Next Article in Journal
Metal-Based Complexes as Pharmaceuticals for Molecular Imaging of the Liver
Previous Article in Journal
Predictive Power of In Silico Approach to Evaluate Chemicals against M. tuberculosis: A Systematic Review

Pharmaceuticals 2019, 12(3), 136; https://doi.org/10.3390/ph12030136

Review
Cellular Mechanisms for Antinociception Produced by Oxytocin and Orexins in the Rat Spinal Lamina II—Comparison with Those of Other Endogenous Pain Modulators
Department of Physiology, Saga Medical School, 5-1-1 Nabeshima, Saga 849-8501, Japan
Received: 17 May 2019 / Accepted: 12 September 2019 / Published: 16 September 2019

Abstract

:
Much evidence indicates that hypothalamus-derived neuropeptides, oxytocin, orexins A and B, inhibit nociceptive transmission in the rat spinal dorsal horn. In order to unveil cellular mechanisms for this antinociception, the effects of the neuropeptides on synaptic transmission were examined in spinal lamina II neurons that play a crucial role in antinociception produced by various analgesics by using the whole-cell patch-clamp technique and adult rat spinal cord slices. Oxytocin had no effect on glutamatergic excitatory transmission while producing a membrane depolarization, γ-aminobutyric acid (GABA)-ergic and glycinergic spontaneous inhibitory transmission enhancement. On the other hand, orexins A and B produced a membrane depolarization and/or a presynaptic spontaneous excitatory transmission enhancement. Like oxytocin, orexin A enhanced both GABAergic and glycinergic transmission, whereas orexin B facilitated glycinergic but not GABAergic transmission. These inhibitory transmission enhancements were due to action potential production. Oxytocin, orexins A and B activities were mediated by oxytocin, orexin-1 and orexin-2 receptors, respectively. This review article will mention cellular mechanisms for antinociception produced by oxytocin, orexins A and B, and discuss similarity and difference in antinociceptive mechanisms among the hypothalamic neuropeptides and other endogenous pain modulators (opioids, nociceptin, adenosine, adenosine 5’-triphosphate (ATP), noradrenaline, serotonin, dopamine, somatostatin, cannabinoids, galanin, substance P, bradykinin, neuropeptide Y and acetylcholine) exhibiting a change in membrane potential, excitatory or inhibitory transmission in the spinal lamina II neurons.
Keywords:
oxytocin; orexin A; orexin B; spinal dorsal horn; synaptic transmission; depolarization; antinociception; patch clamp; rat

1. Introduction

Somatosensory information from the periphery to spinal cord is transmitted through the dorsal root to the dorsal horn whose gray matter is divided into six laminae named I–VI by Rexed [1]. Nociception among this information is transferred by fine myelinated Aδ and unmyelinated C primary-afferent fibers contained in the dorsal root to neurons in the superficial laminae of the dorsal horn, particularly lamina II (substantia gelatinosa), in a mono- and polysynaptic manner ([2,3]; see [4] for review). This nociceptive information flows to the thalamus through a connection with projection neurons in lamina I and deeper laminae of the spinal dorsal horn (see [5] for review), and then to the primary sensory area of the cerebral cortex, producing nociceptive sensation. Since the proposal of the gate-control theory of pain by Melzack and Wall [6], a modulation of synaptic transmission in spinal lamina II neurons is thought to play a crucial role in the regulation of nociceptive sensation (see [4,7] for review). This synaptic modulation mainly occurs at pre- and/or postsynaptic sites of excitatory and inhibitory synapses in spinal lamina II neurons through actions of a number of endogenous substances that are either locally produced/released in the spinal dorsal horn or released from descending neurons originating from higher centers such as the brainstem and hypothalamus.
Consistent with the importance of spinal lamina II neurons, a plastic change in glutamatergic primary-afferent inputs to the neurons occurs in hyperalgesia caused by an intraplantar injection of complete Freund’s adjuvant [8] or ovariectomy [9] in rats. In partial nerve injury rat models compared with sham rats, primary-afferent-evoked inhibitory transmission is inhibited in spinal lamina II neurons; γ-aminobutyric acid (GABA)-synthesizing enzyme expression is reduced in level in the spinal dorsal horn [10]. There is a difference between wild-type and Glra3-/- mice in mechanical and thermal behavior of pain model produced by complete Freund’s adjuvant injection, indicating an involvement of glycine-receptor α3 subunit (see [11] for review) in inflammatory pain [12]. A decrease in expression of the potassium-chloride exporter KCC2 shifts transmembrane chloride gradient and thus causes normally inhibitory anionic synaptic currents to be excitatory in rat spinal dorsal horn neurons; as a result, nociceptive thresholds are markedly reduced ([13]; see [14] for review). Recently, Medrano et al. [15] have suggested that a shift in the reversal potential for chloride is an important component of a loss of inhibitory tone in neuropathic pain mouse models produced by nerve injury.
Many of the targets on which the endogenous analgesics act are membrane proteins including receptors for neurotransmitters. There are many investigations about the effects of the analgesics on synaptic transmission in lamina II neurons by using the whole-cell patch-clamp technique in spinal cord slice preparations dissected from 2- to 8-week-old rodents. Lamina II layer is easily discernable as a translucent band in slices under a binocular microscope with light transmitted from below ([16,17]; see [18,19] for review). Recently, hypothalamus-derived neuropeptides, oxytocin, orexins A and B, that have an ability to alleviate pain when intrathecally administrated, were shown to exhibit a similar synaptic modulation, i.e., inhibitory transmission facilitation following excitatory transmission facilitation, in adult rat spinal lamina II neurons [20,21,22]. Their modulatory actions were either similar to or different from those of the other endogenous pain modulators (opioids, nociceptin, adenosine, adenosine 5’-triphosphate (ATP), noradrenaline, serotonin, dopamine, somatostatin, cannabinoids, galanin, substance P, bradykinin, neuropeptide Y and acetylcholine) in the spinal cord level. These modulators produce a membrane depolarization or hyperpolarization, a facilitation or depression of excitatory transmission, or a facilitation or depression of inhibitory transmission. This review article will mention cellular mechanisms for synaptic modulation produced by oxytocin, orexins A and B and also other endogenous pain modulators’ ones that are related to the present topics, and then discuss a similarity and difference in antinociceptive mechanisms among these modulators.

2. Fast Synaptic Transmission in Spinal Lamina II Neurons

As in the brain, excitatory transmission in the lamina II is mediated by L-glutamate released from nerve terminals; this amino acid originates from primary-afferent fibers and glutamatergic interneurons in the lamina II [17]. On the other hand, inhibitory transmission in lamina II neurons is mediated not only by GABA as in the brain but also by glycine; these amino acids are released from the terminals of GABAergic and/or glycinergic neurons that exist in the spinal dorsal horn or that originate from supraspinal regions such as the rostral ventromedial medulla [23,24,25].
Although fast excitatory transmission in the central nervous system is generally mediated by ionotropic α-amino-3-hydroxy-5-methyl-4-isoxazole propionate (AMPA) and N-methyl-D-aspartate (NMDA) receptor-channels (see [11] for review), spontaneous and electrically-evoked excitatory postsynaptic currents (EPSCs) recorded at –70 mV in lamina II neurons are due to the activation of AMPA receptor-channels, because they are completely blocked by an AMPA receptor-channel antagonist 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX, 10 μM; for example, see [20,26]). NMDA receptor-channels are activated at more positive potentials than –70 mV (see Section 4.11 and Section 4.12). On the other hand, inhibitory postsynaptic currents (IPSCs) are mediated by ionotropic GABAA and glycine receptor-channels ([23]; see [11] for review of the receptor-channels).
L-Glutamate released from the central terminals of primary-afferent Aδ-fibers or C-fibers to lamina II neurons can be examined by measuring monosynaptic EPSCs evoked by stimulating the dorsal root. Aδ-fiber or C-fiber-evoked EPSCs are distinguished on the basis of the conduction velocity of afferent fibers and stimulus threshold to elicit EPSCs; the Aδ-fiber or C-fiber responses, respectively, are considered as monosynaptic in origin when the latency remains constant and there is no failure during repetitive stimulation at 20 Hz for 1 sec, or when failures do not occur during repetitive stimulation at 2 Hz for 10 sec (for example, see [8,27]). On the other hand, L-glutamate released spontaneously from primary-afferent central terminals and glutamatergic interneuron terminals to lamina II neurons can be examined by measuring spontaneous EPSCs (sEPSCs). sEPSCs in lamina II neurons in adult rat spinal cord slices were unaffected in frequency and amplitude by the voltage-gated Na+-channel blocker tetrodotoxin (TTX, 0.5 μM; for example, see [20]), probably because of deafferentiation in the slices used. This result indicates that all of sEPSCs occur without the propagation of action potentials from cell soma, whose neuron is presynaptic to lamina II neurons, to the terminals, resulting in spontaneous releases under our experimental conditions.
GABAergic and glycinergic spontaneous IPSCs (sIPSCs) are, respectively, recorded in the presence of the glycine-receptor antagonist strychnine and the GABAA-receptor antagonist bicuculline. GABAergic sIPSCs are about three-fold longer in duration than glycinergic ones (for example, see [28]). GABAergic and glycinergic sIPSCs were unaffected in frequency and amplitude by TTX (0.5 μM) or a mixture of CNQX (10 μM) and an NMDA receptor-channel antagonist DL-2-amino-5-phosphonovaleric acid (APV, 50 μM; [28]). This result indicates that sIPSCs occur in a manner independent of action potential production, and AMPA and NMDA receptor activation under our experimental conditions.

3. Effects of Hypothalamus-Derived Neuropeptides on Synaptic Transmission in Spinal Lamina II Neurons

3.1. Oxytocin Action

A posterior pituitary hormone oxytocin (a 9-amino acid peptide) has various actions including social interaction and antinociception, other than milk ejection during lactation and uterine contraction during parturition [29,30,31,32]. There is much evidence showing that oxytocin plays a role in regulating nociceptive transmission to the spinal dorsal horn from the periphery. First, there are oxytocin-immunoreactive fibers to the spinal superficial dorsal horn (SDH) from the hypothalamic paraventricular nucleus [33,34], and oxytocinergic axons make synaptic contacts with spinal SDH neurons [35]. Second, oxytocin-binding sites densely exist in the spinal dorsal horn [36,37,38,39,40]. Third, the electrical stimulation of the anterior part of the hypothalamic paraventricular nucleus increased oxytocin concentration in cerebrospinal fluid and produced antinociception [41]. Somatic noxious stimulation activated hypothalamic paraventricular oxytocinergic neurons projecting to the spinal dorsal horn [42]. Intraperitoneal or intrathecal administration of oxytocin reportedly produced antinociception in rats [43,44].

3.1.1. Action of Oxytocin on Holding Current

In 67% of the adult (6–8 weeks old) rat lamina II neurons tested, oxytocin (0.5 μM) produced an inward current at –70 mV (membrane depolarization); the peak amplitude of this current averaged to be 12.6 pA. Remaining all neurons except for neurons (1%) exhibiting an outward current had no effect on holding currents. The peak inward current was concentration-dependent with a half-maximal effective concentration (EC50) value of 0.022 μM. This activity of oxytocin was slow in recovery after its washout. This desensitization may be due to an internalization of oxytocin receptors [29]. Oxytocin-induced inward current was resistant to TTX (0.5 μM), indicating no involvement of action potential production [20]. Moreover, the inward current persisted in Krebs solution containing CNQX (10 μM) and nominally Ca2+-free Krebs solution, indicating a direct action of oxytocin and also no involvement of Ca2+ entry from extracellular solution.
Oxytocin current was mimicked by an oxytocin-receptor agonist TGOT ([Thr4,Gly7]-oxytocin; 0.5 μM) and was reduced in amplitude by an oxytocin-receptor antagonist dVOT ([d(CH2)51,Tyr(Me)2,Thr4,Orn8,des-Gly-NH29]-vasotocin; 1 μM), indicating an involvement of oxytocin receptors. This result is consistent with the localization of oxytocin receptors in the adult rodent spinal SDH [45,46,47]. Consistent with the fact that oxytocin receptors are coupled to guanosine 5’-triphosphate (GTP)-binding proteins (G proteins), oxytocin activity disappeared 60 min after the whole-cell configuration using patch-pipette solutions containing guanosine 5'-O-(2-thiodiphosphate) (GDP-β-S, 1 mM; which has an ability to block the actions of G proteins) [20].
Oxytocin receptors trigger Gq/11, Gs or Gi/o protein-mediated cellular signaling cascades [32]. Oxytocin-induced inward current was decreased in amplitude by U-73122 (10 μM; an inhibitor of phospholipase C (PLC) coupled to Gq/11 protein [48]), indicating an involvement of PLC. Phosphatidylinositol 4,5-bisphosphate hydrolysis caused by PLC leads to the production of two second messengers, diacylglycerol (DAG) that activates protein kinase C (PKC) and inositol 1,4,5-triphosphate (IP3) that releases Ca2+ from intracellular Ca2+ stores. Oxytocin activity was suppressed by 2-aminoethoxydiphenyl borate (200 μM; an IP3-induced Ca2+-release inhibitor [49]) but not dantrolene (10 μM; a Ca2+-induced Ca2+-release inhibitor [50]), chelerythrine (10 μM; a PKC inhibitor [51]) and dibutyryl cyclic-AMP (1 mM), a membrane-permeable analogue of cyclic-AMP (whose intracellular concentration is regulated by Gs and Gi/o protein). Current–voltage relation for the oxytocin current reversed at negative potentials more than the equilibrium potential for K+ or around 0 mV. The oxytocin current was decreased in amplitude in high-K+ (10 mM), low-Na+ (decreased by 117 mM) or Ba2+ (1 mM)-containing Krebs solution. These results indicate that the inward current is due to an alteration in membrane permeabilities to K+ and/or Na+, which is possibly mediated by PLC (whose activation is due to α subunit of Gq/11 protein) and IP3-induced Ca2+ release [20]. The oxytocin current may be mediated by closed K+ channels, as seen in muscarine-sensitive K+ channels [52], and opened Na+ channels; the former idea is supported by a sensitivity of oxytocin current to Ba2+. Breton et al. [53] have reported a closure of A-type and delayed-rectifier K+-channels produced by TGOT in young rat lamina II neurons. A possibility cannot be ruled out in lamina II neurons that not only Gq/11 but also Gs or Gi/o protein is activated by oxytocin and thus an interaction among activations of their G proteins occurs (see [32]).
Oxytocin is known to activate not only oxytocin receptors but also vasopressin receptors with a lower efficacy [54]. Vasopressin receptors are classified into the V1A, V1B and V2 subtypes; antinociception produced by the systemic administration of oxytocin in the mouse is reportedly mediated by vasopressin V1A receptors [39]. Although a vasopressin V1A-receptor antagonist (d(CH2)51,Tyr(Me)2, Arg8)-vasopressin (1 μM) diminished the peak amplitude of oxytocin (0.5 μM) current in lamina II neurons, there was no correlation in amplitude between responses of a vasopressin-receptor agonist [Arg8]-vasopressin (0.5 μM) and oxytocin (0.5 μM) [20]. Therefore, the lamina II oxytocin response appeared to be not mediated by vasopressin V1A receptors. This idea corresponds to the observation that the labeling of spinal SDH layers by [125I]vasopressin antagonist is weaker in extent than that of [125I]oxytocin antagonist in the adult rat [46].
In lamina II neurons where oxytocin (0.5 μM) produced an inward current, noradrenaline (20 μM), serotonin (40 μM) or adenosine (1 mM) elicited an outward current while (-)-nicotine (100 μM) or carbamoylcholine (10 μM) produced an inward current at –70 mV ([20]; see below).

3.1.2. Action of Oxytocin on Excitatory Transmission

Oxytocin (0.5 μM) did not affect the frequency and amplitude of sEPSC, and monosynaptically-evoked primary-afferent Aδ-fiber and C-fiber EPSC amplitude in adult rat lamina II neurons [20]. On the other hand, TGOT is reported to increase the spontaneous release of L-glutamate from nerve terminals in spinal SDH neurons of young (2–4 weeks old) rats [55]. This difference between adult and young rats appeared to be due to a developmental change in synaptic modulation produced by oxytocin. Indeed, in young (11-21 postnatal days) rats, oxytocin (0.5 μM) produced not only an inward or outward current at –70 mV but also presynaptically inhibited or facilitated spontaneous excitatory transmission, depending on the neurons tested [56]. It has been also reported that there is a difference between adult male and female rats in synaptic modulation produced by oxytocin (0.5 μM) in lamina II neurons [56].
Although an oxytocin-induced L-glutamate release increase has been reported in neonate rat spinal SDH neurons in culture [36], Robinson et al. [57] have found an inhibition by oxytocin of primary-afferent-evoked excitatory transmission in adult mouse spinal SDH neurons. There seems to be a difference in excitatory transmission modulation produced by oxytocin among distinct animal species and also ages.

3.1.3. Action of Oxytocin on Inhibitory Transmission

Oxytocin (0.5 μM) increased the frequency of GABAergic and glycinergic sIPSC with a small increase in its amplitude in >90% of the adult rat lamina II neurons tested. The extents of the GABAergic sIPSC frequency and amplitude increase averaged 438% and 66%, respectively, and those of the glycinergic ones 578% and 35%, respectively. These facilitatory effects were slow in recovery after washout of oxytocin, as seen in its depolarized effect. The activity of oxytocin was concentration-dependent; EC50 values for it to increase GABAergic and glycinergic sIPSC frequency were 0.024 and 0.038 μM, respectively. These activities were mimicked by TGOT (0.5 μM), depressed by dVOT (1 μM) and TTX (0.5 μM), indicating an involvement of oxytocin receptors and action potential production [20]. Breton et al. [55] also reported spontaneous GABAergic transmission enhancement produced by TGOT in young rat spinal SDH neurons, although glycinergic transmission was not examined.
These results indicate that oxytocin produces a membrane depolarization in lamina II neurons by activating oxytocin receptors, which increases the neuronal activity of the neurons, leading to the enhancement of inhibitory transmission, a possible mechanism for antinociception [20]. This idea is supported by the observation that the EC50 values (0.024-0.038 μM) for sIPSC frequency increase are closed to that (0.022 μM) for depolarization production. On the other hand, Breton et al. [55] have proposed the idea that an increase in the spontaneous release of L-glutamate onto GABAergic neurons, produced by oxytocin-receptor activation, results in GABAergic transmission enhancement, leading to antinociception.
Recently, oxytocin has been shown to increase intracellular Ca2+ concentration and hyperpolarize membranes in cultured rat capsaicin-sensitive dorsal root ganglion (DRG) neurons [58]. Moreover, oxytocin is reported to activate transient receptor potential vanilloid-1 (TRPV1) channels [59]. TRPV1 channel activation produced by capsaicin results in sEPSC frequency increase and monosynaptically-evoked primary-afferent C-fiber EPSC amplitude reduction in lamina II neurons ([60,61]; for review, see [62]). Since oxytocin (0.5 μM) did not affect sEPSC frequency and C-fiber EPSC amplitude (see above), oxytocin at this concentration did not appear to activate TRPV1 channels located in primary-afferent central terminals in the lamina II.

3.2. Orexins Action

There is much evidence demonstrating that hypothalamic neuropeptides, orexin A (hypocretin 1, 33-amino acid peptide) and orexin B (hypocretin 2, 28-amino acid peptide; [63,64]), play a pivotal role in not only arousal/wakefulness (for review, see [65,66,67]) but also in inhibiting nociceptive transmission in the spinal dorsal horn. For instance, orexinergic fibers in the hypothalamus project to the spinal dorsal horn in rodents [68], orexins A and B exist in the rat spinal cord, albeit the latter is more highly located than the former [69], and the rat spinal cord expresses G protein-coupled orexin-1 and orexin-2 receptors [70,71] that are activated by orexins A and B [64]. Orexin-1 receptors bind more effectively (by about 100 fold) orexin A than orexin B while orexin-2 receptors bind orexins A and B with a comparable affinity [64]. Intrathecal administration of orexins A and B produces antinociception in rodents [72,73,74].

3.2.1. Action of Orexin A on Holding Current

In 18% of the adult rat lamina II neurons examined, orexin A (0.05 μM) produced an inward current at –70 mV (membrane depolarization) that was not accompanied by a change in spontaneous excitatory transmission. On the other hand, 19% of the lamina II neurons did not alter holding currents while exhibiting an increase in sEPSC frequency. Both of these orexin A actions were elicited in 50% of the lamina II neurons. Remaining neurons (13%) did not respond to orexin A. This variability possibly results from a heterogeneity in orexin A receptor expression among different neurons in the lamina II such as islet, central, medial-lateral, radial or vertical type neurons [75]. The peak amplitude of the inward current averaged to be 5.9 pA. The inward current production was repeated, and was concentration-dependent with an EC50 value of 0.0045 μM [21].
The inward current produced by orexin A was reduced in amplitude by an orexin-1 receptor antagonist SB334867 (N-(2-methyl-6-benzoxazolyl)-N'-1,5-naphthyridin-4-yl urea; 1 μM) but not an orexin-2 receptor antagonist JNJ10397049 (N-(2,4-dibromophenyl)-N'-[(4S,5S)-2,2-dimethyl-4 -phenyl-1,3-dioxan-5-yl]-urea; 1 μM; see [76] for their antagonists) and TTX (0.5 μM), indicating an involvement of a direct activation of orexin-1 but not orexin-2 receptors by orexin A without action potential production [21].

