To identify the primary voltage-gated sodium channel in subchondral sensory fibers that responsible for mechanical hypersensitivity during OA progression, we tested the expression levels of different sensory related sodium channel Na_Vs in OA mice post anterior cruciate ligament transection (ACLT). The transcription levels of mRNAs that encode Na_Vs in DRG including Na_V1.1, Na_V1.2, Na_V1.3, Na_V1.6, Na_V1.7, Na_V1.8, Na_V1.9 were measured by qPCR using mRNA isolated from mouse ipsilateral L3-5 DRGs one month post-ACLT or sham surgery. Compares to the sham-operated group, the expression of mRNA encoding Na_V1.8 increased 2.5 folds in the ACLT group as the highest upregulation among all the Na_Vs. The mRNAs encoding Na_V1.7 and Na_V1.9 showed moderate upregulation in the ACLT group relative to that of the Sham group while the changes of Na_V1.1 Na_V1.2 Na_V1.3 or Na_V1.6 were not detected (Figure 1a). Therefore, we further investigated Na_V1.8 protein expression in the immune-histological analysis of OA subchondral bone. The intensity of Na_V1.8 immunofluorescence in subchondral bone was also elevated about 2 to 3 folds in OA mice compared to sham- operated mice one- or two-months post-surgery (Figure 1b and c). To examine whether the increase of Na_V1.8 expression is limited to in a certain subtype(s) of the DRG neuron that innervates subchondral bone in OA mice, Na_V1.8 was co-stained with different markers for sensory nerve subtypes based on the current classification of DRG neurons (Usoskin et al., 2015). The expression rate of Na_V1.8 in total nerve fibers (labeled by pan neuron marker PGP9.5) innervated in subchondral bone significantly increased post-ACLT (Figure 1—figure supplement 1a,f). Moreover, the elevated Na_V1.8 expression was highly co-localized with the peptidergic nociceptor marked by calcitonin gene-related peptide (CGRP) (Brain et al., 1985; Figure 1—figure supplement 1b,f) and mechanosensitive low-threshold mechanoceptors (labeled by PIEZO2) (Eijkelkamp et al., 2013). The expression of Na_V1.8 was also slightly elevated in the synovium (Figure 1—figure supplement 1f 1 hr, j). Both western blot analysis (Figure 1f) and immunostaining (Figure 1—figure supplement 1f 1 g, i) of ipsilateral lumbar 3–5 DRG confirmed the upregulation of Na_V1.8 expression at the DRG level. We then further validated whether DRG neurons with upregulated Na_V1.8 expression directly innervates fibers in the subchondral bone. We injected a neurophilic fluorescent dye (DiI) into the subchondral bone to label the distal nerve fibers (Ferreira-Gomes et al., 2010). We found that the Na_V1.8⁺ neurons labeled by DiI significantly increased in the DRG neurons of OA rats relative to sham-operated rats (Figure 1g,h), indicating that DiI was transported into DRG neurons by the sensory fibers innervated subchondral bone in a retrograde manner. These results suggest that the expression of pain-related sodium channel Na_V1.8 is upregulated in DRG neurons and their axons that innervate subchondral bone during progression. We then examined the association between Na_V1.8 neuronal activity and OA pain. Von Frey test showed that the hind paw withdrawal threshold (HPWT) dropped nearly 60% and maintained at this level throughout the two month-period post ACLT relative to the sham-operated group, suggesting that development of mechanical allodynia in OA mice (Figure 1g; Chen et al., 2017). To assess the potential role of Na_V1.8 in DRG neuronal excitability, we used an in vivo DRG imaging in Pirt^GCaMP3fl/-mice that we recently developed. In this genetically targeted mice, genetic-encoded Ca²⁺ indicator GCaMP3 is specifically expressed in >95% of all DRG neurons under the control of the Pirt promoter. In the Pirt^GCaMP3fl/-mice, the excitability of the nociceptive neurons in DRG can be visualized by fluorescence signals of calcium influx. The number of excited DRG neurons ipsilateral to the surgery significantly increased in OA mice compared to sham-operated mice, and importantly, administration of Na_V1.8 inhibitor (A-803467)(Jarvis et al., 2007) blunted the signal in DRG (Figure 1h and i). To validate the excitation of DRG neurons related to Na_V1.8, we performed patch-clamp in the DRG neurons that were isolated from mice that underwent ACLT or sham surgery. The action potential number and Na_V1.8 current density significantly elevated in ACLT mice relative to sham-operated mice, and the elevations were blocked by A-803467 (Figure 1j–l, Figure 1—figure supplement 1f 1 k and l). Thus, the activation of Na_V1.8 mediates OA pain related DRG neuron hypersensitivity.

