Microplastic-Induced Macrophage Dysfunction Drives Lung Tumor Progression through Glutathione Imbalance

Abstract

Microplastics (MPs) are emerging contaminants whose immunological consequences remain poorly defined. Here, we investigated MP-induced immune responses using bone marrow-derived macrophages and a lung tumor model to delineate how MPs modulate tumor immunity. MPs triggered TLR2- and TLR4-dependent signaling pathways in macrophages, which initiated AP-1 signaling and lysosomal destabilization, followed by mitochondrial depolarization and excessive reactive oxygen species production. Despite NRF2 pathway activation, GPX1 and GPX3 were selectively suppressed, revealing a paradoxical uncoupling of glutathione metabolism that precipitated macrophage ferroptosis. In vivo, orally ingested MPs accumulated across multiple organs. In the lungs of tumor-bearing mice, MP exposure led to a time-dependent remodeling of the immune microenvironment, characterized by marked infiltration of M1-like macrophages and functional impairment of lymphocytes at later stages, which was accompanied by increased tumor burden. These findings identify an immune–redox–ferroptosis axis driven by glutathione imbalance and suggest redox disruption as a mechanistic link between microplastic exposure and tumor progression.

The dramatic increase in global plastic production and consumption has triggered plastic waste accumulation at an alarming level. Once released into the environment, plastic waste degrades physically, chemically, and biologically, forming microplastics (MPs, ≤5 mm) and nanoplastics (NPs, ≤1 μm). These particles are widely dispersed across terrestrial, marine, and atmospheric environments, raising concerns about human exposure through ingestion, inhalation, or direct bloodstream entry via medical devices. − MPs are present in the human tissues and circulation, representing potential health risks associated with their systemic toxicity. , Polystyrene MPs are extensively used in industrial and biomedical applications, positioning them in the primary focus of toxicity research. − Polystyrene MPs trigger inflammatory responses and induce tissue damage in multiple organs, including the lungs, liver, brain, and intestines. − These pathological outcomes are supposed to arise from aberrant immune cell activation, yet the molecular processes by which immune cells recognize polystyrene MPs and initiate downstream signaling pathways remain poorly defined.

Macrophages are pivotal in pathogen defense, immune regulation, and tissue homeostasis, initiating inflammation and coordinating programmed cell death pathways (including apoptosis, pyroptosis, and necroptosis) in response to stress signals. , MP exposure markedly increases reactive oxygen species (ROS) levels in macrophages, promoting inflammation and cellular dysfunction. To mitigate oxidative stress, cells rely on antioxidant defense systems, with the crucial role of glutathione metabolism. Glutathione is a key antioxidant that directly scavenges ROS or modulates oxidative balance through enzymes, such as glutathione peroxidase. Although glutathione dysregulation-associated ferroptosis has been reported in other cell types, the impact of MPs on the glutathione axis in macrophages remains poorly defined. Therefore, a comprehensive understanding of how the underlying mechanisms of MP-induced oxidative stress affect macrophage function is essential for elucidating the broader immunological consequences of MP exposure. Importantly, it remains unresolved whether dysregulation of glutathione metabolism specifically drives ferroptotic cell death in macrophages and contributes to immune-metabolic remodeling.

Previous studies have reported that MP exposure can promote tumor progression in various tissues, including the intestine, liver, and breast, with oxidative stress, chronic inflammation, and immune suppression proposed as major underlying mechanisms. − Notably, although ingested MPs are known to accumulate extensively in the lung during systemic distribution, evidence supporting their role in promoting lung tumorigenesis and remodeling the lung tumor immune microenvironment remains limited.

In this study, we aimed to investigate how MPs affect the function of macrophages and delineate the molecular cascade in which organellar stress-induced ROS imbalance disrupts glutathione homeostasis and culminates in ferroptosis. Moreover, by using two-photon intravital and three-dimensional organ imaging, we directly visualize real-time macrophage-MP interactions across multiple tissues and integrate these dynamic observations with mechanistic analyses. Notably, we demonstrate that MPs reprogram the lung tumor immune microenvironment by enhancing M1-like macrophage infiltration and amplifying inflammatory signaling, thereby fostering conditions favorable for tumor progression. These insights not only shed light on how MP-induced immune perturbations shape tumor immunity but also provide a critical foundation for designing and implementing future therapeutic interventions.

Results

BMDMs Phagocytose MPs

The characterization of polystyrene MP is presented in Figure A-D. Scanning electron microscopy (SEM) revealed that the particles were highly spherical with smooth surfaces (Figure A). Dynamic light scattering (DLS) analysis demonstrated optimal dispersion and colloidal stability in deionized water, with diameters ranging from approximately 937.0 to 1,025.4 nm (Figure B). A zeta potential of −39.92 mV indicated electrostatic stabilization in aqueous suspension (Figure C). Chemical characterization by Fourier transform infrared (FTIR) spectroscopy further confirmed the identity of polystyrene, as evidenced by characteristic vibrational peaks at 3,026 and 2,923 cm^–1 corresponding to aromatic C–H stretching, 1,641, 1,493, and 1,453 cm^–1 corresponding to aromatic CC stretching, and 748 and 697 cm^–1 corresponding to C–H out-of-plane bending (Figure D). These spectral features are consistent with the known vibrational modes of polystyrene, indicating the preservation of its chemical structure. Although nominally ∼1 μm, these submicron particles occupy the micro–nano interface, a dimensional regime where nanomaterials typically exhibit high surface-to-volume ratios and strong biointerface interactions.

1.

BMDMs sense, internalize, and transport MPs. (A) Scanning electron microscopy (SEM) image showing the morphology of MPs. Scale bar = 5 μm; scale bar (magnified) = 1 μm. (B) Size distribution of MPs measured by dynamic light scattering (DLS), showing a monodisperse peak at ∼1000 nm. (C) ζ-potential of MPs measured by laser Doppler electrophoresis, showing a surface charge −39.92 mV. (D) Fourier-transform infrared (FTIR) spectra of MPs confirm characteristic absorption peaks for polystyrene. (E) BMDMs were allowed to migrate on fibronectin-coated confocal dishes in the presence or absence of MPs for 30 min at 37 °C. Migration was tracked under each condition. x-y projections were measured at 50-μm intervals, then BMDM velocity and displacement rate were analyzed. Representative data were obtained from at least three independent experiments. (F) Representative images present BMDMs contacting and engulfing MPs (red) during migration. Yellow arrowheads: MP-interacting cells. Scale bars = 10 μm. (G) Transmission electron microscopy analysis revealed MP engulfment and intracellular damage in BMDMs migrating on fibronectin with MP exposure. Yellow arrowheads: MPs internalized by phagocytes. Red dotted squares (lower panels): magnified views of the upper panels. Scale bars = 5 μm (upper panels) or 1 μm (lower panels). (H) SEM analysis showed MP surface adherence to BMDMs during migration on fibronectin with MP exposure, revealing MP adherence to cell surfaces. Scale bars = 2 μm. (I) Actin-GFP-transfected BMDMs were allowed to migrate on MPs (red) and fibronectin-coated confocal dishes. Representative images show actin-GFP-expressing BMDMs transporting surface-adherent MPs (red) during migration. Yellow arrowheads: macrophage pseudopodia capturing and retracting MPs (red) toward the cell body. Scale bars = 10 μm. The data represent the results of at least three independent experiments and are expressed as the mean ± SEM, using unpaired two-tailed t tests. ***p < 0.001, **p < 0.01.

We investigated whether polystyrene MP exposure influenced bone marrow-derived macrophage (BMDM) cellular responses. Based on the dose–response analysis of MP-induced cellular stress, we selected 50 μg of MPs for further study, a concentration below the cytotoxic EC50 (82.56 μg) that elicited oxidative and inflammatory responses without substantial loss of cell viability (Figure S1A,B,D,E). Although this dose exceeds estimated single-exposure levels, it was empirically determined and widely used in nanotoxicology to model cumulative deposition and to ensure measurable immunological perturbations. − Over 90% of the BMDMs vigorously bound to polystyrene MPs while migrating on a fibronectin-coated surface (Figure S1C). Moreover, the migration speed and displacement extent of MP-treated BMDMs were significantly higher than those of control cells (Figure E and Video S1). BMDMs exhibited enhanced migratory activity toward polystyrene MPs. Sequentially, BMDMs strongly attached MPs to their cell surface in a sweeping manner (Figure F and Video S2). We further explored whether BMDMs endocytose polystyrene MPs. Transmission electron microscopy (TEM) analysis revealed morphological damage in the MP-treated BMDMs, including disrupted plasma membrane boundaries. Furthermore, spherical MPs appeared in the cytoplasm, indicating internalization (Figure G). SEM analysis confirmed MP adherence on the BMDM surface (Figure H).

