Asthma and Inflammation Transcriptionally Upregulate Aryl hydrocarbon receptor in Airway Smooth Muscle via p38/JNK-AP1 Signaling

Abstract

Background and Purpose:

The aryl hydrocarbon receptor (AhR) is a ligand-activated transcription factor which maintains cellular homeostasis. AhR in airway fibroblast, epithelial and immune cells inhibit inflammatory responses. Nevertheless, its expression and role in airway smooth muscle (ASM), an airway structural cell indispensable in asthma pathophysiology, are obscure. This study uncovers the AhR’s expression, underlying mechanisms, and activity in ASM during inflammation and asthma.

Experimental Approach:

Cultured primary human non-asthmatic and asthmatic ASM cells were treated with TNFα/IL-13, with/without pharmacological inhibitors targeting PI3K, p38MAPK, JNK, NFkB, and AP1. AhR expression was analyzed using RNA-sequencing, confocal, qPCR, and immunoblotting. AP1 specific role was confirmed using c-JUN (AP1) siRNA and ChIP-qPCR. AhR activity was determined by Nanoluciferase in AhR agonist treated cells.

Key Results:

We found ubiquitous expression of AhR in ASM with upregulation in asthmatic ASM. TNFα increased AhR expression, whereas IL-13 did not alter its expression. Furthermore, p38 and JNK inhibition significantly reduced AhR expression with TNFα exposure, whereas PI3K inhibition had no effect. AP1 inhibition and c-JUN knockdown significantly downregulated AhR expression, while NFkB inhibition showed no effect. TNFα promoted c-JUN binding to AhR promoter and increased AhR mRNA expression. Additionally, AhR agonists significantly increased the xenobiotic response element (XRE) activity, CYP1B1, and AhR nuclear expression. TNFα exposure reduced XRE activity and AhR nuclear expression.

Conclusion and Implications:

Our findings suggest inflammation and asthma transcriptionally upregulates AhR through p38/JNK-AP1 pathway in ASM, identifying a potential therapeutic target for modulating AhR and its downstream effects in asthma.

Introduction

Asthma is one of the most common chronic inflammatory diseases in the world characterized by airway inflammation, remodelling, and hyperresponsiveness, leading to various respiratory symptoms such as breathlessness, wheezing, and airflow obstruction (Prakash, 2013; M. I. Reza et al., 2024). In the United States there are around 25 million active cases of asthma with approximately 10.6 deaths per million (Data, 2021). These numbers indicate the severity of this chronic disease. In recent years, despite significant advances being made in asthma research, current therapeutics are unable to mitigate airway thickening and remodelling, thereby patients experience poor control. This indicates an urgent need for further in-depth studies in asthma pathophysiology, explicitly on airway remodelling. Recent studies have widely explored the involvement of airway smooth muscle cells (ASM) in asthma pathophysiology (Prakash, 2013). ASM cells are oriented in the airway circumference where they regulate airway tone, however their excess contraction results in reduction in airway lumen diameter, a major challenge in asthma (M. I. Reza et al., 2024), as it causes shortness of breath and wheezing. Moreover, the hypercontractile response of ASM facilitates irregular bronchoconstriction and blockage of airflow in response to small stimulus and causes airway hyperresponsiveness (Prakash, 2013). Besides ASM contribution in bronchoconstriction, it plays a paramount role in airway remodeling by increasing airway wall thickness (Ambhore et al., 2018; Balraj et al., 2024). Apart from proliferation, extracellular matrix production and deposition in ASM play a substantial role in the progression of airway remodeling (Ambhore et al., 2019).

It is well known that asthma is influenced by multiple factors including genetics, infection, diet, and environmental exposures (Baxi & Phipatanakul, 2010), with the immune response and subsequent regulatory mechanisms influencing pathophysiology, disease progression, and severity. Aryl hydrocarbon receptor (AhR) is a conserved ligand-activated transcription factor expressed in various cells and tissues (Kim et al., 2024; Opitz, Holfelder, Prentzell, & Trump, 2023). It serves as a sensor for environmental toxins, pollutants, host and microbial metabolites that controls cellular and physiological processes through complex transcriptional programs that are ligand- and cell-type specific (Poulain-Godefroy et al., 2020). In addition, AhR has many physiological and homeostatic functions in different tissues and organs, including development, pluripotency, proliferation, regeneration, and senescence (Claudia Rejano-Gordillo et al., 2022). Recent studies suggest that knockout of AhR may exacerbate inflammatory responses in lung diseases (Alessandrini et al., 2022), indicating anti-inflammatory properties of AhR in airway inflammation. Similarly, Pariano M et al demonstrated that AhR agonism prevents hypoxia-induced inflammation, restores immune homeostasis, and improves lung function (Pariano et al., 2023). These findings highlight the importance of AhR in airway disease. In addition, there are a few reports exploring the role of AhR in immune cells (Drew R Neavin, Duan Liu, Balmiki Ray, & Richard M Weinshilboum, 2018), fibroblasts (Shi et al., 2021), and airway epithelium (Alessandrini et al., 2022; Hu et al., 2021). However, AhR expression and its role in ASM per se especially during inflammation and asthma is unexplored. Therefore, in this study, we explored AhR expression in human ASM and underlying mechanisms which regulate AhR expression during inflammation and asthma. We selected tumor necrosis factor-α (TNFα) and interleukin-13 (IL-13) as representative cytokines based on their distinct roles in asthma pathogenesis. TNFα is a key mediator of T helper 1 (Th1)-type inflammation that is elevated in severe asthma and drives airway hyperresponsiveness and remodeling (Ford, Reza, Ruwanpathirana, Sathish, & Britt, 2025; Luo, Hu, Xu, & Dong, 2022). IL-13 represents the T helper 2 (Th2) cytokine that mediates IgE production and mucus hypersecretion- hallmark features of allergic asthma (Ford et al., 2025; Luo et al., 2022). This dual cytokine approach allows us to examine AhR regulation across both major inflammatory axes in asthma. We observed asthma and pro-inflammatory cytokine TNFα, but not IL-13, increased AhR expression through p38 mitogen-activated protein kinase (p38MAPK) and c-Jun N-terminal kinase (JNK) and subsequently their downstream transcription factor activator protein-1 (AP1) in ASM cells, indicating involvement of biased inflammatory signaling in regulating AhR expression. Accordingly, our study suggests inflammation and asthma transcriptionally upregulate AhR in ASM through p38/JNK and AP1 pathway.

