C57BL/6 mice (male, 6–8 weeks, 20–25 g) were obtained from Liaoning Changsheng Biological Center, Shenyang, China. The mice were housed in a centralized location and kept under standard laboratory conditions (5 animals per individual cage, 24°C ± 2°C, and 50%–70% humidity under a 12/12-h light/dark cycle) with free access to water and food. The procedures used in the animal experiments were approved by the Animal Care and Ethics Committee of China Medical University (Issue number: KT2022115). Utmost care was taken to ensure the welfare of the rodents, and their usage was kept to a minimum.

Complete Freund’s adjuvant (F5881) was purchased from Sigma‒Aldrich Canada Ltd. S1P (HY-108496, 5 µL, 100 Um), SK1-I (HY-119016), FTY720 (HY-12005), siponimod (HY-12355), and KRP-203 (HY-13660) were purchased from MCE (Mair et al., 2011). Inflammatory pain was induced by intraplantar injection of 20 μL CFA in the left hind paws. For intragastric administration, the mouse was fixed in an upright position, and the gavage needle was gently inserted into the mouth and then the esophagus along the posterior wall of the pharynx to stimulate the mice to swallow. The needle was quickly inserted into approximately 3–4 cm, and the liquid was injected. After intragastric administration, the gavage needle was gently removed to observe whether there was any leakage, and attention was given to the health status of the mice. For intrathecal injection, mice were fixed, and the intervertebral space of L5-L6 was used as the injection position to align the needle along the midline of the spine. The needle was inserted gently vertically, and when touching the bone, the angle was slowly reduced to approximately 30°, the needle was slid into the intervertebral space, and the liquid was injected. Placement was confirmed by a lateral tail flick as the needle entered the subarachnoid space.

All behavioral tests were performed in an individual Plexiglas chamber on an elevated metal mesh floor under stable room temperature and humidity. Mice were habituated to the environment for at least 1 h before the experiments. The paw withdrawal response frequency (PWF) was measured to assess mechanical allodynia using Dixon’s updown method. A 0.07- or 0.4-g von Frey filament was applied ten times with 10-s time intervals in the plantar surface of a hind paw until it bent into an S shape for a duration of ≤ 6 s. A positive reaction was recorded as rapid paw withdrawal, licking or shaking of the paws. The percentage of paw withdrawal responses after each filament stimulus was counted as the frequency. The paw withdraw latency (PWL) was used to assess thermal hyperalgesia. Mice were placed on the glass plate of the Hargreaves apparatus. A radiant heat source through a keyhole was applied to the dorsal surface of the hind paw (maximum time of 20 s to eliminate any tissue injury), and the latency to lift, lick, or withdraw the paw was determined for all the mice. All behavioral tests were performed in a blinded manner.

Whole blood samples collected in the mouse serum separation tube were placed at room temperature for 2 h and then centrifuged at 1,000 × g for 20 min, and the supernatant was obtained for follow-up experiments. The mouse tissues were washed with precooled PBS (0.01 m, pH = 7.4), the residual blood was removed, and the tissues were cut into pieces. The chopped tissue and the corresponding volume of PBS were placed into the grinder to split the tissue cells. The homogenate was centrifuged at 5,000 × g for 10 min, and the supernatant was taken for follow-up experiments. The specific determination method was performed according to the ELISA kit (Echelon Biosciences). Fifty microliters of standard working solution or sample was added to each well, and then 50 μL of biotin conjugate working solution was added immediately and incubated at 37°C for 60 min. The liquid was discarded, and 200 μL of washing buffer was added to each well and washed 3 times. After drying, 100 μL HRP enzyme was added to each well and incubated at 37°C for 60 min. The liquid was discarded, and 200 μL of washing buffer was added to each well and washed 5 times. After drying, 90 μL TMB was added to each well and incubated at 37°C for 20 min. Fifty microliters of terminator solution was added to each well, and the 450 nm OD value was read immediately.

Total RNA was extracted from ipsilateral hind paw and L4-6 spinal cord tissue using TRIzol reagent (Invitrogen). cDNA was synthesized from total RNA (1 μg) using Hifair II 1st strand cDNA Synthesis SuperMix (Yeasen, China). The reverse transcription procedure was performed at 25°C for 25 min and 42°C for 30 min followed by 85°C for 5 min. The sequences of the PCR primers (Sango Biotech, Shanghai, China) are listed in Table 1. qPCR was carried out at 95°C for 2 min followed by 40 cycles of 95°C for 15 s and 60°C for 1 min. qPCR was conducted using the SYBR-Green System (Yeasen, China). Relative gene expression was calculated using the 2^−ΔΔCT method, with GAPDH as the internal control and reference gene.