We then examined the mechanism of upregulation of Na_V1.8 expression during OA progression. To examine whether excessive PGE2 contributes to the upregulation of Na_V1.8, we firstly performed immunostaining of cyclooxygenase 2 (Cox2) in subchondral bone sections. Cox2 expression was significantly increased in subchondral bone and primarily in osteocalcin positive osteoblastic cells post ACLT mice compared with sham-operated mice (Figure 2a and b). Consistently, PGE2 concentration in subchondral bone increased about three times in OA mice relative to sham-operated mice (Figure 2c, Figure 2—figure supplement 1a–d). To examine if elevated PGE2 upregulates the expression of Na_V1.8 for the mechanical allodynia in OA, we generated osteoblast specific Cox2 deficient mice (Cox2_Oc^-/- mice) by crossbreeding Cox2^fl/fl mice with Bglap-Cre mice. PGE2 concentration in subchondral bone was significantly lower in Bglap-Cre::Cox2^fl/fl mice compared with Cox2^fl/fl mice post ACLT (Figure 2h). Notably, the Na_V1.8 immunofluorescence intensity in subchondral bone immunostaining was also significantly reduced in Bglap-Cre::Cox2^fl/fl mice compared with Cox2^fl/fl mice (Figure 2d and i). In addition, the number of Na_V1.8⁺ neurons in DRG was also significantly decreased (Figure 2e and j) in the Bglap-Cre::Cox2^fl/fl mice relative to Cox2^fl/fl mice. We then investigated whether a decrease of PGE2 alleviates OA pain. We crossed Bglap-Cre::Cox2^fl/fl mice with Pirt ^GCaMP3fl/- mice to measure the DRG neuronal excitability in Bglap-Cre::Cox2^fl/fl::Pirt ^GCaMP3fl/- mice post ACLT. Pirt-GCaMP3 DRG imaging showed that the number of excited neurons was significantly reduced in Bglap-Cre::Cox2^fl/fl::Pirt^GCaMP3fl/- mice compared with Cox2^fl/fl::Pirt ^GCaMP3fl/- mice post ACLT (Figure 2f and k). Moreover, the single neuron excitability in ipsilateral L4 DRG was functionally tested by whole cell patch clamp electrophysiology. The whole cell current clamp revealed that the action potential firing number was significantly decreased in Bglap-Cre::Cox2^fl/fl mice compared with Cox2^fl/fl mice after ACLT (Figure 2g and l). Concurrently, the Na_V1.8 current density was reduced for about 40% recorded by the whole-cell voltage-clamp (Figure 2g and m). The mechanical allodynia was simultaneously attenuated in Bglap-Cre::Cox2^fl/flmice relative to Cox2^fl/fl mice as revealed by the Von Frey behavior test (Figure 2n). Catwalk analysis also showed that the Maximal Contact At and Maximal Intensity of ipsilateral hind paw was significantly higher in Bglap-Cre::Cox2^fl/fl mice compared with Cox2^fl/fl mice post ACLT (Figure 2o). Thus, PGE2 derived from osteoblastic cells stimulates the pain hypersensitivity in OA mice likely by upregulating of Na_V1.8 in subchondral nociceptive neurons.

