Loss of FCHSD1 leads to amelioration of chronic obstructive pulmonary disease

Significance

Chronic obstructive pulmonary disease (COPD/emphysema) is a life-threatening disorder with high morbidity and mortality that is prevalent worldwide. It is characterized by destruction of the alveolar wall and a decline in lung function. However, few palliative therapies are currently available. We show that FCH and double SH3 domains 1 (Fchsd1) knockout mice are protected against airspace enlargement induced by elastase. FCHSD1 deficiency enhanced nuclear translocation of NRF2 and attenuated elastase-induced reduction in SIRT1 levels, which reduced inflammation and apoptosis of lung cells. These data indicate that FCHSD1 promotes the initiation phase of emphysema development and indicate potential therapeutic targets for COPD.

Abstract

Chronic obstructive pulmonary disease (COPD/emphysema) is a life-threatening disorder and there are few effective therapies. Cigarette smoke-induced oxidative stress, airway inflammation, and apoptosis of lung cells have been reported to be involved in the pathogenesis of COPD/emphysema and lead to alveolar septal destruction. Here we show that the expression level of FCH and double SH3 domains 1 (FCHSD1) was drastically increased in mice in response to elastase instillation, an experimental model of COPD. FCHSD1 is a member of the F-BAR family with two SH3 domains. We found that Fchsd1 knockout (Fchsd1^−/−) mice were protected against airspace enlargement induced by elastase. Elastase-instilled lungs of Fchsd1^−/− mice showed reduced inflammation and apoptosis compared with WT mice. We also found that elastase-induced reduction of Sirtuin 1 (SIRT1) levels, a histone deacetylase reported to protect against emphysema, was attenuated in the lungs of Fchsd1^−/− mice. Furthermore, FCHSD1 deficiency enhanced nuclear translocation of nuclear factor-like 2 (NRF2), a redox-sensitive transcription factor, following H₂O₂ stimulation. Conversely, Fchsd1 overexpression inhibited NRF2 nuclear translocation and increased the reduction of SIRT1 levels. Notably, FCHSD1 interacted with NRF2 and SNX9. Our results show that FCHSD1 forms a multicomplex with NRF2 and SNX9 in the cytosol that prevents NRF2 from translocating to the nucleus. We propose that FCHSD1 promotes initiation of emphysema development by inhibiting nuclear translocation of NRF2, which leads to down-regulation of SIRT1.

Chronic obstructive pulmonary disease (COPD) is a common disease worldwide with high morbidity and mortality. It is characterized by destruction of the alveolar wall and a decline in lung function (1). Pulmonary emphysema, a major component of COPD, is caused by cigarette smoking and other environmental risk factors. Emerging evidence suggests that cigarette smoke-induced oxidative stress has a critical role in the initiation phase of emphysema development. It enhances inflammation and extracellular matrix proteolysis and then enhances apoptosis of alveolar cells and septal destruction (2, 3). Oxidative stress is also involved in the development of elastase-induced emphysema (4, 5). In the progression phase, disrupted alveolar maintenance triggers apoptosis and autophagy. Disordered autophagy increases cellular stress, such as endoplasmic reticulum stress, which leads to apoptosis (6, 7). Self-amplifying injury loops involving apoptosis, oxidative stress, and inflammation further enhance disease progression (8). In addition, accelerated cell senescence in the COPD lung leads to decreased cell proliferation, which increases inflammation and reduces cell regeneration (9). However, current therapies for COPD are palliative and do not affect pulmonary cellular maintenance (10).

The acute inflammatory response of lung cells—such as epithelial cells, endothelial cells, fibroblasts, and alveolar macrophages—in COPD pathogenesis involves many intracellular signaling pathways (8). In these pathways, the activation of signaling proteins is regulated by a variety of intracellular trafficking activities and involves interactions with adaptor complexes. Molecular translocation events are thought to be driven by the generation of new protein–protein binding interactions or by second messenger-mediated binding interactions between signaling proteins and subcellular structures, while the manner by which each protein translocates from one site to another is not fully resolved (11).

Among the molecules involved in intracellular trafficking, FCH and Bin-Amphiphysin-Rvs (F-BAR) proteins belong to a class of cytosolic proteins that is primarily associated with endocytosis and vesicle trafficking (12). F-BAR proteins often contain SH3 domains, which mediate multiple interactions with endocytic, cytoskeletal, and signaling proteins (13, 14); therefore, it seems reasonable that F-BAR/SH3–containing proteins might be associated with certain diseases by modulating molecular trafficking. In fact, although the mechanisms remain unclear, emerging evidence indicates that these proteins are associated with various diseases. For example, loss of WRP is linked to impaired learning and memory in mice (15). Xiao et al. showed that FCHSD2 is a substrate for phosphorylation by ERK1/2 and regulates EGF receptor endocytic trafficking. Furthermore, the expression level of FCHSD2 is positively correlated with higher lung cancer patient survival rates (16). To investigate the molecular mechanism of COPD, we searched for COPD-related proteins, focusing on F-BAR/SH3–containing proteins. We identified the FCH domain and double SH3 domains 1 (FCHSD1). FCHSD1 is a mammalian ortholog of Drosophila Nervous Wreck (Nwk) and belongs to the F-BAR protein family (17). Like other F-BAR proteins, FCHSD1 is associated with certain diseases and signaling proteins (18, 19). However, to the best of our knowledge, no published study has determined the function of FCHSD1 in respiratory disease.

