TRESK channel contribution to nociceptive sensory neurons excitability: modulation by nerve injury

It is long known that peripheral axon injury produces sensory neuron hyperecitability [7, 26]. In order to study changes in background conductances contributing to this phenomena, we recorded a few neurons in L4-L5 DRG to confirm that they were hyperexcitable 3 weeks after sciatic nerve axotomy. Intracellular recordings in small-medium DRG neurons (soma: 26.6 ± 0.6 μm; range 22-30) in the excised intact ganglia showed an increased soma excitability measured as the number of axon potentials fired by a normalized 1-s depolarizing pulse 2.5× the 20 ms spike threshold current. Axotomized neurons fired more spikes (6.5 ± 2.4 spikes; mean ± SEM; n = 8) compared with contralateral uninjured neurons (1.9 ± 0.5 spikes; n = 8; p < 0.05, unpaired t-test; Figure 1A). In addition to the increased excitability, injured neurons had a higher R_in (624.5 ± 159.7 vs. 141.4 ± 47.8 MΩ; p < 0.001, unpaired t-test), without significant differences in resting membrane potential (RMP -54.9 ± 4.1 vs. - 55.9 ± 2.9 mV) or action potential threshold (0.62 ± 0.1 vs. 0.67 ± 0.3 nA). These results are in general agreement with previous reports of excitability of injured and uninjured sensory neurons in DRG ganglia [7, 10, 27, 28]. In parallel, we studied whether neuronal hyperexcitability produced by injury can be due to gene expression changes in background K_2P channels that are known to be expressed in DRG neurons [17, 29]. Injured DRG neurons did not show significant changes in the transcripts for TREK-1, TREK-2, TASK-1 or TRAAK (1.22 ± 0.23; 1.05 ± 0.07; 0.80 ± 0.06; 0.79 ± 0.23 fold-change, respectively) but displayed a significant reduction in TRESK transcripts (0.46 ± 0.07 vs. contralateral side; p < 0.001; Figure 1B; n = 4 independent animals). As previously described, injury increases the expression of the α2δ1 subunit of the voltage-dependent calcium channel [30], thus we used its expression as a positive control. In the same samples, the transcript for this subunit was increased by 5-fold when compared to the contralateral uninjured side (Cacna2d1: 5.00 ± 0.47; p < 0.001; Figure 1B). Sham surgery did not show significant changes in the expression of any of the tested genes when compared to the contralateral uninjured side (TREK-1: 0.83 ± 0.21; TREK-2: 0.83 ± 0.24; TASK-1: 0.87 ± 0.31; TRAAK: 0.89 ± 0.31; TRESK: 0.98 ± 0.36; Cacna2d1: 1.06 ± 0.14; n = 4; Figure 1B). When axotomized and sham animal groups where compared, only TRESK (p < 0.05) and Cacna2d1 (p < 0.001) showed significant differences on channel expression. Changes in opposing directions were seen when comparing the expression of TREK-1 in axotomized and sham animals (Figure 1B) that were close but not statistically significant (p = 0.09).

