Abstract

   This study investigates the size-dependent mechanisms by which
   polystyrene (PS) microplastics (MPs) cause intestinal epithelial
   injury, focusing on the differential effects of nanoscale (100 nm) and
   microscale (10 μm) particles. Using both cellular and animal models, we
   found that 100 nm MPs are internalized by intestinal epithelial cells
   via endocytosis, triggering oxidative stress and ferroptosis, while
   10 μm MPs primarily induce epithelial damage through mechanical
   disruption and metabolic reprogramming. Proteomic analyses revealed
   that small MPs significantly upregulate ferroptosis-related genes,
   including Fosl1 and components of the p53 pathway. Functional studies
   showed that Fosl1 promotes p53 transcription, suppresses Slc7a11, and
   contributes to ferroptosis. Mechanistically, ferroptosis was
   characterized by intracellular iron accumulation, glutathione (GSH)
   depletion, GPX4 inactivation, and excessive lipid peroxidation.
   Treatment with Ferrostatin-1 or Fosl1 knockdown alleviated ferroptosis
   and epithelial injury. In contrast, large MPs activated the
   mechanosensitive YAP pathway, leading to cytoskeletal remodeling and a
   metabolic shift from oxidative phosphorylation to anaerobic glycolysis.
   This metabolic reprogramming was associated with increased
   inflammation. These findings demonstrate that microplastic particle
   size critically determines toxicity mechanisms—ferroptosis for small
   particles and YAP-mediated metabolic disruption for large ones—and
   suggest potential therapeutic targets for mitigating
   microplastic-induced intestinal damage.

Graphical abstract

   [52]graphic file with name 12951_2025_3545_Figa_HTML.jpg

Supplementary Information

   The online version contains supplementary material available at
   10.1186/s12951-025-03545-1.

   Keywords: Nanoplastics, Microplastics, Ferroptosis, Fosl1, Immunogenic
   cell death, YAP signaling, Metabolic reprogramming

Introduction

   The increasing accumulation of plastic waste has become a significant
   global environmental issue, drawing widespread concern. As an
   inexpensive and highly versatile synthetic material, plastic has found
   widespread application across numerous sectors, including packaging,
   household goods, construction materials, and medical devices. The
   remarkably slow decomposition of plastic results in its long-term
   persistence in natural ecosystems, thereby intensifying environmental
   concerns. Currently, over 400 million tons of plastic are produced
   annually [[53]1], resulting in the accumulation of vast quantities of
   plastic waste that cause significant harm to ecosystems and
   biodiversity [[54]2, [55]3].

   In recent years, microplastics (MPs) and nanoplastics (NPs), which
   arise from the degradation or fragmentation of larger plastic debris,
   have been increasingly detected in diverse environmental matrices,
   including oceans, freshwater systems, soil, air, and even the food
   chain. These particles can be ingested by marine organisms and
   terrestrial animals, bioaccumulate through trophic transfer, and
   ultimately enter the human body via seafood, drinking water, and
   agricultural products [[56]4].

   Microplastics and nanoplastic particles are degradation by-products of
   plastic, which are ubiquitously distributed in the environment and have
   the potential to enter organisms via the food chain. The most common
   microplastics in the environment are polyethylene (PE), polyvinyl
   chloride (PVC), polypropylene (PP), polyethylene Terephthalate (PET)
   and polystyrene (PS) [[57]5]. Microplastics generally refer to plastic
   particles smaller than 5 mm [[58]6, [59]7]. While a universal
   definition for nanoplastics (NPs) remains elusive, most studies
   describe them as plastic particles ranging from 1 to 1000 nanometers in
   size [[60]2, [61]8, [62]9].

   Recent research has shifted from a purely environmental perspective to
   growing concern about the potential impacts of MPs and NPs on
   biological systems. Owing to their small size and physicochemical
   characteristics, these particles are capable of interacting with cells
   and tissues, thus posing risks to biological health [[63]10, [64]11].
   MPs and NPs are primarily ingested through the oral route and tend to
   accumulate in the gastrointestinal (GI) tract over time. Once in the
   intestine, they directly interact with intestinal epithelial cells,
   compromising epithelial barrier integrity, disrupting immune balance,
   and affecting gut health. These disruptions can trigger inflammatory
   responses, damage mucosal structures, and potentially promote
   tumorigenesis, contributing to a cascade of gastrointestinal disorders
   [[65]12–[66]15].

   As the first point of contact for ingested MPs and NPs, intestinal
   epithelial cells play a vital role in maintaining gut barrier function
   and immune homeostasis [[67]16–[68]18]. Consequently, significant
   attention has been directed toward understanding the relationship
   between gut health and microplastic pollution. The intestinal barrier
   not only acts to prevent the invasion of harmful substances but also
   plays a crucial role in maintaining normal immune function [[69]18].
   The intestinal barrier prevents the translocation of harmful substances
   while also orchestrating immune responses. Due to their larger size,
   MPs generally cannot enter cells via endocytosis. Instead, they are
   thought to damage epithelial integrity by direct physical compression,
   inducing membrane rupture and triggering mechanical stress [[70]19].
   Therefore, the damage caused by MPs and NPs is likely mediated
   primarily through physical interactions with intestinal epithelial
   cells. Such contact may compromise the intestinal barrier through
   multiple mechanisms, including mechanical disruption, activation of
   immune responses, and induction of oxidative stress [[71]14]. In
   contrast, due to their smaller size and larger surface area,
   nanoplastics are more readily absorbed by the body and can enter cells
   [[72]20], triggering toxic reactions such as oxidative stress, DNA
   damage, and cell apoptosis [[73]21]. Notably, nanoplastics can enter
   cells directly via endocytosis by intestinal epithelial cells,
   activating intracellular oxidative stress responses and causing
   cellular damage [[74]22–[75]26]. Furthermore, nanoplastic particles may
   alter the gut microbiome, further disrupting intestinal immune function
   and metabolic stability [[76]27–[77]29].

   Once internalized, MPs and NPs may undergo limited biodegradation and
   are thought to persist in tissues, especially the liver, intestines,
   and lymphoid organs. The intracellular handling of NPs may involve
   autophagic or lysosomal pathways, while MPs may be retained within the
   gut lumen or taken up by macrophages [[78]30]. However, the metabolic
   fate and clearance mechanisms of these particles remain poorly
   characterized, necessitating further investigation.

   Notably, micro/nanomaterial-mediated toxicity has emerged as a critical
   toxicological concept. These materials can induce organ-specific
   dysfunction, inflammatory responses, and even systemic immune
   alterations depending on their size, shape, and surface properties.
   Such findings underscore the need to understand how nanoscale and
   microscale plastics exert differential biological effects [[79]31].

   Despite the expanding literature on microplastic-induced gut toxicity,
   few studies have systematically compared how particle size influences
   specific mechanisms of intestinal damage. Understanding the
   size-dependent toxicological pathways of MPs and NPs is critical for
   accurate risk assessment and the identification of targeted
   intervention strategies. This study addresses this gap by directly
   comparing PS particles across a size gradient (100 nm to 10 μm), which
   allows for a controlled evaluation of cellular and molecular responses
   to different plastic scales. Emphasizing this comparative size-based
   approach is a novel aspect of our work, as most previous studies have
   focused on a single plastic type or size. Using both in vivo (mouse)
   and in vitro (intestinal epithelial cell) models, we administered
   particles of 100 nm, 500 nm, 1 μm, 5 μm, and 10 μm via oral gavage for
   a 6-week period. We employed cell viability assays, transmission
   electron microscopy, flow cytometry, and proteomics to investigate the
   resulting inflammatory and molecular changes.The results indicated that
   100 nm nanoplastics induced ferroptosis [[80]32, [81]33]in intestinal
   epithelial cells. Following ferroptosis, cellular antigens were
   released and subsequently recognized by dendritic cells (DCs) [[82]34,
   [83]35], which presented the antigens to CD8 + T cells [[84]36],
   leading to an increased infiltration of both dendritic cells and
   CD8 + T cells into the intestinal mucosa. This exacerbated inflammation
   and further promoted gut injury. Proteomic sequencing revealed that
   Fosl1 expression was upregulated, suggesting its role in regulating P53
   [[85]37] transcription and participating in the ferroptosis process.
   The ferroptosis inhibitor Ferrostatin-1 [[86]38, [87]39] was used to
   detect ROS, MDA and Western blot on cell experiments, and in vivo mice
   experiments, to further validate the occurrence and effect of
   ferroptosis. In contrast, the damage induced by larger microplastic
   particles appears to be associated with metabolic reprogramming.
   Proteomics and non-targeted energy metabolomics analyses suggested that
   glycolysis plays a role in this process. Additionally, larger
   microplastic particles may induce mechanical changes at the cellular
   level, such as compression-induced injury, which upregulates the key
   protein YAP [[88]40, [89]41] and drives a shift in metabolism from
   oxidative phosphorylation to anaerobic glycolysis. Intervention with
   the YAP inhibitor Verteporfin [[90]42] demonstrated that suppressing
   YAP translocation to the cell nucleus could potentially reverse
   inflammation and alleviate gut damage induced by microplastics.

   This study has elucidated the potential mechanisms underlying gut
   inflammation induced by microplastics of different sizes, highlighting
   that microplastics and nanoplastics exert distinct, size-specific
   mechanisms that contribute to gut damage. These findings offer
   important theoretical insights into the molecular mechanisms underlying
   microplastic-induced intestinal injury, and propose potential
   therapeutic targets for the prevention and mitigation of such damage.

Materials and methods

Materials

   Two types of polystyrene (PS) microplastics (Jiangsu Zhichuan, China)
   were used in this study. Conventional PS particles with sizes of
   100 nm, 500 nm, 1 μm, 5 μm, and 10 μm were applied in animal and
   cell-based toxicological studies. Red fluorescent PS microplastics
   (Ex = 520 nm, Em = 580 nm) of the same sizes were used for visualizing
   uptake, accumulation, and distribution in vivo and in vitro. Particle
   morphology and size were confirmed by scanning electron microscopy
   (SEM) and laser particle size analysis. Chemical composition was
   validated using Fourier-transform infrared spectroscopy (FTIR).

