Acid-sensing ion channels modulate bladder nociception

Abstract

Sensitization of neuronal pathways and persistent afferent drive are major contributors to somatic and visceral pain. However, the underlying mechanisms that govern whether afferent signaling will give rise to sensitization and pain are not fully understood. In the present report, we investigated the contribution of acid-sensing ion channels (ASICs) to bladder nociception in a model of chemical cystitis induced by cyclophosphamide (CYP). We found that the administration of CYP to mice lacking ASIC3, a subunit primarily expressed in sensory neurons, generates pelvic allodynia at a time point at which only modest changes in pelvic sensitivity are apparent in wild-type mice. The differences in mechanical pelvic sensitivity between wild-type and Asic3 knockout mice treated with CYP were ascribed to sensitized bladder C nociceptors. Deletion of Asic3 from bladder sensory neurons abolished their ability to discharge action potentials in response to extracellular acidification. Collectively, the results of our study support the notion that protons and their cognate ASIC receptors are part of a mechanism that operates at the nerve terminals to control nociceptor excitability and sensitization.

NEW & NOTEWORTHY Our study indicates that protons and their cognate acid-sensing ion channel receptors are part of a mechanism that operates at bladder afferent terminals to control their function and that the loss of this regulatory mechanism results in hyperactivation of nociceptive pathways and the development of pain in the setting of chemical-induced cystitis.

INTRODUCTION

Normal bladder function and responses to bladder inflammation and injury depend on sensory inputs from primary afferent fibers located in the urothelium as well as in the subjacent lamina propria and bladder musculature (1–4). These fibers express a myriad of ligand- and mechano-gated ion channels and receptors that convert chemical and mechanical stimuli into electrical signals (2, 5, 6). How these signals are processed and integrated in the afferents that convey information about bladder filling and noxious events to the central nervous system remains uncertain.

Acid-sensing ion channels (ASICs) are voltage-insensitive cation-permeable channels expressed in neurons that respond to extracellular acidification (7). Four genes (Accn1, Accn2, Accn3, and Accn4) that encode for six ASIC subunits and splice variants have been identified in rodents (ASIC1a, ASIC1b, ASIC2a, ASIC2b, ASIC3, and ASIC4) (8–17). ASIC subunits associate to form homo- and heterotrimers with biophysical properties that vary with respect to agonist affinity, single-channel conductance, rate of desensitization, rate of recovery from desensitization, and cation selectivity (11, 16, 18–21). ASIC1a, ASIC2a, ASIC2b, and ASIC4 are expressed throughout the central nervous system and peripheral nervous system, whereas ASIC1b and ASIC3 are primarily expressed in sensory neurons (9, 11, 12, 21–23). Although in sensory and most central nervous system neurons proton-gated currents are mediated by ASIC heteromers (7, 18, 24–27), loss of specific subunits alters a subset of sensory modalities (mechanosensation and nociception) at the periphery (7, 28–33) and learning and memory (34). A number of studies have reported an increase in sensory responses at single-fiber and whole animal levels in ASIC subunit-null mice (7, 28, 32, 33, 35–40). In this regard, we recently showed that the intravesical pressure required to trigger voiding is lower and that the voided volume is smaller in global Asic3-null mice than in wild-type (WT) littermates (7). These findings raise the question of how missing a ligand-gated channel expressed at nerve terminals can increase sensory signaling.

The urinary bladder is one of the major sources of visceral pain. The consensus is that normal micturition depends on mechanosensitive Aδ fiber afferents that respond to bladder distension in the physiological range (2, 3) and that C fibers have high thresholds and respond to bladder distension only at elevated pressure (41–43). However, a population of C fibers that respond to bladder distension in the physiological range of pressures has also been reported (44–48). Visceral pain is the result of sensitization of neuronal pathways (i.e., primary sensory neurons and spinal ascending neurons) involved in the transmission of information from internal organs to the supraspinal level. Alternatively (or in addition), it can arise from the dysregulation of descending pathways that modulate spinal nociceptive transmission (49–51). Compelling evidence indicates that in models of cystitis induced by cyclophosphamide (CYP), bladder pain is driven, at least in part, by sensitized C afferent fibers (52–54). Here, we investigated the role of ASICs in bladder nociception in chemical-induced cystitis. Our results revealed that ASICs play a central role in bladder nociception by regulating the excitability and sensitization of C nociceptors.

MATERIALS AND METHODS

Reagents

All chemicals were purchased from Sigma-Aldrich (St. Louis, MO) unless otherwise specified.

Animals

All experimental procedures were approved by the Institutional Animal Care and Use Committee of the University of Pittsburgh. Mice were randomly assigned to control and treated groups. Histological and physiological analyses were performed with the researcher blinded to the genotype and treatment of mice. Asic3 knockout (KO) (Asic3^tm1Wsh/J) mice were obtained from Jackson Labs (Bar Harbor, ME). Age-matched C57BL/6J (WT) littermates were used as control animals. Mice were bred and housed at the University of Pittsburgh under 12:12-h light-dark cycles with free access to food and water. Physiological experiments were conducted with mice between the ages of 12 and 24 wk. Animals were euthanized by CO₂ inhalation followed by a thoracotomy.

