Cannabis has been used as a pleasure-inducing drug and traditional medicine for thousands of years, and since the 2 cannabinoid receptors CB11,2 and CB23 were identified, the endocannabinoid signaling system has been a focus of medical research and has been considered a potential therapeutic target.4 Endocannabinoids mimic the pharmacological actions of the psychoactive principle agent in marijuana, Δ9-tetrahydrocannabinol, and regulate multiple physiological functions, such as analgesia, regulation of food intake, immunomodulation, inflammation, addictive behavior, epilepsy, and others.5
Anandamide, the ethanolamide of arachidonic acid, was the first endocannabinoid isolated from the brain6; it acts as a partial agonist on CB1 receptors, with a lesser effect on CB2 receptors.7 Several groups have shown an analgesic effect of exogenous anandamide through the CB1 receptor in acute,8–10 persistent inflammatory,11–13 and neuropathic pain models.14,15 CB1 receptors are distributed throughout the pain pathways of the central nervous system (CNS), including the periaqueductal gray, amygdala, and spinal trigeminal tract,16,17 and the peripheral nervous system including the dorsal root ganglion (DRG),18 suggesting an analgesic effect of anandamide via CB1 receptors. However, anandamide may also act on other ion channels consisting of pain signaling pathways, including voltage-gated Ca2+ channels, TASK1 channels, 5-HT3 receptor, rectifying K+ channels, and N-methyl-D-aspartate receptors19–24; thus, the mechanisms of the analgesic effects of anandamide remain unclear.
Voltage-gated sodium channels play an essential role in action potential initiation and propagation in excitable nerve and muscle cells. Nine distinct pore-forming α subunits (Nav1.1–Nav1.9), which are associated with auxiliary β subunits, have been identified,25,26 and each has a different pattern of development and localization as well as distinct physiological and pathophysiological roles. Sodium channel α subunits expressed in DRG (Nav1.7, Nav1.8, Nav1.9) are believed to play crucial roles in inflammatory and neuropathic pain and are considered potential targets of these conditions.27–30 Previous studies have shown that anandamide inhibits sodium channel function in the brain through the inhibition of veratridine-dependent depolarization of synaptosomes31 and suppresses tetrodotoxin-sensitive (TTX-S) and tetrodotoxin-resistant (TTX-R) sodium currents in rat DRG neurons.32 These results suggest that sodium channels are potential targets for anandamide. However, the precise mechanisms of anandamide on each α subunit are still unknown. It is of great importance to clarify these mechanisms because each α subunit has a difference of 20% to 50% in amino acid sequence in the transmembrane and extracellular domains and therefore has different physiological functions. Here, we explored the effects of anandamide on several sodium channel α subunits, including Nav1.2, that is expressed primarily in the CNS; Nav1.6 that is expressed in the CNS and DRG neurons; and Nav1.7 and Nav1.8 that are expressed in DRG neurons.
This study was approved by the Animal Research Committee of the University of Occupational and Environmental Health.
Adult female Xenopus laevis frogs were obtained from Kyudo Co., Ltd. (Saga, Japan). Anandamide was purchased from Sigma-Aldrich (St. Louis, MO). Rat Nav1.2 α subunit cDNA was a gift from Dr. W. A. Catterall (University of Washington, Seattle, WA). Rat Nav1.6 α subunit cDNA was a gift from Dr. A. L. Goldin (University of California, Irvine, CA). Rat Nav1.7 α subunit cDNA was a gift from G. Mandel (Oregon Health and Science University, Portland, OR). Rat Nav1.8 α subunit cDNA was a gift from Dr. A. N. Akopian (University of Texas Health Science Center, San Antonio, TX), and human β1 subunit cDNA was a gift from Dr. A. L. George (Vanderbilt University, Nashville, TN).
cRNA Preparation and Oocyte Injection
After linearization of cDNA with ClaI (Nav1.2 α subunit), NotI (Nav1.6, 1.7 α subunit), XbaI (Nav1.8 α subunit), and EcoRI (β1 subunit), cRNAs were transcribed by using SP6 (1.8 α, β1 subunit) or T7 (Nav1.2, 1.6 1.7 α subunit) RNA polymerase from the mMESSAGE mMACHINE kit (Ambion, Austin, TX). Preparation of X. laevis oocytes and cRNA microinjection were performed as described previously.33 Briefly, stage IV to VI oocytes were manually isolated from a removed portion of ovary. Next, oocytes were treated with collagenase (0.5 mg/mL) for 10 minutes and placed in modified Barth’s solution (88 mmol/L NaCl, 1 mmol/L KCl, 2.4 mmol/L NaHCO3, 10 mmol/L HEPES, 0.82 mmol/L MgSO4, 0.33 mmol/L Ca(NO3)2, and 0.91 mmol/L CaCl2, adjusted to pH 7.5), supplemented with 10,000 U penicillin, 50 mg gentamicin, 90 mg theophylline, and 220 mg sodium pyruvate per liter (incubation medium). Nav α subunit cRNAs were coinjected with β1 subunit cRNA at a ratio of 1:10 (total volume was 20–40 ng/50 nL) into Xenopus oocytes (all α subunits were coinjected with the β1 subunit). Injected oocytes were incubated at 19°C in incubation medium, and 2 to 6 days after injection, the cells were used for electrophysiological recordings.
