Neuropathic pain results from damage to, or dysfunction of, the peripheral or central nervous system. Thermal hyperalgesia and tactile allodynia are well-documented symptoms of neuropathic pain in both the clinical setting and in animal models. Although neuropathic pain responds to opioids, the results are unsatisfactory and adjuvant drugs are often needed. Naloxone is demonstrated to have dual effects. At classic doses, naloxone causes antagonism of opioid effects, whereas at ultra-low doses, it enhances the antinociceptive effect of morphine.1,2 In the present study, we tried to elucidate the mechanism of the augmentation by ultra-low dose naloxone of morphine antinociception in partial sciatic nerve–transected (PST) rats.
Neuropathic pain is associated with enhanced synaptic excitatory signal transduction in the spinal cord dorsal horn, which may explain the thermal hyperalgesia and tactile allodynia produced.3,4 Glutamate is the major excitatory neurotransmitter in the superficial dorsal horn of the spinal cord5,6 and its concentration in the synaptic cleft depends on the dynamics of glutamate release from presynaptic nerve terminals and clearance by glutamate transporters, present on both glial cells and neurons.6–10 Changes in glutamate transporter expression and regulation of synaptic glutamate homeostasis in the spinal cord have critical roles in both the induction and maintenance of neuropathic pain.11
The use of opioids for neuropathic pain is often discouraged because of their ineffectiveness, potential for tolerance induction, risk of addiction, and limiting side effects.12 Nerve injuries alter opioid receptor function and the subsequent intracellular signaling cascade, which reduce the antinociceptive potency of morphine on neuropathic pain.13 At high doses (microgram range), naloxone inhibits the analgesic effect of morphine, but at ultra-low doses, it enhances the analgesic effect of morphine by blocking excitatory opioid receptor signaling in dorsal root ganglion neurons, inhibiting enkephalin release, and inhibiting neuroinflammation in microglia.1,2,14,15 Nonetheless, there is controversy about the clinical relevance of the proposed mechanisms of naloxone function.16–18 PST in rats provides a peripheral nerve injury neuropathic pain model for assessing the clinical relevance of intrathecal treatment of neuropathic pain. The primary objective of this study was to examine the effects of ultra-low dose naloxone plus morphine treatment on the antinociceptive response and on the expression of glutamate transporters in the PST rat spinal cord dorsal horn, because glutamate transporter expression might have therapeutic implications, especially in the regulation of excitatory synaptic transmission by maintaining the homeostasis of extracellular glutamate levels.
This study protocol was reviewed and approved by the Animal Care and Use Committee of the National Defense Medical Center and conformed to the guidelines of the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publication No. 80-23, revised 1996). Male Wistar rats weighing 280 to 320 g were housed individually with soft bedding on a 12-hour night/day cycle with free access to food and water at all times. All efforts were made to minimize the number of animals used and their suffering.
Intrathecal Catheterization and Establishment of the Neuropathic Pain Animal Model
Under 2% to 2.5% isoflurane anesthesia, rats were implanted with an intrathecal catheter and, in some cases, a microdialysis probe.19,20 The intrathecal catheter and microdialysis probe were constructed as described previously.21 The rat was placed prone in a stereotaxic frame and the cisternal membrane was exposed. A polyethylene catheter (PE-10 tubing, 8.0 cm) or a microdialysis probe was inserted through a small puncture in the membrane and threaded caudally to reach the lumbar enlargement of the spinal cord. The rostral end of the catheter was exteriorized at the top of the head and the wound was closed with sutures. All rats then underwent either PST or sham operation.22 In the PST rats, the left sciatic nerve was exposed at the midthigh level and a Prolene 7-0 (Ethicon, Somerville, NJ) ligature was placed through the midpoint of the nerve just cranially to the branch running to the musculus biceps femoris, resulting in half of the nerve being transected in the ventrocranial direction up to the ligature. In the sham-operated rats, the nerve was exposed, and then the wound was closed with sutures. Thermal and tactile hypersensitivity were tested in separate rats on days 1, 4, 7, 10, 14, 18, and 21 after PST. Rats displaying any sign of motor deficit were excluded from the study; in addition, approximately 10% of PST rats did not develop neuropathic pain and were also excluded. On day 10 after PST, 15 ng naloxone (group n), 15 μg naloxone (group N), 10 μg morphine (group M), 15 ng naloxone plus 10 μg morphine (group Mn), 15 μg naloxone plus 10 μg morphine (group MN), or saline (group s) was injected via the intrathecal catheter in a 10-μL volume, followed by 10 μL of saline to flush the catheter. After drug administration, the rats were randomly assigned to undergo thermal or tactile hypersensitivity testing or cerebrospinal fluid (CSF) dialysate collection.
