Life Sciences 136 (2015) 79–86
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Excitatory effect of Neurotropin® on noradrenergic neurons in rat locus coeruleus Hisashi Okai a,⁎, Ryohei Okazaki a, Minoru Kawamura a, Megumu Yoshimura b a b
Department of Pharmacological Research, Institute of Bio-Active Science, Nippon Zoki Pharmaceutical Co., Ltd., 442-1, Kinashi, Kato, Hyogo 673-1461, Japan Graduate School of Health Sciences, Kumamoto Health Science University, Kumamoto 861-5598, Japan
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Article history: Received 19 December 2014 Received in revised form 14 May 2015 Accepted 4 June 2015 Available online 2 July 2015 Keywords: Neurotropin Neuropathic pain Locus coeruleus Descending pain inhibitory system Noradrenaline In vivo patch clamp
a b s t r a c t Aims: Although the clinical use of Neurotropin® as an analgesic for chronic pain has been firmly established, its analgesic mechanism is still unclear. In this study, we investigate the direct effects of Neurotropin using an electrophysiological method. Main methods: Blind patch-clamp recordings were made from rat locus coeruleus (LC) and periaqueductal gray (PAG) neurons in brainstem slices of normal rats. The effects of intracerebroventricular (icv) injection of Neurotropin on nociceptive transmission were recorded from spinal substantia gelatinosa (SG) neurons in fifth lumbar spinal nerve-ligated (L5-SNL) rats using an in vivo patch-clamp method. Key findings: Neurotropin (0.2–1.0 NU/mL) dose-dependently increased the firing rate in noradrenergic LC neurons of normal rats. Under the voltage-clamp condition, Neurotropin induced an inward current in 90% of LC neurons that was not affected by tetrodotoxin or an injection of GDP-β-S (G protein inhibitor) through recording pipettes. In contrast, Neurotropin had no effects on all PAG neurons tested. Using in vivo patch-clamp recordings, the icv injection of Neurotropin inhibited both frequency and amplitude of pinch-evoked excitatory postsynaptic currents of SG neurons in L5-SNL rats. These results suggest that Neurotropin directly excites the descending noradrenergic LC neurons and inhibits nociceptive transmission in the spinal dorsal horn. Significance: This study is the first direct demonstration that Neurotropin activates the noradrenergic descending pain inhibitory systems, and this would reinforce the usefulness of Neurotropin in the treatment of human neuropathic pain. © 2015 Elsevier Inc. All rights reserved.
1. Introduction Neurotropin, a non-protein extract from the inflamed skin of rabbits inoculated with vaccinia virus, is widely used in Japan and China as an analgesic for chronic pain such as postherpetic neuralgia and other neuropathic pain. A double-blind placebo-controlled cross-over study is currently underway at the National Institute of Nursing Research in the United States to investigate the effectiveness of Neurotropin in the treatment of fibromyalgia. Neurotropin is recommended as a secondline treatment in the guideline for managing neuropathic pain in Japan [29]. Analgesic mechanisms of Neurotropin have been examined mainly in a chronic stress model — namely, SART (Specific Alternation of Rhythm in Environmental Temperature)-stressed animals [7,8,12,20], which exhibit a chronic hyperalgesic state due to repeated periods of cold stress. The analgesic effect of systemic administration of Neurotropin on SART-stressed rats is antagonized by inhibition of spinal α2 adrenoceptors or serotonergic 5-HT3 receptors, but not opioid μ
⁎ Corresponding author. E-mail address: [email protected] (H. Okai).
http://dx.doi.org/10.1016/j.lfs.2015.06.013 0024-3205/© 2015 Elsevier Inc. All rights reserved.
receptors [14]. These results suggest that Neurotropin activates the monoaminergic descending pain inhibitory systems, but not opioidergic neurons in SART-stressed rats. Neurotropin ameliorates chronic inflammation-induced hyperalgesia and L5 spinal nerve ligation (L5-SNL)-induced mechanical allodynia, and these analgesic effects are also depressed by intrathecally administered α2 adrenoceptor antagonist or 5-HT3 receptor antagonist [17,22,28]. Furthermore, the anti-allodynic effect of Neurotropin is more marked following intracerebroventricular (icv) administration compared to intrathecal administration [22]. Therefore, it may be presumed that Neurotropin activates the supraspinal site of monoaminergic descending pain inhibitory systems. The midbrain periaqueductal gray (PAG) projects to the locus coeruleus (LC) and rostral ventromedial medulla (RVM) [2]. Therefore, the analgesic effect produced by stimulation of PAG is mediated via the activation of LC and/or RVM. It is well known that the nucleus raphe magnus (NRM) [5], which is the main nucleus in the RVM, contains 5-HT and other inhibitory transmitters such as GABA and glycine [1,11,25]. Based on behavioral studies [14], the antinociceptive effects of Neurotropin might be a result of activation of the supraspinal LC, PAG and/or NRM. Electrophysiologically, however, it is still unclear whether Neurotropin acts on one, or all three, brain areas.
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Previously, we demonstrated that Neurotropin directly excites NRM neurons in brainstem slices of normal rats [21]. Our present study, therefore, examined the direct effects of Neurotropin on PAG and LC neurons. Since it has previously been shown that systemic administration of Neurotropin causes a dose-dependent inhibition of hyperalgesia in the adjuvant-induced arthritic rat but has no significant effects on the nociceptive threshold in normal rats [17], L5-SNL chronic pain model rats were used for in vivo examinations. In vivo patch-clamp recordings from spinal SG neurons were performed to examine the effects of icv administration of Neurotropin on excitatory inputs from the receptive fields. 2. Materials and methods 2.1. Drugs Neurotropin was prepared by the Nippon Zoki Pharmaceutical Co., Ltd. (Osaka, Japan). [D-Ala2, N-MePhe4, Gly-ol]-enkephalin (DAMGO) was purchased from Sigma-Aldrich (St. Louis, MO). 2-Amino-3-(3hydroxy-5-methyl-isoxazol-4-yl) propanoic acid (AMPA) was purchased from Funakoshi Co., Ltd. (Tokyo, Japan). Tetrodotoxin (TTX), a sodium channel blocker, was purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). These drugs were dissolved or diluted in Krebs solution before use. Guanosine-5′-O-(2-thiodiphosphate) (GDP-β-S, G protein inhibitors) was purchased from Sigma-Aldrich (St. Louis, MO).
