Nicotine Elevated Intracellular Ca²⁺ in Rat Airway Smooth Muscle Cells via Activating and Up-Regulating α7-Nicotinic Acetylcholine Receptor

It is well documented that tobacco use is the most important risk factor for the development of chronic obstructive pulmonary disease (COPD), which is characterized with profound airway remodeling, including airway smooth muscle hypertrophy and hyperplasia[1,2,3,4]. However, the role of the major active component of tobacco, nicotine, in COPD is still unclear. The primary molecular target of nicotine is the nicotinic acetylcholine receptors (nAChRs), which are ligand-gated channels composed of homo- or heteropentamer of various subunits (α1-α10; β1-β4) and operate as cation entry pathways across the plasma membrane [5,6,7]. Nicotine interacts with various nAChR subtypes with different affinities (1 to 130 µM) [8,9].

Activation of nAChR elicits Ca²⁺ response in part through direct Ca²⁺ permeation via the receptor channel [10]. In addition, nAChR-mediated membrane depolarization can activate voltage-dependent Ca²⁺ channels (VDCCs) to further augment Ca²⁺ signals generated by nAChRs [11,12,13]. These two mechanisms are complementary physiologically, such that Ca²⁺ entry through the inwardly rectifying nAChR channels is predominant under either resting or hyperpolarized conditions, whereas Ca²⁺ influx through VDCCs occurs at more depolarizing potentials (>-40 mV) [13,14,15]. In addition to extracellular Ca²⁺ entry, Ca²⁺ release from intracellular stores may also play a crucial role in defining nAChR mediated Ca²⁺ responses. Activation of Ca²⁺ stores following stimulation of nAChR contributes to the long-lasting Ca²⁺ signals neuronal cells [12,16,17,18,19,20]. Specific nAChR subtypes are associated with defined Ca²⁺ mobilization pathways. For example, nAChRs that contain α3 and/or β2 subunits in brain and ganglionic preparations are associated predominantly with Ca²⁺ signals mediated by depolarization and activation of VDCCs [12,18,19], whereas α7-nAChRs generate robust Ca²⁺ transients which reflect Ca²⁺ entry through the nAChR and Ca²⁺-induced-Ca²⁺ release (CICR) [12,18,21]. Functional coupling between α7-nAChRs and ryanodine receptors has been shown in hippocampal astrocytes, where α7-nAChR-mediated Ca²⁺ signals arise primarily from CICR through ryanodine receptors [21,22].

α7-nAChR subunits form functional homo-pentameric nAChR which has a much higher Ca²⁺ permeability than other nAChRs [23,24,25]. α7-nAChR is activated by agonists such as choline and nicotine [23,26] and blocked by antagonists such as α-bungarotoxin and methyllycaconitine (MLA) [27,28]. Furthermore, α7-nAChR exists in a variety of cell types including neuron [23], bronchial epithelial cells [29], aortic endothelial cells [29], macrophage [29], and artery smooth muscle cells (ASMCs) [30]. However, the expression and function of α7-nAChR in the airway smooth muscle cells remain unclear.

In the present study, we tested the hypothesis that nicotine modulates ASMC functions through the activation of nAChRs. We sought to identify the nAChR subtypes expressed in rat ASMCs and characterize the effects of chronic and acute nicotine exposure on Ca²⁺ mobilization in ASMCs. Our results herein demonstrated that (1) multiple functional nAChRs are expressed in rat ASMCs; (2) nicotine exposure upregulates α7-nAChR expression and function; and (3) the biological effects of nicotine may resulted from α7-nAChR mediated increase in [Ca²⁺]_i. These results suggest that α7-nAChRs may be the major target of nicotine in ASMCs, and its upregulation during nicotine exposure may lead to elevated [Ca²⁺]_i, providing a novel mechanism for airway remodeling.

