Introduction

Plastic products are now indispensable in global manufacturing and consumer sectors due to their cost-effective and versatile applications. However, their exponentially growing global production (estimated at 400 million metric tons annually), combined with extremely slow biodegradation kinetics, has precipitated an environmental crisis1. Plastic waste is now infiltrating terrestrial and aquatic ecosystems at alarming rates2. Current projections indicate that global plastic production and associated waste streams will continue to rise until large-scale implementation of biodegradable alternatives is achieved3,4. Conventional plastics degrade environmentally through the synergistic effects of UV-induced photodegradation and mechanical weathering, eventually breaking into micro- (1–5,000 μm) and nano-sized (< 1 μm) particles5,6. Emerging toxicological evidence reveals that microplastics (MPs) can breach biological barriers and accumulate in vital organs (particularly in the liver) in various species, including mammals and humans7,8. This bioaccumulation is worsened by trophic transfer within food webs, where MPs adsorb environmental contaminants and pathogens, amplifying their ecotoxicological risks through biomagnification processes9,10.

Accumulating evidence demonstrates that MPs bioaccumulation in multiple organ systems induces oxidative stress (OS) and cytotoxic effects through reactive oxygen species (ROS)-mediated pathways11,12. Specifically, exposure to polystyrene MPs (PS-MPs) in mice significantly depleted hepatic glutathione (GSH) reserves, elevated malondialdehyde13 levels, and suppressed activities of antioxidant enzymes (SOD, CAT)14. Another study found that PS-MPs increased hepatotoxicity and lipotoxicity in human liver, indicating potential risks for related diseases15. The liver, functioning as the primary metabolic hub for xenobiotic clearance, exhibits particular vulnerability to MPs-induced toxicity16,17. Regarding hepatic pathology, MPs exposure has been shown to disrupt lipid homeostasis in murine models, leading to hepatic steatosis through PPARγ-mediated dysregulation of fatty acid β-oxidation18,19.

Emerging evidence indicates that systemic translocation of PS-MPs into hepatocytes via circulatory distribution initiates OS-mediated cytotoxicity. This occurs primarily through mitochondrial ROS overproduction and endoplasmic reticulum stress-induced GSH depletion20. As a redox-regulated programmed cell death mechanism, apoptosis is critically modulated by OS signaling. During PS-MPs exposure, ROS overproduction triggers the opening of the mitochondrial permeability transition pore (MPTP), activating the intrinsic apoptotic pathway. This is characterized by an elevated Bax/Bcl-2 ratio and subsequent activation of the caspase-9/caspase-3 cascade21,22. Intriguingly, the tumor suppressor p53 exhibits dual redox regulatory roles under OS: it suppresses Cyclooxygenase-2 (Cox-2)-mediated pro-inflammatory ROS generation while paradoxically enhancing antioxidant enzyme (e.g., SOD, CAT) transcription through NRF2 pathway activation23. OS also serves as the key molecular link connecting MPs to ferroptosis. Ferroptosis, an iron-dependent oxidative cell death process distinct from apoptosis, manifests through GSH/glutathione peroxidase 4 (GPX4) axis collapse and iron overload-driven Fenton reactions, which amplify lipid peroxidation24,25. This oxidative cascade has been implicated in diverse pathologies, ranging from neurodegenerative disorders to chemoresistant cancers. Emerging evidence suggests that ferroptosis activation (via solute carrier family 7 member 11 (SLC7A11) inhibition) may counteract tumorigenesis by eliminating malignant clones26,27. Accumulating in vivo evidence confirms MP-induced ferroptosis. However, research on whether MPs can trigger ferroptosis in vitro remains limited. The toxicological research highlights OS as a predominant focus for investigating MPs-driven apoptotic and ferroptotic pathways, with many studies prioritizing redox imbalance as a key mechanism underlying pollutant-associated cellular outcomes.