To examine whether EP4 at sensory neurons is the primary receptor that propagates PGE2 signals in upregulating Na_V1.8and OA pain, we specifically knocked out EP4, the skeletal pain related receptor for PGE2 (Yoshida et al., 2002), in peripheral sensory nerves by crossbreeding Advillin-Cre (Avil-Cre) (Zurborg et al., 2011) mice with Ptger4 ^fl/fl mice (Ptger4 is the gene that encodes EP4 receptor). Consistently with Bglap-Cre::Cox2^fl/fl mice, the intensity of Na_V1.8 immunofluorescence was significantly reduced in Avil-Cre::Ptger4 ^fl/fl mice compared with Ptger4 ^fl/fl mice post-ACLT (Figure 3a,d, Figure 2—figure supplement 1e–h). We then further confirm this finding in DRG neurons that cultured in PGE2. We found that Na_V1.8 protein expression was significantly reduced by siRNA against EP4 in western blot analysis of levels from the cell lysates. Knocking-down the expression of EP1-EP3 did not have a significant effect on Nav1.8 expression (Figure 2—figure supplement 1i). To determine the effect of conditional deletion of EP4 on DRG neuronal excitability, we generated Avil-Cre::Ptger4 ^fl/fl::Pirt - GCaMP3^fl/- and Ptger4 ^fl/fl::Pirt ^GCaMP3fl/- ACLT mice. In vivo ipsilateral L4 DRG Pirt ^GCaMP3 imaging demonstrated significantly dampened excitability in the Ptger4 ^fl/fl:: Pirt - GCaMP3^fl/- mice relative to the control group (Figure 3b and e). The patch-clamp analysis further revealed that the DRG neuronal hypersensitivity and Na_V1.8 currents were significantly reduced in Avil-Cre::Ptger4 ^fl/fl mice relative to Ptger4 ^fl/fl mice post-ACLT (Figure 3c,f,g). Moreover, both Von Frey test and catwalk analysis showed attenuation of OA pain when EP4 is conditionally deleted in sensory neurons (Figure 3h and i). Thus, the EP4 receptor expressed in DRG neurons is responsible for the propagation of subchondral PGE2-induced upregulation of Na_V1.8 and neuronal excitability in OA mice.

To investigate the mechanism of PGE2 stimulated upregulation of Na_V1.8 expression, RT-qPCR was performed with mRNA isolated from primary DRG neurons treated with PGE2. The result showed that Na_V1.8 transcription levels were significantly elevated at 6 and 12 hr after incubation with PGE2 (Figure 4a). Moreover, PGE2 stimulated phosphorylation of protein kinase A (PKA) (Gold et al., 1998) and cAMP response element-binding protein (Creb1) (Lonze and Ginty, 2002; Figure 4b). Notably, the effect of PGE2 and Forskolin, a cAMP stimulant in the upregulation of Na_V1.8 protein expression was dampened by Creb1 inhibitor 666–15 (Figure 4c), indicating PGE2 stimulates Nav1.8 expression through the PKA-Creb1 signaling pathway. Consistently, the neuronal excitability and Na_V1.8 current density stimulated by PGE2 was abolished by the application of PKA inhibitor (PKI) or CREB inhibitor 666–15 (Figure 4d–f). Co-immunofluorescence staining further demonstrated that PKA levels significantly increased in Na_V1.8 positive DRG neurons in ACLT mice compared with sham-operated mice (Figure 4g and h). To examine the mechanism of PGE2-induced Na_V1.8 transcription, we performed chromatin immunoprecipitation (ChIP) assay with three potential pCreb1-binding elements in the Na_V1.8 promoter. ChIP assay revealed that pCreb1 binds to the Na_V1.8 promoter at binding site two to stimulate transcription of Na_V1.8 gene (Figure 4i–k). Taken together, our findings reveal that PGE2 induces transcription of Na_V1.8 by stimulation phosphorylation PKA and pCreb1, which directly binds to Na_V1.8 promoter.

