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. 2026 May 13;4(9):1999–2012. doi: 10.1021/envhealth.5c00830

Microplastic-Induced DRP1-BAX Interaction Contributes to Mitochondrial Apoptosis in Lung Epithelial Cells

Xiaoqi Hu †,‡, Xinyi Yuan †, Xue Cao †, Jingran Su †, Ping Zhang †, Yuting Guo †, Fang Zhang †,*, Wenjun Ding †,*
PMCID: PMC13595378  PMID: 42775187

Abstract

As pervasive environmental pollutants, microplastics­(MPs) have been identified in marine ecosystems, food, air, and drinking water, prompting serious consideration of their probable adverse effects on public health. Emerging evidence associates MPs with respiratory risks, yet their toxic mechanisms in lung tissue remain poorly elucidated. In this investigation, C57BL/6n mice were exposed to 10 mg/kg polystyrene microplastics (PS-MPs) with varying sizes (0.2 μm, 1 μm) individually or combined every 2 days for 14 days. The results exhibited that PS-MPs exposure induced the size-dependent respiratory dysfunction in mice, characterized by oxidative stress and pulmonary epithelial apoptosis. In vitro experiments revealed that PS0.2 internalization of MLE-12 cells triggered apoptosis via increasing BAX-mediated cytochrome C release and caspase-3 activation. Mechanistically, PS0.2 promoted DRP1-dependent mitochondrial fission, leading to membrane potential collapse and respiratory chain impairment. The administration of Mdivi-1/si-DRP1 inhibition of DRP1 effectively attenuated mitochondrial fragmentation and apoptosis. Additionally, molecular docking analysis and coimmunoprecipitation revealed that DRP1 directly interacts with BAX to facilitate oligomerization of this pro-apoptotic protein, promoting the release of cytochrome C. These findings establish a novel DRP1-BAX signaling axis in MPs-induced pulmonary toxicity and the first molecular link between PS-MP size and mitochondrial apoptosis via DRP1-BAX oligomerization into environmental particle-related respiratory pathogenesis.

Keywords: microplastics, pulmonary toxicity, apoptosis, mitochondrial function, size-effect

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1. Introduction

Over the past decades, the ubiquitous utilization of plastic products has redefined modern lifestyle, offering unparalleled convenience. However, this progress has come at a steep cost: the emergence of a global environmental crisismicroplastic pollution. This pervasive contamination has posed a critical public health crisis to human health and ecosystem integrity. Microplastics (MPs, 0.1 μm–5 mm) have pervasively colonized global biomes, extending from the deepest ocean trenches to alpine snowpacks, and from bustling urban centers to remote polar regions, demonstrating unparalleled mobility through atmospheric and hydrodynamic vectors. , The omnipresence of MPs has created multiple exposure pathways for humans, primarily through inhalation, ingestion, and dermal contact. , Scientists have gradually realized that MPs are common in the air. The World Health Organization has estimated that human inhaled exposure could reach 3000 particles per day. Workers from fiberglass-reinforced plastics manufacturing exhibited a higher prevalence of chronic cough and elevated proportions of neutrophils and lymphocytes in induced sputum samples. Convincing evidence has proved MPs existence in the lungs from autopsies, and lung cancer resection candidates or patients received lung volume reduction surgery, even bronchoalveolar lavage fluid (BALF) from the lower human airways and sputum. However, the current understanding of the mechanisms underlying MP-induced respiratory damage remains insufficient, particularly regarding their long-term accumulation effects, particle size-dependent toxicity, and potential synergistic effects with other environmental pollutants.

A critical barrier in MP health risk assessment stems from the methodological constraints in detecting and quantifying airborne MPs, especially submicrometer-scale particles, which hinders the accurate estimation of human exposure doses. Notably, the pathogenic potential of inhaled MPs demonstrates a size-dependent correlation. Mechanistic investigations have revealed that MPs with diameters within the 5–30 μm primarily deposit in nasopharyngeal regions through impaction, while those within the 1–5 μm μm tend to settle in the terminal bronchioles due to gravitational sedimentation. In contrast, ultrafine MPs (<1 μm) predominantly accumulate in the alveoli via diffusion. , This size-dependent deposition pattern is particularly concerning given that most MPs detected in BALF from adult European populations are microfibers, with a mean length of 1.73 ± 0.15 mm. These findings underscore critical demands in evaluating the differential toxicity of MPs based on their size distribution. Particle size dictated their deposition patterns, among MPs > 5 μm are trapped in upper airways, 1–5 μm deposit in bronchioles via sedimentation, while submicron PS-MPs (0.1–1 μm) efficiently reach alveoli through Brownian diffusion. We specifically selected 0.2 μm which representing ultrafine MPs and 1 μm threshold for alveolar deposition to probe size-dependent toxicity.

Apoptosis of pulmonary epithelial cells serves as a pivotal driver in the pathological sequelae of most respiratory diseases. Mitochondria, often referred to as the cellular powerhouses, are central to the regulation of apoptosis and cellular homeostasis. Contemporary research has elucidated that dysregulation of mitochondrial dynamics, characterized by imbalances in fission and fusion processes, critically contributes to pulmonary pathogenesis. Mitochondrial membrane fusion is primarily mediated by the outer membrane GTPases mitofusin-1 (MFN1) and mitofusin-2 (MFN2) in concert with optic atrophy 1 (OPA1), which collectively facilitate the biogenesis of reticular mitochondrial architecture. Conversely, mitochondrial fission is regulated by fission 1 (FIS1), which recruits the cytosolic dynamin-related protein 1 (DRP1) to the mitochondrial surface, initiating the division of mitochondria. Emerging evidence has suggested that nanoplastics could disrupt mitochondrial function by inducing excessive production of mitochondrial reactive oxygen species (ROS), altering mitochondrial membrane potential (MMPs), and impairing mitochondrial respiratory chain in bronchial epithelial BEAS-2B cells. Furthermore, nanoplastics have been revealed to induce pulmonary inflammation via the p38/NF-κB pathway, a process closely associated with mitochondrial dysfunction. Given that the equilibrium between mitochondrial fission and fusion is essential for maintaining mitochondrial homeostasis, metabolic adaptation, and apoptosis. While prior studies linked PS-MPs to pulmonary inflammation, the size-dependent molecular mechanisms governing mitochondrial apoptosis remain unexplored. Critically, no evidence exists on whether submicron PS-MPs directly modulate mitochondrial dynamics through DRP1-BAX crosstalk, which could be a potential hallmark differentiating plastic particles from other insoluble particulates.

Therefore, to systematically investigate the pulmonary toxicity of MPs and their potential role in pulmonary diseases, this study employed the integrated in vivo (C57BL/6N murine model) and in vitro (MLE-12 alveolar epithelial cell line) approaches to evaluate a ubiquitous polymer with extensive human exposure, polystyrene (PS), in two environmentally relevant diameters: PS0.2 (0.2 μm in diameter) and PS1 (1 μm in diameter). The inclusion of a combined exposure group (PS0.2+PS1) mimiced real-world scenarios where MPs existed as polydisperse mixtures. The study focused on uncovering the molecular mechanisms underlying MP-induced lung injury by examining how PS-MPs disrupted mitochondrial dynamics and triggered apoptosis in pulmonary epithelial cells. The study hypothesized that PS0.2 exhibited higher pulmonary toxicity than PS1 due to enhanced alveolar deposition and mitochondrial fission via DRP1-BAX interaction, aiming to establish the first evidence that submicron PS-MPs activate a DRP1-dependent mitochondrial apoptotic axis, providing mechanistic basis for regulating ultrafine plastic emissions.

