Dezocine alleviates cancer-induced bone pain and suppresses intraosseous tumor growth in renal cell carcinoma via the NAMPT/NAD+ axis

Abstract

Bone metastasis–associated pain is a major cause of morbidity in advanced renal cell carcinoma (RCC), and effective therapies that concurrently mitigate pain and intraosseous tumor burden remain limited. Here, we evaluated dezocine as a potential dual-action agent and explored the involvement of the NAMPT/NAD⁺ axis. In 786-O cells, dezocine dose-dependently suppressed proliferation (IC₅₀ = 73.6 μM), clonogenicity and increased apoptosis, accompanied by reduced NAMPT expression and a decreased NAD⁺/NADH ratio. Pharmacological rescue with the NAMPT activator SBI-797812 partially reversed these effects. In vivo, using an intra-tibial RCC cell inoculation model that recapitulates osteolysis and cancer-induced bone pain (CIBP), dezocine alleviated mechanical allodynia and thermal hyperalgesia, and reduced osteolytic bone destruction and intraosseous tumor progression; co-administration of SBI-797812 partially attenuated these effects (n = 6). Tumor tissues from dezocine-treated mice showed decreased Ki67 and NAMPT, increased cleaved caspase-3, and enhanced AMPK phosphorylation, consistent with metabolic stress–associated apoptosis. Collectively, these findings suggest that dezocine may provide dual analgesic and antitumor benefits in intraosseous RCC lesions, at least in part by suppressing the NAMPT/NAD⁺ axis, supporting further translational evaluation for bone metastasis–associated pain management.

Introduction

Renal cell carcinoma (RCC) is the most common kidney malignancy, with clear cell RCC (ccRCC) accounting for 70%–80% of cases. ¹ Due to its asymptomatic early course, most patients are diagnosed at an advanced stage. About 20%–35% of patients with RCC will develop distant metastases, with the skeletal system being a frequent site. ² Bone metastases affect approximately one-third of patients with metastatic RCC and are predominantly osteolytic, leading to symptomatic skeletal-related events (SREs) such as severe pain, pathologic fractures, and spinal cord compression, and are associated with shorter survival. ³ Bone metastasis reflects advanced disease and is a major driver of morbidity and impaired quality of life in RCC. ⁴ Cancer-induced bone pain (CIBP) is a complex pain state with both inflammatory and neuropathic components. Tumor-driven osteolysis and local acidosis, neurotrophic signaling, and spinal glial activation/central sensitization collectively contribute to ongoing and breakthrough pain. ⁵ Current management of RCC bone metastasis–related pain is multidisciplinary, combining systemic anticancer therapy with localized radiotherapy/surgery, bone-targeting agents, and analgesics. ⁶ However, pain control remains suboptimal for many patients, and opioid dose escalation is often limited by adverse effects and long-term safety concerns, highlighting the need for safer and mechanism-informed interventions.

Dezocine is an opioid analgesic with a distinct pharmacological profile and has been discussed as having a favorable safety profile in clinical use, warranting exploration for complex pain states with neuropathic features. ⁷ Preclinical evidence also suggests that dezocine has direct antitumor effects beyond analgesia, inhibiting proliferation, migration, and inducing apoptosis in breast, ovarian, and cervical cancer models.^8–10 The mechanisms underlying these effects appear to be closely linked to their interference with tumor cell metabolic reprogramming, particularly by modulating the NAMPT/NAD⁺ (Nicotinamide phosphoribosyltransferase/Nicotinamide adenine dinucleotide) axis. NAMPT, the rate-limiting enzyme in the NAD⁺ salvage pathway, sustains fundamental cellular processes including energy metabolism, DNA repair, and signal transduction.^11,12 In various solid tumors, including RCC, NAMPT is frequently overexpressed, which fuels tumor bioenergetics, promotes progression and metastasis, and contributes to therapy resistance.^13,14 Consequently, NAMPT is increasingly recognized as a druggable metabolic vulnerability in cancer.

