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NEUROPHARMACOLOGY
Department of Anesthesiology, Pennsylvania State University College of Medicine, Milton S. Hershey Medical Center, Hershey, Pennsylvania
Received November 1, 2004; accepted January 5, 2005.
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Abstract |
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; Naguib and Yaksh, 1994; Hood et al., 1997; Ma et al., 2001; Duttaroy et al., 2002). The analgesic effect is mediated by muscarinic receptors because the muscarinic antagonist atropine blocks the antinociceptive effects produced by both muscarinic agonists or acetylcholinesterase inhibitors (Zhuo and Gebhart, 1991; Naguib and Yaksh, 1994). Receptor autoradiography and immunocytochemistry studies have shown that the highest density of muscarinic receptors in the spinal cord is located in the superficial lamina in both rats and humans (Yamamura et al., 1983; Scatton et al., 1984; Villiger and Faull, 1985; Li et al., 2002). Molecular cloning studies have revealed five molecularly distinct muscarinic acetylcholine receptors referred to as M1 to M5 (Caulfield, 1993; Wess, 1996). The five muscarinic receptor subtypes are all linked to different types of G proteins. The odd-numbered muscarinic subtypes (M1, M3, and M5) are selectively linked to Gq/11 proteins, whereas the even-numbered subtypes (M2 and M4) are preferentially coupled to the pertussis toxin (PTX)-sensitive Gi/o family (Felder, 1995; Wess, 1996; Caulfield and Birdsall, 1998). Although previous studies have shown that M2,M3, and M4 muscarinic receptor subtypes are present in the spinal cord dorsal horn (Hoglund and Baghdoyan, 1997; Yung and Lo, 1997; Duttaroy et al., 2002), the possible physiological functions of these individual subtypes are not fully understood.
GABA is the most important inhibitory neurotransmitter in the mammalian central nervous system. Stimulation of spinal GABA release and GABAB receptors is an important analgesic mechanism of muscarinic agonists in spinal dorsal horn (Baba et al., 1998; Li et al., 2002; Chen and Pan, 2003). The spinal lamina II neurons are under the tonic control of glutamatergic excitatory and GABAergic/glycinergic inhibitory inputs. Glutamate released from primary afferents is a major excitatory neurotransmitter, which conveys nociceptive information to neurons in the superficial lamina (Yoshimura and Jessell, 1990). Muscarinic receptor activation increases GABA release in spinal dorsal horn (Baba et al., 1998). We have shown that acetylcholine can inhibit glutamate release, and inhibition of glutamate release is at least partly mediated by spinal GABA release (Li et al., 2002). Also, muscarinic agents inhibit dorsal horn projection neurons through GABAB receptors (Chen and Pan, 2004). However, the subtypes of muscarinic receptors in potentiation of synaptic GABA release in the spinal cord are not clear. In the present study, we used different pharmacological interventions to determine possible muscarinic receptor subtypes that are involved in the spinal cord GABA release.
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Materials and Methods |
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Intrathecal Treatment with PTX. To determine the involvement of M2 and M4 subtypes in the stimulatory effect of oxotremorine-M on synaptic GABA release, a group of rats was pretreated with intrathecal PTX to inactivate inhibitory Gi/o proteins (Wong et al., 1992; Fields and Casey, 1997; Chen and Pan, 2004). Intrathecal catheters were inserted in rats anesthetized using 2% halothane. The catheters (polyethylene-10 tubing) were inserted through an incision in the cisternal membrane and advanced 4.5 cm caudal so that the tip of each catheter was positioned at the lumbar spinal level. Rats were injected with intrathecal 2 µg of PTX 5 to 7 days before the final electrophysiology experiments.
