Polyethylene terephthalate microplastics impair erectile function through macrophage mediated cGAS-STING ferroptosis

Summary

Microplastic pollution is a global concern, yet its impact on male reproductive health remains unclear. We assessed chronic polyethylene terephthalate (PET) microplastic exposure using human corpus cavernosum (CC) tissues, a rat model, and cell assays. MPs were quantified in CC from 10 patients; those with erectile dysfunction (ED) showed a higher MP burden, with PET predominant. In rats, chronic PET-MP exposure dose-dependently impaired erectile function, increased fibrosis, and reduced smooth muscle. Mechanistically, PET-MPs localized to macrophage mitochondria, causing depolarization and ROS generation, mtDNA leakage, cGAS-STING activation, and macrophage ferroptosis. This ferroptotic signaling amplified inflammation, promoted M1 polarization, and triggered endothelial-to-mesenchymal transition, leading to vascular dysfunction and ED. Depleting macrophages or inhibiting cGAS-STING or ferroptosis reduced inflammation and partially rescued erectile responses. Together, these data identify a cGAS-STING-ferroptosis axis linking environmental MP exposure to ED and suggest upstream innate-immune and ferroptosis pathways as therapeutic targets.

Graphical abstract

Highlights

• •PET microplastics accumulate in the human corpus cavernosum and are associated with ED
• •Chronic PET exposure impairs erectile function and promotes penile fibrosis in rats
• •PET activates macrophage mtDNA-cGAS-STING signaling and induces ferroptosis
• •Macrophage ferroptosis drives EndMT and vascular dysfunction to worsen ED

Reproductive medicine; Environmental health

Introduction

Erectile dysfunction (ED) is a prevalent condition that significantly impairs the sexual quality of life for both men and women, with an overall incidence of approximately 40% in men.^1,2 Penile erection is primarily driven by smooth muscle relaxation in the corpus cavernosum (CC), mediated by endothelial nitric oxide (NO) signaling.^3,4 Consequently, endothelial dysfunction is considered a major pathophysiological contributor to ED. In addition to traditional risk factors such as diseases,^5,6,7 medications,⁸ and unhealthy lifestyle habits,⁹ increasing evidence suggests that environmental pollutants also contribute significantly to the pathogenesis of ED. These pollutants include organic toxins,^10,11,12 heavy metals,^13,14 radiation,¹⁵ and plastic-derived compounds.¹⁶

Among these pollutants, microplastics (MPs) have emerged as a pressing global concern. In 2023, global plastic production exceeded 4.1 billion tons, and without effective waste control, annual plastic pollution is projected to surpass 1 billion tons by 2060.^17,18 Discarded plastics are progressively degraded into MPs and nanoplastics (NPs).^19,20 MPs are defined as plastic particles smaller than 5 mm in diameter.²¹ These particles can release additives such as stabilizers, plasticizers, colorants, and flame retardants,^22,23 and may further degrade into NPs with diameters below 1 μm.^19,24 MPs can enter the human body via inhalation,^19,25,26 ingestion,^19,27,28 skin contact,²⁹ or intravenous injection,³⁰ exerting widespread systemic effects on organs such as the lungs,³¹ kidneys,³² blood vessels,³³ and reproductive organs.³⁴ These effects are largely mediated by oxidative stress, inflammatory responses, pyroptosis, and DNA damage.^31,32,34 Recent studies have shown that MPs promote tissue fibrosis, endothelial dysfunction, and redox imbalance, ultimately contributing to ED.¹⁶ Furthermore, several plastic polymers, including polyethylene terephthalate (PET), polystyrene (PS), polyvinyl chloride, and polypropylene, have been detected in human penile tissue.³⁵ Among these, PET appears to be the predominant polymer, yet its specific role in the pathogenesis of ED remains poorly understood.

Macrophages are abundant immune cells in the penile CC and serve as key mediators of inflammation, tissue remodeling, and immune surveillance.^36,37 Macrophage activation is primarily triggered by either pathogen-associated molecular patterns (PAMPs) or damage-associated molecular patterns (DAMPs).^38,39 DAMPs, often generated during cellular injury, can promote macrophage polarization toward the pro-inflammatory M1 phenotype, leading to sterile inflammation.⁴⁰ Among DAMP-associated signals, mitochondrial DNA (mtDNA) is particularly vulnerable to environmental insults because of its unique structure and subcellular localization compared with nuclear DNA.⁴¹ MPs can be internalized into the cytoplasm of macrophages without encapsulation by lipid bilayers, thereby inducing cytotoxicity, disrupting mitochondrial integrity, and promoting mtDNA release.^42,43,44,45 The extruded mtDNA activates the cyclic GMP-AMP synthase (cGAS)–stimulator of interferon genes (STING) pathway, triggering inflammatory cascades and ferroptosis,^44,46,47 thereby establishing a vicious cycle of cellular injury within the local microenvironment. However, the specific mechanisms through which macrophages modulate the local microenvironment during pollutant-induced ED remain poorly understood.

This study aimed to investigate whether PET exposure induces mitochondrial damage in CC cells, leading to the release of mtDNA and subsequent activation of the cGAS-STING signaling pathway in macrophages. It was hypothesized that this activation promotes M1 polarization, induces macrophage ferroptosis, and triggers a pro-inflammatory response, ultimately resulting in endothelial dysfunction and the development of ED. Elucidating this mechanism may provide novel insights into how plastic-derived environmental pollutants contribute to ED.

Results

Polyethylene terephthalate-microplastics accumulation and exposure impair erectile function and induce fibrotic remodeling in penile tissue

MP composition was profiled in CC tissues obtained from penile carcinoma margins with preserved erectile function (Penile-CA, n = 4) and from patients with diabetes and ED undergoing inflatable penile prosthesis implantation (ED-IPP, n = 6). Baseline characteristics of the cohort are shown in Table S2. Controls (n = 4) were older patients with penile cancer with preserved erectile function, whereas patients with ED (n = 6) were generally younger but had long-standing diabetes and poorer glycemic control. BMI and most comorbidities were broadly comparable between groups, with only a few patients showing additional vascular risk factors such as hypertension, smoking, or alcohol use. Multiple polymer types were detected in all samples, with inter-individual variability in relative abundance; PET accounted for a substantial fraction in most cases. PET burden was significantly higher in ED-IPP than in Penile-CA tissues (Figures 1A and 1B; Tables S1 and S2). The PET-MPs used for experimental exposure were characterized prior to use; SEM and disc-centrifuge particle sizing revealed irregular morphology with a modal diameter of approximately 4–5 μm (Figures S1A and S1B).