Similar to the isolation of pulmonary arterial smooth cells described previously [31], bronchi were dissected from lungs of male Sprague Dawley rats (body wt 300-500 g). The epithelium was removed by rubbing the luminal surface with a cotton swab. The bronchi were allowed to recover for 30 min in cold (4°C) physiological salt solution (PSS) that contained (in mM) 130 NaCl, 5 KCl, 1.2 Mg Cl₂, 10 HEPES, and 10 glucose. It was followed by 20 min in reduced-Ca²⁺ PSS (20 µM CaCl₂) at room temperature. The tissue was then digested at 37°C for 20 min in reduced-Ca²⁺ PSS containing collagenase (type I, 1,750 U/ml), papain (9.5 U/ml), bovine serum albumin (2 mg/ml), and dithiothreitol (1 mM). After resuspension and trituration of the digested tissue, cells were plated onto 25-mm coverslips (for fluorescent microscopy) or 10-cm petri dishes (for molecular biological measurements) and incubated for 3-6 days in smooth muscle growth medium 2 (Clonetics, Walkersville, MD) containing 5% serum in a humidified atmosphere of 5% CO₂-95% air at 37°C. Twenty-four hours before an experiment, the concentration of serum in culture media was decreased to 0.5% to stop cell growth.

Cellular purity of the cultures was assessed by examining morphological appearance under phase-contrast microscopy and immunofluorescence staining for α-actin under fluorescence microscopy. For the latter, ASMCs were treated with a primary monoclonal antibody raised against smooth muscle α-actin (Sigma, St. Louis, MO), Cy3-conjugated secondary antibody (excitation λ = 550 nm, emission λ = 570 nm; Jackson ImmunoResearch, West Grove, PA), and a nuclear stain (YO-PRO-1, excitation λ= 488 nm, emission λ = 509 nm; Molecular Probes, Eugene, OR). Cells were examined under a Zeiss LSM-510 inverted laser-scanning confocal fluorescence microscope with a Zeiss Plan-Neofluor ×40 oil immersion objective (Atlanta, GA). For each determination, we inspected at least 1,000 cells in at least 40 randomly selected fields. Coverslips not exposed to the primary antibody, but otherwise treated similarly, served as controls.

As previously described [32], after incubation with 5 µM fura-2 (Molecular Probes) for 60 min at 37°C under an atmosphere of 5% CO₂-95% air, coverslips with rat ASMCs were mounted onto a closed polycarbonate chamber clamped in a heated aluminum platform (PH-2; Warner Instrument, Hamden, CT) on the stage of a Nikon TSE 100 Ellipse inverted microscope (Melville, NY). The chamber was perfused at 0.5 ml/min with Krebs-Ringer bicarbonate (KRB) solution, which contained (in mM) 118 NaCl, 4.7 KCl, 0.57 MgSO₄, 1.18 KH₂PO4, 25 NaHCO₃, 2.5 CaCl₂, and 10 glucose. KRB solution was equilibrated with 16% O₂-5% CO₂ at 38°C in heated reservoirs and led via stainless steel tubing and a manifold to an in-line heat exchanger (SF-28, Warner Instrument), which rewarmed the perfusate just before it entered the cell chamber. The temperature of the heat exchanger and chamber platform was controlled by a dual-channel heater controller (TC-344B, Warner Instrument). This system maintained temperature at 37°C and oxygen tension at 112 ± 2.0 mmHg at the coverslip. After perfusing the chamber for 10 min to remove extracellular dye, ratiometric measurement of fura-2 fluorescence at 60-s intervals was performed using a collimated light beam from a xenon arc lamp filtered by interference filters at 340 and 380 nm and focused onto rat ASMCs visualized with a ×20 fluorescence objective (Super Fluor 20; Nikon, Torrance, CA). Light emitted from the cells at 510 nm was returned through the objective and detected by a cooled charge-coupled device imaging camera. An electronic shutter (Vincent Associates, Rochester, NY) was used to minimize photobleaching. Protocols were executed, and data were collected on-line with InCyte software (Intracellular Imaging, Cincinnati, OH). [Ca²⁺]_i was calculated from fura-2 fluorescence ratios (F340/F380) using linear regression between adjacent points on a calibration curve generated by measuring F340/F380 in at least seven calibration solutions with [Ca²⁺] between 0 and 610 nM (Molecular Probes). Because the behavior of fura-2 in solution may differ from that in cells, calculated [Ca²⁺]_i should be considered as an estimate of the actual [Ca²⁺]_i.

