Impaired TRIM16-Mediated Lysophagy in Chronic Obstructive Pulmonary Disease Pathogenesis

Lysosomes contain more than 60 different hydrolytic enzymes and have been recognized as the degradative end point for intracellular and exogenous cargo (1). Recent advances show that the lysosome is not simply a site for processing cellular waste but also an organelle for regulating essential cellular processes, including differentiation, metabolic response, stress resistance, and cell fate. The lysosome functions via storage of amino acids and ions and also modulation of signaling pathways, gene expression, and autophagy (a machinery for lysosomal self-degradation) (2–4). It has been reported that lysosomal function declines with aging, and lysosomal dysfunction is causally linked to the pathogenesis of neurodegenerative diseases and a variety of age-related disorders (1–4). Intriguingly, lysosomal pH stability reflecting lysosomal function has been implicated in determining life span in yeast (5, 6). Along with functional impairment of the lysosome, increase in lysosomal storage of highly cross-linked aggregates can be associated with further disruption of basic lysosomal functions, including trafficking and autolysosome formation, suggesting the existence of a vicious cycle, resulting in progressive lysosomal dysfunction, especially in the setting of aging-related pathology (1, 2).

Chronic obstructive pulmonary disease (COPD), a representative aging-related pulmonary disorder, is a leading cause of death worldwide and is mainly induced by long-term cigarette smoke (CS) exposure (7). Regulated cell death and accelerated cellular senescence evoked by CS exposure have been widely implicated in the COPD pathogenesis through impaired cell repopulation and the prolonged inflammation resulting from senescence-associated secretory phenotype (8–15). We have reported that insufficient autophagic degradation is involved in accelerating cellular senescence in COPD pathogenesis, and increased lipofuscin in epithelial cells was demonstrated in COPD lungs (8–11, 16). Lipofuscin has been recognized as a characteristic biomarker of cellular senescence and reflects the lysosomal accumulation of nondegradable aggregates composed of oxidized proteins, lipids, and metals (17, 18). Lipofuscin formation is orchestrated by the magnitude of oxidative damage, efficiency of mitochondrial repair and proteasomal systems, and functional integrity of the lysosomes (18). Accordingly, it is plausible that aging-associated and CS-enhanced functional deterioration of lysosomes is causally linked to accelerated cellular senescence accompanied by lipofuscin formation during COPD pathogenesis.

Lysosomal membrane permeabilization (LMP) is indicative of a damaged lysosome and is triggered by various endogenous and exogenous stressors, resulting in cell damage through the leaking of hydrolases into the cytosol (19, 20). Although the entire mechanism for LMP remains unclear, oxidative stress has been known to induce LMP, and high levels of iron in lysosomes are responsible for membrane destabilization through Fenton-type reaction-mediated toxic hydroxyl radical formation. It is likely that CS with high levels of reactive species can be a major cause of LMP in COPD lungs (21). LMP induces the specific autophagic response known as lysophagy to eliminate the damaged lysosomes (22, 23). The protective role of lysophagy in maintaining lysosomal integrity has been demonstrated in hyperuricemic nephropathy using proximal tubule-specific Atg5-deficient mice, a known lysosome damaging model (23). Among the 10 human galectins, a recent paper showed that galectin-3 can be responsible for damaged lysosomes and has a pivotal role in conducting lysophagy (20, 24). During the process of LMP, β-galactosides located within the lumen of the lysosome become exposed to the cytosol, resulting in galectin-3 binding to ruptured lysosomes (19, 25). During lysophagy, the cooperative role of galectin-3 and tripartite motif protein (TRIM) 16 in recognizing damaged lysosomes and selectively mobilizing autophagy regulators to the lysosome has been demonstrated (20, 24, 25). However, induction of LMP by CS exposure and the involvement of TRIM16-mediated lysophagy in the regulation of lysosomal functional integrity remains uncertain in the context of COPD pathogenesis.

In the current study, we examine CS-induced LMP by means of galectin-3 localization in the lysosome. Involvement of TRIM16-mediated lysophagy in regulating CS extract (CSE)–induced lysosomal dysfunction and cellular senescence are evaluated in in vitro models using human bronchial epithelial cells (HBEC). Participation of galectin-3–TRIM16–mediated lysophagy in COPD pathogenesis is further examined in COPD lung tissues and in HBEC isolated from COPD patients.

