluorescence staining of AT2 cells (labeled with AT2 marker SPC, green) and intracellular zinc ions (labeled with the zinc-specific probe Fluozin-3 AM, green) in lung tissues of Slc39a1 icKO or Control mice following LPS instillation. White arrows denote colocalization of SPC and intracellular zinc ions. Nuclei were counterstained with DAPI (blue). Scale bar, 10 μm. E Mt2 mRNA expression in lung tissues of tamoxifen-induced Slc39a1 icKO and Control mice, with or without LPS treatment, was quantified by qPCR (n = 3 mice per group). F, G Total cell counts (F) and protein concentrations (G) in BALF from experimental mice groups were quantified (n = 3 or 7 mice per group). H Representative H&E staining showing alveolar pathology and inflammatory cell infiltration in lungs of Slc39a1 icKO and control mice. Scale bar, 100 μm. I Lung injury scores were quantified from histology (n = 5 mice per group). Data are presented as mean ± S.D. E–G and I One-way ANOVA, followed by Tukey’s multiple comparisons test. Source data are provided as a Source Data file.

AT2-specific overexpression of SLC39A1 offers therapeutic benefits for ALI

To further evaluate the biological function of AT2 SLC39A1 in ALI, we utilized AAV5-SPC-Slc39a1-ZsGreen (referred to as AAV-Slc39a1) to specifically overexpress SLC39A1 in AT2 cells via intratracheal instillation²¹. AAV5-SPC-Vector-ZsGreen (referred to as AAV-Control) served as the control with an equal virus titer (Fig. 4A). Immunofluorescence staining, western blot and immunochemical staining confirmed the AT2-specific overexpression of SLC39A1 in mice treated with AAV5-Slc39a1 (Fig. 4B–E). qPCR results indicated a further upregulation of Mt2 mRNA expression in the lung tissues of mice treated with AAV-Slc39a1, suggesting increased zinc uptake (Fig. 4F). Mice administrated AAV-Slc39a1 showed lower IL-6 levels, cell counts and protein concentrations in BALF than those receiving AAV-Control following LPS exposure (Fig. 4G–I). Furthermore, AAV-Slc39a1 treatment improved lung pathological changes (Fig. 4J, K). To further explore the therapeutic potential of zinc, we examined whether zinc supplementation could alleviate ALI. In contrast to zinc chelation with TPEN (as shown in Figs. S3, S4), the administration of ZnSO₄ significantly ameliorated lung injury in both LPS-challenged mice (Fig. S5) and LPS-treated BEAS-2B cells (Fig. S6). Notably, although zinc supplementation reduced lung injury in wild-type mice, this protective effect was completely abolished in Slc39a1 icKO mice (Fig. 5A–E), indicating that SLC39A1-mediated zinc uptake is essential for protection against ALI.

Fig. 4. AT2-specific Slc39a1 overexpression ameliorates LPS-induced ALI.

A C57BL/6 J mice received intratracheal instillation of 6.4 × 10¹⁰ vector genomes (vg) of either AAV5-SPC-Vector-ZsGreen (AAV-Control) or AAV5-SPC-Slc39a1-ZsGreen (AAV-Slc39a1). Twenty-one days post-instillation, mice underwent intratracheal administration of LPS (2 mg/kg) or vehicle control. Lung tissues and bronchoalveolar lavage fluid (BALF) were collected 1 day post-LPS exposure for assessment of lung injury severity. B Representative fluorescence staining of AT2 cells (SPC, red) and AAV-SPC-ZsGreen (green) in murine lung sections. White arrows indicate colocalization of SPC signal and AAV5-SPC-Slc39a1-ZsGreen. Nuclei counterstained with DAPI (blue). Scale bar, 20 μm. C Murine lung sections immunostained for AT2 cells (SPC, red) and SLC39A1 (purple). Arrows denote SPC/SLC39A1 co-localization. DAPI nuclear counterstain (blue). Scale bar, 10 μm. D, E SLC39A1 expression in AAV-transduced lungs. D Western blot analysis. E Representative IHC staining (Scale bars, 100 μm, main; 10 μm, insets). Groups: AAV-Control vs. AAV-Slc39a1. F qPCR quantification of Mt2 transcripts in lungs from AAV-Control-transduced vs. AAV-Slc39a1-transduced mice with or without LPS challenge (n = 3 independent biological replicates). G–I IL-6 concentrations (G), total cell counts (H), and protein concentrations (I) in bronchoalveolar lavage fluid (BALF) from AAV-Control-transduced versus AAV-Slc39a1-transduced mice with or without LPS challenge (n = 4, 6, or 10 mice per group, as indicated in panels). J Representative H&E staining showing alveolar pathology and inflammatory cell infiltration in lungs of AAV-Control-transduced and AAV-Slc39a1-transduced mice with or without LPS challenge. Scale bar, 100 μm. K Corresponding lung injury scores quantified from histology (n = 5 mice per group). Data are presented as mean ± S.D. F–I, K One-way ANOVA, followed by Tukey’s multiple comparisons test. Source data are provided as a Source Data file.

Fig. 5. SLC39A1 governs the protective effects of zinc supplementation in ALI.

A Slc39a1 icKO or Control mice received 15-day ZnSO₄ (25 mM) supplemented or standard drinking water, followed by intratracheal LPS instillation (2 mg/kg). Lung tissues and bronchoalveolar lavage fluid (BALF) were collected 24 h post-exposure for assessment of lung injury severity. B Representative H&E staining illustrating alveolar pathology and inflammatory cell infiltration in murine lungs, as schematized in panel (A). Scale bar, 200 μm or 50 μm (as indicated). C Corresponding lung injury scores quantified from histology (n = 5 mice per group). D, E Total cell counts (D) and protein concentrations (E) in BALF from experimental groups; corresponding to panel (A) schematic (n = 7 mice per group). Data are presented as mean ± S.D. C–E One-way ANOVA, followed by Tukey’s multiple comparisons test. Source data are provided as a Source Data file.

AT2 SLC39A1 controls the activation of autophagy machinery

Recently, zinc has emerged as a factor associated with improved autophagy machinery^22,23, promoting our investigation into the potential impact of SLC39A1 on autophagy. Transmission electron microscopy (TEM) results revealed an accumulation of autophagic vacuole structures in lung tissues of mice upon LPS treatment (Fig. 6A). qPCR analysis showed a significant increase in mRNA expression levels of various autophagy-related genes, such as Ulk1, Atg3, Atg4b, Atg4d, Atg5, Becn1, Atg7, Lc3b, Atg10, Atg12, Atg13, Atg14 and Atg16l1 in response to LPS treatment (Fig. 6B). A similar trend was observed in the protein expression levels in lung tissues of mice (Fig. 6C), rat primary AT2, A549 and BEAS-2B cells (Fig. S7A–C) exposed to LPS. RNA-seq analysis further revealed that the autophagic response was blunted in the lung tissues of Slc39a1 icKO mice compared with controls following LPS challenge (Fig. 6D). AT2-specific depletion or overexpression of SLC39A1 either suppressed or further promoted the upregulation of ULK1, ATG7, ATG5 and LC3B in LPS-treated lung tissues of mice (Fig. 6E, F) or A549 cells (Fig. 6G, H). LPS treatment also induced the accumulation of autophagic vacuoles in BEAS-2B cells (Fig. S7D). Furthermore, inhibition of vacuolar H⁺-ATPase (V-ATPase) with bafilomycin A1 (BafA1) further enhanced the accumulation of LC3B-II in these cells (Fig. S7E). Consistent with this, BafA1 treatment increased the number of LC3-EGFP puncta in BEAS-2B cells stably overexpressing LC3-EGFP, whereas treatment with the PI3K inhibitor wortmannin (Wort) reduced LC3-EGFP puncta formation (Fig. S7F, G), suggesting that LPS-induced upregulation of LC3B-II reflects enhanced autophagic flux rather than impaired autophagic degradation. These results indicate that SLC39A1 plays a key role in regulating autophagy activation during ALI.