Total RNA was prepared from rat ASMCs, as previously described [33]. Briefly, samples were placed in cold TRIzol reagent (50-100 mg tissue/ml) and homogenized at 20,000 rpm (Ultra-Turrax T-25). After incubation at 30°C for 5 min and addition of chloroform (0.2 ml/ml TRIzol), samples were centrifuged at 8,000 g at 4°C for 15 min. The upper aqueous phase of the supernatant was mixed with 100% isopropanol (0.5 ml/ml TRIzol), incubated at 30°C for 10 min, and recentrifuged at 4°C and 12,000 g for 10 min. The clear gel-like RNA precipitate was washed with 1 ml of 70% alcohol, dissolved in diethyl pyrocarbonate water (1 µg/µl), and stored at -70°C. The quality of the RNA isolate was determined from the ratio of absorbance at 260 nm to that at 280 nm (>1.7) and the integrity of the samples was judged by examining 28S and 18S ribosomal RNA bands appearing on electrophoresis of denatured RNA samples through an 1% agarose formaldehyde gel.

RT was performed with the First-Strand cDNA Synthesis kit (Pharmacia Biotech, Austin, TX), as previously described (27). The resultant cDNA was subjected to PCR by adding 4 µl of the First Strand cDNA reaction mixture to a 50-µl PCR reaction mixture, which consisted of 1 pM of each PCR primer, 10 mM Tris_HCl (pH 8.3), 50 mM KCl, 2 mM MgCl₂, 200 µM each dNTP, and 2 units Taq DNA polymerase. Primer sequences specific for rat nAChRs, or β-actin were designed using Primer 5 software and are listed in Table 1. The fidelity and specificity of these oligonucleotides were confirmed using the Basic Local Alignment Search Tool program. cDNA was amplified in a DNA thermal cycler (model 2400 GeneAmp PCR System; Perkin-Elmer, Foster City, CA). The mixture was annealed at 56-58°C (1 min), extended at 72°C (2 min), and denatured at 94°C (1 min) for 31-35 cycles. It was followed by a final extension at 72°C (10 min) to ensure complete product extension. The PCR products were electrophoresed through a 1.5% agarose gel, and the amplified cDNA bands were visualized by ethidium bromide staining.

cDNA was quantified by real-time qPCR using QuantiTect SYBR Green PCR Master Mix (Qiagen) in a iCyclerIQ real-time PCR detection system (Bio-Rad) using the following conditions: 95°C for 15 min and 45 cycles, each at 94°C for 15 s, 56°C for 20 s, and 72°C for 20 s. The volume of each real-time qPCR reaction mixture was 25 µl containing 300 nM forward and reverse primers and cDNA template from 50 ng RNA. Identity of the qPCR products was confirmed by 1) a single sharp peak in the melting curve performed after cDNA amplification; 2) a single band of the expected size resolved by agarose gel electrophoresis; and 3) the correct DNA sequence. Melting curves were performed at 95°C for 1 min and 55°C for 1 min, followed by 80 increments of 0.5°C at 10-s intervals. Real-time qPCR detection threshold cycle values were generated by iCyclerIQ software. Relative concentrations of each transcript were calculated using the Pfaffl method [34]. Mean efficiencies of amplification for each gene were used to quantify expression of α7-AChR relative to β-actin in the same sample.