that was equilibrated with 95% O2 and 5% CO2 at 37 ± 1 °C. Drugs and Krebs solution were applied to the slices by perfusion via a three-way stopcock without any changes in the perfusion rate or temperature. 2.5. Patch-clamp recordings Blind whole-cell patch-clamp recordings were made from normal rat LC, ventrolateral PAG, and SG neurons in the spinal cord with patch electrodes (tip resistances: 6–12 MΩ). The patch pipettes were filled with a solution containing (mM): K-gluconate 135, KCl 5, CaCl2 0.5, MgCl2 2, EGTA 5, HEPES 5, ATP-Mg 5 and pH 7.2 for excitatory postsynaptic currents (EPSCs) recordings, or containing (mM): cesium sulfate 110, CaCl2 0.5, MgCl2 2, EGTA 5, HEPES 5, ATP-Mg 5 and pH 7.2 for inhibitory postsynaptic currents (IPSCs) recordings. GDP-β-S was added to the solutions in patch pipettes at 2 mM. Signals were acquired with a patch-clamp amplifier (Axopatch 200B amplifier, Molecular Devices, CA, USA). Data were digitized with an A/D converter (digidata1400, Molecular Devices), stored on a personal computer using a data acquisition program (pClamp10.2, Molecular Devices, CA, USA), and analyzed with Clampfit software (clampfit10.2, Molecular Devices, CA, USA) and AxoGraph X (AxoGraph Scientific, CA, USA). In the voltage-clamp mode, the holding potentials were held at −70 mV (at which glycine-/GABA-mediated IPSCs were negligible) or 0 mV (at which glutamate-mediated EPSCs were negligible). Frequencies of action potentials were monitored in the zero current-clamp mode. 2.6. In vivo patch-clamp recordings
2.2. Animals Six-week-old male SD rats were purchased from Kyudo Co., Ltd. (Saga, Japan). The animals were housed under a 12-h light–dark cycle with free access to food and water. At the time of the experiment, the animals ranged in age from 7 to 11 weeks. All experimental procedures were approved by the Ethics Committee on Animal Experiments at Kyushu University (Fukuoka, Japan), or an ethical committee on animal experiments at the Nippon Zoki Pharmaceutical Co., Ltd. (Kato, Japan), and performed in accordance with the ethical guidelines for investigations of experimental pain in conscious animals issued by the International Association for the Study of Pain [31]. 2.3. Surgical preparation and behavioral testing
In vivo patch-clamp recordings were performed according to the method of Furue et al. [6]. Briefly, after rats were deeply anesthetized with urethane (1.0–1.2 g/kg, i.p.), the lumbar enlargement was exposed by a thoraco-lumbar laminectomy at the T12 to L2 levels. Then, rats were placed in a stereotaxic apparatus (Model ST-7, Narishige, Tokyo, Japan). After the dura mater was cut and reflected, a dorsal root that entered the spinal dorsal horn covering the recording region was gently moved aside with a glass retractor to expose the lateral edge of the SG. The pia-arachnoid membrane was then cut with fine scissors to make a small window to allow a patch electrode to enter the SG. The surface of the exposed spinal cord was irrigated with Krebs solution equilibrated with 95% O2–5% CO2 at 37 ± 1 °C. The noxious pinch stimuli were applied to the ipsilateral receptive field of the recorded neuron using a clamp applying 200 g of force. The clamp had previously been calibrated using a pull tension gauge.
L5-SNL model rats were prepared according to the method of Kim and Chung [16]. Briefly, the right L5 spinal nerve was isolated and tightly ligated with a 5–0 silk suture under pentobarbital (40 mg/kg, i.p.) anesthesia. After surgery, rats were housed in a cage in groups of three with free access to food and water and allowed to recover over 14 days. Right paw mechanical allodynia was confirmed before in vivo patch-clamp recording by measuring the hindpaw withdrawal threshold in response to application of von Frey filaments using the up–down method [3,4]. Only those rats which showed a withdrawal threshold of less than 4 g were used for in vivo patch-clamp recording. Behavioral tests were performed before the patch-clamp recordings at 14–28 days after nerve ligation.
For icv injection, the shaved scalp was incised to expose the skull, and a hole was drilled into the skull above the right lateral ventricle (distance from interaural: anterior, +8.2 mm; lateral, +1.3 mm; vertical, +6.0 mm) according to the stereotaxic coordinates of the atlas of Paxinos and Watson [24]. Then, using a 30-gauge needle tip inserted into the right lateral ventricle and a CMA/100 microinjection pump (CMA Microdialysis AB, Stockholm, Sweden), 15 μL of each drug solution was applied over 1 min.
2.4. Brain slice preparations
2.8. Immunohistochemistry
Rat brain slice preparations were obtained according to the method of Nakatsuka et al. [18]. Rats were deeply anesthetized with urethane (1.2 g/kg, i.p.), and then decapitated without delay. The whole brain was removed and placed in an ice-cold, pre-oxygenated Krebs solution (in mM: NaCl 117, KCl 3.6, CaCl2 2.5, MgCl2 1.2, NaH2PO4 1.2, NaHCO3 25, and D-glucose 11). After removal of the dura mater, the brainstem, including LC or PAG, was coronally sliced at 400 μm thickness using a microslicer. The slices were then placed in a recording chamber and perfused continuously at a rate of 10–15 mL/min with Krebs solution
Recorded LC neurons were labeled with neurobiotin (0.2% in the electrode solution; Vector Laboratories, CA, USA) diffused from the patch pipettes after whole-cell configuration. After the electrophysiological recordings, brain slices were immersed in 4% paraformaldehyde/0.1 mM phosphate-buffered saline (PBS) overnight, and embedded in paraffin. Slices were sectioned coronally (8 μm) using a microtome (Daiwa Koki, K.K., Saitama, Japan), and incubated with anti-tyrosine hydroxylase antibody (1:200 dilution in PBS; Enzo Life Sciences, Inc., NY, USA) overnight at 4 °C. After rinsing in PBS, the sections
2.7. Icv injection of Neurotropin
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were incubated with FITC-conjugated AffiniPure Donkey Anti-rabbit IgG (dilution 1:100; FITC-IgG; Jackson Immuno Research Laboratories, Inc., West Grove, PA) and Texas Red® dye-conjugated streptavidin (Jackson Immuno Research Laboratories, Inc., West Grove, PA) for 6 h. The sections were mounted on gelatinized slides and photographed with a fluorescence microscope (TE2000, Nikon, Tokyo, Japan). 2.9. Statistical analysis All numerical data were provided as means ± standard error of the mean (SEM). The statistical significance between two groups was determined by Student's t-test or the paired t-test. Statistical significance between more than two groups was analyzed using Dunnett's multiple comparison tests. p values less than 0.05 were considered to be significant.
