All protocols were approved by the Animal Care and Use Committee of Renji Hospital, Shanghai Jiao Tong University School of Medicine, Shanghai, China (Chairman: Dr. Huili Dai) on 20 August 2018 (Permit Number: RJ 2018–0820). All procedures followed the guidelines of the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals (Department of Health and Human Services, NIH Publication No. 86–23, revised 1985) and the policies of the International Association for the Study of Pain regarding the use of laboratory animals. Efforts were made to minimize suffering due to surgery and to reduce the overall number of animals. All experiments were performed on male Sprague Dawley rats (weight: 200 to 250 g) that were housed in an animal facility, and provided with ad libitum water and standard laboratory food pellets. Rats (n = 162) were habituated to their environment (22 to 24 °C; 50 to 60% relative humidity; 12-h light/dark cycle) for 3 days before the experiments.
To specifically knock-down the expression of L-type calcium channels in the L4/5 DRG, which are responsible for the transmission of nociceptive information and thus conduct pain perception in the plantar incision model, we performed direct DRG microinjection with the recombinant adeno-associated virus 2/5 (AAV2/5) with Cav1.2 (Cacna1c) shRNA. DRG microinjection was performed as described previously, with minor modification [29, 56]. Briefly, a midline incision was made in the lower back of the lumbar spine to reveal L4 and/or L5 articular processes, which were then removed with a bone rongeur. After the DRG was exposed, a viral solution (1.62 × 1013 Vector Genomes (V.G.)/mL, 2 μL) was injected into two sites of L4 and L5 DRG using a glass micropipette connected to a Hamilton syringe. The pipette was removed 10 min after injection. The surgical field was rinsed with sterile saline and the skin incision was closed with sutures. The injected rats displayed no sign of paresis or other abnormalities, indicating that immune responses to the viral injections were minimal.
The viral solution consisted of AAV2/5-H1-shRNA (Cacna1c)-CAG-EGFP or AAV2/5-H1-NC_shRNA-CAG-EGFP-WPRE-pA (Taitool Bioscience Co.Ltd., Shanghai, China). The viral vector was pAAV2/5-H1-shRNA-CAG-EGFP . H1 was the promoter for shRNA, and CAG was the promoter for EGFP. The supplemental materials provide detailed viral vector mapping and sequencing data. The sequence of Cav1.2(Cacna1c) shRNA was: 5′- TCCCCgCCATTTTCACCATTGAAATTTTCAAGAGAAATTTCAATGGTGAAAATGGcTTTTT-3′. AAV2/5-H1-NC_shRNA-CAG-EGFP-WPRE-pA was used as a negative control to eliminate the influence of other interfering factors. Each recombinant AAV2/5 was locally injected into L4 and L5 DRG 21 days before SD, because AAV2/5 requires about 3 weeks before beginning gene expression after injection, and maintains relatively long-term gene transcriptional expression ability as an episome . To confirm the positive and control AAVs effectively infected the DRG neurons, frozen sections of L4 and L5 microinjected DRG was observed to detect the presence of abundant green fluorescence (EGFP). A non-injected DRG was used to exclude non-specific emission and to account for background fluorescence (Additional file 1: Figure S4e). There were 5 to 10 rats per group.
Postsurgical pain model
The plantar incision (PI) surgery was performed as previously described . Rats were anesthetized with 2% isoflurane, with 0.8–1.0 L/min oxygen delivered via a nose cone. The surface of the left hind paw was prepared under sterile conditions. Then a 1-cm longitudinal incision was made with a surfical blade through the skin and fascia of the plantar aspect of the foot, starting 0.5 cm from the proximal edge of the heel and extending toward the toes. The origins and insertions of muscles remained intact, and the flexor muscle was elevated and incised. After hemostasis with gentle pressure, the skin was sutured with 5–0 nylon thread and the wound was covered with bacitracin ointment. After surgery, the animals were allowed to recover in their cages. Typically, the wounds healed well within 5 to 6 days. In all experiments, the contralateral paw was used as a control. Rats in a sham control group received anesthesia but no surgery.
