Anti-PD-1 treatment impairs opioid antinociception in rodents and nonhuman primates

Abstract

Emerging immunotherapies with monoclonal antibodies against programmed cell death protein-1 (PD-1) have shown success in treating cancers. However, PD-1 signaling in neurons is largely unknown. We recently reported that dorsal root ganglion (DRG) primary sensory neurons express PD-1 and activation of PD-1 inhibits neuronal excitability and pain. Opioids are mainstay treatments for cancer pain, and morphine produces antinociception via mu opioid receptor (MOR). Here we report that morphine antinociception and MOR signaling require neuronal PD-1. Morphine-induced antinociception following systemic or intrathecal injection was compromised in Pd1^−/− mice. Morphine antinociception was also diminished in wild-type mice after intravenous or intrathecal administration of Nivolumab, a clinically used anti-PD-1 monoclonal antibody. In mouse models of inflammatory, neuropathic, and cancer pain, spinal morphine antinociception was compromised in Pd1^−/− mice. MOR and PD-1 are co-expressed in sensory neurons and their axons in mouse and human DRG tissues. Morphine produced antinociception by 1) suppressing calcium currents in DRG neurons, 2) suppressing excitatory synaptic transmission and 3) inducing outward currents in spinal cord neurons; all of these actions were impaired by PD-1 blockade in mice. Loss of PD-1 also enhanced opioid-induced hyperalgesia and tolerance and potentiates opioid-induced microgliosis and long-term potentiation in the spinal cord in mice. Finally, intrathecal infusion of Nivolumab inhibited intrathecal morphine-induced antinociception in nonhuman primates. Our findings demonstrate that PD-1 regulates opioid receptor signaling in nociceptive neurons, leading to altered opioid-induced antinociception in rodents and nonhuman primates.

One-sentence summary:

PD-1regulates opioid signaling in primary sensory and spinal cord neurons and plays a critical role in opioid antinociception.

INTRODUCTION

Programmed cell death protein-1 (PD-1) is a member of the immunoglobulin gene superfamily, initially identified in T cells upon programmed cell death, with Pd1 mRNA expression restricted to the thymus (1). Mice lacking Pd1 develop auto-immune disease, suggesting an immune suppressive role of PD-1 (2). PD-L1, the ligand for PD-1, is highly expressed in many cancers and associated with mortality in cancer patients (3, 4), establishing a role of PD-1 in cancer-induced immune suppression. Inhibition of the interaction between PD-1 and PD-L1, known as an immune checkpoint blockade, enhances T-cell responses to produce antitumor activity (5). Emerging immune therapies using anti-PD-1 monoclonal antibodies have shown success in treating various cancers such as melanoma (4, 5). PD-1 is also expressed by melanoma cells, promoting tumor growth (6).

Despite extensive studies of PD-1 in non-neuronal cells, the nature of PD-1 signaling in neurons is largely unknown. Is PD-1 a neuromodulator or a neuro-checkpoint inhibitor? We recently showed that primary sensory neurons of dorsal root ganglion (DRG) also express functional PD-1 receptor and that activation of PD-1 by PD-L1 inhibits neuronal excitability and pain in mice (7). PD-L1 is produced by non-malignant tissues including DRG and spinal cord (7), implicating a physiological role of PD-L1. Furthermore, Pd1-deficient mice exhibit increased excitability in DRG neurons and increased pain sensitivity. In a mouse model of melanoma, cancer pain is masked by PD-L1, produced from melanoma cells and acting on skin nerve terminals (7).

Opioids are mainstay treatments for cancer pain (8-10). Morphine produces antinociception via mu opioid receptor (MOR) (11, 12), which is expressed in the central nervous system (CNS: brain and spinal cord) and peripheral nervous system (PNS: DRG and nerve) (11, 13). MOR mediates both the beneficial effects (such as antinociception) and the unwanted side effects of morphine (such as hyperalgesia, tolerance, withdrawal responses) (11, 14). This study was undertaken to investigate the interactions between PD-1 and MOR in the PNS and CNS. Our findings demonstrated that PD-1 is required for morphine antinociception and MOR signaling in DRG and spinal cord neurons in mice. Nivolumab, an FDA-approved anti-PD-1 monoclonal antibody, also inhibited morphine-induced antinociception in nonhuman primates after spinal administration.

RESULTS

Loss of PD-1 results in a reduction in morphine antinociception in mice

We first examined whether morphine antinociception would be altered in Pd1 knockout (Pd1^−/−) mice after subcutaneous injection (1, 3, and 10 mg/kg, s.c). Tail-flick testing in wild-type (WT) mice revealed a rapid (<0.5 h) and dose-dependent increase in tail-flick latency (TFL) that lasted more than 3 h after morphine treatment (P < 0.0001, Fig. 1A and fig. S1, A-C). The duration of the antinociceptive effect was reduced in Pd1^−/− animals (P < 0.0001, Fig. 1A and fig. S1, A-C). Hot-plate testing also showed an impairment of morphine-induced antinociception in Pd1^−/− mice (P < 0.0001, Fig. 1A; and fig. S1, D-F). The dose-response curve showed a right-shift in morphine antinociception in Pd1^−/− mice compared to WT animals (P < 0.05, tail-flick and hot-plate tests, Fig. 1B). Intrathecal morphine administration (2 nmol, i.t.) also elicited marked antinociception in tail-flick and hot-plate tests in WT mice; but this action was compromised in Pd1^−/− mice (P < 0.0001, tail-flick and hot-plate tests, Fig. 1C). Collectively, these results suggest that morphine antinociception requires PD-1 via both peripheral and central actions.

Fig. 1. Morphine antinociception is diminished in mice lacking PD-1.

(A) Subcutaneous morphine antinociception (s.c., 10 mg/kg) in WT mice and Pd1^−/− mice in tail-flick (left) and hot-plate (right) tests. Saline injection in WT mice and Pd1^−/− mice was included as vehicle. (B) Dose-responses and ED₅₀ values of s.c. morphine antinociception in tail-flick (left) and hot-plate (right) tests in WT and Pd1^−/− mice. *P < 0.05, ****P < 0.0001, WT vs. Pd1^−/−, unpaired two-tailed t-test. (C) Intrathecal morphine antinociception (i.t., 2 nmol) in WT and Pd1^−/− mice in tail-flick (left) and hot-plate (right) tests. Saline injection in WT and Pd1^−/− mice was included as vehicle. (D) Intrathecal antinociception of DAMGO (MOR agonist), DPDPE (DOR agonist) or U69593 (KOR agonist) in tail-flick test. (A, C, D) *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, WT vs. Pd1^−/−, two-way ANOVA, followed by Bonferroni’s post hoc test. (E-G) Intrathecal morphine antinociception (2 nmol) in LLC-induced bone cancer pain (E), CFA-induced inflammatory pain (F), and SNL-induced neuropathic pain (G) in WT and Pd1^−/− mice. n.s., no significance, *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, vs. pre-injection baselines. ^##P < 0.01, ^###P < 0.001, ^####P < 0.0001, WT vs. Pd1^−/−, two-way ANOVA, followed by Bonferroni’s post hoc test. Data are Mean ± SEM. Sample sizes are indicated in brackets.

