Spinal microRNA-134-5p targets glutamate receptor ionotropic kainate 3 to modulate opioid induced hyperalgesia in mice

Abstract

Background: Fentanyl and its analogs are extensively used for pain relief. However, their paradoxically pronociceptive effects often lead to increased opioids consumption and risk of chronic pain. Compared to other synthetic opioids, remifentanil has been strongly linked to acute opioid hyperalgesia after exposure [remifentanil-induced hyperalgesia (RIH)]. The epigenetic regulation of microRNAs (miRNAs) on targeted mRNAs has emerged as an important pathogenesis in pain. The current research aimed at exploring the significance and contributions of miR-134-5p to the development of RIH. Methods: Both the antinociceptive and pronociceptive effects of two commonly used opioids were assessed, and miRNA expression profiles in the spinal dorsal horn (SDH) of mice acutely exposed to remifentanil and remifentanil equianalgesic dose (RED) sufentanil were screened. Next, the candidate miRNA level, cellular distribution, and function were examined by qPCR, fluorescent in situ hybridization (FISH) and Argonaute-2 immunoprecipitation. Furthermore, bioinformatics analysis, luciferase assays, miRNA overexpression, behavioral tests, golgi staining, electron microscopy, whole-cell patch-clamp recording, and immunoblotting were employed to investigate the potential targets and mechanisms underlying RIH. Results: Remifentanil induced significant pronociceptive effects and a distinct miRNA-profile from sufentanil when compared to saline controls. Among top 30 differentially expressed miRNAs spectrum, spinal miR-134-5p was dramatically downregulated in RIH mice but remained comparative in mice subjected to sufentanil. Moreover, Glutamate Receptor Ionotropic Kainate 3 (Grik3) was a target of miR-134-5p. The overexpression of miR-134-5p attenuated the hyperalgesic phenotype, excessive dendritic spine remodeling, excitatory synaptic structural plasticity, and Kainate receptor-mediated miniature excitatory postsynaptic currents (mEPSCs) in SDH resulting from remifentanil exposure. Besides, intrathecal injection of selective KA-R antagonist was able to reverse the GRIK3 membrane trafficking and relieved RIH. Conclusion: The miR-134-5p contributes to remifentanil-induced pronociceptive features via directly targeting Grik3 to modulate dendritic spine morphology and synaptic plasticity in spinal neurons.

Introduction

Opioids are the cornerstone for clinical perioperative analgesia, moderate-to-severe pain treatment, and cancer pain relief. ¹ Strikingly, increasing opioid prescriptions incur high morbidity of debilitating central nervous system (CNS) complications, including dependence, tolerance, and opioid-induced hyperalgesia (OIH), which enables weak stimuli to elicit intense pain sensitization and pain chronification.^2,3 It has been shown that even after short-term or single-dose use, opioids lead to an increased risk of OIH over 6 months, indicating that opioids can produce persistent alterations in the pain processing pathway, well beyond the duration of opioid exposure.^4,5 However, OIH is often concealed by injury or medication and lacks simple diagnostic tests and criteria in clinical settings; thus this clinical entity may occur much more frequently than generally reported, and the limited attention to OIH compromises patients’ pain control and safety. ⁶ Notably, remifentanil exhibits the most obvious pronociceptive effects compared to other opioids, whereas another extensively used opioid, sufentanil, was not found to correlate with hyperalgesia in patients, indicating distinct mechanisms involved in remifentanil-induced hyperalgesia (RIH).^7,8 Consequently, an unbiased comparison between remifentanil and sufentanil may help to illuminate the mechanisms underlying the progression and pathogenesis of RIH.

Experimental data suggests that a broad spectrum of small non-coding RNAs, especially microRNAs (miRNAs) that regulate the gene post-transcriptional inhibition by inducing degradation of the complementary messenger RNA (mRNA), are emerging as attractive candidates for structural and functional neuronal changes during various neurological diseases, including epilepsy, Alzheimer’s disease (AD), depression, and chronic pain. ⁹ Consistently, the dysregulation of miRNAs within pain-related neural regions, especially in the spinal dorsal horn (SDH), have been implicated in the processes of nociceptive input transmission in mammals. ¹⁰ Further, a few miRNAs changes have been unequivocally validated by clinical trials, and miRNA-targeted therapeutics have showed some promising preclinical results.^11,12 Most relevantly, studies on morphine-affected rodents revealed that abnormal expression of neuronal miRNAs in the brain, such as miR-let-7, miR-339, and miR-124, is associated with the development of OIH.^13,14 However, there is no report regarding miRNA expression changes that are induced by remifentanil exposure and their role in RIH. Targeting specific miRNAs changes may define potential biomarkers for diagnostic or therapeutic targets in RIH.

In the current research, we evaluated the distinct antinociceptive and pronociceptive effects of remifentanil and sufentanil, and further analyzed the differential expression profiles of miRNAs in the SDH of remifentanil and remifentanil equianalgesic dose- (RED-) sufentanil exposure mice versus saline controls using an objective and reliable high-throughput microarray. We discovered that a candidate miRNA, miR-134-5p, was down-regulated in remifentanil but not in sufentanil exposed mice, and further provided evidence that the interaction between miR-134-5p and its target genes Glutamate Receptor Ionotropic Kainate 3 (Grik3) modulated the pain-like behavior induced by remifentanil. Moreover, the overexpression of miR-134-5p in SDH reversed spinal dendritic spine remodeling and Kainate receptor mediated-synaptic plasticity. Thus, our study provides a spectrum of spinal miRNA changes after two commonly used opioid treatments and specifically identifies the precise physiological functions of miR-134-5p/Grik3 as a promising target for the prevention and treatment of RIH, and may provide the insight into improving clinical pain management with opioid analgesics.

Materials and methods

Animals

In this study, 2 to 3-month-old male C57BL/6J mice were raised in an artificially controlled 14-h/10-h light-dark environment with food and water ad libitum. All animals were purchased from Beijing Huafukang Bioengineering Co., Ltd. All animal use protocols were reviewed and approved by the Institutional Animal Care and Use Committee of Tianjin Medical University. All animal behavior tests were applied in strict accordance with the International Association for the Study of Pain directives.

Opioid-induced hyperalgesia model

OIH was induced in mice by four intermittent injections of remifentanil [10 μg/kg, 100 μl per injection, intravenous (i.v.); RenFu, China] or sufentanil (0.5 μg/kg, 100 μl per injection, i.v.; RenFu) at 15-min intervals, resulting in a cumulative dose of 40 μg/kg or 2 μg/kg, respectively. The dose of opioids was selected based on the EC₅₀ value as described in the antinociceptive assay. This injection method established a nonspontaneous pain model and did not induce opioid tolerance.^15,16 Control mice received the same amount of normal saline. In all instances, regardless of the opioid dose, the total injection amount was kept constant at 0.4 mL to avoid acute heart failure.