Figure 1.

Characterization of microplastics in human penile tissues and PET-MP-induced functional and histological alterations in a rat model

(A) Proportional distribution of common MP types in penile tissue samples from patients with Penile-CA and ED-IPP.

(B) Quantification of PET levels in penile tissues from the two groups.

(C) Representative intracavernosal pressure (ICP) and mean arterial pressure (MAP) traces.

(D) ICP/MAP ratio in the control, L-MP, and H-MP groups.

(E) Masson’s trichrome staining of CC tissue; scale bars, 150 μm.

(F) Quantification of smooth muscle-to-collagen ratio.

(G) Western blot analysis of eNOS, p-eNOS, TGF-β1, and α-SMA.

(H) IF staining of eNOS, TGF-β1, and α-SMA in penile tissue; scale bars, 50 μm. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001.

Functionally, rats exposed to PET-MPs exhibited a dose-dependent reduction in erectile responses during cavernous nerve stimulation, as shown by representative ICP traces and group quantitative analysis (Figures 1C and 1D). Histological assessment revealed increased collagen deposition and decreased smooth-muscle content, consistent with fibrotic remodeling (Figures 1E and 1F). At the molecular level, eNOS and p-eNOS were reduced, whereas TGF-β1 was upregulated and α-SMA was downregulated in CC tissues from PET-exposed rats (Figure 1G; densitometry in Figures S1C–S1E). Immunofluorescence (IF) confirmed these alterations (Figure 1H). In this context, α-SMA primarily labels cavernosal smooth muscle cells rather than myofibroblasts; therefore, its decrease reflects the depletion of corporal smooth muscle, consistent with replacement by collagen and fibrotic remodeling. Collectively, these findings demonstrate that elevated PET burden is associated with impaired erectile function and fibrotic remodeling of penile tissue.

Polyethylene terephthalate-microplastics exposure triggers macrophage-mediated inflammation in penile tissue

To investigate the inflammatory response induced by PET-MPs in penile tissue, the expression of pro-inflammatory cytokines was first examined. ELISA and Western blot analyses (Figures 2A and 2B; densitometry in Figures S2A–S2C) demonstrated that PET-MP exposure significantly increased the levels of interleukin (IL)-1β, IL-6, and tumor necrosis factor (TNF)-α in a dose-dependent manner. PET-MPs were detected within CD68-positive macrophages, with stronger co-localization observed in the H-MP group, indicating enhanced phagocytic uptake and activation (Figure 2C). Furthermore, IF staining for M1 (CD86) and M2 (CD163) polarization markers revealed that PET-MP exposure induced a shift toward M1 polarization, particularly in the H-MP group (Figures 2D and S2D). To verify the role of macrophages in ED pathogenesis, clodronate-liposomes were administered to deplete macrophages in vivo. Macrophage depletion with clodronate-liposomes reduced CD68⁺ cell abundance (Figure S2E), partially restored erectile responses (Figures 2E and S2F), and decreased cytokine production (Figure 2F). These findings indicate that PET-MP-induced pro-inflammatory macrophages contribute to ED, and that their depletion alleviates inflammation and functional impairment, consistent with prior evidence showing that MPs promote M1 polarization and inflammatory activation.^48,49,50,51

Figure 2.

PET-MPs induce macrophage-mediated inflammation and contribute to ED

(A) ELISA analysis of pro-inflammatory cytokines IL-1β, IL-6, and TNF-α in CC tissue.

(B) Western blot analysis shows the expression of IL-1β, IL-6, and TNF-α.

(C) IF staining for CD68 (green) and PET particles (red) in penile tissue; scale bars, 50 μm.

(D) M1 and M2 polarization of macrophages assessed by dual IF staining; scale bars, 50 μm.

(E) ICP/MAP measurement in the clodronate + H-MP group and PBS + H-MP groups.

(F) ELISA of IL-1β, IL-6, and TNF-α after macrophage depletion. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001.

Polyethylene terephthalate-microplastics exposure impairs mitochondrial function and redox balance in penile tissue

To investigate the impact of PET-MPs on penile tissue, transcriptomic sequencing was performed on CC samples. Gene Ontology enrichment analysis revealed that exposure to PET-MPs significantly downregulated numerous mitochondrial-related genes, including those involved in the inner mitochondrial membrane, respiratory chain complexes, and adenosine triphosphate metabolism pathways, indicating mitochondrial dysfunction (Figure 3A). Further gene set enrichment analysis demonstrated marked the suppression of the respiratory electron transport chain, mitochondrial membrane components, and oxidative phosphorylation, accompanied by the upregulation of the NF-κB signaling pathway, indicative of an activated inflammatory response (Figure 3B). Fluorescence imaging using MitoTracker revealed PET-MPs within the cytoplasm in close spatial proximity to mitochondria (Figure S3). This proximity indicates co-localization but does not establish physical interaction or recognition by mitochondria. To examine the source of oxidative stress, cells were treated with NAC (a broad-spectrum antioxidant) and Mito-TEMPO (a mitochondria-targeted antioxidant), and total reactive oxygen species (ROS) levels were measured using the DCFH-DA probe. PET-MP exposure significantly increased cellular ROS levels, which were partially reversed by NAC treatment (Figure 3C), suggesting oxidative stress involvement. MitoSOX staining further confirmed that PET-MP exposure markedly elevated mitochondrial-specific ROS production, which was alleviated by both NAC and Mito-TEMPO (Figure 3D). In addition, JC-1 staining demonstrated a loss of mitochondrial membrane potential, with a dose-dependent decrease in the red/green fluorescence ratio (Figures 3E and 3F). Collectively, these results indicate that PET-MP exposure disrupts mitochondrial homeostasis in macrophages by elevating mitochondrial ROS production and inducing mitochondrial membrane depolarization.

Figure 3.

Effects of PET-MP exposure on mitochondrial function and oxidative stress in macrophages

(A) GO enrichment analysis of downregulated genes in PET-exposed CC tissue.

(B) GSEA plots show the enrichment of mitochondrial and inflammatory pathways.

(C) Total intracellular ROS levels detected by DCFH-DA staining and flow cytometry.

(D) Mitochondrial ROS levels assessed by MitoSOX staining and flow cytometry.

(E) JC-1 staining to evaluate mitochondrial membrane potential; scale bars, 50 μm.

(F) Quantification of red/green fluorescence ratio from JC-1 staining. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001.