Given that intraosseous tumor burden and osteolysis are key drivers of CIBP, we hypothesized that dezocine may confer dual benefits by suppressing RCC growth via NAMPT/NAD⁺ modulation and thereby attenuating downstream bone destruction–associated pain. We assessed the effects of dezocine on RCC cell proliferation, clonogenicity, and apoptosis, and elucidated its impact on the NAMPT/NAD⁺ axis in vitro. Subsequently, using an intra-tibial RCC inoculation model that recapitulates osteolysis and cancer-induced bone pain, we evaluated the effects of dezocine on tumor burden, osteolysis, and pain-related behaviors in vivo. To test this hypothesis and the specific involvement of the NAMPT/NAD⁺ axis, we employed the pharmacological activator SBI-797812 for rescue experiments in vitro and in vivo. ¹⁵ The present study focused on clarifying the critical role of the NAMPT/NAD⁺ axis in mediating the effects of dezocine, providing new experimental evidence for mechanistic research and therapeutic strategy development in RCC bone metastasis.

Materials and methods

Cell culture

The human renal clear cell carcinoma cell line 786-O and the mouse renal carcinoma cell line Renca were purchased from Zhejiang Ruyao Biotechnology Co., Ltd. All cell lines were authenticated by short tandem repeat (STR) profiling and confirmed to be free of mycoplasma contamination. The cells were cultured in RPMI-1640 medium (11875093, Gibco) supplemented with 10% fetal bovine serum (FS201-02, TransGen Biotech) and 1% penicillin-streptomycin (15140148, Gibco), and maintained at 37°C with 5% CO₂.

Cell viability assay

CCK-8 Assay: Cells were seeded into 96-well plates at approximately 5 × 10³ cells per well. After overnight attachment, the cells were treated with varying concentrations of dezocine (10, 20, 40, 80, 160 µM). At 24, 48, and 72 h post-treatment, 10 µL of CCK-8 reagent (CA1210, Solarbio) was added to each well, followed by incubation for 2 h in the incubator. Wells containing culture medium and CCK-8 reagent, but without cells, were used as blanks for background subtraction. The absorbance of each well was measured at 450 nm using a microplate reader, and the relative cell viability was calculated accordingly. Each experiment was independently repeated three times (n = 3, biological replicates). Data are presented as mean ± SEM.

The concentration range of dezocine (10–160 µM) was selected based on preliminary dose-finding experiments and previous studies reporting antitumor effects of dezocine in other cancer cell lines,^8,10 which demonstrated activity at 20–100 µM. The IC50 value at 48 h (73.6 µM) was used as a reference to select lower concentrations (20, 40 µM) for functional assays (colony formation, migration, invasion, and apoptosis), minimizing cytotoxicity while capturing biological effects.

Colony formation assay

Cells were seeded at a density of 500 cells per well into 6-well plates. After attachment, the cells were treated with dezocine at concentrations of 20, 40, and 80 µM. The culture was continued for 10–14 days, with periodic replacement of the drug-containing medium, until visible colonies formed in the control group. At the end of the experiment, the medium was removed, and the cells were fixed with methanol for 15 min, followed by staining with 0.1% crystal violet solution (C8470, Solarbio) for 30 min. After drying, the colonies in each well were counted and photographed for statistical analysis.

Wound healing and invasion assays

Wound-healing assay: 786-O cells were seeded in 6-well plates and cultured to 90%–100% confluence in complete medium. A sterile 200-µL pipette tip was used to create a linear scratch across the cell monolayer. Detached cells were gently washed away with PBS, and the remaining cells were incubated in serum-free RPMI-1640 medium containing the indicated concentrations of dezocine (10, 20, 40 µM). Representative images of the scratch area were captured at 0 h and 48 h post-scratch using an inverted microscope (100× magnification). The wound closure area (or migration ratio) was quantified by measuring the remaining cell-free area using ImageJ software (National Institutes of Health, Bethesda, MD, USA). The residual wound area (%) was calculated as: (area at 48 h/area at 0 h) × 100%. Each experiment was independently repeated three times (n = 3 biological replicates).

Invasion assay: Invasion was assessed using Matrigel-coated Transwell inserts (8-µm pore size, Corning). Matrigel (diluted 1:8 in serum-free medium, 50 µL per insert) was applied to the upper chamber and allowed to solidify. 786-O cells (5 × 10⁴ in 200 µL serum-free medium) were seeded into the upper chamber, and the lower chamber was filled with 600 µL medium containing 10% FBS as a chemoattractant. After 48 h of incubation, non-invading cells on the upper membrane surface were removed with a cotton swab. Invading cells on the lower surface were fixed with methanol and stained with 0.1% crystal violet. Five randomly selected fields per insert were photographed under a microscope (200× magnification) and counted. Data are presented as the average number of invading cells per field. All assays were performed in triplicate.