All electrical recordings were performed at room temperature (23°C). Oocytes were placed in a 100 μL recording chamber and perfused at 2 mL/min with Frog Ringer’s solution containing 115 mmol/L NaCl, 2.5 mmol/L KCl, 10 mmol/L HEPES, 1.8 mmol/L CaCl2, pH 7.2, by using a peristaltic pump (World Precision Instruments Inc., Sarasota, FL). Recording electrodes were prepared with borosilicate glass by using a puller (PP-830, Narishige group company, Tokyo, Japan), and microelectrodes were filled with 3 mol KCl/0.5% low-melting-point agarose with resistances between 0.3 and 0.5 MΩ. The whole-cell voltage clamp was achieved through these 2 electrodes by using a Warner Instruments model OC-725C (Warner, Hamden, CT). Currents were recorded and analyzed by using pCLAMP 7.0 software (Axon Instruments, Foster City, CA), and the amplitude of expressed sodium currents was typically 2 to 15 μA. Transients and leak currents were subtracted by using the P/N procedure. Anandamide stocks were prepared in dimethylsulphoxide (DMSO) and diluted in Frog Ringer’s solution to a final DMSO concentration not exceeding 0.05%. Anandamide was then perfused for 5 to 10 minutes to reach equilibrium.
The voltage dependence of activation was determined by using 50-millisecond depolarizing pulses from a holding potential causing maximal current, Vmax (–90 mV for Nav1.2 and Nav1.6 or –100 mV for Nav1.7 and Nav1.8), and from a holding potential causing half-maximal current, V1/2 (from approximately –40 mV to –70 mV) to 50 mV in 10 mV increments. Normalized activation curves were fitted to the Boltzmann equation: G/Gmax = 1/(1 + exp(V1/2 – V)/k), where G is the voltage-dependent sodium conductance, Gmax is the maximal sodium conductance, G/Gmax is the normalized fractional conductance, V1/2 is the potential at which activation is half maximal, and k is the slope factor. The G value for each oocyte was calculated by using the formula G = I/(Vt – Vr), where I is the peak sodium current, Vt is the test potential and Vr is the reversal potential. The Vr for each oocyte was estimated by extrapolating the linear ascending segment of the current voltage relationship (I–V) curve to the voltage axis. To measure steady-state inactivation, currents were elicited by a 50-millisecond test pulse to –20 mV for Nav1.2 and Nav1.6 or –10 mV for Nav1.7 or +10 mV for Nav1.8 after 200 milliseconds (500 milliseconds for only Nav1.8) prepulses ranging from –140 mV to 0 mV in 10 mV increments from a holding potential of Vmax. Steady-state inactivation curves were fitted to the Boltzmann equation: I/Imax = 1/(1 + exp(V1/2 – V)/k), where Imax is the maximal sodium current, I/Imax is the normalized current, V1/2 is the voltage of half-maximal inactivation, and k is the slope factor. To investigate a use-dependent sodium channel block of anandamide, currents were elicited at 10 Hz by a 20-millisecond depolarizing pulse of –20 mV for Nav1.2 and Nav1.6 or –10 mV for Nav1.7 or +10 mV for Nav1.8 from a V1/2 holding potential in both the absence and presence of 30 μmol/L anandamide. Peak currents were measured and normalized to the first pulse and plotted against the pulse number. Data were fitted to the monoexponential equation INa = exp(–τuse·n) + C, where n is pulse number, C is the plateau INa, and τuse is the time constant of use-dependent decay.
All values are presented as the mean ± SEM (n = 5–8). The n values refer to the number of oocytes examined. Each experiment was performed with oocytes from at least 2 frogs. Control sodium current recorded in absence of anandamide was assigned a value of 100%. Data were statistically evaluated by paired t test by using GraphPad Prism software (GraphPad Software, Inc., San Diego, CA). Hill slope and half-maximal inhibitory concentration values were also calculated by using this software.