Opioids induce antinociception through the μ, δ, and κ opioid receptors (MOR, DOR, and KOR respectively) (11). Intrathecal injection of specific agonists DAMGO (MOR), DEDPE (MOR), and U69593 (KOR) each evoked antinociception in WT mice (Fig. 1D and fig. S2, A-C). Pd1^−/− mice only exhibited reduction in DAMGO but not DPDPE or U69593 induced antinociception (P = 0.0030 and 0.0487 for DAMGO in tail-flick and hot-plate test, respectively, Fig. 1D and fig. S2, A-C). Thus, PD-1 primarily affects the MOR-mediated antinociception.

Opioids are a mainstay treatment for cancer pain, which often manifests after bone metastasis (15). We assessed whether morphine would attenuate cancer pain in WT and Pd1^−/− mice using a bone cancer model (16). Femoral inoculation of Lewis lung cancer cells resulted in severe cancer pain, as indicated by reductions in mechanical threshold (Fig. 1E). Spinal morphine administration completely reversed tumor-induced mechanical allodynia in WT mice (P < 0.0001, Fig. 1E and fig. S2D). However, the anti-allodynic effect of morphine was largely compromised in Pd1^−/− mice (P < 0.0001, Fig. 1E). The anti-hyperalgesia effect of morphine in Hargreaves test was also compromised in Pd1^−/− mice (P < 0.0001, fig. S2, E and F). Furthermore, i.t. morphine evoked anti-allodynic effects in inflammatory pain and neuropathic pain in WT mice, induced by complete Freund’s adjuvant (CFA) and spinal nerve ligation (SNL), respectively; also in these cases, morphine’s antinociception was compromised in Pd1^−/− mice (P < 0.0001 in both CFA and SNL models, Fig. 1, F and G).

Anti-PD-1 treatment with Nivolumab diminishes morphine antinociception in wild-type mice

Next, we tested whether impaired morphine antinociception in Pd1^−/− mice could be recapitulated by treatment of Nivolumab, a human anti-PD-1 monoclonal antibody previously shown to evoke allodynia in WT naive mice (7). Systemic pre-treatment with Nivolumab (10 mg/kg, i.v.), given 1 h prior to the morphine injection, reduced morphine antinociception in the tail-flick and hot-plate tests (P < 0.0001, tail-flick and hot-plate tests, Fig. 2, A and B, fig. S3, A and B). Furthermore, spinal pretreatment with Nivolumab (1 μg, i.t., given 30 min prior to morphine) also decreased morphine antinociception (2 nmol, i.t., P < 0.0001 for both tail-flick and hot-plate tests, Fig. 2C). Dose-response analysis revealed increases in ED₅₀ values of morphine antinociception in Pd1^−/− mice and Nivolumab-treated mice, as compared to WT mice (fig. S3C). ED₅₀ values (mg/kg) were: 2.8 (talk-flick test) and 4.1 mg/kg (hot-plate test) in WT mice, 5.5 (tail-flick) and 9.3 (hot-plate) in Pd1^−/− mice, and 8.2 (tail-flick) and 8.2 (hot-plate) in Nivolumab-treated mice (fig. S3C). To assess for the possibility of immune modulation in this process, we further tested the effects of spinal pretreatment with Nivolumab on morphine antinociception in CB-17 mice with deficiency in T cells and B cells. Nivolumab also decreased morphine antinociception (2 nmol, i.t.) in adaptive CB-17 immune cell-deficient mice (P = 0.0117 and 0.0069 for tail-flick test and hot-plate test, respectively, fig. S3D).

Fig. 2. Morphine antinociception is decreased in WT mice after pretreatment with anti-PD-1 antibody Nivolumab.

(A) Subcutaneous morphine antinociception (s.c., 10 mg/kg) after intravenous pretreatment with Nivolumab (i.v., 10 mg/kg) in tail-flick (top) and hot-plate (bottom) tests. Nivolumab (Nivo) or control human IgG4 (IgG4) was i.v. injected 60 min prior to s.c. morphine injection. (B) Dose-response curve of morphine induced antinociceptive effects (MPE% at the peak time point) in Nivolumab (i.v., 10 mg/kg) pretreated mice. (C) Spinal antinociception of morphine following intrathecal injection (2 nmol) in mice pretreated with IgG4 or Nivolumab (i.t., 1 μg) in tail-flick (top) and hot-plate (bottom) tests. Nivolumab or IgG4 was i.t. injected 30 min prior to i.t. morphine injection. (A,C) *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, morphine IgG4 vs. morphine Nivolumab, two-way ANOVA, followed by Bonferroni’s post hoc test. Data are Mean ± SEM. Sample sizes are shown in brackets.

In addition to male mice, morphine antinociception was also decreased in female Pd1^−/− mice (P = 0.0155 for tail-flick test, P = 0.0070 for hot-plate test) and blunted by Nivolumab in female WT mice (P = 0.0025 for tail-flick test, P = 0.0102 for hot-plate test, fig. S4, A-D). Taken together, our data demonstrate that Pd1 deletion or PD-1 blockade causes reduction in morphine antinociception in naïve animals and also in animals with inflammatory, neuropathic, and cancer pain. These results also suggest a functional interaction between PD-1 and MOR receptors.

PD-1 is co-expressed with MOR and regulates MOR function in DRG neurons

To determine the mechanisms by which PD-1 regulates opioid antinociception, we examined the interaction of PD-1 and MOR at the cellular level. In the DRG, MOR is mainly expressed by small-diameter nociceptive neurons (13). Using a sensitive RNAscope assay, we observed a high percentage of co-localization of Pd1 and mu-opioid receptor (Oprm1) mRNAs in DRG neurons (Fig. 3, A and B). To determine the mRNA expression of Pd1 and Oprm1 in individual DRG neurons, we quantified the number of fluorescence-labeled puncta in positive neurons, showing that approximately 45% neurons express Pd1 mRNA, 40% neurons express Oprm1 mRNA, and 30% DRG neurons expressed both mRNAs (Fig. 3C). Among Pd1⁺ cells, ~ 60% co-expressed Oprm1 (Fig. 3C).

Fig. 3. PD-1 and MOR are co-expressed in mouse DRG neurons and have functional interaction.

(A, B) In situ hybridization (ISH, RNAscope) images of Oprm1 mRNA (A) and Pd1 mRNA (B) in mouse DRG neurons of WT mice. Sixteen pairs of numbers show 16 neurons co-expressing both mRNAs. White starts (A) and yellow stars (B) show single-labeled neurons. Scale bar, 20 μm. (C) The percentage of positive neurons for Pd1 and Oprm mRNA expression and co-expression in DRG. 1244 neurons from 3 mice were analyzed. (D) Double staining showing co-localization of PD-1 and MOR in DRG. Yellow arrows indicate double-labeled neurons. Lower panels, enlarged boxes show co-expression of both receptors on the cell surface indicated by yellow arrows. Scale bars, 10 μm. Right panel, PD-1 immunostaining with a PD-1 blocking peptide. Blue DAPI staining labels cell nuclei. Scale, 10 μm. (E) Proximity ligation assay (PLA) shows positive signals of PD-1/MOR interaction in cytoplasm (arrows) and axons (arrowheads) of cultured DRG neurons (9 images from two repeats). (F) Co-IP showing PD-1/MOR interaction in DRG. DRG lysate was immunoprecipitated with control IgG or MOR antibody and immunoblotted with PD-1 or MOR antibody. This experiment was repeated in triplicate. (G, H) Morphine (10 μM) effects on calcium currents in small-diameter DRG neurons (<25 μm) of WT and Pd1^−/− mice. (G) Traces of calcium currents. (H) Time-course of relative calcium currents. *P < 0.05, **P < 0.01, ****P < 0.0001, Pd1^−/− vs. WT group, two-way ANOVA, followed by Bonferroni’s post hoc test. (I) Double staining for MOR and PD-1 in human dorsal root. PD-1 was labeled by Nivolumab (1 mg/ml). Arrows indicate the double-labeled nerve fibers. Scale bars, 250 μm (left) and 50 μm (right). Data are Mean ± SEM. Sample sizes are indicated in brackets.

