Animals

All animal care and experimental methods complied with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and were preapproved by the Institutional Animal Care and Use Committee at the Torrey Pines Institute for Molecular Studies (Port St. Lucie, FL, USA). Consistent with these guidelines, ongoing statistical testing of data collected was used to minimize the number of animals used, within the constraints of necessary statistical power. Animal studies are reported in compliance with the ARRIVE guidelines (Kilkenny et al., 2010; McGrath and Lilley, 2015).

Animals were group‐housed (three to five mice per cage, depending on group allocation) in plastic cages with conventional bedding and provided with food and water ad libitum under a 12 h day/night cycle (lights on at 07:00 h). The colonies for Oprl1‐eGFP‐homozygous (NOP^eGFP/eGFP; NOP‐eGFP) mice on a C57/BL6/J background have been routinely maintained in‐house (Ozawa et al., 2015). To match the genetic background, male and female C57BL6/J wild‐type mice (4 weeks old) were obtained from the Jackson Laboratory (Bar Harbor, ME, USA). All behavioural studies were conducted in C57BL6/J wild‐type mice from the Jackson Laboratory in order to minimize the potential impact of the C‐terminally attached eGFP on the functionality of NOP receptors, though the NOP‐eGFP mice showed a normal nociceptive response to the thermal stimulation in the tail flick tests (Ozawa et al., 2015). Rather than using wild‐type littermates of the NOP‐eGFP mice, we used commercially available C57BL6/J mice in order to test sufficient number of animals of the same age. This also confers consistency on behavioural experiments. As we observed consistent changes in NOP receptors when measuring protein in NOP‐eGFP mice and mRNA in wild‐type mice, there are no anticipated problems in using wild‐type mice to conduct the pain behavioural experiments in the current study.

Spinal nerve ligation

SNL surgery at L4 and L5 mouse spinal nerves, which most closely mimics L5/L6 ligation in rats (Kim and Chung, 1992), was carried out in 5–8‐week‐old mice (male NOP‐eGFP mice, 20–25 g; female and male C57BL6/J wild‐type, 13–20 g), following previously reported procedures (Ye et al., 2015). Under isoflurane anaesthesia (5% for induction and 2% for maintenance in 10% O₂), animals were placed in a prone position. A dorsal midline incision was made to expose the lumbar L6 transverse process. Cutting of L6 transverse process allowed visualization of the L5 spinal nerve and the L3 and L4 spinal nerves joining each other. L4 and L5 spinal nerves were then isolated and tightly ligated with 6–0 silk suture. Muscles were sutured using absorbable sutures and skin tightly glued. In sham‐operated animals, the surgical procedure was identical to the SNL surgery except that spinal nerves were not ligated. Body temperature was maintained at 37°C throughout the surgery using a heating pad. At the end of the procedure, mice were monitored until full recovery from anaesthesia. Two weeks after surgery, we applied gentle tactile stimuli to the ipsilateral hindpaw of un‐anaesthetized mice in order to define the tactile allodynia established by SNL using a set of von Frey filaments as described below.

Immunohistochemistry

Two and four weeks after surgery, sham‐operated and SNL NOP‐eGFP mice (N = 4 male mice per group) were anesthetized with an overdose of isoflurane for 1 min and transcardially perfused in 4% paraformaldehyde in PBS. Anaesthetic depth was ensured by testing the pedal withdrawal reflex (pinching on both hindpaws) before perfusion. The DRG (L4, L5 and L6) and spinal cord (lumbar cord) were dissected and cryoprotected in 30% sucrose in PBS. Tissues were then frozen in OCT (Sakura Finetek, INC., Torrence, CA, USA) and stored in −80°C until used. DRG from the ipsilateral and contralateral sides were collected separately. DRG from the same animal group were then pooled for cryoprotection and frozen in the same mould. Tissue sections (40 μm for spinal cord; and 10 μm for DRG) were prepared by using a cryostat (Leica Biosystems, Buffalo Grove, IL, USA) and blocked with PBS containing 5% normal donkey serum and 0.3% Triton X‐100 for 1 h at room temperature. The sections were then incubated with primary antibodies, indicated in each figure, at 4°C, overnight. For the chicken anti‐GFP antibody, the incubation was performed at 37°C for 2 h. After extensive washing with PBS containing 1% normal donkey serum and 0.3% Triton X‐100, sections were incubated with appropriate secondary antibody conjugated to AlexaFluor (1:1000) for 2 h at room temperature.

