Research Article

Vagal innervation is required for pulmonary function phenotype in Htr4−/− mice

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Human genome-wide association studies have identified over 50 loci associated with pulmonary function and related phenotypes, yet follow-up studies to determine causal genes or variants are rare. Single nucleotide polymorphisms in serotonin receptor 4 (HTR4) are associated with human pulmonary function in genome-wide association studies and follow-up animal work has demonstrated that Htr4 is causally associated with pulmonary function in mice, although the precise mechanisms were not identified. We sought to elucidate the role of neural innervation and pulmonary architecture in the lung phenotype of Htr4−/− animals. We report here that the Htr4−/− phenotype in mouse is dependent on vagal innervation to the lung. Both ex vivo tracheal ring reactivity and in vivo flexiVent pulmonary functional analyses demonstrate that vagotomy abrogates the Htr4−/− airway hyperresponsiveness phenotype. Hyperpolarized 3He gas magnetic resonance imaging and stereological assessment of wild-type and Htr4−/− mice reveal no observable differences in lung volume, inflation characteristics, or pulmonary microarchitecture. Finally, control of breathing experiments reveal substantive differences in baseline breathing characteristics between mice with/without functional HTR4 in breathing frequency, relaxation time, flow rate, minute volume, time of inspiration and expiration and breathing pauses. These results suggest that HTR4’s role in pulmonary function likely relates to neural innervation and control of breathing.

pulmonary function is a readily and reliably measurable index of the physiological state of the lungs that is used clinically to diagnose and follow the progression of chronic obstructive lung disease (COPD), asthma, and other lung conditions. Pulmonary function is also an independent predictor of mortality in the general population (1, 48). Heritable contributions to pulmonary function may be as high as 54% in humans (24, 50). Genome-wide association studies (GWAS) provide an unbiased, comprehensive approach to identifying genetic variants associated with disease states. GWAS have identified nearly 50 loci associated with pulmonary function in humans (16, 35, 42, 47, 49). While GWAS are extremely successful in identifying genetic loci associated with disease, identifying the causal genes or variants generally requires other techniques including more costly experimental animal studies.

In GWAS meta-analysis, two groups simultaneously identified and cross-replicated single nucleotide polymorphisms (SNPs) within the serotonin (5-hydroxytryptamine) receptor 4 (HTR4) gene related to pulmonary function in humans (16, 35). Additional studies further implicated HTR4 in the related clinical phenotypes of airway obstruction and COPD (43, 51). Recently, we demonstrated that HTR4 is causally related to pulmonary function using a murine Htr4-knockout model (Htr4−/−). Htr4−/− animals displayed increased whole lung (R) and airway resistance (Rn) at baseline as well as increased airway hyperresponsiveness (AHR) to inhaled methacholine (20). However, precisely how HTR4 modulates pulmonary function remains to be elucidated.

Serotonin receptors (HTRs) encompass a class of seven different receptor classes that bind the neurotransmitter serotonin. All but one of the genes within this family are G protein-coupled receptors found within a wide range of tissues throughout the body that activate excitatory or inhibitory secondary signaling cascades (30). In human and murine brain, HTR4 localizes to the olfactory tubercule, basal ganglia, hippocampus, and pre-Bötzinger complex, which generates respiratory rhythm (7, 36). HTR4 is peripherally expressed in most of the gastrointestinal tract, vasculature, adrenal glands, urinary tract, and heart (3, 18). Furthermore, HTR4 is also expressed, albeit at lower levels, in lung, airway epithelial cells, and smooth muscle cells of adults (2, 11) and temporally in the developing human fetal lung (19).

Breathing is tightly regulated by the nervous system, thus ensuring appropriate tissue oxygenation by sensing changes in blood pH and gas composition and other environmental inputs (15). The trachea and bronchi are richly innervated by autonomic and sensory nerve fibers (23). The majority of efferent nerve fibers reach these loci via the vagus nerve. Inspiration activates vagal afferent nerves sending impulses to the brain stem where parasympathetic reflex bronchial smooth muscle contraction is initiated (21). Interrupted parasympathetic nerve activity (e.g., by surgical or chemical vagotomy or with muscarinic receptor antagonists) induces dilation of the airways in mice (5, 6, 10), and vagal innervation is required for antigen-induced airway hyperresponsiveness in multiple rodent models (28, 37, 39, 45). Thus vagotomy prevents feedback to the central nervous system and is a useful tool to elucidate the role of this neural input on pulmonary function phenotypes.

