Morphine is a powerful analgesic for treating moderate to severe pain. However, long-term morphine administration induces tolerance, which hampers its clinical use. Morphine tolerance is a complex physiological response, which includes opioid receptor uncoupling,1 and endocytosis/desensitization,2 increased binding of β-arrestin to opioid receptor,3 glutamatergic receptor activation,4 and neuroinflammation.5,6 The glutamatergic receptor system, especially the N-methyl-D-aspartate receptor (NMDAR), is tetrameric hetero-oligomers consisting of the essential NR1 subunit and ≥1 modulatory NR2A–D and NR3 subunit. Activation of spinal NMDARs has a crucial role in the development of morphine tolerance7,8 and neuropathic pain.9 Pharmacological blockade of NMDARs or disruption of the NR1 subunit gene significantly attenuates morphine tolerance.10,11 The postsynaptic density (PSD) protein family, particularly PSD-95, is critical for anchoring NMDAR NR2 subunits in the postsynaptic membrane, which triggers many physiological and pathological responses.12,13
Resveratrol (trans-3,4′,5-trihydroxystilbene) is a natural polyphenolic compound found in a large number of plants as components of the human diet, including peanuts, mulberries, grapes, and red wine. Evidence suggests that resveratrol has several beneficial biological effects such as antioxidation,14 neuroprotection,15,16 anticancer, antiinflammation, and cardioprotection.17 It has no known toxic side effects.18 Resveratrol suppresses synthesis of proinflammatory mediators by inhibiting cyclooxygenase and lipoxygenase pathways, which implies that resveratrol may have analgesic activity.19–21 Moreover, in a carrageenan-induced inflammatory model, resveratrol reversed thermal hyperalgesia in rats.22 In addition, resveratrol is found to inhibit nitric oxide and tumor necrosis factor (TNF)-α expression in diabetic and spinal nerve ligation rats.16,23 These results suggest that resveratrol has potential in the management of pain in patients who develop morphine tolerance. The exact effects and mechanisms of acute resveratrol treatment to enhance morphine's antinociception after morphine tolerance development remain unclear. The present study is the first assessment of acute resveratrol treatment on the antinociceptive effect of morphine in morphine-tolerant rats and its possible mechanism in influencing PSD-95/NMDAR expression in morphine-tolerant rats.
Animal Preparation and Intrathecal Drug Delivery
The use of rats in this study conformed to the Guiding Principles in the Care and Use of Animals of the American Physiology Society and was approved by the National Defense Medical Center Animal Care and Use Committee. Male Wistar rats (350–400 g) were anesthetized with phenobarbital (65 mg/kg, intraperitoneally) and implanted with 2 intrathecal catheters. The catheter was inserted via the atlantooccipital membrane down to the spinal cord segments L5, L6, and S1-3 that are relative to the tail flick reflex.24 One intrathecal catheter was connected to a mini-osmotic pump (Alzet; DURECT Corporation, Cupertino, CA) for infusion of saline (1 μL/h) or morphine (15 μg/h) for 5 days (performed at the rate of 1 μL/h). After catheterization (day 0), all rats were then returned to their home cages, each rat being housed individually and maintained on a 12-hour light/dark cycle with food and water freely available. Rats with neurological deficits were excluded. On day 5 after morphine tolerance developed, the intrathecal catheter for saline or morphine infusion was cut, and the rats were intrathecally injected with saline (5 μL), dimethyl sulfoxide (DMSO) (5 μL), resveratrol (7.5, 15, 30, or 60 μg in 5 μL DMSO), or ifenprodil (5, 10, or 20 μg in 5 μL saline). Thirty minutes later, a single dose of morphine (15 μg in 5 μL saline, intrathecally) was injected and its antinociceptive effect was measured. All drugs were purchased from Sigma (St. Louis, MO), and delivered intrathecally in 5 μL and flushed by 5 μL of saline. Preliminary results showed no abnormal motor function after intrathecal injection of test drugs (data not shown).
