Uncovering a novel role of nAChRs in oxidative stress-mediated vascular dysfunction in COPD

Abstract

Tobacco smoke is the main risk factor for the development of chronic obstructive pulmonary disease (COPD). Despite current therapies alleviate symptoms there are limitations in the efficacy of treatments to curb its cardiovascular morbidities, particularly vascular dysfunction and the development of pulmonary hypertension. Our previous studies demonstrate that cigarette smoke directly contributes to pulmonary arterial dysfunction. Nevertheless, a further characterization of the molecular basis involved is needed for more effective targeted treatment. We have performed in vitro analysis with human pulmonary artery smooth muscle cells (hPASMC) challenged with cigarette smoke extract, and in vivo approaches of tobacco exposure in murine models and transgenic mice. Furthermore, we extended our analysis to include hPASMCs from COPD patients compared to non-COPD individuals, as well as pulmonary arteries from human tissue samples. These approaches allowed us to explore the molecular pathways contributing to the harmful effects from oxidative stress, calcium dysregulation and disruptions to the contractile machinery of pulmonary artery smooth muscle cells. Interestingly, these effects were triggered by the activation of nicotinic acetylcholine receptors (nAChRs) in these cells. Additionally, we demonstrated that nAChR antagonists or α7 nAChR deletion in a murine model effectively protected pulmonary artery function from damage. Most importantly, α7 nAChR expression in pulmonary arteries of COPD patients rose with disease severity and showed an inverse correlation with respiratory function. These findings have important clinical implications, indicating that nAChR-targeted tailored antagonists could be a promising therapeutic strategy for COPD-related vascular dysfunction.

1. Introduction

Chronic obstructive pulmonary disease (COPD) is a complex respiratory pathology characterized by chronic bronchitis, emphysema, small airway obstruction, and persistent respiratory symptoms. Environmental factors, such as the rising of fine particulate matter in the environment and cigarette smoking (CS), exert a substantial influence on the development and progression of COPD [1]. Some patients with COPD also experience alterations in resistance vessel structure mainly due to pulmonary artery (PA) remodeling, which in turn contributes to elevated pulmonary artery pressure (PAP) and the development of pulmonary hypertension (PH). This, in turn, results in right ventricular dysfunction and, in many cases, to the death of the patient [2].

Despite its profound impact on public health, curative and preventive therapies for COPD-related PH are still lacking. Therefore, it is imperative to acquire a comprehensive understanding of the mechanisms underpinning this condition. While the direct effects of CS on the airways are well-documented, its impact on the pulmonary vasculature remains incompletely characterized. Conventionally, vascular remodeling was believed to result from alveolar hypoxia attributed to emphysema and airway obstruction. However, emerging evidence challenges this view, as elevated PAP only moderately correlates with hypoxemia and airflow limitation, and there is no correlation between elevated PAP and emphysema severity [3,4]. Additionally, previous studies show that mild COPD patients and smokers without airway obstruction exhibit media thickening in their pulmonary arteries, suggesting the involvement of other unknown mechanisms, presumably triggered by CS, in the development of PH [5]. Our previous studies demonstrate that cigarette smoke extract (CSE) directly impairs pulmonary arterial vasoconstriction and vasodilation, strongly suggesting intrinsic alterations in the smooth muscle cell (SMC) layer. This was due in part to CSE-mediated dysregulation in the levels and activity of the voltage-dependent K⁺ channel Kv7.4 in PA [6].

Oxidative stress, understood as accumulation of ROS, can be caused not only by oxidizing substances such as tobacco, but also by an excessive production of these species or by a decrease in antioxidant defenses of the cells. This condition has been closely associated with COPD pathogenesis, as evidenced by increased oxidative stress markers in exhaled breath condensates and lung tissues of COPD patients compared to control subjects [7,8]. Similarly, alterations of the redox balance when glutathione availability is decreased [9], or in dysregulated antioxidant response [10] explains why treatment with NAC, a glutathione precursor, is a widespread therapy for COPD patients [[11], [12], [13]]. The excessive ROS production triggers a redox imbalance that contributes to organelle damage and disrupted homeostasis [14], as we have previously shown to occur with the NO-sGC signaling pathway in CSE-challenged human pulmonary artery smooth muscle cells (hPASMC) [10]. These findings suggest that CS-induced changes in the PASMC may contribute to the development of PH in susceptible individuals. Furthermore, we have shown that CS induces mitochondrial dysfunction linked to a pulmonary artery vasodilation impairment [10]. However, despite pulmonary artery contractile impairment has also been proved, its molecular mechanisms remain unclear and its relationship with oxidative stress should be considered.

Calcium ions (Ca²⁺) and ROS function as intracellular signaling molecules with interactive pathways. On the one hand, ROS can activate various calcium channels located in the plasma membrane, endoplasmic reticulum (ER), and mitochondria, leading to an increase in intracellular Ca²⁺ levels [15,16]. On the other hand, elevated intracellular Ca²⁺ levels can activate various ROS-generating enzymes, including NADPH oxidases and mitochondrial enzymes, leading to increased ROS production [17]. Considering the involvement of ROS in the development of PH, it would be important to know whether the crosstalk between ROS and calcium plays a role, as well as the upstream mechanisms that generate this signal. In this respect, nicotinic acetylcholine receptors (nAChRs) are ligand-gated ion channels found in the nervous system and various tissues. They play a key role in neurotransmission by responding to acetylcholine and nicotine, leading to ion flux and cellular signaling [18]. Recently, the expression of these channels, in particular the α4 and α7 nAChR, have been demonstrated in the airway respiratory tract [19,20]. Modulation of these receptors is being explored as a potential therapeutic target for respiratory conditions. In this respect, some authors have highlighted the direct effects of nicotine on mitochondria. Although its mechanism of action remains unclear, data point to an involvement of nicotinic channels in these processes [21]. In line with this, previous reports suggest that calcium homeostasis may be affected during exposure to CS due to the influx of calcium and sodium ions through nAChRs located in the plasmatic and the mitochondrial membrane [20,22,23]. Thus, it is plausible that signaling through these receptors would play an important role in the effects of smoking on the pulmonary artery through an increase in ROS and Ca²⁺.