Mice were deeply anesthetized with 1% sodium pentobarbital and transcardially perfused with 20 mL of 0.9% cold saline followed by 4% paraformaldehyde in 0.1 M phosphate-buffered saline (PBS, pH 7.4) via the cardiovascular system. After perfusion, the L5 spinal segments were removed and postfixed overnight at 4°C. The next day, the segments were dehydrated in a sucrose gradient (20% and 30% sucrose in PBS) at 4°C for 24 h until they completely sank to the bottom. Serial frozen 30-µm spinal cord sections were cut on a coronal plane using a cryostat and prepared for immunofluorescence staining. After washing with phosphate-buffered saline, the sections were blocked with 0.1% BSA for 1 h at room temperature, followed by incubation overnight at 4°C with the primary antibodies rabbit anti-S1PR1 (1:200; Proteintech # 55133-1-AP), goat anti-IBA-1 (1:200; Abcam # ab289874), mouse anti-GFAP (1:200; Proteintech # 60190-1-Ig), and mouse anti-NeuN (1:200; Abcam # ab279296) diluted in PBS. The sections were incubated with the corresponding secondary antibodies for 1 h after washing three times with PBS at room temperature. The stained sections were mounted with 4′, 6-diamidino-2-phenylindole (DAPI) (Abcam #ab104139), and images were captured using a fluorescence microscope (Leica).

L4-6 spinal cord tissues were homogenized and lysed on ice for 30 min in 100 μL RIPA lysis buffer solution (Beyotime Biotechnology) containing protease and phosphatase inhibitors. The homogenates were centrifuged at 12,000 rpm for 15 min at 4°C. The protein samples were quantified using the BCA protein assay (Beyotime Biotechnology). The proteins were separated on a 10% SDS‒PAGE gel (Bio-Rad) and transferred to PVDF membranes, which were then blocked with 5% BSA in TBS-Tween. The blots were probed with antibodies against S1PR1 (1:1,000, Proteintech # 55133-1-AP), STAT3 (1:1,000, Cell Signaling # 9139), phosphorylated STAT3 (1:1,000, Cell Signaling #9131), ERK (1:1,000, Cell Signaling #9102), phosphorylated ERK (1:1,000, Cell Signaling #4370), phosphorylated P38 (1:1,000, #4511), p38 (1:1,000, #9212), and GAPDH (1:10,000, Zhongshan Golden Bridge Biotechnology #TA-08). The blots were incubated with horseradish peroxidase-labeled rabbit anti-mouse (1:5,000, Abcam #ab6789) or goat anti-rabbit (1:5,000, Abcam #ab6721) secondary antibodies. The protein bands were visualized with enhanced chemiluminescence reagents (Sigma), and the band densities were analyzed using ImageJ software. All experiments had at least four biological replicates, and representative blots are presented in the figures.

The data are expressed as the mean ± standard error of the mean (SEM). Statistical analysis was performed using Prism GraphPad 6.0. For the behavioral tests, the differences between groups were analyzed using two-way repeated-measures ANOVA followed by Bonferroni’s post hoc test. For immunofluorescence, RT‒PCR, and Western blotting, differences between groups were compared by one-way ANOVA with Tukey’s post hoc test. For all comparisons, p < 0.05 indicated statistical significance.

The CFA model was selected as a chronic nociceptive pain model. We found that the content of S1P in the serum of CFA mice was significantly increased, especially on days 1 and 3. In the dorsal root ganglion (DRG), the level of S1P increased from day 1 to day 7, and in the spinal cord, the expression of S1P began to increase 3 days after CFA model induction (Figure 1A). To further verify the role of S1P in pain, exogenous S1P was administered through intrathecal injection, and the mice showed transient mechanical and thermal hyperalgesia (Figures 1B–D). S1P regulates the development of neuroinflammation in the nervous system (Wang et al., 2021). Neuroinflammation is typically characterized by the activation of microglia and astrocytes (Kasatkina et al., 2021). Through immunofluorescence detection, we found that administration of S1P activated microglia and astrocytes in the dorsal horn of the spinal cord in the CFA model, which was closely related to nociceptive pain (Figures 1E–G). These results indicated that pathological pain could cause an increase in S1P expression and that S1P may participate in the occurrence of nociceptive pain.

S1P is a signaling lipid synthesized by sphingosine kinases (SPHK1 and SPHK2) (Jozefczuk et al., 2020; Sukocheva et al., 2020). Since S1P is a pain-inducing substance, does the inhibition of S1P synthesis reduce the nociceptive pain caused by CFA? We found that administration of the SPHK1 inhibitor SK1-I (10 mg/kg) reduced the content of S1P in the serum, DRG and spinal cord in the CFA model (Figure 2A). Intraperitoneal administration of SK1-I (10 mg/kg) reversed the nociceptive pain caused by CFA (Figures 2B–D). SK1-I inhibited the activation of microglial cells and astrocytes in the spinal dorsal horn 3 days after CFA model induction (Figures 2E–G).

