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Am J Physiol Lung Cell Mol Physiol 290: L526-L533, 2006. First published October 21, 2005; doi:10.1152/ajplung.00340.2005
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Thromboxane A2 induces airway constriction through an M3 muscarinic acetylcholine receptor-dependent mechanism

Irving C. Allen,1 John M. Hartney,1 Thomas M. Coffman,3 Raymond B. Penn,2 Jürgen Wess,4 and Beverly H. Koller1

1Curriculum in Genetics and Molecular Biology, University of North Carolina at Chapel Hill, Chapel Hill, North Carolina; 2Center For Human Genomics, Wake Forest University Health Science Center, Winston-Salem, North Carolina; 3Division of Nephrology, Duke University Medical Center, Durham, North Carolina; and 4Laboratory of Bioorganic Chemistry, National Institute of Diabetes and Digestive and Kidney Diseases, Bethesda, Maryland

Submitted 3 August 2005 ; accepted in final form 20 October 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Thromboxane A2 (TXA2) is a potent lipid mediator released by platelets and inflammatory cells and is capable of inducing vasoconstriction and bronchoconstriction. In the airways, it has been postulated that TXA2 causes airway constriction by direct activation of thromboxane prostanoid (TP) receptors on airway smooth muscle cells. Here we demonstrate that although TXA2 can mediate a dramatic increase in airway smooth muscle constriction and lung resistance, this response is largely dependent on vagal innervation of the airways and is highly sensitive to muscarinic acetylcholine receptor (mAChR) antagonists. Further analyses employing pharmacological and genetic strategies demonstrate that TP-dependent changes in lung resistance and airway smooth muscle tension require expression of the M3 mAChR subtype. These results raise the possibility that some of the beneficial actions of anticholinergic agents used in the treatment of asthma and chronic obstructive pulmonary disease result from limiting physiological changes mediated through the TP receptor. Furthermore, these findings demonstrate a unique pathway for TP regulation of homeostatic mechanisms in the airway and suggest a paradigm for the role of TXA2 in other organ systems.

nerve; prostanoid; bronchoconstriction; asthma; vagus

THROMBOXANE A2 (TXA2) was originally identified as the active agent in extracts of human platelets and was subsequently shown to be capable of initiating the contraction of both vascular and airway smooth muscle (ASM) (21). TXA2 synthesis is initiated by the oxidation of arachidonic acid by either prostaglandin G/H synthase-1 (or cyclooxygenase-1; COX-1) or prostaglandin G/H synthase-2 (COX-2) (21). The PGH2 produced by the cyclooxygenases is metabolized to TXA2 by thromboxane synthase, an enzyme expressed by many cell types. In the lung, for example, TXA2 is produced by a number of cells, including the epithelia, smooth muscle, and resident macrophages (39, 56). In aqueous solutions, TXA2 is rapidly hydrolyzed to TXB2, a stable and inactive metabolite (21). The short half-life of TXA2 suggests that it functions in an autocrine/paracrine fashion and that its actions are limited to tissues in proximity to the source of its synthesis. Because of the instability of TXA2, most experimental studies of TXA2 biology have utilized stable TXA2 mimetics such as U-46619 (8, 9, 32).

The actions of TXA2, as well as those of other prostanoids, are mediated through binding to specific heterotrimeric G protein-associated prostanoid receptors. With the cloning of the thromboxane prostanoid (TP) receptor and the generation of a mouse line lacking expression of this gene, all of the in vivo actions of TXA2 studied to date have been shown to be dependent on expression of this single G protein-coupled receptor (30, 40, 45, 52). Most evidence supports the coupling of this receptor to the Gq family of proteins, and many of its physiological actions have been attributed to Gq-mediated activation of phospholipase C and increases in intracellular calcium concentration ([Ca2+]i). In addition to ASM, many other tissues and cell types in the mouse have been reported to express TP receptors, including cells of the immune system and epithelial cells (11, 13, 22, 37, 38, 49, 54, 56). TP receptor expression on neurons has also been implied from pharmacological studies (26, 29).

Exposure of human tracheal rings to U-46619 results in cumulative, concentration-dependent contractions (3). In studies examining isolated human bronchial smooth muscle, U-46619 was shown to be 300 times more potent as a constricting agent than other prostanoids. These ex vivo studies were consistent with studies in humans (27) and other animals (34) demonstrating that inhalation of U-46619 results in rapid bronchoconstriction (12). Furthermore, some studies have even suggested that other constricting prostanoids, such as PGF2 and PGD2, may mediate constriction by binding the TP receptor (10, 14).