3.2.2. Action of Orexin A on Excitatory Transmission

The sEPSC frequency increase produced by orexin A (0.05 μM) averaged 54% in extent, and was not accompanied by a change in sEPSC amplitude. The sEPSC frequency increase was repeated and was concentration-dependent with an EC50 value of 0.030 μM [21]. The sEPSC frequency increase produced by orexin A was diminished in extent by SB334867 (1 μM) but not JNJ10397049 (1 μM) and TTX (0.5 μM), indicating an involvement of a direct activation of orexin-1 but not orexin-2 receptors by orexin A without action potential production [21].
Jeon et al. [77] have reported a similar inward current and spontaneous excitatory transmission enhancement produced by orexin A in young rat lamina II neurons.

3.2.3. Action of Orexin A on Inhibitory Transmission

In 51% of the adult rat lamina II neurons examined, orexin A (0.05 μM) enhanced GABAergic spontaneous transmission that was observed in the presence of strychnine (1 μM). The extents of sIPSC frequency and amplitude increases averaged 119% and 33%, respectively. The orexin A activity was repeated. The GABAergic transmission enhancement was suppressed in extent by SB334867 (1 μM) and TTX (0.5 μM) but not JNJ10397049 (1 μM), indicating an involvement of an activation of orexin-1 but not orexin-2 receptors by orexin A and action potential production [21].
In 79% of the lamina II neurons examined, orexin A (0.05 μM) enhanced glycinergic spontaneous transmission that was observed in the presence of bicuculline (20 μM). The extents of sIPSC frequency and amplitude increases averaged 85% and 38%, respectively. The glycinergic transmission enhancement was decreased in extent by SB334867 (1 μM) and TTX (0.5 μM) but not JNJ10397049 (1 μM), indicating an involvement of an activation of orexin-1 but not orexin-2 receptors by orexin A and action potential production [21].
The GABAergic and glycinergic transmission enhancements are suggested to occur as a result of the production of action potentials by membrane depolarization and increased L-glutamate release through orexin-1 receptors activated by orexin A [21].

3.3. Orexin B Action

3.3.1. Action of Orexin B on Holding Current

As with orexin A, orexin B (0.05 μM) produced an inward current at –70 mV (membrane depolarization) and/or sEPSC frequency increase in adult rat lamina II neurons. In 16% of the neurons tested, orexin B produced an inward current with no change in spontaneous excitatory transmission. On the other hand, 18% of the lamina II neurons did not alter holding currents while producing sEPSC frequency increase. Both of these orexin B actions were elicited in 32% of the lamina II neurons. Remaining neurons (34%) did not respond to orexin B. Although such a variability in response is possibly due to a heterogeneity of lamina II neurons expressing orexin B receptors [75], it has been reported in the young rat that lamina II neurons exhibiting orexin B activity are greater in proportion in radial or vertical neurons than central or unclassified neurons [78]. The peak amplitude of the inward current in adult rats averaged to be 6.5 pA. The inward current production was repeated and was concentration-dependent with an EC50 value of 0.020 μM [22]. This value was four-fold larger than that of orexin A while being comparable to that (0.022 μM) for oxytocin to produce an inward current (see above). Although ionic mechanisms for the inward current produced by orexin B could not be investigated due to its small amplitude, it was suggested that orexin B-induced inward current in young rats may result from a change in membrane permeability to multiple ions [78]. Orexins-induced inward current has been generally attributed to inhibition of K+ channels, stimulation of Na+/Ca2+-exchanger and/or activation of non-selective cation channels (see [79] for review).
The inward current produced by orexin B was resistant to SB334867 (1 μM) and TTX (0.5 μM) while being sensitive to JNJ10397049 (1 μM), indicating an involvement of a direct activation of orexin-2 but not orexin-1 receptors by orexin B without action potential production [22].

3.3.2. Action of Orexin B on Excitatory Transmission

The sEPSC frequency increase produced by orexin B (0.05 μM) averaged 71% in extent, and was not accompanied by a change in sEPSC amplitude. As seen in orexin A activity, the sEPSC frequency increase was repeated and was concentration-dependent with an EC50 value of 0.039 μM [22]. This value was almost comparable to that of orexin A (see above).
The sEPSC frequency increase produced by orexin A was suppressed in extent by JNJ10397049 (1 μM) but not SB334867 (1 μM) and TTX (0.5 μM), indicating an involvement of a direct activation of orexin-2 but not orexin-1 receptors by orexin B without action potential production [22]. This result is consistent with the observations that orexin B is much more potent for orexin-2 than orexin-1 receptors [64] and that orexin-2 receptors exist at high densities in the rat spinal SDH [70].
Grudt et al. [78] have reported that orexin B increases sEPSC frequency with no change in dorsal root-evoked EPSC amplitudes in young rat spinal lamina II neurons.

3.3.3. Action of Orexin B on Inhibitory Transmission

In 71% of the adult rat lamina II neurons examined, orexin B (0.05 μM) enhanced glycinergic spontaneous transmission that was observed in the presence of bicuculline (20 μM). The extents of sIPSC frequency and amplitude increases averaged 110% and 54%, respectively. The orexin B activity was repeated. The glycinergic transmission enhancement was reduced in extent by JNJ10397049 (1 μM) and TTX (0.5 μM) but not SB334867 (1 μM), indicating an involvement of an activation of orexin-2 but not orexin-1 receptors by orexin B and action potential production [22]. It is suggested that the glycinergic transmission enhancement may occur as a result of the production of action potentials by membrane depolarization and increased L-glutamate release through orexin-2 receptors activated by orexin B.
On the other hand, in 24 out of the 26 lamina II neurons tested, orexin B (0.05 μM) had no effect on GABAergic spontaneous transmission that was observed in the presence of strychnine (1 μM). The remaining two neurons exhibited a spontaneous GABAergic transmission enhancement. This result was quite different from that of orexin A (see above; [22]). The result that orexin B enhances glycinergic but not GABAergic transmission is consistent with the observation that strychnine but not bicuculline depresses enhanced inhibitory transmission produced by orexin B in young rats [78]. This difference in orexin B activity between GABAergic and glycinergic transmission was distinct from the activity of oxytocin that enhanced both GABAergic and glycinergic transmission (see above).
In lamina II neurons sensitive to orexin B (0.05 μM), orexin A (0.05 μM) produced an inward current at –70 mV and sEPSC frequency increase with extents comparable to those of orexin B. Moreover, orexin A (0.05 μM) increased glycinergic sIPSC frequency and amplitude with similar extents to those of orexin B (0.05 μM) in the same neuron [22]. This observation that individual lamina II neurons are sensitive to both orexins A and B may be consistent with the fact that both orexins A and B are produced from a precursor peptide and that orexins A and B co-localize in the cat hypothalamus and brain stem [80].
As shown above, there was a difference between orexins A and B in synaptic modulation in lamina II neurons. Orexin A produced a membrane depolarization about four-fold more effectively than orexin B; orexin A enhanced both GABAergic and glycinergic transmission while orexin B only glycinergic transmission. These results may explain a difference between orexins A and B in antinociceptive effects, that is, more effectiveness of orexin A than orexin B [72,73].
Orexin B-sensitive lamina II neurons responded to oxytocin (0.5 μM) with the production of an inward current at –70 mV, but did not exhibit any change in spontaneous excitatory transmission following oxytocin application, indicating clearly a difference in synaptic modulation between orexins and oxytocin [22].
Although orexin B activities similar to those of our studies were reported in young rats [78], orexin B-responsive neurons appeared to be larger in proportion in adult (66–71%) than young (12–29%) rats, albeit orexin B was used at a higher concentration (1 μM) in young rats. There appeared to be a developmental change in orexin B activities.
Post- and presynaptic orexins’ actions similar to those in rat spinal lamina II neurons have been found in other preparations, such as rodent histaminergic tuberomammillary [81], laterodorsal tegmental [82], median preoptic nucleus [83], pedunculopontine tegmental [84], rostral ventrolateral medulla [85] and orexin neurons [86].

4. Effects of Other Endogenous Pain Modulators on Synaptic Transmission in Spinal Lamina II Neurons

4.1. Opioid Actions

Opioids activate three subtypes of opioid receptors, μ-, δ- and κ-type, leading to voltage-gated Ca2+ channel inhibition, inwardly-rectifying K+ channel activation (both of which are due to βγ subunit) or adenylate cyclase inhibition (which is mediated by α subunit) through Gi/o protein activation (for review, see [87]). These opioid receptors are expressed in the spinal SDH, especially lamina II in rats ([88,89]; for review, see [90]). Opioid-binding sites were partially reduced in number in the dorsal horn after the disruption of primary afferents by dorsal rhizotomy [88] or the pretreatment with capsaicin [91], indicating the localization of opioid receptors in the dorsal horn. The rat lamina II reportedly contains endogenous opioid peptides such as enkephalins [92,93], endomorphin-1 (Tyr-Pro-Trp-Phe-NH2) and endomorphin-2 (Tyr-Pro-Phe-Phe-NH2; which is different by only one amino acid from endomorphin-1) [94,95], the latter two of which bind to the μ-opioid receptor with a high affinity compared to δ- and κ-opioid receptors (for review, see [96]). Furthermore, endomorphin-2-like substances were released from the rat spinal dorsal horn in response to electrical stimulation applied to the dorsal root entry zone [97]. Intrathecal administration of opioids produced a powerful analgesia in rats ([98]; for review, see [99]). Opioids administrated into the lamina II in anesthetized cats suppressed an excitation of deeper dorsal horn neurons caused by noxious peripheral stimuli without a change in their responses to innocuous stimuli such as touch [100].
μ- and δ-Type opioid-receptor agonists [(D-Ala2, N-Me-Phe4, Gly5-ol)enkephalin (DAMGO) and (D-Pen2, D-Pen5)enkephalin (DPDPE), respectively; each 1 μM] reduced the peak amplitude of monosynaptically-evoked Aδ-fiber EPSC and also the frequency of miniature EPSC (mEPSC) recorded in the presence of TTX without a change in its amplitude in 70–100% of the adult rat lamina II neurons tested. The actions of DAMGO and DPDPE were not seen in the presence of μ- and δ-opioid receptor antagonists [D-Phe-Cys-Tyr-D-Trp-Arg-Thr-Pen-Thr-NH2 (CTAP) and naltrindole, respectively; each 1 μM], respectively, indicating the presence of μ- and δ-type opioid receptors involved in inhibiting the release of L-glutamate from primary-afferent central terminals and from interneuron terminals [101]. Monosynaptic C-fiber-evoked EPSC was decreased in peak amplitude by DAMGO more effectively than monosynaptic Aδ-fiber-evoked one [102]. On the other hand, κ-type opioid-receptor agonist D-(5 α,7 α,8 β)-(+)-N-methyl-N-[7-(1-pyrrolidinyl)-] -oxaspiro[4,5] dec-8-yl]benzeneacetamide (U-69593; 1 μM) reduced monosynaptic Aδ-fiber EPSC amplitude and mEPSC frequency in only 30% of the lamina II neurons tested [101]. A similar inhibitory effect of U-69593 (0.3 μM) on mEPSC frequency has been demonstrated in young rat lamina II neurons [103]. In about 50% of the adult rat lamina II neurons examined, endomorphin-1 and endomorphine-2 (each 1 μM) activated inwardly-rectifying K+ channels, resulting in an outward current at –70 mV (membrane hyperpolarization). Such a paucity of responsive neurons may be due to a heterogeneity in μ-opioid receptor expression among different lamina II neurons [75]. These endomorphin actions were concentration-dependent with almost the same EC50 value (0.19-0.21 μM) and inhibited by CTAP (1 μM), indicating the activation of μ-opioid receptors in postsynaptic neurons ([104]; for review, see [105]). Yajiri and Huang [106] reported an inhibition of primary-afferent Aδ-fiber-evoked excitatory transmission by endomorphin-1 or endomorphin-2, although their actions were not compared in extent with each other. Endogenous opioids other than endomorphins are reported to hyperpolarize membranes [107]. Although not only μ- but also δ- and κ-opioid receptor agonists exhibited a membrane hyperpolarizing effect in rat lamina II neurons, this effect in individual neurons was distinct in responsiveness among these agonists; μ agonist was effective in a higher proportion of the lamina II neurons tested compared to other agonists [108]. Regarding inhibitory transmission, spontaneous and focally-evoked transmissions mediated by GABA and glycine were not affected by DAMGO (1 μM), where the focal stimulation was performed by using an electrode put within 150 μm of the neuron recorded [101]. In young rat lamina II neurons, both endomorphin-1 and endomorphin-2 are reported to hyperpolarize membranes [94] and to inhibit excitatory transmission [94,109]. Kerchner and Zhuo [110] have reported that DAMGO (1 μM) reduces evoked IPSC amplitude and the frequency of miniature IPSC (mIPSC) recorded in the presence of TTX in rat dorsal horn neurons; it is unknown why there is a difference between their results and ours.

4.2. Nociceptin Action

Nociceptin (a 17-amino acid peptide), which is also known as orphanin FQ, activates G protein-coupled opioid receptor-like1 (ORL1) receptors (recently called nociceptin opioid peptide receptors, NOP receptors) that are similar in structure to the opioid receptors but do not bind opioids. The activation of the NOP receptors results in an inhibition of voltage-gated Ca2+ channels, an activation of inwardly-rectifying K+ channels (both of which by βγ subunit) or an inhibition of adenylate cyclase (by α subunit) by activating Gi/o protein, as seen in opioid receptor activation [111,112]. In spite of such a similarity between the NOP and opioid receptors, these receptors exhibit remarkable differences in receptor functions such as phosphorylation, desensitization and internalization and in their modulatory functions of nociceptive transmission (see [113,114] for review). NOP receptors are densely distributed in the SDH of the adult rat spinal cord [115]. Nociceptin peptide and pre-pronociceptin mRNA [116,117] are densely distributed in the SDH of the rodent spinal cord. Both nociceptin and NOP receptor immunoreactivities were upregulated in rat DRG neurons after nerve injury and inflammation [118]. Furthermore, nociceptin-like substances are reported to be released from the rat spinal dorsal horn in response to electrical stimulation applied to the dorsal root entry zone [119]. According to behavioral studies in adult rats, intrathecal administration of nociceptin produced an antinociceptive effect [120,121,122] and attenuated hyperalgesia in a model of sciatic-nerve injury [123] as well as of inflammation by carageenan injection [124].
In adult rat lamina II neurons, nociceptin (1 μM) activated an inwardly-rectifying K+ channel, resulting in membrane hyperpolarization; this activity was concentration-dependent with an EC50 value of 0.23 μM [125]. Nociceptin (1 μM) also reduced the peak amplitudes of monosynaptic Aδ-fiber and C-fiber EPSCs evoked in lamina II neurons by stimulating the dorsal root, where C-fiber EPSCs (57% peak amplitude reduction) were more sensitive to nociceptin than Aδ-fiber ones (30%). Since nociceptin did not affect a response of lamina II neurons to bath-applied AMPA, this action was presynaptic in origin, i.e., due to a decrease in the release of L-glutamate from primary-afferent central terminals [126]. These nociceptin actions were inhibited by a nociceptin precursor nocistatin (1 μM) and an NOP receptor antagonist, 1-[(3R, 4R)-1-cyclooctylmethyl-3-hydroxymethyl-4-piperidyl]-3-ethyl-1,3-dihydro-2H-benzimidazol-2-one (CompB or J-113397; 1-3 μM [127]), indicating that the activation of NOP receptor leads to a decrease in the excitability of lamina II neurons and thus to antinociception. Like DAMGO, nociceptin (1 μM) did not affect spontaneous and focally-evoked inhibitory transmissions mediated by GABA and glycine in lamina II neurons [126]. Excitatory transmission inhibition, the lack of changes in inhibitory transmission and membrane hyperpolarization produced by nociceptin have been demonstrated in young rat lamina II neurons [128,129,130].
Although nociceptin exhibits modulatory actions similar to those of opioids in lamina II neurons, nociceptin but not opioids inhibit T-type voltage-gated Ca2+ channels in rat DRG neurons, suggesting a difference between the two neuropeptides in modulating primary-afferent-evoked glutamatergic transmission (see [131] for review).

4.3. Adenosine Action

Adenosine activates three subtypes of G protein-coupled metabotropic receptors as classified into A1-, A2- (A2A-, A2B-) and A3-types. The A1 and A2 adenosine receptors (Gi/o- and Gs-protein coupled ones, respectively) are primarily coupled to adenylate cyclase in a negative and positive manner, respectively, while the A3 one activates an IP3/DAG system through PLC (by α subunit of Gq/11 protein); the A1 receptor also opens K+ channels or closes Ca2+ channels (by α subunit) through Gi/o protein activation (for review, see [132]). The presence of A1 adenosine receptors at a high density in the spinal dorsal horn has been demonstrated in terms of A1 agonist-binding sites [133], A1 adenosine receptor mRNAs [134] and proteins [135]. Adenosine in the lamina II would be either released from neurons and/or glial cells or produced as a result of a cleavage by ectonucleotidases of ATP released from them. In support of a role of adenosine in the lamina II, this region has a high density of the rat equilibrative nucleoside transporter (rENT1) that controls the extracellular level of nucleosides such as adenosine [135]. Behavioral studies have demonstrated that an intrathecal administration of adenosine analogues produces antinociception in the hot plate and tail flick tests [136]. The antinociceptive effect of adenosine has been reported to be due to the activation of the A1 adenosine receptor ([137,138]; for review, see [139]). Recent studies have demonstrated an effectiveness of adenosine A3 receptor activation in alleviating neuropathic pain in rodents [140]; this antinociceptive effect appears to be due to an inhibition of enhanced microglial activation in the spinal dorsal horn [141].
In adult rat lamina II neurons, adenosine concentration-dependently produced an outward current at –70 mV (membrane hyperpolarization) and reduced the frequency of sEPSC without a change in its amplitude; their EC50 values were 177 and 277 μM, respectively [142,143]. The outward current was due to the activation of K+ channels that were sensitive to Ba2+ (100 μM) and 4-aminopyridine (5 mM) while being resistant to tetraethylammonium (TEA; 5 mM; [143]). Monosynaptic Aδ-fiber and C-fiber-evoked EPSCs were depressed in amplitude by adenosine (100 μM) with a comparable extent (26% and 27%, respectively, in the same neuron). EC50 value for adenosine in reducing Aδ-fiber EPSC peak amplitudes was 217 μM, a value similar to those of outward current and sEPSC frequency reduction produced by adenosine [144]. All of the adenosine actions were mimicked by an A1 adenosine-receptor agonist, N6-cyclopentyladenosine (1 μM), and blocked by an A1 antagonist, 8-cyclopentyl-1,3-dipropylxanthine (1 μM), indicating an involvement of the A1 adenosine receptor [142,143]. Spontaneous and focally-evoked GABAergic and glycinergic IPSCs were also suppressed in frequency and amplitude, respectively, by adenosine (100 μM) through the activation of presynaptic A1 adenosine receptors. EC50 values for adenosine in reducing evoked GABAergic and glycinergic IPSC amplitude were 14.5 and 19.1 μM, respectively ([145]; see [146] for review). These findings are consistent with the presence of the A1 adenosine receptor in the spinal dorsal horn. It may be of interest to note an enhancement of adenosine A1 receptor sensitivity at excitatory but not inhibitory synapses in the lamina II in a rat partial nerve-injury model of neuropathic pain [147]. This result suggests that neuropathic pain can alter the modulatory effect of adenosine on excitatory transmission in lamina II neurons.
In young hamster spinal lamina II neurons, it has been demonstrated that adenosine inhibits glutamatergic excitatory transmission by a pre- and postsynaptic mechanism; the latter action is due to the activation of K+ channels [148].

4.4. ATP Action

ATP activates P2 receptors classified into two subtypes, ionotropic P2X receptors and G protein-coupled metabotropic P2Y receptors; seven P2X receptors (P2X1-P2X7) and eight P2Y receptors (P2Y1, P2Y2, P2Y4, P2Y6, P2Y11, P2Y12, P2Y13 and P2Y14) have been characterized [149,150]. ATP in the lamina II would originate from primary-afferent central terminals or intrinsic neurons and/or glia cells in the spinal dorsal horn, generally being released from the cytoplasm to extracellular space through a nucleoside transporter.
In young hamster lamina II neurons, ATP (1-5 mM) induced a fast inward current (membrane depolarization) in a manner sensitive to a P2X- and P2Y-receptor antagonist, suramin (0.5 mM), and potentiated the peak amplitude of dorsal root-evoked EPSC [151]. A P2X- and P2Y-receptor antagonist, pyridoxal-phosphate-6-azophenyl-2',4'-disulfonic acid (PPADS; 10 μM), presynaptically reduced the peak amplitude of dorsal root-evoked EPSCs in young rat SDH neurons [152], although PPADS at 50 μM did not affect spontaneous and electrically-evoked EPSCs [153]. With respect to inhibitory transmission, ATP (10 μM) enhanced the release of glycine from nerve terminals in dissociated rat lamina II neurons attached with synaptic buttons in a manner sensitive to PPADS (10 μM) while resistant to N-ethylmaleimide (3 μM; a sulfhydryl alkylating agent having an ability to block G-protein functions), indicating an involvement of P2X receptors [154]. Intrathecal administration of PPADS did not produce any antinociceptive effects when examined by the tail-flick test [152]. These results demonstrate that ATP may play some role in modulating nociceptive transmission in the SDH, albeit this being unclear.