We next tested whether the deletion of Na_V1.8⁺ neurons could attenuate OA pain. Scn10a -Cre mice were crossed with Rosa26^iDTRfl/fl mice to generate Scn10a -Cre:: Rosa26^iDTRfl/fl mice. In these mice, the Na_V1.8⁺ neurons underwent apoptosis upon receiving the injection of diphtheria toxin (Buch et al., 2005). The ablation of Na_V1.8⁺ neurons had no effect on the articular cartilage deterioration, as shown by similar OARSI scores between Scn10a -Cre:: Rosa26^iDTRfl/fl mice and Rosa26^iDTRfl/fl mice post-ACLT (Figure 5a,e). The efficacy of specific neuron ablation was evidenced by a significant reduction of Na_V1.8⁺ signals at both subchondral bone and DRG level (Figure 5b,c,f,g). Consistently, the DRG hypersensitivity was reduced as indicated by a decreased AP firing (Figure 5d and h). We further investigated whether the ablation of Na_V1.8⁺ sensory neurons reduces the pain in OA mice by catwalk gait analysis (Lakes and Allen, 2016). The results showed that max intensity, which reflected mechanical pain sensitivity (Kameda et al., 2017), was increased in Scn10a -Cre:: Rosa26^iDTRfl/fl mice (Figure 5i). Similarly, the Von Frey test displayed a significant reduction of HPWT in Scn10a -Cre:: Rosa26^iDTRfl/fl mice compared with Rosa26^iDTRfl/fl mice after ACLT (Figure 5j). Taken together, the Scn10a -Cre:: Rosa26^iDTRfl/fl mice indicates that specific ablation Na_V1.8 can alleviate OA pain in OA mice.

We previously showed that inhibition of excessive TGF-β activity attenuated OA progression by restringing the coupling of subchondral bone remodeling (Figure 6—figure supplement 1a–n, Figure 6—figure supplement 2 4a-e). We have developed a small molecule drug by conjugating TGF-β type I receptor kinase inhibitor (TβR1I) covalently with alendronate through a metabolically cleavable carbamate linkage (Qin et al., 2018). The conjugate is effectively delivered to the bone surface where TβR1I is released by cleavage of the carbamate linkage in vivo. (Figure 6—figure supplement 3a). Administration of the conjugate in human MSCs effectively inhibited TGF-β signaling evidenced by a significant reduction of pSMAD2/3 (Figure 6—figure supplement 3b and c). As excessive PGE2 production and subsequent upregulation of Nav1.8 are triggered by abnormal bone remodeling, we investigated whether conjugate treatment can alleviate OA pain by downregulates the activity of Nav1.8. As expected, the articular cartilage degeneration was attenuated with a weekly injection of conjugate 100 ug/kg in ACLT mice compared with the vehicle group, with a significant improvement of the OARSI score (Figure 6a and g). In concurrent with the cartilage protection, the subchondral bone microarchitecture was improved in μCT analysis of ACLT mice treated with the conjugate treatment compared with the vehicle group (Figure 6b and h, Figure 6—figure supplement 3f). As shown in Figure 6c and i, phosphorylation of Smad2/3 was effectively inhibited by the conjugate in the subchondral bone. The number of TRAP⁺ osteoclastic cells and Osterix⁺ pre-osteoblast were reduced (Figure 6—figure supplement 3e and g, Figure 6d and j). As a result, the BML in tibial subchondral bone was significantly reduced in the conjugate treated group (Figure 6f) further indicating that coupling of the osteoclast bone resorption and osteoblastic bone formation were restored.

Finally, we examined whether the conjugate effect on the improvement of subchondral bone structure and articular cartilage degeneration could also relieve OA pain. Interestingly, subchondral PGE2 concentration was significantly reduced in ACLT mice with the conjugate treatment relative to the vehicle group (Figure 6l). Importantly, the expression of Na_V1.8 was also reduced in both subchondral bone (Figure 6e and k), and ipsilateral lumbar DRG (Figure 6m). Moreover, the electrophysiological tests demonstrate that data showed that conjugate treatment blunted the upregulation of DRG neuron activity (Figure 6n and o) and Na_V1.8 currents (Figure 6n and p) in ACLT mice. The effect of the conjugate on joint pain related behaviors were examined in the Catwalk test. HPWT, maximal contact AT and swing phase were significantly ameliorated in ACLT mice with the administration of conjugate relative to the vehicle group (Figure 6q and r). These data suggest that alendronate-TβR1I conjugate relieves OA pain by modifying the disease. This was likely achieved by the decrease of PGE2 in the improvement of subchondral bone structure.