Building on our prior work demonstrating altered pulmonary function in mice following knockout of the Htr4 gene, we sought to gain mechanistic insight into how HTR4 functions to regulate pulmonary function. We employed stereology and hyperpolarized (HP) 3He magnetic resonance (MR) imaging (MRI) to evaluate the role of HTR4 on pulmonary architecture and development. We also used flexiVent lung procedures coupled with additional techniques including plethysmography and isolated tracheal ring studies to investigate the role of neural innervation on our previously observed Htr4−/− pulmonary phenotype.



Htr4−/− (B6.129P2-Htr4tm1Dgen/J) and wild-type (WT; C57BL/6J) mice were purchased from The Jackson Laboratory (Bar Harbor, ME) and maintained as Het × Het breeders to generate sibling WT and Htr4−/− mice that were genotyped per the Jackson Laboratory protocol. Male mice were between 18 and 22 wk of age for flexiVent studies. Male and female mice aged 9–25 wk were used for whole body plethysmography experiments. Male mice between 14 and 21 wk of age were used for tracheal ring experiments. All animal work described in this study was conducted according to NIH guidelines and were approved by the National Institute of Environmental Health Sciences (NIEHS) animal care and use committee.

Lung function assessment.

For HTR2 antagonist experiments, 12 mg/kg ketanserin (S006, Sigma, St. Louis, MO) in 0.9% saline (S8776 Sigma [certificate 7647-14-5 (< 0.005 U/ml endotoxin)]) was given intraperitoneally 15 min before lung function analysis with flexiVent as previously described (20). In brief, urethane-anesthetized mice (1.5 g/kg ip) were tracheotomized with a 19-gauge stainless steel cannula and injected with pancuronium bromide (0.8 mg/kg ip) to paralyze the diaphragm, preventing autonomous breathing. Mice were then connected to a computer-controlled ventilation device that also performed lung function “snapshot” and “quickprime” perturbations at baseline and following methacholine administration.

Surgical vagotomy was completed on a separate cohort of anesthetized animals as previously described (9). Briefly, mice were prepared for vagotomy as described above with the addition of the following step. A piece of surgical thread was passed underneath both the right and left vagus nerves that run parallel to the trachea before pancuronium bromide injection and subsequent mechanical ventilation. After cessation of autonomous breathing, both vagus nerves were severed with scissors passed under the thread and the standard methacholine dose response was conducted. Sham mice underwent isolation of vagus nerves with surgical thread without severance.

RNA and quantitative PCR.

The whole caudal lobe from each mouse was homogenized and RNA was isolated and purified following manufacturer’s instructions (RNeasy Kit 74104 Qiagen, Hilden, Germany). Following purification of RNA, cDNA was created with equal amounts of template RNA and subjected to real-time RT-PCR analysis using TaqMan-based assay [Applied Biosystems/Life Technologies, Carlsbad, CA: Htr1a (Mm00434106_s1), Htr1b (Mm00439377_s1), Htr2a (Mm00555764_m1), Htr2b (Mm00434123_m1), Gapdh (Mm99999915_g1)]. Relative expression was calculated with the ΔΔCt method using Gapdh as the housekeeping gene and WT expression levels set to 1.


