IL-5Rα–Mediated Signaling in Eosinophils May Contribute to the Epithelial–Mesenchymal Transition in Eosinophilic Chronic Rhinosinusitis With Nasal Polyps
Article information
Abstract
Background and Objectives
Eosinophilic chronic rhinosinusitis with nasal polyps (ECRSwNP) is characterized by type 2 inflammation, prominent eosinophil accumulation, and enhanced epithelial–mesenchymal transition (EMT). Although interleukin (IL)-5 plays a central role in eosinophil differentiation and activation, its involvement in driving EMT in human nasal epithelial cells (HNECs) remains unclear. This study investigated IL-5 receptor α (IL-5Rα)–mediated signaling and EMT profiles in ECRSwNP, aiming to determine whether IL-5–activated eosinophil-like cells promote EMT in HNECs.
Methods
Nasal tissues from healthy controls, non-ECRSwNP patients, and ECRSwNP patients (n=12 per group) were analyzed using hematoxylin and eosin staining, quantitative real-time polymerase chain reaction, immunohistochemistry, and Western blotting. HL-60 promyelocytic leukemia cells were differentiated into eosinophil-like cells and subsequently stimulated with IL-5. HNECs were co-cultured with undifferentiated HL-60 cells, differentiated HL-60 cells, or IL-5–stimulated differentiated HL-60 cells using a Transwell system. Expression of EMT-related markers was assessed by Western blotting and immunofluorescence.
Results
ECRSwNP tissues demonstrated markedly increased eosinophil infiltration, elevated IL-4 and IL-5 transcript levels, and significantly higher expression of IL-5Rα, phosphorylated Janus kinase 2 (JAK2), and phosphorylated signal transducer and activator of transcription 5 (STAT5) compared with controls and non-ECRSwNP tissues. EMT-associated changes, characterized by reduced E-cadherin and increased vimentin expression, were most pronounced in ECRSwNP. In vitro, IL-5 stimulation enhanced IL-5Rα expression, activated JAK2/STAT5 signaling, and increased secretion of eosinophil-associated mediators in differentiated HL-60 cells. Co-culture with IL-5-stimulated eosinophil-like cells induced EMT in HNECs, whereas unstimulated HL-60 cells exerted minimal effects.
Conclusion
IL-5Rα-mediated signaling is upregulated in ECRSwNP and may contribute to EMT through eosinophil-dependent mechanisms. IL-5–activated eosinophils promote EMT-related changes in HNECs, providing a mechanistic link between type 2 inflammation and epithelial remodeling. These findings suggest that IL-5Rα signaling may represent a potential therapeutic target for modulating tissue remodeling in ECRSwNP.
INTRODUCTION
Chronic rhinosinusitis (CRS) is defined as inflammation of the nasal cavity and paranasal sinuses lasting more than 12 weeks and has a substantial negative impact on patients’ quality of life [1]. According to a meta-analysis, CRS affects approximately 8.71% of the global population, and its prevalence has increased over time [2]. Traditionally, CRS has been classified into two phenotypes, CRS with nasal polyps (CRSwNP) and CRS without nasal polyps (CRSsNP), based on endoscopic findings. However, more recent approaches emphasize endotypes defined by underlying pathogenic mechanisms, allowing CRSwNP to be further subdivided into type 2 and non–type 2 CRS. This endotype-based classification is increasingly applied to guide patient management and to predict disease risk and prognosis [1]. Type 2 CRS is characterized by persistent eosinophilic inflammation and frequent disease recurrence even after surgery and is closely associated with comorbid asthma and other type 2 inflammatory diseases [3,4].
Recent studies have increasingly focused on the pathogenesis of CRS, particularly mechanisms underlying tissue remodeling. Among these mechanisms, epithelial–mesenchymal transition (EMT) has emerged as a key process contributing to epithelial barrier dysfunction and chronic inflammation [5,6]. EMT is a biological process in which polarized epithelial cells acquire mesenchymal characteristics. This process is initiated by disruption of tight junctions, leading to decreased expression of epithelial markers such as E-cadherin, occludin, claudin, and zonula occludens-1, along with increased expression of mesenchymal markers including N-cadherin, vimentin, α-smooth muscle actin, collagen, fibronectin, and other extracellular matrix proteins. As a consequence, cells lose polarity and cell–cell adhesion while acquiring migratory and invasive properties [6].