In the inhibitor study, 10 μg DL-threo-β-benzyloxyaspartic acid (DL-TBOA) was injected intrathecally into the PST rats 40 minutes after intrathecal coadministration of 15 ng naloxone plus 10 μg morphine; then, the tactile paw withdrawal threshold was examined at 30-minute intervals for 120 minutes and at 3-minute intervals from 40 to 60 minutes after intrathecal DL-TBOA injection.
Behavior Test for Tactile Allodynia
The sensitivity of the plantar surface to touch was assessed using an automated von Frey Dynamic Plantar Anesthesiometer (Ugo Basile, Comerio, Italy), which does not produce tissue damage.23,24 According to the Kyoto protocol of the International Association for the Study of Pain, Basic Pain Terminology, the dynamic plantar anesthesiometer produces non-noxious tactile stimuli.25 Each rat was placed in an individual plastic cage (25 cm long × 10 cm wide × 14 cm high) with a wire mesh floor and acclimatized to the cage for 15 minutes before each test session, and then a paw withdrawal response was elicited by applying an increasing force using a blunt-end metal filament (0.5 mm in diameter) focused on the middle of the plantar surface of the hindpaw. The force was initially below the detection threshold, then was increased from 1 to 50 g in 1-g steps over 20 seconds, then held at 50 g for a further 10 seconds. The rate of force increase was 2.5 g/s. The threshold was recorded as the force eliciting the hindpaw removal reflex, which was recorded as the mean of 3 measurements at 1-minute intervals.
Behavior Test for Thermal Hyperalgesia
Heat sensitivity was examined using a Hargreaves radiant heat apparatus (7371; Ugo Basile; infrared setting, 80). The rat was placed in a plastic cage (23 cm long × 18 cm wide × 14 cm high) and allowed to acclimatize to the cage for 30 minutes before the behavioral test. The cage was then placed on a glass plate above the plantar test apparatus and a movable noxious heat source placed directly under the plantar surface of the hindpaw. When activated, the apparatus applied a continuous infrared heat stimulus to the plantar surface and a distinctive paw withdrawal reflex was elicited, which stopped an automated timer (based on infrared reflection). The hindpaws were tested alternately, with a 5-minute interval between consecutive tests. Three measurements of latency were averaged for each hindpaw in each test session. The baseline threshold was between 8 and 10 seconds in normal rats and a 22-second cutoff time was used to prevent tissue damage.
We calculated the area under the curve (AUC) to measure the analgesic effect of treatment. The AUC for the paw withdrawal threshold versus time was calculated by trapezoidal approximation over the total observation period (180 minutes). The AUC for the cutoff (22 seconds in the thermal plantar test, 50 g in the dynamic plantar anesthesiometer test) minus the AUC for the intrathecal saline group (group s) was averaged and defined as 100%. The AUC for the test group was converted into a percentage using the following equation: percentage effect = 100 × [(AUC for the treatment group − AUC for the saline group)/(AUC of cutoff − AUC for the saline group)].