Materials and methods

Chemicals, drugs and antibodies

AhR antibody was obtained from Thermo Scientific (Cat# MA1–514; Waltham, MA, USA, RRID:AB_2273723) and Cell Signaling Technologies (Cat# 83200S; Danvers, MA, USA, RRID:AB_2800011). α-SMA antibody was procured from Millipore Sigma (Cat# A2547; Burlington, MA, USA, RRID:AB_476701) and Cell Signaling Technologies (Cat# 19245; Danvers, MA, USA, RRID:AB_2734735). β-actin antibody was purchased from Applied Biological Materials (Cat# G043; Richmond, BC, Canada, RRID:AB_2631287). c-JUN and Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) antibodies were purchased from Cell Signaling Technologies (Cat# 9165S, RRID:AB_2130165 and 5174, RRID:AB_10622025; Danvers, MA, USA). E-Cadherin antibody was obtained from R and D biosystems (Cat# AF748; Minneapolis, MN, USA, RRID:AB_355568). Alexa flour secondary antibodies, Dulbecco’s modified eagle medium (DMEM/F12), fetal bovine serum (FBS), and antibiotic-antimycotic (AbAm), Dulbecco’s phosphate buffer saline (DPBS) was obtained from Thermo Scientific (Waltham, MA, USA). IRDye secondary antibodies were obtained from Li-Cor Biosciences (Lincoln, NE, USA). TNFα and IL-13 were procured from R&D systems (Minneapolis, MN, USA). SR11302, Wortmannin, SB203580, and SP 600125 were obtained from Bio-Techne Corporation (Minneapolis, MN, USA). 2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD) was obtained from AccuStandard (Cat# D-404N; New Haven, CT, USA). SN50 was procured from Santa Cruz Biotechnology, Inc (Dallas, TX, USA). XRE-pNL1.3[secNLuc] was a gift from Severine Degrelle & Thierry Fournier (Addgene plasmid # 182295). All other chemicals or antibodies were obtained from Sigma-Aldrich (St. Louis, MO, USA) unless otherwise specified.

Human tissue and ASM cells

Lung tissues and ASM cells were isolated from healthy and mild to moderate asthma patients visited for thoracic surgery at Mayo Clinic (Saint Marys Hospital, Rochester, MN) as previously reported.(Sathish et al., 2012) Approval was received from Mayo Clinic IRB. In brief, to isolate primary ASM cells, airway samples were epithelium denuded and then subjected to enzymatic digestion as per the manufacturer’s instructions (Worthington Biochemical). The isolated cells were cultured in DMEM/F12 medium containing 10% FBS and 1% AbAm and incubated at 37°C temperature, 5% CO2, and 95% air. For studies, ASM cells with maximum 5^th passage were used. The phenotype of ASM cells were periodically characterized by checking the expression of ASM specific markers such as caldesmon and α-SMA through immunoblotting. Additionally, to ensure that the cells are free from mycoplasma, periodically mycoplasma tests was performed through polymerase chain reaction (PCR) using kit (Cat #G238, Applied Biological Materials, Richmond, BC, Canada).

Treatments

Serum deprived (24h) human non-asthmatic and asthmatic ASM cells were exposed to pro-inflammatory cytokines TNFα (20 ng/mL)(M. I. Reza et al., 2024) and IL-13 (50 ng/mL)(M. I. Reza et al., 2024; Sathish et al., 2011) for 6, 12, 24, and 48h for expression studies. For mechanistic studies, serum deprived cells were pre-treated with either wortmannin (PI3K inhibitor: 50 nM)(Aravamudan et al., 2017), SB203580 (p38 inhibitor: 600nM)(Aravamudan et al., 2017), SP600125 (JNK inhibitor: 20 μM)(Moynihan et al., 2008), SR11302 (AP1 inhibitor 10 μM)(Aravamudan et al., 2017), and SN50 (nuclear factor kappa B, NFkB inhibitor: 20 μM) (Aravamudan et al., 2017; Sathish et al., 2011) for 2h with/without TNFα exposure for 24h.

siRNA transfection

c-JUN Knockdown was done using small interfering RNA (siRNA) as previously described(Ambhore et al., 2019; M. I. Reza, Syed, Singh, Husain, & Gayen, 2021). Briefly, both human non-asthmatic and asthmatic ASM cells were cultured on 6 well or 100 mm plate. Once ~70% confluent, cells were transfected using silencer pre-designed c-JUN siRNA (Cat# AM16708, 20 nM) and Lipofectamine 3000 (Thermo Scientific, Waltham, MA, USA)) in serum free DMEM-F12 media (without antibiotic). Following 6h of incubation, 20% FBS containing culture media (without antibiotic) was added and incubated for 48h. Subsequently, cells were serum deprived for 24h and treated with TNFα for 24h. Specific siRNA knockdown efficiency was verified by immunoblotting.

Laser capture microdissection

LCM was performed as previously established in our lab (Loganathan, Pandey, Ambhore, Borowicz, & Sathish, 2019). In brief, human lung tissue was fixed using Carnoy’s solution, paraffin embedded, and sectioned to 10 μm onto RNAse-free slides. Following deparaffinization with xylene and ethanol, ASM regions (0.3 or 1.0 mm²) were identified by reference to corresponding H&E-stained sections, microdissected using a Zeiss AxioImager Z1 PALM Microbeam LCM system (Zeiss, Thornwood, NY, USA), and catapulted into RNAse-free microtubes containing 0.5 ml PicoPure^® RNA extraction buffer. Total ribonucleic acid (RNA) was isolated using the PicoPure^® RNA Isolation Kit (Thermo Scientific, Waltham, MA, USA), and subsequently complementary DNA (cDNA) synthesized. Quantitative PCR (qPCR) was utilized to measure AhR messenger RNA (mRNA) expression.

RNA‑Sequencing (RNA-Seq) transcriptome profiling

The RNA-Seq data was analyzed as reported previously(M. I. Reza et al., 2024). In brief, RNA-Seq data of non-asthmatic and asthmatic human ASM cells was obtained from our own online deposited NCBI database (GEO GSE119579)(Fong et al., 2018). The data was processed and analyzed at Center for computationally assisted science and technology (CCAST), North Dakota State University. Quality control (QC) of data was done using FastQC v0.11.8 and MultiQC v1.9. Next, reads were mapped using STAR aligner v2.7.5a. Furthermore, to quantify the reads and obtain raw counts, we used quantMode gene counts flag. Subsequently, raw counts were transformed to counts per million (CPM) using edgeR and genes with CPM below 0.5 in more than 50% of samples were filtered out (considered low-expression). Additionally, we performed post-mapping QC with MultiQC and edgeR.