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neurons were stained with tyrosine hydroxylase (Fig. 2B). All recorded neurons injected with neurobiotin (n = 10) were positive to tyrosine hydroxylase and, therefore, noradrenergic (Fig. 2C), and nine out of ten neurons were responsive to 1 NU/mL of Neurotropin (Fig. 2D). The Neurotropin-induced inward current was not affected by pretreatment with TTX (1 μM, Fig. 3A, B). The frequency and amplitude of miniature EPSCs (mEPSCs) were also not affected by the application of Neurotropin (Fig. 3C, D). Immediately after establishment of the whole-cell recording configuration with pipettes containing GDP-β-S, DAMGO (1 μM) induced an outward current and Neurotropin induced an inward current (Fig. 4A). After 30 min, GDP-β-S had diffused into LC neurons, blocking the DAMGO-induced outward currents (4.3 ± 2.8% compared to the current just after establishment of the wholecell configuration, n = 7) whereas Neurotropin-induced inward currents were unaffected (Fig. 4B, C). 3.2. Effects of Neurotropin on ventrolateral PAG neurons
3. Results 3.1. Effects of Neurotropin on LC neurons In the current-clamp mode, spontaneous action potentials were observed in all LC neurons tested in vitro. Bath-applied Neurotropin concentration-dependently increased the frequency of action potentials. At concentrations of 0.2, 0.5 and 1.0 NU/mL of Neurotropin, the frequency of action potentials was increased to 165.2 ± 32.8% (n = 6), 380.0 ± 15.2% (n = 6), and 771.2 ± 273.5% (n = 6), respectively (Fig. 1A). Conversely, after Neurotropin was washed off the samples, the frequency of action potentials decreased temporarily, canceling the previous increase in frequency of action potentials for a few seconds. This might have been due to a hyperpolarizing response by Neurotropin following depolarization but further analysis of this was beyond the scope of the present study. In the voltage-clamp mode at a holding potential of −70 mV, Neurotropin (0.2–1.0 NU/mL) induced an inward current in a clear concentration-dependent manner (Fig. 1B). At 0.2, 0.5 and 1.0 NU/mL, Neurotropin induced inward currents of 13.6 ± 2.1, 36.6 ± 7.4 and 80.4 ± 17.4 pA, respectively (p b 0.0001, n = 10 at all concentrations, Fig. 1C). Blind patch-clamped neurons in the LC area were injected with neurobiotin (Fig. 2A). After the analysis of responses,
To confirm whether or not Neurotropin specifically depolarized LC neurons, patch-clamp recordings were made from ventrolateral PAG neurons. None of the recorded neurons showed spontaneous action potentials. Bath-applied Neurotropin did not induce any currents in all neurons tested (n = 10, Fig. 5A, B). Neurotropin also did not alter the frequency or the amplitude of mEPSC and mIPSC (n = 10, Fig. 5C, D). Thus, the excitatory effect of Neurotropin was specific to LC neurons, and not mediated by activation of PAG neurons. 3.3. Effects of icv injection of Neurotropin on nociceptive transmission in the dorsal horn of L5-SNL rats as evaluated by the in vivo patch-clamp method Patch-clamp recordings from SG neurons in spinal cord slice preparations reveal that bath-applied noradrenaline depresses the amplitude of EPSCs evoked by dorsal root stimulation. This effect is mediated by the activation of presynaptic α2 adrenoceptors [15]. It is, therefore, possible that the activation of LC neurons might depress peripheral nociceptive inputs in the spinal dorsal horn, resulting in antinociception. Since this idea could not be tested by in vitro experiments, we made in vivo patch-clamp recordings from SG neurons to clarify whether icv injection of Neurotropin affects nociceptive transmission in the spinal
Fig. 1. Excitatory effect of Neurotropin on normal rat LC neurons. A: In the current-clamp mode, Neurotropin (0.2–1.0 NU/mL) concentration-dependently increased spontaneous firing in rat LC neurons. B: In the voltage-clamp mode at a holding potential of −70 mV, Neurotropin (0.2–1.0 NU/mL) concentration-dependently induced inward currents in rat LC neurons. C: A summary of the quantitative analysis of Neurotropin (0.2–1.0 NU/mL, n = 10)-induced inward currents in rat LC neurons in the voltage-clamp mode. Each column shows the mean ± SEM (n = 10). *p b 0.05 vs Krebs solution (Dunnett's multiple comparison tests).
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Fig. 2. Effect of Neurotropin on noradrenergic neurons in normal rat coronal brain slices including LC area. A: Recorded LC neurons were identified by injecting the recording cells with neurobiotin and staining with Texas Red®-coupled streptoavidin. B: Noradrenergic neurons were identified by staining for tyrosine hydroxylase immunohistochemically. C: Merged image shows that the recorded neuron is noradrenergic. D: Neurotropin (1.0 NU/mL) induced inward current in rat LC neurons that were immunohistochemically identified as noradrenergic neurons. The column shows the means ± SEM (n = 9).
dorsal horn. L5-SNL rats were used as a chronic pain model for in vivo patch-clamp recordings. First, the behavioral changes in L5-SNL rats were assessed by measuring the hind paw withdrawal threshold. All
L5-SNL rats showed hypersensitivities to von Frey filaments compared to those prior to L5 spinal nerve ligation (14.9 ± 0.1 g to 2.6 ± 0.1 g, p b 0.0001, n = 27). In our previous study, we reported that
Fig. 3. Effect of tetrodotoxin (TTX) on Neurotropin-induced inward current and mEPSCs in normal rat LC neurons. A: In the voltage-clamp mode at a holding potential of −70 mV, Neurotropin induced inward currents in rat LC neurons without (left) or with (right) TTX. B: Each column shows Neurotropin-induced inward currents in rat LC neurons with or without TTX. TTX had no effect on Neurotropin-induced inward current (n = 10). C, D: Each column shows the relative frequency (C) and amplitude (D) of mEPSCs after treatment with Neurotropin compared to baseline. TTX had no effect on the frequency or amplitude of Neurotropin-induced mEPSCs in rat LC neurons. Each column shows the mean ± SEM (n = 10).
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Fig. 4. Effect of GDP-β-S on Neurotropin-induced inward current in normal rat LC neurons. A: GDP-β-S was applied to LC neurons through inclusion in the patch pipette solution. Immediately after establishment of the whole-cell configuration, DAMGO induced outward current and Neurotropin induced inward current (upper). After 30 min, GDP-β-S had diffused into LC neurons, blocking the DAMGO-induced outward current compared to just after establishment of the whole-cell configuration whereas Neurotropin-induced inward currents were unaffected (lower). B: Each column shows the relative amplitude of DAMGO- and Neurotropin-induced currents just after and 30 min after establishment of the whole-cell configuration (n = 7). Each column shows the mean ± SEM. *p b 0.05 vs. just after establishment of the whole-cell configuration (paired t-test).
Fig. 5. Effects of Neurotropin on normal rat ventrolateral PAG neurons. A: Neurotropin had no effect on ventrolateral PAG neurons. AMPA was used for confirmation of the condition of recorded neurons. B: Each column shows Krebs solution and Neurotropin-induced current. C: Each column shows the relative frequency and amplitude of the mEPSCs of Krebs solution and Neurotropin compared to baseline. D: Each column shows the relative frequency and amplitude of the mIPSCs of Krebs solution and Neurotropin compared to baseline. Each column shows the mean ± SEM (n = 10). *p b 0.05 vs. Krebs solution (Student's t-test).
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icv injection of Neurotropin (400 mNU/rat) significantly increased hind paw withdrawal thresholds (2.5 ± 0.2 g to 11.9 ± 1.3 g, p = 0.0003, n = 9) in L5-SNL rat [22]. Next, to observe the effects of Neurotropin on nociceptive responses in SG neurons, mechanical pinch stimuli were delivered using a clamp applying 200 g of force to the receptive fields, eliciting a barrage of large amplitude EPSCs in L5SNL rats (Fig. 6A). Under the current-clamp condition, these synaptic inputs were large enough to increase spike firings (not shown). Icv
injection of Neurotropin (300 NU/rat) significantly decreased the relative frequency of eEPSCs to 57.2 ± 11.2% (p = 0.002, n = 9) in the peak at 15 min after icv injection and decreased the relative amplitude of eEPSCs to 59.4 ± 6.0% (p = 0.04, n = 9) in the peak at 30 min after the injection (Fig. 6C). On the other hand, DAMGO (0.5 nmol/rat) significantly decreased the relative frequency to 69.1 ± 7.3% (p = 0.0006, n = 9) and the relative amplitude to 60.5 ± 7.6% (p = 0.005, n = 9) of eEPSCs in the peak at 5 min after icv injection (Fig. 6C).