Animal experimental design

   Four-week-old male BALB/c mice (SPF grade) were obtained from the Anhui
   Experimental Animal Center (Hefei, China) and housed under standardized
   conditions (temperature: 22–26 °C; 12-hour light/dark cycle). After a
   one-week acclimatization period, mice were randomly assigned into
   experimental groups based on the study objectives, with 5 mice per
   group as follows:

Size-dependent toxicity evaluation (n = 30, 5 per group)

   Control (PBS), and five PS microplastic exposure groups (100 nm,
   500 nm, 1 μm, 5 μm, 10 μm). Each treatment group received 200 µL of
   1 mg/mL PS microplastics (equivalent to 0.1 mg/day) by oral gavage
   daily for 7 weeks; the control group received an equal volume of PBS.

Ferroptosis Inhibition experiment (n = 25, 5 per group)

   Control, 100 nm MPs, 10 μm MPs, 100 nm MPs + Ferrostatin-1, and 10 μm
   MPs + Ferrostatin-1. Ferrostatin-1 (1 mg/kg) was administered via
   intraperitoneal injection every other day for 7 weeks, concurrent with
   MP gavage.

YAP Inhibition experiment (n = 15, 5 per group)

   Control, 10 μm MPs, and 10 μm MPs + VTPF. VTPF (5 mg/kg) was
   intraperitoneally injected twice per week for 7 weeks, along with daily
   gavage of 10 μm MPs (0.1 mg/day).

   Across all experiments, prior to the final gavage, mice were weighed
   and fasted for 8 h. They were then euthanized via cervical dislocation.
   Blood samples and gastrointestinal tissues—including the stomach,
   cecum, and colon—were collected for subsequent histological,
   biochemical, and immunological analyses. All animal protocols were
   reviewed and approved by the Animal Ethics Committee of Anhui Medical
   University and adhered to institutional guidelines for animal welfare.

Cell culture and SiRNA transfection

   The human colonic epithelial cell line NCM460 was obtained from Wuhan
   Warner Biological Technology. Cells were maintained in RPMI 1640 medium
   (Gibco, USA) supplemented with 10% fetal bovine serum, 100 IU/mL
   penicillin, and 100 µg/mL streptomycin at 37 °C in a humidified 5% CO₂
   incubator. Upon reaching > 90% confluence, cells were passaged and
   seeded into 6-, 24-, or 96-well plates for subsequent experiments.
   Fosl1-specific siRNAs and YAP-specific siRNAs were transiently
   transfected into NCM460 cells using the jetPRIME transfection reagent,
   following the manufacturer’s instructions. The siRNA oligonucleotide
   sequences were as follows: Fosl1-siRNA-1: 5′-GCUCAUCGCAAGAGUAGCA-3′ and
   Fosl1-siRNA-2: 5′-GAGCUGCAGUGGAUGGUAC-3′ and YAP-siRNA-1:
   5′-GCCATTAAAGGCAGCTGTTC-3′ and YAP-siRNA-2: 5′-AGCACTGTGCCAGGTATCAC-3′.
   Briefly, siRNAs were diluted in jetPRIME buffer, mixed with the
   transfection reagent, and incubated at room temperature for 10 min
   before being added to the cells. After 12 h of transfection, the medium
   was replaced with fresh complete culture medium, and the cells were
   incubated for an additional 48 h. Cells were then harvested for protein
   extraction, and the knockdown efficiency of Fosl1 and YAP was confirmed
   by Western blot analysis.

Characterization of polystyrene microplastics

   Polystyrene (PS) microplastic particles of five sizes (100 nm, 500 nm,
   1 μm, 5 μm, and 10 μm) were purchased from ZhiChuan Technology
   (Jiangsu, China). The morphology and dispersion of particles were
   examined using scanning SEM at both low and high magnifications.
   Particle size distributions were analyzed with a laser particle size
   analyzer. The chemical composition of all PS microplastics was
   confirmed using FTIR.

Uptake of fluorescently labeled PS microplastics of different sizes by normal
Colon epithelial cells

   NCM460 cells were seeded into 24-well plates and cultured for 24 h to
   ensure stable growth. Red fluorescent polystyrene (PS) microplastics of
   various sizes were added at 1 mg/mL and co-incubated with the cells for
   another 24 h. After incubation, wells were rinsed with PBS to remove
   non-internalized particles. Red fluorescence and DAPI-stained nuclei
   were visualized using an Olympus inverted fluorescence microscope.

Histopathological staining of tissues

   Colon inflammation was evaluated by histopathological analysis. Tissues
   were fixed in 4% paraformaldehyde, dehydrated with graded ethanol,
   embedded in paraffin, and sectioned at 4 μm thickness. Hematoxylin and
   eosin (H&E) and Alcian Blue–Periodic Acid–Schiff (AB-PAS) staining were
   performed using commercial kits (Solarbio, Beijing, China). Stained
   sections were imaged using a Leica DM6B upright microscope.

Detection of inflammatory cytokines by ELISA

   Inflammatory cytokines (TNF-α, IL-1β, IL-6, and IL-10) in colon tissues
   were quantified using ELISA kits (Meibo Biotechnology, Jiangsu, China)
   following the manufacturer’s instructions. Optical density was measured
   at 450 nm using a BioTek microplate reader, and cytokine concentrations
   were calculated from standard curves.

Cell viability assay

   NCM460 cells were plated in 96-well plates and incubated for 24 h to
   promote adhesion. The medium was then replaced with RPMI 1640
   containing polystyrene (PS) microplastics of different sizes and
   concentrations. After 24 h of exposure, CCK-8 reagent was added and
   incubated for 2 h. Absorbance at 450 nm was recorded to assess cell
   viability.

Apoptosis detection

   Cells were harvested using EDTA-free trypsin, washed with PBS, and
   centrifuged at 800 rpm for 5 min. Pellets were resuspended in 500 µL
   binding buffer and transferred to flow tubes. Annexin V-FITC and PI (5
   µL each) were added and gently mixed. After 10 min of dark incubation
   at room temperature, samples were analyzed within 1 h on a Beckman
   Coulter flow cytometer. FITC and PI signals were collected via FL1 and
   FL3, and data were processed using FlowJo.

Live/dead staining assay

   NCM460 cells were seeded into 24-well plates and exposed to PS
   microplastics of various sizes. Calcein AM/PI staining was prepared per
   instructions. After removing the medium, cells were washed with PBS and
   incubated with 250 µL stain at 37 °C in the dark for 30 min. Green
   (Ex/Em = 494/517 nm) and red (Ex/Em = 535/617 nm) fluorescence were
   observed under a microscope.

Proteomics sequencing

   NCM460 cells were lysed in protease inhibitor buffer and disrupted by
   sonication on ice. Following centrifugation at 12,000 rpm (4 °C,
   15 min), the supernatant was collected and protein concentration was
   measured using a BCA assay (Biyuntian, Shanghai). Approximately 100 µg
   of protein was reduced with DTT, alkylated with IAM, and separated via
   SDS-PAGE. Protein bands of interest were excised and digested with
   trypsin. Peptides were desalted using C18 columns, eluted with 0.1%
   formic acid, and dried under nitrogen gas.

   Samples were analyzed using an Easy-nLC 1200 system coupled to a Q
   Exactive HF mass spectrometer (Thermo Scientific). Peptides were
   reconstituted in 0.1% formic acid and loaded onto a C18 column (150 mm
   × 75 μm, 3 μm) for gradient elution with 0.1% formic acid and 80%
   acetonitrile at 300 nL/min. Mass spectra were acquired in positive ion
   mode across 350–1800 m/z.

   Proteomic data were processed using Proteome Discoverer v2.4 with the
   UniProt database. Differential protein expression was quantified using
   label-free or TMT-based strategies. Functional enrichment (GO and KEGG)
   was performed to explore pathways involved in oxidative stress,
   metabolism, and cell regulation.

Western blotting

   Cells were lysed on ice in buffer containing PMSF and extraction
   reagent. Protein levels were determined using a BCA kit. Equal amounts
   were resolved by 10% SDS-PAGE and transferred onto PVDF membranes.
   After blocking with 5% non-fat milk for 1 h at room temperature,
   membranes were incubated overnight at 4 °C with primary antibodies
   targeting Fosl1, P53, Slc7a11, Gpx4, YAP, and GAPDH. Following TBST
   washes, HRP-labeled secondary antibodies were applied for 1 h at room
   temperature. Signals were detected via enhanced chemiluminescence using
   a Tanon imaging system (Shanghai, China).

Mitochondrial membrane potential detection

   JC-1 staining was prepared per instructions. For positive control,
   cells were treated with 10 µM CCCP for 20 min. After removing the
   medium, cells in 24-well plates were rinsed with PBS and incubated with
   1 mL complete medium plus 1 mL JC-1 solution at 37 °C for 20 min.
   Meanwhile, 1× JC-1 buffer was prepared on ice.

   Staining solution was removed, and cells were washed twice with cold
   buffer. Fresh medium was added, and fluorescence was observed under an
   Olympus inverted microscope. Red fluorescence (Ex/Em = 525/590 nm)
   indicated normal mitochondrial potential; green (Ex/Em = 490/530 nm)
   reflected depolarization.

Cell ROS detection

   DCF-DA was diluted 1:1000 in serum-free medium to a final concentration
   of 10 µM, following the manufacturer’s instructions. After removing the
   original medium, cells were incubated with DCF-DA at 37 °C for 20 min.
   Excess dye was removed by washing three times with serum-free medium.
   Rosup was used as a positive control to induce oxidative stress.
   Fluorescence was observed using an Olympus inverted microscope
   (Ex/Em = 488/525 nm).

Tissue Immunofluorescence (IF)

   Paraffin-embedded mouse colon tissues were cut into 3 μm sections and
   deparaffinized with distilled water. Endogenous peroxidase activity was
   blocked using AR1108 (Boster, Wuhan) for 10 min at room temperature,
   followed by PBS rinses. Antigen retrieval was carried out via microwave
   heating. Sections were then incubated in 5% BSA (Biosharp, Hefei) for
   2 h at room temperature. Primary antibodies—CD11b (1:200, Huabio), CD8
   (1:200, Proteintech), and MHC II (1:200, Huabio)—were applied and
   incubated overnight at 4 °C. After returning to room temperature,
   sections were washed with PBS and gently blotted. FITC-labeled
   secondary antibodies (BL033A, Biosharp) were added and incubated for
   2 h in the dark. Following PBS washes, nuclei were counterstained with
   DAPI in anti-fade mounting medium (P0131, Beyotime). Fluorescence
   images were acquired using a Leica upright fluorescence microscope.