Immunofluorescence

Immunofluorescence was performed as previously reported(Ambhore et al., 2024; M. I. Reza et al., 2024; M. I. Reza, Syed, Kumariya, et al., 2021). In brief, for lung tissues 6 μm-thick sections were placed on slides, baked at 56°C for 2h, deparaffinized with xylene, and gradually rehydrated with ethanol (from 100% to 70%). Furthermore, we performed antigen retrieval using citrate buffer (pH 6.0), and sections were permeabilized with 0.1% Triton X-100. Subsequently, to reduce non-specific binding, sections were blocked with 10% goat serum and incubated overnight at 4°C with E-Cadherin (diluted 1:100), αSMA (diluted 1:100), with or without AhR antibody (Cat# MA1–514, diluted 1:100). The next day, sections were washed with PBS+0.1% Tween (PBS-T), and respective species-specific Alexa flours (AF-647 for E-cadherin, AF-568 for AhR, and AF-488 for αSMA, each diluted 1:200) were applied for 2 hours at room temperature. After additional washing with PBS-T, sections were counterstained with 4’,6-diamidino-2-phenylindole (DAPI) and visualized on a Zeiss-LSM900 with Airyscan2 confocal microscope (Oberkochen, Baden-Württemberg, Germany).

For immunofluorescence on ASM cells, fixation was done with 4% paraformaldehyde, followed by permeabilization with 0.1% Triton X-100 and blocking with 10% goat serum. Further, cells were incubated overnight at 4°C with or without AhR antibody (1:100). The next day, cells were rinsed with PBS-T, followed by staining with Alexa Fluor-568 (1:200) and Phalloidin for 1 hour at room temperature. After final washes with PBS-T, cells were counterstained with DAPI gel and observed using the Zeiss-LSM900 with Airyscan2 confocal microscope. All the fluorescence images were captured under consistent conditions. To quantify AhR subcellular localization, nuclear and cytoplasmic regions of interest (ROIs) were manually delineated by circular selection (3 cells/field, minimum 30 cells/group) using Zeiss Zen 3.0 software. Mean fluorescence intensity (MFI) values for each ROI were recorded and normalized to the respective area (MFI/μm²).

Immunoblotting

We performed immunoblotting on non-asthmatic and asthmatic ASM cells following the previously established protocol(Balraj et al., 2024; Singh et al., 2021). In brief, cells were scrapped in PBS and centrifuged. Obtained cell pellets were dissolved in 1X cell lysis buffer (+protease and phosphatase inhibitors cocktail), vortexed three times with 10 minutes interval between each vortex on ice, and centrifuged to obtain cell lysates. The total protein in lysates was quantified by DC Protein Assay Kit (BioRad, Hercules, CA, USA). 30–40 μg protein was loaded and separated on 4–15% gradient gels (Criterion Gel System; BioRad, Hercules, CA, USA). Further, rapid transfer technique (Trans-Blot Turbo; BioRad, Hercules, CA, USA) was used to transfer the protein from gel to polyvinylidene difluoride (PVDF) membrane (0.22-μm). Next, membrane was dried and incubated with 5% BSA at room temperature to reduce the non-specific binding. Membrane was incubated overnight with gentle rocking using species specific primary antibodies at 4°C, antibody was recovered, and the membrane was washed three times (10 minutes each wash) with Tris-buffered saline+0.1% Tween (TBS-T). Subsequently, membrane was incubated with respective species specific IRDye secondary antibodies (diluted 1:10,000) for 1h at room temperature. After additional washing with TBS-T, membrane was scanned using Li-Cor Odyssey XL System (Lincoln, NE, USA). To quantify the bands, densitometry analysis was done using Image Studio v.5.2 software. We normalized the protein of interest intensity with housekeeping protein (β-actin or GAPDH) intensity. Fold change was calculated relative to the vehicle-treated group, which was set as 1.

Real time quantitative polymerase chain reaction (RT-qPCR)

Total RNA was isolated following manufacturer’s protocol by Quick-RNA MiniPrep Kit (Zymo Research, Irvine, CA, USA) and quantified using Synergy HTX Multi-Mode Plate Reader (BioTek Instruments, Winooski, VT, USA). To avoid genomic DNA contamination, DNAse I treatment was performed. Further, 1 μg RNA was reverse transcribed to complementary DNA (cDNA) using OneScript plus cDNA Synthesis Kit (Applied Biological Materials, Richmond, BC, Canada). Finally, cDNA was loaded on the 96 well plate (Thermo Scientific, Waltham, MA, USA) with respective primers (IDT) and BlasTaq 2X qPCR MasterMix (Applied Biological Materials, Richmond, BC, Canada), and qPCR was run on QuantStudio 3 (Thermo Scientific, Waltham, MA, USA). Relative expression of mRNA was normalized to the housekeeping gene S16. Fold changes were calculated using the ΔΔCt method, with vehicle-treated cells used as the reference condition and set as 1. The primers used for qPCR are listed in table 1.

Table 1.

List of qPCR primers.

Gene	Primer
AhR	Forward: GTCGTCTAAGGTGTCTGCTGGA Reverse: CGCAAACAAAGCCAACTGAGGTG
CYP1A1	Forward: GATTGAGCACTGTCAGCAGAAGC Reverse: ATGAGGCTCCAGGAGATAGCAG
CYP1B1	Forward: GCCACTATCACTGACATCTTCGG Reverse: CACGACCTGATCCAATTCTGCC
S16	Forward: GCTTTCCTTTTCCGGTTGCG Reverse: CACGGATGTCTACACCAGC

Chromatin-Immunoprecipitation-qPCR (ChIP-qPCR)

Chromatin was isolated using Pierce Chromatin Pre-Module (Cat# 26158, Thermo Scientific, Waltham, MA, USA) as per the manufacturer’s instruction. Cells were crosslinked by 1% formaldehyde, lysed, and digested by Micrococcal nuclease (MNase) digestion. Extracted chromatin size was verified by agarose gel electrophoresis. Furthermore, ChIP was performed using High-Sensitivity ChIP Kit (Cat #ab185913, Abcam, Cambridge, UK). In brief, chromatin was immunoprecipitated with c-JUN antibody, and subsequently crosslinking was reversed, DNA was released, and purified. Further, the purified ChIP DNA was processed for qPCR (as described in RT-qPCR section) using primers targeting AhR promoter regions (Forward: CCAGCATGAGGTATCTAAGGATTTA; Reverse: CCAGTAACATGTGGAAGAGAGTAA). The Ct value of AhR was normalized to IgG and the fold enrichment was calculated as described by manufacturer’s protocol.