Normal and COPD airways were collected from the first through the fourth order bronchi from pneumonectomy and lobectomy specimens from resections performed for primary lung cancer, as previously described (26). Informed consent was obtained from all surgical participants as part of an approved ongoing research protocol by the ethical committee of The Jikei University School of Medicine (23-153(5443)). HBECs were isolated with protease treatment, and freshly isolated HBECs were plated onto rat tail collagen type I–coated (10 μg/ml) dishes and incubated overnight, then the medium was changed to bronchial epithelial growth medium (BEGM; LONZA, Tokyo, Japan). Cultures were characterized immunohistochemically using anti-cytokeratin Abs (Lu-5; BioCare Medical, Concord, CA) and anti-vimentin (Sigma-Aldrich, Tokyo, Japan). HBEC showed >95% positive staining with anti-cytokeratin and <5% positive staining with anti-vimentin Ab (data not shown). HBEC were serially passed and used for experiments until passage three. Most experiments were performed with HBEC from non-COPD patients. The bronchial epithelial cell line BEAS-2B was cultured in RPMI 1640 (catalog no. 11875-093; Gibco Life Technologies) with 10% FCS (catalog no. 26140-079; Life Technologies) and penicillin–streptomycin (catalog no. 15070-063; Life Technologies). The Abs used were rabbit anti-TRIM16 (catalog no. ab72131; Abcam; catalog no. sc-398851; Santa Cruz Biotechnology; and catalog no. NBP1-84921; Novus Biologicals), rabbit anti–galectin-3 (catalog no. 12733; Cell Signaling Technology and catalog no. ab53082; Abcam), rabbit anti-LAMP1 (catalog no. 9091; Cell Signaling Technology and catalog no. sc-20011; Santa Cruz Biotechnology), rabbit anti–p-histone H2A.X (Ser¹³⁹) (catalog no. 2577S; Cell Signaling Technology), rabbit anti-phospho–epidermal growth factor receptor (EGFR) (catalog no. sc-57542; Santa Cruz Biotechnology), rabbit anti-EGFR (catalog no. 4267S; Cell Signaling Technology), rabbit anti-MAP1LC3B (catalog no. NB600-1384; Novus Biologicals), mouse anti-MAP1LC3B (catalog no. M152-3; MBL International), rabbit anti-CDKN1A (catalog no. 2947; Cell Signaling Technology), rabbit anti-p62 (catalog no. PM045; MBL International), rabbit anti-ubiquitin (catalog no. BML-PW8810; Enzo Life Sciences), and mouse anti-ACTB (catalog no. A5316; Sigma-Aldrich). Epidermal growth factor (EGF) (catalog no. E9644; Sigma-Aldrich), bafilomycin A1 (Baf A1) from Streptomyces griseus (Sigma-Aldrich, catalog no. B1793), Torin1 (catalog no. 4247; Tocris Bioscience), 3-methyladenine (3MA) (Sigma-Aldrich, catalog no. M9281), Hoechst 33258 (Sigma-Aldrich, catalog no. B2883), PROTEOSTAT Aggresome detection kit (Enzo Life Sciences, catalog no. ENZ-51035), LysoTracker Red (catalog no. L7528; Invitrogen), pepstatin A (catalog no. 4397; Peptide Institute), E-64-d (catalog no. 4321-v; Peptide Institute), CM-H2DCFDA (catalog no. C6827; Molecular Probes/Life Technologies), collagen, type I solution from rat tail (catalog no. C3867; Sigma-Aldrich), leucine-leucine-O-methyl ester (LLOMe) (catalog no. 16008; Cayman Chemical), potassium hexacyanoferrate (III) (catalog no. 167-03722; FUJIFILM Wako), iron (III) chloride, anhydrous (catalog no. 01510; Yoneyama Yakuhin Kogyo), Nuclear fast red-aluminum sulfate solution 0.1% for microscopy (catalog no. 100121; Merk), and cycloheximide (catalog no. 037-20991; FUJIFILM Wako) were purchased.