2. Materials and Methods

2.1. Characterizations of PS-MPs

PS-MPs (0.2 and 1 μm) were obtained from the Tianjin Baseline ChromTech Research Centre in China. Transmission electron microscope (TEM) and scanning electron microscope (SEM) were used to observe the size and morphology of PS-MPs in distilled water. The confocal laser microscope (LSM 880, Zeiss, Germany) was applied to observe the fluorescence intensity of the green fluorescence-labeled PS-MPs. The composition of the particles was confirmed by the Fourier transform infrared spectrum (Frontier FT-IR PerkinElmer Co., Ltd.). The size distribution of PS-MPs was analyzed by a granularity analyzer (Zetasizer Nano ZS90, Malvern, UK). PS-MPs were sonicated for 10 min before each experiment.

2.2. Animal Experiment

All the experimental procedures were approved by the Animal Care and Use Committee of the University of Chinese Academy of Sciences (Approval document number: UCAS-A-2021–11–29). Eight-week-old C57BL/6 male mice were purchased from Beijing Vital River Animal Technology Laboratory. The mice were kept in a constant environment with a 12-h light/dark cycle and unrestricted access to food and water. After 1 week of adaptation, mice were randomly divided into four groups (n = 10): PS0.2, PS1, PS0.2+PS1­(MIX), and control group. The mice were administered 10 mg/kg body weight of different particles or distilled water in a volume of 20 μL by intratracheal instillation once every 2 days. Two weeks later, tidal volume (VT), respiratory rate (RR), and minute ventilation (MV) were measured using a whole-body plethysmograph (WBP-4MR, TOW, China) to evaluate the lung function of mice.

2.3. H&E, Immunohistochemical and Immunofluorescent Staining

Lung tissues were fixed in 4% paraformaldehyde and then embedded in paraffin blocks. The 5-μm-sections were stained with hematoxylin and eosin (H&E) staining kits. In immunohistochemical (IHC) staining, the sections were performed with antigen retrieval after deparaffinization and rehydration, and specific antibodies and diaminobenzidine (DAB) were incubated to assess protein localization. In immunofluorescent (IF) staining tests, the tissue slices were incubated specific antibodies overnight after being placed in a sodium citrate buffer for antigen retrieval. Then, the sections were treated with Alexa Fluor secondary antibodies. A quantitative analysis of the staining tests was performed using ImageJ software. The reagents used for IHC and IF staining were listed in Table S1.

2.4. Cell Culture and Treatment

The MLE-12 cell line was purchased from the American Type Culture Collection (ATCC, USA) and cultured in high-glucose Dulbecco’s modified Eagle medium supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin at 37 °C under a humidified atmosphere of 5% CO2. The cells were exposed to PS-MPs at varying concentrations of 0, 10, 20, 40, 80, and 160 μg/mL PS-MPs for 24 h. Cell Counting Kit-8 (CCK-8, CK001, Lablead.Inc., Beijing) was used to determine the cell viability of MLE-12 cells. In addition, MLE-12 cells were pretreated with 10 μmol/L Drp1 inhibitor Mdivi-1 for 2 h, and exposed to 80 μg/mL PS0.2 for 24 h.

2.5. Western Blotting Analysis

The detailed protocols for protein extraction and Western blots were described previously. The primary antibodies used were listed in Table S2.

2.6. Apoptosis Measurement

Following exposure to PS0.2 or Mdivi-1, MLE-12 cells were harvested, washed twice with phosphate-buffered saline, and resuspended. Cells were stained with Annexin V-FITC/PI in the dark at 25 °C for 15 min. Apoptotic subpopulations were quantified using a BD FACSCelesta flow cytometer with standard quadrant analysis.

2.7. Detection of Mitochondrial Respiratory Function and Membrane Potential

After PS0.2 exposure for 24 h, MLE-12 cells were trypsinized and resuspended for cell counting. The suspension was carefully loaded into the O2k chamber of Oroboros O2k (Oroboros Instruments, Innsbruck, Austria), and oligomycin (Omy, 2.5 μmol/L), uncoupling agent FCCP (titrated in 0.5 μmol/L steps), rotenone (Rot, 0.5 μmol/L), and antimycin A (AA, 2.5 μmol/L) were added to the chamber through an injection system to probe basal respiration, ATP production, and maximal respiratory capacity of the mitochondrion. The JC-1 kit was used to measure mitochondrial membrane potential.

2.8. Co-immunoprecipitation

To access the interaction of BAX and DRP1, MLE-12 cells were lysed with lysis buffer for immunoprecipitation (W0013, LABLEAD, China) after PS0.2 exposure. Taking a little supernatant for protein denaturation as input group.20 μL Protein A/G-agarose beads (P0233, LABLEAD, China) and 1 μg IgG-antibody were added into the supernatant at 4 °C for 1 h to detect nonspecific binding, and the agarose beads were separated as the IgG group. Then, the supernatant was coincubated with 20 μL beads and BAX or DRP1 antibody at 4 °C overnight to capture the antibody–antigen complex. The cold washing buffer was used to elute beads and analyzed by Western blot.

2.9. Molecular Docking

Protein–protein docking simulations were performed using the HDOCK web server (http://hdock.phys.hust.edu.cn/), integrating rigid-body docking, conformational sampling, and binding free energy scoring. The platform enables comprehensive analysis of protein interaction interfaces by identifying critical amino acid residues within a 5 Å interaction radius, a standard cutoff for defining molecular recognition events. Structural coordinates for BAX (UniProt ID Q07813) and DRP1 (UniProt ID Q8K1M6) were retrieved from the UniProt database and subjected to rigid-body docking simulations.

2.10. Statistical Analysis

Quantitative data were presented as means ± standard error of means (SEM). The data were analyzed using GraphPad Prism 9 software. Intergroup differences were assessed using a two-tailed unpaired Student’s t test or a one-way ANOVA with Bonferroni post hoc correction. *P < 0.05 was defined as statistically significant.

3. Results

3.1. Physicochemical Characteristics of PS-MPs

TEM and SEM results displayed that PS-MPs particles presented a regular spherical shape with a smooth surface. The particles with the fluorescent tag could be detected under the excitation at 488 nm (Figure A). PS0.2 and PS1 were distributed around 200 and 1000 nm, respectively. Fourier-transform infrared spectroscopy showed The absorption bands observed at 2800–3000 cm–1 were attributed to C–H stretching vibrations of aliphatic chains, while the peak at 1602 cm–1 corresponded to aromatic ring skeletal vibration, which confirmed the chemical components of microplastic particles were polystyrene (Figure B). However, the size distribution of MIX group indicated apparent aggregation of PS0.2 and PS1 (Figure C).

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Physicochemical characteristics of PS-MPs. (A) The TEM and SEM obtained the PS-MPs size and morphology, while a confocal laser microscope detected the fluorescence of PS-MPs. (B) The composition of MPs was analyzed by Fourier-transform infrared spectroscopy. (C) The size distribution of different MPs was given by the granularity analyzer.

