Abstract

   Microplastics (MPs) are widespread environmental pollutants that can
   enter the human body through the food chain, potentially leading to
   lung damage. However, the underlying mechanisms responsible for this
   damage remain unclear. Ducks, a commonly consumed poultry species in
   China, are particularly susceptible to MPs exposure due to their
   farming environment. In this study, Shaoxing ducklings were
   administered two distinct concentrations of polystyrene microplastics
   (PS-MPs) (1 mg/L and 100 mg/L) via oral route, alongside a control
   group, over a period of four weeks to establish an in vivo model for
   evaluating the effects of microplastic exposure in ducks.
   Simultaneously, rat type II alveolar epithelial (RLE-6TN) cells were
   exposed to different concentrations of PS-MPs (0, 10, 100, and
   500 µg/mL) for 48 h, thereby constructing an in vitro exposure model.
   Our results showed that PS-MPs caused pathological damage, inflammatory
   cell infiltration, and activation of the LPS/TLR4 inflammatory pathway
   in the lung. Further analysis revealed that PS-MPs disrupted the
   tricarboxylic acid (TCA) cycle and inhibited oxidative phosphorylation.
   Mechanistic investigation demonstrated that PS-MPs induced
   mitochondrial dysfunction and consequent excessive mitophagy. This
   study investigates the mechanisms by which PS-MPs contribute to
   mitochondrial dysfunction and mitophagy, potentially exacerbating lung
   inflammation, offering valuable insights for mitigating the toxic
   effects of PS-MPs on human and animal health.

Graphical Abstract

   [38]graphic file with name 12951_2025_3503_Figa_HTML.jpg

Supplementary Information

   The online version contains supplementary material available at
   10.1186/s12951-025-03503-x.

   Keywords: Duck, Polystyrene microplastics, Mitochondrion, TCA cycle,
   Mitophagy

Introduction

   The use of plastic products significantly improves the quality of human
   life. However, the convenience of “white garbage” has led to widespread
   plastic waste, raising global concerns about serious environmental
   issues [[39]1, [40]2]. Microplastics (MPs), defined as particles with
   diameters smaller than 5 mm, are a common component of plastic
   materials [[41]3]. MPs are particularly concerned about their extensive
   distribution, strong adsorption properties, and resistance to
   degradation, making them a growing threat to both health and
   environment [[42]4, [43]5]. Ingestion of MPs has been increasingly
   reported across various animal species, where these particles
   accumulate in multiple organs such as the liver, heart, nervous system,
   and intestines, causing hepatotoxicity, cardiotoxicity, neurotoxicity,
   and gastrointestinal toxicity [[44]6–[45]10]. Among these species,
   ducks are especially vulnerable due to their feeding habits and the
   contamination of aquatic environments with MPs. Subsequently, ducks
   contaminated with MPs could enter the human diet, resulting in the
   bioaccumulation of MPs in the human body. By ingesting MPs from
   polluted water and food sources, these particles accumulate in various
   organs, especially the lungs, where they can cause severe damage.
   Despite the growing awareness of microplastic pollution, research into
   the molecular mechanisms underlying the toxic effects of MPs
   accumulation in the lungs remains limited.

   Mitochondria play a key role in cellular energy metabolism, redox
   balance, and programmed cell death [[46]11, [47]12]. Moreover, it is
   also the key regulator of inflammatory responses, driven by both its
   structural components and the diverse metabolic products it generates
   [[48]13]. Under the condition of dysfunction, mitochondria display
   fragmentation and membrane depolarization, leading to the production of
   large amounts of reactive oxygen species (ROS) and activating the
   mitochondrial cell death pathway [[49]14]. To counteract this damage,
   cells initiate mitophagy, a specialized autophagy that selectively
   degrades damaged, dysfunctional or surplus mitochondria, thereby
   safeguarding cellular homeostasis [[50]15]. Excessive accumulation of
   ROS in the mitochondria can trigger abnormal mitophagy, which
   ultimately results in cell death and contributes to the development of
   diseases [[51]16]. Remarkably, the accumulation of MPs in the body can
   cause oxidative damage to the heart, liver, and skeletal muscle,
   resulting in significant toxic effects [[52]9, [53]17, [54]18].
   However, whether mitochondrial dysfunction triggers mitophagy in
   MPs-induced lung injury remains unclear. We hypothesize that
   MPs-induced lung damage occurs via a pathogenic cascade in which
   mitochondrial dysfunction triggers ROS overproduction, dysregulating
   mitophagy and perpetuating a self-amplifying cycle of oxidative stress
   and chronic inflammation.

   In the mitochondrion, the TCA cycle serves as a pivotal metabolic
   pathway that plays a critical role in regulating mitochondrial function
   and managing oxidative stress in eukaryotic cells [[55]19]. The
   metabolites in the TCA cycle are widely recognized for their roles in
   energy metabolism, hypoxic response, and immunity regulation [[56]19].
   Numerous observations suggest that TCA cycle disruption is closely
   linked to mitochondrial dysfunction, promoting chain-dependent ROS
   production and ultimately leading to cell death [[57]20–[58]22].
   However, it remains unclear whether TCA cycle imbalance occurs under
   MPs exposure and contributes to MPs-induced lung cell death.
   Consequently, there is a strong interest in exploring whether MPs
   exposure disrupts TCA cycle balance, then accelerating mitochondrial
   dysfunction in the lung.

   Ducks, as a species of waterfowl, inhabit aquatic environments such as
   rivers and ponds, accessing water for drinking and sourcing food
   primarily from zooplankton and fish. Consequently, ducks exhibit a
   higher risk of exposure to and accumulation of MPs in comparison to
   other poultry species. To better understand the potential mechanism
   behind the toxic effect of PS-MPs, a duck model for PS-MPs (5 μm) was
   used to illustrate their effect on lung injury. We conducted
   metabolomics, and succinylomics to explore the hazards of PS-MPs
   exposure in the lung. Additionally, transmission electron microscopy
   (TEM), flow cytometry, oxygen consumption rate (OCR) analysis,
   immunofluorescence, qPCR, and western blot were utilized to determine
   the molecular regulatory mechanism involved in PS-MPs-exposed lung
   injury both in vivo and in vitro. These results provide new theories
   for comprehensively understanding the mechanisms of toxicity caused by
   MPs in birds and mammals.