Sleep disturbance procedure
Rats were intermittently deprived of REM sleep using the small-platform method, as described previously [19, 37]. In brief, a small platform (15 cm high, 5 cm diameter) that was fixed to the center of a plastic water tank cage (45 × 53 × 72 cm) was surrounded with water (5 cm deep). At the onset of sleep, the muscular atonia caused the body to contact the water, thus awaking the animal. Each rat was placed individually on a platform within a plastic water tank cage, and was housed therein for 6 h per day for 3 consecutive days during the daytime before and after the surgery (6 days total), with food and water supplied ad libitum. Control rats were placed in groups in plastic cages in the same environment. For most experiments, rats were divided into 4 groups (5 to 10 rats per group): sham; sham+ SD; incision; and incision+SD. Some experiments employed additional treatments with nifedipine, (an L-type channel sensitive calcium channel blocker) or viral injections, as indicated in the text.
von Frey filaments (mechanical stimulation)
The von Frey filament test was performed each day from 1 day before surgery to 15 days after surgery. Each rat was habituated in a small (7.5 × 15 × 15 cm) plastic cage with air vents at the top for at least 30 min before testing. Mechanical sensitivity was determined with a series of von Frey filaments (2.0 to 26 g) that were applied to the plantar surface of the left and right hind paws. Each filament was tested five times in increasing order from the lowest force. Between individual measurements, von Frey filaments were applied at least 3 s after the rats had returned to their initial resting state. The minimal force that led to either a rapid paw withdrawal and/or an escape attempt in at least 3 of the 5 stimulations was determined as the threshold of the mechanical response.
Laser heat pain (thermal stimulation)
Each rat was habituated for 30 min in a small plastic cage (7.5 × 15 × 15 cm) with air vents at the top on a glass plate. Laser heat was applied by aiming a beam of light through a hole in the light box, through the glass plate, to the middle of the plantar surface of each hind paw. When the animal lifted its foot, the light beam was turned off. The time from stimulation to foot withdrawal (latency) was measured. Each trial was repeated three times at 10-min intervals for each hind paw, and a cut-off time of 20 s was used to avoid tissue damage.
Nifedipine (N7634, Sigma), a small molecule L-type channel-sensitive calcium channel blocker which is widely used in clinical practice, was dissolved in a vehicle solution of 95% sterile saline and 5% DMSO. Then, intraperitoneal injections (15 mg/kg; a concentration that produces an antinociceptive effect in rats ) were given 1 h before the behavioral tests. Nifedipine works within 10 min after administration, has maximal effect in 1 to 2 h, and its effect lasts 4 to 6 h. Normal saline (NS) was injected into control rats. Each injection was given in a volume less than 1.0 mL on days 8 and 9 after incision surgery.
Western blot analysis was performed as previously described, with minor modification [3, 56]. In brief, bilateral L4–6 DRG were collected, rapidly frozen, and homogenized in chilled SDS lysis buffer (P0013G, Beyotime). The crude homogenate was centrifuged at 4 °C for 15 min at 12,000 g. The supernatant was collected and the pellet (nuclei and debris) was discarded. Protein concentration was measured, and the samples were then heated at 100 °C for 15 min and electrophoresed in SDS-PAGE. The proteins were then transferred onto polyvinylidene fluoride (PVDF) membranes (IPVH00010, Immobilon-P). The membranes were blocked with 1% bovine serum albumin (BSA) at 4 °C overnight, and then incubated with rabbit anti-Cav1.2 antibody (L-type; 1:200, ACC-003, Alomone), rabbit anti-Cav2.1 antibody (P/Q-type; 1:200, ACC-001, Alomone), rabbit anti-Cav2.2 antibody (N-type; 1:200, ACC-002, Alomone), rabbit anti-Cav2.3 antibody (R-type; 1:200, ACC-006, Alomone), rabbit anti-Cav3.2 antibody (T-type; 1:200, ACC-001, Alomone), mouse anti-Egr1 antibody(1:200, sc-101,033, Santa Cruz), or rabbit beta tubulin antibody (1:3000, AB0039, Abways) at 4 °C overnight under gentle agitation. Beta tubulin was used as a loading control. The membranes were washed and then incubated with a horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (1:2000, A0208, Beyotime) or a horseradish peroxidase-conjugated goat anti-mouse secondary antibody (1:2000, A0216, Beyotime) for 1 h at room temperature. The blots were developed using the ECL Plus detection system. Band density was measured using Image J software.
