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. 2014 Jun;85(6):839–848. doi: 10.1124/mol.113.091272

Sensory Nerve Terminal Mitochondrial Dysfunction Induces Hyperexcitability in Airway Nociceptors via Protein Kinase C

Stephen H Hadley 1, Parmvir K Bahia 1, Thomas E Taylor-Clark 1,✉
PMCID: PMC4014670  PMID: 24642367

Abstract

Airway sensory nerve excitability is a key determinant of respiratory disease-associated reflexes and sensations such as cough and dyspnea. Inflammatory signaling modulates mitochondrial function and produces reactive oxygen species (ROS). Peripheral terminals of sensory nerves are densely packed with mitochondria; thus, we hypothesized that mitochondrial modulation would alter neuronal excitability. We recorded action potential firing from the terminals of individual bronchopulmonary C-fibers using a mouse ex vivo lung-vagal ganglia preparation. C-fibers were characterized as nociceptors or non-nociceptors based upon conduction velocity and response to transient receptor potential (TRP) channel agonists. Antimycin A (mitochondrial complex III Qi site inhibitor) had no effect on the excitability of non-nociceptors. However, antimycin A increased excitability in nociceptive C-fibers, decreasing the mechanical threshold by 50% and increasing the action potential firing elicited by a P2X2/3 agonist to 270% of control. Antimycin A–induced nociceptor hyperexcitability was independent of TRP ankyrin 1 or TRP vanilloid 1 channels. Blocking mitochondrial ATP production with oligomycin or myxothiazol had no effect on excitability. Antimycin A–induced hyperexcitability was dependent on mitochondrial ROS and was blocked by intracellular antioxidants. ROS are known to activate protein kinase C (PKC). Antimycin A–induced hyperexcitability was inhibited by the PKC inhibitor bisindolylmaleimide (BIM) I, but not by its inactive analog BIM V. In dissociated vagal neurons, antimycin A caused ROS-dependent PKC translocation to the membrane. Finally, H2O2 also induced PKC-dependent nociceptive C-fiber hyperexcitability and PKC translocation. In conclusion, ROS evoked by mitochondrial dysfunction caused nociceptor hyperexcitability via the translocation and activation of PKC.

Introduction

A large subset of sensory nerves in the somatosensory and visceral systems is nociceptive; it is adapted to detect noxious and potential noxious stimuli and then trigger sensations, behaviors, and reflexes that are intended to diminish future noxious threats. Nevertheless, nociceptor excitability is not static and, in damaged and inflamed tissues, nociceptors can become hyperexcitable such that subthreshold stimuli inappropriately evoke nociceptive responses or superthreshold stimuli evoke inappropriately large responses. The bronchopulmonary airways are densely innervated by nociceptive vagal C-fibers, whose activation leads to the initiation of reflexes (cough, hypersecretion, bronchospasm) and sensations (dyspnea, itch, urge to cough) (Carr and Undem, 2003). Airway nociceptive reflexes are exacerbated in inflammatory and infectious airway disease (Riccio et al., 1996; Carr and Lee, 2006; Sabogal et al., 2005; Nassenstein et al., 2007), most clearly observed clinically as an excessive and hyperresponsive cough in respiratory disease (Jesenak et al., 2009; Morice, 2010). As such, there is considerable interest in understanding the mechanisms through which inflammation and other noxious stimuli modulate nociceptor excitability.

Increasing evidence suggests that mitochondrial dysfunction contributes to inflammation in multiple tissues via the actions of mitochondrially evoked reactive oxygen species (ROS) (Reddy, 2011; Sena and Chandel, 2012). DNA mutations leading to aberrant mitochondrial protein function/expression have been known for many years to dramatically impact ATP production and ROS signaling. Recently, it was shown that numerous signaling events can also increase mitochondrial ROS production, for example, hypoxia (Bell et al., 2007), tumor necrosis factor-α (Corda et al., 2001), neurotrophins via p75NTR (Pehar et al., 2007), transforming growth factor-β (Michaeloudes et al., 2011), and Toll-like receptors (West et al., 2011). This is likely of particular importance for the peripheral terminals of sensory nerves, which contain a high density of mitochondria (Hung et al., 1973; von During and Andres, 1988). We have previously shown an interaction between nerve terminal mitochondria and nerve terminal function in a study that demonstrated that mitochondrial ROS can cause overt bronchopulmonary C-fiber activation (action potential discharge) via the selective gating of transient receptor potential (TRP) ankyrin 1 (TRPA1) (Nesuashvili et al., 2013). It is not yet known whether mitochondrial ROS can also modulate neuronal excitability.

To study the effect of mitochondrial dysfunction and ROS on bronchopulmonary C-fiber excitability, we have used antimycin A, a selective inhibitor of the Qi site on complex III in the mitochondrial electron transport chain. This selective inhibitor stimulates mitochondrial superoxide formation and the subsequent production of numerous ROS (Turrens et al., 1985; Gyulkhandanyan and Pennefather, 2004; Tretter et al., 2007; Stowe and Camara, 2009). Cellular oxidation can modulate the function of numerous proteins, including ion channels and enzymes (Thannickal and Fanburg, 2000; Stowe and Camara, 2009). In particular, protein kinase C (PKC) can be activated by ROS independently of Ca2+ or diacylglycerol (DAG) (Gopalakrishna and Anderson, 1989; Knapp and Klann, 2000). PKC activation is known to induce nociceptor hyperexcitability in somatosensory neurons via its phosphorylation of voltage- and ligand-gated ion channels (Cesare et al., 1999; Amadesi et al., 2004; Baker, 2005; Wu et al., 2012). In this study, we found that mitochondrial ROS induced hyperexcitability of nociceptive bronchopulmonary C-fibers via oxidative activation and translocation of PKC.

Materials and Methods

All experiments were approved by the University of South Florida Institutional Animal Care and Use Committee.

Bronchopulmonary C-Fiber Extracellular Recordings

Male C57BL/6J mice (The Jackson Laboratory, Bar Harbor, ME) were killed by CO2 asphyxiation, followed by exsanguination. The innervated isolated lung preparation was prepared, as previously described (Nassenstein et al., 2008; Nesuashvili et al., 2013). Briefly, the airways and lungs with their intact extrinsic innervation (vagus nerve including vagal ganglia) were taken and placed in a dissecting dish containing Krebs bicarbonate buffer solution composed of the following (mM): 118 NaCl, 5.4 KCl, 1.0 NaH2PO4, 1.2 MgSO4, 1.9 CaCl2, 25.0 NaHCO3, and 11.1 D-glucose, and equilibrated with 95% O2 and 5% CO2 (pH 7.3–7.4) and containing 3 μM indomethacin. Connective tissue was trimmed away, leaving the trachea and lungs with their intact nerves. The airways were then pinned to the larger compartment of a custom-built two-compartment recording chamber that was lined with silicone elastomer. A vagal ganglion was pulled into the adjacent compartment of the chamber through a small hole and pinned. Both compartments were separately superfused with Krebs bicarbonate buffer (37°C). A sharp glass electrode was pulled by a Flaming Brown micropipette puller (P-87; Sutter Instruments, Novato, CA) and filled with 3 M NaCl solution. The electrode was inserted and placed near the cell bodies of vagal ganglion. The recorded action potentials were amplified (Microelectrode AC amplifier 1800; A-M Systems, Everett, WA), filtered (0.3 kHz low cutoff and 1 kHz high cutoff), and monitored on an oscilloscope (TDS1002B; Tektronix, Beaverton, OR). The scaled output from the amplifier was captured and analyzed by a Macintosh computer using NerveOfIt software (Phocis, Baltimore, MD). Action potential discharge was quantified off-line and recorded in 1-second bins. The background activity was usually either absent or less than 2 Hz.

Nerve Fiber Characterization.