Fig. 6. SLC39A1 orchestrates zinc-mediated autophagy in LPS-induced ALI.

A Representative transmission electron microscopy (TEM) images of lung tissues from mice treated with vehicle or LPS (2 mg/kg) for 24 h. Yellow arrows denote autophagic vacuoles. Scale bar, 500 nm. B, C Temporal profiling of autophagy genes in LPS-challenged lungs of mice. B qPCR analysis of mRNA expression levels. (C) Western blot analysis of protein expression levels. Time points: 0, 6, 12, and 24 h (n = 3 mice per group). D RNA-seq analysis of lung tissues from Control vs. Slc39a1 icKO mice ± LPS (24 h) revealed differentially expressed genes (DEGs) enriched in autophagy-related pathways (n = 3 mice per group). E, F Western blot analysis of autophagy-related proteins (ULK1, ATG5, ATG7, LC3B-I/II) in lungs of: (E) Slc39a1 icKO vs Control mice, (F) AAV-Slc39a1 vs AAV-Control mice, following ± LPS treatment. G, H Western blot analysis of autophagy-related proteins (ULK1, ATG5, ATG7, LC3B-I/II) in (G) SLC39A1-knockdown and (H) SLC39A1-overexpressing A549 cells after ± LPS treatment. Data are presented as mean ± S.D. B One-way ANOVA, followed by Tukey’s multiple comparisons test. Source data are provided as a Source Data file.

The SLC39A1-zinc-autophagy axis defines a critical pathway contributing to LPS-induced ALI

To determine the role of SLC39A1-mediated autophagy activation in ALI, Lc3b knockout mice (Lc3b^−/−) and their littermate controls were utilized. The Lc3b^−/− mice exhibited almost undetectable expression of LC3B protein (Fig. 7A), leading to increased lung injury degrees (Fig. 7B–G) and a higher mortality rate (Fig. 7H) compared to their littermate controls when exposed to LPS. Notably, zinc supplementation failed to alleviate lung injury in Lc3b^−/− mice following LPS exposure (Fig. 7I–M), indicating that zinc supplementation primarily ameliorates LPS-induced ALI through activation of the autophagy machinery. Mirroring the outcomes of SLC39A1 modulation, suppression or enhancement of autophagy was achieved by modulating zinc uptake (via TPEN or ZnSO4) in lung tissues of LPS-treated mice (Fig. S8A–D). To further clarify the hierarchical relationship between SLC39A1 and autophagy, we employed a combination of AAV vectors for Lc3b knockdown and Slc39a1 icKO mice (Fig. 7N). The efficiency of Lc3b knockdown was validated in the murine lung epithelial cell line MLE-12 (Fig. S9). Strikingly, our data demonstrate that both Slc39a1 icKO and AAV-mediated AT2-specific Lc3b knockdown independently exacerbated lung injury; however, administration of AAV-shLc3b to Slc39a1 icKO mice did not further potentiate injury phenotypes beyond those observed with either intervention alone (Fig. 7O–R). This genetic epistasis indicates that SLC39A1 and autophagy operate within the same linear pathway, with SLC39A1 functioning upstream of autophagy activation.

Fig. 7. Zinc-activated autophagy via SLC39A1 confers protection in LPS-induced ALI.

A LC3B protein expression in lungs of WT versus Lc3b^−/− mice post-LPS challenge (1 mg/kg, 24 h) was analyzed by western blot analysis (n = 4 mice per group). B-D Total cell count (B), protein concentrations (C), and MPO contents (D) in the BALF of WT versus Lc3b^−/− mice post-LPS challenge were determined (n = 4 mice per group). E Representative H&E staining illustrating alveolar pathology and inflammatory cell infiltration in lungs of WT versus Lc3b^−/− mice post-LPS challenge. Scale bar, 200 μm or 50 μm (as indicated). F Corresponding lung injury scores quantified from histology (n = 5 mice per group). G Lung edema in WT vs. Lc3b^−/− mice post-LPS challenge was quantified by wet-to-dry weight ratio (n = 5 mice per group). H The mortality of WT vs. Lc3b^−/− mice post-LPS challenge was record (n = 21 mice in WT + LPS group and n = 18 in Lc3b^−/− +LPS group). I Lc3b^−/− mice received drinking water ± ZnSO₄ (25 mM) for 15 d, followed by intratracheal LPS administration (1 mg/kg). Lung tissues and BALF were collected 24 h post-LPS challenge for lung injury assessment. J–M Quantification of lung injury indices corresponding to panel (I): (J) Lung edema (wet-to-dry weight ratios); (K) BALF total cell counts; (L) BALF protein concentrations; and (M) BALF MPO contents (n = 5 mice per group). N After sequential intratracheal instillation of AAV-shNC or AAV-shLC3 followed by tamoxifen induction, Slc39a1 icKO or Control mice received vehicle or LPS via intratracheal administration. Lung tissues and BALF were collected 24 h post-exposure for assessment of lung injury severity. O-R Quantification of lung injury indices corresponding to panel (N): (O) alveolar pathology determined by H&E staining; (P) Corresponding lung injury scores quantified from histology; (Q) Total cell counts in BALF; and (R) Protein concentrations in BALF (n = 5 mice per group). Data are presented as mean ± SD. B–D, F, G, J–M Unpaired two-tailed Student’s t-test. P–R One-way ANOVA, followed by Tukey’s multiple comparisons test. Source data are provided as a Source Data file.

SLC39A1-mediated zinc uptake controls autophagy through activating transcription factor MiT/TFE family

Given that SLC39A1-mediated zinc uptake controls the activation of autophagy machinery at the transcriptional level during ALI, we next shifted our focus to examining changes in the critical autophagic transcription factor MiT/TFE family, which is known to play a crucial role in transcriptionally regulating autophagy machinery^24–27. This family comprises four members, TFEB, TFE3, MITF and TFEC, all sharing conserved basic helix-loop-helix (bHLH) and leucine zipper (Zip) domains²⁸. The mRNA expression levels of Tfeb, Tfe3, and Mitf were upregulated over time in the lung tissues of mice exposed to LPS, whereas Tfec remained unchanged (Fig. 8A). Similarly, protein expression levels of TFEB, TFE3, and MITF exhibited an increasing trend in both lung tissues of mice and various epithelial cells exposed to LPS (Fig. 8B; Fig. S10A–C). SLC39A1 and zinc uptake were proved to directly influence the activation of MiT/TFE family. AT2-specific depletion or overexpression of SLC39A1 either suppressed or further promoted the upregulation of TFEB, TFE3, and MITF in LPS-treated mice (Fig. 8C, D) or A549 cells (Fig. S10D, E). Similarly, zinc depletion with TPEN diminished, while zinc supplementation increased, the expression levels of TFEB, TFE3 and MITF in mice (Fig. S11A, B) or BEAS-2B cells exposed to LPS (Fig. S11C, D). Following LPS treatment, TFEB, TFE3, and MITF translocated to the nucleus in BEAS-2B cells (Fig. S11E). This translocation was significantly inhibited by the knockdown of SLC39A1 or zinc chelator TPEN, and enhanced by zinc supplementation (Fig. 8E and Fig. S11E), indicating that zinc can modulate the function of the MiT/TFE family. Additionally, it has been reported that inhibition of mTOR results in the dephosphorylation and subsequent nuclear translocation of the MiT/TFE family²⁶. Interestingly, the knockdown of SLC39A1 or zinc chelator TPEN failed to affect the mTOR inhibitor Torin1-induced nuclear translocation of the MiT/TFE family (Fig. S12A, B), indicating that mTOR signaling may not be involved in the activation of the MiT/TFE family by SLC39A1-mediated zinc uptake.