The lungs were chemically fixed with Karnovsky fixative via tracheal inflation at 25 cm of pressure for 1 h, tied off and stored in fixative at 4 degrees. Lung volume was estimated by volume displacement (38) and the lungs were sampled for light microscopic analysis by systematic uniform random sampling, including the generation isotropic uniform random sections with the orientator (27). For light microscopic evaluations, the lungs were embedded in paraffin, cut in 5-µm sections, and stained with hematoxylin and eosin (H&E). Small pieces of lung tissue with known dimensions were further coembedded in paraffin and the tissue shrinkage in these pieces was estimated after embedding and sectioning. The average shrinkage in paraffin embedding was calculated as 78.84% and all stereological surface area and number quantifications were corrected by this shrinkage factor. The histological sections of the lungs were scanned and digitalized with an Olympus Slide Scanner (VS110, Olympus, Chelmsford, MA) at ×20 magnification and the stereological evaluations were subsequently performed with the newCast software system (Visiopharm, Hoersholm, Denmark). All measurements and calculations were performed as previously described in detail (4), including the assessment of parenchymal, nonparenchymal, alveolar, and septal volume; alveolar surface area; and septal thickness as well as surface area and epithelial thickness in the conducting airways.

Hyperpolarized 3He magnetic resonance imaging.

Ventilation distribution (29) of WT and Htr4−/− mice was assessed by HP 3He MRI. Protocols were approved by the Duke University Institutional Animal Care and Use Committee. Mice were anesthetized with a 75 mg/kg intraperitoneal dose of pentobarbital, and ventilated through a tracheostomy tube with 75% N2-25% O2 (31). For ventilation MRI, 3He gas (Spectra Gases, Alpha, NJ) polarized to ~30% (IGI.9600.He, MITI, Durham, NC) was substituted for N2. During MRI, electrocardiogram, airway pressure, and rectal temperature were continuously monitored.

HP 3He MRI was done on a 2-Tesla magnet (Oxford Instruments, Oxford, UK) controlled by a GE EXCITE 12.0 console. Mice were placed into a cradle that fit into a 65.8 MHz birdcage 3He coil and aligned in the center of the magnet bore. To minimize motion artifacts and gradient-induced signal attenuation, all imaging used respiratory-gated radial acquisitions acquired with the following parameters: slice-selective three-lobe sinc pulse, TR/TE = 8/0.88 ms, BW = 31.25 kHz, FOV = 2.5 × 2.5 × 2.5 cm3, matrix = 1283, and a total of 10,001 radial views acquired at 18 views per breath with a variable flip angle of 9–90° for constant magnetization.

3D 3He ventilation MR images were segmented by a linear-binning method (17) to identify four intensity clusters, of which the volume of the lowest intensity cluster, representing unventilated regions of the thoracic cavity, was used to calculate the ventilation defect percentage (VDP). Additionally, ventilation heterogeneity was characterized using the coefficient of variation (CV) of the intensity histogram.

Tracheal ring assay.

Mouse ex vivo tracheal contractile responses to smooth muscle agonists were performed as previously described (5, 40). In brief, male mice aged 15–21 wk were euthanized via exsanguination (vena cava puncture) following anesthetization by sodium pentobarbital. The trachea was isolated between larynx and the first bronchial bifurcation, and immediately placed in Krebs-Henseleit solution on ice, where connective tissue and fat were removed under dissection microscope. For each mouse, a 3-mm section of trachea was cut and hung on two L-shaped platinum hooks in a glass organ bath (5 ml volume; Radnoti Glass Technology, Monrovia, CA) filled with Krebs-Henseleit solution (37°C) continuously bubbled with 95% O2 and 5% CO2. The lower hook was anchored to the bottom of the glass water bath, and the upper hook was attached to a force transducer connected to a PowerLab data acquisition and analysis system (ADInstruments, Colorado Springs, CO). Tracheal ring segments were equilibrated for 45–60 min, with bath solution changes every 15 min. During equilibration, the resting tension was adjusted to 0.7–0.8 g. Following equilibration, 67 mM potassium chloride (KCl) was administered to induce contraction, followed by three washes and one additional KCl challenge. Rings that failed the KCl challenge were discarded from analysis. After KCl challenge, a cumulative administration of carbachol (C4382, Sigma) was given to obtain a series of concentrations from 10−8 to 3×10−5 M. Each concentration was given until a maximum response was achieved. The recorded contraction force was analyzed using LabChart software (ADInstruments).

Control of breathing.