3.2. PS-MPs Exposure Impaired Pulmonary Function in a Size-Dependent Manner

To assess the pulmonary toxicity of PS-MPs, mice received intratracheal instillation of PS0.2, PS1, or a binary mixture following the regimen in Figure A. Following exposure, the tidal volume of mice in the ps0.2, ps1, and MIX groups was significantly reduced compared to the control group, and the PS0.2 group was also significantly reduced compared to the PS1 group (Figure B). In addition, the minute ventilation of the three groups was also significantly lower than that of the control group (Figure C), while the respiratory rate remained unchanged (Figure D). The lung-to-body weight ratio was markedly increased in the PS0.2 group (Figure E), and elevated total protein levels in BALF were detected in both the PS0.2 and MIX groups (Figure F). Fluorescence imaging confirmed the PS-MPs accumulation in lung sections, with a notably higher accumulation of PS0.2 particles compared to PS1 (Figure G). Histopathological analysis of lung sections using H&E staining revealed thickened alveolar septa and increased inflammatory cell infiltration in exposed mice (Figure H). Notably, the PS0.2 group exhibited more pronounced histopathological changes, higher BALF protein levels, and greater pulmonary function impairment compared to the PS1 group, likely due to the enhanced deposition of smaller particles in the lung. Collectively, these findings indicated that PS-MPs induced pulmonary dysfunction in a size-dependent manner, with smaller particles (PS0.2) exerting more severe effects.

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PS-MPs induced pulmonary dysfunction of mice. (A) Schematic of mice PS-MPs exposure modeling. (B) The tidal volume of mice. (C) The respiratory rate of mice. (D) The minute ventilation of mice. (E) The ratio of mouse lung to body weight. (F) Total protein levels in BALF. (G) The deposition of PS-MPs in lungs. (H) The H&E staining of the lungs. *P < 0.05, **P < 0.01, ***P < 0.001 vs. Control, ### P < 0.001 vs. PS1 group.

3.3. PS-MPs Induced Apoptosis in the Lung

To assess lung injury induced by PS-MP, apoptotic responses were evaluated in murine lung tissues using TUNEL assays, IHC analysis, and immunoblotting. TUNEL-positive cells were significantly increased in the PS0.2 and MIX groups while PS1-exposed lungs showed no statistical difference (Figure A,B). Additionally, molecular profiling demonstrated the upregulated expression of pro-apoptotic BAX protein in PS0.2 and MIX groups, which is concurrent with decreased antiapoptotic BCL-2 levels across all exposure groups (Figure E–G). Notably, PS0.2 exposure specifically elevated the cleaved caspase-3/total caspase-3 ratio (Figure H), suggesting the activation of the caspase-dependent apoptosis. All the data collectively demonstrated that PS0.2, due to its smaller size and greater alveolar deposition, significantly induces apoptosis in mouse lung tissues, highlighting a size-dependent effect of PS-MPs on pulmonary cell death.

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PS-MPs induced apoptosis in the lung. (A) IHC staining of caspase-3 on lung sections with caspase-3. (B) Quantitative analysis of IHC. (C) Tunel staining of lung tissue. Red fluorescence indicated TUNEL-positive cells in the lung, and white arrows indicated apoptotic cells. (D) Quantitative analysis of TUNEL. (E) The representative images of BAX, BCL-2 and the cleaved/total caspase-3 in lungs. Quantitative analysis of protein expression of (F) BAX, (G) BCL-2 and (H) The cleaved/total caspase-3 ratio. *P < 0.05, **P < 0.01, ***P < 0.001 vs Control, #P < 0.05, ###P < 0.001 vs PS1 group.

3.4. PS0.2 not PS1 Exposure Induced Apoptosis of MLE-12 Cells

To clarify the particle-size dependency of PS-MPs-induced apoptosis, MLE-12 cells were used to detect apoptotic levels in vitro. Initial dose–response screening via CCK-8 assay identified 80 μg/mL as the threshold concentration for both PS0.2 and PS1groups, eliciting significant cytotoxicity compared to untreated controls (Figure A). The internalization of PS-MPs into MLE-12 cells was observed with a laser confocal microscope in Figure B. Interestingly, the massive green fluorescence aggregation was seen in the cells of both PS0.2 and MIX groups, while the rare fluorescence appeared in the PS1 group. Moreover, the Western blotting result demonstrated that PS0.2 and the MIX particles reduced the BCL-2 level and increased the expression of BAX and the Caspase-3 ratio. These results indicated that PS0.2 and MIX particles entered the MLE-12 cells and induced apoptosis (Figure C–F). Additionally, the dose effect of PS0.2 on apoptosis was analyzed after MLE-12 cells were exposed to PS0.2 at 40, 80, and 160 μg/mL. Flow cytometry and immunoblotting were applied to detect the apoptosis of MLE-12 cells treated with these concentrations. Compared to the control group, cell apoptosis significantly increased at 80 and 160 μg/mL (Figure G–L). All these findings indicated that the PS0.2 could enter alveolar epithelial cells and result in apoptosis of pulmonary epithelial cells.

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PS-MPs exposure decreased cell viability and induced MLE-12 cell apoptosis. (A) Viability of MLE-12 cells measured by CCK-8 assay after exposure to different-sized PS-MPs at 0, 10, 20, 40, 80, and 160 μg/mL for 24 h. (B) Internalization of PS-MPs labeled with Green fluorescence in MLE-12 cells treated with 80 μg/mL PS-MPs stained with Tubulin for the cytoskeleton (Red) and DAPI for the nucleus (Blue). (C) The representative images of BAX, BCL-2 and the cleaved/total caspase-3 in MLE-12 cells. Quantitative analysis of protein expression of (D) BAX, (E) BCL-2 and (F) The cleaved/total caspase-3 ratio. (G) The images of flow cytometry, the cells were stained with PI and Annexin after PS0.2 exposure. (H) Quantitative analysis of flow cytometry. (I) The representative images of BAX, BCL-2 and the cleaved/total caspase-3 in MLE-12 cells after PS-MPs exposure at 0, 40, 80, and 160 μg/mL for 24 h. Quantitative analysis of protein expression of (J) BAX, (K) BCL-2 and (L) The cleaved/total caspase-3 ratio. *P < 0.05, **P < 0.01, ***P < 0.001 vs. Control.

3.5. PS0.2 Induced Mitochondrial Abnormality

To investigate the mechanism of PS0.2-induced apoptosis in MLE-12 cells, the related indicators of mitochondria were detected. The results showed a notable increase of DRP1 and FIS1, and a decrease of OPA1, MFN1 and MFN2 (Figure A,B), demonstrating an imbalance of mitochondrial dynamics after exposure to PS0.2. Meanwhile, the laser confocal microscope was used to observe mitochondrial morphology in MLE-12 cells stained with MitoTracker dye and anti-DRP1. The images showed that PS0.2 exposure significantly increased DRP1 and thus aggravated mitochondrial fragmentation (Figure C,D). Moreover, mtROS level and mitochondrial membrane potential (MMP) were further detected. PS0.2 induced a significant increase of mtROS and the loss of MMP (Figure E–G). Additionally, the O2K cell energy metabolism analysis system was applied to test the effects of PS0.2 on mitochondrial respiration. A significant decrease was seen in basal respiration, ATP production, maximal respiration and proton leak (Figure H–I). Taken together, PS0.2 exposure caused mitochondrial fragmentation and dysfunction of MLE-12 cells.

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PS0.2 induced mitochondrial fragmentation and dysfunction of MLE-12 cells. (A) Confocal microscopy observed mitochondrial morphological changes in MLE-12 cells treated with 80 μg/mL. Green fluorescent PS-MPs stained with TOM20 and DAPI. (B) Quantitative analysis of mitochondrial morphological changes. (C) Observation of the distribution of PS-MPs in MLE-12 cells and changes in mitochondrial morphology and structure using TEM, red arrows indicate mitochondria and green arrows indicate PS-MPs. (D) The representative images of protein level of FIS1, DRP1, OPA1, MFN1 and MFN2. (E) Quantitative analysis of protein levels. (F) MLE-12 cells were stained with mitochondrial probes Mito-Tracker Red and anti-DRP1 antibodies. White arrows indicate the colocalization. (G) Quantitative analysis of fluorescence intensity of DRP1. (H) The mtROS levels of MLE-12 cells stained with MitoSOX. (I) Statistical analysis of fluorescence intensity. (J) The mitochondrial membrane potential changes of MLE-12 cells stained with JC-1. (K) Quantitative analysis of fluorescence intensity. (L) The level of mitochondrial oxidative phosphorylation. (M) The quantitative analysis. *P < 0.05, **P < 0.01, ***P < 0.001 vs. Control.