The cytotoxicity of MPs exhibits size-dependent internalization kinetics, with intracellular accumulation rates inversely proportional to particle diameter. This is due to the enhanced endocytic uptake efficiency of NPs compared to larger MP particles28,29. Numerous studies have demonstrated that NPs can induce genotoxicity and cytotoxicity through multiple pathways30,31. However, existing research has primarily focused on MPs of ≤ 20 μm to model biodistribution patterns32,33. However, recent studies reveal that large-sized MPs (40–500 μm) exist in a significant proportion (42.6 ± 15.9% to 54.6 ± 15.5%) of human fecal and placental samples34. Given the high environmental persistence of large-sized MPs and their bioaccumulation in animal organ systems, a systematic evaluation of their hepatotoxicity becomes critical. Moreover, research comparing the cytotoxicity of pristine versus surface-modified large-sized MPs remains scarce. To address this gap, the present study established a cell model to explore the hepatotoxicity of PS-MPs and amino-modified polystyrene MPs, OS-mediated cytotoxicity of pristine and PS-NH2-MPs (50 μm) under laboratory-relevant exposure conditions (0–0.5 mg/mL, 48 h). Since ROS are closely linked to ferroptosis development, this study preliminarily examines changes in intracellular ROS levels and their impact on ferroptosis following exposure to large-sized MPs. Current regulatory frameworks face significant challenges35, and this study provides a critical theoretical foundation for addressing them.

Materials and methods

Characterization of MPs

PS-MPs were selected based on their environmental prevalence in daily life, mature commercial production protocols, and well-established surface modification techniques. PS-MPs (50 μm) and PS-NH2-MPs (50 μm) were purchased from Suzhou Zhiyi Microsphere Technology Co., Ltd. (KBsphere, China). Morphology of the particles was confirmed by a scanning electron microscope (SEM, Hitachi, S-4800, Japan). The surface chemistry of the particles was characterized by Fourier-transform infrared spectroscopy (FTIR, IRXROSS, Japan). Experimental concentration of MPs referred to some exposure concentrations reported on MPs toxicology22,36,37. MPs (25 mg) and F12 medium (1 mL) were mixed in an Eppendorf Tubes® 3810X (Eppendorf, Germany) to form a 25 mg/ml suspension. The suspension was then diluted with F12 medium to 0.05, 0.1, 0.2, and 0.5 mg/mL, respectively. The selected concentrations cover conventional in vitro testing brackets (0.05–0.2 mg/mL) for direct literature alignment, with extension to 0.5 mg/mL accounting for particle-size-dependent response modulation–essential for resolving delayed toxicity phenotypes in large-sized MPs systems.

Cell culture and treatment

AML12 (alpha mouse liver 12) cell line was obtained from HyCyte (China), and cultured according to the study of Cao et al.36. After digestion, AML12 cells were inoculated into 96-well or 6-well plates (NEST, China) at 5 × 103 cells or 4.5 × 106 cells per well, respectively, and incubated at 37 ℃ with 5% CO2. After 24 h, AML12 cells were then cultured with PS-MPs and PS-NH2-MPs (0, 0.05, 0.1, 0.2, 0.5 mg/mL) for following experiments.

Cell viability assay

Cell viability was assessed using the Cell Counting Kit-8 (CCK-8) after 24 and 48 h exposure to PS-MPs or PS-NH2-MPs (0–0.5 mg/mL), following the protocol of Wu et al.38. Briefly, treated cells in 96-well plates were incubated with 10 µL CCK-8 reagent per well at 37 °C for 2 h. Absorbance was then measured at 450 nm using a microplate reader. Viability was calculated as relative to untreated control cells.

Oxidative stress measurement

After 48 h exposure to PS-MPs or PS-NH2-MPs (0–0.5 mg/mL), AML12 cells were harvested and lysed using an ultrasonic disruptor. The supernatant was collected by centrifugation (1,000 × g, 15 min, 4 °C) for OS marker analysis. Total protein content was quantified with a BCA assay kit. Levels of GSH, MDA, CAT, and SOD were detected with the assay kit (Nanjing Jian cheng, Bioengineering Institute, China) following manufacturer protocols and established methods36,38. And the levels of ferrous iron (Fe2+) were detected by microplate reader according the guide of Ferrous Ion Content Assay Kit (Solarbio, China).