Pain behavioral testing

All tests were conducted between 10:00 a.m. and 2:00 p.m. in a temperature-controlled room (22°C). The mice were acclimated 2 h daily to the testing apparatus for 3 days prior to the baseline threshold test, and the baseline threshold was tested 1 day before the modeling. All behavioral studies were performed by the same investigator who blinded to the treatments.

von frey test

The mice were acclimated for 2 h on a test platform with a grid spacing of 1 mm², covered by a plexiglass box of 10 × 7 × 40 cm. The paw withdrawal threshold (PWT) was measured using von Frey filaments (0.16, 0.4, 0.6, 1, and 2 g, Stoelting, 58011) in an up-down manner, beginning with 0.16 g stimulation of the central hind paw surface. ¹⁷ Licking or withdrawal during the 2-s stimulus was considered a positive response.

Tail-immersion test

The mice’s bodies were gently immobilized in the tester’s hands with 2 cm of the tail tips exposed and immersed in a water bath at 48°C. The time from tail insertion to retraction was recorded as tail withdrawal latency (TWL), and a cutoff time was set at 30-s to prevent tissue damage. This test was performed only once to prevent behavioral sensitization from multiple harmful stimuli.

Antinociceptive assay

The antinociceptive ability was evaluated as previously reported. ¹⁸ After saline or opioid i.v. Injection, the mice were subjected to the tail immersion test. Antinociceptive ability was defined as the percent inhibition of the TWL. The antinociceptive ability of opioids was measured using a dose-response curve, and the dose producing a 50% reduction in TWL (ED₅₀) was compared between remifentanil and sufentanil to obtain the ratio of equivalent dose.

Rotarod testing

The mice were acclimated for the apparatus for 30 min. After that, a 4–45 r/min over 300 s accelerating protocol was applied to assess locomotor activity. This test was performed in 3 daily sessions to get an average fall latency.

MicroRNA extraction, library preparation, and sequencing

The total RNA from L3-L5 spinal cord samples was extracted using Trizol (Thermo, 15596026). The purity, concentration, and integrity of RNA samples were tested using Nanodrop (Thermo) to select qualified samples for transcriptome sequencing. The small RNA library was prepared via the following steps. Briefly, 3′ SR and 5′ SR adaptors were ligated, and reverse transcription was conducted to synthesize the first chain. PCR amplification and size selection was then performed. PAGE gel was used to screen the fragment, and small RNA libraries were cut from the gel and recovered. Finally, PCR products were purified using the AMPure XP beads, and library quality was assessed with Bioanalyzer. The clustering of the index-coded samples was performed on a cBot Cluster Generation System using TruSeq PE Cluster Kit v4-cBot-HS (Illumia) according to the manufacturer’s instructions. After cluster generation, the library was sequenced on an Illumina HiSeq platform and single-end reads were generated.

Bioinformatic analysis

Raw FASTQ data were first processed through in-house Perl scripts. Clean reads were obtained by removing reads containing adapter, ploy-N and low-quality read from raw data. Reads were trimmed and cleaned by removing sequences <18 nt or >30 nt. At the same time, Q20, Q30, GC-content, and sequence duplication levels of the clean data were calculated. All the downstream analyses were based on high-quality clean data. The clean reads were mapped to the Silva database, GtRNAdb database, Rfam database, and Repbase database, respectively to filter ribosomal RNA, transfer RNA, small nuclear RNA, small nucleolar RNA and other ncRNA and repeats. The remaining reads were used to detect known miRNA and novel miRNA predicted by comparisons with Genome and known miRNAs from miRBase. Randfold tools soft was employed for novel miRNA secondary structure prediction. Gene function was annotated and reviewed based on the following databases: Nr (NCBI non-redundant protein sequences), Pfam (Protein family), KOG/COG (Clusters of Orthologous Groups of proteins), Swiss-Prot (A manually annotated and reviewed protein sequence database), KEGG (KEGG Ortholog database), and GO (Gene Ontology). Differential expression analysis was performed using the DESeq2 R package (1.10.1). GO enrichment analysis of the differentially expressed genes (DEGs) was conducted using GOseq R packages. KOBAS software was used to test the statistical enrichment of DEGs in KEGG pathways.

Quantitative real-time polymerase chain reaction

Total RNA was extracted from L3-L5 spinal cord tissue samples using the miRNeasy Mini Kits (Qiagen, 217004). For miRNAs detection, the poly(A) tailing method was employed during reverse transcription using a miScript Reverse Transcription Kit (Qiagen, 218160). A miScript SYBR Green Polymerase Chain Reaction (PCR) kit (Qiagen, 218073) was used for amplification, and the reaction temperature was set according to the instructions. The sequences of primers were as follows: miR-7116-5p (5′-GCGCGTGAAGACATCAGGAA′),miR-539-3p(5′-GCGCGCATACAAGGATAATTT′),miR-7066-5p(5′-GCGTGGGTTGGGAAATGAG′), miR-500-5p (5′-CGAATCCTTGCTATCTGGGTG′), miR-466h-3p (5′-CGCGTACGCACGCACAC′), miR-134-5p (5′-UGUGACUGGUUGACCAGAGGGG′), miR-7029-3p(5′-CGCGTGTTGGGGACATTTT′), miR-8111 (5′-GCGACCGGGCATGGTAGT′), miR-6944-3p (5′-CGCGTAACTCTTCCCTTGTGC′), miR-8103 (5′-GCGCGTCTCCTGTTCTCTGT′). RNU6(5′-ATTGGAACGATACAGAGAAGATT′) was used as the internal control. For GRIK3 detection, GRIK3 (forward 5′‐GCTGGTCTGCACTGAACTCT‐3′ and reverse 5′‐AAAGGGCATCCCCTGAATGG‐3′); and GAPDH (forward 5′‐AGGTGAAGGTCGGAGTCAAC‐3′ and GAPDH reverse 5′‐CGCTCCTGGAAGATGGTGAT‐3′) were used. The expression of miRNAs and GRIK3 was transformed into relative multiple changes using the 2^−∆∆CT method.

In situ hybridization and Immunohistochemistry

The mice were deeply anesthetized and transcardially perfused with pre-cooled PBS following 4% paraformaldehyde. The hydraulic pressure method was employed to get the entire spinal cord. The L3-L5 spinal cord or DRG sections was then dissected and dehydrated in 30% and 20% sucrose for 24 h respectively. The tissues were then frozen in O.C.T. and cut into 5-μm frozen sections using a cryostat (Leica Biosystems).

A digoxin-labeled locked nucleic acid detection probe for miR-134-5p was purchased from Exiqon. After triple washing with PBS, the sections were fixed with 4% paraformaldehyde for 10 min and then treated with protease for 15 min. The sections were rinsed in the hybridization buffer for 1 h at 48°C, followed by labeling with miR-134-5p or scrambled probe diluted 1:200 with hybridization buffer at 48°C overnight. The sections were stringently rinsed twice with warmed 2× saline sodium citrate (SSC) for 10 min at 37°C and in 0.2× SSC for 20 min at room temperature, then treated with blocking buffer for 1h, incubated with mouse anti-digoxigenin antibody (1:500, Roche 11093274910) for 2 h, and rinsed in PBS. Finally, slides were mounted with medium and covered.