Effects of Anandamide on Peak Na+ Inward Currents
Currents were elicited by using a 50-millisecond depolarizing pulse to –20 mV for Nav1.2 and Nav1.6 or –10 mV for Nav1.7 or +10 mV for Nav1.8 applied every 10 seconds from Vmax or V1/2 holding potential in both the absence and presence of 10 μmol/L anandamide (Fig. 1); anandamide was applied for 10 minutes. Anandamide inhibited the peak INa induced by all α subunits more potently at V1/2 than Vmax. Anandamide reduced the peak INa induced by Nav1.2, Nav1.6, Nav1.7, and Nav1.8 by 46 ± 4, 49 ± 3, 37 ± 2, and 27 ± 2 at V1/2, respectively, and 7 ± 2, 6 ± 1, 9 ± 1, and 21 ± 5% at Vmax, respectively (Fig. 2). Inhibition of anandamide at V1/2 was statistically significant in all α subunits, but those at Vmax were not statistically significant except for the suppression in Nav1.8 by paired t test. Because suppression at V1/2 was potent, we examined the concentration-response relation for anandamide inhibition of the peak INa induced by Nav1.2, Nav1.6, Nav1.7, and Nav1.8 at V1/2 holding potential (Fig. 3). The peak current amplitude in the presence of anandamide was normalized to that in the control, and the effects of anandamide were expressed as percentages of the control. Nonlinear regression analyses of the dose-response curves yielded half-maximal inhibitory concentration values and Hill slopes of 17 ± 3 μmol/L and 0.74 ± 0.04 for Nav1.2, 12 ± 1 μmol/L and 0.79 ± 0.08 for Nav1.6, 27 ± 3 μmol/L and 0.52 ± 0.06 for Nav1.7, 40 ± 14 μmol/L and 0.71 ± 0.10 for Nav1.8, respectively (Fig. 3).
Effects of Anandamide on Sodium Current Activation
We examined the effects of anandamide on 4 α subunits of sodium current activation. Voltage dependence of activation was determined by using 50-millisecond depolarizing pulses from a holding potential of Vmax to 50 mV in 10 mV increments or from a holding potential of V1/2 to 50 mV in 10 mV increments for Nav1.2, Nav1.6, Nav1.7, and Nav1.8. Activation curves were derived from the I-V curves (see Methods); anandamide (30 μmol/L) was applied for 5 minutes. The peak INa was reduced by anandamide at Vmax and V1/2 holding potentials with all subunits (Fig. 4). Anandamide shifted the midpoint of steady-state activation (V1/2) in a depolarizing direction at both holding potentials for all subunits (Fig. 5). These shifts were small (1.9–3.8 mV) but statistically significant (Table 1).
Effects of Anandamide on Sodium Current Inactivation
The effect of anandamide on steady-state inactivation was also investigated. Currents were elicited by a 50-millisecond test pulse to –20 mV for Nav1.2 and Nav1.6 or –10 mV for Nav1.7 or +10 mV for Nav1.8 after 200 millis econds(500 milliseconds for only Nav1.8) prepulses ranging from –140 mV to 0 mV in 10 mV increments from a holding potential of Vmax. Steady-state inactivation curves were fitted to the Boltzmann equation (see Methods); anandamide (30 μmol/L) was applied for 5 minutes. Anandamide significantly shifted the midpoint of steady-state inactivation (V1/2) in the hyperpolarizing direction by 5.2, 5.0, 4.1, and 6.3 mV in Nav1.2, Nav1.6, Nav1.7, and Nav1.8, respectively (Fig. 6, Table 1).
Use-Dependent Block of Sodium Currents by Anandamide
We investigated the use-dependent block of sodium currents by anandamide. Currents were elicited at 10 Hz by a 20-millisecond depolarizing pulse of –20 mV for Nav1.2 and Nav1.6 or –10 mV for Nav1.7 or +10 mV for Nav1.8 from a V1/2 holding potential in both the absence and presence of 30 μmol/L anandamide. Peak currents were measured and normalized to the first pulse and plotted against the pulse number (Fig. 7, A–D). Data were fitted by the monoexponential equation (see Methods); anandamide was applied for 5 minutes. Anandamide significantly reduced the plateau INa amplitude of Nav1.2, Nav1.6, and Nav1.7 from 0.74 ± 0.02 to 0.66 ± 0.03, 0.88 ± 0.01 to 0.66 ± 0.02, and 0.73 ± 0.03 to 0.57 ± 0.04, respectively (Fig. 7E), demonstrating a use-dependent block, whereas anandamide did not reduce the plateau INa amplitude of Nav1.8 (from 0.86 ± 0.03 to 0.84 ± 0.04).