Genotyping

Tissue harvested from an ear punch or a tail snip was placed in a 1.5-mL tube and resuspended in 200 µL of Donghuis’s buffer [50 mM Tris base (pH 8.0), 50 mM KCl, 2.5 mM EDTA, 0.45% IGEPAL CA-630 (Nonidet P-40), and 0.45% Tween 20] containing 1.6 units of proteinase K and incubated overnight at 55°C. Thereafter, the tube was incubated at 94°C for 10 min and centrifuged at 21,000 g for 10 min, and the supernatant was saved for quantitative PCR analysis. The following primers were used for the quantification of DNA copies by quantitative PCR: Asic3 exon 1, forward 5′-CAGCTGTACTCCTGTCGCTG-3′ and reverse 5′-GGCGCAGGGGATTGATGTTA-3′; and β-actin, forward 5′-GGCTGTATTCCCCTCCATCG-3′ and reverse 5′-CCAGTTGGTAACAATGCCATGT-3′. Quantitative PCR amplification reactions were run in duplicate as previously described (7). Gene expression was determined by the ΔC_T method (where C_T is threshold cycle) with β-actin as the reference gene.

CYP-Induced Cystitis

CYP solution (16 mg/mL) was prepared in saline (0.9% NaCl) and sterilized by filtration through a 0.45-µm PVDF filter. Mice were injected intraperitoneally with the CYP solution (80 mg/kg) every other day for up to a week as previously described (55). Control animals were injected with saline. Experimental procedures were conducted a day after the last dose of saline or CYP.

Tissue Edema Quantification

To assess bladder edema, urinary bladders were harvested, carefully dried with filter paper to eliminate any fluids, and weighed to obtain the wet mass (WM). To determine the dry mass (DM), samples were placed in an incubator at 55°C until constant weight was reached (typically after 5 days). The percentage of water tissue content was calculated with the following equation:

$$\mathrm{\%}\text{ Water content}=\frac{\mathrm{WM}-\mathrm{DM}}{\mathrm{WM}}\times 100$$

Assessment of Mechanical Sensitivity in the Pelvic Region

Mice treated with saline or CYP were placed in modular cages (Bioseb, Pinellas Park, FL) on an elevated wire mesh platform. Mice were acclimatized for at least 1 h before the test. Withdrawal thresholds to von Frey filaments (Touch Test Sensory Evaluators, North Coast Medical, Gilroy, CA) applied to the pelvic area were estimated with the up-down method described by Chaplan and colleagues (56). von Frey filaments were applied on the lower abdominal area close to the urinary bladder for 1–3 s, with intervals between stimuli of 15 s. Testing in the pelvic area was initiated with a von Frey filament with a calibrated force of 0.16 g. Abdominal withdrawal (either contraction of the abdominal musculature or postural retraction of the abdomen) and licking or scratching in the pelvic area in response to von Frey filament application were considered a positive response. When a negative response was observed, the next stronger filament was applied. When a positive response was observed, the next weaker stimulus was applied. After the response threshold was first crossed, three additional filaments were applied that varied sequentially up or down based on the animal response. The resulting pattern of positive and negative responses was tabulated, and the 50% response threshold was calculated with the following equation:

$$\text{50\% Withdrawal threshold }(\mathrm{g})=\frac{{10}^{(X_{\mathrm{f}}+k\mathrm{\delta })}}{10,000}$$

where X_f is the value of the final positive von Frey filament used, k is the tabular value for the pattern of positive/negative responses (56), and δ is the mean difference (in log units) between stimuli.

Histopathology and Image Analysis

Urinary bladders from mice treated with saline or CYP were harvested from euthanized animals through an abdominal incision. The tissue was then fixed in neutral buffered formalin [4% (wt/vol) paraformaldehyde, 45.8 mM Na₂HPO₄, and 30 mM NaH₂PO₄·H₂O] for 30 min at 37°C, and each animal’s bladder was then cut into five to seven ∼2-mm-wide strips. Tissue was maintained in neutral buffered formalin for at least 24 h before being embedded in paraffin. Approximately five-micrometer paraffin sections were cut, mounted on slides, and then stained with hematoxylin and eosin. Images were captured with a Leica DM6000B upright microscope (fitted with a ×20 HC PL-APO, 0.80 numerical aperture objective) outfitted with a Gryphax Prokyon digital camera (Jenoptik, Jupiter, FL) and interfaced to an Apple iMac computer running Gryphax software (Jenoptik). The images were exported in TIFF format and compiled in Adobe Illustrator 2020.

Video-Monitored Void Spot Assay

To evaluate micturition in awake, freely moving mice, we used a modified version of the standard void spot assay that incorporates video monitoring, akin to the setup described in Ref. 57. The setup consists of an upper compartment, which houses two mouse cages side by side, each made of acrylic sheets with an ultraviolet-transmitting acrylic bottom (dimensions of 37 × 25 × 20 cm), and a lower compartment, which houses ultraviolet tube lights (ADJ; model T8-F20BLB24, 24 in.). Mouse cages were furnished with the following: an igloo-shaped sleeping chamber, an Eppendorf tube as a “play toy,” and a dish with standard mouse chow and water in the form of HydroGel (ClearH₂O, Westbrook, ME). The bottom of the cages was covered with Cosmos blotting paper (catalog no. 10422-1005, Blick Art Materials, Galesburg, IL). The lower compartments have reflective mirrored walls, which evenly illuminate the chromatography paper from below. Each mouse was monitored by wide-angle cameras (Logitech C930e), one camera positioned above the cage and another camera mounted at the base of the lower compartment. Mice were routinely housed in a facility with 12:12-h light-dark cycles, with 7:00 AM being zeitgeber time (ZT) = 0 (start of the light cycle). Mice were introduced into the cages between ZT = 10 and ZT = 11, and micturition behavior was evaluated during the dark cycle for a 6-h period from ZT = 12 to ZT = 18. The streamed video was captured with an Apple iMac computer running SecuritySpy (Ben Software) at 1 frame/s with a 1,920 × 1,080-pixel resolution. Voiding events were identified by visual inspection of the movies. The void spot area was computed from video images with ImageJ Fiji, as we have previously described (7). Note that the voiding spots become invisible over time. Because of the large number of voiding events observed in mice treated with CYP, the mean voided volume was computed for 10 random voiding events. The parameters evaluated in this study were volume per void (in μL) and number of voids during the 6-h period. Void spot volume was estimated from a calibration curve generated with known amounts of urine.

