From September 2024 to September 2025, patients who underwent functional endoscopic sinus surgery (FESS), septoplasty, or inferior turbinoplasty at Daejeon St. Mary’s Hospital were enrolled in this study. Patients who declined consent for nasal tissue sampling during surgery, as well as those with cystic fibrosis, primary ciliary dyskinesia, immunodeficiency, coagulation disorders, or pregnancy or breastfeeding, were excluded. A total of 36 subjects were included and categorized into three groups: healthy controls (n=12), ECRSwNP (n=12), and non-ECRSwNP (n=12). ECRSwNP was defined as the presence of ≥70 eosinophils per high-power field (HPF, ×400 magnification) in five representative hematoxylin and eosin (H&E)–stained sections, based on criteria proposed by Nakayama et al. [10] and Tokunaga et al. [11]. These studies demonstrated that this cutoff was significantly associated with an increased risk of polyp recurrence and disease refractoriness and accordingly adopted it as a defining criterion for ECRSwNP in their patient classification. Control tissues were obtained from the uncinate process or inferior turbinate, whereas tissues from ECRSwNP and non-ECRSwNP patients were collected from nasal polyp specimens obtained during FESS. Collected nasal tissues were either flash-frozen for molecular analysis or fixed in 10% neutral-buffered formalin, processed, and embedded in paraffin (FFPE) for histological examination. The study protocol was approved by the Institutional Review Board of Daejeon St. Mary’s Hospital (IRB policy No. DC24TISI0055, 03 September 2024). Written informed consent was obtained from all participants prior to surgical intervention and tissue collection, in accordance with the principles of the Declaration of Helsinki.

FFPE nasal tissue sections were cut at a thickness of 4 μm, deparaffinized in xylene, and rehydrated through a graded ethanol series. Standard H&E staining was performed to evaluate tissue morphology and inflammatory cell infiltration. Bright-field images were acquired at ×200 magnification (scale bar=100 μm) using an Olympus BX51 microscope (Olympus, Tokyo, Japan). Eosinophil infiltration was quantified by counting eosinophils in five randomly selected, non-overlapping HPFs (×400 magnification) per section. Results are expressed as the mean number of eosinophils per HPF.

Total RNA was extracted from human nasal mucosal tissues and HL-60 cells using the Easy-BLUEG^TM Total RNA Extraction Kit (iNtRON Biotechnology, Seongnam, Republic of Korea) according to the manufacturer’s instructions. RNA concentration and purity were measured using a NanoDrop One spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Complementary DNA (cDNA) was synthesized from 1 μg of total RNA using Reverse Transcriptase Premix (Elpis Biotech, Daejeon, Republic of Korea). Quantitative real-time polymerase chain reaction (qRT-PCR) was performed on an ABI 7500 FAST Real-Time PCR System (Applied Biosystems, Foster City, CA, USA) using gene-specific primers and SYBR Green Master Mix (Applied Biosystems, Warrington, UK). For human tissues, transcripts of IL-4, IL-5, IL-17, interferon gamma (IFN-γ), IL-5RA, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH) were analyzed, whereas IL-5RA, C-C chemokine receptor type 3 (CCR3), and GAPDH were assessed in HL-60 cells. Each 20-μL reaction contained 10 μL of SYBR Green Master Mix, 0.5 μM of each primer, and 2 μL of cDNA. The thermal cycling protocol consisted of initial denaturation at 95°C for 10 minutes, followed by 40 cycles of 95°C for 15 seconds and 60°C for 1 minute. Relative mRNA expression levels were calculated using the 2⁻ΔΔCt method, with GAPDH serving as the internal control. Primer sequences were as follows: IL-4 (forward: 5′-CGAGTTGAC CGTAACAGACAT-3′, reverse: 5′-CGTCT TTAGCCTTTC CAAGAAG-3′); IL-5 (forward: 5′-AGCTG CCTACGTGTAT GCCA-3′, reverse: 5′-GCAGTGCCAAGG TCTCTTTCA-3′); IL-17 (forward: 5′-TCAACCCGATT GTCCACCAT-3′, reverse: 5′-GAGTTTAGTCCGAAAT GAGGCTG-3′); IFN-γ (forward: 5′-ATTCGGTAACT GACTTGAATGTCC-3′, reverse: 5′-CTCTTCGACCTC GAAACAGC-3′); IL-5RA (forward: 5′-CCTTCTCTTC CAGCTTTGCAC-3′, reverse: 5′-A GTCCTTGACGCACACAACA-3′); CCR3 (forward: 5′-GC CATTTCGGACCTG CTCTT-3′, reverse: 5′-TCCGCTCA CAGTCATTTCCA-3′); and GAPDH (forward: 5′-AACTT TGGCATTGTGGAA GG-3′, reverse: 5′-ACACATTGGG GGTAGGAACA-3′).