To measure conduction velocity, an electrical stimulation (S44; Grass Instruments, Quincy, MA) was applied to the center of the receptive field. The conduction velocity of the individual bronchopulmonary afferent was calculated by dividing the distance along the nerve pathway by the time delay between the shock artifact and the action potential evoked by electrical stimulation. Mechanical stimulation was determined using manual application of calibrated von Frey fibers to the center of the receptive field. C-fibers were also characterized for their expression of TRPA1 and TRP vanilloid 1 (TRPV1) channels by their response to the selective TRPA1 agonist allyl isothiocyanate (AITC; 300 μM), and to the selective TRPV1 agonist capsaicin (1 μM). Chemical stimuli were intratracheally applied as a 1-ml bolus over 10 seconds. A response to either mechanical or chemical stimuli was considered positive if the number of action potentials in any 1-second bin was twice the average background response. All excitatory chemical treatments were separated by at least a 15-minute wash. Due to potential heterologous desensitization, capsaicin was only given at the end of each experiment. Only those bronchopulmonary fibers that had a positive response to both mechanical and chemical stimuli were included in the study. Due to a lack of TRPV1 channels in bronchopulmonary C-fibers from TRPV1−/− mice, nociceptive status was assessed solely by conduction velocity and the response to TRPA1 agonists (Kollarik et al., 2003; Nassenstein et al., 2008).

Determination of Hyperexcitability via Changes in Mechanical Threshold.

The threshold for mechanical pressure required to evoke action potential discharge was determined by progressively decreasing the weight of von Frey fiber applied to the lung receptive field until no response was evoked. The least force required to evoke mechanically induced action potential firing was then repeated once more to confirm threshold reproducibility (Riccio et al., 1996). The lungs were then treated with antimycin A (20 μM). Ten minutes later, von Frey fibers were used to determine the mechanical threshold again following mitochondrial modulation. Hyperexcitability was quantified by comparing the mechanical threshold before and after mitochondrial modulation (paired analysis). Due to the irreversible inhibition of complex III by antimycin (Turrens et al., 1985; Stowe and Camara, 2009), only one antimycin A treatment was used per preparation.

Determination of Hyperexcitability via Changes in Response to Chemical Stimulus.

We determined the action potential discharge response to α,β-methylene ATP (α,β-mATP; P2X2/3 agonist, 30 μM) intratracheally applied as a 1-ml bolus over 10 seconds (Nassenstein et al., 2010). This submaximal dose is sufficient to activate all P2X2/3 expressing bronchopulmonary C-fibers (Nassenstein et al., 2010). The lungs were then treated with antimycin A (20 μM) or other mitochondrial modulators. Ten minutes later, the action potential discharge response to α,β-mATP (30 μM) was again determined. Hyperexcitability to α,β-mATP treatment was quantified in raw form in two ways, as follows: either by comparing the total number of action potentials discharged within 120 seconds or comparing the peak response (maximum number of action potentials that occurred within any 1-second bin) before and after mitochondrial modulation (both paired analyses). Due to variations in responsiveness to α,β-mATP between bronchopulmonary C-fibers, we also normalized each fiber’s second response to α,β-mATP to its first α,β-mATP response, as follows: (total number of action potentials discharged in second response/total number of action potentials discharged in first response) × 100. This normalized quantification of excitability could then be compared between different treatment groups in our mechanistic studies (unpaired analysis). Due to the irreversible inhibition of complex III by antimycin (Turrens et al., 1985; Stowe and Camara, 2009), only one antimycin A treatment was used per preparation.

Knockout Mice

Male TRPV1−/− (The Jackson Laboratory; B6.129X1-Trpv1tm1Jul/J) were mated with female TRPV1−/−. Genotype of offspring was confirmed by polymerase chain reaction.

Dissociation of Mouse Vagal Ganglia

Male C57BL/6J mice were killed by CO2 asphyxiation, followed by exsanguination. Vagal ganglia were immediately isolated and enzymatically dissociated using previously described methods (Nesuashvili et al., 2013). Isolated neurons were plated onto poly(D-lysine)– and laminin-coated coverslips, incubated at 37°C in L-15 (supplemented with 10% fetal bovine serum), and used within 24 hours.

PKC Translocation Immunocytochemistry

Dissociated mouse vagal neurons on glass coverslips were treated with either antimycin A (10 μM), phorbol-12-myristate-13-acetate (PMA; 300 nM), or H2O2 (300 μM) by direct addition to culture media. Neurons were then fixed 2 minutes later using fresh 4% paraformaldehyde to preserve PKC subcellular location. In some cases, antioxidants were applied to media prior to antimycin A, as follows: 90 minutes prior for tempol (1 mM); 15 minutes prior for MnTMPyP [manganese(III) tetrakis(1-methyl-4-pyridyl)porphyrin; 50 μM]. Neurons were then permeabilized using phosphate-buffered saline containing 0.1% Triton X-100 for 10 minutes at room temperature and blocked using phosphate-buffered saline with 0.0025% Tween 20 (PBST) containing 2% BSA for 45 minutes at room temperature. Cells were stained using a pan-PKC primary antibody (H-300; Santa Cruz Biotechnology, Santa Cruz, CA). Antibody concentrated to 0.5 mg/ml was diluted 1:50 in blocking buffer and left on cells overnight at 4°C. Cells were washed three times using PBST and then incubated with a CY5-labeled anti-rabbit secondary antibody (Life Technologies, Carlsbad, CA) diluted 1:1000 in blocking buffer and left on cells for 1 hour at room temperature. After three washes with PBST, coverslips were mounted onto glass slides using VECTASHIELD(R) HardSet Mounting Medium with 4′,6-diamidino-2-phenylindole (Vector Laboratories, Burlingame, CA). Imaging was performed using a Leica SP2 confocal microscope with a 63×objective, as follows: 4′,6-diamidino-2-phenylindole excitation of 405 nm, emission range of 415–480 nm; Cy5 excitation of 633 nm, emission range of 625–735 nm. Analysis of staining intensity was performed using ImageJ. A straight line was drawn across each cell in a region excluding the nucleus, and the Multi-plot function was used to plot the profile of the staining. The ratio of the peak staining at the edge of the cell to the average staining across the center of the cell was generated to give the membrane/cytosolic ratio.

Statistical Analysis

Data were analyzed using GraphPad Prism (GraphPad Software, La Jolla, CA). Where appropriate, paired or unpaired Student’s t test was used. A P value less than 0.05 was taken as significant. All data were expressed as mean ± S.E.M.

Results

Antimycin A Decreases Mechanical Thresholds in Nociceptive C-Fibers.

To study the effect of mitochondrial modulation on the excitability of bronchopulmonary C-fiber terminals, we have used a mouse ex vivo lung-vagal ganglia preparation. In this preparation, the action potential discharge evoked in individual nerves by either mechanical or chemical stimuli to the airways can be quantified. Previous studies have shown that the majority of fibers innervating the lung are slowly conducting and express TRPA1 and TRPV1 (Nassenstein et al., 2008; Nesuashvili et al., 2013). This subset of nerves is thought to serve as nociceptors in the mouse airways. In this study, we characterized bronchopulmonary sensory fibers based on their conduction velocity and their responsiveness to the selective TRPA1 and TRPV1 agonists (AITC and capsaicin, respectively). Bronchopulmonary fibers were considered nociceptive if they expressed TRPV1. As described before (Kollarik et al., 2003), TRPV1 expression was largely confined to fibers with conduction velocities <0.7 m/s.