CSF Dialysate Collection and Measurement of Excitatory Amino Acids
The microdialysis probe was connected to a syringe pump (CMA-100, Acton, MA) and perfused with Ringer solution (8.6 mg/mL sodium chloride, 0.33 mg/mL calcium chloride, and 0.3 mg/mL potassium chloride). Before the first CSF dialysate collection for each rat, the probe was washed for 30 minutes with Ringer solution. CSF dialysates were collected on the day before PST and on days 1, 4, 7, 10, 14, 18, and 21 after PST. For the drug treatment study on day 10 after PST, a CSF sample was collected before drug injection (basal level) and another 6 samples were collected at 30, 60, 90, 120, 150, and 180 minutes after drug administration. Each sample was collected for 30 minutes at a flow rate of 5 μL/min into a polypropylene tube on ice, which was then frozen at −80°C until assayed. CSF dialysates were analyzed for amino acids. Samples were derivatized by o-phthalaldehyde and 3-mercaptopropionic acid in borate buffer (Agilent part no. 5061-3335; Santa Clara, CA) using a high-performance liquid chromatograph (1100; Agilent) equipped with a reverse-phase ZORBAX Eclipse AAA column (4.6 × 150 mm2, 3.5 μm) and fluorescent detector (Gilson model 121, set at 428 nm) as described previously.26,27 External standards (156.25, 312.5, 625, 1250, and 2500 μmol/L authentic amino acids) are run at the beginning and the end of each sample group. Peak heights will be normalized to the standard peaks and then quantified based on a linear relationship between peak height and amounts of corresponding standards.
Spinal Cord Preparation and Western Blotting Analysis
All rats received behavior tests of tactile allodynia and thermal hyperalgesia, and rats were killed immediately after completing the tests. The rats were rapidly decapitated and the left dorsal quadrant of the lumbar spinal cord enlargement was removed and stored at −80°C until used. The spinal cord samples were homogenized in ice-cold lysis buffer (50 mM Tris, pH 7.5, 150 mM NaCl, 2% Triton X-100, 100 μg/mL phenylmethylsulfonyl fluoride, 1 μg/mL aprotinin), and centrifuged at 100,000 g for 30 minutes at 4°C. The supernatant was collected as the sample and the protein amount was determined using the Bradford protein assay. The proteins were denatured by heating at 90°C for 10 minutes in an equal volume of reducing sample buffer and separated on 10% sodium dodecylsulfate–polyacrylamide gel electrophoresis gels, then transferred to polyvinylidene difluoride membranes (0.22 μM pore size; Millipore, Bedford, MA). The membranes were blocked for 1 hour at room temperature with blocking buffer (5% milk in 20 mM Tris-HCl, pH 7.4, 0.1% Tween 20, 137 mM NaCl), then incubated overnight at 4°C with polyclonal rabbit antibodies against rat GLAST (anti-GLAST; 1:1000; producing a band of approximately 67 kDa), rat GLT-1 (anti-GLT-1; 1:1000; producing a band at approximately 66 kDa), or rat excitatory amino acid (EAAC1 [anti-EAA]C1; 1:1000; producing a band at approximately 69 kDa) (all from Alpha Diagnostic International, Inc., San Antonio, TX) diluted in blocking buffer, then incubated for 1 hour at room temperature with horseradish peroxidase–conjugated goat antirabbit immunoglobulin G antibodies (1:2000; Jackson ImmunoResearch Laboratories, Inc., West Grove, PA), and diluted in blocking buffer. After reaction for 1 minute with electrochemiluminescence solution (Amersham, Arlington Heights, IL), bound antibody was visualized using a chemiluminescence imaging system (Syngene, Cambridge, UK). After probing for 1 of the 3 glutamate transporter proteins, the blots were incubated for 18 minutes at 56°C in stripping buffer (62.6 mM Tris-HCl, pH 6.7, 2% sodium dodecylsulfate, 100 mM mercaptoethanol) and reprobed with mouse monoclonal anti–β-actin antibody (1:10,000; producing a band at approximately 43 kDa; Sigma Chemical Co., St. Louis, MO) diluted in blocking buffer as the loading control. Each Western blot analysis was performed 3 times for each rat, with 6 rats per group. The density of each specific band was measured using a computer-assisted imaging analysis system (Gene Tools Match Software; Syngene).