Retrograde Labeling of Bladder Sensory Neurons

Bladder afferent neurons were labeled with the fluorescent retrograde axonal tracer 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiI, Invitrogen, Waltham, MA), as previously described (7, 58–60). Briefly, mice were anesthetized with isoflurane, and the bladder was exposed through an abdominal incision (∼1 cm in length). DiI [5% (wt/vol) in DMSO] was injected at three or four sites (total volume: 12–15 µL) in the bladder wall. The muscle layer and skin incision were individually closed with 5.0 PDO absorbable monofilament surgical suture (AD Surgical, Sunnyvale, CA). Postoperative analgesia was provided by subcutaneous administration of ketoprofen (5 mg/kg; Zoetis, Parsippany-Troy Hills, NJ). Ampicillin (10 mg/kg; Boehringer Ingelheim Vetmedica, Duluth, GA) was administered to prevent infections.

Isolation of Bladder Sensory Neurons

Seven to ten days after the surgical procedure, dorsal root ganglia (L₆–S₂) from two or three mice were collected and placed in a cell culture dish containing Neurobasal-A medium (Invitrogen). Dorsal root ganglia were cut into small fragments and then incubated in a cell culture flask containing 5 mL of Neurobasal-A medium supplemented with 10 mg of collagenase type 4 (Worthington Biochemical, Lakewood, NJ) and 5 mg of trypsin (Worthington Biochemical) for 30 min at 37°C. Tissue fragments were gently triturated with a fire-polished glass pipette, and the cell suspension was centrifuged at 460 g for 5 min. The pellet, containing the somas of sensory neurons, was resuspended in complete neuro media (Neurobasal-A media supplemented with 5% B27 supplement, 0.5 mM l-glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin). The centrifugation and resuspension steps were repeated three times. Finally, the pellet was resuspended in 1.5 mL of complete neuro media, and the suspension was plated on coverslips coated with ornithine (Sigma) and laminin (Invitrogen) inside a six-well tissue culture plate. After 2 h at 37°C, warm complete neuro media were added to the wells. For long-term experiments, the complete neuro media were supplemented with 100 ng/mL of nerve growth factor (Invitrogen).

Patch-Clamp Experiments

Current- and voltage-clamp experiments were performed between 2 and 8 h (acute) or between 24 and 72 h after plating, respectively. Whole cell patch-clamp recordings were performed with the amphotericin B perforated patch technique. Glass coverslips containing DiI-labeled dorsal root ganglion neurons were transferred to a chamber mounted on the stage of a Nikon Ti inverted microscope equipped with a Sedat Quad set (Chroma Technology, Brattleboro, VT), a PhotoFluor II metal halide light source (89 North, Burlington, VT), a Lambda 10-3 filter-wheel system (Sutter Instrument, Novato, CA), and an ORCA-Flash 2.8 camera (Hamamatsu, Bridgewater Township, NJ). The Sedat Quad set contains a combination of 10 excitation, emission, and band-pass filters that permit the detection of fluorescence emission from different fluorophores including DiI. To identify DiI-labeled neurons, we used an excitation filter (ET555/25x, Chroma Technology). Current- and voltage-clamp recordings were carried out at room temperature with a PC-505B patch-clamp amplifier (Warner Instruments, Hamden, CT), and data were captured with a Digidata 1440 A acquisition system using pCLAMP 10 (Molecular Devices, San Jose, CA). Signals were low-pass filtered at 1 kHz (4-pole Bessel filter) and digitized at 5 kHz. Micropipettes were pulled from borosilicate glass capillary tubes (Warner Instruments) with a PP-830 puller (Narishige, Amityville, NY). Fire-polished micropipettes with a tip resistance of 1.5–3 mΩ were used for patch-clamp recordings. Pipette filling solution contained (in mM) 145 KCl, 1 MgCl₂, 0.1 CaCl₂, 1 EGTA, and 10 HEPES (pH 7.2). Amphotericin B was added to the pipette solution to a final concentration of 120 µg/mL. The extracellular bath solution contained (in mM) 135 NaCl, 5 KCl, 1 MgCl₂, 2.5 CaCl₂, 10 glucose, and 10 HEPES (pH 7.4 or 8) or MES (pH 6.5). After a whole cell configuration was established in voltage-clamp mode, the membrane potential was clamped at −60 mV and the cell capacitance was obtained from the software. A gravity-fed perfusion system (AutoMate Scientific, Berkeley, CA) and a perfusion pencil positioned near the cell were used for rapid fluid delivery.