Results

PS-MPs caused lung injury in ducks

   To investigate the toxicity of PS-MPs exposure to the lung, we detected
   the distribution of PS-MPs in lung tissues by scanning electron
   microscopy (SEM). We found that PS-MPs were widely distributed
   throughout the lung tissues with increasing doses of PS-MPs treatment
   (Fig. [59]1A–C). Compared to the control (CK) group, PS-MPs were
   significantly detected in lung tissues of exposed ducks, confirming the
   pulmonary deposition of microplastics. HE staining revealed
   dose-dependent structural lung injury, with exposed groups exhibiting
   obvious vascular congestion (characterized by erythrocyte infiltration
   into pulmonary capillaries) and dense inflammatory cell infiltration
   (Fig. [60]1D–F). These findings establish a direct correlation between
   PS-MPs accumulation and escalating pulmonary damage, underscoring the
   pathological consequences of microplastic exposure in ducks.

Fig. 1.

   [61]Fig. 1
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   PS-MPs distributed in the lung and triggered lung injury in ducks. A–C
   SEM examination (scale bar = 50 μm). Yellow arrows point to PS-MPs. D–F
   HE staining (scale bar = 400 μm and 100 μm, respectively)

PS-MPs exacerbated lung inflammation through the LPS/TLR4 pathway

   We explored the molecular pathways modification underlying inflammation
   induced by PS-MPs. PS-MPs markedly elevated LPS levels in the lung
   tissues of ducks compared to the CK group (p < 0.05, Fig. [63]2A), with
   almost two fold enhancement under high level PS-MPs exposure. Notably,
   higher TLR4 and MyD88 levels were observed in PS-MPs group, indicating
   that LPS overproduction in the lungs activated the TLR4 pathway and
   increased the phosphorylation of NF-κB and IκBα (p < 0.05,
   Fig. [64]2B). The lung tissues of ducks exhibited a notable elevation
   in the mRNA expression levels of MyD88, NF-κB, and IκBα in response to
   low and high doses of PS-MPs, in a dose-dependent manner (p < 0.05,
   Fig. [65]2C). Additionally, PS-MPs markedly elevated the release of
   IL-1β, IL-6, and TNF-α (p < 0.05, Fig. [66]2D–F) and promoted the
   protein expression level of pro-IL-1β and TNF-α (p < 0.05,
   Fig. [67]2G). Immunofluorescence (IF) analysis of lung sections
   revealed elevated TNF-α expression levels in PS-MPs-exposed ducks
   (Fig. [68]2H). Similarly, both mRNA and protein levels indicated that
   PS-MPs induced the expression of inflammatory mediators iNOS and COX2
   in the lungs (p < 0.05, Fig. S1). These results showed that PS-MPs
   exacerbated lung damage by activating LPS/TLR4 pathway.

Fig. 2.

   [69]Fig. 2
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   PS-MPs caused LPS overproduction and inflammatory response in the lung
   of ducks. A LPS levels (n = 10). B The relative expression levels of
   TLR4-related proteins (TLR4, MyD88, p-NF-κB, NF-κB, p-IκBα and IκBα) (n
   = 3). C The relative mRNA expression levels of TLR4-related genes
   (MyD88, NF-κB and IκBα) (n = 6). D-F The levels of IL-1β, IL-6 and
   TNF-α in the lung (n = 10). G The relative protein expression levels of
   pro-IL-1β and TNF-α (n = 3). H IF staining for TNF-α in the lung (scale
   bar = 50 μm). Different letters (a-c) indicate statistically
   significant differences (p < 0.05)

The metabolite profile of lung tissues indicated that PS-MPs potentially
induced mitochondrial dysfunction

   Given the crucial role of LPS/TLR4 activation in lung injury with or
   without PS-MPs, non-targeted metabolic profiles of the lungs were
   conducted. In the positive mode, 820 metabolites were identified, while
   422 metabolites were identified in the negative mode across all
   samples. To better understand the metabolic differences in the lung
   tissues based on PS-MPs exposure, principal component analysis (PCA)
   and orthogonal partial least squares discriminant analysis (OPLS-DA)
   were performed. PCA results showed that the PC1 explained 20.4% of the
   variations in positive mode and 17.1% in negative mode (Fig. [71]3A).
   OPLS-DA score plot analysis further confirmed the clear separations
   between lung tissues from the CK and HMPs groups in both positive mode
   and negative mode (Fig. [72]3B). Meanwhile, the explained variation
   (R2Y) and prediction capability (Q2) values in both modes were > 0.9
   and > 0.7, respectively, indicating the reliability of the model’s
   results (Fig. S2A).

Fig. 3.

   [73]Fig. 3
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   PS-MPs altered the composition of metabolites of the lung in ducks. A
   The PCA plot in positive and negative mode. B The OPLS-DA score plot in
   positive and negative mode. C Volcano plot of the differentially
   expressed metabolites in positive and negative modes. D KEGG pathway in
   positive and negative mode. E The common pathways contribute to the
   difference of energy metabolism in PS-MPs-exposed ducks. Red dot
   indicated upregulated metabolites, green dot indicated downregulated
   metabolites

   To identify differential metabolites in the lungs after PS-MPs
   exposure, we set a variable importance in projection (VIP) score > 1
   and p < 0.05 as thresholds. Compared to the CK group, 30 upregulated
   and 292 downregulated metabolites were screened in the positive mode,
   while 12 upregulated and 136 downregulated metabolites were observed in
   the negative mode (Fig. [75]3C, Fig. S2B, C). Functional enrichment
   analysis revealed that the affected metabolic pathways were primarily
   involved in amino acids biosynthesis, alanine, aspartate and glutamate
   metabolism, pyruvate metabolism and arginine biosynthesis
   (Fig. [76]3D). Notably, L-lactate, S-lactoylglutathione, phosphoenol
   pyruvate, succinate exhibited a notable decrease in response to high
   doses of PS-MPs, while L-glutamate represented an elevation in high
   PS-MPs group (Fig. [77]3E, Fig. S2D). These metabolites highlighted a
   key metabolic pathway network impacting pyruvate metabolism
   (downregulation in L-lactate and S-lactoylglutathione), glycolysis
   (downregulation in phosphoenol pyruvate), TCA cycle and oxidative
   phosphorylation (downregulation in succinate), arginine biosynthesis
   (upregulation in L-glutamate) following PS-MPs exposure (Fig. [78]3E,
   Fig. S2D). These findings suggested that PS-MPs exposure could disrupt
   mitochondrial metabolism.