Here, we evaluated the role of FCHSD1 in elastase-induced emphysema and addressed whether it might regulate the initiation or progression phase of COPD and lead to lung tissue damage and emphysematous destruction.

Results

Level of FCHSD1 in the Lung Is Increased in Mice Exposed to Elastase: Generation of Fchsd1^−/− Mice.

To identify emphysema-related molecules, we first investigated the expression of FCH family proteins in mouse lung after elastase treatment. We found up-regulated levels of FCHSD1 in lungs in the early response stage to elastase treatment, whereas the levels of other FCH family proteins—such as CIP4, TOCA1, FER, WRP, and FCHSD2—did not change (Fig. 1A). We also found that FCHSD1 protein was increased in mouse type II alveolar epithelial cells (MLE-12 cells) after H₂O₂ stimulation (Fig. 1B). Conversely, FCHSD1 levels did not change after LPS instillation into the lungs (Fig. 1C). Interestingly, without stimulation, high levels of FCHSD1 were present in lungs especially in fibroblasts or epithelial cells, but not other tissues (Fig. 2 A and B). This suggested that FCHSD1 may have an important role in the lung. To determine the role of FCHSD1 in the development of pulmonary emphysema, we generated Fchsd1 knockout (Fchsd1^−/−) mice, which were born at the expected Mendelian ratio (SI Appendix, Fig. S1 A–C). There was no significant difference in immune cell fraction in the spleen, bone marrow, blood, or lung (SI Appendix, Fig. S2), in epithelial cell or fibroblast fraction in the lung (SI Appendix, Fig. S2D), or in the gross appearance and histological morphology of these organs between Fchsd1^−/− and WT mice (SI Appendix, Fig. S1 D and E).

Fig. 1.

Level of Fchsd1 in the lung is increased in COPD model mice. (A) The level of FCHSD1 and other FCH family proteins was determined by immunoblotting of mouse lung 7 d after intratracheal injection of elastase. Data are representative of at least two independent experiments. (B) The level of FCHSD1 and other FCH family proteins was determined by immunoblotting of MLE-12 cells exposed to 200 μM H₂O₂ for 12 h. Data are representative of at least two independent experiments. (C) The level of FCHSD1 in the lung was determined by immunoblotting 1 and 3 d after intratracheal injection of LPS. Data are representative of at least two independent experiments.

Fig. 2.

FCHSD1 deficiency ameliorates elastase-induced airspace enlargement but does not alter LPS-induced inflammation and acute lung injury. (A) The level of FCHSD1 was determined by immunoblotting of whole-tissue homogenates from various nonstimulated mouse tissues. Data are representative of at least two independent experiments. (B) The level of FCHSD1 was determined by immunoblotting of primary cells sorted from nonstimulated mouse lungs. Data are representative of at least two independent experiments. (C–F) Morphological analysis of lungs after intratracheal injection of elastase. Lungs were collected for analysis 36 d after elastase injection. MRI images (C) and H&E-stained images (D) are representative of experiments from four separate mice. Original magnification, 4×. (Scale bars, 500 μm.) (E) MLI was determined by analysis using ImageJ software. (F) Percentage of emphysema area was determined using BZ-X Analyzer software. Both E and F were measured in H&E-stained sections of total lung. Data are expressed as the mean ± SEM. n = 4 per group (elastase), n = 3 per group (PBS). Data are representative of at least two independent experiments. (G) Survival of Fchsd1^−/− and WT mice after intratracheal injection of LPS (10 mg/kg). n = 7 per group. Data are representative of at least two independent experiments. (H and I) Neutrophil number (H) and cytokine concentrations (I) in BAL fluid from Fchsd1^−/− and WT mice 24 h after intratracheal treatment with LPS or PBS. n = 4 per group (LPS), n = 3 per group (PBS). Data are expressed as the mean ± SEM. Data are representative of at least two independent experiments. **P < 0.01; n.s., not significant (P > 0.05).

FCHSD1 Deficiency Protects against Pulmonary Emphysema.

We exposed Fchsd1^−/− and WT mice to elastase. MRI analysis revealed that Fchsd1^−/− mice showed smaller areas of low opacity in the lung compared with WT mice (Fig. 2C). Consistent with this, histological analysis clearly indicated that Fchsd1^−/− mice showed a significantly decreased mean linear intercept (MLI) and emphysematous area compared with WT mice (Fig. 2 D–F). However, no difference in survival or lung inflammation was observed after LPS-induced acute lung injury between Fchsd1^−/− and WT mice (Fig. 2 G–I). These results indicate that FCHSD1 might have a crucial role in the pathogenesis of experimental elastase-induced pulmonary emphysema.

FCHSD1 Enhances Inflammation and Cell Death in Mouse Lung Exposed to Elastase.