Immunohistochemistry and Image Analysis
All rats received behavior tests of tactile allodynia and thermal hyperalgesia, and rats were killed immediately after completing the tests. Rats were deeply anesthetized and perfused intracardially with 0.15 M phosphate buffer (pH 7.2), followed by 4% paraformaldehyde in the same buffer. The lumbar spinal cord enlargements were removed, rinsed in phosphate buffer, and cryoprotected in 10% sucrose in phosphate buffer for 24 hours and 30% sucrose in phosphate buffer for 24 hours. They were then embedded in optimal cutting temperature compound (Sakura Finetek, Inc., Torrance, CA) and frozen at −80°C. Sections (5 μm) were cut from the cranial end of the dorsal root entry zone of the L4 spinal nerve to the caudal end of the L5 root entry zone, air-dried on microscope slides for 30 minutes at room temperature, and fixed by immersion in ice-cold acetone/methanol (1:1) for 1 minute. After 3 washes in ice-cold phosphate-buffered saline (PBS), the sections were blocked by incubation in blocking solution (10% normal goat serum in PBS) at room temperature for 1 hour, then were incubated sequentially with fluorescein isothiocyanate-labeled mouse monoclonal antirat glial fibrillary acidic protein (GFAP) antibody (1:100, green fluorescence; Molecular Probes, Eugene, OR) and unlabeled polyclonal rabbit antirat GLT-1 or antirat GLAST antibodies (1:100; Alpha Diagnostic International, Inc.) (both diluted in PBS) for 16 hours at 4°C, and finally incubated for 40 minutes at room temperature with Qdot 655 goat F(ab′)2 antirabbit immunoglobulin G antibodies (1:200, red fluorescence; Quantum Dot Corp., Hayward, CA) in blocking buffer. Images were captured using an Olympus BX 50 fluorescence microscope (Olympus Optical, Tokyo, Japan) and a Delta Vision disconsolation microscopic system operated using SPOT software (Diagnostic Instruments, Inc., Sterling Heights, MI). The fraction of the total area stained positive for glutamate transporters in laminae I and II or laminae III to V of the dorsal horn was calculated using NIH Image J software (Bethesda, MD), and the fractional areas in the 3 rats in each group were averaged.
Differences in the hindpaw withdrawal thresholds between PST and sham-operated animals were assessed using the Wilcoxon Mann-Whitney U test (2-tailed) with correction for repeated testing. Data for the relative density of Western blotting bands were analyzed by 1-way analysis of variance followed by multiple comparisons with the Student-Newman-Keuls post hoc test. Data for the fractional areas of glutamate transporters were compared using the Student t test. All data are presented as means ± SEMs. P values <0.05 are considered statistically significant.
Effect of PST on Thermal Hyperalgesia and Tactile Allodynia
PST induced rapid and persistent reduction of the paw withdrawal thresholds to heat and tactile stimuli. As shown in Figure 1A, the paw withdrawal threshold to radiant heat decreased significantly from day 4 to day 21, with a maximal reduction of 37%. Mechanical allodynia was seen from day 1 to day 21, with a maximal reduction of 56% (Fig. 1B). In contrast, no change was observed in the withdrawal thresholds in the thermal or tactile test for the sham-operated hindpaw or the nonoperated contralateral hindpaw. Thus, PST induced thermal hyperalgesia and tactile allodynia.