Primary cultures of ASMCs were washed with phosphate-buffered saline and homogenized in HEPES buffer at 20,000 rpm (Ultra-Turrax T-25)., total protein concentration in the homogenate was determined by the BCA protein assay (Lowry method) as described previously [35]. Homogenate proteins were separated by SDS-PAGE calibrated with prestained protein molecular weight markers (Bio-Rad, Hercules, CA). Separated proteins were transferred to nitrocellulose membranes (Hybond-C, Bio-Rad). After blocked with 5% nonfat dry milk in Tris-buffered saline and 0.1% Tween 20, membranes were incubated with affinity-purified polyclonal antibodies specific for α7-AChR (Santa Cruz Biotechnology, Santa Cruz, CA). The membranes were then washed and incubated with anti-rabbit horseradish peroxidase-conjugated IgG for 1 hour, and an enhanced luminol-linked chemiluminescence detection system (Pierce Biotechnology, Rockford, IL) was used to detect the bound antibody.

We selected two sites in the α7-AChR mRNA sequence as siRNA targets based on principles described previously [36]. The targeted α7-AChR sequences, based on which the siRNAs were chemically synthesized by Invitrogen (Carlsbad, CA), were 5'-UCUCCUGAACUGGUGUGCAUGGUUU-3' (siRNA-1) and 5'- AAACCAUGCACACCAGUUCAGGAGA-3' (siRNA-2). The negative control siRNAs were purchased from Invitrogen. In vitro transfections of ASMCs with 10 µg/ml siRNA or control sequence were performed using Lipofectamine 2000 (Invitrogen, Carlsbad, CA) following manufacturer's protocols. The efficiency of α7-AChR suppression was determined by Western blot, and the Ca²⁺ fluorescence experiments were preformed 72 hours after siRNA transfection.

The following drugs used in this study were obtained from Sigma: nicotine, methyllycaconitine (MLA), α-bungarotoxin. Activated pervanadate solution was prepared freshly for each experiment and made by mixing a stock solution of vanadate with H₂O₂ in an equal molar ratio. Fura-2-AM (Molecular Probes) was prepared on the day of the experiment as a 2.5 mM stock solution in dimethyl sulfoxide (DMSO).

Data are expressed as means ± SEM, where n is the number of experiments performed, and the number of cells in each experiment ranged between 20 and 40, as indicated in results section and the figure legends. Statistical comparisons were performed using Student's t-test. Differences were considered to be significant when P < 0.05.

ASMCs cultured from rat pulmonary bronchi were phase bright and assumed a spindle-shaped appearance with cytoplasmic projections extending from a larger central area that contained the nucleus (Fig. 1A). Immunostaining for smooth muscle α-actin was positive in >95% of cells (Fig. 1B). None of the cells were positive when staining was performed without the primary anti-α-actin antibody.

It has been reported that nicotine treatment raises resting [Ca²⁺]_i in other cells [12,37], but whether nicotine exerts its effect in ASMCs is unclear. To examine this possibility, ASMCs were cultured in the presence of nicotine. As depicted in Fig. 2B, a noticeable increase in baseline [Ca²⁺]_i was observed in nicotine (100 µM) treated culture cells after exposure periods of 24 to 72 hour. The nicotine effect on resting [Ca²⁺]_i was concentration-dependent, with significant induction at concentrations from 10^-6 to 10^-4 M after 48 hours (Fig. 2A). The most robust and consistent effect was seen at 10^-4 M. A higher concentration of nicotine (10^-3 M) caused a decline in the change of baseline [Ca²⁺]_i, possibly due to inactivation of nAChRs or cytotoxicity. These results for the first time suggest that nicotine plays a role in Ca²⁺ homeostasis of ASMCs.