Polyethylene terephthalate-microplastics exposure induces mitochondrial DNA leakage and cyclic GMP-AMP synthase-stimulator of interferon genes activation in macrophages

To assess mtDNA leakage following PET-MP exposure, IF staining revealed a marked increase in extranuclear mtDNA within macrophages from both L-MP and H-MP groups, appearing as cytosolic puncta adjacent to mitochondria (Figure 4A). Quantitative polymerase chain reaction further confirmed a dose-dependent elevation of cytosolic levels of mtDNA genes ND1, Cytb, and D-loop (Figure 4B). Consistent with impaired mitochondrial fitness, PET-MP exposure decreased the expression of PPARα, PGC-1α, and SDHB, while upregulating the DNA sensors cGAS and STING (Figures 4C and 4D). These changes were further confirmed by IF staining, which showed increased cGAS and STING expression in PET-exposed macrophages, particularly in the H-MP group (Figures 4E and 4F). Collectively, PET-MPs induce mtDNA release and activate the cGAS-STING pathway in macrophages, thereby linking mitochondrial stress to innate immune signaling.

Figure 4.

PET-MP exposure induces mtDNA leakage and activates the cGAS-STING pathway

(A) IF staining shows the co-localization of dsDNA (green) and mitochondria (red) in macrophages; scale bars, 50 μm.

(B) qPCR analysis of cytosolic mtDNA markers (ND1, Cytb, and D-loop).

(C) Western blot analysis of mitochondrial function-related proteins (PPARα, PGC-1α, and SDHB) and DNA-sensing proteins (cGAS and STING).

(D) Densitometric quantification of Western blot bands normalized to β-tubulin.

(E and F) IF staining of STING and cGAS in macrophages; scale bars, 50 μm. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001.

Polyethylene terephthalate-microplastics induce cyclic GMP-AMP synthase-stimulator of interferon genes-mediated ferroptosis in macrophages

To investigate the involvement of cGAS-STING signaling in PET-MP-induced ED, PET-MP-exposed macrophages were compared with cells co-treated with the cGAS inhibitor RU.521. Differential expression analysis identified 249 differentially expressed genes (71 upregulated and 178 downregulated; Figure S4A). KEGG pathway enrichment revealed ferroptosis among the most significantly altered pathways (Figure 5A). A ferroptosis-focused heatmap showed the downregulation of GPX4, SLC7A11, and FTH1, accompanied by the upregulation of TFRC and NCOA4 following PET-MP exposure (Figure 5B). Western blot analysis confirmed a dose-dependent reduction in GPX4 and SLC7A11 together with increased NCOA4 in PET-MP-treated cells, with erastin serving as a positive control (Figure 5C; densitometry in Figure S4D). Biochemical markers supported these transcriptomic findings: malondialdehyde levels were significantly elevated (Figure 5D), whereas glutathione levels were reduced (Figure 5E), indicating lipid peroxidation and impaired antioxidant capacity. Concurrently, intracellular Fe²⁺ concentrations were elevated (Figure 5F), as further verified by FerroOrange imaging (Figure 5H). TEM revealed classical ferroptotic features in PET-treated macrophages, including mitochondrial shrinkage and loss of cristae (Figure 5G). Flow cytometry additionally demonstrated a dose-dependent increase in total cellular ROS, with erastin producing the highest signal (Figures S4B and S4C). Mechanistically, treatment with either Ferrostatin-1 or RU.521 attenuated PET-MP-induced signaling, as evidenced by reduced cGAS and STING expression and partial restoration of GPX4 and NCOA4 levels (Figure 5I; densitometry in Figure S4E). Consistent with the in vitro data, IF staining of rat CC showed decreased GPX4 and increased NCOA4 in PET-MP-exposed tissues (Figure 5J), further supporting the activation of NCOA4-dependent ferroptosis in vivo. Collectively, these results indicate that PET-MPs precipitate cGAS-STING-driven ferroptosis through iron accumulation, NCOA4-mediated ferritinophagy, and oxidative stress, whereas the inhibition of the cGAS-STING pathway partially reverses this process.

Figure 5.

PET-MP exposure induces ferroptosis in macrophages via the cGAS-STING pathway

(A) KEGG pathway enrichment analysis based on DEGs from macrophage samples treated with PET-MP and PET+RU.521.

(B) Expression of ferroptosis-related genes from transcriptomic data.

(C) Western blot analysis of NCOA4, GPX4, and SLC7A11 in macrophages.

(D) MDA levels in different treatment groups.

(E) GSH levels in different treatment groups.

(F) Intracellular Fe²⁺ concentration.

(G) TEM images of macrophage mitochondria; scale bars, 5 μm (left column); scale bars, 1 μm (right column).

(H) Fe²⁺ staining of macrophages.

(I) Western blot analysis after treatment with Fer-1 or RU.521.

(J) IF staining of GPX4 and NCOA4 in CC tissue after PET-MP exposure; scale bars, 50 μm. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001.

Polyethylene terephthalate-microplastics promote EndMT via cyclic GMP-AMP synthase-driven macrophage ferroptosis and inflammatory signaling

To determine whether PET-MPs reprogram the endothelial phenotype through macrophage-mediated inflammation, EndMT and endothelial function were assessed. In rat CC, PET-MP exposure decreased CD31 and increased vimentin expression (Figure 6A). Western blot analysis confirmed that co-treatment with PET-MPs and macrophages reduced E-cadherin and CD31 levels while upregulating α-SMA and FSP1 in endothelial cells (Figure 6B; densitometry in Figure S5A). Functionally, PET-MPs enhanced endothelial cell migration and wound closure, with the MP + Mφ group showing the greatest increase in motility; these changes were partially attenuated by co-treatment with a TGF-β type I receptor kinase inhibitor (TRKi, SB-431542) (Figures 6C–6F). In contrast, tube formation assays revealed that PET-MP exposure impaired the tube-forming ability of endothelial cells, and macrophage co-culture further reduced vascular branching and lumen formation, whereas TRKi partially restored network complexity (Figures 6G and 6H). Similar phenotypic changes were observed in CCECs, where PET-MP exposure downregulated CD31 and upregulated vimentin expression (Figure 6I). Western blot analysis further revealed the activation of the TGF-β/Smad signaling pathway, decreased GPX4 expression, and alterations in EndMT-related markers in MP + Mφ-treated cells. Both the ferroptosis inhibitor Fer-1 and TRKi partially reversed these effects, normalizing E-cadherin, CD31, α-SMA, and GPX4 expression and reducing TGF-β, Smad2/3, and p-Smad2/3 activation (Figure 6J; densitometry in Figures S5B and S5C). Together with the macrophage data, these findings indicate that PET-MPs promote EndMT through macrophage ferroptosis and TGF-β/Smad-dependent inflammatory signaling downstream of cGAS-STING activation. The partial rescue by TRKi suggests that TGF-β signaling acts as a key effector that links macrophage ferroptosis to endothelial dysfunction in PET-MP-induced ED.