4.5. Noradrenaline Action

Adrenoceptors, which are activated by noradrenaline, are classified into at least three subtypes of α1 (1A-, 1B-, 1D-types), α2 (2A-, 2B-, 2C-types) and β (1-, 2-, 3-types), all of which are G protein-coupled metabotropic receptors. The α1 adrenoceptors activate an IP3/DAG system through PLC (by α subunit of Gq/11 protein). On the other hand, the α2 adrenoceptors either inhibit adenylate cyclase (by α subunit of Gi/o protein) or regulate the activation of K+ or Ca2+ channels through βγ subunit of Gi/o protein, leading to their opening and closing, respectively. All of the β adrenoceptors activate adenylate cyclase (by α subunit of Gs protein; for review, see [155]). The α1A/D and α1B receptor mRNAs are present in the rat spinal cord, albeit being at a low density [156]. The α2A and α2C adrenoceptors are reported to exist in primary-afferent C-fiber central terminals and interneurons, respectively, in the SDH [157]. The rat SDH does not express β1 and β2 mRNAs [158]. There is a descending noradrenaline-containing fiber pathway from cell groups designated A5, A6 (nucleus locus ceruleus) and A7 (subceruleus) in the pons to the spinal dorsal horn [159,160]. Electrical stimulation of this pons region results in behavioral analgesia [161]. This noradrenaline pathway is also activated by systemically-administrated opioids [162] or electrical stimulation of the midbrain periaqueductal gray region [163]. Intrathecal administration of noradrenaline itself is known to have an antinociceptive effect when assessed by the tail-flick and hot-plate tests [164,165].
Noradrenaline acts on both pre- and postsynaptic sites in the spinal dorsal horn. In adult rat lamina II neurons, noradrenaline (10 μM) induced an outward current at –70 mV (membrane hyperpolarization); this effect was mimicked by an α2 agonist, clonidine (10 μM), and was blocked by an α2 antagonist, yohimbine (0.5 μM), indicating that this current response is due to the activation of α2 adrenoceptors [166,167]. Noradrenaline (10-100 μM) also enhanced the release of GABA and glycine to lamina II neurons from inhibitory neurons; this action was mimicked by an α1 agonist, phenylephrine (10-60 μM), and was inhibited by an α1 antagonist, prazocin (0.5 μM), and an α1A antagonist, 2-(2,6-dimethoxyphenoxyethyl)aminomethyl-1,4-benzodioxane (WB-4101; 0.5 μM), indicating an involvement of α1, possibly α1A adrenoceptors [166,168,169]. Noradrenaline (50 μM) did not affect sEPSC frequency and amplitude, but inhibited the release of electrically-evoked L-glutamate to lamina II neurons from primary-afferent Aδ-fiber and C-fiber central terminals, where Aδ-fiber EPSCs (50% peak amplitude reduction) were more sensitive to noradrenaline than C-fiber ones (28%) [170]. These inhibitory actions were mimicked by clonidine (10 μM) and an α2A agonist, oxymetazoline (10 μM), and was blocked by yohimbine (1 μM), indicating an involvement of α2, possibly α2A adrenoceptors [170]. A similar inhibitory action of clonidine has been reported for excitatory transmission in neurons existing in the outer layer of the lamina II [171]. All of the noradrenaline actions result in a decrease in the excitability of lamina II neurons. A β-adrenoceptor agonist, isoproterenol (40 μM), did not affect the inhibitory and excitatory transmission [166,170].

4.6. Serotonin Action

Serotonin (5-hydroxytryptamine, 5-HT) receptors are G protein-coupled metabotropic receptors except for the 5-HT3 receptor which is a cation-permeable channel. The metabotropic 5-HT receptors are composed of at least 5-HT1 (A-, B-, D-, E-, F-types), 5-HT2 (A-, B-, C-types), 5-HT4, 5-HT5 (A-, B-types), 5-HT6 and 5-HT7 receptors. The 5-HT1 and 5-HT5 receptors are negatively coupled to adenylate cyclase (by α subunit of Gi/o protein) while the 5-HT4, 5-HT6 and 5-HT7 receptors activate adenylate cyclase (by α subunit of Gs protein). The 5-HT2 receptor activates an IP3/DAG system through PLC (by α subunit of Gq/11 protein; for review, see [172]). The 5-HT receptors are expressed within the spinal cord [173,174]; particularly, binding sites for (+)-hydroxy-2-(di-n-propylamino)tetralin (8-OH-DPAT, an agonist specific to the 5-HT1A and 5-HT7 receptors) are expressed in the SDH including lamina II [174]. There are descending inhibitory serotonergic systems from the medullary raphe nuclei in the brainstem to the spinal dorsal horn [175,176]. Electrical stimulation of the nucleus raphe magnus releases 5-HT in the spinal dorsal horn [177]. Intrathecal application of 5-HT and 8-OH-DPAT resulted in antinociception when estimated using the tail-flick test [178].
In adult rat lamina II neurons, 5-HT (40 μM) induced either outward or inward currents at –70 mV, indicating a heterogeneity in 5-HT receptor subtype expression among different lamina II neurons [75,179]. The former (membrane hyperpolarization) was mimicked by 8-OH-DPAT (10 μM), and was completely blocked by a selective 5-HT1A receptor antagonist, WAY 100635 (10 μM), indicating an involvement of 5-HT1A receptors [180]. On the other hand, the latter (membrane depolarization) was observed in a small number of the neurons tested, and was mimicked by a 5-HT3 receptor agonist, 1-(m-chlorophenyl)-biguanide (mCPBG; 30 μM), indicating an involvement of 5-HT3 receptors [180]. When examined in morphologically-identified neurons, the 5-HT-induced outward currents were produced in vertical (21/34), small islet (11/34) and radial cells (2/34) while 5-HT-induced inward currents in islet (1/5) and small islet cells (4/5), indicating that 5-HT produces a membrane hyperpolarization in excitatory neurons, because most vertical cells are glutamatergic [180]. 5-HT (40 μM) also inhibited the release of L-glutamate to lamina II neurons from primary-afferent Aδ-fiber and C-fiber central terminals as noradrenaline did; inhibitions of Aδ-fiber and C-fiber responses were comparable in extent (each 39%). The action of 5-HT on C-fiber responses was mimicked by 8-OH-DPAT, but was not blocked by WAY 100635, indicating that 5-HT receptors existing in primary-afferent C-fiber central terminals appear to be a 5-HT1A-like type that is neither the 5-HT1A- nor the 5-HT7-type [9]. With respect to inhibitory transmission, 5-HT (100 μM) and mCPBG (30 μM) increased mIPSC frequency and amplitude in lamina II neurons; its amplitude increase was sensitive to TTX (1 μM). This result indicates that the inhibitory transmission enhancement is mediated by action potential production occurring as a result of 5-HT3 receptor-mediated depolarization [180]. Fukushima et al. [181] have reported spontaneous GABAergic transmission enhancement mediated by 5-HT3 receptors in adult mouse spinal SDH neurons. Moreover, 5-HT3 receptor activation in the spinal cord is shown to increase the release of GABA by using the microdialysis method [182].

4.7. Dopamine Action

Dopamine receptors, which are activated by dopamine, are classified into two subtypes of D1-like (D1 and closely-related D5) and D2-like (D2, closely-related D3 and D4) receptors, all of which are G protein-coupled metabotropic receptors. The D1 and D5 receptors activate adenylate cyclase (by α subunit of Gs protein) while the D2, D3 and D4 receptors are negatively coupled to adenylate cyclase (by α subunit of Gi/o protein; [183]). Both D1-like and D2-like receptors exist in the rat spinal cord at a high density [184]. There are descending dopaminergic pathways to the spinal dorsal horn from the periventricular posterior region (A11) of the hypothalamus in rats [185,186,187,188]. Intrathecally-administrated apomorphine (a D2 receptor agonist) inhibited thermally and chemically evoked noxious responses in rats ([189]; see [190] for review).
In adult rat lamina II neurons, dopamine concentration-dependently produced an outward current at –70 mV (membrane hyperpolarization; EC50 = 77.8 μM) in a manner resistant to TTX and CNQX and sensitive to intracellular GDP-β-S and extracellular Ba2+. The outward current produced by dopamine was mimicked by a D2-like receptor agonist quinpirole (30 μM) and depressed in amplitude by a D2-like receptor antagonist sulpiride (30 μM), indicating K+ channel opening (possibly by βγ subunit of Gi/o subunit) through D2-like receptor activation [191,192]. On the other hand, dopamine (100 μM) did not affect the frequency and amplitude of mEPSCs recorded in the presence of TTX [191].

4.8. Somatostatin Action

Somatostatin (a 14-amino acid peptide) receptors (SSTRs) are classified into at least five subtypes named SSTR1-SSTR5, all of which are coupled to G protein, resulting in inhibiting adenylate cyclase (by α subunit), opening K+ channels or closing Ca2+ channels (by βγ subunit) through Gi/o protein activation (see [193] for review). The SDH including lamina II contains SSTR-like immunoreactivity for the SSTR1-SSTR3 [194]. Somatostatin is expressed in rat DRG neurons [195] and somatostatin-positive fibers are localized at the highest density in the rat lamina II [196]. Intrathecal administration of somatostatin produced antinociceptive responses to noxious heat in cats [197] and inhibited nociceptive responses to subcutaneous formalin in rats [198].
In adult rat lamina II neurons, somatostatin produced an outward current at –60 mV (membrane hyperpolarization) in a manner resistant to TTX and in a concentration-dependent manner with an EC50 value of 0.82 μM [199,200]. Such a response to somatostatin (1 μM) was seen in about 50% of the neurons tested, indicating a heterogeneity in somatostatin receptor expression among different lamina II neurons [75]. The somatostatin activity was sensitive to intracellular Cs+ and TEA, and extracellular Ba2+, indicating an involvement of K+ channels. Consistent with this idea, somatostatin current exhibited an inwardly-rectifying property and reversed at a potential close to the equilibrium potential for K+. With respect to synaptic transmission, somatostatin (1 μM) had no effect on mEPSC frequency and amplitude, monosynaptic dorsal root-evoked EPSC amplitudes, and the frequency and amplitude of GABAergic and glycinergic mIPSCs [199]. A similar outward current produced by somatostatin has been reported in juvenile rat lamina II neurons [201]. It has not been examined what kinds of SSTRs are involved in the somatostatin-induced outward currents.

4.9. Cannabinoid Action

Cannabinoid receptors, which are activated by cannabinoids having an ability to alleviate pain, are classified into two subtypes, Gi/o protein coupled CB1 and CB2 receptors [202,203]. Intrathecal administration of a prototypical cannabinoid D9-tetrahydrocannabinol resulted in antinociception in the tail-flick test in adult rats [204]. A similar antinociceptive effect was produced by a mixed CB1/CB2 receptor agonist WIN55,212-2 (WIN-2) [205]. Such behavioral results are possibly mediated by CB1 receptors in the spinal dorsal horn, because localization of CB1 receptors has been demonstrated there by in situ hybridization [206], agonist binding [207] and immunohistochemistry [208]. A selective CB1-receptor antagonist SR141716A facilitated nociceptive responses of rat spinal dorsal horn neurons [209].
In adult rat lamina II neurons, an endocannabinoid N-arachidonoylethanolamide (anandamide; 0.01-10 μM) and WIN-2 (5-10 μM) had no effect on holding currents at –70 mV, sEPSC frequency and amplitude while reducing monosynaptically-evoked Aδ-fiber and C-fiber EPSC amplitudes; the former reduction (32% reduction by 10 μM anadamide) was larger than the latter (17% reduction) [210]. Although anandamide is thought to be an endogenous ligand of TRPV1 channels in vascular preparations [211], the anadamide activity in lamina II neurons was due to the activation of cannabinoid receptors but not TRPV1 channels, because the actions of anandamide on excitatory transmission were quite different from those of capsaicin [60,61]. A similar inhibitory action of anandamide on excitatory transmission has been shown in juvenile rat lamina II neurons [212]. With respect to inhibitory transmission, anandamide (10 μM) reduced focally-evoked GABAergic and glycinergic IPSC amplitudes in a manner sensitive to SR141716A (5 μM) in adult rat lamina II neurons [213]. Since anandamide (10 μM) reduced the frequency of GABAergic and glycincergic sIPSC without a change in the amplitude, its activities on evoked inhibitory transmission were pre- but not post-synaptic in origin [213]. Similar actions on inhibitory transmission were induced by WIN-2 (5 μM) and an endogenous cannabinoid-receptor agonist 2-arachydonoyl glycerol (20 μM; [214]) [213].

4.10. Galanin Action

Galanin (a 29/30-amino acid peptide), which was first extracted from porcine upper intestines [215], was reported to extensively exist in the peripheral and central nervous systems (see [216] for review). Galanin serves as a neurotransmitter or neuromodulator in various physiological functions such as feeding and pain (see [216,217,218] for review). There are three subtypes of G protein-coupled metabotropic receptor (GalR1, 2, 3: coupled to Gi/o; Gi/o or Gq/11; and Gi/o protein; respectively) for galanin [216]. There is much evidence for the idea that galanin plays a role in regulating nociceptive transmission to the spinal dorsal horn from the periphery. First, galanin immunoreactivity, GalR1, 2, 3 mRNAs and proteins are located in the rat DRG and the spinal dorsal horn [219,220,221,222,223,224,225]. The expression of galanin was upregulated in DRG neurons after nerve injury and in dorsal horn neurons after inflammation [216,218]. For example, peripheral inflammation induced by the injection of carrageenan into the hindpaw of rats increased the number of galanin mRNA-positive neurons in the spinal SDH [226]. Second, intrathecally administrated galanin modulated nociceptive responses in rats [227,228,229,230]. Transgenic mice overexpressing galanin in a population of DRG neurons exhibited nociceptive responses different from those of wild-type controls [231]. The intrathecal administration of galanin produced such a biphasic effect, as nociception at low doses and antinociception at high doses [229,230].
In adult rat lamina II neurons, galanin (0.03 μM) increased the frequency of sEPSC without a change in its amplitude, indicating a presynaptic effect; this action was concentration-dependent with an EC50 value of 2.0 nM. This effect reduced in extent in Ca2+-free or a voltage-gated Ca2+-channel blocker La3+ (30 μM)-containing Krebs solution, and was mimicked by a GalR2/R3 agonist galanin 2-11 [229] but not a GalR1 agonist M617 (galanin(1-13)-Gln14-bradykinin (3-9)amide [232]; each 0.03 μM). Galanin also produced in a concentration-dependent way an outward current at –70 mV (membrane hyperpolarization) with an EC50 value of 44 nM, a value larger than that for sEPSC frequency increase. This outward current was mimicked by M617 but not galanin 2-11 (each 0.1 μM). Moreover, galanin (0.1 μM) reduced monosynaptically-evoked Aδ-fiber and C-fiber EPSC amplitudes; the former reduction (35%) was larger than the latter (12%). A similar action was produced by galanin 2-11 but not M617 (each 0.1 μM). Spontaneous and focally-evoked inhibitory transmissions mediated by GABA and glycine were unaffected by galanin (0.1 μM). These results indicate that galanin at lower concentrations enhances the spontaneous release of L-glutamate from nerve terminals due to increased intracellular Ca2+ concentration by Ca2+ entry from external solution following GalR2/R3 activation while galanin at higher concentrations also produces a membrane hyperpolarization by activating GalR1 receptors. Moreover, galanin reduces L-glutamate release onto lamina II neurons from primary-afferent central terminals by activating GalR2/R3 receptors [233]. These effects could contribute to at least a part of the biphasic behavioral effect of galanin. Alier et al. [234] have reported an inhibition by galanin of excitatory transmission evoked in lamina II neurons by stimulating the dorsal root entry zone in young adult rats.

4.11. Substance P Action

Substance P (a 11-amino acid peptide) is a member of tachykinin family together with neurokinins A and B. Neurokinin (NK) receptors, which are activated by tachykinins, are classified into three subtypes of NK-1, NK-2 and NK-3, all of which are G-protein coupled metabotropic receptors, leading to the activation of PLC (by α subunit of Gq/11 protein). Agonists most sensitive to the NK-1, -2 and -3 receptors are substance P, neurokinin A and neurokinin B (all of which are small peptides sharing a common amino acid sequence at their carboxy terminal), respectively (for review, see [235]). It is well-known that the activation of the NK1 receptor by substance P depolarizes membranes of dorsal horn neurons, leading to nociception [236] and that further such depolarization serves as a relief of NMDA receptors from a voltage-dependent block by Mg2+ (see [11]) which in turn results in sensitization called wind-up, a progressive increase in the number of action potentials evoked per stimulus in response to a repetitive stimulation of primary-afferent C-fibers [237,238,239].
Although the lamina II has the highest density of substance P-containing primary-afferent C-fiber terminals [240], this peptide seemed not to be involved in synaptic modulation in adult rat lamina II neurons, because substance P (1 μM) did not induce any responses [241] and a repeated stimulation (20 Hz for 1 sec) of primary-afferent C-fibers which was expected to release neuropeptides such as substance P did not produce any slow synaptic responses [26]. This result is consistent with the observation that there are few NK1 receptor-like immunoreactive neurons in the lamina II [242]. It may be possible that substance P released from the C-fiber endings acts on dendrites of neurons whose cell bodies exist in IV/V laminae deeper than the lamina II, resulting in membrane depolarization in deep dorsal horn neurons, as reported previously [243]. Projection neurons in the lamina I have a depolarizing response to substance P that plays a role in the induction of synaptic plasticity occurring there [244].

4.12. Bradykinin Action

Bradykinin (a 9-amino acid peptide) is locally produced from kininogens by kallikreins and kininases at the site of the injured and inflamed tissue. Bradykin receptors are classified into two subtypes of injury-induced B1 receptor and constitutively-expressed B2 receptor, both of which are G-protein coupled metabotropic receptors, leading to the activation of PLC (by α subunit of Gq/11 protein; see [245] for review). Although the B1 and B2 receptors are located in the peripheral terminals of primary-afferent neurons and involved in peripheral sensitization (reduction in threshold for receiving nociceptive stimuli) ([246]; see [247] for review), there is evidence showing that bradykinin plays a role in modulating nociceptive transmission in the spinal dorsal horn. Intrathecal administration of a B2-receptor antagonist reduced the second phase of nociceptive responses to intraplantar formalin in rats [248] and intrathecally-administrated B1- and B2-receptor agonists produced thermal hyperalgesia measured by the hot-plate test in mice [249].
In adult rat lamina II neurons, bradykinin but not a B2-receptor agonist des-Arg9-bradykinin (each 10 μM) increased both bath-applied AMPA and NMDA responses that were measured at –70 and –40 mV (at the latter potential, a relief of blockade by Mg2+ of NMDA receptor-channels is expected; see [11]), respectively; the increases in the AMPA and NMDA responses were reduced in extent by intracellular GDP-β-S. These results indicate an increase in a sensitivity of AMPA and NMDA receptors to L-glutamate, caused by B1 receptor activation by bradykinin. Consistent with this postsynaptic effect, bradykinin (10 μM) increased the peak amplitude of monosynaptically-evoked primary-afferent Aδ-fiber and C-fiber EPSCs. Moreover, bradykinin but not des-Arg9-bradykinin (each 10 μM) increased the frequency and amplitude of mEPSC recorded in the presence of TTX, indicating not only postsynaptic but also presynaptic facilitatory actions of bradykinin, mediated by B1 receptors. This action was not accompanied by a change in holding currents at –70 mV. Such a glutamatergic transmission enhancement could explain the observation that intrathecal administration of bradykinin produced thermal hypersensitivity that was suppressed by an NMDA-receptor antagonist MK-801 [250].

4.13. Neuropeptide Y Action

Neuropeptide Y (a 36-amino acid peptide; C-terminal amidated one), which extensively exists in the peripheral and central nervous systems [251], plays a role in various physiological functions such as feeding and pain ([252,253]; see [254] for review). Neuropeptide Y receptors, which are activated by neuropeptide Y, are classified into at least six subtypes of Y1-Y6, all of which are G protein (Gi/o protein)-coupled metabotropic receptors [255]. There is much evidence for the idea that neuropeptide Y plays a role in regulating nociceptive transmission to the spinal dorsal horn from the periphery. First, neuropeptide Y-like immunoreactivity is densely located in the laminae I-II [256], Y1 receptors are expressed in small-sized DRG neurons and the spinal dorsal horn [257], and Y2 receptors are located in large-sized DRG neurons in rats [258]. Second, intrathecally-administrated neuropeptide Y produced antinociception in the hot-plate test in rats [259]. Third, Y1 receptor-knockout mice exhibited hyperalgesia in response to acute thermal, cutaneous and visceral chemical stimuli, while completely lacking the analgesic effects of neuropeptide Y [260].
In 33% of the adult rat lamina II neurons examined, neuropeptide Y (1 μM) produced an outward current at –60 mV (membrane hyperpolarization) in a manner resistant to TTX. Such a paucity of responsive neurons also shows a heterogeneity in neuropeptide Y receptor expression among different lamina II neurons [75]. The neuropeptide Y activity was sensitive to intracellular Cs+ and TEA, and extracellular Ba2+, indicating an involvement of K+ channels. Consistent with this idea, neuropeptide Y current exhibited an inwardly-rectifying property and reversed at a potential close to the equilibrium potential for K+. Moreover, neuropeptide Y activity was sensitive to intracellular GDP-β-S, was mimicked by a Y1-receptor agonist [Leu31,Pro34]-neuropeptide Y (1 μM) and suppressed by a Y1-receptor antagonist BIBP 3226 (1 μM), indicating Y1 receptor activation. With respect to synaptic transmission, neuropeptide Y (1 μM) had no effect on sEPSC frequency and amplitude, monosynaptic dorsal root-evoked EPSC amplitudes, GABAergic and glycinergic mIPSC frequency and amplitude, and focally-evoked GABAergic and glycinergic IPSC amplitudes [261].