Intracellular ROS levels were measured using the fluorescent probe DCFH-DA (Biosharp, China). After removing the culture medium, cells were washed twice with PBS and incubated with 10 µM DCFH-DA in serum-free medium at 37 °C for 30 min (protected from light). Fluorescence was visualized under a microscope (excitation/emission: 488/525 nm) and quantified using ImageJ software. Three random fields per well were analyzed, with fluorescence intensity normalized to untreated controls. The assay was performed according to the protocol manufacturer (Biosharp, China).

Flow cytometry

Apoptosis and intracellular ROS were quantified by flow cytometry (CytoFLEX, Beckman, USA) after exposure to PS-MPs (0.5 mg/mL) or PS-NH2-MPs (0–0.5 mg/mL) for 48 h. For apoptosis analysis, cells stained with Annexin V-FITC/PI (Beyotime, China) were excited at 488 nm, with emissions detected through 525/40 nm (FITC) and 610/20 nm (PI) filters following established methods38. For ROS measurement, cells incubated with 10 µM DCFH-DA in serum-free medium (37 °C, 30 min, dark) were analyzed at 488 nm excitation/525 nm emission. Fluorescence intensity was expressed as the geometric mean of DCF signal (ROS) or quadrant gating (apoptosis), normalized to untreated controls. Data were processed using CytExpert software with three independent replicates.

RT-qPCR

Total RNA was extracted from AML12 cells using RNAiso Plus reagent (Takara, Japan), with RNA purity verified spectrophotometrically (A260/A280 = 1.8–2.0). cDNA was synthesized using the PrimeScriptTM FAST RT reagent Kit with cDNA Eraser (Takara, Japan). RT-qPCR was performed in triplicate using TB Green™ Premix Ex Taq™ II (Takara) on a 7500 RT-qPCR system. Each 20 µL reaction contained: 10 µL premix, 0.8 µL forward/reverse primers (10 µM; sequences in Table 1), 2 µL cDNA, and 6.4 µL nuclease-free water. Cycling conditions: 40 cycles of 95 °C for 5 s and 60 °C for 30 s; melt curve analysis (65–95 °C, 0.5 °C increments). Data were analyzed via the 2−ΔΔCt method with ACTB as reference gene, ensuring primer specificity and amplification efficiency.

Table 1 Sequences of primers for quantitative real-time PCR.
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Western blot analysis

Proteins were extracted using RIPA buffer with protease inhibitors. but uniform concentration was 20 µg. After centrifugation (15,000 × g, 15 min, 4 °C), supernatants were quantified by BCA assay. Samples (20 µg/lane) were separated on 10% SDS-PAGE gels and transferred to PVDF membranes via semi-dry transfer. The membranes were blocked with Western Quick Block Kit (GenScript, America) at 25℃ for 10 min, and then incubated with the primary antibodies (according to the recommended does in the instructions of the manufacturer) overnight (Table 2). The secondary antibodies Goat anti mouse IgG, Goat anti ribbit IgG (1:1,000, Invitrogen, China) were kept with membranes at 25 ℃ for 3 h. β-actin is an internal control. The protein bands were visualised using a DAB Substrate kit (Solarbio, China) following the guidelines of the manufacturer and quantified with ImageJ software. Data were normalized to β-actin and analyzed using Prism 9.0.

Table 2 Antibodies for Western blot assay.
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Statistical analysis

Statistical analysis were performed by using Prism 9.0 software (America) and all data were assessed by using analysis of variance (ANOVA) between the different groups. The data were derived from at least three separate experiments, and the results were presented as mean ± standard deviations. A P value of less than 0.05 was accepted statistically significant.

Additional methodological details and instrument specifications are provided in the Supplementary Material.

Results

Imaging characterization of PS-MPs and PS-NH2-MPs

Characterization of PS-MPs has been shown in Fig. 1. SEM found that almost all of the PS-MPs and PS-NH2-MPs were uniform spherical (Fig. 1A), and the average particle size of the PS-MPs and PS-NH2-MPs was 50 μm, consistent with the size specifications supplied by the manufacturer. The successful amine functionalization of PS-NH2-MPs (Fig. S1) ensured valid comparison with pristine PS-MPs. Also, the coexistence of cells and MPs was shown in Fig. 1B, and the 50 μm MPs were larger than the AML12 cells (Fig. 1B).