The LC3B cDNA was the kind gift of Dr. N. Mizushima (Tokyo University, Tokyo, Japan) and Dr. T. Yoshimori (Osaka University, Osaka, Japan) and was cloned into the pEGFP-C1 vector (4). TRIM16 expression vector (pRP[Exp]-EGFP/Puro-CMV > hTRIM16) and control vector were designed and purchased (VectorBuilder). Small interfering RNA (siRNA) targeting TRIM16 (catalog nos. s20873, s20874), LGALS3 (catalog nos. s8148, s8149), ATG5 (catalog nos. s18159, s18160), and negative control siRNAs (catalog nos. AM4635, AM4641) were purchased from Life Technologies. Specific knock-downs of TRIM16, and LGALS3 were validated using two different siRNA. Transfections of HBEC and BEAS-2B cells were performed using the Neon Transfection System (catalog no. MPK5000; Invitrogen/Life Technologies) using matched optimized transfection kits (catalog no. MPK10096; Invitrogen/Life Technologies).

CSE was prepared as previously described (4). Forty milliliters of CS were drawn into the syringe and slowly bubbled into sterile serum-free cell culture media in 15-ml BD Falcon Tubes. One cigarette was used for the preparation of 10 ml of solution. The CSE solution was filtered (0.22 μm; catalog no. SLGS033SS; Merk Millipore) to remove insoluble particles and was designated as a 100% CSE solution.

HBEC grown on 6- to 12-well culture plates was lysed in RIPA buffer (catalog no. 89900; Thermo Fisher Scientific) with protease inhibitor mixture (catalog no. 11697498001; Roche Diagnostics) and 1 mM sodium orthovanadate, or lysed with Laemmli sample buffer. Western blotting was performed as previously described (24). For each experiment, equal amounts of total protein were resolved by 7.5–15% SDS/PAGE. After SDS/PAGE, proteins were transferred to a PVDF membrane (catalog no. ISEQ00010; Millipore), and incubation with specific primary Ab was performed for 2 h at 37°C or 24 h at 4°C. After washing several times with PBS with Tween 20, the membrane was incubated with anti-rabbit IgG, HRP-linked secondary Ab (catalog no. 7074; Cell Signaling Technology) or anti-mouse IgG, HRP-linked secondary Ab (catalog no. 7076; Cell Signaling Technology) followed by chemiluminescence detection (catalog no. 34080; Thermo Fisher Scientific and catalog no. 1705061; Bio-Rad Laboratories) with the ChemiDocTM Touch imaging system (Bio-Rad Laboratories, Hercules, CA).

BEAS-2B were transfected with control or TRIM16 expression vector, and LLOMe and CSE treatment were started 48 h posttransfection. Immunoprecipitation was performed as previously described (14). Cells were lysed in immunoprecipitation (IP) buffer containing 20 mM Tris (pH 7.5), 137 mM NaCl, 1 mM MgCl2, 1 mM CaCl2, 1% NP-40, 10% glycerol, supplemented with 1:100 Halt Protease and Phosphatase Inhibitor Cocktail (, catalog no. 78440; Thermo Fisher Scientific) and Benzonase (catalog no. 70746-4; Novagen) at 12.5 U/ml. The lysates were rotated at 4°C for 30 min. The supernatant was incubated with Ab-conjugated beads (catalog no. 28944008; GE Healthcare) and rotated at 4°C overnight. The IP was washed and collected by magnet five times with IP buffer and boiled with SDS. Samples were analyzed by Western blotting.

Immunohistochemical staining was performed as previously described with minor modification on the paraffin-embedded lung tissues (24). The relative ratio of positively stained areas in airway epithelial cells was calculated by using ImageJ. Immunofluorescence staining was also performed as previously described (24). BEAS-2B cells were transfected with pEGFP-LC3, and CSE treatment was started 48 h posttransfection. Baf A1 (200 nM) treatment was started 6 h before fixation to clearly demonstrate the autophagosome formation of GFP-LC3 “dots,” which result from BafA1 prevention of lysosomal degradation. After a 24 h treatment with CSE, BEAS-2B cells were fixed with 4% paraformaldehyde for 15 min followed by permeabilization with 0.03% Triton X (catalog no. 160-24751; Wako Pure Chemical) for 60 min. After blocking with 1% BSA (catalog no. A2153; Sigma-Aldrich) for 60 min, the primary and secondary Abs were applied according to the manufacturer’s instructions. Fluorescence microscopy analysis of p-histone H2AX was performed in HBEC (Olympus BX60 microscope; Olympus, Tokyo, Japan and BZ-X700; Keyence, Tokyo, Japan). Confocal laser scanning microscopy analysis of galectin-3, lysosomal-associated membrane protein 1 (LAMP1), and EGFP-LC3 in BEAS-2B cells and HBECs was also performed (LSM 880; Carl Zeiss, Tokyo, Japan).