We purchased C57BL/6J (WT) 3 months old male mice from Jackson Laboratories. We purchased Sprague Dawley (SD) 3 months old male rats from Charles River company. To develop the mechanical instability related OA model, we used ACLT surgery (Malfait and Little, 2015). Briefly, after ketamine and xylazine anesthesia, the left ACL was surgically transected and sham operations were performed on other groups of mice. For the time-course experiments, mice were euthanized at 4, 8 or 12 weeks after surgery (n = 6 per group).

The Rosa26^iDTRfl/fl mice were purchased from Jackson Laboratory. The Advillin-Cre (Avil-Cre) and Pirt^GCaMP3 mouse strain were kindly provided by Xingzhong Dong (The Johns Hopkins University). The Bglap-Cre mice were provided by Thomas J. Clemens (The Johns Hopkins University). The Cox2^fl/fl mice were kindly provided by Harvey Herschman (University of California, Los Angeles). The Ptger4 ^fl/fl mice were provided by Brian L. Kelsall (the National Institutes of Health). The Scn10a-Cre mice were kindly provided by Yun Guan (The Johns Hopkins University). Heterozygous Bglap-Cre mice were crossed with a Cox2^fl/fl mouse; the offspring were intercrossed to generate the following genotypes: WT, Bglap-Cre, Cox2^fl/fl, Bglap-Cre::Cox2^fl/fl. Cox2^fl/fl or Bglap-Cre::Cox2^fl/fl mice were further crossed with Pirt^GCaMP3fl/- mice to generate Cox2^fl/fl::Pirt^GCaMP3fl/- mice or Bglap-Cre::Cox2::Pirt^{GCaMP3 fl/-} mice for in vivo GCaMP3 DRG imaging. Heterozygous Avil-Cre mice were crossed with Ptger4^fl/fl mice. The offspring were intercrossed to generate the following genotypes: wild type (referred as ‘WT’ in the text), Avil-Cre (Cre recombinase expressed driven by Advillin promoter), Ptger4^fl/fl, Avil-Cre:: Ptger4^fl/fl (conditional deletion of EP4 receptor in Advillin lineage cells). Ptger4^fl/fl or Avil-Cre:: Ptger4^fl/fl mice were further crossed with Pirt-GCaMP3 ^fl/- mice to generate Ptger4^fl/fl::Pirt-GCaMP3^fl/- mice or Avil-Cre:: Ptger4^fl/fl::Pirt^{GCaMP3 fl/} mice for in vivo GCaMP3 DRG imaging. Heterozygous Scn10a-Cre mice were crossed with the Rosa26^iDTRfl/f mouse; the offspring were intercrossed to generate the following genotypes: WT, Scn10a-Cre::Rosa26^iDTRfl/fl, Scn10a-Cre::Rosa26^iDTRfl/fl mice. We injected 12-week-old Scn10a-Cre:: Rosa26^iDTRfl/fl or Rosa26^iDTRfl/fl mice with 1 μg/kg DTX intraperitoneally three times per week after ACLT for 4 weeks. We obtained femurs, tibiae and DRG from the mice after euthanasia. For conjugate injections, we used intraperitoneal injection method and 1 mg/kg per week dosage according to previous toxicological experiments. All animals were maintained at the animal facility of The Johns Hopkins University School of Medicine. All the experimental protocols were approved by the Animal Care and Use Committee of The Johns Hopkins University (Protocol number: Mo18M308).

After approval by the Institutional Review Board of The Johns Hopkins Hospital,, we collected tibial plateau specimens from eight individuals with osteoarthritis that underwent total knee arthroplasty. The knee joints from three healthy young adults underwent lower limb amputations after trauma serves as healthy controls. The demographic data of patients were collected. The samples were used to perform histology and immunohistochemistry after decalcification.