This study aims to delve deeper into the molecular mechanisms underpinning the development of PH secondary to COPD. We hypothesize that cigarette smoke may be signaling in hPASMC and directly contributing to PA dysfunction in patients with COPD and COPD-related PH. Our results proved that tobacco exposure promoted oxidative stress, elevation of the intracellular calcium and alterations in the contractile machinery that explain CSE-induced alterations on PA reactivity after cigarette smoke exposure. These effects are promoted through nAChR signaling stimulated by nicotine present in the CSE. These results unveiled, for the first time, the expression of these receptors in human pulmonary arteries. Furthermore, we conducted thorough research to investigate potential pharmacological applications aimed at alleviating the symptoms associated with COPD-related vascular dysfunction.

2. Materials and methods

2.1. Cell culture and treatments

Primary hPASMC were obtained from ScienCell Research Laboratories (#3110). The cells were grown in culture according to the specifications recommended by the manufacturer in SMC complete medium (ScienCell, #1101) containing 2 % fetal bovine serum (ScienCell, #0010), 1X SMC growth serum (ScienCell, #2352 and #1152), 100 U/mL penicillin and 100 μg/mL streptomycin (ScienCell, #0503). Cells were grown to 90 % maximum confluence, maintained at 37 °C in a humidified atmosphere of 5 % CO2 and used for a maximum number of eight passages. In the indicated experiments, cells were pre-treated with 15 μM n-acetylcysteine (NAC) for 2 h (Sigma, A7250) prior to co-treatment with 15 % CSE. Nicotine (Sigma Aldrich, N3876) was used at different concentrations (100 nM, 250 nM, 1 μM, 2,5 μM and 10 μM) for 30 min. As a positive control for nAChRs activation, cells were treated with different concentrations (100 nM, 250 nM, 1 μM, 10 μM and 50 μM) of the agonist N-[(3R)-1-Azabiciclo[2.2.2]oct-3-yl]furo[2,3-c]pyridine-5-carboxamide (PHA-543613, from Sigma, PZ0135) for 30 min. Other treatments were as follows: 5 μM cytochalasin B (cytB) (Merck, C6762) for 2 h, 10 μM mecamylamine hydrochloride (Me) (Sigma Aldrich, M9020) for 2 h, and 100 nM alpha bungarotoxin (α-bt) (Alomone, #B-100) for 30 min, and then CSE treatment for the indicated time. Human α3-nAChR (sc-37055) and α7-nAChR siRNA (sc-42532) were purchased from Santa Cruz, USA. Control fluorescent scRNA (SIC005) was purchased from Sigma-Aldrich. Transfection of siRNA was performed according to the specifications recommended by the manufacturer using a commercial transfection reagent (sc-29528) from Santa Cruz, USA. Cells were then analyzed by RT-qPCR, WB or flow cytometry.

2.2. CSE preparation

CSE was prepared from commercial Kentucky 3R4F cigarettes (University of Kentucky Lexington, KY, 9.4 mg tar and 0.73 mg nicotine per cigarette). The smoke from the complete consumption of five cigarettes was continuously bubbled into 150 mL of PBS using VACU- SAFE aspiration system (INTEGRA Biosciences AG) at a vacuum pump flow rate of 8 L/min. CSE solution was filtered through a 0.20 μm-pore system, immediately aliquoted and kept at −80 °C until used. To ensure a similar preparation amongst different batches, CSE concentration was measured spectrophotometrically at 320 nm wavelength. This solution was considered to be 100 % CSE and was diluted to obtain the desired concentrations for each experiment. To avoid the exposure to volatile substances from the smoke, untreated and CSE-challenged cells were always kept in different incubators.

2.3. RT-qPCR analysis

Cells were grown to 90 % confluence in culture plates and total RNA was isolated using TRI Reagent (Molecular Research Center Inc., TR118). RNA (1 μg) was reverse transcribed to complementary DNA (cDNA) with MultiScribeTM Reverse Transcriptase (Applied Biosystems, 4308228) in a final volume of 20 μL. cDNA (1 μL) was amplified with 1 μM of the specific primer pairs and quantitative PCR was performed using GoTaq® qPCR Master Mix (Promega, A6001). Data was analyzed by the 2^-ΔΔCt method using StepOne Plus software (Applied Biosystems). All values were controlled with HPRT1 gene expression levels, and normalyzed to arbitrary units among different replicates. THP-1 cell line was used as a positive control for nAChRs α3, α4 and α7 expression. Primers for PCR were as follows in 5′–3′ orientation: HPRT1 (fwd: ATTGTAATGACCAGTCAACAGGG, rev: GCATTGTTTTGCCAGTGTCAA), CHRNA3 (fwd: CAGAGTCCAAAGGCTGCAAG, rev: AGAGAGGGACAGCACAGCAT), CHRNA4 (fwd: CCTCGGCCTGTCCATCGCTCA, rev: AAGACGGTGAGCGACAGCAC), CHRNA7 (fwd: CAATGGAGAATGGGACCTAGTG, rev: GCAGCATGAAGACGGTAAGA). Electrophoresis was performed through agarose gels (0.7 % agarose) prepared in tris-acetate (40 mM)/EDTA (10 mM) buffer and 5 μl of GelStar Nucleic Acid Stain 10,000 x (Lonza, 50535). 20 μl per sample resulting from PCR were mixed with Blue/Orange loading buffer loading dye 6x (PROMEGA, G190A) and loaded onto gel. Electrophoresis was run at 120V and gel bands were photographed with a transilluminator (Uvitec, BTS-20.M).