Concentrations of TXA2 and other prostanoids in the airway are elevated in a number of lung diseases including asthma and chronic obstructive pulmonary disease (COPD) (15, 41, 42). In addition to higher concentrations, airway sensitivity to these lipid mediators is also increased in disease states (44). Although TXA2, and possibly other lipid mediators, can mediate their effects by engaging the TP receptor on ASM, the mechanism by which TP receptor activation leads to bronchoconstriction is not well defined. Here we address this question directly using both genetic and pharmacological approaches in conjunction with both in vivo and ex vivo studies of mouse airways. Similar to observations in other species including humans, we show that inhaled TXA2 leads to an increase in lung resistance and that TXA2 can also mediate constriction of tracheal rings ex vivo. Analysis of M3 muscarinic acetylcholine receptor (mAChR) subtype knockout mice showed that the in vivo airway constrictor responses to TXA2 are primarily dependent on parasympathetic innervation of the lungs and the presence of functional M3 mAChRs.


   METHODS
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 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
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Experimental animals. All studies were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and were approved by the Institutional Animal Care and Use Committee of the University of North Carolina at Chapel Hill (protocol no. 05-163). Mice deficient in the TP receptor were generated and genotyped by Southern blot analysis or PCR as previously described (52). The 129 TP–/– mice were obtained by breeding chimeras generated from TP+/– 129/Olac embryonic stem cells directly with 129/SvEv females. Heterozygous animals were crossed for two consecutive generations to 129/SvEv animals to reduce the contribution of the 129/Olac substrain. The generation of mAChR3–/– mice has been described previously (58). The mAChR3–/– and corresponding mAChR+/+ control mice were maintained on a 129/SvEv (50%) x CF1 (50%) mixed genetic background. All experiments were carried out using 8- to 12-wk-old mice.

Measurement of airway reactivity in intubated mice. Mice were anesthetized with 70–90 mg/kg pentobarbital sodium (American Pharmaceutical Partners, Los Angeles, CA), tracheostomized, and mechanically ventilated at a rate of 300 breaths/min, a tidal volume of 6 ml/kg, and a positive end-expiratory pressure (PEEP) of 3–4 cmH2O with a computer-controlled small animal ventilator (Scireq, Montreal, Canada). Once ventilated, mice were paralyzed with 0.8 mg/kg pancuronium bromide. Dynamic lung resistance (RL) and dynamic compliance (Cdyn) were determined by transducing airway pressure and airflow using a precisely controlled piston during a single inspiration and expiration with an amplitude of 150 µl and a period of 1 s (SnapShot). The single-compartment equation of motion (Eq. 1) allows determination of RL and Cdyn (25)

Formula 1(1)
where P(t) = pressure at time t, R = resistance, Formula 1(t) = flow at time t, C = compliance, V(t) = volume at time t, and P0 = resting pressure (PEEP).

Five separate baseline measurements of parameters were taken before exposure to U-46619. After baseline assessments, aerosols of (15S)-hydroxy-9,11-epoxymethanoprosta-5Z,13E-dienoic acid (U-46619; Cayman Chemical, Ann Arbor, MI), at doses indicated (generally 10–6 M to 10–3 M), were delivered via nebulizer through a side port in the ventilator circuit for 30 s at a rate of 200 breaths/minute. Because a response to U-46619 was observed only after 10–4 M and 10–3 M doses, these concentrations were used in lieu of the full range of doses in some experiments. After U-46619 exposure, RL was measured every 10 s for 3 min. To determine the contribution of mAChRs to the U-46619 response, mice received either an intraperitoneal injection of atropine sulfate (10 µM/kg; American Pharmaceutical Partners) or an intravenous injection of hexahydro-sila-difenidol p-fluorohydrochloride (4-DAMP; 4 ng/g, Sigma Chemical, St. Louis, MO). Similar to other studies measuring airway resistance using these techniques (4, 23, 53), data are presented as percent baseline RL.

A surgical vagotomy was conducted to assess the contribution of intact parasympathetic innervation. While the mice were anesthetized, the vagal nerves on each side of the esophagus were identified under a dissecting microscope and dissected free of the fascia and carotid artery. A silk suture was passed under each vagus to allow easy identification and severing of the nerve in the ventilated animals. Mice were then tracheostomized and placed on a ventilator (Flexivent). After mice were treated with the paralytic agent, five snapshots were recorded. In the experimental animals, after establishment of basal lung mechanics, both vagal nerves were severed. Postvagotomy baseline RL and Cdyn were then reassessed, and the mice were challenged with U-46619.