To further investigate BMDM cytoskeletal morphology in the real-time polystyrene MP adherence process, we transfected BMDMs with actin-green fluorescent protein (GFP), allowing for BMDM morphology and motility visualization. Actin polymerization led to pseudopod extension, which bound to the polystyrene MPs and pulled them into the cell body (Figure I and Video S3). These results strongly suggest that BMDMs recognize, migrate toward, and eventually endocytose MPs, during which actin polymerization occurs.

BMDM Recognition of MPs via TLR2/4 Initiates Inflammatory Cascades

MPs are nonbiological particles without conventional recognition motifs. We therefore investigated the mechanism to drive MPs engulfment by BMDMs. Using live-cell actin-GFP imaging in BMDMs above, we found robust cytoskeletal remodeling and pseudopod formation directed toward MPs, demonstrating that BMDMs actively target these particles despite their nonbiological nature. To elucidate the upstream recognition pathways involved in this response, we analyzed the expression of pattern-recognition receptors. We found that BMDMs exposed to MPs exhibited increased expression of Toll-like receptor (TLR) 2 and TLR4 protein, similar to the pattern observed upon lipopolysaccharide (LPS) stimulation. Moreover, downstream signaling component expression was increased compared to control, particularly that of those related to activator protein 1 (AP-1), including Toll-interleukin-1 receptor domain-containing adaptor protein (TIRAP), myeloid differentiation primary response 88 (MyD88), tumor necrosis factor receptor-associated factor 6 (TRAF6), phosphorylated-c-Jun N-terminal kinase (P-JNK), and phosphorylated c-Jun (P-c-Jun) (Figure A). These findings suggest that MPs exploit macrophage recognition pathways reminiscent of pathogen-associated molecular patterns, notably through TLR2/4–AP-1 signaling.

2.

MPs induce inflammation by recognizing TLR2/4 in BMDMs. (A) Representative Western blot images of BMDMs treated with MPs or LPS (positive control) for 24 h. TLR2, TLR4, TIRAP, MyD88, TRAF6, P-JNK, and P-c-Jun protein expression levels, analyzed by Western blotting. β-actin expression was used as a loading CTL. (B) Relative mRNA expression levels of Il-1β, Il-6, and Tnf-α, measured in the presence of MP. Gapdh was used for normalization. (C) Representative flow cytometry plots presenting BMDM M1 and M2 polarization upon MP treatment. The M1 and M2 macrophage populations were quantified as CD86⁺CD206^– and CD86^–CD206⁺, respectively. (D) Representative immunofluorescence images of BMDMs treated with or without MPs for 24 h, stained with MPO (green), CitH3 (red), and DAPI (blue). Scale bars, 10 μm. Fluorescence intensity-based MPO and CitH3 quantification in BMDMs. a.u., arbitrary unit. The data represent at least three independent experiments and are expressed as the mean ± SEM, using one-way ANOVA. ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05.

Activation of this signaling cascade has been implicated in the induction of inflammatory responses. We therefore examined whether its activation was associated with increased expression of pro-inflammatory cytokine mRNA levels. The mRNA levels of interleukin (Il ) -1β, Il-6, and tumor necrosis factor (Tnf)-α were significantly elevated in BMDMs exposed to MPs compared with controls, confirming that this pathway drives a pro-inflammatory transcriptional response (Figure B).

To examine how MP exposure affects macrophage polarization, we analyzed phenotypic marker expressions, revealing an increased proportion of CD86-positive cells (M1 marker), whereas CD206-positive cell proportions (M2 marker) did not change considerably upon MP treatment (Figure C). Because M1 polarization has been associated with macrophage extracellular trap (MET) formation through cellular structural damage and nuclear envelope breakdown. , We further performed immunofluorescence analysis to assess myeloperoxidase (MPO) and citrullinated histone H3 (CitH3), two key components released during MET formation. In polystyrene MP-treated BMDMs, MPO and CitH3 surrounded internalized MP-containing BMDMs, showing a significant increase in MET formation (Figure D). Our findings suggest that MPs act as pathogen-associated molecular pattern (PAMP)-like stimuli and may serve as potent promoters of pro-inflammatory macrophage programming.

Lysosomal Destabilization Drives Mitochondrial Dysfunction in MP-Exposed Macrophages

TLR-mediated recognition of pathogens typically induces phagocytosis, followed by degradation of pathogens through the phagolysosomal pathway. In light of the pathogen-mimetic behavior of MPs, we investigated whether MP uptake similarly engages and perturbs lysosomal function in BMDMs. We first monitored galectin-3, which redistributes from a diffuse cytosolic pattern to discrete puncta upon lysosomal membrane permeabilization. Confocal imaging revealed a significant increase in galectin-3 puncta in MP-treated BMDMs compared with controls (Figure A). To further evaluate lysosomal rupture, we examined the spatial relationship between lysosome-associated membrane protein 1 (LAMP1), a membrane marker, and cathepsin B (CTSB), a luminal protease. Whereas CTSB colocalized with LAMP1 in control cells, MP exposure caused cytosolic and perinuclear dispersion of CTSB, indicating leakage of luminal contents (Figure B). Lysotracker fluorescence, indicative of intact and acidified lysosomes, was substantially reduced upon MP exposure (Figure C).

3.

MP exposure triggers both lysosomal and mitochondrial dysfunction in BMDMs. (A) Representative confocal images and quantification of galectin-3 puncta in BMDMs following MP exposure. Galectin-3 puncta (red, white arrowheads). Nuclei stained with DAPI (blue). Scale bars: 10 μm. (B) Representative confocal images of LAMP1 (red) and cathepsin B (CTSB, green) in BMDMs or without MP treatment. Nuclei stained with DAPI (blue). Scale bars: 10 μm. (C) Flow cytometric analysis of lysotracker fluorescence in BMDMs. (D) Representative flow cytometry dot plots showing mitochondrial membrane potential in BMDMs stained with JC-1 dye. The proportions of JC-1 aggregates (red) and monomers (green) were quantified. (E) Fluorescent confocal images of mitochondrial activity in MP-treated and untreated BMDMs, stained with MitoTracker Red CMXRos. Scale bars = 15 μm. The MitoTracker fluorescence intensity was quantified to calculate mean fluorescence intensity (MFI). (F) Flow cytometry analysis of mitochondrial ROS levels in BMDMs using MitoSOX staining. The figure contains representative MitoSOX fluorescence intensity histograms and quantifications. (G) Mitochondrial ATP levels were measured using a luminescence-based assay in MP-treated and untreated BMDMs. a.u., arbitrary unit. The data represent at least three independent experiments and are expressed as the mean ± SEM, using unpaired two-tailed t tests. ****p < 0.0001, ***p < 0.001, **p < 0.01.

Given the well-established lysosome–mitochondria crosstalk, we next assessed whether lysosomal disruption propagated to mitochondrial energetics by measuring mitochondrial membrane integrity with the 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) dye. The proportion of JC-1 aggregates in MP-treated groups was lower than that in control, whereas the proportion of JC-1 monomers increased (Figure D). Furthermore, we compared the fluorescence intensity of MitoTracker Red CMXRos, which stains mitochondria in a membrane potential-dependent manner. MP-treated cells showed reduced mean fluorescence intensity compared with that in control, thus indicating mitochondrial damage caused by MP exposure (Figure E). Subsequently, we assessed mitochondrial reactive oxygen species (ROS) levels using MitoSOX. MPs significantly increased ROS within the mitochondria (Figure F). Furthermore, mitochondrial adenosine triphosphate (ATP) production was substantially reduced in the MP-treated condition compared with that in control (Figure G). Therefore, MP uptake in BMDMs triggered lysosomal destabilization that coincided with mitochondrial depolarization, oxidative stress, and metabolic collapse, suggesting a cascade that compromises the bioenergetic and redox capacity essential for macrophage effector functions.

MP Exposure Uncouples NRF2 and GPX1/3, Disrupting Glutathione Balance

To determine how lysosomal and mitochondrial injury reprograms macrophage gene expression, we performed bulk RNA-seq on MP-exposed BMDMs. A total of 149 genes were differentially expressed (Padj <0.05; log₂ fold-change >1 or < −1), including 70 upregulated and 79 downregulated genes (Figure A). Gene Ontology enrichment analysis revealed significant changes in biological processes related to stress response, detoxification, and oxidative stress (Figure B). In line with these findings, MP exposure markedly increased intracellular levels of ROS and nitric oxide (NO) (Figure C,D).

4.