Fig. 8. SLC39A1-mediated zinc influx activates autophagy via MiT/TFE family in ALI.

A, B C57BL/6 J mice were intratracheally administered LPS (2 mg/kg), and lung tissues were collected at 6, 12, and 24 h post-administration. Expression levels of MiT/TFE family transcription factors were assessed by (A) qPCR (n = 3 mice per group) or (B) western blot (n = 3 mice per group). C, D Protein expression levels were analyzed by western blot in Slc39a1 icKO mice and their controls (C), as well as AAV-Slc39a1 mice and AAV-Control mice (D), with all groups exposed to either Vehicle or LPS as indicated. E BEAS-2B cells stably expressing indicated protein were transfected with SLC39A1-targeting siRNA or negative control siRNA (siNC) for 48 h, and fluorescence signals were quantified 24 h post-treatment with LPS. AcGFP signals (green) and DAPI-stained nuclei (blue) are shown; scale bar, 25 μm. F Schematic diagram illustrating the purification process of GST and its fusion proteins in HEK293T cells and the subsequent zinc-resin pull-down assay. G, H Western blot analysis (G) and commassie blue staining (H) were applied to determine the purity and specificity of indicated protein (red arrows). The asterisk marks the cleaved GST moiety. I Western blot analysis of indicated protein eluted from resin in the presence or absence of zinc. J Schematic diagram of the construction of stable A549 cell lines overexpressing indicated protein; followed by inductively coupled plasma mass spectrometry (ICP-MS) analysis of zinc content in the pulled-down GST-fusion proteins after treatment with or without LPS. K Western blot analysis of indicated protein expression in the A549 cell lines presented in (J). L ICP-MS analysis of zinc content in the pulled-down GST-fusion proteins (TFE3, TFEB, MITF) after treatment with or without LPS in the A549 cell lines presented in (J) (n = 5 replicates for each group). M Crystal structure of the human TFE3 bHLH-LZ domain (residues 359–399) in complex with Zn²⁺ (PDB ID: 7F09). Data are presented as mean ± SD. A One-way ANOVA, followed by Tukey’s multiple comparisons test. L Unpaired two-tailed Student’s t-test. Source data are provided as a Source Data file.

Zinc is known to bind to and modulate the activity of various proteins²⁹, prompting us to hypothesize that zinc may directly interact with and activate MiT/TFE family transcription factors. To test this, we expressed and purified GST, GST-TFE3, GST-TFEB, and GST-MITF proteins in HEK293T cells using the pEBG eukaryotic expression system coupled with GSH magnetic beads (Fig. 8F–H). Using Zn²⁺-IMAC chromatography, we assessed the zinc-binding ability of these proteins. The results demonstrated that GST-TFE3, GST-TFEB, and GST-MITF, but not the GST control, specifically bound to zinc-charged resin, indicating direct zinc binding to TFE3, TFEB, and MITF (Fig. 8I). We further established stable A549 cell lines expressing GST-TFE3, GST-TFEB, or GST-MITF. Following treatment with LPS or vehicle, purified proteins were subjected to ICP-MS analysis, which revealed that LPS stimulation significantly enhanced zinc binding to TFE3, TFEB, and MITF (Fig. 8J–L). Additionally, the crystal structure of the human TFE3 HLH-Lz domain (PDB: 7F09), resolved at 2.6 Å, revealed two zinc ions coordinated within this domain by residues such as R365 and E368³⁰, suggesting the presence of multiple zinc-binding sites in TFE3 (Fig. 8M). Collectively, these results provide strong evidence that zinc can directly bind to members of the MiT/TFE protein family.

Loss of Tfe3 aggravates LPS-induced ALI

To further confirm the involvement of the MiT/TFE family in ALI, Tfe3 knockout (Tfe3^−/y) mice were generated. Deletion of Tfe3 (Fig. 9A), similar to the deletion of Lc3b, led to worsened lung injury (Fig. 9B–F) and reduced activation of autophagy machinery (Fig. 9G). Consistently, the knockdown of TFE3, TFEB, or MITF in BEAS-2B cells (Fig. S13A) led to suppressed activation of the autophagy machinery (Fig. S13B), increased IL-8 levels (Fig. S13C), and elevated oxidative stress markers GCLM and HMOX1 (Fig. S13D). Importantly, the aggravated lung injury resulting from Tfe3 loss could not be reversed by zinc supplementation, indicating that the protective role of zinc is dependent on TFE3 (Fig. 9H–L). These findings suggest that zinc-mediated activation of the TFE3 is crucial for autophagy machinery and mitigation of ALI.

Fig. 9. SLC39A1-mediated zinc influx attenuates LPS-induced ALI by regulating TFE3-dependent autophagy.

A TFE3 protein expression in lung tissues from Tfe3^−/y mice and controls treated with LPS (2 mg/kg) was analyzed by western blot (n = 3 mice per group). B Representative H&E-stained sections show alveolar pathology and inflammatory cell infiltration in lungs from WT and Tfe3^−/y mice under Vehicle or LPS treatment. Scale bar, 100 μm. C Corresponding lung injury scores quantified from histology (n = 5 mice per group). D-F Total cell count (D), protein concentration (E), and IL-6 levels (F) in BALF from WT and Tfe3^−/y mice after Vehicle/LPS treatment were quantified (n = 4 or 5 mice per group). G Protein expression of ULK1, ATG5, ATG7, and LC3B-I/II in lungs of WT or Tfe3^−/y mice after 24 h LPS treatment was analyzed by western blot (n = 3 mice per group). H Tfe3^−/− mice received drinking water ± ZnSO₄ (25 mM) for 15 d, followed by intratracheal LPS administration (2 mg/kg). Lung tissues and BALF were collected 24 h post-LPS challenge for lung injury assessment. I-L Quantification of lung injury indices corresponding to panel (H): Representative H&E-stained sections show alveolar pathology and inflammatory cell infiltration in lungs (I); Corresponding lung injury scores quantified from histology (J); BALF total cell counts (K); and BALF protein concentrations (L) (n = 5 mice per group). C–F One-way ANOVA, followed by Tukey’s multiple comparisons test. J–L Unpaired two-tailed Student’s t-test. Source data are provided as a Source Data file.