Measurement of tension development in tracheal rings. Mice were killed by inhalation of CO2. The tracheae were then rapidly excised in 3- to 4-mm segments, cleaned of superficial fat and connective tissue, and placed in Krebs-Henseleit solution (53). These segments were mounted between two triangular stainless steel hooks, put into double-jacketed glass organ baths containing Krebs-Henseleit solution (maintained at a pH of 7.40–7.45), and continuously gassed with carbogen (5% O2 and 5% CO2). The upper support for the tracheal segment was attached, via silk thread, to an FT03 isometric transducer (Astro-Med, West Warwick, RI), and force generation was recorded with an MP 100WS system (BIOPAC Systems). The rings were equilibrated in the respective buffered solutions for 30 min at a predetermined resting tension based on experimental calibration (~0.5 g) (18, 47, 51). Rings were then preconstricted with methacholine (MCh; 10–8 M), allowed to equilibrate for 10 min, washed three to four times, and reequilibrated to 0.5 g. A dose-response curve was then generated with either MCh (10–8 M to 10–3 M) or U-46619 (10–10 M to 10–3 M) (50).

To assess the contribution of muscarinic receptors, atropine (10–6 M) was added directly to the buffers of selected rings 15 min before experimental challenges. The response to U-46619 (50–500 nM) was examined as well as the response to a single dose of MCh (100 µM) delivered immediately after the final dose of U-46619. The response to this application of MCh is shown in the figures. The rings were then washed three to four times, and a final dose of MCh (100 µM) was added to ensure tissue viability (data not shown). To assess the contribution of the M3 mAChR, rings from mAChR3–/– and mAChR3+/+ mice were preconstricted with serotonin (5-HT, 20 mM), allowed to equilibrate for 10 min, washed three to four times, and reequilibrated to 0.5 g. The response to U-46619 (50 and 100 nM) was examined as was the response to a single dose of 5-HT (20 mM) delivered immediately after the final dose of U-46619. The response to this application of 5-HT is shown in the figures.

All experiments and dissections were conducted in the presence of indomethacin (10–6 M; Sigma-Aldrich, St. Louis, MO) to prevent endogenous prostaglandin release. Similar to other studies using these techniques (36, 53, 57), data are presented as mg developed tension/mg tissue weight.

Ovalbumin sensitization and airway challenge. Groups of mice were sensitized by intraperitoneal injection of 20 µg of ovalbumin (OVA; Grade V, Sigma) emulsified in 2.25 mg of aluminum hydroxide (Sigma) in a total volume of 200 µl on days 1 and 14. Mice were challenged (45 min) via the airways with OVA (1% in saline) for 3 days (days 21–23) using a jet nebulizer (TSI Jet Neb). Control mouse groups received the two OVA immunizations but were challenged with aerosolized saline. Airway mechanics were assessed 24 h after the final aerosol OVA or saline challenge (day 24).

After airway assessments, mice were killed, and ~1 ml of blood was collected by heart stick. Blood was allowed to coagulate, and the serum was collected. Total IgE levels were determined by ELISA (ICN Biomedicals). Bronchoalveolar lavage (BAL) was performed five times with 1.0 ml of sterile Hanks' buffered saline solution each time. Cells present in the BAL fluid were determined using a hemacytometer. The recovered BAL fluid was centrifuged to remove cells, and the IL-13 levels in the supernatant were determined using ELISA (R&D Systems).

Statistical analysis. Data are presented as means ± SE. A random effects model followed by Tukey's multiple-comparison test was utilized to assess dose-response data. ANOVA followed by Tukey's multiple-comparison test was performed on complex data sets. Statistical significance for single data points was assessed by Student's two-tailed t-test. A P value of <0.05 was considered statistically significant.


   RESULTS
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 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
TXA2 increases airway constriction in a TP receptor-dependent manner. TXA2 has been shown to cause bronchoconstriction in a number of species. Because of the short half-life of TXA2, experimental investigation into TXA2/TP receptor-dependent effects typically employs the synthetic agonist U-46619. Preliminary experiments verified that under our experimental conditions and with doses of U-46619 generally used in in vivo experiments, all of the actions of this mimetic were dependent on the expression of the TP receptor. U-46619 at concentrations up to 1 mM failed to elicit an increase in RL from mice in which the gene encoding the TP receptor was disrupted by homologous recombination (TP–/–) (Fig. 1A). In contrast, RL in wild-type mice increased in response to 100 µM and 1 mM U-46619 (Fig. 1A). Conversely, MCh challenge resulted in similar increases in RL in wild-type and TP–/– mice (Fig. 1B). We next examined the ability of U-46619 to constrict mouse trachea. A very steep dose response was observed: U-46619 elicited no significant increase in tension at doses <10 nM, whereas maximal responses were achieved with 100 nM of this agonist (data not shown). Consistent with our in vivo data, ex vivo tension generation in tracheal ring preparations was similar in rings from wild-type and TP–/– mice stimulated with MCh, whereas U-46619 caused significant tension generation in rings from wild-type mice but not in rings isolated from TP–/– mice (Fig. 1C).