MP exposure impairs ROS detoxification in BMDMs through the dysregulation of the glutathione pathway. (A) Heatmap showing gene expression patterns with differential expressions. (B) Gene ontology (GO) enrichment analysis of DEGs in MP-treated BMDMs. Top stress-related pathways, including response to oxidative stress and detoxification, are ranked by the normalized enrichment score (NES). (C) ROS production in MP-treated BMDMs, quantified relative to CTL. (D) Nitrite production in MP-treated BMDMs, quantified relative to CTL. (E) Gene set enrichment analysis (GSEA) of the oxidative stress response gene set based on total RNA-seq data from MP-treated BMDMs compared with that from untreated CTL cells. The enrichment score curve and NES indicate substantial up-regulation of the oxidative stress response genes in MP-treated BMDMs. (F) Heatmap showing the differential expression of oxidative stress response-involved genes in MP-treated BMDMs compared with that of untreated CTL cells. Genes up- and down-regulated by MP exposure are marked in red and blue, respectively. The most substantially up- and down-regulated glutathione metabolism-related genes are highlighted. Glutathione metabolism-related genes, such as Gclm, Slc7a11, Nqo1, Gclc, Gpx1, and Gpx3, are highlighted. (G) Volcano plot depicting differential expressions of glutathione metabolism-related genes in MP-treated BMDMs compared with that in CTL cells. (H) Representative Western blot images showing the expression of NRF2 and that of its downstream antioxidant-related targets (GCLM, SLC7A11, NQO1, and GCLC) in BMDMs treated with MPs for 24 h. (I) Glutathione-dependent antioxidant enzyme GPX1 and GPX3 expression was analyzed using β-actin as a loading CTL. (J) GSH/GSSG ratio and total glutathione level quantifications in MP-treated BMDMs relative to the CTL cells. a.u., arbitrary unit. The data represent at least three independent experiments and are expressed as the mean ± SEM, using unpaired two-tailed t tests. ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05.

To delineate redox regulatory pathways affected by MPs, we performed gene set enrichment analysis (GSEA). It revealed that MP exposure-up-regulated genes were significantly associated with oxidative stress responses, among which the most up- and down-regulated genes following MP treatment were both associated with glutathione metabolism (Figure E-G). Excessive ROS and NO production typically activate glutathione metabolism, which acts as a compensatory defense against oxidative stress. In this process, nuclear factor erythroid 2-related factor 2 (NRF2) is initially activated and subsequently up-regulates the expression of antioxidant-related genes, such as Gclm, Slc7a11, Nqo1, and Gclc. , Measurement of protein expression levels revealed a considerable increase in the expression of NRF2, glutamate cysteine ligase modifier (GCLM), solute carrier family 7 member 11 (SLC7A11), NAD­(P)H quinone dehydrogenase 1 (NQO1), and glutamate-cysteine ligase catalytic subunit (GCLC) in response to MP treatment (Figure H). Glutathione peroxidase (GPX)­1 and GPX3 play critical roles in glutathione metabolism by neutralizing excessively generated oxidative species. These enzymes reduce hydrogen peroxide (H₂O₂) to water (H₂O) and oxygen (O₂) through redox reactions involving glutathione. However, in BMDMs treated with MPs, the protein and mRNA expression levels of GPX1 and GPX3 decreased (Figures I and S1G,H). Furthermore, both the total glutathione levels and the glutathione (GSH)/glutathione disulfide (GSSG) ratio were lower in the MP-treated condition than in control (Figure J). These findings suggest that although NRF2-driven antioxidant responses are activated, MPs paradoxically suppress GPX1 and GPX3, thereby disrupting glutathione homeostasis and sustaining redox imbalance in macrophages.

Glutathione Imbalance Mediates Ferroptotic Death in MP-Exposed Macrophages

MPs impair lysosome and mitochondria in BMDMs, thus inhibiting glutathione metabolism that neutralizes excessive ROS production. Consequently, ROS continuously accumulate within the cells and may trigger an inflammatory response that could ultimately lead to cell death. To investigate this mechanism, we examined whether MP exposure induces cell death and, if so, subsequently determined the type of cell death that occurs. Live-cell migration imaging revealed that BMDMs extended their pseudopodia to make contact with MPs. As migration progressed, the cells contracted and eventually died (Figure A and Video S4). We conducted apoptosis and necrosis assays to determine the type of cell death. MP-treated BMDMs exhibited a notable increase in necrotic cell death compared with that of control cells (Figure B). These necrotic features are consistent with regulated necrosis, raising the possibility that MPs engage ferroptotic pathways.

5.

MP-induced ferroptosis in BMDMs through oxidative stress and iron accumulation. (A) Representative images showing BMDMs engulfing MPs (red) during migration, followed by cell death over time. Yellow arrowheads: cells that captured MPs and subsequently underwent cell death. Scale bars = 10 μm. (B) Representative flow cytometry dot plots of annexin V and DAPI staining in MP-treated and untreated BMDMs. Necrotic cell death was quantified as Annexin V⁺ DAPI⁺; live cells were Annexin V^– DAPI^–. (C) KEGG pathway enrichment of up-regulated genes in MP-treated BMDMs, highlighting the ferroptosis pathway. (D) The fluorescent confocal images showing intracellular iron accumulation in MP-treated and untreated BMDMs, stained with FerroOrange. Scale bars = 15 μm. The FerroOrange fluorescence intensity was quantified to calculate MFI. (E) The relative MFI of oxidized C11-BODIPY was measured in the presence of MPs. (F) Ferroptosis-associated GPX4, FTL, and FTH1 expression level analysis. β-actin was used as a loading CTL. a.u., arbitrary unit. The data represent at least three independent experiments and are expressed as the mean ± SEM, using unpaired two-tailed t tests. ****p < 0.0001, ***p < 0.001, **p < 0.01.

Gene expression analysis using Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment further revealed that genes up-regulated following MP treatment were closely associated with the ferroptosis pathway (Figure C). Ferroptosis is an iron-dependent form of cell death, driven by dysregulated intracellular iron homeostasis. , To investigate iron accumulation in BMDMs following MP treatment, we used the Fe²⁺-selective fluorescent probe FerroOrange and observed a pronounced increase in iron ions within the cells (Figure D). Furthermore, lipid peroxidation, a hallmark of ferroptosis, was also observed, as indicated by an increase in oxidized boron-dipyrromethene (BODIPY)-C11 fluorescence in the MP-treated condition (Figure E and S1F). Consistent with ferroptotic features, the protein expression levels of GPX4, a key inhibitor of ferroptosis, and iron-regulatory proteins ferritin light chain (FTL) and ferritin heavy chain 1 (FTH1) were decreased in MP-treated cells (Figure F). These results strongly suggest that MPs activate the ferroptosis pathway, which plays a critical role in MP-induced cell death.

Ingested MPs Promote Lung Tumor Progression

Orally ingested MPs are known to distribute to multiple organs, with the lung representing a prominent site of deposition due to its extensive vascularization and dense macrophage population. To evaluate the systemic distribution and immunological effect in vivo, we orally administered polystyrene MPs to wild-type and CX3CR1-GFP mice. Accumulation of MPs across organs was visualized using 3D whole-organ imaging, and their interaction with CX3CR1-GFP⁺ macrophages was monitored via two-photon intravital microscopy. Red fluorescence-labeled MPs were found embedded in the lung parenchyma, showing conspicuous accumulation following oral exposure (Figure A and Video S5). We next performed H&E staining to evaluate the effect of MP exposure on lung tissue integrity. Histological analysis revealed that MP-exposed lungs displayed a markedly thickened alveolar wall and diffuse alveolar injury compared with controls (Figure B). In addition, MP-exposed tissues exhibited increased nuclear density in the alveolar walls, consistent with enhanced infiltration of innate immune cells. Moreover, two-photon intravital imaging of the lungs showed red fluorescent MPs distributed throughout the tissue, and a subset of GFP-positive macrophages exhibited yellow fluorescence due to colocalization with MPs, thus indicating phagocytic uptake (Figure C and Video S6). Notably, we observed cell–cell interactions wherein one macrophage actively migrated toward an MP-laden macrophage and established direct contact with the MP-containing region, which was reminiscent of a “knocking” behavior (Figure C and Video S6). Such dynamic behaviors, rarely captured in vivo, suggest that MPs actively engage intercellular communication circuits rather than being passive contaminants. Collectively, these findings demonstrate that ingested MPs can accumulate in the lung, where they interact with immune cells and alter their behavior. Specifically, MPs promoted cell–cell interactions and enhanced phagocytic activity, indicating that MPs are not merely passive contaminants but may dysregulate pulmonary immune responses.

6.