Figure 6.

PET-MPs induce EndMT via macrophage ferroptosis and activation of the cGAS-STING pathway

(A) IF staining of CD31 and vimentin in CC tissue; scale bars, 50 μm.

(B) Western blot analysis of E-cadherin, CD31, α-SMA, and FSP1 expression in endothelial cells cultured with PET-MP alone (MP) or PET-MP plus macrophages (MP + Mφ).

(C) Transwell migration assay shows enhanced endothelial cell migration; scale bars, 50 μm.

(D) Quantification of migrated cells in the Transwell assay.

(E) Scratch wound-healing assay shows increased wound closure; scale bars, 100 μm.

(F) Quantification of wound-healing rates.

(G) Tube formation assay shows impaired angiogenesis; scale bars, 100 μm.

(H) Quantification of vascular junctions in the tube-formation assay.

(I) IF staining of CD31 and vimentin expression in endothelial cells; scale bars, 50 μm.

(J) Western blot analysis of TGF-β, Smad2/3 phosphorylation, E-cadherin, and GPX4 expression. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗p < 0.001.

Discussion

Globally, the incidence of ED continues to rise and has become a major burden on men’s health. Although vascular, neural, and endocrine factors are traditionally regarded as the principal causes of ED, accumulating evidence suggests that environmental pollutants are also important pathogenic contributors. In particular, MP pollution has been linked to several core pathophysiological processes relevant to ED, including vascular endothelial injury,⁵² disturbances in glucose and lipid metabolism,^{53,54,55,56,57,58,59} heightened systemic inflammation,^60,61 and structural and functional damage to the nervous system.^62,63 Clinically, substantial amounts of PET have been detected in penile tissues from patients undergoing prosthesis implantation for ED,^35,64 reinforcing the proposed association between environmental MPs and ED.

This study systematically demonstrated, for the first time, that exposure to environmentally prevalent PET-MPs directly induces ED in male animals. Mechanistically, PET-MP exposure triggered mitochondrial stress, mtDNA leakage, and activation of the cGAS-STING pathway in macrophages, leading to ferroptosis, excessive cytokine release, and endothelial dysfunction within the CC. These findings align with previous reports showing that MP exposure elicits immune-inflammatory responses and fibrotic remodeling across organs⁶⁵ and extend those observations to penile tissue. Consistent with translational relevance, multiple polymer types were detected in human CC samples, with a higher MP burden observed in patients with ED than in controls.

Macrophages serve as key amplifiers in this process. MP exposure induces the infiltration of M1-polarized macrophages,⁵⁰ which have been implicated in inflammation within the liver and testis.^51,66 A marked accumulation and M1 polarization of macrophages were observed within the CC following PET-MP exposure, accompanied by robust release of pro-inflammatory cytokines, including TNF-α, IL-6, and IL-1β.^67,68,69,70 These cytokines inhibit vasodilation and damage smooth-muscle function, serving as major contributors to ED.^71,72,73 Macrophage depletion using clodronate liposomes mitigated inflammation and improved erectile responses,⁷⁴ indicating that macrophages act as both targets and drivers of PET-MP toxicity. Phagocytic uptake of PET-MPs and cellular debris^75,76,77,78 likely facilitates cytosolic DNA accumulation and activates the cGAS-STING pathway.⁷⁹ Ferroptosis-related signals were predominantly localized to macrophage-rich regions, further underscoring their central role in mediating cell death and cytokine amplification.

The present data further clarify the upstream role of cGAS-STING. PET-MP exposure increased mitochondrial ROS, damaged mtDNA, and promoted cytosolic mtDNA leakage, thereby activating cGAS-STING and initiating local immune-inflammatory responses. Long-term exposure to MPs or NPs induces the cytosolic accumulation of double-stranded DNA in various organs, activates the STING pathway, and upregulates downstream pro-inflammatory cytokines, contributing to hepatic fibrosis,⁸⁰ pulmonary fibrosis,⁴⁷ cardiomyocyte senescence,⁸¹ esophageal and gastric mucosal inflammation,^82,83 spermatogenic disorders,⁸⁴ and aberrant activation of macrophage immune responses.^79,85 Downstream, cGAS-STING promotes ferroptosis, characterized by iron-catalyzed lipid peroxidation, GPX4 suppression, and iron overload,⁸⁶ through increased mtROS and mitochondrial-membrane injury.^46,87 STING has been reported to translocate to mitochondria, interact with MFN1/2 to elevate mtROS,⁸⁸ and cooperate with NCOA4-dependent ferritinophagy to exacerbate ferroptosis and inflammation.⁸⁹ Furthermore, IRF3, an STING effector, has been shown to modulate ferroptosis.⁹⁰ Consistent with these mechanistic links, our data indicate that the PET-MP-induced activation of cGAS-STING is required for NCOA4-dependent ferritinophagy and ferroptosis in macrophages. PET-MP exposure upregulated NCOA4, increased Fe²⁺ accumulation and lipid peroxidation, and suppressed GPX4, whereas the pharmacological inhibition of cGAS with RU.521 or blockade of ferroptosis with Ferrostatin-1 attenuated cGAS-STING activation and partially normalized these ferroptosis markers. Collectively, these findings indicate that cGAS-STING signaling acts not only as a key upstream driver of inflammation but also as a direct regulator of metabolic imbalance and ferroptosis. As the pharmacologic modulation of the cGAS-STING pathway is being actively explored as a therapeutic strategy in cardiovascular, hepatic, neurologic, and sepsis-associated diseases,^91,92,93,94 our data further support cGAS-STING as a plausible upstream target for intervention in environmentally induced ED, a clinically complex and multifactorial condition.

