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There is increasing interest in RNA interference in pain research using the intrathecal route to deliver small-interfering RNA (siRNA). An interferon (IFN) response is a common side-effect of siRNA. However, the IFN response in the spinal cord after intrathecal administration of siRNA remains unknown. We hypothesized that high doses of siRNAs can elicit off-target analgesia via releasing IFN-α. We investigated the IFN response and its role in regulating pain sensitivity in the spinal cords after intrathecal administration of siRNAs.
Male Sprague–Dawley rats were given intrathecal injections of non-targeting (NT) siRNAs or IFN-α and tested for complete Freund's adjuvant (CFA)-induced mechanical allodynia and heat hyperalgesia. IFN-α in the spinal cord after injection of NT siRNAs was measured by western blotting and immunohistochemical staining.
IFN-α was up-regulated in the spinal cord after intrathecal treatment of NT siRNAs. Intrathecal injection of NT siRNAs, at high doses of 10 or 20 μg, reduced CFA-induced inflammatory pain (P<0.05). Intrathecal application of IFN-α inhibited pain hypersensitivity in inflamed rats and produced analgesia in naïve rats (P<0.05). Notably, the anti-nociceptive effects elicited by NT siRNAs and IFN-α were reversed by IFN-α neutralizing antibody and naloxone.
Our data suggest that (i) intrathecal administration of high doses of siRNA (≥10 μg) induced up-regulation of IFN-α in the spinal cord and produced analgesic effects through IFN-α, and (ii) IFN-α's analgesic effect is mediated via opioid receptors. Caution must be taken to avoid IFN-α-mediated analgesic effects of siRNAs in pain research.
Interferons (IFNs) were discovered as natural anti-viral substances produced during viral infections and were initially characterized for their ability to ‘interfere’ with viral replication, slow cell proliferation, and profoundly alter immunity. There are two major types of IFNs: type-I contains IFN-α and IFN-β, two of the major members, and also IFN-ω, IFN-τ, IFN-δ, IFN-κ, and IFN-, whereas type-II contains only IFN-γ.1 Recent studies showed that IFN-γ was produced by astrocytes in the central nervous system (CNS) and enhanced neuropathic pain by stimulation of IFN-γ receptors specifically expressed in spinal microglia.2,3
RNA interference is widely used to study gene function and develop new therapeutic reagents.4,5 This strategy was also explored for the treatment of persistent pain.6,7 Effective delivery of small-interfering RNAs (siRNAs) to the CNS remains to be a challenge. Systemic delivery of unmodified siRNAs to the CNS is currently precluded, since siRNAs do not readily cross the brain–blood barrier and are rapidly degraded in vivo by endonucleases. However, intrathecal (spinal) injection of siRNAs into the spinal fluid, together with several delivery-assisting reagents such as lipofectamine, iFect, and polyethyleneimine (PEI), has been shown to knockdown gene expression in the dorsal root ganglion and spinal cord and modulate pain sensitivity.6–10 Of note, siRNAs have been shown to induce type-I IFN responses.11,12 It is virtually unknown whether siRNA can trigger IFN-α responses in the spinal cord to regulate pain sensitivity. We examined whether intrathecal siRNAs could induce IFN-α expression in the spinal cord and whether siRNA could modulate pain via IFN-α.
To avoid knocking down the expression of specific genes, we used non-targeting (NT) siRNA and RNA-induced silencing complex (RISC)-free siRNA. Because GU-rich siRNA was shown to induce profound IFN-α expression in immune cells,11 we also tested the effects of GU-rich siRNA on inflammatory pain. NT siRNA, GU-rich siRNA,11 and RISC-free siRNA were synthesized, purified, and annealed by Dharmacon Research Incorporation (Lafayette, CO, USA). An NIH-blast search was conducted to ensure that no gene was being targeted for NT siRNA. Based on these sequences, the following double-stranded siRNAs were carefully designed. The effects and sequences of these siRNA are described in Figure 1a and and11b. The siRNAs were dissolved in RNase-free water at 1 μg μl−1 as stock solutions. The siRNA was mixed with PEI (Fermentas Inc., Glen Burnie, MD, USA), 10 min before injection, to increase cell membrane penetration and reduce degradation.10 PEI was dissolved in 5% glucose. Relative amounts of RNA to carrier were as follows: six equivalents of PEI nitrogen per RNA phosphate, which was 0.18 μl of PEI solution per microgram of RNA. The final injection solution of siRNA (0.25 μg μl−1) contained 5% sucrose.