Image analysis and quantification

Images were collected under a Leica TCS SP5II confocal microscope and LAS AF Lite software (Leica Microsystems) or an A1R confocal microscope and NIS‐Elements Imaging software (images shown in Figure 1C, D; Nikon, Melville, NY, USA). All of the images directly compared were collected using the same laser intensities for the corresponding colour channels. Image analysis was carried out by using Fiji (National Institute of Health) by an experimenter, blinded to group conditions. Brightness and contrast were adjusted equally among the images compared when necessary. Images from four sections per animal were collected for analysis. Cell counts, immunoreactive intensities and percentage obtained for each section were averaged. To investigate the effect of chronic pain on NOP receptor distribution in areas of the spinal cord relevant for nociceptive processing, immunofluorescent measurements were carried out at the ipsilateral and contralateral spinal laminae I to IIo shown in Figure 1A. Using Fiji, the region of interest (ROI) was manually drawn with freehand line tool, and the intensity and size of the regions were measured using the ROI manager. The fluorescent intensity obtained from each ROI was then divided by its own area size. Values [NOP‐eGFP immunoreactivity (%)] for ipsilateral and contralateral sides of SNL mice and for contralateral side of sham‐operated mice were calculated relative to ipsilateral of sham group. To ensure NOP‐eGFP‐positive DRG neurons, background measurement was carried out at three low immunoreactive areas on the DRG in each image and the intensity was averaged. All DRG neurons that exceeded twice the background intensity were counted as NOP‐eGFP positive. To quantify the immunoreactivity of NOP‐eGFP in DRG neurons, NOP‐eGFP‐positive DRG neurons were first selected, and their fluorescent intensity and size were measured using the freehand line tool and ROI manager in Fiji. Fluorescent intensities and sizes were summed separately, and the immunoreactivity of NOP‐eGFP per image was calculated as [(total immunoreactive intensity of NOP‐eGFP‐positive cells)/(total area size of NOP‐eGFP‐positive cells)]. Immunoreactivity of NOP‐eGFP for ipsilateral and contralateral sides of SNL mice were then compared with the corresponding side of sham‐operated mice and represented as a percentage of sham group.

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Figure 1

Changes in the distribution of NOP‐eGFP receptors in SNL mice. NOP‐eGFP receptor distribution was investigated 14 days post SNL of the L4/L5 spinal nerves. (A) Representative images show that NOP‐eGFP (green) is concentrated in the inner part of lamina II (IIi), where PKCγ interneurons (red) are located in the L4 and L5 ipsilateral spinal cord of the SNL mice (observed in yellow), while the immunoreactivity in laminae I and II outer (IIo) is decreased. In contralateral spinal cord, slightly lower immunoreactivity of NOP‐eGFP was observed in SNL mice, but it is not significant compared to sham‐operated mice. No changes were observed in the L6 segment. CGRP (blue), a marker for the spinal laminae I and IIo. All of the representative images were prepared from the same sham‐operated or SNL mice. Roman numerals in zoomed images for the ipsilateral side indicate lamina. White arrowhead depicts the position of the lateral spinal nucleus. Ip, Ipsilateral; C, Contralateral. (B) Quantification of immunoreactivity of NOP‐eGFP in the ipsilateral and contralateral laminae I and IIo. N = 4 mice per group. (C) Representative images of the L4 spinal dorsal horns 4 weeks after surgery. As in the results in panel (A), a decrease in NOP‐eGFP immunoreactivity was observed in the ipsilateral spinal cord of SNL mice. Note that images for ipsilateral and contralateral were obtained from the same section. In the same section, CGRP staining was also carried out as a reference to verify the effect of SNL in the ipsilateral spinal dorsal horn. (D) Representative high‐power images of the ipsilateral L4 spinal cord derived from sham‐operated and SNL mice. Right panel, an expanded image of the red square in the merged images. No PKCγ interneurons co‐expressed NOP‐eGFP. (E) Effect of SNL on mechanical allodynia measured by von Frey (N = 4 for sham and N = 5 for SNL male NOP‐eGFP mice). After initial von Frey testing, one mouse from SNL group was eliminated from the immunostaining experiments because it did not develop mechanical allodynia. For all quantifications, data shown are means ± SEM. Scale bars: (A) and (C), 100 μm; (D), 50 μm.