Fig. 6. Effect of intracerebroventrically administered Neurotropin on pinch-evoked EPSCs in dorsal horn neurons of L5-SNL rats as evaluated by the in vivo patch clamp method. A: Typical trace of pinch (hatched bar)-evoked EPSCs. B: The trace shows control (pretreatment) and 5 min after icv injection of Neurotropin. Neurotropin (right) suppressed the pinch-evoked EPSCs, compared with control (left). C: Pinch stimulation was added to the receptive fields of recorded neurons at pretreatment, and at 5, 15 and 30 min after icv injection of saline, Neurotropin (300 mNU/rat) or DAMGO (0.5 nmol/rat). The relative frequency and relative amplitude of eEPSCs, as compared with those of the control, were reduced by icv injection of Neurotropin and DAMGO. Each column shows the mean ± SEM (n = 9, respectively). *p b 0.05 vs control (Student's t-test).
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4. Discussion In the present study, we examined the mechanisms of the analgesic effect of Neurotropin on normal rat LC and PAG neurons in vitro. In addition, in vivo patch-clamp recordings were also made from SG neurons in L5-SNL chronic pain model rats to further confirm the antinociceptive effect of Neurotropin. In vitro experiments showed that Neurotropin has excitatory effects on LC but not PAG neurons. The membrane depolarization induced by Neurotropin in LC neurons might be mediated by the direct activation of non-selective cationic channels because the depolarization was not affected by TTX or injection of GDP-β-S. In addition, there is supporting evidence that Neurotropin-induced membrane depolarization in NRM neurons has a reversal potential of near 0 mV [21]. It is, therefore, possible that the depolarization in LC is mediated by the direct activation of non-selective cationic channels. The increased firing rate of LC neurons evoked by Neurotropin may cause a further release of noradrenaline in the spinal dorsal horn, inhibiting the release of glutamate from the fine primary afferents [15]. In fact, in vivo patch-clamp recordings showed that the amplitude of EPSCs evoked by pinch stimuli applied to the receptive fields was depressed by icv injection of Neurotropin. Consistent with the present results, previous behavioral studies have also shown that the effect of Neurotropin is in part inhibited by intrathecally administered α2 adrenoceptor antagonists. Furthermore, systemic and icv, but not intrathecal, administration of Neurotropin significantly inhibited mechanical allodynia in L5-SNL rats [22]. Hayashida et al. have reported that microinjection of gabapentin into the LC increases the nociceptive threshold and, using microdialysis and immunohistochemical staining in SNL rats, have demonstrated the analgesic mechanisms of gabapentin [9,27]. Similarly, we have also shown that microinjection of Neurotropin into the LC in SART-stressed rats has an anti-nociceptive effect (unpublished data). Our future studies will examine the effect of microinjection of Neurotropin into the LC in SNL rats. It has been reported that optoactivation of LC neurons produces not only antinociceptive but also pronociceptive changes in thermal nociception, and that these actions are mediated by distinct subpopulations of neurons in the LC [10]. It is, therefore, possible that Neurotropin activates both antinociceptive and pronociceptive LC neurons.
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Since electrical or chemical stimulation of LC neurons induces an antinociceptive effect [13,30], the net effect of activation of most LC neurons might be antinociceptive. In the present study, icv injection of Neurotropin produced antinociception and depressed EPSCs evoked by noxious pinch stimuli at the spinal cord level. Under these conditions, however, it was difficult to test the effect of Neurotropin on different subpopulations of neurons in the LC. Since LC receives inputs from PAG neurons [2], the effect of Neurotropin might be a result of activation of PAG neurons. Neurotropin, however, had no effect on PAG neurons. This finding suggests that activation of monoaminergic pain inhibitory neurons by Neurotropin does not arise from PAG neurons, but from direct activation of LC and/or NRM neurons. Although we could not examine in vivo the effect of α2 adrenoceptor antagonists on antinociception in the spinal cord due to the difficulty in keeping patch recordings from single SG neurons for long enough to test the antagonists on icv injection of Neurotropin, the present slice experiments and previous behavioral studies as well as the effects of noradrenaline on synaptic responses in SG in slice preparations lend support to the notion that the analgesic effect of Neurotropin is mediated by direct activation of LC and/or NRM neurons [15,22]. In contrast, microinjection of opioid into ventrolateral PAG produced analgesia, and this analgesic effect was mediated via the activation of NRM neurons [23]. The spinal cord slice experiments reported previously reveal that noradrenaline inhibits the release of glutamate from primary Aδ and C afferents through the activation of α2 adrenoceptors expressed at the primary afferent terminals [15]. Notwithstanding the presynaptic site of the noradrenaline action, the frequency of EPSCs was also depressed. One possible explanation for this may be that the released noradrenaline not only inhibits the release of glutamate but also hyperpolarizes the membrane at bifurcations of primary afferent terminals, resulting in cessation of propagating action potentials to the releasing sites [19]. Alternatively, since some of the pinch-evoked EPSCs are polysynaptic, their polysynaptic circuits may be blocked by the noradrenaline which is released as a result. It is yet to be confirmed if the pinch-evoked barrage of EPSCs is mono- or poly-synaptic, or a mixture of both. As mentioned in the Introduction, Neurotropin is a non-protein extract from the inflamed skin of rabbits inoculated with vaccinia
Fig. 7. Schematic representation of activation of the descending pain inhibitory system by Neurotropin. Neurotropin directly activates LC and NRM neurons, but not PAG neurons. Therefore, noradrenaline and serotonin may be released to the synapse on dorsal horn neurons that transmit the pain signal from peripheral receptive fields. Noradrenaline and serotonin may inhibit transmission of the pain signal pre- and post-synaptically in the spinal dorsal horn.
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virus. Therefore, it is not known if this depolarizing effect of Neurotropin is mediated by unknown receptor(s) or non-specifically activates nonselective ionic channels. The fact that Neurotropin depolarized LC and NRM but not PAG neurons, and the fact that the effect was dosedependent, supports the notion that Neurotropin interacts with specific receptor(s) in LC and NRM neurons. Identification of the receptor(s), though, is beyond the scope of the present study. On the other hand, Sonohata et al. [26] reported that perfusion of noradrenaline onto the surface of the spinal cord not only reduces pinch-evoked EPSCs but also produces outward currents. These findings, however, are inconsistent with the results in our current study in which no outward current was observed, suggesting that the noradrenaline which was released from the descending pain inhibitory system reached the presynaptic sites but not the postsynaptic membrane of SG neurons. In conclusion, electrophysiological investigations using in vitro and in vivo patch-clamp recordings clarified that Neurotropin directly activates LC but not PAG neurons, resulting in a reduction in pain transmission at the level of the spinal dorsal horn (Fig. 7). 5. Conclusions Our finding revealed that Neurotropin directly excites the descending noradrenergic LC neurons and inhibits nociceptive transmission in the spinal dorsal horn. This study is the first direct demonstration that Neurotropin activates the noradrenergic descending pain inhibitory systems, and this would reinforce the usefulness of Neurotropin in the treatment of human neuropathic pain. Conflict of interest statement Three of the authors are employed by the Nippon Zoki Pharmaceutical Co., Ltd. in Japan.