WT and Htr4−/− mice were assessed for baseline breathing parameters and control of breathing perturbations using noninvasive whole-body plethysmography (Buxco, Wilmington, NC). Mice were acclimated to breathing chambers for 60 min/day for 3 days without measurements. For each experimental day, mice were placed in breathing chamber for 15 min before recording 30 min of baseline parameters, given 5 min of gas-treatment perturbation [hypoxic (10% O2), hypercapnic (8% CO2), or both (10% O2, 8% CO2)], followed by breathing room air for at least 30 min. Mice were treated with one perturbation per day over the course of 3 days.

Statistical analysis.

For assessment of genotype differences in methacholine induced airway hyperresponsiveness and in the tracheal ring assay, Proc GLM (SAS Version 9.3) was used to calculate dose-adjusted least squares means and standard errors of the means (SE) by genotype. Values were reported as significant if P value for genotype was < 0.05 after accounting for the dose effect. Genotype differences for quantitative PCR were assessed with two-tailed Student’s t-test. For control of breathing parameter analysis, recorded parameter values were averaged by minute and Proc GLM (SAS 9.3) was used with repeated measures to assess differences across various time windows adjusting for age, experiment day, and chamber. To avoid chamber bias, mice were randomized by genotype and sex to chambers 1–8. Plotted traces are least squares means and associated standard errors. For gas perturbation assessment (5 min) and recovery (30 min), the percent deviation of each time point from the average of the last 5 min before gas administration was used.


Airway hyperresponsiveness in Htr4−/− mice is not related to Htr2a/b.

Of the seven serotonin receptors, all but one act as G protein-coupled receptors and share similar motifs. In addition to HTR4, HTR1B and HTR2A/B are associated with cardiopulmonary function humans and mice (12, 22). To examine whether Htr4−/− mice have compensatory upregulation of other serotonin receptor family members contributing to the Htr4−/− phenotype, we assessed, via real-time qRT-PCR on whole lung tissue, the main isoform transcript levels of lung Htr1 and Htr2 in naive WT and Htr4−/− mice. While expression of Htr1 was not significantly altered in Htr4−/− mice, these mice exhibited increased Htr2a expression (Fig. 1A). Following HTR2 blockade by ketanserin, a selective HTR2 antagonist, differences in pulmonary function between WT and Htr4−/− mice persisted. Indeed, Htr4−/− mice had increased lung resistance (R), tissue resistance (G), elastance (E), tissue elastance (H) and decreased compliance (C) compared with WT mice (Fig. 1B). Thus, while Htr2a is upregulated in Htr4−/− mice, this upregulation is not responsible for the observed pulmonary function differences.

Fig. 1.

Fig. 1.Serotonin receptor family dysregulation in Htr4−/− mice does not contribute to pulmonary phenotype. A: relative mRNA expression of serotonin receptor family members Htr1a, Htr1b, Htr2a, and Htr2b in wild-type (WT) and Htr4−/− animals (n ≥ 4/genotype; *P < 0.05). B: assessment of airway hyperresponsiveness to methacholine (MCh) after treatment with ketanserin. Whole lung resistance (R), Newtonian resistance (Rn), tissue resistance (G), elastance (E), and tissue elastance (H) were higher and compliance (C) lower in Htr4−/− animals relative to WT. Least-squares means and SE are plotted after adjusting for age and dose (n ≥ 16/genotype; *P < 0.05).

Htr4−/− mice have no discernible differences in airway architecture.

Previously, we reported no discernible gross histopathological differences in lung histology between Htr4−/− and WT mice (20). To examine whether more subtle anatomic alterations exist in the airway structures of Htr4−/− mice that may explain the observed physiological abnormalities, we conducted extensive stereology assessment on lungs from Htr4−/− and WT mice. The total volume and the proportion of nonparenchymal volume were the same in Htr4−/− and WT mice (Fig. 2A). In the distal lung, there were no genotypic differences in alveolar volume and surface area (Fig. 2, B and D), or in septal volume and thickness (Fig. 2, B and C). Likewise, epithelial thickness and the surface area of conducting airways were not significantly different between genotypes (Fig. 2, E and F). These data suggest that the functional changes in the Htr4−/− phenotype are not due to alterations in lung architecture.

Fig. 2.