Immediately after euthanasia, we resected and fixed the animals knee joints or DRG in 10% buffered formalin for 24 hr, decalcified them in 0.5 M ethylenediaminetetraacetic acid (EDTA, pH 7.4) for 14 d and embedded them in paraffin or gelatin solution (20% D-sucrose, 2% Polyvinylpyrrolidone (PVP) and 8% gelatin in PBS). Four-micrometer sagittal oriented sections of the medial compartment of left knees were processed for hematoxylin and eosin, safranin orange and fast green and Tartrate-resistant acid phosphatase (TRAP) staining (Sigma). For immunohistology and immunofluorescence, slides (4 μm for immunohistology, 20 μm for DRG, 60 μm for knee immunofluorescence) were incubated with antigen retrieval buffer (Dako, S169984-2) at 96°C for 15 min, gradually cooled to room temperature and washed with tris-buffered saline with Tween (TBST). After blocking, the slides were incubated with primary antibodies overnight at 4°C. Secondary antibody (1:200) was used to incubate the samples for 1 hr at room temperature. For immunohistochemical staining, a horseradish peroxidase–streptavidin detection system (Dako) was used to detect immunoactivity, followed by counterstaining with hematoxylin (Sigma-Aldrich). For immunofluorescence, the fluorescent conjugated secondary antibody (1:200) was applied. The photographs of the immunohistology sections were recorded by light microscopy (DP71 microscope camera, Olympus) and analyzed by OsteoMeasure XP software (OsteoMetrics). We calculated OARSI scores as previously described (Glasson et al., 2010). The OARSI scores were evaluated by two independent graders and the averages were taken. For the immunofluorescence, the photographs were shot under laser confocal microscopy (Zeiss, LSM 780) and Zen 2.2 software.

The mice knees were scanned using high-resolution μCT (SkyScan 1275, Bruker microCT) as previously described (Zhen et al., 2013). The scanner was set at a voltage of 65 kVp, a current of 153 μA and a resolution of 5.7 μm per pixel. We reconstructed and analyzed outcomes using NRecon v1.6, and CTAn v1.9, respectively. Three-dimensional reconstructions were done by CTVol v2.0 (Bruker microCT). We defined the region of interest to cover the trabecular part of the medial compartment of tibial subchondral bone, and five consecutive images from the medial tibial plateau were used for 3-dimensional reconstruction. We analyzed 3D parameters as following: TV (total tissue volume; containing both trabecular and cortical bone), BV/TV (trabecular bone volume per tissue volume) and Tb.Pf (trabecular pattern factor).

We performed in vivo μMRI studies on a horizontal 9.4T Bruker Biospec preclinical scanner according to our previous protocol (Zhen et al., 2013). Briefly, we showed subchondral BML by T2-weighted scanning with 2D RARE (rapid acquisition with relaxation enhancement) sequence, a TE/TR (echo time/repetition time) of 15.17 ms/3,000 ms, 30 slices at 0.35 mm thickness, 1.75 cm ×1.75 cm field of view (FOV) with a matrix size of 256 × 128. The fat suppression was done in T2-weighted imaging with a chemical shift selective fat saturation pulse tuned to the fat resonant frequency.

Bilateral lumbar DRGs were harvested from 4 week male WT mice. For DRG neuron culture medium, MEM was supplemented with 5% fetal bovine serum (Gibco), 2X penicillin and streptomycin solution (Gibco), 1X GlutaMAX (Thermo Fisher), 20 μM 5-fluoro-2-deoxyuridine (Sigma-Aldrich) and 20 μM uridine (Sigma-Aldrich). DRG neurons were digested and dissociated with 1 mg/ml collagenase D (Roche) for 90 min and then 1X TrypLE Express solution (Thermo Fisher) for 15 min. The dissociated DRG neurons were placed on a precoated dish with 100 μg / ml poly-D-lysine (thermal fisher) and 10 μg / ml laminin (thermal fisher). 100 ng / ml Nerve growth factor (R and D) was applied to maintain the neuronal activity. After 24 hr incubation, PGE2 (1 μM) or PBS were applied to stimulate the DRG neurons. In vitro RNA interference was performed using commercially available RNAi products from Thermal Scientific (s72365, s72370, s72373, and s72375) and the protocol was followed by the manufacture’s instruction. Briefly, after neuron seeding for 24 hr, media were replaced for the cells to be prepared for transfection. Lipofectamine RNAi MAX (13778100, Invitrogen) was diluted in OptiMEM (31985062, Thermal Fisher) and incubated for 5 min, then mixed with siRNAs or scramble control RNAs for five mins. Diluted DNA and Lipofectamine RNAi MAX were mixed and incubated at room temperature for 20 min and then used to transfect the DRG neurons. The medium was replaced 10 hr following transfection and neurons were harvested 24 hr after transfection. Similarly, the human GFP labeled MSC was purchased from Cyagen and is cultured in MEM with 10% fetal bovine serum (Gibco), 1X penicillin and streptomycin solution (Gibco).

