Endometriosis and vulvodynia affect 10% to 28% of females of reproductive age,21,31,53,69 contributing to a combined economic burden of more than $140 billion/year in the United States alone.64,76 Dyspareunia, also known as vaginal hyperalgesia or painful intercourse, is one of the most debilitating symptoms experienced by women with endometriosis and vulvodynia. Despite this, the aetiology and pathogenesis of chronic pain associated with endometriosis- and vulvodynia-induced dyspareunia remains to be identified.25,40,67,79
The first step in the pain pathway is the primary sensory afferents that project from peripheral tissues to the central nervous system (CNS). The colon, bladder, and female reproductive organs all receive input from pelvic sensory afferent pathways.75 Although many studies have characterised the properties of sensory afferent nerves innervating the colon and bladder,27 little is known about how mechanical stimuli is detected and transmitted from female reproductive organs to the CNS. Thirty years ago, Berkley's group showed that pelvic and hypogastric afferents innervating the rat female reproductive tract fired action potentials to a variety of mechanical and chemical stimuli.3,4,6,55 Hormonal variations occurring across the estrous cycle, pregnancy, and labour also modulate the mechanosensitivity of these afferents.43,44,54 Remarkably, additional studies exploring the mechanosensory properties of sensory afferents innervating female reproductive organs are lacking.
Modulation of ion channels and receptors expressed by peripheral sensory afferent nerves has shown promise in the treatment of chronic pain arising from visceral organs.10,28,37,47,57,58 However, only a limited number of studies have investigated the expression of nociceptive ion channels within sensory neurons projecting to female reproductive organs. Amongst these, ion channel transient receptor potential vanilloid 1 TRPV1,19,70 purinergic P2X3,19 the voltage-gated sodium (NaV) channel NaV1.8,41 and the voltage-gated potassium channels (Kv) KV6.4 and KV2.141 have been identified, but function was not explored.
Recent evidence shows that NaV channels have potential in the management of acute and chronic visceral pain.22,36,37,50,58 NaV channels are important determinants of sensory neuron excitability, having an essential role in the generation and propagation of action potentials and the transduction of sensory stimuli.17,18,20,22 The NaV channel family consists of 9 isoforms (NaV1.1 − NaV1.9) that are distinguished by their relative sensitivity to the neurotoxin tetrodotoxin (TTX), characterised as either TTX-sensitive (NaV1.1-NaV1.4, NaV1.6, and NaV1.7) or TTX-resistant (TTX-R) (NaV1.5, NaV1.8, and NaV1.9).17,18 Numerous channelopathies in SCN9A (NaV1.7), SCN10A (NaV1.8), and SCN11A (NaV1.9) have been demonstrated to alter pain sensitivity in humans.20,71
Given the lack of knowledge on the sensory afferent pathways innervating female reproductive organs, the aims of this study were: (1) to examine the mechanosensory properties of sensory afferents innervating the mouse vagina; (2) to determine the expression profile of NaV channels contained within these afferents, (3) to determine their contribution to vaginal afferent excitability, and (4) to determine how pharmacological modulation of these channels alters nociceptive signalling and ultimately regulation of vaginal pain sensitivity in vivo.
The Animal Ethics Committees of the South Australian Health and Medical Research Institute (SAHMRI), and Flinders University approved all experiments involving animals (ethics number SAM342). All experiments conformed to the relevant regulatory standards and the ARRIVE guidelines. Virgin female C57BL/6J mice at 8 to 13 weeks of age were used and acquired from an in-house C57BL/6J breeding program (JAX strain #000664; originally purchased from The Jackson Laboratory, breeding barn MP14; Bar Harbor, ME) within SAHMRI's specific and opportunistic pathogen-free animal care facility. Mice were group housed (maximum 5 mice per cage) within individual ventilated cages (IVCs), which were filled with coarse chip dust-free aspen bedding (PURA; Cat#—ASPJMAEB-CA, Niederglatt, Switzerland). These cages were stored on IVC racks in specific housing rooms within a temperature-controlled environment of 22°C and a 12-hour light/12-hour dark cycle. Mice had free access to LabDiet JL Rat and Mouse/Auto6F chow (Cat# 5K52, St. Louis, MO) and autoclaved reverse osmosis purified water. It has been reported that an extended absence of male pheromones leads to a state of anestrus (Lee–Boot effect).45 In this current study, female mice were group housed in IVC cages and the littermate male mice were separated at weaning. All female mice were housed in IVC racks devoid of male animals with their cages cleaned in distinct Tecniplast cage cleaning stations. All female mice used in this study were virgin (never been mated). Although vaginal lavage or other cytology tests to confirm cycle stage were not performed in the current study, visual examination of the vaginal opening and lumen suggested no obvious signs of estrous.
2.2. Pharmacological modulators
Veratridine, a nonselective agonist of NaV channels, was purchased from Sigma-Aldrich (North Ryde, Australia). Tetrodotoxin (TTX), a selective blocker of TTX-sensitive NaV channels, was purchased from Tocris Biosciences (Australia).
2.3. Ex vivo afferent recording preparation from pelvic nerves innervating the female reproductive tract
On the day of experimentation, mice were humanely killed by CO2 inhalation, and the whole female reproductive tract was removed and afferent recordings from pelvic nerves innervating the vagina area were performed. Briefly, intact female reproductive organs (vagina and uterus) were removed along with the attached neurovascular bundle containing pelvic and pudendal nerves (Fig. 1A). The whole tissue was transferred to ice-cold Krebs solution (in mM: 117.9 NaCl, 4.7 KCl, 25 NaHCO3, 1.3 NaH2PO4, 1.2 MgSO4 (H2O)7, 2.5 CaCl2, and 11.1 D-glucose); and after further dissection, the distal and central portions of the female reproductive organs were opened longitudinally (Fig. 1B). The tissue was pinned flat, mucosal side up, in a specialised organ bath consisting of 2 adjacent compartments generated from clear acrylic (Danz Instrument Service, Adelaide, South Australia, Australia), the floors of which were lined with Sylgard (Dow Corning Corp., Midland, MI) (Fig. 1B). The neurovascular bundle containing the pelvic nerve was extended from the tissue compartment into the recording compartment where they were laid onto a mirror. A movable wall with a small “mouse hole” (to allow passage of the nerves) was lowered into position and the recording chamber filled with paraffin oil.9 The organ compartment was superfused with Krebs solution, bubbled with carbogen (95% O2 and 5% CO2) at a temperature of 34°C. The pelvic nerve was dissected away from the neurovascular bundle and from the nerve sheath surrounding the nerve under a dissecting microscope. Using fine forceps, the nerve trunk was teased apart into 6 to 10 bundles, which were individually placed onto a platinum recording electrode (Fig. 1B). A separate platinum reference electrode rested on the mirror in a small pool of Krebs solution adjacent to the recording electrode. Action potentials, generated by mechanical stimuli applied to the afferent's receptive field, were recorded by a differential amplifier, filtered, and sampled (20 kHz) using a 1401 interface (Cambridge Electronic Design, Cambridge, United Kingdom).