To further investigate the effect of injury on background channels, we compared whether "in vitro axotomy" produced by dissociation of DRG neurons rendered similar effects than in vivo axotomy (sciatic nerve transection). Despite the fact that dissociated DRGs may have several differences with neurons axotomized in the animal, others have reported that dissociation of DRGs produces neuronal hyperexcitability [11, 31], thus we asked whether similar changes underlie this hyperexcitable state. As expected, small and medium-sized neurons (soma: 23.2 ± 0.5 μm; range 16-27) axotomized in the animal and then dissociated (Axo+Diss group) were hyperexcitable (9.7 ± 3.1 spikes; n = 15; Figure 2A). However, uninjured but dissociated neurons (Diss group), also showed marked hyperexcitability (12.3 ± 3.7 spikes, n = 16), confirming that neuronal dissociation alone also induces a significant increase in cell excitability. No significant differences were obtained for R_in, RMP, AP amplitude, AP duration or AHP amplitude, although Diss neurons had a slightly lower action potential threshold compared with neurons in the Axo+Diss group (0.06 ± 0.01 vs. 0.15 ± 0.03 nA; p < 0.01; Figure 2A). To assess whether injury modifies total background currents in DRG neurons, we performed whole-cell patch clamp recordings in both groups of neurons using a ramp protocol and in the presence of 2 μM TTX. Current measurements at -110 mV (Axo+Diss: -2.7 ± 0.9 pA/pF; Diss: -4.8 ± 0.7 pA/pF; n = 7; Figure 2B) or at +50 mV (38.1 ± 6.2 and 39.7 ± 6.8 pA/pF; n = 7) did not show significant differences between groups. In addition, using a protocol to minimize activation of voltage-gated transient K⁺ outward currents [18], outward and inward currents measured at the end of the depolarizing step and the hyperpolarizing ramp (arrows; Figure 2C) also failed to elicit significant differences between Axo+Diss (-5.7 ± 2.3 pA/pF at -25 mV and 4.6 ± 1.0 at -135 mV; n = 7) and Diss neurons (-11.3 ± 2.5 pA/pF at -25 mV and 6.2 ± 1.9 pA/pF at - 135 mV; n = 5), despite the fact that Diss neurons appeared to have a slightly higher outward current. Because patch clamp recordings were done at room temperature (~22°C) in small- and medium-sized neurons were TRESK is preferentially and abundantly expressed [18, 29] and due to the fact that other background channels present in DRGs are mostly inactivated at this temperature [19, 32], the recorded background currents should be mainly carried by TRESK channels.

To correlate the effects of injury with the expression of background channels, we next tested an injury marker described to be highly up-regulated in DRG neurons [33‐37]. In in vivo axotomized neurons, ATF3 expression was increased by 25-fold compared with contralateral uninjured cells (Figure 3A; p < 0.001; n = 4 animals). In contrast, ATF3 expression did not show significant differences between Axo+Diss and Diss neurons (Figure 3B; n = 3 independent dissociated cultures), meaning that this transcript was now upregulated in both groups. Similarly, the differential expression of Cacna2d1previously found in the ganglia (Figure 1B) was greatly reduced, with no significant differences between groups (compare data on Figure 1B and 3B). In a similar fashion, the reduction in TRESK expression found in vivo was greatly diminished in the dissociated neurons, but a still significantly decreased expression was found between the Axo+Diss and the Diss group (0.72 ± 0.06; n = 4; p < 0.05; Figure 3B). All together, this data suggest that neuron dissociation produces similar changes to those caused by directly injuring the cells in the living animal, as clearly shown by the expression of injury markers Cacna2d1 and ATF3, and confirm downregulation of TRESK channel expression by nerve injury.

To determine the participation of TRESK in the excitability of sensory neurons, we took advantage of the reported blocking activity of alkylamides like hydroxy-α-sanshool on this channel [23, 24]. IBA, an alkylamide derivative [25, 38] was first tested on HEK293 cells co-transfected with rat TRESK and eGFP to verify its blocking activity on this channel. 100 μM IBA application blocked total background current by 19.6 ± 3.3% (p < 0.01; n = 6) and 69.5 ± 8.7% at 500 μM (p < 0.001; n = 7; Figure 4A, B), which returned to basal values after washing the drug. IBA did not show significant effects in control cells transfected with eGFP (Figure 4B). As previously described [39], lamotrigine 100 μM also blocked TRESK-mediated background currents by 46.9 ± 8.1% (p < 0.01; n = 6). In addition, IBA modified resting membrane voltage of TRESK transfected HEK cells (Figure 4C, D). IBA application produced a mean depolarization of 8.3 ± 3.2 mV (p < 0.05 vs. vehicle; n = 4) that returned to RMP after several seconds (Figure 4D). In a similar way, lamotrigine induced a transient depolarization of 11.2 ± 4.5 mV (p < 0.05; n = 4). We also tested the effect of IBA on the other major K_2P channels expressed in DRGs. At the maximum concentration assayed (500 μM), IBA produced an increase in TREK-1 and TREK-2 currents (32.1 ± 9.4% and 34.8 ± 15.8% respectively; p < 0.05; n = 5; Figure 4B) and did not show significant effects on TRAAK (0.33 ± 21.1%; n = 4).