Construction of Intrathecal Catheter
Spinal microdialysis probe construction was modified and adapted from our previous study.25–27 An intrathecal catheter was constructed using an 8-cm PE5 tube (0.008-in. inner diameter, 0.014-in. outer diameter; Spectranetics, Colorado Springs, CO) and a 3.5-cm silastic tube (Dow Corning, Midland, MI). The silastic tube was inserted into the PE5 tube and the joint was sealed with epoxy resin and silicon rubber. The dead space of the intrathecal catheter was approximately 8 μL.
Tail-flick latency was measured using the hot water immersion test (52°C ± 0.5°C). The baseline latency was approximately 2 ± 0.04 seconds with a cutoff time of 10 seconds. Rats were placed in plastic restrainers for drug injection and antinociception assessment. The percentage of the maximal possible antinociceptive effect (% MPE) was calculated as follows: (maximum latency − baseline latency)/ (cut of latency − baseline latency) × 100. Latency less than baseline or >10 seconds was assigned MPE values of 0% or 100%, respectively.
Spinal Cord Sample Preparation and Western Blotting Analysis
After drug treatment, as described in the animal preparation and intrathecal drug delivery section, rats were killed by exsanguination under isoflurane (ABBOTT; Abbott Laboratories Ltd., Queenborough, Kent, UK) anesthesia, and laminectomy was performed at the lower edge of the 12th thoracic vertebra. The lumbar enlargement of the spinal cord was immediately removed and stored at −80°C until used for Western blotting. The dorsal portion of the lumbar spinal cords was fractionated into cytosolic, membrane, and nuclear fractions using a cytoplasmic, nuclear, and membrane compartment protein extraction kit, as recommended by the manufacturer (BioChain Institute, Inc., Hayward, CA). The membrane and cytosolic fractions were checked for specificity by Western blotting with mouse antirat epidermal growth factor receptor (1:2000; MBL, Naka-Ku, Nagoya, Japan) and antirat α-tubulin (1:5000; Laboratory Frontier, Seodaemun-gu, Seoul, Korea) antibodies, respectively. The protein concentration of the samples was determined by the bicinchoninic acid assay (Pierce; Thermo Fisher Scientific Inc., Waltham, MA) using bovine serum albumin as the standard. Samples containing 15 μg of protein were adjusted to a similar volume with loading buffer (10% sodium dodecyl sulfate, 20% glycerin, 125 mM Tris, 1 mM EDTA, 0.002% bromphenol blue, 10% β-mercaptoethanol) and the proteins denatured by heating at 95°C for 5 minutes, separated on 10% sodium dodecyl sulfate–polyacrylamide gels, and transferred onto nitrocellulose membranes (Bio-Rad, Hercules, CA). The membranes were blocked with 5% nonfat milk in Tris-Tween buffer saline (50 mM Tris-HCL, 154 mM NaCl, and 0.05% Tween 20; pH 7.4) and incubated overnight at 4°C with polyclonal rabbit antirat NR1, NR2A, or NR2B (all 1:1000 dilution in 5% nonfat milk in Tris-Tween buffer saline; from Chemicon, Temecula, CA) or monoclonal mouse antirat PSD-95 (1:5000 dilution in 5% nonfat milk in Tris-Tween buffer saline) (all from Millipore) antibodies. The membranes were then incubated for 1 hour at room temperature with the corresponding horseradish peroxidase–conjugated donkey antirabbit or antimouse immunoglobulin G antibodies, as appropriate (1:2000 in 5% nonfat milk in Tris-Tween buffer saline; all from Chemicon). Membrane bound secondary antibodies were detected using Chemiluminescenceplus reagent (Perkin Elmer Life and Analytical Sciences, Waltham, MA) and visualized using a chemiluminescence imaging system (Syngene, Cambridge, UK). The optic density of each specific band was measured using a computer-assisted imaging analysis system (Gene Tools Match software; Syngene).