3.6. DRP1 Inhibitor Mdivi-1 Attenuated PS0.2-Induced Apoptosis

To explain the causal role of mitochondrial fission in apoptosis, a mitochondrial fission inhibitor, Mdivi-1, was applied to pretreat MLE-12 cells before PS0.2 exposure. As shown in Figure A-B, Mdivi-1 pretreatment effectively alleviated PS0.2-induced mitochondrial length reduction and network destruction. Meanwhile, Mdivi-1 decreased the apoptosis of PS0.2-treated MLE-12 cells (Figure C,D). Additionally, PS0.2 could promote the translocation of BAX and DRP1 from the cytoplasm to mitochondria, and the release of cytochrome C from mitochondria to the cytoplasm (Figure E–H). However, treating with Mdivi-1 could significantly decrease the level of BAX and DRP1 in mitochondria under PS0.2 exposure (Figure E–H). These results implied that DRP1-mediated BAX translocation may regulate PS0.2-induced apoptosis.

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DRP1 inhibitor Mdivi-1 attenuated PS0.2-induced apoptosis. MLE-12 cells were treated with/without Mdivi-1 (10 μM) and 80 μg/mL PS0.2. (A) The effect of Mdivi-1 on PS0.2-induced mitochondrial deformation stained with TOM20. (B) Quantitative statistics of mitochondrial mean branch length. (C) The images of flow cytometry. (D) Quantitative analysis of flow cytometry.(E) BAX, DRP1 and cytochrome-C distribution and levels in total cell, cytoplasm and mitochondria, TOM40 and GAPDH were used as internal references. Quantitative analysis of (F) BAX, (G) DRP1, (H) cytochrome-C. *P < 0.05, **P < 0.01, ***P < 0.001 vs. Control.

3.7. DRP1 Interacted with BAX to Promote Its Oligomerization in PS0.2-Induced Apoptosis

To elucidate the mechanisms of DRP1-mediated apoptosis, we examined apoptosis-related proteins in mitochondrial fractions after DRP1 knockdown (si-DRP1). si-DRP1 treatment significantly attenuated PS0.2-induced BAX mitochondrial translocation but did not alter mitochondrial BCL-2 levels (Figure A-B). Molecular docking simulations predicted stable DRP1-BAX interactions via hydrogen bonds (e.g., BAX-Thr42/DRP1-Gly586-Gly587, BAX-Arg78/DRP1-Tyr455) and hydrophobic forces, supported by a high-confidence binding score – 281.20, 93% (Figure C), and the coimmunoprecipitation validated this interaction (Figure D). Mechanistically, inactive BAX exists as cytoplasmic monomers or loosely bound to the mitochondrial outer membrane (OMM), but cellular damage triggers its conformational activation and oligomerization on the OMM. Nondenaturing gel electrophoresis revealed that PS0.2 promoted BAX dimer formation, whereas si-DRP1 suppressed this oligomerization (Figure E). Quantitative confocal microscopy further confirmed that PS0.2 enhanced BAX mitochondrial colocalization (Figure F-G) and DRP1-BAX interaction (Figure H–I), both of which were inhibited by si-DRP1. Collectively, these results demonstrated that PS0.2 induced apoptosis by promoting DRP1-BAX interaction, which facilitated BAX mitochondrial translocation, conformational activation, and oligomerization, thereby activating the mitochondrial apoptotic pathway.

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PS0.2 drived mitochondrial apoptosis via promoting DRP1-BAX interactions. The representtative images (A) and quantitative analysis (B) of BAX, DRP1, cytochrome-c and BCL-2 in mitochondria of MLE-12 cells treated with si-DRP1 and PS0.2. (C) HDOCK predicted the docking sites of BAX and DRP1. Green represented BAX, blue represented DRP1 protein, rod-shaped structures represented interacting amino acid residues, yellow dashed line represented hydrogen bonds. (D) The reciprocal coimmunoprecipitation displayed the interaction between BAX and DRP1. (E) Oligomerization of the BAX in MLE-12 cells following si-DRP1 and PS0.2 treatment for 24 h. Co-localization images (F) and quantitative analysis (G) of BAX and mitochondria (TOM20) in MLE-12 treated with 80 μg/mL PS0.2. White arrows indicate the colocalization. Co-localization images (H) and quantitative analysis (I) of BAX and DRP1. Data were mean ± SEM, *P < 0.05, **P < 0.01, ***P < 0.001 vs. Control.

4. Discussion

The increasing environmental burden of microplastics (MPs) poses a significant threat to respiratory health. Epidemiological studies have linked occupational exposure to plastic particulates, such as polyvinyl chloride and nylon fibers, to a higher incidence of pulmonary diseases. , Animal models using tracheal instillation, a primary in vivo method, have shown that exposure to polystyrene microplastics (PS-MPs) of various sizes (1–5 μm, 10–20 μm) can induce lung inflammation, oxidative stress, apoptosis, exacerbate pulmonary fibrosis, and aggravate asthma. , While the toxicological outcomes of PS-MPs are documented, the precise molecular mechanisms driving pulmonary dysfunction, particularly in a size-dependent manner, remain incompletely defined. Our data confirm that PS-MP exposure impairs pulmonary function, evidenced by reduced tidal volume and minute ventilation in mice. More importantly, we identify a clear size-dependency: PS0.2 (0.2 μm) induced significantly more severe pulmonary injury and epithelial apoptosis than PS1 (1 μm). This aligns with the principle that particle toxicity is governed by properties like size, shape, and surface chemistry. Submicron particles (<1 μm) are more readily internalized by alveolar epithelial cells and have a greater potential for systemic distribution, , explaining the greater alveolar deposition and cellular uptake of PS0.2 observed in our study compared to PS1. Notably, in mixed exposures (PS0.2+PS1), toxicity might dominate by the PS0.2 component without evidence of synergy or antagonism, a finding potentially related to particle aggregation altering bioavailability. Thus, a thorough physicochemical characterization of MPs in biological fluids could better correlate aggregation state with bioavailability and toxic outcome.

The toxicity of particles is related to their chemical composition, particle size and shape, surface charge and adhered contaminants, etc. Existing studies have mechanically fragmented high-abundance polyethylene (PE), polystyrene (PS), polyethylene terephthalate (PET), and polyvinyl chloride (PVC) microplastics in the atmosphere and administered them to mice and mouse lung epithelial TC-1 cells, respectively. The results showed that different types of microplastics caused varying degrees of ferroptosis due to their surface charges and hydrophobicity. Notably, the particle-size effect, a critical determinant of biological impact, has been extensively documented in marine ecosystems, with 1–5 μm MPs penetrating the intestinal barriers of echinoderms. − Recent research has pointed out the joint toxicity and synergism of PS0.5 and PS0.05, exhibiting synergistic intestinal toxicity through ROS-mediated apoptosis in mammals. However, whether there is a size-dependent effect of PS on pulmonary toxicity needs further clarification. Inhalation studies have demonstrated that 100 nm PS-MPs could cause lung epithelial cell senescence and lung inflammation. Meanwhile, chronic exposure to PS-MPs in 1–5 μm through intranasal instillation induced severe inflammatory responses, apoptosis and collagen deposition in mice. Our research chose 0.2 and 1 μm PS-MPs to explore the size effect and combined toxicity. In the lung tissue sections, we found more PS0.2 deposited in alveoli than PS1 and more severe lung injury in the PS0.2 group. In vitro experiments also showed a great amount of PS0.2 particles entered MLE-12 cells while PS1 almost adhered to the surface of the cell membrane. These results explained the size-dependent effect of PS-MPs in lung injury. Furthermore, no synergistic or antagonistic effects were observed in mixed PS-MP exposures.