FFPE nasal tissue sections (4 μm) were deparaffinized and rehydrated, followed by antigen retrieval in 10 mM citrate buffer (pH 6.0) using microwave heating for 15 minutes. After cooling, sections were treated with 3% hydrogen peroxide for 10 minutes to block endogenous peroxidase activity. Samples were then blocked with 5% bovine serum albumin (BSA) for 30 minutes and incubated overnight at 4°C with primary antibodies against IL-5Rα (Abcam, Cambridge, UK), E-cadherin (Abcam, Cambridge, UK), and vimentin (Cell Signaling Technology, Danvers, MA, USA). Following phosphate-buffered saline (PBS) washes, sections were incubated with a biotinylated secondary antibody for 1 hour and subsequently visualized using the Vectastain ABC Kit and ImmPACT Vector Red substrate (Vector Laboratories, Burlingame, CA, USA). Nuclei were counterstained with hematoxylin, and slides were mounted with coverslips. Images were captured at ×400 magnification (scale bar=50 μm) using an Olympus BX51 microscope. Quantitative analysis was performed using ImageJ software (National Institutes of Health, Bethesda, MD, USA). The percentage of positive cells for each marker was calculated as the ratio of immunopositive cells to total nucleated cells in five randomly selected HPFs (×400). All analyses were conducted in a blinded manner by two independent observers, and mean values were used for statistical analysis.

Nasal mucosal tissues from healthy controls, ECRSwNP patients, and non-ECRSwNP patients, as well as HL-60 cells under various experimental conditions, were used for protein analysis. Samples were lysed on ice for 30 minutes in lysis buffer (Elpis Biotech, Daejeon, Republic of Korea) supplemented with protease and phosphatase inhibitor cocktails (Roche Diagnostics, Mannheim, Germany). Lysates were centrifuged at 13,000 rpm for 15 minutes at 4°C, and supernatants were collected. Protein concentrations were determined using a bicinchoninic acid assay kit (Pierce, Rockford, IL, USA). Equal amounts of protein (20 μg) were separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis and transferred onto nitrocellulose membranes (Bio-Rad Laboratories, Hercules, CA, USA). Membranes were blocked with 5% skim milk in Tris-buffered saline containing 0.1% Tween-20 for 1 hour at room temperature and incubated overnight at 4°C with primary antibodies against IL-5Rα, phosphorylated Janus kinase 2 (p-JAK2; Tyr1007/1008), total Janus kinase 2 (JAK2), phosphorylated signal transducer and activator of transcription 5 (p-STAT5; Tyr694), total signal transducer and activator of transcription 5 (STAT5), and E-cadherin (all from Abcam, Cambridge, UK; 1:1,000 dilution), as well as vimentin and GAPDH (both from Cell Signaling Technology, Danvers, MA, USA; 1:1,000 dilution). After three washes with TBST, membranes were incubated with horseradish peroxidase–conjugated secondary antibodies (Invitrogen, Carlsbad, CA, USA) for 2 hours at room temperature. Protein bands were visualized using an enhanced chemiluminescence detection system (Millipore, Burlington, MA, USA) and imaged with a ChemiDoc XRS+ system (Bio-Rad Laboratories, Hercules, CA, USA). Band intensities were quantified using ImageJ software. Relative protein expression levels were normalized to GAPDH for total proteins or to the corresponding total protein for phosphorylated forms (p-JAK2/JAK2 and p-STAT5/STAT5).