Tissues were collected from a separate group of for immunofluorescence studies. Rats were subjected to perfusion with 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS), followed by 4% PFA in PBS post-fixation overnight. The L4–6 DRG were cryo-protected in a 20% sucrose solution overnight, and then in a 30% sucrose solution. The tissues were dissected and processed (section thickness: 20 μm) for immunofluorescence staining as previously described [14, 31]. Sections were intensively washed with PBS, and then treated with an immunostain blocking buffer (P0102, Beyotime) for 1 h at room temperature before staining. The primary antibody was rabbit anti-Cav1.2 antibody (1:50, ACC-003, Alomone). Double staining used the rabbit anti-Cav1.2 antibody with an antibody against FITC-conjugated isolectin B4 (marker for small non-peptidergic neurons ; 10 μg/mL, L2895, Sigma); an antibody against mouse anti-CGRP (marker for small peptidergic neurons ; 1:100, ab81887, Abcam); an antibody against mouse anti-neurofilament 200 (marker for medium/large neurons ; 1:500, N0142, Sigma); and mouse anti-Egr1 antibody to detect whether Egr-1 and Cav1.2 are present in the same cell (1:200, sc-101,033sc-101033, Santa Cruz). All sections were then incubated with either anti-mouse IgG conjugated to Alexa Fluor® 488 (1:1000, 4408, Cell Signaling) or anti-rabbit IgG conjugated to Alexa Fluor® 594 (1:1000, 8889, Cell Signaling). Conjugated antibodies were also used for nuclear staining with DAPI. Images were taken with a fluorescence microscope (Olympus, DP80) and processed using Image J software. Three to four slices per DRG per rat were counted to eliminate the uneven distribution of large, medium and small neurons due to the irregular shape of DRG . There were 3 to 4 rats per treatment group.
Whole-cell patch clamp recording
DRG neuron culture
Acute dissociated L4–5 DRG neurons were prepared as previously described . Rats were divided into groups as described above, and then euthanized with isoflurane. The L4–5 DRG were collected in cold DMEM/F12 medium (12634–010, Gibco) with 10% fetal bovine serum (10099-141, Gibco), 100 U/mL Penicillin, and 100 μg/mL Streptomycin (15140-122, Gibco), and then treated with an enzyme solution (5 mg/mL dispase and 1 mg/mL collagenase type I) in HBSS (14025-076, Gibco). Neurons were dissociated after trituration, resuspended in mixed DMEM/F12, and then plated onto 5 mm diameter coverslips that were coated with 50 μg/mL poly-D-lysine (P0296, Sigma). The DRG neurons were incubated at 95% O2 and 5% CO2, and at 37 °C.