Accumulating evidence has indicated that MPs exert detrimental effects on multiple organs in murine models, including the brain, liver, heart and intestine, which were mostly attributed to oxidative stress and apoptotic cascades. Programmed cell death fundamentally operates via extrinsic (death receptor/caspase-8), intrinsic mitochondrial (caspase-9), and ER stress (caspase-12) pathways. Our study demonstrated that PS0.2 exposure promoted the expression of pro-apoptotic BAX and reduced antiapoptotic BCL-2 both in vivo and in vitro, as well as the caspase-3 activation. We also observed the reduction of mitochondrial membrane potential and the suppression of mitochondrial respiration, indicating that mitochondria played a critical role in apoptotic regulation. Current mechanistic models posit that mitochondrial outer membrane permeabilization (MOMP) facilitates cytochrome c efflux into the cytosol, thereby initiating apoptosome-mediated caspase cascade reaction, culminating in irreversible apoptotic execution. BCL-2 is located at the mitochondrial outer membrane (MOM) and inhibits BAX activity, while BAX is usually in the cytoplasm and translocates to the MOM under cellular stress. However, the caspase-3 activation was only observed in the PS0.2 group, suggesting that PS0.2 induced mitochondria-related apoptosis, while PS1 had no significant effect on apoptosis in the lung.

Physiological mitochondrial homeostasis in healthy cells is maintained through tightly regulated fusion-fission cycles, a dynamic remodeling process critical for preserving metabolic plasticity and genomic stability. Mitochondrial fusion could promote oxidative phosphorylation and enhance ATP production. Mitochondrial fission usually ensures the correct distribution of mitochondrial DNA in cells and removes the damaged mitochondria to restore vitality. However, sustained mitochondrial fission drives pathogenic progression via dysregulated mitophagy coupled with apoptotic/necroptotic activation, as demonstrated in neurodegenerative and ischemic pathologies. Studies have demonstrated that Drp1 translocates from the cytoplasm to mitochondria during the early stages of apoptosis, driving mitochondrial fragmentation and cytochrome C release, activating the caspase cascade and triggering apoptosis. Our data showed that PS0.2 exposure dysregulated mitochondrial dynamics in MLE-12 alveolar epithelium, manifesting as imbalanced fission/fusion ratios. Moreover, the pretreatment of Mdivi-1 and si-DRP1 significantly decreased the PS0.2-induced mitochondrial fragmentation and restored the expression of BAX, thus reducing the occurrence of apoptosis. These results indicated that apoptosis induced by PS0.2 depended on Drp1-mediated mitochondrial fission.

Toxicity studies on particulate matter, including Silica nanoparticles, silver nanoparticles, and PM2.5, have revealed that these particles promote DRP1-mediated mitochondrial fission, leading to apoptosis. However, whether DRP1 directly mediates apoptosis and the underlying molecular mechanisms remain unclear. Notably, such environmental stressors may trigger DRP1-BAX interactions by inducing mitochondrial membrane damage, similar to hypoxia-induced DRP1-BAX-PiC complex formation in mPTP dysregulation. While these studies highlight the critical role of DRP1 in apoptosis, the mechanisms underlying mitochondrial fission and the actions of pro-apoptotic molecules remain to be elucidated. It is reported that the pro-apoptotic protein BAX and the dynamin-like protein DRP1 colocalize at mitochondria during apoptosis to mediate MOMP and fragmentation. The GTPase domain of DRP1, containing conserved GTP-binding motifs, is responsible for hydrolyzing GTP to provide energy, driving mitochondrial membrane constriction and fission, and regulating the oligomerization of DRP1 and membrane-binding capacity through conformational changes. BAX features a BH3 domain in the N-terminal domain, enabling BAX activation and oligomerization.

According to our molecular docking prediction, the N-terminal domain of BAX played a critical role in binding to the GTPase and helical stalk regions of DRP1, enhancing their membrane activity and apoptotic function. In the BAX-Thr42 and DRP1-Gly586/Gly587, the two short hydrogen bonds (2.6 Å and 2.4 Å) likely stabilized the conformation of DRP1’s GTPase active site, consistent with literature reports that DRP1 drives BAX activation via GTP hydrolysis. Additionally, the BAX-Arg78 and DRP1-Tyr455 had two hydrogen bonds (2.9 Å and 1.5 Å), suggesting a high-affinity interaction at this site, potentially facilitating DRP1-induced conformational remodeling of BAX. This finding aligns with the conclusion that the hydrogen bond network is critically involved in modulating the activity of the DRP1 GTPase domain. While our molecular docking analysis predicted stable interactions via residues such as BAX-Thr42/DRP1-Gly586-Gly587 and BAX-Arg78/DRP1-Tyr455, future studies employing site-directed mutagenesis of these key residues will be crucial to definitively confirm their necessity for the DRP1-BAX interaction and the subsequent activation of the mitochondrial apoptotic pathway.

Current toxicological studies show that PS-MPs induce various cytotoxic effects in respiratory epithelia, such as inflammation, apoptosis, and ferroptosis in BEAS-2B cells, , as well as ROS formation and cytokine secretion in primary lung cells. Our work identifies DRP1/BAX-mediated mitochondrial apoptosis as central to PS-MP-induced pulmonary toxicity, raising questions about its interplay with ferroptosis and pyroptosis. Mitochondrial dysfunction, including excessive fission, increased permeability, and elevated ROS, is recognized as an integrative hub initiating multiple cell death programs. We find that PS0.2 triggers DRP1 activation, BAX oligomerization, and cytochrome c release, indicating mitochondrial damage acts as an initiating node for divergent signaling. Ferroptosis, driven by iron-dependent lipid peroxidation, shares a common instigator with this apoptosis, excessive mitochondrial ROS, which directly promotes phospholipid peroxidation. Similarly, mitochondrial dysfunction and mtDNA release activate the NLRP3 inflammasome, the canonical pyroptosis mediator. PS-MP-induced mitochondrial ROS bursts and cardiolipin externalization may serve as danger signals for NLRP3 assembly, suggesting DRP1/BAX-mediated perturbation primes the cell for inflammasome activation. Thus, future research should explore the crosstalk between these death pathways in MP-induced lung injury. Determining the sequence of key events, including DRP1 phosphorylation, BAX activation, lipid peroxidation and GSDMD cleavage will clarify their hierarchical or parallel relationships.