Immunocytochemistry and Image Analysis
Rats were killed by exsanguination under isoflurane anesthesia, and the lumbar enlargement (L5-S3) of the spinal cord was immediately removed and embedded in optimal cutting temperature compound (Sakura Finetek USA Inc., Torrance, CA). Sections (5 μm) were fixed by immersion in ice-cold acetone/methanol (1:1) for 5 minutes. After washing in ice-cold phosphate-buffered saline, sections were incubated with fluorescein isothiocyanate–labeled mouse monoclonal antirat CD11b/c (OX42; marker of microglial) or fluorescein isothiocyanate–labeled mouse monoclonal antirat glial fibrillary acidic protein (marker of astrocyte; all dilution in phosphate-buffered saline with Triton X-100 containing 2% normal goat serum; Serotec, Oxford, UK) at 4°C overnight. The images were captured using an Olympus BX 50 fluorescence microscope (Olympus Optical, Tokyo, Japan) and a Delta Vision disconsolation microscopic system operated by SPOT software (Diagnostic Instruments Inc., Sterling Heights, MI). Controls without primary antibody were run to confirm the specificity of staining.
Immunoprecipitation of PSD-95/NR1 and NR2B Subunits Complex
To determine the assembly of PSD-95, NR1 and NR2B subunits, coimmunoprecipitation was performed by using immobilized anti–PSD-95 antibody. Anti–PSD-95 antibody (1:50; Cell Signaling, Danvers, MA) was covalently cross-linked to Dynabeads® protein A (Invitrogen, Carlsbad, CA) according to the manufacturer's instructions. The PSD-95/NR1 and NR2B complexes were isolated by incubating 200 μg of spinal cord dorsal horn membrane proteins solubilized in cytoplasmic, nuclear, and membrane Compartmental Protein Extraction Kit extraction buffer with 50 μL of Dynabeads protein A for 1 hour at room temperature. The incubation performed with normal mouse serum was used as a negative control. Dynabeads were precipitated using a magnet, and then the beads were extensively washed with phosphate-buffered saline. Precipitated proteins were eluted with 50 μL sodium dodecyl sulfate– containing sample buffer, and 20 μL of the samples were used for Western blots as described above in spinal cord sample preparation and Western blotting analysis.
Quantitative Real-Time Polymerase Chain Reaction
Total RNA was extracted using TRIzol reagent (Invitrogen). The first-strand cDNA synthesis reaction was performed using 1 μg of Dnase-treated total RNA, 50 ng of random hexamer primer, 0.5 mM dNTP mix, 10 mM dithiothreitol, 1× RT buffer, and 200 U of Superscript III™ reverse transcriptase (Invitrogen) in a total volume of 20 μL. The reaction was performed at 25°C for 10 minutes, followed by 50°C for 50 minutes, and terminated at 85°C for 5 minutes. Real-time polymerase chain reaction was performed using the Applied Biosystems Prism 7500 Sequence Detection System with FastStart Universal SYBR Green Master Mix (ROX; Roche Applied Science, Mannheim, Germany) gene-specific primers, and diluted cDNA. The thermal cycle conditions were 10 minutes at 95°C, 2-step polymerase chain reaction for 40 cycles of 95°C/15 seconds, and, finally, incubation at 60°C for 1 minute. Fluorescent data were acquired during each extension phase. After 40 cycles, a melting curve was generated to verify primer specificities. All samples were tested in triplicate. The amplification data were analyzed using Applied Biosystems Prism Sequence Detection Software version 1.1. To compare the relative expression levels using different treatments, the expression of the gene of interest was normalized to that of the GAPDH control using the ΔΔCT method recommended by the manufacturer. The primer sequences were as follows: TNF-α (NM_012675; forward: 5′-CACCGGCAAGGATTCCAA-3′, reverse: 5′-CACTCAGGCATCGACATTCG-3′), interleukin (IL)- 1β (NM_031512; forward: 5′-AGCCTTTGTCCTCTGCCAAGT-3′, reverse: 5′-CCAGAATGTGCCACGGTTTT-3′), IL-6 (NM_031512; forward: 5′-TGTTCTCAGGGAGATCTTGGAAAT-3′, reverse: 5′-CATCGCTGTTCATACAATCAGAATT-3′).