Experiments were performed on adult male Sprague–Dawley rats (200–260 g). The experiments followed the Ethical Guidelines of the International Association for the Study of Pain13 and ARRIVE guidelines.14 All animal procedures performed in this study were approved by the Animal Care Committee of Harvard Medical School, Boston, MA, USA.
For a single intrathecal injection, the spinal cord was punctured under brief sevoflurane anaesthesia using a 30 G needle between L5 and L6 of the spinal column to deliver the reagents (40 μl) into the cerebral spinal fluid. Immediately after the needle entry into the subarachnoid space (change in resistance), a brisk tail-flick was observed after the needle puncture.15
To produce inflammatory pain, we injected 100 μl of complete Freund's adjuvant (CFA) into a hind paw of a rat. To test the effect of siRNA on CFA-induced allodynia, six groups of rats (n=6 in each group) received intrathecal injection of 5, 10, or 20 μg of NT-siRNA, 10 μg of naked NT-siRNA, 10 μg of RISC-free siRNA, or 10 μg GU-rich siRNA (per group), 1 day before injection of 100 μl CFA into the left hind paw of rats. Two control groups of rats (n=6 each group) received intrathecal injections of either 1.8 μl of PEI or 40 μl of saline 1 day before CFA injection. Mechanical allodynia was tested at 6 and 24 h after injection of CFA. We also tested the effect of NT-siRNA on CFA-induced heat hyperalgesia. Heat hyperalgesia was tested at 6 h after intrathecal injection of 10 μg NT-siRNA and at 24, 48, and 72 h after injection of CFA (n=8 each group).
To test the analgesic effects and dose–responses of IFN-α, four groups of rats (n=6 in each group) received intrathecal injection of 10, 25, or 100 ng of IFN-α 1 day after injection of CFA. The control group of rats (n=6) received intrathecal injection of 40 μl saline 1 day after injection of CFA in the left hind paw. Paw withdrawal threshold was recorded before and 1 day after injection of CFA. Then, mechanical allodynia was tested at 30 min, 1, 2, 4, and 24 h after injection of 10, 25, or 100 ng of IFN-α or saline (control). Heat hypersensitivity was tested at 1 h after intrathecal injection of 100 ng IFN-α or 40 μl saline without further injection of CFA (n=6). To test for the reversing effect of IFN-α neutralizing antibody on the analgesic effect produced by siRNA or IFN-α, the rats received intrathecal injections of 10 μg of NT-siRNA (n=5) or 100 ng IFN-α (n=6), and injection of CFA in the left hind paw 1 day before injection of the IFN-α antibody. Mechanical allodynia was tested 30 min, 1, and 3 h after intrathecal injection of 30 ng of IFN-α antibody. Similarly, to test for the reversing effect of naloxone on the analgesic effects produced by siRNA or IFN-α, the rats received intrathecal injection of 10 μg NT-siRNA (n=5) or 100 ng IFN-α (n=6) and injection of CFA in the left hind paw 1 day before injection of 20 nmol of naloxone. Mechanical allodynia was tested 30 min, 1, 2, and 4 h after intrathecal injection of 20 nmol of naloxone.
For western blotting of IFN-α (n=6 in each group), the L4–5 spinal cord segments were harvested 1 day after intrathecal injection of 10 μg NT-siRNA and intradermal injection of CFA into the left hind paws of the rats. The control group of rats received intrathecal injection of 1.8 μl PEI.