In the present study, we demonstrated that anandamide suppresses the Nav1.2, Nav1.6, Nav1.7, and Nav1.8 α subunits in a concentration-dependent manner. half-maximal inhibitory concentration values ranged from 12 μmol/L (Nav1.6) to 40 μmol/L (Nav1.8). Wiley et al.34 have reported that IV administration of anandamide produce a dose-dependent antinociceptive effect in the tail-flick test with mice, and the 50% effective dose (ED50) of that was 15 mg/kg. They also have shown that the plasma concentration of anandamide was 4.96 μg/mL (14.3 μmol/L) when 10 mg/kg of anandamide was administered, suggesting that half-maximal inhibitory concentration values used in the present study are pharmacologically relevant and are close to the plasma concentration exhibiting an antinociceptive effect by anandamide. We also demonstrated that anandamide has more potent inhibitory effects on sodium currents at V1/2 holding potential (inactivated state) than at Vmax holding potential (resting state) in a manner similar to that of local anesthetics on sodium channels. Therefore, the analgesic effects of anandamide may be mediated through sodium channel blockade. The present results are partially consistent with previous reports that anandamide suppresses TTX-S veratridine-dependent depolarization of synaptosomes, the binding of batrachotoxin to sodium channels, and TTX-S sustained repetitive firing in cortical neurons31 and inhibits TTX-S and TTX-R sodium currents in a concentration-dependent manner in rat DRG neurons.32 However, their precise mechanisms of anandamide on several sodium channel α subunits have not yet been investigated. Considering that Nav1.6 was distributed in both CNS and DRG neurons, and that Nav1.8 was distributed in DRG neurons, our results are consistent with a previous study showing that anandamide inhibited sodium currents with half-maximal inhibitory concentration values of 5.4 μmol/L for the TTX-S current and 38 μmol/L for the TTX-R current in DRG neurons,32 suggesting that TTX-S and TTX-R currents in DRG neurons may represent Nav1.6 and Nav1.8 currents, respectively. Because Nav1.6 is expressed in both the brain and DRG, and anandamide suppressed Nav1.6 function most potently among the 4 α subunits, the effect of anandamide on Nav1.6 may be the most important.
The effects of anandamide on channel gating, including activation and inactivation, demonstrated common characteristics among the 4 α subunits we studied. Anandamide shifted the midpoint of steady-state activation (V1/2) in a depolarizing direction at both V1/2 and Vmax holding potentials for all α subunits, and the shifts were significant, although the shifts were small (approximately 4 mV). Anandamide also significantly shifted the midpoint of steady-state inactivation (V1/2) in the hyperpolarizing direction (approximately 7 mV) for all α subunits. These results suggest that both inhibition of activation and the enhancement of inactivation are common mechanisms of sodium current inhibition by anandamide for Nav1.2, Nav1.6, Nav1.7, and Nav1.8. A combination of effects on both activation and inactivation might produce sufficient effects to suppress sodium currents although each effect is small. Inhibition by anandamide at Vmax holding potential for Nav1.2, Nav1.6, and Nav1.7 was small and not significant, whereas that for Nav1.8 was significant (Fig. 1), indicating that resting-channel block is one of the important mechanisms of anandamide inhibition for only Nav1.8. Anandamide exhibited use-dependent block with repetitive stimuli for Nav1.2, Nav1.6, and Nav1.7 but not Nav1.8. The presence of use-dependent block by anandamide suggests the possibility of open-channel block and the ability to slow the recovery time from blocks that are seen with amitriptyline.35 Sodium channel blockers such as local anesthetics, tricyclic antidepressants, and volatile anesthetics have been shown to shift the voltage dependence of steady-state inactivation with no effect on activation and exhibit use-dependent block.35–39 Our results show that anandamide shows a negative shift in the voltage dependence of inactivation and use-dependent block except for Nav1.8 that are seen with other sodium channel blockers yet also shifts the steady-state activation in a depolarizing direction, suggesting that it may have different binding sites or allosteric conformational mechanisms for these sodium channel antagonists. Moreover, a resting-channel block, not an open-channel block, for Nav1.8 may be a key for exploring the mechanism of sodium channel inhibition by anandamide in detail.