Figure 1
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  		Fig. 1. Changes in airway physiology of wild-type and TP–/– mice in response to the thromboxane A2 (TXA2) analog U-46619 and methacholine (MCh). A: change in lung resistance (RL) of TP+/+ and TP–/– mice in response to U-46619. After baseline measurements, the airways were exposed to aerosolized vehicle followed by increasing doses of aerosolized U-46619 (10–6 M to 10–3 M). The percent change in RL from baseline increased significantly in the wild-type mice exposed to 10–4 M and 10–3 M U-46619 compared with no increase in RL in TP–/– mice. TP+/+ mice, n = 34; TP–/– mice, n = 13 (*P < 0.05; **P < 0.005). B: TP–/– and TP+/+ mice demonstrated similar increases in RL after exposure to MCh. TP+/+ and TP–/– mice were subjected to aerosolized vehicle and 1 bolus dose of aerosolized MCh (50 mg/ml = 255.1 mM). The percent increase from baseline was similar for both TP+/+ and TP–/– mice. TP+/+ mice, n = 5; TP–/– mice, n = 5 (*P, #P < 0.05). C: contraction of TP–/– and TP+/+ tracheal rings. Excised tracheal rings from both TP+/+ and TP–/– mice were challenged with a single dose of U-46619 (1 µM), and the change in tension was recorded. This was followed by treatment with MCh (100 µM). Ring tension was significantly increased after addition of U-46619 in rings from TP+/+ mice, whereas no increase was observed in rings from TP–/– mice. Ring tension was significantly increased in response to MCh in both groups. TP+/+ mice, n = 3; TP–/– mice, n = 3 (**P < 0.007; #P, *P < 0.05).

 
Vagotomy significantly attenuates U-46619-induced increases in RL. The simplest explanation for the observed changes in RL and ex vivo tension generation is that binding of U-46619 to TP receptors expressed by ASM directly activates Gq-coupled pathways leading to increases in [Ca2+]i and that this constriction is independent of the activity of the parasympathetic innervation of the airways. To test this model, we examined the ability of U-46619 to elicit an increase in RL in animals in which the vagus nerves were surgically severed just proximal to the nodose ganglion. Sham-operated animals, treated identically to the experimental animals except for the final severing of the nerve, served as controls. Baseline RL was assessed for 5 min before and 5 min after vagotomy. Baseline RL was higher in the mice in which the surgeries had been performed compared with values generally observed for mice on the 129/SvEv background, likely reflecting local mediator release during the surgical procedure. As expected, the RL of the sham-operated animals increased in response to U-46619. However, the magnitude of the percent change in RL was smaller than that previously measured in wild-type 129 animals. This likely reflects the higher baseline RL in the surgically manipulated animals. A slight but significant drop in the baseline RL (Fig. 2A) and an increase in airway compliance (data not shown) were observed after vagotomy. This change in baseline resistance was expected, as similar changes have been observed in other species after vagotomy (6, 7, 33). The drop in RL reflects the loss of basal parasympathetic tone in the airways. After establishment of postvagotomy baseline RL, mice were challenged with U-46619 (10–4 M and 10–3 M). Although U-46619 still elicited an increase in RL in the vagotomized animals, the response was significantly attenuated compared with that observed in the sham-operated animals (P < 0.05, Fig. 2B). These data suggest that in vivo, TXA2 functions in part via an indirect neurally mediated mechanism to induce ASM reactivity. This neurally mediated mechanism functions in a concerted manner with other mechanism(s) that are not dependent on vagal innervation of the airways.


Figure 2
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  		Fig. 2. Vagotomy significantly attenuates change in airway resistance induced by U-46619 challenge. A: parasympathetic tone in mouse lung. Mice were anesthetized, and the vagal nerve was dissected free of the fascia and carotid artery. Mice were tracheostomized and placed on a ventilator (Flexivent). After treatment with a paralytic agent, 5 snapshots were recorded. In 1 group of animals, after establishment of basal lung mechanics, the vagus nerves were severed. The percent change in RL from the precut baseline decreased significantly in vagotomized mice compared with the surgical controls. TP+/+ mice, n = 10; vagotomized TP+/+ mice, n = 9 (*P < 0.05). B: the response to U-46619 is attenuated in vagotomized mice. After the baseline assessment described above, vagotomized mice and the surgical controls were exposed to vehicle for 20 s, and airway reactivity was assessed every 10 s for the next 3 min. Mice were then exposed to increasing doses of the TXA2 analog U-46619, and the change in lung mechanics after each dose was recorded. The response of the vagotomized mice was significantly attenuated at the highest dose of U-46619 (*P < 0.05). Importantly, a significant response to U-46619 could still be measured in the vagotomized animals.