Fig. 2.Htr4 does not impact the development of pulmonary architecture. Stereological assessment of lung volume (A), distal lung volume (B), septal thickness (C), alveolar surface area (D), epithelial thickness (E), and conducting airway surface area (F) in Htr4−/− and WT mice. Data are presented as means ± SE (n = 8/genotype).

Ventilation distribution is unaltered in Htr4−/− mice.

To examine whether Htr4 deficiency alters ventilation and gas distribution, we utilized hyperpolarized 3He MRI to visualize lung inflation. Htr4−/− mice exhibited no differences in the VDP or ventilation heterogeneity (CV) as compared with WT (Fig. 3). This suggests that HTR4’s effects on pulmonary function are not related to defects in either lung inflation or gas distribution.

Fig. 3.

Fig. 3.Pulmonary inflation is not impacted in Htr4−/− animals. Hyperpolarized 3He MR ventilation images and corresponding linear-binning maps in representative WT and Htr4−/− mice. The ventilation defect percentage (VDP) and ventilation heterogeneity (CV) were unchanged in Htr4−/− mice compared with WT. Data are presented as means ± SE (n ≥ 4/genotype).

Genotype differences are abolished in ex vivo tracheal ring assay.

Knowing that Htr4 is expressed in portions of the brain responsible breathing rhythmogenesis, we hypothesized that the mechanism underlying Htr4−/− changes in pulmonary function may be related to neural innervation. Removal of the trachea from the animal eliminates its neural innervation, thus allowing for assessment of basal airway response apart from the response regulated by the autonomic nervous system. We did not observe genotype differences in carbachol-induced smooth muscle contraction of tracheal rings isolated from Htr4−/− and WT mice (Fig. 4). This suggests that neural innervation may play a role in the observed pulmonary phenotypes in Htr4−/− mice.

Fig. 4.

Fig. 4.No genotype differences in ex vivo tracheal ring contraction. Tracheal rings of WT and Htr4−/− mice were given increasing concentrations of the bronchoconstrictor carbachol (Cch). Means and standard errors are plotted after normalization to KCl; n ≥ 12/genotype; P = 0.35 for genotype after accounting for dose.

Genotype differences are abolished by in vivo vagotomy.

Ex vivo tracheal ring contraction experiments remove other in vivo inputs of nutrients and localized neurotransmitter signaling from assayed tracheal rings. Thus, to validate the tracheal assay findings in vivo, we performed vagotomy followed by flexiVent assessment of pulmonary function. With the exception of a reduction in tissue elastance (H) in Htr4−/− mice, baseline pulmonary function parameters were not changed by vagotomy (Fig. 5). As expected, a bilateral vagotomy blunted methacholine-induced AHR (horizontal black bars represent WT responses for mice with sham vagotomy; Fig. 6). Furthermore, following vagotomy, the genotype differences in baseline pulmonary function parameters that we previously reported (whole lung resistance, tissue resistance, compliance, elastance, and tissue elastance) were abolished (Fig. 6). These data, in concert with the ex vivo tracheal ring results, indicate that neural innervation is a critical component of HTR4’s regulation of pulmonary function.

Fig. 5.

Fig. 5.The effects of vagotomy on baseline pulmonary function parameters were minimal. Baseline lung function parameters [whole lung resistance (R), Newtonian resistance (Rn), tissue resistance (G), compliance (C), elastance (E), tissue elastance (H)] were examined on flexiVent in WT and Htr4−/− mice. Least squares means and standard errors of Sham vs. Vagotomy adjusting for age. *P < 0.05; Sham (n = 4 WT, n = 5 Htr4−/−); Vagotomy (n = 9 WT, n = 10 Htr4−/−).

Fig. 6.

Fig. 6.Vagotomy ablates the pulmonary phenotype of Htr4−/− animals. Mice were vagotomized and then pulmonary function was assessed via flexiVent before and after methacholine (MCh). Whole lung resistance (R), Newtonian resistance (Rn), tissue resistance (G), compliance (C), elastance (E), and tissue elastance (H) were not significantly different between Htr4−/− and WT mice. Horizontal black bars indicate response at 50 mg/ml of methacholine with sham vagotomy surgery for WT mice (n = 4). Least-squares means and SE are plotted after adjusting for age (n ≥ 9/genotype).