For Sudan black staining, lipofuscin staining was performed in cell cultures as previously described (17). For the Schmorl method, lipofuscin staining was performed with potassium ferricyanide and Nuclear fast red on the paraffin-embedded lung tissues.

Aggresome staining was performed using HBEC grown on 12-well culture plates according to the manufacturer’s instructions (catalog no. ENZ-51035; Enzo Life Sciences). Aggresome staining was also performed on frozen-embedded lung tissues.

Measurement of the pH of lysosomes was performed using HBEC grown on 12-well culture plates with LysoTracker Red (catalog no. L7528, Invitrogen) staining.

HBEC and BEAS-2B cells were used. BEAS-2B cells were starved for 12 h with RPMI without FCS at 37°C. The serum-starved cells were then preincubated for 30 min in the presence of cycloheximide (20 μg/ml) before incubation with EGF (100ng/ml) at 37°C for the indicated times. The cells were then washed with ice-cold PBS and lysed, followed by SDS-PAGE and Western blot analysis or immunofluorescence staining. Bafilomycin A1 (0.17 μM) was added when the cells were incubated with RPMI (27).

Senescence-associated β-galactosidase (SA-β-Gal) staining was performed using HBEC grown on 12-well culture plates according to the manufacturer’s instructions (catalog no. CS0030; Sigma-Aldrich).

The MTT assay was performed according to the manufacturer’s instructions (catalog no. 11465007001; Roche).

HBEC were transfected with control siRNA or TRIM16 siRNA. CSE (1%) treatment was started 24 h posttransfection and HBEC were washed three times with PBS after a 48-h treatment. To collect the conditioned medium, HBEC were incubated in BEGM for 48 h. IL-8, IL-6, and IL-1β were measured with a Human IL-8/CXCL8 Quantikine ELISA Kit (catalog no. D8000C; R&D Systems), a Human IL-6 Quantikine ELISA Kit (catalog no. D6050; R&D Systems), and a Human IL-1β Quantikine ELISA Kit (catalog no. DLB50; R&D Systems) according to the manufacturer’s instructions, respectively.

Lung tissues from pneumonectomy and lobectomy specimens were fixed with 2% glutaraldehyde/0.1M PB (pH7.4) and dehydrated with a graded series of ethanol. Fixed tissues were then embedded in epoxy resin. Ultrathin sections were stained with uranyl acetate and lead citrate and observed with the Hitachi H-7500 transmission electron microscope (Hitachi, Tokyo, Japan). Mitochondria number per one μm² of cell area in small airway epithelial cells were evaluated by counting 10 image fields (10000 X) for each sample.

C57BL/6J (catalog no. 25031; CLEA Japan, Tokyo, Japan) mice were purchased and were maintained in the animal facility at The Jikei University School of Medicine. All experimental procedures are approved by The Jikei University School of Medicine Animal Care Committee. Mice were exposed to control air (three males and three females) or CS (three males and three females) 5 d/wk over a 6 mo period (inExpose; SCIREQ Scientific Respiratory Equipment). The lungs were removed at 6 mo and were used for histological examination. For histological examination, the lungs were fixed overnight in 10% buffered formalin, embedded in paraffin, and sections were stained with H&E according to conventional protocols for histopathological evaluation. Immunohistochemistry was performed as previously described (28).

Data are shown as the average (± SEM) taken from at least three independent experiments. Comparisons between two different groups were determined by Student t test for parametric data or Mann–Whitney U test for nonparametric data. ANOVA was used for multiple comparisons, and Bonferroni post hoc tests were used to test for statistical significance. Significance was defined as p < 0.05. The statistical software used was Prism v.5 (GraphPad Software, San Diego, CA).