 
Atropine attenuates U-46619-induced ASM constriction. Previous studies investigating the ability of TXA2/U-46619 to meditate direct actions on smooth muscle cells utilized ex vivo preparations of either tracheal rings or strips of ASM prepared from tracheae or bronchi. In our studies, tracheal rings were isolated from wild-type and TP–/– mice and stimulated with increasing doses of U-46619 (Fig. 3A). Rings from untreated TP–/– mice were included in the study to control for nonreceptor-mediated actions of U-46619 (data not shown). As expected, U-46619 increased tension in the rings from wild-type mice. No response was observed in the TP–/– animals. To determine whether the response observed with the wild-type preparations was dependent on muscarinic cholinergic pathways, we next examined the ability of atropine (10–6 M), a nonselective mAChR antagonist, to block this response. At all stimulating concentrations of U-46619, pretreatment of rings with atropine significantly attenuated tension generation. However, U-46619 at doses >100 nM provoked a significant change in tension in the atropine-treated rings over baseline (P < 0.005, Fig. 3A). As expected, in the presence of atropine, addition of MCh to rings contracted with 500 nM U-46619 did not evoke a further increase in tension (Fig. 3A). To ensure that the atropine-pretreated tracheal rings were still capable of MCh-induced contraction, the rings were washed multiple times to remove the atropine. MCh was subsequently added back to the ring buffer. In this case, MCh increased tension in all groups of mice, regardless of previous treatment protocol or genotype (data not shown).


Figure 3
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  		Fig. 3. Atropine attenuates U-46619-mediated changes in airway smooth muscle (ASM) constriction and RL. A: atropine partially blocks the contractile response to U-46619. Excised tracheal rings from TP+/+ mice were incubated in either the presence or absence of atropine (10–6 M), challenged with increasing doses of U-46619 (50–500 nM), and challenged with a bolus dose of MCh (100 µM). Atropine significantly reduced the average tension during all doses tested. TP+/+ mice with atropine, n = 17; TP+/+ mice, n = 11 (**P < 0.005). B: atropine attenuates U-46619-mediated changes in RL. The change in RL in response to U-46619 in intubated TP+/+ mice pretreated with atropine (10 µM/kg ip), TP+/+ mice pretreated with saline, and TP–/– mice was determined. Mice were subjected to aerosolized vehicle followed by increasing doses of aerosolized U-46619 (from 10–6 to 10–3 M; 10–6 and 10–5 M data not shown). After the U-46619 challenge, the response to aerosolized MCh (50 mg/ml = 255.1 mM) was measured (data not shown). A dose-dependent increase in RL was observed in the TP+/+ mice in response to U-46619, and this response was absent in the TP–/– animals. Pretreatment with atropine dramatically attenuated the response of TP+/+ mice to U-46619. As expected, atropine pretreatment inhibited the MCh response (data not shown). Atropine-pretreated TP+/+ mice, n = 10; saline-pretreated TP+/+ mice, n = 10; TP–/– mice, n = 11 (§P < 0.05; *P < 0.05; **P < 0.005).

 
Atropine inhibits U-46619-induced increases in RL. We next evaluated the ability of atropine to block U-46619-mediated increases in RL. TP–/– and wild-type mice were pretreated with either atropine (10 µM/kg) or saline. Untreated TP–/– mice were included in the study to control for TP receptor-independent effects of U-46619. Mice were intubated, baseline RL was established, and the mice were then exposed to vehicle followed by aerosols of U-46619 (10–4 M and 10–3 M). As expected, a significant increase in RL was observed in the saline-pretreated wild-type mice exposed to U-46619 (Fig. 3B). The U-46619 response in the atropine-pretreated mice was significantly attenuated compared with the saline-pretreated animals (Fig. 3B). As previously observed, TP–/– mice failed to respond to U-46619 (Fig. 3B).