The identification of mitochondrial dysfunction as a central event in MP-induced pulmonary toxicity, as demonstrated in our study, logically points to mitochondrial-targeted therapeutic strategies as a promising intervention avenue. Among these, mitochondrial-targeted antioxidants represent a compelling candidate for mitigating MP-associated injury. MitoQ (mitoquinone mesylate) is a ubiquinone derivative conjugated to a lipophilic triphenylphosphonium cation, which enables its selective accumulation within mitochondria driven by the organelle’s negative membrane potential. Emerging research on PS-NPs have shown that MitoQ pretreatment can attenuate NP-induced mitochondrial dysfunction, insulin resistance, and cellular damage in various models. This aligns with the broader literature where MitoQ has demonstrated protective effects against mitochondrial oxidative stress in models of neurodegenerative, cardiovascular, and metabolic diseases. In our specific pathway, where PS-MPs activate DRP1, promote BAX oligomerization, and induce cytochrome c release, the application of MitoQ could theoretically intercept the cascade upstream by neutralizing the mtROS that may act as both a trigger and an amplifier of these events. Beyond MitoQ, other pharmacological agents targeting mitochondrial dynamics or general oxidative stress have shown relevance. Our own data with the DRP1 inhibitor Mdivi-1 and si-DRP1 directly prove that inhibiting excessive mitochondrial fission alleviates PS-MP-induced apoptosis. This suggests that combinatorial approaches, targeting both the fission machinery (e.g., with Mdivi-1) and its oxidative consequences (e.g., with MitoQ) could yield synergistic benefits. Furthermore, broad-spectrum antioxidants like N-acetylcysteine (NAC) have been reported to reduce oxidative stress and inflammation in some MP exposure models, although their lack of mitochondrial specificity may limit their efficacy against core mitochondrial lesions.

However, our study had several limitations. First, two specific sizes of PS-MPs (0.2 and 1 μm) were representative but did not fully capture the diversity of microplastic sizes and shapes in environmental pollution. Future studies should include a wider variety to better reflect real-world conditions. Future microplastic models for research should go beyond the PS-MPs model and consider the toxicity of weathered/aged microplastics, irregularly shaped microplastics such as fiber microplastics, as well as other common polymer microplastics. For instance, by mechanically crushing and adding sodium dodecyl sulfate (SDS) as a surfactant to prepare environmentally relevant microplastics with morphology and size closer to real-world conditions, the accuracy of environmental risk assessments for microplastics can be effectively improved. In addition, our study used a fixed mixing ratio (assuming a 1:1 mass ratio at a total dose of 10 mg/kg) and no interactions were found. Not considering conducting gradient dose experiments, i.e., changing the PS0.2: PS1 mass ratio while maintaining a constant total mass. This will test whether there is a threshold for the dominant toxicity of PS0.2, or whether it is linearly related to its proportion in the mixture, which has a direct impact on the risk assessment of MP aerosols in polydisperse systems. Tackling these gaps in future research will be key to fully understanding how microplastics harm human health.

5. Conclusion

In summary, our study provided novel insights into the size-dependent pulmonary toxicity of microplastics, particularly highlighting the distinct mechanisms by which PS0.2 and PS1 induce lung injury. A key finding was that PS0.2 primarily induced mitochondria-dependent apoptosis via Drp1-mediated fission, shedding light on the toxicity of nanosized microplastics. This mechanism was reinforced by the protective effect of Mdivi-1 and si-DRP1, highlighting the critical role of mitochondrial dynamics in PS-MP toxicity. Our study also found that PS0.2, but not PS1, strongly activates caspase-3 and impairs mitochondrial function, offering clear evidence of size-dependent effects, which is a less explored area in microplastic research. Furthermore, the absence of synergistic or antagonistic effects in mixed exposures underscores the dominant role of smaller particles in driving toxicity, consistent with recent findings on microplastic behavior in the environment.

Supplementary Material

eh5c00830_si_001.pdf (63.3KB, pdf)

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/envhealth.5c00830.

  • Reagents used for IHC and IF staining and primary antibodies used in Western blot assay (PDF)

∥.

X.H. and X.Y. contributed equally to this work. X.H.: Methodology, data analysis, writing. X.Y.: Methodology, validation, investigation. X.C. and J.S. assisted in animal experiments. P.Z.: Preparation and data processing of TEM samples. Y.G.: Guidance on mitochondrial morphology photography and analysis. F.Z.: Conceptualization, resources, writing. W.D.: Supervision, project administration, funding acquisition.

This study was supported by the Fundamental Research Funds for the Central Universities and the Chinese Academy of Sciences/State Administration of Foreign Experts Affairs International Partnership Program for Creative Research Teams.

Ethics approval and consent to participate: Animal experiments were performed following the principles of laboratory animal care (NIH publication no. 85–23, revised 1985) and approved by the University of Chinese Academy of Sciences Animal Care and Use Committee (Approval document number: UCAS-A-2021–11–29).

The authors declare no competing financial interest.

Published as part of Environment & Health special issue “Micro- and Nano-plastics: What’s Hidden Beneath”.