We first studied the effect of antimycin-induced mitochondrial modulation on the mechanical threshold for bronchopulmonary fibers using punctate stimuli of the lung tissue with von Frey fibers. All bronchopulmonary fibers are directly activated by punctate force (Kollarik et al., 2003; Nassenstein et al., 2008) in a P2X-independent manner (Weigand et al., 2012), and in this study a maximum von Frey stimulus (1.4 g) evoked peak action discharge of 16.8 ± 1.7 Hz. In this work, we determined the minimum force required to evoke action potential discharge in each fiber using the rationale that an increase in excitability would decrease the threshold for mechanically induced action potential discharge (Riccio et al., 1996). Each fiber’s threshold was stable over the duration of experimentation (Fig. 1A). Antimycin A (20 μM) evoked significant action potential discharge in TRPA1-expressing bronchopulmonary fibers via a ROS-dependent TRPA1 activation—this mechanism is described in our previous study (Nesuashvili et al., 2013). The direct activation of these TRPA1-expressing nerves lasted from 2 to 7 minutes; other fibers were not overtly activated. We tested the mechanical threshold 10 minutes following antimycin A treatment and compared with the mechanical threshold previous to antimycin A treatment. Antimycin A (20 μM) decreased the threshold for mechanical stimulation by approximately 50% in nociceptive neurons (P < 0.005, n = 13; Fig. 1, B and C). Seven of these 13 fibers were TRPA1-expressing nociceptors, but there was no correlation of expression of TRPA1 and the antimycin A–induced threshold decrement (data not shown). However, antimycin A had no effect on the threshold for mechanical stimulation for faster conducting, non-nociceptive bronchopulmonary fibers (P > 0.05, n = 6; Fig. 1C), which do not express TRPA1 or TRPV1.

Fig. 1.

Fig. 1.

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Antimycin A decreases the threshold required for mechanical stimulation in nociceptive C-fibers. (A) Representative trace of action potential discharge elicited by von Frey fibers [a = 0.4 g (threshold); b = 0.16 g (subthreshold)] in a nociceptive bronchopulmonary C-fiber [defined by conduction velocity (CV) and sensitivity to capsaicin (data not shown)]. Inserts show magnified timescale of responses. (B) Representative trace of mechanical threshold determination in a nociceptive bronchopulmonary C-fiber before (control) and 10 minutes after treatment with antimycin A (20 μM). Application of von Frey stimulus denoted by arrows. (C) Mean ± S.E.M. mechanical threshold for action potential discharge before (control, hatched bars) and 10 minutes after treatment with antimycin A (20 μM, black bars) in nociceptive and non-nociceptive bronchopulmonary C-fibers, as defined by sensitivity to TRPV1 agonist (capsaicin, 1 μM) and conduction velocity (Kollarik et al., 2003). Significant difference in mechanical sensitivity after antimycin A, *P < 0.005.

Antimycin A Increases Excitability to Chemical Stimuli in Nociceptive C-Fibers.

The standardization of manually operated von Frey fiber mechanical stimulation of C-fibers can be challenging, particularly in a three-dimensional tissue, such as the lung. We therefore extended our study of mitochondrial modulation-induced hyperexcitability to determine its effects on bronchopulmonary C-fiber responses to chemical stimuli. Previous studies showed that the vast majority of mouse bronchopulmonary sensory nerves are activated by the selective P2X2/3 agonist α,β-mATP and that this response is reproducible (Kollarik et al., 2003; Nassenstein et al., 2010). Consistent with these reports, 30 μM α,β-mATP activated >90% of bronchopulmonary C-fibers regardless of their conduction velocity or TRP channel expression. To confirm the reproducibility of P2X2/3 receptor responses, we determined the action potential discharge to repeated treatment of 30 μM α,β-mATP in 18 nociceptive C-fibers (separated by 15 minutes). There was no significant difference in either the total number of action potentials evoked (first, 64.2 ± 9.5; second, 62.7 ± 11.1; Fig. 2B) or the peak response (first, 17.7 ± 2.3 Hz; second, 15.9 ± 2.21 Hz; Fig. 2C).

Fig. 2.

Fig. 2.

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Antimycin A increases α,β-mATP–induced responses in nociceptive C-fibers. (A) Representative traces of action potential discharge evoked by 10-second challenge with α,β-mATP (P2X2/3 agonist, 30 μM) in a nociceptive bronchopulmonary C-fiber before (control) and 10 minutes after treatment with antimycin A (20 μM). Quantification of action potential discharge is shown below each trace. (B) Mean ± S.E.M. total action potential discharge evoked by α,β-mATP (30 μM) before (control, hatched bars) and 10 minutes after treatment with either vehicle (white bar) or antimycin A (20 μM, black bars) in nociceptive and non-nociceptive bronchopulmonary C-fibers. Significant difference in second α,β-mATP–induced responses after antimycin A, *P < 0.01. (C) Mean ± S.E.M. peak action potential discharge evoked by α,β-mATP (30 μM) before (control, hatched bars) and 10 minutes after treatment with either vehicle (white bar) or antimycin A (20 μM, black bars) in nociceptive and non-nociceptive bronchopulmonary C-fibers. Significant difference in second α,β-mATP–induced responses after antimycin A, *P < 0.01. CV, conduction velocity.

Ten minutes after treatment with 20 μM antimycin A, the response to 30 μM α,β-mATP was significantly increased in nociceptive C-fibers (n = 24; Fig. 2A), as assessed by the total number of action potentials evoked (first, 105 ± 34.9; second, 220 ± 79.4; P < 0.01) (Fig. 2B) and the peak response (first, 15.4 ± 1.5 Hz; second, 20.5 ± 1.7 Hz; P < 0.01) (Fig. 2C). Similar to our mechanical threshold data, antimycin A failed to induce hyperexcitability to α,β-mATP in non-nociceptive bronchopulmonary fibers (n = 9).

The action potential discharge to α,β-mATP varied significantly from nerve to nerve, as follows: of 189 fibers examined in the present study, the total number of action potentials evoked ranged from 16 to 280, and the peak discharge ranged from 3 to 37. To account for the intrinsic variability in neuronal responses, we quantified excitability by the change in the response to the second α,β-mATP treatment (i.e., after antimycin A) as a percentage of each individual fiber’s response to the first α,β-mATP treatment. As such, antimycin increased nociceptor excitability to 276 ± 54% (n = 24) compared with vehicle, which had no effect on excitability (105 ± 14%, n = 18, P < 0.005) (see Figs. 3B and 4, A and B).

Fig. 3.

Fig. 3.

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Antimycin A–induced nociceptor hyperexcitability to α,β-mATP is independent of TRPA1 and TRPV1. (A) Mean ± S.E.M. response to second application of α,β-mATP (30 μM) normalized to response to first application of α,β-mATP prior to antimycin A (20 μM). Bronchopulmonary C-fibers were characterized by conduction velocity and sensitivity to TRPV1 agonist (capsaicin, 1 μM) and TRPA1 agonist (AITC, 300 μM) (Kollarik et al., 2003; Nassenstein et al., 2008). Significant increase in second α,β-mATP–induced responses after antimycin A, *P < 0.05. (B) Mean ± S.E.M. response to second application of α,β-mATP (30 μM) normalized to response to first application of α,β-mATP prior to either vehicle (white bar) or antimycin A (20 μM, black bars) in nociceptive C-fibers. The role of TRPV1 in hyperexcitability was determined using pretreatment with the selective TRPV1 antagonist iodoresiniferatoxin (IRTX; 1 μM) or by using C-fibers from TRPV1−/− mice. Significant increase in second α,β-mATP–induced responses after antimycin A compared with vehicle, *P < 0.05. CV, conduction velocity.

Fig. 4.

Fig. 4.

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Antimycin A–induced nociceptor hyperexcitability to α,β-mATP is dependent on mitochondrial ROS and PKC. (A) Mean ± S.E.M. response to second application of α,β-mATP (30 μM) normalized to response to first application of α,β-mATP prior to either vehicle (white bar), antimycin A (20 μM, black bar), oligomycin (10 μM, dark gray bar), or myxothiazol (500 nM, light gray bar) in nociceptive C-fibers. Significant increase in second α,β-mATP–induced responses compared with vehicle, *P < 0.01. (B) Mean ± S.E.M. response to second application of α,β-mATP (30 μM) normalized to response to first application of α,β-mATP prior to either vehicle (white bar) or antimycin A (20 μM, black bars) in nociceptive C-fibers. The roles of ROS and PKC in hyperexcitability to α,β-mATP were determined using pretreatment with DTT (1 mM), NAC (1 mM), GSH (1 mM), myxothiazol (500 nM), BIM I (1 μM), or BIM V (1 μM). Significant increase in second α,β-mATP–induced responses after antimycin A compared with vehicle, *P < 0.05. Significant reduction in antimycin A–induced hyperexcitability to α,β-mATP, #P < 0.05.