4.14. Phospholipase A2 Activation Action

Phospholipase A2 (PLA2) is thought to play a pivotal role in a variety of physiological functions including nociception through the production of arachidonic acid which is one of fatty acids released from the sn-2 position of membrane phospholipids by PLA2 activation (for review, see [262]). The arachidonic acid is involved in regulating neuronal functions as a result of the synthesis of eicosanoids such as prostanoids (for review, see [263]) or without conversion to metabolites (for review, see [264]). The spinal cord contains the small molecular-weight secreted PLA2 (sPLA2) and the large molecular-weight cytosolic PLA2 (for review, see [265] and [266]).
Melittin (a 26-amino acid basic peptide; a major component of the bee venom [267]) is known to be an in vitro activator of sPLA2 with no effect on cytosolic PLA2 [268,269,270]. In adult rat lamina II neurons, melittin (1 μM) did not change holding current at –70 mV while increasing the frequency and amplitude of sEPSC. This frequency increase was concentration-dependent with an EC50 value of 1.1 μM. Melittin activity disappeared in the presence of a selective PLA2 inhibitor 4-bromophenacyl bromide (4-BPB, 10 μM; [271]) while being unaffected by TTX (0.5 μM), a cyclooxygenase inhibitor indomethacin (100 μM) and a lipoxygenase inhibitor nordihydroguaiaretic acid (NDGA; 100 μM), indicating an involvement of sPLA2 activation and possibly arachidonic acid but not its metabolites produced by cyclooxygenase and lipoxygenase [272]. Melittin (1 μM) also increased the frequency and amplitude of sIPSC; the effect of melittin on GABAergic but not glycinergic sIPSC was not seen in the presence of TTX (0.5 μM) and CNQX (10 μM). EC50 values for melittin to increase glycinergic sIPSC frequency and amplitude were 0.73 and 0.64 μM, respectively [28]. 4-BPB, another PLA2 inhibitor aristolochic acid (100 μM; [273]), and NDGA (100 μM) but not indomethacin (100 μM) inhibited the facilitatory action of melittin on glycinergic transmission. These results indicate that the GABAergic transmission enhancement is mediated by glutamate-receptor activation and neuronal activity increase, possibly owing to excitatory transmission enhancement, while the glycinergic one is due to PLA2 and subsequent lipoxygenase activation [28]. The GABAergic transmission enhancement was attributed to actions of acetylcholine (ACh) and noradrenaline which activate ACh receptors (AChRs; nicotinic and muscarinic types; see below) and α1 adrenoceptors, respectively ([169]; see [274] for review). With respect to arachidonic acid’s metabolites, prostaglandin E2 (10 μM) is reported to produce a membrane depolarization with no effect on excitatory transmission in a minority of the adult rat lamina II neurons examined [275]. In mouse spinal SDH neurons, prostaglandin E2 increased mEPSC frequency [276]. There appeared to be a difference in prostaglandin E2 activity between animals used.

4.15. Acetylcholine Action

AChRs are classified into ionotropic nicotinic AChRs (nAChRs) and metabotropic muscarinic AChRs (mAChRs). The nAChRs in the central nervous system are composed of either a combination of α (2-6 types) and β (2-4 types) subunits or a homomer of α7-α9 subunits (for review, see [277]), while the mAChRs are composed of M1, M2, M3, M4 and M5 receptors. The M1, M3 and M5 receptors activate an IP3/DAG system through PLC (by Gq/11 protein), while the M2 and M4 receptors inhibit adenylate cyclase (by Gi/o protein; [278]). The adult rat lamina II contains α3, α4, α5 and β2 subunit mRNAs [279,280]. A high density of mAChRs which bind [3H]-pirenzepine has been demonstrated in the lamina II by autoradiographic studies in adult rats [281]. Choline acetyltransferase (an enzyme which synthesizes ACh)-immunoreactive neurons in the lamina III are frequently presynaptic to lamina II neurons [282] and this enzyme co-localizes with GABA in the spinal dorsal horn [283], suggesting that ACh in the lamina II originates from inhibitory neurons in the spinal dorsal horn. Intrathecal administration of a mAChR agonist carbamoylcholine or ACh esterase inhibitors (reversible ones: neostigmine and physostigmine; irreversible one: echothiophate) produced analgesia to noxious thermal stimuli in rats [284,285]. On the other hand, intrathecal application of nAChR agonists, nicotine, cytisine, A-85380 and epibatidine, resulted in nociception but the latter two drugs also produced antinociception in a manner sensitive to a broad-spectrum nAChR antagonist, mecamylamine, when evaluated by the paw-withdrawal test to heat, suggesting the presence of a type of nAChRs involved in antinociception in the spinal dorsal horn [286,287].
In adult rat lamina II neurons, (-)-nicotine (100 μM) or carbamoylcholine (10 μM) elicited an inward current at –70 mV (membrane depolarization) [20,288,289]. ACh also has an ability to enhance the release of GABA and glycine from the terminals of GABAergic and glycinergic neurons by the activation of nAChRs and mAChRs. (-)-Nicotine (100 μM) and (-)-cytisine (20 μM) enhanced the frequency and amplitude of GABAergic and glycinergic sIPSCs in a manner sensitive to mecamylamine (5 μM; [169,289]). Baba et al. [288] have demonstrated that carbamoylcholine (10 μM) and neostigmine (10 μM; which is expected to increase the concentration of ACh in the synaptic cleft) increase the frequency and amplitude of GABAergic sIPSC (also see [169]). These results suggest that antinociception produced by ACh is mediated by inhibitory neurotransmitters such as GABA and glycine released by the activation of nAChRs and mAChRs.

5. Similarity and Difference among Endogenous Neuromodulators in Antinociceptive Mechanisms at Cellular Levels in the Spinal Lamina II

Table 1 demonstrates a comparison of synaptic modulation produced by oxytocin, orexins A and B with those of other endogenous pain neuromodulators in rodent lamina II neurons. Membrane depolarization produced by oxytocin [20], and membrane depolarization and/or enhanced spontaneous L-glutamate release produced by orexins A and B [21,22], in lamina II neurons could increase the membrane excitability of these neurons. This depolarizing effect was distinct from those of analgesic neuropeptides (endomorphins [104], nociceptin [125,128], somatostatin [199,200], galanin [233] and neuropeptide Y [261]), adenosine [143], noradrenaline [167], 5-HT [9,180] and dopamine [191,192], which produced membrane hyperpolarization in lamina II neurons, leading to antinociception. Although oxytocin, orexins A and B produced no change or increase in L-glutamate release from nerve terminals in the lamina II [20,21,22], the spontaneous or electrically evoked release of L-glutamate from nerve terminals was inhibited by endomorphins [104,106], nociceptin [126], adenosine [142,144], noradrenaline (evoked release; [170]), 5-HT [9], cannabinoids (evoked release; [210]) and galanin (evoked release; [233]), resulting in antinociception. The idea that reduced primary-afferent central terminal L-glutamate release to lamina II neurons is involved in antinociception is supported by the observation that sEPSC frequency is increased by the activation of TRPV1 channels whose inhibitors produce antinociception when intrathecally administrated (for review, see [62,290,291]). Bradykinin, intrathecal administration of which produced nociception, increased L-glutamate release from nerve terminals and a sensitivity to L-glutamate of AMPA and NMDA receptors, both of which enhanced glutamatergic transmission in lamina II neurons [250]. Such a sensitivity increase of AMPA and NMDA receptors in lamina II neurons was produced by interleukin-1β [292] that produced a measure of nociception, i.e., wind-up (see above), when administrated intrathecally [293].
As with a GABAB-receptor agonist baclofen [27,241,294,295] and a metabolite of opioid tramadol, O-desmethyltramadol [296,297,298], clinically-used analgesics that produce antinociception by intrathecal administration generally hyperpolarize membranes and inhibit the release of L-glutamate from nerve terminals in the lamina II.
Interestingly, oxytocin, orexins A and B enhanced spontaneous inhibitory transmission [20,21,22]. As a facilitation of this transmission decreases the membrane excitability of lamina II neurons, this enhancement could account for the antinociceptive effect of oxytocin [43,44], orexins A and B [72,73,74], as suggested for antinociception produced by noradrenaline [166,168,169] and 5-HT [180,181]. Since noradrenaline and 5-HT also hyperpolarize membranes and reduce L-glutamate release from nerve terminals in the lamina II [9,167,170,180], these actions also could contribute to their antinociceptive actions together with enhanced inhibitory transmission. Consistent with the idea that GABAergic transmission enhancement is involved in antinociception produced by oxytocin, bicuculline suppressed antinociceptive responses produced by the electrical stimulation of hypothalamic paraventricular nucleus or oxytocin application [299].
On the other hand, nAChR and mAChR activation produced by ACh leads to a membrane depolarization, which in turn facilitates spontaneous inhibitory transmission in lamina II neurons, resulting in antinociception [169,288,289]. Thus, the antinociceptive mechanism of oxytocin appears to be similar to that of ACh. Lamina II neurons sensitive to oxytocin exhibited a membrane depolarization in response to (-)-nicotine and carbamoylcholine [20]. As a portion of lamina II neurons is sensitive to both oxytocin and orexin B [22], these hypothalamic neuropeptides and ACh may produce antinociception in a synergistic manner.
With respect to spontaneous inhibitory transmission, oxytocin and orexin A enhanced both GABAergic and glycinergic transmission [20,21], while orexin B enhanced glycinergic but not GABAergic transmission in the majority of lamina II neurons examined [22]. In general, both GABAergic and glycinergic transmission in lamina II neurons are modulated by many pain modulators in a similar manner (no change with DAMGO [101], nociceptin [126], somatostatin [199], galanin [233] and neuropeptide Y [261]; inhibition with adenosine [145] and anandamide [213]). On the other hand, GABAergic transmission was affected by noradrenaline in a distinct manner from that of glycinergic transmission [168] and PLA2 activation produced by melittin resulted in glycinergic but not GABAergic transmission enhancement in the presence of TTX [28,169]. 5-HT3 receptor activation in the spinal cord increased the release of GABA but not glycine, measured by the microdialysis method [182]. There are neurons exhibiting synaptically-evoked glycine but not GABA responses in the adult rat SDH [300]. It remains to be investigated why there is a difference in synaptic modulation by endogenous pain modulators between GABAergic and glycinergic transmission in lamina II neurons.
Since neuropathic or inflammatory pain can drastically change the level of endogenous neuromodulator or the function of receptor for neuromodulator, as seen in the cases of nociceptin (see [113] for review), adenosine and galanin (see Section 4.2, Section 4.3 and Section 4.10), such changes also will have to be taken into consideration in pain neuromodulators’ effects on nociceptive transmission.
Oxytocin, orexins A and B exclusively activate oxytocin, orexin-1 and orexin-2 receptors, respectively, while many of other antinociceptive modulators activate several receptor subtypes whose activation results in distinct synaptic modulation. Therefore, pain therapy by oxytocin or orexins in the spinal cord level may have an advantage over other antinociceptive modulators in activating only one kind of receptor leading to inhibitory transmission enhancement.

6. Conclusions

The present review article demonstrated that the antinociceptive effects of hypothalamic neuropeptides (oxytocin, orexins A and B) are produced by membrane depolarization and/or increased spontaneous release of L-glutamate from nerve terminals, both of which result in action potential production that leads to enhanced spontaneous inhibitory transmission by activating their specific receptors in the adult rat spinal lamina II. Such a mechanism was partly similar to those of the other analgesics in the lamina II. There will be a particular spinal dorsal horn circuitry that produces a net inhibition of the projection neurons transferring nociceptive information to the brain as a result of synaptic modulation produced by the hypothalamic neuropeptides in the lamina II. Such a circuitry remains to be revealed.
It is likely that not only the spinal lamina II but also other nervous systems are involved in the antinociceptive effects of oxytocin, orexins A and B. For instance, it is likely that oxytocin-containing hypothalamic neurons project to neurons of deep layers of the spinal cord, leading to antinociception (see [301]). This idea may possibly also apply to the case of orexins, because an abundant distribution of orexins is present in both superficial and deep layers in the lumbar segment of the rat spinal cord [69]. Alternatively, orexin A possibly acts on brain stem neurons including locus coeruleus or periaqueductal gray neurons that activate the descending pain inhibitory pathway to the spinal dorsal horn, leading to antinociception [302,303]. In future, it would be necessary to examine the involvement of neural pathways other than the hypothalamus-spinal lamina II pathway in antinociception produced by oxytocin and orexins.

Funding

My studies about oxytocin’s and orexins’ actions were supported by JSPS KAKENHI Grant Numbers 24500461 and 15K08673, respectively. No funding supported the preparation of this manuscript.

Acknowledgments

I want to thank the reviewers of this article whose many suggestions were helpful in improving it.

Conflicts of Interest

The author declares no conflict of interest.