Voltage and Current Clamp

ASIC currents were evoked at a membrane potential of −60 mV by a drop in extracellular pH from 8.0 to 6.5, unless otherwise noted. The time constant of desensitization (τ) was estimated by fitting the current decay to a single-exponential function. For current-clamp experiments, only cells that had a resting membrane potential more negative than −40 mV and generated action potentials with a distinct overshoot higher than 0 mV in response to depolarizing current injections were studied. To determine the passive and active membrane properties, action potentials were evoked by injecting a series of 4-ms rectangular depolarizing current pulses of increasing intensity. The following passive and active membrane properties of bladder sensory neurons were computed: membrane capacitance, resting membrane potential, action potential rheobase, action potential threshold, action potential amplitude, action potential duration at 0 mV, and magnitude of hyperpolarization below the resting membrane potential (7, 61). The action potential rheobase and threshold are defined as the minimal depolarizing current injection necessary to evoke an action potential and the maximal membrane potential depolarization obtained in the absence of an action potential, respectively (7, 61). Tetrodotoxin (TTX; 1 µM) was used to discriminate sensory neurons of C and Aδ origin. Electrophysiological data were analyzed with Clampfit (Molecular Devices).

Statistical Analysis

Data are expressed as means ± SE (n), where n equals the number of independent experiments analyzed. Parametric or nonparametric tests were used as appropriate. P values of <0.05 were considered statistically significant. Statistical comparisons were performed with GraphPad 8 (GraphPad Software, San Diego, CA).

RESULTS

Sensitized Afferents Contribute to Referred Pelvic Allodynia in Mice With Chemically Induced Cystitis

CYP is an alkylating agent used to treat an array of cancers and as an immune suppressor in nephrotic syndrome and granulomatosis with polyangiitis and after organ transplantation (62). Cystitis, which ensues as the CYP metabolite acrolein accumulates in the urinary bladder, is one of the most common side effects observed in patients treated with CYP (63, 64). We established a model of chemical cystitis induced by CYP administration as previously stated by Boudes and colleagues (55). This mouse model of chemical cystitis is characterized by a mild inflammatory response in bladder tissue, organ hyperactivity, and pelvic allodynia (55). In good agreement with the report of Boudes and colleagues (55), our results showed that the administration of CYP (80 mg/kg) to WT mice every other day for a week caused pelvic allodynia (Fig. 1, A and B). A growing body of evidence indicates that spontaneous afferent activity and enhanced responses to physiological stimuli (i.e., afferent sensitization) contribute to the manifestation of pain behaviors in experimental animals (54, 65–71). To assess whether the referred pelvic allodynia seen in WT mice treated with CYP is driven by sensitized primary afferents, we examined the electrical properties and firing to sustained stimulation of acutely isolated bladder sensory neurons. Neurons were classified based on the sensitivity of the action potential to tetrodotoxin (TTX) as TTX resistant (TTX-R) or TTX sensitive (TTX-S). Functional studies have indicated that most bladder C fiber afferents are sensitive to capsaicin and discharge action potentials that are insensitive (resistant) to tetrodotoxin (TTX-R), whereas most Aδ fiber afferents are insensitive to capsaicin and sensitive to TTX (TTX-S) (2, 3, 54). This is in good agreement with the expression of TTX-R voltage-gated Na⁺ (Na_v) channel subunits (i.e., Na_v1.8 and Na_v1.9) in NF-200 negative sensory neurons (72). Table 1 shows the passive and active electrical properties of acutely isolated bladder sensory neurons from WT mice treated with saline or CYP. Consistent with sensitization, acutely isolated bladder sensory neurons with TTX-R action potentials from mice treated with CYP had a lower rheobase and more negative action potential threshold than those from control mice (Table 1). To examine the excitability of bladder sensory neurons, we applied suprathreshold current pulses equivalent to 1, 1.5, 2, 2.5, and 3 times the rheobase for 500 ms with 4-s intervals (Fig. 1C). Concordant with sensitization, neurons with TTX-R action potentials from mice treated with CYP discharged more spikes in response to electrical stimulation than their control counterparts (Fig. 1D). In summary, these findings indicate that the referred pelvic allodynia seen in the setting of CYP-induced chemical cystitis is mediated, at least in part, by sensitized afferents with TTX-R action potentials.

Fig. 1.

Mouse model of chemical cystitis induced by cyclophosphamide (CYP) administration. A: experimental protocol for CYP administration. Female mice were injected intraperitoneally with saline [control (Ctrl)] or CYP (80 mg/kg). B: 50% withdrawal threshold (g) to von Frey filaments applied to the pelvic area for wild-type (WT) mice treated with saline or CYP (n = 6 for each group). **P < 0.01 (Mann–Whitney test). C: CYP treatment sensitizes bladder sensory neurons with tetrodotoxin-resistant (TTX-R) action potentials. Representative tracings of action potential firing in response to suprathreshold stimulation of bladder sensory neurons with TTX-R action potentials from WT mice treated with saline (top) or CYP (middle). Action potentials were evoked by depolarizing current pulses equal to 1, 1.5, 2, 2.5, and 3 times the rheobase (Rh). The current pulse protocol is shown at bottom. D: stimulus-response relationships for sensory neurons from WT mice treated with saline or CYP (n = 12−37). *P < 0.05, **P < 0.01 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). TTX-S, tetrodotoxin sensitive.

Table 1.

Effect of CYP administration on passive and active electrical properties of bladder sensory neurons

	Saline		CYP
	TTX-R	TTX-S	TTX-R	TTX-S
Membrane capacitance, pF	31 ± 2	36 ± 3	25 ± 2*	43 ± 4
Resting membrane potential, mV	−59 ± 1	−59 ± 2	−54 ± 1	−55 ± 2
AP rheobase, pA	688 ± 112	560 ± 197	155 ± 46†	591 ± 151
AP threshold, mV	−25 ± 1	−23 ± 1	−31 ± 1†	−24 ± 1
AP duration, ms	3.3 ± 0.1	2.3 ± 0.2	3.6 ± 0.4	3.7 ± 0.2
AP amplitude, mV	103 ± 2	96 ± 3	104 ± 2	96 ± 4
Magnitude of afterhyperpolarization, mV	−12 ± 1	−9 ± 2	−19 ± 2*	−15 ± 3

Values are means ± SE; n = 12–37. Wild-type mice received 4 doses of saline or cyclophosphamide (CYP), and sensory neurons were isolated a day after the last dose as described in materials and methods. Neurons were classified based on the sensitivity of the action potential (AP) to tetrodotoxin (TTX) as TTX sensitive (TTX-S) or TTX resistant (TTX-R). APs were evoked by 4-ms current pulses. Statistically significant differences compared with controls (saline): *P < 0.05 and †P < 0.001 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test).