To define which cell types express miR-134-5p, the above sections were subsequently blocked with 0.3% Triton X-100 for 10 min and 5% goat serum for 1 h and then incubated with primary antibodies overnight at 4°C. The following primary antibodies were used: anti-NeuN (1:200, Abcam 177487), anti-GFAP (1:200, Cell Signaling Technology 12389), anti-Iba-1 (1:200, Abcam 178847), and anti-CGRP (1:200, Abcam 81887). After incubation, sections were rinsed three times in PBS, and subsequently incubated with corresponding fluorescence-labeled secondary antibody for 1 h at room temperature. Images were captured using a fluorescent microscope (Olympus), and three sections of L3-L5 SDH or DRG from 3 mice were analyzed for fluorescence quantification. All analysis was performed using ImageJ software in a blinded manner.

Argonaute-2 immunoprecipitation

Individual L3-L5 spinal cord tissues for each group were triturated, homogenized in immunoprecipitation buffer (300 mm NaCl, 5 mm MgCl₂, 0.1% NP-40, and 50 mm Tris-HCl, pH 7.5), and centrifuged. Then, the lysate was incubated with Argonaute-2 antibodies (1:50, Cell Signaling Technology C34C6) at 4°C overnight. On the following day, protein A agarose beads (Santa Cruz Biotechnology) were added, mixed, incubated for 1 h at 4°C, and then centrifuged. The supernatant was discarded. The sediment was processed for miRNA q-RT PCR, and the procedure was performed as previously mentioned. All samples were performed in triplicate.

Luciferase assay

HEK293 cells obtained from the Cell Bank of the Chinese Academy of Sciences (Shanghai, China) were co-transfected with miR-134-5p (100 nM) or the scrambled control and wild type (pMIR-R-Grik3-WT) or the mutant (pMIR-R- Grik3-MUT) 3′-UTR plasmids (Obio, H13564, H13565). After 48 h, the cells were harvested and Firefly and Renilla luciferase activities were measured using a dual-luciferase reporter assay kit (Promega, E1910). The Firefly luminescence/Renilla luminescence ratio values were used for normalization and analysis. The experiments were performed in triplicate.

Golgi staining

The mice were deeply anesthetized, and the fresh L3-L5 spinal dorsal horn was quickly dissected without perfusion. The tissues were immersed in a mixture of solutions A and B from FD rapid glogistain kit (FD NeuroTechnologies, PK401) for 2 weeks, and then the tissues were transferred into solution C for 72 h. The tissues were then embedded in agarose and sectioned at 100 μm with a vibrating microtome (Leica, Wetzlar) for staining as follows. The sections were rinsed with distilled water for 4 min twice, dipped in a mixture of solutions D and E and distilled water (1:1:2) for 10 min, and then rinsed twice again in distilled water for 4 min. Subsequently, the sections were dehydrated in 50%, 75%, and 95% ethanol for 4 min according to the concentration gradient and rinsed four times with 100% ethanol for 4 min. Finally, the sections were soaked thrice in xylene for 4 min and then covers lipped with resin. The entire experiment was performed in the dark at room temperature. The images were acquired using a microscope (Olympus). The total length of dendrites, the number of primary branches, and the number and morphology of dendritic spines were analyzed. Dendritic spines of three individual sections were analyzed for each condition.

Electron microscopy

To examine the ultrastructure of synapses in SDH, we followed a previously described method using transmission electron microscopy. ¹⁹ L3-L5 spinal dorsal horn tissue samples were quickly dissected without perfusion and carefully dissected into 1 mm³ cubes and placed in 4% glutaraldehyde at 4°C for 2 h. Next, the sample fixation, dehydration, and embedding were performed by an experienced technician to avoid the potential risk of biohazard reagent exposure. Sections at 50 nm were examined under a transmission electron microscope (Philips). Asymmetric synapses were located by visual identification of dendrites under the electron microscope (EM). 20 micrographs were randomly captured from each tissue, and a blind analysis of morphological parameters using ImageJ software was performed. Three SDH tissues from each group were used for EM analysis.

Spinal cord slice preparation and whole-cell patch-clamp recording of miniature excitatory postsynaptic currents (mEPSCs)

The L4-L5 spinal cord segments were removed by hydraulic extrusion and sliced into transverse slices (350 μM) using a vibrating microslicer (Leica) in oxygenated (95% O₂, 5% CO₂) sectioning buffer (120 mM CholineCl, 2.5 mM KCl, 0.5 mM CaCl₂, 7 mM MgCl₂, 1.25 mM NaH₂PO₄, 26 mM NaHCO₃, and 25 mM glucose) at 4°C. Next, the slices were incubated in a preoxygenated artificial cerebrospinal fluid solution (ACSF; 126 mM NaCl, 2 mM MgCl₂, 3.5 mM KCl, 1.25 mM NaH₂PO₄, 26 mM NaHCO₃, 2 mM CaCl₂, and 10 mM D-glucose, pH 7.4) at 34°C for 30 min and recovery at room temperature (25 ± 1°C) for at least 1 h before recording. Slices were transferred to a recording chamber under perfusion ACSF (2 mL/min, 32–34°C). Spinal cord neurons were visualized with infrared optics using an upright fixed microscope equipped with a 40× water-immersion lens (FN1, Nikon) and CCD monochrome video camera (IR-1000, DAGE-MTI). Patch pipettes (resistance of 3–7 mΩ) were prepared by a horizontal pipette puller (P-1000; Sutter Instruments). For Kainate receptor mediated-mEPSC recording, spinal cord neurons were held at −70 mV in the present of 20 μM bicuculline methiodide (BMI) and 1 μM TTX, 5 μM strychnine, 10 μM CNQX, and 50 μM AP-5, with the pipette solution with pipette solution (130 mM KCl, 0.5 mM CaCl₂, 10 mM HEPES, 2 mM MgCl₂, 10 mM EGTA, 2 mM Mg-ATP, 0.3 mM Na-GTP, pH 7.3). Clampfit nine software (Axon Instruments) were used for data acquisition and analysis.

Western blot analysis

The mice were anesthetized with 3% sevoflurane and sacrificed. The whole spinal cord was blown out with cold PBS from the tail end to the head end. Total protein was extracted from the L3–L5 dorsal horn using RIPA (Thermo, 89900), Membrane proteins were extracted using the Compartment Protein Extraction kit (Thermo, 89842), and the protein content was measured using the bicinchoninic acid assay (Thermo, A53225). After 10% SDS-PAGE gel electrophoresis and membrane transfer, the PVDF membrane of the transformed protein was placed into TBST containing 5% skimmed milk powder, sealed at room temperature for 2 h, and then incubated with the following primary antibodies: GRIK3 (1:100, ab183035) and β-actin (1:2,000, Sigma–Aldrich SAB5500001). The membranes were then incubated with horseradish peroxidase-conjugated secondary antibodies (KPL, 074-1506), and the bands were visualized using Supersignal HRP Chemiluminescence Substrates (Millipore, WBKLS100). The images were captured using Alpha FluorChem FC3 and analyzed using bundled software (Alpha Innotech).