Effect of Intrathecal Injection of Ultra-Low Dose Naloxone on the Effect of Morphine
On day 10 after PST, intrathecal injection of 10 μg morphine (group M) restored the radiant heat paw withdrawal threshold to the basal level seen in sham-operated rats from 30 to 90 minutes after injection (Fig. 2A), but only had intermediate effect on the tactile threshold (Fig. 2B) between sham-operated and saline control rats. Intrathecal injection of 15 ng naloxone plus 10 μg morphine (group Mn) reversed the hindpaw withdrawal threshold to radiant heat to the basal level seen in sham-operated rats for a longer period, from 30 to 150 minutes after injection (Fig. 2A). Surprisingly, for tactile allodynia, ultra-low dose naloxone plus morphine had an antiallodynia effect and restored the tactile threshold to the baseline (39.15 ± 0.84 g) from 30 to 90 minutes after injection (Fig. 2B). Intrathecal injection to PST rats with either 15 μg (group N) or 15 ng (group n) naloxone alone did not affect the AUC for thermal hyperalgesia or tactile allodynia compared with the saline group (group s) over the 180-minute observation period after drug treatment (Fig. 2, C and D). In contrast, injection of 15 ng naloxone and 10 μg morphine produced an antihyperalgesia and antiallodynia effect in PST rats (35.1% and 55.4% increases, respectively, in the AUC compared with group s) and had a much better effect than morphine alone (18.7% and 32.9% increase, respectively, compared with group s) or morphine plus high dose naloxone (12.1% and 13.4% increase, respectively, compared with group s) (Fig. 2, C and D). Thus, intrathecal injection of ultra-low dose naloxone enhanced the effect of morphine on the reversal of thermal hyperalgesia and tactile allodynia in PST rats.
Effect of Intrathecal Coadministration of Ultra-Low Dose Naloxone Plus Morphine on EAA Concentrations in the CSF Dialysates
CSF dialysates were collected from the lumbar regions of the rat spinal cords. After PST, glutamate and aspartate concentrations in the CSF dialysates significantly increased from day 1 to day 21, with no significant difference between the results for the different days (Fig. 3, A and B). On day 10 after PST, CSF dialysates were collected every 30 minutes for 3 hours and analyzed for EAAs. No significant difference in EAA concentrations was found among groups s, N, n, M, and MN at any time point, whereas in group Mn, both EAAs decreased significantly at 60, 90, and 120 minutes and the Glu concentration decreased significantly at 150 minutes after intrathecal drug injection (Fig. 3, C and D). Thus, intrathecal coadministration of ultra-low dose naloxone plus morphine reduced EAA concentrations in the CSF dialysates of PST rats.
Effect of Ultra-Low Dose Naloxone on Glutamate Transporter Expression in the Spinal Cord Dorsal Horn of PST Rats
The time course of glutamate transporter expression in the rat spinal cord dorsal horn after PST was examined by Western blotting. PST induced significant downregulation of the glial glutamate transporters GLAST and GLT-1, but not the neuronal glutamate transporter EAAC1, on the PST side of the spinal cord dorsal horn (Fig. 4). In fact, there was an increase in EAAC1 expression on days 1 and 4 after PST, followed by a gradual decrease to the baseline level on day 21. There was no difference in glutamate transporter expression in the contralateral spinal cord dorsal horn between PST and sham-operated rats or between sham-operated and naïve rats (data not shown).
The effects of drug treatment on glutamate transporter expression were examined on day 10 after PST. Intrathecal administration of 15 ng naloxone plus 10 μg morphine restored GLT-1 and GLAST expressions to the basal level, but had no effect on EAAC1 expression (Fig. 5, A and B). There was no significant change in glutamate transporter expression with any of the other treatments.
In the immunohistochemical study, attenuation of GLT-1 and GLAST immunostaining in the spinal cord dorsal horn ipsilateral to the PST was observed (Fig. 6, Aa and Ba); quantitative analyses revealed a significant reduction in GLT-1 and GLAST immunostaining in laminae I and II of the spinal cord dorsal horn ipsilateral to the PST (Fig. 6, Ae and Be) and restoration of GLT-1 and GLAST expression by intrathecal injection of 15 ng naloxone plus 10 μg morphine (Fig. 6, Ac and Bc) in laminae I and II (Fig. 6, Ae and Be). On the contralateral side to the PST, there was no apparent change after intrathecal treatment (Fig. 6, Ad and Bd). No obvious change in glutamate transporters was seen in laminae III to V on either side of the spinal cord dorsal horn of PST rats. In laminae I and II of the spinal dorsal horn on the PST side, GFAP immunoreactivity showed hypertrophy of astrocytes after PST (Fig. 6, Ag and Bg), with a decrease in GLT-1 and GLAST immunoreactivity (Fig. 6, Af and Bf). Furthermore, GLT-1 and GLAST were colocalized with astrocytes (Fig. 6, Ah and Bh). Intrathecal injection of 15 ng naloxone plus 10 μg morphine restored GLT-1 and GLAST expression in laminae I and II on the ipsilateral side of the spinal dorsal horn to the PST (Fig. 6, Ak and Bk) and GLT-1 and GLAST were colocalized with astrocytes (Fig. 6, Ak and Bk).