Loss of ASIC3 Exacerbates Nociceptive Responses in Mice Treated With CYP

To assess the role of ASICs in bladder nociception, we used a global Asic3 KO mouse line (33). We have previously shown that a fraction of the sensory neurons innervating the urinary bladder exhibit proton-gated currents and that deletion of the Asic3 subunit from these neurons abolished their ability to discharge action potentials in response to extracellular acidification (7). This finding indicates that ASIC function in bladder sensory neurons depends on the ASIC3 subunit. Therefore, to determine whether loss of ASIC3 alters somatic sensitivity in the pelvic area, we measured the 50% mechanical withdrawal threshold (g) to von Frey filaments in WT and global Asic3 KO mice. Of note, both female and male Asic3 KO mice treated with CYP exhibited an exaggerated response to von Frey filaments applied to the pelvic area 5 days after the initial dose of CYP, a time point at which only modest changes in pelvic sensitivity were apparent in WT mice (Fig. 2, B and C). Note that four doses of CYP (7 days of treatment) were required to observe a change in pelvic sensitivity of similar magnitude in WT mice (Fig. 1). Bladder water content, a surrogate of edema, was significantly higher in mice treated with CYP than control mice, but no differences were seen between WT and Asic3 KO mice (Fig. 2D). We did not observe inflammatory cells in paraffin-embedded bladder sections stained with hematoxylin and eosin of mice from either group treated with CYP or control mice (Fig. 2E). Therefore, we conclude that the inflammatory process does not account for the difference in pelvic sensitivity observed between the two groups. These results support the notion that ASIC signaling exerts an inhibitory effect on bladder nociceptive pathways.

Fig. 2.

Loss of acid-sensing ion channel subunit 3 (Asic3) enhances pelvic nociceptive responses to von Frey filaments. A: experimental protocol for cyclophosphamide (CYP) administration. Mice were injected intraperitoneally with saline [control (Ctrl)] or CYP (80 mg/kg). B: 50% withdrawal threshold (g) to von Frey filaments applied to the pelvic area for female wild-type (WT) and Asic3 knockout (KO) mice treated with saline (Ctrl) or CYP (n = 15–20). **P < 0.01, ***P < 0.001 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). C: 50% withdrawal threshold (g) to von Frey filaments applied to the pelvic area for male WT and Asic3 KO mice treated with saline or CYP (n = 10–14). ***P < 0.001 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). D: tissue water content of urinary bladders from male WT and Asic3 KO mice treated with saline or CYP (n = 12–18). *P < 0.05, **P < 0.01 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). E: hematoxylin and eosin-stained bladder sections from WT and Asic3 KO mice treated with saline or CYP (n = 4 for each group). SM, smooth muscle; UT, urothelium. Note that sections from mice treated with CYP presented edema.

CYP Treatment Increases Voiding Frequency

To assess voiding function in awake female mice with chemical-induced cystitis, we used a video-monitored void spot assay. As shown in Fig. 3A, left, WT and Asic3 KO mice that received saline exhibited a characteristic voiding pattern with large voids near a border. In contrast, mice treated with CYP exhibited an irritated bladder phenotype with a high number of voiding events of small volume (Fig. 3A, right). Mice treated with CYP from both groups exhibited higher voiding activity and a smaller mean volume per void than control mice (Fig. 3, C and D). No significant differences in these parameters were observed between WT and Asic3 KO mice that received CYP.

Fig. 3.

Cyclophosphamide (CYP) treatment increases voiding frequency. A: representative images showing voiding spots from wild-type (WT) and acid-sensing ion channel subunit 3 (Asic3) knockout (KO) mice treated with saline [control (Ctrl)] or CYP. Voiding events are marked with a red arrow. B: representative examples of voluntary urination behavior during a 6-h period for WT and Asic3 KO mice treated with saline or CYP. C and D: voiding function in awake WT and Asic3 KO mice treated with saline or CYP evaluated with a video-monitored void spot assay. Number of voids (C) and mean volume per void (D) were computed as described in materials and methods (n = 9 or 10 independent mice per group). **P < 0.01 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test).