We next addressed whether the protection afforded by FCHSD1 deficiency against emphysema is associated with decreased inflammation at an early stage. Fchsd1^−/− mice showed protection against acute inflammation induced by elastase, including reduced numbers of total infiltrating cells and neutrophils, and a reduced concentration of tumor necrosis factor (TNF)-α in bronchoalveolar lavage (BAL) fluid compared with similarly treated WT mice (Fig. 3 A–C). Consistent with this result, induction of interleukin-6 (Il6) and Tnfa mRNA was reduced in Fchsd1^−/− mouse lung fibroblasts (MLFs) compared with WT MLFs in response to H₂O₂-induced oxidative stress (Fig. 3E). Conversely, ectopic expression of Fchsd1 in MLE-12 cells by retroviral infection led to increased Il6 and Tnfa mRNA levels following H₂O₂ stimulation compared with cells infected with empty vector (Fig. 3H). However, production of inflammatory cytokines, such as IL-6 and TNF-α, in response to Toll-like receptor (TLR) ligands or induction of Il6 and Tnfa mRNA in response to H₂O₂ stimulation were not different between Fchsd1^−/− and WT alveolar macrophages (SI Appendix, Fig. S3). Furthermore, TUNEL staining of the lungs after elastase treatment revealed reduced death of alveolar structural cells in Fchsd1^−/− mice compared with WT mice (Fig. 3D). Similarly, expression of cleaved caspase 3 was reduced in Fchsd1^−/− MLFs compared with WT MLFs after H₂O₂ treatment (Fig. 3F). In addition, acetylation of p53 at Lys382, which is a marker of apoptosis, was decreased in Fchsd1^−/− MLFs compared with WT MLFs (Fig. 3G). Conversely, retroviral expression of Fchsd1 in MLE-12 cells led to increased cleaved caspase 3 expression and acetylation of p53 at Lys382 in response to H₂O₂ compared with cells infected with empty vector (Fig. 3 I and J).

Fig. 3.

FCHSD1 enhances lung inflammation and apoptosis in mice exposed to elastase. (A) Cell influx into BAL fluid of Fchsd1^−/− and WT mice after intratracheal treatment with elastase determined by flow cytometry. n = 4 per group. Data are expressed as the mean ± SEM. Data are representative of at least two independent experiments. (B) Neutrophil number in BAL fluid of Fchsd1^−/− and WT mice 24 h after intratracheal treatment with elastase determined by flow cytometry. n = 4 per group. Data are expressed as the mean ± SEM. Data are representative of at least two independent experiments. (C) TNF-α concentration in BAL fluid of Fchsd1^−/− and WT mice 24 h after intratracheal treatment with elastase measured by ELISA. n = 4 per group. Data are expressed as the mean ± SEM. Data are representative of at least two independent experiments. (D) Lung sections of Fchsd1^−/− and WT mice collected 24 h after elastase or PBS administration were subjected to TUNEL staining. TUNEL⁺ cells (brown) in the lung are shown. Original magnification, 10×. (Scale bars, 200 μm.) Data are representative of at least two independent experiments. (E) Il6 and Tnfa mRNA levels in Fchsd1^−/− and WT MLFs after treatment with 275 μM H₂O₂ for indicated times. Data are expressed as the mean ± SEM (n = 3). Data are representative of at least two independent experiments. (F) Immunoblot analysis of Fchsd1^−/− and WT MLFs after treatment with 250 μM H₂O₂ for 12 h. Data are representative of at least two independent experiments. (G) Immunoblot analysis of Fchsd1^−/− and WT MLFs after treatment with 50 μM H₂O₂ for 24 h. Data are representative of at least two independent experiments. (H) Il6 and Tnfa mRNA levels in MLE-12 cells transfected with retroviral Fchsd1 or empty control vector in response to 250 μM H₂O₂ for indicated times. Data are expressed as the mean ± SEM of duplicates (n = 2). Data are representative of at least two independent experiments. (I) Immunoblot analysis of MLE-12 cells transfected with retroviral Fchsd1 or empty control vector after treatment with 100 μM H₂O₂ for 12 h. Data are representative of at least two independent experiments. (J) Immunoblot analysis of MLE-12 cells transfected with retroviral Fchsd1 or empty control vector after treatment with 250 μM H₂O₂ for 3 h. Data are representative of at least two independent experiments. *P < 0.05; **P < 0.01; n.s., not significant (P > 0.05).

Collectively, these findings indicated that FCHSD1 enhances inflammation and cell death after elastase or H₂O₂ stimulation.

Loss of FCHSD1 Led to Elevation of Sirtuin 1 Levels in Lungs Exposed to Elastase and in Cells Exposed to H₂O₂.

To define the function of FCHSD1 in acute lung inflammation and apoptosis, including p53 acetylation in response to elastase, we investigated the level of Sirtuin 1 (SIRT1) in lungs after elastase treatment. Consistent with previous observations (20), intratracheal injection of elastase led to a marked decrease in SIRT1 levels in the lung (Fig. 4A). This was attenuated in Fchsd1^−/− mice compared with WT mice 24 h after intratracheal elastase treatment (Fig. 4A). Similarly, FOXO3a and peroxisome proliferator–activated receptor-γ (PPARγ) levels, target molecules of deacetylation by SIRT1, were higher in Fchsd1^−/− mice than in WT mice exposed to elastase (Fig. 4A). Reduction of SIRT1 levels were attenuated in Fchsd1^−/− MLFs compared with WT MLFs in response to H₂O₂ (Fig. 4B). Conversely, retroviral expression of Fchsd1 in MLE-12 cells led to reduced SIRT1 levels in response to very low concentrations of H₂O₂, whereas SIRT1 levels were not reduced in response to low H₂O₂ concentrations in cells infected with empty vector (Fig. 4C). Together, these results confirmed a key role of FCHSD1 in the reduction of SIRT1 levels in response to oxidative stress.