EMT is known to play a more prominent pathogenic role in CRSwNP than in CRSsNP [6]. Among patients with CRSwNP, eosinophilic CRSwNP (ECRSwNP), which represents a type 2 endotype, exhibits a greater degree of EMT than non-ECRSwNP. Accumulating evidence suggests that eosinophils may actively contribute to this process [7,8].
Among the key cytokines involved in type 2 inflammation, interleukin (IL)-4 has been shown to induce EMT in human nasal epithelial cells (HNECs) through the signal transducer and activator of transcription 6 (STAT6)/interferon regulatory factor 4 (IRF4) signaling pathway [8].
IL-5, another central type 2 cytokine, plays a critical role in the growth, differentiation, and activation of eosinophils, which serve as key effector cells in ECRSwNP [9]. However, whether IL-5 can similarly induce EMT in HNECs has not been previously investigated. Therefore, in this study, we aimed to determine whether IL-5 receptor α (IL-5Rα)–mediated signaling contributes to EMT in ECRSwNP.
METHODS
Study subjects
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.
H&E staining and eosinophil quantification
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.
Quantitative real-time polymerase chain reaction
Total RNA was extracted from human nasal mucosal tissues and HL-60 cells using the Easy-BLUEGTM 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′).
Immunohistochemistry and quantitative analysis
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.
Western blot 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).
Cell culture and differentiation
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.
HNEC culture and co-culture with differentiated HL-60 cells
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.
Immunofluorescence staining and quantitative analysis
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.
Enzyme-linked immunosorbent assay
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.
Statistical analysis
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.
RESULTS
Increased eosinophil infiltration and upregulated type 2 cytokine mRNA expression in ECRSwNP
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).
Histological and transcriptional features of ECRSwNP and non-ECRSwNP. A: H&E-stained nasal mucosal sections from a healthy control, an ECRSwNP patient, and a non-ECRSwNP patient (×200 magnification; scale bar=100 μm). B: Quantification of eosinophil infiltration per HPF in H&E-stained sections shown in panel (A). C and D: Relative mRNA expression of type 2 cytokines (IL-4 and IL-5) determined by qRT-PCR and normalized to GAPDH. E and F: Relative mRNA expression of type 1 and type 3 cytokines (IL-17 and IFN-γ) analyzed by qRT-PCR, with GAPDH used as the internal reference. Results are expressed as the mean±SEM of 12 patient samples per group. Statistical significance was determined by one-way ANOVA followed by the Tukey post hoc test. *p<0.05. ECRSwNP, eosinophilic chronic rhinosinusitis with nasal polyps; H&E, hematoxylin and eosin; ns, not significant; HPF, high-power field; IL, interleukin; IFN-γ, interferon gamma; qRT-PCR, quantitative real-time polymerase chain reaction; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; SEM, standard error of the mean; ANOVA, analysis of variance.
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.
Upregulation of IL-5RA and associated downstream pathways in ECRSwNP
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).
Upregulation of IL-5Rα and activation of downstream signaling pathways in ECRSwNP. A: IHC staining of IL-5Rα in nasal mucosal tissues from a healthy control, an ECRSwNP patient, and a non-ECRSwNP patient. Positive signals are indicated by red chromogen (×400 magnification; scale bar=50 μm). B: Quantification of IL-5Rα–positive cells, expressed as the percentage of positively stained cells in the IHC images shown in panel (A). C: IL-5RA mRNA expression analyzed by qRT-PCR and normalized to GAPDH. D: Representative Western blot analysis of IL-5Rα and downstream phosphorylated signaling proteins in nasal mucosal tissue lysates. GAPDH was used as a loading control; two representative samples per group are shown. E–G: Densitometric quantification of IL-5Rα and phosphorylated signaling proteins. Band intensities were normalized to GAPDH for IL-5Rα (E) and to the corresponding total protein for phosphorylated forms (F and G). Results are expressed as the mean±SEM of 12 patient samples per group. Statistical significance was determined by one-way ANOVA followed by the Tukey post hoc test. *p<0.05. IL-5Rα, interleukin-5 receptor α; ECRSwNP, eosinophilic chronic rhinosinusitis with nasal polyps; ns, not significant; p-JAK2, phosphorylated Janus kinase 2; JAK2, Janus kinase 2; p-STAT5, phosphorylated signal transducer and activator of transcription 5; STAT5, signal transducer and activator of transcription 5; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; IHC, immunohistochemistry; qRT-PCR, quantitative real-time polymerase chain reaction; SEM, standard error of the mean; ANOVA, analysis of variance.
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.