SLC39A1 impedes AT2 apoptosis and pyroptosis by promoting zinc-mediated mitophagy

Activation of cell death pathways is known to contribute to epithelial injury, a key feature of ALI/ARDS in humans^31,32. Since mitochondria can initiate cell death when impaired or dysfunctional, their elimination through autophagy, also known as mitophagy, has been shown to inhibit cell death³³. Based on this, we hypothesize that SLC39A1 may induce mitophagy to suppress cell death in the lung epithelium during ALI. TEM analysis showed that the absence or overexpression of Slc39a1 exacerbated or reduced the accumulation of damaged mitochondria in the lung tissues of mice exposed to LPS (Fig. 10A, B). Similarly, mitochondrial proteins ATP5B and TIM23 increased in Slc39a1-deficient mice (Fig. 10C) but decreased in Slc39a1-overexpressing mice (Fig. 10D). This trend was also observed in SLC39A1-knockdown or overexpressing A549 cells (Fig. S14A, B). To determine whether SLC39A1 modulates cell death pathways, we performed RNA sequencing analysis. Whereas LPS induced a significant apoptotic and pyroptotic response in control lungs, this response was even more pronounced in Slc39a1 icKO mice (Fig. 10E). Immunofluorescence staining using TUNEL and cleaved-Caspase 3 antibody indicated increased apoptosis with Slc39a1 depletion, whereas its overexpression reversed these effects (Fig. 10F, G). Similarly, western blot analysis results demonstrated that the absence or overexpression of Slc39a1 led to alterations in the expression levels of PARP/cleaved PARP both in lung tissues of mice (Fig. 10H, I) and in A549 cells (Fig. S14C, D) treated with LPS. In addition to its role in apoptosis, SLC39A1 significantly regulates pyroptosis in lung epithelial cells during ALI. Depletion of Slc39a1 further upregulated the expression of pyroptotic markers, including LDH in BALF (Fig. 10J), as well as NLRP3, ASC, GSDME, and GSDMD-NT in lung tissues (Fig. 10L) and A549 cells (Fig. S14E). Conversely, overexpression of Slc39a1 suppressed the expression of these pyroptotic markers both in vivo and in vitro (Fig. 10K, M; Fig. S14F). These results indicate that SLC39A1 expression is positively correlated with mitophagy but negatively correlated with apoptosis and pyroptosis. Overall, our findings suggest that SLC39A1 triggers a zinc-MiT/TFE-autophagy axis to inhibit apoptosis and pyroptosis in the lung epithelium, thereby ameliorating acute lung injury (Fig. 10N).

Fig. 10. SLC39A1 enhances mitophagy to inhibit apoptotic and pyroptotic cell death in AT2 cells during ALI.

A, B Representative TEM images of lungs from (A) Slc39a1 icKO, (B) AAV-Slc39a1, and corresponding control mice after 24 h LPS treatment. Red arrows denote autophagic vacuoles, yellow arrows mark aberrant mitochondria, and green arrows indicate lamellar bodies in AT2 cells. Scale bars, 1 μm or 500 nm (as indicated). C, D Expression of ATP5B and TIM23 in lung tissues from (C) Slc39a1 icKO, D AAV-Slc39a1, and corresponding control mice treated with Vehicle/LPS was analyzed by western blot. E RNA-seq analysis of lung tissues from Control mice and Slc39a1 icKO mice after 24 h Vehicle/LPS treatment revealed differential gene enrichment in apoptosis and pyroptosis pathways (n = 3 per group). F, G Representative fluorescence staining for (F) TUNEL (green) and (G) cleaved caspase-3 (red) in lung tissues from Slc39a1 icKO, AAV-Slc39a1, and control mice after Vehicle/LPS treatment. Nuclei counterstained with DAPI (blue). Scale bar, 50 μm. H, I Expression of PARP and cleaved-PARP (cl-PARP) in lungs from (H) Slc39a1 icKO, (I) AAV-Slc39a1, and control mice after Vehicle/LPS treatment was analyzed by western blot. J, K LDH activity in BALF from Slc39a1 icKO, AAV-Slc39a1, and control mice treated with Vehicle or LPS (n = 5 or 10 mice per group). L, M Expression of NLRP3, ASC, GSDME and GSDMD-NT in lungs from (H) Slc39a1 icKO, (I) AAV-Slc39a1, and control mice after Vehicle/LPS treatment was analyzed by western blot. N Schematic diagram illustrating that SLC39A1-mediated zinc uptake suppresses apoptosis and pyroptosis in AT2 cells by activating the MiT/TFE family and promoting the autophagic clearance of damaged mitochondria. J, K One-way ANOVA, followed by Tukey’s multiple comparisons test. Source data are provided as a Source Data file.

Discussion

Despite considerable advancements in both clinical and basic research on ALI/ARDS, the quest for specific therapeutic drugs remains challenging³⁴. The current study presented here unveiled several key findings: i) a comprehensive evaluation of the dynamic alterations in zinc transporters in ALI, highlighting the upregulation of AT2 SLC39A1 as a novel mechanism for epithelial protection; ii) providing evidence suggesting that AT2 SLC39A1 triggers the transcriptional activation of mitophagy to eliminate aberrant mitochondria by enhancing zinc uptake; iii) an illustration of how zinc directly binds to the MiT/TFE family, initiating the transcriptional activation of mitophagy to inhibit AT2 apoptosis and pyroptosis. This study is the first to establish a direct connection between the zinc transporter SLC39A1 and ALI, indicating that targeting AT2 SLC39A1 could offer a promising avenue for preventing or alleviating ALI. While Major et al. recently reported an intrinsic protective mechanism mediated by endothelial aryl hydrocarbon receptor activity in influenza virus infection-induced ALI³⁵, our study emphasizes the upregulation of zinc transporter SLC39A1 in alveolar type II epithelial cells as an additional intrinsic protective mechanism in lung epithelium, together shed the light on the role of non-immune cells to promote host protection in ALI/ARDS.

The SLC39A1 gene was first cloned in 1999 by Lioumi et al³⁶. The SLC39A1 protein is ubiquitously expressed and shares a high homology, ~90%, between human and mice. Its biological significance has been extensively studied in cancer research. For instance, SLC39A1 has facilitated rapid zinc uptake in prostate cancer and leukemia cells^37–39. Additionally, SLC39A1-positive fibroblasts have been known to promote zinc uptake and transfer it to lung cancer cells, leading to chemotherapy resistance⁴⁰. Furthermore, SLC39A1, possibly in conjunction with SLC39A3, has been implicated in regulating platelet zinc homeostasis and function⁴¹. To our best knowledge, our study firstly identified that SLC39A1 as the crucial zinc transporter in ALI. Intriguingly, SLC39A1-mediated zinc uptake may serve as a universal protective mechanism in various ALI models, including those induced by LPS, acid aspiration, and CLP operation, indicating that targeting SLC39A1 could hold promise for treating the highly heterogeneous ALI/ARDS in diverse scenarios. Intriguingly, SLC39A1-dependent modulation of zinc uptake was observed exclusively under LPS challenge, whereas overexpression alone exerted minimal impact. This finding aligns with growing evidence that phosphorylation serves as a pivotal regulatory mechanism for zinc transporters. For example, Taylor et al.⁴² reported that CK2-mediated phosphorylation of the endoplasmic reticulum zinc transporter SLC39A7 triggers zinc release, downstream signaling activation, and cell migration-all of which are abolished upon CK2 inhibition or mutagenesis of SLC39A7. Moreover, in silico analyses predict that all 24 human zinc transporters possess conserved phosphorylation sites, suggesting that post-translational modification may represent a widespread mechanism governing their function⁴³. Based on these insights, we hypothesize that LPS enhances SLC39A1-mediated zinc transport through a dual regulatory scheme comprising transcriptional induction and post-translational modification-likely involving phosphorylation. Although our data are consistent with this model, the precise molecular details underlying this regulation require further elucidation.