Plasmid transfection and luciferase assay

Bacterial agar stab of XRE-pNL1.3[secNLuc] was processed as described(Ambhore et al., 2019; Degrelle, Ferecatu, & Fournier, 2022). In brief, the stab was streaked onto agar plate, single colonies harvested and incubated in LB broth overnight. Next, the plasmid was isolated and purified using ZymoPure II Plasmid MidiPrep Kit (Cat# D4201, Irvine, CA, USA). XRE-pNL1.3[secNLuc] was verified by gel electrophoresis. Furthermore, both non-asthmatic and asthmatic ASM cells were cultured in 96 well plate (5000 cells per well). At 50% confluency, cells were transiently transfected with purified 200 ng/well XRE-pNL1.3[secNLuc] plasmid using Lipofectamine 3000 (Thermo Scientific, Waltham, MA, USA) in serum free DMEM-F12 media (without antibiotic). Following 6h of incubation, 20% FBS containing culture media (without antibiotic) was added and incubated for 24h. Subsequently, cells were serum deprived for 24h and treated with either AhR agonists TCDD (10 nM)(Blankenship & Matsumura, 1997) and 6-formylindolo[3,2-b]carbazole (FICZ: 10 nM)(Manzella et al., 2018) +/− AhR antagonist CH223191 (10 μM)(Manzella et al., 2018) with/without TNFα exposure for 24h. Further, secreted Nano luciferase was measured as per the instructions provided by Nano-Glo^® Luciferase Assay System (Cat# N1110, Promega, Madison, WI, USA). Cells were placed on orbital shaker at 100 RPM for 2 minutes to remove concentration gradients. Next, 75 μL cell supernatant was dispensed in another opaque 96-well plate and one volume (75 μL) of Nano-Glo^® Luciferase assay reagent was added to each well. After 3 minutes of incubation, luminescence was recorded using Synergy HTX Multi-Mode Plate Reader (BioTek Instruments, Winooski, VT, USA). The luminescence was normalized with vehicle and presented as % xenobiotic response element (XRE) activity.

Statistical Analysis

Shapiro-Wilk normality test was used to test normal distribution. Subsequently, for comparing two groups, parametric unpaired Student’s T-test was applied and for more than two groups, one-way ANOVA followed by Tukey post hoc test was applied. Data was considered statistical significance with P values less than 0.05. All values are expressed as means ± SEM.

Nomenclature of Targets and Ligands

Key protein targets and ligands in this article are hyperlinked to corresponding entries in http://www.guidetopharmacology.org, and are permanently archived in the Concise Guide to PHARMACOLOGY 2023/24 (Alexander, Fabbro, Kelly, Mathie, Peters, Veale, Armstrong, Faccenda, Harding, Davies, Annett, et al., 2023; Alexander, Fabbro, Kelly, Mathie, Peters, Veale, Armstrong, Faccenda, Harding, Davies, Beuve, et al., 2023; Alexander, Kelly, et al., 2023).

Results

AhR is abundantly expressed in human ASM

To assess AhR expression in human ASM, we performed immunofluorescence of AhR in fixed human lung tissue sections. ASM layer was identified with α-SMA and epithelium layer with E-cadherin. Additionally, we also determined AhR expression in enzymatically isolated ASM cells. Interestingly, we found notable expression of AhR in ASM and epithelium layer of human airway tissue (Figure 1A) and explicitly in ASM cells (Figure 1B). To gain further insight into subcellular localization of AhR in baseline condition, we quantified its nuclear and cytoplasmic expression using immunofluorescence images in ASM cells. As shown in Figure 1C, the MFI of AhR was significantly (p=0.03) higher in the cytoplasm compared to the nucleus showing a more pronounced cytoplasmic (inactive) accumulation.

Figure 1.

Immunofluorescence showing baseline AhR expression in human (A) lung sections and (B) ASM cells. (C) Mean fluorescence intensity (minimum 30 individual cells per group) of AhR protein in nucleus and cytoplasm of ASM cells. Images were quantified using Zeiss Zen 3.0 software as described in Methods. α-SMA (AF-488) shows smooth muscle specific colocalization. Arrow indicates AhR in ASM. Scale bar: 10 μm. Inset 2 μm. Data were analyzed using unpaired Student’s T-test. Data represented mean ± SEM (N = 6 each, non-asthmatic 3M/3F, independent donor samples); *p<0.05.

AhR is upregulated in asthmatic ASM

Here, we determined AhR expression in asthmatic samples. We performed the transcriptomic analysis of RNA sequencing data of non-asthmatic and asthmatic human ASM cells (NCBI GEO GSE119579). It showed AhR pathway genes [AhR, AhR nuclear translocator (ARNT), ARNT2, AhR nuclear translocator-like protein (ARNTL), and ARNTL2, Cytochrome P450 enzymes CYP1A1, and CYP1B1 are expressed in ASM, with slightly greater expression in some asthmatic samples (Figure 2A and B). Although transcriptomic analysis reflected marginally upregulated expression of a number of AhR pathway genes in asthmatic ASM cells, the current study specifically focused on AhR due to its key regulatory role and our overall objective of ascertaining its role in airway inflammation. The other genes’ expression profiles may be secondary changes or parallel regulatory mechanisms, which are worth investigating in the future. Furthermore, we verified AhR mRNA and protein expression in non-asthmatic and asthmatic ASM cells. We observed significant upregulation of AhR mRNA (p=0.0024) and protein (p<0.001) in asthmatic ASM cells (Figure 2C to E). Separately, we isolated the ASM layer of lung tissue using LCM and measured the AhR mRNA expression. As speculated, we observed significantly higher expression of AhR mRNA (p=0.0178) in ASM layer of asthmatic lung tissue (Figure 2F).

Figure 2.