PS-MPs disrupted tricarboxylic acid cycle (TCA) cycle metabolism

   To determine whether PS-MPs exposure directly affected TCA cycle
   metabolism, we conducted an absolute quantitative metabolomic analysis
   in lung tissues (Fig. [79]4A). Interestingly, we found that PS-MPs
   markedly reduced the abundance of key TCA cycle intermediates,
   including pyruvate, α-ketoglutarate, succinate, fumarate, malate, and
   oxaloacetate, indicating significant impairment of the TCA cycle
   (Fig. [80]4B, [81]C). This disruption aligned with concurrent
   reductions in energy-related metabolites (ATP, cyclic AMP, GTP, GDP)
   and redox cofactors (NADP^+, NAD^+), collectively pointing to profound
   dysfunction in oxidative phosphorylation (Fig. [82]4B, [83]C). These
   results indicated that PS-MPs exposure disrupted the balance of the TCA
   cycle and oxidative phosphorylation metabolism.

Fig. 4.

   [84]Fig. 4
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   The effect of PS-MPs exposure on the TCA cycle and oxidative
   phosphorylation metabolism in ducks. A Schematic diagram of TCA cycle.
   B Heatmap of differential metabolites. C The levels of metabolites in
   the TCA cycle and oxidative phosphorylation. *p < 0.05 vs CK group

Succinylomics analysis revealed that TCA cycle-related proteins were involved
in response to PS-MPs exposure

   To further explore whether PS-MPs could disrupt the TCA cycle and
   contribute to lung damage, we analyzed protein succinylation of lung
   tissues treated with or without PS-MPs. We identified 1489
   succinylation sites across 521 proteins, with 1376 sites and 458
   proteins being quantifiable (p < 0.05, Fig. [86]5A, Table S1). Among
   these, 67 succinylated proteins were localized to mitochondria
   (Fig. [87]5B, Table S2). Comparison between HMPs and CK group revealed
   that 13 succinylation sites across 13 proteins were significantly
   upregulated, while 51 succinylation sites across 44 proteins were
   significantly downregulated (Fig. [88]5C, Table S3). Notably, 12 out of
   the 57 differentially succinylated proteins were localized to
   mitochondria (Fig. [89]5D, Table S4).

Fig. 5.

   [90]Fig. 5
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   Succinylation of the TCA cycle-related protein was involved in the
   PS-MPs-exposed lung injury. A The numbers of succinylated sites and the
   corresponding proteins detected in PS-MPs-exposed lung tissues. B The
   distribution of succinylated proteins. C The numbers of differential
   succinylated sites and the corresponding differential proteins. D The
   distribution of differential succinylated proteins. E KEGG, GO-BP, and
   GO-MF analysis of differential succinylated proteins. F The
   differential succinylated sites and the corresponding proteins in the
   TCA cycle. Red dot indicated significantly upregulated succinylated
   sites. Green dot indicated significantly downregulated succinylated
   sites. G Succinylated profiling identified differential succinylated
   sites of TCA cycle-related proteins (IDH1 and MDH2). H Pattern diagram
   of the succinylated sites of IDH1 and MDH2

   Pathway enrichment analysis, including Kyoto Encyclopedia of Genes and
   Genomes (KEGG), Gene Ontology-Biological Process (GO-BP), and
   GO-Molecular Function (MF), showed significant enrichments in the
   process such as oxaloacetate metabolism, fumarate metabolism, NAD^+
   activity, and ADP binding (Fig. [92]5E, Tables S5, S6). Among the
   differentially succinylated proteins, we observed that the
   succinylation levels of IDH1 at lysine 245 (K245) and MDH2 at lysine
   204 (K204) were significantly decreased, whereas the succinylation of
   MDH2 at lysine 182 (K182) was significantly increased following PS-MPs
   treatment (Fig. [93]5F–H). This evidence highlighted the potential of
   PS-MPs to impair the mitochondrial function and contributed to lung
   damage through alterations in protein succinylation within the TCA
   cycle.

PS-MPs triggered mitochondrial dysfunction and oxidative stress

   Based on these results, we inferred that PS-MPs exposure could impair
   lung mitochondrial function. To explore this, we assayed the ATP levels
   and mtDNA copies in the lungs under PS-MPs exposure. Compared with CK
   ducks, the lungs of PS-MPs-treated ducks showed a significant decrease
   in ATP production with a dose-dependent manner (p < 0.05, Fig. [94]6A)
   and mtDNA levels were also decreased in PS-MPs-treated lungs compared
   to CK lung (p < 0.05, Fig. [95]6B). Then, we observed the mitochondrial
   morphology by TEM. The lungs of the PS-MPs ducks displayed significant
   mitochondrial abnormalities including swelling and broken or absent
   cristae (Fig. [96]6C). We also analyzed the expression of key
   mitochondrial regulators at both the protein and mRNA levels. PS-MPs
   ducks markedly inhibited the SIRT1, SIRT3, and TFAM levels (p < 0.05,
   Fig. [97]6D, [98]E). Furthermore, we investigated the expression of
   markers associated with mitochondrial dynamics. The OPA1, MFN1, and
   MFN2 levels were significantly downregulated (p < 0.05, Fig. [99]6F,
   [100]G). To assess the extent of oxidative stress, we detected the ROS
   level and antioxidant enzyme activity. PS-MPs markedly elevated the ROS
   and MDA levels, while the activities of SOD and CAT were substantially
   reduced compared to the CK group (p < 0.05, Fig. [101]6H–K). These
   results indicated that excessive PS-MPs exacerbated mitochondrial
   damage and oxidative stress response in the duck lungs.