Differential expression of EMT markers 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).
Differential expression of EMT markers in ECRSwNP and non-ECRSwNP. A: IHC staining of E-cadherin (top row) and vimentin (bottom row) in nasal mucosal tissues from a healthy control, an ECRSwNP patient, and a non-ECRSwNP patient. Positive signals are indicated by red chromogen (×400 magnification; scale bar=50 μm). B and C: Quantification of E-cadherin– and vimentin–positive areas from the IHC staining shown in panel (A). D: Western blot analysis of E-cadherin and vimentin in nasal mucosal tissue lysates from control, ECRSwNP, and non-ECRSwNP groups. GAPDH served as the loading control, and two representative samples are shown for each group. E and F: Densitometric quantification of E-cadherin (E) and vimentin (F) normalized to GAPDH. Results are expressed as the mean±SEM of 12 patient samples per group. Statistical significance was determined by one-way ANOVA followed by the Tukey post hoc test. *p<0.05. ECRSwNP, eosinophilic chronic rhinosinusitis with nasal polyps; ns, not significant; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; EMT, epithelial–mesenchymal transition; IHC, immunohistochemistry; SEM, standard error of the mean; ANOVA, analysis of variance.
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.
IL-5 induces eosinophilic differentiation, effector function, and JAK2–STAT5 signaling in HL-60 cells
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.
Differentiation of HL-60 cells into eosinophil-like cells and IL-5–induced effector molecule release and JAK2/STAT5 pathway activation. A: Relative mRNA expression of IL-5RA in undifferentiated HL-60 cells (HL-60), butyric acid–differentiated eosinophil-like HL-60 cells (Diff. HL-60), and differentiated HL-60 cells further stimulated with IL-5 (Diff. HL-60 + IL-5). Expression levels were analyzed by qRT-PCR and normalized to GAPDH. B and C: Secretion of eosinophil-associated mediators from differentiated HL-60 cells in response to IL-5 stimulation. Butyric acid–differentiated HL-60 cells were treated with vehicle or 10 ng/mL IL-5 for 24 hours and culture supernatants were analyzed by ELISA for ECP (B) and TGF-β1 (C). D: Representative Western blot analysis of IL-5Rα and downstream phosphorylated signaling proteins in HL-60, Diff. HL-60, and Diff. HL-60 + IL-5 groups. GAPDH was used as a loading control. E–G: Densitometric quantification of IL-5Rα and phosphorylated signaling proteins in HL-60, Diff. HL-60, and Diff. HL-60 + IL-5 groups. Band intensities were normalized to GAPDH for IL-5Rα (E) and to the corresponding total protein for phosphorylated forms (F and G). Results are expressed as the mean±SEM of three independent experiments (n=3). Statistical significance was determined by one-way ANOVA followed by the Tukey post hoc test. *p<0.05. IL-5Rα, interleukin-5 receptor α; IL-5, interleukin-5; ECP, eosinophil cationic protein; ns, not significant; TGF-β1, transforming growth factor β1; p-JAK2, phosphorylated Janus kinase 2; JAK2, Janus kinase 2; p-STAT5, phosphorylated signal transducer and activator of transcription 5; STAT5, signal transducer and activator of transcription 5; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; qRT-PCR, quantitative real-time polymerase chain reaction; ELISA, enzyme-linked immunosorbent assay; SEM, standard error of the mean; ANOVA, analysis of variance.
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.
IL-5Rα–mediated signaling contributes to EMT in HNECs
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).
Co-culture with IL-5–stimulated eosinophil-like HL-60 cells promotes EMT in HNECs. A: Western blot analysis of E-cadherin and vimentin in HNECs co-cultured with undifferentiated HL-60 cells (HNECs + HL-60), butyric acid–differentiated HL-60 cells (HNECs + Diff. HL-60), and differentiated HL-60 cells further stimulated with IL-5 (HNECs + Diff. HL-60 + IL-5). GAPDH served as the loading control. B and C: Densitometric quantification of E-cadherin (B) and vimentin (C) normalized to GAPDH from the Western blot analysis shown in panel (A). D: Immunofluorescence staining of HNECs co-cultured with undifferentiated HL-60 cells (HNECs + HL-60), butyric acid–differentiated HL-60 cells (HNECs + Diff. HL-60), or IL-5–stimulated, butyric acid–differentiated HL-60 cells (HNECs + Diff. HL-60 + IL-5). E-cadherin (green, upper panels) and vimentin (red, lower panels) were visualized, with nuclei counterstained by DAPI (blue). Images were acquired at ×200 magnification (scale bar=100 μm). E and F: Quantitative analysis of immunofluorescence intensity for E-cadherin (E) and vimentin (F), expressed as fold change relative to the HNECs + HL-60 group. Results are expressed as the mean±SEM of three independent experiments (n=3). Statistical significance was assessed by one-way ANOVA followed by the Tukey post hoc test. *p<0.05. HNECs, human nasal epithelial cells; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; ns, not significant; IL-5, interleukin-5; EMT, epithelial–mesenchymal transition; DAPI, 4′,6-diamidino-2-phenylindole; SEM, standard error of the mean; ANOVA, analysis of variance.