CD4/CD8 detection

   CD4⁺ and CD8⁺ T cell subsets were assessed by flow cytometry. Spleens
   were collected and mechanically dissociated to isolate lymphocytes,
   followed by red blood cell lysis using a commercial buffer. Cells were
   washed with PBS and transferred into flow tubes. Samples were incubated
   at 37 °C for 4 h with fluorophore-labeled antibodies: PB450-CD45,
   FITC-CD4, APC-CD8, and PerCP/Cyanine5.5-CD3. After staining, cells were
   washed and resuspended in staining buffer. Flow cytometric analysis was
   conducted using a Beckman Coulter system, and data were analyzed with
   FlowJo software.

DC cell detection

   Dendritic cell (DC) subsets were analyzed by flow cytometry. Splenic
   lymphocytes were isolated, washed with PBS, and transferred into flow
   tubes. Cells were incubated at 37 °C for 4 h with fluorophore-labeled
   antibodies: PB450-CD45, PE-CD11c, FITC-MHC II, APC-CD80, and
   PE/Cyanine7-CD86. Following staining, cells were washed and resuspended
   in staining buffer. Flow analysis was performed on a Beckman Coulter
   cytometer, and data were processed using FlowJo.

MDA (Malondialdehyde) content detection

   After cell harvesting and centrifugation, 1 mL of extraction buffer was
   added, and samples were sonicated. Lysates were centrifuged at 8000 g,
   4 °C for 10 min, and the supernatant was collected and kept on ice. MDA
   working solution was prepared per the manufacturer’s protocol. A 200 µL
   aliquot of supernatant was transferred to a microcuvette or 96-well
   plate. Absorbance at 532 nm and 600 nm was measured using a
   spectrophotometer. ΔA values were calculated by subtracting blank
   readings from sample absorbance. MDA levels were determined using the
   standard formula.

Cellular glutathione (GSH) detection

   NCM460 cells were seeded in 96-well plates and incubated for 24 h.
   After medium removal, PS microplastics of different sizes diluted in
   RPMI-1640 were added and incubated for another 24 h. Cells were rinsed
   once with PBS to eliminate residual particles. Then, 150 µL of
   glutathione detection reagent was added per well. Absorbance at 412 nm
   was measured every 5 min over a 25 min period using a microplate
   reader. Glutathione concentration was determined based on a standard
   curve.

Dual-luciferase reporter assay

   Culture cells in good growth condition, and plate them into a 24-well
   culture plate the day before plasmid transfection. On the day of
   transfection, carry out plasmid transfection according to the
   experimental design. After 24 h of transfection, observe the expression
   of the fluorescently labeled gene in the cells using a fluorescence
   microscope. Then, treat the cells with the Dual-Luciferase Reporter
   Assay Kit (RG027, Biyuntian) and perform luciferase expression
   detection.

ChIP-qPCR (Chromatin Immunoprecipitation and Quantitative PCR)

   First, add adherent cells and suspension cells to culture medium with
   1% formaldehyde and PBS solution respectively for cross-linking,
   incubating at room temperature for 10 min, and then add glycine to
   terminate the cross-linking. For adherent cells, wash with pre-chilled
   PBS and scrape the cells; for suspension cells, centrifuge to collect
   them. After adding lysis buffer, incubate the cells on ice and sonicate
   them to break up the chromatin. Next, mix the lysate with Protein A/G
   magnetic beads, and incubate for 3 h. Use a magnetic rack to separate
   and wash the beads. Then, elute the immunocomplexes from the beads with
   elution buffer, and perform cross-link reversal in a 65 °C water bath,
   adding RNase A and Proteinase K for further treatment. Finally, purify
   the target DNA through DNA extraction and perform qPCR to
   quantitatively analyze the Ct values and compare differences across
   groups.

Molecular Docking

   Molecular docking was performed to investigate the potential
   interaction between Fosl1 (UniProt ID: [91]P15407) and DNA-bound TP53
   (Gene ID: 7157). Protein and nucleic acid docking was conducted using
   AlphaFold3 ([92]https://alphafoldserver.com). Among the predicted
   complexes, the structure with the highest ranking score was selected
   for downstream analysis.

   Protein preparation was performed in PyMOL v2.4, including removal of
   water molecules and non-essential ligands, followed by hydrogen atom
   addition for structure stabilization. The resulting protein–DNA docking
   models were visualized using PyMOL to analyze spatial conformations and
   potential binding interactions between Fosl1 and TP53.

Energy metabolism sequencing

   A 50 mg (± 2.5 mg) sample was extracted with 500 µL of pre-chilled 70%
   methanol (− 20 °C), vortexed for 3 min, and centrifuged at 12,000 rpm,
   4 °C for 10 min. After a second centrifugation to remove proteins, 200
   µL of the supernatant was filtered and stored at − 20 °C for analysis.
   Metabolite profiling was performed using UHPLC–MS/MS on a Waters
   ACQUITY H-Class system with a BEH Amide column (1.7 μm, 100 × 2.1 mm).
   The mobile phases consisted of 10 mM ammonium acetate with 0.3% ammonia
   (A) and 90% acetonitrile (B). Separation was achieved with a 0.40
   mL/min flow rate at 40 °C and a 2 µL injection volume under the
   following gradient: 0–1.2 min (5:95), 8 min (30:70), 9–11 min (50:50),
   11.1–15 min (5:95). MS analysis was carried out on a QTRAP^®
   6500 + equipped with an ESI source at 550 °C (+ 5500 V/−4500 V; curtain
   gas: 35 psi) in MRM mode using optimized acquisition settings.
   Metabolites were identified via spectral comparison with the MWDB and
   quantified based on MRM peak areas using external calibration curves.

Atomic force microscopy (AFM)

   Atomic force microscopy (AFM) measurements were performed using a
   NT-AIST AFM system (HORIBA, Japan) operating in force mapping mode,
   equipped with high-precision pyramidal probes (MikroMasch, USA). The
   cantilevers used had a nominal spring constant of 0.5 N/m. After each
   measurement session, the AFM tip was cleaned with ethanol and
   sterilized under UV light to remove potential contaminants. Both
   surface topography and mechanical property (Young’s modulus) maps were
   acquired to characterize the biological samples. Topographical scans
   were conducted over a 30 μm × 30 μm area, while force mapping for
   Young’s modulus was performed over a 10 μm × 10 μm region, during which
   400 force–indentation curves were collected and analyzed for modulus
   calculation.

Actin-Tracker Red-Rhodamine

   Cells were washed with PBS and fixed in 3.7% formaldehyde for 20 min at
   room temperature. After fixation, they were rinsed three times with
   0.1% Triton X-100 in PBS (5 min each). Actin-Tracker Red was diluted
   1:40–1:200 in PBS containing 1–5% BSA and 0.1% Triton X-100. A 200 µL
   volume of staining solution was applied per slide and incubated in the
   dark for 30–60 min at room temperature. Slides were then washed three
   times with PBS/Triton X-100 at 5-min intervals. Fluorescence was imaged
   using a Leica upright microscope.

Cell Immunofluorescence

   NCM460 cells were cultured on sterilized coverslips in 12-well plates
   and incubated for 12 h to allow attachment. After medium removal, cells
   were washed with PBS and fixed in 4% paraformaldehyde for 10 min at
   room temperature, followed by three PBS rinses. Permeabilization was
   performed using 0.2% Triton X-100 in PBS for 10 min, then washed again.
   Blocking was done with 5% BSA in PBS for 30 min at room temperature.
   Cells were incubated with primary antibodies for 1 h in the dark,
   followed by three 5-min PBS washes. Nuclei were stained with DAPI
   (1:1000 in PBS) for 1 min in the dark and rinsed. Coverslips were
   mounted with 90% glycerol, and fluorescence was imaged using a Leica
   upright microscope.

Cell lactate content detection

   To quantify intracellular lactate, 200 µL of PBS was added to each cell
   sample and homogenized. Lysates were centrifuged at 10,000 × g, 4 °C
   for 10 min, and supernatants were collected and kept on ice. Part of
   the supernatant was used for protein measurement to normalize lactate
   levels. Lactate assay reagents and standards were prepared following
   the manufacturer’s protocol. Standards were serially diluted from 0 to
   7 mmol/L. For detection, 5 µL of each standard or sample was added to a
   96-well plate, followed by 100 µL of enzyme solution and 20 µL of
   reagent III. After 10 min incubation at 37 °C, 180 µL of reagent IV was
   added, and the plate was shaken for 5 s. Absorbance at 530 nm was
   recorded using a microplate reader, and lactate levels were determined
   from the standard curve.

ATP content detection

   Cells were lysed on ice or at 4 °C using 200 µL of lysis buffer per
   well (6-well plates). Lysates were pipetted to mix and centrifuged at
   12,000 × g, 4 °C for 5 min. Supernatants were collected for ATP
   analysis. ATP standards ranging from 0.01 to 10 µM were prepared for
   calibration. The assay reagent was diluted 1:9 to generate the working
   solution. A total of 100 µL of working solution was added to each well
   or tube and incubated at room temperature for 5 min. Then, 20 µL of
   sample or standard was added, gently mixed, and incubated for at least
   2 s. Luminescence was detected using a chemiluminescence reader, and
   ATP concentrations were calculated from RLU values using the standard
   curve.

Extracellular acidification rate (ECAR) detection

   To evaluate ECAR, cells were trypsinized, neutralized with
   serum-containing medium, washed with saline, and pelleted by
   centrifugation. After removing the supernatant, cells were resuspended
   in Reagent 1 as instructed. A 100 µL cell suspension was dispensed into
   each well of a black 96-well plate; blank wells received 100 µL of
   Reagent 1. Plates were incubated at 37 °C in the dark for 30 min.
   Subsequently, 100 µL of ECAR working solution was added. Fluorescence
   (Ex/Em = 490/535 nm) was recorded every 2–5 min for 100–120 min using a
   microplate reader. ECAR was determined by calculating the fluorescence
   change rate (ΔF/ΔT) over time.

Statistical analysis

   Data are presented as mean ± SEM. Statistical analysis was performed
   using GraphPad Prism 10.1. One-way ANOVA followed by Tukey’s post hoc
   test was used for group comparisons. p < 0.05 was considered
   statistically significant.

Results

Characterization of PS microplastics and uptake of microplastics of different
sizes by intestinal epithelial cells

   As characterized by SEM and FTIR, the PS microplastics used in this
   study were spherical and chemically consistent (Fig. [93]1A–C).