The M3 mAChR receptor-preferring antagonist 4-DAMP significantly attenuates U-46619-induced increases in RL. ASM expresses the Gq/11-coupled M3 mAChR, and both pharmacological studies and studies with mice lacking this receptor indicate that airway constriction in response to cholinergic agents is mediated primarily through this receptor (19, 43, 46, 48). The muscarinic antagonist 4-DAMP preferentially binds to the M3 mAChR receptor (5); however, it also displays high affinity for M1, M4, and M5 mAChRs (5). Therefore, we determined the lowest dose of 4-DAMP that completely blocks MCh-induced changes in airway resistance in the mouse. This dose of 4-DAMP (4 ng/g; data not shown), when delivered by intravenous injection before challenge with U-46619, significantly inhibited U-46619-induced increases in RL, with values never significantly exceeding those recorded for the TP–/– mice (Fig. 4).


Figure 4
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  		Fig. 4. The M3 muscarinic acetylcholine receptor (mAChR)-preferring antagonist hexahydro-sila-difenidol p-fluorohydrochloride (4-DAMP) attenuates U-46619-mediated RL. The change in RL in response to U-46619 in intubated TP+/+ mice pretreated with 4-DAMP (4 ng/g; iv injection), TP+/+ mice treated with vehicle, and TP–/– mice was determined. A dose-dependent increase in RL was observed in the TP+/+ mice in response to U-46619, and this response was absent in the TP–/– animals. Pretreatment with 4-DAMP inhibited the response of wild-type mice to U-46619, and no significant increase in RL was observed in this group over that measured in TP–/– animals. 4-DAMP-pretreated TP+/+ mice, n = 10; vehicle-pretreated TP+/+ mice, n = 21; TP–/– mice, n = 3 (§P < 0.05; #P < 0.05; *P < 0.005).

 
Mice lacking the M3 mAChR demonstrate a significant decrease in U-46619-induced ASM constriction and RL. To further demonstrate the role of muscarinic cholinergic pathways in TXA2-mediated changes in airway constriction, we examined the response to U-46619 in mice lacking the M3 mAChR (mAChR3–/–). Tracheal rings from mAChR3–/– and age/strain-matched control mice were exposed to constricting doses of U-46619. A significant increase in tension was observed in the wild-type control animals after exposure to 50 and 100 nM of agonist. Consistent with the ability of atropine to modulate contraction of the tracheal rings, response of the mAChR3–/– tracheae was dramatically attenuated at both concentrations of U-46619 examined. However, a small but significant increase in constriction was still observed in mAChR3–/– tracheal rings in response to U-46619 (Fig. 5A). To verify that the altered response of the mAChR3–/– rings did not reflect a fundamental defect in smooth muscle function as a consequence of loss of this receptor and to verify that the preparations used in this study were viable, we examined the response of these same tracheae to serotonin. A robust increase in tension was observed after treatment of both wild-type and mAChR3–/– trachea with this constricting agent.


Figure 5
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  		Fig. 5. ASM constriction and RL after U-46619 treatment is attenuated in mice lacking the M3 mAChR (mAChR3–/–). A: tracheal rings from mAChR3–/– mice demonstrated attenuated contractile responses to U-46619. Excised tracheal rings from mAChR3+/+ and mAChR3–/– mice were exposed to U-46619 (50 and 100 nM), and changes in tension were recorded. Ring tension was significantly increased after addition of U-46619 in rings from mAChR3+/+ mice. A small but significant increase in average tension was observed in mAChR3–/– mice at both U-46619 doses. To verify the viability of the rings, these manipulations were followed by treatment with a constricting dose of serotonin (5-HT, 20 mM). No difference in 5-HT-mediated constriction was observed between the mAChR3+/+ and mAChR3–/– tracheal rings. mAChR3+/+ mice, n = 7; mAChR3–/– mice, n = 8 (**P < 0.005; #P < 0.05; {ddagger}P < 0.05). B: the change in RL in response to U-46619 in intubated mAChR3+/+ mice was significantly attenuated. Mice were subjected to a 20-s aerosolization of increasing doses of U-46619 (10–4 to 10–3 M). A dose-dependent increase in RL was observed in the mAChR3+/+ mice in response to U-46619, and this response was significantly attenuated in the mAChR3–/– animals. mAChR3+/+ mice, n = 5; mAChR3–/– mice, n = 7 (*P < 0.05; **P < 0.005).

 
We next examined the ability of thromboxane to increase RL in mice lacking the M3 mAChR. As seen in Fig. 5B, loss of the M3 mAChR significantly attenuates the U-46619-induced increases in RL. More than 80% of the response to U-46619 (10–3 M) was lost in the mAChR3–/– mice. However, consistent with our findings in experiments examining the effects of atropine, mAChR3–/– mice showed a small but significant increase in RL in response to U-46619 (Fig. 5B). These data suggest a predominant role for the M3 mAChR in TXA2-mediated airway constriction.