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.
DISCUSSION
In this study, we demonstrated that ECRSwNP, characterized by eosinophilic inflammation and upregulated IL-5Rα–mediated signaling, exhibited a greater degree of EMT compared with healthy controls. Furthermore, using a co-culture model incorporating eosinophil-like cells, we showed that IL-5Rα–mediated signaling in eosinophils may contribute to EMT in HNECs, which may help explain the enhanced EMT observed in the ECRSwNP group.
Eosinophils play a pivotal role in the pathogenesis of ECRSwNP, and IL-5 is essential for their differentiation, survival, migration, and activation [9]. IL-5 exerts its biological effects by binding to IL-5Rα, leading to receptor dimerization with the common β chain and initiation of multiple intracellular signaling cascades [12,13]. Among these downstream pathways, IL-5Rα activation triggers JAK2, which subsequently phosphorylates STAT5. p-STAT5 then translocates to the nucleus and induces transcription of target genes involved in eosinophil survival and effector functions, including degranulation [13–16].
Among type 2 cytokines implicated in ECRSwNP, Chen et al. [8] demonstrated that IL-4 promotes EMT in HNECs via the STAT6/IRF4 signaling pathway. Furthermore, IL-4–induced M2 macrophages were shown to enhance EMT progression in HNECs, suggesting that both direct and indirect IL-4–mediated mechanisms contribute to epithelial remodeling.
With respect to IL-5, another key type 2 cytokine, Chen et al. [17] recently reported that IL-5RA is upregulated in lung epithelial cells during the development of pulmonary fibrosis. Their study demonstrated that epithelial IL-5RA promotes EMT through activation of the JAK2/STAT3 signaling pathway. These findings suggest that IL-5RA, beyond its classical role in eosinophils, may directly contribute to fibrotic remodeling in lung epithelial cells.
However, to date, no study has directly investigated the effect of IL-5 on EMT in HNECs. To our knowledge, the present study is the first to demonstrate that IL-5, in the presence of eosinophils, contributes to EMT in HNECs. This finding highlights a novel mechanism by which IL-5 may influence epithelial remodeling in ECRSwNP through eosinophil-mediated crosstalk within the tissue microenvironment. We demonstrated that IL-5 stimulation significantly increased TGF-β1 secretion from eosinophil-like cells. Given that TGF-β1 is a well-established inducer of EMT in nasal epithelial tissue, it is plausible that IL-5–activated eosinophils may promote EMT via paracrine release of TGF-β1 [18,19]. Although this mechanism has yet to be directly validated, it provides a potential explanation for the epithelial changes observed in our co-culture system.
Nevertheless, several limitations of this study should be acknowledged. First, although we observed increased activation of the IL-5Rα/JAK2/STAT5 pathway in ECRSwNP tissues and in IL-5–stimulated Diff. HL-60 cells, we did not perform pathway-blocking experiments to determine whether inhibition of individual signaling components attenuates EMT. As a result, we were unable to conclusively establish a causal role for this pathway in EMT induction and instead demonstrated an association between IL-5Rα signaling and EMT-related changes in HNECs. Second, the eosinophil-like cells used in this study were differentiated from HL-60 cells using sodium butyrate and therefore do not fully represent mature human eosinophils with respect to phenotype and function. In particular, these cells may not recapitulate mature eosinophils in terms of granule protein composition, receptor expression, or degranulation dynamics, which may limit the physiological translatability of the co-culture findings. Finally, given the in vitro nature of this study, inherent limitations exist in extrapolating these results to the in vivo setting, where the tissue microenvironment is considerably more complex.
In light of these limitations, future studies are warranted to validate our findings in vivo, to determine whether EMT is mediated through the IL-5/JAK2/STAT5 signaling pathway using targeted inhibition approaches, and to confirm these effects using primary human eosinophils rather than eosinophil-like cells derived from HL-60.