Fig. 1.

   [94]Fig. 1
   [95]Open in a new tab

   Characterization of PS microplastics and uptake of microplastics of
   different sizes by intestinal epithelial cells. A SEM images of five
   different diameters of PS microplastics at low and high magnifications.
   Scale bar = 2 μm. (B) Laser particle size analysis of the average sizes
   and distribution of the five different sizes of PS microplastics. (C)
   FTIR spectrum analysis of PS microplastics. (D) Co-culturing five
   different sizes of red fluorescent PS microplastics with NCM460 cells
   for 24 h, observing the uptake of PS microplastics by cells. Images
   show DAPI channel, red fluorescence channel, and merged images from top
   to bottom. Scale bar = 100 μm. (E) Co-culturing five different sizes of
   PS microplastics with NCM460 cells for 24 h, observing the uptake of PS
   microplastics by cells using TEM images at low and high magnifications.
   Scale bar = 2 μm

   To examine whether microplastics of different sizes could be
   internalized by human intestinal epithelial cells (NCM460), we cultured
   NCM460 cells in vitro. The results indicated that microplastics sized
   100 nm and 500 nm were readily engulfed by NCM460 cells and accumulated
   in large quantities within the cells. In contrast, 1 μm and 5 μm
   microplastics were minimally internalized, and 10 μm microplastics were
   not taken up by the cells at all (Fig. [96]1D). To further confirm
   these observations, we employed transmission electron microscopy (TEM)
   to observe the uptake of microplastics by the cells. TEM imaging
   revealed that NCM460 cells ingested a significant number of 100 nm and
   500 nm microplastic particles, while only a few 1 μm and 5 μm
   microplastic particles were internalized, and no internalization of
   10 μm particles was observed (Fig. [97]1E).

   Additionally, we examined the effects of microplastic uptake on
   cellular integrity. In the 100 nm, 500 nm, and 1 μm experimental
   groups, the continuity and integrity of the cell membranes remained
   largely intact, although some damage to the mitochondria was noted. In
   contrast, in the 5 μm and 10 μm experimental groups, the integrity of
   the cell membranes was compromised, while mitochondrial morphology
   remained relatively normal, suggesting that mechanical damage to the
   cells occurred to a certain extent.

Long-term exposure to PS microplastics induces Colon inflammation in mice

   After a one-week acclimation, BALB/c mice were randomly divided into
   six groups: one control and five polystyrene microplastic (PS-MP)
   exposure groups. The treated groups received daily oral gavage of
   1 mg/mL PS-MP suspension, while controls received equal volumes of PBS.
   After seven weeks of treatment and care, mice were sacrificed, and
   colon tissues, serum, and fecal samples were collected for ELISA, H&E
   staining, and other biochemical assays (Fig. [98]2A).

Fig. 2.

   [99]Fig. 2
   [100]Open in a new tab

   Long-term exposure to PS microplastics induces colon inflammation in
   mice. A Schematic diagram of the treatment process for microplastic and
   control groups. (B) DAI scores during the gavage period for the
   microplastic and control groups. (C) Changes in body weight gain curves
   for the microplastic and control groups during the gavage period. (D)
   Comparison of colon weight between control and microplastic-treated
   groups. (E) Comparison of colon length between control and
   microplastic-treated groups. (F) Anatomical diagram of the intestines
   and H&E staining of colon tissue in the microplastic and control
   groups. (G) Expression levels Four inflammatory factors in the
   microplastic and control groups (ELISA). *p < 0.05, **p < 0.01,
   ***p < 0.001, ****p < 0.0001

   To evaluate colon inflammation, mice were dynamically monitored using a
   simplified Disease Activity Index (DAI) scoring system (Fig. [101]2B)
   while body weight changes were recorded (Fig. [102]2C, Additional file
   1: Fig. [103]S1). At baseline, no significant differences in body
   weight were observed between the microplastic-exposed and control
   groups (P > 0.05). However, during the gavage period, mice exposed to
   microplastics exhibited a slower rate of weight gain compared to
   controls.By the end of the study, mice exposed to microplastics showed
   a significant reduction in body weight compared to controls (P < 0.05).

   Further analysis of colon weight and length revealed a significant
   reduction in the microplastic-exposed groups compared to controls, with
   the most pronounced effect observed in the 100 nm group (P < 0.05,
   Fig. [104]2D and E, Additional file 2: Fig. [105]S2). Additionally, H&E
   staining of colon tissue sections indicated marked inflammatory cell
   infiltration across all microplastic-treated groups (Fig. [106]2F).
   ELISA analysis further demonstrated a significant increase in
   pro-inflammatory cytokines (TNF-α, IL-1β, and IL-6) alongside a notable
   reduction in the anti-inflammatory cytokine IL-10 (P < 0.05,
   Fig. [107]2G).

   These results indicate that long-term exposure to PS microplastics
   impairs body weight gain and induces colon inflammation in mice.

Microplastics induce Colon epithelial cell death

   The previous results suggested that long-term exposure to PS
   microplastics induces colon inflammation in mice. To further
   investigate the mechanisms underlying cellular damage caused by
   microplastics, we selected the NCM460 cell line for in vitro analysis.
   First, to determine the appropriate concentration of PS microplastics
   for the experiment, we performed CCK-8 cell proliferation and toxicity
   assays. As shown in Fig. [108]3A, treatment with PS microplastics of
   varying sizes exhibited concentration-dependent cytotoxicity. At a
   concentration of 1 mg/mL, PS microplastics caused mild damage to NCM460
   cells, with similar cell viability across all particle size groups.
   Specifically, cell viability was 84%, 80.8%, 78.2%, 77.4%, and 70% for
   the 100 nm, 500 nm, 1 μm, 5 μm, and 10 μm groups, respectively.
   However, at higher concentrations, all groups exhibited significant
   cell damage, with a noticeable decrease in cell viability.

Fig. 3.

   [109]Fig. 3
   [110]Open in a new tab

   Microplastics induce colonic epithelial cell death. (A) CCK-8 cell
   proliferation and toxicity assay. (B) Annexin V-FITC flow cytometry
   apoptosis detection. (C) Calcein-AM/PI live-dead cell fluorescence
   staining. Scale bar = 100 μm. (D) Model diagram of NCM460 cells
   ingesting PS microplastics of different sizes. *p < 0.05, **p < 0.01,
   ***p < 0.001, ****p < 0.0001

   To avoid excessive cell death caused by high concentrations of PS
   microplastics and to better explore the size-dependent effects of
   microplastics on cellular damage, we selected 1 mg/mL as the baseline
   concentration for subsequent experiments. Thus, in the following cell
   experiments, the Control group served as the control, and the PS
   microplastic concentration in all size groups was 1 mg/mL.

   Figure [111]3B presents the apoptosis analysis, indicating an apoptosis
   rate of 0.42% in the control group, compared to 8.42% (100 nm group),
   6.14% (500 nm group), 5.86% (1 μm group), 6.41% (5 μm group), and
   14.25% (10 μm group). Figure [112]3C illustrates Calcein-AM/PI
   live-dead cell fluorescence staining, where live cells exhibited green
   fluorescence (Calcein-AM), while dead cells displayed red fluorescence
   (PI). The red-to-green fluorescence ratio served as an indicator of
   cell damage. In the control group, minimal red fluorescence was
   observed, indicating normal cell viability. In contrast, microplastic
   exposed groups exhibited increased red fluorescence, suggesting varying
   degrees of cytotoxicity. Notably, the 100 nm and 10 μm groups showed a
   more pronounced increase in red fluorescence compared to the 500 nm,
   1 μm, and 5 μm groups, indicating that cell damage was most severe in
   the 100 nm and 10 μm groups. Furthermore, the substantial size
   difference between 100 nm and 10 μm polystyrene microplastics may
   contribute to their distinct biological effects.

   Based on these results, we observed that PS microplastics smaller than
   1 μm were more readily taken up by NCM460 cells, whereas PS
   microplastics larger than 1 μm were less efficiently internalized. The
   most notable effects were observed with the 100 nm and 10 μm PS
   microplastics. Therefore, we selected 100 nm and 10 μm PS microplastics
   as representative particle sizes for further investigation into their
   potential mechanisms of cellular damage (Fig. [113]3D).

Proteomics reveals mechanistic differences in Colon inflammation induced by
different sizes of PS microplastics

   To further explore the mechanisms by which PS microplastics of
   different sizes damage intestinal epithelial cells, we selected 100 nm
   and 10 μm particles as representative sizes. These particles were
   co-cultured with NCM460 cells for 24 h, after which proteomic
   sequencing analysis was conducted (Fig. [114]4A-H). The results
   revealed significant differences in gene expression between the NP
   group (100 nm PS microplastics) and the MP group (10 μm PS
   microplastics) (Fig. [115]4A). Venn diagram analysis (Fig. [116]4B)
   showed that 29 differentially expressed genes were common to both
   groups, while the NP100 and MP10 groups had 62 and 389 unique
   differentially expressed genes, respectively. This suggests that PS
   microplastics of different sizes may induce cellular damage through
   distinct molecular mechanisms.

Fig. 4.

   [117]Fig. 4
   [118]Open in a new tab

   Proteomics reveals mechanistic differences in colon inflammation
   induced by different sizes of PS microplastics. (A) Volcano plots
   illustrating differential gene expression in NP10 vs. NC, NP100 vs.
   NP10, and MP10 vs. NC comparisons. Red and blue dots indicate
   upregulated and downregulated genes, respectively. Significantly
   changed genes are annotated. (B) Venn diagram showed the overlap of
   three groups of differential genes: NP_10 vs. NC, NP_100 vs. NP_10,
   MP_10 vs. NC. (C)Heatmap displaying expression profiles of DEGs in
   NP100, MP10, and NC groups. Color gradients reflect expression levels
   (red: high, blue: low). Hierarchical clustering revealed group-specific
   gene expression patterns. (D) KEGG pathway enrichment analysis. The
   KEGG enrichment bubble map showed the signaling pathway of significant
   enrichment of differential genes. (E) Reactome pathway enrichment
   analysis. Reactome bubble map showed the enrichment of differential
   genes in the Reactome pathway. (F) WikiPathways enrichment analysis.
   Bubble plot illustrates significant enrichment of DEGs across
   biological pathways. The X-axis shows the Rich Factor (DEG ratio per
   pathway), Y-axis displays pathway names. Bubble size denotes gene
   count; color indicates p-value.(G) Gene function network analysis,
   where the network maps showed significant associations between
   enrichment functions and pathways. Nodes represent enrichment
   functions, and lines indicate the degree of similarity or gene sharing
   between functions or pathways. (H) Summary bar chart of significant
   function, which summarizes the functional and biological processes of
   significant enrichment

   Heatmap analysis (Fig. [119]4C) identified the Fosl1 gene, which is
   associated with ferroptosis, as significantly upregulated in the NP100
   group. Other genes related to cellular stress and metabolism, such as
   HSPB1 and MAPKAPK5, were also elevated, indicating that smaller-sized
   microplastics may induce oxidative stress and inflammatory responses.
   In contrast, the MP 10 group exhibited significant changes in
   metabolism-related genes, such as a marked increase in the expression
   of GPAT3 and FGD3. Additionally, MP 10 group showed alterations in
   genes related to mitochondrial function and energy metabolism,
   including COX15 and MPC2, suggesting a link to energy metabolism
   remodeling and cell structural damage in intestinal epithelial cells.