Several groups have evaluated antinociception by exogenous anandamide via CB1 receptors.8–10 Indeed, a recent review has shown that activation of both CB1 and CB2 receptors reduces nociceptive processing in acute and chronic animal models of pain.40 Alternatively, other investigators have suggested that anandamide produces antinociception through a CB1-independent mechanism. For example, anandamide antinociception is not blocked by pretreatment with the selective CB1 antagonist SR141716A.41 Rapid metabolism of anandamide to arachidonic acid has been shown to be one of the reasons for the failure of SR141716A to antagonize the effects of anandamide; in experiments, the ability of SR141716A to reverse anandamide antinociception was improved (but not completely) when anandamide metabolism to arachidonic acid was inhibited with coadministration of an amidase inhibitor, phenylmethylsulfonyl fluoride.42 That study also demonstrated that cyclooxygenase did not alter the effects of anandamide, whereas it blocked the effects of arachidonic acid, suggesting a pain-inhibitory effect of anandamide by noncannabinoid mechanisms. Another recent study suggested that anandamide induced antinociception by stimulating endogenous norepinephrine release that activated peripheral adrenoceptors inducing antinociception, although whether the effect was caused through cannabinoid receptors remains unknown.43
This study indicates that sodium channel inhibition by anandamide is independent of signaling through cannabinoid receptors, because in recombinant experiments such as our present examination, the effects on channels or receptors can be excluded except for that expressed in membranes. Previous reports also indicate a direct effect of anandamide on sodium channels by demonstrating that sodium channel-related activities by anandamide in the brain may be independent of the presence of AM 251 (a CB1 antagonist),31 AM 251, AM 630 (a CB2 antagonist) and capsazepine (a vanilloid receptor type 1 antagonist) do not interfere with anandamide suppression of sodium currents in DRG.32 Therefore, we believe that the effects of anandamide on Nav1.2, Nav1.6, Nav1.7, and Nav1.8 α subunits are direct. Taken together, to the best of our knowledge, this is the first direct evidence to demonstrate the inhibitory effects and its mechanisms on neuronal sodium channel α subunits in recombinant experiment systems.
Several sodium channel α subunits are believed to be involved in the pathogenesis of inflammatory and neuropathic pain. Mutations in Nav1.7 have been linked to inherited pain syndromes, including inherited erythromelalgia, that is characterized by episodes of burning pain, erythema, mild swelling in the hands and feet,44 and paroxysmal extreme pain disorder (PEPD), which is characterized by severe rectal, ocular, and mandibular pain.45 Recently, anandamide has been reported to inhibit resurgent current of wild-type Nav1.7 and the PEPD mutants expressed in transfected human embryonic kidney 293 cells, and this inhibition was suggested as a therapeutic target for PEPD patients.46 Nav1.8 has demonstrated its ability to carry most current underlying the upstroke of the action potential in nociceptive neurons,47 and the use of Nav1.8 knockdown rats after antisense oligodeoxynucleotide treatment has demonstrated a role for Nav1.8 in inflammatory pain,48 whereas Nav1.8 expression has been reported to increase in nerves proximal to injury sites in patients with chronic neuropathic pain.49 In an infraorbital nerve injury model of rats, the level of Nav1.6 protein was significantly increased proximal to the lesion site, suggesting a role of Nav1.6 in neuropathic pain conditions.50 However, these α subunits highly expressed in normal DRG have been reported to show diverse expression in DRG of inflammatory and neuropathic pain models. Nav1.7 mRNA and protein increased in DRG after peripheral inflammation induced by carrageenan,51,52 whereas Nav1.7 protein decreased in the injured DRG after spared nerve injury in animals.53 Nav1.8 mRNA and protein increased in DRG neurons of rodents after injection of carrageenan into a hindpaw,51,54,55 and yet peripheral nerve injury down-regulates Nav1.8 mRNA and protein expression in the injured DRG.29,53,56 Based on this evidence, suppression of sensory neuron sodium channel function by anandamide may be an important mechanism independent of the cannabinoid receptor. Because of the limitations of our experiments, further investigation is warranted to extrapolate our findings into clinical practice.
In conclusion, anandamide at pharmacologically relevant concentrations inhibited sodium currents of Nav1.2, Nav1.6, Nav1.7, and Nav1.8 α subunits expressed in the Xenopus oocytes with differences in the effects on sodium channel gating. These results provide a better understanding of the mechanisms underlying the analgesic effects of anandamide, but further studies are needed to clarify the relevance of sodium channel inhibition by anandamide to analgesia.
Name: Dan Okura, MD.
Contribution: This author helped data collection, data analysis, and manuscript preparation.
Attestation: Dan Okura approved the final manuscript and attests to the integrity of the original data and the analysis reported in this manuscript.
Name: Takafumi Horishita, MD, PhD.
Contribution: This author helped study design, data collection, data analysis, and manuscript preparation.
Attestation: Takafumi Horishita approved the final manuscript and attests to the integrity of the original data and the analysis reported in this manuscript, and also is the archival author.
Name: Susumu Ueno, MD, PhD.