The release of cellular contents and inflammatory cytokines such as TNF-α, IL-1β, and IL-6 during ferroptosis exacerbates local inflammation and causes severe endothelial dysfunction, manifested by EndMT.^95,96,97 PET-MPs decreased endothelial markers (CD31, eNOS) and increased mesenchymal markers (α-SMA, vimentin), indicating EndMT.⁹⁸ EndMT is a process in which endothelial cells, under inflammatory or stress stimuli, acquire fibroblast-like characteristics, contributing to vascular remodeling and fibrosis.⁹⁹ It is considered a key mechanism in vascular-related diseases such as ED.^95,100 The present data further demonstrate that PET-MP-induced macrophage inflammation, through the release of TGF-β and the generation of reactive oxygen species, creates a pro-inflammatory microenvironment that drives endothelial cells to lose their phenotype and acquire mesenchymal features.^101,102 EndMT leads to the replacement of CC smooth muscle by collagen and other matrix proteins, reducing vascular compliance, consistent with the fibrotic and endothelial injury phenotype observed after high-dose nanoplastic exposure.¹⁶ The cGAS-STING pathway may also participate in EndMT regulation, as previous studies have shown that mtDNA-induced cGAS-STING signaling directly promotes mesenchymal transition in endothelial cells, resulting in endothelial dysfunction.¹⁰³ In PET-MP-induced ED, macrophage-mediated inflammation via the cGAS-STING pathway induces ferroptosis, which amplifies inflammatory signaling to endothelial cells, triggers EndMT and endothelial injury, and ultimately results in ED. This provides a novel perspective on the contribution of endothelial injury and EndMT to ED, highlighting that environmental pollutants can indirectly destabilize the endothelial phenotype and compromise vascular structural integrity through immune-driven inflammation. Targeted interventions, including RU.521, Fer-1, and clodronate liposomes, designed to block the cGAS-STING pathway, inhibit ferroptosis, or deplete pro-inflammatory macrophages, significantly alleviated PET-MP-induced tissue inflammation and ED pathology, further consolidating the mechanistic pathway proposed in this study.

In conclusion, this study systematically demonstrates that PET-MP exposure induces mitochondrial dysfunction and ROS accumulation in macrophages, leading to mtDNA release and activation of the cGAS-STING pathway. This cascade promotes M1 polarization, ferroptosis, and inflammatory cytokine production, which in turn drives endothelial dysfunction and EndMT in cavernosal endothelial cells, ultimately resulting in the structural and functional impairment of penile tissue and the development of ED. These findings highlight macrophage-endothelial crosstalk via cGAS-STING-driven ferroptosis as a central mechanism by which environmental MPs damage the male reproductive system and suggest upstream innate immune signaling and ferroptosis pathways as plausible therapeutic targets for environment-related ED.

Limitations of the study

This study has several limitations. First, all experiments were conducted in a rat model, and the exposure doses and regimens used cannot fully reproduce the chronic, low-dose, mixed-polymer MP exposure that humans typically experience. Moreover, we examined PET as a single representative polymer and did not evaluate other types of MPs or their potential interactions, which may limit the generalizability of our findings. Second, although human ED is usually driven by a combination of vascular, neural, endocrine, and psychological factors, this work mainly focused on cavernosal macrophage ferroptosis and vascular dysfunction and did not directly investigate neurogenic or endocrine mechanisms. Emerging experimental evidence suggests that micro- and nanoplastics can cross biological barriers, induce neuroinflammation, and disrupt endocrine homeostasis, and these systemic effects will need to be specifically explored in future studies. Third, the human CC cohort in this study consisted of patients with penile cancer with preserved erectile function, and men with long-standing diabetic ED, and did not include patients with non-diabetic ED or patients with diabetic without ED as separate comparison groups. Therefore, the current human data cannot determine whether increased cavernosal PET-MP accumulation is mainly attributable to diabetes itself, ED-related vascular alterations, or their combined effects, and these human findings are better regarded as exploratory support for the mechanisms identified in the animal and in vitro models.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Fujun Zhao (zhaofujun2005@sjtu.edu.cn).

Materials availability

This study did not generate new unique reagents. All antibodies, chemicals, commercial kits, software, and instrumentation are listed in the key resources table.

Data and code availability

Raw RNA-seq reads have been deposited in the NCBI Sequence Read Archive (SRA) under the BioProjects PRJNA1364359 and PRJNA1364739. No custom code was used. Additional information required to reanalyze the data reported in this article is available from the lead contact upon request. Source data for other figures will also be provided upon request.

Acknowledgments

This work was supported by the National Key Research and Development Program of China (Grant No. 2022YFC2702701), the National Natural Science Foundation of China (Grant Nos. 82371634 and 82300870), and the Natural Science Foundation of Shanghai (Grant No. 22ZR1450800).

Author contributions

Conceptualization and study design: S.L., W.W., and Y.Z.; methodology: S.L., Y.Z., S.Li., and H.J.; investigation: S.L., H.J., S.L., L.W., and P.D.; formal analysis and visualization: S.L., W.W., and Y.Z.; data curation: S.L., Y.Z., and J.L.; writing – original draft: S.L., W.W., and Y.Z.; writing – review and editing: J.L., C.J., and F.Z.; resources: J.L. and F.Z.; funding acquisition: C.J. and F.Z.; supervision: F.Z.

Declaration of interests

The authors declare that they have no conflict of interest.