All data are presented as mean ± SEM. Statistical analysis was performed using SigmaStat 3.0 software (SYSTAT Software Inc., San Jose, CA). Tail-flick latencies were analyzed using 2-way (time and treatment) analysis of variance (ANOVA) followed by subsequent 1-way ANOVA (at each time of the experiment) with a post hoc Student-Newman-Keuls test. For immunoreactivity data, the intensity of each test band was expressed as the optical density relative to that of the average optical density for the corresponding control band. For statistical analysis, immunoreactivity was analyzed using 1-way ANOVA, followed by multiple comparisons with the Student-Newman-Keuls post hoc test. A significant difference was defined as a P value <0.05.
Resveratrol Enhances the Antinociceptive Effect of Morphine in Morphine-Tolerant Rats
Similar to our previous study,28,29 morphine challenge (15 μg, intrathecally) produced a significant antinociceptive effect in vehicle-infused rats (100% MPE) but not in morphine-tolerant rats (20% MPE) (Fig. 1A). In contrast, morphine challenge only produced 20% of MPE in morphine-tolerant rats. Resveratrol (30 μg) alone did not produce an antinociceptive effect in either saline- or morphine-infused rats (data not shown). However, pretreatment with resveratrol 30 minutes before the morphine challenge enhanced morphine's antinociceptive effect in morphine-tolerant rats in a dose-dependent manner, with a maximal effect at 60 minutes (Fig. 1B). As shown in Figure 1B, resveratrol (30 μg) significantly improved morphine-induced antinociception in morphine-tolerant rats with an MPE of up to 63%. A higher dose of resveratrol (60 μg) did not further enhance the antinociceptive effect of resveratrol (30 μg) treatment in morphine-tolerant rats.
Resveratrol Reverses the Up-Regulation of NMDAR NR1/NR2B Subunit Expression in Synaptosomal Membrane of Morphine-Tolerant Rat Spinal Cords
Western blot analysis of spinal dorsal horn cytosolic extracts showed no significant difference in NR1, NR2A, and NR2B subunit expression among groups (Fig. 2, A and B). In contrast, as shown in Figure 2B, increasing NR1 and NR2B subunit expression was observed in the synaptosomal membrane of morphine-tolerant rat spinal cords, and this was reversed by resveratrol (30 μg) pretreatment before the morphine challenge; α-tubulin and epidermal growth factor receptor were used to confirm the identity of the cytosolic and membrane fractions.
NR1/NR2B Subunit Antagonist Ifenprodil Attenuates the Antinociceptive Tolerance of Morphine
As shown in Figure 3, on day 5, 3 hours after discontinuation of the morphine infusion, a morphine challenge (15 μg) did not produce an antinociceptive effect in morphine-tolerant rats, whereas a significant antinociceptive effect was seen in the saline control group rats (Sal + Sal). Treatment with ifenprodil (10 μg, intrathecally) 30 minutes before the morphine challenge attenuated antinociceptive tolerance in morphine-tolerant rats; ifenprodil alone did not produce any antinociceptive effect in either saline- or morphine-infused rats (data not shown). The result indicates that suppression of NMDAR activity maintained the antinociceptive effect of morphine. A higher dose of ifenprodil (20 μg) did not further enhance the antinociceptive effect in morphine-tolerant rats.