Effect of Glutamate Transporter Blockade on the Antihyperalgesia and Antiallodynia of Ultra-Low Dose Naloxone Plus Morphine
DL-TBOA is a nontransportable broad-spectrum glutamate transporter blocker and inhibits extracellular glutamate uptake by membrane glutamate transporters.28 As shown in Figure 7, intrathecal DL-TBOA treatment immediately reduced the paw withdrawal threshold in PST rats and the effect lasted for 15 minutes. Moreover, 40 minutes after intrathecal injection of 15 ng naloxone plus 10 μg morphine, intrathecal injection of 10 μg DL-TBOA completely abolished the ultra-low dose naloxone plus morphine effect of reversing the tactile allodynia in PST rats; the effect lasted for 15 minutes. Intrathecal saline had no effect on mechanical allodynia.
In the present study, we found that (1) PST decreased expression of the glial glutamate transporters GLT-1 and GLAST in the rat spinal cord dorsal horn, but not of the neuronal glutamate transporter EAAC1; (2) ultra-low dose naloxone plus morphine administration restored glial GLT-1 and GLAST expression in PST rats and alleviated the PST-induced thermal hyperalgesia and tactile allodynia; (3) the accumulation of EAAs in the CSF of PST rats was reduced by intrathecal ultra-low dose naloxone plus morphine treatment; (4) astroglial glutamate transporter levels in laminae I and II of the spinal dorsal horn were altered by PST and intrathecal ultra-low dose naloxone plus morphine treatment; and (5) the glutamate transport blocker DL-TBOA abolished the antiallodynia effect of ultra-low dose naloxone plus morphine. These results suggest that glial glutamate transporters have a key role in maintaining EAA homeostasis and that the inhibition of their downregulation is responsible for the antihypersensitivity effect of intrathecal ultra-low dose naloxone plus morphine administration in the nociceptive assessments.
The PST rat model allows nerve lesions to be produced without introducing foreign material, shows a reduction in thermal and tactile thresholds, and allows examination of the effect of drugs on animal behavior after endoneurial injury.29 PST induces endoneurial inflammation with less of an epineurial component, in contrast to the chronic nerve-constriction injury model, which causes persistent epineurial inflammation. Both PST and chronic nerve-constriction injury induce nerve injury and neuroinflammation, which result in neuropathic pain, with a reduction in the pain threshold and increased hypersensitivity to both noxious and non-noxious stimuli.30 Our present study demonstrated that PST induced rapid and persistent painful neuropathy, as does the chronic nerve-constriction injury model. After recovery from surgery-related inflammation, PST induced a chronic neuropathic pain state in rats, with thermal hyperalgesia and tactile allodynia, which are similar to clinical neuropathic pain presentations.31 Painful peripheral neuropathy is associated with anatomical changes in the spinal cord dorsal horn, including increased expression of GFAP, an indicator of reactive gliosis after central nervous system injury, and decreased expression of μ-opioid receptors and glial glutamate transporters.11,32,33 In our present study, reduced expression of the glial glutamate transporters GLT-1 and GLAST, but not the neuronal glutamate transporter EAAC1, was observed in the ipsilateral spinal cord dorsal horn of the PST rats both by Western blotting and immunohistochemical analysis. Moreover, the time course of the change in glial glutamate transporter expression in the spinal cord dorsal horn matched that of the emergence of hypersensitivity to noxious thermal and non-noxious tactile stimuli. An initial upregulation of neuronal EAAC1 was seen on days 1 and 4 after PST; this served as a protective mechanism to minimize the adverse consequences of nerve injury–induced glutamate accumulation in the synapse, which overstimulates postsynaptic glutamatergic receptors, but is unable to prevent the development of neuropathic pain, as suggested previously.11