STAR★Methods

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
TGF-β	Abcam	Cat# ab92486; RRID: AB_10561953
α-SMA	Abcam	Cat#ab7817; RRID: AB_307611
eNOS	Cell Signaling Technology	Cat#9572; RRID: AB_10693928
p-eNOS (Ser1177)	Cell Signaling Technology	Cat#9571; RRID: AB_329837
β-Tubulin	Proteintech	Cat#10068-1-AP; RRID: AB_2288035
IL-1β	Cell Signaling Technology	Cat#12703; RRID: AB_2687522
IL-6	Cell Signaling Technology	Cat#12912; RRID: AB_2715528
TNF-α	Abcam	Cat#ab6671; RRID: AB_305641
PPARα	Abcam	Cat#ab8934; RRID: AB_306004
PGC-1α	Abcam	Cat#ab54481; RRID: AB_881987
SDHB	Proteintech	Cat#10620-1-AP; RRID: AB_2146320
cGAS	Cell Signaling Technology	Cat#31659; RRID: AB_2799180
STING	Cell Signaling Technology	Cat#13647; RRID: AB_2732796
GPX4	Abcam	Cat#ab125066; RRID: AB_11156945
E-cadherin	Cell Signaling Technology	Cat#3195; RRID: AB_10562570
Smad2	Cell Signaling Technology	Cat#5339; RRID: AB_10699035
Smad3	Cell Signaling Technology	Cat#9523; RRID: AB_331576
p-Smad2	Cell Signaling Technology	Cat#3108; RRID: AB_490941
p-Smad3	Cell Signaling Technology	Cat#9520; RRID: AB_331037
CD31	Abcam	Cat#ab28364; RRID: AB_726362
FSP1	Abcam	Cat#ab218163; RRID: AB_2868454
CD86	Abcam	Cat#ab53004; RRID:AB_869910
CD163	Proteintech	Cat# 16646-1-AP, RRID: AB_10639860
CD68	Proteintech	Cat# 28058-1-AP, RRID: AB_2881133
Vimentin	Proteintech	Cat# 10366-1-AP, RRID: AB_2265789
TOM20	Proteintech	Cat#11802-1-AP; RRID:AB_2207530
anti-dsDNA	Abcam	Cat #ab27156; RRID:AB_2923262
Biological samples
Human corpus cavernosum tissues	This paper;	N/A
Chemicals, peptides, and recombinant proteins
Polyethylene terephthalate microplastics (PET-MPs)	Zhichuan BioTech	N/A
N-Acetyl-L-cysteine (NAC)	Sigma-Aldrich	Cat#A7250
Mito-TEMPO	Sigma-Aldrich	Cat#SML0737
RU.521 (cGAS inhibitor)	Invitrogen	Cat#TCI-S3335
Ferrostatin-1 (Fer-1)	MedChemExpress	Cat#HY-100579
Erastin	MedChemExpress	Cat#HY-15763
SB-431542 (ALK5 inhibitor)	Selleckchem	Cat#S1067
TRIzol reagent	Invitrogen	Cat#15596018
Triton X-	Beyotime	Cat#ST795
4% paraformaldehyde	Beyotime	Cat#P0099
MitoTracker	Beyotime	Cat#C1048
DAPI	Servicebio	Cat#G1012
Critical commercial assays
BCA Protein Assay Kit	Beyotime	Cat# P0012
RIPA lysis buffer	Beyotime	Cat#P0013B
Masson’s Trichrome Staining Kit	Servicebio	Cat#G1006
ECL kit	NCM Biotech	Cat#P10100
ELISA kits (TNF-α )	Jianglai Bio	Cat#JL10539
ELISA kits ( IL-1β)	Jianglai Bio	Cat#JL18442
ELISA kits (IL-6 )	Jianglai Bio	Cat#JL20896
MDA Assay Kit	Beyotime	Cat#S0131S
GSH Assay Kit	Jiancheng Bioengineering	Cat#A006-2
Tissue Iron Colorimetric Assay	Jiancheng Bioengineering	Cat#A039-2
JC-1 Mitochondrial Membrane Potential Assay	Beyotime	Cat#C2006
DCFH-DA (cellular ROS)	Beyotime	Cat#S0033S
MitoSOX™ Red (mitochondrial ROS)	Invitrogen	Cat#M36007
QIAamp DNA Mini Kit	QIAGEN	Cat#51304
FerroOrange	Cell Signaling Technology	Cat#36104
Deposited data
Raw RNA-seq reads (rat corpus cavernosum)	SRA	BioProject: PRJNA1364359;
Raw RNA-seq reads (rat macrophages)	SRA	BioProject: PRJNA1364739;
Experimental models: Cell lines
Primary rat bone-marrow–derived macrophages (BMDMs)	Procell	CP-R141
Rat corpus cavernosum endothelial cells (CCECs, primary)	Procell	CP-R133
Experimental models: Organisms/strains
Rat: Sprague–Dawley, male, 8-week (250 ± 15 g)	Shanghai Laboratory Animal Center	N/A
Oligonucleotides
Primers in this paper, see STAR Methods for RNA extraction and quantitative PCR	This paper	N/A
Software and algorithms
ImageJ	NIH	https://imagej.nih.gov/ij/
GraphPad Prism v9.0	GraphPad Software	https://www.graphpad.com/
FlowJo	FlowJo, LLC	https://www.flowjo.com/
F-Search (py-GC/MS library search)	Frontier Laboratories	https://frontier-lab.com/
DESeq2 (RNA-seq)	Bioconductor	https://bioconductor.org/packages/DESeq2/

Experimental model and study participant details

Human tissues

All procedures involving human tissue were approved by the Ethics Committee of Shanghai General Hospital (Permit No. 2023-108) and were conducted in accordance with the Declaration of Helsinki. Written informed consent was obtained from all CC donors after they were fully informed of the study objectives and procedures. Four normal CC samples were collected from the surgical margins of penile carcinoma resections. All patients in this group reported preserved erectile function, including stimulus-induced and morning erections. An additional six CC samples from patients with diabetes and ED were obtained via biopsy during inflatable penile prosthesis implantation. All human participants were male. Clinical, and baseline characteristics for all patients providing CC samples are summarized in Table S2.

Animals

All animal procedures were approved by the Institutional Animal Care and Use Committee of Shanghai General Hospital (Approval No. 2024AW012) and complied with relevant international guidelines for laboratory animal care and use. Eight-week-old male Sprague–Dawley rats (250 ± 15 g; Shanghai Laboratory Animal Center, China) were housed under controlled conditions (18–24°C, 40–70% humidity, 12-h light/dark cycle) with free access to food and water. Rats were randomly allocated to nine groups (n = 10 per group): control, low- and high-dose PET exposure, low- and high-dose fluorescent PET exposure, and the corresponding low- and high-dose PET or fluorescent PET groups combined with PBS- or clodronate-liposome injection.

Human microplastic exposure was estimated at 0.1–5 g/week for a 60 kg adult,¹⁰⁴ corresponding to human equivalent daily doses of 0.238 mg/kg (low) and 11.90 mg/kg (high). Animal equivalent doses were derived by body surface area normalization (Animal equivalent dose (AED) = Human equivalent dose (HED) × Km; Km for rats = 6.2),¹⁰⁵ yielding 1.48 mg/kg and 73.81 mg/kg. PET-MPs (Zhichuan BioTech, China) were characterized by scanning electron microscopy and particle size analysis and administered in drinking water at 0, 10, or 100 mg/L to mimic environmentally relevant exposure. The PET-MP exposure concentrations of 10 mg/L and 100 mg/L used in this study fall within the extrapolated rat-equivalent dose range and are consistent with dose levels commonly employed in rodent micro- and nanoplastic toxicology studies.^{106,107,108,109,110}

To assess the contribution of macrophages, rats in selected groups received PBS- or clodronate-liposomes (Yesen Biotechnology; PBS-liposomes 40338ES05/10, clodronate-liposomes 40337ES05/10) via tail vein injection. Clodronate-liposomes were given as a 50 mg/kg loading dose followed by 25 mg/kg every 7 days for three additional injections,^111,112 with PBS-liposomes administered on the same schedule. After 4 weeks of exposure, erectile function testing, histological analysis, molecular assays, and transcriptomic profiling were performed.