To further explore these effects, IBA was tested on currents recorded from DRG neurons in culture. Total current was recorded in the presence of TTX and using a ramp protocol (as in Figure 2B). As shown in Figure 4E, bath application of 400 μM IBA blocked part of the total current (36.6 ± 10.1% at +45 mV; n = 6; p < 0.01 vs. basal current). The effect of this compound was also tested using a protocol as in Figure 2C. IBA blocked part of the current elicited by a depolarizing pulse to -25 mV (30.8 ± 7.5%; n = 6; p < 0.05 vs. basal current; Figure 4F) and induced a small but non-significant decrease at -135 mV (3.8 ± 3%; n = 6). In summary, experiments show that TRESK-mediated background currents are transiently blocked by IBA in a dose-dependent manner. In DRG neurons, native currents are also blocked by IBA, despite a possible potentiation of other K_2P channels. This suggests that the blocking effect of IBA on TRESK is more important than the activation of other channels, or that TRESK has a higher contribution to the background currents in DRGs.

We next tested whether IBA applied to cultured DRG neurons increased intracellular calcium by inhibiting K_2P channels, as demonstrated for hydroxy-α-sanshool [23, 24]. Simultaneous recording of membrane voltage and intracellular Ca²⁺ in DRG neurons showed that IBA application (100 μM) produces a transient membrane voltage depolarization (40.06 ± 4.16 mV) in 8 of 11 neurons tested (RMP -48.8 ± 1.5 mV) accompanied by an increase in intracellular Ca²⁺ (1.79 ± 0.14 Fluorescence ratio R/Ro; Figure 5A). Action potential firing was also visible in these neurons upon membrane voltage depolarization (8.5 ± 3.3 spikes; Figure 5A, inset). Three neurons did not modify their membrane voltage or intracellular calcium in response to IBA application. These results confirm that inhibition of K_2P by IBA depolarizes membrane potential and activates Ca²⁺ entry through voltage-dependent Ca²⁺ channels. Intracellular calcium recordings showed that 100 μM IBA activated 49.3% of DRG neurons with a mean Ca²⁺ peak of 1.81 ± 0.06 (n = 101), while the vehicle used to dissolve the drug did not produce significant effects (Figure 5B and 5C). Among the neurons sensitive to IBA, 71.1% also responded to 1 μM capsaicin (R/Ro: 2.05 ± 0.09; n = 57), thus, as illustrated in experiments shown in Figure 5C, we identified three different subsets of neurons: 1) those selectively responding to IBA, 2) those responding only to capsaicin and 3) neurons that responded to both compounds. IBA-sensitive and non-sensitive DRG neurons showed a similar soma distribution (Figure 5D). 81.1% of the IBA-sensitive cells had soma diameters smaller than 30 μm, corresponding to small- and medium-sized DRG neurons. In the IBA-insensitive group, 77.9% of the neurons had somas <30 μm. According to this, 55.8% of small- and medium-sized neurons (<30 μm; 63/113 neurons) responded to IBA, while in large diameter neurons, the percentage of response to IBA was 41.9% (13/31 neurons). In summary, these data show that an important subpopulation of IBA-sensitive neurons are small diameter neurons (mainly nociceptors), most of them expressing TRPV1 (~70%) but not all (~30%).

A total of 45 C-nociceptor units with good signal to noise ratio were recorded from the sciatic nerve of 6 rats. Of these, 24 were classified as mechano-sensitive and 21 as mechano-insensitive C-nociceptors (Type 1A and 1B of [40], respectively). Intracutaneous injection of IBA (2 μl of 1% IBA) with a 26 Gauge needle induced abundant bursts of ongoing spontaneous activity in 11 nociceptor units, all of them belonging to the mechano-insensitive class, which represents a 52.3% of all the recorded mechano-insensitive C-nociceptors (Figure 6). The time to activation of the units was variable, ranging from immediately after the injection (Figure 6B, unit in red), to several minutes after it (Figure 6B, units in blue and grey). Surprisingly, IBA injection did not activate any of the mechano-sensitive units, despite the fact that some of them had short-lasting response to the needle insertion, indicating that the injection site coincided with their receptive field (not illustrated). This finding is highly suggestive of a selective effect of IBA on the mechano-insensitive C-nociceptor class.