Immunohistochemistry also showed high degree of co-localization of PD-1 and MOR immunoreactivity (IR) in DRG neurons (Fig. 3D, fig. S5A). Co-localization was especially observed on the cell surface of small-diameter DRG neurons (Fig. 3D). Specificity of PD-1-IR was confirmed by a lack of staining after blocking peptide treatment (Fig. 3D) and was also tested previously in Pd1^−/− mice (7). MOR expression did not change in Pd1^−/− mice (P = 0.3637 vs. WT, fig. S5, B-C). Furthermore, we observed co-localization of PD-1 and MOR in axons of the mouse spinal nerve (fig. S5D), indicating axonal transport of PD-1 and MOR from cell bodies to peripheral nerve axons.

We further examined possible PD-1/MOR interaction using proximity ligation assay (PLA) in dissociated DRG neurons (17) and co-immunoprecipitation (Co-IP) in DRG tissue. PLA analysis revealed positive fluorescence signals on cell bodies and axons in 44 % of cultured mouse DRG neurons (Fig. 3E and fig. S5E), and staining was absent in negative control with omission of primary antibody or following treatment with blocking peptide (fig. S5E). Co-IP experiment demonstrated that anti-MOR antibody pulled down PD-1 in mouse DRG lysates (8.7-fold of control IgG, Fig. 3F). These results indicate close proximity of PD-1 and MOR and possible interaction between these two receptors in native DRG neurons.

Opioids inhibit pain by suppressing the function of calcium channels in primary afferent neurons (18). Accordingly, we recorded calcium currents in dissociated small-diameter DRG neurons (<25 μm) in WT and Pd1^−/− mice using whole-cell patch clamp (18). Morphine (10 μM) produced a 35% reduction in calcium currents in WT DRG neurons (Fig. 3, G and H). However, this reduction was blunted in DRG neurons from Pd1^−/− mice (P < 0.0001, Fig. 3, G and H). Thus, PD-1 is a required signaling element for morphine to suppress the calcium channel function in mouse DRGs.

We previously showed that PD-1-IR is expressed on the surface of human DRG neurons (7). We further examined PD-1 and MOR expression in human dorsal roots. Co-expression of PD-1 and MOR was observed on the nerve axons of the human dorsal roots (Fig. 3I). The Co-IP assay also suggested a possible interaction of MOR and PD-1 in human dorsal root lysates (fig. S5F).

PD-1 disruption impairs suppression of nociceptive transmission by morphine in spinal cord neurons

Inhibition of calcium channels in the central terminals of DRG neurons by opioid treatment also inhibits neurotransmitter release in the spinal cord dorsal horn (SDH) via presynaptic regulation (11). Double staining in SDH revealed PD-1 colocalization with MOR in axons and axonal terminals (fig. S6A) and with CGRP, a neuropeptide that is present in primary afferent central terminals (fig. S6B). The results suggest that spinal PD-1 may originate from DRG neurons.

To test the function of PD-1 in SDH synaptic transmission, we prepared spinal cord slices and recorded spontaneous excitatory post-synaptic currents (sEPSCs) in outer lamina II neurons (Fig. 4, A-E). These interneurons are predominantly excitatory and form a nociceptive circuit with primary C-afferents and lamina I projection neurons (19). Bath perfusion of WT spinal cord slices with morphine (10 μM) led to a marked inhibition of sEPSC frequency (44.8% inhibition, P < 0.0001, Fig. 4, A and C). However, in Pd1^−/− SDH neurons, the degree of morphine-induced inhibition was diminished (22.7%, P = 0.0068, Fig. 4, B, D and E). Morphine (10 μM) caused a mild inhibition of sEPSC amplitude (10.7%) in WT mice (P = 0.0013) but had no such inhibition in Pd1^−/− mice (P = 0.09561, Fig. 4, A-E). No significant difference was observed in the sEPSC amplitude in Pd1^−/− cells compared to WT after morphine treatment (P = 0.2692, Fig. 4E). Morphine suppressed sEPSC frequency with an IC₅₀ of 1.07 μM and 15.24 μM in WT and Pd1^−/− samples, respectively (fig. S7, A-C). DAMGO’s (0.5 μM) inhibition on sEPSCs was also diminished in Pd1^−/− neurons compared to WT cells (P = 0.0154, fig. S7, D and E), suggesting a specific involvement of MOR. Thus, Pd1 deletion diminishes MOR-mediated suppression of spinal nociceptive synaptic transmission.

Fig. 4. Morphine’s inhibition on spinal cord nociceptive neurons in WT and Pd1^−/−mice and the effects of Nivolumab.

(A-E) Morphine’s effects on spinal excitatory synaptic transmission in lamina II neurons of spinal cord slices in WT and Pd1^−/− mice. (A, B) Recording traces of spontaneous excitatory postsynaptic currents (sEPSCs) and cumulative histograms of the inter-event interval and amplitude of sEPSCs in neurons from WT (A) and Pd1^−/− (B) mice, before (Control) and after morphine treatment (10 μM). The histograms were examined for 1 min before morphine treatment (338 events in WT mice and 415 events in Pd1^−/− mice) and 1 min after morphine washout (85 events in WT mice and 235 events in Pd1^−/− mice). D indicates distance between two curves. (C, D) sEPSCs frequency and amplitude before and after morphine (10 μM) treatment in WT and Pd1^−/− mice. (E) Percentage changes in sEPSCs frequency (upper) and amplitude (lower) in WT and Pd1^−/− mice. Unpaired two-tailed t-test. (F, G) Morphine-induced outward currents in lamina II neurons of spinal cord slices of WT and Pd1^−/− mice. (F) Traces of outward currents in WT and Pd1^−/− mice. (G) Left, chart distribution of neurons with and without outward current. Right, amplitude of morphine-induced outward currents in WT and Pd1^−/− mice. (H) Effects of control IgG (left) and Nivolumab (right) on morphine’s inhibition of sEPSCs in lamina II neurons of spinal cord slices of WT mice. Top, recording traces of sEPSCs. Bottom, sEPSCs frequency and amplitude. (I) Percentage changes in sEPSCs frequency (top) and amplitude (bottom) showing the effects of IgG or Nivolumab on morphine’s inhibition of sEPSCs. (A,B) Kolmogorov Smirno Test; (C,D,H) paired two-tailed t-test; (E,G,I) unpaired two-tailed t-test. Data are Mean ± SEM. Sample sizes are indicated in brackets.