Spinal Cord Slice Preparations. Rats were anesthetized with 2% halothane in O2, and the lumbar segment of the spinal cord was rapidly removed through a limited laminectomy. The rats then were killed by inhalation of 5% halothane. The segment of lumbar spinal cord was immediately placed in an ice-cold sucrose artificial cerebrospinal fluid (aCSF) presaturated with 95% O2 and 5% CO2. The sucrose aCSF contained: 234 mM sucrose, 3.6 mM KCl, 1.2 mM MgCl2, 2.5 mM CaCl2, 1.2 mM NaH2PO4, 12.0 mM glucose, and 25.0 mM NaHCO3. The tissue was then placed in a shallow groove formed in a gelatin block and glued on the stage of a vibratome (Technical Product International, St. Louis, MO). Transverse spinal cord slices (350 µm) were cut in the ice-cold sucrose aCSF and then preincubated in Krebs' solution oxygenated with 95% O2 and 5% CO2 at 34°C for at least 1 h before they were transferred to the recording chamber. The Krebs' solution contained: 117.0 mM NaCl, 3.6 mM KCl, 1.2 mM MgCl2, 2.5 mM CaCl2, 1.2 mM NaH2PO4, 11.0 mM glucose, and 25.0 mM NaHCO3. Recordings of postsynaptic currents were performed using the whole-cell voltage-clamp method, as we described previously (Li et al., 2002; Pan et al., 2002). The lamina II has a distinct translucent appearance and can easily be distinguished under the microscope. In this study, efforts were made to record neurons in the outer zone of lamina II (Li et al., 2002; Pan and Pan, 2004). The neurons located in the dorsal one-third of lamina II in the spinal slice were identified under a fixed stage microscope (BX50WI; Olympus, Tokyo, Japan) with differential interference contrast/infrared illumination. The electrode for the whole-cell recordings was triple pulled from borosilicate glass capillaries with a puller (P-97; Sutter Instrument Company, Novato, CA). The impedance of the pipette was 4 to 7 M
when filled with internal solution containing: 110 mM Cs2SO4, 5 mM KCl, 2.0 mM MgCl2, 0.5 mM CaCl2, 5.0 mM HEPES, 5.0 mM EGTA, 5.0 mM ATP-Mg, 0.5 mM Na-GTP, and 1 mM guanosine 5'-O-(2-thiodiphosphate) (GDP-
-S), and 10 mM lidocaine N-ethyl bromide (QX314), adjusted to pH 7.2 to 7.4 with 1 M CsOH (290–320 mOsm). GDP--S was added to the internal solution to block the possible postsynaptic effect mediated by muscarinic agonists through G proteins (Li et al., 2002; Pan et al., 2002). QX314 was added to the internal solution to suppress the action potential generation from the recorded cell. The slice was placed in a glass-bottomed chamber (Warner Instrument, Hamden, CT) and fixed with parallel nylon threads supported by a U-shaped stainless steel weight. The slice was continuously perfused with Krebs' solution at 5.0 ml/min at 34°C maintained by an inline solution heater and a temperature controller (TC-324; Warner Instrument).
Electrophysiological Recordings. Recordings of postsynaptic currents began 5 min later, after whole-cell access was established and the current reached a steady state. The input resistance was monitored, and the recording was abandoned if it changed more than 15% (Li et al., 2002; Pan et al., 2002). Signals were recorded using an amplifier (MultiClamp700A; Axon Instruments Inc., Union City, CA) at a holding potential of 0 mV, filtered at 1 to 2 kHz, digitized at 10 kHz, and stored in Pentium computer with pCLAMP 9.0 (Axon Instruments Inc.). All spontaneous inhibitory postsynaptic currents (sIPSCs) were recorded in the presence of 2 µM strychnine, a glycine receptor antagonist, and 20 µM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX). To record the miniature inhibitory postsynaptic currents (mIPSCs), 1 µM tetrodotoxin (TTX) was added to the perfusion solution.
Oxotremorine-M, himbacine, MT-3 toxin, 4-diphenylacetoxy-N-methylpiperidine methiodide (4-DAMP), GDP--S, atropine, strychnine, CNQX, and bicuculline were obtained from Sigma-Aldrich (St. Louis, MO). PTX was purchased from List Biological Laboratories Inc. (Campbell, CA). TTX and QX314 were obtained from Alomone Labs (Jerusalem, Israel). Drugs were dissolved in Krebs' solution and perfused into the slice chamber using syringe pumps.