References

  1. Rexed, B. The cytoarchitectonic organization of the spinal cord in the cat. J. Comp. Neurol. 1952, 96, 415–495. [Google Scholar] [CrossRef] [PubMed]
  2. Kumazawa, T.; Perl, E.R. Excitation of marginal and substantia gelatinosa neurons in the primate spinal cord: Indications of their place in dorsal horn functional organization. J. Comp. Neurol. 1978, 177, 417–434. [Google Scholar] [CrossRef] [PubMed]
  3. Sugiura, Y.; Lee, C.L.; Perl, E.R. Central projections of identified, unmyelinated (C) afferent fibers innervating mammalian skin. Science 1986, 234, 358–361. [Google Scholar] [CrossRef] [PubMed]
  4. Merighi, A. The histology, physiology, neurochemistry and circuitry of the substantia gelatinosa Rolandi (lamina II) in mammalian spinal cord. Prog. Neurobiol. 2018, 169, 91–134. [Google Scholar] [CrossRef] [PubMed]
  5. Willis, W.D., Jr.; Coggeshall, R.E. Sensory Mechanisms of the Spinal Cord, 2nd ed.; Plenum: New York, NY, USA, 1991. [Google Scholar]
  6. Melzack, R.; Wall, P.D. Pain mechanisms: A new theory. Science 1965, 150, 971–979. [Google Scholar] [CrossRef] [PubMed]
  7. Todd, A.J. Neuronal circuitry for pain processing in the dorsal horn. Nat. Rev. Neurosci. 2010, 11, 823–836. [Google Scholar] [CrossRef] [PubMed]
  8. Nakatsuka, T.; Park, J.-S.; Kumamoto, E.; Tamaki, T.; Yoshimura, M. Plastic changes in sensory inputs to rat substantia gelatinosa neurons following peripheral inflammation. Pain 1999, 82, 39–47. [Google Scholar] [CrossRef]
  9. Ito, A.; Kumamoto, E.; Takeda, M.; Takeda, M.; Shibata, K.; Sagai, H.; Yoshimura, M. Mechanisms for ovariectomy-induced hyperalgesia and its relief by calcitonin: Participation of 5-HT1A-like receptor on C-afferent terminals in substantia gelatinosa of the rat spinal cord. J. Neurosci. 2000, 20, 6302–6308. [Google Scholar] [CrossRef] [PubMed]
  10. Moore, K.A.; Kohno, T.; Karchewski, L.A.; Scholz, J.; Baba, H.; Woolf, C.J. Partial peripheral nerve injury promotes a selective loss of GABAergic inhibition in the superficial dorsal horn of the spinal cord. J. Neurosci. 2002, 22, 6724–6731. [Google Scholar] [CrossRef] [PubMed]
  11. Kumamoto, E. The pharmacology of amino-acid responses in septal neurons. Prog. Neurobiol. 1997, 52, 197–259. [Google Scholar] [CrossRef]
  12. Harvey, V.L.; Caley, A.; Müller, U.C.; Harvey, R.J.; Dickenson, A.H. A selective role for α3 subunit glycine receptors in inflammatory pain. Front. Mol. Neurosci. 2009, 2, 14. [Google Scholar] [CrossRef] [PubMed]
  13. Coull, J.A.M.; Boudreau, D.; Bachand, K.; Prescott, S.A.; Nault, F.; Sik, A.; de Koninck, P.; de Koninck, Y. Trans-synaptic shift in anion gradient in spinal lamina I neurons as a mechanism of neuropathic pain. Nature 2003, 424, 938–942. [Google Scholar] [CrossRef] [PubMed]
  14. Kohno, T. A role of spinal inhibition in neuropathic pain. In Cellular and Molecular Mechanisms for the Modulation of Nociceptive Transmission in the Peripheral and Central Nervous Systems; Kumamoto, E., Ed.; Research Signpost: Kelara, India, 2007; pp. 131–145. [Google Scholar]
  15. Medrano, M.C.; Dhanasobhon, D.; Yalcin, I.; Schlichter, R.; Cordero-Erausquin, M. Loss of inhibitory tone on spinal cord dorsal horn spontaneously and nonspontaneously active neurons in a mouse model of neuropathic pain. Pain 2016, 157, 1432–1442. [Google Scholar] [CrossRef] [PubMed]
  16. Yang, K.; Li, Y.-Q.; Kumamoto, E.; Furue, H.; Yoshimura, M. Voltage-clamp recordings of postsynaptic currents in substantia gelatinosa neurons in vitro and its applications to assess synaptic transmission. Brain Res. Protoc. 2001, 7, 235–240. [Google Scholar] [CrossRef]
  17. Yoshimura, M.; Nishi, S. Blind patch-clamp recordings from substantia gelatinosa neurons in adult rat spinal cord slices: Pharmacological properties of synaptic currents. Neuroscience 1993, 53, 519–526. [Google Scholar] [CrossRef]
  18. Fürst, S. Transmitters involved in antinociception in the spinal cord. Brain Res. Bull. 1999, 48, 129–141. [Google Scholar] [CrossRef]
  19. Zeilhofer, H.U.; Wildner, H.; Yévenes, G.E. Fast synaptic inhibition in spinal sensory processing and pain control. Physiol. Rev. 2012, 92, 193–235. [Google Scholar] [CrossRef]
  20. Jiang, C.-Y.; Fujita, T.; Kumamoto, E. Synaptic modulation and inward current produced by oxytocin in substantia gelatinosa neurons of adult rat spinal cord slices. J. Neurophysiol. 2014, 111, 991–1007. [Google Scholar] [CrossRef]
  21. Wang, C.; Fujita, T.; Kumamoto, E. Modulation by orexin A of spontaneous excitatory and inhibitory transmission in adult rat spinal substantia gelatinosa neurons. Biochem. Biophys. Res. Commun. 2018, 501, 100–105. [Google Scholar] [CrossRef]
  22. Wang, C.; Fujita, T.; Kumamoto, E. Orexin B modulates spontaneous excitatory and inhibitory transmission in lamina II neurons of adult rat spinal cord. Neuroscience 2018, 383, 114–128. [Google Scholar] [CrossRef]
  23. Yoshimura, M.; Nishi, S. Primary afferent-evoked glycine- and GABA-mediated IPSPs in substantia gelatinosa neurones in the rat spinal cord in vitro. J. Physiol. 1995, 482, 29–38. [Google Scholar] [CrossRef] [PubMed]
  24. Antal, M.; Petkó, M.; Polgár, E.; Heizmann, C.W.; Storm-Mathisen, J. Direct evidence of an extensive GABAergic innervation of the spinal dorsal horn by fibres descending from the rostral ventromedial medulla. Neuroscience 1996, 73, 509–518. [Google Scholar] [CrossRef]
  25. Kato, G.; Yasaka, T.; Katafuchi, T.; Furue, H.; Mizuno, M.; Iwamoto, Y.; Yoshimura, M. Direct GABAergic and glycinergic inhibition of the substantia gelatinosa from the rostral ventromedial medulla revealed by in vivo patch-clamp analysis in rats. J. Neurosci. 2006, 26, 1787–1794. [Google Scholar] [CrossRef] [PubMed]
  26. Nakatsuka, T.; Ataka, T.; Kumamoto, E.; Tamaki, T.; Yoshimura, M. Alteration in synaptic inputs through C-afferent fibers to substantia gelatinosa neurons of the rat spinal dorsal horn during postnatal development. Neuroscience 2000, 99, 549–556. [Google Scholar] [CrossRef]
  27. Ataka, T.; Kumamoto, E.; Shimoji, K.; Yoshimura, M. Baclofen inhibits more effectively C-afferent than Aδ-afferent glutamatergic transmission in substantia gelatinosa neurons of adult rat spinal cord slices. Pain 2000, 86, 273–282. [Google Scholar] [CrossRef]
  28. Liu, T.; Fujita, T.; Nakatsuka, T.; Kumamoto, E. Phospholipase A2 activation enhances inhibitory synaptic transmission in rat substantia gelatinosa neurons. J. Neurophysiol. 2008, 99, 1274–1284. [Google Scholar] [CrossRef] [PubMed]
  29. Gimpl, G.; Fahrenholz, F. The oxytocin receptor system: Structure, function, and regulation. Physiol. Rev. 2001, 81, 629–683. [Google Scholar] [CrossRef] [PubMed]
  30. Lee, H.-J.; Macbeth, A.H.; Pagani, J.H.; Young, W.S., 3rd. Oxytocin: The great facilitator of life. Prog. Neurobiol. 2009, 88, 127–151. [Google Scholar]
  31. Raggenbass, M. Vasopressin- and oxytocin-induced activity in the central nervous system: Electrophysiological studies using in-vitro systems. Prog. Neurobiol. 2001, 64, 307–326. [Google Scholar] [CrossRef]
  32. Stoop, R. Neuromodulation by oxytocin and vasopressin. Neuron 2012, 76, 142–159. [Google Scholar] [CrossRef]
  33. Cechetto, D.F.; Saper, C.B. Neurochemical organization of the hypothalamic projection to the spinal cord in the rat. J. Comp. Neurol. 1988, 272, 579–604. [Google Scholar] [CrossRef] [PubMed]
  34. Sawchenko, P.E.; Swanson, L.W. Immunohistochemical identification of neurons in the paraventricular nucleus of the hypothalamus that project to the medulla or to the spinal cord in the rat. J. Comp. Neurol. 1982, 205, 260–272. [Google Scholar] [CrossRef] [PubMed]
  35. Rousselot, P.; Papadopoulos, G.; Merighi, A.; Poulain, D.A.; Theodosis, D.T. Oxytocinergic innervation of the rat spinal cord. An electron microscopic study. Brain Res. 1990, 529, 178–184. [Google Scholar] [CrossRef]
  36. Jo, Y.-H.; Stoeckel, M.-E.; Freund-Mercier, M.-J.; Schlichter, R. Oxytocin modulates glutamatergic synaptic transmission between cultured neonatal spinal cord dorsal horn neurons. J. Neurosci. 1988, 18, 2377–2386. [Google Scholar] [CrossRef]
  37. Liu, X.; Tribollet, E.; Ogier, R.; Barberis, C.; Raggenbass, M. Presence of functional vasopressin receptors in spinal ventral horn neurons of young rats: A morphological and electrophysiological study. Eur. J. Neurosci. 2003, 17, 1833–1846. [Google Scholar] [CrossRef] [PubMed]
  38. Reiter, M.K.; Kremarik, P.; Freund-Mercier, M.J.; Stoeckel, M.E.; Desaulles, E.; Feltz, P. Localization of oxytocin binding sites in the thoracic and upper lumbar spinal cord of the adult and postnatal rat: A histoautoradiographic study. Eur. J. Neurosci. 1994, 6, 98–104. [Google Scholar] [CrossRef] [PubMed]
  39. Schorscher-Petcu, A.; Sotocinal, S.; Ciura, S.; Dupré, A.; Ritchie, J.; Sorge, R.E.; Crawley, J.N.; Hu, S.-B.; Nishimori, K.; Young, L.J.; et al. Oxytocin-induced analgesia and scratching are mediated by the vasopressin-1A receptor in the mouse. J. Neurosci. 2010, 30, 8274–8284. [Google Scholar] [CrossRef] [PubMed]
  40. Uhl-Bronner, S.; Waltisperger, E.; Martínez-Lorenzana, G.; Condés-Lara, M.; Freund-Mercier, M.J. Sexually dimorphic expression of oxytocin binding sites in forebrain and spinal cord of the rat. Neuroscience 2005, 135, 147–154. [Google Scholar] [CrossRef]
  41. Martínez-Lorenzana, G.; Espinosa-López, L.; Carranza, M.; Aramburo, C.; Paz-Tres, C.; Rojas-Piloni, G.; Condés-Lara, M. PVN electrical stimulation prolongs withdrawal latencies and releases oxytocin in cerebrospinal fluid, plasma, and spinal cord tissue in intact and neuropathic rats. Pain 2008, 140, 265–273. [Google Scholar] [CrossRef]
  42. Condés-Lara, M.; Rojas-Piloni, G.; Martínez-Lorenzana, G.; Rodríguez-Jiménez, J. Paraventricular hypothalamic oxytocinergic cells responding to noxious stimulation and projecting to the spinal dorsal horn represent a homeostatic analgesic mechanism. Eur. J. Neurosci. 2009, 30, 1056–1063. [Google Scholar] [CrossRef]
  43. Lundeberg, T.; Uvnäs-Moberg, K.; Ågren, G.; Bruzelius, G. Anti-nociceptive effects of oxytocin in rats and mice. Neurosci. Lett. 1994, 170, 153–157. [Google Scholar] [CrossRef]
  44. Yang, J.; Yang, Y.; Chen, J.-M.; Liu, W.-Y.; Wang, C.-H.; Lin, B.-C. Central oxytocin enhances antinociception in the rat. Peptides 2007, 28, 1113–1119. [Google Scholar] [CrossRef] [PubMed]
  45. Moreno-López, Y.; Martínez-Lorenzana, G.; Condés-Lara, M.; Rojas-Piloni, G. Identification of oxytocin receptor in the dorsal horn and nociceptive dorsal root ganglion neurons. Neuropeptides 2013, 47, 117–123. [Google Scholar] [CrossRef] [PubMed]
  46. Tribollet, E.; Barberis, C.; Arsenijevic, Y. Distribution of vasopressin and oxytocin receptors in the rat spinal cord: Sex-related differences and effect of castration in pudendal motor nuclei. Neuroscience 1997, 78, 499–509. [Google Scholar] [CrossRef]
  47. Wrobel, L.; Schorscher-Petcu, A.; Dupré, A.; Yoshida, M.; Nishimori, K.; Tribollet, E. Distribution and identity of neurons expressing the oxytocin receptor in the mouse spinal cord. Neurosci. Lett. 2011, 495, 49–54. [Google Scholar] [CrossRef] [PubMed]
  48. Kobrinsky, E.; Mirshahi, T.; Zhang, H.; Jin, T.; Logothetis, D.E. Receptor-mediated hydrolysis of plasma membrane messenger PIP2 leads to K+-current desensitization. Nat. Cell Biol. 2000, 2, 507–514. [Google Scholar] [CrossRef]
  49. Maruyama, T.; Kanaji, T.; Nakade, S.; Kanno, T.; Mikoshiba, K. 2APB, 2-aminoethoxydiphenyl borate, a membrane-penetrable modulator of Ins(1,4,5)P3-induced Ca2+ release. J. Biochem. 1997, 122, 498–505. [Google Scholar] [CrossRef] [PubMed]
  50. Ohta, T.; Ito, S.; Ohga, A. Inhibitory action of dantrolene on Ca-induced Ca2+ release from sarcoplasmic reticulum in guinea pig skeletal muscle. Eur. J. Pharmacol. 1990, 178, 11–19. [Google Scholar]
  51. Herbert, J.M.; Augereau, J.M.; Gleye, J.; Maffrand, J.P. Chelerythrine is a potent and specific inhibitor of protein kinase C. Biochem. Biophys. Res. Commun. 1990, 172, 993–999. [Google Scholar] [CrossRef]
  52. Brown, D.A.; Adams, P.R. Muscarinic suppression of a novel voltage-sensitive K+ current in a vertebrate neurone. Nature 1980, 283, 673–676. [Google Scholar] [CrossRef]
  53. Breton, J.-D.; Poisbeau, P.; Darbon, P. Antinociceptive action of oxytocin involves inhibition of potassium channel currents in lamina II neurons of the rat spinal cord. Mol. Pain 2009, 5, 63. [Google Scholar] [CrossRef]
  54. Chini, B.; Manning, M.; Guillon, G. Affinity and efficacy of selective agonists and antagonists for vasopressin and oxytocin receptors: An “easy guide” to receptor pharmacology. Prog. Brain Res. 2008, 170, 513–517. [Google Scholar]
  55. Breton, J.-D.; Veinante, P.; Uhl-Bronner, S.; Vergnano, A.M.; Freund-Mercier, M.J.; Schlichter, R.; Poisbeau, P. Oxytocin-induced antinociception in the spinal cord is mediated by a subpopulation of glutamatergic neurons in lamina I-II which amplify GABAergic inhibition. Mol. Pain 2008, 4, 19. [Google Scholar] [CrossRef]
  56. Jiang, C.-Y.; Fujita, T.; Kumamoto, E. Developmental change and sexual difference in synaptic modulation produced by oxytocin in rat substantia gelatinosa neurons. Biochem. Biophys. Rep. 2016, 7, 206–213. [Google Scholar] [CrossRef]
  57. Robinson, D.A.; Wei, F.; Wang, G.D.; Li, P.; Kim, S.J.; Vogt, S.K.; Muglia, L.J.; Zhuo, M. Oxytocin mediates stress-induced analgesia in adult mice. J. Physiol. 2002, 540, 593–606. [Google Scholar] [CrossRef]
  58. Gong, L.; Gao, F.; Li, J.; Li, J.; Yu, X.; Ma, X.; Zheng, W.; Cui, S.; Liu, K.; Zhang, M.; et al. Oxytocin-induced membrane hyperpolarization in pain-sensitive dorsal root ganglia neurons mediated by Ca2+/nNOS/NO/KATP pathway. Neuroscience 2015, 289, 417–428. [Google Scholar] [CrossRef]
  59. Nersesyan, Y.; Demirkhanyan, L.; Cabezas-Bratesco, D.; Oakes, V.; Kusuda, R.; Dawson, T.; Sun, X.; Cao, C.; Cohen, A.M.; Chelluboina, B.; et al. Oxytocin modulates nociception as an agonist of pain-sensing TRPV1. Cell Rep. 2017, 21, 1681–1691. [Google Scholar] [CrossRef]
  60. Yang, K.; Kumamoto, E.; Furue, H.; Yoshimura, M. Capsaicin facilitates excitatory but not inhibitory synaptic transmission in substantia gelatinosa of the rat spinal cord. Neurosci. Lett. 1998, 255, 135–138. [Google Scholar] [CrossRef]
  61. Yang, K.; Kumamoto, E.; Furue, H.; Li, Y.-Q.; Yoshimura, M. Action of capsaicin on dorsal root-evoked synaptic transmission to substantia gelatinosa neurons in adult rat spinal cord slices. Brain Res. 1999, 830, 268–273. [Google Scholar] [CrossRef]
  62. Kumamoto, E.; Fujita, T.; Jiang, C.-Y. TRP channels involved in spontaneous L-glutamate release enhancement in the adult rat spinal substantia gelatinosa. Cells 2014, 3, 331–362. [Google Scholar] [CrossRef]
  63. De Lecea, L.; Kilduff, T.S.; Peyron, C.; Gao, X.-B.; Foye, P.E.; Danielson, P.E.; Fukuhara, C.; Battenberg, E.L.F.; Gautvik, V.T.; Bartlett, F.S., II; et al. The hypocretins: Hypothalamus-specific peptides with neuroexcitatory activity. Proc. Natl. Acad. Sci. USA 1998, 95, 322–327. [Google Scholar] [CrossRef] [PubMed]
  64. Sakurai, T.; Amemiya, A.; Ishii, M.; Matsuzaki, I.; Chemelli, R.M.; Tanaka, H.; Williams, S.C.; Richardson, J.A.; Kozlowski, G.P.; Wilson, S.; et al. Orexins and orexin receptors: A family of hypothalamic neuropeptides and G protein-coupled receptors that regulate feeding behavior. Cell 1998, 92, 573–585. [Google Scholar] [CrossRef]
  65. Gao, X.-B.; Horvath, T. Function and dysfunction of hypocretin/orexin: An energetics point of view. Annu. Rev. Neurosci. 2014, 37, 101–116. [Google Scholar] [CrossRef] [PubMed]
  66. Leonard, C.S.; Kukkonen, J.P. Orexin/hypocretin receptor signalling: A functional perspective. Br. J. Pharmacol. 2014, 171, 294–313. [Google Scholar] [CrossRef] [PubMed]
  67. Sakurai, T. The neural circuit of orexin (hypocretin): Maintaining sleep and wakefulness. Nat. Rev. Neurosci. 2007, 8, 171–181. [Google Scholar] [CrossRef] [PubMed]
  68. Van den Pol, A.N. Hypothalamic hypocretin (orexin): Robust innervation of the spinal cord. J. Neurosci. 1999, 19, 3171–3182. [Google Scholar] [CrossRef] [PubMed]
  69. Date, Y.; Mondal, M.S.; Matsukura, S.; Nakazato, M. Distribution of orexin-A and orexin-B (hypocretins) in the rat spinal cord. Neurosci. Lett. 2000, 288, 87–90. [Google Scholar] [CrossRef]
  70. Cluderay, J.E.; Harrison, D.C.; Hervieu, G.J. Protein distribution of the orexin-2 receptor in the rat central nervous system. Regul. Pept. 2002, 104, 131–144. [Google Scholar] [CrossRef]
  71. Hervieu, G.J.; Cluderay, J.E.; Harrison, D.C.; Roberts, J.C.; Leslie, R.A. Gene expression and protein distribution of the orexin-1 receptor in the rat brain and spinal cord. Neuroscience 2001, 103, 777–797. [Google Scholar] [CrossRef]
  72. Cheng, J.-K.; Chou, R.C.-C.; Hwang, L.-L.; Chiou, L.-C. Antiallodynic effects of intrathecal orexins in a rat model of postoperative pain. J. Pharmacol. Exp. Ther. 2003, 307, 1065–1071. [Google Scholar] [CrossRef]
  73. Mobarakeh, J.I.; Takahashi, K.; Sakurada, S.; Nishino, S.; Watanabe, H.; Kato, M.; Yanai, K. Enhanced antinociception by intracerebroventricularly and intrathecally-administered orexin A and B (hypocretin-1 and -2) in mice. Peptides 2005, 26, 767–777. [Google Scholar] [CrossRef] [PubMed]
  74. Yamamoto, T.; Nozaki-Taguchi, N.; Chiba, T. Analgesic effect of intrathecally administered orexin-A in the rat formalin test and in the rat hot plate test. Br. J. Pharmacol. 2002, 137, 170–176. [Google Scholar] [CrossRef] [PubMed]
  75. Grudt, T.J.; Perl, E.R. Correlations between neuronal morphology and electrophysiological features in the rodent superficial dorsal horn. J. Physiol. 2002, 540, 189–207. [Google Scholar] [CrossRef] [PubMed]
  76. Lebold, T.P.; Bonaventure, P.; Shireman, B.T. Selective orexin receptor antagonists. Bioorg. Med. Chem. Lett. 2013, 23, 4761–4769. [Google Scholar] [CrossRef] [PubMed]
  77. Jeon, Y.; Park, K.B.; Pervin, R.; Kim, T.W.; Youn, D.-h. Orexin-A modulates excitatory synaptic transmission and neuronal excitability in the spinal cord substantia gelatinosa. Neurosci. Lett. 2015, 604, 128–133. [Google Scholar] [CrossRef] [PubMed]
  78. Grudt, T.J.; van den Pol, A.N.; Perl, E.R. Hypocretin-2 (orexin-B) modulation of superficial dorsal horn activity in rat. J. Physiol. 2002, 538, 517–525. [Google Scholar] [CrossRef] [PubMed]
  79. Kukkonen, J.P.; Leonard, C.S. Orexin/hypocretin receptor signalling cascades. Br. J. Pharmacol. 2014, 171, 314–331. [Google Scholar] [CrossRef] [PubMed]
  80. Zhang, J.-H.; Sampogna, S.; Morales, F.R.; Chase, M.H. Co-localization of hypocretin-1 and hypocretin-2 in the cat hypothalamus and brainstem. Peptides 2002, 23, 1479–1483. [Google Scholar] [CrossRef]