Opioids also activate potassium channels to generate outward currents and hyperpolarization in neurons (20). Morphine (10 μM) evoked outward currents in 36.7% (5/14) SDH lamina II neurons of WT mice with an amplitude of 15.8 ± 1.5 pA (Fig. 4, F and G). Morphine-evoked outward currents were significantly smaller in Pd1^−/− mice (7.8 ± 1.8 pA; P = 0.0104), despite a similar response rate (30.7% of 13 neurons, Fig. 4, F and G). Additionally, potassium currents were blocked by Tertiapin-Q, a high affinity blocker for inward-rectifier K⁺ channels, suggesting an involvement of G protein-gated inwardly rectifying potassium (GIRK) K⁺ channels (fig. S7F).

We next investigated whether Nivolumab would alter the effects of morphine on SDH neurons of WT mice. Perfusion of spinal cord slices with Nivolumab at a very low concentration (100 ng/ml ≈ 0.7 nM) significantly reduced the morphine’s inhibition of sEPSC frequency (P = 0.0004, compared with human IgG4 control, Fig. 4, H and I). Nivolumab at 0.7 nM without morphine did not affect sEPSC frequency but did increase sEPSC frequency at higher concentrations (P = 0.0229 and P = 0.0351 for 2.1 and 10.5 nM, respectively, fig. S8, A-F). Treatment with control IgG4 did not affect the morphine-induced inhibition of sEPSC (Fig. 4H and fig. S8). Nivolumab (0.7 nM) also suppressed morphine’s inhibition on sEPSC amplitude (P = 0.0257, Fig. 4I).

PD-L1 and morphine act together to inhibit synaptic transmission and nociception

As a major ligand of PD-1, PD-L1 produces antinociception in mouse models of pathological pain (7). A dose response comparison showed that PD-L1 and morphine inhibit sEPSC frequency at IC₅₀ of 0.60 nM and 1.07 μM, respectively (fig. S7C and fig. S9, A and B). PD-L1 failed to inhibit sEPSCs in SDH neurons lacking Pd1 (fig. S9C). In naïve animals, Pd1 deficiency led to an increase in sEPSC frequency (P < 0.0001) but not amplitude (P = 0.7208) (fig. S9D), indicating that PD-1 is an endogenous negative regulator of synaptic transmission in mouse SDH neurons.

We further assessed whether PD-L1 and morphine would act together to modulate nociception. At very low concentrations, PD-L1 (0.075 nM) and morphine (0.01 μM) each produced mild inhibition of sEPSC frequency (5% and 8%, respectively), but co-application of PD-L1 and morphine at these same low concentrations produced a much greater inhibition (40%) of sEPSCs, suggesting a collaborative enhancement of these two compounds in inhibition of sEPSC at low concentrations (P < 0.0001, Fig. 5A-D). This enhancement was also observed in behavioral testing. Intrathecal PD-L1 (0.075 nmol) failed to produce any antinociception, and intrathecal morphine (0.75 nmol) only produce very mild and transient antinociception. However, intrathecal co-application of PD-L1 and morphine at these low doses produced a much greater degree of antinociception (P < 0.0001 for both tail-flick and hot-plate tests, Fig. 5E).

Fig. 5. PD-L1 and morphine produce collaborative inhibition of spinal synaptic transmission and nociception in WT mice.

(A-C) Top, sEPSC traces showing the effects of PD-L1 (A), morphine (B), and PD-L plus morphine (C) in lamina II neurons of spinal cord slices. The upper and lower traces in each panel were obtained from the same neurons. Bottom, sEPSCs frequency and amplitude. Date were analyzed by paired two-tailed t-test. (D) Percentage changes in sEPSCs frequency (left) and amplitude (right) under the treatment of PD-L1, morphine, and PD-L1 plus morphine. (E) Antinociception of PD-L1 plus morphine following intrathecal administration in tail-flick test (left) and hot-plate test (right). PD-L1 or saline was injected 30 min prior to morphine injection. (A-C), paired two-tailed t-test; (D) one-way AVONA; (E) two-way ANOVA, followed by Bonferroni’s post hoc test. *P < 0.05, **P < 0.01, ***P < 0.0001, ****P < 0.0001, saline + morphine vs. PD-L1 + morphine. Data are Mean ± SEM. Sample sizes are indicated in brackets.

Next, we asked whether PD-L1 would inhibit nociception via opioid receptors. Perfusion of spinal cord slices with the opioid receptor antagonist naloxone (1 μM) had no effects on basal sEPSC frequency and amplitude (fig. S10, A and B), but completely blocked the inhibitory actions of morphine (10 μM, fig. S10, C and D). Naloxone also blocked the PD-L1’s inhibition of sEPSC frequency and amplitude (Fig. S10, E and F). Furthermore, intrathecal injection of PD-L1 (10 μg) reduced mechanical hypersensitivity in experimental bone cancer in mice, but this antinociceptive effect was abrogated by intrathecal naloxone (5 nmol, Fig. S11, A-C). To determine if PD-L1 could directly activate MOR, we utilized a cell line which lacks PD-1 expression but has high expression of MOR and promiscuous Gq proteins coupled to calcium signaling. At all the concentrations (30-3000 ng/ml), PD-L1 failed to activate MOR to trigger Ca²⁺ responses (fig. S11, D and E). As a positive control, DAMGO (0.5 μM) evoked a robust Ca²⁺ increase, which was blocked by naloxone (fig. S11, D and E). Thus, PD-L1 may indirectly interact with MOR to modulate synaptic transmission and nociception.

Loss of PD-1 enhances opioid-induced hyperalgesia and spinal long-term potentiation

Opioid-induced hyperalgesia/allodynia not only occurs after chronic exposure but may also happen after acute treatment (21). Intrathecal DAMGO (0.5 nmol) elicited much faster mechanical allodynia in Pd1^−/− mice, which was evident at 3 h in von Frey testing using both threshold (P < 0.0001, Fig. 6A) and frequency measurements (P = 0.0031, 0.4 g, Fig. 6B). Opioid withdrawal has been shown to induce long-term potentiation (LTP), a critical form of synaptic plasticity underlying the pathogenesis of pain (21, 22). We recorded for LTP in lamina II neurons in spinal cord slices from WT and Pd1^−/− mice following electrical stimulation of the attached dorsal root. Perfusion of DAMGO (0.5 μM, 3 min) in WT neurons initially induced a depression of evoked EPSCs (eEPSCs) during the perfusion, and this depression was diminished in Pd1-deficient neurons (P = 0.0488, Fig. 6, C-E). After DAMGO withdrawal, the depression phase was followed by a recovery phase in WT and Pd1^−/− mice, but the duration of recovery was shortened in Pd1-deficient neurons (P = 0.0102, vs. WT, Fig. 6, C, D, and F). After the recovery phase, WT neurons developed a moderate LTP (20% increase in eEPSC amplitude, Fig. 6, C, D, G). In sharp contrast, LTP was much greater in Pd1^−/− mice (P = 0.0463, Fig. 6G). Additionally, LTP occurred more rapidly in Pd1^−/− mice (Fig. 6D). Together, these findings suggest loss of PD-1 might cause more fast and robust development of opioid-induced hyperalgesia/allodynia, as a result of enhanced LTP in SDH neurons.