Fig. 1
Fig. 1
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Characterization of PS-MPs. (A) SEM image of PS-MPs. (B) Microscope image of PS-MPs with cells. Scale bar: 50 μm.

Cell viability

Firstly, cell viability assay was conducted to study the cytotoxicity induced by PS-MPs and PS-NH2-MPs (50 μm). As shown in Fig. 2, there were no significant effects on the viability of AML12 cells exposed to PS-MPs and PS-NH2-MPs at concentrations 0–0.2 mg/mL for 48 h (P > 0.05). Similarly, AML12 cells also showed no significant reduction on viability after 24 h exposure. Compared to the control group, significant cytotoxicity was observed at the highest tested concentration (0.5 mg/mL) of both PS-MPs and PS-NH2-MPs after 48 h exposure (P < 0.05). Notably, surface functionalization with amino groups (PS-NH2-MPs) resulted in no significant difference (P > 0.05) in inhibitory activity compared to pristine PS-MPs.

Fig. 2
Fig. 2
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Cell viability after exposure to PS-MPs. Statistical trend of cell viability after exposure with PS-MPs at 0, 0.05, 0.1, 0.2 and 0.5 mg/mL for (A) 24 h and (B) 48 h. n = 9. Compared to control: 0 mg/mL; Statistics are calculated as mean ± SD; ANOVA followed by Tukey test was used to assess differences groups; *P < 0.05, **P < 0.01.

Oxidative stress

As shown in Figs. 3 and 4, OS biomarkers were detected in MPs-exposed cells. Notably, MDA levels were significantly reduced following exposure with 0.1 mg/mL PS-MPs (P < 0.05; Fig. 3B). GSH, CAT and SOD activities showed no significant alterations across MPs-treated groups compared to control (P > 0.05; Fig. 3A, D). At the highest exposure concentration (0.5 mg/mL), both PS-MPs and PS-NH2-MPs significantly increased intracellular ROS levels compared to controls, as demonstrated by fluorescence imaging quantification (P < 0.05; Fig. 4A) and flow cytometric analysis (P < 0.01; Fig. 4B). Critically, PS-NH2-MPs did not alter cellular antioxidant capacity compared to PS-MPs (P > 0.05).

Fig. 3
Fig. 3
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Effects of AML12 cells exposure to PS-MPs on oxidative stress. (A-D) The condition of cellular oxidative stress used GSH, MDA, SOD and CAT assay kits to detect after exposure with PS-MPs at 0, 0.05, 0.1, 0.2 and 0.5 mg/mL for 48 h. n = 3. Compared to control; Statistics are calculated as mean ± SD; ANOVA followed by Tukey test was used to assess differences groups; *P < 0.05, **P < 0.01.

Fig. 4
Fig. 4
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ROS levels of AML12 cells exposure to PS-MPs. (A) DCFH-DA staining was followed by fluorescence microscopy to detect the levels of ROS. The green fluorescent dots indicate ROS generation. (B) Detected ROS levels by flow cytometry after treatment. Scale bar: 50 μm. n = 3. Compared to control; Con: control group, 0 mg/mL PS-MPs; PS: 0.5 mg/mL PS-MPs; PS-NH2: 0.5 mg/mL PS-NH2-MP; Statistics are calculated as mean ± SD; *P < 0.05, **P < 0.01.

Apoptosis analysis

Flow cytometry-based quantification of apoptosis is presented in Fig. 5. The analysis revealed a dose-dependent increase in percentage of apoptotic cells following PS-MPs exposure, with statistical significance observed exclusively at the highest concentration (0.5 mg/mL; P < 0.05). Compared to pristine PS-MPs, surface functionalization with amino groups (PS-NH2-MPs) did not exacerbate apoptotic responses (P > 0.05).