Intrathecal injection

The intrathecal injection was performed between the levels of L4 and L5 using a 30-G needle linked to a Hamilton syringe, and 10 μL of the reagent was given when the reflexive tail flick was observed. The doses of AgomiR-134 and ACET were selected based on a previous study.^20,21

For the pharmacological blockade of GRIK3, a selective GRIK3 antagonist ACET (Tocris, 2728) was injected intrathecally before the first opioid injection. For miR-134 overexpression, Scramble or AgomiR-134 (QIAGEN) or AntagomiR-134 (Exiqon) was injected into the L4 and L5 lumbar vertebral space 3, 2, 1 day before the first opioid injection.

Statistical analysis

All statistical analyses were performed with SPSS 21.0 software (SPSS). All animals were randomly assigned to experimental conditions, and all data were included and analyzed by a blinded researcher. All data were expressed as means ± standard error of mean (SEM). The sample size was calculated as previously described. The Shapiro-Wilk test was used for determining the normality of data distribution, and parametric statistics were applied. The homogeneity of variance was validated using the Levene test. For the dose-response antinociceptive assay, the best-fit line was generated following a nonlinear regression analysis based on the inhibition percentage (%IP) for each mouse, where %IP = [(control times − drug-induced times)/(control times)] × 100. The ED₅₀ values of opioid-induced mechanical and thermal hyperalgesia were calculated by nonlinear regression analysis with a sigmoidal dose–response equation (variable slope). The statistical analyses of behavioral, real-time qPCR, Ago-2 immunoprecipitation were performed by two-way analysis of variance (ANOVA) with Bonferroni posthoc comparisons. The results of Western blot analysis, Golgi staining, electron microscopic, and whole-cell patch-clamp recording data were analyzed using one-way ANOVA with Bonferroni posthoc comparisons. A p value <0.05 was considered statistically significant. The F values were provided when the ANOVA was performed.

Results

Distinct pronociceptive effects between remifentanil and sufentanil based on equianalgesic dose

To experimentally investigate the different pronociceptive effects between remifentanil and sufentanil, we first measured the antinociceptive dose-response curves after sufentanil or remifentanil intravenous injection and obtained calculated antinociceptive ED₅₀ of 9.282 μg/kg remifentanil and 0.468 μg/kg sufentanil, doses that were used for subsequent experiments (Figure 1(a)). Then a mouse model of OIH was established through tail intravenous injection of opioids for four consecutive doses (15-min intervals), which can elicit nonspontaneous pain but would not induce opioid tolerance.^15,16 Mechanical allodynia and thermal hyperalgesia were tested until 7 days after injection (Figure 1(b)). Compared to mice treated with saline, the remifentanil-exposed mice displayed mechanical pain behaviors from 6 h to 5 days post-injection, and peak effects were observed at day 2 (Figure 1(c)). Thermal hyperalgesia was elicited within 3 h after administration and returned to baseline by day 5 (Figure 1(d)). However, mice subjected to RED-sufentanil showed less mechanical and thermal hyperalgesia; the PWT decreased in 6 h (Figure 1(c)) and the TWL decreased slightly in 3 and 6 h (Figure 1(d)).

Figure 1.

Distinct pronociceptive effects and differentially expressed microRNAs genes between remifentanil and sufentanil after consecutive injection in mice. (a) Dose-response curves for antinociceptive effects of remifentanil (intravenous; 2.5, 5, 10, 20, 40 μg/kg) and sufentanil (intravenous; 0.1, 0.2, 0.4, 0.8, 1.6 μg/kg). Nonlinear regression analysis generated best-fit lines based on each mouse’s percentage inhibition flick. All data are means ± SEM (n = 8/doses) and analyzed by One-way ANOVA with Bonferroni post-hoc test. (b) Experimental design to test opioid-induced hyperalgesia according to the timeline shown. (c) Time course of the mechanical threshold force required to elicit responses 50% of the time and (d) thermal threshold latency required to tail flick for the remifentanil and sufentanil groups. All data are means ± SEM (n = 8). *p < 0.01, **p < 0.001, ***p < 0.0001 versus saline group, and analyzed by Two-way ANOVA with Bonferroni post hoc test. (e,f) The miRNAs up-regulated or down-regulated by more than 2-fold in the remifentanil (top 5 changed miRNAs listed) and sufentanil groups compared to the saline group (p < 0.05). (g) Heatmap highlights most significantly changed miRNAs among three different groups (n = 3). (h, i, j) miRNA relative expression validated by qPCR. *p < 0.01, **p < 0.001, ***p < 0.0001 versus saline group, and analyzed by Two-way ANOVA with Bonferroni post-hoc test. All data are means ± SEM (n = 6).

Guided by previous reports that pronociceptive effects of remifentanil are determined by dose rather than by infusion duration, ²² we next examined whether this dose-response effect exists in our remifentanil and sufentanil consecutive injection model. As expected, remifentanil induced dose-dependent pronociceptive effects in von Frey and tail immersion tests, with calculated ED₅₀ of 10.66 (95% CI, 10.06–11.17) and 9.03 (8.23–10.01) μg/kg respectively, which lasted longer with higher doses (F (4, 35) = 119.7, p < 0.0001, and F (4, 35) = 212.1, p < 0.0001, two-way ANOVA, sFigure 1(a) and (b)). As a comparison, calculated pronociceptive ED₅₀ of mechanical and thermal hyperalgesia after RED-sufentanil injection were 3.14 (95% CI, 2.78–3.59) and 2.72 (2.42–3.11) μg/kg respectively (F (4, 35) = 133.9, p < 0.0001, and F (4, 35) = 205.6, p < 0.0001, two-way ANOVA, sFigure 1(c) and (d)), which are much higher than its antinociceptive ED₅₀ (0.468 μg/kg). Collectively, our detailed behavioral tests identify that remifentanil possesses significant pronociceptive effects than sufentanil in mice.

Identification of differentially expressed miRNAs after remifentanil or sufentanil exposure in the spinal cord

To identify the spinal modulator critical for the different pain phenotypes induced by the two opioids, we analyzed the differential expression profiles of miRNAs in the SDH of remifentanil and RED-sufentanil exposure mice 1 day after injection, when remifentanil mice showed significant hyperalgesia, but sufentanil mice showed normal responses to painful stimuli. After sequencing the control and opioid-treated L3-L5 spinal cord tissues using an Illumina HiSeq high-throughput sequencing technique, we found that the remifentanil group showed 20 significantly overexpressed miRNAs and 10 down-regulated miRNAs (fold change>2.0, p < 0.05, Figure 1(e)), compared to saline-injected mice. For the differentially expressed miRNAs in sufentanil group, 15 were upregulated and 21 downregulated in SDH as compared with control mice (Figure 1(f)). Venn diagrams reveal the overlap of differential miRNAs between the opioid groups as compared with the control group. Only one intersectional upregulated-microRNAs and five intersectional downregulated-microRNAs are observed, indicating that miRNA expression profiles vary markedly as a function of different opioid treatments. In the heatmap of 30 differentially expressed microRNAs in the remifentanil group, miR-134-5p showed the second most significant decrease in the remifentanil-treated group, while retaining no statistical difference in sufentanil mice (Figure 1(g)). Given to previous research demonstrated the neuronal miR-134-5p decreased in other nociceptive models, we speculated the miR-134-5p remains functional regulatory role in RIH.