2.3.1. Mechanosensory profile of pelvic afferents innervating the vagina
Receptive fields tested in this study were limited to the vagina (above vaginal opening and below cervix) and were identified by systematically stroking the mucosal surface of the vagina with a stiff brush to activate all subtypes of vaginal mechanoreceptors. To study baseline mechanosensory properties of the pelvic afferents innervating a particular receptive field within the vagina, 3 distinct mechanical stimuli were tested: (1) static probing with calibrated von Frey hairs (vfh; 2 g force; applied 3 times for a period of 3 seconds); (2) mucosal stroking of the vaginal surface with calibrated vfh (10-1000 mg force; applied 10 times each); or (3) circular stretch (5 g; applied for a period of 1 minute). Stretch was applied using a claw made from bent dissection pins attached to the tissue adjacent to the afferent receptive field and connected to a cantilever system with thread.9 Weights were applied to the opposite side of the cantilever system to initiate stretch. Only circular, and not longitudinal, stretch was tested in this study. Once baseline mechanosensitivity was tested, a small chamber was then placed onto the mucosal surface of the vagina, surrounding the afferent receptive field. Residual Krebs solution within the chamber was aspirated and the NaV channel pharmacological modulators, veratridine (50 µM) and TTX (0.5 µM), were applied in separate experimental preparations for 5 minutes each. At the concentration used in this study, veratridine has been shown to activate TTX-sensitive (TTX-S) and TTX-R NaV channels,23,28 whereas TTX has been showed to selectively inhibit TTX-S NaV channels.17,28 Mechanical sensitivity of the afferent receptive field was then retested in response to the 3 distinct mechanical stimuli previously tested.
2.3.2. Statistical analysis of afferent recording data
Action potentials were analysed offline using the Spike 2 (version 5.21) software (Cambridge Electronic Design) and discriminated as single units based on distinguishable waveforms, amplitudes, and durations. Data are expressed as mean ± SEM. n = the number of afferents recorded, N = the number of animals used for those specific experiments. Data were statistically compared using Prism 7 software (GraphPad Software, San Diego, CA) and, where appropriate, were analysed using paired or unpaired Student t test and one- or two-way analysis of variance with Bonferroni post hoc tests. Differences were considered statistically significant at *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.
2.4. Identification and culture of dorsal root ganglia neurons innervating the vagina
Retrograde labelling was performed to identify DRG neurons innervating the vagina. Cholera toxin subunit B (0.5%) conjugated to Alexa Fluor 488 (diluted in 0.1 M phosphate buffer, pH = 7.35; Invitrogen, Carlsbad, CA) was injected (2 µL/injection site) at 5 different sites into the vaginal wall. Injections were made using a Hamilton syringe attached to a 23-gauge needle. A tunnelling method was used, whereby the tip of the needle is inserted into the vaginal wall, ∼0.2 cm from the vaginal opening and tunnelled an additional 0.2 cm in a cranial direction within the vaginal wall. The tracer is then expelled as the needle is gradually extracted from the needle tract. This ensures that a greater area of the vaginal wall is exposed to the dye compared to localised injections at the needle insertion points. Mice were then allowed to recover, housed individually, and closely monitored. All mice recovered quickly and showed no adverse signs. Four days after vaginal tracing, mice were humanely euthanised through CO2 inhalation and lumbosacral (LS; L5-S1) dorsal root ganglia (DRG) were removed. Dorsal root ganglia were digested in Hanks balanced salt solution (HBSS; pH 7.4; Life Technologies, Waltham, MA, #14170161) containing 2.5 mg/mL collagenase II (GIBCO, Thermo Fisher Scientific, Waltham, #17101015) and 4.5 mg/mL dispase (GIBCO, Thermo Fisher Scientific, #17105041) at 37°C for 30 minutes. The collagenase–dispase solution was aspirated and replaced with HBSS containing collagenase (4.5 mg/mL) only for 10 minutes at 37°C. After subsequent washes in HBSS, DRG were mechanically disrupted and cells dissociated in 600 µL complete Dulbecco's Modified Eagle Media ([DMEM; GIBCO, Thermo Fisher Scientific, #11995065]; 10% Fetal Calf Serum [Invitrogen, Thermo Fisher Scientific, Waltham, MA]; 2 mM L-glutamine [GIBCO, Thermo Fisher Scientific, #25030081], 100 µM MEM nonessential amino acids [GIBCO, Thermo Fisher Scientific, #11140076], 100 mg/mL penicillin/streptomycin [GIBCO, Thermo Fisher Scientific, #15070063], and 96 µg/L nerve growth factor-7S [Sigma-Aldrich, N0513-0.1 MG]) via trituration through fire-polished Pasteur pipettes of descending diameter, and centrifuged for 1 minute at 50g.1,8,14–16,36,50 Neurons were resuspended in 300 µL complete DMEM and spot-plated (20 µL) onto 13-mm coverslips coated with laminin (20 μg/mL; Sigma-Aldrich, #L2020) and poly-D-lysine (800 μg/mL; Thermo Fisher Scientific). Coverslips were incubated at 37°C in 5% CO2 for 2 to 3 hours to allow neurons to adhere before flooding with 1.7 mL complete DMEM. Neurons were collected for single-cell reverse transcription-polymerase chain reaction (RT-PCR) 12 to 24 hours after dissociation or were recorded using patch-clamp electrophysiology 20 to 48 hours after dissociation.
2.5. Reverse transcription-polymerase chain reaction of individual vagina-innervating dorsal root ganglia neurons
2.5.1. Neuron collection and lysis
Dissociated DRG neurons from vagina-traced mice were perfused with bath solution and picked into the end of an air-filled wide-aperture borosilicate glass pipette fabricated in the P-97 (Sutter Instruments, Novato, CA) pipette puller 12 to 24 hours after dissociation. A separate pipette was used to sample bath solution to assess RNA contamination. The end of the glass pipette containing the collected neuron was broken into a sterile Eppendorf tube containing 10 μL of lysis buffer with DNase (Thermo Fisher, TaqMan Gene Expression Cells-to-CT Kit, cat # AM1728). Incubation with lysis buffer occurred for 5 to 10 minutes at room temperature, followed by addition of 1 μL DNase stop solution and incubation for a further 5 to 10 minutes at room temperature. Lysates were frozen on dry ice and stored at −80°C until cDNA synthesis was performed.