TUNEL assay

Apoptosis was assessed using the TUNEL assay. Coverslips were placed in 24-well plates, and 786-O cells were seeded onto the coverslips and allowed to adhere overnight. The cells were then treated with different concentrations of dezocine for 48 h. Fixation and subsequent steps were performed according to the instructions of the TUNEL detection kit (AFIHC030, AiFangBio). Fluorescence microscopy was employed for observation, with apoptotic nuclei exhibiting green fluorescence (FITC-labeled) under an excitation wavelength of 488 nm. At least five randomly selected fields were analyzed to count the total number of cells and TUNEL-positive cells. The apoptosis rate was calculated and analyzed statistically.

Western blot assay

Protein expression was analyzed by Western blot. Tibial lesion tissues (intraosseous tumor with surrounding bone) were dissected from euthanized mice, snap‑frozen in liquid nitrogen, and stored at –80°C. The tissues were homogenized in RIPA lysis buffer containing protease and phosphatase inhibitors. Total protein was extracted, and its concentration was determined using the BCA method. Equal amounts of protein were separated by 10% SDS‑PAGE and transferred onto PVDF membranes (0.45 µm pore size, Millipore) using a wet transfer system at 100 V for 90 min at 4°C. The membranes were blocked with 5% non‑fat milk in TBST for 1 h at room temperature and then incubated overnight at 4°C with the following primary antibodies: NAMPT (1:200, CQA2092, Cohesion Biosciences), cleaved Caspase‑3 (1:1000, ET1602‑47, Huabio), AMPK (1:1000, ET1608‑40, Huabio), p‑AMPK (1:500, ET1612‑72, Huabio), and GAPDH (1:1000, ab8245, Abcam) as an internal control. After primary antibody incubation, membranes were probed with an HRP‑conjugated secondary antibody (D110058, Sangon) for 1 h at room temperature. Protein bands were visualized using a chemiluminescence imaging system after incubation with enhanced chemiluminescence (ECL) reagent, and their intensities were quantified by densitometry using ImageJ software.

Measurement of NAD⁺/NADH ratio

The intracellular levels of NAD⁺ and NADH were measured using a commercial NAD⁺/NADH Assay Kit (S0175, Beyotime). For cellular assays, cells were treated with 40 μM dezocine, 10 μM SBI-797812, or their combination for 48 h before collection. Cells were lysed, and the lysate was divided into two aliquots. One aliquot was used to measure total NAD (NADt, the sum of NAD⁺ and NADH) by converting all NAD⁺ to NADH with alcohol dehydrogenase, followed by detection of the generated formazan at 450 nm. The other aliquot was heated at 60°C for 30 min to specifically decompose NAD⁺, leaving NADH intact for measurement. For tissue assays, snap-frozen tibial lesion tissues (intraosseous tumor with surrounding bone) were weighed, homogenized on ice in the provided extraction buffer, and processed identically. Absorbance at 450 nm was measured for all samples using a microplate reader. The concentration of NAD⁺ was calculated by subtracting NADH from NADt, and the NAD⁺/NADH ratio was derived from these values. Results are expressed as a percentage relative to the control group (set as 100%). All assays were performed with three replicate wells per group.

Establishment of animal models and grouping

Male BALB/c mice aged 6–7 weeks were purchased from Charles River Laboratories and housed under standard laboratory conditions (temperature: 22 ± 2°C; relative humidity: 60% ± 10%; 12-h light/dark cycle) with free access to water and standard rodent chow. To avoid confounding the assessment of cancer-induced bone pain, no peri- or post-procedural analgesics were administered outside the designated experimental treatments. All mice were monitored daily for signs of distress, such as body weight loss exceeding 20%, reduced mobility, hunched posture, or failure to eat or drink. In strict accordance with the approved IACUC protocol, humane endpoints were enforced; any animal meeting these criteria was euthanized immediately and excluded from subsequent analyses. The animal use protocol was reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) of Zhejiang Huitong Test & Evaluation Technology Group Co., Ltd. (Approval No. HT-2025-LWFB-0054), in full compliance with the Chinese national standard GB/T 35892-2018 ‘Guidelines for Ethical Review of Experimental Animal Welfare’. All experiments were conducted at the Laboratory Animal Center of the same institution, which holds the Laboratory Animal Use License (SYXK (Zhe) 2022-0041). All procedures were performed in accordance with the AVMA Guidelines for the Euthanasia of Animals (2020 edition) and the ARRIVE guidelines.