Given that our previous findings had shown that mitochondrial ROS evoked action potential discharge in nociceptors in a TRPA1-dependent manner, we compared the antimycin A–induced excitability in nociceptors that specifically expressed TRPA1 (n = 8) with those nociceptors that did not express TRPA1 (n = 16). Antimycin A increased excitability in both subsets of nociceptive C-fibers (Fig. 3A), indicating that TRPA1 expression was not required for mitochondrial modulation–associated hyperexcitability to α,β-mATP. There was a trend toward a greater increase in excitability to α,β-mATP in TRPA1-negative C-fibers—perhaps due to mild nonspecific tachyphylaxis occurring following substantial TRPA1-mediated depolarization—although this did not reach statistical significance (P = 0.19).

Our data demonstrating that antimycin A evoked hyperexcitability only in nociceptors also led us to test the hypothesis that TRPV1, which is expressed exclusively in almost all nociceptors, was involved. Using a pretreatment of the selective TRPV1 antagonist iodoresiniferatoxin (1 μM, n = 8) (Wahl et al., 2001), we found that antimycin A–induced sensitization to α,β-mATP in nociceptors was not reduced (Fig. 3B). In addition, we complemented these studies using ex vivo lung-vagal tissue from TRPV1−/− mice. In this study, genetic ablation of TRPV1 failed to reduce antimycin A–induced nociceptor hyperexcitability to α,β-mATP (n = 14). We therefore conclude that, despite the exclusivity of hyperexcitability in the nociceptive population, TRPV1 is not required for antimycin A–induced hyperexcitability to α,β-mATP.

Comparisons with Other Mitochondrial Modulators.

The biochemical response to antimycin A is well characterized in isolated mitochondria and numerous cell types. Antimycin A selectively inhibits the Qi site on complex III in the electron transport chain in the mitochondria, resulting in superoxide formation and the subsequent production of numerous ROS, mitochondrial membrane depolarization, and a decrease in ATP production (Turrens et al., 1985; Gyulkhandanyan and Pennefather, 2004; Tretter et al., 2007; Stowe and Camara, 2009). Other specific complex inhibitors have also been described, in particular oligomycin and myxothiazol. Oligomycin inhibits complex V (also known as ATP synthase) and thus prevents ATP production by the oxidative phosphorylation machinery (Sipos et al., 2003; Hool and Corry, 2007). Inhibition of complex V also evokes a mild hyperpolarization of the mitochondrial membrane potential but has little effect on superoxide production in most systems. Ten minutes following oligomycin (10 μM) treatment, there was no significant change in action potential discharge in response to α,β-mATP (n = 8; Fig. 4A). Myxothiazol is a selective inhibitor of the Qo site on complex III. Inhibition of Qo causes mitochondrial membrane potential depolarization and a decrease in ATP production but only evokes very mild superoxide production (Turrens et al., 1985; Gyulkhandanyan and Pennefather, 2004; Tretter et al., 2007). Ten minutes following myxothiazol (500 nM) treatment, there was also no change in action potential discharge to α,β-mATP (n = 7; Fig. 4A). These data suggest that antimycin A–induced nociceptor hyperexcitability to α,β-mATP is not specifically due to a decrease in ATP production.

Involvement of ROS and PKC in Antimycin A–Induced Hyperexcitability to α,β-mATP.

We hypothesized that antimycin A–induced nociceptor hyperexcitability to α,β-mATP was mediated by the actions of mitochondrially derived ROS. Many biologic actions of ROS are due to their ability to oxidize protein residues (e.g., cysteine), which can be reversed by a reducing agent such as dithiothreitol (DTT) or by quenching with a cysteine antioxidant, such as N-acetyl-cysteine (NAC) or glutathione (GSH). DTT is membrane permeable. NAC is slowly actively transported across plasma membranes (Raftos et al., 2007) and has previously been shown to quench intracellular ROS pathways (Hongpaisan et al., 2004; Takahashi et al., 2011). GSH is membrane impermeable and is not subject to active transport as NAC is (Raftos et al., 2007). We tested the contribution of oxidation to antimycin A–induced hyperexcitability by treating the lungs with DTT (1 mM) or NAC (1 mm) or GSH (1 mM) for 10 minutes prior to and during antimycin A (20 μM) treatment. Both DTT (n = 4) and NAC (n = 8) completely abolished the antimycin A–induced increase in excitability to α,β-mATP (P < 0.01 compared with antimycin A alone), whereas GSH (n = 4) failed to inhibit the antimycin A–induced hyperexcitability to α,β-mATP [P < 0.005 compared with vehicle (no antimycin), P = 0.17 compared with antimycin A alone] (Fig. 4B). Based on the differential distribution of GSH (extracellular) and NAC (extracellular and intracellular), these data suggest that intraneuronal ROS contribute to the antimycin A–induced hyperexcitability to α,β-mATP.

Selective inhibition of Qo using myxothiazol is known to substantially reduce subsequent superoxide production evoked by inhibition of the Qi site (antimycin A) (Turrens et al., 1985; Gyulkhandanyan and Pennefather, 2004; Tretter et al., 2007). We have previously shown that myxothiazol pretreatment prior to antimycin A greatly reduced antimycin A–induced TRPA1 activation (Nesuashvili et al., 2013). In this study, we pretreated the lungs with myxothiazol (500 nM) 3 minutes prior to antimycin A (20 μM) treatment. Myxothiazol prevented the development of antimycin A–induced nociceptor hyperexcitability to α,β-mATP (Fig. 4B), as follows: for antimycin alone the response to the second α,β-mATP was 276 ± 54% (n = 24); with myxothiazol pretreatment of antimycin A the response to the second α,β-mATP was 128 ± 41% (n = 9, P < 0.05). These findings suggest that ROS evoked from mitochondrial complex III are required for antimycin A–induced hyperexcitability to α,β-mATP.

Numerous studies of dorsal root ganglion (DRG) nociceptors involved in somatosensory pain have shown that PKC is a powerful regulator of nociceptor excitability (Cesare et al., 1999; Baker, 2005; Wu et al., 2012). Interestingly, there has been increasing evidence that many PKC isoforms can be activated by cellular oxidants (e.g., superoxide and H2O2) independently of either Ca2+ or lipid mediators, such as DAG (Gopalakrishna and Anderson, 1989; Knapp and Klann, 2000; Cosentino-Gomes et al., 2012). We hypothesized that mitochondrial ROS induces airway nociceptor hyperexcitability to α,β-mATP via activation of protein kinase C. For these studies, we used a selective PKC inhibitor called bisindolylmaleimide I (BIM I; also known as GF 109203X and Gö 6850) and its inactive analog BIM V (also known as Ro 31-6045). These compounds are membrane permeable and differ in only one side chain. BIM I is a potent PKC inhibitor with an IC50 below 500 nM, whereas BIM V has an IC50 for PKC >100 μM (Toullec et al., 1991; Davis et al., 1992). When the lungs were pretreated with BIM I (1 μM) for 10 minutes prior to and during antimycin A (20 μM) treatment, there was no increase in nociceptor hyperexcitability, as follows: the response to the second α,β-mATP was only 136 ± 16% (n = 14, P < 0.01 compared with antimycin A alone) (Fig. 4B). However, pretreatment with the inactive analog BIM V (1 μM) for 10 minutes prior to and during antimycin A (20 μM) treatment failed to prevent antimycin A–induced nociceptor hyperexcitability, as follows: the response to the second α,β-mATP was 231 ± 56% [n = 20, P < 0.05 compared with vehicle (no antimycin), P = 0.28 compared with antimycin A alone] (Fig. 4B). These data suggest that antimycin A–induced nociceptor hyperexcitability to α,β-mATP is dependent on the actions of PKC.