Fig. 6.

   [102]Fig. 6
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   PS-MPs caused the mitochondrial function damage. A ATP levels (n = 10).
   B mtDNA copy number levels (n = 6). C TEM examination (scale bar = 500
   nm and 200 nm, respectively). D The relative protein expression levels
   of SIRT1, SIRT3, and TFAM (n = 3). E The relative mRNA expression
   levels of SIRT1, SIRT3, and TFAM (n = 6). F The relative protein
   expression levels of OPA1, MFN1, and MFN2 (n = 3). G The relative mRNA
   expression levels of OPA1, MFN1, and MFN2 (n = 6). H ROS levels (n
   = 10). I MDA levels (n = 10). J, K The activities of SOD and CAT (n
   = 10). Different letters (a-c) indicate statistically significant
   differences (p < 0.05)

PS-MPs enhanced mitochondrial damage and led to ROS overproduction in RLE-6TN
cells

   To further explore the toxicity of PS-MPs on mitochondrial damage in
   vitro, RLE-6TN cells were utilized as our model. First, the excessive
   PS-MPs on the mitochondrial abundance were assessed. Interestingly,
   PS-MPs treatment (100 and 500 μg/mL PS-MPs) significantly reduced mtDNA
   copy numbers in RLE-6TN cells (p < 0.05, Fig. [104]7A). In addition,
   PS-MPs exposure decreased mitochondrial respiratory capacity,
   indicating lower basal and maximal respiration levels, ATP production
   across different PS-MPs concentrations (p < 0.05, Fig. [105]7B). A
   reduction in mitochondrial membrane potential (MMP) was also observed
   in cells exposed to high doses of PS-MPs, consistent with an increase
   in ROS production (p < 0.05, Fig. [106]7C, [107]D). We further examined
   PS-MPs on the mitochondrial function in RLE-6TN cells. As shown in
   Fig. [108]7E and [109]F, high dose of PS-MPs significantly reduced the
   protein expression of the mitochondrial regulator (SIRT1, SIRT3, and
   TFAM) as well as mitochondrial fusion markers (OPA1, MFN1, and MFN2)
   (p < 0.05). Overall, our findings suggested that PS-MPs induced
   mitochondrial dysfunction and triggered ROS overproduction in vitro,
   exacerbating oxidative stress in RLE-6TN cells.

Fig. 7.

   [110]Fig. 7
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   PS-MPs impaired the mitochondrial function and activated ROS production
   in RLE-6TN cells. A mtDNA copy number levels (n = 3). B Measurement of
   mitochondrial oxygen consumption ratio (OCR). C Measurement of
   mitochondrial membrane potential (MMP). D ROS levels (n = 3). E The
   relative protein expression levels of SIRT1, SIRT3, and TFAM (n = 3). F
   The relative protein expression levels of OPA1, MFN1 and MFN2 (n = 3).
   Different letters (a-d) indicate statistically significant differences
   (p < 0.05)

PS-MPs induced mitophagy both in vivo and in vitro

   Mitophagy is responsible for removing damaged mitochondria. Thereby, we
   wondered whether mitophagy was activated by PS-MPs. Indeed, we observed
   that both protein and mRNA levels of LC3, ATG7, and BECN1 in lung
   tissues were significantly increased, while p62 was obviously decreased
   with increasing doses of PS-MPs exposure (p < 0.05, Fig. [112]8A,
   [113]B). These findings were further supported by IF staining of LC3
   and p62 (Fig. [114]8C, [115]D). Additionally, the expression levels of
   Parkin showed a concentration-dependent increase (p < 0.05,
   Fig. [116]8E, [117]F).

Fig. 8.

   Fig. 8
   [118]Open in a new tab

   PS-MPs mediated mitophagy. A The relative protein expression levels of
   LC3, ATG7, BECN1, and p62 (n = 3). B The relative mRNA expression
   levels of LC3, ATG7, BECN1 and p62 (n = 6). C, D IF staining for LC3
   and p62 in the lung (scale bar = 50 μm). E The relative protein
   expression levels of Parkin (n = 3). F The relative mRNA expression
   levels of Parkin (n = 6). Different letters (a-c) indicate
   statistically significant differences (p < 0.05)

   We then examined the effect of PS-MPs on mitophagy in vitro. As shown
   in Fig. [119]9A, we found the high dose of PS-MPs treatment contributed
   to an obvious increase in the protein levels of LC3, ATG7, and BECN1,
   alongside a reduction in p62 level, consistent with the IF staining
   results (p < 0.05, Fig. [120]9B, [121]C). Meanwhile, high dose of
   PS-MPs treatment significantly elevated Parkin protein level (p < 0.05,
   Fig. [122]9D). TUNEL assay was further determined that PS-MPs-exposed
   RLE-6TN cells directly led to cell death, which was markedly decreased
   by Mdivi-1 (p < 0.05, Fig.S3A). In addition, RLE-6TN cells exposed to
   PS-MPs presented much higher level of cleaved caspase 3 compared to
   untreated RLE-6TN cells, which was reversed by Mdivi-1 (p < 0.05, Fig.
   S3B). Collectively, our data suggested that increased mitophagy
   occurred under PS-MPs-induced mitochondrial dysfunction.

Fig. 9.