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Excitatory effect of Neurotropin® on noradrenergic neurons in rat locus coeruleus Hisashi Okai a,⁎, Ryohei Okazaki a, Minoru Kawamura a, Megumu Yoshimura b a b
Department of Pharmacological Research, Institute of Bio-Active Science, Nippon Zoki Pharmaceutical Co., Ltd., 442-1, Kinashi, Kato, Hyogo 673-1461, Japan Graduate School of Health Sciences, Kumamoto Health Science University, Kumamoto 861-5598, Japan
a r t i c l e
i n f o
Article history: Received 19 December 2014 Received in revised form 14 May 2015 Accepted 4 June 2015 Available online 2 July 2015 Keywords: Neurotropin Neuropathic pain Locus coeruleus Descending pain inhibitory system Noradrenaline In vivo patch clamp
a b s t r a c t Aims: Although the clinical use of Neurotropin® as an analgesic for chronic pain has been firmly established, its analgesic mechanism is still unclear. In this study, we investigate the direct effects of Neurotropin using an electrophysiological method. Main methods: Blind patch-clamp recordings were made from rat locus coeruleus (LC) and periaqueductal gray (PAG) neurons in brainstem slices of normal rats. The effects of intracerebroventricular (icv) injection of Neurotropin on nociceptive transmission were recorded from spinal substantia gelatinosa (SG) neurons in fifth lumbar spinal nerve-ligated (L5-SNL) rats using an in vivo patch-clamp method. Key findings: Neurotropin (0.2–1.0 NU/mL) dose-dependently increased the firing rate in noradrenergic LC neurons of normal rats. Under the voltage-clamp condition, Neurotropin induced an inward current in 90% of LC neurons that was not affected by tetrodotoxin or an injection of GDP-β-S (G protein inhibitor) through recording pipettes. In contrast, Neurotropin had no effects on all PAG neurons tested. Using in vivo patch-clamp recordings, the icv injection of Neurotropin inhibited both frequency and amplitude of pinch-evoked excitatory postsynaptic currents of SG neurons in L5-SNL rats. These results suggest that Neurotropin directly excites the descending noradrenergic LC neurons and inhibits nociceptive transmission in the spinal dorsal horn. Significance: This study is the first direct demonstration that Neurotropin activates the noradrenergic descending pain inhibitory systems, and this would reinforce the usefulness of Neurotropin in the treatment of human neuropathic pain. © 2015 Elsevier Inc. All rights reserved.
1. Introduction Neurotropin, a non-protein extract from the inflamed skin of rabbits inoculated with vaccinia virus, is widely used in Japan and China as an analgesic for chronic pain such as postherpetic neuralgia and other neuropathic pain. A double-blind placebo-controlled cross-over study is currently underway at the National Institute of Nursing Research in the United States to investigate the effectiveness of Neurotropin in the treatment of fibromyalgia. Neurotropin is recommended as a secondline treatment in the guideline for managing neuropathic pain in Japan [29]. Analgesic mechanisms of Neurotropin have been examined mainly in a chronic stress model — namely, SART (Specific Alternation of Rhythm in Environmental Temperature)-stressed animals [7,8,12,20], which exhibit a chronic hyperalgesic state due to repeated periods of cold stress. The analgesic effect of systemic administration of Neurotropin on SART-stressed rats is antagonized by inhibition of spinal α2 adrenoceptors or serotonergic 5-HT3 receptors, but not opioid μ
⁎ Corresponding author. E-mail address: [email protected] (H. Okai).
http://dx.doi.org/10.1016/j.lfs.2015.06.013 0024-3205/© 2015 Elsevier Inc. All rights reserved.
receptors [14]. These results suggest that Neurotropin activates the monoaminergic descending pain inhibitory systems, but not opioidergic neurons in SART-stressed rats. Neurotropin ameliorates chronic inflammation-induced hyperalgesia and L5 spinal nerve ligation (L5-SNL)-induced mechanical allodynia, and these analgesic effects are also depressed by intrathecally administered α2 adrenoceptor antagonist or 5-HT3 receptor antagonist [17,22,28]. Furthermore, the anti-allodynic effect of Neurotropin is more marked following intracerebroventricular (icv) administration compared to intrathecal administration [22]. Therefore, it may be presumed that Neurotropin activates the supraspinal site of monoaminergic descending pain inhibitory systems. The midbrain periaqueductal gray (PAG) projects to the locus coeruleus (LC) and rostral ventromedial medulla (RVM) [2]. Therefore, the analgesic effect produced by stimulation of PAG is mediated via the activation of LC and/or RVM. It is well known that the nucleus raphe magnus (NRM) [5], which is the main nucleus in the RVM, contains 5-HT and other inhibitory transmitters such as GABA and glycine [1,11,25]. Based on behavioral studies [14], the antinociceptive effects of Neurotropin might be a result of activation of the supraspinal LC, PAG and/or NRM. Electrophysiologically, however, it is still unclear whether Neurotropin acts on one, or all three, brain areas.
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Previously, we demonstrated that Neurotropin directly excites NRM neurons in brainstem slices of normal rats [21]. Our present study, therefore, examined the direct effects of Neurotropin on PAG and LC neurons. Since it has previously been shown that systemic administration of Neurotropin causes a dose-dependent inhibition of hyperalgesia in the adjuvant-induced arthritic rat but has no significant effects on the nociceptive threshold in normal rats [17], L5-SNL chronic pain model rats were used for in vivo examinations. In vivo patch-clamp recordings from spinal SG neurons were performed to examine the effects of icv administration of Neurotropin on excitatory inputs from the receptive fields. 2. Materials and methods 2.1. Drugs Neurotropin was prepared by the Nippon Zoki Pharmaceutical Co., Ltd. (Osaka, Japan). [D-Ala2, N-MePhe4, Gly-ol]-enkephalin (DAMGO) was purchased from Sigma-Aldrich (St. Louis, MO). 2-Amino-3-(3hydroxy-5-methyl-isoxazol-4-yl) propanoic acid (AMPA) was purchased from Funakoshi Co., Ltd. (Tokyo, Japan). Tetrodotoxin (TTX), a sodium channel blocker, was purchased from Wako Pure Chemical Industries, Ltd. (Osaka, Japan). These drugs were dissolved or diluted in Krebs solution before use. Guanosine-5′-O-(2-thiodiphosphate) (GDP-β-S, G protein inhibitors) was purchased from Sigma-Aldrich (St. Louis, MO).