Lastly, we determined whether PKC activation or ROS was sufficient to evoke vagal bronchopulmonary C-fiber hyperexcitability in the absence of mitochondrial dysfunction by antimycin A. PMA is a potent and selective activator of PKC. As before in the antimycin A studies, the nociceptor action potential response to 30 μM α,β-mATP was determined. Then the lungs were treated with 3 μM PMA, and the response to α,β-mATP was determined 2 minutes later. As expected, PMA induced nociceptor hyperexcitability to α,β-mATP, as follows: the response to the second α,β-mATP was 267 ± 111% (n = 11, P < 0.05 compared with vehicle) (Fig. 5). Using a similar protocol, we found that exogenously applied H2O2 also induced airway nociceptor hyperexcitability to α,β-mATP, as follows: 2 minutes after H2O2 (1 mM), the response to the second α,β-mATP was 207 ± 35% (n = 15, P < 0.01 compared with vehicle) (Fig. 5). Consistent with the mechanism of antimycin A–induced hyperexcitability to α,β-mATP, H2O2-induced hyperexcitability to α,β-mATP was prevented by pretreatment with the selective PKC inhibitor BIM I (1 μM), as follows: the response to the second α,β-mATP was only 89 ± 8% (n = 5, P < 0.005 compared with H2O2 alone) (Fig. 5).

Fig. 5.

Fig. 5.

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H2O2 increases α,β-mATP–induced responses in nociceptive C-fibers via PKC. Mean ± S.E.M. response to second application of α,β-mATP (30 μM) normalized to response to first application of α,β-mATP prior to either vehicle (white bar), PMA (3 μM, gray bar), or H2O2 (1 mM, black bars) in nociceptive C-fibers. The roles of PKC in H2O2-induced hyperexcitability to α,β-mATP were determined using pretreatment with BIM I (1 μM). Significant increase in second α,β-mATP–induced responses compared with vehicle, *P < 0.05. Significant reduction in H2O2-induced hyperexcitability to α,β-mATP, #P < 0.005.

Some of the airway C-fibers (8 of 16) used in the H2O2 hyperexcitability studies expressed TRPA1 (as determined by sensitivity to AITC). Although not the focus of this study, TRPA1 has been shown to be activated by H2O2 in vitro (Andersson et al., 2008). Consistent with those findings, H2O2 robustly activated TRPA1-expressing C-fibers (peak discharge of 6.4 ± 1.8 Hz) compared with those C-fibers that did not express TRPA1 (peak discharge of 0.6 ± 0.6 Hz). H2O2-induced C-fiber activation was short-lived, and the response was absent prior to the second α,β-mATP stimulus. As with antimycin A–induced hyperexcitability to α,β-mATP, the H2O2-induced hyperexcitability to α,β-mATP was unaffected by the presence or absence of TRPA1 (data not shown).

Antimycin A Induces PKC Translocation in Vagal Neurons.

Critical to PKC activity is its subcellular location (Cosentino-Gomes et al., 2012). Typically, PKC isoforms remain in the cytosol while inactive. Upon activation, PKC translocates to the plasma membrane, in which it engages in phosphorylation of a wide variety of protein targets. Given that previous studies demonstrated multiple isoforms of PKC heterogeneously expressed throughout the somatosensory DRG neuronal population (Cesare et al., 1999; Hayase et al., 2007; Wu et al., 2012; Zhang et al., 2012) and that PKC isoform expression has not yet been characterized in vagal neurons, we decided in this study to observe global PKC translocation using a pan-PKC antibody that would detect all PKC isoforms. All vagal neurons were immunoreactive for PKC with no fluorescent signal observed in the absence of primary antibody (data not shown). In control conditions, the vast majority of PKC immunoreactivity was located in the cytosol (n = 225) (Fig. 6). As expected, there was a significant increase in PKC immunoreactivity at the plasma membrane 2 minutes after PMA (300 nM) treatment (n = 216, P < 0.0001 compared with control), indicating that activation of PKC causes translocation to the plasma membrane. Consistent with our electrophysiological data at the airway vagal nerve terminal, antimycin A (10 μM) induced significant PKC translocation to the plasma membrane (n = 134, P < 0.0005 compared with control). Antimycin A–induced PKC translocation was abolished by the pretreatment of the neurons with a combination of the antioxidants tempol (1 mM) and MnTMPyP (50 μM) (n = 41, P < 0.05 compared with antimycin alone), indicating a critical role of mitochondrial ROS in these responses. Finally, we found that exogenously applied H2O2 (300 μM) was sufficient to induce significant PKC translocation to the membrane (n = 173, P < 0.005 compared with control). Overall, these data indicate that mitochondrial ROS induce PKC translocation, consistent with a role of PKC in mitochondrial ROS-induced airway nociceptor hyperexcitability.

Fig. 6.

Fig. 6.

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Antimycin A induces PKC translocation to the plasma membrane via ROS in vagal neurons. (A–E) Representative confocal images of PKC (red) and nuclei (4′,6-diamidino-2-phenylindole, blue) in dissociated vagal sensory neurons. Scale denotes 25 μm. In dissociated ganglia preparations, neurons are typically >20 μm in diameter; other non-neuronal cell types are also present that are small with substantial nuclei (Nassenstein et al., 2010). (A) Control conditions. (B) After PMA (300 nM). (C) After antimycin A (10 μM). (D) After antimycin A (10 μM), pretreated with tempol (1 mM) and MnTMPyP (50 μM). (E) After H2O2 (300 μM). (F) Mean ± S.E.M. PKC membrane/cytosolic ratio for experiments described in A–E. Significant increase in membrane/cytosolic ratio compared with vehicle, *P < 0.005. Significant reduction in membrane/cytosolic ratio compared with antimycin A alone, #P < 0.05.

Discussion

Inflammation alters mitochondrial function (Sena and Chandel, 2012). Given that peripheral sensory nerve terminals are densely packed with mitochondria (Hung et al., 1973; von During and Andres, 1988), we investigated the effect of mitochondrial modulation on sensory nerve terminal excitability. In this study, ROS evoked by mitochondrial dysfunction induced hyperexcitability to both mechanical and chemical stimuli in nociceptive bronchopulmonary C-fibers but had little effect on the excitability of non-nociceptive nerves. Nociceptive nerves protect the innervated organ through the initiation of reflexes and behavioral responses (Carr and Undem, 2003), and hyperexcitability contributes to aberrant signs and symptoms in inflammatory and infectious states (Meyer et al., 1996; Carr and Lee, 2006). Thus, we present evidence of a novel mechanism by which nocifensive airway responses can be inappropriately modulated.

There are two subtypes of mouse vagal bronchopulmonary C-fibers: slowly conducting C-fibers express various nociceptor receptors such as TRPV1 (Kollarik et al., 2003) and are thought to be synonymous with nociceptive afferents in humans that mediate cough, dyspnea, and reflex bronchospasm (Carr and Undem, 2003); faster conducting C-fibers do not express TRPV1 and are considered to be non-nociceptive—perhaps involved in regulation of eupnic breathing. We previously demonstrated that antimycin A, a Qi site inhibitor of the electron transfer chain complex III that induces production of mitochondrial ROS (Stowe and Camara, 2009), caused the activation of nociceptive C-fibers via the activation of another nociceptive ion channel TRPA1 (Nesuashvili et al., 2013). This activation typically lasted 2–7 minutes. In this study, we found that antimycin A evoked a profound increase in nociceptive C-fiber excitability as determined by decreased threshold for punctate mechanical stimulation and by increased action potential discharge to α,β-mATP, an agonist for P2X2/3 ion channels. Antimycin A failed to increase non-nociceptive nerve excitability. Unlike antimycin A–induced C-fiber activation (Nesuashvili et al., 2013), antimycin A–induced hyperexcitability to α,β-mATP was not dependent on the expression of TRPA1. Despite the exclusivity of hyperexcitability in the TRPV1-expressing nociceptive population, TRPV1 was also not required for antimycin A–induced hyperexcitability to α,β-mATP. We briefly investigated the effect of 1 mM H2O2 on nociceptor activation and found that activation correlated with sensitivity to the TRPA1 agonist AITC. This is consistent with other studies using similar doses (Andersson et al., 2008), although studies using 120 mM H2O2 suggest that, along with TRPA1, P2X channels also contribute to H2O2-induced C-fiber activation (Lin et al., 2013).