Furthermore, the expression changes in this set of miRNAs were validated by qPCR in another series of control and opioid-treated mice. Among the 10 miRNAs showing significant expression changes, seven were confirmed by qPCR analysis as showing more than 50% level-changes in the spinal cord tissue of remifentanil mice ((F (2, 15) = 44.75, p < 0.0001, and F (2, 15) = 16.46, p < 0.0001, two-way ANOVA, Figure 1(h) and (i)). These results indicate that the real-time qPCR data were consistent with that of the miRNA-seq analysis regarding expression levels of these seven miRNAs. Consequently, we further postulate that miR-134-5p might be involved in the significant pronociceptive effects of remifentanil.

Evaluation of spinal miR-134-5p expression, distribution, and function in mice injected with remifentanil

The time course of spinal miR-134-5p expression changes was validated by real-time qPCR. The miR-134-5p expression robustly decreased at 6 h after remifentanil injection and maintained until day 5 (F (1, 10) = 277.1, p < 0.0001, two-way ANOVA, Figure 1(j)). Meanwhile, miRNA-specific in situ hybridization was further used to label and confirm the changes in miR-134-5p expression in the spinal cord. The image showed decreased miR-134-5p positive signals after remifentanil exposure, while largely unchanged in the RED-sufentanil group (Figure 2(a)). Further combined with immunofluorescence staining confirmed that miR-134-5p was highly expressed in spinal dorsal neurons (NeuN⁺) and rarely colocalized with microglia (Iba-1⁺) or astrocytes (GFAP⁺) after remifentanil treatment (Figure 2(b)). Additionally, the level of miR-134-5p did not differ in L3-L5 dorsal root ganglion (DRG) (Figure 2(c)).

Figure 2.

Remifentanil inhibits the total and functional miR-134-5p expression in SDH neurons. (a) Representative FISH images of miR-134-5p expression in the L3-L5 SDH of different groups on day 2 after exposure (n = 3), Scale bar: 100 μm. (b) Cell-specific localization of miR-134-5p, combined miR-134-5p (red) ISH and subsequent immunofluorescence for NeuN (left, green) or GFAP (middle, green) or Iba1 (right, green) in SDH sections from remifentanil mice (n = 3), Scale bar: 100 μm. (c) Representative FISH images of miR-134-5p expression levels in L3-L5 peptidergic CGRP-expressing DRG neurons of remifentanil mice on day 2 after the i.v. injection (n = 3), Scale bar: 100 μm. (d) Representative bond for Argonaute-2 (Ago2) protein from saline-treated mice and mice 6 h (left) or 24 h (right) after remifentanil/sufentanil exposure. Input, total lysates; IP, immunoprecipitates. (e) Quantification of total expression of Ago2 protein in SDH total lysates of opioids- or saline- treated mice. (f) Recruitment of miR-134-5p by Ago2 to miRISCs in SDH from control mice and opioid exposure mice. ***p < 0.0001 versus saline group, and analyzed by Two-way ANOVA with Bonferroni post hoc test. All data are means ± SEM (n = 6).

Given these observations of a specific decreased expression of spinal total miR-134-5p on developmental consistency to RIH, we further investigated if spinal functional miR-134-5p, which binds to the Argonaute2 (Ago2) protein and directly inhibits its complementary mRNAs, ⁹ was synchronously downregulated after remifentanil treatment. The results showed no significant differences in Ago2 expression in total lysates (input) nor in Ago2 immunoprecipitation at 6 h or 1 day after opioids exposure (F (2, 15) = 0.1267, p = 0.8819, two-way ANOVA, Figure 2(d) and (e)). The level of miR-134-5p bound to Ago-2 in the control mice was the highest, whereas the level was much lower in the remifentanil group. However, the administration of RED-sufentanil did not reduce functional miR-134-5p expression compared with the control group (F (2, 30) = 127.8, p < 0.0001, two-way ANOVA, Figure 2(f)). Taken together, our results suggest that acute remifentanil exposure could induce down-regulated expression in spinal miR-134-5p of neurons in mice.

Pharmacological targeting of spinal miR-134-5p regulates remifentanil-induced hyperalgesia

To specifically assess the importance of miR-134-5p in the pathogenesis of RIH, three intrathecal injections of AgomiR-134-5p or scrambled sequences as control were used to upregulate the expression level of spinal miR-134-5p on days −3, −2, and −1 prior to remifentanil exposure (Figure 3(a)). The qPCR and FISH results showed that Agomir-134-5p intrathecal pretreatment prevented the downregulation of miR-134-5p at days 1, 3, and 5 after remifentanil exposure and maintained on day 5 (F (2, 45) = 520.6, p < 0.0001, two-way ANOVA, and F (2, 6) = 41.74, p = 0.0003, one-way ANOVA, Figure 3(b) and (c)). Locomotor and remifentanil antinociception were insusceptible to pharmacological modulation of miR-134-5p in mice (F (2, 21) = 0.1878, p = 0.8301, and F (2, 21) = 0.8408, p = 0.4454, one-way ANOVA, Figure 3(d) and (e)). Pain behavioral tests showed that the onset of mechanical allodynia (F (2, 21) = 17.86, p < 0.0001, two-way ANOVA, Figure 3(f)) and thermal hyperalgesia (F (2, 21) = 53.75, p < 0.0001, two-way ANOVA, Figure 3(g)) were delayed, and the severity was reduced, with a dose-dependent effect in mice pretreated with 0.5 nmol or 1 nmol AgomiR-134-5p, compared with mice that underwent remifentanil and scrambled sequences injection. To further reveal the role of miR-134-5p in modulating pain hypersensitivity, we injected AntagomiR-134-5p or a scramble into the lumbar enlargement of naïve mice. AntagomiR-134-5p induced spontaneous pain, mechanical allodynia, and thermal hyperalgesia from 3 h to 9 h after injection in naïve mice (Data not shown). These behavioral findings collectively support that miR-134-5p contributes to nociceptive behaviors after remifentanil exposure.