that was equilibrated with 95% O2 and 5% CO2 at 37 ± 1 °C. Drugs and Krebs solution were applied to the slices by perfusion via a three-way stopcock without any changes in the perfusion rate or temperature. 2.5. Patch-clamp recordings Blind whole-cell patch-clamp recordings were made from normal rat LC, ventrolateral PAG, and SG neurons in the spinal cord with patch electrodes (tip resistances: 6–12 MΩ). The patch pipettes were filled with a solution containing (mM): K-gluconate 135, KCl 5, CaCl2 0.5, MgCl2 2, EGTA 5, HEPES 5, ATP-Mg 5 and pH 7.2 for excitatory postsynaptic currents (EPSCs) recordings, or containing (mM): cesium sulfate 110, CaCl2 0.5, MgCl2 2, EGTA 5, HEPES 5, ATP-Mg 5 and pH 7.2 for inhibitory postsynaptic currents (IPSCs) recordings. GDP-β-S was added to the solutions in patch pipettes at 2 mM. Signals were acquired with a patch-clamp amplifier (Axopatch 200B amplifier, Molecular Devices, CA, USA). Data were digitized with an A/D converter (digidata1400, Molecular Devices), stored on a personal computer using a data acquisition program (pClamp10.2, Molecular Devices, CA, USA), and analyzed with Clampfit software (clampfit10.2, Molecular Devices, CA, USA) and AxoGraph X (AxoGraph Scientific, CA, USA). In the voltage-clamp mode, the holding potentials were held at −70 mV (at which glycine-/GABA-mediated IPSCs were negligible) or 0 mV (at which glutamate-mediated EPSCs were negligible). Frequencies of action potentials were monitored in the zero current-clamp mode. 2.6. In vivo patch-clamp recordings
2.2. Animals Six-week-old male SD rats were purchased from Kyudo Co., Ltd. (Saga, Japan). The animals were housed under a 12-h light–dark cycle with free access to food and water. At the time of the experiment, the animals ranged in age from 7 to 11 weeks. All experimental procedures were approved by the Ethics Committee on Animal Experiments at Kyushu University (Fukuoka, Japan), or an ethical committee on animal experiments at the Nippon Zoki Pharmaceutical Co., Ltd. (Kato, Japan), and performed in accordance with the ethical guidelines for investigations of experimental pain in conscious animals issued by the International Association for the Study of Pain [31]. 2.3. Surgical preparation and behavioral testing
In vivo patch-clamp recordings were performed according to the method of Furue et al. [6]. Briefly, after rats were deeply anesthetized with urethane (1.0–1.2 g/kg, i.p.), the lumbar enlargement was exposed by a thoraco-lumbar laminectomy at the T12 to L2 levels. Then, rats were placed in a stereotaxic apparatus (Model ST-7, Narishige, Tokyo, Japan). After the dura mater was cut and reflected, a dorsal root that entered the spinal dorsal horn covering the recording region was gently moved aside with a glass retractor to expose the lateral edge of the SG. The pia-arachnoid membrane was then cut with fine scissors to make a small window to allow a patch electrode to enter the SG. The surface of the exposed spinal cord was irrigated with Krebs solution equilibrated with 95% O2–5% CO2 at 37 ± 1 °C. The noxious pinch stimuli were applied to the ipsilateral receptive field of the recorded neuron using a clamp applying 200 g of force. The clamp had previously been calibrated using a pull tension gauge.
L5-SNL model rats were prepared according to the method of Kim and Chung [16]. Briefly, the right L5 spinal nerve was isolated and tightly ligated with a 5–0 silk suture under pentobarbital (40 mg/kg, i.p.) anesthesia. After surgery, rats were housed in a cage in groups of three with free access to food and water and allowed to recover over 14 days. Right paw mechanical allodynia was confirmed before in vivo patch-clamp recording by measuring the hindpaw withdrawal threshold in response to application of von Frey filaments using the up–down method [3,4]. Only those rats which showed a withdrawal threshold of less than 4 g were used for in vivo patch-clamp recording. Behavioral tests were performed before the patch-clamp recordings at 14–28 days after nerve ligation.
For icv injection, the shaved scalp was incised to expose the skull, and a hole was drilled into the skull above the right lateral ventricle (distance from interaural: anterior, +8.2 mm; lateral, +1.3 mm; vertical, +6.0 mm) according to the stereotaxic coordinates of the atlas of Paxinos and Watson [24]. Then, using a 30-gauge needle tip inserted into the right lateral ventricle and a CMA/100 microinjection pump (CMA Microdialysis AB, Stockholm, Sweden), 15 μL of each drug solution was applied over 1 min.
2.4. Brain slice preparations
2.8. Immunohistochemistry
Rat brain slice preparations were obtained according to the method of Nakatsuka et al. [18]. Rats were deeply anesthetized with urethane (1.2 g/kg, i.p.), and then decapitated without delay. The whole brain was removed and placed in an ice-cold, pre-oxygenated Krebs solution (in mM: NaCl 117, KCl 3.6, CaCl2 2.5, MgCl2 1.2, NaH2PO4 1.2, NaHCO3 25, and D-glucose 11). After removal of the dura mater, the brainstem, including LC or PAG, was coronally sliced at 400 μm thickness using a microslicer. The slices were then placed in a recording chamber and perfused continuously at a rate of 10–15 mL/min with Krebs solution
Recorded LC neurons were labeled with neurobiotin (0.2% in the electrode solution; Vector Laboratories, CA, USA) diffused from the patch pipettes after whole-cell configuration. After the electrophysiological recordings, brain slices were immersed in 4% paraformaldehyde/0.1 mM phosphate-buffered saline (PBS) overnight, and embedded in paraffin. Slices were sectioned coronally (8 μm) using a microtome (Daiwa Koki, K.K., Saitama, Japan), and incubated with anti-tyrosine hydroxylase antibody (1:200 dilution in PBS; Enzo Life Sciences, Inc., NY, USA) overnight at 4 °C. After rinsing in PBS, the sections
2.7. Icv injection of Neurotropin
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were incubated with FITC-conjugated AffiniPure Donkey Anti-rabbit IgG (dilution 1:100; FITC-IgG; Jackson Immuno Research Laboratories, Inc., West Grove, PA) and Texas Red® dye-conjugated streptavidin (Jackson Immuno Research Laboratories, Inc., West Grove, PA) for 6 h. The sections were mounted on gelatinized slides and photographed with a fluorescence microscope (TE2000, Nikon, Tokyo, Japan). 2.9. Statistical analysis All numerical data were provided as means ± standard error of the mean (SEM). The statistical significance between two groups was determined by Student's t-test or the paired t-test. Statistical significance between more than two groups was analyzed using Dunnett's multiple comparison tests. p values less than 0.05 were considered to be significant.