To further examine the nAChR subtypes that maybe involved in the altered Ca²⁺ homeostasis, the expression of various nAChR subunits (α1-7 and β1-4) in rat ASMCs was determined using RT-PCR. As shown in Fig. 3A, the expression of α2-7, β2, and β3 nAChR RNA were clearly detected in rat ASMCs with α7 being the most abundant nAchR subtype followed by β2, α5, α6, β3, α4, α3, and α2. The transcripts of α1, β1, and β4 were not detected in ASMCs. Since α7-nAChR is most abundantly expressed and possesses the unique property of the highest Ca²⁺ permeability among the nAChR subtypes [38,39,40], we focused our attention on the role of α7-nAChR in nicotine-induced Ca²⁺ response.

We evaluated the effect of nicotine exposure on α7-nAChR expression in the cultured ASMCs. RT-PCR analysis revealed a time-dependent nicotine (10 µM) induction of α7-nAChR mRNA, with a maximal increase after 48 hours of culture (Fig. 3B and C). The effect of nicotine on α7-nAChR expression was concentration-dependent (10^-8 to 10^-4 M); the mRNA level was approximately doubled at 10 and 100 µM nicotine (Fig. 3D and E). Further increase in nicotine concentration reduced the increase in α7-nAChR expression in ASMCs (data not shown). The observation of nicotine-induced α7-nAChR upregulation was confirmed by quantitative real-time RT-PCR (Fig. 3E), which showed nicotine treatment at 10 µM for 48 hours caused a 1.7-fold increase in α7-nAChR mRNA expression compared to the control. Consistent with the changes observed in α7-nAChR mRNA, α7-nAChR protein level was increased in nicotine treated cells in a concentration-dependent fashion, with 2-fold increase at 10^-5 M compared to the control (Fig. 4A and 5B). The stimulatory effect of nicotine on α7-nAChR protein expression in ASMCs was maximal at 48 hours of treatment (Fig. 4C and 4D). These results clearly suggest that nicotine can auto-regulate the expression of its own receptors in ASMCs.

In addition to the chronic effect of nicotine exposure on resting [Ca²⁺]_i, acute exposure of nicotine evoked a rapid increase in [Ca²⁺]_i in ASMCs. Nicotine-induced Ca²⁺ transient is concentration-dependent (Fig. 5A and B). At a lower concentration of 10^-5 M, nicotine caused a mild sustained Ca²⁺ increase with an average ∆[Ca²⁺]_i of 50.6 ± 5.1 nM, whereas 10^-4 M nicotine evoked a robust biphasic Ca²⁺ response with a peak ∆[Ca²⁺]_i of 330.4 ± 28.5 nM. The nicotine-induced Ca²⁺ response was further examined in cultured ASMCs with or without pretreatment with 1 µM nicotine for 48 hours. Acute nicotine-induced Ca²⁺ transients were significantly enhanced in ASMCs after 48 hours of nicotine treatment as compared to control ASMCs (Fig. 5C and D). The peak Δ[Ca²⁺]_i induced by 50 µM nicotine was 320.3 ± 31.1 nM in nicotine-pretreated ASMCs and 191.2 ± 23.4 nM in control ASMCs. There was no significant difference between the sustained Ca²⁺ response of the nicotine-treated (145.2 ± 13.9 nM) and the control (121.7 ± 14.1 nM, P > 0.05) ASMCs. These results suggest that chronic exposure to a low level of nicotine enhances the nAChR mediated peak Ca²⁺ response, consistent with the upregulation of α7-nAChR expression in the nicotine-treated ASMCs.