Previous reports have shown that injection of IBA activates wide-dynamic range spinal neurons that receive convergent input from nociceptors thus implying that inhibition of K_2P channels by alkylamides can trigger painful sensations [25]. In contrast, hydroxy-α-sanshool topically applied to the hindpaw failed to elicit any flinching or guarding behaviors [24]. Since our data shows that IBA activates a subset of small- and medium-sized sensory neurons that are thought to be involved in nociception, we next asked whether injection of IBA in the hindpaw evokes any nocifensive behavior. We injected 2 μl of 0.1% or 1% IBA and recorded the flinching and licking behavior during 10 min after the injection. At 0.1%, IBA produced a significant increase in flinching (16.6 ± 5.9 flinches; n = 14; p < 0.05) compared to vehicle injection (1.73 ± 0.5; n = 11; Figure 7A), showing a higher increase in the initial minutes after injection and declining towards baseline after 6 min (Figure 7B). The same concentration also produced a significant increase in licking behavior (4.4 ± 1.4 licks; n = 14; p < 0.05) compared with vehicle (0.18 ± 0.13; n = 11) and showed a similar temporal pattern. The effect was more robust with 1% IBA, with mean values of 46.9 ± 13.7 flinches (n = 11; p < 0.01; Figure 7A, B) and 20.3 ± 6.6 licks (n = 11; p < 0.01; Figure 7A, C). This time, the effect was more sustained and both flinching and licking behaviors lasted for more than 10 min (Figure 7B, C). Abundant guarding behavior was also observed during this time period (not shown).

In a different set of animals, threshold of evoked mechanical pain in response to von Frey hairs was measured before and 2 min after hindpaw injection of 1% IBA (Figure 7D). Vehicle injection did not produce any significant change in mechanical pain threshold (from 15 ± 0 to 13.7 ± 1.2 g; n = 8). In contrast, 1% IBA produced a marked decrease in mechanical pain sensitivity (13.4 ± 1.1 to 4.25 ± 1.8 g; n = 8), which was statistically significant compared with the basal value (p < 0.001) or with the control group (p < 0.001). All together, the results obtained suggest that peripheral blocking of background K⁺ currents produces nocifensive behaviors in the animal.

We used siRNA to reduce TRESK expression in lumbar DRG neurons in order to confirm the implication of TRESK channels in pain modulation. In vivo TRESK silencing led to a mRNA decrease in lumbar DRGs comparable to that found after axotomy (42.5% mean decrease in mRNA level; silenced vs. the group injected with control siRNA; data not shown; n = 10). When mechanical sensitivity was studied, paw withdrawal thresholds did not show significant differences in the baseline values (Pre) between groups. After siRNA injections, paw withdrawal threshold was unaltered in animals injected with the control siRNA (from 21.4 ± 1.5 to 22.3 ± 0.7 g; n = 6; Figure 8A). In TRESK-silenced animals, paw withdrawal threshold was significantly decreased (18.9 ± 0.8 to 15.1 ± 0.8; n = 10) when compared with the baseline level previous to siRNA injection (p < 0.01) or with the control group after control siRNA injection (p < 0.001). In contrast, paw withdrawal threshold to thermal stimulation did not significantly change neither in the control (from 14.4 ± 1.0 to 13.9 ± 1.3 s; Figure 8B) nor in the silenced group (15.0 ± 1.0 to 12.7 ± 1.2 s), despite a small tendency to decrease in the latter, suggesting that reduced TRESK expression may have a greater effect on the detection of pain in response to mechanical stimuli.