Intrathecal drug administration

By following the method of Hylden and Wilcox (1980), intrathecal (i.t.) administration of N/OFQ (3 and 10 nmol) or vehicle was performed, using doses based on a previously reported study (Rizzi et al., 2015). Briefly, an awake mouse was held firmly by the pelvic girdle in one hand. A disposable 30 gauge 0.5 inch needle attached to a 10 μL Hamilton syringe was then inserted into the vertebrae between L5 and L6 of the mouse spinal column. A characteristic tail flick was confirmed as the penetration of the subarachnoid space. N/OFQ was administered in a total volume of 5 μL per mouse. Changes in each pain modality described below were determined 10 min after the injection of N/OFQ in SNL mice as compared to sham‐operated mice. For the antagonist test, 10 mg·kg⁻¹ of http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=1693 was i.p. administered to mice 30 min prior to N/OFQ or vehicle (Toll et al., 2009; Khroyan et al., 2011).

Assessment of tactile allodynia

Tactile allodynia was measured in mice that received either SNL or sham surgery as hindpaw withdrawal response to a set of von Frey filaments with forces of 0.04, 0.07, 0.16, 0.4, 0.6, 1.0 and 2.0 g (Stoelting, Wood Dale, IL, USA) using the up/down method of Chaplan et al. (1994). Prior to the test, mice were habituated for 30 min in individual chambers placed on a wire‐mesh floor. The test began with a von Frey filament that had a buckling weight of 0.4 g. The von Frey filament was applied to the plantar surface of the ipsilateral hindpaw (left hindpaw) of the animal with continuous pressure for approximately 5 s. A positive response was recognized as a sharp withdrawal of the testing hindpaw. If the animal lifted its paw, the next filament with lower force was then applied. If the animal did not lift its paw, the next filament with higher force was used. Each response was recorded, and the experiment ended once the animal had made five responses after the initial positive response. The 50% paw withdrawal threshold was calculated (Dixon, 1980). If the animal made four consecutive positive responses, a score of 0.04 g was assigned, whereas if the animal had five consecutive negative responses, then a score of 2.0 g was assigned.

Assessment of heat and cold hyperalgesia

Heat hyperalgesia was measured using a hot/cold plate (IITC, Woodland Hills, CA, USA) with the surface temperature maintained at 55°C. Mice were placed on the hot plate, and the latency for withdrawal of the hindpaw or jumping from the metal plate was measured. A cut‐off latency of 30 s was used to avoid tissue damage. For cold hyperalgesia, mice were placed on the metal plate with the surface temperature maintained at 2°C. The number of flicks of the ipsilateral hindpaw was counted during an observation period of 5 min.