Fig. 5
Fig. 5
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Percentage of apoptotic cells analysis in AML12 cells. (A) Percentage of apoptotic cells was measured by flow cytometry after exposure to PS-MPs at 0, 0.05, 0.1, 0.2 and 0.5 mg/mL for 48 h. (B) Percentage of apoptotic cells was measured by flow cytometry after exposure to PS-MPs and PS-NH2-MPs at 0.5 mg/mL. n = 3. Compared to control; Con: control group; 0 mg/mL PS-MPs; PS: 0.5 mg/mL PS-MPs; PS-NH2: 0.5 mg/mL PS-NH2-MPs; Statistics are calculated as mean ± SD; *P < 0.05, **P < 0.01.

Apoptosis analysis in AML12 cells is shown in Fig. 6. Western blot analysis of apoptosis-related proteins in AML12 cells revealed no significant changes in p53, Bcl-2 or Bax expression levels between MPs-treated groups and untreated controls (P > 0.05; Fig. 6A, B). In contrast, transcriptional profiling demonstrated distinct apoptotic responses: Caspase-9 mRNA expression was significantly upregulated in PS-MPs-exposed cells (P < 0.05; Fig. 6C), while Bax mRNA levels were significantly higher in PS-MPs-exposed cells than in those exposed to PS-NH2-MPs (P < 0.05).

Fig. 6
Fig. 6
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Apoptosis analysis in AML 12 cells. (A-B) Make Western blot analysis of P53, Bcl-2 and Bax after exposure to PS-MPs for 48 h. The blots have been cropped. Original blots are presented in Supplementary Fig. S2. (C) RT-qPCR analysis of P53, Bcl-2, Bax, CytC, Caspase-9, Caspase-3 and Cox-2 mRNA after exposure to PS-MPs for 48 h. n = 3. Compared to control; Con: control group, 0 mg/mL PS-MPs; PS: 0.5 mg/mL PS-MPs; PS-NH2: 0.5 mg/mL PS-NH2-MPs; Statistics are calculated as mean ± SD; *P < 0.05, **P < 0.01.

Ferroptosis analysis

Ferroptosis analysis in AML12 cells is shown in Fig. 7. The intracellular Fe2+ content exhibited a non-significant upward trend with increasing MPs concentrations (P > 0.05; Fig. 7A). Western blot analysis revealed no statistically significant changes in glutathione synthetase (GSS), GPX4 and SLC7A11 protein levels in AML12 cells exposed to 0.5 mg/mL MPs (P > 0.05; Fig. 7C, D). Similarly, RT-qPCR analysis demonstrated no statistically significant differences in the mRNA expression levels of GPX4, SLC7A11, ACSL4 and FTH1 in MP-treated groups compared to controls (P > 0.05; Fig. 7B).

Fig. 7
Fig. 7
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Ferroptosis analysis in AML 12 cells. (A) Ferrous ion detection after exposure to PS-MPs at 0, 0.05, 0.1, 0.2 and 0.5 mg/mL for 48 h. (B) RT-qPCR analysis of GPX4, SLC7A11, ACSL4 and FTH1 mRNA after exposure to PS-MPs for 48 h. (C-D) Western blot analysis of GSS, GPX4 and SLC7A11. The blots have been cropped. Original blots are presented in Supplementary Fig. S2. n = 3. Compared to control; Con: control group, 0 mg/mL PS-MPs; PS: 0.5 mg/mL PS-MPs; PS-NH2: 0.5 mg/mL PS-NH2-MPs; Statistics are calculated as mean ± SD; *P < 0.05, **P < 0.01.

Discussion

Plastics are widely utilized in multiple industries, and their production has seen continuously1. While extensive toxicological research has prioritized MPs (≤ 20 μm) and NPs due to their enhanced cellular internalization capacity32,39. Emerging evidence reveals limitations of small particles: (1) 2 μm PS-MPs only induced ROS elevation at high concentrations (≥ 0.1 mg/mL) without viability loss in AML12 cells22; (2) internalizable small particles (1–10 μm) exhibit higher cytotoxicity than larger particles (50–100 μm) at same concentrations40. This paradox highlights the necessity to evaluate non-internalizable large-sized MPs (20–700 μm), which constitute a substantial proportion of plastic pollutants in ecosystems and human biological samples34,41. The key finding of this study reveals that exposure to large-sized PS-MPs (50 μm) induces ROS generation in AML12 cells without triggering ferroptosis, challenging the NPs-centric toxicity models and suggesting risk assessments focused on extracellular particle-cell interactions rather than relying solely on internalization-driven toxicity models.

