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Mrgprs on vagal sensory neurons contribute to bronchoconstriction and airway hyperresponsiveness

Asthma, accompanied by lung inflammation, bronchoconstriction, and airway hyperresponsiveness, is a significant public health burden. Here we report that G protein-coupled receptor Mrgprs are expressed in a subset of vagal sensory neurons innervating the airway and mediates cholinergic bronchoconstriction and airway hyperresponsiveness. These findings provide novel insights into the neural mechanisms underlying the pathogenesis of asthma.

receptor, is expressed in dorsal root ganglion (DRG) sensory neurons innervating the skin and mediates non-histaminergic itch 6,7 . We found that MrgprC11 is specifically expressed in vagal ganglia besides DRG ( Supplementary Fig. 1a). Vagal ganglia consist of two different ganglia, termed nodose and jugular ganglia, with distinct embryonic origins: nodose neurons are derived from the epibranchial placode, while jugular neurons are derived from the neural crest 8 . Using Wnt1-Cre;R26-lsl-LacZ mice, in which all the neural crest derived neurons are expressing LacZ, we found that MrgprC11 is selectively expressed in jugular sensory neurons, but not in nodose sensory neurons (Fig. 1a-c, Supplementary Fig. 1c-e and Supplementary Fig. 2). Intriguingly, hMrgprX1, the functional orthologous of MrgprC11, is also selectively expressed in jugular ganglion ( Supplementary Fig. 1b).
MrgprC11 immunoreactivity was detected in 6.15% (84/1365) of jugular sensory neurons and in 2.00% (84/4190) of total vagal sensory neurons. MrgprC11 + jugular sensory neurons were of small diameter (average 200.27 ± 9.24 μm 2 ), positive for nociceptor marker TRPV1 (Fig. 1d-f), and negative for myelinated neuron marker neurofilament 200kD ( Supplementary Fig. 3). We performed retrograde tracing to determine whether MrgprC11 + jugular sensory neurons innervate the airway. We observed a total of 750 airway-innervating vagal sensory neurons labeled by retrograde tracer CTB-488 (cholera toxin b subunit conjugated with Alexa 488), and 64 of them were positive for MrgprC11 ( Fig. 1g-i), demonstrating that MrgprC11 + jugular neurons innervate the airway. However, among the 201 airway-innervating DRG neurons, none of them was positive for MrgprC11 ( Fig. 1j-l). Therefore, we concluded that MrgprC11 + primary sensory neurons in different sensory ganglia exhibit different peripheral innervation patterns: MrgprC11 + jugular sensory neurons innervate the airway, whereas MrgprC11 + DRG sensory neurons innervate the skin (Fig.  1m) 7,9 . These results allow us to attribute the physiological responses in the airway induced by the activation of MrgprC11 to jugular sensory neurons in the following experiments.
To elucidate the function of MrgprC11 + jugular sensory nerves in the airway, we performed head-out body-plethysmography. Aspiration of Bam8-22 into the airway evoked a change in the respiratory pattern characterized by an increase in the amplitude of the respiratory waveform and a slight but significant increase in the respiratory rate ( Fig. 1n-p and Supplementary Fig. 7a-c). In contrast, Bam8-22 did not induce any change in respiration in Mrgpr-clusterΔ −/− mice (Fig. 1o, p, Supplementary Fig. 7a-c). The Bam8-22-induced respiratory effect was similar to that observed upon the administration of methacholine, a bronchoconstrictor acting on muscarinic acetylcholine receptor (mAChR) in airway smooth muscle (Fig. 1n, q, r and Supplementary Fig. 7d-e). Both Bam8-22-and methacholineinduced respiratory effects were blocked by ipratropium bromide, a mAChR blocker ( Fig.  1q-r, Supplementary Fig. 7d-e). Administration of capsaicin to activate TRPV1 + vagal sensory neurons induced a different respiratory response characterized by a cessation in breathing that persisted for an average of 5.66 ±0.81 s, followed by a secondary phase with diminished frequency and prolonged braking after each breath (Fig. 1n). This is a prototypic response induced by the administration of respiratory sensory irritants such as cigarette smoke and cinnamaldehyde into the respiratory tract [11][12][13][14] . It is interesting that although MrgprC11 + jugular sensory neurons are expressing TRPV1, the respiratory reflexes evoked by Bam8-22 and capsaicin are quite different. These results suggest that activation of a few afferents by Bam8-22 versus a large population by capsaicin can produce nonlinear effects in lung responses.
The similarity between Bam8-22 and methacholine-induced respiratory responses led us to the hypothesis that Bam8-22 may induce bronchoconstriction, which we tested by measuring airway mechanics. Retro-orbital intravenous injection of Bam8-22 significantly increased lung resistance (R L ) in WT mice but not in Mrgpr-clusterΔ −/− mice ( Fig. 2b and Supplementary Fig. 8a-b), demonstrating that Bam8-22 induces bronchoconstriction. Stimulation of a subset of vagal sensory neurons caused release of acetylcholine from parasympathetic nerve fibers, which in turn acts on the mAChR in airway smooth muscle to induce bronchoconstriction (Fig. 2a) 15 . This parasympathetic cholinergic pathway is the major neural mechanism controlling bronchoconstriction in the lung 15 . The Bam8-22induced increase in lung resistance was completely blocked by ipratropium bromide (Fig.  2c) and vagotomy surgery (Fig. 2d), which severs the vagus nerve, demonstrating that Bam8-22 induced cholinergic bronchoconstriction. Bam8-22-induced bronchonconstriction was not affected by the deletion of TRPA1 or TRPV1 ( Supplementary Fig. 8c). Bam8-22 did not cause tracheal smooth muscle contraction (Fig. 2e), ruling out the possibility that Bam8-22 directly stimulates airway smooth muscle. Taken together, these data suggest that Bam8-22 activates MrgprC11 + jugular sensory neurons that in turn stimulates the parasympathetic system and induces cholinergic bronchoconstriction.