2.4. Western blot analysis

Cells were grown to 90 % confluence in culture plates in the presence or absence of CSE for 24 h and lysates were prepared in non-reducing 2X Laemmli buffer, boiled at 95 °C for 10 min in the presence of 20 mM DTT, electrophoretically separated by SDS-PAGE and transferred onto 0.45-μm nitrocellulose membranes (GE Healthcare Life Sciences, 10600003). Total protein bands were reversibly stained with Fast Green FCF (Sigma, F7252) and imaged on ImageQuant LAS 4000 or Amersham ImageQuant 800 (GE Healthcare Life Sciences) for total protein quantification. Transferred proteins were incubated overnight at 4 °C with specific primary antibodies. Horseradish peroxidase conjugated secondary antibodies were added for 1 h at room temperature and protein signal was then visualized using Immobilon Forte (Millipore, WBLUF0500) on ImageQuant LAS 4000 or Amersham ImageQuant 800. Specific protein bands intensity was quantified by densitometry using ImageJ 1.51 software (U. S. National Institutes of Health, Bethesda, Maryland, USA) and normalized to the intensity of Fast Green FCF staining from each complete gel lane. Antibodies for western blotting were as follows: α3 nAChR (sc-58605, Santa Cruz Biotechnology), α7 nAChR (ab216485, Abcam), MLC (3672S, Cell Signaling), p-MLC Thr18/Ser19 (3674, Cell Signaling), 2055p-PKCε (Ser729) (sc-12355-R, Santa Cruz Biotechnology), MLCK (ab314185, Abcam), β1 Integrin (550034, BD), TOM22 (1C9-2, Miltenyi Biotec), TFAM (ab131607, Abcam), anti-mouse-HRP (P0260, Dako), anti-rabbit-HRP (32460, Invitrogen), anti-rat-HRP(AP136P, Sigma-Aldrich).

2.5. Determination of total and mitochondrial O2^- and intracellular calcium

Total and mitochondrial superoxide levels, as well as intracellular calcium levels, were analyzed with fluorescent probes 10 μM dihydroethidium (DHE) (Invitrogen, D11347), 5 μM MitoSOX™ Red (Thermo Fisher Scientific, M36008) and 1 μM Fluo-4 AM (ThermoFisher Scientific, F14201) respectively. Afterwards, cells were trypsinized and quantified by flow cytometry.

2.6. Immunofluorescence

hPASMC were seeded onto fibronectin (20 μg/mL) coated 13-mm glass coverslips and then incubated under the desired experimental conditions. Cells were then fixed (20 min at 4 °C) with 4 % paraformaldehyde in PBS, and permeabilized (10 min RT) with 0,3 % saponine in 0,2 % BSA/PBS. Afterwards cells were labeled with p-MLC Thr18/Ser19 (3674, Cell Signaling) and vinculin (V9131, Sigma) followed by Alexa Fluor™ 488 and 568 labeled secondary antibodies and Phalloidin Alexa Fluor™ 647 (Invitrogen, A22287) staining. Cells mounted in coverslips with Faramount Aqueous Medium were imaged with a Leica DMR microscope using an immersion 40× objective (HCX PL APO 100X/1.40–0.7 oil CS, Leica) and illuminated with a mercury lamp (ebq 100, Lightning & Electronics JENA). Ten images per sample were collected using Leica DCF360FX camera and LAS V4.1 software. The coherency coefficient was quantified on the same images using Orientation J, a software package comprising various plugins designed for ImageJ and Fiji. The Vector Field software assessed the vector modulus, typically indicative of coherence (Advances in Anatomy, Embryology and Cell Biology, vol. 219, Springer International Publishing, ch. 3, 2016). Human pulmonary artery segments from surgical procedures performed at the University Hospital La Princesa were used to analyze the expression of α7 nAChR. Tissue was completely embedded into OCT compound prior to cryostat sectioning. Cut frozen sections at 5 μm were mounted on poly-l-lysine-coated histological slides and immunofluorescence staining was performed. Samples were fixed with 4 % formaldehyde and permeabilized with 0,1 % and 0,3 % Tx-100 in PBS for 10 min each. After blocking with 10 % BSA for 1h at RT, samples were incubated overnight at 4 °C with primary α7 nAChR (ab216485, Abcam) antibody, primary anti-smooth muscle actin, α-SMA (M0851, Dako) or with anti-CD31 (AF3628, R&D Systems) and subsequently incubated for 1h with secondary antibody goat anti-rabbit 647, goat anti-mouse 647 (ab150115, Abcam) or chicken anti-goat 647 (A-21469, Thermofisher) respectively, used all of them at 1:400. An anti-fade mounting media with DAPI (Life Science Inc., St. Petersburg, FL, USA) was used to fix the coverslip to a slide. The slides were examined using the THUNDER Imager 3D fluorescence microscope (Leica Microsystems) and processed with LASX software. Lung sections were also counterstained with Nuclear Fast red-aluminum sulfate solution 0,1 % (1.00121, Sigma-Aldrich). Tissues were imaged with a Nikon Eclipse Ci-L microscope using an air 10× objective (E Plan 10X/0.65). At least 4 images per sample were collected using Nikon NIS-Elements software and color deconvolution plugin.

2.7. Cytoskeleton recovery assay

The actin cytoskeleton was destabilized by incubation for 2h with cytB (Sigma-Aldrich, C6762) (5 μM) followed by a 1h recovery period, during which, cells were cultured without or with 15 % CSE. Conditions requiring pretreatment with any drug (NAC, α-bt and Me) were treated before adding cytB. Treatment was withdrawn during destabilization with cytB and reinstatement during the recovery period. Finally, the cells were fixed, labeled with Phalloidin Alexa Fluor™ 568 (Invitrogen, A12380) and analyzed by flow cytometry. The mean fluorescence intensity (MFI) result of cells whose cytoskeleton had been destabilized with cytB was normalized with respect to cells that had received the same treatment but without cytB, in order to obtain the percentage of repolymerized F-actin.

2.8. Mitochondrial extraction

Mitochondria were isolated with the mitochondrial isolation kit from Miltenyi Biotec (130-094-532) according to the specifications recommended by the manufacturer. The eluent was analyzed by WB using antibodies against the proteins of interest (α3 and α7 nAChRs). In parallel, the mitochondria suspension was analyzed by flow cytometry with an APC and FITC-A labeled secondary antibody.