Human promyelocytic leukemia HL-60 cells (ATCC, Manassas, VA, USA) were cultured in RPMI-1640 medium (Gibco, Grand Island, NY, USA) supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin. Cells were maintained at 37°C in a humidified incubator containing 5% CO₂, and the culture medium was replaced every 2–3 days. To generate differentiated eosinophil-like HL-60 cells (Diff. HL-60), cells were treated with 0.5 mM sodium butyrate (Sigma-Aldrich, St. Louis, MO, USA) for 7 days, with medium changes performed every 48 hours. Successful differentiation was verified by qRT-PCR, demonstrating a marked increase in IL-5RA mRNA expression relative to undifferentiated HL-60 cells, consistent with acquisition of an eosinophil-like transcriptional profile. For activation, Diff. HL-60 cells were stimulated with recombinant human IL-5 (PeproTech, Rocky Hill, NJ, USA; 10 ng/mL) for 24 hours, generating the Diff. HL-60 + IL-5 group. Untreated HL-60 cells served as undifferentiated controls.

Primary HNECs were purchased from PromoCell GmbH (Heidelberg, Germany) and cultured in Bronchial Epithelial Cell Growth Medium (PromoCell) supplemented with the manufacturer’s recommended growth factors. Cells were maintained at 37°C in a humidified atmosphere containing 5% CO₂, and passages 3–5 were used for all experiments. For co-culture experiments, HNECs were seeded in 6-well plates at a density of 2×10⁵ cells per well and cultured until reaching 80%–90% confluence. Diff. HL-60 cells and IL-5-stimulated Diff. HL-60 cells (Diff. HL-60 + IL-5), prepared as described above, were added to HNEC cultures at a 1:2 ratio (HNECs:HL-60 cells) using Transwell inserts (0.4-μm pore size; Corning Inc., Corning, NY, USA), allowing paracrine interaction without direct cell–cell contact.

Experimental groups were as follows: 1) HNECs + HL-60 (undifferentiated HL-60 cells), 2) HNECs + Diff. HL-60 cells, and 3) HNECs + Diff. HL-60 + IL-5 cells.

After 48 hours of co-culture, HNECs were collected for immunofluorescence staining.

HNECs co-cultured with HL-60 cells under the indicated experimental conditions were subjected to immunofluorescence staining to evaluate epithelial and mesenchymal marker expression. After 48 hours of co-culture, HNECs were fixed with 4% paraformaldehyde for 15 minutes at room temperature, permeabilized with 0.1% Triton X-100 for 10 minutes, and blocked with 4% BSA in PBS for 2 hours. Cells were then incubated for 1 hour at room temperature with primary antibodies against E-cadherin (Abcam;1:200) and vimentin (Cell Signaling Technology;1:200). Following three washes with PBS, cells were incubated for 30 minutes in the dark with Alexa Fluor–conjugated secondary antibodies (Invitrogen). Nuclei were counterstained with 4′,6-diamidino-2-phenylindole (DAPI) for 5 minutes, and coverslips were mounted using an anti-fade mounting medium. Fluorescence images were acquired using a BX53 fluorescence microscope (Olympus) at ×200 magnification under identical exposure settings for all samples. E-cadherin signals were visualized in green, vimentin in red, and nuclei in blue. Quantitative fluorescence intensity analysis was performed using ImageJ software. Fluorescence intensity values were normalized to the control group (HNECs co-cultured with undifferentiated HL-60 cells; HNECs + HL-60) and expressed as fold change relative to control. All image analyses were conducted in a blinded manner by two independent observers, and the mean of their measurements was used for statistical analysis.

