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Anesthetic Pharmacology: Research Report

The Endocannabinoid Anandamide Inhibits Voltage-Gated Sodium Channels Nav1.2, Nav1.6, Nav1.7, and Nav1.8 in Xenopus Oocytes

Okura, Dan MD*; Horishita, Takafumi MD, PhD*; Ueno, Susumu MD, PhD; Yanagihara, Nobuyuki PhD; Sudo, Yuka PhD§; Uezono, Yasuhito MD, PhD; Sata, Takeyoshi MD, PhD*

Author Information
doi: 10.1213/ANE.0000000000000070
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Abstract

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.

METHODS

This study was approved by the Animal Research Committee of the University of Occupational and Environmental Health.

Materials

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.

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(V1/2V)/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/(VtVr), 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(V1/2V)/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.

Data Analysis

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.

RESULTS

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).

Figure 1
Figure 1:
Inhibitory effects of anandamide on peak sodium inward currents in Xenopus oocytes expressing Nav1.2, Nav1.6, Nav1.7, and Nav1.8 α subunits with β1 subunits at 2 holding potentials. Representative traces are shown. Sodium currents were evoked by 50-millisecond depolarizing pulses to –20 mV for Nav1.2 and Nav1.6 or –10 mV for Nav1.7 or +10 mV for Nav1.8 from Vmax holding potential (upper panel) or V1/2 holding potential (lower panel) in both the absence and presence of 10 μmol/L anandamide; anandamide was applied for 10 minutes.
Figure 2
Figure 2:
Inhibitory effects of anandamide on peak sodium inward currents in Xenopus oocytes expressing Nav1.2, Nav1.6, Nav1.7, and Nav1.8 α subunits with β1 subunits at 2 holding potentials. Percent inhibition of sodium current of anandamide was calculated. Open columns represent the effect at Vmax holding potential, and closed columns indicate the effect at V1/2 holding potential. Anandamide inhibited 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. Data are represented as the mean ± SEM (n = 5–7). **P < 0.01, compared with the control (based on paired t test).
Figure 3
Figure 3:
Concentration-response curves for anandamide suppression of sodium currents elicited by 50-millisecond depolarizing pulses to –20 mV for Nav1.2 and Nav1.6 or –10 mV for Nav1.7 or +10 mV for Nav1.8 from V1/2 holding potential. The peak current amplitude in the presence of anandamide was normalized to that in the control, and the effects of anandamide are expressed as percentages of the control. Half-maximal inhibitory concentration values and Hill slopes were 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, and 40 ± 14 μmol/L and 0.71 ± 0.10 for Nav1.8, respectively. Data are represented as the mean ± SEM (n = 5–8). Data were fit to the Hill slope equation to give the half-maximal inhibitory concentration values and Hill slopes. Half-maximal inhibitory concentration values and Hill slopes were calculated by using GraphPad Prism.

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).

Table 1
Table 1:
Effects of Anandamide on Activation and Inactivation in Oocytes Expressing Nav1.2, Nav1.6, Nav1.7, and Nav1.8 α Subunits with β1 Subunits
Figure 4
Figure 4:
Effects of anandamide on I-V curves of sodium currents in oocytes expressing Nav1.2 (A), Nav1.6 (B), Nav1.7 (C), and Nav1.8 (D) α subunits with β1 subunits. Currents were elicited by using 50-millisecond depolarizing steps between –80 and 60 mV in 10 mV increments from a Vmax holding potential (left panel) and elicited by using 50-millisecond depolarizing steps between –60 and 60 mV in 10 mV increments from a V1/2 holding potential (right panel); anandamide (30 μmol/L) was applied for 5 minutes; upper panel, representative INa traces from oocytes expressing Nav1.2, Nav1.6, Nav1.7, and Nav1.8 with β1 subunits in both the absence and presence of 30 μmol/L anandamide; lower panel, effects of anandamide on representative I-V curves elicited from Vmax holding potential (left panel) and V1/2 holding potential (right panel) (closed circles, control; open circles, anandamide). Peak currents were normalized to the maximal currents observed from –20 to +10 mV. Data are represented as the mean ± SEM (n = 5–8).
Figure 5
Figure 5:
Effects of anandamide on channel activation in oocytes expressing Nav1.2 (A), Nav1.6 (B), Nav1.7 (C), and Nav1.8 (D) α subunits with β1 subunits from Vmax holding potential (left panels) or V1/2 holding potential (right panels). Closed circles represent control; open circles indicate the effect of anandamide. Data are expressed as the mean ± SEM (n = 5–8). Activation curves were fitted to the Boltzmann equation; V 1/2 is shown in 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).

Figure 6
Figure 6:
Effects of anandamide on inactivation curves in oocytes expressing Nav1.2 (A), Nav1.6 (B), Nav1.7 (C), and Nav1.8 (D) α subunits with β1 subunits. 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-millisecond (500-millisecond for only Nav1.8) prepulses ranging from –140 mV to 0 mV in 10 mV increments from a holding potential of Vmax; anandamide (30 μmol/L) was applied for 5 minutes; right panel, representative INa traces in both the absence and presence of anandamide; left panel, effects of anandamide on inactivation curves (closed circles, control; open circles, anandamide). Steady-state inactivation curves were fitted to the Boltzmann equation, and the V 1/2 values are shown in Table 1. Data are expressed as the mean ± SEM (n = 6–8).

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).

Figure 7
Figure 7:
Use-dependent block of sodium channel on Nav1.2, Nav1.6, Nav1.7, and Nav1.8 α subunits with β1 subunits 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; anandamide was applied for 5 minutes. Peak currents were measured and normalized to the first pulse and plotted against the pulse number (A, Nav1.2; B, Nav1.6; C, Nav1.7; D, Nav1.8). Closed circles represent control; open circles indicate the effect of anandamide. Data were fitted to the monoexponential equation, and values for fractional block of the plateau of normalized INa are shown in (E). Data are expressed as the mean ± SEM (n = 5–6). *P < 0.05 and **P < 0.01, compared with the control (paired t test).

DISCUSSION

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.

DISCLOSURES

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.

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