(A) Heatmap and (B) bar graph of RNA Seq showing normalized read counts of AhR and its binding partners in non-asthmatic and asthmatic human ASM cells. (C) mRNA and (D and E) protein expression of AhR in non-asthmatic and asthmatic ASM cells. (F) LCM-assisted AhR mRNA expression in isolated ASM layer from human lung tissue section of non-asthmatic and asthmatic samples. Fold change in mRNA and protein expression was calculated relative to vehicle, using S16 and β-actin as internal controls for qPCR and immunoblotting, respectively. Data were analyzed using unpaired Student’s T-test. Data represented mean ± SEM (N = 6–9 each, non-asthmatic 3–5M/3–4F, asthmatic 4–5M/2–4F, independent donor samples); *p<0.05, **p<0.01, ***p<0.001.

Although AhR upregulation in asthmatic and inflamed ASM cells is observed, its function in airway hyperresponsiveness, ASM contractility, proliferation, and remodeling remains to be established. Mechanistically, AhR activation has been shown to control proliferation, contractile protein expression, and inflammatory reactions in various cell types, such as fibroblasts and vascular smooth muscle cells (Lehmann et al., 2011; C. Rejano-Gordillo et al., 2022). Our recent findings show AhR activation diminishes mitogen-induced ASM proliferation and extracellular matrix (ECM) (M. Reza et al., 2023; M. Reza et al., 2024), possibly by inhibiting pro-remodeling pathways such as TGF-β/SMAD/ERK (Ojiaku, Yoo, & Panettieri, 2017). AhR also modulates cytoskeletal dynamics and calcium homeostasis (Sondermann et al., 2023), which are critical for ASM contractility. As TNFα is a potent inducer of ASM inflammation and remodeling, and that we observed its specific role in upregulating AhR, it is plausible that TNFα-induced AhR expression serves as a compensatory or regulatory mechanism to limit excessive ASM proliferation or ECM during inflammation. Altogether, these findings support a model where AhR, when appropriately activated, could negatively regulate airway remodeling and hyperresponsiveness. However, further mechanistic studies are required to determine whether AhR can directly modulate ASM contractility and AHR in vivo.

TNFα, but not IL13, upregulates AhR in non-asthmatic ASM cells

We determined AhR mRNA expression in non-asthmatic ASM cells following pro-inflammatory cytokine exposure at different time points. As asthma involves both Th1 and Th2 inflammation, we used TNFα (Th1-associated) and IL13 (Th2-associated) exposures to model ASM responses during Th1 and Th2 inflammation, respectively. qPCR analysis showed time dependent (6h, 12h, 24h, and 48h) increase in AhR mRNA expression following TNFα treatment with maximum increase was seen at 24h (P<0.001). In contrast, IL-13 exposure did not significantly alter AhR mRNA levels at any time point. Similarly, immunoblotting demonstrated significantly enhanced AhR protein expression following 24h (p=0.0015) and 48h (p=0.0282) of TNFα, while no significant changes were observed with IL13 exposure (Figure 3B and C). These data confirmed that TNFα, but not IL13, upregulates AhR in non-asthmatic ASM cells.

Figure 3.

(A) AhR mRNA expression in non-asthmatic ASM cells during pro-inflammatory cytokines (TNFα: 20 ng/mL and IL13: 50 ng/mL) exposure at different time points (6h, 12h, 24h, and 48h). AhR protein expression in non-asthmatic ASM cells during pro-inflammatory cytokines exposure at (B) 24h and (C) 48h. Fold change in mRNA and protein expression was calculated relative to vehicle, using S16 and β-actin as internal controls for qPCR and immunoblotting, respectively. Data were analyzed using one-way ANOVA followed by Tukey’s post hoc test. Data represented mean ± SEM (N = 6–9 each, non-asthmatic 3–5M/3–4F, asthmatic 4–5M/2–4F, independent donor samples); *p<0.05, **p<0.01, ***p<0.001.

p38 and JNK, not PI3K are involved in TNFα mediated upregulation of AhR in non-asthmatic and asthmatic ASM cells

To understand the mechanisms of TNFα-mediated AhR upregulation, we explored the AhR expression following PI3K, p38, and JNK inhibition in the presence of TNFα in ASM cells. Interestingly, p38 and JNK inhibition significantly blunted AhR expression in non-asthmatic (p=0.003, Figure 4A and D) and asthmatic (p=0.002 and 0.036, Figure 4B and E) ASM cells under baseline conditions. Notably, p38 and JNK inhibition significantly blocked the TNFα effect on AhR expression (p=0.02 and p<0.001, Figure 4C and D). Conversely, PI3K inhibition did not alter AhR expression in either non-asthmatic (+/− TNFα, Figure 4A and C) or asthmatic ASM cells (Figure 4B). These data indicate TNFα increases AhR through p38 and JNK pathways in both non-asthmatic and asthmatic ASM cells.

Figure 4.

AhR protein expression following PI3K (Wortmannin: 50 nM) and p38 (SB203580 600 nM) inhibition in (A) non-asthmatic and (B) asthmatic ASM cells. (C) AhR protein expression following PI3K and p38 inhibition with TNFα exposure (20 ng/mL for 24h) in non-asthmatic ASM cells. AhR protein expression following JNK inhibition (SP 600125: 20 μM) in (D) non-asthmatic ASM cells with TNFα exposure (20 ng/mL for 24h) and (E) asthmatic ASM cells. Fold change in protein expression was calculated relative to vehicle, using β-actin as internal control. Data were analyzed using unpaired Student’s T-test or one-way ANOVA followed by Tukey’s post hoc test. Data represented mean ± SEM (N = 6–9 each, non-asthmatic 3–5M/3–4F, asthmatic 4–5M/2–4F, independent donor samples); *p<0.05, **p<0.01, ***p<0.001.

Interestingly, we observed that JNK inhibitor selectively reduced basal AhR expression in asthmatic ASM cells, but not in non-asthmatic cells. This selective sensitivity might reflect that basal JNK signaling is more pronounced in asthmatic ASM, possibly due to the pro-inflammatory and oxidative stress environment in asthmatic airways (Alrashdan et al., 2012; Lee et al., 2006). Whereas, comparatively inactive JNK signaling in non-asthmatic ASM cells may be the underlying reason for unchanged AhR expression with JNK inhibition. These findings highlight the context-dependent AhR regulation.