Htr4/ mice control of breathing dysregulation.

Vagal afferents are responsible for communicating many factors associated with pulmonary function and control of inspiration/expiration to the brain, more specifically the pre-Bötzinger complex. Noninvasive whole body plethysmography was used to monitor breathing parameters of WT and Htr4−/− mice at baseline; during exposure to hypoxic, hypercapnic, and hypoxic/hypercapnic air; and during recovery to these perturbations. Baseline breathing parameters were altered in Htr4−/− mice (Table 1). Htr4−/− mice had decreased breathing frequency (f), flow rates (EF50), minute volume (MV) and peak expiratory flow (PEF) coupled with increased end expiratory pause (EEP), peak inspiratory flow (PIF), relaxation time to expire constant volume (RT), and time of expiration (Te) and inspiration (Ti). The genotype effects were similar in magnitude between males and females and always in the same direction, but most statistical differences by genotype were observed in males due to slightly higher variances in females. Baseline tracings for plethysmography parameters f, MV, Te, and PEF are shown in Fig. 7 and further illustrate the genotype differences summarized in Table 1. There were no substantive genotype differences in response to any of the three gas conditions or in the recovery period following exposure (data not shown). Ultimately, these data further support neural system involvement in the Htr4−/− pulmonary phenotype.

Table 1. Control of breathing parameters in WT and Htr4/ mice

SexParameter (units)30 min mean (WT)30 min mean (Htr4−/−)Htr4−/− vs. WTP Value
FemaleEEP (ms)47.32952.0484.71890.001
MaleEEP (ms)60.43366.6976.26400.153
FemaleEF50 (ml/s)0.28250.2724−0.01010.072
MaleEF50 (ml/s)0.17970.1721−0.00760.024
FemaleEIP (ms)4.78794.80510.01730.975
MaleEIP (ms)4.99815.03520.03710.055
FemaleFrequency (bpm)401.26393.90−7.36310.287
MaleFrequency (bpm)309.65302.73−6.91470.013
FemaleMV (ml/m)88.40888.045−0.36250.080
MaleMV (ml/m)56.98055.885−1.09500.021
FemalePEF (ml/s)4.69494.6495−0.04540.118
MalePEF (ml/s)3.18913.1016−0.08760.024
FemalePIF (ml/s)6.76066.90040.13980.257
MalePIF (ml/s)4.54264.55980.01720.033
FemaleRpef (ml/s)0.27240.2657−0.00680.198
MaleRpef (ml/s)0.26270.2538−0.00880.315
FemaleRT (s)0.07310.07630.00320.052
MaleRT (s)0.09150.09580.00430.012
FemaleTe (s)0.13000.13530.00530.135
MaleTe (s)0.16540.17500.00970.026
FemaleTi (s)0.06270.06290.00020.926
MaleTi (s)0.08180.08290.00110.026
FemaleTV (ml)0.20610.21020.00410.172
MaleTV (ml)0.17350.17410.00060.354

The average minute differences in least squares means from repeated-measures regression across 30 min of baseline breathing between genotypes (Htr4−/− vs. WT) are reported for the following parameters: end-expiratory pause (EEP), expiratory flow at 50% exhale (EF50), end-inspiratory pause (EIP), breathing frequency, minute volume (MV), peak expiratory flow (PEF), peak inspiratory flow (PIF), ratio of start of expiration to PEF over time of expiration (Rpef), relaxation time needed to expire constant volume (RT), time of expiration (Te), time of inspiration (Ti), and tidal volume (TV). Regression analysis was adjusted for age, genotype, and age*genotype and run separately by sex (n ≥ 15 per sex per genotype).

Fig. 7.

Fig. 7.Baseline control of breathing parameters are altered in Htr4−/− mice. Least squares means from repeated measures regression adjusted for age, genotype, and age*genotype are displayed for breathing frequency, (A) minute volume (B), expiration time (C), and peak expiratory flow (D) are plotted [n ≥ 12/genotype (males)].