Fig. 6. DAMGO induced hyperalgesia and spinal long-term potentiation is enhanced in Pd1^−/− mice.

(A, B) Paw withdrawal threshold (A) and paw withdrawal frequency to 0.16g filament (B) change after DAMGO (0.5 nmol, i.t.) in WT and Pd1^−/− mice. *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, WT vs. Pd1^−/−, two-way ANOVA, followed by Bonferroni’s post hoc test. (C-G) LTP of EPSC amplitude in lamina II neurons of spinal cord slices following DAMGO withdrawal in WT and Pd1^−/− mice. (C) Recording traces of evoked-EPSC before, during, and after DAMGO (0.5 μM, 3 min) in SDH neurons of WT and Pd1^−/− mice. (D) Time course of evoked EPSC amplitude of WT and Pd1^−/− neurons. Two-way ANOVA. (E) The effects of DAMGO on evoked-EPSC amplitude, compared to the baseline, in WT and Pd1^−/− mice during the drug perfusion. (F) Latency (min) of recovery of DAMGO-induced depression of evoked-EPSC amplitude, compared to the baseline (100%) in WT and Pd1^−/− mice. (G) Maximum potentiation (% of baseline) of evoked-EPSC amplitude at 30-36 min after DAMGO (0.5 μM) perfusion in WT and Pd1^−/− mice. (E-G) Unpaired two-tailed t-test. Data are Mean ± SEM. Sample sizes are indicated in brackets.

Loss of PD-1 enhances chronic opioid-induced tolerance and microgliosis in SDH

Chronic morphine exposure (6 daily injections, 10 mg/kg, s.c.) induced antinociceptive tolerance in WT and Pd1^−/− mice in tail-flick testing (Fig. 7A, left) and hot-plate testing (Fig. 7B, left). The maximum possible effect (MPE) analysis revealed that morphine evoked tolerance in WT mice on day 4 in tall-flick testing and on day 3 in hot-plate testing (P < 0.0001, Fig. 7, A and B, right). In Pd1^−/− mice, the induction was faster: morphine induced tolerance on day 2 in both tall-flick and hot-plate tests (P = 0.0163 and 0.0256, respectively, Fig. 7, A and B, right).

Fig. 7. Chronic morphine induced tolerance and microgliosis in SDH is potentiated in Pd1^−/− mice.

(A, B) Tail-flick (A) and hot-plate (B) tests showing the latency (left) and maximum possible effect (%MPE, right) of morphine antinociception using a 6-day treatment paradigm (10 mg/kg, subcutaneous, once daily) in WT and Pd1^−/− mice. *P < 0.05, ***P < 0.001, vs. Day 1 baseline antinociception, two-way ANOVA. Morphine antinociception was tested 30 min after each injection. (C, D) Microglial activation (microgliosis) in SDH of WT and Pd1^−/− mice after 4-day morphine exposure (10 mg/kg, subcutaneous, once daily). (C) Low magnification (Top) and high magnification (Bottom) images of IBA1 immunostaining. Scales, 500 μm. (D) Quantification of IBA1 immunoreactivity in SDH. **P < 0.01, unpaired two-tailed t-test, WT vs. Pd1^−/− mice. Data are Mean ± SEM. Sample sizes are indicated in brackets.

Since microglial activation in SDH after chronic opioid exposure regulates opioid tolerance (23, 24) and PD-1 was implicated in microglia signaling (25), we investigated microglia activation (microgliosis) in SDH of WT and Pd1^−/− mice following 4-day morphine exposure (10 mg/kg, s.c., daily). This course of morphine treatment did not trigger microgliosis in WT mice, as revealed by IBA-1 immunostaining, but induced a marked microgliosis in Pd1^−/− mice (P = 0.0040, Fig. 7, C and D). Thus, enhanced microglial activation in the SDH plays an active role in the facilitation of opioid-induced tolerance in PD-1 deficiency.

We examined additional adverse effects of opioids, including constipation and withdrawal response in WT and Pd1^−/− mice. Gastrointestinal transit (GIT) assay revealed increased baseline GI movement in Pd1^−/− compared to WT mice (P = 0.0190, fig. S12A). Subcutaneous morphine (10 mg/kg) induced a substantial reduction of GI movement in WT mice, and this deficit was improved in Pd1^−/− mice (P = 0.0265, fig. S12A). We also tested morphine induced withdrawal response following chronic morphine exposure (26) by examining naloxone-induced (2 mg/kg, i.p.) jumping response for 30 min in WT and Pd1^−/− mice. Naloxone-induced jumping was potentiated in Pd1^−/− mice within the first 10 min (P = 0.0001 and P = 0.0039, fig. S12B), suggesting a moderate modulation of opioid withdrawal response by PD-1.

Spinal infusion of Nivolumab blocks morphine antinociception in nonhuman primates

Nonhuman primates (NHP) have been tested for new pain therapeutics including opioids due to their phylogenetic proximity to humans (27). We examined the actions of Nivolumab on nociception and morphine antinociception in NHPs. To minimize immunomodulatory response of this human antibody and target the spinal cord pain circuit, we delivered drug via a catheter implanted into the intrathecal cerebrospinal space of NHP (Fig. 8A) (28). Tail flick testing in a 46 °C water bath revealed a dose-dependent hyperalgesia following intrathecal infusion of Nivolumab (10, 30 and 100 μg) in NHPs (Fig. 8A, B). The induction of hyperalgesia, corresponding to a reduction in tail flick latency, was very rapid, peaking within 1 h, suggesting a possible neuronal hyperactivity (Fig. 8B). This hyperalgesia was maintained at 6 h but fully recovered at 24 h (P < 0.0001, n = 5 NHPs, Fig. 8B). By contrast, the isotope control serum (IgG4) had no effect on thermal threshold even at the highest dose (100 μg, i.t., Fig. 8B). Thus, Nivolumab is sufficient to produce hyperalgesia in NHPs (85% reduction of thermal latency at the dose of 100 μg, Fig. 8A, B).

Fig. 8. Morphine antinociception is blocked following pretreatment with Nivolumab in nonhuman primates.

(A) Paradigm of intrathecal injection, drug treatment, and tail-flick test in 46 °C water bath for testing thermal hyperalgesia. (B) Thermal threshold changes after intrathecal Nivolumab (10, 30 and 100 μg). ****P < 0.0001, compared with control human IgG4 treatment, two-way ANOVA, followed by Bonferroni’s post hoc test. (C) Paradigm of intrathecal injection, drug treatment, and tail-flick test in 50°C water bath for testing morphine antinociception. (D-F) The effects of intrathecal pretreatment with Nivolumab (30 and 100 μg) on spinal antinociception of morphine (30 μg). (D) Tail withdrawal latency before and after Nivolumab administration. (E) %MPE, ****P < 0.0001, IgG4 + morphine vs. Nivolumab (Nivo) + morphine, two-way ANOVA, followed by Bonferroni’s post hoc test. (F) Area under the curve (AUC) of %MPE; P = 0.0002, P < 0.0001, one-way ANOVA, followed by Bonferroni’s post hoc test. n = 5 nonhuman primates per group. Nivolumab or control antibody (human IgG4) was i.t. injected 60 min prior to i.t. morphine injection. Data are Mean ± SEM.