   KEGG pathway enrichment analysis further elucidated the potential
   mechanisms of action for the two sizes of PS microplastics
   (Fig. [120]4D). Differential genes in the NP group were predominantly
   enriched in ferroptosis and P53 signaling pathways, indicating that
   smaller-sized PS microplastics may induce ferroptosis through lipid
   peroxidation. In contrast, differential genes in the MP group were
   notably enriched in metabolic pathways, such as the TCA cycle and
   oxidative phosphorylation, suggesting that larger-sized PS
   microplastics may induce cell death by modulating glycolysis and
   metabolic remodeling.

   Reactome pathway analysis (Fig. [121]4E) supported these findings,
   showing that genes in the NP100 group were significantly enriched in
   programmed cell death and P53 regulation pathways, while those in the
   MP10 group were concentrated in fatty acid metabolism and mitochondrial
   function pathways. WikiPathways analysis (Fig. [122]4F) further
   highlighted the association of the NP100 group with ferroptosis, while
   the MP10 group’s genes were enriched in inflammatory response and
   glycolysis pathways. Gene functional enrichment network analysis
   (Fig. [123]4G) indicated that smaller-sized PS microplastics are
   involved in ferroptosis and P53 signaling, whereas larger-sized PS
   microplastics mediate cell damage through fatty acid metabolism and
   glycolysis abnormalities following mechanical damage.

   Finally, the functional enrichment bar chart (Fig. [124]4H) quantified
   the enrichment levels of significant pathways, underscoring the close
   association of the NP100 group with ferroptosis and P53 signaling
   regulation, as well as the significant enrichment of the MP10 group in
   glycolysis, fatty acid metabolism, and extracellular matrix binding.
   These results suggest that smaller-sized PS microplastics may cause
   cellular damage through lipid peroxidation and ferroptosis, while
   larger-sized PS microplastics likely affect cell function through
   mechanical damage and metabolic remodeling.

Small-sized NP-mediated in vitro ferroptosis

   The proteomic analysis suggested that small-sized PS nanoplastics may
   primarily induce ferroptosis through lipid peroxidation, while
   larger-sized PS microplastics are likely to induce cell apoptosis due
   to metabolic alterations. To validate that nanoplastics can induce
   ferroptosis in vitro, we assessed the expression of ferroptosis-related
   protein markers under treatment with different sizes of microplastics
   (Control, 100 nm, 500 nm, 1 μm, 5 μm, 10 μm). Western blot analysis
   showed that 100 nm and 500 nm PS microplastics significantly
   downregulated the expression of ferroptosis-related proteins SLc7a11
   and Gpx4, while the 1 μm group exhibited a non-significant downward
   trend. In contrast, no significant changes were observed in the
   larger-sized microplastic groups (5 μm and 10 μm) (Fig. [125]5A).

Fig. 5.

   [126]Fig. 5
   [127]Open in a new tab

   NPs mediated ferroptosis in vitro. (A) Western blotting analysis of
   ferroptosis-related proteins. (B) Reactive oxygen species (ROS) assay.
   Scale bar = 200 μm. (C) JC-1 mitochondrial membrane potential assay.
   Scale bar = 200 μm.*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

   JC-1 staining was conducted to evaluate mitochondrial membrane
   potential (MMP) alterations, serving as an indicator of oxidative
   stress and ferroptosis-associated mitochondrial dysfunction. As shown
   in Fig. [128]5B, the Control group predominantly exhibited red
   fluorescence, with minimal green fluorescence, indicative of intact
   mitochondrial function. In contrast, the CCCP-positive control group,
   known to induce mitochondrial dysfunction, displayed a predominance of
   green fluorescence with minimal red fluorescence, confirming MMP loss.
   In the 100 nm, 500 nm, and 1 μm groups, a decrease in red fluorescence
   and a corresponding increase in green fluorescence were observed
   compared to the Control group, suggesting mitochondrial membrane
   potential disruption. This effect was less pronounced in the 5 μm and
   10 μm groups, further indicating that smaller-sized PS microplastics
   induce greater mitochondrial dysfunction.

   Additionally, we assessed reactive oxygen species (ROS) levels using a
   ROS assay, where the intensity of green fluorescence corresponds to ROS
   levels within the cells. The results in Fig. [129]5C showed that the
   Control group exhibited minimal green fluorescence, while the ROS
   inducer (Rosup) positive control group displayed abundant green
   fluorescence. Among the microplastic-treated groups, the 100 nm,
   500 nm, and 1 μm groups exhibited strong green fluorescence, indicating
   significantly elevated ROS levels. Conversely, the 5 μm and 10 μm
   groups showed only a slight increase in fluorescence compared to the
   Control group, suggesting that ROS levels were substantially elevated
   only in the small-sized PS microplastic-treated groups.

Ferrostatin-1 alleviates colon inflammation induced by NPs in vivo

   In vitro experiments suggested that NPs can induce ferroptosis,
   contributing to colon inflammation. However, it remained unclear
   whether this mechanism occurs in vivo and whether ferroptosis
   inhibition could mitigate NP-induced intestinal damage. Ferrostatin-1
   (Fer-1), a well-established ferroptosis inhibitor, is known to suppress
   lipid peroxidation and oxidative stress. To test its protective effects
   in vivo, Fer-1 was administered to mice following NP exposure.

   After one week of acclimatization, BALB/c mice were randomly assigned
   to five groups: a control group, two NP exposure groups (100 nm and
   10 μm), and two NP exposure groups receiving intraperitoneal Fer-1.
   Mice received daily oral gavage of 1 mg/mL PS microplastics, with or
   without Fer-1, for seven weeks. Colon tissues, serum, and fecal samples
   were collected for ELISA, H&E staining, and multiplex
   immunofluorescence (Fig. [130]6A).

Fig. 6.

   [131]Fig. 6
   [132]Open in a new tab

   Ferrostatin-1 alleviates colon inflammation induced by NPs in vivo. (A)
   Diagram of the treatment process for microplastic and control groups of
   mice. (B) DAI scores during the gavage process in the microplastic and
   control groups. (C) Body weight growth curves of microplastic and
   control group mice during gavage. (D) Comparison of colon weight in
   microplastic and control groups. (E) Comparison of colon length in
   microplastic and control groups. (F) Expression levels of Four
   inflammatory factors in microplastic and control groups (ELISA). (G)
   Anatomical diagram and H&E staining of colonic tissue in microplastic
   and control groups. (H) Multiplex immunofluorescence. (I) Flow
   cytometry analysis of CD4/CD8 ratio and DC cells. *p < 0.05,
   **p < 0.01, ***p < 0.001, ****p < 0.0001

   NP exposure significantly reduced body weight, colon length and weight,
   and increased DAI scores and pro-inflammatory markers, indicating
   intestinal inflammation. In contrast, Fer-1 treatment effectively
   improved these parameters (Figs. [133]6B–F, Additional file3:
   Fig.[134]S3,Additional file4: Fig.[135]S4 ), suggesting mitigation of
   NP-induced injury.

   Histopathological analysis (Fig. [136]6G) revealed extensive
   inflammatory infiltration in NP-treated mice, which was notably
   alleviated by Fer-1. However, Fer-1 had little effect on large-sized
   MP-induced inflammation, indicating that ferroptosis may not be
   involved in the damage caused by larger particles.

   To investigate ferroptosis-mediated immune activation, we analyzed
   colon tissues via multiplex immunofluorescence and flow cytometry.
   NP-treated mice showed increased infiltration of dendritic cells (DCs)
   and CD8⁺ T cells, alongside a reduced CD4/CD8 ratio and elevated DC
   (CD80⁺CD86⁺) counts—hallmarks of excessive immune activation. Fer-1
   reversed these changes (Figs. [137]6H–I), indicating that ferroptosis
   contributes to immunogenic cell death (ICD) in the colon.

   Together, these findings demonstrate that small-sized NPs induce
   intestinal inflammation through ferroptosis-mediated ICD, and that
   inhibition of ferroptosis by Fer-1 can effectively alleviate this
   damage.

Fosl1 regulates P53 transcription to promote nano-plastics-Induced
ferroptosis

   Proteomic analysis revealed significant gene expression changes in the
   NP-treated group, particularly upregulation of Fosl1, Cox-15, and CCN1.
   Functional enrichment analysis showed that these differentially
   expressed proteins were enriched in the ferroptosis and p53 signaling
   pathways. Among them, Fosl1—a member of the FOS transcription factor
   family and a core component of the AP-1 complex—was identified as a key
   regulatory molecule potentially linking NP exposure to ferroptosis.
   Given its involvement in cell proliferation, apoptosis, and stress
   responses, Fosl1 was selected for further investigation.

   To assess its role in ferroptosis, we conducted in vitro experiments
   with five treatment groups: Control, 100 nm, 100 nm + siFosl1, 10 μm,
   and 10 μm + siFosl1. Western blotting (Fig. [138]7A, Additional
   file5:Fig.S5) confirmed upregulation of Fosl1 in the 100 nm NP group,
   along with downregulation of Slc7a11 and Gpx4 upon Fosl1 knockdown—an
   effect not observed in the 10 μm group.

Fig. 7.