The role of α7-nAChR in the nicotine-induced Ca²⁺ response was further examined pharmacologically in ASMCs, 100 µM nicotine elicited a biphasic response with a peak transient response of 293.1 ± 31.1 nM followed by a sustained phase of 142.3 ± 15.6 nM (n = 4, measured at 500-700 s after nicotine application) in control ASMCs. MLA (20 nM), a competitive antagonist specific for α7-nAChR, reversibly inhibited nicotine-induced Ca²⁺ response by 62.1± 2.2% (n = 5, P<0.05) in the peak and 50.2± 1.5% (P<0.05) in the sustained Ca²⁺ response (Fig. 6A,B). Another specific nAChR antagonist α-bungarotoxin (10 µM) also significantly inhibited the nicotine-induced Ca²⁺ response (Fig. 6C, D). The peak Ca²⁺ responses were 220.3 ± 15.4 nM (n = 5) and 96.5 ± 11.2 nM (n = 5, P< 0.05), and the sustained Ca²⁺ responses were 113.5 ± 15.6 nM (n = 5) and 61.8 ± 9.6 nM (n = 5, P<0.05) in the absence and presence of α-bungarotoxin, respectively. To further examine if α7-nAChR indeed mediates nicotine-induced Ca²⁺ response in ASMCs, ASMCs were transfect with α7-nAChR specific siRNA and scrambled control. siRNA transfection specifically decreased α7-nAChR protein expression after 72 hours by 65.4±1.8% (n=3), whereas negative control sequence did not substantially change the expression of the protein (Fig. 7A and B) Furthermore, nicotine-induced Ca²⁺ response was significantly abrogated in ASMCs transfected with α7-nAChR specific siRNA (Fig. 7C). The peak Ca²⁺ responses were 250.2 ± 23.4 nM (n = 4) and 61.3 ± 8.2 nM (n = 4, P< 0.05) in ASMCs transfected with α7-nAChR specific siRNA and scrambled sequence, respectively ( Fig. 7D), while the sustained Ca²⁺ responses were unaffected (control: 102.2 ± 11.6 nM; siRNA: 98.2 ± 10.4 nM). These results clearly suggest that α7-nAChR is responsible for a major component of the nicotine-induced Ca²⁺ response in ASMCs.

Chronic obstructive pulmonary disease (COPD) is an inflammatory airway disease characterized by different patterns of airway remodeling [41]. The decrease in lung function in COPD is associated with an increase in bronchial smooth muscle [42], which is likely the most important abnormality responsible for the airway narrowing in response to broncho-constricting stimuli [43]. The mechanisms underlying such remodeling of smooth muscle remain largely unclear; and the role of nicotine in this process has not been explored. In this study, we found that chronic nicotine treatment of ASMCs causes an increase in baseline [Ca²⁺]_i, which is an important determinant of airway smooth muscle growth and proliferation [44], in a concentration- and time-dependent fashion. We further found that rat ASMCs express multiple nAChR subtypes, including α2-α7, as well as β2 and β3. This is consistent with a previous report using immunohistochemistry showing that α4 and α7-nAChR proteins are expressed in mouse tracheal smooth muscle [45]. Among the nAChRs in ASMCs, α7-nAChR is one of the most abundant subtypes and possesses the unique functional property of high Ca²⁺ permeability [38,39,40]. Therefore, one may speculate that nicotine stimulates Ca²⁺ influx via α7-nAChR and perhaps other nAChRs to activate Ca²⁺-dependent pathways including CaM kinase II [13,46] to play a contributing role in the proliferation and contraction of ASMCs.

The pathological effect of nicotine on ASMC functions is supported by the observation that nicotine exposure increased α7-nAChR expression in ASMCs. This is consistent with previous studies showing that chronic exposure to nicotine up-regulate the expression of nAChRs, including α7-nAChR [47,48,49], in neuronal cells. The increase in α7 mRNA level in ASMCs suggests that nicotine, by binding to its receptors, enhances α7-nAChR transcription. Similar to our observation, chronic nicotine exposure causes the increase in mRNA levels of several nAChR subtypes in rat bladder [50]. In neurons, some nAChR subtypes undergo up-regulation, changes in stoichiometry and increase in active-state (functional up-regulation) after chronic nicotine exposure [48]. Up-regulation of α7-nAChRs at the post-transcriptional levels has also been reported in neuronal cell cultures [49]. The transcriptional and post-transcriptional regulation of nAChRs by nicotine could be related to differences between neurons and smooth muscle cells. A recent study suggest that nicotine-induced alterations of native neuronal nicotinic receptors are caused by multiple mechanisms [51], and the mechanisms could vary for different nAChR subtypes [52]. Hence, the regulatory mechanisms responsible for nicotine-induced nAChR expression in ASMCs require further investigations.