Fig. 4.

Loss of FCHSD1 led to elevation of SIRT1 levels in the lung after elastase administration. The levels of indicated proteins were determined by immunoblotting of mouse lung. (A) Immunoblot analysis of lungs from Fchsd1^−/− and WT mice 24 h after intratracheal treatment with elastase. Data are representative of at least two independent experiments. (B) Immunoblot analysis of Fchsd1^−/− and WT MLFs after treatment with 275 μM H₂O₂ for 1 h. Data are representative of at least two independent experiments. (C) Immunoblot analysis of MLE-12 cells transfected with retroviral Fchsd1 or empty control vector after treatment with H₂O₂ for the indicated times. Data are representative of at least two independent experiments.

FCHSD1 Negatively Regulates Nuclear Import of Nuclear Factor-Like 2 in Response to H₂O₂-Induced Oxidative Stress.

With regard to the elastase-induced reduction of SIRT1, we next investigated the intracellular localization of nuclear factor–like 2 (NRF2), a redox-sensitive transcription factor, in response to oxidative stress. Immunoblot analysis of cytosol and nuclear fractions showed that the nuclear import of NRF2 was enhanced in Fchsd1^−/− MLFs compared with WT MLFs in response to H₂O₂ treatment (Fig. 5A), consistent with the immunocytochemistry results (Fig. 5C). Conversely, retroviral expression of Fchsd1 in MLE-12 cells led to a reduction in the nuclear import of NRF2 when similarly treated with H₂O₂ compared with cells infected with empty vector (Fig. 5 B and D). However, the mRNA level of Nrf2 was not altered by retroviral expression of Fchsd1 in MLE-12 cells following H₂O₂ treatment compared with cells infected with empty vector (SI Appendix, Fig. S4A). Similarly, it was also not different between Fchsd1^−/− and WT MLFs in response to H₂O₂ stimulation or between lungs from Fchsd1^−/− and WT mice after elastase treatment (SI Appendix, Fig. S4 B and C). These results indicate that FCHSD1 has a crucial role in nuclear translocation of NRF2 but does not affect the transcript level of Nrf2.

Fig. 5.

FCHSD1 negatively regulates nuclear import of NRF2 in response to oxidative stress caused by H₂O_2. (A) Immunoblot analysis of nuclear and cytosol fractions of Fchsd1^−/− and WT MLFs left untreated or treated with 275 μM H₂O₂ for 10 min. Data are representative of at least two independent experiments. (B) Immunoblot analysis of nuclear and cytosol fractions of MLE-12 cells transfected with retroviral Fchsd1 or empty control vector left untreated or treated with 250 μM H₂O₂. Data are representative of at least two independent experiments. (C) Immunofluorescence detection of NRF2 (pink) in Fchsd1^−/− and WT MLFs left untreated or treated with 275 μM H₂O₂ for 10 min. Original magnification, 62×. (Scale bars, 10 μm.) Data are representative of at least two independent experiments. (D) Immunofluorescence detection of NRF2 (red) and GFP (green) in MLE-12 cells transfected with retroviral Fchsd1 or empty control vector left untreated or treated with 250 μM H₂O₂ for 10 min. Original magnification, 62×. (Scale bars, 10 μm.) Data are representative of at least two independent experiments.

FCHSD1 Interacts with NRF2 and SNX9.

FCHSD1 contains two SH3 domains, which mediate multiple interactions with other proteins. To further determine the function of FCHSD1 in regulating nuclear translocation of NRF2, we focused on exploring the roles of Sorting Nexin 9 (SNX9), a key interactor of FCHSD1 (21), in endocytosis and intracellular trafficking. To confirm and extend the identification of SNX9 as a FCHSD1-interacting protein, and to confirm whether FCHSD1 directly binds to NRF2, we performed coimmunoprecipitation experiments for FCHSD1, SNX9, and NRF2 in MLE-12 cells. Surprisingly, we found that FCHSD1 interacts with SNX9 and also with NRF2 (Fig. 6A). We next addressed whether H₂O₂ simulation alters these interactions. FCHSD1 binding to NRF2 and SNX9 was inhibited in response to H₂O₂ treatment (Fig. 6B). Furthermore, FCHSD1 did not associate with KEAP1 (Fig. 6B). In parallel, binding of SNX9 to NRF2 and Importin 8, one of the β-importins that interacts with SNX9, was promoted after H₂O₂ stimulation (Fig. 6C). FCHSD1 did not interact with SIRT1 or the indicated importins (SI Appendix, Fig. S5A). SNX9 and NRF2 did not bind to the importins (SI Appendix, Fig. S5 B and C). These results show that FCHSD1 interacts with NRF2 and SNX9 in the cytosol and prevents NRF2 from translocating to the nucleus. Furthermore, in response to H₂O₂ simulation, FCHSD1 dissociates from the complex and nuclear translocation of NRF2 occurs in association with increased NRF2–SNX9 and SNX9–Importin 8 interactions.