ASICs Control Afferent Sensitization

Sensitization of primary visceral afferents is expressed as spontaneous activity, an increase in response magnitude and/or a decrease in response threshold (73). To assess whether the referred allodynia observed in Asic3 KO mice treated with CYP is driven by sensitized primary afferents, we examined the electrical properties and firing evoked by sustained electrical stimulation of acutely isolated bladder sensory neurons from WT and Asic3 KO mice treated with three doses of saline or CYP. Consistent with sensitization, bladder sensory neurons with TTX-S action potentials isolated from both WT and Asic3 KO mice treated with CYP had a more positive resting membrane potential, lower rheobase, and more negative action potential threshold than their control counterparts that received saline (Table 2). No changes in passive or active electrical properties of bladder sensory neurons with TTX-R action potentials were observed among WT mice treated with saline or CYP. In contrast, bladder sensory neurons with TTX-R action potentials from Asic3 KO mice treated with CYP exhibited a lower rheobase and more negative action potential threshold than their control counterparts. Subsequently, we evaluated whether CYP treatment alters action potential firing during sustained stimulation. As shown in Fig. 4, C and D, sensory neurons with TTX-R and TTX-S action potentials from both WT and Asic3 KO mice treated with saline displayed a phasic pattern of action potential firing in response to suprathreshold stimulation. CYP treatment promoted the tonic firing of sensory neurons with TTX-S action potentials in both groups (Fig. 4C). Strikingly, we found that the administration of CYP to Asic3 KO mice resulted in the sensitization of bladder sensory neurons with TTX-R action potentials (Fig. 4D). Note that, independently of the treatment (saline or CYP), a similar proportion of bladder sensory neurons responded to TTX in WT and Asic3 KO mice (Fig. 4B). In summary, these results support the notions that 1) ASICs regulate the sensitization of a cluster of nociceptive neurons, 2) the regimen of CYP used in our study promotes the sensitization of Aδ fibers at early time points and of C fibers at later time points in WT mice, 3) differences in pelvic sensitivity between WT and Asic3 KO mice treated with CYP are mediated by sensitized C fibers, and 4) based on the referred pelvic allodynia seen in Asic3 KO mice, loss of ASIC3 results in the sensitization of high-order neurons and interneurons in the spinal cord.

Table 2.

Effect of CYP administration on passive and active electrical properties of bladder sensory neurons from WT and Asic3 KO mice

	WT				Asic3 KO
	Saline		CYP		Saline		CYP
	TTX-R	TTX-S	TTX-R	TTX-S	TTX-R	TTX-S	TTX-R	TTX-S
Membrane capacitance, pF	31 ± 2	42 ± 3	30 ± 2	37 ± 5	30 ± 2	40 ± 5	27 ± 2	33 ± 2
Resting membrane potential, mV	−55 ± 2	−60 ± 2	−54 ± 1	−50 ± 2d	−56 ± 2	−57 ± 3	−53 ± 2	−50 ± 1f
AP rheobase, pA	484 ± 66	587 ± 91	419 ± 90	157 ± 73e	500 ± 121	596 ± 107	141 ± 31b	134 ± 42d
AP threshold, mV	−26 ± 1	−23 ± 1	−27 ± 1	−26 ± 1c	−25 ± 1	−23 ± 1	−30 ± 1b	−28 ± 1c
AP duration, ms	3.8 ± 0.3	2.8 ± 0.3	3.8 ± 0.5	3.5 ± 0.5	3.8 ± 0.4	2.5 ± 0.4	2.7 ± 0.2	3.1 ± 0.6
AP amplitude, mV	98 ± 3	100 ± 5	97 ± 3	91 ± 5	101 ± 5	97 ± 6	95 ± 4	85 ± 5
Magnitude of afterhyperpolarization, mV	−15 ± 2	−7 ± 2	−16 ± 1	−18 ± 3c	−13 ± 3	−11 ± 3	−23 ± 2a	−22 ± 3

Values are means ± SE; n = 10–22. Wild-type (WT) and acid-sensing ion channel subunit 3 (Asic3) knockout (KO) mice received 3 doses of saline or cyclophosphamide (CYP), and sensory neurons were isolated a day after the last dose as described in materials and methods. Neurons were classified based on the sensitivity of the action potential (AP) to tetrodotoxin (TTX) as TTX sensitive (TTX-S) or TTX resistant (TTX-R). APs were evoked by 4-ms current pulses. Statistically significant: ^aP < 0.05, Asic3 KO CYP vs. WT CYP or Asic3 KO saline; ^bP < 0.001, Asic3 KO CYP vs. WT CYP or Asic3 KO saline; ^cP < 0.05, WT saline vs. WT CYP or Asic3 KO saline vs. Asic3 KO CYP; ^dP < 0.01, WT saline vs. WT CYP or Asic3 KO saline vs. Asic3 KO CYP; ^eP < 0.001, WT saline vs. WT CYP; ^fP < 0.01, Asic3 KO saline vs. Asic3 KO CYP (Kruskal-Wallis test followed by Dunn’s multiple comparisons test).

Fig. 4.

Acid-sensing ion channel subunit 3 (ASIC3) controls the sensitization of bladder nociceptors. Mice received 3 doses of saline [control (Ctrl)] or cyclophosphamide (CYP) every other day, and sensory neurons were isolated a day after the last dose. Patch-clamp experiments were conducted within 2–8 h after isolation. A: representative tracings of action potential firing in response to suprathreshold stimulation of acutely isolated bladder sensory neurons with tetrodotoxin-resistant (TTX-R) action potentials from wild-type (WT) (top) and Asic3 knockout (KO) (middle) mice. The current pulse protocol is shown at bottom. Rh, rheobase. B: percentage of bladder sensory neurons from WT and Asic3 KO with TTX-R and TTX-sensitive (TTX-S) action potentials (n = 27–37). C: stimulus-response relationships for sensory neurons with TTX-S action potentials from mice treated with saline or CYP (n = 10–15). *P < 0.05, Asic3 KO control vs. Asic3 KO CYP; and +P < 0.001, WT control vs. WT CYP (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). D: stimulus-response relationships for sensory neurons with TTX-R action potentials from WT and Asic3 KO mice treated with saline or CYP (n = 15–22). **P < 0.01, +P < 0.001, Asic3 KO CYP vs. Asic3 KO control or WT CYP (Kruskal-Wallis test followed by Dunn’s multiple comparisons test).