Resveratrol Inhibits the Increase of Postsynaptic Membrane PSD-95 Expression and Down-Regulates PSD-95/NR1 and PSD-95/NR2B Coassembly in Morphine-Tolerant Rat Spinal Cords
In Figure 4, the density of the PSD-95 band on immunoblots of postsynaptic membrane fraction from the saline-infusion plus DMSO-injected group is defined as 1. Resveratrol treatment alone had no effect on PSD-95 expression. Long-term morphine infusion increased PSD-95 expression in synaptosomal membrane and this effect was inhibited by resveratrol pretreatment before the morphine challenge. PSD-95 provides a physical structure for NMDARs anchoring at the postsynaptic membrane, and for PSD-95 coassembling with NR1 and NR2B. As shown in Figure 5, an increase of coassembly of 3 proteins was seen in the morphine-tolerant rat lumbar spinal cord dorsal horn. Resveratrol pretreatment before the morphine challenge significantly inhibited the increase of PSD-95, NR1, and NR2B expression and coassembly in rats receiving long-term intrathecal morphine infusion.
Resveratrol Suppresses Proinflammatory Cytokine mRNA Expression and Glial Activation
We observed a significant increase of TNF-α, IL-1β, and IL-6 mRNA expression in morphine-tolerant rat spinal cords, and this was significantly attenuated by resveratrol pretreatment (Fig. 6). Furthermore, in an immunohistochemical study (Fig. 7), the morphology of both astrocyte and microglia was altered from ramified shape to ameboid shape (Fig. 7, A, B, D, and E); also, a strong immunoreactivity was observed in astrocyte and microglia in the dorsal spinal cord of morphine-tolerant rats (Fig. 7, B and E). These changes indicate that astrocyte and microglia were more activated in morphine-tolerant spinal cord than those in the saline-infusion plus DMSO-injected rats (Fig. 7, A and D), and pretreatment with resveratrol (30 μg) significantly suppressed the activation of astrocyte and microglia (Fig. 7, C and F).
In the present study, we found that resveratrol treatment maintained the antinociceptive effect of morphine, inhibited morphine-induced proinflammatory cytokines expression, and suppressed glial activation in morphine-tolerant rat spinal cords. Moreover, long-term morphine infusion induced NMDAR NR1 and NR2B subunit up-regulation in synaptosomal membrane of morphine-tolerant rat lumbar spinal cords, which was suppressed by the resveratrol treatment. The NMDAR antagonist ifenprodil also produced an effect similar to resveratrol. In addition, resveratrol inhibited PSD-95/NR1/NR2B complex expression in morphine-tolerant rat spinal cords. Taken together, regulation of PSD-95 expression by resveratrol is the key to NMDAR NR1 and NR2B subunit expression on the synaptosomal membrane and the subsequent opioid signaling.
Resveratrol rapidly altered the expression of NMDAR NR1 and NR2B subunits on postsynaptic membrane, which was associated with the amount of PSD-95; PSD-95 protein complex was demonstrated to dynamically regulate postsynaptic function and activity, including regulation of synaptic plasticity, synaptogenesis, learning, and memory.30,31 Studies have suggested that PSD-95 is critical for NMDAR NR2A and NR2B subunit anchoring on the postsynaptic membrane, thus triggering physiology and pathology alterations.12,13 Binding of PSD-95 to the NMDAR NR2B C-terminal serine/threonine-X-valine motif was demonstrated to inhibit receptor endocytosis from the neuron surface, and stabilize NMDARs on the cell surface.32,33 These studies suggest that PSD-95 has a crucial role in NMDAR complex trafficking, membrane targeting, and internalization. In the present study, a significant increase in PSD-95/NR1/NR2B immunoprecipitation in morphine-tolerant rats was inhibited by resveratrol treatment. It suggests that low PSD-95 expression results in the loss of NMDAR NR1 and NR2B subunit stability on the postsynaptic membrane, thus inhibiting the subsequent intracellular signaling cascade.