Cell culture and treatments

Primary rat bone marrow–derived macrophages (BMDMs) and corpus cavernosum endothelial cells (CCECs) were obtained from Procell Life Science & Technology (Wuhan, China) and cultured under standard conditions (37°C, 5% CO₂). BMDMs were maintained in Macrophage Complete Medium (Procell), and CCECs were cultured in Endothelial Cell Medium (ECM; ScienCell, USA).

To evaluate the paracrine effects of macrophages on endothelial cells, BMDMs were treated with PET-MPs (50 or 100 μg/mL) for 24 h.^79,113,114 Culture supernatants were collected, centrifuged to remove cellular debris, and used to stimulate CCECs for an additional 24 h.

In selected experiments, BMDMs were pretreated for 1 h with pharmacological inhibitors, including N-acetyl-L-cysteine (NAC, 3 mM; Sigma-Aldrich, A7250), Mito-TEMPO (20 μM; Sigma-Aldrich, SML0737),¹¹⁵ RU.521 (5 μM; Invitrogen, TCI-S3335) (Haarer et al., 2023), Ferrostatin-1 (Fer-1, 2 μM; MedChemExpress, HY-100579),^116,117 Erastin (10 μM; MedChemExpress, HY-15763),¹¹⁶ or SB-431542 (10 μM; Selleck, S1067),¹¹⁸ a selective transforming growth factor (TGF)-β type I receptor (ALK5) inhibitor. All treatment concentrations were selected based on previously published studies.

Method details

Microplastic detection by pyrolysis–GC/MS

Approximately 0.2 g of human CC tissue was treated with 10% KOH solution (three times the tissue volume) at 40°C under continuous agitation for 72 h.^119,120 The digested mixture was transferred into ultracentrifuge tubes, mixed with absolute ethanol, and centrifuged at 100,000 × g for 4 h. The resulting precipitate, containing MPs, was washed three times with ethanol, dried, and subjected to pyrolysis.

Pyrolysis–gas chromatography/mass spectrometry (Py–GC/MS) was performed using an Agilent 7890 GC/5977 MS system coupled with a Frontier PY-3030D pyrolyzer interfaced to a Shimadzu GCMS-QP2010 Plus. Separation was achieved on an SH-Rxi-5MS capillary column (30 m × 0.25 mm × 0.25 μm, Shimadzu). For each sample, the weighed precipitate was placed into a stainless-steel crucible and pyrolyzed at 550°C under high-purity helium (5.0 grade) as the carrier gas (1.0 mL/min). The GC temperature program was: initial 40°C (2 min), ramp to 320°C at 20°C/min, hold for 14 min. The MS was operated in selected ion monitoring (SIM) mode.

Identification and quantification targeted 12 common polymer types, including polymethyl methacrylate (PMMA), polyethylene (PE), polyvinyl chloride (PVC), polyamide 66 (PA66), polyethylene terephthalate (PET), polycarbonate (PC), acrylonitrile–butadiene–styrene (ABS), polypropylene (PP), thermoplastic polyurethane (TPU), polyamide 6 (PA6), styrene–butadiene (SB), and polystyrene (PS). Mass spectral data were processed using F-Search software, and polymer concentrations (μg/g tissue) were determined via external calibration curves constructed from reference standards.

Quality control followed a plastic-free protocol: laboratory coats were 100% cotton, nitrile gloves were used, all glassware was rinsed with Milli-Q water and covered with aluminum foil, and plastic consumables were strictly avoided.

Microplastic characterization

The morphology and particle size of PET-MPs were analyzed using scanning electron microscopy (SEM) and centrifugal particle size (CPS) analysis. For SEM, samples were mounted on aluminum stubs, gold-coated, and imaged using a ZEISS scanning electron microscope (Germany) at 10.00 kV with a working distance of 7.2 mm and 2,000× magnification (SE2 detector). For particle size distribution, PET particles were dispersed in deionized water by sonication and analyzed using a CPS Disc Centrifuge (CPS Instruments, USA) under a sucrose gradient at room temperature.

Assessment of erectile function

As previously described,¹²¹ rats were anesthetized with 5% pentobarbital sodium. The major pelvic ganglion and cavernous nerve were exposed via a lower abdominal incision. A 25 G butterfly needle filled with 250 U/mL heparinized saline was inserted into the CC to record intracavernous pressure (ICP), while the cavernous nerve was stimulated using a hook electrode (5 V, 20 Hz, 5 ms pulse width, 60 s duration). A second 25 G needle was inserted into the right carotid artery to monitor mean arterial pressure (MAP). Signals were recorded using a physiological recording system (MP160, BIOPAC Systems). The ICP/MAP ratio was calculated as an index of erectile function.

Western blotting

Total protein was extracted from cells and penile tissues using RIPA lysis buffer supplemented with phenylmethylsulfonyl fluoride (PMSF). Protein concentration was measured using a BCA assay kit (Beyotime, China). Equal amounts of protein (30 μg) were separated on 10% SDS-PAGE gels at 120 V for 2 h and transferred onto NC membranes at 400 mA for 1 h.

Membranes were blocked with 5% non-fat milk at room temperature for 2 h and incubated overnight at 4°C with primary antibodies against TGF-β (Abcam, ab92486, 1:1000), α-SMA (Abcam, ab7817, 1:5000), eNOS (Cell Signaling Technology, CST #9572, 1:1000), p-eNOS (CST #9571, 1:1000), β-tubulin (Proteintech, 10068-1-AP, 1:5000), IL-1β (CST #12703, 1:1000), IL-6 (CST #12912, 1:1000), TNF-α (Abcam, ab6671, 1:1000), PPARα (Abcam, ab8934, 1:1000), PGC-1α (Abcam, ab54481, 1:1000), SDHB (Proteintech, 10620-1-AP, 1:2000), cGAS (CST #31659, 1:1000), STING (CST #13647, 1:1000), GPX4 (Abcam, ab125066, 1:1000), E-cadherin (CST #3195, 1:1000), Smad2 (CST #5339, 1:1000), Smad3 (CST #9523, 1:1000), p-Smad2 (CST #3108, 1:1000), p-Smad3 (CST #9520, 1:1000), CD31 (Abcam, ab28364, 1:1000), and FSP1 (Abcam, ab218163, 1:1000). After washing, membranes were incubated with HRP-conjugated secondary antibodies for 1 h at room temperature. Bands were visualized using an ECL kit (NCM Biotech, Suzhou, China), and signal intensity was quantified using ImageJ software.