The rats were killed by decapitation under deep anaesthesia (intraperitoneal pentobarbital 120 mg kg−1). The L4–5 spinal cord segments were quickly removed after the rat was decapitated, and homogenized with a hand-held pellet pestle in T-PER Tissue Protein Extraction Reagent (Pierce, Rockford, IL, USA) [25 mM bicine, 150 mM sodium chloride (pH 7.6)] containing protease inhibitors (Protease Inhibitor Cocktail, Calbiochem, Darmstadt, Germany) [100 mM 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride, 80 μM aprotinin, crystalline, 5 mM bestatin, 1.5 mM E-64 protease inhibitor, 2 mM leupeptin, and 1 mM pepstatin A]. After being placed on ice for 30 min, the homogenate was centrifuged at 10 000g for 15 min at 4°C. The supernatant was collected and assayed for protein content using the Bio-Rad DC Protein Assay Kit (Bio-Rad Laboratories, Hercules, CA, USA) and stored at −20°C until further use. Protein samples (30 μg per lane) were boiled under denaturing conditions for 10 min, separated on sodium dodecyl sulphate–polyacrylamide gel electrophoresis gels (5–10% gradient gel) with running buffer and molecular weight standards, as suggested by the manufacturer. Finally, they were transferred to polyvinylidene difluoride filters (Immobilon-P, Millipore, Bedford, MA, USA). After protein transfer, the polyvinylidene difluoride membranes were blocked with 5% non-fat milk in phosphate-buffered saline (PBS) for 1 h and then incubated overnight at 4°C with polyclonal antibody against IFN-α (1:500, R&D Systems Inc., NC, USA) and mouse monoclonal anti-β-tubulin (1:2000, Santa Cruz Biochemicals, Inc., Santa Cruz, CA, USA) (diluted) in 0.1% Tween 20–PBS. The membranes were washed for 30 min in washing buffer at room temperature. Blots were incubated for 1 h at room temperature with horseradish peroxidase-conjugated donkey anti-goat immunoglobulin G (diluted 1:5000 in 5% milk-PBS; Jackson ImmunoResearch, West Grove, PA, USA) and horseradish peroxidase-linked sheep anti-mouse IgG (dilution 1:5000 in 5% milk-PBS; Amersham Biosciences, Arlington Heights, IL, USA). They were visualized by incubation with chemiluminescent solution (Immobilon Western Chemiluminescent HRP Substrate; Millipore, Billerica, MA, USA). The membranes were washed in washing buffer for another 30 min; bands were visualized on the UVP BioSpectrum 500 Imaging System (UVP, Upland, CA, USA).
For immunohistochemical staining, the L4–5 spinal cord segments were harvested 1 day after intrathecal injection of 10 μg NT-siRNA and intradermal injection of CFA into the left hind paws of the rats. The control group of rats received intrathecal injection of 1.8 μl of PEI.
Rats (n=3 in each group) were deeply anaesthetized with isoflurane and perfused through the ascending aorta with saline followed by 4% paraformaldehyde with 1.5% picric acid in 0.16 M phosphate buffer, and the L4–5 spinal cord segments were collected and post-fixed in the same fixative overnight. Spinal cord sections (30 μm free-floating sections) were cut using a cryostat and processed for immunofluorescence. Tissue sections were blocked with 2% goat serum, and incubated overnight at 4°C with primary antibody consisting of rabbit anti-IFN-α antibody (1:1000, R&D Systems). The sections were then incubated for 1 h at room temperature with FITC-conjugated secondary antibodies (1:300, Jackson ImmunoResearch Laboratories Inc., West Grove, PA, USA). The specificity of immunostaining and antibodies was tested by (i) omission of the primary antibody and (ii) absorption of antibodies with respective peptide antigens. The stained sections were examined with a Nikon fluorescence microscope, and images were captured with a CCD Spot camera.
The rats were habituated to the testing environment daily for at least 2 days before baseline testing. For testing mechanical sensitivity, animals were put into boxes on an elevated metal mesh floor and allowed 30 min for habituation before examination. The plantar surface of each hind paw was stimulated with a series of von Frey hairs with logarithmically incremental stiffness (Stoelting Co., Wood Dale, IL, USA), presented perpendicular to the plantar surface (3–5 s for each hair). The 50% paw withdrawal threshold was determined using Dixon's up–down method.16 Heat sensitivity was tested by radiant heat using Hargreaves apparatus (Stoelting Co., Wood Dale, IL, USA) and the data were presented as paw withdrawal latency (PWL). The radiant heat intensity was adjusted so that basal PWL before inflammation was between 10 and 12 s, with a cut-off of 20 s to prevent tissue damage.