Loss of ASIC3 Alters Proton Signaling in Sensory Neurons

To assess the effect of CYP treatment in ASIC function, we examined the currents evoked by extracellular acidification in bladder sensory neurons harvested from WT and Asic3 KO mice treated with saline or CYP. Bladder sensory neurons were classified based on the response elicited by extracellular acidification as nonresponsive or responsive with a transient or sustained response (Fig. 5) (7). Extracellular acidification evoked a transient inward current characteristic of ASICs in ∼25–30% of the bladder sensory neurons harvested from both WT and Asic3 KO mice treated with saline or CYP (Fig. 5, A and D, and Fig. 6A). About 50% of the bladder sensory neurons exhibited a sustained response to extracellular acidification with an increase in current that did not decay over time (Fig. 5, B and E, and Fig. 6A). A significant proportion (22–25%) of bladder sensory neurons from both WT and Asic3 KO mice were insensitive to extracellular acidification (Fig. 5, C and F, and Fig. 6A). In good agreement with our previous study (7), proton-evoked currents were significantly larger in bladder sensory neurons from Asic3 KO mice than from WT mice treated with saline (Fig. 6B). CYP administration caused a reduction in the magnitude of the current evoked by acidification in neurons isolated from Asic3 KO mice (Fig. 6B). As expected, and consistent with previous studies (7, 33), the current decay (desensitization) was slower in bladder sensory neurons from Asic3 KO mice than from WT mice (Fig. 6C). CYP treatment accelerated ASIC desensitization in WT mice. Since the time course of desensitization depends on the channel’s subunit composition (18, 26, 74), we conclude that bladder sensory neurons express multiple ASIC subunits and that loss of ASIC3 results in the assembly of ASIC1/2 heterotrimers with slower rates of desensitization.

Fig. 5.

Proton-evoked currents in bladder sensory neurons. Representative tracings show the 3 distinctive currents evoked by extracellular acidification in bladder sensory neurons from wild-type (WT) mice (A–C) and acid-sensing ion channel (ASIC) subunit 3 (Asic3) knockout (KO) mice (D–F). Dorsal root ganglia were harvested from mice 7–10 days after injection of 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiI) in the bladder wall. Sensory neurons were isolated and cultured as described in materials and methods. Whole cell currents were evoked by a change in extracellular pH from 8.0 to 6.5. Sensory neurons were classified on the basis of their response to extracellular acidification as responsive with a transient increase in current (ASIC; A and D), responsive with a sustained current increase (B and E), and nonresponsive (C and F).

Fig. 6.

Loss of acid-sensing ion channel (ASIC) subunit 3 (ASIC3) ablates proton signaling in bladder sensory neurons. Mice received 3 doses of saline [control (Ctrl)] or cyclophosphamide (CYP) every other day, and sensory neurons were isolated a day after the last dose. Patch-clamp experiments were conducted within 24–72 h after isolation. A: percentage of bladder sensory neurons from wild-type (WT) and Asic3 knockout (KO) mice with ASIC currents (n = 25–98). B: peak current density of ASIC current in bladder sensory neurons from WT and Asic3 KO mice (n = 9–29). *P < 0.05, **P < 0.01 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). Currents were evoked by a drop in extracellular pH from 8.0 to 6.5. C: time constant of desensitization (τ) of ASIC currents in sensory neurons from WT and Asic3 KO mice treated with saline or CYP (n = 9–29). **P < 0.01, ***P < 0.001 (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). D: representative tracings of ASIC currents in bladder sensory neurons from WT and Asic3 KO mice treated with CYP. Whole cell currents were repeatedly evoked by a change in extracellular pH from 8.0 to 6.5 with an interstimulus interval of 1 min. E: ASICs in sensory neurons from Asic3 KO mice exhibit tachyphylaxis. ASIC currents were evoked repeatedly by a drop in extracellular pH from 8.0 to 6.5 with an interstimulus interval of 1 min in sensory neurons harvested from WT or Asic3 KO mice. Proton-evoked peak currents were normalized to the amplitude of the 1st peak (n = 6–11). *P < 0.05, WT CYP vs. Asic3 KO CYP; and **P < 0.01, WT Ctrl vs. Asic3 KO Ctrl and WT CYP vs. Asic3 KO CYP (Kruskal-Wallis test followed by Dunn’s multiple comparisons test). F: representative voltage recordings illustrating the effect of extracellular acidification on bladder sensory neurons from WT and Asic3 KO mice treated with CYP. The area labeled a is expanded in the inset. Note that bladder sensory neurons from WT mice, but not those from Asic3 KO mice, discharged action potentials in response to extracellular acidification. G: percentage of bladder sensory neurons from WT and Asic3 KO mice treated with CYP that discharged action potentials in response to extracellular acidification (n = 8–11).

ASICs in naive mouse sensory neurons desensitized rapidly at acidic pH and transitioned to an activable state upon pH neutralization within seconds (18). It is noteworthy that we found that repetitive pH stimulation of bladder sensory neurons from Asic3 KO mice resulted in a progressive attenuation of the peak current (Fig. 6, D and E). In contrast, and consistent with previous studies, proton-evoked currents in neurons from WT mice do not decay upon repetitive stimulation. Previous studies have shown that repeated stimulation of cells expressing ASIC1a results in a gradual attenuation of proton-evoked current, a process referred as tachyphylaxis (75–77). The adaptation to repetitive stimulation seen in sensory neurons from Asic3 KO mice further indicates that they express ASIC1/2 heterotrimers.