We also observed similar Bam8-22-induced effects in guinea pigs, another animal species that is frequently used to study respiratory reflex and to model respiratory diseases. gpMrgprx1 (XM_003462608), the predicted Mrgprc11 ortholog, is expressed in airwayinnervating jugular and nodose neurons (Supplementary Fig. 9a-b). Bam8-22 activated guinea pig vagal sensory neurons in a gpMrgprx1-dependent way ( Supplementary Fig. 9ce). Excitingly, intravenous injection of Bam8-22 in guinea pigs induces robust bronchoconstriction (Fig. 2f, Supplementary Fig. 9f). However, whether Bam8-22 also activates vagal-cholinergic reflex to induce bronchoconstriction in guinea pigs, as it does in mice, requires further investigations.
Anaphylaxis is an acute, potentially life-threatening allergic reaction mediated by mast cell activation. Anaphylactic symptoms in the respiratory system include coughing, airway swelling, and bronchoconstriction 16 . Retro-orbital injection of OVA resulted in a pronounced increase in lung resistance in OVA-sensitized WT mice (Fig. 3a). This effect was significantly inhibited in sensitized Mrgpr-clusterΔ −/− mice ( Fig. 3a and Supplementary   Fig. 8d), suggesting that Mrgprs mediate anaphylactic bronchoconstriction. Anaphylactic bronchoconstriction is evoked mainly by mast cell mediators acting on both airway smooth muscle and sensory nerves 17 . Interestingly, neuropeptide FF (NPFF), an MrgprC11 agonist that can be released by activated mouse bone marrow-derived mast cells 18 , also induced bronchoconstriction (Fig. 3b, Supplementary Fig. 8e). Additional studies are required to determine whether Mrgprs play a role in the sensitization of mast cells, their activation, or both.
Influenza virus infection causes characteristic effects within the respiratory tract, including airway inflammation and airway hyperresponsiveness. It is a major risk factor for exacerbation of asthma and causes significant morbidity and mortality annually 19 . We infected WT and Mrgpr-clusterΔ −/− mice with Influenza A virus (IAV) to examine whether sensory nerves contribute to IAV-induced respiratory symptoms. Both WT and Mrgpr-clusterΔ −/− mice displayed robust immune responses after inoculation (Fig. 3d, Supplementary Fig. 10a-b). However, we observed significantly lower IAV-induced airway hyperresponsiveness in Mrgpr-clusterΔ −/− mice compared to WT mice (Fig 3c), demonstrating that Mrgprs are required for IAV-induced airway hyperresponsiveness. The deletion of Mrgpr genes did not affect the airway hyperresponsiveness in a house dust miteinduced allergic asthma model ( Supplementary Fig. 11). These data suggest that distinct mechanisms exist for the development of airway hyperresponsiveness under different pathological conditions. We next asked whether direct stimulation of MrgprC11 + jugular sensory neurons could induce airway hyperresponsiveness in the absence of lung inflammation. We pretreated WT and Mrgpr-clusterΔ −/− mice with Bam8-22 to activate MrgprC11 + jugular sensory neurons. 5 min after Bam8-22 treatment, when the Rrs has returned to their baseline level, the airway responsiveness to MCh were measured in WT and Mrgpr-clusterΔ −/− mice. We found that pretreatment with Bam8-22 caused a remarkable increase in airway responsiveness in WT mice, but not in Mrgpr-clusterΔ −/− mice (Fig. 3e). As expected, the enhanced airway sensitivity was abolished by vagotomy in WT mice (Fig. 3f). These data demonstrated that activation of MrgprC11 + jugular sensory neurons can potentiate airway responsiveness in the absence of inflammation. Airway responsiveness to MCh is the result of both direct action on airway smooth muscle and muscle contraction-induced vagal reflex 15,20 . Our data, which is consistent with previous studies 3,4 , supports the hypothesis that the activation of vagal Cfiber by Bam8-22 may lead to the sensitization of the vagal afferent-brainstemparasympathetic reflex pathway and enhance airway responsiveness. Full length of the DNA gel images are presented in Supplementary Fig. 12.
In summary, we demonstrated that MrgprC11 + jugular sensory neurons, constituting only 2% of vagal sensory neurons, mediate cholinergic bronchoconstriction and play a role in the development of airway hyperresponsiveness in mice. Therapeutic approaches targeting this small population will avoid the possible side effects of antagonizing all vagal nociceptors, which play important roles in maintaining the physiological functions of multiple inner organs. Future studies investigating the endogenous agonists of MrgprC11 in the lung, the peripheral/central axonal projections of MrgprC11 + neurons, and the neural circuit in which these neurons reside will further reveal the neural mechanisms of respiratory diseases. The Mrgpr-clusterΔ −/− mice exhibited decreased anaphylactic bronchoconstriction and reduced airway hyperresponsiveness after influenza virus infection. Since 12 Mrgpr genes were deleted in the Mrgpr-clusterΔ −/− mice, the phenotypes observed may not be mediated solely by MrgprC11. Whether the other Mrgpr genes are involved in the tested models requires further investigation.