Finally, we tested whether Nivolumab would impair morphine antinociception in NHPs as in rodents. Nivolumab was given 1 h prior to morphine injection, and tail withdrawal testing was conducted in a 50 °C water bath to test morphine antinociception (29) (Fig. 8C). Intrathecal infusion of morphine (30 μg) evoked full antinociception for > 3 h, as shown by tail withdrawal latency (Fig. 8D), MPE (Fig. 8E), and area under the curve (AUC) (Fig. 8F). Intrathecal pretreatment with Nivolumab (30 and 100 μg) produced a dose-dependent inhibition of morphine antinociception in NHPs (P < 0.001, n = 5 animals), whereas IgG4 (100 μg, i.t.) had no effect (Fig. 8D-F). Nivolumab (100 μg) largely blocked morphine antinociception in NHPs (> 85% reduction, P < 0.001, n = 5, Fig. 8F). These data strongly indicate that anti-PD-1 treatment could block morphine antinociception in NHPs.

DISCUSSION

We have demonstrated a previously unrecognized role of PD-1 in regulating opioid antinociception in rodents and NHPs, with several lines of cellular, functional, and behavioral evidence demonstrating a crosstalk between PD-1 and MOR. First, PD-1 and MOR are co-localized in DRG neurons, spinal nerve axons, and spinal cord axonal terminals. The proximity and potential interaction between these two molecules were further evidenced by Co-IP and PLA experiments. Second, morphine’s antinociception in tail-flick and hot-plate testing in naïve mice as well as its anti-allodynic actions in inflammatory, neuropathic, and bone cancer pain models are all diminished in Pd1^−/− mice. Third, morphine’s antinociceptive cellular actions, including suppression of calcium currents in DRG neurons, inhibition of sEPSCs in SDH neurons, and induction of outward currents in SDH neurons, were all compromised in Pd1^−/− mice. Morphine’s signaling deficits in Pd1^−/− mice can be recapitulated by Nivolumab treatment in adult wild-type mice, arguing against developmental regulation in Pd1^−/− mice. Future studies are warranted to determine how PD-1 interacts with MOR at the cellular and molecular levels.

We also tested for adverse effects from opioid treatment, including hyperalgesia/allodynia, tolerance, and withdrawal response in WT and Pd1^−/− mice. These excitatory adverse effects of opioids (morphine and DAMGO) are all potentiated in Pd1^−/− mice. In contrast, constipation, an inhibitory adverse effect of opioid, was improved in Pd1^−/− mice. Pd1 deficiency or blockade in WT mice also increased basal synaptic transmission (EPSC) and LTP in SDH neurons, whereas exogenous PD-L1 suppressed EPSC in these neurons. Together, these observations indicate an overwhelming inhibitory action of PD-1 on neurons. Our previous study showed that activation of PD-1 by PD-L1 activates the phosphatase SHP-1 in nociceptors to suppress neuronal excitability via inhibition of sodium channels and activation of potassium channels (7). Thus, PD-1 may serve as a neuronal suppressant in the PNS and CNS. It remains to be investigated if PD-1 acts as a pan-neuronal inhibitor in different brain regions.

There are two caveats to this study. First, although we revealed neuronal modulation by PD-1 in opioid-induced antinociception and hyperalgesia, we did not rule out the contribution of PD-1-mediated immune modulation to these opioid-induced processes. PD-1 is highly expressed by T cells and antigen presenting cells such as dendritic cells and macrophages to inhibit immunity (30). However, PD-1 in microglia acts to inhibit the function of these CNS immune cells (25). Our data showed that loss of PD-1 potentiated chronic morphine induced microgliosis in the SDH, contributing to enhanced opioid tolerance in Pd1^−/− mice. In addition, PD-1 expressed on other immune cell types such as T cells and macrophages may also play a role in opioid tolerance. However, the acute effect of PD-1 on opioid antinociception is very likely mediated by neuronal PD-1, given its rapid actions within a few minutes to tens of minutes. To determine the specific contribution of each cell type in morphine antinociception and tolerance, conditional knockout mice with Pd1 deletion in different neuronal and immune cell types are needed. Second, the downstream signaling of PD-1 remains to be determined. The phosphatase SHP-1 is required for PD-1 to regulate the activities of sodium channels and potassium channels and pain sensitivity (10). It will be of great interest to investigate the involvement of SHP in opioid antinociception and tolerance.

Nivolumab’s approval for treating various cancers (31, 32) and the use of opioids as mainstay treatments for cancer pain (10, 33) render major clinical implications for our findings in this study. We found co-localization and interactions between MOR and PD-1 in human dorsal root tissue and demonstrated Nivolumab blockade of spinal morphine nociception in nonhuman primates. Thus, anti-PD-1 immunotherapy may interfere with opioid antinociception in cancer patients by disrupting the PD-1-MOR interaction in peripheral nerve and DRG, as well as in the spinal cord particularly with disruption of the blood-brain-barrier in some cancer patients. Additionally, PD-L1 might be used to treat clinical pain and enhance opioid antinociception in non-cancer patients. A recent case report showed that PD-1 immune therapy elicited severe itch (pruritus) in an 88-year-old woman with a 7-month history of lung adenocarcinoma, and remarkably, treatment with naloxone resulted in substantial relief within 1 hour (34), suggesting a correlation of PD-1 and MOR in human. This finding also suggested that naloxone may inhibit proinflammatory cytokine production in peripheral mononuclear cells in this patient (34). Future clinical trials are warranted to test morphine-induced antinociception and morphine-induced pruritus in patients receiving anti-PD-1 immune therapy and explore the underlying immune and neuronal modulations.

MATERIALS AND METHODS

Study Design

This study was designed to evaluate the interactions between PD-1 and MOR in the PNS and CNS. First, we examined the role of PD1 in modulating opioid antinociception in tail-flick and hot-plate tests and in mouse models of bone cancer pain, inflammatory pain and neuropathic pain by using Pd1^−/−mice and/or the anti-PD1 antibody Nivolumab. Second, we checked the expression patterns of PD1 and MOR using RNAscope and immunohistochemistry, and the interaction between PD1 and MOR was tested by PLA assay and Co-IP. Third, we evaluated how PD-1 regulates opioid signaling using electrophysiology recordings in dissociated DRG neurons and spinal cord slices. Fourth, we assessed the side effects of morphine including hyperalgesia, tolerance, GIT, and withdrawal responses. Finally, we tested whether Nivolumab would impair morphine antinociception in NHPs. The sample size and power calculation was determined based on our experience with the experimental models, anticipated biological variables, previous literature (7, 35), and differences were analyzed using two tailed t-test, one-way ANOVA, and two-way ANOVA. All data were included in analyses. All experiments have been replicated successfully and all results reported have been reliably reproduced. Sample sizes and replicates are shown in the figure legends. Animals were randomly assigned into different cages and groups before experiments. In behavior tests, the investigators were blinded to drug administrations and group assignments.