Quantification of NOP receptor mRNA expression

Total RNA for the spinal cord and DRG neurons was isolated from wild‐type mice that received SNL or sham surgery (2 and 6 weeks after the surgery) using RNAeasy Mini kit (Qiagen, Hilden, Germany). mRNA analysis for the 6‐week group was carried out with mice (three female and three male) previously used for pharmacological testing. Mice were deeply anesthetized with isoflurane and killed by cervical dislocation. The lumbar portion of the spinal cord (from L4 to L5 segments) as well as ipsilateral and contralateral L4‐L5 DRG was dissected on ice. L4 and L5 DRG for each animal were pooled together for analysis. For dissecting the spinal cord, the spinal column was carefully removed from neck to tail in order to expose the dorsal side. The cranial and the caudal ends as well as L4 and L5 segments of the spinal column were marked for reference. The lumbar cord was carefully removed from the spinal column. The spinal cord was carved at the centre towards the caudal end by following the midline reference in order to separate ipsilateral and contralateral portions and the L4‐L5 were then excised. Total RNA was subjected to reverse transcription with a QuantiTect Reverse Transcription Kit (Qiagen). To measure mRNA levels, real‐time quantitative RT‐PCR was performed with the iCycler IQ5 multicolor Real‐Time PCR detection system (Bio‐Rad Laboratories, Hercules, CA, USA) using QuantiFast SYBR Green PCR Kit (Qiagen); 95°C for 3 min to activate the HotStart enzyme followed by 45 cycles of amplification and quantification (10 s at 95°C; 30 s at 60°C) each with a single FAM fluorescence measurement. Melting curve analysis (60–95°C) was also performed in order to verify reaction specificity for each PCR products. Gene‐specific primers for NOP receptor and GAPDH (an endogenous reference gene) were designed by the NCBI primer‐blast. GAPDH was used as a reference gene to normalize NOP receptor mRNA expression due to the previous evidence showing stable expression in DRG and spinal cord between normal and nerve‐injured animals (Laedermann et al., 2014; Jiang et al., 2015). All of the primers were purchased from Eurofins (Louisville, KY, USA). Primers for NOP receptor and GAPDH were as follows: NOP_F459, 5′‐ATGTCATCCTCAGGCACACC‐3′; NOP_R573, 5′‐AAATGGCCAGAAGCCCAGAA‐3′, GAPDH_F581, 5′‐GTGGAAGGGCTCATGACCAC‐3′; and GAPDH_R698, 5′‐ATGCAGGGATGATGTTCTGG‐3′. Data were calculated as the mean ΔCt relative to the house‐keeping gene GAPDH. The ΔΔCt method (Livak and Schmittgen, 2001) was employed to calculate percentage expression of SNL mice relative to sham‐operated mice [(relative NOP mRNA level for individual mouse)/(averaged relative NOP mRNA level of sham‐operated mice)×100].

Experimental design

Assessment of tactile allodynia was carried out in all the mice used for molecular and behavioural analysis, at 2 weeks after surgery. Animals that were not deemed healthy after surgery were excluded from the experiments. Prior to pharmacological tests, 8 and 10 mice from each group were randomly selected and subjected to cold and hot plate, respectively, in order to measure the basal response to each pain modality. Beginning 3 weeks after surgery, mice received i.t. injections of N/OFQ (3 or 10 nmol) or vehicle and were examined on each behavioural test described above. The behavioural tests were carried out in the following order: mechanical allodynia (3 weeks post‐surgery), heat hyperalgesia (4 weeks post‐surgery) and cold hyperalgesia (5 weeks post‐surgery). After all the behavioural tests, tissues from six mice per group were dissected for the RNA preparation. Additionally, in a separate cohort of SNL mice, i.t N/OFQ (10 nmol) was examined in the presence or absence of SB‐612111 (10 mg·kg⁻¹, i.p.) for mechanical allodynia (3 weeks post‐surgery) and heat hyperalgesia (4 weeks post‐surgery). The exact number of animals tested in different treatments in the study is indicated in the Methods section or in the figure legends. A sample size for behavioural experiments was determined by a power analysis done with G*Power 3.1.9.2. Animals in each group (sham‐operated and SNL mice; female and male) were randomized before treatments. All pain behavioural experiments were conducted by experimenters blinded to the condition and treatments used.