Animals
All experiments were performed with approval from the Johns Hopkins University Animal Use and Care Committee and the Georgia Institute of Technology Animal Use and Care Committee. The generation of Mrgpr-clusterΔ −/− mice was described previously 7 and they had been backcrossed to C57Bl/6J mice for at least ten generations. Wnt1-Cre and ROSA26lsl-LacZ mouse lines were purchased from Jackson Laboratory. Pirt-Cre line was generated in Dong's Laboratory. ROSA26-lsl-GCaMP6 line was kindly provided by Dwight Bergles at Johns Hopkins University. 3 weeks old male animals were used for Ca 2+ imaging assay. 2-3 months old males were used for all other experiments. 200-250 grams male Hartley Guinea pigs were purchased from Charles River. All the animals were housed in the vivarium with a 12-hr-light/dark cycle, and all the behavioral tests were performed from 9 a.m. to 1 p.m. in the light cycle. The housing group was 5 at maximum for mice and 2 at maximum for guinea pigs.

Retrograde tracing of sensory neurons innervating the airway
Mice were anesthetized with pentobarbital (70 mg/kg) and the ventral upper tracheal region was exposed and pierced with a microinjecting needle attached to a Hamilton syringe. Retrograde tracer CTB-488 (cholera toxin b subunit conjugated with Alexa 488, 20 μl, 1 mg/ ml)) was directly injected into the tracheal lumen. Vagal ganglia and DRG were collected 7 days after the injection to perform immunohistochemical analysis. Male Hartley guinea pigs (250 to 300 g) were anesthetized with ketamine/xylazine (50 mg/kg ketamine and 2.5 mg/kg xylazine). DiI (0.2%, 200 μl; in 10% DMSO and 90% saline, invitrogen D282) was delivered into the airway as previously described 21 . Briefly, a tracheotomy was performed and the caudal part of the trachea was exposed. DiI was injected into the lumen of the trachea with a syringe needle.

RT-PCR
Total RNA was extracted from various tissues using Trizol reagent (Invitrogen) according to the manufacturer's instructions. Mouse tissues were collected from 2-3 months old C57Bl/6 male mice. Reverse transcription was carried out using the Superscript III first-strand synthesis system (Invitrogen). PCR conditions for mouse Mrgpr genes, hMrgprX1, hTRPV1 and hGAPDH: 95 °C for 3 min, 40 cycles of 30 s at 95 °C, 30 s at 55 °C and 60 s at 72 °C.
All the PCR results were confirmed by DNA sequencing.