  81. Eriksson, K.S.; Sergeeva, O.; Brown, R.E.; Haas, H.L. Orexin/hypocretin excites the histaminergic neurons of the tuberomammillary nucleus. J. Neurosci. 2001, 21, 9273–9279. [Google Scholar] [CrossRef] [PubMed]
  82. Burlet, S.; Tyler, C.J.; Leonard, C.S. Direct and indirect excitation of laterodorsal tegmental neurons by hypocretin/orexin peptides: Implications for wakefulness and narcolepsy. J. Neurosci. 2002, 22, 2862–2872. [Google Scholar] [CrossRef]
  83. Kolaj, M.; Coderre, E.; Renaud, L.P. Orexin peptides enhance median preoptic nucleus neuronal excitability via postsynaptic membrane depolarization and enhancement of glutamatergic afferents. Neuroscience 2008, 155, 1212–1220. [Google Scholar] [CrossRef] [PubMed]
  84. Kim, J.; Nakajima, K.; Oomura, Y.; Wayner, M.J.; Sasaki, K. Electrophysiological effects of orexins/hypocretins on pedunculopontine tegmental neurons in rats: An in vitro study. Peptides 2009, 30, 191–209. [Google Scholar] [CrossRef] [PubMed]
  85. Huang, S.-C.; Dai, Y.-W.E.; Lee, Y.-H.; Chiou, L.-C.; Hwang, L.-L. Orexins depolarize rostral ventrolateral medulla neurons and increase arterial pressure and heart rate in rats mainly via orexin 2 receptors. J. Pharmacol. Exp. Ther. 2010, 334, 522–529. [Google Scholar] [CrossRef] [PubMed]
  86. Yamanaka, A.; Tabuchi, S.; Tsunematsu, T.; Fukazawa, Y.; Tominaga, M. Orexin directly excites orexin neurons through orexin 2 receptor. J. Neurosci. 2010, 30, 12642–12652. [Google Scholar] [CrossRef] [PubMed]
  87. Minami, M.; Satoh, M. Molecular biology of the opioid receptors: Structures, functions and distributions. Neurosci. Res. 1995, 23, 121–145. [Google Scholar] [CrossRef]
  88. Besse, D.; Lombard, M.C.; Zajac, J.M.; Roques, B.P.; Besson, J.M. Pre- and postsynaptic distribution of μ, δ and κ opioid receptors in the superficial layers of the cervical dorsal horn of the rat spinal cord. Brain Res. 1990, 521, 15–22. [Google Scholar] [CrossRef]
  89. Gouardères, C.; Beaudet, A.; Zajac, J.-M.; Cros, J.; Quirion, R. High resolution radioautographic localization of [125I]FK-33-824-labelled mu opioid receptors in the spinal cord of normal and deafferented rats. Neuroscience 1991, 43, 197–209. [Google Scholar] [CrossRef]
  90. Mansour, A.; Fox, C.A.; Akil, H.; Watson, S.J. Opioid-receptor mRNA expression in the rat CNS: Anatomical and functional implications. Trends Neurosci. 1995, 18, 22–29. [Google Scholar] [CrossRef]
  91. Gamse, R.; Holzer, P.; Lembeck, F. Indirect evidence for presynaptic location of opiate receptors on chemosensitive primary sensory neurones. Naunyn-Schmiedeberg’s Arch. Pharmacol. 1979, 308, 281–285. [Google Scholar] [CrossRef]
  92. Hunt, S.P.; Kelly, J.S.; Emson, P.C. The electron microscopic localization of methionine-enkephalin within the superficial layers (I and II) of the spinal cord. Neuroscience 1980, 5, 1871–1890. [Google Scholar] [CrossRef]
  93. Merchenthaler, I.; Maderdrut, J.L.; Altschuler, R.A.; Petrusz, P. Immunocytochemical localization of proenkephalin-derived peptides in the central nervous system of the rat. Neuroscience 1986, 17, 325–348. [Google Scholar] [CrossRef]
  94. Wu, S.Y.; Dun, S.L.; Wright, M.T.; Chang, J.-K.; Dun, N.J. Endomorphin-like immunoreactivity in the rat dorsal horn and inhibition of substantia gelatinosa neurons in vitro. Neuroscience 1999, 89, 317–321. [Google Scholar] [CrossRef]
  95. Zadina, J.E.; Hackler, L.; Ge, L.-J.; Kastin, A.J. A potent and selective endogenous agonist for the μ-opiate receptor. Nature 1997, 386, 499–502. [Google Scholar] [CrossRef] [PubMed]
  96. Horvath, G. Endomorphin-1 and endomorphin-2: Pharmacology of the selective endogenous μ-opioid receptor agonists. Pharmacol. Ther. 2000, 88, 437–463. [Google Scholar] [CrossRef]
  97. Williams, C.A.; Wu, S.Y.; Dun, S.L.; Kwok, E.H.; Dun, N.J. Release of endomorphin-2 like substances from the rat spinal cord. Neurosci. Lett. 1999, 273, 25–28. [Google Scholar] [CrossRef]
  98. Yaksh, T.L.; Rudy, T.A. Analgesia mediated by a direct spinal action of narcotics. Science 1976, 192, 1357–1358. [Google Scholar] [CrossRef] [PubMed]
  99. Cousins, M.J.; Mather, L.E. Intrathecal and epidural administration of opioids. Anesthesiology 1984, 61, 276–310. [Google Scholar] [PubMed]
  100. Duggan, A.W.; Hall, J.G.; Headley, P.M. Suppression of transmission of nociceptive impulses by morphine: Selective effects of morphine administered in the region of the substantia gelatinosa. Br. J. Pharmacol. 1977, 61, 65–76. [Google Scholar] [CrossRef] [PubMed]
  101. Kohno, T.; Kumamoto, E.; Higashi, H.; Shimoji, K.; Yoshimura, M. Actions of opioids on excitatory and inhibitory transmission in substantia gelatinosa of adult rat spinal cord. J. Physiol. 1999, 518, 803–813. [Google Scholar] [CrossRef]
  102. Ikoma, M.; Kohno, T.; Baba, H. Differential presynaptic effects of opioid agonists on Aδ- and C-afferent glutamatergic transmission to the spinal dorsal horn. Anesthesiology 2007, 107, 807–812. [Google Scholar] [CrossRef]
  103. Wrigley, P.J.; Jeong, H.-J.; Vaughan, C.W. Dissociation of μ- and δ-opioid inhibition of glutamatergic synaptic transmission in superficial dorsal horn. Mol. Pain 2010, 6, 71. [Google Scholar] [CrossRef] [PubMed]
  104. Fujita, T.; Kumamoto, E. Inhibition by endomorphin-1 and endomorphin-2 of excitatory transmission in adult rat substantia gelatinosa neurons. Neuroscience 2006, 139, 1095–1105. [Google Scholar] [CrossRef] [PubMed]
  105. Fujita, T.; Nakatsuka, T.; Kumamoto, E. Opioid receptor activation in spinal dorsal horn. In Cellular and Molecular Mechanisms for the Modulation of Nociceptive Transmission in the Peripheral and Central Nervous Systems; Kumamoto, E., Ed.; Research Signpost: Kerala, India, 2007; pp. 87–111. [Google Scholar]
  106. Yajiri, Y.; Huang, L.-Y.M. Actions of endomorphins on synaptic transmission of Aδ-fibers in spinal cord dorsal horn neurons. J. Biomed. Sci. 2000, 7, 226–231. [Google Scholar] [CrossRef] [PubMed]
  107. Yoshimura, M.; North, R.A. Substantia gelatinosa neurones hyperpolarized in vitro by enkephalin. Nature 1983, 305, 529–530. [Google Scholar] [CrossRef] [PubMed]
  108. Eckert, W.A., III; Light, A.R. Hyperpolarization of substantia gelatinosa neurons evoked by μ-, κ-, δ1-, and δ2-selective opioids. J. Pain 2002, 3, 115–125. [Google Scholar] [CrossRef] [PubMed]
  109. Wu, S.-Y.; Ohtubo, Y.; Brailoiu, G.C.; Dun, N.J. Effects of endomorphin on substantia gelatinosa neurons in rat spinal cord slices. Br. J. Pharmacol. 2003, 140, 1088–1096. [Google Scholar] [CrossRef] [PubMed]
  110. Kerchner, G.A.; Zhuo, M. Presynaptic suppression of dorsal horn inhibitory transmission by μ-opioid receptors. J. Neurophysiol. 2002, 88, 520–522. [Google Scholar] [CrossRef] [PubMed]
  111. Meunier, J.-C.; Mollereau, C.; Toll, L.; Suaudeau, C.; Moisand, C.; Alvinerie, P.; Butour, J.-L.; Guillemot, J.-C.; Ferrara, P.; Monsarrat, B.; et al. Isolation and structure of the endogenous agonist of opioid receptor-like ORL1 receptor. Nature 1995, 377, 532–535. [Google Scholar] [CrossRef]
  112. Reinscheid, R.K.; Nothacker, H.-P.; Bourson, A.; Ardati, A.; Henningsen, R.A.; Bunzow, J.R.; Grandy, D.K.; Langen, H.; Monsma, F.J., Jr.; Civelli, O. Orphanin FQ: A neuropeptide that activates an opioidlike G protein-coupled receptor. Science 1995, 270, 792–794. [Google Scholar] [CrossRef]
  113. Schröder, W.; Lambert, D.G.; Ko, M.C.; Koch, T. Functional plasticity of the N/OFQ-NOP receptor system determines analgesic properties of NOP receptor agonists. Br. J. Pharmacol. 2014, 171, 3777–3800. [Google Scholar] [CrossRef]
  114. Winters, B.L.; Christie, M.J.; Vaughan, C.W. Electrophysiological actions of N/OFQ. Handb. Exp. Pharmacol. 2019, 254, 91–130. [Google Scholar] [PubMed]
  115. Anton, B.; Fein, J.; To, T.; Li, X.; Silberstein, L.; Evans, C.J. Immunohistochemical localization of ORL-1 in the central nervous system of the rat. J. Comp. Neurol. 1996, 368, 229–251. [Google Scholar] [CrossRef]
  116. Houtani, T.; Nishi, M.; Takeshima, H.; Nukada, T.; Sugimoto, T. Structure and regional distribution of nociceptin/orphanin FQ precursor. Biochem. Biophys. Res. Commun. 1996, 219, 714–719. [Google Scholar] [CrossRef] [PubMed]
  117. Neal, C.R., Jr.; Mansour, A.; Reinscheid, R.; Nothacker, H.-P.; Civelli, O.; Watson, S.J., Jr. Localization of orphanin FQ (nociceptin) peptide and messenger RNA in the central nervous system of the rat. J. Comp. Neurol. 1999, 406, 503–547. [Google Scholar] [CrossRef]
  118. Chen, Y.; Sommer, C. Nociceptin and its receptor in rat dorsal root ganglion neurons in neuropathic and inflammatory pain models: Implications on pain processing. J. Peripher. Nerv. Syst. 2006, 11, 232–240. [Google Scholar] [CrossRef] [PubMed]
  119. Williams, C.A.; Wu, S.Y.; Cook, J.; Dun, N.J. Release of nociceptin-like substances from the rat spinal cord dorsal horn. Neurosci. Lett. 1998, 244, 141–144. [Google Scholar] [CrossRef]
  120. Erb, K.; Liebel, J.T.; Tegeder, I.; Zeilhofer, H.U.; Brune, K.; Geisslinger, G. Spinally delivered nociceptin/orphanin FQ reduces flinching behaviour in the rat formalin test. Neuroreport 1997, 8, 1967–1970. [Google Scholar] [CrossRef]
  121. Hao, J.-X.; Wiesenfeld-Hallin, Z.; Xu, X.-J. Lack of cross-tolerance between the antinociceptive effect of intrathecal orphanin FQ and morphine in the rat. Neurosci. Lett. 1997, 223, 49–52. [Google Scholar] [CrossRef]
  122. Xu, X.-J.; Hao, J.-X.; Wiesenfeld-Hallin, Z. Nociceptin or antinociceptin: Potent spinal antinociceptive effect of orphanin FQ/nociceptin in the rat. Neuroreport 1996, 7, 2092–2094. [Google Scholar]
  123. Yamamoto, T.; Nozaki-Taguchi, N.; Kimura, S. Effects of intrathecally administered nociceptin, an opioid receptor-like1 (ORL1) receptor agonist, on the thermal hyperalgesia induced by unilateral constriction injury to the sciatic nerve in the rat. Neurosci. Lett. 1997, 224, 107–110. [Google Scholar] [CrossRef]
  124. Yamamoto, T.; Nozaki-Taguchi, N.; Kimura, S. Effects of intrathecally administered nociceptin, an opioid receptor-like1 (ORL1) receptor agonist, on the thermal hyperalgesia induced by carageenan injection into the rat paw. Brain Res. 1997, 754, 329–332. [Google Scholar] [CrossRef]
  125. Luo, C.; Kumamoto, E.; Furue, H.; Yoshimura, M. Nociceptin-induced outward current in substantia gelatinosa neurones of the adult rat spinal cord. Neuroscience 2001, 108, 323–330. [Google Scholar] [CrossRef]
  126. Luo, C.; Kumamoto, E.; Furue, H.; Chen, J.; Yoshimura, M. Nociceptin inhibits excitatory but not inhibitory transmission to substantia gelatinosa neurones of adult rat spinal cord. Neuroscience 2002, 109, 349–358. [Google Scholar] [CrossRef]
  127. Ozaki, S.; Kawamoto, H.; Itoh, Y.; Miyaji, M.; Iwasawa, Y.; Ohta, H. A potent and highly selective nonpeptidyl nociceptin/orphanin FQ receptor (ORL1) antagonist: J-113397. Eur. J. Pharmacol. 2000, 387, R17–R18. [Google Scholar] [CrossRef]
  128. Lai, C.C.; Wu, S.Y.; Dun, S.L.; Dun, N.J. Nociceptin-like immunoreactivity in the rat dorsal horn and inhibition of substantia gelatinosa neurons. Neuroscience 1997, 81, 887–891. [Google Scholar] [CrossRef]
  129. Liebel, J.T.; Swandulla, D.; Zeilhofer, H.U. Modulation of excitatory synaptic transmission by nociceptin in superficial dorsal horn neurones of the neonatal rat spinal cord. Br. J. Pharmacol. 1997, 121, 425–432. [Google Scholar] [CrossRef]
  130. Zeilhofer, H.U.; Muth-Selbach, U.; Gühring, H.; Erb, K.; Ahmadi, S. Selective suppression of inhibitory synaptic transmission by nocistatin in the rat spinal cord dorsal horn. J. Neurosci. 2000, 20, 4922–4929. [Google Scholar] [CrossRef]
  131. Moran, T.D.; Abdulla, F.A.; Smith, P.A. Cellular neurophysiological actions of nociceptin/orphanin FQ. Peptides 2000, 21, 969–976. [Google Scholar] [CrossRef]
  132. Dunwiddie, T.V.; Masino, S.A. The role and regulation of adenosine in the central nervous system. Annu. Rev. Neurosci. 2001, 24, 31–55. [Google Scholar] [CrossRef]
  133. Geiger, J.D.; Labella, F.S.; Nagy, J.I. Characterization and localization of adenosine receptors in rat spinal cord. J. Neurosci. 1984, 4, 2303–2310. [Google Scholar] [CrossRef]
  134. Reppert, S.M.; Weaver, D.R.; Stehle, J.H.; Rivkees, S.A. Molecular cloning and characterization of a rat A1-adenosine receptor that is widely expressed in brain and spinal cord. Mol. Endocrinol. 1991, 5, 1037–1048. [Google Scholar] [CrossRef]
  135. Ackley, M.A.; Governo, R.J.M.; Cass, C.E.; Young, J.D.; Baldwin, S.A.; King, A.E. Control of glutamatergic neurotransmission in the rat spinal dorsal horn by the nucleoside transporter ENT1. J. Physiol. 2003, 548, 507–517. [Google Scholar] [CrossRef]
  136. Holmgren, M.; Hedner, J.; Mellstrand, T.; Nordberg, G.; Hedner, Th. Characterization of the antinociceptive effects of some adenosine analogues in the rat. Naunyn-Schmiedeberg’s Arch. Pharmacol. 1986, 334, 290–293. [Google Scholar] [CrossRef]
  137. Karlsten, R.; Gordh, T.; Hartvig, P.; Post, C. Effects of intrathecal injection of the adenosine receptor agonists R-phenylisopropyl-adenosine and N-ethylcarboxamide-adenosine on nociception and motor function in the rat. Anesth. Analg. 1990, 71, 60–64. [Google Scholar] [CrossRef]
  138. Sawynok, J.; Sweeney, M.I.; White, T.D. Classification of adenosine receptors mediating antinociception in the rat spinal cord. Br. J. Pharmacol. 1986, 88, 923–930. [Google Scholar] [CrossRef]
  139. Sawynok, J. Adenosine receptor activation and nociception. Eur. J. Pharmacol. 1998, 317, 1–11. [Google Scholar] [CrossRef]
  140. Chen, Z.; Janes, K.; Chen, C.; Doyle, T.; Bryant, L.; Tosh, D.K.; Jacobson, K.A.; Salvemini, D. Controlling murine and rat chronic pain through A3 adenosine receptor activation. FASEB J. 2012, 26, 1855–1865. [Google Scholar] [CrossRef]
  141. Terayama, R.; Tabata, M.; Maruhama, K.; Iida, S. A3 adenosine receptor agonist attenuates neuropathic pain by suppressing activation of microglia and convergence of nociceptive inputs in the spinal dorsal horn. Exp. Brain Res. 2018, 236, 3203–3213. [Google Scholar] [CrossRef]
  142. Lao, L.-J.; Kumamoto, E.; Luo, C.; Furue, H.; Yoshimura, M. Adenosine inhibits excitatory transmission to substantia gelatinosa neurons of the adult rat spinal cord through the activation of presynaptic A1 adenosine receptor. Pain 2001, 94, 315–324. [Google Scholar] [CrossRef]
  143. Liu, T.; Fujita, T.; Kawasaki, Y.; Kumamoto, E. Regulation by equilibrative nucleoside transporter of adenosine outward currents in adult rat spinal dorsal horn neurons. Brain Res. Bull. 2004, 64, 75–83. [Google Scholar] [CrossRef]
  144. Lao, L.-J.; Kawasaki, Y.; Yang, K.; Fujita, T.; Kumamoto, E. Modulation by adenosine of Aδ and C primary-afferent glutamatergic transmission in adult rat substantia gelatinosa neurons. Neuroscience 2004, 125, 221–231. [Google Scholar] [CrossRef]
  145. Yang, K.; Fujita, T.; Kumamoto, E. Adenosine inhibits GABAergic and glycinergic transmission in adult rat substantia gelatinosa neurons. J. Neurophysiol. 2004, 92, 2867–2877. [Google Scholar] [CrossRef]
  146. Kumamoto, E.; Fujita, T. Role of adenosine in regulating nociceptive transmission in the spinal dorsal horn. In Recent Research Developments in Physiology; Pandalai, S.G., Ed.; Research Signpost: Kelara, India, 2005; Volume 3, pp. 39–57. [Google Scholar]
  147. Imlach, W.L.; Bhola, R.F.; May, L.T.; Christopoulos, A.; Christie, M.J. A positive allosteric modulator of the adenosine A1 receptor selectively inhibits primary afferent synaptic transmission in a neuropathic pain model. Mol. Pharmacol. 2015, 88, 460–468. [Google Scholar] [CrossRef]
  148. Li, J.; Perl, E.R. Adenosine inhibition of synaptic transmission in the substantia gelatinosa. J. Neurophysiol. 1994, 72, 1611–1621. [Google Scholar] [CrossRef]
  149. Abbracchio, M.P.; Burnstock, G.; Boeynaems, J.-M.; Barnard, E.A.; Boyer, J.L.; Kennedy, C.; Knight, G.E.; Fumagalli, M.; Gachet, C.; Jacobson, K.A.; et al. International union of pharmacology LVIII: Update on the P2Y G protein-coupled nucleotide receptors: From molecular mechanisms and pathophysiology to therapy. Pharmacol. Rev. 2006, 58, 281–341. [Google Scholar] [CrossRef]
  150. Khakh, B.S.; North, R.A. P2X receptors as cell-surface ATP sensors in health and disease. Nature 2006, 442, 527–532. [Google Scholar] [CrossRef]
  151. Li, J.; Perl, E.R. ATP modulation of synaptic transmission in the spinal substantia gelatinosa. J. Neurosci. 1995, 15, 3357–3365. [Google Scholar] [CrossRef]
  152. Li, P.; Calejesan, A.A.; Zhuo, M. ATP P2X receptors and sensory synaptic transmission between primary afferent fibers and spinal dorsal horn neurons in rats. J. Neurophysiol. 1998, 80, 3356–3360. [Google Scholar] [CrossRef]
  153. Gu, J.G.; Bardoni, R.; Magherini, P.C.; MacDermott, A.B. Effects of the P2-purinoceptor antagonists suramin and pyridoxal-phosphate-6-azophenyl-2', 4'-disulfonic acid on glutamatergic synaptic transmission in rat dorsal horn neurons of the spinal cord. Neurosci. Lett. 1998, 253, 167–170. [Google Scholar] [CrossRef]
  154. Rhee, J.-S.; Wang, Z.-M.; Nabekura, J.; Inoue, K.; Akaike, N. ATP facilitates spontaneous glycinergic IPSC frequency at dissociated rat dorsal horn interneuron synapses. J. Physiol. 2000, 524, 471–483. [Google Scholar] [CrossRef]
  155. Bylund, D.B.; Eikenberg, D.C.; Hieble, J.P.; Langer, S.Z.; Lefkowitz, R.J.; Minneman, K.P.; Molinoff, P.B.; Ruffolo, R.R., Jr.; Trendelenburg, U. International Union of Pharmacology nomenclature of adrenoceptors. Pharmacol. Rev. 1994, 46, 121–136. [Google Scholar]
  156. Pieribone, V.A.; Nicholas, A.P.; Dagerlind, Å.; Hökfelt, T. Distribution of α1 adrenoceptors in rat brain revealed by in situ hybridization experiments utilizing subtype-specific probes. J. Neurosci. 1994, 14, 4252–4268. [Google Scholar] [CrossRef]
  157. Stone, L.S.; Broberger, C.; Vulchanova, L.; Wilcox, G.L.; Hökfelt, T.; Riedl, M.S.; Elde, R. Differential distribution of α2A and α2C adrenergic receptor immunoreactivity in the rat spinal cord. J. Neurosci. 1998, 18, 5928–5937. [Google Scholar] [CrossRef]
  158. Nicholas, A.P.; Pieribone, V.A.; Hökfelt, T. Cellular localization of messenger RNA for beta-1 and beta-2 adrenergic receptors in rat brain: An in situ hybridization study. Neuroscience 1993, 56, 1023–1039. [Google Scholar] [CrossRef]
  159. Hagihira, S.; Senba, E.; Yoshida, S.; Tohyama, M.; Yoshiya, I. Fine structure of noradrenergic terminals and their synapses in the rat spinal dorsal horn: An immunohistochemical study. Brain Res. 1990, 526, 73–80. [Google Scholar] [CrossRef]