AP1, not NFkB is important in TNFα mediated upregulation of AhR in non-asthmatic and asthmatic ASM cells

p38 and JNK regulate transcription factors NFkB and AP1 expressions to modulate the expression of target genes. AP1 inhibition blunted AhR expression in non-asthmatic (p=0.056, Figure 5A) and asthmatic (p=0.013, Figure 5B) ASM cells under baseline (no treatment) conditions. Additionally, AP1 inhibition significantly blocked the TNFα effect on AhR expression (p=0.02, Figure 5C). Conversely, NFkB inhibition did not alter AhR expression in either non-asthmatic (in the presence of TNFα, Figure 5A and C) or asthmatic (Figure 5B) ASM cells. Furthermore, we knocked down the AP1 major subunit c-JUN using respective siRNA in ASM cells. Following the knockdown confirmation (Figure 5D), AhR expression was determined in the presence of TNFα. Similar to pharmacological inhibition, molecular inhibition of AP1 by c-JUN siRNA also significantly downregulated the AhR expression in both non-asthmatic (p=0.0028, Figure 5E) and asthmatic (p=0.0031, Figure 5F) ASM cells. Additionally, c-JUN knockdown also blunted the effect of TNFα on AhR expression (p=0.009) in non-asthmatic ASM cells (Figure 5E). These data indicate TNFα increases AhR through p38 and JNK mediated upregulation of AP1 pathways in both non-asthmatic and asthmatic ASM cells.

Figure 5.

AhR protein expression following NFkB (SN50: 20 μM) and AP1 (SR11302: 10 μM) inhibition in (A) non-asthmatic and (B) asthmatic ASM cells. (C) AhR protein expression following NFkB and AP1 inhibition with TNFα exposure (20 ng/mL for 24h) in non-asthmatic ASM cells. (D) c-JUN knockdown confirmation (20 nM c-JUN siRNA for 48h). AhR protein expression in c-JUN knockdown (E) non-asthmatic ASM cells with TNFα exposure (20 ng/mL for 24h) and (F) asthmatic ASM cells. Fold change in protein expression was calculated relative to vehicle, using β-actin as internal control. Data were analyzed using unpaired Student’s T-test or one-way ANOVA followed by Tukey’s post hoc test. Data represented mean ± SEM (N = 6–9 each, non-asthmatic 3–5M/3–4F, asthmatic 4–5M/2–4F, independent donor samples); *p<0.05, **p<0.01, ***p<0.001.

AP1 positively regulates AhR transcription in ASM cells

We next elucidated AP1 transcriptional regulation of AhR. Firstly, we performed sequence analysis of the promoter region up to 10 KB upstream of initiation codon ATG on AhR gene for putative binding sites. Interestingly, AhR promoter region demonstrated multiple putative binding sites for AP1 (Figure 6A) and NFκB (supplementary figure S2). However, functional validation using specific inhibitors demonstrated that only AP1 inhibition significantly reduced AhR expression, while NFκB inhibition had no effect (Figure 5), indicating that AP1, but not NFκB, is a key regulator of AhR expression in ASM cells under inflammatory conditions. Accordingly, we verified if AP1 interacts with AhR promoter region. IgG (negative control) showed no binding while RNA polymerase (positive control) showed significant binding to the GPADH promoter (Figure 6B). Interestingly, we observed significant increase (p=0.002) in c-JUN binding to AhR promoter in TNFα exposed ASM cells (Figure 6C); however, binding was reduced by AP1 inhibition. Furthermore, we verified AhR mRNA expression following TNFα and AP1 inhibition to elucidate the effect of AP1 binding on AhR transcription. As previously observed, TNFα exposure significantly increased (p<0.001) AhR mRNA expression (Figure 6D). Interestingly, AP1 inhibition significantly reduced (p=0.0017) AhR mRNA expression (Figure 6D). Moreover, c-JUN knockdown significantly downregulated (p=0.01) AhR mRNA (Figure 6E). However, TNFα exposure in the c-JUN knockdown ASM cells failed to induce AhR mRNA expression (Figure 6E). Together, these data demonstrated AP1 interacts with AhR promoter and positively regulates its transcription.

Figure 6.

(A) Locations of transcription factor AP1 binding sites on AhR promoter region. Locations are distances upstream of the initiation codon (ATG). ChIP-qPCR of (B) RNA polymerase binding to GAPDH promoter region and (C) AP1 binding to AhR promoter region. AP1 transcriptional regulation of AhR was confirmed by determining AhR mRNA expression in TNFα (20 ng/mL for 24h) and AP1 inhibitor (SR11302: 10 μM for 24h) treated (D) normal and (E) c-JUN knockdown (20 nM c-JUN siRNA for 48h) ASM cells. Fold change in mRNA expression was calculated relative to vehicle, using S16 as internal control for qPCR. Data were analyzed using unpaired Student’s T-test or one-way ANOVA followed by Tukey’s post hoc test. Data represented mean ± SEM (N = 3–6 each, non-asthmatic 1–3M/2–3F, independent donor samples); **p<0.01, **p<0.01, ***p<0.001.

TNFα does not alter AhR activity in ASM cells

Recent studies have identified a diverse array of endogenous and exogenous compounds capable of modulating AhR signaling. For instance, aryl‐containing metabolite like 1‐aminopyrene (Miao et al., 2020) and dietary components such as flavonoids (e.g., quercetin, kaempferol) and tryptophan derivatives (e.g., kynurenine) exhibit AhR agonist properties (Chen, Wang, Fu, Yin, & Xu, 2023; Kaiser, Parker, & Hamrick, 2020). In the context of asthma, these ligands may influence ASM function by altering inflammatory responses or remodeling pathways. In this study, we used well known AhR activators TCDD and FICZ. We transfected the ASM cells with XRE-pNL1.3 vector bearing secNluc Tag and conducted nanoluciferase assay. We found AhR agonists TCDD and FICZ significantly increased the XRE activity in both non-asthmatic (p=0.0011 and 0.0057, Figure 7A) and asthmatic (p<0.001, Figure 7B) ASM cells, with more pronounced effect in asthmatics. On the other hand, AhR antagonist CH-223191 abrogated the TCDD and FICZ effect on XRE activity (Figure A and B) in both cell types. Interestingly, TNFα did not increase XRE activity, indeed it produced slight reduction (p=0.27, Figure 7C). Besides, TCDD and FICZ significantly increased XRE activity even in the presence of TNFα (p<0.001, Figure 7C). As cytoplasmic AhR is inactive while nuclear AhR is active, we visualized AhR nuclear vs cytoplasmic localization by Airscan confocal microscopy and calculated the ratio of nuclear to cytoplasmic AhR MFI. Interestingly, TCDD and FICZ showed significantly increased nuclear to cytoplasmic ratio (p=0.007 and 0.037, Figure 7D and E) while TNFα slightly reduced (p=0.089, Figure 7D and E). These data indicate TNFα only increases AhR cytoplasmic expression (Inactive AhR) but not activity. Further, we evaluated the mRNA expression of downstream genes, CYP1A1 and CYP1B1, in ASM cells treated with TNFα, TCDD, and FICZ. The AhR agonists significantly increased CYP1B1 expression (Figure 7F), while CYP1A1 (Figure S1) levels remained mostly unchanged. Interestingly, TNFα caused a modest increase in both CYP’s, but the differences were not statistically significant, suggesting while TNFα increases overall AhR levels in the cytoplasm, it does not activate the AhR downstream pathway, and any minor increase in CYP’s might be due to other alternative mechanisms such as NFκB or MAPK, previously demonstrated to regulate CYP’s during inflammation (Smerdova et al., 2014). These results highlight an important distinction between AhR expression and activation, particularly during inflammation. The cytoplasmic sequestration of AhR following TNFα treatment is likely a ligand-independent induction that is not followed by its activation. In the absence of ligand, AhR is retained within the cytoplasm and bound to chaperone proteins such as Hsp90 and is therefore transcriptionally inactive (Tsuji et al., 2014). Nuclear translocation also requires conformational change induced by specific agonists permitting its dissociation from the cytosolic complex (McGuire, Whitelaw, Pongratz, Gustafsson, & Poellinger, 1994; Tsuji et al., 2014). TNFα-induced cytosolic sequestration of AhR might therefore be a poised but inactive condition, perhaps sensitizing cells for subsequent ligand-mediated activation. This distinction between expression and activation is significant in the explanation of AhR’s role in asthma, where ligand availability may shape cellular responses.