Our mechanistic studies suggest that antimycin A evokes airway nociceptor hyperexcitability to α,β-mATP via the actions of ROS and PKC, largely independent of mitochondrial ATP production. Antimycin A, oligomycin (complex V inhibitor), and myxothiazol (complex III Qo site inhibitor) all decrease ATP production, but only antimycin A causes significant mitochondrial superoxide production (Stowe and Camara, 2009) and only antimycin A evoked significant nociceptor hyperexcitability in this study. Furthermore, myxothiazol, which prevents antimycin A–induced ROS production, prevented antimycin A–induced hyperexcitability, indicating that the hyperexcitability was specifically due to ROS produced from the complex III Qo site. It should be noted that we have not measured ROS or ATP production from the nerve terminals in this study. The potency of oligomycin, antimycin A, and myxothiazol used in their single concentrations on ROS or ATP production cannot be confirmed, and, thus, caution must be employed in drawing conclusions. Nevertheless, the concentrations were chosen from previous studies showing the selective effects of inhibition of ATP synthase (oligomycin), complex III Qi site (antimycin A), and complex III Qo site (myxothiazol) on ROS production, ATP production, and oxygen consumption (Turrens et al., 1985; Sipos et al., 2003; Gyulkhandanyan and Pennefather, 2004; Hool and Corry, 2007; Tretter et al., 2007; Stowe and Camara, 2009). The role of ROS in this pathway was confirmed by the inhibition of antimycin A–induced hyperexcitability by the antioxidant NAC (detoxifies ROS) and the reducing agent DTT (reverses sulfhydryl group oxidation). NAC is actively transported across the plasma membrane unlike the antioxidant GSH (Raftos et al., 2007). GSH (only detoxifies extracellular ROS) failed to inhibit antimycin A–induced hyperexcitability, indicating that intracellular (i.e., intraneuronal) ROS contributed significantly to this phenomenon. It is not known whether superoxide or other downstream ROS (e.g., H2O2) are the end effectors of mitochondrial modulation–induced hyperexcitability, but the inhibition by DTT would argue against a significant role of lipid peroxidation products, which react with proteins in a DTT-insensitive manner (Andersson et al., 2008).

Antimycin A and H2O2 caused nociceptor hyperexcitability to α,β-mATP that was sensitive to the selective PKC inhibitor BIM I but not to its inactive analog BIM V. Selectivity is a relative term for kinase inhibitors, but at 1 μM BIM I inhibits PKC without affecting protein kinase A; and the other off-target effects of bisindolylmaleimide compounds are shared by both BIM I and BIM V (Toullec et al., 1991; Davis et al., 1992). That PKC activation could lead to nociceptor hyperexcitability was further demonstrated by PMA, an activator of PKC, which produced hyperexcitability of a similar magnitude to antimycin A and H2O2. PMA and the activation of PKC have been shown previously to induce hyperexcitability in vagal nociceptive neurons (Kagaya et al., 2002; Ikeda et al., 2005; Gu and Lee, 2006, 2009; Matsumoto et al., 2007). PKC translocation from the cytosol to the plasma membrane is an indicator of PKC activation (Cosentino-Gomes et al., 2012). Our electrophysiological studies were supported by confocal imaging in dissociated neurons, indicating that antimycin A and H2O2 both induced PKC translocation. Antimycin A–induced PKC translocation was prevented by a combination of tempol (superoxide dismutase mimetic) and MnTMPyP (combined superoxide dismutase and catalase mimetic), indicating that ROS were required for this effect.

There are 15 PKC isoforms, which are divided into three groups based upon sensitivity to Ca2+ and DAG, as follows: classic isoforms (α, β, and γ) require both Ca2+ and DAG for activation; novel isoforms (δ, ε, η, and θ) require DAG but not Ca2+ for activation; and atypical isoforms (ι/λ and ζ) require neither Ca2+ nor DAG for activation. To date there has not been any characterization of PKC isoform expression in vagal sensory neurons, although PKC expression in somatosensory DRG neurons indicates that, at a minimum, β1, β2, δ, ε, and ζ are expressed in complex combinatorial patterns (Cesare et al., 1999; Wu et al., 2012). Similar to our findings, PKC has repeatedly been shown to play a role in inflammation-induced hyperexcitability in nociceptive DRG (Cesare et al., 1999; Baker, 2005; Wu et al., 2012; Zhang et al., 2012). It is known that PMA binds to the same enzyme subsite as DAG, and thus PMA activates only classic and novel PKC isoforms, suggesting that PKC expression is not limited to atypical isoforms in vagal sensory nerves. Although ROS activate PKC by the oxidation of cysteines in a pair of zinc fingers associated with the DAG binding site (Gopalakrishna and Anderson, 1989; Knapp and Klann, 2000), this does not preclude the contribution of atypical isoforms that are activated by the same mechanism on their single zinc finger site (that does not bind DAG) (Cosentino-Gomes et al., 2012). Further studies are required to elucidate which PKC isoform is involved in mitochondrial ROS-induced bronchopulmonary C-fiber hyperexcitability.

Another question raised by these studies is how activation of PKC causes hyperexcitability in vagal bronchopulmonary C-fibers. Activated PKC isoforms are capable of modifying neuronal excitability via the phosphorylation of numerous cellular targets, including ion channels and transporters. In particular, PKC has been shown to induce hyperexcitability via tetrodotoxin-resistant voltage-gated Na+ channels in both vagal (Kagaya et al., 2002; Ikeda et al., 2005; Matsumoto et al., 2007) and DRG neurons (Baker, 2005; Hayase et al., 2007; Wu et al., 2012), indicating possible candidates for mitochondrial ROS-induced bronchopulmonary C-fiber hyperexcitability. However, given that tetrodotoxin-resistant voltage-gated Na+ channels are expressed in both nociceptive and non-nociceptive neurons (Kwong et al., 2008), further work is needed to elucidate the mechanism highlighted in the present study and its apparent exclusivity to nociceptive C-fibers.

In summary, we have found that antimycin A causes a robust increase in bronchopulmonary C-fiber excitability via the actions of mitochondrial ROS and PKC. Mitochondrial ROS are known to increase excitability in CNS neurons (Nozoe et al., 2008; Yin et al., 2010), to activate vagal nociceptive neurons (Nesuashvili et al., 2013), and to regulate Ca2+ signaling in DRG nociceptive neurons (Kim and Usachev, 2009). Airway inflammatory disease is associated with mitochondrial dysfunction and mitochondrial ROS production (Mabalirajan et al., 2008; Aguilera-Aguirre et al., 2009; Reddy, 2011), and a wide variety of inflammatory signaling has been shown to increase mitochondrial ROS (Corda et al., 2001; Bell et al., 2007; Pehar et al., 2007; Michaeloudes et al., 2011; West et al., 2011). Given the mitochondrial density within sensory nerve terminals, it is possible that inflammation may induce nociceptor hyperexcitability via the production of mitochondrial ROS. It is now increasingly clear that, in addition to initiating apoptosis, mitochondrial dysfunction and ROS production also contribute to more subtle physiologic signaling (Sena and Chandel, 2012), and it remains to be determined whether chronic mitochondrial ROS in sensory nerve terminals contribute to hyper-reflexia in airway disease.