After 1 week of acclimatization, a CIBP model was established by intra-tibial implantation of Renca cells. Briefly, mice were anesthetized with isoflurane. Subsequently, Renca cells (1 × 10⁵ cells/mouse) were injected into the right tibial bone marrow cavity. The injection site was sealed with bone wax to prevent cell reflux, thereby establishing an intra-tibial RCC implantation model. ¹⁶ The Control group received an injection of an equivalent volume of physiological saline. After the procedure, mice were placed on a heating pad for warmth. Upon successful modeling, mice were randomly assigned into five groups (n = 6 per group) based on body weight: Control group, Model group, Dezocine group (30 mg/kg), Dezocine + SBI-797812 group (30 mg/kg + 10 mg/kg), and SBI-797812 alone group (10 mg/kg). Starting from day 3 post-modeling, drugs or corresponding vehicles were administered intraperitoneally once daily for 25 consecutive days (until day 28). Dezocine (30 mg/kg) and SBI-797812 (10 mg/kg) were each dissolved in a vehicle containing 10% DMSO + 90% saline. The injection volume was standardized at 10 mL/kg body weight. All groups (Control, Model, Dezocine, Dezocine + SBI, and SBI-alone) received an equal volume of the same vehicle (10% DMSO in saline). For the Control and Model groups, vehicle alone was administered; for the Dezocine group, vehicle containing dezocine; for the SBI-alone group, vehicle containing SBI-797812; and for the combination group, vehicle containing both drugs. All injections were performed at the same time each day (between 9:00 and 10:00 AM) to minimize circadian variations.

The dose of dezocine (30 mg/kg, intraperitoneal, once daily) was chosen based on previous in vivo studies demonstrating analgesic and antitumor efficacy of dezocine in rodent models,^8,10 as well as our preliminary dose-ranging study (10, 20, 30, 40 mg/kg), which indicated that 30 mg/kg produced optimal pain relief without observable sedation or motor impairment. The dose of SBI-797812 (10 mg/kg) was selected according to the original study describing this NAMPT activator ¹⁵ and our pilot experiments.

All surgical procedures (intra-tibial inoculation and μCT imaging) were performed under inhalation anesthesia with isoflurane (R510-22, RWD Life Science). Induction was carried out at 3%–5% isoflurane in 100% oxygen at a flow rate of 0.6–1 L/min, and anesthesia was maintained at 1%–2% isoflurane delivered via a nose cone. Respiratory rate and pedal withdrawal reflex were monitored throughout the procedure to ensure an adequate depth of anesthesia.

At the experimental endpoint (day 28 post-inoculation) or immediately upon meeting predefined humane endpoint criteria (e.g. body weight loss >20%, severe lethargy, inability to eat or drink), mice were euthanized by cervical dislocation while under deep anesthesia (5% isoflurane), followed by confirmation of respiratory and cardiac arrest. This method conforms to the AVMA Guidelines for the Euthanasia of Animals (2020 edition). All euthanasia procedures were performed by trained personnel in a dedicated facility.

Behavioral pain assessment

Mechanical Pain Threshold Test (von Frey Test): Mechanical paw withdrawal threshold (PWT) was measured using von Frey filaments on days 0 (baseline), 4, 7, 11, 14, 17, 21, 24, and 28 post‑inoculation (or until the predefined humane endpoint). Testing was performed 1 h after drug administration. Mice were placed on a metal mesh platform and acclimated for 30 min. A series of von Frey filaments with ascending forces was applied vertically to the mid-plantar surface of the ipsilateral (right) hind paw for 6–8 s. The minimal force required to elicit a paw withdrawal or licking response was recorded. The 50% mechanical PWT (paw withdrawal threshold) was calculated using the up-down method.