Figure 7.

XRE luciferase activity following AhR activation by TCDD (10 nM for 24h) and FICZ (10 nM for 24h) +/− AhR antagonist CH-223191(10 μM for 24h) in (A) non-asthmatic and (B) asthmatic ASM cells. (C) XRE luciferase activity following TCDD and FICZ treatment +/− CH-223191 with TNFα exposure in non-asthmatic ASM cells. (D) representative images and (E) mean florescence intensity of nuclear to cytoplasmic expression of AhR following TNFα, TCDD, and FICZ exposure in non-asthmatic ASM cells. Images were quantified using Zeiss Zen 3.0 software. The nuclear-to-cytoplasmic ratio were derived from area-normalized MFI values of circular ROIs (≥30 cells/group) as described in methods. (F) CYP1B1 mRNA expression following TNFα, TCDD, and FICZ exposure in non-asthmatic ASM cells. Fold change in mRNA expression was calculated relative to vehicle, using S16 as internal control for qPCR. Data were analyzed using one-way ANOVA followed by Tukey’s post hoc test. Data represented mean ± SEM (N = 6 each, non-asthmatic 3M/3F, asthmatic 4M/2F, independent donor samples); *p<0.05, **p<0.01, ***p<0.001.

Discussion

Inflammation is an immune response to injury and is beneficial under normal circumstances(Laroux, 2004). However, an aberrant immune response to non-pathogenic stimuli leads to a chronic inflammatory response involved in the pathogenesis of various diseases(Bennett, Reeves, Billman, & Sturmberg, 2018; Laroux, 2004). For example, asthma involves chronic inflammatory responses and airway inflammation (Luo et al., 2022; Wang et al., 2018). In ASM, pro-inflammatory cytokines instigate the hypercontractile phenotype by upregulating the receptors for inflammatory mediators, and promoting ASM contraction, contributing to airway hyperresponsiveness (Wang et al., 2018). AhR has been widely explored in various inflammatory diseases(Drew R Neavin et al., 2018). Multiple studies have explored the role of AhR in modulating lung inflammation(Beamer & Shepherd, 2013), however, most studies focused on immune cells(Drew R Neavin et al., 2018), fibroblasts(Shi et al., 2021), and airway epithelium (Alessandrini et al., 2022; Hu et al., 2021). Although ASM is pivotal in airway inflammation and asthma and plays a critical role in the pathophysiology of this disease, to our knowledge there is no data about AhR expression or function in ASM. Therefore, in this study, we first show AhR expression, its underlying mechanisms, and activity during inflammation and asthma in human ASM cells.

To unveil if AhR is expressed in human ASM or not, we performed immunofluorescence of AhR in fixed human lung tissue sections and enzymatically isolated ASM cells. Interestingly, we found prominent expression of AhR in ASM which was comparable to its expression in neighbouring epithelium. After confirming the AhR baseline expression in human ASM, we demonstrated its expression during asthma and inflammation. Interestingly, we noted increased expression of AhR gene, mRNA, and protein expression in asthmatic ASM cells. Interestingly, we observed differential effect of pro-inflammatory cytokines on AhR expression. Broadly, Th1 cells mediated Type 1 (primarily by TNFα) and Th2 mediated type 2 (primarily by IL-13) inflammation balances immune response in various inflammatory diseases, including asthma(Luo et al., 2022). Specially in asthma, both type 1 and 2 inflammation contribute to ASM hypercontractility and remodelling.

In this study, we found that AhR expression in ASM cells is selectively upregulated by TNFα, but not by IL-13, suggesting that Th1 inflammation governs AhR induction not Th2. This aligns with the notion that type 2 immune response is viewed as compensatory mechanism of type 1 immune response, and vice versa (Bennett et al., 2018; Laroux, 2004). Explicitly, in asthma, Th2 mediated inflammation has been considered a primary player in disease pathogenesis (Luo et al., 2022). Thereby, we speculate that in asthmatic ASM cells, AhR upregulation by TNFα may serve as a compensatory or regulatory mechanism to mitigate excessive Th2-mediated responses. Indeed, several studies demonstrated that AhR activation alleviates lung inflammation by reducing Th2 cytokines, thus justifying our hypothesis (Luebke, Copeland, Daniels, Lambert, & Gilmour, 2001; Tarkowski, Kur, Nocuń, & Sitarek, 2010). Thus, TNFα-driven AhR upregulation may represent a protective adaptation in asthma. Moreover, future studies could explore the role of other Th1 cytokines, interferon gamma (IFNγ), and Th2 cytokines, IL4 and IL6, on regulating AhR expression in ASM during inflammation and asthma.