First, we examined the effect of CSE on lysosome function by means of LysoTracker Red staining, evaluating lysosomal acidification. Although no clear alteration was demonstrated by LAMP1 staining, CSE treatment (1.0% for 24 h) clearly attenuated positivity of LysoTracker Red staining in HBEC (Fig. 1A). Accumulation of lipofuscin, which was obvious after 48 h, was demonstrated by Sudan black B staining during CSE exposure (Fig. 1B). Lysosomal membrane-damaging agent LLOMe is generally used for inducing LMP. In line with previous findings, LLOMe (1mM) induced formation of galectin-3 puncta in the cytosol, which were clearly colocalized with lysosomes, as determined by confocal microscopic evaluation of LAMP1 staining (Fig. 1C). CSE also induced colocalization of galectin-3 puncta and LAMP1, indicating that CSE can induce LMP with concomitantly impaired lysosomal function in HBEC (Fig. 1C).

Lysophagy, a lysosome-selective form of autophagy, has been implicated in the elimination of damaged lysosomes, including those with LMP (20, 24). To clarify the involvement of lysophagy in regulating CSE-induced LMP, the effects of 3MA, an inhibitor of autophagosome formation, Baf A1, an inhibitor of fusion between autophagosomes and lysosomes, and Torin 1, an autophagy inducer were evaluated, respectively. Autophagy inhibitors 3MA and Baf A1 showed enhanced LMP as measured by galectin-3 and LAMP1 colocalization, which was clearly reduced by Torin 1–mediated autophagy (Fig. 1D). To further elucidate the participation of lysophagy, colocalization between galectin-3 puncta and EGFP-LC3 dots representative of autophagosome formation was examined. Both LLOMe and CSE treatments induced colocalization of galectin-3 and EGFP-LC3 dots in the presence of Baf A1 in control siRNA-treated HBEC (Fig. 1E). siRNA knockdown of ATG5, essential for autophagosome formation, clearly abolished EGFP-LC3 dot formation but enhanced accumulation of galectin-3 dots (Fig. 1E), indicating that lysophagy is responsible for eliminating CSE-induced ruptured lysosomes in HBEC.

Changes in galectin-3 and TRIM16 protein levels during CSE exposure were evaluated in HBEC. TRIM16 protein levels peaked at 24 h, but galectin-3 protein levels gradually increased until 48 h after CSE exposure (Fig. 2A). To elucidate the involvement of TRIM16 in eliminating CSE-induced ruptured lysosomes, siRNA-mediated TRIM16 knockdown experiments were performed. Efficient knockdown was demonstrated at the protein level (Fig. 2C). Autophagy flux in response to CSE exposure was reduced by TRIM16 knockdown, which was shown by decreased LC3B-II conversion and accumulation of p62 and ubiquitinated proteins in HBEC (Fig. 2B, 2C). Autophagy flux was further examined by using BEAS2B cells expressing EGFP-LC3B. CSE-induced EGFP-LC3B dots reflecting autophagosome formation were clearly suppressed by TRIM16 knockdown (Fig. 2D), suggesting that TRIM16 is at least partly responsible for conducting autophagy flux in response to CSE exposure. A specific role for TRIM16 in regulating LMP through lysophagy during CSE exposure was demonstrated by increased colocalization between galectin-3 and LAMP1 in the setting of TRIM16 knockdown (Fig. 2E) and direct association between galectin3 and TRIM16 in CSE and LLOMe-treated BEAS-2B cells, as shown by IP assay (Fig. 2F).

Next, the effect of impaired lysophagy on lysosome function was examined by TRIM16 knockdown. TRIM16 knockdown clearly enhanced CSE-induced lysosomal dysfunction, which was demonstrated by further reduction of LysoTracker Red staining (Fig. 3A). Lysosomal dysfunction is a part of the mechanism for accumulation of aggresomes comprising aggregated misfolded proteins. CSE treatment induced aggresome formation, which was clearly enhanced by TRIM16 knockdown (Fig. 3B). Impaired lysophagy by TRIM16 knockdown can be associated with enhanced cellular senescence in response to CSE exposure in HBEC, which was demonstrated by accumulation of lipofuscin, positive staining of SA-β-Gal and histone H2AFX, increased protein level of the senescence-associated cyclin-dependent kinase inhibitor p21, and reduced cell proliferation (Fig. 3C–F, Supplemental Fig. 1). To further characterize cellular senescence, inflammatory cytokine expression of senescence-associated secretory phenotype SASP was evaluated. CSE treatment induced IL-8, IL-6, and IL-1β secretion, and significant enhancement by TRIM16 knockdown was demonstrated in IL-8 and IL-1β (Fig. 3G). These data suggest that TRIM16-mediated lysophagy is involved in regulating the functional integrity of the lysosome and in cellular senescence during CSE exposure.