Immunofluorescence

Immunofluorescence (IF) staining of cultured cells and paraffin-embedded penile tissue was performed following previously established procedures.¹²² After fixation, permeabilization, and blocking, samples were incubated overnight at 4°C with primary antibodies against eNOS, TGF-β, α-SMA, CD86, CD163, CD68, dsDNA, STING, cGAS, CD31, and Vimentin at the manufacturer-recommended dilutions. For mitochondrial labeling, MitoTracker dye was applied to cultured cells prior to fixation. After incubation with appropriate fluorophore-conjugated secondary antibodies and DAPI counterstaining, fluorescence images were acquired using a BX43 fluorescence microscope (Olympus, Tokyo, Japan).

Flow cytometric detection of ROS and mitochondrial ROS

Intracellular ROS were measured using DCFH-DA (Beyotime, China), and mitochondrial ROS were assessed with MitoSOX Red (Invitrogen, USA), following the manufacturers’ protocols. Cells were incubated with 10 μM DCFH-DA or 5 μM MitoSOX Red at 37°C for 30 min in the dark, washed with PBS, and analyzed by flow cytometry (CytoFLEX, Beckman Coulter, USA). Data were quantified using FlowJo software.

Assessment of oxidative stress and iron content

Oxidative stress in CC tissues and BMDMs were evaluated by measuring malondialdehyde (MDA) and glutathione (GSH) using commercial kits (Beyotime Biotechnology, Shanghai, China for MDA; Jiancheng Bioengineering Institute, Nanjing, China for GSH) according to the manufacturers’ protocols. Total protein concentrations were used for normalization. Tissue iron content was measured using a colorimetric assay kit (Jiancheng Bioengineering Institute, Nanjing, China).

Masson’s trichrome staining

Paraffin-embedded penile tissue sections (5 μm) were stained using a Masson’s trichrome staining kit (G1006, Servicebio, Wuhan, China) according to the manufacturer’s instructions. Collagen fibers were stained blue and smooth muscle red. Images were acquired using a light microscope (Leica Microsystems, Germany), and the smooth muscle/collagen ratio was quantified with ImageJ software.

Transmission electron microscopy (TEM)

Cells were treated with PET-MPs for 24 h, then fixed with 3% glutaraldehyde and 1% osmium tetroxide. After dehydration and embedding, ultrathin sections were cut, stained with uranyl acetate and lead citrate, and examined under a transmission electron microscope (Hitachi H-7650, Japan).

Transcriptomic analysis

Total RNA was extracted from rat CC or macrophage samples using TRIzol reagent (Invitrogen), and RNA integrity was assessed with an Agilent 2100 Bioanalyzer. RNA-seq library preparation and sequencing were performed by OE Biotech Co., Ltd. (Shanghai, China). After library construction and quality control, paired-end sequencing was carried out on an Illumina platform. Raw reads were quality-filtered and aligned to the Rattus norvegicus reference genome. Differential gene expression (DEG) was analyzed using DESeq2, with significance thresholds set at |log₂FC| > 1 and adjusted p < 0.05. Functional enrichment analyses, including Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analyses, were conducted to interpret the biological relevance of differentially expressed genes (DEGs).

JC-1 staining

Mitochondrial membrane potential was assessed using JC-1 dye (Beyotime Biotechnology, China). Treated cells were incubated with JC-1 working solution at 37°C for 20 min in the dark, washed twice with JC-1 buffer, and imaged using a fluorescence microscope (Leica Microsystems, Germany). Red/green fluorescence ratios were used to evaluate mitochondrial depolarization.

Mitochondrial DNA quantification

Cells were seeded at 1 × 10⁶ cells/well in 6-well plates. After treatment, cells were lysed with 1% NP-40 on ice for 20 min. Supernatants were collected by centrifugation (16,000 × g, 15 min, 4°C), and cytosolic DNA was extracted using the QIAamp DNA Mini Kit (QIAGEN, Germany).^123,124,125

qPCR was performed using SYBR Green Master Mix (Bio-Rad) on a 7500 Real-Time PCR System (Applied Biosystems, USA). Primers targeted mtDNA regions (CytB, ND1, D-loop), with 18S rRNA as the nuclear reference gene. Relative mtDNA abundance was calculated using the 2ˆ–ΔCt method. The primers used were:

CytB-F: GCTTTCCACTTCATCTTACCATTTA

CytB-R: TGTTGGGTTGTTTGATCCTG

ND1-F: TATCTCAACCCTAGCAGAAA

ND1-R: TAACGCGAATGGGCCGGCTG

D-loop-F: AATCTACCATCCTCCGTGAAACC

D-loop-R: TCAGTTTAGCTACCCCCAAGTTTAA

18S-F: TAGAGGGACAAGTGGCGTTC

18S-R: CGCTGAGCCAGTCAGTGT

Endothelial functional assays

To assess angiogenic capacity, CCECs (2 × 10⁴ cells/well) were seeded onto 96-well plates precoated with Matrigel (Corning, 356234, USA) and cultured in ECM medium. After 6 h, capillary-like networks were imaged using an inverted microscope (BX43; Olympus, Tokyo, Japan) and quantified morphologically.

Cell migration was evaluated using a Transwell assay. CCECs (5 × 10⁴ cells) in serum-free ECM were placed in the upper chamber of inserts (8.0 μm pores; Corning), with ECM containing 10% FBS in the lower chamber as chemoattractant. After 24 h, migrated cells were fixed, stained with crystal violet, and counted under a microscope.

For the wound healing assay, a uniform scratch was introduced into confluent CCECs in 6-well plates using a sterile 200 μL pipette tip. Cells were rinsed with PBS and incubated in serum-free ECM. Images were acquired at 0 and 24 h to evaluate wound closure.

Quantification and statistical analysis

All image quantifications, including fluorescence intensity and Western blot densitometry, were performed using ImageJ software (NIH, USA). Statistical analyses were performed using GraphPad Prism 9.5 (GraphPad Software, USA). Data are presented as mean ± standard deviation (SD). For comparisons between two groups, an unpaired two-tailed Student’s t test was used when assumptions of normality and homogeneity of variance were satisfied; otherwise, the Mann–Whitney U test was applied. For multiple-group comparisons, one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test was used. A p value < 0.05 was considered statistically significant. Statistical significance was denoted as ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001.

Published: February 17, 2026