Historically, zinc has been hailed as vital for the function of the immune system, earning its reputation as a guardian of the immune response⁴⁴. Zinc supplementation ameliorated endotoxin-induced murine lung injury by reducing neutrophil recruitment and activity⁴⁵, whereas dietary zinc deficiency potentiated ventilator-induced lung injury in mice⁴⁶. Zinc was previously thought to inhibit phosphatases in a nonspecific manner, leading to sustained activation of kinases such as ERK and Akt⁴⁷. Our research unveiled that the pivotal role of SLC39A1 in facilitating zinc uptake in lung epithelial cells. Deficiency in AT2 SLC39A1 or treatment with the zinc chelator TPEN significantly reduced zinc uptake, consequently heightening lung injury both in vivo and in vitro. Conversely, overexpression of AT2-specific SLC39A1 or zinc supplementation improved lung injury outcomes. Overall, our study underscores the importance of maintaining zinc homeostasis in lung epithelial cells, complementing the existing understanding of zinc as a critical regulator in immune cell function⁴⁴.

The role of zinc as a positive regulator of autophagy has been supported by a growing body of evidence^22,23. For instance, SLC39A7 was responsible for regulating zinc uptake within cardiomyocyte mitochondria, activating PINK1/Parkin-mediated mitophagy and mitigating myocardial reperfusion injury⁴⁸. Additionally, the zinc-ERK-Beclin 1 signaling pathway was implicated in the above process⁴⁹. Furthermore, SLC39A1 had been shown to interact with key proteins involved in mitochondrial division and calcium transport, thereby modulating zinc uptake in mitochondria and facilitating the removal of damaged mitochondria through mitophagy in HeLa cells⁵⁰. Our study reveals that AT2 SLC39A1-mediated zinc uptake primarily triggers the transcriptional activation of autophagy machinery, particularly mitophagy. The elimination of dysfunctional mitochondria via mitophagy may result in decreased apoptosis and pyroptosis in AT2 cells, thereby contributing to the preservation of lung epithelial integrity and the mitigation of lung injuries. Autophagy is considering as a double-edged sword in various human diseases. It is reported that certain pulmonary pathogens can exploit the autophagic process to evade immune surveillance and enhance their replication, thereby promoting pathogen proliferation and aggravating inflammation⁵¹. Nevertheless, the prevailing view in the field, strongly supported by findings from our study using Lc3b^−/− or Tfe3^−/y mice, indicates that autophagy acts predominantly as a protective mechanism in ALI. We hypothesize that a plausible explanation for this apparent paradox lies in the dual role of autophagy: while its impairment may restrict pathogen replication, it simultaneously weakens the host’s capacity to mitigate inflammation, oxidative stress, and related signaling cascades, ultimately leading to a more severe ALI phenotype.

Building upon our findings regarding the role of AT2 SLC39A1-mediated zinc uptake in transcriptionally activating autophagy at the transcriptional level, we aim to delve deeper into the mechanisms through which zinc transcriptionally triggers mitophagy. Recent research had unveiled the capacity of zinc to enhance autophagic flux and lysosomal function by activating TFEB in glioma cells. TFEB is recognized as a powerful regulator of autophagy machinery and lysosomal biogenesis²⁵. TFEB can be upregulated and activated by LPS, subsequently leading to the expression of downstream factors ATG5/ATG9B/BNIP3 in dendritic cell⁵². Moreover, TFEB regulated mitophagy process to adapt to metabolic stress in pancreatic beta cells⁵³, or in conjunction with TFE3, influencing the muscle mass in amyotrophy by regulating mitophagy⁵⁴. Additionally, TFEB, TFE3 and MITF might facilitate Parkin-mediated mitochondrial clearance and concurrently be regulated by PINK1/Parkin⁵⁵. Our results suggested that TFEB, TFE3 and MITF, besides TFEC, were upregulated in the lung tissues of mice and AT2 cells in response to LPS. Based on these observations, we hypothesize that zinc might directly bind to TFEB/TFE3/MITF and activate autophagy machinery at the transcriptional level. This hypothesis stems from the known ability of metal ions to bind to specific proteins and regulate essential biological processes^56–58. As anticipated, our findings establish a causal relationship between AT2 SLC39A1 and its mediated zinc uptake and the expression and activation of TFEB, TFE3, and MITF. Furthermore, zinc-resin pull-down and ICP-MS analyses confirmed a direct interaction between these transcription factors and zinc. Together, these results indicate that the MiT/TFE family functions as a downstream effector of zinc uptake mediated by AT2 SLC39A1.

The current study has several limitations that should be acknowledged. First, the mechanism underlying the predominant upregulation of SLC39A1 in AT2 cells remains unclear and warrants further investigation. Second, zinc’s capacity to bind nearly 10% of the human proteome underscores its extensive involvement in cellular signaling and biological complexity²⁹. Although more than 20 transcription factors (including the MiT/TFE family, FOXO1/3, and CREB) have been linked to autophagy^24,25,27,59, the extent and manner in which zinc modulates these factors remain largely unexplored. Third, while our results indicate that zinc exerts protective effects primarily through autophagy activation, we cannot rule out contributions from other signaling pathways downstream of zinc uptake. Furthermore, although autophagy is activated to clear damaged mitochondria, it may also target other impaired organelles or proteins, suggesting a broader regulatory scope. Therefore, our study provides only a partial elucidation of the mechanisms through which epithelial SLC39A1 protects against ALI/ARDS. Finally, identifying potent agonists of SLC39A1 could facilitate the translation of these findings into clinical applications for the treatment of ALI/ARDS.

Methods

Human studies

Human lung tissues and sections were collected from ten non-acute respiratory distress syndrome (ARDS) patients (lung tumor patients) and six ARDS patients at the Department of Pulmonary and Critical Care Medicine, the First Affiliated Hospital of Chongqing Medical University, Chongqing, China. The use of human samples adhered to the principles outlined in the Declaration of Helsinki and was approved by the Institutional Review Boards of the First Affiliated Hospital of Chongqing Medical University, Chongqing, China (Approval Number: 2020-147-2). Written informed consent were obtained from all participants or their surrogates. Patients with ARDS met the criteria outlined in the Berlin definition and underwent transbronchial lung cryobiopsy (TBLC) for more accurate diagnosis^60,61. The procedure of TBLC was conducted according to our previous study⁶². In brief, under general anesthesia, the 5.9 mm flexible bronchoscope was interposed into the opening of corresponding target lobar bronchus according to the HRCT image. Then the 2.4-mm cryoprobe (ERBE Germany, 20416-032) was advanced through the working channel of the flexible bronchoscope towards the pleura until resistance was met where it was withdrawn ~20 mm, and the cryoprobe freeze for 3 to 6 sec. The flexible bronchoscope, cryoprobe, and biopsy specimen were removed en bloc through the airway. The probe with the frozen biopsy sample was submerged into room temperature saline to speed thawing and removal of sample. Additionally, the tumor-adjacent non-malignant lung tissues (distance>5 cm) from non-ARDS patients donors were obtained during surgery. The clinical information of ARDS patients and non-ARDS patients is shown in Table S1 and S2.