All experimental procedures were carried out in accordance with the recommendations of the International Association for the Study of Pain (IASP) and were reviewed and approved by the University of Barcelona Animal Care Committee (Ref. 5336, 5406). Adult male Sprague-Dawley rats (Harlan; 100-150 g) were kept at 22°C with free access to food and water in an alternating 12 h light and dark cycle. Rats were anesthetized with isoflurane and a small incision in the skin was made to separate the muscle and expose the sciatic nerve, that was transected proximal to the bifurcation into the tibial and peroneal divisions as previously described [6, 7]. To avoid nerve regeneration, a 3 mm segment of the nerve was removed. The same procedure was performed in sham animals without transecting the nerve. To prevent foot mutilation, Mordex^® (Lab. URGO, Hernani, Spain) was applied to the operated foot. Daily inspections on operated animals were done to observe possible autotomy, which was scored according to the scale described by Wall et al. [66]. None of the animals used in the study attained a score of more than 5. After surgery, animals were kept for 3 weeks to allow the development of neuronal hyperexcitability due to axotomy. After this period, animals were anesthetized with isofluorane, killed by decapitation and DRGs (L4 and L5) from injured and contralateral uninjured sides were removed for neuronal culture or for RNA extraction.

L4 and L5 DRG were collected in cold phosphate buffered saline (PBS) with glucose, cleaned with iridectomy scissors under an stereoscopic microscope and incubated in phosphate buffered saline (PBS, Sigma) supplemented with 10 mM glucose, 10 mM Hepes, 100 U.I./mL penicillin, 100 μg/mL streptomycin, and collagenase type IA (4-5 mg/ml, Sigma) for 60 min at 37°C with gentle shaking. Digested ganglions were gently triturated with head-polished Pasteur pipettes; collagenase was inhibited by adding the solution to 10 ml of Dulbecco's Modified Eagle's Medium (DMEM) containing 10% fetal bovine serum and the mixture was centrifuged at 1000 rpm for 5 min. The pellet was suspended in DMEM plus 10% fetal bovine serum, 100 mg/mL L-glutamine, 100 U.I./mL penicillin, 100 μg/mL streptomycin and the suspension was plated on glass coverslips treated with poly-L-lysine/laminin and placed in culture dishes in an incubator at 37°C and 95% air, 5%CO₂. No NGF or other growth factors were added. Cells were used for electrophysiological recording within 3-48 h of plating.

DRG neurons obtained as described above were plated on 25 mm diameter glass coverslips (VWR Scientific Inc., Philadelphia, PA) and used 24-48 h thereafter. Cells were loaded with 5 μM fura-2/AM (Calbiochem, San Diego, CA) for 45-60 min at 37°C in incubation buffer (140 mM NaCl, 4.3 mM KCl, 1.3 mM CaCl₂, 1 mM MgCl₂, 10 mM glucose, 10 mM HEPES, at pH 7.4 with NaOH). Coverslips with fura-2 loaded cells were transferred into an open flow chamber (1 ml incubation buffer) mounted on the heated stage of an inverted Olympus IX70 microscope using a TILL monocromator as a source of illumination. Pictures were taken with an attached cooled CCD camera (Orca II-ER, Hamamatsu Photonics, Japan) and were digitized, stored and analyzed on a PC computer using Aquacosmos software (Hamamatsu Photonics, Shizuoka, Japan). After a stabilization period, image pairs were obtained alternately every 4 s at excitation wavelengths of 340 (λ1) and 380 nm (λ2; 10 nm bandwidth filters) in order to excite the Ca²⁺ bound and Ca²⁺ free forms of this ratiometric dye, respectively. The emission wavelength was 510 nm (120-nm bandwidth filter). Typically, 5-10 cells were present in a field and [Ca²⁺]_i values were calculated and analyzed individually for each single cell from the 340- to 380-nm fluorescence ratios at each time point. Several experiments with cells from different primary cultures were used in all the groups assayed.