The evidence highlights PS-MPs as potent hepatotoxic agents. Recent studies have documented their systemic accumulation and capacity to induce OS through hepatic redox imbalance11,42,43. While NPs (< 1 μm) are readily internalized by cells and disseminated in the circulation to drive inflammation and apoptosis via intracellular ROS cascades44,45, our findings reveal a different toxicological profile for large-sized MPs (50 μm). In alignment with size-dependent uptake limitations46, images confirmed minimal phagocytic internalization of 50 μm PS-MPs by AML12 hepatocytes (Fig. 1B), consistent with prior reports demonstrating preferential cellular exclusion of particles32,33,47. Although the concentrations of PS-MPs and PS-NH2-MPs are significantly higher than environmental levels, dose-response analysis demonstrated that hepatocyte viability was reduced only at 0.5 mg/mL (P < 0.05; Fig. 2B), accompanied by ROS hypergeneration (Fig. 4) without ferroptotic reaction (Fig. 7).

While smaller MPs are well-documented to deplete intracellular antioxidants like GSH and SOD via lysosomal disruption and mitochondrial ROS overproduction48,49, our findings reveal a distinct antioxidant adaptation response to large-sized MPs (50 μm). Exposure to 0.5 mg/mL PS-MPs/PS-NH2-MPs induced significant ROS elevation (P < 0.05; Fig. 4B), paradoxically reduced MDA level (P > 0.05; Fig. 3). The observed MDA reduction at doses (0.1 mg/mL) aligns with reports that larger MPs (> 20 μm) preferentially induce membrane-associated ROS activation rather than intracellular lipid peroxidation11, contrasting sharply with NPs-driven MDA surges38. These size-dependent mechanistic disparities corroborate findings reported by Meng et al., where antioxidant enzyme responses varied significantly across MPs size fractions50. Previous studies have consistently shown that smaller MPs generally induce stronger toxicity than larger particles across cell types, with key manifestations including elevated ROS production and compromised antioxidant defenses, which collectively disrupt cellular homeostasis and may activate ferroptosis pathways51,52. However, our current research reveals a unique antioxidant adaptation response to large-sized MPs (50 μm). While our experimental concentrations (0.5 mg/mL) exceed environmentally relevant MP levels documented in mammalian livers53, the observed OS and adaptive antioxidant responses provide valuable reference angles for extrapolating chronic low-dose risks under real-world exposure scenarios benchmarks for extrapolating chronic low-dose risks under real-world exposure scenarios. These findings collectively redefine MPs toxicity paradigms, emphasizing that particles-cell surface interaction modalities, not merely intracellular invasion, induce hepatotoxic effects.

Despite unchanged protein expression of p53, Bcl-2 and Bax (Fig. 6), the significant upregulation of Caspase-9 mRNA (P < 0.05; Fig. 6C) suggests mitochondrial pathway priming without full apoptotic commitment. This parallels studies showing that ROS activates the ASK1-JNK pathway to promote Caspase-9 transcription, while p53 has dual roles in ROS scavenging (via SOD/CAT induction) and apoptotic regulation54,55.

Importantly, cell type significantly influences MP toxicity outcomes. Macrophages are susceptible to ferroptosis partly because they have low levels of the antioxidant GSH. In contrast, AML12 cells possess natural antioxidant defenses. For example, their catalase activity is 3.2 times higher than that in macrophage-like RAW264.7 cells22. This inherent capacity likely explains their resistance to MP-induced lipid peroxidation, even when ROS levels rise. These distinct vulnerability profiles across cell types necessitate customized risk assessment frameworks.