Secretion of eosinophil-associated mediators was quantified using commercially available enzyme-linked immunosorbent assay (ELISA) kits. Diff. HL-60 cells were treated with vehicle or recombinant human IL-5 (10 ng/mL) for 24 hours, after which culture supernatants were collected and centrifuged at 1,000×g for 10 minutes to remove cellular debris. Concentrations of eosinophil cationic protein (ECP), transforming growth factor-β1 (TGF-β1), eosinophil peroxidase (EPO), and IL-13 were measured using ELISA kits (R&D Systems, Minneapolis, MN, USA) according to the manufacturer’s instructions. Absorbance was measured at 450 nm with a reference wavelength of 570 nm using a microplate reader (Bio-Rad Laboratories, Hercules, CA, USA). Analyte concentrations were calculated from standard curves and normalized to viable cell counts.

All experiments were performed in at least three independent replicates. Data are presented as mean±standard error of the mean. Statistical comparisons among groups were performed using one-way analysis of variance, followed by the Tukey post hoc test. Associations between tissue IL-5Rα expression and EMT marker levels across individual patient samples were assessed using Spearman’s rank correlation analysis. All statistical analyses were conducted using GraphPad Prism 5 (GraphPad Software, La Jolla, CA, USA), and a p-value <0.05 was considered statistically significant.

To confirm the type 2 inflammatory profile of the study subjects, histological and transcriptional analyses were performed. H&E staining demonstrated markedly increased eosinophil infiltration in nasal polyp tissues from patients with ECRSwNP compared with those from non-ECRSwNP patients and healthy controls (Fig. 1A). Quantitative assessment under HPFs revealed a significant increase in eosinophil counts in ECRSwNP tissues (81.82±2.28 cells/HPF) relative to non-ECRSwNP tissues (11.87±5.84 cells/HPF; p<0.001) and control tissues (1.44±0.19 cells/HPF; p<0.001) (Fig. 1B).

Relative mRNA expression levels of representative type 2 cytokines (IL-4 and IL-5), a type 1 cytokine (IFN-γ), and a type 3 cytokine (IL-17) were quantified by qRT-PCR and normalized to GAPDH. IL-4 and IL-5 expression levels were significantly elevated in the ECRSwNP group compared with both the non-ECRSwNP group and healthy controls (both p< 0.001), whereas IL-17 and IFN-γ expression levels were significantly higher in the non-ECRSwNP group than in both the ECRSwNP group and controls (both p<0.001) (Fig. 1C–F). These results validate that the ECRSwNP group exhibits a type 2–dominant inflammatory profile, consistent with previous characterizations.

Immunohistochemistry (IHC) staining for IL-5Rα in nasal mucosal tissues from a healthy control, an ECRSwNP patient, and a non-ECRSwNP patient is shown in Fig. 2A. Quantitative analysis of IL-5Rα-positive cells demonstrated a significant increase in ECRSwNP tissues (25.17%±2.55%) compared with control tissues (2.61%±0.25%; p<0.001) and non-ECRSwNP tissues (4.90%±0.81%; p<0.001) (Fig. 2B). Consistent with the IHC findings, IL-5RA mRNA expression, as determined by qRT-PCR and normalized to GAPDH, was significantly elevated in ECRSwNP tissues compared with both control tissues (p<0.001) and non-ECRSwNP tissues (p=0.001) (Fig. 2C).

To further substantiate the IHC and qRT-PCR findings, Western blot analysis was performed to evaluate IL-5Rα protein expression and activation of downstream signaling molecules. This analysis revealed distinct differences in both IL-5Rα expression and phosphorylation patterns of IL-5Rα-associated signaling proteins in ECRSwNP tissues compared with the other groups (Fig. 2D). Quantitative densitometric analysis demonstrated that IL-5Rα expression, as well as the ratios of phosphorylated to total JAK2 (p-JAK2/JAK2) and STAT5 (p-STAT5/STAT5), were all significantly increased in ECRSwNP tissues relative to control and non-ECRSwNP tissues. Corresponding p-values were 0.001 and 0.003 for IL-5Rα, 0.002 and 0.031 for p-JAK2/JAK2, and <0.001 and <0.001 for p-STAT5/STAT5, respectively (Fig. 2E–G). These findings indicate that the IL-5Rα–mediated JAK2/STAT5 signaling pathway is activated in ECRSwNP.