Ingested MPs accumulate in the lung, promoting tumor progression in mice. (A) The 3D organ images of lung showing the distribution of MPs (red) in MP-exposed and nonexposed mice. White arrowhead: MPs. Scale bars = 1500 μm. (B) Representative lung sections stained with H&E from nonexposed and MP-exposed mice. (C) Time-lapse two-photon intravital images of mouse lung showing tissue-resident macrophages (green) interacting with MPs (red) over time following 1 week of oral gavage. White arrowheads: macrophages in contact with MPs. Following 1 week of oral administration of MPs, mice were intravenously injected with LLC cells (designated as day 0) to induce lung metastasis. Oral gavage of MPs was continued three times per week throughout the experimental period. (D) Body weight changes in LLC and LLC+MP groups during tumor progression. (E) Quantification of RFP-positive LLC tumor cells in lung tissues by flow cytometry at the indicated time points following intravenous LLC injection. (F) Representative images of whole lung tissues isolated from tumor-bearing mice with or without MP exposure and quantification of the number of lung tumor nodules in LLC and LLC+MP mice at day 14. White arrowheads: metastatic tumor nodules. (G) Whole-organ 3D images of lungs from tumor-bearing mice subjected to 1-week oral MP administration and concurrent intravenous injection of LLC-RFP cells. The figure presents metastatic LLC-RFP cancer cells (red) and autofluorescent tissue structures (green), potentially associated with MP exposure. White arrowheads: regions with extensive tumor formation. Scale bars = 1000 μm. (H) Representative H&E-stained lung sections from CTL, LLC and LLC+MP mice, showing increased tumor burden in MPs-treated mice. Scale bars = 500 μm. (I,J) Quantitative real-time PCR analysis of hypoxia and angiogenesis-related genes in lung tissues, including Hif1α (I) and Vegfa (J) normalized to LLC controls. All the data represent at least four independent experiments and are expressed as the mean ± SEM, using unpaired two-tailed t tests. **p < 0.01, *p < 0.05.

Persistent perturbation of pulmonary immune responses is tightly associated with chronic inflammation, a key driver of lung cancer progression. − We therefore assessed whether MP accumulation in the lung contributes to tumor progression by establishing an in vivo model of MP exposure. Mice were orally administered MPs for 1 week prior to intravenous injection of red fluorescent protein (RFP)-fused Lewis lung carcinoma (LLC) cells (designated as day 0), followed by continued MP administration throughout the experimental period, and lung tumor progression was monitored at the indicated time points. Systemic changes associated with MP exposure and tumor progression were first evaluated by monitoring body weight over time (Figure D). From day −7 to day 7, no significant differences in body weight were observed between the LLC and LLC+MP groups, whereas body weight was significantly reduced in the LLC+MP group at day 14 (Figure D). To directly assess lung tumor burden over time, the proportion of LLC cells in the lung was quantified by flow cytometry. While no significant differences were detected between the LLC and LLC+MP groups at early time points (day 0 to day 7), a significant increase in the proportion of LLC cells was observed in the LLC+MP group at day 14 (Figure E). Consistent with the flow cytometric findings, gross examination of lung tissues revealed a significantly higher number of metastatic tumor nodules in the LLC+MP group compared with LLC controls at day 14 (Figures F and S2A). To further visualize tumor burden throughout the lung, 3D imaging of cleared lung tissues revealed more extensive distribution of RFP-positive tumor masses in MP-exposed mice (Figure G and Video S7).

Histological analysis further revealed enhanced tumor progression in the lungs of MP-exposed mice. H&E-stained lung sections showed more extensive tumor progression and disrupted alveolar wall in the LLC+MP group compared with LLC controls (Figure H). Given the increased tumor burden and histological progression, the expression of hypoxia- and angiogenesis-related genes was measured in lung tissues. Quantitative analysis revealed that the expression levels of Hif1α and Vegfa were significantly increased in the LLC+MP group (Figure I,J). These findings suggest that MP exposure disrupt pulmonary immune homeostasis, thereby contributing to the establishment of a pro-tumorigenic microenvironment.

MPs Reshape Lung Tumor Microenvironment

MP exposure has been implicated in tumor progression in various organs, but its role in shaping the lung tumor microenvironment remains poorly understood. We therefore investigated whether MPs remodel the lung immune microenvironment to promote tumor progression. To assess the impact of MPs on the immune microenvironment, immune cell populations in lung tissues from both experimental groups were analyzed by flow cytometry (Figure S2B). We first assessed temporal changes in total immune cell infiltration by quantifying CD45⁺ immune cells in the lung. From day 0 to day 7, the absolute number of CD45⁺ immune cells remained comparable between the LLC and LLC+MP groups. In contrast, at day 14, an increase in the total number of CD45⁺ immune cells was observed in MP-exposed mice (Figure B).

7.

MP exposure alters the lung immune microenvironment during tumor progression. (A) Total number of CD45⁺ immune cells in lung tissues from LLC and LLC+MP mice at the indicated time points. (B) Quantification of lung immune cell numbers, including IM, AM, NK cells, CD8 T cells, and CD4 T cells, in LLC and LLC+MP lung tissues at day 14. (C) Relative mRNA expression levels of Ccl2 in lung tissues from LLC and LLC+MP mice. (D,E) Quantification of mean fluorescence intensity (MFI) of CD86 (M1-associated marker) and CD206 (M2-associated marker) in IM in the lungs of tumor-bearing mice at day 14. (F) Quantification of intracellular ROS levels in lung tissues using DCFDA-based flow cytometry. (G) Immunoblot analysis of GPX4, GPX1, and GPX3 expression in lung tissues from LLC and LLC+MP mice with or without ferroptosis inhibitor (Fer-1) treatment. β-actin was used as a loading control. (H-J) Quantification of GPX4 (H), GPX1 (I), and GPX3 (J) protein expression normalized to β-actin. (K) Representative H&E-stained lung sections from LLC+MP mice with or without Fer-1 treatment. Scale bars = 500 μm. All the data represent at least four independent experiments and are expressed as the mean ± SEM, using unpaired two-tailed t tests and one-way ANOVA. ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05.

To further characterize immune cell populations during the early phase of tumor progression (day 0–7), we analyzed major immune cell subsets in the lung. No significant differences were observed in the numbers of infiltrating macrophages (IM), alveolar macrophages (AM), CD8 T cells, and CD4 T cells between the two groups. However, natural killer (NK) cells showed an increase in MP-exposed mice during this early phase (Figure S2C-E). At day 14, the number of IM showed the marked increase among all immune subsets analyzed and was significantly enhanced in MP-exposed mice compared with LLC controls (Figure B). Consistent with the marked expansion of IM, expression of the macrophage-recruiting chemokine C–C motif chemokine ligand 2 (Ccl2) was significantly elevated in lung tissues from MP-exposed mice (Figure C). Moreover, the numbers of NK cells and CD8 T cells were also increased (Figure C). At day 14, among IM, the M1 phenotype was significantly increased following MP exposure, whereas the M2 phenotype remained unchanged (Figure D,E). A similar pattern was observed in AM, in which the M1 phenotype was significantly increased in MP-exposed mice, while the M2 phenotype showed no significant change (Figure S3A,B). Additionally, quantitative analysis of lung tissues revealed significantly elevated mRNA expression levels of the tumor-associated cytokines Il-10 and transforming growth factor beta 1 (Tgf- β1) in MP-exposed mice (Figure S3C,D).

Given the increased numbers of NK and T cell populations at day 14, we further assessed their functional states within the lung tumor microenvironment. Flow cytometric analysis revealed increased Annexin V staining in NK cells, CD8 T cells, and CD4 T cells from MP-exposed mice, consistent with elevated levels of cell death (Figure S3E). Furthermore, Ki-67 expression was reduced in NK cells, whereas no significant difference was observed in T cell populations (Figure S3F). In addition, intracellular cytokine staining demonstrated increased interferon gamma (IFN-γ) production in NK cells from MP-exposed mice (Figure S3G). Moreover, expression of Granzyme B, a key effector molecule for cytotoxic function, was increased in CD8 T cells and CD4 T cells at day 14 (Figure S3H).