Data Analysis. Data are presented as means ± S.E. The sIPSCs and mIPSCs were analyzed off-line with a peak detection program (MiniAnalysis; Synaptosoft, Decatur, GA). Measurements of the amplitude and frequency of sIPSCs and mIPSCs were performed over a period of at least 1 min during control, drug application, and recovery. For each analysis, 300 to 1500 events were included. The mIPSCs and sIPSCs were detected by the fast rise time of the signal over an amplitude threshold above the background noise. The amplitude detection threshold was typically 5 to 6 pA. We manually excluded the event when the noise was erroneously identified as the mIPSCs or sIPSCs by the program. The background noise level was typically constant throughout the recording of a single neuron. The cumulative probability of the amplitude and interevent interval of mIPSCs/sIPSCs was compared using the Kolmogorov-Smirnov test, which estimates the probability that two cumulative distributions are similar. The effects of oxotremorine-M on the amplitude and frequency of sIPSCs and mIPSCs were determined by paired two-tailed Student's t test or one-way analysis of variance. Neurons were considered to be responsive to oxotremorine-M if the frequency of sIPSCs was altered >20%. P < 0.05 was considered to be statistically significant.
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Results |
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Effect of Oxotremorine-M on sIPSCs and mIPSCs. The baseline frequency of sIPSCs ranged from 0.41 to 3.07 Hz (1.47 ± 0.23 pA), whereas the amplitude of sIPSCs during control was 12.31 ± 0.87 pA (from 8.15–22.47 pA; n = 16). To determine the concentration-dependent effect of oxotremorine-M on sIPSCs, 1, 3, 5 and 10 µM oxotremorine-M, a general specific muscarinic receptor agonist, was perfused in an accumulative fashion to the tissue chamber. Each concentration was applied for a duration of 3 min. In a total of 20 lamina II neurons, oxotremorine-M at the concentration of 1 µM had no significant effect on sIPSCs. Oxotremorine-M increased sIPSCs in a concentration-dependent manner (Fig. 1). At 3 µM, oxotremorine-M significantly increased the frequency (ranging from 1.86–7.33 Hz; 5.58 ± 0.43 pA) of sIPSCs in all 16 neurons examined. Among the 16 neurons tested, 3 µM oxotremorine-M also significantly increased the amplitude of sIPSCs from 12.87 ± 1.56 to 17.13 ± 2.12 pA in eight cells. In another eight neurons, 3 µM oxotremorine-M did not significantly increase the amplitude of sIPSCs (11.75 ± 0.83 to 11.42 ± 1.07 pA). The sIPSCs were completely blocked by 20 µM bicuculline, indicating that they were mediated by GABAA receptors (Fig. 1A).
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The stimulatory effect of 3 µM oxotremorine-M on sIPSCs was completely blocked by the specific muscarinic antagonist atropine (2 µM, n = 9; Fig. 2, A and B). Atropine was applied 3 to 4 min before applying the agonist. In six separate lamina II neurons, we determined whether application of the same concentration of oxotremorine-M produces a reproducible effect on sIPSCs. The initial effect of oxotremorine-M on sIPSCs can be completely washed out in 10 min. Repeat application of 3 µM oxotremorine-M caused a similar increase in the amplitude and frequency of sIPSCs (Fig. 2C).
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In another 10 lamina II neurons, the effect of 3 µM oxotremorine-M on mIPSCs was tested in the presence of 1 µM TTX. Oxotremorine-M did not significantly alter the frequency and amplitude of mIPSCs in all 10 cells tested (Fig. 3).
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Effect of Oxotremorine-M on sIPSCs in PTX-Treated Rats. To determine the contribution of M2 and M4 subtypes to the effect of oxotremorine-M on synaptic GABA release to lamina II neurons, rats were pretreated with intrathecal 2 µg of PTX to inactivate inhibitory Gi/o proteins 5 to 7 days before final electrophysiology experiments. In five rats pretreated with PTX, a total of 27 lamina II neurons were tested. Oxotremorine-M (1–10 µM) had no significant effect on the frequency of sIPSCs in 19 of 27 cells (Fig. 4). Furthermore, although 3 to 10 µM oxotremorine-M significantly increased the frequency of sIPSCs in a concentration-dependent manner in the remaining eight neurons (Fig. 4), its potentiating effect was significantly reduced compared with that in untreated rats. Oxotremorine-M only slightly increased the amplitude (from 12.92 ± 0.73 to 18.58 ± 1.36 pA, P > 0.05) of sIPSCs in only three of eight neurons tested.