Figure 3.

miR-134 mimics prevents and reverses hyperalgesia in mice subjected to remifentanil. (a) Intrathecal administration protocol of AgomiR-134 or scramble sequence, followed by behavioral testing and experiments at the indicated time points. (b) miR-134 relative expression in the SDH in remifentanil ± AgomiR-134 group on day 1, 3, 5 after i.v. injections. *p < 0.01, **p < 0.001, ***p < 0.0001 versus remifentanil group, and analyzed by Two-way ANOVA with Bonferroni post hoc test. All data are means ± SEM (n = 6). (c) Representative fluorescence ISH signals showing effects of AgomiR-134 pre-treatment on miR-134 expression within the SDH 1 day after remifentanil exposure, Scale bar: 100 μm. (d) Measurement of locomotor function in an accelerating rotarod test after the last AgomiR-134-5p administration. (e) Antinociceptive MPE for remifentanil antinociception from comparing first and last AgomiR-134-5p administration (n = 8). ***p < 0.0001 versus remifentanil group and analyzed by One-way ANOVA with Bonferroni post-hoc test. All data are means ± SEM (n = 8). (f, g) Time course of the mechanical allodynia and thermal hyperalgesia in scramble-treated and AgomiR134-5p-treated mice with RIH. *p < 0.01, **p < 0.001, ***p < 0.0001 versus remifentanil group, and analyzed by Two-way ANOVA with Bonferroni post-hoc test. All data are means ± SEM (n = 8).

Identifying miR-134-5p target genes

Prediction through bioinformatics databases, including miRSearch, miRDB, HOCTAR, DIANA, and miRWalk, 3 of them intersected and indicated that the Glutamate Receptor Ionotropic Kainate 3 (Grik3) gene encoding GRIK3, which is implicated in excitatory synaptic potentiation and mainly abundant in the murine spinal dorsal horn, might be the target of miR-134-5p (Figure 4(a)).^23,24 The results showed an exact match between them in Grik3 3’-UTR 7462-7481 bp downstream from the stop codon. Notably, the target sites of Grik3 genes are conserved between humans and rodents (Figure 4(b)). Furthermore, the binding sites of mmu-miR-134-5p on the target gene (Grik3) in the 3‘-UTR region and the inhibitory effect were detected by the luciferase assay. The assay showed that mmu-miR-134-5p could regulate the luciferase activity with 3’-UTR in Grik3, with a reduction of 58.76%. However, after mutation of the binding site, this regulatory relationship was significantly weakened (Figure 4(c)), with a rebound of 19.45%, confirming that mmu-miR-134-5p directly targets the 3’-UTR regions of Grik3 mRNA and negatively modulates Grik3 expression. Nevertheless, mmu-miR-134-5p still had a mild regulatory effect on the luciferase activity in Grik3-mutated 3’-UTR (Figure 4(c)), indicating the presence of other atypical binding sites.

Figure 4.

miR-134 targets Grik3 and pharmacological inhibition of GRIK3 induces anti-hyperalgesic effects. (a, b) miR-134-5p directly targets the Grik3, the predicted overlapping binding pairing sequence (red) alignment of miR-134-5p in mouse and human Grik3 3′UTR. (c) Activity of luciferase with the Grik3 3′-UTR or mutation in the 3′-UTR in HEK293T cells co-transfected with control microRNA or a miR134-5p mimics. The Renilla/firefly activity values were used for normalization and analyzed by Student’s t test, ***p < 0.001 versus control group. All data are means ± SEM (n = 6).(d, e) Spinal Grik3-mRNA and GRIK3-protein relative expression in mice underwent remifentanil with AgomiR-134 or Scramble sequence injection. All the data are means ± SEM (n = 6) and analyzed by one-way ANOVA with Bonferroni post hoc comparisons. *p < 0.01, ***p < 0.001 versus saline. (f, g) Time course of the mechanical allodynia and thermal hyperalgesia in vehicle-treated and GRIK3 antagonists ACET-treated mice with RIH. *p < 0.01, **p < 0.001, ***p < 0.0001 versus remifentanil group, and analyzed by Two-way ANOVA with Bonferroni post-hoc test. All data are means ± SEM (n = 8). (h) Representative western blotting bonds for GRIK3 membrane trafficking after remifentanil/ACET exposure. ***p < 0.0001 versus remifentanil group, and analyzed by Two-way ANOVA with Bonferroni post-hoc test. All data are means ± SEM (n = 6).

Subsequently, we investigated whether the expression level of Grik3 is regulated by miR-134-5p in vivo. AgomiR-134 (1 nM) or scramble was intrathecally injected into the mice before a 3-day remifentanil exposure. We found significant inhibitory effects of miR-134-5p overexpression on Grik3 mRNA expression levels, and increased mRNA expression of Grik3 by remifentanil exposure was also repressed by miR-134-5p overexpression (F (3, 20) = 17.10, p < 0.0001; one-way ANOVA, Figure 4(d)). At the protein level, miR-134-5p overexpression did not affect the expression of Grik3 in control mice, whereas protein levels were significantly reduced by miR-134-5p overexpression in remifentanil-treated mice compared with both control and hyperalgesic mice injected with scramble. In addition, remifentanil-induced up-regulation of Grik3-protein was blocked by miR-134-5p overexpression (F (3, 20) = 9.014, p = 0.0006; one-way ANOVA, Figure 4(e)). Overall, these results suggest that miR-134-5p contributes to the remifentanil induced Grik3 down-regulation, but not basal expression levels.

Pharmacological inhibition of GRIK3 induce antiallodynic-like behavioral responses

We subsequently investigated the antiallodynic effect of ACET, a potent and selective kainate receptor antagonist, in our current RIH model. We intrathecally injected ACET into the mice before remifentanil exposure, and then measured mechanical and thermal pain behaviors. Remifentanil exposure mice infused with ACET (100 nM) showed increased PWT (F (2, 21) = 16.85, p < 0.0001; two-way ANOVA, Figure 4(f)) and longer latency to hot water in tail immersion (F (2, 21) = 32.31, p < 0.0001; two-way ANOVA, Figure 4(g)) compared with vehicle-infused remifentanil mice. Furthermore, the immunoblotting results demonstrated the membrane trafficking of spinal GRIK3 was increased after remifentanil exposure, and it was blocked by ACET (F (2, 15) = 20.95, p < 0.0001; one-way ANOVA, Figure 4(h)). These results suggest that the inhibition of GRIK3 can block pain-like behaviors induced by remifentanil via limiting GRIK3 transportation.

miR-134-5p modulates spinal dendritic spine remodeling in mice subjected to remifentanil-induced hyperalgesia

To determine the potential roles of miRNAs in hyperalgesia progression, we performed GO analysis using the STRING system. DEGs in remifentanil mice were significantly involved in regulating glutamatergic synapses (Figure 5(a)), whereas these miRNAs were associated with both glutamatergic synapses and GABAergic synapses in sufentanil mice (Figure 5(b)), indicating that remifentanil mainly targets the regulation of glutamatergic synapses. Indeed, miR-134-5p was previously recognized as the modulator of synaptic plasticity and crucial for the functional strengthening of glutamate receptor-mediated excitatory synapses,^25,26 which may contribute to the hyperalgesia observed in remifentanil mice.

Figure 5.

miR-134 is involved in remifentanil-induced augment of spine dendritic morphology. (a, b) Terms showed DEGs between the miRNAs and control probes in remifentanil group or sufentanil group. (c) Representative Golgi staining images of dendritic spines from SDH neurons, Scale bar:10 μm. (d) Convex hull analysis of dendritic complexity and dendritic morphology. (e) Schematic diagrams of thin, mushroom, and stubby dendritic spines respectively. (left) The changes in dendritic spine density of various types (right). **p < 0.001, ***p < 0.0001 versus saline group or remifentanil group, analyzed by One-way ANOVA with Bonferroni post-hoc test. All data are means ± SEM (n = 3 sections).