   [123]Fig. 9
   [124]Open in a new tab

   PS-MPs aggravated mitophagy in RLE-6TN cells. A The relative protein
   expression levels of LC3, ATG7, BECN1, and p62 (n = 3). B, C IF
   staining for LC3 and p62 in RLE-6TN cells (scale bar = 50 μm). D The
   relative protein expression levels of Parkin (n = 3). Different letters
   (a-d) indicate statistically significant differences (p < 0.05)

Discussion

   Microplastic residues in water and farmland have emerged as potential
   health risks and were also identified and accumulated within the lung
   tissues in both human and animals [[125]23–[126]25]. As waterfowl,
   ducks inhabit rivers and ponds, drink from these water sources. This
   lifestyle makes ducks more prone to encounter and ingest microplastics
   compared to other poultry species. In our study, we found that PS-MPs
   exposure to duck could accumulate in the lungs, leading to distinct
   pathological changes, including blood permeation into pulmonary
   capillaries and infiltration of inflammatory cells (Fig. [127]1D–F).
   Additionally, we observed an increase in LPS levels (Fig. [128]2A). LPS
   triggers inflammatory responses by binding to TLR4, activating the
   adaptor MyD88, and initiating NF-κB signaling pathways, which in turn,
   boost the release of critical pro-inflammatory mediators [[129]26,
   [130]27]. Consistent with these findings, we observed the upregulation
   of TLR4, MyD88, phosphorylated NF-κB, and phosphorylated IκBα in the
   lung tissues, resulting in elevating TNF-α, IL-6, and IL-1β
   (Fig. [131]2B–H). Our findings confirmed that PS-MPs exacerbate lung
   damage by activating the LPS/TLR4 pathway. Notably, previous studies
   have shown that MPs disrupt intestinal barrier integrity via the
   NF-κB/NLRP3/IL-1β/MLCK pathway and compromise reproductive function via
   the miR-199a-5p/HIF-1α and CNR1/CRBN/YY1/CYP2E1 signaling axes
   [[132]28, [133]29]. Together, these findings underscore the systemic
   toxicity of PS-MPs across multiple organ systems, with our study
   specifically elucidating the lung damage. Given the evolutionary
   conservation of inflammatory and stress-response pathways, these
   results raise critical concerns about the potential human health risks
   posed by microplastic pollution, particularly in populations with
   prolonged environmental or dietary exposure.

   Given the crucial role in regulating inflammatory responses utilizing
   multiple signals, that can drive chronic inflammation-associated
   diseases, such as periodontitis and osteoarthritis, in response to cell
   stress and loss of homeostasis [[134]30], we wondered whether
   mitochondria could act as potential inducers contributing to lung
   inflammation. Strikingly, exposure to PS-MPs led to significant
   impairment in critical mitochondrial pathways, notably the TCA cycle
   and OXPHOS system, as evidenced by pathway activity profiles
   (Fig. [135]3E). Absolute quantitative metabolomics analysis further
   corroborated these findings, demonstrating pronounced reductions in key
   metabolites associated with both the TCA cycle and OXPHOS system
   (Fig. [136]4B, [137]C). These results collectively confirm a systemic
   disruption in mitochondrial energy metabolism, compromising cellular
   energy production. Moreover, succinylomics analysis revealed that key
   enzymes of the TCA cycle were markedly modified in the PS-MPs-exposed
   group (Fig. [138]5F–H). Previous studies also showed that environmental
   hazards can disrupt the TCA cycle, leading to proteotoxic stress
   [[139]31]. Besides, disruption of the TCA cycle triggers mitochondrial
   damage by activating retrograde signaling pathways, which communicate
   metabolic dysfunction and induce profound metabolic reprogramming
   [[140]32, [141]33]. These findings strongly suggest that mitochondrial
   function is severely compromised under PS-MPs exposure.

   Mitochondria is widely recognized as the center of energy metabolism,
   playing an important role in ATP production and maintaining normal
   cellular functions [[142]34]. We assessed ATP levels in both tissual
   and cellular samples, and as expected, ATP production was significantly
   reduced (Figs. [143]6A, [144]7B), directly indicating mitochondrial
   dysfunction in response to PS-MPs exposure. This is consistent with
   observed mitochondrial morphological changes, such as swelling and
   disrupting or absent cristae in the PS-MPs group (Fig. [145]6C). Next,
   we explored whether the self-protection mechanisms including
   mitochondrial biogenesis and fusion were activated to preserve the
   mitochondrial function. However, mitochondrial biogenesis was
   significantly impaired, as evidenced by the notable reduction in TFAM
   protein levels. Additionally, we observed significant downregulation of
   OPA1, MFN1, and MFN2 proteins, essential for mitochondrial fusion.
   SIRT3, which promotes mitochondrial fusion by deacetylating OPA1
   [[146]35], was also downregulated, further suggesting that
   mitochondrial fusion was inhibited (Fig. [147]6D, [148]F).
   Mitochondrial fusion is critical for maintaining the mitochondrial
   function, as it helps dilute damaged mitochondrial DNA, redistribute
   metabolic intermediates, and improve mitochondrial respiration
   [[149]36]. However, in this case, these self-repair pathways were
   disrupted, indicating that PS-MPs caused profound mitochondrial
   dysfunction. This prompted investigation into whether oxidative stress
   threshold has been exceeded. Our hypothesis was supported by
   significant increases in ROS production and MDA levels (Fig. [150]6H,
   [151]I). ROS, a critical mediator of stress-related signaling pathways,
   directly reflects cellular oxidative stress. Similarly, elevated MDA, a
   hallmark of lipid peroxidation, correlates with ROS overproduction. The
   marked rise in both biomarkers signifies severe oxidative damage,
   aligning with prior studies where PS-MPs induced ROS accumulation in
   ovarian granulosa cells and colon tissues [[152]37, [153]38]. Notably,
   key antioxidant enzymes, such as superoxide dismutase (SOD) and
   catalase (CAT), were significantly suppressed under PS-MP exposure,
   diverging from earlier findings [[154]39]. These enzymes are essential
   for maintaining redox balance by scavenging ROS, and their depletion
   likely exacerbates oxidative injury [[155]40–[156]42]. Combine with the
   increased ROS and MDA levels, along with decreased antioxidant activity
   (e.g., SOD and CAT) (Fig. [157]6H–K). This evidence showed that
   mitochondrial dysfunction promotes the accumulation of ROS, amplifies
   oxidative stress, and impairs antioxidant defenses. Critically,
   pre-treatment with the antioxidant N-acetylcysteine (NAC) or oxidative
   stress inhibitors AM251 substantially attenuated ROS generation and
   inflammatory cascades, alleviating PS-MP-induced toxicity [[158]38,
   [159]43]. These findings collectively identify oxidative stress as a
   central mediator of PS-MP toxicity, linking mitochondrial impairment to
   systemic cellular damage.