ASICs are considered bona fide proton sensors that mediate the responses to acidification in sensory neurons. To assess whether CYP treatment alters proton signaling in bladder afferents, we exposed sensory neurons isolated from WT and Asic3 KO mice treated with CYP to a pulse of acidic pH under current-clamp conditions. Consistent with our previous study (7), 64% of bladder sensory neurons with ASIC currents from WT mice treated with CYP, but none of those harvested from Asic3 KO mice, discharged action potentials in response to extracellular acidification (Fig. 6, F and G). This result demonstrates that loss of ASIC3 ablates proton signaling in bladder afferents.

DISCUSSION

ASICs are depolarizing channels that mediate responses to extracellular acidification at afferent nerve terminals. Numerous studies have reported an increase in mechanosensitivity (7, 32, 33, 35–39) and enhanced pain behaviors (present study and Refs. 28, 40) in ASIC subunit-null mice. However, until now, it was unclear how loss of a proton sensor can increase sensory responses in ASIC subunit-null mice. Here, we provide direct evidence that ASIC signaling exerts an inhibitory effect on bladder nociceptive pathways, delaying afferent sensitization in the face of injury induced by CYP administration.

Understanding the process of afferent sensitization is important, as persistent afferent drive increases the excitability of spinal ascending neurons, a process referred as to “central sensitization” (50), which underlies referred somatic hypersensitivity and cross-organ sensitization. The results of our study revealed that three doses of CYP prompt the sensitization of bladder sensory neurons with TTX-S action potentials (i.e., Aδ fibers) in WT mice, but that four doses cause the sensitization of sensory neurons with TTX-R action potentials (i.e., C fibers). Of major significance, only three doses of CYP were necessary to induce pelvic hypersensitivity in Asic3 KO mice. The sensitization of sensory neurons of C origin in WT and Asic3 KO mice correlates temporally with the emergence of pelvic allodynia. In essence, we conclude that the differences in mechanical pelvic sensitivity seen between WT and Asic3 KO mice treated with CYP (3 doses) are mediated by sensitized C nociceptors. Remarkably, even though WT mice developed pelvic hypersensitivity with four doses of CYP, the degree of sensitization (e.g., number of action potentials discharged in response to suprathreshold stimulation) of bladder sensory neurons with TTX-R action potentials in this group did not reach that seen in sensitized neurons from Asic3-null mice. Taken as a whole, our findings are consistent with ASICs regulating the excitability and, as a result, the sensitization of a population of bladder nociceptors.

Voiding behaviors were not influenced by the genotype of the mice. However, as expected, both WT and Asic3 KO mice treated with CYP exhibited increased voiding activity compared with control mice. Patch-clamp experiments showed that CYP treatment sensitizes to the same extent neurons with TTX-S action potentials in both WT and Asic3 KO mice. This suggests that the observed changes in voiding behavior between mice treated with CYP and control mice are driven by sensitized bladder afferents with TTX-S action potentials (i.e., Aδ fibers). In summary, the results of our study indicate that ASICs do not contribute to the irritative voiding behavior seen in mice treated with CYP. Our findings are consistent with the notion that voiding and nociceptive responses to noxious stimuli are mediated by different populations of bladder sensory neurons, Aδ and C, respectively.

Even though acid-gated currents were first observed in sensory neurons in the early 1980s (78–80), we still do not understand the function of ASICs in afferent fibers. Likewise, it remains unclear how protons are generated in the proximity of nerve terminals to trigger sensory responses. Consistent with previous reports (7, 33), our study revealed that deletion of ASIC3 from sensory neurons does not ablate ASIC currents, but it altered their magnitude and rate of desensitization. These findings indicate that ASICs are assembled as heterotrimers in bladder sensory neurons. Unexpectedly, CYP treatment altered the magnitude and rate of desensitization of ASIC currents. This could be the result of changes in the expression of ASIC subunits and consequently assembly of channels with different stoichiometry and biophysical properties or alternatively posttransductional modifications that alter channel function. Of major significance, the results of our study showed that bladder sensory neurons from Asic3 KO mice are unable to discharge action potentials in response to a drop in extracellular pH, consistent with the notion that ASICs serve as bona fide proton sensors in afferents. In summary, our findings indicate that protons and their cognate ASIC receptors encompass a tuning mechanism at nerve terminals that regulate afferent excitability and that the loss of this mechanism results in aberrant firing in the face of injury and consequently in afferent sensitization. Therefore, we posit that peripherally restricted allosteric modulators that target ASICs can be used to treat bladder conditions that present with pain, such as interstitial cystitis/bladder pain syndrome.

Perspectives and Significance

Our study shows that ASICs are necessary for the response of bladder sensory neurons to extracellular acidification and provides evidence that they regulate the transmission of nociceptive information arising from nociceptors innervating the urinary bladder. However, we cannot explain how the loss of proton signaling increases neuronal excitability in the face of injury induced by CYP. Hence, further studies are needed to define at the molecular level the mechanism by which ASIC activation tunes afferent firing. To the best of our knowledge, this is the first study that discloses a mechanism mediated by a ligand-gated ion channel that operates in afferents to regulate excitability and sensitization.

GRANTS

This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants R01DK084060 and RO1DK119183, by the Physiology and Model Systems Core of the Pittsburgh Center for Kidney Research (P30DK079307), and by the Urology Care Foundation Research Scholar Award Program (to N.M.).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

N.M. and M.D.C. conceived and designed research; performed experiments; analyzed data; interpreted results of experiments; prepared figures; drafted manuscript; edited and revised manuscript; and approved final version of manuscript.