We used Pirt-GCaMP3^f/- mice in DRG imaging. In order to monitor the activity of large populations of DRG neurons in intact live animals, Dr Xinzhong Dong’s Lab developed an in vivo imaging technique by using Pirt^GCaMP3 genetically engineered mice, in which the genetic-encoded Ca²⁺indicator GCaMP3 is specifically expressed in >95% of all DRG neurons by Pirt promoter (Kim et al., 2008; Kim et al., 2014b).After surgical exposure of ipsilateral L4 DRG, in vivo imaging was immediately performed. Similarly as previously described (Miller et al., 2018), the animals were maintained under inhalation anesthesia with assisted ventilation through endotracheal incubation. A laser scanning confocal microscope (Leica LSI microscope system) with a water immersed lens was used to capture the fluorescent signals. Live images were acquired at 10 frames with 600 Hz in frame-scan mode per 6–7 s, at depths below the dura ranging from 0 to 70 µm. 25 g of direct compression was applied to the ipsilateral knee after ACLT or sham surgery using a rodent pincher (IITC Life Science) to stimulate DRG neuronal firing. The duration of the mechanical force application maintained 15–30 s after 40–50 s of baseline imaging and the activated neuron number was counted and analyzed.

Whole-cell current-clamp recordings were performed to perform the action potential of DR neurons according to the previous study (Bierhaus et al., 2012). Only small and medium-sized DRG neurons with a resting membrane potential more negative than −40 mV were recorded. The extracellular solution contained (in mM): NaCl 140, KCl 4, CaCl₂ 2, MgCl₂ 1; HEPES 10, NaOH 4.55, glucose 5 (pH 7.4, 300–310 mOsm/kg H₂O). The pipette solution contained (in mM): KCl 135, MgCl₂ 0.1, Mg-ATP 1.6, HEPES 10, EGTA 2, (pH 7.3 at 25°C, adjusted with NaOH). The voltage was firstly clamped at −60 mV. For action potential stimulation, the frequency is by 2 × and 3 × rheobase and ramp current stimulation (0.1, 0.3, 0.5, and 1.0 nA/sec ramp current).

To measure the TTX resistant Na_V1.8 currents in DRG neurons, the voltage-clamp technique was used. For recordings on DRG neurons, the extracellular solution contained (mM): NaCl 60, KCl 3, Choline-Cl 80, CaCl₂ 0.1, MgCl₂ 0.1, HEPES 10, tetraethylammonium chloride 10, glucose 10 and CdCl₂ 0.1 (pH adjusted to 7.4, 300–310 mOsm/kg H₂O) TTX (1 uM) and TC-N 1572 (1.6uM) were applied to the solution to block TTX sensitive sodium current and Na_V1.9 currents. The pipette solution contained (mM) CsF 140, EGTA 5, MgCl 1, and HEPES 10, glucose 10 (pH 7.4, osmolarity 285–295 mOsm/kg H₂O). Only cells with an initial seal >1 GΩ were recorded. The Na_V1.8 currents were recorded responding to potential from –70 to +50 mV in 10 mV increments. The maximal current densities (pA/pF) were calculated and analyzed.