Fig. 6.

FCHSD1 interacts with NRF2 and SNX9, but not with Importin 8, while SNX9 interacts with NRF2 and Importin 8. (A) MLE12 cells were transfected with Flag-tagged FCHSD1, Myc-tagged SNX9, and HA-tagged NRF2, or empty vector and immunoprecipitated (IP) with anti-Flag resin before western analysis for bound NRF2 or SNX9. In the bottom, whole-cell lysate (WCL) was immunoblotted for the indicated targets. Data are representative of at least two independent experiments. (B) Immunoblot analysis of MLE-12 cells expressing Flag-tagged FCHSD1 or empty vector and immunoprecipitation with anti-Flag resin in response to oxidative stress caused by 250 μM H₂O₂ treatment for the indicated times. Data are representative of at least two independent experiments. (C) Immunoblot analysis of MLE-12 cells expressing Flag-tagged SNX9, or empty vector and immunoprecipitation with anti-Flag resin in response to oxidative stress caused by 250 μM H₂O₂ treatment for the indicated times. Data are representative of at least two independent experiments. (D) In the nonstimulated state, FCHSD1 forms complexes with NRF2 and SNX9 in the cytosol and prevents NRF2 from translocating to the nucleus. NRF2 is also bound to KEAP1, its endogenous inhibitor, and is targeted for proteasomal degradation. In response to oxidative stress, KEAP1 is modified and stabilizes NRF2. Then, FCHSD1 dissociates from the complex and SNX9 bound to nuclear membrane-associated Importin 8 targets NRF2 to the nuclear pore.

Discussion

Our findings indicate that FCHSD1 promotes the initiation phase of emphysema development by inhibiting nuclear translocation of NRF2 and then enhancing the reduction of SIRT1 levels. In a mouse model of emphysema induced by intratracheal injection of elastase, we found that deletion of Fchsd1 protected against elastase-induced airspace enlargement, which is a characteristic feature of COPD/emphysema. Furthermore, we showed that FCHSD1 aggravated inflammation, apoptosis and down-regulation of SIRT1. With regard to the underlying mechanism, we identified protein interactions among FCHSD1, NRF2, and SNX9, and found that FCHSD1 negatively regulates nuclear translocation of NRF2 in response to oxidative stress.

Interestingly, deletion of Fchsd1 did not protect against LPS stimulation. Consistent with this observation, FCHSD1 levels were not altered by LPS injection. This finding is in contrast to previous studies that showed protection by NRF2 and SIRT1 against LPS-induced acute lung inflammation (22–24). It is possible that the FCHSD1-SNX9-NRF2 system only has a small effect on the proinflammatory response triggered by TLR ligand stimulation in immune cells such as alveolar macrophages because the levels of the component proteins may be lower in these cells than in other cells. Further studies are needed to determine the detailed distribution of these proteins and the differentiated cell types in which the FCHSD1-SNX9-NRF2 system has a critical role in various stress responses.

NRF2 is a basic leucine zipper (bZIP) transcription factor. Following oxidative or electrophilic stress, it detaches from its cytosolic inhibitor, KEAP1, translocates to the nucleus and binds to antioxidant response element–containing genes (25). NRF2 is critical in determining susceptibility to lung inflammation, oxidative stress, and alveolar cell apoptosis, and disruption of the Nrf2 gene in mice aggravates emphysema (25–27). Although the mechanism of its translocation into the nucleus is not entirely elucidated, it has been shown that NRF2 has nuclear localization signal motifs and that some importins associate with it (28). Our findings revealed that SNX9 binds to NRF2 and Importin 8 in response to oxidative stress, while FCHSD1 binds to SNX9 and NRF2. This observation indicates that FCHSD1 negatively regulates NRF2 nuclear translocation via complex formation with SNX9 and NRF2. This role of FCHSD1 may be independent of KEAP1 because we did not find direct interaction between FCHSD1 and KEAP1 during H₂O₂ stimulation. We also showed that FCHSD1 expression was increased in response to oxidative stress, which is expected to act as a brake to stop the excessive oxidative stress response. Such regulation may contribute to maintaining redox homeostasis because excessive activation of NRF2 causes reductive stress and leads to disorders such as cardiomyopathy and liver injury (29, 30).

We found that administration of elastase in vivo and H₂O₂ stimulation in vitro reduced the level of SIRT1. SIRT1 is a NAD⁺-dependent type III histone deacetylase, and it also deacetylates many other proteins. In response to stimulation, such as DNA damage or oxidative stress, proteins such as p53 and FOXO proteins are deacetylated, which impairs apoptosis (31). SIRT1 also plays a critical role in many pathological processes, including inflammation, stress resistance, and cellular senescence/aging (32, 33). Recently, emerging evidence has shown that SIRT1 levels are reduced in the lungs of patients and mice with COPD, which suggests the involvement of SIRT1 in the pathogenesis of COPD (20, 34, 35). Consistent with this, oxidative stress, such as that caused by H₂O₂, decreases the level of SIRT1 in vitro (34, 36). These findings suggest that FCHSD1 enhances alveolar inflammation and apoptosis by reducing SIRT1, which aggravates emphysema, although the possibility that decreased nuclear translocation of NRF2 by FCHSD1 may also directly contribute to the process cannot be excluded.

