Nitrous oxide (N2O), one of the most common drugs used in anesthetic practice, has strong analgesic action. The mechanism of antinociceptive action of N2O has been extensively investigated. It activates noradrenergic neurons in the brainstem (1) and increases noradrenaline release in the brain and the spinal cord (2,3). Antagonism of adrenergic receptors (4) or depletion of noradrenaline (3) in the spinal cord decreases the antinociceptive effect of N2O. Furthermore, lesioning of brainstem noradrenergic neurons also inhibits N2O antinociception (1). These findings suggest that N2O exerts its antinociceptive effect, at least partly, via the activation of descending noradrenergic inhibitory pathways. Moreover, it was demonstrated that the activation of the descending inhibitory pathways by N2O is mediated by opioid peptide release in the periaqueductal gray area (PAG) of the midbrain (5).
Nociceptin-orphanin FQ and its endogenous ligand of the opioid receptor-like 1 receptor (NOP) have high structural homologies with classical opioid peptides and receptors, respectively, and are widely expressed in the central nervous system (6). Although physiological roles of the nociceptin system have been examined from a variety of aspects, many issues remain to be clarified. We (7) have previously reported that the minimum anesthetic alveolar concentrations for volatile anesthetics (halothane, isoflurane, and sevoflurane) did not differ between nociceptin receptor knockout (NOP−/−) mice and wild-type (NOP+/+) mice, in contrast to the report that the potency of sevoflurane was reduced in μ-opioid receptor knockout mice compared with wild-type mice (8). There are nociceptin-containing nerve terminals and NOP in the PAG (9,10), in which N2O induces opioid peptide release. Therefore, the nociceptin system may affect the antinociceptive action of N2O. In the present study, we investigated whether the nociceptin system is involved in the antinociceptive action of N2O by using NOP−/− mice and their littermates. We first examined the behavioral difference between NOP−/− and NOP+/+ mice by the acetic acid-induced writhing test under N2O. Then we measured the plasma adrenocorticotropic hormone (ACTH) concentrations to examine the difference of stress hormonal response. Finally, we evaluated immediate early gene expression (c-Fos) of the spinal cords after N2O exposure to examine the difference of N2O-induced activation of the descending inhibitory neurons.
The study protocol of the animal experiments was approved by the Animal Experiment Committee of Kyoto University. NOP−/− mice, with a mixed genetic background of C57BL and 129 strains, were developed by the means of gene targeting (11) and back-crossed repeatedly to the C57BL strain. We studied male NOP−/− mice and NOP+/+ mice (8–10 wk old; 20–30 g). These animals were obtained by intercrossing NOP+/− heterozygotes and genotyped by polymerase chain reaction of tail biopsy tissue. The animals were housed in a controlled environment (25°C ± 1°C) and had free access to food and water. The room lights were on between 7 am and 7 pm. All experiments were performed between 9 am and 5 pm to avoid circadian effects.
Visceral pain was scored according to the method described previously (12). Each mouse was initially placed in an individual polypropylene chamber (22 cm in diameter and 12 cm in height) and allowed 30 min to acclimatize to laboratory surroundings. The mouse was then exposed to either 30% oxygen/70% nitrogen or 30% oxygen/70% N2O continuously delivered from an anesthetic machine (Aika, Ichikawa Shiseido, Tokyo, Japan) into the chamber through an inflow port, and the gas was exhausted through an outflow port. Total gas flow rate was 6 L/min. Concentrations of oxygen, carbon dioxide, and N2O in the chamber were continuously monitored with an infrared analyzer (Capnomac Ultima, Datex-Ohmeda, Helsinki, Finland). Thirty minutes after the beginning of the exposure, 0.6 mL of 0.9% acetic acid was injected into the peritoneal cavity, and the intensity of nociception was quantified by counting the total number of writhings occurring between 0 and 30 min after the injection. The writhing response consists of a contraction of the abdominal muscles together with a stretching of the hindlimbs. The effect of this treatment was reversible. A warm blanket (Animal Blanket Controller ATB-1100, Nihon Kohden, Tokyo, Japan) was used to keep the rectal temperature to 38°C ± 0.5°C. The investigator who observed the writhing response was blinded to the NOP status of the animal.