Subsequently, we used the Golgi staining analysis to confirm the functional effects of miR-134-5p on spinal dendritic outgrowth and morphology changes in RIH mice. This was done using mice from the previous AgomiR dose-response studies. Compared with the control mice, there was significant augmentation in the dendritic outgrowth and complexity in remifentanil-treated mice (F (3, 20) = 36.61, p < 0.0001, and F (3, 20) = 61.55, p < 0.0001, one-way ANOVA, Figure 5(c) and (d)). Morphology and number of dendritic spines were involved in the synaptic plasticity changes under pain maintenance, the overall density of dendritic spines increased after remifentanil treatment. Besides, there was significant contribution of remifentanil to the shapeshifting of dendritic spines of SDH neurons. The density of thin and mushroom dendritic spines, which have been explored to play an important role in neuron activity, increased significantly (Figure 5(e)). Strikingly, miR-134-5p pre-treatment exhibited an inhibitory effect on remifentanil-induced dendritic spine rearrangement, Agomir-134 (i.t.) reversed the increase in dendritic spine density and shapeshifting (F (3, 20) = 68.91, p < 0.0001, two-way ANOVA, Figure 5(e)).

miR-134-5p modulates spinal synaptic plasticity in mice subjected to remifentanil-induced hyperalgesia

The abnormal dendritic spine structure we observed could induce aberrant synaptic structure. To pursue this speculation, EM was employed for detailed analysis of the ultrastructure of the asymmetric (excitatory) synapses from different groups within the spinal dorsal horn (Figure 6(a)). Morphometric analysis of asymmetric synapses at the SDH of remifentanil mice showed that all the synaptic parameters varied significantly between remifentanil mice and control mice (Figure 6(b)). In remifentanil mice, asymmetric synapses had thicker and longer PSDs, smaller synaptic cleft distances, and more presynaptic vesicles (Figure 6(b)). As expected, miR-134-5p overexpression reversed the excitatory synapses structural plasticity in mice subjected to RIH (F (3, 20) = 476.4, p < 0.0001; F (3, 20) = 31.18, p < 0.0001; F (3, 20) = 96.49, p < 0.0001; F (3, 20) = 12.56, p < 0.0001, F (3, 20) = 86.50, p < 0.0001; one-way ANOVA, Figure 6(b)).

Figure 6.

miR-134 is involved in remifentanil-induced augment of excitatory synaptic plasticity. (a) Representative EM images of asymmetric synapses in SDH on day 4 after remifentanil i.v. injections. Green indicates presynaptic area and pink outlines postsynaptic area, Scale bar: 500 nm. (b) Quantification of morphological synaptic parameters of the asymmetric synapses in SDH: presynaptic area, total number of synaptic vesicles, synaptic cleft distance, PSD length, and PSD thickness at day 4. *p < 0.01, **p < 0.001, ***p < 0.0001 versus saline group or remifentanil group, analyzed by One-way ANOVA with Bonferroni post-hoc test. All data are means ± SEM (n = 3 sections). (c) Representative traces of KAR–mediated miniature excitatory postsynaptic currents in the dorsal horn neurons of spinal cord slices after remifentanil incubation (4 nM) and the effects of miR-134-5p mimics (Agomir-134, 5 µM). (d, e) The amplitude (d) and interevent interval (e) of miniature excitatory postsynaptic currents in the spinal dorsal horn neurons under different conditions were evaluated. ***p < 0.0001 versus saline group or remifentanil group, analyzed by one-way ANOVA with Bonferroni post-hoc comparisons. All data are means ± SEM (n = 6 sections).

Given the findings of significant excitatory synapses structural plasticity changes, the whole-cell patch-clamp recordings were used to confirm the functional changes. The representative traces of Kainate receptor-mediated mEPSCs in the spinal cord dorsal horn neurons after different interventions are shown in Figure 6(c). Both the frequency and the amplitude increased in remifentanil exposed mice. Strikingly, Agomir-134 application apparently inhibited the increased amplitude and frequency of current induced by remifentanil (F (2, 39) = 386.4, p < 0.0001, and F (2, 39) = 118.8, p < 0.0001, one-way ANOVA, Figure 6(c) and (d)). These results show that miR-134-5p is an important regulator of the morphology and plasticity of excitatory synapses in the development of RIH.

Discussion

Our findings demonstrate that remifentanil could induce more significant hyperalgesia, compared to sufentanil with an equivalent analgesic dose in mice. Diminished spinal miR-134-5p mediates this distinct pronociceptive effect and contributes to the changes in the dendritic structure and synaptic plasticity of spinal cord neurons, which are essential for the generation of remifentanil induced central sensitization and pain hypersensitivity. Besides, the regulatory role of miR-134-5p is partly achieved via targeting Grik3 mRNAs, and eventually potentiating the kainite receptor mediated mEPSCs. Moreover, pharmacological inhibition of GRIK3 had anti-allodynic effects. Collectively, our findings provide evidence that decreasing spinal miR-134-5p resulted in a state of RIH deterioration and synaptic plasticity excess via partly ensuing upregulation of Grik3 (Figure 7). Targeting at miR-134-5p and GRIK3 may shed light on the management of RIH.

Figure 7.

Schematic illustration for the proposed mechanism of remifentanil-induced hyperalgesia regulation by miR-134-5p/GRIK3 pathway. Left panel: remifentanil sets up a cascade of events that lead to decreased levels of miR-134-5p and increased expression of GRIK3, which ultimately causes opioid induced hyperalgesia. Right panel: detailed cascade reaction in spinal synapse after treatment with remifentanil, the procedure that mature miR-134-5p generated and assembled into the RISCs was inhibited, this leads to overexpression of complementary mRNAs (Grik3). Increased expression of GRIK3 was transported to postsynaptic membrane, which triggers structural and functional modifications of excitatory synapses, and finally induced dendritic spine rearrangement and pain exacerbation. RISC=RNA-induced silencing complex.