   [139]Fig. 7
   [140]Open in a new tab

   Fosl1 regulates P53 transcription to promote nano-plastics-induced
   ferroptosis. (A) Western blot detection of ferroptosis-related
   proteins. (B) TEM images of mitochondrial changes in NCM460 cells in
   the Control, 100 nm, and 100 nm + siFOSL1 groups at low and high
   magnifications. Scale bar = 2 μm. (C) ROS detection. Scale bar
   = 200 μm. (D) Fluorescent quantification of ROS detection. (E) MDA
   content detection. (F) GSH content detection. (G) GSSG content
   detection. (H) JC-1 mitochondrial membrane potential detection. Scale
   bar = 200 μm. (I) Quantification of JC-1 fluorescence in mitochondrial
   membrane potential detection. (J) Dual-luciferase assay. (K) Chromatin
   immunoprecipitation (ChIP)-qPCR experiment. (L) Molecular docking
   analysis. Hydrogen bonds are represented in yellow. FOSL1 and DNA-TP53
   are represented in blue and gray, respectively. *p < 0.05, **p < 0.01,
   ***p < 0.001, ****p < 0.0001

   TEM (Fig. [141]7B) revealed typical mitochondrial damage in the NP
   group, including condensed membranes, shrunken volume, disrupted outer
   membranes, and loss of cristae. Knockdown of Fosl1 mitigated these
   structural abnormalities, suggesting a protective effect on
   mitochondrial integrity.

   We further evaluated ferroptosis-related biomarkers: ROS
   (Fig. [142]7C), mitochondrial membrane potential (JC-1, Fig. [143]7H),
   malondialdehyde (MDA, Fig. [144]7E), and glutathione (GSH/GSSG,
   Fig. [145]7F and G). Compared to the 100 nm NP group, the
   100 nm + siFosl1 group showed significantly reduced ROS, restored
   mitochondrial potential, decreased MDA levels, and a shift in GSH/GSSG
   ratio—indicating reduced oxidative stress and ferroptosis. These
   changes were not seen in the 10 μm MP groups, suggesting a
   size-specific effect.

   To determine whether Fosl1 directly regulates P53 transcription, we
   conducted dual-luciferase reporter assays (Fig. [146]7J) and ChIP-qPCR
   (Fig. [147]7K). Results confirmed direct binding of Fosl1 to the P53
   promoter. Additionally, molecular docking using AlphaFold3
   (Fig. [148]7L) supported this interaction, providing a structural basis
   for Fosl1-mediated transcriptional regulation.

   Collectively, these findings demonstrate that Fosl1 promotes
   ferroptosis in NP-induced colon inflammation by enhancing P53
   transcription and suppressing Slc7a11 and Gpx4. In contrast, these
   effects were absent in the MP group, further highlighting the
   size-dependent mechanism of toxicity.

Non-targeted energy metabolomics reveals MPs induce colon inflammation via
metabolic reprogramming

   Proteomic analyses revealed that differentially expressed proteins in
   the 10 μm microplastic-treated group were predominantly enriched in
   metabolic pathways, particularly the tricarboxylic acid (TCA) cycle and
   oxidative phosphorylation. To further explore the impact of large
   polystyrene microplastics on intestinal epithelial cell metabolism, an
   energy metabolomics analysis was conducted on colon tissues from both
   the control and 10 μm microplastic-treated groups. The analysis focused
   on key metabolic pathways, including glycolysis, the TCA cycle, and
   oxidative phosphorylation, to elucidate potential metabolic
   alterations.

   Energy metabolomics analysis, as demonstrated by principal component
   analysis (PCA) (Fig. [149]8A), revealed a clear separation between the
   MP-treated group and the control group, indicating that MP exposure
   significantly altered the cellular metabolic profile. Further analysis
   using volcano plots of differential metabolites (Fig. [150]8B)
   identified notable metabolic shifts induced by MP exposure.
   Specifically, early-stage glycolytic metabolites, such as
   fructose-1,6-bisphosphate, were significantly upregulated, whereas
   late-stage glycolytic intermediates, including phosphoenolpyruvate and
   2-phospho-D-glycerate, were markedly downregulated. These findings
   suggest an increase in glycolytic flux accompanied by a metabolic
   blockade in late-stage glycolysis.

Fig. 8.

   [151]Fig. 8
   [152]Open in a new tab

   Non-targeted energy metabolomics reveals MPs induce colon inflammation
   via metabolic reprogramming. (A) Principal component analysis (PCA) of
   sample groups. Component 1 and Component 2 represent the first and
   second principal components, respectively, with percentages indicating
   the variance explained. Each point denotes a sample, and colors
   represent sample groupings. (B) Each dot represents a metabolite. Red
   and blue indicate upregulated and downregulated metabolites,
   respectively; gray denotes non-significant changes. (C) Bar plot of
   differential metabolites. The X-axis shows logFC values; the Y-axis
   lists metabolites. Red and blue bars indicate up- and downregulated
   metabolites, respectively. (D) Heatmap of metabolite expression. The
   horizontal axis shows sample names, and the vertical axis lists
   metabolites. Expression values are standardized, with red indicating
   higher and blue indicating lower expression levels. (E) KEGG pathway
   enrichment analysis based on differential metabolites. The Rich Factor
   indicates the ratio of differential to total annotated metabolites
   within each pathway; higher values reflect greater enrichment. (F)
   Violin plot illustrating data distribution. The box shows the
   interquartile range, the thin line represents the 95% confidence
   interval, the horizontal line indicates the median, and the outer
   contour reflects density

   Moreover, a significant upregulation of oxaloacetate, a key
   intermediate in the TCA cycle, indicates that cells may attempt to
   compensate for the glycolytic blockade by enhancing TCA cycle activity.
   However, the diminished late-stage glycolysis alongside the increased
   TCA cycle activity suggests a potential shift toward anaerobic
   metabolism.

   To further illustrate the changes in metabolite abundance, we presented
   the variations in key metabolites between the treatment and control
   groups using bar charts (Fig. [153]8C), heatmaps (Fig. [154]8D), and
   violin plots (Fig. [155]8F). These comparative analyses revealed that
   in the large-particle PS microplastic treatment group,
   glycerol-3-phosphate and phosphoenolpyruvate were significantly
   reduced, while fructose-1,6-bisphosphate and metabolites from the
   pentose phosphate pathway, such as D-erythrose 4-phosphate, were
   significantly elevated. These changes suggest that late-stage
   glycolytic metabolites were significantly downregulated, and cells may
   adapt to metabolic stress by enhancing early glycolysis and the pentose
   phosphate pathway.

   Overall, the trend suggests a metabolic shift from aerobic to anaerobic
   pathways, characterized by increased anaerobic metabolic activity.
   Metabolic pathway enrichment analysis (Fig. [156]8E) further
   highlighted that the metabolic disturbances induced by MPs were
   predominantly concentrated in glycolysis/gluconeogenesis, the TCA
   cycle, and the pentose phosphate pathway. These findings indicate that
   MPs exposure significantly disrupts cellular energy metabolism, driving
   cells to shift from efficient aerobic metabolism to less efficient but
   faster anaerobic metabolism.

Microplastics regulate metabolic reprogramming via YAP1-Mediated crosstalk of
cellular mechanics signal transduction and glycolysis

   To further investigate the metabolic reprogramming induced by large
   polystyrene (PS) microplastics, we integrated metabolomics and
   proteomics data and mapped the changes in glucose metabolism
   (Fig. [157]9A). The results revealed that multiple key proteins and
   metabolites involved in glucose metabolism were altered in the colon
   following 10 μm PS microplastic treatment. Specifically, the
   microplastic-treated group exhibited significant metabolic
   abnormalities in the glycolysis pathway. The early glycolytic
   metabolite fructose-1,6-bisphosphate (F1,6BP) was significantly
   upregulated, while late-stage glycolytic metabolites such as
   2-phospho-D-glycerate (2PG) and phosphoenolpyruvic acid (PEP) were
   significantly downregulated, suggesting a blockade in the later stages
   of glycolysis. Additionally, pyruvate and lactic acid levels were
   significantly elevated, indicating a shift toward enhanced anaerobic
   metabolism to compensate for increased energy demands. Meanwhile, the
   accumulation of oxaloacetate in the TCA cycle, coupled with the
   reduction of succinate and α-ketoglutarate (α-KG), suggested a decline
   in TCA cycle activity. These findings imply that oxidative
   phosphorylation pathways were likely impaired due to mitochondrial
   damage, and glucose metabolism shifted toward anaerobic glycolysis,
   resulting in decreased ATP production.

Fig. 9.

   [158]Fig. 9
   [159]Open in a new tab

   Microplastics regulate metabolic reprogramming via YAP1-mediated
   crosstalk of cellular mechanics signal transduction and glycolysis. (A)
   Metabolic pathway map. (B) Cell morphologies and moduli of Control
   group and 10 μm group photographed by Atomic Force Microscopy.
   Scale = 2 μm. (C) Low and high power images of cells in the Control
   group and 10 μm group were taken by transmission electron microscopy
   (TEM). Scale = 2 μm. (D) Cell F-Actin staining in Control group and
   10 μm group. Scale = 200 μm. (E) Annexin V-FITC flow cytometry for
   apoptosis of cells in Control group and 10 μm group. (F) Mitochondrial
   apoptosis-related proteins were detected by Western Blotting. (G)
   Immunofluorescence staining of YAP protein. Scale = 200 μm. (H) The
   cell morphology and modulus of 10 μm group and 10 μm + si-YAP group
   were photographed by Atomic Force Microscopy. Scale = 2 μm. (I) Low and
   high power images of cells in the 10 μm group and 10 μm + si-YAP group
   were taken by transmission electron microscopy (TEM). Scale = 2 μm. (J)
   F-Actin staining in 10 μm group and 10 μm + si-YAP group.
   Scale = 200 μm. (K) Annexin V-FITC flow cytometry for apoptosis of
   cells in 10 μm group and 10 μm + si-YAP group. (L) Lactic acid content
   detection (lactic acid colorimetry). (M) Extracellular acidification
   rate detection. (N) ATP content detection. *p < 0.05, **p < 0.01,
   ***p < 0.001, ****p < 0.0001

   This phenomenon indicates that MPs exposure may drive a shift in cell
   metabolism from aerobic to anaerobic, though the underlying molecular
   mechanisms remain unclear. Based on the observed changes, we
   hypothesized that alterations in glucose metabolism could be driven by
   mechanical stress at the cellular level. To test this, we validated the
   relationship between cellular mechanics and anaerobic metabolism.
   Atomic force microscopy (AFM) analysis (Fig. [160]9B) demonstrated a
   significant increase in the Young’s modulus of cells exposed to 10 μm
   microplastics compared to the Control group. Transmission electron
   microscopy (TEM) (Fig. [161]9C) further showed local deformations of
   the cell membrane and mitochondrial swelling in the 10 μm group,
   suggesting mechanical damage induced by MPs.