To assess the extent of EMT among the three groups—ECRSwNP, non-ECRSwNP, and healthy controls—we performed IHC staining for E-cadherin, an epithelial marker, and vimentin, a mesenchymal marker (Fig. 3A). Quantitative analysis of the positively stained area revealed distinct expression patterns among groups. E-cadherin expression was significantly reduced in both ECRSwNP and non-ECRSwNP tissues compared with control tissues (both p<0.001), indicating epithelial barrier disruption. However, no significant difference in E-cadherin expression was observed between the ECRSwNP and non-ECRSwNP groups (p=0.684) (Fig. 3B). In contrast, vimentin expression was markedly increased in both ECRSwNP and non-ECRSwNP tissues compared with control tissues (both p<0.001), reflecting enhanced mesenchymal features. Notably, the ECRSwNP group exhibited a significantly greater vimentin-positive area than the non-ECRSwNP group (p<0.001) (Fig. 3C).

To further evaluate protein-level differences, Western blot analysis was performed to assess E-cadherin and vimentin expression in tissue samples from control, non-ECRSwNP, and ECRSwNP groups (Fig. 3D). Densitometric quantification normalized to GAPDH generally supported the IHC findings for group-wise changes relative to controls. E-cadherin expression was significantly decreased in both ECRSwNP and non-ECRSwNP tissues compared with controls (control vs. ECRSwNP, p=0.003; control vs. non-ECRSwNP, p<0.001), whereas no significant difference was observed between the ECRSwNP and non-ECRSwNP groups (p=0.698) (Fig. 3E). Conversely, vimentin expression was significantly increased in both ECRSwNP and non-ECRSwNP tissues compared with control tissues (control vs. ECRSwNP, p<0.001; control vs. non-ECRSwNP, p<0.001). However, vimentin expression did not differ significantly between the ECRSwNP and non-ECRSwNP groups (p=0.101) (Fig. 3F).

To support the proposed mechanistic link between IL-5Rα signaling and EMT-related changes in human tissues, correlation analyses were performed across individual patient samples using IHC-derived positivity values (Supplementary Fig. 1 in the online-only Data Supplement). When all samples were pooled (n=36), IL-5Rα positivity showed a strong inverse correlation with E-cadherin positivity (Spearman’s ρ=−0.946, p< 0.001) (Supplementary Fig. 1A in the online-only Data Supplement) and a strong positive correlation with vimentin positivity (Spearman’s ρ=0.915, p<0.001) (Supplementary Fig. 1B in the online-only Data Supplement). Within the ECRSwNP subgroup, significant correlations in the expected directions were also observed (E-cadherin: ρ=−0.839, p<0.001; vimentin: ρ=0.882, p=0.001; n=12), providing supportive human evidence that higher tissue IL-5Rα expression is associated with EMT-like marker patterns in this phenotype.

To investigate the process by which HL-60 cells differentiate into eosinophil-like cells and become activated upon IL-5 stimulation, we compared IL-5RA mRNA expression among three groups: undifferentiated HL-60 cells, butyric acid–induced Diff. HL-60 cells, and IL-5-treated Diff. HL-60 cells. qRT-PCR analysis normalized to GAPDH demonstrated a stepwise increase in IL-5RA expression across the three conditions. IL-5RA mRNA levels were significantly higher in Diff. HL-60 cells than in undifferentiated HL-60 cells (p=0.003) and were further elevated in Diff. HL-60 cells following IL-5 stimulation compared with unstimulated Diff. HL-60 cells (p=0.011) (Fig. 4A). To further characterize the eosinophil-like phenotype beyond IL-5RA expression, we assessed CCR3 expression and eosinophil-associated effector mediators. Butyric acid treatment significantly increased CCR3 mRNA expression and enhanced secretion of EPO and IL-13, which were further augmented following IL-5 stimulation (Supplementary Fig. 2 in the online-only Data Supplement), supporting the development of a more comprehensive and IL-5-responsive eosinophil-like phenotype.