   In addition to impair mitochondrial biogenesis and reduce mitochondrial
   fusion, we observed a significant reduction in mtDNA copies
   (Figs. [160]6B, [161]7A), a direct marker of mitochondrial abundance.
   Therefore, it makes sense to anticipate that damaged mitochondria will
   be removed by the process of mitophagy, which is the activation of a
   quality control mechanism. To keep the mitochondria intact, mitophagy
   is necessary because it maintains the proper ratio of healthy to damage
   mitochondria, hence regulating the quality of mitochondria [[162]44]. A
   prior study demonstrated that mitophagy is intricately associated with
   the generation of ROS [[163]45]. Furthermore, the ROS-mediated
   mitophagy induced by PS-MPs contributes to kidney damage [[164]46]. In
   our study, we systematically investigated whether mitophagy occurred in
   the lungs following PS-MPs exposure. We found that ATG7 and BECN1 were
   significantly upregulated. BECN1 is a key component of autophagy,
   involved in the formation, extension, and maturation of autophagosomes
   [[165]47, [166]48]. Additionally, the LC3 II/LC3 I ratio, a
   well-established marker of mitophagy, was significantly increased under
   PS-MPs treatment, along with a notable downregulation of p62. It was
   reported that cell-specific p62 ablation led to excessive IL-1β
   production, which was associated with hypersensitivity to
   endotoxin-induced shock [[167]49]. Activated mitophagy could reduce p62
   levels as well [[168]50]. Meanwhile, IF staining consistently confirmed
   the upregulation of LC3 and the downregulation of p62. Furthermore, we
   observed a significant upregulation of Parkin, a protein closely
   involved in mitophagy [[169]51]. At the cellular level, these findings
   were further validated in RLE-6TN cells. Collectively, our results
   demonstrated strong activation of mitophagy in response to PS-MPs
   exposure-induced mitochondrial dysfunction. While mitophagy is
   necessary to preserve the quality of the mitochondria, abnormal or
   excessive mitophagy can cause an excess of ROS in the mitochondria,
   which can lead to cell death [[170]52]. Notably, pharmacological
   inhibition of autophagy partially relieves PS-MP-induced mitochondrial
   and lysosomal damage [[171]37], suggesting that PS-MPs drives
   pathological mitophagy, disrupting mitochondrial homeostasis and
   exacerbating lung inflammation in ducks. In addition, this evidence
   showed that MPs penetrated the lungs to reduce the viability of lung
   cells by inducing oxidative stress and causing cell death [[172]53].
   However, to address current biological limitations, future studies
   should integrate primary duck lung cells with RLE-6TN cell models to
   further elucidate the mechanisms underlying PS-MP-induced mitochondrial
   dysfunction. Furthermore, the dose-dependent effects were evident:
   low-dose PS-MPs exposure caused less prominent histopathological
   changes, oxidative stress, and mitophagy activation compared to
   high-dose groups. Intriguingly, despite high-dose PS-MPs exposure,
   ducks exhibited no obvious clinical symptoms and maintained normal
   survival rates, raising concerns about potential human consumption of
   PS-MPs-contaminated poultry products. Previous studies suggested that
   MPs caused serve damage and inflammation in poultry products, such as
   heart, liver, muscle and intestine [[173]10, [174]54, [175]55]. While
   such meat may appear safe, PS-MPs bioaccumulation in tissues could pose
   long-term risks to human health through dietary exposure. Notably, NAC,
   an effective antioxidant, markedly reduced polystyrene nanoplastics
   (PS-NPs)-triggered oxidative damage in lung epithelial cells by
   restoring mitochondrial function and inhibiting cell death [[176]56,
   [177]57], as well as Mdivi-1 [[178]58]. Although antioxidants (e.g.,
   NAC) and autophagy inhibitors (e.g., Mdivi-1) show therapeutic
   potential in mitigating plastics toxicity, these interventions were not
   explored in the current study. Nevertheless, our mechanistic insights
   into the mitochondrial dysfunction-inflammatory axis provide a
   foundation for developing targeted strategies to counteract
   microplastic-related health risks in humans.

   Taken together, our findings verified that PS-MPs disrupted
   mitochondrial homeostasis, leading to excessive mitophagy and LPS/TLR4
   pathway activation, which potentially contributed to lung injury. Our
   study provides new insights into the mechanism behind the damage in the
   lung, highlighting the critical role of mitochondrial dysfunction under
   PS-MPs exposure. Future research should aim to explore the effects of
   PS-MPs on other organ systems and investigate the specific mechanisms
   and potential therapeutic strategies by which PS-MPs interacts with
   cellular and tissue responses.

Materials and methods

Preparation of animals and experimental design

   PS-MPs (Unibead, 5 μm in size) were obtained from Tianjin BaseLine
   Scientific Co., Ltd. (Tianjin, China). The characterization of PS-MPs
   was measured using electron microscopy (Zeiss, Germany) and SEM (Fig.
   S4). One-day-old Shaoxing ducklings (n = 60), procured from Hanchao
   Poultry Company in Hangzhou, China, were randomly assigned to three
   experimental groups, each comprising 20 individuals: (1) The control
   group (CK) received standard drinking ultrapure water without PS-MPs;
   (2) the low-dose group (LMPs) was administered drinking ultrapure water
   containing a concentration of 1 mg/L of PS-MPs; and while (3) the
   high-dose group (HMPs) received drinking ultrapure water with a
   concentration of 100 mg/L of PS-MPs. The exposure doses of PS-MPs were
   selected based on previous toxicological studies of PS-MPs in mice and
   poultry [[179]9, [180]18]. In these previous studies, 1 mg/L, 10 mg/L
   and 100 mg/L were used to feed the chicken [[181]9]. Considering the
   previous research reported the 1 mg/L PS-MPs is an environment-related
   concentration [[182]59, [183]60]. Therefore, we chose moderate doses of
   1 mg/L and 100 mg/L for this study. After one week of the adaptation
   period, the ducklings were maintained on these treatments for four
   weeks. During the experimental period, ducks were raised in cage, and
   kept at room temperature with flowing air. The ducks were fed ad
   libitum with the same formula diet and subjected to a standard light
   regimen of 17 h light (17L:7D). All experimental ducks were healthy and
   were not administered any antibiotic treatments during the experiment.
   At five weeks old, the animals were euthanized (approved number:
   2024ZAASLA019), lung tissues were collected and stored in 4%
   formaldehyde or at −80°C for further analysis.