   To further explore the effects of mechanical stress on the
   cytoskeleton, we performed phalloidin staining (Additional
   file6:Fig.[162]S6). In the Control group, actin filaments appeared
   clear and intact, whereas in the 10 μm group, actin filaments were
   disordered, broken, and rearranged, indicating that mechanical damage
   may affect the cytoskeletal structure. We also assessed apoptosis using
   flow cytometry. As shown in Fig. [163]9D, the apoptosis rate in the
   Control group was 0.78%, whereas in the 10 μm group, it increased
   significantly to 11.2%, suggesting a marked increase in apoptosis
   following MPs exposure. Western blotting of mitochondrial
   apoptosis-related proteins (Fig. [164]9E) revealed no significant
   differences in GAPDH expression across groups. However, the expression
   levels of Bax, Caspase-9, and Caspase-3 were lower in the Control group
   and increased in the 10 μm group.

   Yes-associated protein (YAP) is a well-established regulator of cell
   mechanics and has been implicated in glycolytic metabolism. Under
   physiological conditions, YAP remains phosphorylated in the cytoplasm,
   where it interacts with inhibitory factors, maintaining an inactive
   state. However, upon dephosphorylation, YAP translocates from the
   cytoplasm to the nucleus, a process known as nuclear translocation,
   which is critical for its functional activation. Given its role in
   cellular mechanics, this study focused on YAP involvement in damage
   processes induced by large-sized microplastics. Western blot analysis
   revealed a significant upregulation of YAP expression in the 10 μm
   microplastic group (Additional file 7: Fig. [165]S7). To confirm YAP
   nuclear translocation, immunofluorescence staining was performed
   (Fig. [166]9F). In the Control group, YAP localization was
   predominantly cytoplasmic. In contrast, in the 10 μm treatment group, a
   marked increase in nuclear YAP-associated fluorescence was observed,
   indicating a significant nuclear translocation of YAP following MP
   exposure.

   To further investigate the role of YAP in microplastic-induced
   mechanical damage, we knocked down YAP in the MPs treatment group
   (Additional file7:Fig.[167]S7). AFM imaging (Fig. [168]9G) showed a
   significant reduction in the Young’s modulus of the 10 μm + si-YAP
   group compared to the 10 μm group, suggesting that microplastics
   regulate mechanotransduction through YAP. However, TEM (Fig. [169]9H)
   and phalloidin staining (Additional file6:Fig.[170]S6) indicated that
   YAP knockdown only partially alleviated the damage to cell membrane
   integrity and cytoskeletal disruption, suggesting that the primary role
   of YAP is in signaling regulation rather than directly affecting
   membrane structure. Flow cytometry revealed that the apoptosis rate in
   the 10 μm group was 12.64%, whereas in the 10 μm + si-YAP group, it
   decreased to 8.62% (Fig. [171]9I), indicating that YAP knockdown
   alleviated apoptosis. Western blotting confirmed this result
   (Fig. [172]9E), showing that apoptosis-related proteins Bax, Caspase-9,
   and Caspase-3 were downregulated in the 10 μm + si-YAP group compared
   to the 10 μm group.

   To explore the role of YAP in energy metabolism and metabolic
   reprogramming, we analyzed lactate content, extracellular acidification
   rate (ECAR), and ATP levels in the Control, 10 μm, si-YAP, and
   10 μm + si-YAP groups (Fig. [173]9J-L). The results showed that lactate
   levels and ECAR were significantly elevated in the 10 μm group,
   indicating enhanced anaerobic metabolism, while ATP levels were
   significantly reduced, suggesting mitochondrial dysfunction and a shift
   toward anaerobic glycolysis. However, in the 10 μm + si-YAP group,
   these indicators were alleviated, with lactate and ECAR levels
   decreasing, and ATP levels significantly restored. These findings
   reveal that MPs regulate mechanosignaling via YAP, contributing to
   metabolic reprogramming and leading to cell apoptosis due to low ATP
   production. Moreover, YAP regulation can reverse the metabolic changes
   and alleviate cellular damage induced by MPs exposure.

YAP inhibitor ameliorates microplastic-induced colon inflammation

   To explore the role of YAP in microplastic (MP)-induced colon
   inflammation and assess its potential as a therapeutic target, an in
   vivo experiment was conducted using the YAP-specific inhibitor VTPF.
   VTPF is a widely used YAP inhibitor that prevents YAP nuclear
   translocation by suppressing its activity, thereby disrupting
   downstream YAP-related signaling pathways. Mice were assigned to three
   experimental groups: Control, 10 μm microplastic exposure (10 μm
   group), and 10 μm microplastic exposure with VTPF treatment
   (10 μm + VTPF group). The 10 μm group received weekly oral gavage of
   microplastics, while the 10 μm + VTPF group was administered both 10 μm
   PS microplastics via gavage and intraperitoneal injections of VTPF. The
   experiment lasted for 7 weeks (Fig. [174]10A). Physiological parameters
   were continuously monitored in all groups throughout the study.

Fig. 10.

   [175]Fig. 10
   [176]Open in a new tab

   YAP inhibitor ameliorates microplastic-induced colon inflammation. (A)
   Schematic of the treatment protocol for microplastic and control
   groups. (B) DAI scores during the gavage process for microplastic and
   control groups. (C) Body weight growth curve changes during gavage in
   the microplastic and control groups. (D) Comparison of colon mass
   between microplastic and control groups. (E) Comparison of colon length
   between microplastic and control groups. (F) Expression levels of Four
   inflammatory factors in microplastic and control groups (ELISA). (G)
   Anatomical diagrams of the intestinal structures and H&E staining of
   colon tissues in the microplastic and control groups. (H) Schematic of
   mechanical sensing and metabolic reprogramming induced by 10 μm PS
   microplastics. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

   Compared to the Control group, mice exposed to 10 μm microplastics
   showed significant weight loss, increased Disease Activity Index (DAI)
   scores, shortened intestinal length, and reduced intestinal mass. In
   contrast, VTPF treatment exerted protective effects, evidenced by
   weight gain, markedly lower DAI scores, and restoration of intestinal
   length and mass to near-control levels (P < 0.05, Fig. [177]10B-E,
   Additional file 8: Fig. [178]S8).

   To further assess the anti-inflammatory effect of VTPF, we performed
   ELISA on intestinal tissues from euthanized mice (Fig. [179]10F). The
   results demonstrated that in the 10 μm + VTPF group, pro-inflammatory
   cytokine levels were significantly reduced, while the anti-inflammatory
   cytokine IL-10 showed partial recovery (P < 0.05). Additionally,
   histological examination of the colonic tissue via H&E staining
   (Fig. [180]10G) revealed no significant inflammation in the Control
   group, while typical pathological features of colitis were observed in
   the 10 μm group. In contrast, intestinal inflammation was significantly
   reduced in the 10 μm + VTPF group. These findings confirm that the YAP
   inhibitor VTPF can mitigate microplastic-induced intestinal
   inflammation, offering significant protection against
   microplastic-induced intestinal damage in mice.

   To explore the molecular mechanisms underlying large-particle
   microplastic-induced intestinal damage, we constructed a mechanistic
   model, as depicted in Fig. [181]10H. Upon contact with the cell
   membrane, large-particle PS microplastics mechanically stimulate YAP
   activation, leading to nuclear translocation. The translocated YAP then
   regulates the expression of glycolysis-related genes, such as LDHA and
   PKM (validated by Western blotting, shown in the supplementary figure),
   promoting a shift towards glycolysis.

Discussion

   With the escalation of microplastic pollution, increasing research has
   focused on its impacts on ecological and human health [[182]43].
   Studies have shown that microplastics can damage intestinal epithelial
   cells and impair barrier integrity through oxidative stress,
   inflammation, and immune dysregulation [[183]44–[184]46]. However,
   microplastics span a wide size range—from nanometers to micrometers—and
   their toxicological mechanisms may differ accordingly. Despite this,
   current research has primarily addressed plastic types and exposure
   levels, with limited attention to particle size. Given that particle
   size affects uptake, immune recognition, and potentially gut microbiota
   composition, it likely plays a key role in intestinal toxicity. Yet,
   its contribution to microplastic-induced intestinal inflammation
   remains insufficiently understood.

   This study aims to address the current knowledge gap by investigating
   how microplastic particle size influences the molecular mechanisms of
   intestinal injury. In our study, we selected 100 nm and 10 μm
   polystyrene particles as representative sizes for nanoplastics and
   microplastics, respectively. This decision was based on two key
   considerations: first, these sizes are among the most commonly detected
   in environmental samples and widely used in experimental studies,
   providing consistency and comparability with existing literature;
   second, they effectively reflect the biological differences between
   cellular-uptake-prone nanoparticles and mechanically disruptive
   microparticles. Therefore, while a broader range of particle sizes
   might offer additional insights, our focus on these two typical sizes
   was intended to highlight the distinct toxicity mechanisms associated
   with each class. Using both cell and animal models, we found that
   nano-scale polystyrene microplastics primarily induce damage through
   ferroptosis, whereas larger micro-scale particles act via mechanical
   disruption and metabolic reprogramming. Furthermore, we identified two
   therapeutic targets and demonstrated that intervention with
   Ferrostatin-1 and VTPF significantly alleviated colon injury in vivo.
   These findings offer new mechanistic insights and suggest potential
   strategies for treating microplastic-induced intestinal damage.

   The core finding of this study is that microplastics of different sizes
   induce intestinal injury via distinct molecular pathways. Smaller
   particles, owing to their larger surface area, more readily penetrate
   the intestinal epithelial barrier [[185]47]. Once internalized, they
   impair mitochondrial function and disrupt iron homeostasis, leading to
   oxidative stress and intracellular iron accumulation, which promotes
   lipid peroxidation and activates ferroptosis [[186]48]. Ferroptosis is
   a regulated form of cell death driven by iron-dependent lipid
   peroxidation [[187]49, [188]50]. The Slc7a11 system imports cystine for
   glutathione (GSH) synthesis, while Gpx4 reduces lipid peroxides
   [[189]51–[190]53]. GSH depletion inactivates Gpx4, causing ROS
   accumulation and ferroptosis initiation.