For each animal, RNA from L4-L5 DRG pairs (axotomized and contralateral control) was extracted with Trizol (Sigma, Madrid) and first-strand cDNA was transcribed using the RETROscript kit (Ambion). qPCR experiments were performed in an AbiPrism 7300 using the TaqMan Universal PCR MasterMix (Applied Biosystems) with primers obtained from TaqMan Gene Expression assays: Rn00597042_m1 (TREK-1); Rn00576558_m1 (TREK-2); Rn00583727_m1 (TASK-1); Rn00587450_m1 (TRAAK); Rn99999916_s1 (GADPH); Rn00563784_m1 (ATF3); Rn00563853_m1 (CaCna2d1). A Custom Taqman Gene expression assay (Applied Biosystems) was designed for rat TRESK using the following primers: forward: TGCACAGTGTTCAGCACAGT; Reverse: CATATAGCATGCACAGGAACTTACC. Amplification of GADPH transcripts was used as a standard for normalization of all qPCR experiments and gene fold-expression was assessed with the ΔΔC_T method ipsilateral vs. contralateral side. Experiments were performed in quadruplicate.

Electrophysiological recordings were performed with a patch-clamp amplifier (Axopatch 200B, Molecular Devices, Union City, CA) and restricted to small and medium DRG neurons (<30 μm; <45 pF), which largely correspond to nociceptive neurons. Patch electrodes were fabricated in a Flaming/Brown micropipette puller P-97 (Sutter instruments). Electrodes had a resistance between 4-7 MΩ when filled with intracellular solution (in mM): 97.5 K⁺-gluconate, 32.5 KCl, 1 MgCl₂, 5 EGTA, 10 HEPES at pH 7.2 and 300 mOsm/Kg. An artificial cerebrospinal fluid (ACSF) was used as bath (in mM): 125 NaCl, 2.5 KCl, 0.5 CaCl₂, 2.5 MgCl₂, 1.25 NaH₂PO₄, 26 NaHCO₃, 10 glucose, 2 Na-piruvate, 3 myo-inositol, 0.5 ascorbic acid at pH 7.4 and 310 mOsm/Kg. Membrane currents were recorded in the whole-cell patch clamp configuration, filtered at 2 kHz, digitized at 10 kHz and acquired with pClamp 9 software and in the presence of 2 μM TTX. Data was analyzed with Clampfit 9 (Molecular Devices) and Prism 4 (GraphPad Software, Inc., La Jolla, CA). Series resistance was always kept below 30 MΩ and compensated at 70-80%. All recordings were done at room temperature (22-23°C). When studying the excitability of neurons in culture, after achieving the whole-cell configuration, the amplifier was switched to current-clamp bridge mode. Only neurons with a resting membrane voltage below -50 mV were considered for the study. To study neuronal excitability, we examined the resting membrane potential (RMP); action potential (AP) current threshold elicited by 20 ms depolarizing current pulses in 0.05-0.1 nA increments; whole-cell input resistance (R_in) was calculated on the basis of the steady-state I-V relationship during a series of 100-ms hyperpolarizing currents delivered in steps of 0.01-0.02 nA from 0.2 to 0.1 nA; AP amplitude (measured from RMP to AP peak; AP duration (measured at 50% AP amplitude); AHP (measured from RMP to peak hyperpolarization). Repetitive discharge was measured by counting the spikes evoked by 1-s, intracellular pulses of depolarizing current normalized to 2.5 times the AP threshold current. In some experiments (Figure 5A), calcium imaging and recording of membrane voltage was simultaneously performed. Cells were loaded first with fura-2 as described previously. Next, the whole-cell patch clamp configuration was achieved and the amplifier was switched to current-clamp bridge mode to record membrane voltage. Despite a slow decrease of fura-2 fluorescence values due to cell content dialysis, the short duration of the recording and the ratiometric measurement with fura-2 compensated for this effect.

Intracellular recordings in the excised ganglion were performed using an Axoclamp2B amplifier (Molecular Devices, Union City, CA) in the bridge-mode configuration. Axotomized or control intact L4 or L5 ganglia were treated with collagenase IA 4 mg/ml for 30 min at 37°C and then transferred to a recording chamber mounted in the stage of an upright BX50-WI microscope (Olympus, Japan). The ganglion was fixed with a nylon mesh that allowed the passage of the recording electrode through the mesh fibers. Pipettes filled with 3M K⁺-acetate had a resistance of 80-130 MΩ. Recordings were performed in the ganglion bathed in artificial cerebrospinal fluid (ACSF) solution at room temperature (22-23°C). Only neurons that had a resting membrane potential below -50 mV and a R_in of more than 50 MΩ were included in the study.