TP receptor-mediated changes in RL are enhanced in the allergic inflamed airway but remain sensitive to atropine. Allergic airway inflammation was induced in wild-type and TP–/– mice. All mice were immunized via intraperitoneal injection with OVA/alum. To induce allergic lung disease, mice were challenged with aerosols of OVA. Control animals were exposed to aerosols of saline. Twenty-four hours after the final exposure to antigen or saline, mice were intubated and exposed to increasing doses of U-46619. As depicted in Fig. 6, the response to both 10–4 M and 10–3 M U-46619 was substantially increased in OVA-challenged wild-type mice compared with saline-exposed control animals. Thus, as observed in humans, the bronchoconstricting actions of TXA2 are dramatically enhanced in the inflamed airway (1). No significant change in RL in response to U-46619 was observed in TP–/– mice, confirming that even in the inflamed airway, the response to U-46619 remains dependent on TP receptor expression (Fig. 6). We next determined whether, similar to the response in naïve mice, the response of mice with allergic lung disease to U-46619 was sensitive to atropine. Alternatively, we considered the possibility that the M3 receptor-independent TP response is amplified in the inflamed lung. To distinguish between these two possibilities, a group of OVA-challenged mice and their corresponding saline-challenged controls received a single intraperitoneal treatment of atropine just before intubation and measurement of airway mechanics. As seen in Fig. 6, the response of the mice with allergic lung disease to U-46619 was dramatically reduced. The response remaining in these animals was not significantly greater than the response to U-46619 measured in atropine-treated naïve animals. Thus the mechanism by which U-46619 mediates the increased responsiveness in the diseased animal is similarly dependent on an intact muscarinic cholinergic pathway(s).


Figure 6
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  		Fig. 6. Atropine inhibits U-46619-mediated changes in resistance in the inflamed lung. Atropine inhibits U-46619-mediated changes in RL after challenges with 1% ovalbumin (OVA). The change in RL in response to U-46619 in intubated TP+/+ mice, TP+/+ mice pretreated with atropine (10 µM/kg ip), TP+/+ mice treated with saline, and TP–/– mice was determined. Mice were subjected to a 30-s aerosolization of increasing doses of U-46619 (10–4 and 10–3 M), followed by 3 min of response measurements. A dose-dependent increase in RL was observed in the TP+/+ mice and in the OVA-challenged TP+/+ mice in response to U-46619, and this response was absent in the TP–/– animals. Pretreatment with atropine inhibited the response of the wild-type mice to U-46619. Furthermore, no significant increase in RL was observed in the atropine-pretreated group over that measured in TP–/– animals. TP+/+ mice, n = 11; atropine-pretreated TP+/+ mice, n = 6; saline-pretreated TP+/+ mice, n = 24; TP–/– mice, n = 6 (*P < 0.05; #P < 0.05; {lozenge}P < 0.05; {ddagger}P < 0.05).

 
At the end of the measurements of lung mechanics, as described above, the induction of allergic airway disease was verified by determining serum IgE levels, cellularity of BAL fluid, and histopathology of the lungs. Increases of similar magnitude were observed in total BAL fluid cell counts (3.3 x 106 total cells in OVA-treated animals vs. 1 x 105 in saline-treated animals) and airway eosinophilia (~80% of cells present were eosinophils) between the OVA-challenged TP–/– and wild-type mice (data not shown). Likewise, a similar increase in IgE levels (2,500 ng/ml in OVA vs. 225 ng/ml in saline) and IL-13 levels (550 pg/lung after OVA challenge vs. undetectable in saline-treated) was observed between the OVA-challenged TP–/– and wild-type mice (data not shown).


   DISCUSSION
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
TXA2 is a potent mediator of human airway constriction. The expression of the TP receptor on ASM, combined with the early demonstration of the ability of TXA2 to mediate constriction of vascular smooth muscle, suggested that TXA2-induced increases in airway resistance are mediated through direct actions of TXA2 on ASM. However, results from the present study demonstrate that in both the naïve and inflamed airway, the constrictor effect of TXA2 is dependent on the activity of muscarinic cholinergic pathways.

We utilized both pharmacological and genetic approaches to establish the role of the cholinergic pathways in TP receptor-mediated changes in airway and RL. U-46619-mediated increases in RL were largely attenuated when animals were treated with the nonselective muscarinic receptor antagonist atropine or the M3 receptor-preferring antagonist 4-DAMP. Consistent with these findings, U-46619 caused only minimal increases in RL in both mAChR3–/– mice (suggesting a primary role for the M3 mAChR subtype) and vagotomized mice (suggesting a dependence on parasympathetic innervation).