We next examined whether MP exposure alters the redox status in the lung tumor microenvironment at day 14. Flow cytometric analysis revealed significantly elevated intracellular ROS levels in lung tissues from LLC+MP mice compared with LLC controls (Figure F). Immunoblot analysis of ferroptosis-related antioxidant enzymes in lung tissues showed that the protein levels of GPX4, a central regulator of ferroptosis, were reduced in the LLC+MP group compared with LLC controls (Figure G,H). Moreover, treatment with the ferroptosis inhibitor ferrostatin-1 (Fer-1) modestly restored GPX4 protein levels in MP-exposed mice (Figure G,H). Consistent with impaired glutathione-dependent antioxidant defense, the protein expression levels of GPX1 and GPX3, were also markedly decreased in the LLC+MP group compared with LLC controls (Figure G,I,J). This pattern was similar to the in vitro findings observed in MP-treated macrophages, suggesting that MP exposure impairs glutathione metabolism in vivo. In addition, Fer-1 treatment partially restored GPX1 and GPX3 expression in MP-exposed tumor-bearing mice (Figure G,I,J). H&E staining of lung sections revealed that tumor burden appeared attenuated in LLC+MP mice treated with Fer-1 compared with untreated LLC+MP mice (Figure K). These findings support that MP exposure is associated with extensive immune remodeling, oxidative stress, and ferroptosis-related alterations in the lung tumor microenvironment.

Discussion

This study demonstrates that MPs, beyond being passive environmental pollutants, act as immunologically active agents that perturb immune homeostasis and could potentially contribute to tumorigenesis. By integrating in vitro and in vivo studies, we reveal a novel pathway through which MPs trigger innate immune activation, lysosomal damage, redox imbalance, and regulated cell death. The 1 μm particles used in this study bridge the micro- and nanoplastics, and their aggregation-driven physicochemical features likely underlie the observed immune activation. Our findings reveal that MP exposure induces glutathione imbalance and macrophage dysfunction, leading to immune-ferroptosis remodeling and an immunosuppressive microenvironment associated with lung tumor progression. −

We observed that macrophages recognize MPs through TLR2 and TLR4, thereby triggering actin polymerization and phagocytosis. Although previous studies have implicated the importance of nuclear factor kappa B pathway in MP-induced inflammation, our findings demonstrate a distinct activation of the AP-1 signaling axis, indicating a previously unrecognized immune-sensing mode specific to environmental particles. This signaling difference is likely attributable to the unique physicochemical MP properties (e.g., size and composition) and the functional heterogeneity among cell types. Furthermore, AP-1 signaling activation upon MP recognition might promote M1-like macrophage polarization, consistent with our observation of increased CD86 expression in vitro. The promotion of actin polymerization and MET formation further highlights the multifaceted innate immune responses elicited by MP exposure, beyond classical phagocytosis.

Various cell types have been reported to exhibit increased ROS generation upon MP exposure. , In line with previous studies, we found that MP exposure led to increased ROS levels in BMDMs. Notably, while ROS increased in an MP concentration-dependent manner, the key glutathione peroxidases GPX1 and GPX3 displayed a biphasic response showing an increase at low doses (0–10 μg) but a reduction at ≥50 μg. This pattern suggests that antioxidant defense programs may be adaptively induced under mild oxidative stress. However, beyond a certain exposure threshold, the detoxification capability of the glutathione-dependent system may become functionally limited, which results in insufficient ROS neutralization.

Moreover, total RNA sequencing analysis demonstrated that MP exposure modulates the expression of genes involved in glutathione metabolism in BMDMs, with both up-regulation and down-regulation observed. Although MP exposure up-regulated antioxidant genes and proteins, including NRF2, GCLC, GCLM, NQO1, and SLC7A11, the expression of key GSH peroxidases, such as GPX1 and GPX3, was paradoxically down-regulated at both the transcript and protein levels. This indicates that the enzymatic system essential for effective ROS detoxification remains functionally impaired. This selective GPX enzyme suppression was accompanied by intracellular GSH level reductions, most likely due to both impaired synthesis and increased consumption in response to oxidative stress. Lysosomal and mitochondrial damage can cascade to other organelles, including the cytosol and endoplasmic reticulum, which support GPX1 and GPX3 synthesis. Such impairment may suppress GPX production and amplify redox imbalance. − In summary, these findings suggest that MPs disrupt both the biosynthetic and effector arms of the GSH system, thereby undermining redox homeostasis and promoting sustained oxidative stress in BMDMs. We propose that this dysfunction initiates a pathological feedback loop, in which impaired antioxidant defense exacerbates ROS accumulation, thereby amplifying MP-induced immunotoxicity.

MP exposure promotes ferroptosis and necrotic cell death in BMDMs, leading to inflammatory amplification and oxidative injury, as revealed by live-cell imaging. Among these, the necrotic pathway appears to amplify local inflammation via pro-inflammatory cytokine release, which in turn activates neighboring immune cells. Concurrently, excessive ROS-induced lipid peroxidation triggers ferroptosis, resulting in the loss of membrane integrity and dysfunction of intracellular organelles. These findings implicate a feedback loop that leads to oxidative injury, which consequently promotes immune cell death and ultimately exacerbates inflammation and amplifies ROS generation.

The systemic accumulation of orally administered MPs in multiple organs, including the brain, lungs, intestines, and lymph nodes, suggests that these particles can translocate across epithelial barriers and disseminate through the circulatory or lymphatic systems, with potential implications for multiorgan toxicity and immune modulation. Among these organs, the lung has been reported as the second most prominent site of MP deposition despite oral exposure, underscoring its unexpected vulnerability. Accumulation in these organs has been linked to pathological alteration, among which tumorigenic effects have been documented in the liver, intestine, and other organs. However, their contribution to lung cancer has remained largely unexplored. This limited understanding may reflect the prevailing perception that lung tumorigenesis is primarily driven by smoking and air pollution, leading to an underestimation of MPs as potential contributors. Our study addresses this gap by demonstrating that MPs accumulate in the lung, disrupt immune homeostasis, and exacerbate tumor burden, thereby identifying MPs as an additional environmental factor in pulmonary carcinogenesis.

In support of this, our in vivo data indicate that MP exposure is associated with a progressive reshaping of the lung tumor microenvironment and an increased lung tumor burden. The expansion of IMs was the most prominent immune change at day 14, and these IMs exhibited an M1-like inflammatory phenotype. While acute inflammatory macrophage activation can support antitumor immunity, persistent or excessive inflammatory polarization may become maladaptive and contribute to tumor-promoting inflammation, a recognized driver of tumor progression. − These findings suggest that MP exposure promotes a macrophage-centered immune remodeling program in established tumors.

Despite increased infiltration of NK and CD8 T cells at late stages, the MP-exposed lung tumor microenvironment exhibited features of functional immune dysregulation rather than effective antitumor immunity. The coexistence of enhanced cell death susceptibility and imbalanced cytotoxic signaling suggests that MP-induced inflammatory and oxidative stress may drive partial immune activation while simultaneously promoting activation-induced cell death or functional exhaustion of cytotoxic lymphocytes. These results highlight that numerical expansion of lymphocyte populations does not necessarily translate into improved antitumor immunity in MP-exposed lungs. MP exposure markedly disrupted redox homeostasis in tumor-bearing lungs, characterized by elevated ROS levels and reduced expression of the ferroptosis-regulating antioxidant enzymes GPX4, GPX1, and GPX3. Pharmacological inhibition of ferroptosis with ferrostatin-1 partially restored GPX expression and was associated with reduced lung tumor burden, supporting a functional link between MP-induced oxidative stress, ferroptosis-related dysregulation, and tumor progression. These findings suggest that MPs promote a tumor-permissive microenvironment through immunometabolic and redox imbalance, rather than through a nonspecific inflammatory response alone. Although our results are consistent with emerging evidence linking environmental pollutants with increased cancer risk, further clinical investigation will be required to define the relevance of MP exposure to human lung tumor progression.

Despite these insights, a few critical questions should be further addressed. Our studies might not fully reflect the complex immune interactions present in humans. Further research is required to determine whether pharmacological or genetic modulation of glutathione-related pathways can mitigate the sustained oxidative stress induced by MP exposure. Moreover, the potential heterogeneity in immune responses driven by variations in MP size, surface charge, or polymer composition was not fully addressed in this study. Consequently, future studies should focus on developing more refined human exposure models, elucidating the interactions among different immune cell types in response to MPs, and exploring targeted approaches to disrupt the feedback loops identified in this study.

By connecting cellular mechanisms to systemic outcomes, our findings offer a conceptual framework for understanding the health risks associated with MP exposure. Given the pervasive presence of MPs in the environment, these results raise important public health concerns and underscore the need for future therapeutic and regulatory strategies to mitigate the immunological and oncogenic risks posed by MPs.

Conclusion

In this study, we demonstrated that MPs are not passive environmental contaminants but active immunomodulatory agents that perturb macrophage redox balance, promote ferroptosis, and lung tumor progression. Our research provides a new clue for understanding the aberrant role of MPs in macrophage-mediated immune response and underscores the necessity to address immunological consequences in various chronic diseases, including cancer.