References

  1. Vethaak A. D., Legler J.. Microplastics and human health. Science. 2021;371(6530):672–674. doi: 10.1126/science.abe5041. [DOI] [PubMed] [Google Scholar]
  2. Luo H., Liu C., He D., Xu J., Sun J., Li J., Pan X.. Environmental behaviors of microplastics in aquatic systems: A systematic review on degradation, adsorption, toxicity and biofilm under aging conditions. J. Hazard Mater. 2022;423(Pt A):126915. doi: 10.1016/j.jhazmat.2021.126915. [DOI] [PubMed] [Google Scholar]
  3. Jenner L. C., Sadofsky L. R., Danopoulos E., Rotchell J. M.. Household indoor microplastics within the Humber region (United Kingdom): Quantification and chemical characterisation of particles present. Atmos. Environ. 2021:259118512. doi: 10.1016/j.atmosenv.2021.118512. [DOI] [Google Scholar]
  4. Prata J. C., da Costa J. P., Lopes I., Duarte A. C., Rocha-Santos T.. Environmental exposure to microplastics: An overview on possible human health effects. Sci. Total Environ. 2020:702134455. doi: 10.1016/j.scitotenv.2019.134455. [DOI] [PubMed] [Google Scholar]
  5. Vasse G. F., Melgert B. N.. Microplastic and plastic pollution: impact on respiratory disease and health. Eur. Respir Rev. 2024;33(172):230226. doi: 10.1183/16000617.0226-2023. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Chen C. H., Tsai P. J., Wang Y. F., Pan C. H., Hung P. C., Ho J. J., Perng D. W., Nemery B., Guo Y. L.. Respiratory health effects of the fiberglass-reinforced plastic lamination process in the yacht-building industry. Scand J. Work Environ. Health. 2021;47(1):62–69. doi: 10.5271/sjweh.3924. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Amato-Lourenço L. F., Carvalho-Oliveira R., Júnior G. R., Dos Santos Galvão L., Ando R. A., Mauad T.. Presence of airborne microplastics in human lung tissue. J. Hazard Mater. 2021:416126124. doi: 10.1016/j.jhazmat.2021.126124. [DOI] [PubMed] [Google Scholar]
  8. Jenner L. C., Rotchell J. M., Bennett R. T., Cowen M., Tentzeris V., Sadofsky L. R.. Detection of microplastics in human lung tissue using μFTIR spectroscopy. Sci. Total Environ. 2022:831154907. doi: 10.1016/j.scitotenv.2022.154907. [DOI] [PubMed] [Google Scholar]
  9. Baeza-Martínez C., Olmos S., González-Pleiter M., López-Castellanos J., García-Pachón E., Masiá-Canuto M., Hernández-Blasco L., Bayo J.. First evidence of microplastics isolated in European citizens’ lower airway. J. Hazard Mater. 2022:438129439. doi: 10.1016/j.jhazmat.2022.129439. [DOI] [PubMed] [Google Scholar]
  10. Huang S., Huang X., Bi R., Guo Q., Yu X., Zeng Q., Huang Z., Liu T., Wu H., Chen Y., Xu J., Wu Y., Guo P.. Detection and Analysis of Microplastics in Human Sputum. Environ. Sci. Technol. 2022;56(4):2476–2486. doi: 10.1021/acs.est.1c03859. [DOI] [PubMed] [Google Scholar]
  11. Prata J. C.. Airborne microplastics: Consequences to human health? Environ. Pollut. 2018:234115–126. doi: 10.1016/j.envpol.2017.11.043. [DOI] [PubMed] [Google Scholar]
  12. Bakand S., Hayes A., Dechsakulthorn F.. Nanoparticles: a review of particle toxicology following inhalation exposure. Inhal. Toxicol. 2012;24(2):125–135. doi: 10.3109/08958378.2010.642021. [DOI] [PubMed] [Google Scholar]
  13. Luo D., Chu X., Wu Y., Wang Z., Liao Z., Ji X., Ju J., Yang B., Chen Z., Dahlgren R., Zhang M., Shang X.. Micro- and nano-plastics in the atmosphere: A review of occurrence, properties and human health risks. J. Hazard Mater. 2024:465133412. doi: 10.1016/j.jhazmat.2023.133412. [DOI] [PubMed] [Google Scholar]
  14. Larson-Casey J. L., He C., Carter A. B.. Mitochondrial quality control in pulmonary fibrosis. Redox Biol. 2020:33101426. doi: 10.1016/j.redox.2020.101426. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Prakash Y. S., Pabelick C. M., Sieck G. C.. Mitochondrial Dysfunction in Airway Disease. Chest. 2017;152(3):618–626. doi: 10.1016/j.chest.2017.03.020. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Lin S., Zhang H., Wang C., Su X. L., Song Y., Wu P., Yang Z., Wong M. H., Cai Z., Zheng C.. Metabolomics Reveal Nanoplastic-Induced Mitochondrial Damage in Human Liver and Lung Cells. Environ. Sci. Technol. 2022;56(17):12483–12493. doi: 10.1021/acs.est.2c03980. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Woo J. H., Seo H. J., Lee J. Y., Lee I., Jeon K., Kim B., Lee K.. Polypropylene nanoplastic exposure leads to lung inflammation through p38-mediated NF-κB pathway due to mitochondrial damage. Part Fibre Toxicol. 2023;20(1):2. doi: 10.1186/s12989-022-00512-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Adebayo M., Singh S., Singh A. P., Dasgupta S.. Mitochondrial fusion and fission: The fine-tune balance for cellular homeostasis. FASEB J. 2021;35(6):e21620. doi: 10.1096/fj.202100067R. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Hu X., Su J., Chen M., Tu Y., Wu C., Cao X., Yuan X., Zhang F., Ding W.. Macrophage-derived exosomal TNF-α promotes pulmonary surfactant protein expression in PM(2.5)-induced acute lung injury. Sci. Total Environ. 2023:892164732. doi: 10.1016/j.scitotenv.2023.164732. [DOI] [PubMed] [Google Scholar]
  20. Kern D. G., Kern E., Crausman R. S., Clapp R. W.. A retrospective cohort study of lung cancer incidence in nylon flock workers, 1998–2008. Int. J. Occup Environ. Health. 2011;17(4):345–351. doi: 10.1179/107735211799041814. [DOI] [PubMed] [Google Scholar]
  21. Miller A., Teirstein A. S., Chuang M., Selikoff I. J., Warshaw R.. Changes in pulmonary function in workers exposed to vinyl chloride and polyvinyl chloride. Ann. N.Y. Acad. Sci. 1975:24642–52. doi: 10.1111/j.1749-6632.1975.tb51079.x. [DOI] [PubMed] [Google Scholar]
  22. Cao J., Xu R., Geng Y., Xu S., Guo M.. Exposure to polystyrene microplastics triggers lung injury via targeting toll-like receptor 2 and activation of the NF-κB signal in mice. Environ. Pollut. 2023:320121068. doi: 10.1016/j.envpol.2023.121068. [DOI] [PubMed] [Google Scholar]
  23. Lu K., Lai K. P., Stoeger T., Ji S., Lin Z., Lin X., Chan T. F., Fang J. K., Lo M., Gao L., Qiu C., Chen S., Chen G., Li L., Wang L.. Detrimental effects of microplastic exposure on normal and asthmatic pulmonary physiology. J. Hazard Mater. 2021:416126069. doi: 10.1016/j.jhazmat.2021.126069. [DOI] [PubMed] [Google Scholar]
  24. Xu J. L., Lin X., Wang J. J., Gowen A. A.. A review of potential human health impacts of micro- and nanoplastics exposure. Sci. Total Environ. 2022;851(Pt 1):158111. doi: 10.1016/j.scitotenv.2022.158111. [DOI] [PubMed] [Google Scholar]
  25. Banerjee A., Shelver W. L.. Micro- and nanoplastic induced cellular toxicity in mammals: A review. Sci. Total Environ. 2021;755(Pt 2):142518. doi: 10.1016/j.scitotenv.2020.142518. [DOI] [PubMed] [Google Scholar]
  26. Vethaak A. D., Leslie H. A.. Plastic Debris Is a Human Health Issue. Environ. Sci. Technol. 2016;50(13):6825–6826. doi: 10.1021/acs.est.6b02569. [DOI] [PubMed] [Google Scholar]
  27. Ji Y., Chen L., Wang Y., Zhang J., Yu Y., Wang M., Wang X., Liu W., Yan B., Xiao L., Song X., Lv C., Chen L.. Realistic Nanoplastics Induced Pulmonary Damage via the Crosstalk of Ferritinophagy and Mitochondrial Dysfunction. ACS Nano. 2024;18(26):16790–16807. doi: 10.1021/acsnano.4c02335. [DOI] [PubMed] [Google Scholar]
  28. Pyl M., Taylor A., Oberhänsli F., Swarzenski P., Hussamy L., Besson M., Danis B., Metian M.. Size-dependent transfer of microplastics across the intestinal wall of the echinoid Paracentrotus lividus. Aquat toxicol. 2022:250106235. doi: 10.1016/j.aquatox.2022.106235. [DOI] [PubMed] [Google Scholar]