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neurons were stained with tyrosine hydroxylase (Fig. 2B). All recorded neurons injected with neurobiotin (n = 10) were positive to tyrosine hydroxylase and, therefore, noradrenergic (Fig. 2C), and nine out of ten neurons were responsive to 1 NU/mL of Neurotropin (Fig. 2D). The Neurotropin-induced inward current was not affected by pretreatment with TTX (1 μM, Fig. 3A, B). The frequency and amplitude of miniature EPSCs (mEPSCs) were also not affected by the application of Neurotropin (Fig. 3C, D). Immediately after establishment of the whole-cell recording configuration with pipettes containing GDP-β-S, DAMGO (1 μM) induced an outward current and Neurotropin induced an inward current (Fig. 4A). After 30 min, GDP-β-S had diffused into LC neurons, blocking the DAMGO-induced outward currents (4.3 ± 2.8% compared to the current just after establishment of the wholecell configuration, n = 7) whereas Neurotropin-induced inward currents were unaffected (Fig. 4B, C). 3.2. Effects of Neurotropin on ventrolateral PAG neurons
3. Results 3.1. Effects of Neurotropin on LC neurons In the current-clamp mode, spontaneous action potentials were observed in all LC neurons tested in vitro. Bath-applied Neurotropin concentration-dependently increased the frequency of action potentials. At concentrations of 0.2, 0.5 and 1.0 NU/mL of Neurotropin, the frequency of action potentials was increased to 165.2 ± 32.8% (n = 6), 380.0 ± 15.2% (n = 6), and 771.2 ± 273.5% (n = 6), respectively (Fig. 1A). Conversely, after Neurotropin was washed off the samples, the frequency of action potentials decreased temporarily, canceling the previous increase in frequency of action potentials for a few seconds. This might have been due to a hyperpolarizing response by Neurotropin following depolarization but further analysis of this was beyond the scope of the present study. In the voltage-clamp mode at a holding potential of −70 mV, Neurotropin (0.2–1.0 NU/mL) induced an inward current in a clear concentration-dependent manner (Fig. 1B). At 0.2, 0.5 and 1.0 NU/mL, Neurotropin induced inward currents of 13.6 ± 2.1, 36.6 ± 7.4 and 80.4 ± 17.4 pA, respectively (p b 0.0001, n = 10 at all concentrations, Fig. 1C). Blind patch-clamped neurons in the LC area were injected with neurobiotin (Fig. 2A). After the analysis of responses,
To confirm whether or not Neurotropin specifically depolarized LC neurons, patch-clamp recordings were made from ventrolateral PAG neurons. None of the recorded neurons showed spontaneous action potentials. Bath-applied Neurotropin did not induce any currents in all neurons tested (n = 10, Fig. 5A, B). Neurotropin also did not alter the frequency or the amplitude of mEPSC and mIPSC (n = 10, Fig. 5C, D). Thus, the excitatory effect of Neurotropin was specific to LC neurons, and not mediated by activation of PAG neurons. 3.3. Effects of icv injection of Neurotropin on nociceptive transmission in the dorsal horn of L5-SNL rats as evaluated by the in vivo patch-clamp method Patch-clamp recordings from SG neurons in spinal cord slice preparations reveal that bath-applied noradrenaline depresses the amplitude of EPSCs evoked by dorsal root stimulation. This effect is mediated by the activation of presynaptic α2 adrenoceptors [15]. It is, therefore, possible that the activation of LC neurons might depress peripheral nociceptive inputs in the spinal dorsal horn, resulting in antinociception. Since this idea could not be tested by in vitro experiments, we made in vivo patch-clamp recordings from SG neurons to clarify whether icv injection of Neurotropin affects nociceptive transmission in the spinal
Fig. 1. Excitatory effect of Neurotropin on normal rat LC neurons. A: In the current-clamp mode, Neurotropin (0.2–1.0 NU/mL) concentration-dependently increased spontaneous firing in rat LC neurons. B: In the voltage-clamp mode at a holding potential of −70 mV, Neurotropin (0.2–1.0 NU/mL) concentration-dependently induced inward currents in rat LC neurons. C: A summary of the quantitative analysis of Neurotropin (0.2–1.0 NU/mL, n = 10)-induced inward currents in rat LC neurons in the voltage-clamp mode. Each column shows the mean ± SEM (n = 10). *p b 0.05 vs Krebs solution (Dunnett's multiple comparison tests).
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Fig. 2. Effect of Neurotropin on noradrenergic neurons in normal rat coronal brain slices including LC area. A: Recorded LC neurons were identified by injecting the recording cells with neurobiotin and staining with Texas Red®-coupled streptoavidin. B: Noradrenergic neurons were identified by staining for tyrosine hydroxylase immunohistochemically. C: Merged image shows that the recorded neuron is noradrenergic. D: Neurotropin (1.0 NU/mL) induced inward current in rat LC neurons that were immunohistochemically identified as noradrenergic neurons. The column shows the means ± SEM (n = 9).
dorsal horn. L5-SNL rats were used as a chronic pain model for in vivo patch-clamp recordings. First, the behavioral changes in L5-SNL rats were assessed by measuring the hind paw withdrawal threshold. All
L5-SNL rats showed hypersensitivities to von Frey filaments compared to those prior to L5 spinal nerve ligation (14.9 ± 0.1 g to 2.6 ± 0.1 g, p b 0.0001, n = 27). In our previous study, we reported that
Fig. 3. Effect of tetrodotoxin (TTX) on Neurotropin-induced inward current and mEPSCs in normal rat LC neurons. A: In the voltage-clamp mode at a holding potential of −70 mV, Neurotropin induced inward currents in rat LC neurons without (left) or with (right) TTX. B: Each column shows Neurotropin-induced inward currents in rat LC neurons with or without TTX. TTX had no effect on Neurotropin-induced inward current (n = 10). C, D: Each column shows the relative frequency (C) and amplitude (D) of mEPSCs after treatment with Neurotropin compared to baseline. TTX had no effect on the frequency or amplitude of Neurotropin-induced mEPSCs in rat LC neurons. Each column shows the mean ± SEM (n = 10).
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Fig. 4. Effect of GDP-β-S on Neurotropin-induced inward current in normal rat LC neurons. A: GDP-β-S was applied to LC neurons through inclusion in the patch pipette solution. Immediately after establishment of the whole-cell configuration, DAMGO induced outward current and Neurotropin induced inward current (upper). After 30 min, GDP-β-S had diffused into LC neurons, blocking the DAMGO-induced outward current compared to just after establishment of the whole-cell configuration whereas Neurotropin-induced inward currents were unaffected (lower). B: Each column shows the relative amplitude of DAMGO- and Neurotropin-induced currents just after and 30 min after establishment of the whole-cell configuration (n = 7). Each column shows the mean ± SEM. *p b 0.05 vs. just after establishment of the whole-cell configuration (paired t-test).
Fig. 5. Effects of Neurotropin on normal rat ventrolateral PAG neurons. A: Neurotropin had no effect on ventrolateral PAG neurons. AMPA was used for confirmation of the condition of recorded neurons. B: Each column shows Krebs solution and Neurotropin-induced current. C: Each column shows the relative frequency and amplitude of the mEPSCs of Krebs solution and Neurotropin compared to baseline. D: Each column shows the relative frequency and amplitude of the mIPSCs of Krebs solution and Neurotropin compared to baseline. Each column shows the mean ± SEM (n = 10). *p b 0.05 vs. Krebs solution (Student's t-test).
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icv injection of Neurotropin (400 mNU/rat) significantly increased hind paw withdrawal thresholds (2.5 ± 0.2 g to 11.9 ± 1.3 g, p = 0.0003, n = 9) in L5-SNL rat [22]. Next, to observe the effects of Neurotropin on nociceptive responses in SG neurons, mechanical pinch stimuli were delivered using a clamp applying 200 g of force to the receptive fields, eliciting a barrage of large amplitude EPSCs in L5SNL rats (Fig. 6A). Under the current-clamp condition, these synaptic inputs were large enough to increase spike firings (not shown). Icv
injection of Neurotropin (300 NU/rat) significantly decreased the relative frequency of eEPSCs to 57.2 ± 11.2% (p = 0.002, n = 9) in the peak at 15 min after icv injection and decreased the relative amplitude of eEPSCs to 59.4 ± 6.0% (p = 0.04, n = 9) in the peak at 30 min after the injection (Fig. 6C). On the other hand, DAMGO (0.5 nmol/rat) significantly decreased the relative frequency to 69.1 ± 7.3% (p = 0.0006, n = 9) and the relative amplitude to 60.5 ± 7.6% (p = 0.005, n = 9) of eEPSCs in the peak at 5 min after icv injection (Fig. 6C).