The regulatory role of SIRT1-NRF2 cross-talk in various processes has aroused researchers’ interests. SIRT1 can regulate the level and activity of NRF2 (37), while NRF2 can regulate the level of SIRT1 (38, 39). Since our data indicate that FCHSD1 interacts with NRF2 but not with SIRT1, we conclude that NRF2 promotes SIRT1 levels and that FCHSD1 regulates this process. However, the underlying mechanism by which NRF2 regulates SIRT1 levels though FCHSD1–NRF2 interaction still needs further investigation.

Emerging evidence shows that activation of NRF2 is a promising therapeutic strategy. Indeed, bardoxolone methyl, the most potent known activator of the NRF2 pathway, reduces the serum creatinine concentration in patients with type 2 diabetes mellitus and chronic kidney disease (40). However, although it improves renal function, the phase 3 BEACON trial failed because bardoxolone methyl did not reduce the risk of end-stage renal disease, and many adverse events, including death from cardiovascular causes, were observed (41). SIRT1 is also considered a promising pharmaceutical target for the treatment of various diseases. For example, a potent SIRT1 activator, resveratrol, has been shown to extend the life span of various species (42) and to protect against metabolic disease (43). However, further studies are needed before it can be used clinically for certain diseases. The lung-specific expression and activity of FCHSD1 indicates that inhibition of FCHSD1 may contribute to a synergistic effect on NRF2- or SIRT1-targeted treatment, such as with bardoxolone methyl or resveratrol, and lower the risk of systemic adverse events.

In conclusion, our data provide insight into the role of FCHSD1 in stress-response sensing and as an integrator of oxidative stress. FCHSD1 reduced nuclear translocation of NRF2 and increased the reduction of SIRT1 levels, which are all ultimately involved in the pathogenesis of pulmonary emphysema. Our findings may also lead to a specific therapeutic strategy to ameliorate/halt the progression of emphysema via inhibition of FCHSD1.

Materials and Methods

Animal Husbandry and Ethics.

Male mice (8- to 12-wk-old) were housed individually in ventilated cages in a temperature and light-regulated room in a specific pathogen-free facility and received food and water ad libitum. All studies received local ethics review board approval and were performed in accordance with the guidelines of the animal care and use committee of the Research Institute for Microbial Diseases at Osaka University.

Generation of Fchsd1^−/− Mice.

Oocytes and sperm were harvested from C57BL6/J mice and cultured together for 3 h in modified human tubal fluid medium. The fertilized eggs were washed using M2 medium and then 30 zygotes were collected and washed with Opti-MEM I three times to remove the serum in the medium. The zygotes were then electroporated using a CUY21EDIT II electroporator with a F501PT1-10 platinum plate electrode (length: 10 mm; width: 3 mm; height: 0.5 mm; gap: 1 mm) (BEX). The electrode was filled with 5 µL Opti-MEM I containing a mixture of Cas9 protein and single-guide RNA. After electroporation, the zygotes were immediately collected from the electrode chamber and washed with M2 medium four times. The eggs were then cultured in MWM medium at 37 °C in a 5% CO₂ incubator until the two-cell stage. These eggs were then transferred to the oviducts of pseudopregnant females on the day of the vaginal plug.

Experimental Emphysema.

Groups of 8- to 12-wk-old male mice were anesthetized, and porcine pancreatic elastase (Fujifilm Wako Pure Chemical Corporation) at 4.2 U/kg body weight or saline alone were given intratracheally in a volume of 50 μL. Mice were killed at the indicated time points by terminal anesthesia for the collection of lungs and BAL fluid. Lungs samples were fixed for morphological evaluation.

Flow Cytometry and FACS.

Cell suspensions were prepared by sieving and pipetting. Cells were washed in FACS buffer (0.5% bovine serum albumin [BSA] and 2 mM EDTA in PBS, pH 7.2), then incubated with antibodies for 20 min followed by washing twice with FACS buffer. Data were acquired on a flow cytometer (FACS CantoII, BD Bioscience) and analyzed using FlowJo (Tree Star) (44). Cells were sorted using a FACS Aria III (BD Bioscience). The following antibodies for flow cytometry and cell sorting were purchased from BioLegend: anti–Ly-6C (HK1.4), anti–Ly-6G (1A8), anti-Mac1 (M1/70), anti-CD45.2 (104), anti-CD11c (N418), anti-CD3e (145-2C11), anti-CD4 (GK1.5), anti-F4/80 (BM8), anti-SiglecF (S17007L), anti-CD8 (53-6.7), anti-CCR3 (J073E5), anti-B220 (RA3-6B2), anti-EpCAM (G8.8), anti-CD31 (MEC13.3), and anti-CD140a (APA5).

BAL and Analysis by Flow Cytometry and ELISA.