Glial glutamate transporters are responsible for the regulation of excitatory synaptic transmission by controlling the synaptic glutamate concentration.6,9,10 The wrapping of neurons by astroglia has a significant role in regulating glutamatergic neurotransmission, which controls synaptic glutamate clearance by glutamate transporters. Excessive glutamate increases excitatory signal transmission to postsynaptic neurons via NMDA (N-methyl-D-aspartic acid) and AMPA (α-amino-3-hydroxyl-5-methyl-4-isoxazole-propionate) receptors. Downregulation of spinal glutamate transporters contributes to various abnormal pain states, such as neuropathic pain and morphine tolerance.34 In our present study, enhancement of excitatory synaptic transmission in the spinal neuronal networks was responsible for the hypersensitivity of pain signaling, which underlies neuropathic pain development. Several theories are proposed to explain the mechanisms of glial glutamate transporter downregulation. In neurons, μ-opioid receptors on presynaptic glutamatergic terminals and their activation reduce the probability of glutamate release,35 and it is, therefore, proposed that a reduction in μ-opioid receptors in the superficial dorsal horn after peripheral nerve injury reduces the opportunity for μ-opioid receptor activation, thus increasing the probability of glutamate release and subsequent opioid insensitivity.36–38 Furthermore, this neuronal alteration changes glutamate transporter expression in astrocytes.39 Although the influence of neurons on astrocytes still needs to be investigated, the reduction in glial glutamate transporter expression after PST could be partly caused by the downregulation of μ-opioid receptors in neurons. In astrocytes, various factors, such as epidermal growth factor, tumor necrosis factor-α, and G protein–coupled receptors, regulate glial glutamate transporter expression.40 In addition, μ-opioid receptors in astrocytes could also have a pivotal role in glial glutamate transporter regulation.40 Induction of GLT-1 expression in astrocytes is partially mediated by the extracellular signal–regulated protein kinase (ERK), and ERK phosphorylation enhances GLT-1 expression in rat astrocytes.41–43 Acute μ-opioid receptor agonist binding to μ-opioid receptors activates the ERK phosphorylation cascade, with a peak at 2 to 10 minutes.44,45 This suggests that the rapid turnover of GLT-1 expression is induced by μ-opioid receptor activation, because morphine insensitivity and subsequent GLT-1 downregulation in astrocytes is associated with the downregulation of μ-opioid receptors in peripheral nerve-injured rats.36 GLAST expression is regulated by neuron-glia communication and the intracellular calcium concentration; excitatory signals trigger cytosolic calcium release from the endoplasmic reticulum and cause downregulation of GLAST expression in astrocytes.46,47 Increased μ-opioid receptor activation could, therefore, upregulate glial glutamate transporter expression. Our present findings suggest that downregulation of the glial glutamate transporters GLT-1 and GLAST could be closely associated with the downregulation or inhibition of μ-opioid receptor activity after peripheral nerve injury.36 This contributes to the hypersensitivity to noxious thermal stimuli and non-noxious tactile stimuli and the poor response to morphine (Fig. 1, A and B) by retarding the clearance of EAAs (Fig. 3, A and B), resulting in enhancement of excitatory synaptic transmission.