Each mouse was placed in an individual plastic restrainer and anesthetized with 0.8% halothane/70% N2O during the experiment. Inspired concentrations of each anesthetic (total gas flow, 6 L/min) were continuously monitored with an infrared analyzer. The rectal temperature was controlled to 38°C ± 0.5°C using a heat lamp and a blanket. Thirty minutes after the beginning of the exposure to inhaled anesthetics, 0.9% acetic acid (0.6 mL) was injected intraperitoneally. The mouse was decapitated, and the blood samples were taken just before the injection and at 30 and 60 min after the injection. After separation, plasma was stored at −80°C until analysis. The plasma ACTH concentrations were measured by radioimmunoassay according to the method described previously (13).
We examined the c-Fos positive cells in the spinal cords of 10 NOP−/− and 10 NOP+/+ mice. Either a NOP−/− or NOP+/+ mouse was initially placed in an individual polypropylene chamber and allowed 30 min to acclimatize to laboratory surroundings. The mouse was then exposed to either 30% oxygen/70% nitrogen or 30% oxygen/70% N2O (total gas flow, 6 L/min). Inspired concentrations of each anesthetic were continuously monitored with an infrared analyzer. After 90 min of exposure to each gas, the mouse was anesthetized with intraperitoneal pentobarbital (100 mg/kg) and transcardially perfused with 20 mL of phosphate-buffered saline (PBS) 0.1 M, followed by perfusion with 20 mL of 4% paraformaldehyde in PBS 0.1 M. The mouse was decapitated, and the entire spinal cord was expelled by rapid injection of PBS 0.1 M into the spinal canal at the sacral vertebral level. The spinal cord was stored in 30% sucrose in PBS 0.1 M overnight at 4°C. A portion of the spinal cord at the lumbar level (approximately L5) was sliced into 30-μm-thick sections with a cryotome (Bright Otf Cryostat, Bright Instrument Company, Huntingdon, England) at −20°C, and the slices were collected in PBS 0.1 M as free-floating sections.
Sections were first incubated for 1 h in a blocking solution consisting of 2% skim milk and 0.1% Triton-X in PBS 0.1 M (Buffer 1). They were incubated overnight with rabbit anti-c-Fos antibody (1:20,000; Santa Cruz Biotechnology, Santa Cruz, CA) in Buffer 1 at 4°C. The sections were rinsed with PBS 0.1 M and were incubated for 1 h with biotinylated goat anti-rabbit immunoglobulin (1:200; Vector Laboratories, Burlingame, CA) in Buffer 1. The sections were rinsed with PBS 0.1 M and were incubated for 1 h with avidin-biotin-peroxidase complex (Vector Laboratories) in 1.5% normal goat serum in PBS 0.1 M. Visualization of the immunohistochemical reaction was achieved by incubation with 3,3′-diaminobenzidine (DAB) with nickel-ammonium sulfate, to which hydrogen peroxide was added (DAB kit; Vector Laboratories). After the staining procedure was completed, the sections were rinsed in PBS 0.1 M followed by distilled water and placed on slide glasses, which were dehydrated in 100% ethanol and cleared in 100% xylene, and cover slips were placed.
Using a DAB staining, c-Fos–positive cells were identified by dense black nuclear staining under bright-field microscope (Olympus Model BX60; Olympus Optical, Tokyo, Japan). Randomly selected sections were photographed using a digital camera. The number of c-Fos–positive cells was counted for each area of the spinal cord (i.e., L1–2 [superficial area], L3–4 [nucleus proprius area], L5–6 [neck area], and L7–10 [ventral area]), according to the method previously described (14). Five photos were taken from each mouse, and the mean number of c-Fos–positive cells per section was calculated. The investigator was blinded to the genetic type of the animal and the treatment that the experimental groups had received.
The data were analyzed by unpaired t-test (acetic acid-induced writhing test) and two-way analysis of variance followed by the post hoc test (measurement of plasma ACTH and immunohistochemistry). P < 0.05 was considered to be statistically significant.
Frequent abdominal contractions were observed after intraperitoneal injection of acetic acid both in NOP+/+ and NOP−/− mice. Figure 1 demonstrates that the acetic acid-induced writhings were not significantly different between the two groups of mice under 30% oxygen/70% nitrogen. Exposure to N2O significantly decreased the number of writhings in NOP+/+ mice (80.5% decrease) but not in NOP−/− mice.