Mice were analyzed at indicated time points. After making an incision in the trachea, a plastic cannula was inserted and airspaces were lavaged with 1 mL saline. This procedure was performed three times. After centrifugation, supernatant was used for cytokine measurement, and cells were used for cell determination. The cells were treated with red blood cell lysis buffer (Sigma-Aldrich) and washed with FACS buffer. Cell determination data of Mac1⁺ Ly6c⁺ Ly6g⁺ neutrophils were acquired using a FACS Canto II flow cytometer and analyzed by FlowJo software as described above. Commercial ELISA kits (R&D Biosystems) were used to measure total TNF-α and IL-6 in the indicated samples, in accordance with the manufacturer’s instructions.

LPS-Induced Lung Inflammation and Injury Model.

Male mice (8 to 12 wk of age) were intratracheally instilled with LPS from Salmonella minnesota R595 (10 mg/kg body weight; Invivogen). After 24 h, BAL fluid was collected for cell counts and analysis of cytokines. For mortality studies, mice were monitored every day for up to 15 d.

Cell Isolation and Culture.

Primary lung fibroblasts (MLFs) were isolated from WT and Fchsd1^−/− mice using trypsin digestion, and the cells were grown in DMEM supplemented with 10% fetal calf serum (no. 10270-106; Gibco), 100 U/mL penicillin G, 100 μg/mL streptomycin (both from Nacalai Tesque), and 2-mercaptoethanol (no. 21417-52; Nacalai Tesque) in a 5% CO₂, 95% O₂ atmosphere at 37 °C. Mouse type II alveolar epithelial cells (MLE-12) were cultured in DMEM Ham’s F-12 medium (no. 08460-95; Nacalai Tesque) supplemented with 2% fetal calf serum (no. 10270-106; Gibco), 100 U/mL penicillin G, 100 μg/mL streptomycin (both from Nacalai Tesque), 0.005 mg/mL insulin (no. 2585; Gibco), 0.01 mg/mL transferrin (no. T5391; Sigma Aldrich), 30 nM sodium selenite (no. S9133; Sigma Aldrich), 10 nM hydrocortisone (no. H0135; Sigma Aldrich), 10 nM β-estradiol (no. E2257; Sigma Aldrich), 10 mM Hepes (no. H0887; Sigma Aldrich), and 2 mM l-glutamine (no. 250030; Gibco). Alveolar macrophages represented >95% of cells in the BAL of unchallenged C57BL/6 mice and were recovered as described above. MLFs were used up to passage 3, while MLE-12 cells were used up to passage 10. The treatments used were: H₂O₂ 50 to 275 μM (no. 7722-84-1; Santoku Chemical Industries), LPS 1 μg/mL (no. tlrl-smlps; Invivogen), and Pam3CSK4 1 μg/mL (no. tlrl-pms; Invivogen). Primary lung cells were prepared as described previously (45). Briefly, lungs were excised from mice and cut into small pieces, then cells were dissociated from lung tissues with collagenase I (1 mg/mL; Sigma). The single-cell suspensions were stained with antibodies as described above, and then epithelial cells (defined as lung CD45⁻ CD31⁻ EpCAM1⁺ cells), fibroblasts (defined as lung CD45⁻ CD31⁻ EpCAM1⁻ CD140a⁺ cells), endothelial cells (defined as lung CD45⁻ CD31⁺ EpCAM1⁻ CD140a⁻ cells), and macrophages (defined as lung CD45⁺ CD11c⁺ Mac1⁺ cells) were sorted using a FACS Aria III.

Construction of Expression Plasmids.

Fchsd1, Nrf2, and Snx9 cDNAs were obtained by PCR from a mouse cDNA library. Fchsd1 cDNAs were cloned into pLZR-ires-GFP for retrovirus production. Flag-tagged Fchsd1, Flag-tagged Nrf2, HA-tagged Nrf2, Flag-tagged Snx9, and Myc-tagged Snx9 were cloned into pcDNA3.1(+) for immunoprecipitation.

Retroviral Transduction.

MLE-12 cells were transduced with retroviral supernatant. Virus was produced using PlatE packaging cells transfected with pLZR-ires-GFP containing a full-length Fchsd1 cDNA or empty vector. After transduction, GFP⁺ cells were sorted using a FACS Aria III (44).

Quantitative Reverse-Transcription PCR.

Total RNA was isolated using an RNA purification kit (no. 1828665; Roche) or TRIzol (no. 15596018; Thermo Fisher Scientific) and reverse transcription was performed with ReverTraAce (Toyobo) according to the manufacturer’s instructions. Quantitative reverse-transcription PCR (RT-qPCR) was performed using real-time PCR Master mix (Toyobo), and fluorescence from the TaqMan probe for each gene was detected with a 7500 real-time PCR system (Applied Biosystems). To determine the relative induction of mRNA, the mRNA level of each gene was calculated using the standard curve method normalized to 18S. Commercially available gene-specific primers and probe sets were obtained from Integrated DNA Technologies (Coralville). The primers and probes used in the study were: Il6 (Mm.PT.58.10005566), Tnfa (Mm.PT.58.12575861), and Nrf2 (Mm.PT.58.29108649).

Western Blotting.