Reagents

Mouse PD-L1 (#ab130039) and human IgG4 (#ab90286) were purchased from Abcam. Nivolumab (OPDIVO), a humanized anti-PD-1 antibody, was from Bristol-Myers Squibb. Morphine Sulfate was obtained from WEST-WARD pharmaceuticals. Naloxone (#1453005), Complete Freund’s Adjuvant (CFA, #F5881), DAMGO (#E7384), DPDPE (#E3888), and U69593 (#U103) were obtained from Sigma Aldrich. Tertiapin-Q (#1316) was obtained from Tocris. Fura2-AM (#F14185) was from Invitrogen.

Animals/Mice

Pd1 (Pdcd1) knockout mice with a C57BL/6 background and NOD.CB-17-Prkdc^scid mice were purchased from the Jackson Laboratory (Stock #021157 and #001303) and maintained at the Duke animal facility (7). Young mice (aged 5–7 weeks of both sexes) were used for electrophysiological studies in the spinal cord and DRG neurons. Adult male mice (aged 8–10 weeks), including knockout mice and corresponding wild-type control mice, as well as some CD1 mice (Charles River), were used for behavioral and pharmacological studies. Female mice were also used for behavioral tests in Fig 1E, G; Fig 6, A and B; Fig 7, A and B; and fig. S4. Mice were group-housed on a 12-hour light/12-hour dark cycle at 22 ± 1 °C with free access to food and water. Animals were randomly assigned to each group. Sample sizes were estimated based on our previous studies for similar types of behavioral, biochemical, and electrophysiological analyses (7, 35). Two to five mice were housed in each cage. Animal experiments were conducted in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals and approved by Duke University Institutional Animal Care and Use Committee.

Animals/Nonhuman primates

Five adult male and female rhesus monkeys (Macaca mulatta), 9.3–13.8 kg, were kept at an indoor facility accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International (Frederick, MD, USA). They were individually housed in species-specific rooms with environmental controls set to maintain 21–25°C, 40–60% relative humidity and a 12 h light (06:30-18:30)/12 h dark cycle. Their daily diet consisted of approximately 20–28 biscuits (Purina Monkey Chow; Ralston Purina Co., St. Louis, MO, USA), fresh fruit and water ad libitum. Small amounts of primate treats and various cage-enrichment devices were supplied as forms of environmental enrichment. All monkeys have been previously trained in the warm water tail-withdrawal assay. All animal care and experimental procedures were conducted in accordance with the Guide for the Care and Use of Laboratory Animals as adopted and promulgated by the US National Institutes of Health (Bethesda, MD, USA) and approved by the Institutional Animal Care and Use Committee in Wake Forest University School of Medicine (Winston-Salem, NC, USA). This study is reported in accordance with the ARRIVE guidelines for reporting experiments involving animals (36).

Mouse pain models and behavioral testing

Inflammatory pain: peripheral inflammation was induced by intraplantar injection of 20 μL CFA (Sigma) in a hind paw under brief anesthesia with isoflurane.

Neuropathic pain: To produce spinal nerve ligation (SNL), animals were anesthetized with isoflurane and the L6 transverse process was removed to expose the L4 and L5 spinal nerves. The L5 spinal nerve was then isolated and tightly ligated with 6–0 silk thread.

Bone cancer pain: Lewis lung carcinoma cell line (LLC1) was obtained from ATCC. Before the inoculation, the cancer cells were digested with 0.25% trypsin and made into a suspension of 1×10⁸/ml cells in PBS. Mice were anesthetized with 3% isoflurane, and a 0.5–1 cm superficial incision was made near the knee joint to expose the patellar ligament. Then a 25-gauge needle was inserted at the site of the intercondylar notch of the left femur into the femoral cavity. The needle was connected with a 10 μL microinjection syringe containing 2 μL suspension of 2 × 10⁵ tumor cells, as well as 2 μL absorbable gelatin sponge solution for the closure of the injection site. The contents of the syringe were slowly injected into the femoral cavity for a duration of 2 min. To further prevent leakage of tumor cells, the outside injection site was sealed with silicone adhesive (Kwik-Sil, World Precision Instruments). The wound was then closed with silk sutures and re-swabbed.

Drug injection in mice

For intravenous injection, anti-PD-1 antibody (Nivolumab, 10 mg/kg in 100 μl saline) or control antibody (human IgG4) was administered into the tail vein of mouse. For intrathecal injection, spinal cord puncture was made by a Hamilton microsyringe (Hamilton) with a 30-G needle between the L5 and L6 level to deliver reagents (5 μl) to the cerebral spinal fluid. For subcutaneous injection, morphine (1, 3, 10, 30 mg/kg in 250 μl saline) was injected beneath the back skin with a 30-G needle.

Behavioral tests of pain in mice

All animals were habituated to testing environment for at least 2 days before the baseline testing. All the tests were concluded blindly.

Tail-flick test. Mice were gently held by hand with a terry glove. The exposed distal part of the tail (3 cm) was immersed into 50°C hot water. The tail-flick latency was defined as the time required for a mouse to flick or remove its tail out of hot water. A maximum cut-off value of 15 seconds was set to avoid thermal injury. Tail-flick latency was assessed before and after drug injection. Data are also expressed as the maximum possible effect (MPE), calculated as MPE (%) = 100 × [(postdrug response – baseline response)/(cutoff response − baseline response)]. The MPE (%) data from each animal were also converted to area under the curve (AUC).

Hot-plate test. For all the experiments, hot-plate test was conducted after tail-flick test. Mice were placed on the hot plate at 53 °C and the reaction time was scored when animal began to exhibit signs of pain avoidance such as jumping or paw licking. A maximum cut-off value of 40 seconds was set to avoid thermal injury.

Mechanical sensitivity. For CFA induced inflammatory pain, SNL induced neuropathic pain, and bone cancer pain, as well as morphine-induced hyperalgesia, mechanical sensitivity was assessed by von Frey hairs. Animals were habituated to the testing environment daily for at least 2 d before baseline testing. Animals were confined in boxes placed on an elevated metal mesh floor and the hind paws were stimulated with a series of von Frey hairs with logarithmically increasing stiffness (0.02–2.56g, Stoelting), presented perpendicularly to the central plantar surface. We determined the 50% paw withdrawal threshold by the up-down method. To determine mechanical allodynia, we also tested paw withdrawal frequency using a low-threshold filament (0.4 g).

Thermal sensitivity was tested using Hargreaves radiant heat apparatus (IITC Life Science). The basal paw withdrawal latency was adjusted to 9–12 s, with a cutoff of 20 s to prevent tissue damage.

Opioid-induced tolerance in mice

Chronic morphine treatment was used to induce antinociceptive tolerance. Briefly, mice were given s.c. injection of 10 mg/kg morphine daily for 6 days. The tail-flick and hot plate latency was tested before and 1 hour after each morphine injection. The MPE (%) value was calculated to compare the antinociceptive effects and tolerance development. Additional WT and Pd1^−/− mice were used to examine microgliosis 4 days after morphine treatment.