Animals

Our study was conducted exclusively in male mice. Consequently, it remains unclear whether these findings are generalizable to females. This choice of sex was based primarily on practical considerations: at the same age, male and female mice differ substantially in body weight, which might introduce variability in response to injury or treatment. Moreover, there is currently no conclusive evidence suggesting that sex significantly influences the progression of acute lung injury in this model. All animal operations were performed in accordance with the protocols and ethical conduct approved by the Animal Care and Use Committee of Chongqing Medical University (Approval number: IACUC-CQMU-2023-0088) and in compliance with the ARRIVE guidelines. Slc39a1^flox/flox mice were obtained from Cyagen Biosciences (Suzhou, China). Slc39a1^flox/flox mice were cross-bred with Sftpc-CreERT2 mice to generate AT2-Slc39a1 knockout mice (Slc39a1 icKO). To induce the AT2 specific knockout of SLC39A1, Slc39a1 icKO mice were intraperitoneally injected with tamoxifen for 5 consecutive days. After 14 days to eliminate the effects of tamoxifen, Slc39a1 icKO mice and their corresponding Control mice were then intratracheally instilled with LPS (2 mg/kg). Lc3b^−/− mice were originally obtained from the Jackson Laboratory (Stock Number: 009336), and the characteristics of Lc3b^−/− mice were described in our previous study⁶³. Tfe3^−/y mice were purchased from GemPharmatech (Jiangsu, China). For mouse genotyping, genomic DNA was extracted from the murine tail-tips and primer sequences for PCR are listed in Table S3. Wild-type C57BL/6 J mice were purchased from and raised in the Experimental Animal Center of Chongqing Medical University (Animal certificate number: SCXK, Chongqing, 2012-0001). AAV5-SPC-Slc39a1-ZsGreen (abbreviated as AAV-Slc39a1) and its control AAV5-SPC-Vector-ZsGreen (AAV-Control), as well as AAV2/5-SPC-mir30-Map1lc3b-mCherry (abbreviated as AAV-shLc3b) and its corresponding control AAV2/5-SPC-mCherry (AAV-shNC), were all obtained from Hanbio Co., Ltd. (Shanghai, China). C57BL6/J mice were intratracheally instilled with about 6.4 × 10¹⁰ vg AAV. 21 d after the AAV instillation, the mice were subjected to LPS (2 mg/kg) intratracheal administration.

Acute lung injury murine model

LPS intratracheal instillation-induced acute lung injury (ALI) murine model was performed according to the procedure described in our pervious study⁶⁴. In brief, after anesthetizing by 1% pentobarbital sodium, the mice were placed on their back on a 40° slope. The trachea was intubated by a BD insyte catheter Q5 with a shortened needle. LPS (1 mg/kg or 2 mg/kg in indicated experiments) were dissolved in 50 μL 0.9% sodium chloride and were instilled followed by 150 μL air using a syringe. The mice were kept in a head-up position until spontaneous breathing resumed. Subsequently, the animals were carefully placed onto the 37 °C heating plate until they recovered from anesthesia. After LPS exposure, the mice did not show any severe sign of physical symptoms, such as lethargy, respiratory distress, or poor feeding. The mice were sacrificed 24 h after LPS treatment. The cecal ligation and puncture (CLP) model was performed according the protocol described previously with slight modification⁶⁵. Male C57BL/6 mice aged 9-10 weeks were anesthetized intraperitoneally with by 1% pentobarbital sodium, and the cecum was exposed and ligatured at its external third, the followed with the puncture by a 21-gauge needle. The cecum was then returned to the peritoneal cavity. Sham-operated animals underwent identical laparotomy, and the cecum was exposed but not ligated or punctured. Mice received 1 mL saline subcutaneously for resuscitation on the 37 °C heating plate⁶⁶. The lung tissues were obtained 12 h after the operation. Acid aspiration-induced ALI murine model was conducted according to the protocol described in previous study⁶⁷. Briefly, anaesthesized mice were intratracheally instilled with HCl (pH 1.5; 2 mL/kg) and ventilated for 3 h. The lung tissues were obtained 6 h after acid aspiration. For pharmacological interventions, the zinc chelator TPEN (10 mg/kg) was administered intraperitoneally (i.p.) 1 h prior to LPS exposure. Alternatively, ZnSO₄ (25 mM) was provided in the drinking water for 15 days before LPS treatment.

Bronchoalveolar lavage (BAL) fluids

The assessment of the degree of lung injury in were performed according to the procedures described in our previous studies^68,69. In brief, after 24 h of intratracheal instillation of LPS, bronchoalveolar lavage (BAL) fluids were collected from lungs by flushing the lungs with 0.9% sterile physiological saline. Afterwards, the collected lavage washes were immediately centrifuged at 2000 × g for 10 min at 4 °C, then the supernatant was collected. The protein concentrations in the cell-free supernatant were further tested by the method of bicinchoninic acid. The BALF cell pellets after centrifugation were resuspended in 1 mL of normal saline. The total number of cells in the resuspension solution was counted with TC20™ Automated Cell Counter (Bio-Rad, USA). The concentrations of IL-6, IL-1β, TNF-α, MPO and LDH activity in BALF were detected using enzyme-linked immunosorbent (ELISA) assay according to the manufacturer’s instructions. The absorbance was measured at 450 nm using Multiskan Go spectrophotometer (Thermo Fisher Scientific).

Assessment of pulmonary edema

The assessment of lung edema was conducted as previous description⁶⁹. In short, the separated lung tissues were put into the weighed micro-centrifuge tubes. Firstly, the wet weights of lung tissues were measured. Then the dry weights of lung tissues were weighed after getting dried in the oven at 60 °C for 48 h. The wet-to-dry weight ratio of lung tissues was calculated as the mass of wet lung divided by the mass of dry lung.

Hematoxylin & Eosin staining

H&E was performed according to the protocol described previously⁶⁸. Briefly, after indicated treatment, the lung tissues used for pathological observation were soaked and fixed in 4% paraformaldehyde. Hematoxylin-Eosin Staining of the lung tissue was performed with paraffin slices according to standard protocols. The representative images were captured by a microscope (Olympus IX53, Tokyo, Japan). According to the results of H&E staining to obtain lung injury scores, forty random high-power fields (400× magnification) were scored in a blinded manner for each experimental group. The selection of random fields usually involves continuous random displacements from the current position (each at least one high-power field length). In order to generate lung injury scores, according to the correlation of each feature, the sum of the five independent variables covering neutrophils in the alveolar space, neutrophils in the interstitial space, hyaline membrane, protein fragments that fill the alveolar space, and thickening of the alveolar space were weighted and then normalized to the calculated number of fields. The resulting damage score was a continuous value between 0 and 1 (including 1)^70,71.