All data were expressed as mean (standard deviation) (sd). Differences between the groups were compared using Student's t-test or one-way analysis of variance with the Bonferroni post hoc test. The criterion for statistical significance was P<0.05. The density of specific bands from western blotting was measured using Image-Pro Plus analysis software (MediaCybernetics, Silver Spring, MD, USA), and the results were expressed as the ratio of N-methyl-d-aspartate receptor type 1 immunoreactivity to β-tubulin immunoreactivity.
The mechanism by which RNAi inhibits gene expression is shown in Figure 1a. To test the off-target anti-nociceptive effect of siRNA, three NT siRNAs were used, including NT-siRNA, RISC-free siRNA, and GU-rich siRNA (Fig. 1b). CFA-induced allodynia was attenuated by all three types of siRNAs delivered intrathecally and the anti-allodynia effects of NT-siRNA were dose-dependent: the lower dose (5 μg) was ineffective and the highest dose (20 μg) almost blocked allodynia completely (Table 1). In contrast, PEI or a naked siRNA (without PEI) had no effect on allodynia (Table 1). Because the anti-allodynic effects were similar among 10, 20 μg NT-siRNA, RISC-free siRNA, and GU-rich-siRNA treatment groups, we used 10 μg NT-siRNA for the subsequent behavioural, biochemical, and histochemical studies. Of note, CFA-induced heat hypersensitivity (heat hyperalgesia) was also suppressed by NT-siRNA (Fig. 1c).
IFN responses were reported as common adverse effects of siRNAs.11,12 Remarkably, IFN-α neutralizing antibody reversed NT-siRNA-induced anti-nociception in CFA-inflamed rats (Fig. 1d), indicating an essential role for IFN-α in mediating siRNA-produced off-target analgesic effects.
To further support the idea that IFN-α regulates pain sensitivity, CFA-induced mechanical allodynia (Fig. 2a) was reversed by intrathecal administration of IFN-α, in a dose-dependent manner (Fig. 2b). IFN-α also increased paw withdrawal threshold in naïve animals (Fig. 2c), indicating an analgesic role for IFN-α. Furthermore, the IFN-α's anti-allodynic effect was reversed by intrathecal administration of the IFN-α neutralizing antibody (Fig. 2d), suggesting a specific effect of IFN-α.
Activation of opioid receptors is implicated in IFN-α effects in the brain.15 Notably, intrathecal administration of the opioid receptor antagonist naloxone (20 nmol) reversed the anti-allodynia effects produced by both IFN-α and NT-siRNA in the CFA model (Fig. 3). Therefore, siRNAs are likely to produce pain relief by producing IFN-α, which may activate spinal opioid receptors.
Finally, we examined IFN-α expression in the spinal cord. Western blotting analysis showed an increase in the spinal cord IFN-α level after intrathecal treatment with NT-siRNA (Fig. 4a). NT-siRNA also increased IFN-α immunoreactivity in the dorsal horn (Fig. 4b).
We demonstrated in this study that off-target anti-nociception/analgesia induced by IFN-α in the spinal cord is a novel effect of siRNAs. Although both the siRNA-induced IFN-α responses and IFN-α-induced analgesic effects have been reported,11,12,17 there are no studies demonstrating that siRNAs produce analgesic effects through the IFN-α response in the spinal cord. Thus, this is the first report demonstrating the IFN-α-mediated analgesic effects of siRNAs in the spinal cord. This off-target analgesia is dose-dependent and requires a high intrathecal dose (≥10 μg).9,10 It is generally believed that siRNA-induced off-target effects are sequence-dependent.11,12 For example, siRNAs with GU-rich immune stimulatory sequences are very potent in inducing IFN-α expression.11 The siRNA-induced off-target analgesia shown in this study was also sequence-dependent (Fig. 1b).