GWAS have identified variants in HTR4 as potential contributors to genetic variability in pulmonary function (16, 25, 41). Our group followed up these findings in murine models of pulmonary function and demonstrated a distinct pulmonary phenotype associated with genetic disruption of Htr4: increased basal airway resistance and increased airway hyperresponsiveness (20). In the present study, we sought to provide mechanistic insights into how HTR4 affects pulmonary function. The findings presented here suggest that HTR4 is required for proper neuronal signaling between the brain and lungs, and that HTR4 participates in inspiration/expiration rhythmogenesis and control of breathing.

Most previous research on Htr4, a serotonin receptor gene, has focused on the nervous system. Seven serotonin receptor gene family members with multiple isoforms per member are expressed within the nervous system and in select organs; many have similar functions and conserved motifs in humans (30). In the present study we investigated the possibility that other serotonin receptors may be coordinately regulated in Htr4−/− mice, perhaps as an adaptive or compensatory change. We focused on Htr1 and Htr2 due to their previously identified roles in modulating cardiopulmonary function. HTR1B has been implicated in pulmonary arterial contraction in humans (22), and HTR2A has been shown to mediate vasoconstriction in rats (26). Furthermore, ketanserin, a selective 5HT2A-receptor antagonist (to a lesser extent 2B), has been used to treat pulmonary arterial hypertension in the elderly (12). In Htr4−/− mice, we did observe an increase in Htr2a expression, but blocking Htr2a signaling did not abrogate the previously identified Htr4 phenotype, suggesting that while other serotonin receptors may be dysregulated in Htr4−/− mice, this dysregulation does not contribute to the Htr4−/− pulmonary phenotype. Treatment with ketanserin did not affect baseline pulmonary function parameters (data not shown). However, it is worth noting that animals treated with ketanserin displayed attenuated responses to methacholine-induced bronchoconstriction compared with our previous paper (20). Although it is beyond the scope of this article, these data suggest that Htr2a/b may also be involved in methacholine-induced AHR in mice as has been reported in guinea pigs (39).

We also examined pulmonary architecture in Htr4−/− and WT animals. Previously, our group reported that lungs from Htr4−/− and WT animals did not differ in either size or histopathological characteristics. Here, we sought to discern whether differences existed by genotype in micropulmonary architecture that were not observable with our prior methods. Despite assessing multiple stereology parameters including parenchymal and nonparenchymal contributions to lung volume, alveolar and septal volume, and distal lung vs. conducting airway metrics, our data indicated that disruption of Htr4 does not result in alterations of pulmonary architecture. This is not surprising, as modest human variations in pulmonary function within the normal range are not associated with discernible structural differences in the lung or airways (33).

Even without airway architecture differences, physiological differences in inflation characteristics could result in changes in pulmonary function. We used state-of-the-art technology to assess pulmonary inflation characteristics by combining MRI with a nonradioactive hyperpolarized helium isotope that provides quantitative information regarding the dynamic phenomenon of gas replenishment of the lung. This technique is effective in identifying changes in baseline inflation (44). Our observed lack of differences in inflation characteristics between Htr4−/− and WT lungs suggests that disruption of Htr4 has no effect on pulmonary inflation and gas distribution in adult mice.

Neural input into pulmonary function extends into the conducting airways and controls airway smooth muscle tone. Htr4 is expressed in the vagal nerve afferents specific to the lung (32), suggesting that the disruption of Htr4 may impact both breathing regulation and airway smooth muscle tone. Our work presented here suggests that HTR4 plays a role in maintaining proper airway smooth muscle tone via neuronal innervation. Tracheal rings (which lack neural innervation) from mice with/without functional HTR4 reacted similarly to increasing concentrations of carbachol. Likewise, flexiVent assessment of pulmonary function following the severing of the vagus nerve abolished the Htr4−/− phenotype. Combined, these two experiments provide convincing evidence that HTR4 functions through neural innervation to regulate pulmonary function. If the airway afferents are important in the effect of Htr4 disruption on pulmonary function, in theory there should be no effect of the knockout on control of breathing parameters following vagotomy. Unfortunately, because vagotomy causes mice to slowly cease the spontaneous breathing required to assess these parameters, such data cannot be collected. Vagal innervation is necessary for a complete response to antigen-induced AHR in guinea pigs (39), rats (37), mice (28, 45). With the exception of slightly reduced tissue elastance (H) in Htr4−/− mice (Fig. 5), we only saw the effect of vagotomy in methacholine-induced AHR (Fig. 6). This is consistent with previously referenced work and may highlight differences in rodent and human physiology.