2.9. Cell senescence assay

Cell senescence was measured with the Invitrogen™ CellEvent™ Senescence Green Flow Cytometry Assay Kit (C10840). Cells were treated with or without 15 % CSE and the specified pre-treatments (Me or α-bt) for 48 h. After this time, cells were trypsinized and washed 2 times with 1X PBS. Then, fixed in 4 % paraformaldehyde for 10 min at room temperature, and stained with the CellEvent™ Senescence Green Probe diluted 1/1000 in CellEvent™ Senescence Buffer for 2 h at 37 °C. Cells were washed in 1X PBS with 1 % BSA, then resuspended in 1X PBS and analyzed by flow cytometry.

2.10. Vascular reactivity measurement

PA from C57BL/6J wild-type or α7 knockout mice were carefully dissected free of surrounding tissue and cut into rings (1.8–2 mm length). Then, PA were pre-treated with 10 μM mecamylamine (Sigma,#M9020) for 2 h, 100 nM α-bungarotoxin (Alomone, #B-100) for 30 min or 15 μM N-acetylcysteine, NAC (Sigma, A7250) for 2 h. Afterwards, rings were overnight-exposed to 15 % CSE. Vessel segments were mounted on a wire myograph in the presence of Krebs physiological solution. Vascular reactivity was recorded by an isometric force transducer and a displacement device coupled with a digitalization and data acquisition system (PowerLab, Paris, France). Preparations were firstly stimulated by rising the K⁺ concentration of the buffer to 80 mM in exchange for Na⁺. Vessels were washed three times and allowed to recover before a new stimulation. Thereafter, PA were stimulated with serotonin (5-HT, 10⁻⁵ M, Sigma) and the relaxation induced by the endothelium-dependent vasodilator acetylcholine (ACh, from 10⁻⁹ to 10⁻⁴ M, Sigma) or the sGC stimulator riociguat (from 10⁻⁹ to 10⁻⁵ M) were assessed. The relaxant responses to ACh or riociguat were expressed as percentage of decrease in 5-HT-induced precontraction.

2.11. Animal studies

Animals supplied by Janvier Laboratories were maintained under standardized conditions and supplied with standard diet and water ad libitum. For in vivo CS exposure, 11-week-old male C57BL/6J WT mice were divided into the following groups: air-exposed (n = 5); air-exposed + mecamylamine (n = 5); CS-treated (n = 5) and CS-treated + mecamylamine (n = 5). CS-treated mice were daily exposed to ten filter-free 3R4F reference cigarettes (University of Kentucky, Lexington, KY), divided into two sessions per day, five cigarettes each and 8 h apart, for 2 weeks. Mainstream CS was generated by an exposure system where combustion of the cigarette was drawn into the mice chambers via a peristaltic pump (KD Scientific Inc.) following the previously described protocol [6]. Control mice were administered filtered air in an identical chamber. Mecamylamine was administered subcutaneous at a dose of 2 mg/kg one day before starting CS treatment and subsequently every other day. At the end of the treatments, animals were euthanized and lungs (left lobule) were immediately fixed in 4 % formaldehyde and OCT-embedded to assess tissue immunostaining. PA from the right lobules were dissected free of surrounding tissue and prepared for vascular reactivity immediately after euthanasia.

2.12. Patient samples

Lung tissue samples were obtained in collaboration with the thoracic surgery unit from University Hospital La Princesa. Residual lung pulmonary arteries were obtained from oncological lobectomies from patients who had not received chemotherapy or radiation within 4 years before surgery or from COPD patients undergoing lung resection categorized as moderate, mild or no COPD according to GOLD classification. The protocol for tissue collection was approved by the ethic committee of the Hospital La Princesa (5159). Samples from patients with respiratory conditions other than COPD have been excluded from the study.

2.13. Smooth muscle cell isolation from human pulmonary arteries

Pulmonary arteries from patients were firstly denuded to remove the adventitial layer. The artery was then enzymatically treated with a solution containing 1.125 mg/mL collagenase (C7657, Sigma), 0.1 mg/mL elastase (E1250, Sigma) and 1 mg/mL albumin to remove the intimal layer. Finally, it was cut into pieces and cultured by the hanging drop method until the pieces dry. They were then soaked in culture media (as specified in cell culture method) and amplified to 90 % confluence. The culture cells were aliquoted and frozen in liquid nitrogen until use.

2.14. Study approval

All animal experiments and procedures were carried out in accordance with the guidelines of the European Community Council Directive (86/609/EEC) and the Spanish Royal Decree 53/2013 based on the European Union normative (2010/63/UE). Animal protocols were approved by the Committee for Research and Ethics of the Universidad Autónoma of Madrid (PROEX 322/14).

The experiments with human samples followed the principles set out in the WMA Declaration of Helsinki and the Department of Health and Human Services Belmont Report. All patients gave written informed consent before inclusion in the study and remained anonymous in the context of this study. Human procedures were supervised and approved by the Committee for Research and Ethics at University Hospital La Princesa (#approval code 5159).

2.15. Statistics

Data are presented as the mean ± s.e.m. Two-sided unpaired Student's t-test were used to compare two groups and one-way ANOVA tests followed by Tukey's post-hoc test were used when comparing three or more groups, according with the conditions of normality and homoscedasticity. In case the assumptions of normality and homoscedasticity were not accomplished, Wilcoxon test was used to compare two groups and non-parametrical Kruskal–Wallis test was used to compare three or more groups followed by Wilcoxon post-hoc test to compare between groups. For the correlation analysis, Pearson's correlation was used unless stated otherwise. No statistically significant differences were considered to be present when p values were larger than 0.05. No statistical methods were used to predetermine sample size. Statistical and graphical analyses were performed with R version 4.3.3.