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In another four rats pretreated with PTX, we determined whether the remaining effect of oxotremorine-M on sIPSCs is mediated by the M3 subtype in the spinal cord. In 12 of 28 lamina II neurons tested, 3 µM oxotremorine-M had no significant effect on both the frequency and amplitude of sIPSCs. In the remaining 16 neurons, 3 µM oxotremorine-M significantly increased the frequency of sIPSCs (P < 0.05; Fig. 5). Among these 16 cells tested, oxotremorine-M significantly increased the amplitude (from 11.09 ± 0.63 to 17.76 ± 2.84 pA) of sIPSCs in five neurons. This effect was completely blocked by subsequent application of 25 nM 4-DAMP (Fig. 5), a relatively selective M3 receptor antagonist (Ehlert, 1996; Yigit et al., 2003). In the preliminary study, we found that 25 nM 4-DAMP is the lowest concentration that completely blocked the stimulatory effect of 3 µM oxotremorine-M on sIPSCs of lamina II cells in M2/M4 double-knockout mice (H. M. Zhang and H. L. Pan, unpublished data).
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Effect of Himbacine on the Stimulatory Action of Oxotremorine-M on sIPSCs. To further determine the role of M2 and M4 receptor subtypes in the effect of oxotremorine-M on sIPSCs, a relatively selective M2 and M4 antagonist, himbacine (Dorje et al., 1991; Miller et al., 1992; Doller et al., 1999) was used. Himbacine was applied 3 to 4 min before applying the agonist. In 15 of 30 (50%) cells tested, the effect of 3 µM oxotremorine-M on the frequency of sIPSCs was completely blocked by 1 µM himbacine (Fig. 6C). In another 15 (50%) cells, the effect of 3 µM oxotremorine-M on the frequency of sIPSCs was significantly attenuated by 1 µM himbacine (Fig. 6, A and D). In the above 30 neurons tested, 3 µM oxotremorine-M significantly increased the amplitude (from 13.74 ± 0.82 to 25.69 ± 2.19 pA) of sIPSCs in 13 cells. Following application of 1 µM himbacine, oxotremorine-M increased the amplitude (from 12.47 ± 1.17 to 16.97 ± 1.45 pA, P < 0.5) of sIPSCs in 7 of 13 cells examined. Furthermore, we determined whether the remaining effect of oxotremorine-M on sIPSCs in the presence of 1 µM himbacine is mediated by the M3 subtype in some cells. In all six lamina II neurons tested, the effect of oxotremorine-M on sIPSCs was blocked completely by subsequent application of 25 nM 4-DAMP (Fig. 6, A and D).
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Effect of MT-3 Toxin on the Stimulatory Action of Oxotremorine-M on sIPSCs. To further delineate the role of the M4 receptor subtype in the effect of oxotremorine-M on sIPSCs, MT-3 toxin, a selective M4 receptor antagonist (Jolkkonen et al., 1994; Ellis et al., 1999), was used. MT-3 toxin (100 nM) was perfused for 4 min followed by 3 µM oxotremorine-M. In 12 of 14 cells tested, MT-3 significantly reduced the effect of 3 µM oxotremorine-M on the frequency of sIPSCs (Fig. 7, A and B). Among these 12 cells, the unblocked effect of oxotremorine-M was completely abolished by further perfusion of 1 µM himbacine in six (50%) cells (Fig. 7C). In another six (50%) cells, the stimulatory effect of oxotremorine-M on the frequency of sIPSCs was only partially attenuated by 1 µM himbacine but was completely eliminated by further application of 25 nM 4-DAMP (Fig. 7D). In the remaining 2 of 14 cells tested, the effect of 3 µM oxotremorine-M on the frequency of sIPSCs was completely blocked by 100 nM MT-3 (from 1.08–1.01 Hz in one cell and from 1.46–1.55 Hz in another cell). Among the 14 cells studied above, 3 µM oxotremorine-M significantly increased the amplitude of sIPSCs from 16.98 ± 2.61 to 25.08 ± 3.99 pA in 10 neurons. Following application of 100 nM MT-3, 3 µM oxotremorine-M only significantly increased the amplitude of sIPSCs from 13.76 ± 1.42 to 19.59 ± 3.17 pA in 4 of 10 neurons.