Previous studies have found that S1P plays a biological role by activating the GPCR signaling pathway through the binding of S1PRs, a homologous receptor on the cell surface. By detecting the mRNA expression of S1PRs in the spinal cord of CFA mice, we found that the mRNA expression level of S1PR1 was significantly increased (Figure 3A). The expression of S1PR1 mRNA was upregulated 1, 3, and 14 days after CFA model establishment (Figure 3B). To further verify the role of S1PR1 in nociceptive pain, we selected several competitive and functional S1PR1 antagonists: FTY720, KRP-203 and siponimod. S1PR1 antagonists were administered daily on day 1 after CFA was established via intragastric (i.g.) administration, and PWF and PWLwere detected on days 0, 1, 3, 5, 7, and 11 of CFA (Figures 3C–K). Irrespective of whether FTY720, KRP-203 or siponimod were provided, PWFs were decreased and PWLs were increased from day 3 after CFA in a dose- and time-dependent manner (Figures 3C–K).

Next, we evaluated mechanical pain and thermal pain in mice after knockdown of S1PR1. The knockdown effect of S1PR1 siRNA was proven by RT‒PCR (Figure 4A). 3 days after CFA, knockdown of S1PR1 via siRNA resulted in markedly decreased PWF and increased PWL, which lasted for 4 h (Figures 4B–D). Western blotting and immunofluorescence also demonstrated that knockdown of S1PR1 reversed the increase in S1PR1 protein levels caused by CFA (Figures 4E–G). Next, we determined the cell location of S1PR1. Immunofluorescence showed that in the CFA model, S1PR1 was specifically expressed in astrocytes, and no obvious colocalization was found in microglia and neurons (Figure 4H).

To determine why S1PR1 was involved in the development of pain, we combined bioinformatics analysis to identify the proteins that interact with it. The protein‒protein interaction network was created and visualized using Cytoscape. STAT3, ERK, and p38 MAPK have been reported to be important pain-causing molecules (Koichi and Noguchi, 2004; Wan et al., 2017), which suggests that the activation of S1PR1 may activate the downstream pain signaling pathway. Previous studies have found that the activation of STAT3, ERK, and p38 MAPK in glial cells promotes the release of inflammatory factors. Studies on the STAT3 and MAPK families in pain indicated that phosphorylation of these proteins was involved in pain sensitivity caused by inflammation and neuropathic pain (Zhao et al., 2021a; Zhao et al., 2021b). Therefore, we speculate that the activation of S1PR1 leads to the upregulation of the STAT3, ERK, and p38MAPK signaling pathways and participates in the development of pain. On this basis, we verified by Western blotting that the phosphorylation of STAT3, ERK, and p38MAPK decreased significantly after S1PR1 was knocked down (Figures 5A–F). Previous studies have found that activation of glial cell S1PR1 contributes to the development of neuropathic pain, specifically knocking out the S1PR1 allele in GFAP-positive astrocytes, and no SNL-induced neuropathic pain occurs (Chen et al., 2019; Doyle et al., 2020a). Increasing evidence has shown that microglia and astrocytes play an important role in the development of inflammatory pain caused by CFA (Ji et al., 2016). In our study, after using siRNA to inhibit S1PR1, the activation of microglia and astrocytes was significantly inhibited (Figures 5G–J). It is speculated that the activation of S1PR1 on glia is involved in the occurrence of pain.

Our previous research found that the expression levels of proinflammatory factors such as IL-1β and TNF-a increased, and the expression level of the anti-inflammatory factor IL-10 decreased. IL-1β and TNF-a are currently widely recognized pain and inflammatory factors that play an important role in the occurrence of neuroinflammation (Pinho-Ribeiro et al., 2017). In the CFA pain model, there was a significant increase in IL-1β mRNA and protein levels (Figures 6A, D, E). Newly generated IL-1β precursors must be hydrolyzed into mature forms by proteases to have pain-inducing effects (Takenouchi et al., 2011; Arman et al., 2020). Recent research shows that during the occurrence and development of pain, the cleavage of IL-1β mainly depends on MMP9 and MMP2 matrix metalloproteinases, and MMP9 cleaves IL-1β to control the earliest stage of pain (<1d). MMP2 cleaves IL-1β, which controls the late stage of development and maintenance of pain (<3d) (Ji et al., 2009). In our study, after inhibiting S1PR1 with siRNA, the expression of MMP2 was inhibited (Figures 6A, B). S1P/S1PR1 induced phosphorylation of NF-κB p65 and activation of NF-κB nuclear translocation (Zheng et al., 2019). We also proved that knockdown of S1PR1 reversed the activation of NF-κB induced by CFA (Figures 6A, C). In addition, the knockdown of S1PR1 inhibited the activation of the inflammatory factors IL-1β, IL-6, TNF-a and iNOS and increased the mRNA levels of the anti-inflammatory factors IL-10 and Arg-1 (Figures 6E–J).