3. Results

3.1. CSE stimulates ROS and Ca²⁺ elevation in hPASMCs via nAChR activation

Considering that the interplay between Ca²⁺ and ROS signaling is an important determinant in pathophysiology, we aimed to explore this signaling cascade in the context of COPD. Our previous studies demonstrate an endogenous increase in ROS following cigarette smoke treatment [6,10]. However, we do not know whether this signal is also accompanied by alterations in intracellular calcium levels. To this end, we analyzed in parallel the dynamics of ROS and calcium production in response to cigarette smoke extract (CSE). Our results showed a parallel increase in cytoplasmic ROS and calcium levels peaking at 30 min and persisting for 24 h (Fig. 1A). Next we investigated whether cell membrane calcium channels trigger this signaling response, with a focus on nicotinic acetylcholine receptors (nAChRs) as plausible mediators. We analyzed the levels of α3, α4 and α7, since the latter two have been identified in the airway respiratory tract [19,20]. Our analysis identified for the first time mRNA expression of α3 and α7 nAChRs in human pulmonary artery smooth muscle cells (hPASMC) which were also reflected at the protein levels. (Fig. 1B). Furthermore, flow cytometry analyses showed that α3 nAChR surface levels remained unaffected while α7 levels were significantly increased in CSE-treated hPASMC (Fig. 1C). We then examined whether these channels were active and if nicotine, or a nAChR agonist PHA stimulated these channels, enhancing Ca²⁺ influx. Notably, both PHA and nicotine increased intracellular Ca²⁺ and ROS levels in a concentration-dependent manner (Fig. 1D). These findings suggest that CSE effects may be mediated through nicotinic receptor activation in hPASMC. To validate this, we examined whether blocking nAChRs could prevent the CSE-induced increases in ROS and intracellular calcium. Specifically, we treated CSE-exposed hPASMC with α-bungarotoxin (α-bt), a selective α7 nAChR antagonist, or mecamylamine (Me), a non-selective nAChR antagonist. Both antagonists significantly prevented CSE-mediated increase in calcium and ROS within 30 min, and this effect persisted for at least 24 h (Fig. 1E and Supplementary Fig. S1A). To assess α3 and α7 nAChR involvement, we used specific siRNAs, which partially reduced the expression of these channels (Supplementary Fig. S1B and Fig. S1C). Interestingly, this reduction in nAChRs levels was sufficient to diminish the CSE-mediated increase in the levels of calcium and partially prevented the increase in ROS, an effect that is more noticeable in the case of α7 (Fig. 1F). These findings suggest that both α3 and α7 nAChRs may contribute to CSE-induced calcium and ROS increases in hPASMCs.

Fig. 1.

Analysis of ROS and calcium levels along with nAChRs levels and activity. (A) Intracellular calcium and total ROS levels were measured with Fluo-4-AM and DHE probes respectively at the indicated times in hPASMCs and analyzed by flow cytometry. (B) Expression of various nAChR subunits in hPASMC or in THP-1 cell line (used as a positive control) was analyzed by PCR and protein levels for α3 and α7 nAChR in hPASMC cell lysates were analyzed by Western blot (n = 3). (C) Cell surface α3 and α7 nAChR expression was analyzed and quantified by flow cytometry in these cells. Data are represented as the mean ± s.e.m of MFI; (n = 6). (D) Intracellular calcium and total ROS levels were measured with Fluo-4-AM and DHE probes respectively and analyzed by flow cytometry in hPASMCs treated for 30 min with the nAChR agonists (PHA or nicotine) at the indicated concentrations. Data are represented as the mean ± s.e.m; (n = 5–6). (E) hPASMC were pre-treated with nAChR antagonists: 100 nM α-bt or 10 μM mecamylamine (Me) and afterwards incubated with 15 % CSE for the indicated times, then intracellular calcium and total ROS levels were measured with Fluo-4-AM and DHE probes respectively (n = 5–19). (F) hPASMC were interfered with specific siRNA for α3 or α7 nAChR. Afterwards cells were treated without (−) or with (+) 15 % CSE for 24 h and intracellular calcium and total ROS levels were measured with Fluo-4-AM and DHE probes respectively (n = 4–5) (G) mtROS were measured with the MitoSOX™ probe and quantified by FACS in hPASMC previously treated with PHA or nicotine at the indicated concentrations for 30 min, or in cells pre-treated for 2 h with 10 μM Me or 30 min with 100 nM α-bt, then treated for 1h with 15 % CSE. Western blot analysis of nAChRs on isolated mitochondria from hPASMC. Representative images of total cell lysate (TL), membrane fraction (MF) and isolated mitochondria (MT) probed with anti-β1 integrin (to detect plasma membrane contamination), anti-α3 nAChR, anti-α7 nAChR, and TFAM (as a mitochondrial control marker), (n = 3). Data are represented as the mean ± s.e.m. Statistical comparison between groups was performed using two-sided unpaired Student's t-test (A, C, G upper), Kruskal-Wallis test followed by Wilcoxon test (E upper, F) or ANOVA test followed by Tukey's test (D lower, G lower); (∗)p < 0.05, (∗∗)p < 0.01, (∗∗∗)p < 0.001, (∗∗∗∗)p < 0.0001, (ns) non-significant.

Since CSE also elevates mitochondrial ROS (mtROS) levels [10], we investigated whether the activation of nAChRs also induced an increase in mitochondrial ROS. Our results proved that PHA and nicotine significantly increased mtROS levels in a concentration-dependent manner with the higher effect at 100 nM PHA or 250 nM nicotine (Fig. 1G). Interestingly, α-bungarotoxin did not prevent the CSE-induced increase in mitochondrial ROS, whereas mecamylamine effectively inhibited the rise in mtROS in CSE-treated hPASMCs (Fig. 1G and Supplementary Fig. S1D). Given that α-bt is not cell-permeant, our findings suggested the presence of nAChRs localized within hPASMC mitochondria. This was confirmed by our analysis, which identified α3 and α7 nAChR expression in isolated mitochondria (Fig. 1G). These results further validate the role of nAChRs in tobacco-induced mitochondrial dysfunction, highlighting their involvement in CSE-induced mtROS production.