Electronic Von Frey hair algesiometer (IITC Life Science) was used to measure the hind paw withdrawal threshold. Before starting the test, mice were separately placed in elevated Plexiglas chambers on metal mesh flooring for 30 mins. A von Frey hair with bending force (0.6 g, 1 g, 1.4 g, 2 g, 4 g) was exerted perpendicular to the plantar surface of the hind paw until it just bent and the hind paw of mice of elevated. The force displayed on the electronic device were recorded. The threshold force required to elicit withdrawal of the paw was determined three times on each hind paw and averaged.

Gait analysis was performed on mice 4 weeks after ACLT by the CatWalk system (Noldus) according to our previous protocol (Zhen et al., 2013). Briefly, each mouse was placed walkway and allowed to allow the free movement from one side to the other side for at least three times. Mice were trained previously in the formal experiment. After the recording of mouse gait, several parameters were generated, and 5 of the most relevant parameters to OA pain were analyzed. (1) stands, (2) maximal contact at. (3) maximal (4) swing and (5) single stance.

Western blotting was performed on the lysates of DRG neuron culture and tibial subchondral bone marrow. The samples were separated by SDS-PAGE gel and transferred onto a nitrocellulose membrane (Bio-Rad Laboratories). After incubation with specific primary and secondary antibodies, signals were detected by an enhanced chemiluminescence kit (Amersham Biosciences). The primary antibodies used are as follow rabbit anti-Na_V1.8 (1:500, ASC-016, Alomone), rabbit anti-CREB (1:2000, #9179, Cell Signaling Technology), rabbit anti-pCREB (1:1000, 9198, Cell Signaling Technology), rabbit anti-PKA c- c-α (1:1000, 4782, Cell Signaling Technology) and rabbit anti-GAPDH (1:1000, 5174, Cell Signaling Technology). The experiments were repeated three times and a representative film was selected.

To measure the concentration of PGE2 in the subchondral bone marrow of mice tibiae, a PGE2 ELISA kit (514010, Cayman) was used according to the manufacturer’s manual. Briefly, we harvest the subchondral bone and then homogenized by ultrasound. The supernatant was aspired after high-speed centrifugation (13,200 g) for 10 mins. The concentration of PGE2 was normalized by total protein concentration using the BCA assay.

The ChIP assay was carried out using the epiquik ChIP Kit (Epigentek catalog number: P-2002–1). Briefly, the cultured lumbar DRG cells were crosslinked with 1% formaldehyde at for 10 min. After the collection of the cell, the sonication was performed until the DNA was broken into fragments with a mean length of 200 bps – 500 bps. The samples were subjected to immunoprecipitation with 2 mg of rabbit antibodies against pCreb1 (CST, 1:50) for 90 min at room temperature and 10% of the sample for immunoprecipitation was used as an input (a positive control). After purification, the DNA fragments were amplified using qRT-PCR with the primers for Na_V1.8 promoter listed in Supplementary Table 2.

Retrograde tracing was performed at 3-month-old male SD rats (Charles River Laboratories) (300–400 g, n = 6 per group) 2 months after ACLT. According to the previous study (Ferreira-Gomes et al., 2010), a 20 mm parapatellar incision was made over the medial side of the left knee. Ipsilateral femoral and tibial subchondral bone were subject to retrograde labeling. We injected 2 μl DiI (Molecular Probes; with 5 mg/ml in N, N dimethylformamide) into the femoral and tibial subchondral bone areas using a Hamilton syringe with a 27-gauge needle. Immediately after injection, bone wax was used to seal the drilling holes to prevent tracer leakage. Animals were euthanized 2 weeks after retrograde injection and the left lumbar DRGs (L3-5) were isolated for fluorescence detection. Twenty sections from each DRG were used for statistical analysis.

Data are presented as means ± standard deviations. Error bars represent standard deviations. We used unpaired or paired two-tailed Student’s t-tests for comparisons between two groups, and one-way ANOVA with Bonferroni post hoc test for multiple comparisons, in comparison between three or more groups, two-way ANOVA with Bonferroni post hoc test were used. All data demonstrated a normal distribution and similar variation between groups. For all experiments, p<0.05 was considered to be significant.