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Discussion |
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Although the spinal muscarinic receptors in antinociception are well established, the subtypes mediating this effect remain unclear. Previous behavioral studies suggest that muscarinic analgesia is mediated through M1 and M2 receptors (Iwamoto and Marion, 1993) or M1 and M3 receptors in the spinal cord (Naguib and Yaksh, 1997). However, the highly M1-selective agonist xanomeline does not produce any analgesic effect (Sheardown et al., 1997). Using muscarinic subtype knockout mice, it has been shown that the M2 and M4 subtypes are responsible for the analgesic effect of muscarinic receptor agonists (Gomeza et al., 1999; Duttaroy et al., 2002). Furthermore, radioligand binding experiments fail to demonstrate the presence of the M1 receptor in the rat spinal cord (Hoglund and Baghdoyan, 1997). Thus, only M2, M3, and M4 muscarinic receptor subtypes have been documented in spinal cord dorsal horn, and the M2 and M3 subtypes are particularly concentrated in the superficial laminae of the spinal cord (Hoglund and Baghdoyan, 1997; Yung and Lo, 1997; Li et al., 2002). Nevertheless, the functional roles of these three muscarinic receptor subtypes in synaptic transmission and muscarinic analgesia remain to be established.
The potential cellular mechanisms of muscarinic analgesia in the spinal cord have been shown in recent studies. For example, carbachol can increase the GABA release to spinal lamina II neurons (Baba et al., 1998). Glutamate released from primary afferents to spinal dorsal horn neurons is important for transmission of nociceptive information (Yoshimura and Jessell, 1990; Pan and Pan, 2004). The excitatory inputs from the primary afferents to spinal dorsal horn neurons are tonically controlled by GABAergic interneurons (Barber et al., 1978; Lu and Perl, 2003; Pan and Pan, 2004). We have shown that acetylcholine inhibits spinal glutamate release, and this inhibitory action is partially mediated by GABA release and activation of presynaptic GABAB receptors in the spinal cord (Li et al., 2002). Furthermore, the antinociceptive effect produced by intrathecal administration of muscarinic agonists is significantly attenuated by the specific GABAB receptor antagonist CGP55845(Li et al., 2002). We also found that muscarinic inhibition of dorsal horn projection neurons is mediated by GABAB receptors, and this effect is through PTX-sensitive Gi/o proteins (Chen and Pan, 2004). Thus, M2 and M4 are the most important subtypes for the inhibitory effect of muscarinic agents on nociception and dorsal horn neurons. In the present study, oxotremorine-M significantly increased the frequency of sIPSCs but not mIPSCs, indicating that oxotremorine-M-induced GABA release is through activation of muscarinic receptors located at somatodendritic sites but not on the presynaptic terminals of GABAergic interneurons in the spinal cord. To determine the role of M2 and M4 subtypes in oxotremorine-M-induced synaptic GABA release, we examined if inactivation of Gi/o proteins with intrathecal pretreatment of PTX altered the effect of oxotremorine-M on sIPSCs. In rats pretreated with PTX, 56% of lamina II neurons failed to respond to oxotremorine-M. Also, in the remaining 44% of neurons, the effect of oxotremorine-M on the frequency of sIPSCs was significantly attenuated compared with that in untreated rats. Furthermore, the relatively selective M2 and M4 subtype antagonist himbacine (Miller et al., 1992; Doller et al., 1999) produced a similar inhibitory effect on oxotremorine-M-induced increase in GABAergic sIPSCs of lamina II neurons. These data are consistent with the behavioral data about the critical role of M2 and M4 receptors in muscarinic analgesia using M2/M4 subtype knockout mice (Duttaroy et al., 2002). Therefore, the spinal M2 and M4 subtypes play a predominant role in increased GABAergic input to spinal dorsal horn neurons by muscarinic agonists.