To further confirm the involvement of TRIM16 in CSE-induced lysophagy, TRIM16 overexpression experiments were performed. TRIM16 overexpression clearly reduced colocalization between galectin-3 and LAMP1, suggesting enhanced lysophagic elimination of damaged lysosomes with LMP (Fig. 4A). TRIM16 overexpression recovered the functional integrity of lysosomes, as shown by increased positivity of LysoTracker Red staining and reduced aggresome accumulation (Fig. 4B, 4C). In contrast to knockdown experiments, TRIM16 overexpression clearly suppressed cellular senescence induced by CSE exposure in HBEC (Fig. 4D–G).

To elucidate the physiological association between long-term CS exposure and lysosomal dysfunction with respect to insufficient lysophagy, a CS-exposed mouse model was used. CS exposure induced accumulation of lipofuscin and aggresomes in airway epithelial cells (Fig. 5A, 5B). Compared with control-exposed mice, immunohistochemical evaluation showed decreased TRIM16 but increased galectin-3 expression levels in airway epithelial cells in CS-exposed mice. Consistently, high-magnification views of galectin-3 staining clearly demonstrated puncta formation in airway epithelial cells of CS-exposed mice that were absent in controls, suggesting the existence of CS-induced LMP (Fig. 5C). Consistent with immunohistochemical evaluation, Western blotting of lung homogenates also showed decreased TRIM16 but increased galectin-3 expression levels in CS-exposed mice (Fig. 5D).

Lysosomal dysfunction and TRIM16-mediated lysophagy were also evaluated using COPD lung tissues. In line with previous findings (8), increased lipofuscin was detected in alveolar and airway epithelial cells in COPD lungs (Figure 6A, Supplemental Fig. 2,). Immunohistochemical evaluation demonstrated increased LAMP1 expression levels in alveolar and airway epithelial cells of COPD lungs (Fig. 6A, Supplemental Fig. 2). Increased aggresomes were also demonstrated in airway epithelial cells in COPD lungs (Fig. 6B). Consistent with LAMP1 expression levels, electron microscopic evaluation showed significantly increased numbers of lysosomes in airway epithelial cells of COPD lungs (Fig. 6C). Decreased TRIM16 but increased galectin-3 expression levels were revealed in airway epithelial cells in COPD lungs by immunohistochemical evaluation, which may reflect insufficient lysophagy with concomitantly increased accumulation of lysosomes with LMP (Fig. 6D). A high-magnification view of galectin-3 staining clearly demonstrated dot formation in airway epithelial cells in COPD lungs (Fig. 6D). In line with immunohistochemical evaluations, Western blotting of lung homogenates also showed decreased TRIM16 and increased galectin-3 expression levels in COPD lungs (Fig. 6E).

To further confirm the involvement of insufficient lysophagy and lysosomal dysfunction in COPD pathogenesis, HBEC were isolated from COPD patients. Patient characteristics are presented in Table I. Compared with HBEC from nonsmokers and non-COPD smokers, Western blotting of HBEC from COPD patients showed significantly decreased TRIM16 but increased LAMP1 and galectin-3 expression levels (Fig. 7A). Positive correlation between TRIM16 expression levels and pulmonary function tests of percentage of forced expiratory volume in 1 s (FEV1.0)/forced vital capacity (FVC) (FEV1.0%) was demonstrated (Fig. 7B). Although no correlation was detected between LAMP1 expression levels (Supplemental Fig. 3), galectin-3 expression levels were positively correlated with smoking index but negatively correlated with FEV1.0%. Intriguingly, there was a negative correlation between TRIM16 and galectin-3 expression levels, suggesting the existence of a causal link between insufficient lysophagy associated with TRIM16 reduction and accumulation of galectin-3–positive–damaged lysosomes (Fig. 7B). Lysosomal dysfunction in COPD HBEC was demonstrated by increased aggresome formation, reduced LysoTracker Red staining, and lipofuscin accumulation, respectively (Fig. 7C, 7E, 7F). Measurement of EGF-stimulated p-EGFR degradation, a functional assay for lysosomes, was also performed. Lysosomal dysfunction in COPD HBEC was further confirmed by delayed p-EGFR degradation (Fig. 7D). These data suggest the involvement of LMP in COPD pathogenesis, which can be at least partly attributed to insufficient lysophagy conferred by reduced TRIM16 (Supplemental Fig. 4).