In this study, we identified differences between genotypes in multiple control of breathing parameters, suggesting HTR4’s involvement in control of breathing. The generation of respiratory rhythm, while not completely understood, is considered to originate in neurons of the pre-Bötzinger complex of the ventrolateral medulla; disruption of synaptic transmission in this region ablates rhythmogenesis (34). These neurons integrate information received from other areas of the brain to regulate inspiratory and expiratory rhythm, maintain eupnea, and adapt breathing to posture and activity level (14). Furthermore, the prevalence of Htr4 expression in the pre-Bötzinger complex (7) and the known modulatory effect of serotonin as excitatory for respiration in whole-cell patch recordings (13) are consistent with our observation of decreased breathing frequency in Htr4−/− mice.

Throughout this study we highlight the role of vagal innervation in the maintenance of pulmonary function, but, as mentioned above, neural regulation of pulmonary function occurs within the central nervous system and extends into the conducting airways, making differentiating between central and peripheral effects difficult. Because HTR4 is highly expressed within the central nervous system and also mediates a tonic positive influence on the firing activity of serotoninergic neurons (8), it is possible our observed effects are a combination of both afferent and efferent signaling. Furthermore, many of the responsibilities of the vagus nerve are shared throughout the nervous system such as chemosensors for gas and pH sensing in the periphery and central nervous system, and thus our results may be diluted by the compensation of these other mechanisms. Pulmonary diseases are accompanied by changes in the entire neural pathway including enhanced activity of primary afferent nerves, increases in synaptic efficacy at secondary nerves in the central nervous system, and changes in the autonomic nerve pathways (46). Thus, while it is difficult to distinguish between the tightly entwined effects of HTR4 within the central and peripheral nervous systems, this does not diminish the importance of our results outlining the role of HTR4 in both control of breathing and neural regulation of airway tone.

In summary, we report here that HTR4 plays a role in maintaining proper airway tone via vagal innervation and affects rhythmogenesis of inspiration/expiration. Because lack of functional HTR4 causes increased airway resistance (20), our data suggest that HTR4 functions to regulate smooth muscle tone via vagal innervation. These data further validate human findings from GWAS and provide mechanistic insight on how the HTR4 locus affects pulmonary function.


This research was supported by the Intramural Research Program of the NIH, NIEHS (Grants Z01 025043, Z01 ES025045, and Z01 ES043012). Work at the Radiology Center at Duke University was supported in part by grant P41 EB015897.


No conflicts of interest, financial or otherwise, are declared by the author(s).


J.H., B.D., D.C.Z., and S.L. conceived and designed research; J.H., H.L., C.B., R.V., and L.M. performed experiments; J.H., H.L., C.B., and R.V. analyzed data; J.H., C.E.N., C.B., B.D., D.C.Z., and S.L. interpreted results of experiments; J.H. and C.E.N. prepared figures; J.H. and C.E.N. drafted manuscript; J.H., C.E.N., H.L., C.B., R.V., L.M., B.D., D.C.Z., and S.L. edited and revised manuscript; J.H., C.E.N., H.L., C.B., R.V., L.M., B.D., D.C.Z., and S.L. approved final version of manuscript.


We thank Grace Kissling at NIEHS for statistical expertise, Julie Foley at NIEHS for assistance, as well as Dallas Hyde, Frank Ventimiglia, and Leialoha Putney at University of California-Davis for their expertise and assistance with stereology.


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  • Address for reprint requests and other correspondence: S. J. London, NIEHS, 111 TW Alexander Dr., PO Box 12233, MD A3-05, Research Triangle Park, NC 27709 (e-mail: ).