Although the M2 receptor appears to be the most predominant subtype in the spinal cord (Yung and Lo, 1997; Duttaroy et al., 2002), there is a small but functionally significant M4 subtype in the spinal cord (Hoglund and Baghdoyan, 1997; Ellis et al., 1999; Duttaroy et al., 2002; Mulugeta et al., 2003). Also, the spinal cord M4 receptors have been shown to be important in the muscarinic analgesia in mice (Ellis et al., 1999). We used MT-3 toxin to further assess the contribution of the M4 subtype to muscarinic agonist-induced synaptic GABA release to spinal dorsal horn neurons. MT-3 has a 40-fold selectivity for the M4 subtype over the M1 subtype and a greater than 500-fold selectivity for the M4 receptor over M2,M3, and M5 receptors (Jolkkonen et al., 1994; Liang et al., 1996). We found that MT-3 significantly reduced the effect of oxotremorine-M on sIPSCs in most cells tested. This suggests that the M4 subtype plays an important role in muscarinic agonist-induced increase in the GABAergic tone in the spinal cord. The signaling mechanisms underlying the potentiating effect of M2 and M4 subtypes on synaptic GABA release in the spinal cord are not clear. The stimulatory action of M2 and M4 muscarinic subtypes on synaptic GABA release is probably due to their coupling to specific types of inhibitory Gi proteins and associated adenylyl cyclase isozymes. For example, activation of the M4 muscarinic receptor inhibits the activity of adenylyl cyclase isozymes I, V, VI, and VIII, but stimulates the activity of adenylyl cyclase isozymes II, IV, and VII (Nevo et al., 1998). It has been shown that the M4 receptor could stimulate cAMP production through activation of adenylyl cyclase II that is coupled to Gi1 or Gi2 proteins (Liu et al., 1999). Therefore, modulation of GABAergic transmission by M2 and M4 receptors is different in the brain and spinal cord.
Another important finding of this study is that the M3 receptor subtype also plays a role in potentiation of GABAergic inputs to spinal dorsal horn neurons by the muscarinic agonist. The presence of M3 receptor subtype in the rat spinal cord has been shown in a radioligand binding study (Hoglund and Baghdoyan, 1997). In the present study, we found that oxotremorine-M still increased sIPSCs in 42% of lamina II neurons in rats pretreated with intrathecal PTX. This effect was completely blocked by the relatively selective M3 antagonist 4-DAMP (Felder, 1995; Ehlert, 1996; Yigit et al., 2003). Similarly, the potentiating effect of oxotremorine-M on GABAergic sIPSCs in the presence of himbacine was also abolished by 4-DAMP. Thus, these data strongly suggest that the M3 subtype also plays a significant role in increased GABAergic inputs to the spinal dorsal horn neurons produced by muscarinic agonists. Further studies are warranted to elucidate the specific role of the spinal M3 subtype in muscarinic analgesia.
In summary, we determined the muscarinic receptor subtypes in mediating the increased spinal GABAergic inputs to rat dorsal horn neurons in the present study. We found that PTX pretreatment and the M2 and M4 receptor antagonist himbacine blocked most of the effect of oxotremorine-M on sIPSCs in the spinal cord. The remaining effect of oxotremorine-M on sIPSCs was abolished by an M3 subtype antagonist 4-DAMP. This converging evidence strongly suggests that M2 and M4 as well as M3 receptor subtypes are all involved in the augmentation of synaptic GABA release to spinal dorsal horn neurons by muscarinic agonists. Therefore, these three muscarinic receptor subtypes likely act synergistically in potentiation of the spinal inhibitory tone and mediate the antinociceptive actions of muscarinic receptor agonists. This new information is important for our understanding of the pharmacological mechanisms underlying muscarinic analgesia in the spinal cord. It should be acknowledged that GABA and glycine are often coreleased to most but not all lamina II neurons (Pan and Pan, 2004). We did not study glycinergic IPSCs in this particular work because strychnine was used to isolate GABAergic IPSCs throughout the study. The effect of muscarinic modulation of spinal glycine release and its implication in the analgesic action of muscarinic agonists should be systematically investigated in future studies.
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Footnotes |
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ABBREVIATIONS: PTX, pertussis toxin; aCSF, artificial cerebrospinal fluid; QX314, lidocaine N-ethyl bromide; GDP--S, guanosine 5'-O-(2-thiodiphosphate); sIPSC, spontaneous inhibitory postsynaptic current; CNQX, 6-cyano-7-nitroquinoxaline-2,3-dione; mIPSC, miniature inhibitory postsynaptic current; TTX, tetrodotoxin; 4-DAMP, 4-diphenylacetoxy-N-methylpiperidine methiodide; CGP55845 (3-N-[1-(5)-3,4-dichlorophenyl)-ethyl]-amino-2-(s)-hydroxypropyl)-benzyl-phosphinic acid.
Address correspondence to: Dr. Hui-Lin Pan, Department of Anesthesiology, H187, Pennsylvania State University College of Medicine, 500 University Drive, Hershey, PA 17033. E-mail: hpan{at}psu.edu
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