To assess the secretory profile of eosinophil-associated mediators, we measured the concentrations of ECP and TGF-β1 using ELISA. No significant differences were observed between undifferentiated HL-60 cells and Diff. HL-60 cells in either ECP (p=0.072) or TGF-β1 levels (p=0.238). In contrast, levels of both ECP and TGF-β1 were significantly increased in Diff. HL-60 cells following IL-5 stimulation compared with the other two groups (p<0.001 for all comparisons), indicating IL-5–induced, activation-dependent upregulation of eosinophil-related secretory function (Fig. 4B and C).

To evaluate the impact of eosinophilic differentiation and IL-5 stimulation on IL-5Rα expression and downstream signaling activation, Western blot analysis was performed using GAPDH as a loading control (Fig. 4D). Densitometric quantification revealed a progressive increase in IL-5Rα protein expression from undifferentiated HL-60 cells to Diff. HL-60 cells (p=0.003), with further elevation following IL-5 stimulation compared with unstimulated Diff. HL-60 cells (p=0.047) (Fig. 4E). Phosphorylation of JAK2 did not differ significantly between undifferentiated and differentiated cells (p=0.733) but was significantly increased following IL-5 stimulation compared with both undifferentiated HL-60 cells (p=0.015) and Diff. HL-60 cells (p=0.037) (Fig. 4F). Similarly, p-STAT5 levels were significantly elevated in Diff. HL-60 cells (p= 0.002) and were further increased in IL-5-stimulated Diff. HL-60 cells (p<0.001) compared with undifferentiated cells. IL-5-stimulated Diff. HL-60 cells also exhibited higher p-STAT5 expression than differentiated cells without stimulation (p= 0.011) (Fig. 4G). Collectively, these results indicate that eosinophilic differentiation of HL-60 cells, together with subsequent IL-5 stimulation, induces IL-5Rα–mediated activation of downstream JAK2/STAT5 signaling pathways.

HNECs were co-cultured with undifferentiated HL-60 cells, Diff. HL-60 cells, or IL-5-stimulated Diff. HL-60 cells, and changes in EMT marker expression were analyzed by Western blot using GAPDH as a loading control (Fig. 5A). Densitometric analysis demonstrated that E-cadherin expression was significantly reduced in HNECs co-cultured with IL-5-stimulated Diff. HL-60 cells compared with those co-cultured with undifferentiated HL-60 cells (p=0.009) or unstimulated Diff. HL-60 cells (p=0.007). Conversely, vimentin expression was significantly increased in the IL-5-stimulated Diff. HL-60 co-culture group relative to the undifferentiated HL-60 (p=0.005) and unstimulated Diff. HL-60 groups (p= 0.043). Notably, no significant differences were observed between the undifferentiated HL-60 and unstimulated Diff. HL-60 co-culture groups for either E-cadherin (p=0.945) or vimentin (p=0.191) (Fig. 5B and C).

To visually confirm the effects of HL-60 differentiation and IL-5 stimulation on EMT marker expression in HNECs, immunofluorescence staining was performed (Fig. 5D). Quantitative analysis of fluorescence images yielded results consistent with the Western blot findings. HNECs co-cultured with IL-5-stimulated Diff. HL-60 cells exhibited significantly lower E-cadherin fluorescence intensity than those co-cultured with undifferentiated HL-60 cells (p=0.004) or Diff. HL-60 cells (p=0.004). In contrast, vimentin fluorescence intensity was significantly higher in the IL-5-stimulated Diff. HL-60 co-culture group compared with the undifferentiated HL-60 (p=0.001) and Diff. HL-60 groups (p=0.003). No significant differences were observed between undifferentiated HL-60 and unstimulated Diff. HL-60 co-cultures for either E-cadherin (p=0.999) or vimentin (p=0.524).

Collectively, these findings indicate that IL-5Rα–mediated signaling in eosinophil-like HL-60 cells promotes EMT in HNECs, as evidenced by coordinated downregulation of E-cadherin and upregulation of vimentin.