In the current study, we show that CSE induces lysosomal damage, with LMP detected by galectin-3 accumulation, and that lysophagy is responsible for maintaining functional integrity of the lysosome during CSE exposure. TRIM16 is involved in conducting lysophagy, and impaired TRIM16-mediated lysophagy is causally associated with lysosomal dysfunction and enhanced cellular senescence. The CS-exposed mouse model and COPD patient sample evaluation indicate that increased LMP in lung epithelial cells, which can be at least partly attributed to impairment of TRIM16-mediated lysophagy, may be responsible for COPD pathogenesis through enhancing cellular senescence.

Lysosomal damage with LMP accompanied by leaking of H⁺ and hydrolases into the cytosol induces a variety of detrimental effects, including caspase-dependent and -independent cell death (19). Although LMP-mediated cell death has been widely implicated in a variety of pathological settings, including bacterial infection, neurodegenerative disorders, and renal disease (21), recent development of the galectin-3 puncta assay allows us to identify the subpopulation of cells exhibiting LMP without cell death, suggesting the existence of various degrees of LMP, potentially associated with inflammation and cellular senescence through inflammasome activation and autophagy inhibition (19, 20). Among the 10 human galectins, galectin-1, -3, -8, and -9 can bind exposed β-galactosidase at the inner lysosomal membrane, signaling downstream events for cell defense (29). Galectin-3 is more specifically involved in the mechanisms for autophagic elimination of damaged lysosomes, suggesting that accumulation of galectin-3 in lysosomes may reflect not only increased LMP but also reduced capacity of lysophagic elimination (25). We showed increased LMP in response to CSE exposure by detecting colocalization of galectin-3 and LAMP1 without an apparent increase in cell death (Fig. 1 and data not shown). CSE-induced LMP was clearly enhanced by autophagy inhibition but reduced by autophagy activation (Fig. 1). Increased galectin-3 puncta of LMP were detected in epithelial cells in both CS-exposed mouse lungs and COPD lungs. We have demonstrated insufficient autophagic degradation in COPD pathogenesis (9, 10, 16), indicating that accumulation of LMP may also reflect insufficient lysophagic elimination of damaged lysosomes in COPD pathogenesis. Although the exact mechanisms for LMP development remain unclear, oxidative stress has been known to induce LMP, and high levels of iron in lysosomes are responsible for membrane destabilization through Fenton-type reactions-mediated toxic hydroxyl radical formation (21). CS contains high levels of reactive oxygen species (ROS) and also induces mitochondrial ROS production, hence it is likely that increased ROS is a major cause of LMP during CS exposure (10).

TRIM16, a TRIM family of E3 ubiquitin ligase, has been shown to be recruited to galectin-3 on damaged lysosomes, and the TRIM16/galectin-3 complex serves as the platform for autophagosome formation during lysophagy (20, 25). We observed enhancement of CSE-induced LMP and lysosomal dysfunction in the setting of TRIM16 knockdown, which were clearly reversed by TRIM16 overexpression in HBEC (Figs. 3, 4). TRIM16 knockdown clearly suppressed autophagy flux, further supporting the notion that TRIM16 is responsible for the elimination of ruptured lysosomes through lysophagy (Fig. 2). We speculate that TRIM16 reduction may impair not only lysophagy but also the entire autophagy flux. Because autophagy flux is governed by the functional integrity of lysosomes, the accumulation of LMP-damaged and dysfunctional lysosomes caused by reduced lysophagy may result in further impairment of autophagy flux, suggesting the existence of a vicious cycle between impaired lysophagy and insufficient autophagy flux during COPD pathogenesis. TRIM16 knockdown also clearly enhanced CSE-induced cellular senescence that was reversed by TRIM16 overexpression, indicating that accumulation of dysfunctional lysosomes by impaired lysophagy is a part of the mechanism of accelerated cellular senescence. Although the effect of impaired lysophagy on cytotoxicity was also evaluated by lactate dehydrogenase assay in the setting of TRIM16 knockdown, no significant increase in cell death was demonstrated during CSE exposure (data not shown), which can be attributed to our experimental conditions using low CSE concentration. Involvement of TRIM16-mediated lysophagy in COPD pathogenesis was shown by reduced TRIM16 protein levels with concomitantly increased dysfunctional lysosomes in airway epithelial cells in CS-exposed mouse and COPD lungs (Figs. 4, 5). A negative correlation between TRIM16 and galectin-3 expression levels was detected in HBEC, further supporting the notion that impaired lysophagy caused by TRIM16 reduction is responsible for accumulation of galectin-3–positive–damaged lysosomes during COPD pathogenesis (Supplemental Fig. 4).