  160. Satoh, K.; Kashiba, A.; Kimura, H.; Maeda, T. Noradrenergic axon terminals in the substantia gelatinosa of the rat spinal cord: An electron-microscopic study using glyoxylic acid-potassium permanganate fixation. Cell Tissue Res. 1982, 222, 359–378. [Google Scholar] [CrossRef]
  161. Yeomans, D.C.; Clark, F.M.; Paice, J.A.; Proudfit, H.K. Antinociception induced by electrical stimulation of spinally projecting noradrenergic neurons in the A7 catecholamine cell group of the rat. Pain 1992, 48, 449–461. [Google Scholar] [CrossRef]
  162. Yaksh, T.L. Pharmacology of spinal adrenergic systems which modulate spinal nociceptive processing. Pharmacol. Biochem. Behav. 1985, 22, 845–858. [Google Scholar] [CrossRef]
  163. Aimone, L.D.; Jones, S.L.; Gebhart, G.F. Stimulation-produced descending inhibition from the periaqueductal gray and nucleus raphe magnus in the rat: Mediation by spinal monoamines but not opioids. Pain 1987, 31, 123–136. [Google Scholar] [CrossRef]
  164. Howe, J.R.; Wang, J.-Y.; Yaksh, T.L. Selective antagonism of the antinociceptive effect of intrathecally applied alpha adrenergic agonists by intrathecal prazosin and intrathecal yohimbine. J. Pharmacol. Exp. Ther. 1983, 224, 552–558. [Google Scholar]
  165. Reddy, S.V.R.; Maderdrut, J.L.; Yaksh, T.L. Spinal cord pharmacology of adrenergic agonist-mediated antinociception. J. Pharmacol. Exp. Ther. 1980, 213, 525–533. [Google Scholar]
  166. Baba, H.; Shimoji, K.; Yoshimura, M. Norepinephrine facilitates inhibitory transmission in substantia gelatinosa of adult rat spinal cord (part 1). Effects on axon terminals of GABAergic and glycinergic neurons. Anesthesiology 2000, 92, 473–484. [Google Scholar] [CrossRef]
  167. Sonohata, M.; Furue, H.; Katafuchi, T.; Yasaka, T.; Doi, A.; Kumamoto, E.; Yoshimura, M. Actions of noradrenaline on substantia gelatinosa neurones in the rat spinal cord revealed by in vivo patch recording. J. Physiol. 2004, 555, 515–526. [Google Scholar] [CrossRef]
  168. Baba, H.; Goldstein, P.A.; Okamoto, M.; Kohno, T.; Ataka, T.; Yoshimura, M.; Shimoji, K. Norepinephrine facilitates inhibitory transmission in substantia gelatinosa of adult rat spinal cord (part 2). Effects on somatodendritic sites of GABAergic neurons. Anesthesiology 2000, 92, 485–492. [Google Scholar] [CrossRef]
  169. Liu, T.; Fujita, T.; Kumamoto, E. Acetylcholine and norepinephrine mediate GABAergic but not glycinergic transmission enhancement by melittin in adult rat substantia gelatinosa neurons. J. Neurophysiol. 2011, 106, 233–246. [Google Scholar] [CrossRef]
  170. Kawasaki, Y.; Kumamoto, E.; Furue, H.; Yoshimura, M. α2 Adrenoceptor-mediated presynaptic inhibition of primary afferent glutamatergic transmission in rat substantia gelatinosa neurons. Anesthesiology 2003, 98, 682–689. [Google Scholar] [CrossRef]
  171. Pan, Y.-Z.; Li, D.-P.; Pan, H.-L. Inhibition of glutamatergic synaptic input to spinal lamina IIo neurons by presynaptic α2-adrenergic receptors. J. Neurophysiol. 2002, 87, 1938–1947. [Google Scholar] [CrossRef]
  172. Raymond, J.R.; Mukhin, Y.V.; Gelasco, A.; Turner, J.; Collinsworth, G.; Gettys, T.W.; Grewal, J.S.; Garnovskaya, M.N. Multiplicity of mechanisms of serotonin receptor signal transduction. Pharmacol. Ther. 2001, 92, 179–212. [Google Scholar] [CrossRef]
  173. Hamon, M.; Gallissot, M.C.; Menard, F.; Gozlan, H.; Bourgoin, S.; Vergé, D. 5-HT3 receptor binding sites are on capsaicin-sensitive fibres in the rat spinal cord. Eur. J. Pharmacol. 1989, 164, 315–322. [Google Scholar] [CrossRef]
  174. Marlier, L.; Teilhac, J.-R.; Cerruti, C.; Privat, A. Autoradiographic mapping of 5-HT1, 5-HT1A, 5-HT1B and 5-HT2 receptors in the rat spinal cord. Brain Res. 1991, 550, 15–23. [Google Scholar] [CrossRef]
  175. Bowker, R.M.; Reddy, V.K.; Fung, S.J.; Chan, J.Y.H.; Barnes, C.D. Serotonergic and non-serotonergic raphe neurons projecting to the feline lumbar and cervical spinal cord: A quantitative horseradish peroxidase-immunocytochemical study. Neurosci. Lett. 1987, 75, 31–37. [Google Scholar] [CrossRef]
  176. Steinbusch, H.W.M. Distribution of serotonin-immunoreactivity in the central nervous system of the rat-cell bodies and terminals. Neuroscience 1981, 6, 557–618. [Google Scholar] [CrossRef]
  177. Sorkin, L.S.; McAdoo, D.J.; Willis, W.D. Raphe magnus stimulation-induced antinociception in the cat is associated with release of amino acids as well as serotonin in the lumbar dorsal horn. Brain Res. 1993, 618, 95–108. [Google Scholar] [CrossRef]
  178. Xu, W.; Qiu, X.-C.; Han, J.-S. Serotonin receptor subtypes in spinal antinociception in the rat. J. Pharmacol. Exp. Ther. 1994, 269, 1182–1189. [Google Scholar]
  179. Lu, Y.; Perl, E.R. Selective action of noradrenaline and serotonin on neurones of the spinal superficial dorsal horn in the rat. J. Physiol. 2007, 582, 127–136. [Google Scholar] [CrossRef]
  180. Abe, K.; Kato, G.; Katafuchi, T.; Tamae, A.; Furue, H.; Yoshimura, M. Responses to 5-HT in morphologically identified neurons in the rat substantia gelatinosa in vitro. Neuroscience 2009, 159, 316–324. [Google Scholar] [CrossRef]
  181. Fukushima, T.; Ohtsubo, T.; Tsuda, M.; Yanagawa, Y.; Hori, Y. Facilitatory actions of serotonin type 3 receptors on GABAergic inhibitory synaptic transmission in the spinal superficial dorsal horn. J. Neurophysiol. 2009, 102, 1459–1471. [Google Scholar] [CrossRef]
  182. Kawamata, T.; Omote, K.; Toriyabe, M.; Yamamoto, H.; Namiki, A. The activation of 5-HT3 receptors evokes GABA release in the spinal cord. Brain Res. 2003, 978, 250–255. [Google Scholar] [CrossRef]
  183. Missale, C.; Nash, S.R.; Robinson, S.W.; Jaber, M.; Caron, M.G. Dopamine receptors: From structure to function. Physiol. Rev. 1998, 78, 189–225. [Google Scholar] [CrossRef]
  184. Levant, B.; McCarson, K.E. D3 dopamine receptors in rat spinal cord: Implications for sensory and motor function. Neurosci. Lett. 2001, 303, 9–12. [Google Scholar] [CrossRef]
  185. Holstege, J.C.; Van Dijken, H.; Buijs, R.M.; Goedknegt, H.; Gosens, T.; Bongers, C.M.H. Distribution of dopamine immunoreactivity in the rat, cat and monkey spinal cord. J. Comp. Neurol. 1996, 376, 631–652. [Google Scholar] [CrossRef]
  186. Skagerberg, G.; Björklund, A.; Lindvall, O.; Schmidt, R.H. Origin and termination of the diencephalo-spinal dopamine system in the rat. Brain Res. Bull. 1982, 9, 237–444. [Google Scholar] [CrossRef]
  187. Ozawa, H.; Yamaguchi, T.; Hamaguchi, S.; Yamaguchi, S.; Ueda, S. Three types of A11 neurons project to the rat spinal cord. Neurochem. Res. 2017, 42, 2142–2153. [Google Scholar] [CrossRef]
  188. Swanson, L.W.; Kuypers, H.G. The paraventricular nucleus of the hypothalamus: Cytoarchitectonic subdivisions and organization of projections to the pituitary, dorsal vagal complex, and spinal cord as demonstrated by retrograde fluorescence double-labeling methods. J. Comp. Neurol. 1980, 194, 555–570. [Google Scholar] [CrossRef]
  189. Jensen, T.S.; Yaksh, T.L. Effects of an intrathecal dopamine agonist, apomorphine, on thermal and chemical evoked noxious responses in rats. Brain Res. 1984, 296, 285–293. [Google Scholar] [CrossRef]
  190. Puopolo, M. The hypothalamic-spinal dopaminergic system: A target for pain modulation. Neural Regen. Res. 2019, 14, 925–930. [Google Scholar] [CrossRef]
  191. Tamae, A.; Nakatsuka, T.; Koga, K.; Kato, G.; Furue, H.; Katafuchi, T.; Yoshimura, M. Direct inhibition of substantia gelatinosa neurones in the rat spinal cord by activation of dopamine D2-like receptors. J. Physiol. 2005, 568, 243–253. [Google Scholar] [CrossRef]
  192. Taniguchi, W.; Nakatsuka, T.; Miyazaki, N.; Yamada, H.; Takeda, D.; Fujita, T.; Kumamoto, E.; Yoshida, M. In vivo patch-clamp analysis of dopaminergic antinociceptive actions on substantia gelatinosa neurons in the spinal cord. Pain 2011, 152, 95–105. [Google Scholar] [CrossRef]
  193. Schindler, M.; Humphrey, P.P.A.; Emson, P.C. Somatostatin receptors in the central nervous system. Prog. Neurobiol. 1996, 50, 9–47. [Google Scholar] [CrossRef]
  194. Von Banchet, G.S.; Schindler, M.; Hervieu, G.J.; Beckmann, B.; Emson, P.C.; Heppelmann, B. Distribution of somatostatin receptor subtypes in rat lumbar spinal cord examined with gold-labelled somatostatin and anti-receptor antibodies. Brain Res. 1999, 816, 254–257. [Google Scholar] [CrossRef]
  195. Tuchscherer, M.M.; Seybold, V.S. Immunohistochemical studies of substance P, cholecystokinin-octapeptide and somatostatin in dorsal root ganglia of the rat. Neuroscience 1985, 14, 593–605. [Google Scholar] [CrossRef]
  196. Hökfelt, T.; Elde, R.; Johansson, O.; Luft, R.; Nilsson, G.; Arimura, A. Immunohistochemical evidence for separate populations of somatostatin-containing and substance P-containing primary afferent neurons in the rat. Neuroscience 1976, 1, 131–136. [Google Scholar] [CrossRef]
  197. Sandkühler, J.; Fu, Q.G.; Helmchen, C. Spinal somatostatin superfusion in vivo affects activity of cat nociceptive dorsal horn neurons: Comparison with spinal morphine. Neuroscience 1990, 34, 565–576. [Google Scholar] [CrossRef]
  198. Chapman, V.; Dickenson, A.H. The spinal and peripheral roles of bradykinin and prostaglandins in nociceptive processing in the rat. Eur. J. Pharmacol. 1992, 219, 427–433. [Google Scholar] [CrossRef]
  199. Jiang, N.; Furue, H.; Katafuchi, T.; Yoshimura, M. Somatostatin directly inhibits substantia gelatinosa neurons in adult rat spinal dorsal horn in vitro. Neurosci. Res. 2003, 47, 97–107. [Google Scholar] [CrossRef]
  200. Nakatsuka, T.; Fujita, T.; Inoue, K.; Kumamoto, E. Activation of GIRK channels in substantia gelatinosa neurones of the adult rat spinal cord: A possible involvement of somatostatin. J. Physiol. 2008, 586, 2511–2522. [Google Scholar] [CrossRef]
  201. Kim, S.J.; Chung, W.H.; Rhim, H.; Eun, S.-Y.; Jung, S.J.; Kim, J. Postsynaptic action mechanism of somatostatin on the membrane excitability in spinal substantia gelatinosa neurons of juvenile rats. Neuroscience 2002, 114, 1139–1148. [Google Scholar] [CrossRef]
  202. Manzanares, J.; Julian, M.D.; Carrascosa, A. Role of the cannabinoid system in pain control and therapeutic implications for the management of acute and chronic pain episodes. Curr. Neuropharmacol. 2006, 4, 239–257. [Google Scholar] [CrossRef]
  203. Pertwee, R.G. Cannabinoid receptors and pain. Prog. Neurobiol. 2001, 63, 569–611. [Google Scholar] [CrossRef]
  204. Lichtman, A.H.; Martin, B.R. Cannabinoid-induced antinociception is mediated by a spinal α2-noradrenergic mechanism. Brain Res. 1991, 559, 309–314. [Google Scholar] [CrossRef]
  205. Hohmann, A.G.; Tsou, K.; Walker, J.M. Cannabinoid modulation of wide dynamic range neurons in the lumbar dorsal horn of the rat by spinally administered WIN55,212-2. Neurosci. Lett. 1998, 257, 119–122. [Google Scholar] [CrossRef]
  206. Mailleux, P.; Vanderhaeghen, J.-J. Distribution of neuronal cannabinoid receptor in the adult rat brain: A comparative receptor binding radioautography and in situ hybridization histochemistry. Neuroscience 1992, 48, 655–668. [Google Scholar] [CrossRef]
  207. Hohmann, A.G.; Briley, E.M.; Herkenham, M. Pre- and postsynaptic distribution of cannabinoid and mu opioid receptors in rat spinal cord. Brain Res. 1999, 822, 17–25. [Google Scholar] [CrossRef]
  208. Hegyi, Z.; Kis, G.; Holló, K.; Ledent, C.; Antal, M. Neuronal and glial localization of the cannabinoid-1 receptor in the superficial spinal dorsal horn of the rodent spinal cord. Eur. J. Neurosci. 2009, 30, 251–262. [Google Scholar] [CrossRef]
  209. Chapman, V. The cannabinoid CB1 receptor antagonist, SR141716A, selectively facilitates nociceptive responses of dorsal horn neurones in the rat. Br. J. Pharmacol. 1999, 127, 1765–1767. [Google Scholar] [CrossRef]
  210. Luo, C.; Kumamoto, E.; Furue, H.; Chen, J.; Yoshimura, M. Anandamide inhibits excitatory transmission to rat substantia gelatinosa neurones in a manner different from that of capsaicin. Neurosci. Lett. 2002, 321, 17–20. [Google Scholar] [CrossRef]
  211. Zygmunt, P.M.; Petersson, J.; Andersson, D.A.; Chuang, H.-h.; Sørgård, M.; Di Marzo, V.; Julius, D.; Högestätt, E.D. Vanilloid receptors on sensory nerves mediate the vasodilator action of anandamide. Nature 1999, 400, 452–457. [Google Scholar] [CrossRef]
  212. Morisset, V.; Urban, L. Cannabinoid-induced presynaptic inhibition of glutamatergic EPSCs in substantia gelatinosa neurons of the rat spinal cord. J. Neurophysiol. 2001, 86, 40–48. [Google Scholar] [CrossRef]
  213. Kawasaki, Y.; Fujita, T.; Yang, K.; Kumamoto, E. Anandamide depresses glycinergic and GABAergic inhibitory transmissions in adult rat substantia gelatinosa neurons. Pharmacol. Pharm. 2015, 6, 103–117. [Google Scholar] [CrossRef]
  214. Sugiura, T.; Kondo, S.; Sukagawa, A.; Nakane, S.; Shinoda, A.; Itoh, K.; Yamashita, A.; Waku, K. 2-Arachidonoylglycerol: A possible endogenous cannabinoid receptor ligand in brain. Biochem. Biophys. Res. Commun. 1995, 215, 89–97. [Google Scholar] [CrossRef]
  215. Tatemoto, K.; Rökaeus, Å.; Jörnvall, H.; McDonald, T.J.; Mutt, V. Galanin—A novel biologically active peptide from porcine intestine. FEBS Lett. 1983, 164, 124–128. [Google Scholar] [CrossRef]
  216. Lang, R.; Gundlach, A.L.; Kofler, B. The galanin peptide family: Receptor pharmacology, pleiotropic biological actions, and implications in health and disease. Pharmacol. Ther. 2007, 115, 177–207. [Google Scholar] [CrossRef]
  217. Hökfelt, T. Galanin and its receptors: Introduction to the Third International Symposium, San Diego, California, USA, 21–22 October 2004. Neuropeptides 2005, 39, 125–142. [Google Scholar] [CrossRef]
  218. Liu, H.-X.; Hökfelt, T. The participation of galanin in pain processing at the spinal level. Trends Pharmacol. Sci. 2002, 23, 468–474. [Google Scholar] [CrossRef]
  219. Brumovsky, P.; Mennicken, F.; O’Donnell, D.; Hökfelt, T. Differential distribution and regulation of galanin receptors- 1 and -2 in the rat lumbar spinal cord. Brain Res. 2006, 1085, 111–120. [Google Scholar] [CrossRef]
  220. Ch’ng, J.L.C.; Christofides, N.D.; Anand, P.; Gibson, S.J.; Allen, Y.S.; Su, H.C.; Tatemoto, K.; Morrison, J.F.B.; Polak, J.M.; Bloom, S.R. Distribution of galanin immunoreactivity in the central nervous system and the responses of galanin-containing neuronal pathways to injury. Neuroscience 1985, 16, 343–354. [Google Scholar] [CrossRef]
  221. Kerekes, N.; Mennicken, F.; O’Donnell, D.; Hökfelt, T.; Hill, R.H. Galanin increases membrane excitability and enhances Ca2+ currents in adult, acutely dissociated dorsal root ganglion neurons. Eur. J. Neurosci. 2003, 18, 2957–2966. [Google Scholar] [CrossRef]
  222. Landry, M.; Bouali-Benazzouz, R.; André, C.; Shi, T.J.S.; Léger, C.; Nagy, F.; Hökfelt, T. Galanin receptor 1 is expressed in a subpopulation of glutamatergic interneurons in the dorsal horn of the rat spinal cord. J. Comp. Neurol. 2006, 499, 391–403. [Google Scholar] [CrossRef]
  223. O’Donnell, D.; Ahmad, S.; Wahlestedt, C.; Walker, P. Expression of the novel galanin receptor subtype GALR2 in the adult rat CNS: Distinct distribution from GALR1. J. Comp. Neurol. 1999, 409, 469–481. [Google Scholar] [CrossRef]
  224. Skofitsch, G.; Jacobowitz, D.M. Galanin-like immunoreactivity in capsaicin sensitive sensory neurons and ganglia. Brain Res. Bull. 1985, 15, 191–195. [Google Scholar] [CrossRef]
  225. Waters, S.M.; Krause, J.E. Distribution of galanin-1, -2 and -3 receptor messenger RNAs in central and peripheral rat tissues. Neuroscience 2000, 95, 265–271. [Google Scholar] [CrossRef]
  226. Ji, R.-R.; Zhang, X.; Zhang, Q.; Dagerlind, Å.; Nilsson, S.; Wiesenfeld-Hallin, Z.; Hökfelt, T. Central and peripheral expression of galanin in response to inflammation. Neuroscience 1995, 68, 563–576. [Google Scholar] [CrossRef]
  227. Cridland, R.A.; Henry, J.L. Effects of intrathecal administration of neuropeptides on a spinal nociceptive reflex in the rat: VIP, galanin, CGRP, TRH, somatostatin and angiotensin II. Neuropeptides 1988, 11, 23–32. [Google Scholar] [CrossRef]
  228. Kuraishi, Y.; Kawamura, M.; Yamaguchi, T.; Houtani, T.; Kawabata, S.; Futaki, S.; Fujii, N.; Satoh, M. Intrathecal injections of galanin and its antiserum affect nociceptive response of rat to mechanical, but not thermal, stimuli. Pain 1991, 44, 321–324. [Google Scholar] [CrossRef]
  229. Liu, H.-X.; Brumovsky, P.; Schmidt, R.; Brown, W.; Payza, K.; Hodzic, L.; Pou, C.; Godbout, C.; Hökfelt, T. Receptor subtype-specific pronociceptive and analgesic actions of galanin in the spinal cord: Selective actions via GalR1 and GalR2 receptors. Proc. Natl. Acad. Sci. USA 2001, 98, 9960–9964. [Google Scholar] [CrossRef] [PubMed]
  230. Wiesenfeld-Hallin, Z.; Villar, M.J.; Hökfelt, T. The effects of intrathecal galanin and C-fiber stimulation on the flexor reflex in the rat. Brain Res. 1989, 486, 205–213. [Google Scholar] [CrossRef]
  231. Holmes, F.E.; Bacon, A.; Pope, R.J.P.; Vanderplank, P.A.; Kerr, N.C.H.; Sukumaran, M.; Pachnis, V.; Wynick, D. Transgenic overexpression of galanin in the dorsal root ganglia modulates pain-related behavior. Proc. Natl. Acad. Sci. USA 2003, 100, 6180–6185. [Google Scholar] [CrossRef]
  232. Lundström, L.; Sollenberg, U.; Brewer, A.; Kouya, P.F.; Zheng, K.; Xu, X.-J.; Sheng, X.; Robinson, J.K.; Wiesenfeld-Hallin, Z.; Xu, Z.-Q.; et al. A galanin receptor subtype 1 specific agonist. Int. J. Peptide Res. Ther. 2005, 11, 17–27. [Google Scholar] [CrossRef]
  233. Yue, H.-Y.; Fujita, T.; Kumamoto, E. Biphasic modulation by galanin of excitatory synaptic transmission in substantia gelatinosa neurons of adult rat spinal cord slices. J. Neurophysiol. 2011, 105, 2337–2349. [Google Scholar] [CrossRef]
  234. Alier, K.A.; Chen, Y.; Sollenberg, U.E.; Langel, Ü.; Smith, P.A. Selective stimulation of GalR1 and GalR2 in rat substantia gelatinosa reveals a cellular basis for the anti- and pro-nociceptive actions of galanin. Pain 2008, 137, 138–146. [Google Scholar] [CrossRef]
  235. Khawaja, A.M.; Rogers, D.F. Tachykinins: Receptor to effector. Int. J. Biochem. Cell. Biol. 1996, 28, 721–738. [Google Scholar] [CrossRef]
  236. Urbán, L.; Randic, M. Slow excitatory transmission in rat dorsal horn: Possible mediation by peptides. Brain Res. 1984, 290, 336–341. [Google Scholar] [CrossRef]
  237. Dickenson, A.H.; Sullivan, A.F. Evidence for a role of the NMDA receptor in the frequency dependent potentiation of deep rat dorsal horn nociceptive neurones following C fibre stimulation. Neuropharmacology 1987, 26, 1235–1238. [Google Scholar] [CrossRef]