Materials and Methods

Polystyrene MPs Characterization

Polystyrene MPs were purchased from Polysciences (08226-15) and Thermo Fisher Scientific (F13083). Plain MPs (Polybead carboxylate 1.0 μm; actual size: 990 nm), and red fluorescent MPs (Fluospheres, polystyrene carboxylate 1.0 μm; actual size: 950 nm, labeled with red; excitation 580 nm/emission 605 nm) were provided as 2% solids (w/v). The hydrodynamic diameter and ζ-potential of MPs were measured using a particle size and ζ-potential analyzer (ELS-Z1000, Otsuka Electronics, Hirakata, Osaka, Japan). Dynamic light scattering (DLS) and electrophoretic light scattering (ELS) modes were used to assess particle size distribution and surface charge, respectively. For Fourier-transform infrared (FTIR) spectroscopy, lyophilized MPs were analyzed using an INVENIO FT-IR spectrometer (Bruker, Billerica, MA, USA) equipped with an ATR accessory. Spectra were collected over the range of 8000–350 cm^–1 to confirm polystyrene-specific absorption bands.

Mice

C57BL/6 mice (Orient Bio) and CX3CR1-GFP mice were maintained in a specific pathogen-free environment at Yonsei University College of Medicine. All animal experiments were approved by the Institutional Animal Care and Use Committee of Yonsei University College of Medicine (Ethics Approval Number 2023-0118).

Macrophage Preparation

Bone marrow (BM) was isolated from the tibia and femur of C57BL/6 mice by flushing. The isolated BM precursors were cultured at 37 °C with 5% CO₂ in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with mouse macrophage colony-stimulating factor (20 ng/mL), 10% fetal bovine serum, and 1% penicillin/streptomycin/amphotericin B (PSA) for 4 days to differentiate into BMDMs.

Cell Cytotoxicity Assay

BMDMs were seeded at a density of 1 × 10⁵ cells/well in a 96-well plate for 24 h. The cells were then treated with 500 μM H₂O₂ and varying concentrations of polystyrene MPs (0, 5, 10, 50, 100, and 500 μg). After incubation for 24 h, methyl thiazole tetrazolium bromide (MTT; Amresco, 0793) was added, and the cells were further incubated for 3 h at 37 °C. After removing the media, dimethyl sulfoxide (DMSO; Sigma, D8418) was added for 15 min and the plate was read with a microplate spectrometer (BioTek, Winooski, VT, USA) at 570 nm. EC50 was calculated in GraphPad PRISM 10.6.1 using nonlinear regression with three parameters.

Electron Microscope

For transmission electron microscopy (TEM) analysis, BMDMs (1 × 10⁶ cells/ml) were seeded in a fibronectin-coated (10 μg/mL) confocal dish and incubated for 24 h at 37 °C with 5% CO₂. After treating BMDMs with 50 μg of polystyrene MPs for 24 h, the MP-exposed BMDMs were fixed for 12 h in Karnovsky’s fixative (0.1 M phosphate buffer [PB; pH 7.4] containing 2% glutaraldehyde and 2% paraformaldehyde). The 0.1 M PB was then used for washing and dehydration in an ascending ethanol series. The cells were embedded using the Poly/Bed 812 Kit (Polysciences) and stained with toluidine blue for observations under an optical microscope. Images were captured using a JEM-1011 TEM instrument (JEOL, Tokyo, Japan). For scanning electron microscopy (SEM) analysis, BMDMs were fixed for 24 h in Karnovsky’s fixative under the same conditions as for TEM and then washed twice for 30 min in 0.1 M PB. The samples were postfixed with 1% OsO₄ for 2 h and dehydrated in an ascending ethanol-gradient series (50–100%) using a Critical Point Dryer (Leica EM CPD300, Wetzlar, Germany). Thereafter, samples were coated with platinum using a sputter coater (Leica EM ACE600) and observed under a field-emission SEM (Merlin, Zeiss, Oberkochen, Germany).

Migration Analysis

For live imaging of BMDMs, cells were seeded in fibronectin-coated confocal dishes with Hanks’ Balanced Salt Solution (HBSS) containing 10% fetal bovine serum (FBS) and were incubated at 37 °C with 5% CO₂ for 24 h. Thereafter, the BMDMs were stimulated with or without polystyrene MPs for 30 min, respectively. BMDM migration was then observed for 30 min using an Eclipse Ti2 fluorescence microscope (Nikon, Tokyo, Japan). Phagocytosis and cell death-image data were analyzed using Volocity software (PerkinElmer, Waltham, MA, USA), and calculations were performed using MATLAB software. For fluorescent live imaging, BMDMs were seeded in 24-well plates at a density of 2 × 10⁵ cells per well. When cells reached 70–90% confluence, they were transfected with the actin-GFP plasmid using Lipofectamine 3000 (Invitrogen, L3000001). Five hours after transfection, the BMDMs were preincubated for 24 h in DMEM medium supplemented with 10% FBS and 1% PSA. MP-treatment conditions were the same as those used for the BMDMs in the previous step. Images were captured under the same conditions as those for nonfluorescent live cell imaging. Migration tracks were analyzed using Volocity and Fiji (NIH) software, and all migration data were obtained from the entire image sequence.

ROS Measurements

To measure intracellular ROS levels, BMDMs were plated in fibronectin-coated confocal dishes. Samples were then treated with or without polystyrene MPs (50 μg) at 37 °C with 5% CO₂ for 24 h. Thereafter, the culture medium was removed, the cells were washed with 1× ROS buffer, and the cells were then incubated for 30 min at 37 °C with 2′, 7′-dichlorofluorescin diacetate (DCFDA, 20 μM; Abcam, ab113851) solution for 45 min at 37 °C in a 5% CO₂ incubator in the dark. The cells were then gently rinsed thrice with 1× ROS buffer. Subsequently, fluorescent-cell imaging was performed for each group using an Eclipse Ti2 fluorescence microscope, and the fluorescent-cell images were analyzed using Volocity and Fiji software. To measure mitochondrial ROS levels, BMDMs were plated in six-well plates with or without polystyrene MPs at 37 °C with 5% CO₂ for 24 h. All samples were harvested and washed in FACS buffer (PBS containing 2% FBS and 2 mM ethylenediaminetetraacetic acid). BMDMs were stained with MitoSox (2.5 μM; Invitrogen, M36009) for 30 min. Samples were measured via flow cytometry analysis using BD FACS LSR II and were analyzed using FlowJo software (TreeStar, Ashland, OR, USA).

Quantification of Nitrite Production

BMDMs were plated in fibronectin-coated confocal dishes. Samples were treated with or without polystyrene MPs (50 μg) at 37 °C with 5% CO₂ for 24 h. After incubation, 50 μL of supernatant was dispensed into 96-well plates, 50 μL of sulfanilamide solution was added to each well, and the wells were incubated for 15 min at 25 °C in the dark. Subsequently, 50 μL of N-1-naphthylethylenediamine dihydrochloride solution (Promega, G2930) was added to each well, and the cells were further incubated for 10 min under identical conditions. Finally, the absorbance values at 540 nm were measured using a microplate spectrometer.

Necrosis and Apoptosis Assays

BMDMs were plated in six-well plates and treated with or without polystyrene MPs at 37 °C with 5% CO₂ for 24 h. Thereafter, the samples were harvested and washed in cold Annexin V-binding buffer (BioLegend, 422201). Flow cytometry was performed using 4′,6-diamidino-2-phenylindole (DAPI; Thermo Fisher, D1306) and fluorescein isothiocyanate (FITC)-annexin V antibodies (BioLegend, 640906). Cells positive for both FITC and DAPI were identified as necrotic, whereas FITC-positive cells were classified as apoptotic. Cells negative for both FITC and DAPI represented live cells. Samples were analyzed by flow cytometry using BD FACS LSR II and evaluated using FlowJo software.