Emerging evidence suggests that NMDAR activation has a crucial role in morphine tolerance,34,35 and blockade of NMDAR function effectively attenuates morphine tolerance.36,37 In our present study, pretreatment with the NMDAR antagonist ifenprodil maintained the antinociceptive effect of morphine in morphine-tolerant rats. NMDARs are highly permeable to Ca2+, and the NR1 subunit of NMDARs is essential for NMDAR function.38 Our present study found that long-term morphine infusion increased NMDAR NR1 and NR2B subunit expression on the postsynaptic membrane and caused morphine tolerance, which agrees with a previous report of an increase of NR1 expression in morphine-tolerant rat spinal dorsal horn.39 It seems that enhancement of NR1 subunit expression at the synapse may strengthen NMDAR-mediated synaptic and intracellular signaling, thus leading to hyperalgesia, and morphine tolerance.11,40
Spinal cord glial activation seems to be a common mechanism that leads to pathological pain and morphine tolerance. Long-term opioid exposure causes synaptic plasticity, which alters the homeostasis between neurons and glial cells and subsequently induces analgesic tolerance.41 Interaction of glial cells–neurons allows glial activation, and induces complicated neurotransmission and expresses various functional receptors, particularly glutamatergic receptors.42,43 Glutamate has a crucial role in excitatory synaptic plasticity and morphine tolerance.28,29 The glutamate-mediated neuron-glial interaction was demonstrated to influence neuronal excitability, synaptic transmission, and central sensitization.44 Coadministration of MK-801 with morphine attenuated glial activation in morphine-tolerant rats, suggesting an involvement of NMDAR-mediated signaling in morphine-induced glial activation.45 Similarly, our present study found that chronic morphine infusion induced glial activation, as shown by glial cell hypertrophy in the dorsal horn of tolerant rat spinal cords, and pretreatment of resveratrol attenuated the morphine infusion–induced glial activation and NMDAR NR1/NR2B subunit expression and reversed PSD-95/NR1 and PSD-95/NR2B coassembly in postsynaptic membrane in morphine-tolerant rats. Based on these results, we suggest that reduction of postsynaptic membrane PSD-95/NMDAR expression by resveratrol treatment may be responsible for attenuating glial activation in morphine-tolerant rat spinal cords. The signaling between neurons–glial cells is complicated. Our present results could not differentiate the main action site of resveratrol on neurons or glial cells; therefore, further in situ hybridization experiments are presently being performed in our lab. In our preliminary observation, we found that morphine induced a morphological change in cultured microglia (BV2), including membrane ruffling and chemotaxis, and NR1 overexpression. Administration of resveratrol or the NMDAR antagonist ifenprodil significantly inhibited morphine-induced BV2 cell activation and migration. These data suggested that NMDAR alteration may modulate several cytoskeleton proteins and influence cell morphology and mobility in morphine-treated microglia. Evidence also demonstrated interactions between NMDARs and α-actinin/actin cytoskeleton.46,47 Taken together, we suggest that (i) resveratrol inhibits morphine-induced glial activation via NMDAR-related signaling pathway, and (ii) NMDARs on glial cell membranes may have a different role from those on neurons.