2.5.2. cDNA synthesis
Synthesis of cDNA was performed using the SuperScript VILO IV ezDNase (Thermo Fisher, cat #11766050) kit according to the manufacturer's instructions. Each cDNA synthesis batch included one reverse transcriptase (RT) negative control to assess genomic DNA contamination. Synthesised cDNA was stored at −20°C until PCR was performed.
2.5.3. Reverse transcription-polymerase chain reaction
For each RT-PCR reaction, 10 μL of PCR Master mix, 0.5 μL of each TaqMan primer (see Table 1 below for details), 8 μL of water, and 1.6 μL cDNA from each sample was tested in singlicate for each target. Tubb3 was used as a neuronal marker, and Gfap was used as a glial marker. RT controls, bath controls, and negative controls (water instead of cDNA) were routinely included in PCR reactions, and a positive control test was performed for each primer using cDNA synthesised from whole DRG RNA. Assays were run for 50 cycles on a 7500 Fast Real-Time PCR System (Applied Biosystems, Victoria, Australia) machine, using 7500 Fast software, v2.0.6. Genes were considered expressed if a complete amplification curve was obtained within 50 cycles.
Table 1 -
TaqMan primers used for qRT-PCR and single-cell RT-PCR, obtained from Life Technologies.
|Hypoxanthine-guanine phosphoribosyltransferase 1 (reference gene)
|Glyceraldehyde 3-phosphate dehydrogenase (reference gene)
|Tubulin, beta 3 class III
RT-PCR, reverse transcription-polymerase chain reaction; qRT-PCR, quantitative RT-PCR.
2.6. Quantitative reverse transcription-polymerase chain reaction of whole lumbosacral dorsal root ganglia and vaginal tissue
2.6.1. Tissue collection
Dorsal root ganglia (L5-S1) and vaginal tissues were collected immediately after euthanasia by CO2 inhalation. For DRG, whole lumbosacral DRG were surgically removed, snap frozen, and stored at −80°C before RNA extraction. Vaginal tissues were collected by removing the whole female reproductive tract and dissecting away the cervix, uterus, and uterine horns. The remaining/sole vaginal tissue was then snap frozen and stored at −80°C before RNA extraction.
2.6.2. RNA extraction
RNA was extracted using the PureLink RNA Micro kit (Invitrogen, Victoria, Australia, cat #12183-016; DRG) or the PureLink RNA Mini kit (Invitrogen, cat #12183018A; vaginal tissue) followed by a DNAse treatment (Life Technologies, cat #12185-010) according to the manufacturer's instructions.
2.6.3. Quantitative reverse transcription-polymerase chain reaction
Quantitative RT-PCR was performed using Express qPCR Supermix (Applied Biosystems, cat # 11785200) with commercially available hydrolysis probes (TaqMan; Life Technologies, see Table 1 above for details) and RNAse-free water (AMBION, Victoria, Australia, cat #AM9916). For each reaction, 10 μL of qPCR SuperMix, 1 μL of TaqMan primer, 2 µL RT enzyme mix, 2 μL of water, and 5 μL of RNA (diluted in RNAse-free H2O to approximately 100 ng/well) from each sample were tested in duplicate for each NaV channel subtype. Hprt and Gapdh were used as endogenous controls for all tissues. Assays were run for 50 cycles on a 7500 Fast Real-Time PCR System (Applied Biosystems) machine, using 7500 Fast software, v2.0.6. Quantity of mRNA is expressed as ΔCt relative to the geometric mean of reference genes Hprt and Gapdh.
2.7. Whole-cell current-clamp and voltage-clamp electrophysiology of vagina-innervating dorsal root ganglia neurons
Dissociated DRG neurons isolated from vagina-traced mice were recorded in current-clamp mode on day 1 after culturing (20-30 hours) and in voltage-clamp mode on day 1 and day 2 after culturing (20-48 hours). Neurons patched in this study were small to medium in size, with an average diameter of 27 ± 1 μm.
2.7.1. Solutions and pipettes
Intracellular current-clamp solution contained (in mM): 135 KCl; 2 MgCl2; 2 MgATP; 5 EGTA-Na; and 10 HEPES-Na; adjusted to pH 7.3. Extracellular (bath) current-clamp solution contained (in mM): 140 NaCl; 4 KCl; 2 MgCl2; 2 CaCl2; 10 HEPES; and 5 glucose; adjusted to pH 7.4. Intracellular voltage-clamp solution contained (in mM): 60 CsF; 45 CsCl; 2 MgCl2; 5 EGTA-Na; 10 HEPES-Cs; 30 TEA-Cl; and 2 MgATP; adjusted to pH 7.2 with CsOH, 280 mOsm. Extracellular (bath) voltage-clamp solution contained (in mM): 70 NaCl; 50 NMDG; 40 TEA-Cl; 4 CsCl; 2 MgCl2; 2 CaCl2; 10 HEPES; and 5 glucose; adjusted to pH 7.4, approximately 300 mOsm. Standard wall borosilicate glass pipettes (OD × ID × length: 1.5 × 0.86 × 7.5 cm, Harvard, cat # 64-0792) were pulled and fire-polished to 3 to 10 MΩ for current-clamp recordings, and 1 to 3 MΩ for voltage-clamp recordings using a P-97 (Sutter Instruments) pipette puller.