   Dysregulation of the p53 tumor suppressor pathway is a hallmark of many
   human cancers [[191]37, [192]54]. p53 plays a central role in
   determining cell fate, including the induction of cell cycle arrest,
   apoptosis, and senescence [[193]55, [194]56]. Recent studies have shown
   that elevated ROS levels can activate p53, which in turn suppresses
   Slc7a11 expression and promotes ferroptosis [[195]57]. It is plausible
   that nanoplastics (NPs) exacerbate intracellular oxidative stress and
   iron accumulation, thereby activating this pathway.

   While most previous studies have focused on the ability of
   microplastics to induce cellular damage through oxidative stress
   [[196]58–[197]60], few have explored the potential link between
   microplastics and ferroptosis in colon injury. In our study, proteomic
   analysis revealed a significant upregulation of Fosl1 in the 100 nm
   exposure group. Functional assays confirmed that Fosl1 knockdown
   reduced ferroptosis-related proteins (Slc7a11 and Gpx4) and decreased
   key markers, including ROS levels, JC-1 depolarization, and MDA
   accumulation.

   To investigate the molecular mechanism underlying Fosl1’s role in
   ferroptosis, we conducted ChIP-qPCR and dual-luciferase assays, which
   demonstrated that Fosl1 could bind to the promoter region of p53,
   thereby promoting p53 expression. This, in turn, downregulated the
   expression of SLc7a11 [[198]22, [199]25, [200]58], further exacerbating
   ferroptosis. Our findings highlight the pivotal role of Fosl1 in the
   ferroptosis induced by NPs. This insight not only provides new
   molecular understanding of microplastic-induced ferroptosis but also
   suggests potential therapeutic targets for future treatment of
   microplastic-induced intestinal damage.

   In vivo, flow cytometry and multiplex immunofluorescence revealed
   increased infiltration of dendritic cells (DCs) and CD8⁺ T cells in the
   colonic tissue of NP-exposed mice. In chronic inflammation, elevated DC
   levels often indicate immune overactivation [[201]61]. Ferroptosis may
   promote antigen release, which is recognized by DCs and presented to T
   cells, enhancing CD8⁺ T cell recruitment. This immune-activating
   cascade characterizes immunogenic cell death (ICD), wherein cell death
   alters the immune microenvironment and triggers cytotoxic responses
   [[202]62, [203]63].

   Flow cytometry and multiplex immunofluorescence results collectively
   demonstrate that nanoparticles trigger a pronounced immune and
   inflammatory response in the intestinal microenvironment, thereby
   promoting immunogenic cell death. Moreover, treatment with the
   ferroptosis inhibitor Ferrostatin-1 [[204]39, [205]64] significantly
   attenuated ferroptosis, which was accompanied by a marked reduction in
   the infiltration of activated DCs and CD8 + T cells within the tissues.
   These findings further support the notion that ferroptosis serves as a
   key initiating event in nanoparticle-induced intestinal damage.
   Although this study demonstrated that nanoplastics-induced ferroptosis
   was associated with enhanced infiltration of dendritic cells and CD8⁺ T
   cells, a definitive diagnosis of immunogenic cell death generally
   requires the assessment of specific DAMPs, such as calreticulin (CRT),
   HMGB1, and extracellular ATP. Although these markers were not directly
   measured in this study, previous reports have indicated that
   ferroptosis can trigger DAMP release, contributing to immune
   activation. Future work will focus on validating these molecular
   hallmarks to further confirm the ICD phenotype triggered by
   nanoplastics [[206]65, [207]66].

   Compared to nanoparticles, larger-sized microplastics are often
   overlooked in research due to their limited ability to be internalized
   by cells [[208]67], resulting in a lack of in-depth studies on their
   effects. Unlike smaller nanoparticles, the damage mechanisms of larger
   microplastics are likely more related to physical mechanical forces or
   direct mechanical effects. In our study, these particles disrupted
   membrane integrity and cytoskeletal structure in NCM460 cells, leading
   to altered cell mechanics and metabolic reprogramming. Energy
   metabolomics combined with proteomic analysis revealed significant
   upregulation of glycolysis-related proteins and metabolites, indicating
   a metabolic shift toward anaerobic glycolysis in response to MP-induced
   mechanical stress.

   The metabolic disturbances caused by large microplastics appear closely
   linked to Yes-associated protein (YAP), a key cellular mechanosensor
   involved in translating mechanical stimuli into gene expression changes
   [[209]68, [210]69]. In our study, MP exposure significantly upregulated
   YAP expression and induced its nuclear translocation, likely activating
   glycolysis-related genes. YAP is recognized as a central regulatory
   node connecting mechanical stress to metabolic reprogramming,
   particularly glucose metabolism [[211]41, [212]70]. Targeting YAP
   offers a promising strategy to mitigate MP-induced cellular damage.
   Knockdown of YAP alleviated abnormal glycolytic activity, restored ATP
   synthesis, and reduced lactate accumulation. Furthermore, treatment
   with VTPF, a pharmacological YAP inhibitor, effectively attenuated
   intestinal injury induced by large-sized MPs. These findings highlight
   that microplastics can induce metabolic reprogramming through
   mechanical stress-mediated YAP activation and suggest that modulating
   this pathway may provide therapeutic benefit (Fig. [213]11).

Fig. 11.

   [214]Fig. 11
   [215]Open in a new tab

   Mechanistic illustration of colonic epithelial cell damage induced by
   micro- and nano-sized microplastics

   Although this study reveals novel mechanisms underlying
   microplastic-induced intestinal damage and identifies effective
   therapeutic strategies, several limitations and research gaps remain.
   First, microplastics vary widely in chemical composition, surface
   properties, and environmental interactions. Most studies, including
   ours, focus on a single type (e.g., polystyrene), potentially
   overlooking the distinct or synergistic effects of other polymers such
   as polyethylene. Future research should systematically compare
   different microplastic types, exposure levels, and their combined
   effects with other environmental pollutants on intestinal health.

   Furthermore, the influence of microplastics on the gut microbiome
   remains insufficiently investigated. The gut microbiome plays a
   critical role in maintaining intestinal health and immune function
   [[216]71, [217]72]. Microplastic exposure may disrupt the structure of
   the gut microbiome, which could subsequently affect gut barrier
   function, immune responses, and metabolism [[218]73, [219]74]. Future
   studies could integrate microbiome analysis with metabolomics
   technologies to investigate the effects of microplastic exposure on the
   gut microbiome and its potential role in intestinal damage.

   The impact of microplastics on gut health is not just a biological
   issue but also a growing public health concern [[220]43, [221]75,
   [222]76]. As environmental exposure increases, microplastics have been
   detected in humans via multiple routes—including the food chain,
   drinking water, and airborne particles—with the gastrointestinal tract
   as a major site of accumulation [[223]77, [224]78]. Microplastics may
   contribute to the development of multiple chronic diseases, including
   intestinal inflammation and metabolic disorders [[225]79]. Although
   this study proposes promising therapeutic strategies for
   microplastic-induced gut injury, their clinical translation requires
   further validation through extensive preclinical and clinical studies.

   Although no unified international standard currently exists for human
   microplastic exposure, previous studies estimate that humans may ingest
   approximately 0.1–5 g of microplastics per week, depending on regional
   environmental levels and dietary habits. Based on body surface area
   conversion, this corresponds to an approximate equivalent exposure dose
   of 0.054–2.71 mg/day in mice [[226]74]. The doses used in this study
   fall within or slightly above this range, particularly modeling a
   high-exposure scenario that may be encountered in contaminated
   environments. Therefore, the exposure concentrations we selected remain
   biologically relevant and offer valuable insights into the potential
   health risks associated with heavy microplastic burdens.

   Moreover, our pre-experiments showed that concentrations below 1 mg/mL
   did not significantly affect cell viability, indicating that these
   doses may fall below the biological threshold for measurable toxicity
   in vitro. This supports our decision to use 1 mg/mL as the lower bound
   in mechanistic studies. Nevertheless, we fully acknowledge the
   importance of exploring long-term, low-dose exposure to better mimic
   chronic human exposure. Future studies will further investigate the
   sub-toxic effects of nanoplastics at environmentally relevant
   concentrations over extended exposure periods.

Conclusion

   This study comprehensively elucidates the size-dependent mechanisms by
   which polystyrene (PS) MPs induce intestinal epithelial damage. Our
   findings demonstrate two distinct and non-overlapping toxicological
   pathways: small-sized MPs (100 nm) trigger ferroptosis-driven immune
   injury, while large-sized MPs (10 μm) induce inflammation through
   mechanical disruption and metabolic reprogramming.

   Specifically, 100 nm MPs were internalized by intestinal epithelial
   cells via endocytosis, leading to intracellular oxidative stress and
   ferroptosis. Proteomic analysis revealed a significant upregulation of
   Fosl1 and enrichment in the ferroptosis and p53 pathways.
   Ferrostatin-1, a ferroptosis inhibitor, significantly alleviated both
   cellular and tissue damage, confirming ferroptosis as a central
   mechanism in small MP-induced intestinal toxicity. Additionally, Fosl1
   knockdown reduced ferroptosis-related damage, further highlighting its
   regulatory role.

   In contrast, 10 μm MPs did not enter cells but caused epithelial injury
   via physical compression, cytoskeletal disruption, and metabolic
   reprogramming. Notably, these large particles activated the YAP
   signaling pathway, which promoted glycolysis and intensified
   inflammation. Inhibition of YAP by Verteporfin (VTPF) significantly
   reversed these effects, validating YAP as a key mediator of
   microplastic-induced intestinal injury in larger particles.

   Together, these results uncover critical particle size-specific
   mechanisms—ferroptosis for nanoscale MPs and YAP-mediated glycolytic
   reprogramming for microscale MPs. These mechanistic distinctions offer
   novel insight into microplastic toxicity, establish Fosl1 and YAP as
   potential therapeutic targets, and provide compelling experimental
   evidence for tailored intervention strategies against
   microplastic-induced gut damage.

   In conclusion, this study not only advances our molecular understanding
   of how MPs of different sizes affect intestinal health, but also
   delivers important theoretical support for microplastic risk
   assessment, environmental regulation, and public health protection.

Electronic supplementary material

   Below is the link to the electronic supplementary material.
   [227]Supplementary Material 1^ (3.4MB, docx)
   [228]Supplementary Material 2^ (4.6MB, docx)

Acknowledgements