Previous studies have shown that chronic morphine administration induces glial activation and proinflammatory cytokine TNF-α, IL-1β, and IL-6 expression in morphine-tolerant rat spinal cords.48,49 Intrathecal coadministration of morphine with a glial metabolic inhibitor50 or a glial modulator48 greatly attenuated morphine tolerance. Therefore, limiting proinflammatory cytokine production by activated microglia and astrocytes should be beneficial for reversal of morphine tolerance. In the present study, chronic morphine administration induced a significant increase in TNF-α, IL-1β, and IL-6 mRNA expression in the dorsal horn of morphine-tolerant rat spinal cords. This increase was suppressed by resveratrol pretreatment, suggesting that resveratrol regulates proinflammatory cytokine expression, at the transcriptional level, by interaction with certain transcription factors (such as NFκp65 protein) and related signaling pathways. Moreover, resveratrol treatment also rapidly reversed the morphology of activated astrocyte and microglia to an inactive ramified state. Similarly, MK-801,28 etanercept,4 and lemnalol51 are also reported to rapidly revert glial cells back to an inactive form in different neuropathic animal models. Pretreatment with the NMDAR antagonist MK-801 30 minutes before morphine challenge significantly suppresses glial activation, which was associated with a reduction of proinflammatory cytokine (TNF-α, IL-1β, and IL-6) mRNA and protein expression in morphine-tolerant rats.28 Etanercept pretreatment for 3 hours attenuated glial activation with a rapid inhibition of morphine-induced proinflammatory cytokines TNF-α, IL-1β, and IL-6 mRNA expression.6 Similarly, acute intrathecal lemnalol treatment for 3 hours significantly reverted glial morphology to resting form by inhibition of TNF-α expression on day 14 in chronic constriction injury rats.51 These results further support the idea that reduction of proinflammatory cytokine expression might be a possible mechanism of resveratrol rapidly attenuating glial activation. Moreover, in the present study, resveratrol pretreatment 30 minutes before morphine challenge significantly inhibited synaptic NMDAR NR1 and NR2B subunit expression and was associated with the recovery of morphine's antinociceptive effect. In contrast, subcutaneous infusion of the NMDAR antagonist LY274614 produced a slow reversal of antinociceptive morphine tolerance over several days.52 The possible reasons for the discrepancy between the study by Tiseo et al. and ours are (a) different routes of test drug administration (subcutaneous infusion of drug may go through some metabolic organs [liver and renal] and the brain-blood barrier), (b) LY274614 is a specific NMDAR antagonist and may not have an antiinflammatory effect, (c) Tiseo et al. found a reversal of antinociceptive tolerance, up to 90% of the antinociceptive effect of morphine, by subcutaneous infusion of LY274614 for 9 days.
In summary, the present study demonstrates that resveratrol maintains the antinociceptive effect of morphine in morphine-tolerant rats by down-regulation of NMDAR expression and suppression of neuroinflammation in morphine-tolerant rat spinal cords. Accordingly, this study provides new evidence that resveratrol has potential as an analgesic adjuvant in clinical pain management, particularly for patients who need long-term morphine administration and for morphine-tolerant patients who require better pain relief.
Name: Ru-Yin Tsai, PhD.
Contribution: This author helped design the study, conduct the study, analyze the data, and write the manuscript.
Attestation: Ru-Yin Tsai 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: Kuang-Yi Chou, PhD.
Contribution: This author helped design the study, conduct the study, analyze the data, and provided equal contribution as first author.
Attestation: Kuang-Yi Chou has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Ching-Hui Shen, MD, PhD.
Contribution: This author helped analyze the data.
Attestation: Ching-Hui Shen has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Chih-Cheng Chien, MD, PhD.
Contribution: This author helped analyze the data.
Attestation: Chih-Cheng Chien has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Wei-Yuan Tsai, MS.
Contribution: This author helped conduct the study and analyze the data.
Attestation: Wei-Yuan Tsai has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Ya-Ni Huang, PhD.
Contribution: This author helped analyze the data.
Attestation: Ya-Ni Huang has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Pao-Luh Tao, PhD.
Contribution: This author helped analyze the data.
Attestation: Pao-Luh Tao has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Yaoh-Shiang Lin, MD.
Contribution: This author helped analyze the data.
Attestation: Yaoh-Shiang Lin has seen the original study data, reviewed the analysis of the data, and approved the final manuscript.
Name: Chih-Shung Wong, MD, PhD.
Contribution: This author helped conduct the study and analyze the data.
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: Quinn Hogan, MD.
This study was performed at the Neuropathic Pain and Translational Research Laboratory, Cathay Medical Research Institute, Cathay General Hospital, Xizhi, New Taipei City, Taiwan.
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