Thermal Pain Sensitivity Test (Hot Plate Test): Thermal withdrawal latency was measured using the hot plate test (55.0 ± 0.5°C) on the same days as mechanical testing (days 0, 4, 7, 11, 14, 17, 21, 24, and 28). A cutoff time of 60 s was set to prevent tissue injury.

All behavioral experiments were performed under randomized and blinded conditions. Drug administration, behavioral recording, and subsequent data analysis were independently conducted by different experimenters who were unaware of the group assignments. Throughout the behavioral tests, no obvious signs of motor impairment, sedation, or general distress were observed in dezocine-treated mice, suggesting that gross sedation or overt motor impairment was unlikely to be the primary driver of the observed behavioral changes.

Radiographic (X‑ray) analysis

On day 28 post-modeling, the right tibiae of mice were scanned using a digital X-ray system (MX-20, Faxitron Bioptics, Tucson, AZ, USA). Prior to imaging, animals were anesthetized with isoflurane and positioned on the imaging platform. Parameters were set as follows: tube voltage 50 kV, tube current 200 μA, exposure time 15 s. Representative radiographic images were acquired to assess osteolytic destruction.

Histopathology and immunohistochemistry

To evaluate tumor growth and related protein expression, collected tibial specimens were fixed in 4% paraformaldehyde for 72 h, followed by decalcification in 10% EDTA decalcification solution for 2–3 weeks. Subsequently, the tissues were dehydrated, cleared, and embedded in paraffin. Sections were cut at a thickness of 4 μm. Hematoxylin and eosin (H&E) staining was performed, and tumor cell infiltration within the bone marrow cavity, as well as bone destruction, was observed under an optical microscope. Immunohistochemical staining was conducted using the standard streptavidin-peroxidase (SP) method. After antigen retrieval, sections were incubated with primary antibodies against Ki67 (1:200, Proteintech) and NAMPT (1:100, CQA2092, Cohesion Biosciences) at 4°C overnight, followed by incubation with HRP-conjugated secondary antibodies at room temperature. Diaminobenzidine (DAB) was used for color development, and hematoxylin was applied for counterstaining. Stained sections were observed and photographed under a microscope.

Statistical analysis

Data are presented as mean ± standard error of the mean (SEM). Single time-point comparisons were analyzed by one-way ANOVA followed by Tukey’s post hoc test where appropriate. For behavioral data collected over time, statistical comparisons were performed using two-way repeated-measures ANOVA. A p value < 0.05 was considered statistically significant.

Results

Dezocine suppresses proliferation, migration, and invasion of 786-O Cells

The concentrations of dezocine used in each assay were determined based on the IC₅₀ values from the CCK-8 assay. For cell viability assays, dezocine was tested over a broad concentration range of 10–160 μM, whereas 20–80 μM was used for colony formation assays and 10–40 μM was used for migration and invasion assays. The CCK-8 assay showed that dezocine reduced cell viability in a concentration- and time-dependent manner. The inhibitory effect on cell viability became more pronounced with prolonged treatment. The calculated IC₅₀ values at 24 h, 48 h, and 72 h were 141.6 μM, 73.6 μM, and 57.4 μM, respectively (Figure 1(a)–(c)). Based on these findings, a treatment duration of 48 h was selected for subsequent experiments, employing various concentrations of dezocine. The colony formation assay further confirmed that dezocine significantly inhibited the clonogenic ability of 786-O cells. The number of colonies decreased markedly with increasing drug concentrations (20, 40, and 80 μM; Figure 1(d) and (e)). Wound-healing assays showed that dezocine increased the residual wound area in a concentration-dependent manner, indicating impaired wound closure and reduced migratory capacity of 786-O cells (Figure 1(f) and (g)). Similarly, in the Matrigel invasion assay, the number of invading cells in the dezocine-treated groups also decreased significantly in a concentration-dependent manner (Figure 1(h) and (i)). Together, these results demonstrated that dezocine inhibited the proliferation, migration, and invasion of 786-O cells in vitro in a concentration-dependent manner.

Figure 1.