Contribution: This author helped conduct of the study and manuscript preparation.
Attestation: Susumu Ueno approved the final manuscript.
Name: Nobuyuki Yanagihara, PhD.
Contribution: This author helped conduct of the study and manuscript preparation.
Attestation: Nobuyuki Yanagihara approved the final manuscript.
Name: Yuka Sudo, PhD.
Contribution: This author helped conduct of the study.
Attestation: Yuka Sudo approved the final manuscript.
Name: Yasuhito Uezono, MD, PhD.
Contribution: This author helped conduct of the study.
Attestation: Yasuhito Uezono approved the final manuscript.
Name: Takeyoshi Sata, MD, PhD.
Contribution: This author helped conduct of the study and manuscript preparation.
Attestation: Takeyoshi Sata approved the final manuscript.
This manuscript was handled by: Marcel E. Durieux, MD, PhD.
1. Devane WA, Dysarz FA 3rd, Johnson MR, Melvin LS, Howlett AC. Determination and characterization of a cannabinoid receptor in rat brain. Mol Pharmacol. 1988;34:605–13
2. Matsuda LA, Lolait SJ, Brownstein MJ, Young AC, Bonner TI. Structure of a cannabinoid receptor and functional expression of the cloned cDNA. Nature. 1990;346:561–4
3. Munro S, Thomas KL, Abu-Shaar M. Molecular characterization of a peripheral receptor for cannabinoids. Nature. 1993;365:61–5
4. Zias J, Stark H, Sellgman J, Levy R, Werker E, Breuer A, Mechoulam R. Early medical use of cannabis. Nature. 1993;363:215
5. Pacher P, Bátkai S, Kunos G. The endocannabinoid system as an emerging target of pharmacotherapy. Pharmacol Rev. 2006;58:389–462
6. Devane WA, Hanus L, Breuer A, Pertwee RG, Stevenson LA, Griffin G, Gibson D, Mandelbaum A, Etinger A, Mechoulam R. Isolation and structure of a brain constituent that binds to the cannabinoid receptor. Science. 1992;258:1946–9
7. Pertwee RG, Ross RA. Cannabinoid receptors and their ligands. Prostaglandins Leukot Essent Fatty Acids. 2002;66:101–21
8. Costa B, Vailati S, Colleoni M. SR 141716A, a cannabinoid receptor antagonist, reverses the behavioural effects of anandamide-treated rats. Behav Pharmacol. 1999;10:327–31
9. Mason DJ Jr, Lowe J, Welch SP. Cannabinoid modulation of dynorphin A: correlation to cannabinoid-induced antinociception. Eur J Pharmacol. 1999;378:237–48
10. Welch SP, Huffman JW, Lowe J. Differential blockade of the antinociceptive effects of centrally administered cannabinoids by SR141716A. J Pharmacol Exp Ther. 1998;286:1301–8
11. Calignano A, La Rana G, Giuffrida A, Piomelli D. Control of pain initiation by endogenous cannabinoids. Nature. 1998;394:277–81
12. Richardson JD, Kilo S, Hargreaves KM. Cannabinoids reduce hyperalgesia and inflammation via interaction with peripheral CB1 receptors. Pain. 1998;75:111–9
13. Guindon J, De Léan A, Beaulieu P. Local interactions between anandamide, an endocannabinoid, and ibuprofen, a nonsteroidal anti-inflammatory drug, in acute and inflammatory pain. Pain. 2006;121:85–93
14. Sagar DR, Kendall DA, Chapman V. Inhibition of fatty acid amide hydrolase produces PPAR-alpha-mediated analgesia in a rat model of inflammatory pain. Br J Pharmacol. 2008;155:1297–306
15. Karbarz MJ, Luo L, Chang L, Tham CS, Palmer JA, Wilson SJ, Wennerholm ML, Brown SM, Scott BP, Apodaca RL, Keith JM, Wu J, Breitenbucher JG, Chaplan SR, Webb M. Biochemical and biological properties of 4-(3-phenyl-[1,2,4] thiadiazol-5-yl)-piperazine-1-carboxylic acid phenylamide, a mechanism-based inhibitor of fatty acid amide hydrolase. Anesth Analg. 2009;108:316–29
16. Tsou K, Brown S, Sañudo-Peña MC, Mackie K, Walker JM. Immunohistochemical distribution of cannabinoid CB1 receptors in the rat central nervous system. Neuroscience. 1998;83:393–411
17. Farquhar-Smith WP, Egertová M, Bradbury EJ, McMahon SB, Rice AS, Elphick MR. Cannabinoid CB(1) receptor expression in rat spinal cord. Mol Cell Neurosci. 2000;15:510–21