HVA calcium channel current recording
Whole-cell patch clamp recording was performed 3 to 8 h after plating. Coverslips were placed in the perfusion chamber. The electrode resistances of the micropipettes ranged from 4 to 6 MΩ. Neurons were voltage-clamped with an Axon 1550B amplifier using Clampex software [30, 56]. The Cold Spring Harbor Protocol was followed to separate HVA calcium current from L-type calcium current . The intracellular pipette solution (pH 7.3 with CsOH, 290 mOsm) contained 110 mM CsCl, 5 mM MgCl2, 10 mM EGTA, 10 mM HEPES, 4 mM Mg-ATP, and 0.1 mM GTP. The extracellular solution (pH 7.3 with TEA-OH, 300 mOsm) contained 5 mM CaCl2, 130 mM tetraethylammonium chloride (TEA-Cl), 0.3 mM TTX, 10 mM HEPES, and 10 mM glucose. Series resistance was compensated by 60 to 80%. After establishment of a giga-Ω seal, the neuron membrane potential was maintained at − 90 mV. An initial depolarizing step was applied to change the holding potential to − 30 mV for 1 s to inactivate all LVA calcium channels. Then, a second depolarizing step to 0 mV for 100 ms was applied so that only HVA calcium channels were activated . Online P/4 leak subtraction was performed to eliminate this effect. All data were stored and analyzed using Clampfit software [30, 56]. To specifically verify the contribution of L-type current to the total HVA current, 1 μm nifedipine was applied to the neurons via bath perfusion during measurements.
Action potential recording
To record the action potential, the recording mode was switched to the current clamp. The extracellular solution (pH 7.38 by NaOH) contained 140 mM NaCl, 4 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, and 5 mM glucose. The intracellular pipette solution (pH 7.38 with KOH, 300 mOsm) contained 135 mM KCl, 3 mM Mg-ATP, 0.5 mM Na2ATP, 1.1 mM CaCl2, 2 mM EGTA, and 5 mM glucose. The resting membrane potential was recorded 3 min after a stable recording was first obtained. Depolarizing currents of 100 to 1400 pA (200-ms duration) were delivered in increments of 100 pA until an action potential (AP) occurred. The injection current threshold was defined as the minimum current needed to evoke an AP. The membrane potential was maintained at the existing resting membrane potential during the current injection. The AP threshold was defined as the first point on the rapidly rising phase of the spike at which the change in voltage exceeded 50 mV/ms, and the AP amplitude was defined as the distance from the peak to the baseline. The membrane input resistance of each cell was obtained from the slope of a steady-state I–V plot in response to a series of hyperpolarizing currents (200-ms duration) that were applied in steps of 100 pA, from 200 pA to − 2000 pA. The after-hyperpolarization amplitude was the distance from the maximum hyperpolarization to the final plateau voltage, and the AP overshoot was the distance from the AP peak to 0 mV. All experiments were performed at room temperature and all data were stored and analyzed using Clampfit software [30, 56].
To examine the effect of Egr-1 on the activity of the Cav1.2 promoter, a luciferase assay was performed. A fragment from the Cacna1c gene promotor region and a fragment from the Egr-1 gene were amplified by PCR from genomic DNA to construct Cacna1c gene reporter plasmids and the Egr-1 gene over-expression plasmids, respectively. The PCR products were ligated into the GV238 vector (containing the firefly luciferase reporter gene) and the GV141 vector (containing the renilla luciferase reporter gene) using KpnI and XhoI restriction sites, respectively. DNA sequencing was performed for verification. HEK-293 T (ATCC) cells were cultured for 1 day in DMEM/F12 (12634–010, Gibco) containing 10% fetal bovine serum (10099-141, Gibco) at 37 °C in a humidified incubator with 5% CO2. Cells were then transferred to a 24-well plate, transfected with the Cacna1c gene reporter plasmids with an empty GV141 vector (control) or with Egr-1 gene over-expression plasmids using X-tremegene HP (Roche), according to the manufacturer’s instructions. Two days after transfection, the cells were collected in a passive lysis buffer. The supernatant was used to measure luciferase activity using the Dual-Luciferase Reporter Assay System (E1910, Promega). Independent transfection experiments were repeated three times. The relative reporter activity was calculated after normalization of firefly fluorescence to renilla fluorescence.
All results are presented as means ± standard errors of the mean (SEMs). Statistical analysis was performed using Prism 7.0 software. A two-tailed, unpaired Student’s t-test and one-way or two-way ANOVA were used as appropriate in multiple-comparisons tests. A P value below 0.05 was considered significant.