Responses to TTX (0.1 mM continually perfused by gravity) were recorded at 1 minute after start of perfusion. For current-clamp recordings, 2 baseline responses were measured (1 minute apart) to assess stability of recording. Only cells with a difference of ≤1 current step in rheobase at baseline were included in the analysis. For voltage-clamp recordings, maximum peak current was monitored until reaching a stable amplitude before proceeding with compound application. Neurons were held at −70 mV for 15 ms, hyperpolarised by a −20 pA current injection for 475 ms, and then held at −70 mV for 100 ms. Stepwise depolarising pulses in increments of 10 pA were applied from holding potential of −70 mV with 2-second repetition intervals to determine the rheobase (minimum amount of current required to fire an action potential). Current–voltage (INa − V) relationships were determined by application of a prepulse to −100 mV (100 ms), followed by a series of step pulses from −65 mV to +60 mV (5-mV increments [100 ms]), before returning to hold at −70 mV (repetition interval of 3 seconds, P/8 leak subtraction). Voltage dependence of steady-state fast inactivation was determined by application of a series of prepulses from −110 to +2.5 mV (7.5-mV increments [100 ms]), then a pulse at −30 or −10 mV depending on current amplitude, followed by hold at −70 mV (50 ms) (3-second repetition interval). Diameter measurement: To determine average neuron diameter, the smallest and largest width of the neuronal soma was measured using the microscope eyepiece reticle calibrated with a stage micrometer and averaged. Exclusion criteria: For current-clamp recordings, neurons were not recorded if the resting membrane potential was more depolarized than −40 mV because this is an indicator of poor cell health. For voltage-clamp recordings, only recordings with voltage error of less than 5 mV were used for analysis.
2.7.3. Data acquisition and analysis
Recordings were amplified with Axopatch 200A, digitised with Digidata 1322A, sampled at 20 kHz, filtered at 5 kHz, recorded with pCLAMP 9 software (Molecular Devices), and analysed in Clampfit 10.3.2 (Molecular Devices), Prism v8.0.0 (GraphPad), and IBM SPSS Statistics v25.
2.8. In vivo vaginal distension activation of spinal cord dorsal horn neurons identification by phosphorylated MAP kinase ERK 1/2 (pERK)
2.8.1. In vivo vaginal distension
Vehicle (saline), veratridine (50 µM), or TTX (0.5 µM) was administered intravaginally through a small cannula inserted into the mice vaginal canal under isoflurane anaesthesia. Immediately after, and still under anaesthesia, a 3-mm length (∼4-mm diameter when fully inflated) balloon catheter was inserted into the vaginal canal. Mice were removed from the isoflurane chamber and, on regaining consciousness, the balloon was distended for 30 seconds to 40 mm Hg applied through a syringe attached to a sphygmomanometer pressure gauge. After 30 seconds, the pressure was released, and the balloon deflated (0 mm Hg) for 10 seconds. This process was repeated 5 times. After the fifth distension, mice were given an anaesthetic overdose (0.125 mL/250 g lethabarb) and by 5 minutes, had undergone transcardial perfuse fixation.
2.8.2. Transcardial perfuse fixation and tissue processing
After anaesthetic overdose, the chest cavity was opened and 0.5 mL of heparin saline was injected into the left ventricle followed by insertion of a 22-gauge needle, attached to tubing and a peristaltic perfusion pump. The right atrium was snipped, allowing for perfusate drainage. Warm saline (0.85% physiological sterile saline) was perfused before ice-cold 4% paraformaldehyde in 0.1 M phosphate buffer (Sigma-Aldrich, St. Louis, MO). After complete perfusion, L6-S2 spinal cord segments (determined by level of DRG root insertion points) were removed and postfixed in 4% paraformaldehyde in 0.1M phosphate buffer at 4°C for 18 to 20 hours. The lowest rib was used as an anatomical marker of T13. After postfixation, spinal cords were cryoprotected in 30% sucrose/phosphate buffer (Sigma-Aldrich) overnight at 4°C and then an additional 24-hour incubation at 4°C in 50% optimal cutting temperature compound (OCT; Tissue-Tek, Sakura Finetek, CA) OCT/30% sucrose/phosphate buffer solution before freezing in 100% OCT. Spinal cords were cryosectioned (10 µm thick) and placed onto gelatin-coated slides for immunofluorescence labelling. Spinal cord segments were serially sectioned and distributed over 6 slides, which were used for autostaining of neuronal activation marker pERK.32
2.8.3. Spinal cord–phosphorylated MAP kinase ERK 1/2 immunofluorescence
The spinal cord dorsal horn neurons activated by vaginal distension (VD) were identified by labelling for neuronal activation marker pERK using primary antibody (pERK; 1:800) in Antibody Diluent (S0809, Agilent DAKO, Santa Clara, CA) with 3,3′-diaminobenzidine (DAB)/horseradish peroxidase (HRP) secondary antibody staining. After air drying for 1 hour, sections were postfixed in formalin with a brief rinse in distilled water before target antigen retrieval with EnVision FLEX TRS Low pH target retrieval solution (citrate buffer, pH 6.1; K8005, Agilent DAKO). Nonspecific binding of secondary antibodies was blocked with Serum-Free Protein Block (X0909, Agilent DAKO). Tissue sections were incubated with primary antisera for 1 hour, washed and incubated for 3 minutes in Envision FLEX Peroxidase-blocking Reagent (GV823, Agilent DAKO), followed by 20-minute incubation with Envision FLEX HRP Polymer (GV823, Agilent DAKO) for HRP binding. Negative controls were prepared as above with the primary antibody omitted. Sections were then washed in wash buffer (GC807, DAKO Omnis, Agilent) before a 10-minute incubation in EnVision FLEX Substrate Working Solution (DAB) for staining. After further wash cycles, slides underwent a 3-minute incubation with haematoxylin ready to use solution (K8018, Agilent DAKO) before being rinsed and removed from the DAKO Omnis, dehydrated in alcohol, and cleared in xylene. Slides were mounted with Dibutylphthalate Polystyrene Xylene (DPX, Sigma-Aldrich) and coverslipped.
3,3′-Diaminobenzidine/HRP-stained slides were imaged using a 3DHistech Panoramic 250 Flash II slide scanner at 40x using the autofocus setting (SAHMRI Histology Slide Scanning Service). Images of individual sections from L6 to S2 regions were taken using digital pathology viewing software (QuPath 0.1.2) and analysed using ImageJ software. The images were not manipulated in any way.