Abbreviations

AITC

allyl isothiocyanate

BIM

bisindolylmaleimide

DAG

diacylglycerol

DRG

dorsal root ganglion

DTT

dithiothreitol

GSH

glutathione

α,β-mATP

α,β-methylene ATP

MnTMPyP

manganese(III) tetrakis(1-methyl-4-pyridyl)porphyrin

NAC

N-acetylcysteine

PBST

phosphate-buffered saline with 0.0025% Tween 20

PKC

protein kinase C

PMA

phorbol-12-myristate-13-acetate

ROS

reactive oxygen species

TRP

transient receptor potential

TRPA1

TRP ankyrin 1

TRPV1

TRP vanilloid 1

Authorship Contributions

Participated in research design: Bahia, Taylor-Clark.

Conducted experiments: Hadley, Bahia.

Performed data analysis: Hadley, Bahia, Taylor-Clark.

Wrote or contributed to the writing of the manuscript: Bahia, Taylor-Clark.

Footnotes

This work was supported by the National Institutes of Health National Heart, Lung, and Blood Institute [Grants R00-HL088520 and R01-HL119802].

References

  1. Aguilera-Aguirre L, Bacsi A, Saavedra-Molina A, Kurosky A, Sur S, Boldogh I. (2009) Mitochondrial dysfunction increases allergic airway inflammation. J Immunol 183:5379–5387 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Amadesi S, Nie J, Vergnolle N, Cottrell GS, Grady EF, Trevisani M, Manni C, Geppetti P, McRoberts JA, Ennes H, et al. (2004) Protease-activated receptor 2 sensitizes the capsaicin receptor transient receptor potential vanilloid receptor 1 to induce hyperalgesia. J Neurosci 24:4300–4312 [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Andersson DA, Gentry C, Moss S, Bevan S. (2008) Transient receptor potential A1 is a sensory receptor for multiple products of oxidative stress. J Neurosci 28:2485–2494 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Baker MD. (2005) Protein kinase C mediates up-regulation of tetrodotoxin-resistant, persistent Na+ current in rat and mouse sensory neurones. J Physiol 567:851–867 [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bell EL, Klimova TA, Eisenbart J, Moraes CT, Murphy MP, Budinger GR, Chandel NS. (2007) The Qo site of the mitochondrial complex III is required for the transduction of hypoxic signaling via reactive oxygen species production. J Cell Biol 177:1029–1036 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Carr MJ, Lee LY. (2006) Plasticity of peripheral mechanisms of cough. Respir Physiol Neurobiol 152:298–311. [DOI] [PubMed] [Google Scholar]
  7. Carr MJ, Undem BJ. (2003) Bronchopulmonary afferent nerves. Respirology 8:291–301 [DOI] [PubMed] [Google Scholar]
  8. Cesare P, Dekker LV, Sardini A, Parker PJ, McNaughton PA. (1999) Specific involvement of PKC-epsilon in sensitization of the neuronal response to painful heat. Neuron 23:617–624 [DOI] [PubMed] [Google Scholar]
  9. Corda S, Laplace C, Vicaut E, Duranteau J. (2001) Rapid reactive oxygen species production by mitochondria in endothelial cells exposed to tumor necrosis factor-alpha is mediated by ceramide. Am J Respir Cell Mol Biol 24:762–768 [DOI] [PubMed] [Google Scholar]
  10. Cosentino-Gomes D, Rocco-Machado N, Meyer-Fernandes JR. (2012) Cell signaling through protein kinase C oxidation and activation. Int J Mol Sci 13:10697–10721 [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Davis PD, Hill CH, Lawton G, Nixon JS, Wilkinson SE, Hurst SA, Keech E, Turner SE. (1992) Inhibitors of protein kinase C. 1. 2,3-bisarylmaleimides. J Med Chem 35:177–184 [DOI] [PubMed] [Google Scholar]
  12. Gopalakrishna R, Anderson WB. (1989) Ca2+- and phospholipid-independent activation of protein kinase C by selective oxidative modification of the regulatory domain. Proc Natl Acad Sci USA 86:6758–6762 [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Gu Q, Lee LY. (2006) Hypersensitivity of pulmonary chemosensitive neurons induced by activation of protease-activated receptor-2 in rats. J Physiol 574:867–876 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Gu Q, Lee LY. (2009) Effect of protease-activated receptor 2 activation on single TRPV1 channel activities in rat vagal pulmonary sensory neurons. Exp Physiol 94:928–936 [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Gyulkhandanyan AV, Pennefather PS. (2004) Shift in the localization of sites of hydrogen peroxide production in brain mitochondria by mitochondrial stress. J Neurochem 90:405–421 [DOI] [PubMed] [Google Scholar]
  16. Hayase F, Matsuura H, Sanada M, Kitada-Hamada K, Omatsu-Kanbe M, Maeda K, Kashiwagi A, Yasuda H. (2007) Inhibitory action of protein kinase Cbeta inhibitor on tetrodotoxin-resistant Na+ current in small dorsal root ganglion neurons in diabetic rats. Neurosci Lett 417:90–94 [DOI] [PubMed] [Google Scholar]
  17. Hongpaisan J, Winters CA, Andrews SB. (2004) Strong calcium entry activates mitochondrial superoxide generation, upregulating kinase signaling in hippocampal neurons. J Neurosci 24:10878–10887 [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Hool LC, Corry B. (2007) Redox control of calcium channels: from mechanisms to therapeutic opportunities. Antioxid Redox Signal 9:409–435 [DOI] [PubMed] [Google Scholar]
  19. Hung KS, Hertweck MS, Hardy JD, Loosli CG. (1973) Ultrastructure of nerves and associated cells in bronchiolar epithelium of the mouse lung. J Ultrastruct Res 43:426–437 [DOI] [PubMed] [Google Scholar]
  20. Ikeda M, Yoshida S, Kadoi J, Nakano Y, Mastumoto S. (2005) The effect of PKC activity on the TTX-R sodium currents from rat nodose ganglion neurons. Life Sci 78:47–53 [DOI] [PubMed] [Google Scholar]
  21. Jesenak M, Babusikova E, Petrikova M, Turcan T, Rennerova Z, Michnova Z, Havlicekova Z, Villa MP, Banovcin P. (2009) Cough reflex sensitivity in various phenotypes of childhood asthma. J Physiol Pharmacol 60 (Suppl 5):61–65 [PubMed] [Google Scholar]
  22. Kagaya M, Lamb J, Robbins J, Page CP, Spina D. (2002) Characterization of the anandamide induced depolarization of guinea-pig isolated vagus nerve. Br J Pharmacol 137:39–48 [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Kim MS, Usachev YM. (2009) Mitochondrial Ca2+ cycling facilitates activation of the transcription factor NFAT in sensory neurons. J Neurosci 29:12101–12114 [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Knapp LT, Klann E. (2000) Superoxide-induced stimulation of protein kinase C via thiol modification and modulation of zinc content. J Biol Chem 275:24136–24145 [DOI] [PubMed] [Google Scholar]
  25. Kollarik M, Dinh QT, Fischer A, Undem BJ. (2003) Capsaicin-sensitive and -insensitive vagal bronchopulmonary C-fibres in the mouse. J Physiol 551:869–879 [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Kwong K, Carr MJ, Gibbard A, Savage TJ, Singh K, Jing J, Meeker S, Undem BJ. (2008) Voltage-gated sodium channels in nociceptive versus non-nociceptive nodose vagal sensory neurons innervating guinea pig lungs. J Physiol 586:1321–1336 [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Lin YJ, Hsu HH, Ruan T, Kou YR. (2013) Mediator mechanisms involved in TRPV1, TRPA1 and P2X receptor-mediated sensory transduction of pulmonary ROS by vagal lung C-fibers in rats. Respir Physiol Neurobiol 189:1–9 [DOI] [PubMed] [Google Scholar]