HEK293T cells cultured in DMEM with 10% FBS were seeded in 35-mm dish 24 h before transfection. Cells were transiently transfected with pEGFP vector alone (control) or cotransfected with: rTRESK-pcDNA3.1 (kindly provided by Dr. S. Yost, University of California-San Francisco), pCD8-mTREK-1, pCD8-hTREK-2 or pCD8-mTRAAK (kindly provided by Dr. F. Lesage, Institut de Pharmacologie Moléculaire et Cellulaire-CNRS, Valbonne, France) using FuGene transfection reagent (Roche). Transfected cells were used for electrophysiological recordings 24-48 h after. Patch clamp recordings were performed as described above. For whole-cell experiments, the solutions used were as follows. Intracellular solution (in mM): 140 KCl, 2.1 CaCl₂, 2.5 MgCl₂, 5 EGTA, 10 HEPES at pH 7.3. Bath solution (in mM): 145 NaCl, 5 KCl, 2 CaCl₂, 2 MgCl₂, 10 HEPES at pH 7.4. Cells were continuously superfused with a microperfusion system during the experiments, which were done at room temperature. When studying the membrane potential, after achieving the whole-cell configuration, the amplifier was switched to current-clamp bridge mode.

Rats were injected with 2 μl of a solution containing 0.1 or 1% IBA or propylene glycol (control vehicle) administered intradermally in the hindpaw using a 10 μl, 26 g Hamilton syringe. The rat behavior was observed and the number of flinching and licking of the paw was recorded every minute for a 10 min period starting immediately after the injection.

To assess mechanical sensitivity, the withdrawal threshold to punctate mechanical stimuli of the hindpaw was determined before and 2 min after 1% IBA injection in the hindpaw (as previously described) by the application of calibrated von Frey filaments (North Coast Medical, Inc. Morgan Hill, CA). The von Frey filaments [3.92, 5.88, 9.80, 19.60, 39.21, 58.82, 78.43, and 147.05 mN; equivalent to (in grams) 0.4, 0.6, 1, 2, 4, 6, 8, and 15] were applied vertically to the plantar surface of the hindpaw and gently pushed to the bending point. The 50% withdrawal threshold was determined using the up-down method as previously described [67]. A brisk hindpaw lift in response to von Frey filament stimulation was regarded as a withdrawal response.

Microneurographic recordings were obtained from 6 Spague-Dawley male rats (weight 200-250 g) anesthetized with ketamine (90 mg/kg) and xylacine (10 mg/kg) injected intraperitoneally. The sciatic nerve was exposed at mid-thigh level and intraneural recordings were performed according to the method recently described in detail elsewhere [57]. In brief, tungsten microelectrodes (200 μm diameter, lacquer-insulated, nominal impedance 1MΩ) were inserted into the sciatic nerve trunk with the aid of a micromanipulator. A subcutaneous reference electrode was inserted outside the nerve trunk. The neural signals were amplified with an isolated, high input impedance amplifier (3+ MicroAmp, FHC, USA), bandpass filtered (maximum range 50-5,000 Hz) and fed to a noise eliminator (Hum Bug, Quest Scientific, North Vancouver, Canada). This signal was then fed to a digital audio-monitor (AM10 audio monitor, Grass Technologies, Astro-Med, Inc., USA). Temperature of the skin was measured with a thermocouple placed on the skin adjacent to the receptive fields of the units under study. Electrical stimuli were triggered, and the responses to electrical stimulation recorded and analyzed with a PC and data acquisition board (National Instruments, PCI-6221, USA). The digitized responses were stored on the hard drive of the PC as raw data for offline analysis. Digital filtering (band pass 0.3-2 kHz) and clamping of the baseline were performed both on-line and during off-line analysis for a better visualization of the action potentials.