RT-PCR with human tissues
Human jugular and nodose ganglia were purchased from Tissue for Research (Tissue4Research.com). The results were repeated 5 times with vagal ganglia collected from 5 different donors. Characteristics and medical history of patients from whom the tissues were collected are shown in the table below. The procedure was approved by Georgia Tech Biological Materials Safeguards Committee. Human lung total RNA was purchased from OriGene.   Fig. 2). Validation data for other antibodies are available from the commercial providers.

Calcium imaging
Calcium imaging was performed as previously described 9 . Briefly, cells (HEK293 cells, vagal sensory neurons from 3-4 weeks old mice, or vagal sensory neurons from 200-250 grams guinea pigs) were cultured on glass coverslips. Cells were loaded with fura 2acetomethoxy ester (Molecular Probes) for 30 min in the dark at room temperature. After washing, cells were imaged at 340 and 380 nm excitation to detect intracellular free calcium and 20% increase in F340/F380 was set as threshold. Average fluorescence ratios (F340/ F380) were calculated from ≥ 20 cells and all the population data are presented as mean ± s.e.m.

Whole-cell current-clamp recordings of vagal sensory neurons
Bam8-22-responding vagal sensory neurons were first identified by calcium imaging before they were patched for current clamp recording. Patch pipettes had resistances of 2-4 MΩ. Action potential measurements were performed with an Axon 700B amplifier and the pCLAMP 9.2 software package (Axon Instruments). Neurons were perfused with 2uM Bam8-22 for 20s. All experiments were performed at room temperature.

Ex vivo vagal ganglia GCaMP imaging
A modification of the method described previously 22 was used for the dissection and tissue preparation. Briefly, mice were sacrificed by CO2 inhalation and exsanguination. The blood from the pulmonary circulation was mostly washed out by in situ perfusion with 20 ml of Krebs-bicarbonate solution (KBS) containing (in mM): 118 NaCl, 5.4 KCl, 1 NaH2PO4, 1.2 MgSO4,1.9 CaCl2, 25 NaHCO3, 11.1 dextrose. The trachea and the lung with intact vagus nerves and vagal ganglia were dissected and placed in a two-compartment tissue chamber (Supplementary Figure 6a). The lung was pinned in one compartment (airway chamber) to receive chemical stimuli. The left vagal ganglia (VG chamber) were pinned in a second compartment for multiphoton imaging. Both compartments were perfused with 37 °C KBS. There is no fluid exchange between the two compartments. The trachea was cannulated with PE tubing to allow the delivery of chemicals onto the airway. The lungs were punctured with a 27-gauge needle (1-2 mm deep, 2-8 per lobe) to allow perfusing KBS to exit the tissue. The surface of the rostral vagal ganglia was imaged by a multiphoton microscope (Scientifica) with a 20× water immersion lens.

Head-out body-plethysmography
Head-out body-plethysmography was performed with anesthetized and spontaneously breathing mice. Mice were anesthetized with urethane and tracheostomy was performed to allow the aspiration of chemicals (2 μl, 1 mM capsaicine, 1 mM MCh, 5 mM Bam8-22) directly into the trachea through the cannula. Delivery of the chemicals directly into the trachea allows us to avoid interference from the upper respiratory tract, which is mainly innervated by trigeminal sensory neurons. The body of the animal was kept in a chamber and the respiratory effort was monitored digitally using a Biopac data acquisition system. Respiratory rate and the amplitude of the respiratory waveform were analyzed offline with AcqKnowledge software.

Airway mechanics
Anesthetized (pentobarbital, 70 mg/kg) animals were tracheotomized, attached to a Flexivent pulmonary mechanics analyzer (SCIREQ), and paralyzed with succinylcholine (2 mg/kg). Anesthetized mice were ventilated at a tidal volume of 9 mL/kg, at a frequency of 150 bpm. Anesthetized guinea pigs were ventilated at a tidal volume of 8 mL/kg, at a frequency of 90 bpm. Positive end-expiratory pressure was set at 3 cm H 2 O. We assessed the airway responsiveness using a single-compartment model to measure the total lung resistance (R L ), a dynamic variable that is largely dependent on the resistance to air flow through the airways and is sensitive to airway smooth muscle contraction. Bam8-22 (50 μl, 10 mg/ml for mice, 400 μl, 20 mg/ml for guinea pigs), NPFF (50 μl, 10 mg/ml), and MCh (50 μl, 30 μg/ml) were administered by retro-orbital IV injection. Airway hyperresponsiveness was determined in response to aerosolized methacholine challenges (0, 1, 3, 10, and 30 mg/ml). The nebulizer was on for 10 seconds to deliver each dose of MCh.