18. Hohmann AG, Herkenham M. Localization of central cannabinoid CB1 receptor messenger RNA in neuronal subpopulations of rat dorsal root ganglia: a double-label in situ hybridization study. Neuroscience. 1999;90:923–31
19. Chemin J, Monteil A, Perez-Reyes E, Nargeot J, Lory P. Direct inhibition of T-type calcium channels by the endogenous cannabinoid anandamide. EMBO J. 2001;20:7033–40
20. Mackie K, Lai Y, Westenbroek R, Mitchell R. Cannabinoids activate an inwardly rectifying potassium conductance and inhibit Q-type calcium currents in AtT20 cells transfected with rat brain cannabinoid receptor. J Neurosci. 1995;15:6552–61
21. Maingret F, Patel AJ, Lazdunski M, Honoré E. The endocannabinoid anandamide is a direct and selective blocker of the background K(+) channel TASK-1. EMBO J. 2001;20:47–54
22. Fan P. Cannabinoid agonists inhibit the activation of 5-HT3 receptors in rat nodose ganglion neurons. J Neurophysiol. 1995;73:907–10
23. Poling JS, Rogawski MA, Salem N Jr, Vicini S. Anandamide, an endogenous cannabinoid, inhibits Shaker-related voltage-gated K+ channels. Neuropharmacology. 1996;35:983–91
24. Mendiguren A, Pineda J. Cannabinoids enhance N-methyl-D-aspartate-induced excitation of locus coeruleus neurons by CB1 receptors in rat brain slices. Neurosci Lett. 2004;363:1–5
25. Catterall WA. From ionic currents to molecular mechanisms: the structure and function of voltage-gated sodium channels. Neuron. 2000;26:13–25
26. Catterall WA, Goldin AL, Waxman SG. International Union of Pharmacology. XLVII. Nomenclature and structure-function relationships of voltage-gated sodium channels. Pharmacol Rev. 2005;57:397–409
27. Wood JN, Boorman JP, Okuse K, Baker MD. Voltage-gated sodium channels and pain pathways. J Neurobiol. 2004;61:55–71
28. Cummins TR, Sheets PL, Waxman SG. The roles of sodium channels in nociception: implications for mechanisms of pain. Pain. 2007;131:243–57
29. Decosterd I, Ji RR, Abdi S, Tate S, Woolf CJ. The pattern of expression of the voltage-gated sodium channels Na(v)1.8 and Na(v)1.9 does not change in uninjured primary sensory neurons in experimental neuropathic pain models. Pain. 2002;96:269–77
30. Wang W, Gu J, Li YQ, Tao YX. Are voltage-gated sodium channels on the dorsal root ganglion involved in the development of neuropathic pain? Mol Pain. 2011;7:16
31. Nicholson RA, Liao C, Zheng J, David LS, Coyne L, Errington AC, Singh G, Lees G. Sodium channel inhibition by anandamide and synthetic cannabimimetics in brain. Brain Res. 2003;978:194–204
32. Kim HI, Kim TH, Shin YK, Lee CS, Park M, Song JH. Anandamide suppression of Na+ currents in rat dorsal root ganglion neurons. Brain Res. 2005;1062:39–47
33. Horishita T, Eger EI 2nd, Harris RA. The effects of volatile aromatic anesthetics on voltage-gated Na+ channels expressed in Xenopus oocytes. Anesth Analg. 2008;107:1579–86
34. Wiley JL, Dewey MA, Jefferson RG, Winckler RL, Bridgen DT, Willoughby KA, Martin BR. Influence of phenylmethylsulfonyl fluoride on anandamide brain levels and pharmacological effects. Life Sci. 2000;67:1573–83
35. Wang GK, Russell C, Wang SY. State-dependent block of voltage-gated Na+ channels by amitriptyline via the local anesthetic receptor and its implication for neuropathic pain. Pain. 2004;110:166–74
36. Ragsdale DS, McPhee JC, Scheuer T, Catterall WA. Molecular determinants of state-dependent block of Na+ channels by local anesthetics. Science. 1994;265:1724–8
37. Osawa Y, Oda A, Iida H, Tanahashi S, Dohi S. The effects of class Ic antiarrhythmics on tetrodotoxin-resistant Na+ currents in rat sensory neurons. Anesth Analg. 2004;99:464–71, table of contents
38. Poyraz D, Bräu ME, Wotka F, Puhlmann B, Scholz AM, Hempelmann G, Kox WJ, Spies CD. Lidocaine and octanol have different modes of action at tetrodotoxin-resistant Na(+) channels of peripheral nerves. Anesth Analg. 2003;97:1317–24