2.8.5. Spinal cord phosphorylated MAP kinase ERK 1/2 neuronal counts
Neuronal counts were analysed from previously saved digital photomicrographs, with only neurons with intact nuclei counted. The number of pERK-immunoreactive (IR) dorsal horn neurons/animal was obtained from a minimum of 6 sections/animal/spinal segment (L6-S2). The total number of pERK-IR neurons from the dorsal horn overall and selectively from dorsal horn regions was then averaged across mice following 40 mm Hg pressures of VD after vehicle, TTX (0.5 µM), or veratridine (50 µM) vaginal infusion before balloon insertion. The selective dorsal horn regions analysed were defined as the superficial dorsal horn (SDH) (laminae LI-II), laminae LIII-V, dorsal gray commissure (DGC), lateral spinal nucleus (LSN), and the sacral parasympathetic nucleus (SPN, only in S1-S2 spinal segments) (Supplemental Figure 1, available at http://links.lww.com/PAIN/B146). Selection of the dorsal horn regions were on based on previous published work and the Allen Spinal Cord Atlas available from https://mousespinal.brain-map.org.42
2.8.6. Statistical analysis of phosphorylated MAP kinase ERK 1/2-immunoreactive
Ordinary one-way analysis of variance followed by Bonferroni multiple-comparison post hoc test was used to compare the number of pERK-IR neurons between different treatments in different regions of the dorsal horn. Differences were considered statistically significant at *P < 0.05, **P < 0.01, and ***P < 0.001. Averaged data values are expressed as mean ± SEM. Figures were prepared in GraphPad 7.02. software (San Diego, CA). N represents number of animals per group, whereas n represents the number of neurons or independent observations.
2.9. Visceromotor response to vaginal distension: assessment of vaginal pain sensitivity in vivo
The visceromotor response (VMR) is a nociceptive brainstem reflex consisting of the contraction of the abdominal muscles in response to noxious distension of hollow organs such as the vagina5,24 and the colorectum.11,13,16,30,38,49,58 We recorded the VMR to VD as an objective measurement of vaginal sensitivity to pain in fully conscious animals.2,5,12,24,46,48,51,52
2.9.1. Surgical implantation of electromyography electrodes
The VMR was objectively assessed by electromyography (EMG) to quantify abdominal muscle contractions in response to VD. To this end, EMG electrodes were implanted in the abdominal musculature at least 3 days before VMR assessment. Mice were anesthetised with isoflurane and a 1-cm incision was made just superior to the right inguinal ligament, exposing the external oblique abdominal muscle. Two Teflon-coated stainless-steel wires (Advent Research Materials Ltd, Oxford, United Kingdom) were sutured into the muscle approximately 5 mm apart. The electrodes were tunnelled subcutaneously and exteriorised at the base of the neck for future access. All mice received prophylactic antibiotics (Baytril 5 mg/kg s.c.) and pain relief (buprenorphine 0.05 mg/kg s.c.). After surgery, animals were single housed to protect the EMG electrodes. Animals were allowed to recover from surgery for at least 3 days before VMR assessment.
2.9.2. Assessing visceromotor responses to vaginal distension
On the day of VMR assessment, animals were briefly sedated with isoflurane, and vehicle (saline), veratridine (50 µM), or TTX (0.5 µM) was administered intravaginally through a small cannula inserted into the vaginal canal. Immediately after, a lubricated 3-mm length (∼4-mm diameter when fully inflated) latex balloon was gently passed through the vagina and inserted up to 1 mm proximal to the vaginal verge. Once in position, the balloon catheter was secured to the base of the tail and connected to a barostat (Isobar 3 Barostat; G&J Electronics, Toronto, Canada) for pressure-controlled rapid inflation of air. Animals were transferred to a restrainer with dorsal access, and the EMG electrodes were relayed to a data acquisition system. Animals were allowed to regain consciousness for at least 10 minutes before the distension sequence was initiated. Distensions were applied by the barostat in a pressure-controlled fashion, ranging from the nonnoxious to the noxious range (20-30-40-60-70-80 mm Hg, 30-second duration, 3-minute interval between consecutive distensions). The corresponding EMG signal was recorded (NL100AK headstage), amplified (NL104), filtered (NL 125/126, Neurolog, Digitimer Ltd, Hertfordshire, United Kingdom, bandpass 50-5000 Hz), and digitised (CED 1401, Cambridge Electronic Design, Cambridge, United Kingdom) to a PC for offline analysis using Spike2 (Cambridge Electronic Design).
2.9.3. Statistical analysis of visceromotor response to vaginal distension data
The analog EMG signal was rectified and integrated. To quantify the magnitude of the VMR at each distension pressure, the area under the curve (AUC) during the distension (30 seconds) was corrected for the baseline activity (AUC for 30-second predistension). Total AUC was quantified by adding the individual AUC at each distension pressure. Visceromotor response data are presented as mean ± SEM, and N represents the number of animals. Visceromotor response data were statistically analysed by generalised estimating equations followed by LSD post hoc test when appropriate using SPSS 23.0. Analysis and figures were prepared in GraphPad Prism 7 Software, San Diego, CA. Differences were considered statistically significant at *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.
3.1. Mechanosensory profile of pelvic sensory afferents innervating the female reproductive tract
The peripheral terminals of sensory neurons innervating visceral organs are equipped to detect a variety of mechanical stimuli.27,57 There is a great body of knowledge illustrating how mechanical stimuli is detected by visceral organs such as the colon9,32,66 and the bladder26,28,29; however, little is known about how mechanical stimuli is sensed by the female reproductive organs. To reduce this knowledge gap, we developed a novel ex vivo preparation to characterise the mechanosensory profile of pelvic afferents innervating the female reproductive tract of the mouse (Fig. 1).
We found that pelvic afferent nerve terminals had small (∼0.5 mm) punctate receptive fields from which action potentials were evoked in response to the application of mechanical stimuli. Receptive fields for these individual afferents were distributed throughout the length of the vagina (Fig. 2A). Overall, the majority of afferents recorded were silent at rest, with only 21.4% (3 of 14) of the afferents showing low rates of spontaneous activity.
To examine the mechanosensory properties of the nerve fibres from a particular receptive field, 3 distinct mechanical stimuli were tested. We examined the effect of (1) static focal compression of the receptive field with a calibrated 2 g vfh (Fig. 2B); (2) fine stroking of the vaginal mucosa surface with calibrated (10-1000 mg) vfh (Fig. 2C); and (3) 5 g circular stretch of the vagina, applied through a cantilever system (Fig. 2D). Interestingly, we found that the entire population of vaginal afferents tested (total of 14 afferents from 8 mice) responded to all 3 mechanical stimuli tested (Fig. 2E). We also found that fine stroking of the vaginal mucosal surface and static probing of the receptive fields readily evoked discharge of action potentials throughout the duration of the stimulus (Figs. 2B and C). Circular stretch elicited a wide dynamic range of responses with some afferents firing action potentials throughout the duration of the stimulus (maintained response), whereas others had a more intense discharge at the start followed by more sporadic firing throughout the duration of the stimulus (adapting response, Supplementary Figure 2A, available at http://links.lww.com/PAIN/B146). In addition, the latency, duration, and magnitude of the responses to circular stretch were quite heterogenous amongst the afferents examined (Supplementary Figure 2B–D, available at http://links.lww.com/PAIN/B146).