Organ bath studies
Organ bath studies were performed as previously described 23 . Briefly, whole tracheas were dissected out from sacrificed mice and placed in oxygenated Krebs-bicarbonate solution containing (in mM): 118 NaCl, 5.4 KCl, 1 NaH2PO4, 1.2 MgSO4,1.9 CaCl2, 25 NaHCO3, 11.1 dextrose. Tracheas were cleaned of connective tissue and tracheal rings (whole or laterally divided in half), were suspended between two tungsten stirrups in 10 ml organ chambers filled with Krebs solution that was warmed to 37 °C and bubbled with 95% O2-5% CO2 to maintain a pH of 7.4. One stirrup was connected to a strain gauge (model FT03; Grass Instruments, Quincy, MA, USA), and tension was recorded on a Grass Model 7 polygraph (Grass Instruments). Preparations were stretched to a resting tension of 0.2 g, and washed with fresh Krebs buffer at 15 min intervals during a 60-min equilibration period. After equilibration, tracheas were challenged with either Bam8-22 (10 μM) or electrical field stimulation (12V, 8HZ, 0.5ms, 10s/100s). At the end of each experiment, all tracheas were maximally contracted with carbamylcholine (1mM). The results are expressed as a percentage of maximum contraction.

IAV infection-induced lung inflammation
Mouse adapted influenza A/California/4/2009 H1N1 was a kind gift from Dr. Sabra Klein in the Department of Molecular Microbiology and Immunology at Johns Hopkins University School of Public Health. 8-12-week old mice were anesthetized with ketamine (100 mg/kg) and xylazine (15 mg/kg) and were infected intranasally with 0.4 50% mouse lethal dose (MLD50) of IAV in 30 μl of DMEM. Mice in the control group were infected with 30ul DMEM medium only. At 5 days post infection, the airway mechanics were examined using the Flexivent system and the bronchoalveolar lavage fluid (BALF) was collected to evaluate lung inflammation. BALF was centrifuged and the cell pellet was resuspended in PBS to perform differential cell counting of leukocytes (macrophages/monocytes, neutrophils, eosinophils, and lymphocytes). The supernatant of BALF was used to examine the levels of TNF-a, IFN-g, and IL-6 using enzyme immunoassay kits purchased from R&D Systems (Minneapolis, MN) according to individual kit instructions. Lungs were inflated and fixed with formalin at 30 cm H2O and lung sections were processed and stained with hematoxylin and eosin.

Mouse model of anaphylaxis
Mice were sensitized with 20 μg of OVA emulsified with 2 mg of Imject Alum Adjuvant in 150 μl PBS by i.p. injection. 21 days after OVA sensitization, mice were anesthetized and challenged with 500ug of OVA in 50 μl of PBS and the airway mechanics were measured using the Flexivent system. The airway resistance increased gradually after the OVA injection, reaching the peak at 50-60 seconds after the injection. To examine the level of OVA-specific IgE, serum of sensitized mice was collected on day 18 (before challenge). The OVA-specific IgE level was determined using the anti-Ovalbumin IgE (mouse) ELISA kit (CaymanChemical, 500840) following the user's instructions.

House dust mite-induced airway allergic inflammation
Mice were sensitized by i.p. injection of 100 μg house dust mite extract in 150 μl PBS on days 1 and 14. Lightly anesthetized mice (isoflurane) were challenged by intranasal instillation of 100 μg house dust mite extract in 20 μl PBS on days 18 and 21. Airway mechanics measurements were performed on day 23.

Data analysis and statistics
Statistical analyses were performed using Prism 7 (GraphPad). We did not use statistical methods to predetermine the sample size. Our sample sizes are comparable to those reported in previous publications. Differences between experimental groups were analyzed with twotailed unpaired Student's t test. Threshold for significance (α) was set at 0.05. Data are reported as mean ± s.e.m.. Normality and equality of variance were analyzed with the Shapiro-Wilk normality test and F-test, respectively. When the sample numbers were too small to calculate, distribution was assumed to be normal. The experimenters were blinded to the genotype of the animals. Animals were assigned randomly to the various experimental groups, and data were collected and processed randomly. N numbers were indicated for all the experiments that required statistical analysis. RT-PCR and immunostaining were performed at least three times with three different animals. Retrograde tracings were performed three times using three animals each time. A Life Sciences Reporting Summary is available online.

Supplementary Material
Refer to Web version on PubMed Central for supplementary material.