Our findings reveal limited ferroptotic engagement in AML12 cells after acute exposure to 50 μm MPs. Although mRNA levels of ferroptosis-related genes (GPX4, ACSL4, SLC7A11) showed slight increases, these changes were not statistically significant (P > 0.05; Fig. 7). FTH1—a critical regulator of iron storage56—did not change significantly (P > 0.05), and MDA levels significantly decreased (P < 0.05). Collectively indicate no activation of the classical ferroptosis pathway under large-sized MP exposure. These results align with the demonstration by Wu et al. that MPs (10 μm) fail to induce apoptosis or ferroptosis below 0.2 mg/mL38. The discordancy between ROS elevation and ferroptosis resistance may reflect compensatory antioxidant adaptations, as shown by no significant changes in GSH levels and SOD/CAT activity trends. Such redox buffering capacity likely mitigates iron-dependent Fenton reactions, preventing lipid peroxidation57. Critically, the size-induced exclusion of intracellular MPs internalization precludes lysosomal iron release—a key ferroptosis trigger in NPs-exposed cells58—thereby confining OS to membrane-proximal ROS generation. This is evidenced by persistent null responses across ferroptosis markers despite significant ROS increase (Fig. 7), demonstrating that 50 μm MPs cannot activate intracellular ferroptosis pathways. Such decoupling of ROS production from ferroptotic execution highlights particle size as the dominant determinant of death mechanism selection, contrasting sharply with dose-dependent ferroptosis in NPs systems38.

The observed biological inertia may be fundamentally attributed to size-dependent internalization barriers. Liu et al. demonstrated that particle endocytosis requires thermodynamic driving forces for membrane wrapping and receptor diffusion to binding sites. For particles exceeding optimal dimensions, the kinetic bottleneck of recruiting sufficient membrane receptors severely limits internalization efficiency. They showed that 50 nm PS particles exhibit greater cellular uptake than 500 nm counterparts under identical exposure conditions29. At our experimental scale (50 μm), Brownian motion becomes negligible and membrane adhesion insufficient for internalization, physically precluding direct intracellular interactions. This stands in stark contrast to NPs (≤ 100 nm), which readily penetrate cellular membranes and consequently induce heightened cytotoxicity31.

Surface modification induces differential toxicological profiles. However, this physicochemical alteration failed to translate into significant OS potentiation, with ROS levels in PS-NH2-MP-exposed cells showing no significant change compared to pristine PS-MPs (P > 0.05). This contrasts with reports that surface amino-modification exacerbates NP toxicity31. For instance, 80 nm PS-NH2 at 200 µg/mL significantly elevated cytotoxicity compared to pristine 80 nm PS (P < 0.05), highlighting size-dependent attenuation of surface chemistry effects52.

Although no significant ferroptosis occurred at laboratory-relevant concentrations (≤ 0.5 mg/mL), ROS accumulation and apoptotic priming suggest potential subclinical hepatotoxicity during chronic exposure. Ecotoxicological studies corroborate this concern, demonstrating a positive correlation between MPs abundance and clinical markers of liver dysfunction in animal hepatic tissues13. Therefore, evaluating the chronic effects of MPs at environmentally realistic exposure levels remains an essential direction for future research. Furthermore, this study did not address MPs interactions with co-pollutants (e.g., heavy metals), which may synergistically amplify OS via redox cycling59,60.

However, technical constraints inherent to large-particle exposures limited assessment of some cytotoxicity indicators. Our findings demonstrate that 50 μm PS-MPs induce membrane-localized oxidative stress without activating canonical ferroptosis pathways, likely due to size-imposed exclusion of intracellular uptake. Future studies should prioritize development of advanced exposure models to resolve size-dependent particle-cell interactions. While integrating single-cell transcriptomics and metabolomics to elucidate heterogeneous cellular responses and identify therapeutic targets for MP-induced hepatic dysfunction.

Conclusion

This study reveals that large-sized PS-MPs (50 μm) induce OS in cells by generating ROS. Surface modification MPs (PS-NH2-MPs) exerted minimal effects on oxidative outcomes, confirming particle size is the dominant determinant of toxicity. Although 0.5 mg/mL MPs (a concentration exceeding environmentally relevant levels) did not induce overt ferroptosis, the observed apoptotic priming and adaptive antioxidant responses indicate potential risks of cumulative hepatotoxicity. These findings highlight the importance of large-sized MP risk assessment, providing critical data for policymakers and environmental health research.