3.2. Mechanically evoked responses of pelvic vaginal afferents can be modulated by targeting voltage-gated sodium (NaV) channels
We have recently determined that NaV channels expressed within the cell bodies of sensory afferents that travel from the colon and bladder to the CNS are potential therapeutic targets for treatment of acute and chronic visceral pain.22,28,36,37,50,58 With this in mind, we used our novel ex vivo afferent recording preparation to determine whether pharmacological modulation of NaV channels could alter mechanosensory properties of pelvic afferents innervating the vagina.
Application of the pan-NaV channel agonist, veratridine, resulted in significant increases in pelvic vaginal afferent responses to focal compression (Fig. 3A), mucosal stroking (Fig. 3B), and circular stretch (Figure 3C; Supplementary Figure 3A, available at http://links.lww.com/PAIN/B146) compared with their normal baseline responses. Veratridine did not evoke action potential firing in the absence of mechanical stimuli and did not significant alter the latency nor the duration of the response of vaginal afferents to circular stretch (Supplementary Figure 3A, available at http://links.lww.com/PAIN/B146).
Exposure of vaginal afferent endings to the neurotoxin TTX (blocker of the TTX-S NaV1.1 − NaV1.4, NaV1.6, and NaV1.7 channels) significantly reduced, but did not completely block, vaginal afferent responses to focal compression (Fig. 3D), mucosal stroking (Fig. 3E), and circular stretch (Figure 3F; Supplementary Figure 3B, available at http://links.lww.com/PAIN/B146). TTX did not significantly alter the latency nor the duration of the response of vaginal afferents to circular stretch (Supplementary Figure 3B, available at http://links.lww.com/PAIN/B146). Veratridine and TTX changed vaginal afferent mechanosensory responses by more than 15% relative to the baseline in 6 of the 7 afferents tested (Supplementary Figure 3C, available at http://links.lww.com/PAIN/B146). Overall, these findings indicate that both TTX-S and TTX-R NaV channels are involved in vaginal afferent responses to mechanical stimulation.
3.3. Pharmacological modulation of NaV channels alters the excitability of dorsal root ganglia sensory neurons innervating the vagina
Given the significant effect of NaV channel modulation on altering pelvic vaginal afferent responses to mechanical stimuli, we investigated the relative expression of these channels within sensory neurons innervating the vagina, and whether pharmacological modulation of these channels could alter the electrophysiological properties of vagina-innervating DRG neurons.
First, we investigated whether NaV channels were indeed expressed by individual DRG sensory neurons innervating the vagina. Using single-cell RT-PCR, we determined mRNA expression of all members of the NaV channel family (NaV1.1-NaV1.9) in 39 individual vagina-innervating DRG neurons, identified by retrograde labelling. Overall, we found that all NaV channel family members were expressed throughout the population of retrogradely labelled vagina-innervating DRG neurons examined (Fig. 4). We found 3 general patterns of expression, with NaV channel transcripts expressed by either a large number (NaV1.7: 100%, NaV1.8: 97%), a moderate number (NaV1.9: 62%; NaV1.6: 59%; NaV1.2: 54%; NaV1.1: 33%), or a small number (NaV1.5: 26%; NaV1.3: 21%; NaV1.4: 13%) of vaginal-innervating DRG neurons (Fig. 4A). The expression and coexpression patterns of NaV channel transcripts were heterogeneous amongst neurons, with only a small proportion of cells expressing the same pattern of NaV channel transcripts with no cells expressing the full repertoire of NaV1.1 to 1.9 (Figs. 4B and C).
We also determined the mRNA expression profile of NaV channel transcripts within the vaginal tissue, with qRT-PCR revealing an overall very low level of abundance (supplemental Figure 4, available at http://links.lww.com/PAIN/B146). NaV1.7 was the most abundantly expressed NaV channel in vaginal tissue, whereas NaV1.2 to 1.6 had even lower abundance (Supplementary Figure 4, available at http://links.lww.com/PAIN/B146). NaV1.1, NaV1.8, and NaV1.9 expression in vaginal tissue was below detection limits (Supplementary Figure 4, available at http://links.lww.com/PAIN/B146). Our qRT-PCR data showed that NaV expression was ∼200- to 1000-fold lower in vaginal tissue than in whole lumbosacral DRG (Supplementary Figure 4, available at http://links.lww.com/PAIN/B146).
We next examined whether inhibition of TTX-sensitive NaV channels could alter electrophysiological properties of isolated vagina-innervating DRG neurons using whole-cell patch-clamp electrophysiology (Fig. 5). At baseline conditions in voltage-clamp mode, vagina-innervating DRG neurons exhibited large voltage-dependent sodium currents, which were significantly reduced in amplitude (by 40.9 ± 9.2%) after a 1-minute incubation with TTX (Figs. 5Ai–iv). Furthermore, in current clamp mode, the threshold for action potential generation (rheobase) was significantly increased (2.1 ± 0.2-fold) in the presence of TTX in 78% (35 of the 45) neurons examined (Figs. 5Bi–iv). These findings reflect a decrease in neuronal excitability of vagina-innervating DRG neurons when TTX-S NaV isoforms are blocked. Overall, these results indicate that modulating NaV channels expressed in sensory afferent neurons that innervate the female reproductive tract alters neuronal excitability, hence their ability to fire action potentials.
3.4. Vaginal distension-evoked activation of dorsal horn neurons within the spinal cord can be manipulated by intravaginal administration of NaV channel modulators
Following our findings of NaV channel modulation ex vivo and in vitro, we investigated whether changes in vaginal afferent sensitivity, induced by pharmacological modulation of NaV channels in the periphery, translate to changes in the nociceptive signal sent to the CNS in vivo. Therefore, we aimed to determine whether increasing or decreasing the peripheral sensitivity of vaginal afferents, induced by either veratridine or TTX, respectively, correspondingly altered neuronal activation within the spinal cord. In separate mice, we performed in vivo VD after infusion of either saline (vehicle control), veratridine, or TTX, and compared the number of phosphorylated-MAP-kinase-ERK-immunoreactive (pERK-IR) dorsal horn neurons in the lumbosacral (LS; L6-S2) spinal cord (Figure 6; Supplementary Figure 1, available at http://links.lww.com/PAIN/B146). After saline infusion, VD resulted in pERK-IR neurons being present within the SDH (laminae LI-II), throughout the deep dorsal horn (LIII-V), and in the DGC in relatively equal proportions (Figs. 6Ai–E; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146). A smaller number of pERK-IR neurons were present within the region of the LSN and the SPN in sacral spinal segments (Figs. 6Ai, and F–G; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146).
