Surgical procedures for producing the radicular pain model
The animals were randomly divided into 3 experimental groups: the RC group, sham operation group, and naïve group. The surgical procedures have been described in detail previously [
39]. In brief, after the rats were anesthetized with intraperitoneal sodium pentobarbital (50 mg/kg), they were placed in the prone position, and a midline dorsal incision was made in the lower back to expose the laminas between the L4 and S1 vertebrae. The paraspinal muscles were retracted to expose the right L5–L6 facet joint. A right L5 hemilaminectomy and an L5–L6 partial facetectomy were performed. In the RC group, the right L5 spinal root was carefully exposed and tightly ligated extradurally by using 8–0 nylon sutures proximal to DRG. The subcutaneous tissue and skin were then closed by using 4–0 nylon sutures. In the RC group, 78 rats (n = 15 for the behavioral studies, n = 63 for the electrophysiological studies, as described below) and in the sham operation group, 62 rats (n = 15 for the behavioral studies, n = 47 for the electrophysiological studies, as described below) underwent the surgical procedure as described above without nerve root constriction. In the naïve group, 80 rats (n = 15 for the behavioral studies, n = 65 for the electrophysiological studies, as described below) were used with any surgical procedures.
Evaluation of mechanical hypersensitivity
To assess the mechanical withdrawal response, the rats were placed in a Plexiglas chamber (IITC Life Science Inc., Woodland Hills, CA, USA, measuring 18 × 25 × 18 cm above a wire mesh floor, which allowed full access to the hind paw, and were acclimatized to the environment for at least 20 minutes before the test. A 3.2-g von Frey filament (Semmes–Weinstein Monofilaments, North Coast Medical Inc., San Jose, CA, USA) was used to produce mechanical tactile stimuli, which were applied to the middle area between the foot pads on the plantar surface of the right (constriction side) and left (contralateral side) hind paws. Each hind paw was probed consecutively by 10 stimulations alternating between the right and left for each set. This was repeated thrice at intervals of at least 10 minutes [
39,
40]. Mechanical sensitivity was evaluated as the response rate of withdrawal responses. Visible lifting of the stimulated hind limb was considered to be a withdrawal response. This procedure was performed 1 day before and 3, 5, 7, 10 and 14 days after surgery.
Evaluation of thermal hyperalgesia
To assess the thermal withdrawal response, the rats were placed in a Plexiglas chamber on a glass platform and allowed to acclimatize for at least 20 minutes before the test. The thermal withdrawal response was measured as the latency of hind paw withdrawals elicited by a radiant heat source (Tail Flick Analgesia Meter, IITC Life Science Inc.), which was moved beneath the portion of the hind paw that was flush against the glass, and thermal stimulation was delivered to that site. Thermal hyperalgesia was evaluated as thermal withdrawal latency, which was defined as the time from the onset of radiant heat to the withdrawal of the tested foot. A cutoff time of 10 seconds was set to prevent tissue damage. Each hind paw was tested 5 times with at least 5-minute intervals, alternating between the right and left. The mean withdrawal latency was calculated from the last 4 measurements. Consequently, the thermal withdrawal latency of each rat was defined as the differential score, which was calculated by subtracting the withdrawal latency of the injured side from that of the noninjured side. Positive and negative scores indicated increased and decreased sensitivity of the ipsilateral hind paw, respectively. This test was performed after evaluating mechanical sensitivity and hyperalgesia.
Effects of the selective TRPA1 antagonist on pain-related behavior
In 15 days after surgery (within 24 hours after inserting the catheter), we performed the von Frey test described above for confirming that the catheter-implanted rats showed no evident motor disturbances such as paralysis. A selective antagonist for TRPA1, HC-030031 (Sigma, St Louis, MO, USA), was used to evaluate the involvement of TRPA1 in mechanical hyperalgesia. The rats were treated with HC-030031 (10 μg/site) or with vehicle [10% dimethylsulfoxide (DMSO) in saline] intrathecally. The mechanical hypersensitivity and hyperalgesia were evaluated using the von Frey test and pin-prick test described above at different time points after HC-030031 treatment (5, 15, and 30, 60, 120, 240 and 480 minutes), and spontaneous pain was estimated using the spontaneous pain scale described above at different time points after HC-030031 treatment (30 minutes and 2 hours). For the von Frey test and pin-prick test, the effects of HC-030031 was assessed as the response rate. Furthermore, the effect was compared with the intrathecal injection efficacy, which was calculated by dividing the withdrawal frequency after HC-030031 treatment by the baseline frequency.