Fig. 6. Effect of intracerebroventrically administered Neurotropin on pinch-evoked EPSCs in dorsal horn neurons of L5-SNL rats as evaluated by the in vivo patch clamp method. A: Typical trace of pinch (hatched bar)-evoked EPSCs. B: The trace shows control (pretreatment) and 5 min after icv injection of Neurotropin. Neurotropin (right) suppressed the pinch-evoked EPSCs, compared with control (left). C: Pinch stimulation was added to the receptive fields of recorded neurons at pretreatment, and at 5, 15 and 30 min after icv injection of saline, Neurotropin (300 mNU/rat) or DAMGO (0.5 nmol/rat). The relative frequency and relative amplitude of eEPSCs, as compared with those of the control, were reduced by icv injection of Neurotropin and DAMGO. Each column shows the mean ± SEM (n = 9, respectively). *p b 0.05 vs control (Student's t-test).
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4. Discussion In the present study, we examined the mechanisms of the analgesic effect of Neurotropin on normal rat LC and PAG neurons in vitro. In addition, in vivo patch-clamp recordings were also made from SG neurons in L5-SNL chronic pain model rats to further confirm the antinociceptive effect of Neurotropin. In vitro experiments showed that Neurotropin has excitatory effects on LC but not PAG neurons. The membrane depolarization induced by Neurotropin in LC neurons might be mediated by the direct activation of non-selective cationic channels because the depolarization was not affected by TTX or injection of GDP-β-S. In addition, there is supporting evidence that Neurotropin-induced membrane depolarization in NRM neurons has a reversal potential of near 0 mV [21]. It is, therefore, possible that the depolarization in LC is mediated by the direct activation of non-selective cationic channels. The increased firing rate of LC neurons evoked by Neurotropin may cause a further release of noradrenaline in the spinal dorsal horn, inhibiting the release of glutamate from the fine primary afferents [15]. In fact, in vivo patch-clamp recordings showed that the amplitude of EPSCs evoked by pinch stimuli applied to the receptive fields was depressed by icv injection of Neurotropin. Consistent with the present results, previous behavioral studies have also shown that the effect of Neurotropin is in part inhibited by intrathecally administered α2 adrenoceptor antagonists. Furthermore, systemic and icv, but not intrathecal, administration of Neurotropin significantly inhibited mechanical allodynia in L5-SNL rats [22]. Hayashida et al. have reported that microinjection of gabapentin into the LC increases the nociceptive threshold and, using microdialysis and immunohistochemical staining in SNL rats, have demonstrated the analgesic mechanisms of gabapentin [9,27]. Similarly, we have also shown that microinjection of Neurotropin into the LC in SART-stressed rats has an anti-nociceptive effect (unpublished data). Our future studies will examine the effect of microinjection of Neurotropin into the LC in SNL rats. It has been reported that optoactivation of LC neurons produces not only antinociceptive but also pronociceptive changes in thermal nociception, and that these actions are mediated by distinct subpopulations of neurons in the LC [10]. It is, therefore, possible that Neurotropin activates both antinociceptive and pronociceptive LC neurons.
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Since electrical or chemical stimulation of LC neurons induces an antinociceptive effect [13,30], the net effect of activation of most LC neurons might be antinociceptive. In the present study, icv injection of Neurotropin produced antinociception and depressed EPSCs evoked by noxious pinch stimuli at the spinal cord level. Under these conditions, however, it was difficult to test the effect of Neurotropin on different subpopulations of neurons in the LC. Since LC receives inputs from PAG neurons [2], the effect of Neurotropin might be a result of activation of PAG neurons. Neurotropin, however, had no effect on PAG neurons. This finding suggests that activation of monoaminergic pain inhibitory neurons by Neurotropin does not arise from PAG neurons, but from direct activation of LC and/or NRM neurons. Although we could not examine in vivo the effect of α2 adrenoceptor antagonists on antinociception in the spinal cord due to the difficulty in keeping patch recordings from single SG neurons for long enough to test the antagonists on icv injection of Neurotropin, the present slice experiments and previous behavioral studies as well as the effects of noradrenaline on synaptic responses in SG in slice preparations lend support to the notion that the analgesic effect of Neurotropin is mediated by direct activation of LC and/or NRM neurons [15,22]. In contrast, microinjection of opioid into ventrolateral PAG produced analgesia, and this analgesic effect was mediated via the activation of NRM neurons [23]. The spinal cord slice experiments reported previously reveal that noradrenaline inhibits the release of glutamate from primary Aδ and C afferents through the activation of α2 adrenoceptors expressed at the primary afferent terminals [15]. Notwithstanding the presynaptic site of the noradrenaline action, the frequency of EPSCs was also depressed. One possible explanation for this may be that the released noradrenaline not only inhibits the release of glutamate but also hyperpolarizes the membrane at bifurcations of primary afferent terminals, resulting in cessation of propagating action potentials to the releasing sites [19]. Alternatively, since some of the pinch-evoked EPSCs are polysynaptic, their polysynaptic circuits may be blocked by the noradrenaline which is released as a result. It is yet to be confirmed if the pinch-evoked barrage of EPSCs is mono- or poly-synaptic, or a mixture of both. As mentioned in the Introduction, Neurotropin is a non-protein extract from the inflamed skin of rabbits inoculated with vaccinia
Fig. 7. Schematic representation of activation of the descending pain inhibitory system by Neurotropin. Neurotropin directly activates LC and NRM neurons, but not PAG neurons. Therefore, noradrenaline and serotonin may be released to the synapse on dorsal horn neurons that transmit the pain signal from peripheral receptive fields. Noradrenaline and serotonin may inhibit transmission of the pain signal pre- and post-synaptically in the spinal dorsal horn.
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virus. Therefore, it is not known if this depolarizing effect of Neurotropin is mediated by unknown receptor(s) or non-specifically activates nonselective ionic channels. The fact that Neurotropin depolarized LC and NRM but not PAG neurons, and the fact that the effect was dosedependent, supports the notion that Neurotropin interacts with specific receptor(s) in LC and NRM neurons. Identification of the receptor(s), though, is beyond the scope of the present study. On the other hand, Sonohata et al. [26] reported that perfusion of noradrenaline onto the surface of the spinal cord not only reduces pinch-evoked EPSCs but also produces outward currents. These findings, however, are inconsistent with the results in our current study in which no outward current was observed, suggesting that the noradrenaline which was released from the descending pain inhibitory system reached the presynaptic sites but not the postsynaptic membrane of SG neurons. In conclusion, electrophysiological investigations using in vitro and in vivo patch-clamp recordings clarified that Neurotropin directly activates LC but not PAG neurons, resulting in a reduction in pain transmission at the level of the spinal dorsal horn (Fig. 7). 5. Conclusions Our finding revealed that Neurotropin directly excites the descending noradrenergic LC neurons and inhibits nociceptive transmission in the spinal dorsal horn. This study is the first direct demonstration that Neurotropin activates the noradrenergic descending pain inhibitory systems, and this would reinforce the usefulness of Neurotropin in the treatment of human neuropathic pain. Conflict of interest statement Three of the authors are employed by the Nippon Zoki Pharmaceutical Co., Ltd. in Japan.
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