Cell culture

   RLE-6TN cells were obtained from Wanwu Biotechnology Co., Ltd (Anhui,
   China). Cells were cultured in Ham’s F-12K medium supplemented with 10%
   FBS (Gibco), 1% ITS, and 5 ng/mL EGF at 37 °C with 5% CO[2]. Upon
   achieving 50% confluency, the cells were treated with various
   concentrations of PS-MPs (0, 10, 100, 500 μg/mL) for 48 h [[184]18]. In
   addition, Mdivi-1 (1 μM, Selleck, USA) was administered on 100 μg/mL
   PS-MPs-treated RLE-6TN cells for 24 h.

Oxidative stress detection

   Lung malondialdehyde (MDA), catalase (CAT), and superoxide dismutase
   (SOD) were measured (n = 10). Briefly, a total of 0.1 g of lung tissue
   was homogenized in normal saline and centrifuged to collect the
   supernatant. The levels of MDA and the activities of CAT and SOD were
   then detected following the manufacturer’s protocols (Solarbio,
   Beijing, China).

Inflammatory cytokine and LPS assays

   A total of 0.1 g of lung tissue was homogenized and centrifuged to
   collect the supernatant. The IL-1β, IL-6, and TNF-α levels (n = 10) in
   the tissue were then measured according to the manufacturer’s
   instructions (mlbio, Shanghai, China). The LPS levels (n = 10) were
   quantified from Beijing Sinouk Institute of Biological Technology
   (Beijing, China).

Reactive oxygen species (ROS) level detection

   A total of 0.05 g of lung tissue was homogenized and centrifuged to
   obtain the supernatant. Then, the supernatant was mixed with 50 μM
   2,7-dichlorofluorescein diacetate (DCFH-DA) solution and incubated at
   37 °C for 30 min. The fluorescence intensity was measured (n = 10)
   using a microplate reader (Tecan, Switzerland). Cells were incubated
   with 5 μM CellROX (Thermo, USA) for 30 min, washed twice with PBS, and
   the intracellular ROS levels (n = 3) were measured by flow cytometry
   (Beckman, USA).

ATP content measurement

   Lung tissue was lysed and centrifuged to collect the supernatant. The
   ATP content in the supernatant was measured (n = 10) according to the
   manufacturer’s instructions (Thermo, MA, USA).

Hematoxylin and eosin (H&E) staining

   Lung tissues were fixed in 4% polyformaldehyde overnight and embedded
   in paraffin wax. Then the tissues were sectioned into 5-μm slices and
   stained with hematoxylin and eosin (H&E). After decolorization in
   alcohol, the stained sections were examined under an upright light
   microscope (Olympus, Japan).

PS-MPs distribution detection

   Lung tissues were fixed in 2.5% glutaraldehyde at 4 ℃ overnight. Then,
   the samples were fixed in a 1% osmium tetroxide (pH 7.4), dehydrated
   with ethanol solutions, and plated onto gold in a vacuum coating
   device. The PS-MPs in the lung were observed with SEM (SU8010; Hitachi,
   Japan).

Mitochondrial morphology observation

   Lung tissues (≈1 mm^3) were fixed in 2.5% glutaraldehyde at 4 ℃
   overnight. The tissues were then post-fixed with 1% osmium tetroxide,
   dehydrated, and embedded in epoxy resin, followed by staining with 2%
   uranyl acetate. Images of the sections were captured by TEM (HT7650;
   Hitachi, Japan).

Detection of the gene expression level by qPCR

   qPCR was performed on a LightCycle 96 Real-Time System (Roche,
   Switzerland) using the TB Green^® Premix Ex Taq (TAKARA, Tokyo, Japan).
   Reaction conditions included one cycle of 95 °C for 30 s, 45 cycles of
   95 °C for 5 s and 60 °C for 20 s. GAPDH was employed as the
   housekeeping gene, and relative expression levels were calculated using
   the 2^−ΔΔCt method [[185]61]. The gene primer sequences are detailed in
   Table S7 and Fig. S5.

Mitochondrial DNA (mtDNA) copies detection

   Relative levels of mtDNA copy number were determined as described by
   Eaton et al. [[186]62]. Briefly, total DNA was extracted from all
   tissues or cells using the QIAamp DNA Mini Kit (Qiagen, Germany). A
   total of 2 ng DNA was used for qPCR analysis. For duck lung tissues (n
   = 6), primers targeting the mitochondrial COX1 (mtCOX1) gene were
   designed to assess mtDNA copy number, with the 18 s rRNA gene serving
   for normalization. For RLE-6TN cells (n = 3), primers for the
   mitochondrial ND5 (mtND5) gene were used to measure the mtDNA copy
   number, with the Rpl13 gene used for normalization. The primer
   sequences for mtDNA detection are provided in Table S8 and Fig. S6.

Protein expression level detection

   Tissues and cell lines were lysed using RIPA buffer (adding PMSF and
   phosphatase inhibitor). Protein concentrations were detected using the
   BCA protein assay kit (Beyotime, China). The methods followed in this
   study were based on our previous work [[187]54]. The band of protein is
   detected by exposing an X-ray film to the membrane. The information on
   primary antibodies is shown in Table S9.