3.2. Oxidative stress disrupts assembly of cytoskeletal proteins and dynamics in CSE-treated hPASMC

Considering our previous findings on ROS involvement in vascular impairment following cigarette smoke exposure [6,10], understanding its effects in the cell contractile apparatus may reveal the mechanism behind CSE-mediated alterations in vascular reactivity. First, we performed fluorescence microscopy with phalloidin, p-MLC, and vinculin staining which revealed disrupted cytoskeletal organization in CSE-treated cells. Actin and myosin bundles appeared diffused, unlike the well-organized structure in control cells. Similarly, vinculin staining showed reduced focal adhesion puncta at the periphery in CSE-treated cells compared to evenly distributed puncta in controls (Fig. 2A). Furthermore, coherency coefficient analysis, which measures how fibers or structures are aligned in a similar direction, indicating an organized, ordered structure, also confirmed a more disorganized actin bundle orientation in CSE-treated cells (Fig. 2A). To gain a more quantitative analysis of CSE effects on these proteins, we performed western blotting to assess myosin light chain (MLC) and its phosphorylated active form (p-MLC), observing no change in total MLC levels, but a significant reduction in p-MLC in CSE-treated cells (Fig. 2B). Notably, cells pre-treated with the antioxidant NAC prevented this reduction, highlighting the role of oxidative stress in CSE-induced contractility impairment. Consistently, myosin light chain kinase (MLCK) and the upstream kinase PKCε were reduced in CSE-treated cells (Fig. 2B). Moreover, actin dynamics experiments showed impaired actin polymerization recovery in CSE-treated cells after cytB destabilization, an effect that was also prevented by NAC (Fig. 2C). These findings suggest that ROS generated by CSE exposure disrupt cytoskeletal dynamics, contributing to alter cell cytoskeletal dynamics.

Fig. 2.

CSE effects on hPASMC contractile machinery. (A) hPASMCs were treated for 24 h with 15 % CSE and the effects on proteins of the contractile machinery were analyzed by fluorescence microscopy. Representative images for the staining with phalloidin-Alexa FluorTM 647 (purple), and immunostaining with p-MLC (cyan) and vinculin (yellow, white arrows pointing to focal adhesion staining) are shown (n = 4), scale bar = 10 μm. Quantification of the coherency coefficient analyzed in 32 random cells from these experiments is shown. (B) hPASMCs were pretreated for 2 h with (+) or without (−) 15 μM NAC and maintained for 24 h in the presence of 15 % CSE. Afterwards, levels of myosin light chain (MLC) and its phosphorylated form (p-MLC), and levels of myosin light chain kinase (MLCK) and phosphorylated protein kinase C (p-PKCε isoform) were analyzed. Quantification of protein levels and representative images from (n = 4–14) are shown. (C) Actin dynamic was examined by means of flow cytometry in cells pretreated with (+) or without (−) 15 μM NAC for 2 h then treated with cytochalasin B for 2 h and afterwards exposed to 15 % CSE for 1 h. The percentage of F-actin re-polymerization was assessed and MFI of recovering cells was adjusted by the ratio between destabilized/non-destabilized cells (n = 4–13). Statistical comparison between groups was performed using two-sided unpaired Student's t-test (A, B lower) and ANOVA test followed by Tukey's test (B upper, C) (∗)p < 0.05, (∗∗)p < 0.01, (∗∗∗∗)p < 0.0001, (ns) non-significant.

3.3. Inhibition of nAChRs prevents cigarette smoke effects on hPASMC and improves PA vascular reactivity

Our previously published results demonstrate that cigarette smoke directly contributes to pulmonary arterial remodeling through increased cell senescence [6]. Therefore, we tested the ability of nAChRs antagonists to prevent these effects, as well as those on cytoskeletal dynamics. The results showed that α-bt and Me were able to prevent CSE-mediated cellular senescence (Fig. 3A). Additionally, the loss in actin cytoskeleton dynamics after CSE treatment was significantly prevented in the presence of these antagonists (Fig. 3B), thus strengthening the link between channel activation, ROS production and cytoskeletal dysfunction. Moreover, Me also prevented the CSE-mediated decrease on MLC kinase levels, although no significant effects were observed in the levels of p-MLC (Fig. 3C).

Fig. 3.

Effects of nAChR inhibition on CSE-mediated alterations on hPASMC. (A) hPASMC were pre-treated with 100 nM α-bungarotoxin (α-bt) or 10 μM mecamylamine (Me) and then exposed to 15 % CSE for 48 h. X-gal staining to quantify β-galactosidase activity was analyzed by flow cytometry. Cell senescence is presented as median fold change in β-Gal MFI and normalized to the levels in non-CSE treated (−); (B) Quantification of F-actin re-polymerization percentage measured by flow cytometry. The re-polymerization analysis was assessed in cells exposed to 15 % CSE in the absence (−) or presence (+) of 100 nM α-bt or 10 μM Me. The MFI of recovering cells was adjusted by the ratio between destabilized/non-destabilized cells. Data are presented as mean ± s.e.m (n = 6–7). (C) Analysis of myosin light chain kinase (MLCK) and p-MLC levels in hPASMC pre-treated with 100 nM α-bt or 10 μM Me and then exposed to 15 % CSE for 24 h. Western blot representative images and protein levels quantification are shown (n = 5–7). Data are represented as the mean ± s.e.m.; Statistical comparison between groups was performed using one-way ANOVA test followed by Tukey's test (A, B, C upper) and two-sided unpaired Student's t-test (C lower) (∗)p < 0.05, (∗∗)p < 0.01, (∗∗∗)p < 0.001, (∗∗∗∗)p < 0.0001.

To assess the role of nAChRs in vascular function, we performed wire myography on isolated mouse pulmonary arteries treated with 15 % CSE for 12 h, with or without mecamylamine. This inhibitor prevented the CSE-induced impairment of endothelium-dependent vasodilation in response to ACh or riociguat, with effects comparable to those observed with α-bt or NAC (Fig. 4A and Supplementary Fig. S2). Since α7 nAChR levels were significantly upregulated by CSE in hPASMC membrane, we examined its role in altered vasodilation. Our findings demonstrated that pulmonary arteries from α7 knockout mice maintained normal vasodilatory responses to acetylcholine or riociguat following CSE exposure (Fig. 4B), suggesting that α7 nAChR contributes to CSE-induced vascular impairment. To extend these findings in vivo, we treated WT mice with or without cigarette smoke and with or without Me. Treatment with Me prevented CS induced vasodilation impairment in response to ACh or riociguat. Furthermore, they also preserved alveolar integrity, preventing smoke-induced destruction (Fig. 4C). These results highlight the critical role of α7 nAChRs in pulmonary artery responses to tobacco smoke, with effects extending beyond the vasculature.