Real-time quantitative PCR analysis

Total RNA was extracted with the qPCR Kits (LS1040, Promega, USA) according to the manufacturer’s instructions. The obtained RNA was reverse transcribed into cDNAs by using a RT Master Mix for qPCR (HY-K0510, MCE). Real-time PCR was conducted with SYBR Green qPCR Master Mix (HY-K0523, MCE). RT-qPCR reactions were performed using the CFX Connect real-time system (Bio-Rad, Hercules, CA, USA). The 2 (-ΔΔCt) method and the Δ normalized method to the expression of TATA-binding protein (TBP), GAPDH or β-actin were used to calculate the relative gene expression. Fold change relative to the mean value was determined by 2 (-ΔΔCt). The specific primers were synthesized by Sangon (Shanghai, China) and the sequences of the primers were shown in Table S3.

Western blot

Western blot analysis was performed according to the previous procedure^64,72. Briefly, total protein from the cells or the lung tissues was extracted with RIPA buffer (ab156034, Abcam) containing protease inhibitors (A32953, Thermo Fisher) and PMSF (Sigma-Aldrich). The protein of the samples was denatured by boiling in water at 95 °C for 5 min. 10–30 μg denatured protein sample was subjected to sodium dodecyl sulfate polyacrylamide gel electrophoresis. After electrophoresis, the protein was then transferred to polyvinylidene fluoride membrane (#1620177, Bio-Rad). The membrane was blocked with 5% skim milk for 1 h at room temperature whereafter at 4 °C overnight incubated with specific primary antibodies, then incubated with horseradish peroxidase-conjugated secondary antibodies. The Molecular Imager Gel Doc XR System (Bio-Rad, Hercules, CA, USA) with enhanced chemiluminescence reagents was used to display bands, the intensities of bands were acquired and analyzed by the ImageJ software (NIH, Bethesda, MD, USA). The antibodies information was listed in Table S4. The full-length blots can be found in the Supplementary Information file.

Immunohistochemistry assay

Immunohistochemistry assay was conducted according to the standard procedures described previously⁶⁹. In brief, after the indicated treatment, the tissue-containing slides were rinsed with phosphate-buffered saline and were blocked with 3% hydrogen peroxide for 10 min. Subsequently, the slides were subjected to the incubation of anti-SLC39A1 antibody at 4 °C overnight. On the next day, biotinylated secondary antibody and horseradish enzyme-labeled streptavidin were incubated at room temperature, respectively. After visualization with diaminobenzidine, the slides were dehydrated in graded alcohols, cleared in xylene, mounted with neutral gum and observed under a microscope (Olympus, IX53, Tokyo, Japan). The antibodies used for IHC were listed in Table S5-S6.

Immunofluorescence

Immunofluorescence staining was performed as previously described⁷². Briefly, frozen lung tissue was thawed at room temperature for 30 min, then washed three times with PBS for 5 min each. Subsequently, the tissue was placed in 1 × citrate acid and heated on high heat for 4 min followed by medium heat for 10 min. The sample was then allowed to cool to room temperature. After another round of washing with PBS three times for 5 min each, the tissue was incubated with 1% BSA in a wet chamber at 37 °C or room temperature for 1 h. The lung tissue was then incubated with the primary antibody overnight at 4 °C. The following day, the tissue was washed with PBS five times for 3 min each. Subsequently, the tissue was incubated with the secondary antibody and DAPI at room temperature for 1 h, followed by another round of washing with PBS three times. Finally, an anti-fluorescence quenching agent was used to cover the sample. For in vitro experiments, cells were seeded on sterilized coverslips in a 24-well plate overnight. After treatment, cells were fixed with 4% paraformaldehyde for 15 min at room temperature, permeabilized with 0.2% Triton X-100 for 10 min, and blocked with a buffer containing 2% BSA and 0.3 M glycine in PBS at room temperature for 1 h. The cells were then incubated with DAPI diluted in the blocking buffer for 1 h. Fluorescence images were captured using a Nikon A1R + /A1 Confocal Microscope (Nikon, Tokyo, Japan). The primary antibodies used in this study are listed in Table S5.

Transmission electron microscopy (TEM)

TEM assays were performed as previously described^69,72. Briefly, the heart perfusion fixed technique was used to collect the specimen of lungs. The physiological saline was initially perfused to clear the blood from lungs, and subsequently the 4% paraformaldehyde and 2.5% (w/v) glutaraldehyde buffer were perfused for the fixation. After perfusion, the lungs were excised and cut into 1 mm³ in size. BEAS-2B cells were centrifuged at 225 × g for 5 min after trypsinization and fixed with 4% glutaraldehyde for 2 h at 4 °C. Then, murine lungs or cells were fixed with 1% osmium tetroxide for 1 h at 4 °C. Subsequently, cells were dehydrated in a graded series of alcohol and acetone, followed by embedment in Epon 816 (Electron Microscopy Sciences, Hatfield, PA, USA). Ultrathin sections were obtained by using a Leica ultramicrotome (Leica Microsystems, Buffalo Grove, IL, USA) and stained with uranyl acetate and lead citrate. TEM images were captured under a JEM-1400Plus transmission electron microscope (JEOL Ltd. Tokyo, Japan).

RNA-seq

Total RNA was extracted from the lungs of mice with indicated treatment using TRIzol® Reagent according the manufacturer’s instructions. Then RNA quality was determined by 5300 Fragment Analyzer (Agilent) and quantified using the ND-2000 (NanoDrop). RNA purification, reverse transcription, library construction and sequencing were performed at Shanghai Majorbio Bio-pharm Biotechnology Co., Ltd. (Shanghai, China) according to the manufacturer’s instructions (Illumina, San Diego, CA). The murine RNA-seq transcriptome library was prepared following Illumina® Stranded mRNA Prep, Ligation from Illumina (San Diego, CA) using 1 μg of total RNA. The messenger RNA was isolated according to polyA selection method by oligo(dT) beads and then fragmented by fragmentation buffer firstly. Secondly double-stranded cDNA was synthesized using a SuperScript double-stranded cDNA synthesis kit (Invitrogen, CA) with random hexamer primers (Illumina). Then the synthesized cDNA was subjected to end-repair, phosphorylation and ‘A’ base addition according to Illumina’s library construction protocol. Libraries were size selected for cDNA target fragments of 300 bp on 2% Low Range Ultra Agarose followed by PCR amplified using Phusion DNA polymerase (NEB) for 15 cycles. After quantified by Qubit 4.0, paired-end RNA-seq sequencing library was sequenced with the NovaSeq 6000 sequencer (2 × 150 bp read length). To identify differential expression genes (DEGs) between two different samples, the expression level of each transcript was calculated according to the transcripts per million reads (TPM) method. Differential expression analysis was performed using the DESeq2. DEGs with |log2FC | ≧1 and FDR ≤ 0.05 were considered to be significantly different expressed genes in Autophagy or Apoptosis geneset. All the RNA-seq data are deposited in NCBI GEO (GSE245055).

Isolation of rat primary alveolar type II epithelial cells (rat pAT2)

The fetal rats were taken out from parturient Sprague-Dawley rat and then put into 75% alcohol. The lungs of fetal rat were obtained and cut into fragments. The fragments of lungs were digested with trypsin at 37 °C for 25 min until the suspension was jelly-like, the serum-containing DMEM was added to terminate the digestion. After well mixture, the suspension was filtered with 100-mesh cell strainer, and the cells were then transferred to 15 mL centrifuge tube for a centrifuge at 200 × g for 5 min. The cell pellet was resuspended and added with red blood cell lysate and 0.1% collagenase to eliminate red cells or digest collagen. After two-round adherent culture (40 min for each round), the adherent fibroblast cells were discarded and the suspension containing AT2 were collected for further culture. The purity of isolated AT2 cells was determined by immunofluorescence staining with SPC antibody, or performed TEM analysis to observe the cellular morphology and lamellar body.