In the present study, morphine had a reduced antihyperalgesia and antiallodynia effect in PST rats, and, not surprisingly, a regular dose of naloxone (15 μg) antagonized the analgesic effect of morphine. In contrast, ultra-low dose naloxone (15 ng) augmented the analgesic effect of morphine. This is compatible with the results of previous in vitro and in vivo studies, in which many mechanisms were proposed to explain this phenomenon.1,2,48,49 Mu-opioid receptor signal transduction is coupled to both Gs and Gi proteins.50 Cotreatment with ultra-low dose naloxone attenuates morphine-induced Gs coupling and Gβγ signaling to adenylyl cyclase and increases Gi/o coupling; ultra-low dose naloxone enhances opioid analgesia by preventing the switch in G protein coupling from Gi/o to Gs.51 Ultra-low dose naloxone supposedly selectively antagonizes excitatory, but not inhibitory, opioid receptor signal transduction in mouse dorsal root ganglia and enhances the antinociceptive effect of morphine.1,14 Moreover, at ultra-low doses, naloxone supposedly binds to a pentapeptide segment of the scaffolding protein filamin A with approximately 200-fold higher affinity than to μ-opioid receptors.48,49 Filamin A interacts with μ-opioid receptors and disrupts the classic opioid-Gi/o protein coupling, switching the coupling to Gs protein. This suggests that ultra-low dose naloxone binds to filamin A and blocks its function in switching inhibitory to excitatory downstream signal transduction.48,51,52 In response to μ-opioid receptor activation by ultra-low dose naloxone plus morphine, glial glutamate transporters were also upregulated to enhance EAA uptake and prevent EAA spillover into the CSF (Fig. 3, C and D). Furthermore, the glutamate transporter blocker DL-TBOA blocks glial glutamate transport function, which temporarily abolishes glial glutamate transporter uptake capacity and allows EAA accumulation in the synaptic cleft.28,53 This enhances excitatory synaptic transmission in the spinal cord and causes hypersensitivity of PST rats to tactile stimuli and the poor response to morphine observed in our present study. As the excessive levels of glutamate and aspartate activate NMDA receptor function, they counteract the effect of morphine. These results suggest that the downregulation of GLT-1 and GLAST has a major role in the hypersensitivity of neuropathic pain in PST rats.
In conclusion, in the present study, a clear association was found between spinal cord dorsal horn astrocyte, glutamate transporter expression and peripheral nerve injury–induced hyperalgesia and allodynia in PST rats. Furthermore, we demonstrated that ultra-low dose naloxone plus morphine treatment restores the antinociceptive effect of morphine in PST rats and that this is accompanied by restoration of GLAST and GLT-1 expression in astrocytes, which enhances EAA reuptake and inhibits synaptic excitatory transmission. Clinical efforts to increase astrocytic expression of glutamate transporters or prevent their downregulation could help to clear accumulated EAAs from the synaptic cleft and minimize synaptic excitatory transmission, thus improving the analgesic effect of morphine in neuropathic pain management. The present results open promising prospects to the development of a new pharmacotherapeutic approach against neuropathic pain.
Assaying dialysate γ-aminobutyric acid A would yield information on the impact of the glial-neuronal shift in glutamate uptake. Because neurons readily metabolize glutamate to γ-aminobutyric acid A while glia metabolize glutamate to glutamine, which neurons take up and release as glutamate, a shift in cellular uptake patterns is likely to have an impact on the balance of excitatory and inhibitory signaling in the spinal cord. The details of these mechanisms need further investigation.
Name: Chih-Ping Yang, MD.
Contribution: This author helped design the study, conduct the study, analyze the data, and write the manuscript.
Attestation: Chih-Ping Yang has seen the original study data, reviewed the analysis of the data, approved the final manuscript, and is the author responsible for archiving the study files.
Name: Chen-Hwan Cherng, MD, DMSc.
Contribution: This author helped analyze the data.
Attestation: Chen-Hwan Cherng has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Ching-Tang Wu, MD.
Contribution: This author helped conduct the study.
Attestation: Ching-Tang Wu has seen the original study data and approved the final manuscript.
Name: Hui-Yi Huang, MS.
Contribution: This author helped conduct the study.
Attestation: Hui-Yi Huang has seen the original study data and approved the final manuscript.
Name: Pao-Luh Tao, PhD.
Contribution: This author helped design the study.
Attestation: Pao-Luh Tao approved the final manuscript.
Name: Chih-Shung Wong, MD, PhD.
Contribution: This author helped design the study, analyze the data, and write the manuscript.
Attestation: Chih-Shung Wong has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
This manuscript was handled by: Tony L. Yaksh, PhD.
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