Longer double-strand RNAs (>30 bp) do not evoke severe side-effects in invertebrates and plants, but induce an IFN response in mammalian cells as a non-specific viral defence by activation of protein kinase R.18 It is now clear that some siRNAs and short hairpin RNAs induce IFN responses in vitro.12,19 In addition to responses mediated by protein kinase R, siRNAs may also activate IFN via toll-like receptors (TLR) TLR7 and TLR8, which recognize siRNAs within the endosome.20 Recently, immunostimulatory sequence motifs were identified that should be avoided when selecting RNA interference target sequences.11,20 In a result reported by Judge and colleagues,11 5′-UGUGU-3′ is the immunostimulatory motif within siRNA, based on the screening of 16 siRNAs. We tested the analgesic effect of GU-rich siRNA using the same sequence described by Judge and colleagues.11 Our previous review7 revealed that various doses of siRNA, from 70 ng to 400 μg daily for 6 days, have been used for modulating pain behaviour. The intrathecal doses of siRNA used in several studies are higher than 5 μg. Thus, IFN-α responses are likely to be induced in these studies. Our results demonstrated that siRNAs with different sequences could trigger off-target analgesia. It is also possible that these off-target effects could be reduced or abolished by modifying the structure of siRNAs at the sense strand termini to significantly reduce immunostimulatory activity.20,21
We also examined IFN-α expression in the spinal cord. Western blotting analysis showed increases in spinal cord IFN-α levels after intrathecal treatment of NT-siRNA (Fig. 4a). NT-siRNA also increased IFN-α immunoreactivity in the dorsal horn (Fig. 4b). In the CNS, microglial cells and/or macrophages were reported to produce IFN-α and IFN-β in human brains.22 Traugott and Lebon2,3 reported that some endothelial cells, macrophages, neurones, and astrocytes also produced type I IFN. In vitro experiments in primary cell cultures showed that astrocytes and microglial cells readily produced IFN-α and IFN-β in response to viral infections or synthetic double-stranded RNA treatment.24,25 In addition, Delhaye and colleagues26 showed that a small percentage of neurones from mice infected in vivo with Theiler's virus or La Crosse virus also produced IFN. Thus, we hypothesized that the source of the IFN-α secreted in our study was at least from astrocytes, microglial cells, and neurones.
How can IFN-α suppress inflammatory pain? Activation of μ-, but not δ- and κ-opioid receptors, is implicated in IFN-α effects in the brain.17 IFN-α shares some pharmacological properties with β-endorphin and produces analgesia via μ-opioid receptors.27 IFN-α inhibited the binding of [3H]-naloxone in an in vivo study,28 demonstrating competition between IFN-α and naloxone for membrane binding sites. In this study, we further demonstrated that intrathecal administration of the opioid receptor antagonist naloxone (20 nmol) reversed the anti-allodynia effects produced by both IFN-α and NT-siRNA in the CFA model (Fig. 3). Therefore, siRNAs produce pain relief by producing IFN-α, which activates spinal opioid receptors.
These findings uncovered a novel role of IFN-α in regulating siRNA-induced off-target analgesic effects and revealed a possible new form of neuronal–glial interaction in the spinal cord that could negatively regulate pain sensitivity. Accumulating evidence suggests that neural–glial interactions in the spinal cord facilitate chronic pain states. For example, spinal glia produce proinflammatory cytokines (e.g. tumour necrosis factor-α, interleukin-1β) and chemokines (e.g. C–C motif ligand 2) to increase the sensitivity of spinal nociceptive neurones,29–31 although these proinflammatory cytokines may impair learning and memory in the brain.32 Neural–glial interaction may also negatively regulate pain sensitivity via anti-inflammatory cytokines such as IL-10.33 Astrocytes are sensitive to develop innate immune responses to lipid-carried siRNAs.34 Although IFN-α can be released from glial cells such as microglia cells, astrocytes, and neurones,22–26 it is highly possible that spinal cord astrocytes are the major source of IFN-α after intrathecal administration of siRNA. In addition to an action on opioid receptors, there may be other mechanisms for the analgesic effects of IFN-α. The detailed mechanisms and pathway of secretion and analgesic effect of IFN-α induced by siRNA in the spinal cord deserve further investigation.
In summary, pain relief by a designed siRNA may not be attributable to target gene knockdown. Although previous studies suggested that some sequences (e.g. GU-rich) were essential for siRNA-induced IFN responses,11,20 our data showed that non GU-rich sequences also produced off-target analgesia at high doses. Thus, caution must be taken when designing siRNAs for target validation in pain research.
This work was supported by NIH grants DE17794, NS54932, and NS67686 to R.R.J., National Science Council Grant 100-2314-B-214-002-MY3 and E-Da Hospital Grant EDPJ 98023, 99038, 100037, and EDAHP 99030, Taiwan.