It has been reported that autophagy inhibition by 3MA induces accumulation of lipofuscin-like material, indicating the participation of autophagy in lipofuscin formation (30, 31). Consistently, we showed that impaired lysophagy mediated by TRIM16 knockdown-enhanced accumulation of lipofuscin in response to CSE exposure in HBEC (Fig. 3). Lipofuscin deposition has been considered a hallmark for cellular senescence, and its formation is associated with impaired functionality and effectiveness of the lysosomes (18). We clearly showed increased lipofuscin in both airway and alveolar epithelial cells in COPD lungs (Fig. 6, Supplemental Fig. 2). Proper functional integrity of lysosomes is critical for lysosome-mediated degradation machineries, including autophagy; hence, lysosomal dysfunction resulting in increased lipofuscin may be associated with accumulation of undigested autophagosomes and autolysosomes (21). Previous papers showed increased autophagosome formation in both alveolar macrophages and lung epithelial cells in COPD lungs (32, 33). Impaired autolysosomal degradation was proposed as the mechanism for autophagosome accumulation in alveolar macrophages. Conversely, increased autophagy flux was postulated in lung epithelial cells (32, 33). Although the exact mechanism for increased autophagosomes in epithelial cells remains uncertain, lipofuscin accumulation suggests that not only increased autophagosome formation but also inappropriate autolysosomal degradation conferred by dysfunctional lysosomes can result in autophagosome accumulation in lung epithelial cells of COPD lungs.

There are several limitations in our present study. First, the functional integrity of lysosomes is balanced not only by degradation but also biogenesis, which can be orchestrated by transcription factor EB (TFEB), a master transcription factor for regulating lysosome biogenesis (4, 34). It has been reported that TFEB accumulates in perinuclear spaces and is dysfunctional as a transcription factor in COPD lung tissues; hence, inappropriate biogenesis may also be responsible for accumulation of lysosomal damage, which remains uncertain at this time point (35, 36). Second, LMP has been implicated in cell death, which also has pivotal role in COPD pathogenesis; hence, participation of TRIM16-mediated lysophagy in regulating cell death should be examined (12). Third, lipofuscin was detected in both alveolar and airway epithelial cells, and accelerated cellular senescence was also demonstrated in alveolar epithelial cells (8). Technical restrictions of the cell culture system did not allow us to determine the involvement of TRIM16-mediated lysophagy in regulating functional integrity of lysosomes and cellular senescence in alveolar epithelial cells. Fourth, a specific role for and physiological participation of TRIM16-mediated lysophagy in COPD pathogenesis should be evaluated by using an epithelial-specific TRIM16-deficient mouse model for COPD development in a future study.

In conclusion, CSE-induced lysosomal damage with LMP- and TRIM16-mediated lysophagy was responsible for regulating the functional integrity of lysosomes and also cellular senescence. An increase in ruptured lysosomes and in lipofuscin accumulation with concomitantly reduced TRIM16 in epithelial cells suggests the participation of impaired lysophagy in COPD pathogenesis. Because functional integrity of lysosomes is crucial for maintaining autophagy flux, a vicious cycle between lysophagy and autophagy flux may be at least partly responsible for inappropriate autophagy during COPD pathogenesis.

We thank Stephanie Cambier (University of Washington, Seattle) for technical support.

The authors have no financial conflicts of interest.