In comparison to other synthetic opioids, remifentanil has a much higher incidence of acute opioid induced hyperalgesia. Several distinct mechanisms have been proposed to initiate the different pronociceptive effects among clinically used opioids with different pharmacological profiles. One previous study revealed that morphine, fentanyl, and remifentanil all enhance synaptic transmission at spinal C-fibers but via fundamentally different mechanisms during OIH. Compared to remifentanil, fentanyl and morphine may additionally activate descending, facilitatory, and serotonergic pathways via extraspinal μ-opioid receptors (MORs). ²⁷ Yet, in another study, hyperalgesia was only induced by chronic administration of fentanyl, but not by morphine, and this distinction is attributed to their different efficiencies in inducing NLRP3-dependent neuroinflammation. ²⁸ Strikingly, the bias caused by properties that are typical for morphine but not for fentanyl cannot be excluded in these studies, such as the broad opioid receptor binding profile, the low potency to induce MOR internalization, and the production of active metabolites.^29–31 Thus, we employed two fentanyl derivatives with similar profiles, remifentanil and sufentanil in our current research. Even short-term remifentanil exposure can cause severe and long-lasting hyperalgesia, and the doses of antinociception (9.282 μg/kg) is very close to the levels for mechanical/thermal hyperalgesia priming (10.66/9.03 μg/kg). Nonetheless, sufentanil induced quite short hyperalgesia (less than 6 h) within the equivalent analgesic dose. Remarkably, the dose of antinociception (0.468 μg/kg) was much lower than the levels for mechanical/thermal hyperalgesia priming (3.14/2.72 μg/kg). These findings from murine models are consistent with clinical observations that repeated opioid administration induces analgesic tolerance and hyperalgesia, depending both on the opioid employed and on the administration schedule.^32–34

Structural plasticity of dendritic spines occurs alongside changes in synaptic plasticity, which requires modifications to the specific kinds of excitatory glutamatergic receptors, including the posttranscriptional process mediated by miRNAs. Notably, the brain-neuronal miRNAs could regulate dendritic spine rearrangements and may thus participate in regulating neural plasticity and behaviors. In the hippocampus, miR-134 is abundantly expressed in the dendrites of neurons/synapses, where it inhibits synaptic functional and structural development in an activity-dependent manner. ³⁵ However, whether spinal miR-134-5p retains the same role in mice with OIH is yet to be clarified. In our and others’ previous work, enhanced synaptic transmission in SDH was proven to be critical for RIH.^36,37 Several in vitro studies reported that downregulation of miR-134 in hippocampal neurons dramatically increased synaptic plasticity, and this role may be achieved through upregulating the membrane trafficking of AMPA receptor subunits GluA2. ³⁸ Likewise, we observed the spinal miR-134-5p deficits and resulting dendritic remodeling and the increased density of mushroom spines in SDH from RIH mice. Besides, the PSD length and the thickness of excitatory synapses in the SDH increased, synaptic cleft narrowed, and the presynaptic space widened. Strikingly, artificially upregulating spinal miR-134-5p reversed remifentanil-induced changes in the number, morphology, and distribution of dendritic spines. Thus, our structural evidence indicates that the downregulation of spinal miR-134-5p pathways is a causal mechanism underlying RIH. However, here is no real-time evidence for synaptic spines in controlling the cellular physiology of RIH states and subsequent pain behaviors. Studies using novel cutting-edge tools such as 2-photon in vivo imaging may uncover the mechanistic relationships between miRNAs-related synaptic dysfunction and pain-like behaviors in RIH.

Our bioinformatics analyses suggested that glutamatergic synapse gene pathways were most likely to be targeted by altered miRNAs in RIH. Accordingly, miR-134-5p was found to directly bind and suppress the expression of Grik3 transcripts. Consistent with the in vitro data, this inverse relationship was validated between miR-134-5p and Grik3 transcripts in mice subjected to remifentanil injection. Cumulative studies support that the excitatory glutamate receptor alterations initiate the cascade reactions of the occurrence and development of OIH.^39,40 Kainate receptors (KA-R) are a subfamily of ionotropic glutamate receptors and serve important roles in mediating excitatory synapse transmission in the spinal cord dorsal horn. ⁴¹ These receptors have been implicated in the pathogenesis of several neurological diseases, such as stroke, Alzheimer's disease, epilepsy, and neuropathic pain. ⁴² Glutamate Receptor Ionotropic Kainate 3 (GRIK3), an important excitatory glutamate receptor, mainly participates in a range of neuroactive ligand-receptor interaction pathways. Accumulating evidence has indicated that GRIK3 is highly expressed in the superficial spinal dorsal horn and is directly involved in the pain pathway, ²⁴ especially in neuropathic pain after spinal cord injury. ⁴³ Moreover, KA-R regulates synaptic transmission at both excitatory and inhibitory synapses in the spinal cord dorsal horn.^44–46 Here we confirmed that both the amplitude and frequency of KA-R-mediated mEPSCs were increased in remifentanil group, miR-134-3p could partly reverse remifentanil induced mEPSCs amplification. Intrathecal injection of ACET, a potent and selective KA-R antagonist, was able to reverse the GRIK3 membrane trafficking and relieved RIH. Therefore, our present study was novel in revealing the changes and potential clinical significance of the kainate receptors GRIK3 in RIH.

Although our current research mainly focused on miR-134-5p, our high-throughput sequencing identified many other miRNAs that may also be involved in RIH pathogenesis and maintenance. Some significant differential expressed miRNAs, including miR-539, miR-500, and miR-466, have already been shown to play an important role in neuropathic pain.^47–49 It should be noted, however, that other miRNAs not selected as candidates here may also be associated with RIH if they influence the expression of highly pronociceptive-related genes. Other strengths of this study include the caudal intravenous injection route, which can provide more stable blood concentration and is more similar to clinical practice, considering individual differences in absorption ability and variations in blood concentration. This study has several limitations. First, the ED₅₀ values of the pronociceptive and antinociceptive effects of opioids may be underestimated, as higher doses were avoided to prevent the accompanying respiratory and circulatory depression. Second, we did not detect sex differences in the development of enduring opioid-induced hyperalgesia on both remifentanil and sufentanil, although different pain phenotypes and underlying mechanisms of OIH across sexes have been previously reported.^50,51 Third, this study focused on the experimental validation of only a subset of the many predicted targets of miR-134-5p, one of the glutamatergic synapse pathways. However, it is possible that other transcripts predicted to be regulated by miR-134-5p are also important to the pathogenesis of RIH, such as MOR and Twist1.^52,53

Taken together, our current study proposes that miR-134-5p contributes to the initiation and maintenance of RIH by fine-tuning the spinal dendritic structure and synaptic plasticity. Our detailed descriptions of the miR-134-5p-associated gene networks involved in remifentanil-induced epigenetic gene regulation, dendritic spine remodeling, and behavioral changes may reveal new aspects of RIH pathophysiology. Moreover, this is the first rodent study of spinal miRNAs-profiles in the development of RIH to date and strongly supports that opioid application alters miRNA expression. At large, these findings indicate that direct miRNA modulation can be a promising preventive and therapeutic strategy. Remarkably, the dose remained within the clinical dosing range after conversion between species, which provides rationale for pursuing further pre-clinical and clinical studies of intraoperative remifentanil and RED-sufentanil on sensory processing. Additionally, our data supports the development of therapeutic strategies for disrupting spinal kainate receptor signaling to maintain adequate long-lasting pain relief while limiting potentially harmful opioid induced hyperalgesia. The design of clinical trials aimed at rigorously evaluating the translational potential of kainate receptor antagonist combinations in appropriate patients with opioid-responsive pain conditions will be essential to the development of safer and more effective pain management strategies.

ORCID iD

Guolin Wang https://orcid.org/0000-0001-7867-2091