Tail-withdrawal test and drug injection in nonhuman primates

The warm water tail-withdrawal assay was used to evaluate nociceptive responses modulated by ligands administered intrathecally (28). These monkeys have been implanted with the intrathecal catheter for drug infusion. They seated in primate restraint chairs and the lower part of their shaved tails (approximately 15 cm) was immersed into a thermal flask containing water maintained at either 46 or 50°C. Water at 46°C was used as non-noxious stimuli to detect allodynic/hyperalgesic responses, and 50°C water was used as an acute noxious stimulus to assess antinociceptive effects (27). Tail-withdrawal latencies were measured at each temperature using a computerized timer by experimenters who were unaware of the experimental conditions. If monkeys did not remove their tails within 20 s (cut-off), the flask was removed and a maximum time of 20 s was recorded. Test sessions began with baseline measurements at each temperature. Subsequent tail-withdrawal latencies were measured at multiple time points after intrathecal administration of human IgG4 (100 μg) or Nivolumab (0, 10, 30, or 100 μg). In addition, a single dose (0, 30, or 100 μg) of Nivolumab or human IgG4 (100 μg) was administered intrathecally 1 h before morphine 30 μg in order to determine if Nivolumab interferes with morphine antinociception.

Whole-cell patch clamp recording in mouse spinal cord slices

Mice of both sexes (5–7 weeks old) were anesthetized with urethane (1.5–2.0 g/kg, i.p.). The lumbosacral spinal cord was quickly removed and placed in ice-cold dissection solution (Sucrose 240 mM, NaHCO₃ 25 mM, KCl 2.5 mM, NaH₂PO₄ 1.25 mM, CaCl₂ 0.5 mM and MgCl₂ 3.5 mM), equilibrated with 95% O₂ and 5% CO₂. The slices) which was saturated with 95% O₂ and 5% CO₂. After spinal extraction under anesthesia, animals were sacrificed by decapitation. Transverse spinal slices (300–400 μm) were prepared using a vibrating microslicer (VT1200s Leica). The slices were incubated at 32℃ for at least 30 min in ACSF (NaCl 126 mM, KCl 3 mM, MgCl₂ 1.3 mM, CaCl₂ 2.5 mM, NaHCO₃ 26 mM, NaH₂PO₄ 1.25 mM and glucose 11 mM), equilibrated with 95% O₂ and 5% CO₂. The slices were placed in a recording chamber and completely submerged and perfused at a flow rate of 2–4 ml/min with ACSF which was saturated with 95% O₂ and 5% CO₂ and maintained at room temperature. Lamina II neurons in lumbar segments were identified as a translucent band under a microscope (BX51WIF; Olympus) with light transmitted from below. Whole-cell voltage-clamp recordings were made from outer lamina II neurons by using patch-pipettes fabricated from thin-walled, fiber-filled capillaries. Patch-pipette solution used to record spontaneous excitatory postsynaptic currents (sEPSCs) contained: K-gluconate 135 mM, KCl 5 mM, CaCl₂ 0.5 mM, MgCl₂ 2 mM, EGTA 5 mM, HEPES 5 mM, Mg-ATP 5 mM (pH 7.3 adjusted with KOH, 300mOsm). The patch-pipettes had a resistance of 8–10 M. The sEPSCs recordings were made at a holding potential (VH) of −70 mV in the presence of 10 μM picrotoxin and 2 μM strychnine. Signals were acquired using an Axopatch 700B amplifier. The data were stored and analyzed with a personal computer using pCLAMP 10.3 software. sEPSC events were detected and analyzed using Mini Analysis Program version 6.0.3. The histograms of sEPSC were examined for the events in 1 min before morphine treatment and in 1 min after morphine washout. The D value was measured by the Kolmogorov-Smirnov test and indicates the distance between two curves (37). Morphine-induced outward currents were recorded in whole-cell patch clamp configuration and analyzed by Clampfit 10.6. Briefly, the current value was defined by the peak amplitude value of an outward current subtracted the amplitude of baseline current under voltage-clamp mode. Additionally, some neurons were treated with Tertiapin-Q (100 nM), a blocker of G-protein-activated inwardly rectifying (GIRK) potassium channels (38). In all cases, n refers to the number of the neurons studied. All drugs were bath applied by gravity perfusion via a three-way stopcock without any change in the perfusion rate.

Spinal long-term potentiation (LTP) of C-fiber-evoked excitatory postsynaptic currents (eEPSCs) was elicited in lamina IIo neurons of spinal cord slices by stimulating the dorsal roots attached to the slices, using whole cell voltage-clamp recording (21, 22). The dorsal root electrical stimulation was performed by an electrode with a constant current source of pulse at an interval of 30 seconds. The stimulus (0.1 ms, 30 s interval, 120–300 μA) was applied at an intensity of 1.2 times of the threshold to elicit EPSCs and prevent a conduction block of action potential in the dorsal root. DAMGO (0.5 μM) was bath-applied for 3 min to induce LTP and amplitudes of individual C fiber-eEPSCs were normalized to pre-drug (control) values. LTP was defined as >20% increase in amplitude after DAMGO treatment.

Statistical analysis

All data were expressed as Mean ± S.E.M, as indicated in the figure legends. Statistical analyses were completed with Prism GraphPad 6.1. Behavioral data were analyzed using two-tailed Student’s t-test (two groups), one-Way or two-Way ANOVA (repeated measures over a time course) followed by post-hoc Bonferroni test. Electrophysiological data were tested using one-way ANOVA (for multiple comparisons) or two-tailed student’s t-test (two groups). The criterion for statistical significance was P < 0.05.

Supplementary Material

Supplemental methods and figures

Fig. S1. Dose-response of subcutaneous morphine antinociception in WT and Pd1^−/− mice.

Fig. S2. Antinociception of MOR, DOR, or KOR agonist in WT and Pd1^−/− mice.

Fig. S3. Morphine antinociception in WT and immune-deficient CB-17 mice pretreated with Nivolumab.

Fig. S4. Morphine antinociception in female Pd1^−/− mice and WT mice pretreated with Nivolumab.

Fig. S5. Proximity ligation assay (PLA) showing PD-1/MOR interaction in cultured mouse DRG neurons and co-expression of PD-1/MOR on mouse dorsal roots and Co-IP analysis in human dorsal roots.

Fig. S6. PD-1 is co-expressed with MOR and CGRP in axonal terminals in mouse spinal dorsal horn.

Fig. S7. Effects of morphine, DAMGO and PD-L1 on sEPSCs in lamina IIo neurons of spinal cord slices in WT and Pd1^−/− mice.

Fig. S8. Anti-PD-1 treatment with Nivolumab increases sEPSC frequency in lamina IIo neurons of spinal cord slices.

Fig. S9. Effects of PD-L1 on sEPSCs and outward currents in lamina IIo neurons of spinal cord slice in WT and Pd1^−/− mice.

Fig. S10. PD-L1-induced inhibition of sEPSC in lamina IIo neurons in spinal cord slice and the effect of naloxone.

Fig. S11. PD-L1-induced antinociception in mice and MOR-mediated calcium signaling in vitro.

Fig. S12. Morphine-induced gastrointestinal transit (GIT) and naloxone-induced withdrawal behavior in WT and Pd1^−/− mice.