Immunofluorescence

BMDMs were plated in fibronectin-coated confocal dishes and treated with or without polystyrene MPs (50 μg) at 37 °C with 5% CO₂ for 24 h. To detect METs, cells were fixed in 4% paraformaldehyde for 20 min, rinsed three times with PBS, permeabilized with 0.1% Triton/PBS for 5 min, and blocked with 1% bovine serum albumin/PBS for 1 h. The cells were then incubated overnight with anti-CitH3 (Abcam, ab5103) and FITC-anti-MPO (Abcam, ab90812) antibodies, followed by staining with Alexa Fluor 555 goat antirabbit Immunoglobulin G (2 μg/mL; Abcam, ab150078) for 2 h at room temperature in the dark. To assess intracellular ferrous ions, cells were stained with FerroOrange (10 μM) in HBSS for 30 min at 37 °C. To assess mitochondrial damage, cells were loaded with MitoTracker RedCMXRos (50 nM; Invitrogen, M7512) for 30 min. To assess lysosomal membrane permeabilization, cells were first stained for galectin-3. Cells were incubated overnight at 4 °C with antigalectin-3 antibody (Santa Cruz Biotechnology, sc-32790), followed by Alexa Fluor 647–conjugated goat antimouse IgG secondary antibody (2 μg/mL; Abcam, ab150115) for 2 h at room temperature in the dark. In a separate experiment, cells were stained for the lysosomal markers LAMP1 (Santa Cruz Biotechnology, sc-20011) and cathepsin B (CTSB; Abcam, ab214428). Cells were stained with Alexa Fluor 647–conjugated goat antimouse IgG (2 μg/mL, Abcam, ab150115) and Alexa Fluor 488–conjugated goat antirabbit IgG (2 μg/mL; Abcam, ab150077) for 2 h at room temperature in the dark. After the respective staining procedures, cells were rinsed with PBS three times and mounted with ProLong Gold Antifade DAPI mounting reagent (Invitrogen, P36931). All images were acquired using a confocal microscope (LSM710, Carl Zeiss, Germany) and analyzed with Volocity and Fiji software.

Western Blot

To compare protein expression levels, BMDMs were cultured for 24 h at 37 °C with 5% CO₂, with or without polystyrene MPs (50 μg) or LPS (250 ng/mL), and were then harvested. For in vivo experiments, mice were orally administered MPs (50 μg) three times per week. After 1 week of MP administration, LLC cells were intravenously injected to induce lung metastasis (designated as day 0). Oral gavage of MPs was continued throughout the experimental period. In Fer-1 treatment groups, the ferroptosis inhibitor ferrostatin-1 (Fer-1) was administered intraperitoneally at a dose of 10 mg/kg twice per week starting from day 0. Lung tissues were harvested on day 14 for protein extraction. To prepare single-cell suspensions, minced tissues were digested with Liberase TM (250 ng/mL) and DNase I (250 μg/mL) for 30 min at 37 °C on a shaker. The isolated cells were washed, treated with red blood cell lysis buffer (Gibco, A1049201), and filtered through a 70-μm strainer. The protein samples were then electrophoresed and transferred to membranes as described previously. Each membrane was incubated overnight at 4 °C with primary antibodies against TLR2 (Cell Signaling Technology, 13744S), TLR4 (Proteintech, 66350–1-Ig), TIRAP (Cell Signaling Technology, 13077), MyD88 (Cell Signaling Technology, 4283), TRAF6 (Cell Signaling Technology, 67591), P-JNK (Cell Signaling Technology, 9251), P-c-Jun (Cell Signaling Technology, 3270), NRF2 (Cell Signaling Technology, 12721), GCLM (Abcam, ab126704), SLC7A11 (Novus Biologicals, NB300–318), NQO1 (Santa Cruz Biotechnology, sc-32793), GCLC (Cell Signaling Technology, 52183), GPX1 (R&D systems, AF3798), GPX3 (R&D systems, AF4199), GPX4 (R&D systems, MAB5457), FTL (Santa Cruz Biotechnology, sc-390558;), FTH1 (Santa Cruz Biotechnology, sc-376594), and β-actin (Cell Signaling Technology, 8457). Each membrane was washed with Tris-buffered saline with 0.05% Tween-20 and incubated with an appropriate secondary antibody for 1 h. The bands were then visualized using Clarity Max Western enhanced chemiluminescence substrate (Bio-Rad, 1705062), and the images were analyzed using ImageJ software.

Mitochondrial Membrane Potential Assay

BMDMs were cultured for 24 h at 37 °C with 5% CO₂, with or without polystyrene MPs, and then harvested. Samples were stained with JC-1 dye (Abcam, ab113850) at a final concentration of 5 μM for 20 min at 37 °C in an incubator. Samples were measured via flow cytometry analysis using BD FACS LSR II and were analyzed using FlowJo software.

ATP Concentration Measurements

BMDMs were seeded in white 96-well plate for 24 h at 37 °C with 5% CO₂, either with or without polystyrene MPs. ATP production in BMDMs was evaluated using an ATP Assay Kit (Abcam, ab113849), following the manufacturer’s instructions. Luminescence-based ATP levels were measured using a Varioskan LUX multimode microplate reader (BioTek, Winooski, Vermont, USA), and the results were analyzed using Thermo Scientific SkanIt Software (Thermo Fisher Scientific, USA).

Assessment of Lipid Peroxidation

BMDMs were cultured for 24 h at 37 °C with 5% CO₂, with or without polystyrene MPs, and harvested. Samples were then loaded with C11-bodipy 581/591 (Invitrogen, D3861) at a final concentration of 5 μM for 30 min at 37 °C in an incubator. Samples were measured via flow cytometry analysis using BD FACS LSR II and were analyzed using FlowJo software.

RNA Sequencing and Bioinformatics

BMDMs were plated in six-well plates and treated with or without polystyrene MPs (50 μg) at 37 °C with 5% CO₂ for 24 h. Total RNA was isolated with Trizol reagent (Invitrogen, 15596018), and libraries were prepared using the Next Ultra II Directional RNA-Seq Kit (NEW ENGLAND BioLabs, Inc., UK) in each condition. Ribosomal RNA (rRNA) was removed using the RIBO COP rRNA depletion kit (LEXOGEN, Inc., Austria). The resulting rRNA-depleted RNAs underwent complementary DNA synthesis, shearing, and indexing using Illumina indexes 1–12, followed by enrichment through polymerase chain reaction (PCR). Library quality was assessed using the Agilent 2100 bioanalyzer, and quantification was performed using the library quantification kit on a Step One Real-Time PCR System (Life Technologies). High-throughput sequencing was conducted as paired-end 100 sequencing on the NovaSeq 6000 (Illumina, Inc., San Diego, CA, USA). The initial steps involved assessing raw data quality using FastQC (v0.11.9), aligning the reads to the human reference genome (GRCh38 from Gencode) using HISAT2 (v2.2.1), and quantifying gene expression levels using HTSeq (v0.11.1). Differential gene expression analysis utilized DESeq2 (v1.42.1), defining significance as |log2 fold change| ≥ 1 and adjusted P-value ≤ 0.05. Unsupervised K-means clustering generated heatmaps for visualization. GSEA for Biological Processes utilized the clusterProfiler package (v4.10.1) in R, ranking genes by fold change from DESeq2 analysis.

Histology

Lungs were harvested and fixed in 4% paraformaldehyde for 24 h at 4 °C. Tissues were then incubated in 30% sucrose in PBS (pH 7.4) at 4 °C until the samples sank. Frozen tissue blocks were prepared by embedding samples in OCT compound (Leica, 3801480), and 10 μm cryosections were obtained using a cryostat. Sections were stained with hematoxylin (Bioghost, EOYA-10-OT-1L) and eosin (Bioghost, HEMH-OT-1L). Images were acquired using an Olympus DP71 microscope and processed with cellSens software (Olympus).

Mouse Tumor Model

LLC cells stably expressing RFP were obtained from AntiCancer Incorporated. Cell lines were maintained in DMEM medium containing 10% FBS and incubated at 37 °C with 5% CO₂. LLC cells (1 × 10⁶ per mouse) were injected intravenously through the tail vein. Mice were sacrificed 2 weeks after the intravenously injection of LLC cells.

Glutathione Measurement

BMDMs were seeded in a white 96-well plate and cultured for 24 h at 37 °C with 5% CO₂, with or without polystyrene MPs. GSH and GSSG levels were quantified using a luminescence-based GSH/GSSG-Glo assay (Promega, V6611) according to the manufacturer’s instructions. Luminescence-based measurements of the GSH/GSSG ratio and total GSH levels were performed using a Varioskan LUX multimode microplate reader, and the results were analyzed with Thermo Scientific SkanIt Software.