  29. Kim S. A., Kim L., Kim T. H., An Y. J.. Assessing the size-dependent effects of microplastics on zebrafish larvae through fish lateral line system and gut damage. Mar. Pollut. Bull. 2022;185(Pt A):114279. doi: 10.1016/j.marpolbul.2022.114279. [DOI] [PubMed] [Google Scholar]
  30. Jeong C. B., Won E. J., Kang H. M., Lee M. C., Hwang D. S., Hwang U. K., Zhou B., Souissi S., Lee S. J., Lee J. S.. Microplastic Size-Dependent Toxicity, Oxidative Stress Induction, and p-JNK and p-p38 Activation in the Monogonont Rotifer (Brachionus koreanus) Environ. Sci. Technol. 2016;50(16):8849–8857. doi: 10.1021/acs.est.6b01441. [DOI] [PubMed] [Google Scholar]
  31. Liang B., Zhong Y., Huang Y., Lin X., Liu J., Lin L., Hu M., Jiang J., Dai M., Wang B., Zhang B., Meng H., Lelaka J. J. J., Sui H., Yang X., Huang Z.. Underestimated health risks: polystyrene micro- and nanoplastics jointly induce intestinal barrier dysfunction by ROS-mediated epithelial cell apoptosis. Part Fibre Toxicol. 2021;18(1):20. doi: 10.1186/s12989-021-00414-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Yang D., Zhu J., Zhou X., Pan D., Nan S., Yin R., Lei Q., Ma N., Zhu H., Chen J., Han L., Ding M., Ding Y.. Polystyrene micro- and nano-particle coexposure injures fetal thalamus by inducing ROS-mediated cell apoptosis. Environ. Int. 2022:166107362. doi: 10.1016/j.envint.2022.107362. [DOI] [PubMed] [Google Scholar]
  33. Li S., Ma Y., Ye S., Tang S., Liang N., Liang Y., Xiao F.. Polystyrene microplastics trigger hepatocyte apoptosis and abnormal glycolytic flux via ROS-driven calcium overload. J. Hazard Mater. 2021:417126025. doi: 10.1016/j.jhazmat.2021.126025. [DOI] [PubMed] [Google Scholar]
  34. Zhou Y., Wu Q., Li Y., Feng Y., Wang Y., Cheng W.. Low-dose of polystyrene microplastics induce cardiotoxicity in mice and human-originated cardiac organoids. Environ. Int. 2023:179108171. doi: 10.1016/j.envint.2023.108171. [DOI] [PubMed] [Google Scholar]
  35. Vandenabeele P., Bultynck G., Savvides S. N.. Pore-forming proteins as drivers of membrane permeabilization in cell death pathways. Nat. Rev. Mol. Cell Biol. 2023;24(5):312–333. doi: 10.1038/s41580-022-00564-w. [DOI] [PubMed] [Google Scholar]
  36. Wei M., Ye Y., Ali M. M., Chamba Y., Tang J., Shang P.. Effect of Fluoride on Cytotoxicity Involved in Mitochondrial Dysfunction: A Review of Mechanism. Front. Vet. Sci. 2022;9:850771. doi: 10.3389/fvets.2022.850771. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Spitz A. Z., Gavathiotis E.. Physiological and pharmacological modulation of BAX. Trends Pharmacol. Sci. 2022;43(3):206–220. doi: 10.1016/j.tips.2021.11.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Quiles J. M., Gustafsson Å.. B. The role of mitochondrial fission in cardiovascular health and disease. Nat. Rev. Cardiol. 2022;19(11):723–736. doi: 10.1038/s41569-022-00703-y. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Tábara L. C., Segawa M., Prudent J.. Molecular mechanisms of mitochondrial dynamics. Nat. Rev. Mol. Cell Biol. 2025;26(2):123–146. doi: 10.1038/s41580-024-00785-1. [DOI] [PubMed] [Google Scholar]
  40. Zhao X., Xu H., Li Y., Liu Y., Li X., Zhou W., Wang J., Guo C., Sun Z., Li Y.. Silica nanoparticles perturbed mitochondrial dynamics and induced myocardial apoptosis via PKA-DRP1-mitochondrial fission signaling. Sci. Total Environ. 2022:842156854. doi: 10.1016/j.scitotenv.2022.156854. [DOI] [PubMed] [Google Scholar]
  41. Chang X., Niu S., Shang M., Li J., Guo M., Zhang W., Sun Z., Li Y., Zhang R., Shen X., Tang M., Xue Y.. ROS-Drp1-mediated mitochondria fission contributes to hippocampal HT22 cell apoptosis induced by silver nanoparticles. Redox Biol. 2023:63102739. doi: 10.1016/j.redox.2023.102739. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Liu X., Zhao X., Li X., Lv S., Ma R., Qi Y., Abulikemu A., Duan H., Guo C., Li Y., Sun Z.. PM­(2.5) triggered apoptosis in lung epithelial cells through the mitochondrial apoptotic way mediated by a ROS-DRP1-mitochondrial fission axis. J. Hazard Mater. 2020:397122608. doi: 10.1016/j.jhazmat.2020.122608. [DOI] [PubMed] [Google Scholar]
  43. Duan C., Kuang L., Hong C., Xiang X., Liu J., Li Q., Peng X., Zhou Y., Wang H., Liu L., Li T.. Mitochondrial Drp1 recognizes and induces excessive mPTP opening after hypoxia through BAX-PiC and LRRK2-HK2. Cell Death Dis. 2021;12(11):1050. doi: 10.1038/s41419-021-04343-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Jenner A., Peña-Blanco A., Salvador-Gallego R., Ugarte-Uribe B., Zollo C., Ganief T., Bierlmeier J., Mund M., Lee J. E., Ries J., Schwarzer D., Macek B., Garcia-Saez A. J.. DRP1 interacts directly with BAX to induce its activation and apoptosis. Embo. 2022;41(8):e108587. doi: 10.15252/embj.2021108587. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Fonseca T. B., Sánchez-Guerrero Á., Milosevic I., Raimundo N.. Mitochondrial fission requires DRP1 but not dynamins. Nature. 2019;570(7761):e34–e42. doi: 10.1038/s41586-019-1296-y. [DOI] [PubMed] [Google Scholar]
  46. Lee E. F., Clarke O. B., Evangelista M., Feng Z., Speed T. P., Tchoubrieva E. B., Strasser A., Kalinna B. H., Colman P. M., Fairlie W. D.. Discovery and molecular characterization of a Bcl-2–regulated cell death pathway in schistosomes. Proc. Natl. Acad. Sci. U.S.A. 2011;108(17):6999–7003. doi: 10.1073/pnas.1100652108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Dong C. D., Chen C. W., Chen Y. C., Chen H. H., Lee J. S., Lin C. H.. Polystyrene microplastic particles: In vitro pulmonary toxicity assessment. J. Hazard Mater. 2020:385121575. doi: 10.1016/j.jhazmat.2019.121575. [DOI] [PubMed] [Google Scholar]
  48. Wu Q., Liu C., Liu D., Wang Y., Qi H., Liu X., Zhang Y., Chen H., Zeng Y., Li J.. Polystyrene nanoplastics-induced lung apoptosis and ferroptosis via ROS-dependent endoplasmic reticulum stress. Sci. Total Environ. 2024:912169260. doi: 10.1016/j.scitotenv.2023.169260. [DOI] [PubMed] [Google Scholar]
  49. Schmidt, A. ; Mühl, M. ; Brito, W. ; Singer, D. ; Bekeschus, S. . Antioxidant Defense in Primary Murine Lung Cells following Short- and Long-Term Exposure to Plastic Particles. Antioxidants (Basel) 2023, 12(2). 227 10.3390/antiox12020227. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Smith R. A., Murphy M. P.. Animal and human studies with the mitochondria-targeted antioxidant MitoQ. Ann. N.Y. Acad. Sci. 2010:120196–103. doi: 10.1111/j.1749-6632.2010.05627.x. [DOI] [PubMed] [Google Scholar]
  51. Tang Y., Suo Y., Sun Z., Wu X., Xing Q., Bai Y.. Microplastics induce insulin resistance by causing mitochondrial dysfunction associated with mROS in skeletal muscle in vitro. Ecotoxicol Environ. Saf. 2025:302118585. doi: 10.1016/j.ecoenv.2025.118585. [DOI] [PubMed] [Google Scholar]
  52. Farshad A., Wehrend A.. MitoQ as a Mitochondria-Targeted Antioxidant in Sperm Cryopreservation: An Updated Review on Its Mechanisms, Efficacy, and Future Perspectives. Antioxidants. 2025;14(11):1350. doi: 10.3390/antiox14111350. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Chen Y., Nan Y., Xu L., Dai A., Orteg R. M. M., Ma M., Zeng Y., Li J.. Polystyrene nanoplastics exposure induces cognitive impairment in mice via induction of oxidative stress and ERK/MAPK-mediated neuronal cuproptosis. Part Fibre Toxicol. 2025;22(1):13. doi: 10.1186/s12989-025-00633-w. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Ji Y., Wang C., Wang Y., Fu L., Man M., Chen L.. Realistic polyethylene terephthalate nanoplastics and the size- and surface coating-dependent toxicological impacts on zebrafish embryos. Environ. Sci.: Nano. 2020;7(8):2313–2324. doi: 10.1039/D0EN00464B. [DOI] [Google Scholar]

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