We then showed that the number of pERK-IR neurons evoked by VD is significantly increased by intravaginal administration of veratridine (Figs. 6Aii and 6B; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146) and decreased by TTX (Figs. 6Aiii and 6B; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146). Further analysis of the different regions of the dorsal horn showed that vaginal infusion of veratridine before distension resulted in significantly more pERK-IR dorsal horn neurons relative to saline infusion within the SDH (Fig. 6C; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146) and the LSN (Fig. 6F; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146). By contrast, compared with vehicle, veratridine treatment did not significantly alter the numbers of pERK-IR neurons within the deep dorsal horn (Fig. 6D, Supplementary Figure 5, available at http://links.lww.com/PAIN/B146), DGC (Fig. 6E; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146), or the SPN (Fig. 6G; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146). In comparison, VD after intravaginal application of TTX resulted in a significant decrease in the total number of pERK-IR dorsal horn neurons relative to saline infusion, with significant reductions specifically within the DGC (Fig. 6E; Supplementary Figure 5, available at http://links.lww.com/PAIN/B146).
These in vivo results indicate that veratridine-induced increases in the peripheral sensitivity of vaginal afferent endings lead to enhanced signalling within the spinal cord. Conversely, vaginal administration of TTX can reduce spinal cord neuronal activation evoked by VD. Interestingly, the changes in the number of pERK-IR neurons in response to veratridine mainly occurred within specific regions of the dorsal horn distinct from those affected by TTX.
3.5. Pain sensitivity to vaginal distension can be modulated by targeting NaV channels intravaginally
We next investigated whether the observed alteration in the signalling in the CNS translates to changes in pain sensitivity evoked by VD in conscious animals. To measure vaginal pain sensitivity in vivo, we measured the VMR to increasing VD pressures by recording electromyography (EMG) activity from electrodes surgically implanted into the abdominal muscles.
Vaginal distension evokes an increase in the VMR, and the degree of VMR is related to the amount of pressure applied to the vagina (Fig. 7). We found that mice administered intravaginally with veratridine displayed pronounced hypersensitivity to VD in vivo. This is indicated by significantly elevated VMRs to VD to all distension pressures compared to mice intravaginally treated with vehicle (Figs. 7A–C). Conversely, intravaginal administration of TTX significantly reduced VMRs to VD compared to mice intravaginally treated with vehicle, suggestive of an analgesic effect (Figs. 7A–C).
Taken together, these findings indicate that pharmacological modulation of NaV channels at the periphery alters sensory signalling at the afferent and spinal cord levels, ultimately translating to modulation of pain sensitivity evoked by VD in vivo.
In this study, we determine how pelvic afferents innervating the mouse vagina are tuned to detect a variety of mechanical stimuli. We also show how activation of these afferents leads to pERK-IR within dorsal horn neurons of the spinal cord and altered responses to VD in vivo. We also identify, for the first time, the complete expression profile of NaV channels within individual retrogradely traced vagina-innervating DRG neurons. Finally, we demonstrate that NaV channel modulators are able to alter 1) the responsiveness of pelvic vaginal afferents to mechanical stimuli; (2) the excitability of isolated vagina-innervating DRG neurons; (3) nociceptive signalling into the spinal cord; and ultimately (4) pain sensation evoked by VD in conscious animals.
The female reproductive tract is innervated by sensory afferents from the hypogastric, pelvic, and pudendal nerves.33,39,43,44,59,62,72,73,77 These afferents have been shown to fire action potentials in response to mechanical and chemical stimuli applied within the uterus, cervix, and vagina of anaesthetised rats.3,4,6,55 Remarkably, further studies examining the mechanosensory properties of these afferents and how their modulation alters the transmission of sensory information to the CNS are lacking.
In the current study, using a single-unit ex vivo recording preparation, we show that pelvic afferents innervating the mouse vagina are tuned to detect a variety of mechanical stimuli. Using a flat sheet preparation, where the vagina was opened longitudinally and pinned flat, we show that the receptive fields of pelvic afferent endings were found scattered throughout the whole length of the vagina. Receptive fields were distributed ipsilaterally to the pelvic nerve branch where the recordings were made. Characterisation of afferent mechanosensitivity showed that 100% of the afferents examined responded to all 3 mechanical stimuli tested, suggesting polymodality. We found that focal compression of the receptive field or gentle stroking of the mucosa surface with calibrated vfh evoked slowly adapting responses, with responses that start and end with the stimuli. Circular stretch of the vagina evoked either a sustained or moderately adapting response. In addition, we showed that baseline responses to stroking stimulation of the mucosa were graded, where action potential frequency increased with increased stroke intensity. Overall, the mechanosensory properties of pelvic afferents innervating the mouse vagina are similar to those found in rats by Berkley's studies ∼30 years ago.3,4,6,55
We also demonstrate, for the first time, a direct role for NaV channels in mediating afferent signalling from the vagina. Specifically, we showed that vaginal afferent mechanosensitivity, neuronal activation within the spinal cord, and pain responses to VD could all be significantly augmented by pan-NaV activation with veratridine. This suggests activation of NaV channels can drive vaginal pain transmission, which is consistent with observations in other visceral organs. For example, intravesical instillation of veratridine significantly enhances bladder afferent sensitivity to graded distension.28 Similarly, in the colon, pan-NaV activation with Pacific ciguatoxin (P-CTX-1) activates colonic afferents and correspondingly neurons within the spinal cord.36 Furthermore, accidental consumption of P-CTX-1 or veratridine in humans manifests as acute and severe abdominal pain.36,60,65
In the current study, we also show that TTX significantly reduces vaginal afferent function and neuroexcitability, whilst reducing NaV currents in vaginal-innervating DRG neurons. Correspondingly, this results in a reduced number of pERK-IR neurons within the dorsal horn of the spinal cord and reduces pain responses to noxious VD. Notably, TTX significantly reduced, but did not abolish these responses, implicating both TTX-S (NaV1.1 − NaV1.4, NaV1.6, and NaV1.7) and TTX-R (NaV1.5, NaV1.8, and NaV1.9) channels in this response.