We anticipate that such investigations will provide deeper insight into the role of IL-5Rα–mediated signaling in EMT-driven tissue remodeling in ECRSwNP and help clarify its potential therapeutic implications.
Supplementary Materials
The online-only Data Supplement is available with this article at https://doi.org/10.18787/jr.2025.00065.
Correlation between tissue IL-5Rα expression and EMT markers across individual patient samples. A: Correlation analysis between IL-5Rα and E-cadherin. Scatter plots illustrate the relationship between the percentage of IL-5Rα-positive area (X-axis) and E-cadherin-positive area (Y-axis) in nasal tissues from healthy controls (green, n=12), non-ECRSwNP patients (purple, n=12), and ECRSwNP patients (orange, n=12). Spearman’s rank correlation analysis showed a significant inverse association when all samples were pooled (overall ρ=-0.946, p<0.001; n=36). In addition, a significant negative correlation was observed particularly in the ECRSwNP group (ρ=-0.839, p<0.001; n=12), supporting an association between higher IL-5Rα positivity and reduced E-cadherin positivity in this phenotype. B: Correlation analysis between IL-5Rα and vimentin. Scatter plots showing the correlation between the percentage of IL-5Rα-positive area (X-axis) and vimentin-positive area (Y-axis) in the same study cohorts (n=12 per group). Spearman’s rank correlation analysis showed a significant positive association when all samples were pooled (overall ρ=0.915, p<0.001; n=36). In addition, a significant positive correlation was observed in the ECRSwNP group (ρ=0.882, p=0.001; n=12), supporting an association between higher IL-5Rα positivity and increased vimentin positivity in this phenotype. Each data point represents an individual patient sample (n=36). Statistical significance was determined using Spearman’s rank correlation coefficient (ρ). These analyses provide supportive human evidence for the proposed mechanistic link between IL-5Rα-mediated signaling and EMT in ECRSwNP. ECRSwNP, eosinophilic chronic rhinosinusitis with nasal polyps; IL-5Rα, interleukin-5 receptor α; EMT, epithelial–mesenchymal transition.
Additional validation of the eosinophil-like phenotype in butyric acid–differentiated HL-60 cells. A: Relative mRNA expression of CCR3 in undifferentiated HL-60 cells (HL-60), butyric acid–differentiated eosinophil-like HL-60 cells (Diff. HL-60), and differentiated HL-60 cells further stimulated with IL-5 (Diff. HL-60 + IL-5). Expression levels were analyzed by qRT-PCR and normalized to GAPDH. B and C: Secretion of eosinophil-associated mediators from differentiated HL-60 cells in response to IL-5 stimulation. Butyric acid–differentiated HL-60 cells were treated with vehicle or 10 ng/mL IL-5 for 24 hours, and culture supernatants were analyzed by ELISA for EPO (B) and IL-13 (C). Results are expressed as the mean±SEM of three independent experiments (n=3). Statistical significance was determined by one-way ANOVA followed by Tukey’s post hoc test. *p<0.05. CCR3, C–C chemokine receptor 3; IL-5, interleukin-5; EPO, eosinophil peroxidase; IL-13, interleukin-13; qRT-PCR, quantitative real-time polymerase chain reaction; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; ELISA, enzyme-linked immunosorbent assay; SEM, standard error of the mean; ANOVA, analysis of variance.
Notes
Availability of Data and Material
The datasets generated or analyzed during the study are available from the corresponding author on reasonable request.
Conflicts of Interest
The authors have no potential conflicts of interest to disclose.
Author Contributions
Conceptualization: Dong Chang Lee. Data curation: Hyunsu Choi, Jeong-Min Oh. Formal analysis: Hyunsu Choi, Jeong-Min Oh. Funding acquisition: Hosung Choi. Methodology: Hyunsu Choi, Sung Won Kim. Project administration: Dong Chang Lee. Resources: Soo Whan Kim, Byung Guk Kim. Software: Hyunsu Choi, Sung Won Kim, Soo Whan Kim. Supervision: Jin Hee Cho. Validation: Byung Guk Kim, Jin Hee Cho. Visualization: Hyunsu Choi, Jeong-Min Oh. Writing—original draft: Hosung Choi. Writing—review & editing: Hosung Choi, Dong Chang Lee.
Funding Statement
This paper was supported by funding from the Korean Rhinologic Society in 2025.
Acknowledgments
None