Immunofluorescence (IF) staining

   For the in vivo assay, lung tissues were fixed in 4% polyformaldehyde,
   embedded in paraffin, and sectioned into 5-μm slices. For the in vitro
   assay, cells were fixed with 4% polyformaldehyde and permeabilized with
   0.5% Triton X-100. Both the tissue sections and cells were blocked with
   5% bovine serum and incubated with primary antibodies against LC3, p62,
   and TNF-α overnight. Afterward, samples were incubated with
   FITC-conjugated IgG (H + L) for 1 h. DAPI was used for nuclear
   staining. Fluorescence images were acquired using a microscope (Zeiss,
   Germany). Details of the primary antibodies are provided in Table S10.

Oxygen consumption rate (OCR) assays

   OCR was measured using Seahorse XFe96 Extracellular Flux Analyzer
   (Agilent, USA) [[188]63]. Briefly, a total of 1 × 10^4 cells treated
   with different concentration of PS-MPs (n = 3) were seeded into the
   XF96 cell culture microplate and incubated overnight. Then, the culture
   medium was replaced with assay buffer, prepared from XF base medium
   supplemented with 10 mM D-glucose, 2 mM sodium pyruvate, and 4 mM
   L-glutamine. The OCR was measured following the sequential addition of
   oligomycin (2 μM), FCCP (0.5 μM), and rotenone/antimycin A (1 μM each).
   Data acquisition and analysis were performed using Seahorse XF96 Wave
   software.

TUNEL staining

   Apoptotic cells were detected using an In Situ Cell Death Detection Kit
   according to its manufacturer’s instructions (Roche, Germany). Briefly,
   the cells were seeded in 12-well plate, fixed with 4% polyformaldehyde
   and permeabilized with 0.5% Triton X-100. Then, TUNEL reaction mixture
   was added to incubated cells for 1 h at 37 °C in a light-resistant
   container. DAPI was used for nuclear staining. Fluorescence images were
   acquired using a microscope (Zeiss, Germany).

Non-targeted metabolomic analysis

   Lung tissues from CK (n = 10) and HMPs (n = 10) ducks were prepared for
   non-targeted metabolomics analysis as described by Li et al. [[189]64].
   Briefly, samples were extracted with methanol and centrifuged to obtain
   the supernatant. The supernatants were then freeze-dried under vacuum
   and re-dissolved in 80% methanol solution containing
   2-chlorobenzalanine. Metabolic profiling was performed using liquid
   chromatography-mass spectrometry (LC–MS) on an ACQUITY UPLC^® HSS T3
   column (Thermo, USA).

Absolute quantification of TCA cycle and oxidative phosphorylation
intermediates based on MRM method

   Lung tissues from CK (n = 10) and HMPs (n = 10) ducks were prepared for
   targeted metabolomics analysis according to Zhang et al. [[190]65].
   Briefly, the samples were processed for metabolite extraction, followed
   by separation of metabolites using ultra-high performance liquid
   chromatography (UHPLC; Agilent). Mass spectrometry analysis was
   performed using an AB SCIEX 5500 QTRAP mass spectrometer (USA).

Quantitative succinylomics detection

   Lung tissues from CK (n = 3) and HMPs (n = 3) ducks were collected for
   succinylomic analysis according to Liu et al. [[191]66]. Briefly, the
   lung tissues were lysed with lysis buffer. Proteins were precipitated
   with trichloroacetic acid, washed with pre-cooled acetone, and
   redissolved in triethylammonium bicarbonate (TEAB). The proteins were
   then digested, reduced, alkylated, and enriched using
   anti-succinyllysine antibody-conjugated agarose beads (PTM-402, PTM
   BIO) for LC–MS/MS analysis (timsTOF HT, Bruker, Germany). Tandem mass
   spectra were searched against the Anas platyrhynchos Uniprot database
   used to search tandem mass spectra. Upregulated and downregulated
   proteins/sites were identified with fold changes > 1.5 or < 0.67,
   respectively. Functional annotations and pathway enrichment of
   differentially succinylated proteins were analyzed using GO and KEGG
   analysis. Subcellular localization was predicted using Wolfpsort
   software.

Statistical analysis

   The results of this study are reported as mean ± standard deviation
   (SD), and graphical representations are generated using GraphPad Prism
   8.0 software (CA, USA). The comparisons in different concentration of
   PS-MPs in vitro and in vivo were fulfilled using one-way ANOVA with
   Tukey tests and statistical comparisons of muti-omics between CK and
   HMPs groups were performed using a t-test analysis. The p value cutoff
   was set as 0.05.

Supplementary Information

   [192]Additional file 1.^ (2.1MB, docx)
   [193]Additional file 2.^ (3.1MB, xlsx)
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   [202]Additional file 11.^ (16.4KB, docx)

Author contributions

   T.T.G., and M.H.K. carried a majority of experiments and analyzed data.
   T.T.G., and M.H.K. wrote the main manuscript. T.T.G., and M.H.K.
   prepared figures. M.C.D. and L.C. provided support for animal
   experiment. T.T.G. and Y.T. analyzed the bioinformatic data. T.T.G. and
   W.W.X. performed the statistical analysis. T.T.G., M.H.K., T.Z. and
   L.Z.L. conceived the study, designed the study, evaluated data, and
   revised the manuscript. All authors read and approved the final
   manuscript.

Funding

   This work was supported by the Key R&D Program of Zhejiang Province
   (2024C02004), Zhejiang Provincial Natural Science Foundation of China
   (LQ23C170001), National Natural Science Foundation of China (32372862),
   and China Agriculture Research System of MOF and MARA (CARS-42–6).

Availability of data and materials

   No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

   All procedures were approved by the Zhejiang Academy of Agricultural
   Sciences Ethics Committee (Approved number: 2024ZAASLA019).

Consent for publication

   All authors have agreed to publish this manuscript.

Competing interests

   The authors declare no competing interests.

Footnotes

   Publisher's Note

   Springer Nature remains neutral with regard to jurisdictional claims in
   published maps and institutional affiliations.

   Tiantian Gu and Minghua Kong contributed equally to this work.

Contributor Information

   Tiantian Gu, Email: gtt19931029@126.com.

   Lizhi Lu, Email: lulizhibox@163.com.

References