Our single-cell RT-PCR data showed distinct and overlapping expression profiles of the various members of the NaV family within individual vagina-innervating neurons. Notably, the majority of the neurons expressed at least one TTX-S (NaV1.7) and one TTX-R (NaV1.8) isoform. This single-cell RT-PCR data indicate that the TTX-S component is most likely attributable to combinations of NaV1.1, NaV1.2, NaV1.6, and NaV1.7, which were found to be expressed in a high proportion of vagina-innervating DRG neurons. Although studies have yet to directly investigate a role for NaV1.1 or NaV1.6 in pelvic vaginal afferents, studies in the colon show that these 2 isoforms play key roles in signalling nociceptive information in response to mechanical stimuli.37,50,58 By contrast, NaV1.7 does not seem to contribute to either colonic or bladder nociception, at least in healthy conditions,35 whilst there are currently no functional data to support a role of NaV1.2 in visceral sensory signalling.22 For the TTX-R component, NaV1.8, and NaV1.9 are likely the main contributors because they are highly expressed in vagina-innervating DRG neurons and contribute to other forms of visceral pain.34,36 By contrast, the other TTX-R subunit NaV1.5 is poorly expressed in vagina-innervating DRG neurons. Furthermore, our QRT-PCR data identified extremely limited expression of NaV channel subtypes within the vaginal tissue, suggesting that the effects of veratridine and TTX on modulating afferent mechanosensitivity occur through an afferent mechanism, and not due to secondary changes within the vaginal tissue.
Our findings here regarding the TTX sensitivity and NaV isoform expression profile of vaginal-innervating DRG neurons contrast sharply with those in other visceral organs. For example, both the sodium current density of bladder-innervating neurons and bladder afferent responses to distension are almost completely abolished by TTX. Correspondingly, intrabladder TTX administration dramatically reduces the number of dorsal horn neurons being activated by bladder distension.28 These findings suggest a large TTX-S component and a small TTX-R component in bladder-innervating DRG neurons. By contrast, vaginal-innervating DRG neurons have a more balanced TTX-S and TTX-R contribution, which is likely borne out by fewer vaginal neurons expressing NaV1.1, NaV1.3, and NaV1.6 compared with bladder-innervating DRG neurons.28 Colon-innervating DRG neurons also demonstrate TTX-S and TTX-R components,7 although more of these neurons express NaV1.1, NaV1.2, NaV1.3, and NaV1.935 than vagina-innervating LS DRG neurons. These findings are important because the complete NaV channel expression profile in afferent neurons innervating female reproductive organs has not previously been reported, with only one study showing NaV1.8 expression in afferents innervating the mouse uterus.41 These findings provide novel alternatives to regulate sensation arising from these visceral organs, as discussed below.
In this study, we show that intravaginal administration of the pan-NaV activator veratridine increases vaginal sensory signalling into the spinal cord (measured by an increase of pERK-IR neurons) in response to vagina distension. Notably, veratridine-induced increases in pERK-IR neurons were localised to the SDH and the LSN. Both of these regions have significant roles in relaying nociceptive signalling into various pain processing centres of brain.63,68 Conversely, TTX reduced the number of pERK-IR neurons. However, this reduction occurred primarily within the DGC which, in sacral spinal regions, is known to be important to signalling visceral sensation within dorsal columns.74 These findings correspond well with our observation that in vivo intravaginal administration of veratridine significantly increases the VMR to VD, whereas TTX significantly reduced these responses compared to vehicle-treated mice. Interestingly, because alterations in the VMR to VD in animals treated with either veratridine or TTX occurred across all distension pressures, this suggests that NaV channels mediate the transmission of both nonnoxious and noxious stimuli.
It is well documented that women with endometriosis and vulvodynia experience dyspareunia twice as often as healthy women.61,78 It is also well known that pelvic afferents innervating other visceral organs, such as the colon and bladder, become hypersensitive to mechanical stimuli in animal models of irritable bowel syndrome13,15,16,56,58 and interstitial cystitis/bladder pain syndrome,26,47 which ultimately leads to chronic pelvic pain in humans. Whether the dyspareunia, experienced by women with endometriosis and vulvodynia, could be explained by vaginal afferents developing hypersensitivity to mechanical stimuli remains to be identified. However, this study provides novel findings advancing the understanding of vaginal sensation that can be used to recognise and explore changes in states of chronic pelvic pain associated with endometriosis and vulvodynia. As recent reports show the potential of targeting ion channels, including NaV channels, for the treatment of acute and chronic visceral pain,10–14,28, 34, 37, 50, 58 a similar strategy to treat pain arising from the female reproductive organs may be possible. This concept is supported by our findings, providing the first direct demonstration for a role of NaV channels in regulating vaginal sensation in vivo.
In conclusion, our findings demonstrate that modulation of the electrophysiological properties of afferents innervating the female reproductive tract, by targeting NaV channels expressed within these afferents, leads to changes in nociceptive signalling sent to the CNS and ultimately regulated vaginal pain sensitivity evoked by VD in vivo. These findings contribute towards the understanding of how mechanical stimuli is detected and transmitted from female reproductive organs and uncover potential molecular targets to investigate as novel therapeutics to manage dyspareunia associated with endometriosis and vulvodynia.
Conflict of interest statement
The authors have no conflicts of interest to declare.
Appendix A. Supplemental digital content
Supplemental digital content associated with this article can be found online at http://links.lww.com/PAIN/B146.
Supplemental video content
A video abstract associated with this article can be found at http://links.lww.com/PAIN/B147.
The authors thank Adelaide University Histology Services for performing the pERK labelling and SAHMRI Histology Slide Scanning Service for imaging services. J. Castro is funded by a National Health and Medical Research Council (NHMRC) of Australia Ideas Grant (APP1181448). S.M. Brierley is a NHMRC R.D. Wright Biomedical Research Fellow (APP1126378) and is funded by NHMRC Australia Project Grants #1083480, #1139366, and #1140297. A.M. Harrington received funding through the Australian Research Council (ARC) Discovery Early Career Research Award (DE130100223). A.M. Harrington and S.M. Brierley received funding by an ARC Discovery Project (DP180101395).
Author contributions: J. Castro, J. Maddern, A. Erickson, A.M. Harrington, L. Grundy, and S.M. Brierley contributed with the experimental design. J. Castro, J. Maddern, A. Erickson, and A. Caldwell performed the experiments described in this article. A.M. Harrington contributed to the design and analysis of the pERK studies. J. Castro made the figures. J. Castro and S.M. Brierley wrote the manuscript with contributions from all authors.
Previous presentation of research: Parts of this work has appeared in presentations at the Australian Pain Society (APS) conference in the Gold Coast, Australia, April 7–10, 2019.
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