Fig. 4.

Effects of nAChRs inhibition on PA reactivity. (A) Vascular responses were analyzed in endothelium-intact PAs from C57BL/6J mice (n = 4 animals) from untreated (CTRL), (n = 6 rings), or overnight treated with 15 % CSE (n = 6 rings), alone or in combination with 10 μM Me (Me + CSE, n = 6 rings). Average values of the endothelium-dependent vasodilation induced by increasing amounts of ACh (left) or increasing amounts of riociguat (right) were assessed and expressed as percentages of relaxation related to the maximum 5-HT-driven vasoconstriction. (B) PAs from α7^−/− C57BL/6J mice (n = 5 animals) were untreated (n = 7 rings) or overnight treated with 15 % CSE (n = 6 rings) and vascular responses were analyzed as in (A). (C) C57BL/6J mice (n = 5 per group) were exposed to air (CTRL; n = 8 rings), cigarette smoke (CS; n = 9 rings), or a combination of cigarette smoke and 2 mg/kg Me (Me + CS; n = 9 rings) over a two-week period, following the specified timeline. Afterwards vascular responses were analyzed as in (A). Lung sections from each animal group were stained with nuclear fast red and analyzed by light microscopy. Representative images from controls, CS treated or Me + CS were selected. Data are represented as the mean ± s.e.m.; Statistical comparison between groups was performed using one-way ANOVA test followed by Tukey's test (A, C) or two-sided unpaired Student's t-test (B) (∗)p < 0.05, (∗∗)p < 0.01, (∗∗∗)p < 0.001 (∗∗∗∗)p < 0.0001.

3.4. α7 nAChR expression in pulmonary arteries of COPD patients increases with disease severity

We next comparatively analyzed the levels of α3 and α7 nAChRs by means of flow cytometry in hPASMCs derived from patients with or without COPD. The analysis of α7 nAChR surface expression revealed a significant increase in the levels of this channel, while the levels of α3 nAChR were significantly lower in cells from COPD patients compared to those in cells from non-COPD individuals (Fig. 5A and Supplementary Tables S1 and S2). These results strongly support our in vivo functional findings and emphasize the significance of these receptors in PA functional impairment. Furthermore, immunofluorescence analysis revealed significantly higher total α7 nAChR levels in pulmonary arteries of moderate COPD patients compared to those in non-COPD individuals or mild COPD patients (Fig. 5B and Supplementary Table S3), further highlighting their role in vascular impairment in COPD patients.

Fig. 5.

Analysis of nAChR protein levels in hPASMC and pulmonary arteries. (A) Cell surface α7 and α3 nAChR levels were analyzed in hPASMC from non-COPD (n = 5–8) and COPD (n = 5–6) patients by FACS. Representative histograms display stained cells (dark colored) and unstained controls (light colored). (B) Lung PA were stained with α7 nAChR (red), CD31 (yellow) and α-SMA (green). α7 protein levels were analyzed and quantified by fluorescence microscopy in PA from patients without (n = 10), mild (n = 3) or moderate COPD (n = 6). At least 6 fields randomly selected from the whole artery were quantified. Data are represented as the mean ± s.e.m.; Statistical comparison between groups was performed using two-sided unpaired Student's t-test (A) or one-way ANOVA test followed by Tukey's test (B) (∗)p < 0.05, (∗∗)p < 0.01.

Finally, to determine whether nAChRs levels correlated with clinical parameters in pulmonary function we analyzed the relationship between these receptors and key clinical indicators such as FEV1, which assesses COPD severity, and FEV1/FVC, which distinguishes COPD patients from non-COPD patients. Our results showed that the levels of α7 nAChR inversely correlated with FEV1 (%) and FEV/FVC (FEV1 %: r = −0,664, p = 0,009; FEV1/FVC: r = −0,501, p = 0,068) (Fig. 6A). Conversely, the levels of α3 nAChR directly correlated with these lung function markers (FEV1 %: r = 0,47, p = 0,144; FEV1/FVC: r = 0,639, p = 0,034) (Fig. 6B). Moreover, these receptors displayed distinct patterns of expression when correlated with cigarette consumption, measured in packs-years. While α7 levels showed a direct correlation (r = 0.508, p = 0.077), α3 levels demonstrated an inverse correlation (r = −0.614, p = 0.059). These findings suggest a potential compensatory regulatory mechanism for these receptors in response to smoking and highlight the important role of both nAChRs in COPD development.

Fig. 6.

Correlation analysis of α3 and α7 nAChR levels with clinical parameters.

Pearson correlations between superficial α7 (A) and α3 (B) nAChR levels in hPASMC from patients with or without COPD and clinical parameters including FEV1 (%), FEV1/FVC and Spearman correlation between α7 (A) and α3 (B) nAChR levels and packs per year day (n = 10–14).

4. Discussion

COPD-related PH is a major health concern with limited treatments and poor outcomes. Our previous studies identified direct effects of tobacco smoke on pulmonary vascular dysfunction, particularly impairing vasodilation [6,10]. Here, we explore the molecular mechanisms driving ROS generation and calcium deregulation in hPASMC after tobacco exposure and their role in reduced pulmonary vascular reactivity. CSE exposure induced oxidative stress, increased intracellular calcium, and altered the contractile machinery in hPASMC. The presence of nicotinic receptors in the human pulmonary artery, along with the effects observed from inhibitor treatments and expression interference, underscores their involvement in tobacco-induced vascular dysfunction.

Previous studies suggest that nicotine [24], tobacco smoke [25], and ROS [26,27] impact actin and tubulin dynamics. Our findings showed that CSE disrupted actin dynamics and MLC phosphorylation in hPASMC, key indicators of contractility changes. These effects were ROS-dependent, as NAC treatment effectively restored function. These results may be explained by ROS-mediated MLCK inhibition [28]. Other authors have highlighted the interplay between ROS and calcium, key signaling molecules in vascular pathologies [29]. Our results show increased intracellular calcium levels correlated with elevated total and mitochondrial ROS in hPASMC after CSE exposure. This suggests calcium as the primary signaling element in CSE-mediated effects, driving redox imbalance and vascular dysfunction. A reciprocal loop between calcium and ROS has been proposed in pulmonary hypertension [30], but their exact role in mitochondrial dysfunction remains unclear. In this respect, prior research links mtROS and calcium, showing how altered calcium channel activity modulates mitochondrial calcium, promoting mtROS generation [31,32]. Our findings align with this, suggesting a potential mtROS-calcium interaction upon CSE exposure, shifting toward a pathological state in COPD-related vascular dysfunction.