The present study also demonstrated for the first time that acute application of nicotine activates concentration-dependent Ca²⁺ response in ASMCs, suggesting that the nAChRs expressed are functional Ca²⁺ pathways in ASMCs. Moreover, chronic nicotine exposure increases the nicotine-induced peak Ca²⁺ response in ASMCs. This clearly suggests that nicotine induced persistent changes in nAChR functions, leading to an increased nAChR dependent activity in ASMCs. This is consistent with the increased expression of α7-nAChR mRNA and protein, and the upregulation is associated with increased functional nAChR channels in ASMCs [53]. The increase in the nicotine-induced Ca²⁺ response in the nicotine-treated cells could also be mediated in part by increased trafficking and membrane insertion of functional nAChRs, which is well-documented in neuronal cells[54]. However, whether chronic nicotine exposure also increases the conductance and activity of individual nAChR in ASMCs cannot be determined without single-channel recording on ASMCs.

The fact that nicotine is capable of inducing acute increase in [Ca²⁺]_i of ASMCs led us to further examine the pathways involved using specific nAChR blockers. Although mechanisms other than nAChR activation cannot be ruled out in the nicotine-induced increase in cytosolic Ca²⁺, the effective inhibition of the Ca²⁺ response by the α7-nAChR antagonists MLA and α-bungarotoxin indicates that α7-nAChR activation plays a major role in this process. The role of α7-nAChR in nicotine-mediated Ca²⁺ signaling was further confirmed in the RNA interference experiments, where ASMCs transfected with siRNA that selectively suppresses the expression of α7-nAChR substantially attenuated the nicotine-induced transient Ca²⁺ response. These results provide strong evidence that α7-nAChR is a major Ca²⁺ pathway mediating the nicotine-induced Ca²⁺ responses in ASMCs cells.

It has to be mentioned, however, that our study is focused mainly on the α7-nAChR because of its abundance in ASMCs and its Ca²⁺ permeability. Our results, however, do not exclude the possible participation of other nAChR subtypes in the nicotine-induced Ca²⁺ response and alterations in Ca²⁺ homeostasis in ASMCs. In fact, the partial inhibition of the sustained phase of nicotine-induced Ca²⁺ response by the α7-nAChR specific antagonist MLA and the persistent appearance of the sustained component in the siRNA transfected cells suggest that other nAChR subtypes expressed in ASMCs are involved particularly in the sustained Ca²⁺ response. The molecular identity of nAChR subtypes responsible for the sustained nicotinic Ca²⁺ response, which can be relevant for ASMCs proliferation, warrant further future investigations.

In conclusion, the present study demonstrates that (1) multiple nAChR are expressed in ASMCs; (2) chronic exposure of ASMCs increases resting [Ca²⁺]_i in ASMCs; (3) the expression of α7-nAChR is enhanced in ASMCs after chronic nicotine exposure; (4) acute application of nicotine elicits Ca²⁺ response in ASMCs and the Ca²⁺ response is enhanced in ASMCs chronically exposed to nicotine; and (5) the acute nicotine-induced Ca²⁺ response is mediated in part via α7-nAChR. These results highlight an important role of α7-nAChR in ASMC and suggest an intriguing mechanism that chronic nicotine exposure may activate a feed-forward mechanism to enhance α7-nAChR dependent effects in ASM contractility and remodeling. Future studies focusing on the relative roles of α7-nAChR and other nAChR subtypes will likely generate further interesting findings in nicotine-induced ASMC proliferation and airway remodeling.

The authors would like to thank Dr. Jian Wang, State Key Laboratory of Respiratory Disease, Guangzhou Institute of Respiratory Diseases, Guangzhou Medical University, for his many helpful comments and suggestions. This work was supported by Natural Science Foundation of Guangdong Province.