Data and statistical analysis

The data and statistical analysis comply with the recommendations on experimental design and analysis in pharmacology (Curtis et al., 2015). All of the results are shown as mean ± SEM. Data were tested for statistical significance using unpaired t‐test or two‐way ANOVA that used two between‐subject factors (‘ligation’ and ‘treatment’ for the N/OFQ antinociceptive effect on SNL‐mediated allodynia, and heat/cold hyperalgesia; ‘pretreatment’ × ‘treatment’ for SB‐612111 test on the N/OFQ‐mediated antinociceptive effects). When interaction of factors was statistically significant (P values less than 0.05), pairwise comparisons were conducted by using Newman–Keuls post hoc test (Statistica 7.0 software). Any data point that was detected as an outlier (Grubbs, α = 0.05, P < 0.05) was excluded from unpaired t‐test or two‐way ANOVA.

Materials

N/OFQ and http://www.guidetopharmacology.org/GRAC/LigandDisplayForward?ligandId=1693, an NOP receptor antagonist, were provided by the National Institute on Drug Abuse Drug Supply Program and were dissolved in sterile PBS and 2% DMSO in 0.5% aqueous hydroxypropyl cellulose (HPC) respectively. All solutions were prepared freshly on the test day.

Expression of NOP receptors in DRG neurons is lowered in SNL mice

We next addressed the expression pattern of NOP‐eGFP receptor in the DRG neurons derived from sham‐operated and SNL mice. Similar to the results seen in the spinal cord, the total NOP‐eGFP receptor expression was decreased in the L4 and L5 ipsilateral DRG of SNL mice, compared to the sham‐operated mice (Figure 2A, C). No changes in NOP‐eGFP immunoreactivity were observed in the L6 ipsilateral DRG or any of the contralateral DRG (Figure 2A–D). In addition, the percentage of small NOP‐eGFP‐positive neurons (100–200 and 200–300 μm²) was reduced in L4 DRG subsequent to SNL surgery (Figure 2G). NOP‐eGFP receptor expression disappeared in about 50% of small‐sized DRG neurons in SNL mice. In contrast, an increase in NOP‐eGFP‐positive cell number was observed in medium‐sized L4 neurons (Figure 2G; 400–500 μm²). Small NOP‐eGFP‐positive DRG neurons (<300 μm²) co‐expressing CGRP (NOP‐eGFP+ CGRP+) were rarely found in L4 ipsilateral DRG of SNL mice (Figure 2A, I), suggesting that the expression of NOP receptors might be down‐regulated specifically in peptidergic C‐nociceptors during chronic pain. A decrease in the number of cells expressing CGRP was selectively observed in the ipsilateral L4 and L5 DRG of SNL mice compared to sham‐operated mice (Figures 2E, F), which also agrees with previous neurochemical analyses conducted in SNL mice (Honore et al., 2000). In L5 DRG, surprisingly, there was not a comparable difference in the percentage of NOP‐eGFP‐positive cells between sham‐operated and SNL mice at any neuron sizes (Figure 2H) nor in the percentage of small NOP‐eGFP+ neurons expressing CGRP. As smaller changes in NOP‐eGFP immunoreactivity were observed in L5 ipsilateral DRG, compared to L4 ipsilateral DRG (Figure 2C), these results might indicate that reduced expression is a function of fewer receptors per neuron rather than certain types of neurons being completely devoid of NOP receptors in L5 DRG.