Cultured cells and primary cells were lysed with lysis buffer (20 mM Tris⋅HCl [pH 7.5], 150 mM NaCl, 1 mM EDTA and 1% [vol/vol] Nonidet P-40) containing cOmplete Mini Protease Inhibitor Mixture (Roche), and tissue samples were homogenized in RIPA buffer (no. 08714; Nacalai Tesque) at 1:10 (wt/vol) then centrifuged to pellet cell/tissue debris. The lysates were separated by standard SDS/PAGE and analyzed by immunoblotting. Antibodies to the following proteins were used: SIRT1 (no. 9475; Cell Signaling Technology), FCHSD1 (no. 23362-1-AP; Proteintech), PPARγ (no. 2443; Cell Signaling Technology), NRF2 (no. 12721; Cell Signaling Technology), Lamin A/C (no. 2032; Cell Signaling Technology), β-tubulin (no. 2128; Cell Signaling Technology), WRP (no. E-AB-11600; Elabscience), TOCA-1 (no. E-AB-19876; Elabscience), FER (no. E-AB-19864; Elabscience), CIP4 (no. E-AB-61483; Elabscience), FCHSD2 (no. orb183665; biorbyt), Importin 8 (no. ab208162; Abcam), Importin α5 (no. 18137-1-AP; Proteintech), SNX9 (no. 15721-1-AP; Proteintech), Importin β1 (no. 51186; Cell Signaling Technology), KPNA2 (no. 14372; Cell Signaling Technology), β-actin (no. sc-47778; Santa Cruz Biotechnology), cleaved caspase-3 (no. 9664; Cell Signaling Technology), caspase-3 (no. 9662; Cell Signaling Technology), and Acetyl-p53 (no. 2570; Cell Signaling Technology).

Histological Analysis.

Each organ was rapidly excised, fixed with 4% paraformaldehyde for 24 h, and embedded in paraffin. Tissue sections (5-μm thick) were prepared using a paraffin microtome, deparaffinized with xylene, dehydrated through a graded ethanol series, and processed for hematoxylin and eosin (H&E) staining. The slides stained with H&E were used to evaluate pulmonary emphysema. Percentages of lung tissue affected by emphysema in whole lung were measured using a BZ-X710 fluorescence microscope (Keyence Corp.) and the images were processed and reconstructed using BZ-X Analyzer software in accordance with the manufacturer’s instructions. All quantitative measurements were performed in comparable areas under the same optical and light conditions. For the evaluation of pulmonary emphysema, the MLI was measured after H&E staining using ImageJ software.

Immunocytochemistry.

Cells were cultured on coverslips, fixed with 4% PFA, permeabilized with 0.2% Triton X-100 in PBS and blocked with 1% BSA in PBS containing 0.1% Tween20. Cells were then incubated with primary and secondary antibodies and imaged with a confocal microscope (LSM 880, Zeiss) (45). Antibodies to the following proteins were used: NRF2 (no. 16396-1-AP; Proteintech) and Alexa Fluor 647–conjugated goat anti-rabbit IgG (no. A21245; Invitrogen), as well as Hoechst33342 (no. H342; Dojindo).

Transfection and Immunoprecipitation.

MLE-12 cells were transfected with pcDNA3.1 (+) encoding Flag-tagged Fchsd1, Flag-tagged Nrf2, HA-tagged Nrf2, Flag-tagged Snx9, and Myc-tagged Snx9 or empty vector using Lipofectamine 2000 (no. 11668027; Thermo Fisher Scientific). Twenty-four hours after transfection, cell lysates were collected. For immunoprecipitation with an anti-Flag antibody, cells were lysed in lysis buffer (20 mM Tris⋅HCl [pH 7.5], 150 mM NaCl, and 1% [vol/vol] Nonidet P-40) containing cOmplete Mini Protease Inhibitor Mixture (Roche). Cell lysates were incubated for 2 h with anti-Flag M2 affinity gel (no. A2220; Sigma Aldrich), and washed in wash buffer (20 mM Tris⋅HCl [pH 7.5] and 150 mM NaCl). Proteins were separated by standard SDS/PAGE and analyzed by immunoblotting.

Magnetic Resonance Imaging.

An AvanceIII BioSpec 117/11 system (Bruker) equipped with a 1H QD coil was used for MRI imaging (46). Mice were anesthetized with sevoflurane during image acquisition and the mouse body temperature was maintained with a warm water circulating tube. An ultrashort echo time imaging method was used for lung imaging with the following parameters: TR/TE = 5/0.1 ms, field of view 3 cm, and slice thickness 0.15 mm. Image matrix was set to 200 × 200 × 200.

Subcellular Fractionation.

For subcellular fractionation, MLFs or MLE-12 cells were plated at 1.2 × 10⁶ cells per dish. Cells were stimulated with 275 μM H₂O₂ (for MLFs) or 250 μM H₂O₂ (for MLE12 cells) for the indicated times. After stimulation, cell extracts were prepared using a Nuclear extract kit (no. 40010; Active Motif), in accordance with the manufacturer’s instructions.

Statistical Analysis.

Data from two independent groups were analyzed by Student’s t test. Survival data were analyzed by the log-rank test. For all tests, a P value of less than 0.05 was considered statistically significant.

Data Availability

All study data are included in the article and SI Appendix.