  28. Mabalirajan U, Dinda AK, Kumar S, Roshan R, Gupta P, Sharma SK, Ghosh B. (2008) Mitochondrial structural changes and dysfunction are associated with experimental allergic asthma. J Immunol 181:3540–3548 [DOI] [PubMed] [Google Scholar]
  29. Matsumoto S, Yoshida S, Ikeda M, Tanimoto T, Saiki C, Takeda M, Shima Y, Ohta H. (2007) Effect of 8-bromo-cAMP on the tetrodotoxin-resistant sodium (Nav 1.8) current in small-diameter nodose ganglion neurons. Neuropharmacology 52:904–924 [DOI] [PubMed] [Google Scholar]
  30. Meyer RA, Raja SN, and Cambell JN (1996) Neural mechanisms of primary hyperalgesia, in Neurobiology of Nociceptors (Belmonte C and Cervero F eds) pp 370–389, Oxford University Press, Oxford. [Google Scholar]
  31. Michaeloudes C, Sukkar MB, Khorasani NM, Bhavsar PK, Chung KF. (2011) TGF-β regulates Nox4, MnSOD and catalase expression, and IL-6 release in airway smooth muscle cells. Am J Physiol Lung Cell Mol Physiol 300:L295–L304. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Morice AH. (2010) The cough hypersensitivity syndrome: a novel paradigm for understanding cough. Lung 188 (Suppl 1):S87–S90 [DOI] [PubMed] [Google Scholar]
  33. Nassenstein C, Kammertoens T, Veres TZ, Uckert W, Spies E, Fuchs B, Krug N, Braun A. (2007) Neuroimmune crosstalk in asthma: dual role of the neurotrophin receptor p75NTR. J Allergy Clin Immunol 120:1089–1096 [DOI] [PubMed] [Google Scholar]
  34. Nassenstein C, Kwong K, Taylor-Clark T, Kollarik M, Macglashan DM, Braun A, Undem BJ. (2008) Expression and function of the ion channel TRPA1 in vagal afferent nerves innervating mouse lungs. J Physiol 586:1595–1604 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Nassenstein C, Taylor-Clark TE, Myers AC, Ru F, Nandigama R, Bettner W, Undem BJ. (2010) Phenotypic distinctions between neural crest and placodal derived vagal C-fibres in mouse lungs. J Physiol 588:4769–4783 [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Nesuashvili L, Hadley SH, Bahia PK, Taylor-Clark TE. (2013) Sensory nerve terminal mitochondrial dysfunction activates airway sensory nerves via transient receptor potential (TRP) channels. Mol Pharmacol 83:1007–1019 [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Nozoe M, Hirooka Y, Koga Y, Araki S, Konno S, Kishi T, Ide T, Sunagawa K. (2008) Mitochondria-derived reactive oxygen species mediate sympathoexcitation induced by angiotensin II in the rostral ventrolateral medulla. J Hypertens 26:2176–2184 [DOI] [PubMed] [Google Scholar]
  38. Pehar M, Vargas MR, Robinson KM, Cassina P, Díaz-Amarilla PJ, Hagen TM, Radi R, Barbeito L, Beckman JS. (2007) Mitochondrial superoxide production and nuclear factor erythroid 2-related factor 2 activation in p75 neurotrophin receptor-induced motor neuron apoptosis. J Neurosci 27:7777–7785 [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Raftos JE, Whillier S, Chapman BE, Kuchel PW. (2007) Kinetics of uptake and deacetylation of N-acetylcysteine by human erythrocytes. Int J Biochem Cell Biol 39:1698–1706 [DOI] [PubMed] [Google Scholar]
  40. Reddy PH. (2011) Mitochondrial dysfunction and oxidative stress in asthma: implications for mitochondria-targeted antioxidant therapeutics. Pharmaceuticals 4:429–456 [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Riccio MM, Myers AC, Undem BJ. (1996) Immunomodulation of afferent neurons in guinea-pig isolated airway. J Physiol 491:499–509 [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Sabogal C, Auais A, Napchan G, Mager E, Zhou BG, Suguihara C, Bancalari E, Piedimonte G. (2005) Effect of respiratory syncytial virus on apnea in weanling rats. Pediatr Res 57:819–825 [DOI] [PubMed] [Google Scholar]
  43. Sena LA, Chandel NS. (2012) Physiological roles of mitochondrial reactive oxygen species. Mol Cell 48:158–167 [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Sipos I, Tretter L, Adam-Vizi V. (2003) Quantitative relationship between inhibition of respiratory complexes and formation of reactive oxygen species in isolated nerve terminals. J Neurochem 84:112–118 [DOI] [PubMed] [Google Scholar]
  45. Stowe DF, Camara AK. (2009) Mitochondrial reactive oxygen species production in excitable cells: modulators of mitochondrial and cell function. Antioxid Redox Signal 11:1373–1414 [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Takahashi N, Kuwaki T, Kiyonaka S, Numata T, Kozai D, Mizuno Y, Yamamoto S, Naito S, Knevels E, Carmeliet P, et al. (2011) TRPA1 underlies a sensing mechanism for O2. Nat Chem Biol 7:701–711 [DOI] [PubMed] [Google Scholar]
  47. Thannickal VJ, Fanburg BL. (2000) Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol 279:L1005–L1028 [DOI] [PubMed] [Google Scholar]
  48. Toullec D, Pianetti P, Coste H, Bellevergue P, Grand-Perret T, Ajakane M, Baudet V, Boissin P, Boursier E, Loriolle F, et al. (1991) The bisindolylmaleimide GF 109203X is a potent and selective inhibitor of protein kinase C. J Biol Chem 266:15771–15781 [PubMed] [Google Scholar]
  49. Tretter L, Takacs K, Hegedus V, Adam-Vizi V. (2007) Characteristics of alpha-glycerophosphate-evoked H2O2 generation in brain mitochondria. J Neurochem 100:650–663 [DOI] [PubMed] [Google Scholar]
  50. Turrens JF, Alexandre A, Lehninger AL. (1985) Ubisemiquinone is the electron donor for superoxide formation by complex III of heart mitochondria. Arch Biochem Biophys 237:408–414 [DOI] [PubMed] [Google Scholar]
  51. von Düring M, Andres KH. (1988) Structure and functional anatomy of visceroreceptors in the mammalian respiratory system. Prog Brain Res 74:139–154 [DOI] [PubMed] [Google Scholar]
  52. Wahl P, Foged C, Tullin S, Thomsen C. (2001) Iodo-resiniferatoxin, a new potent vanilloid receptor antagonist. Mol Pharmacol 59:9–15 [DOI] [PubMed] [Google Scholar]
  53. Weigand LA, Ford AP, Undem BJ. (2012) A role for ATP in bronchoconstriction-induced activation of guinea pig vagal intrapulmonary C-fibres. J Physiol 590:4109–4120 [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. West AP, Brodsky IE, Rahner C, Woo DK, Erdjument-Bromage H, Tempst P, Walsh MC, Choi Y, Shadel GS, Ghosh S. (2011) TLR signalling augments macrophage bactericidal activity through mitochondrial ROS. Nature 472:476–480 [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Wu DF, Chandra D, McMahon T, Wang D, Dadgar J, Kharazia VN, Liang YJ, Waxman SG, Dib-Hajj SD, Messing RO. (2012) PKCε phosphorylation of the sodium channel NaV1.8 increases channel function and produces mechanical hyperalgesia in mice. J Clin Invest 122:1306–1315 [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Yin JX, Yang RF, Li S, Renshaw AO, Li YL, Schultz HD, Zimmerman MC. (2010) Mitochondria-produced superoxide mediates angiotensin II-induced inhibition of neuronal potassium current. Am J Physiol Cell Physiol 298:C857–C865 [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Zhang YH, Kays J, Hodgdon KE, Sacktor TC, Nicol GD. (2012) Nerve growth factor enhances the excitability of rat sensory neurons through activation of the atypical protein kinase C isoform, PKMζ. J Neurophysiol 107:315–335 [DOI] [PMC free article] [PubMed] [Google Scholar]

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