Cells

A549 (CCL-185), BEAS-2B (CRL-3588) and MLE-12 (CRL-2110) were purchased from the American Type Culture Collection, HEK293T/17 cells (#GNHu44) was obtained from National Collection of Authenticated Cell Cultures (Shanghai, China). The cells were cultured in DMEM medium supplemented with 10% FBS (Gibco) and 100 U/mL penicillin/streptomycin at 37 °C with 5% CO₂.

Stable cell line construction

DNA fragments encoding SLC39A1-FLAG, EGFP-LC3 (a gift from Tamotsu Yoshimori⁷³), TFEB-AcGFP, TFE3-AcGFP, MiTF-AcGFP, GST-TFEB, GST-TFE3 and GST-MITF were PCR amplified using the corresponding plasmid templates. Each fragment was subsequently inserted into linearized lentivirus expression vector pCDH-CMV-MCS-EF1-Puro (System Biosciences, #CD510B-1) by pEASY®-Basic SeamLess Cloning and Assembly Kit (TransGen Biotech, #CU201-03). The recombinant plasmid was sequenced and co-transfected with packaging plasmids psPAX2 and pMD2.G into HEK293T/17 cells (National Collection of Authenticated Cell Cultures, #GNHu44). Lentiviral supernatants were collected 48 h after transfection. The lentiviruses were then used to infect BEAS-2B or A549 cells, and stable cell lines were established through two rounds of puromycin selection.

GST-fusion protein purification and zinc affinity binding

Firstly, the human TFE3, TFEB or MITF coding sequence was subcloned into the GST-fusion vector pEBG (conserved in our laboratory) and then transfected into HEK293T cells for protein expression. GST-tagged recombinant protein GST-TFE3/TFEB/MITF were purified from transfected HEK293T cells using Glutathione High-Capacity Magnetic Agarose Beads (Millipore, #G0924-1ML) according to the manufacturers protocol. Briefly, the crude extract containing the target protein was pipetted into a 1.5 mL Eppendorf tube and incubated with 100 μL Glutathione High Capacity Magnetic Agarose Beads on a rotating mixer for 1 h. After washing with 50 mM Tris Buffered Saline (pH8.0) for three times to remove unbonded proteins, the GST-fusion protein was eluted from GSH beads using 10 mM reduced glutathione elution buffer. Subsequently, the binding of the purified GST-TFE3/TFEB/MITF protein to zinc was assessed by immobilized-metal affinity chromatography (IMAC) using Profinity^TM IMAC resin (Bio-Rad, #1560131) according to the manufacturer’s instructions. In brief, the Profinity^TM IMAC resin was equilibrated with sodium acetate and incubated with zinc ions solution (0.1 M of ZnCl₂). Then, the purified GST-TFE3/TFEB/MITF protein was loaded on the Zn²⁺-IMAC resin column, and then the column was washed three times with PBS and eluted with 500 mM imidazole. The eluate proteins were collected and used for western blot detection.

Inductively coupled plasma-mass spectrometry (ICP-MS)

ICP-MS was performed as previously described⁶⁹. A549 cell lines overexpressing GST-TFE3, GST-TFEB, and GST-MITF were treated either with Vehicle or LPS for 12 h. The solution containing GST-fusion protein were collected. Subsequently, 0.5 mL homogenate and 5 mL digestion solution were accurately added in Teflon digestion tank (Sineo Microwave Chemistry Technology Co., LTD, Shanghai, China), and then plated onto an electronic temperature control heating plate (EHC-1; Sineo Microwave Chemistry Technology Co., LTD, Shanghai, China). The temperature was set as 180 °C to make the solution colorless and almost dry. Afterward, the solution was removed from the heating plate, cooled to room temperature, and the volume was set to 5 mL by nitric acid solution. After mixing thoroughly, the level of zinc ion was measured using PerkinElmer NexIONTM 300X ICP-MS (PerkinElmer, Norwalk, CT).

Flow cytometry

Flow cytometry was conducted as previously described⁷⁴. For zinc concentration detection, primary rat AT2 cells or A549 cells were seeded into 12-well plates. After indicated treatment, zinc-specific indicator FluoZin-3, AM (Molecular Probes, Waltham, MA, USA) was added to the cell at a final concentration of 500 nM, followed by 30 min incubation at room temperature in the dark. Thereafter, cells were washed three times using the complete medium, followed by an additional incubation for 30 min at room temperature in the dark to allow complete de-esterification of intracellular AM esters. All FACS experiments were performed on CytoFLEX flow cytometry (Beckman Coulter, Miami, FL, USA) and results were analyzed using FlowJo^TM v10 Software BD Biosciences (San Jose, CA, USA).

siRNA transfection

Knockdown of gene with siRNA was conducted as previously described⁷⁴. The A549 or BEAS-2B cells were seeded in 12-well plates. After 24 h, the siRNA duplex and Lipofectamine RNAiMAX transfection reagent were diluted in Opti-MEM media for 5 min, respectively. Then, the diluted siRNA and Lipofectamine RNAiMAX transfection reagent were mixed, and incubated at room temperature for 15 min. In the end, the siRNA and Lipofectamine RNAiMAX transfection reagent mixture were added into cell plates. After 48 h, Real-time fluorescence quantitative PCR or western blot was used to verify the knockdown efficiency. The sequences of siRNA were listed in Table S3.

Statistics and Reproducibility

Comparisons between two groups were made by two-tailed unpaired Student’s t-test or Mann-Whitney U-test. One-way analysis of variance (ANOVA) followed by Tukey post hoc tests was used to compare the significant differences among groups. Quantitative values are presented as the mean ± SD. All statistical tests were conducted using Prism 9 software (GraphPad Software, San Diego, CA, USA). Exact P values (reported to four decimal places) are presented in the figures. All experiments were repeated at least twice; the number of biological replicates for each experiment is indicated in the corresponding figure legends.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Author contributions

Z.Z. conceived this project; Z.Z. and C.C. designed and supervised the project; Y.L. and S.G contributed to clinical sample collection and provided insightful suggestions; J.Z., K.Z., and Y.F. performed major experiments; J.Z. constructed cell lines and performed zinc-resin pull down; K.Z. performed RNA-seq analysis; G.X. performed TEM experiments; H.L. and N.L. assisted in the animal experiments; L.M. and Z.L. performed IHC analysis and qPCR detection of patients’ samples; Z.Z., J.Z., and K.Z. wrote the manuscript; Z.Z. and C.C. revised the manuscript with inputs from all authors.

Peer review

Peer review information

Nature Communications thanks Stefan Ryter and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Data availability

All data generated or analysed in this manuscript are included in the published article and its Supplementary Information. The RNA sequencing data have been made publicly available in the NCBI Gene Expression Omnibus (GEO) repository with the dataset identifier GSE245055. Any additional materials may be obtained from the corresponding author upon request. Source data are provided with this paper.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-026-72403-x.