39. Ouyang W, Herold KF, Hemmings HC Jr. Comparative effects of halogenated inhaled anesthetics on voltage-gated Na+ channel function. Anesthesiology. 2009;110:582–90
40. Starowicz K, Malek N, Przewlocka B. Cannabinoid receptors and pain. Wiley Interdiscip Rev Membr Transp Signal. 2013;2:121–32
41. Adams IB, Compton DR, Martin BR. Assessment of anandamide interaction with the cannabinoid brain receptor: SR 141716A antagonism studies in mice and autoradiographic analysis of receptor binding in rat brain. J Pharmacol Exp Ther. 1998;284:1209–17
42. Wiley JL, Razdan RK, Martin BR. Evaluation of the role of the arachidonic acid cascade in anandamide’s in vivo
effects in mice. Life Sci. 2006;80:24–35
43. Romero TR, Resende LC, Guzzo LS, Duarte ID. CB1 and CB2 cannabinoid receptor agonists induce peripheral antinociception by activation of the endogenous noradrenergic system. Anesth Analg. 2013;116:463–72
44. Waxman SG, Dib-Hajj S. Erythermalgia: molecular basis for an inherited pain syndrome. Trends Mol Med. 2005;11:555–62
45. Fertleman CR, Ferrie CD, Aicardi J, Bednarek NA, Eeg-Olofsson O, Elmslie FV, Griesemer DA, Goutières F, Kirkpatrick M, Malmros IN, Pollitzer M, Rossiter M, Roulet-Perez E, Schubert R, Smith VV, Testard H, Wong V, Stephenson JB. Paroxysmal extreme pain disorder (previously familial rectal pain syndrome). Neurology. 2007;69:586–95
46. Theile JW, Cummins TR. Inhibition of Navβ4 peptide-mediated resurgent sodium currents in Nav1.7 channels by carbamazepine, riluzole, and anandamide. Mol Pharmacol. 2011;80:724–34
47. Renganathan M, Cummins TR, Waxman SG. Contribution of Na(v)1.8 sodium channels to action potential electrogenesis in DRG neurons. J Neurophysiol. 2001;86:629–40
48. Joshi SK, Mikusa JP, Hernandez G, Baker S, Shieh CC, Neelands T, Zhang XF, Niforatos W, Kage K, Han P, Krafte D, Faltynek C, Sullivan JP, Jarvis MF, Honore P. Involvement of the TTX-resistant sodium channel Nav 1.8 in inflammatory and neuropathic, but not post-operative, pain states. Pain. 2006;123:75–82
49. Black JA, Nikolajsen L, Kroner K, Jensen TS, Waxman SG. Multiple sodium channel isoforms and mitogen-activated protein kinases are present in painful human neuromas. Ann Neurol. 2008;64:644–53
50. Henry MA, Freking AR, Johnson LR, Levinson SR. Sodium channel Nav1.6 accumulates at the site of infraorbital nerve injury. BMC Neurosci. 2007;8:56
51. Black JA, Liu S, Tanaka M, Cummins TR, Waxman SG. Changes in the expression of tetrodotoxin-sensitive sodium channels within dorsal root ganglia neurons in inflammatory pain. Pain. 2004;108:237–47
52. Strickland IT, Martindale JC, Woodhams PL, Reeve AJ, Chessell IP, McQueen DS. Changes in the expression of NaV1.7, NaV1.8 and NaV1.9 in a distinct population of dorsal root ganglia innervating the rat knee joint in a model of chronic inflammatory joint pain. Eur J Pain. 2008;12:564–72
53. Berta T, Poirot O, Pertin M, Ji RR, Kellenberger S, Decosterd I. Transcriptional and functional profiles of voltage-gated Na(+) channels in injured and non-injured DRG neurons in the SNI model of neuropathic pain. Mol Cell Neurosci. 2008;37:196–208
54. Okuse K, Chaplan SR, McMahon SB, Luo ZD, Calcutt NA, Scott BP, Akopian AN, Wood JN. Regulation of expression of the sensory neuron-specific sodium channel SNS in inflammatory and neuropathic pain. Mol Cell Neurosci. 1997;10:196–207
55. Coggeshall RE, Tate S, Carlton SM. Differential expression of tetrodotoxin-resistant sodium channels Nav1.8 and Nav1.9 in normal and inflamed rats. Neurosci Lett. 2004;355:45–8
56. Cummins TR, Waxman SG. Downregulation of tetrodotoxin-resistant sodium currents and upregulation of a rapidly repriming tetrodotoxin-sensitive sodium current in small spinal sensory neurons after nerve injury. J Neurosci. 1997;17:3503–14