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Figure 2

Expression of NOP‐eGFP receptors in the DRG neurons of SNL mice. (A, B) Representative images of L4, L5 and L6 ipsilateral and contralateral DRG neurons derived from sham‐operated and SNL mice. Tissue sections were incubated with anti‐GFP (green) and anti‐CGRP (red) antibodies. Right panel, an expanded image of the merged images. White arrowheads denote the cells co‐expressing NOP‐eGFP and CGRP in small DRG neurons. These cells were rarely observed in the L4 DRG in SNL mice. As DRG tissues were pooled for cryoprotection, the representative images for each section were prepared from all of the animals in the corresponding group. Scale bars, 100 μm. (C) Decreased expression level in NOP‐eGFP receptors was observed in the L4 and L5 ipsilateral DRG neurons of SNL mice compared to sham‐operated mice. NOP‐eGFP expression in the L6 ipsilateral DRG neurons was not affected by SNL. (D) No difference between sham and SNL mice in NOP‐eGFP immunoreactivity was observed in the contralateral DRG neurons. (E, F) Population of the CGRP‐expressing neurons in the L4, L5 and L6 ipsilateral and contralateral DRG. Percentage of CGRP‐positive neurons was calculated by comparing with the total number of DRG neurons. (G, H) Size profiling of NOP‐eGFP+ neurons in L4 and L5 ipsilateral DRG. (I) Population of small NOP‐eGFP‐positive neurons (<300 μm²) expressing CGRP (NOP‐eGFP+ CGRP+) in the L4, L5 and L6 ipsilateral DRG. The percentage of NOP‐eGFP+ CGRP+ neurons was determined by comparing with the total number of small CGRP‐positive neurons; [(number of small NOP‐eGFP+ CGRP+ neurons)/(number of small CGRP+ neurons)×100]. For all quantifications, data shown are means ± SEM. N = 4 mice for each group.

mRNA levels in DRG and spinal cord of wild‐type sham‐operated and SNL mice were also determined (Figure 3). Consistent with a decrease in NOP receptor protein, mRNA levels were also reduced in both ipsilateral DRG and spinal cord of SNL wild‐type mice at 2 and 6 week after the SNL surgery (Figure 3A, C). Changes in mRNA expression level of NOP receptor were not observed in either contralateral spinal cord or DRG (Figure 3B, D). These data together with spinal NOP receptor distribution discussed above suggest that an important role of specific subsets of NOP receptor expressing neurons in the spinal cord and DRG may affect the development of NOP receptor‐based analgesics.

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Figure 3

NOP receptor mRNA expression in the spinal cord and DRG of SNL wild‐type mice. Relative mRNA expression of NOP receptor in the ipsilateral (A, C) and contralateral (B, D) spinal cord and DRG neurons were determined by quantitative RT‐PCR, 2 (A, B) and 6 weeks (C, D) after surgery. (A, B) Two weeks after surgery: Ipsilateral DRG: sham (N = 6, three female and three male mice); SNL (N = 5, three female and two male mice). Ipsilateral spinal cord: sham (N = 5, three female and two male mice); SNL (N = 5, two female and three male mice). Contralateral DRG: sham (N = 6, three female and three male mice); SNL (N = 5, three female and two male mice). Contralateral spinal cord: sham (N = 6, three female and three male mice); SNL (N = 6, three female and three male mice). Outliers eliminated: One male SNL mouse from ipsilateral DRG and contralateral DRG; one sham mouse from ipsilateral spinal cord; one SNL female mouse from ipsilateral spinal cord. (C, D) Six weeks after surgery: Ipsilateral DRG: sham (N = 5, two female and three male mice); SNL (N = 6, three female and three male mice). Ipsilateral spinal cord: sham (N = 6, three female and three male mice); SNL (N = 6, three female and three male mice). Contralateral DRG: sham (N = 6, three female and three male mice); SNL (N = 5, two female and three male mice). Contralateral spinal cord: sham (N = 6, three female and three male mice); SNL (N = 6, three female and three male mice). Outliers eliminated: One sham female mouse from ipsilateral DRG; one SNL female mouse from contralateral DRG. Tissues were collected from the mice 2 and 6 weeks after surgery. Results similar to the immunostaining shown in Figures 1 and ​and22 were observed in the spinal cord and DRG. Data are represented as mean ± SEM. *P < 0.05, significantly different from sham‐operated mice; unpaired t‐test, Welch's correction.

This work was supported by the National Institute on Drug Abuse grant DA023281 to L.T. We thank Shannon Ortiz‐Umpierre and Roman Ramirez for maintaining the colony for NOP‐eGFP mice; Reona Asano, Yoshitaka Iwayama and Masako Nagai for excellent technical assistance; and the Optical Workshop and Light Microscopy Facility Center at the MAX PLANCK Florida Institute and the Nikon Center of Excellence at the Florida Atlantic University Brain Institute for the use of the microscopes.