Electrophysiological procedures
We performed
in vivo patch-clamp recording according to a previously described method [
2,
4,
45‐
48]. The rats were used 11–14 days after surgery when mechanical hypersensitivity and thermal hyperalgesia had fully developed. The examiner was blinded to the 3 groups. The rats were supplied oxygen through a nose cone under anesthesia with intraperitoneal urethane (1.2–1.5 g/kg). Rectal temperature was maintained at 37–38°C using a heating pad. Laminectomy was performed from the T13– L2 levels, after which the rats were placed in a stereotaxic apparatus (Model ST-7, Narishige Group Co, Tokyo, Japan), and the spinal cord was carefully exposed. After removing the dura, the right L5 nerve root was observed microscopically as a swollen area with a darker color than the left L5 and right L4 nerve roots. The pia-arachnoid membranes of the right L5 dorsal root entry zone were cut to create a window large enough to insert the patch electrode. The surface of the spinal cord was irrigated with 95% O
2 5% CO
2-equilibrated Krebs solution (10–15 ml/min) (in mM): 117 NaCl, 3.6 KCl, 2.5 CaCl
2, 1.2 MgCl
2, 1.2 NaH
2PO
4, 11 glucose, and 25 NaHCO
3 at 37.0 ± 0.5°C.
The patch electrodes were pulled from thin-walled borosilicate glass capillaries (outer diameter of 1.5 mm, TW150F-4, World Precision Instruments, Sarasota, FL, USA) by using a puller (p-97, Sutter Instruments, Novato, CA, USA) and were filled with patch-pipette solutions composed of the following (in mM): 135 K-gluconate, 5 KCl, 0.5 CaCl
2, 2 MgCl
2, 5 EGTA, 5 ATP-Mg, 5 HEPES–KOH (pH 7.2) for the EPSC recordings and current-clamp recordings. The recording electrode (resistance range, 8–12 MΩ) was advanced at an angle of 30° into SG. After a gigaohm seal was formed with neurons at a depth of 50–150 μm from the surface of the spinal cord, the membrane patch was ruptured by negative pressure, and then the whole-cell patch-clamp recording was initiated. A patch-clamp amplifier (Axopatch 200B, Axon Instruments, Union City, CA, USA) was used to make recordings. In the voltage-clamp mode, the holding potentials (V
H) were -70 mV, at which glycine-mediated and gamma-aminobutyric acid-mediated inhibitory postsynaptic currents were negligible [
49]. Spontaneous excitatory postsynaptic potentials and action potentials were recorded in the current-clamp mode. Spontaneous and evoked EPSCs were recorded as described later. Whole-cell patch-clamp recordings were made from SG neurons at the L5 segment of the spinal cord in the RC group, sham group, and naïve group. Data from neurons with resting membrane potentials more positive than -50 mV were excluded from this study.
After the electrophysiological experiment, some neurons that have been injected with biocytin during the recording were confirmed to be located in SG as described previously [
2,
4,
45,
47]. An A/D converter (Digidata 1322A, Axon Instruments) was used to digitize the data, which were analyzed by using the Mini Analysis Program version 6.0.7 (Synaptosoft, Fort Lee, NJ, USA).
In the voltage-clamp mode, all SG neurons produced large amplitude (>50 pA) barrages of EPSCs to non-noxious or noxious stimuli, such as a paintbrush or toothed forceps to the skin of the lumbar and pelvic regions or the hind paw [
2,
4,
45,
46]. A paintbrush or toothed forceps applied to the skin of the abdominal wall, lumbar and pelvic regions, hind paw, and tail was used to carefully determine the receptive field of a neuron after repeated stimuli. If evoked EPSCs were not observed during stimulation, that neuron was defined as one with no receptive field.
After determining the receptive field, the responses to non-noxious stimuli were assessed using a puff of air (20 psi) through a pipette (outer diameter of 200 μm) applied repetitively to the center of the receptive field (duration of injection, 5 seconds) using a pico-injector (PLI-100, Harvard Apparatus, Holliston, MA, USA), according to a previously described method [
4,
47]. The air-puff stimuli did not cause pain to the examiners. The noxious pinch stimuli were applied with toothed forceps that were firmly fixed on a rod. A weight (60 g) was placed on the forceps for 5 seconds, as referred to in previous reports [
4,
45,
47]. SG neurons exhibited several response profiles during air-puff or pinch stimuli in the current-clamp mode [
50,
51], which have been described in detail previously [
4,
47]. If the recorded neurons responded maximally to non-noxious stimuli enough to reach the action potential threshold, they were excluded in the present study because they may have been located in the lamina or deeper [
4]. The air-puff and pinch stimuli were repeated thrice to evaluate the evoked activities of neurons. Then mean frequencies and amplitudes of the evoked EPSCs for each SG neuron during the stimulation were calculated by the values of the 3 records.