TNFα initiates inflammatory responses through TNFR1 and TNFR2, which ultimately promotes PI3K/Akt, p38 MAPK, and JNK pathways(Webster & Vucic, 2020; Y.-d. Wu & Zhou, 2010). Accordingly, we explored AhR expression following PI3K, p38, and JNK inhibition with TNFα stimuli. Our data indicated that TNFα increases AhR expression specifically through p38 and JNK pathways, but not PI3K in ASM cells. A recent study revealed that TNFα recruits PI3K/Akt through TNFR2 (Lu, Wang, & Liu, 2022) and this receptor is primarily expressed in immune and endothelial cells (Webster & Vucic, 2020) and marginally in smooth muscle cells(Choi, Dikalova, Stark, & Lamb, 2015; Lamb, Choi, Miller, & Stark, 2020). In addition, TNFα elicits its signalling majorly through TNFR1(Choi et al., 2015) which explains why the PI3K inhibition did not alter AhR expression. Further genetic knockdown/knockout approach is needed to validate involvement of TNFR1 vs TNFR2 in regulating AhR expression in ASM.

Furthermore, p38 and JNK regulate transcription factors NFkB and AP1 expression to modulate the expression of target genes(Y.-d. Wu & Zhou, 2010). We found AP1 inhibition blunts AhR expression in non-asthmatics with TNFα and also in asthmatic ASM cells at baseline conditions. Interestingly, the observed TNFα effect on AhR expression was unaffected by NFkB inhibition. Supporting our data on mesenchymal cells, a recent study demonstrated that Punicalagin, an AhR modulator upregulates AhR expression in macrophage through AP1 pathway rather than NFkB pathway. This effect was not altered by the inhibition of NFkB(Dai et al., 2024), which suggests a similar mechanisms observed in our study. Moreover, our study further confirmed the AP1 role in regulating AhR expression through multiple molecular approaches (siRNA and promoter assay). Both non-asthmatic and asthmatic ASM cells showed significant downregulation of AhR expression when c-JUN siRNA was used to inhibit AP1. These data further validate that inflammation induced ASM AhR expression occurs primarily through p38 and JNK mediated AP1 pathway.

AP1 is a transcription factor known to upregulate transcription of various genes(Z. Wu, Nicoll, & Ingham, 2021). In addition, previous studies reported that AhR promoter region contains binding sites for AP1 (Dai et al., 2024). Our own sequence analysis of AhR promoter region also demonstrated multiple putative binding sites for AP1. This implies that AP1 might facilitate the transcription of AhR gene thereby increasing its expression in response to TNFα. Consistent with the above, our ChIP-qPCR analysis showed binding of c-JUN, a subunit of AP1 to AhR promoter. Besides, AP1 inhibition and c-JUN knockdown reduced AhR mRNA expression in the presence of TNFα, confirming AP1 interacts with AhR promoter and thereby regulates positively its transcription.

Unliganded AhR is bound by the Heat Shock Protein 90 (Hsp90) chaperone complex in the cytoplasm and is functionally inactive(Tsuji et al., 2014). After ligand binding, AhR translocates to the nucleus, becomes active and binds to genomic DNA containing XRE to modulate transcription of target genes(McGuire et al., 1994; Tsuji et al., 2014). Here, we found AhR agonists (TCDD and FICZ) significantly increase XRE activity in both non-asthmatic and asthmatic ASM cells, with more pronounced effect in asthmatics. We assumed, since asthmatic ASM cells had more AhR expression than non-asthmatic, the AhR activation is more pronounced in asthmatic ASM cells. Similarly, AhR agonists increase AhR translocation to nucleus and CYP1B1 expression, suggesting increased activity. Surprisingly, we noted that TNFα slightly reduced XRE activity, promotes cytoplasmic AhR expression, and non-significant increase in both CYP1A1 and CYP1B1, further confirming that TNFα upregulates only AhR expression not its activity in ASM. Notably, the moderate induction of CYPs by TNFα may be via AhR-independent pathways, such as NFκB or MAPK signaling, previously demonstrated to regulate CYP expression during inflammation (Smerdova et al., 2014).

While our study elucidates the mechanism of increased AhR expression in ASM during inflammation and asthma, some limitations should be acknowledged. First, although we demonstrated increased AhR expression in the asthmatic human ASM cells and established the AP1/c-JUN pathway’s role through knockdown experiments, validation in mouse model of asthma, and ASM-specific AP1 or c-JUN knockout mice would strengthen these findings. Second, the translational relevance to clinical settings requires further exploration, interestingly, recent data indicate its involvement in human disease. For example, polymorphisms in the AhR gene have been associated with altered asthma and airway inflammation susceptibility (D. R. Neavin, D. Liu, B. Ray, & R. M. Weinshilboum, 2018). In addition, clinical trials of AhR-modulating drugs (e.g., tapinarof) illustrate the therapeutic potential in inflammatory disease (Nogueira, Rodrigues, Vender, & Torres, 2022). Future clinical trials can evaluate AhR agonists in asthma patients, particularly those with steroid-resistant or severe phenotypes, to determine if they can affect airway hyperresponsiveness and remodeling. Third, while we identified AP1 as a central regulator of AhR, other transcription factors or epigenetic mechanisms can be implicated in AhR dysregulation in asthma. Finally, the functional consequences of TNFα-induced AhR expression without activation must be investigated. These limitations could be overcome in future studies by utilizing conditional knockout mice or airway-specific interventions.

Conclusion

Our current study explores how AhR is regulated in ASM cells during inflammation and asthma. We found that AhR is constitutively present in human ASM cells with enhanced expression in asthmatic ASM. Additionally, TNFα, but not IL13, was responsible for increasing AhR levels in non-asthmatic ASM cells. Furthermore, we discovered that the TNFα-driven increase in AhR involves two key signaling pathways, p38 and JNK, but not PI3K. Here, the transcription factor AP1 plays a central role in this process, while NFκB does not. AP1 not only regulates AhR expression but also promotes its transcription in ASM cells. Interestingly, even though TNFα increases the levels of AhR, it does not affect its activity in ASM. Overall, our findings provide new insights into how AhR is regulated in ASM cells and its potential role in asthma and inflammation. These findings open the door to new approaches for targeting AhR and its pathways in the treatment of asthma and other airway diseases.

Abbreviations:

TNFα

Tumor necrosis factor-α

IL-13

Interleukin-13

PI3K

Phosphatidylinositol 3-kinase

p38MAPK

p38 mitogen-activated protein kinase

JNK

c-Jun N-terminal kinase

NFkB

Nuclear factor kappa B

AP1

Activator protein-1

ChIP-qPCR

Chromatin-immunoprecipitation