Halothane was used to prevent the release of ACTH by stresses other than intraperitoneal acetic acid injection, such as handling, light, and sound. All the mice were sufficiently anesthetized and did not move when they were injected with acetic acid intraperitoneally. When anesthetized with halothane and N2O, the plasma ACTH concentrations in NOP+/+ mice were unchanged at 30 and 60 min after the injection, whereas those of NOP−/− mice were significantly increased at 60 min (335% increase) (Fig. 2).
Mice exposed to 30% oxygen/70% nitrogen were awake and active during the experiment, whereas those exposed to 30% oxygen/70% N2O were excited for the first 15 min of exposure, followed by a relatively calm state. Results from the immunohistochemical study are shown in Fig. 3. There was no significant difference between the numbers of c-Fos–positive cells in the entire gray matter of the spinal cord section of NOP +/+ and NOP−/− mice in the 30% oxygen/70% nitrogen group. Exposure to 70% N2O increased the number of c-Fos–positive cells in NOP+/+ mice. The increase in c-Fos–positive cells was significant in L3–4 and the total (110% and 72% increase, respectively). However, exposure to 70% N2O did not increase the number of c-Fos–positive cells of NOP−/− mice.
In NOP+/+ mice, exposure to N2O significantly reduced the number of writhings induced by intraperitoneal injection of acetic acid, whereas in NOP−/− mice, the number of writhings did not significantly change because of N2O inhalation. Furthermore, it was demonstrated that in NOP−/− mice that N2O has significantly less inhibitory effects on the ACTH release induced by intraperitoneal injection of acetic acid than in NOP+/+ mice. These results together indicate that the antinociceptive action of N2O to the noxious visceral stimulus is affected by the nociceptin system.
The mechanisms of the analgesic action of N2O have been extensively explored. It has been demonstrated that N2O induces opioid peptide release in the PAG of the midbrain, leading to the activation of descending noradrenergic inhibitory neurons, which results in modulation of the nociceptive processing in the spinal cord (5). The descending noradrenergic inhibitory neurons from the brainstem terminate in the spinal cord, mainly in L1–4 (15). It was demonstrated that N2O-induced activation of the descending noradrenergic inhibitory neurons results in c-Fos expression in the γ-aminobutyric acid (GABA)ergic interneurons in the spinal cord, which is suggested to be involved in the antinociceptive action of N2O (16). In the present study, N2O exposure induced c-Fos expression in the spinal cord of NOP+/+ mice in L3–4 and the total spinal cords, which is consistent with the finding in a previous study (16). In contrast, N2O exposure did not increase c-Fos expression in the spinal cord of NOP−/− mice. These results suggest that in NOP−/− mice, the antinociceptive action of N2O via the descending noradrenergic inhibitory neurons is attenuated.
In C57BL/6J mice, prepro-nociceptin messenger (m)RNA and NOP mRNA are expressed in the relay nuclei of the descending inhibitory pathway, including PAG, raphe magnus nucleus, and gigantocellular reticular nucleus (17). The results in the present study may suggest that NOP in these relay nuclei is activated by N2O, leading to the activation of the descending inhibitory system. Further investigations are required to determine whether NOP is actually activated in these relay nuclei by N2O. The mechanism for activation of the descending inhibitory system by NOP also requires investigation.
It was reported that targeted disruption of the nociceptin precursor gene results in an increased plasma corticosterone level in mice in both basal conditions and in response to stress (18), suggesting that the nociceptin system might provide an inhibitory input to the hypothalamus-pituitary-adrenal axis, which constitutes the main integrator of neural processing of stress. This may be consistent with our results that indicate that the effect of N2O to suppress ACTH release in response to noxious visceral stimuli is attenuated in NOP−/− mice. It might be presumed that N2O can activate the endogenous inhibitory input to the hypothalamus-pituitary-adrenal axis, the components of which include nociceptin and NOP.
Although targeted disruption of a gene provides a powerful tool to analyze the function of the gene product, there is a potential problem in this methodology. In NOP−/− mice, compensatory changes may be induced by congenital deficiency of NOP. We cannot completely exclude the possibility that the results obtained in this study are caused by the compensatory changes rather than the defect of NOP function itself. To exclude this possibility, we would need to examine the effects of NOP antagonists on the antinociceptive action of N2O.
In summary, we have demonstrated that N2O exerts less antinociceptive action and induces less c-Fos expression in NOP−/− mice than NOP+/+ mice. It is suggested that the nociceptin system is involved in the antinociceptive action of N2O.
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