Responses were recorded with QTRAC software (^©Institute of Neurology, London, UK), using the facility to determine multiple peak latencies and display them as latency "profile" or raster plot. In the latency raster plots, each peak in the filtered voltage signal that exceeded a specified level is represented by a dot on a plot with latency as the ordinate and elapsed time as the abscissa. Depending on the level chosen, the dots could represent action potentials or noise. For the raster figures shown in this paper, latencies of selected units with adequate signal-to-noise were remeasured from the raw data, so that each dot represents an identified single unit. These remeasured figures are referred to as "modified" raster plots. Activity-dependent slowing of C fibers was assessed using the protocol described by Serra et al. [40, 68]. This consists of a sequence of: 1) baseline stimulation at 0.25 Hz for 3 minutes; 2) 3-min pause; 3) 6-min at 0.25 Hz; 4) 3-min 2 Hz train; 5) return to 0.25 Hz baseline until the latencies return to their original values (see Figure 6). This method differentiates "profiles" of activity-dependent slowing in individual C fibers of humans and rats that correspond to specific functional types of peripheral neurons [40, 68‐70]. Fibers slowing more than 10% during 3-min 2 Hz trains are nociceptors (Type 1 fibers in [68]). Further classification of C-nociceptor type into mechano-sensitive and mechano-insensitive units was achieved by studying the degree of slowing at very low frequencies after a pause [40, 71].

siRNA targeting rat TRESK (s175514; Sense AGAGAUUGGUUGCUCGAGAtt; Antisense UCUCGAGCAACCAAUCUCUca) and siRNA Negative Control #1 (4390843) were purchased from Applied Biosystems (Silencer^® Select Pre-designed siRNA) and injected in rats by intrathecal bolus to the lumbar region of the spinal cord once a day for 3 days. Each 10-μl injection corresponded to 2 μg of siRNA complexed with in vivo-jetPEI transfection reagent (Polyplus-transfection SA, Illkirch, France) following the supplier's suggested protocol. The specificity of the effect was evaluated in lumbar dorsal root ganglia by qPCR. On the same day of the behavioral tests (24 h after the last siRNA injection) L4 and L5 ganglia were removed and immediately frozen in liquid N₂. Later, RNA was extracted, cDNA was prepared and used for qPCR using the same primers and following the procedure previously described.

Heat sensitivity of adult male Sprague-Dawley rats (Harlan) was assessed by measuring hindpaw withdrawal latency from radiant infrared source (Hargreaves Method) using a Ugo Basile (Italy) Model 37370 Plantar test. Rats were acclimated to the experience room for at least 30 min and each measurement was the mean of 5 trials. Withdrawal latency was measured 24 h before and 24 h after the last intrathecal injection. Mechanical sensitivity was assessed by measuring hindpaw withdrawal latency with a Dynamic Plantar Aesthesiometer (37450; Ugo Basile, Italy). Incremental force (0-50 g in 40 s ramp) was applied with a 2 mm diameter metal rod to the paw plantar side. When the rat withdrew its paw, mechanical stimulus stopped automatically and time (s) and force (weight in grams) of paw withdrawal was recorded. Paw withdrawal responses were the average of 5 measures. The same experimenter for a given test performed all behavioral experiments, which was blind to the treatment applied to the animal. All measurements were done in a quiet room, taking great care to minimize or avoid discomfort of the animals.

Isobutylalkenyl amide (IBA) was kindly provided by Givaudan (Cincinnati, OH) and initially diluted in dimethylformamide (DMF) at a 100 mM. For in vitro experiments, IBA stock was subsequently diluted in the appropriate medium and used at the stated concentrations (100-500 μM), similarly to what has been reported for hydroxy-α-sanshool [23]. Final concentration of DMF was 0.1% or below. For in vivo experiments (hindpaw injections), IBA was diluted in propylene glycol and used at 1 or 0.1% as previously reported [25]. Lamotrigine, tetrodotoxin (TTX) and capsaicine (Cap) were purchased from Sigma (Madrid, Spain).