Given the importance of identifying the mechanism that initiates this cascade, nAChRs, calcium-permeable channels, are proposed as key triggers of signaling cascades in response to tobacco exposure in the pulmonary artery. In this context, their expression has been identified in bronchial and respiratory tract [33,34]. The α7 nAChR may protect against chronic inflammation and pulmonary emphysema [[35], [36], [37]], but its effects vary across cell types and nicotine exposure. Some studies highlight their role in airway remodeling and dysfunction [38] suggesting α7 nAChR inhibition as a therapeutic target for airway diseases. Despite smoking associates with vascular risks and nicotine-induced dysfunction [39], little research exists on nAChRs in the pulmonary vasculature. Nicotine-mediated cardiovascular effects vary by cell type, but studies confirm multiple nAChR α-subunits in rat systemic arteries [40,41]. Interestingly, nicotine interacts with nAChRs, particularly α7, contributing to vascular damage in animal models [42]. Our findings support this, indicating α7 and possibly α3 nAChRs mediate nicotinic signaling in human pulmonary arteries. Nicotine reconfigures vascular SMC cytoskeletons, altering α-actin, vimentin, and β-tubulin via α7 nAChR, contributing to dysfunction [43]. Due to its high calcium permeability, the α7 nAChR plays a direct role in regulating intracellular calcium signaling [44]. Our experiments using nicotine, nAChR antagonists (α-bungarotoxin, mecamylamine), and specific α3/α7 interference confirm the role of these receptors in CSE-induced calcium accumulation and oxidative stress in human pulmonary artery smooth muscle cells. Additionally, our data on α3 and α7 nAChR regulation in patient cells suggest compensatory mechanisms favoring α7 for cell signaling in response to cigarette smoke, likely due to chronic smoke-induced receptor desensitization [45]. Additionally, we detected α3 and α7 nAChRs in mitochondria, implicating them in CSE-mediated effects. These receptors, previously identified in the outer mitochondrial membrane, contribute to nicotine-induced depolarization, triggering calcium release from the endoplasmic reticulum and increasing mitochondrial oxidative stress [31,46,47]. Altogether, these findings underscore the reticulum-mitochondria axis as a key mediator in CSE-induced mitochondrial dysfunction in hPASMC, reinforcing the interplay between ROS, calcium, and nAChRs in pulmonary vascular pathophysiology.

While previous studies have suggested the potential of nAChRs agonists in COPD inflammation [33], there is no previous study confirming their expression in human PA. Our current research is also the first to elucidate the biological significance of nAChRs in CSE-induced impairment in human pulmonary vascular function. Activation of these receptors by nicotine triggers calcium signaling pathways and promotes oxidative stress in hPASMC. These events alter the contractile machinery and actin dynamics within these cells, collectively contributing to pulmonary artery dysfunction. These findings suggest that the use of nicotine in smoking cessation therapies may need to be re-evaluated. Most importantly, blocking nAChRs effectively prevented the decrease in vasodilation stimulated by ACh or riociguat in PA challenged with CSE. Our model reflects acute tobacco exposure rather than a classical COPD model, which typically requires several months of exposure before the development of emphysema, this distinction should be acknowledged as a limitation when extrapolating our findings to human disease. However, our experimental design was intentionally chosen to avoid the confounding effects of established emphysema and hypoxia, thereby allowing us to specifically investigate the early and direct impact of tobacco exposure on the pulmonary vasculature. Importantly, our results indicate that vascular dysfunction can occur rapidly following tobacco exposure and independently of hypoxia, providing mechanistic insight that supports previous hypotheses and may have implications for understanding the earliest stages of tobacco-related lung pathology.

In addition, our findings in human samples hold significant clinical relevance as they underscore a link between receptor expression with disease severity and smoking habits, this suggesting that nAChRs, and in particular α7 nAChRs, could be potential therapeutic targets for managing COPD-associated vascular complications. The treatment with inhibitors targeting α7 nAChRs, alongside existing vasodilators like riociguat, could potentially provide a therapeutic alternative for patients with COPD-related vascular dysfunction who do not respond to riociguat alone.

CRediT authorship contribution statement

Rosa Andreu-Martínez: Writing – review & editing, Writing – original draft, Validation, Methodology, Investigation, Formal analysis, Data curation. Onofre Munar-Rubert: Writing – original draft, Validation, Methodology, Investigation, Formal analysis, Conceptualization. Jorge Rodríguez-Pérez: Writing – original draft, Validation, Methodology, Investigation, Formal analysis, Data curation. Noelia López: Methodology, Investigation, Formal analysis, Data curation. Bianca Barreira: Validation, Methodology. Laura Sánchez-Carretero: Methodology. Adele Cardeñosa: Methodology. Ana Marcos-Jiménez: Validation, Methodology. Luis Gandía: Resources. Ramón Moreno-Balsalobre: Resources. Héctor Milián: Resources. Francisco Pérez-Vizcaíno: Writing – review & editing, Methodology. Edgar Fernández-Malavé: Methodology. Germán Peces-Barba: Validation, Methodology. Cecilia Muñoz-Calleja: Writing – review & editing, Formal analysis. Ángel Cogolludo: Writing – review & editing, Validation, Methodology, Formal analysis. María J. Calzada: Writing – review & editing, Writing – original draft, Validation, Supervision, Resources, Investigation, Funding acquisition, Conceptualization.

Role of funders

The funders had no involvement in the study design, data collection, analysis, interpretation, manuscript preparation, or any other aspect of the research.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A. Supplementary data

The following is the Supplementary data to this article.

Data availability

All relevant data supporting the key findings of this study are available within the article and its supplementary information files or from the corresponding author upon reasonable request.