  238. Mendell, L.M. Physiological properties of unmyelinated fiber projection to the spinal cord. Exp. Neurol. 1966, 16, 316–332. [Google Scholar] [CrossRef]
  239. Xu, X.-J.; Dalsgaard, C.-J.; Wiesenfeld-Hallin, Z. Spinal substance P and N-methyl-D-aspartate receptors are coactivated in the induction of central sensitization of the nociceptive flexor reflex. Neuroscience 1992, 51, 641–648. [Google Scholar] [CrossRef]
  240. Barber, R.P.; Vaughn, J.E.; Slemmon, J.R.; Salvaterra, P.M.; Roberts, E.; Leeman, S.E. The origin, distribution and synaptic relationships of substance P axons in rat spinal cord. J. Comp. Neurol. 1979, 184, 331–351. [Google Scholar] [CrossRef]
  241. Yang, K.; Kumamoto, E.; Furue, H.; Li, Y.-Q.; Yoshimura, M. Capsaicin induces a slow inward current which is not mediated by substance P in substantia gelatinosa neurons of the rat spinal cord. Neuropharmacology 2000, 39, 2185–2194. [Google Scholar] [CrossRef]
  242. Bleazard, L.; Hill, R.G.; Morris, R. NK1 receptor and the actions of tachykinin agonists in the dorsal horn of the rat indicates that substance P does not have a functional role on substantia gelatinosa (lamina II) neurons. J. Neurosci. 1994, 14, 7655–7664. [Google Scholar] [CrossRef]
  243. Urbán, L.; Dray, A. Synaptic activation of dorsal horn neurons by selective C-fibre excitation with capsaicin in the mouse spinal cord in vitro. Neuroscience 1992, 47, 693–702. [Google Scholar] [CrossRef]
  244. Ikeda, H.; Heinke, B.; Ruscheweyh, R.; Sandkühler, J. Synaptic plasticity in spinal lamina I projection neurons that mediate hyperalgesia. Science 2003, 299, 1237–1240. [Google Scholar] [CrossRef]
  245. Prado, G.N.; Taylor, L.; Zhou, X.; Ricupero, D.; Mierke, D.F.; Polgar, P. Mechanisms regulating the expression, self-maintenance, and signaling-function of the bradykinin B2 and B1 receptors. J. Cell. Physiol. 2002, 193, 275–286. [Google Scholar] [CrossRef]
  246. Rueff, A.; Dray, A. Sensitization of peripheral afferent fibres in the in vitro neonatal rat spinal cord-tail by bradykinin and prostaglandins. Neuroscience 1993, 54, 527–535. [Google Scholar] [CrossRef]
  247. Couture, R.; Harrisson, M.; Vianna, R.M.; Cloutier, F. Kinin receptors in pain and inflammation. Eur. J. Pharmacol. 2001, 429, 161–176. [Google Scholar] [CrossRef]
  248. Chapman, V.; Dickenson, A.H. The effects of sandostatin and somatostatin on nociceptive transmission in the dorsal horn of the rat spinal cord. Neuropeptides 1992, 23, 147–152. [Google Scholar] [CrossRef]
  249. Ferreira, J.; Campos, M.M.; Araújo, R.; Bader, M.; Pesquero, J.B.; Calixto, J.B. The use of kinin B1 and B2 receptor knockout mice and selective antagonists to characterize the nociceptive responses caused by kinins at the spinal level. Neuropharmacology 2002, 43, 1188–1197. [Google Scholar] [CrossRef]
  250. Wang, H.; Kohno, T.; Amaya, F.; Brenner, G.J.; Ito, N.; Allchorne, A.; Ji, R.-R.; Woolf, C.J. Bradykinin produces pain hypersensitivity by potentiating spinal cord glutamatergic synaptic transmission. J. Neurosci. 2005, 25, 7986–7992. [Google Scholar] [CrossRef]
  251. Tatemoto, K.; Carlquist, M.; Mutt, V. Neuropeptide Y—A novel brain peptide with structural similarities to peptide YY and pancreatic polypeptide. Nature 1982, 296, 659–660. [Google Scholar] [CrossRef]
  252. Taiwo, O.B.; Taylor, B.K. Antihyperalgesic effects of intrathecal neuropeptide Y during inflammation are mediated by Y1 receptors. Pain 2002, 96, 353–363. [Google Scholar] [CrossRef]
  253. Zhang, Y.-X.; Lundeberg, T.; Yu, L.-C. Involvement of neuropeptide Y and Y1 receptor in antinociception in nucleus raphe magnus of rats. Regul. Pept. 2000, 95, 109–113. [Google Scholar] [CrossRef]
  254. Hökfelt, T.; Broberger, C.; Zhang, X.; Diez, M.; Kopp, J.; Xu, Z.-Q.; Landry, M.; Bao, L.; Schalling, M.; Koistinaho, J.; et al. Neuropeptide Y: Some viewpoints on a multifaceted peptide in the normal and diseased nervous system. Brain Res. Brain Res. Rev. 1998, 26, 154–166. [Google Scholar] [CrossRef]
  255. Larhammar, D.; Blomqvist, A.G.; Söderberg, C. Evolution of neuropeptide Y and its related peptides. Comp. Biochem. Physiol. C 1993, 106, 743–752. [Google Scholar] [CrossRef]
  256. Wakisaka, S.; Kajander, K.C.; Bennett, G.J. Increased neuropeptide Y (NPY)-like immunoreactivity in rat sensory neurons following peripheral axotomy. Neurosci. Lett. 1991, 124, 200–203. [Google Scholar] [CrossRef]
  257. Zhang, X.; Bao, L.; Xu, Z.-Q.; Kopp, J.; Arvidsson, U.; Elde, R.; Hökfelt, T. Localization of neuropeptide Y Y1 receptors in the rat nervous system with special reference to somatic receptors on small dorsal root ganglion neurons. Proc. Natl. Acad. Sci. USA 1994, 91, 11738–11742. [Google Scholar] [CrossRef] [PubMed]
  258. Zhang, X.; Shi, T.; Holmberg, K.; Landry, M.; Huang, W.; Xiao, H.; Ju, G.; Hökfelt, T. Expression and regulation of the neuropeptide Y Y2 receptor in sensory and autonomic ganglia. Proc. Natl. Acad. Sci. USA 1997, 94, 729–734. [Google Scholar] [CrossRef] [PubMed]
  259. Hua, X.Y.; Boublik, J.H.; Spicer, M.A.; Rivier, J.E.; Brown, M.R.; Yaksh, T.L. The antinociceptive effects of spinally administered neuropeptide Y in the rat: Systematic studies on structure-activity relationship. J. Pharmacol. Exp. Ther. 1991, 258, 243–248. [Google Scholar] [PubMed]
  260. Naveilhan, P.; Hassani, H.; Lucas, G.; Blakeman, K.H.; Hao, J.-X.; Xu, X.-J.; Wiesenfeld-Hallin, Z.; Thorén, P.; Ernfors, P. Reduced antinociception and plasma extravasation in mice lacking a neuropeptide Y receptor. Nature 2001, 409, 513–517. [Google Scholar] [CrossRef] [PubMed]
  261. Miyakawa, A.; Furue, H.; Katafuchi, T.; Jiang, N.; Yasaka, T.; Kato, G.; Yoshimura, M. Action of neuropeptide Y on nociceptive transmission in substantia gelatinosa of the adult rat spinal dorsal horn. Neuroscience 2005, 134, 595–604. [Google Scholar] [CrossRef] [PubMed]
  262. Dennis, E.A. Diversity of group types, regulation, and function of phospholipase A2. J. Biol. Chem. 1994, 269, 13057–13060. [Google Scholar]
  263. Shimizu, T.; Wolfe, L.S. Arachidonic acid cascade and signal transduction. J. Neurochem. 1990, 55, 1–15. [Google Scholar] [CrossRef]
  264. Katsuki, H.; Okuda, S. Arachidonic acid as a neurotoxic and neurotrophic substance. Prog. Neurobiol. 1995, 46, 607–636. [Google Scholar] [CrossRef]
  265. Samad, T.A.; Sapirstein, A.; Woolf, C.J. Prostanoids and pain: Unraveling mechanisms and revealing therapeutic targets. Trends Mol. Med. 2002, 8, 390–396. [Google Scholar] [CrossRef]
  266. Svensson, C.I.; Yaksh, T.L. The spinal phospholipase-cyclooxygenase-prostanoid cascade in nociceptive processing. Annu. Rev. Pharmacol. Toxicol. 2002, 42, 553–583. [Google Scholar] [CrossRef] [PubMed]
  267. Habermann, E. Bee and wasp venoms. Science 1972, 177, 314–322. [Google Scholar] [CrossRef] [PubMed]
  268. Clark, M.A.; Conway, T.M.; Shorr, R.G.L.; Crooke, S.T. Identification and isolation of a mammalian protein which is antigenically and functionally related to the phospholipase A2 stimulatory peptide melittin. J. Biol. Chem. 1987, 262, 4402–4406. [Google Scholar] [PubMed]
  269. Hassid, A.; Levine, L. Stimulation of phospholipase activity and prostaglandin biosynthesis by melittin in cell culture and in vivo. Res. Commun. Chem. Pathol. Pharmacol. 1977, 18, 507–517. [Google Scholar] [PubMed]
  270. Steiner, M.R.; Bomalaski, J.S.; Clark, M.A. Responses of purified phospholipases A2 to phospholipase A2 activating protein (PLAP) and melittin. Biochim. Biophys. Acta 1993, 1166, 124–130. [Google Scholar] [CrossRef]
  271. Mayer, R.J.; Marshall, L.A. New insights on mammalian phospholipase A2(s); comparison of arachidonoyl-selective and -nonselective enzymes. FASEB J. 1993, 7, 339–348. [Google Scholar] [CrossRef]
  272. Yue, H.-Y.; Fujita, T.; Kumamoto, E. Phospholipase A2 activation by melittin enhances spontaneous glutamatergic excitatory transmission in rat substantia gelatinosa neurons. Neuroscience 2005, 135, 485–495. [Google Scholar] [CrossRef] [PubMed]
  273. Vishwanath, B.S.; Kini, R.M.; Gowda, T.V. Characterization of three edema-inducing phospholipase A2 enzymes from habu (Trimeresurus flavoviridis) venom and their interaction with the alkaloid aristolochic acid. Toxicon 1987, 25, 501–515. [Google Scholar] [CrossRef]
  274. Kumamoto, E.; Liu, T.; Fujita, T.; Yue, H.-Y.; Nakatsuka, T. Role of phospholipase A2 in modulating synaptic transmission in the spinal dorsal horn. In Cellular and Molecular Mechanisms for the Modulation of Nociceptive Transmission in the Peripheral and Central Nervous Systems; Kumamoto, E., Ed.; Research Signpost: Kelara, India, 2007; pp. 113–130. [Google Scholar]
  275. Baba, H.; Kohno, T.; Moore, K.A.; Woolf, C.J. Direct activation of rat spinal dorsal horn neurons by prostaglandin E2. J. Neurosci. 2001, 21, 1750–1756. [Google Scholar] [CrossRef]
  276. Minami, T.; Okuda-Ashitaka, E.; Hori, Y.; Sakuma, S.; Sugimoto, T.; Sakimura, K.; Mishina, M.; Ito, S. Involvement of primary afferent C-fibres in touch-evoked pain (allodynia) induced by prostaglandin E2. Eur. J. Neurosci. 1999, 11, 1849–1856. [Google Scholar] [CrossRef] [PubMed]
  277. Dani, J.A. Overview of nicotinic receptors and their roles in the central nervous system. Biol. Psychiatry 2001, 49, 166–174. [Google Scholar] [CrossRef]
  278. Eglen, R.M.; Choppin, A.; Watson, N. Therapeutic opportunities from muscarinic receptor research. Trends Pharmacol. Sci. 2001, 22, 409–414. [Google Scholar] [CrossRef]
  279. Wada, E.; Wada, K.; Boulter, J.; Deneris, E.; Heinemann, S.; Patrick, J.; Swanson, L.W. Distribution of alpha2, alpha3, alpha4, and beta2 neuronal nicotinic receptor subunit mRNAs in the central nervous system: A hybridization histochemical study in the rat. J. Comp. Neurol. 1989, 284, 314–335. [Google Scholar] [CrossRef] [PubMed]
  280. Wada, E.; McKinnon, D.; Heinemann, S.; Patrick, J.; Swanson, L.W. The distribution of mRNA encoded by a new member of the neuronal nicotinic acetylcholine receptor gene family (α5) in the rat central nervous system. J. Physiol. 1990, 526, 45–53. [Google Scholar] [CrossRef]
  281. Yamamura, H.I.; Wamsley, J.K.; Deshmukh, P.; Roeske, W.R. Differential light microscopic autoradiographic localization of muscarinic cholinergic receptors in the brainstem and spinal cord of the rat using [3H]pirenzepine. Eur. J. Pharmacol. 1983, 91, 147–149. [Google Scholar] [CrossRef]
  282. Ribeiro-da-Silva, A.; Cuello, A.C. Choline acetyltransferase-immunoreactive profiles are presynaptic to primary sensory fibers in the rat superficial dorsal horn. J. Comp. Neurol. 1990, 295, 370–384. [Google Scholar] [CrossRef] [PubMed]
  283. Todd, A.J. Immunohistochemical evidence that acetylcholine and glycine exist in different populations of GABAergic neurons in lamina III of rat spinal dorsal horn. Neuroscience 1991, 44, 741–746. [Google Scholar] [CrossRef]
  284. Abram, S.E.; O’Connor, T.C. Characteristics of the analgesic effects and drug interactions of intrathecal carbachol in rats. Anesthesiology 1995, 83, 844–849. [Google Scholar] [CrossRef] [PubMed]
  285. Abram, S.E.; Winne, R.P. Intrathecal acetyl cholinesterase inhibitors produce analgesia that is synergistic with morphine and clonidine in rats. Anesth. Analg. 1995, 81, 501–507. [Google Scholar] [PubMed]
  286. Khan, I.M.; Buerkle, H.; Taylor, P.; Yaksh, T.L. Nociceptive and antinociceptive responses to intrathecally administered nicotinic agonists. Neuropharmacology 1998, 37, 1515–1525. [Google Scholar] [CrossRef]
  287. Khan, I.M.; Stanislaus, S.; Zhang, L.; Taylor, P.; Yaksh, T.L. A-85380 and epibatidine each interact with disparate spinal nicotinic receptor subtypes to achieve analgesia and nociception. J. Pharmacol. Exp. Ther. 2001, 297, 230–239. [Google Scholar] [PubMed]
  288. Baba, H.; Kohno, T.; Okamoto, M.; Goldstein, P.A.; Shimoji, K.; Yoshimura, M. Muscarinic facilitation of GABA release in substantia gelatinosa of the rat spinal dorsal horn. J. Physiol. 1998, 508, 83–93. [Google Scholar] [CrossRef]
  289. Takeda, D.; Nakatsuka, T.; Papke, R.; Gu, J.G. Modulation of inhibitory synaptic activity by a non-α4β2, non-α7 subtype of nicotinic receptors in the substantia gelatinosa of adult rat spinal cord. Pain 2003, 101, 13–23. [Google Scholar] [CrossRef]
  290. Kumamoto, E.; Fujita, T. Differential activation of TRP channels in the adult rat spinal substantia gelatinosa by stereoisomers of plant-derived chemicals. Pharmaceuticals 2016, 9, 46. [Google Scholar] [CrossRef]
  291. Kumamoto, E. Effects of plant-derived compounds on excitatory synaptic transmission and nerve conduction in the nervous system—Involvement in pain modulation. Curr. Top. Phytochem. 2018, 14, 45–70. [Google Scholar]
  292. Liu, T.; Jiang, C.-Y.; Fujita, T.; Luo, S.-W.; Kumamoto, E. Enhancement by interleukin-1β of AMPA and NMDA receptor-mediated currents in adult rat spinal superficial dorsal horn neurons. Mol. Pain 2013, 9, 16. [Google Scholar] [CrossRef]
  293. Arriagada, O.; Constandil, L.; Hernández, A.; Barra, R.; Soto-Moyano, R.; Laurido, C. Effects of interleukin-1β on spinal cord nociceptive transmission in intact and propentofylline-treated rats. Int. J. Neurosci. 2007, 117, 617–625. [Google Scholar] [CrossRef]
  294. Iyadomi, M.; Iyadomi, I.; Kumamoto, E.; Tomokuni, K.; Yoshimura, M. Presynaptic inhibition by baclofen of miniature EPSCs and IPSCs in substantia gelatinosa neurons of the adult rat spinal dorsal horn. Pain 2000, 85, 385–393. [Google Scholar] [CrossRef]
  295. Kangrga, I.; Jiang, M.C.; Randić, M. Actions of (-)-baclofen on rat dorsal horn neurons. Brain Res. 1991, 562, 265–275. [Google Scholar] [CrossRef]
  296. Koga, A.; Fujita, T.; Totoki, T.; Kumamoto, E. Tramadol produces outward currents by activating μ-opioid receptors in adult rat substantia gelatinosa neurones. Br. J. Pharmacol. 2005, 145, 602–607. [Google Scholar] [CrossRef]
  297. Koga, A.; Fujita, T.; Piao, L.-H.; Nakatsuka, T.; Kumamoto, E. Inhibition by O-desmethyltramadol of glutamatergic excitatory transmission in adult rat spinal substantia gelatinosa neurons. Mol. Pain 2019, 15, 1744806918824243. [Google Scholar] [CrossRef]
  298. Yamasaki, H.; Funai, Y.; Funao, T.; Mori, T.; Nishikawa, K. Effects of tramadol on substantia gelatinosa neurons in the rat spinal cord: An in vivo patch-clamp analysis. PLoS ONE 2015, 10, e0125147. [Google Scholar] [CrossRef]
  299. Condés-Lara, M.; Rojas-Piloni, G.; Martínez-Lorenzana, G.; López-Hidalgo, M.; Rodríguez-Jiménez, J. Hypothalamospinal oxytocinergic antinociception is mediated by GABAergic and opiate neurons that reduce A-delta and C fiber primary afferent excitation of spinal cord cells. Brain Res. 2009, 1247, 38–49. [Google Scholar] [CrossRef]
  300. Keller, A.F.; Coull, J.A.M.; Chéry, N.; Poisbeau, P.; De Koninck, Y. Region-specific developmental specialization of GABA-glycine cosynapses in laminas I-II of the rat spinal dorsal horn. J. Neurosci. 2001, 21, 7871–7880. [Google Scholar] [CrossRef]
  301. Eliava, M.; Melchior, M.; Knobloch-Bollmann, H.S.; Wahis, J.; da Silva Gouveia, M.; Tang, Y.; Ciobanu, A.C.; Triana del Rio, R.; Roth, L.C.; Althammer, F.; et al. A new population of parvocellular oxytocin neurons controlling magnocellular neuron activity and inflammatory pain processing. Neuron 2016, 89, 1291–1304. [Google Scholar] [CrossRef]
  302. Hagan, J.J.; Leslie, R.A.; Patel, S.; Evans, M.L.; Wattam, T.A.; Holmes, S.; Benham, C.D.; Taylor, S.G.; Routledge, C.; Hemmati, P.; et al. Orexin A activates locus coeruleus cell firing and increases arousal in the rat. Proc. Natl. Acad. Sci. USA 1999, 96, 10911–10916. [Google Scholar] [CrossRef]
  303. Ho, Y.-C.; Lee, H.-J.; Tung, L.-W.; Liao, Y.-Y.; Fu, S.-Y.; Teng, S.-F.; Liao, H.-T.; Mackie, K.; Chiou, L.-C. Activation of orexin 1 receptors in the periaqueductal gray of male rats leads to antinociception via retrograde endocannabinoid (2-arachidonoylglycerol)-induced disinhibition. J. Neurosci. 2011, 31, 14600–14610. [Google Scholar] [CrossRef]
Table 1. Comparison of synaptic modulation produced by oxytocin, orexins A and B with those of other endogenous pain neuromodulators in rodent spinal lamina II neurons.
Table 1. Comparison of synaptic modulation produced by oxytocin, orexins A and B with those of other endogenous pain neuromodulators in rodent spinal lamina II neurons.
Endogenous Neuromodulators Resting Membrane PotentialGlutamatergic Excitatory TransmissionGABAergic Spontaneous Inhibitory TransmissionGlycinergic Spontaneous Inhibitory TransmissionReferences
Oxytocin *1DepolarizationNo changeFacilitation
(sensitive to TTX)
Facilitation
(sensitive to TTX)
[20]
Orexin A *1DepolarizationFacilitationFacilitation
(sensitive to TTX)
Facilitation
(sensitive to TTX)
[21]
Orexin B *1DepolarizationFacilitationNo changeFacilitation
(sensitive to TTX)
[22]
Endomorphins *1HyperpolarizationDepressionNo changeNo change[104,106]
Nociceptin *1HyperpolarizationDepressionNo changeNo change[125,126]
Adenosine *1HyperpolarizationDepressionDepressionDepression[142,143,144,145]
ATPFast depolarizationFacilitationFacilitation[151,154]
Noradrenaline *1HyperpolarizationNo change
(spontaneous)
Depression
(evoked)
FacilitationFacilitation[166,167,168,170]
Serotonin (5-HT) *1HyperpolarizationDepolarizationDepressionFacilitationFacilitation[9,180,181]
Dopamine *1HyperpolarizationNo change[191,192]
Somatostatin *1HyperpolarizationNo changeNo changeNo change[199,200]
Cannabinoids *1No changeNo change
(spontaneous)
Depression
(evoked)
DepressionDepression[210,213]
Galanin *2HyperpolarizationFacilitation
(spontaneous)
Depression
(evoked )
No changeNo change[233]
Substance P *3No changeNo change[241]
Bradykinin *3No changeFacilitation[250]
Neuropeptide Y *1HyperpolarizationNo changeNo changeNo change[261]
Phospholipase A2
activator
No changeFacilitationFacilitation
(sensitive to TTX)
Facilitation
(resistant to TTX)
[28,169,272]
Acetylcholine
(nicotinic) *1
DepolarizationNo changeFacilitationFacilitation[20,289]
Acetylcholine
(muscarinic) *1
DepolarizationNo changeFacilitationFacilitation[28,288]
Here, when neurons responsive to neuromodulators exhibit several effects, the main effect of them is shown. *1: neuromodulator that produces antinociception; *2: production of both antinociception and pronociception in a manner dependent on its concentration; *3: neuromodulator that produces pronociception (see the text for its detail). GABA: γ-aminobutyric acid; TTX: tetrodotoxin; ATP: adenosine 5’-triphosphate; −: data are not available, to my knowledge.

© 2019 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).
Back to TopTop