A number of mechanisms that are consistent with our findings can be proposed. For example, stimulation of ASM TP receptors may result in an increase in [Ca2+]i. This increase in Ca2+ is insufficient to mediate significant ASM contraction. However, binding of TXA2 to the TP receptor potentiates the activity of the M3 mAChR, thereby increasing the sensitivity of the M3 mAChR to its cognate physiological ligand ACh. With the loss of vagal tone, the release of ACh diminishes, as does the ability of the muscle to respond to TP receptor activation. A number of possible mechanisms by which TXA2 binding to the TP receptor can alter the activity of a coexpressed Gq-coupled receptor can be envisioned, including the formation of heterodimers or simple additive effects of receptor-specific-induced changes in [Ca2+]i or other second messengers. In support of this mechanism, both TP receptors (16, 24) and muscarinic receptors (31, 55, 59, 60) have been shown to form heterodimers with other G protein-coupled receptors. Implicit in this model is the existence of a basal tone of the ASM and constitutive release of ACh in the unprovoked airway. We report here, consistent with findings in other species (6, 7, 20, 33), that a basal ASM tone can be measured in mice, as indicated by a decrease in RL after vagotomy.

Our data are also consistent with alternate pathways, which also involve constitutive release of ACh from parasympathetic nerves. In this example, the release of ACh from the postsynaptic neurons is enhanced by TXA2 binding to presynaptic TP receptors on the vagus nerve. Activation of these TP receptors on the nerve or nerve terminals results in a substantial increase in ACh release. After vagotomy, this constitutive release is diminished and thus is no longer potentiated by activation of TP receptors on nerve termini. Consistent with this model, TP receptors are expressed throughout the central nervous system and spinal cord. However, there is little information concerning the expression of TP receptors by either sensory or afferent nerves of the airways. This model, in which TP receptor activation mediates constriction by modulating neural activity, is consistent with early studies in dogs (2). These studies showed that TXA2 potentiated vagal nerve neuroeffector transmission in ASM tissue. Contractions of canine tracheal smooth muscle (ex vivo preparations) were induced by electrical field stimulation (EFS) or by ACh in the presence or absence of U-46619. Although U-46619 had no effect on the contractile response of ACh when applied exogenously to smooth muscle, it significantly increased the amplitude of the EFS-evoked contractions in an atropine-sensitive manner (2). These results suggest that, in canines, U-46619 has a prejunctional action stimulating increased ACh release from vagal nerve terminals through TP receptors (2). There is some limited evidence that TXA2 can directly stimulate nerves. Nerve fibers isolated from felines, including unmyelinated vagal afferent nerves from the lung and nerves originating from skeletal muscle of the hindlimb, were reported to respond to U-46619 (28, 29).

Both mechanisms described above are consistent with the observation that atropine decreased the response of tracheal rings to TXA2 and that rings from mice lacking the M3 mAChR have a diminished response to TXA2. Previous tracheal ring studies have demonstrated that after removal of vagal signaling, the concentration of ACh at the neuroeffector junction is directly proportional to its rate of release, rate of diffusion, and rate of enzymatic hydrolysis (20). ACh levels are preserved at neuroeffector junctions, and these stores are capable of maintaining muscarinic tone and effective cholinergic signaling for an extended amount of time after eliminating efferent vagal activity (20).

TXA2 is a potent vaso- and bronchoconstricting agent. We demonstrate here that the full action of TXA2 is dependent on intact parasympathetic innervation and the presence of M3 mAChRs in both the healthy and inflamed airway. These findings also raise the possibility that the TP receptor interacts with muscarinic cholinergic pathways in other biological systems. Anticholinergic agents continue to be used in the treatment of severe asthma and COPD. Our work suggests that these drugs, by blocking M3 receptors, may act downstream of a number of important bronchoconstricting agents, such as TXA2, that mediate their actions through prostanoid receptors.


   GRANTS
TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work is supported by National Heart, Lung, and Blood Institute Grants HL-068141 (to B. H. Koller) and HL-58506 (to R. B. Penn).


   ACKNOWLEDGMENTS
 
The authors thank Dr. Julia Walker, Barbara Lawson, Leigh Jania, MyTrang Nguyen, Julie Ledford, and Se-Wei Wang for technical support and Amy Pace for manuscript preparation.


   FOOTNOTES
 

Address for reprint requests and other correspondence: B. H. Koller, Curriculum in Genetics and Molecular Biology, Univ. of North Carolina at Chapel Hill, Chapel Hill, NC 27599 (e-mail: Treawouns{at}aol.com)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
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 ABSTRACT
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