Real-Time Quantitative Polymerase Chain Reaction

To compare the mRNA expression levels, BMDMs were plated in six-well plates and treated with varying concentrations of polystyrene MPs (0, 5, 10, 50, and 100 μg) at 37 °C with 5% CO₂ for 24 h. Thereafter, mRNA was isolated using Trizol reagent according to the manufacturer’s instructions. The RNA reverse transcription and real-time PCR were conducted using an AccuPower Cycle Script RT premix (Bioneer, K-2044) and Power SYBR Green PCR Master Mix (Applied Biosystems, 4367659), according to the manufacturer’s protocols. The mRNA level was quantified VIA the threshold cycle method. Additionally, mRNA was isolated from the lungs of mice that received an intravenously injection of LLC-RFP, with or without oral gavage of polystyrene MPs, or from C57BL/6 mice, using the same method described above. Primer sequences were used to amplify mouse Il-6, Tnf-α, Il-1β, Il-10, Ccl2, Gpx1, Gpx3, Hif1α, Vegfa, Tgf-β1 and glyceraldehyde-3-phosphate dehydrogenase (Gapdh). The following primer sequences were used: Il-6 forward primer, 5′-GTCCTTC­CTACCCC­AATTTCCA-3′ and Il-6 reverse primer, 5′-TAACGCA­CTAGG­TTTGCCGA-3′. Tnf-α forward primer, 5′-TCTACTG­AACTTCG­GGGTGA-3′ and Tnf-α reverse primer, 5′-CACTTGG­TGGTTT­GCTACGA-3′. Il-1β forward primer, 5′-GAAGAA­GAGC­CCATCC­TCTGT-3′ and Il-1β reverse primer, 5′-TTGTCGTTG­CTTGGT­TCTTCC-3′. Il-10 forward primer, 5′-ACAGC­CGGGAAG­ACAATAAC-3′ and Il-10 reverse primer, 5′-GGCAACCCA­AGTAAC­CCTTA-3′. Ccl2 forward primer, 5′-TGGAAGG­AGTGTG­CATGTTC-3′ and Ccl2 reverse primer, 5′-CAAGAC­ACGAAAA­GGCATGA-3′. Gpx1 forward primer, 5′-CCACCGTG­TATGCCT­TCTCC-3′ and Gpx1 reverse primer, 5′-AGAGAGA­CGCGAC­ATTCTCAAT-3′. Gpx3 forward primer, 5′-GCCAGC­TACTGAGG­TCTGACAGA-3′ and Gpx3 reverse primer, 5′-CAAATGG­CCCAAG­TTCTTCTTG-3′. Hif1α forward primer, 5′-CTCAAAGTCGGACAGCCTCA-3′ and Hif1α reverse primer, 5′-CCCTGCA­GTAGGTT­TCTGCT-3′. Vegfa forward primer, 5′-CGAAGTGG­TGAAGTT­CATGGATG-3′ and Vegfa reverse primer, 5′-TTCTGTT­CAGTCTTT­CCTGGTGAG-3′. Tgf-β1 forward primer, 5′-TTGCTTCA­GCTCC­ACAGAGA-3′ and Tgf-β1 reverse primer, 5′-TGGTT­GTAGAG­GGCAAGGAC-3′. Gapdh forward primer, 5′-CCAATG­TGTCCGTC­GTGGATCT-3′ and Gapdh reverse primer, 5′-GTTGAAGT­CGCAGGAG­ACAACC-3′. The relative gene expression levels of each gene were normalized to Gapdh levels.

Flow Cytometry

BMDMs were plated in six-well plates and treated with or without polystyrene MPs at 37 °C with 5% CO₂ for 24 h. Thereafter, the samples were harvested and washed in FACS buffer (PBS containing 2% FBS and 2 mM ethylenediaminetetraacetic acid). To prevent nonspecific binding, cell suspensions were preblocked with Fc Block (BioLegend, 101302). Samples were stained with allophycocyanin (APC)-CD11b (BioLegend, 101212), AlexaFluor700-F4/80 (BioLegend, 123130), FITC-CD206 (BioLegend, 141703), and BV421-CD86 (BioLegend, 105031) to assess M1 and M2 polarization in response to polystyrene MPs. To assess lysosomal damage, cells were stained with APC/Cy7-CD45 (BioLegend, 103116), BV421-CD11b (BioLegend, 101251), FITC-F4/80 (BioLegend, 123107) or FITC-Ly6G (BioLegend, 127606), and Lyso-tracker Red (200 nM; Thermo Fisher, L7528) in PBS for 15 min at 37 °C. To assess the immune microenvironment, mice were orally administered MPs (50 μg) three times per week. After 1 week of MP administration, LLC cells were intravenously injected to induce lung metastasis (designated as day 0). Oral gavage of MPs was continued throughout the experimental period. Lung tissues were harvested on days 0, 3, 7, and 14. Single-cell suspensions were then prepared by digesting minced tissues with Liberase TM (250 ng/mL) and DNase I (250 μg/mL) for 30 min at 37 °C on a shaker. The isolated cells were washed, treated with red blood cell lysis buffer, and filtered through a 70-μm strainer. To prevent nonspecific binding, cell suspensions were preblocked with Fc Block (BioLegend, 101302). Samples were subsequently stained with APC/Cy7-CD45 (BioLegend, 103116), APC-CD11b (BioLegend, 101212), AlexaFluor700-F4/80 (BioLegend, 123130), FITC-CD206 (BioLegend, 141703), BV421-CD86 (BioLegend, 105031), AlexaFluor700-CD45 (BioLegend,147716), APC/Cy7-CD3 (BioLegend, 100222), BV421-CD4 (BioLegend, 100437), BV510-Nkp46 (BioLegend, 137623), BV605-CD8 (BioLegend, 100744), FITC-Ki-67 (BioLegend, 652409), APC-IFN-r (BioLegend, 113605), FITC-Granzyme B (BioLegend, 372205), FITC-Annexin V (BioLegend, 640906) and DCFDA (20 μM). All samples were measured via flow cytometry analysis using BD FACS LSR II and were analyzed using FlowJo software.

Two-Photon Intravital Imaging

The presence of polystyrene MPs in the lungs, as well as their relationships with macrophages, was investigated. Intravital imaging of the lungs tissues was performed according to previously described methods. We orally administered 50 μg polystyrene MPs (red) three times a week to CX3CR1-GFP mice and imaged their lungs. The sequential images were captured at intervals of 1 min.

3D Whole Organ Imaging

The C57BL/6 mice were divided into two groups: one group was orally administered 50 μg polystyrene MPs (red) three times a week, and the other group served as control. C57BL/6 mice were also prepared by injecting LLC-RFP cells, either with or without oral administration of polystyrene MPs. These mice were anesthetized and fixed by perfusing with PBS and 4% formaldehyde, and the lungs were surgically excised. The excised tissues were fixed in 4% paraformaldehyde at 4 °C overnight. Thereafter, the tissues were incubated with 35% sucrose at 4 °C until they sank. To perform tissue clearing and imaging, we used a clearing kit (Binaree, HRTC-001) and selective plane illumination microscopy (Lightsheet Z.1, Carl Zeiss, Germany), following a previously described method. The 3D whole organ imaging video was made using Imaris software (Bitplane, Oxford Instruments, Zurich, Switzerland) and ImageJ software.

Statistical Analysis

The statistical significance of the differences between two groups was determined using a two-tailed Student’s t test, as previously described. Statistical comparisons were performed using one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test, as indicated in the figure legends for each experiment. The threshold for statistical significance was set at p < 0.05. The data are expressed as the mean ± standard error of the mean (SEM) unless otherwise specified. All experiments were conducted at least in triplicate. Prism software (version 9.0.0, GraphPad Software, Inc., La Jolla, CA, USA) was used to analyze the data from the in vitro and in vivo experiments.

The transcriptomic data of total RNA-seq are under deposition in the Gene Expression Omnibus (GEO) under the accession number: GSE299749.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsnano.5c15425.

• Cellular responses of BMDMs to MPs (Figure S1); Time-dependent alterations in lung immune cell composition associated with MP exposure during early tumor progression (Figure S2); MP exposure modulates functional phenotypes of lung immune cells during tumor progression (Figure S3) (PDF)

• (Video S1) MPs induce migration of mouse BMDMs (MOV)

• (Video S2) Mouse BMDMs capture MPs (MOV)

• (Video S3) Mouse BMDMs bind and drag MPs via actin polymerization (MOV)

• (Video S4) MPs cause cell death of mouse BMDM (MOV)

• (Video S5) 3D organ imaging of lungs (MOV)

• (Video S6) Two-photon intravital imaging of MPs-administered mice lungs (MOV)

• (Video S7) 3D organ imaging of lungs in LLC bearing mice (MOV)

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B.K. and K.-M.P. contributed equally.

Conceptualization: Y.-M.H. Methodology: B.K., K.-M.P., H.L. Investigation: B.K., K.-M.P., H.L, Y.-M.H. Visualization: B.K., K.-M.P. Funding acquisition: B.K., Y.-M.H. Project administration: B.K., Y.-M.H. Supervision: Y.-M.H. Writing – original draft: B.K., K.-M.P., Y.-M.H. Writing – review and editing: B.K., K.-M.P., Y.-M.H.

The authors declare no competing financial interest.