Dezocine inhibits proliferation, clonogenicity, migration, and invasion of 786-O renal cancer cells. (a–c) Cell viability of 786-O cells measured by CCK-8 assay after treatment with dezocine at indicated concentrations (10, 20, 40, 80, 160 µM) for 24, 48, and 72 h. IC₅₀ values were calculated as described. (d) Representative images of colony formation by 786-O cells after treatment with dezocine (20, 40, 80 µM) for 10–14 days (crystal violet staining). (e) Quantitative analysis of the number of colonies from (d). (f) Representative images of 786-O cell migration in wound‑healing (scratch) assays after treatment with indicated concentrations (10, 20, 40 µM) of dezocine; scale bar: 200 µm. (g) Statistical analysis of the residual wound area (%) from (f). (h) Representative images of 786-O cell invasion in Transwell Matrigel invasion assays after dezocine treatment (10, 20, 40 µM); scale bar: 200 µm. (i) Statistical analysis of the relative number of invaded cells from (h). Data represent three independent experiments (n = 3). Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001 versus Control.

Dezocine induces cell apoptosis via suppression of the NAMPT/NAD⁺ axis

TUNEL staining revealed that, compared with the control group, the apoptosis rate in dezocine-treated cells was significantly increased and increased further with higher drug concentrations (Figure 2(a) and (b)). Following 48 h of treatment with 40 μM dezocine, Western blot analysis further showed that dezocine downregulated NAMPT protein expression, a key rate-limiting enzyme in NAD⁺ biosynthesis. Co-treatment with the NAMPT activator SBI-797812 partially reversed this downregulation. Meanwhile, cleaved caspase-3 expression was increased by dezocine, and this increase was attenuated upon co-treatment with SBI-797812 (Figure 2(c)–(e)). After dezocine treatment, the NAD⁺/NADH ratio in 786-O cells was markedly reduced. Co-treatment with SBI-797812 partially reversed this reduction, but did not fully restore the ratio to control levels. Administration of SBI-797812 alone notably increased the NAD⁺/NADH ratio and slightly elevated NAMPT expression, but did not increase cleaved caspase-3 levels under the conditions tested (Figure 2(c)–(f)). These results suggest that dezocine likely activates the caspase-3-dependent apoptosis pathway by suppressing NAMPT expression and reducing NAD⁺ availability, as reflected by the decreased NAD⁺/NADH ratio.

Figure 2.

Dezocine induces cell apoptosis by downregulating the NAMPT/NAD⁺ axis. (a) Apoptosis detected by TUNEL fluorescence staining after treatment with dezocine at 20, 40, or 80 µM for 48 h; green fluorescence indicates apoptotic nuclei; scale bar: 50 µm. (b) Statistical analysis of the percentage of TUNEL-positive cells from (a). (c) Representative Western blot bands showing the expression of NAMPT, cleaved Caspase-3, and the loading control (GAPDH) in 786-O cells treated with 40 µM dezocine, 10 µM SBI-797812, or their combination for 48 h. (d–e) Quantitative densitometric analysis of the protein bands in (c) using ImageJ. (f) Statistical analysis of the intracellular NAD⁺/NADH ratio in 786-O cells after different treatments (dezocine 40 µM, SBI-797812 10 µM, 48 h). Data represent three independent experiments (n = 3). Data are presented as mean ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001 versus Control; ##p < 0.01, ###p< 0.001 versus Dezocine.

Dezocine alleviates cancer-induced bone pain and suppresses bone destruction

Experimental results demonstrated that, compared to the control group, mice in the model group exhibited significant mechanical allodynia and thermal hyperalgesia. Monotherapy with dezocine significantly reversed these behavioral changes, as evidenced by an increased mechanical withdrawal threshold and prolonged thermal withdrawal latency. However, when combined with the NAMPT activator SBI-797812, the analgesic effect of dezocine was partially attenuated, but not completely abolished, as evidenced by a moderate decrease in mechanical withdrawal threshold and thermal withdrawal latency compared with the dezocine-alone group (Figure 3(a) and (b)). X-ray images revealed marked osteolytic destruction and tumor‑associated bone expansion in the tibiae of the model group mice. Dezocine monotherapy visibly reduced bone destruction. In contrast, co-administration with SBI-797812 attenuated the bone-protective effect of dezocine, resulting in a rebound in the degree of bone destruction (Figure 3(c)). H&E staining showed that representative images suggested reduced tumor cell infiltration within the bone tissue in the dezocine-treated group compared with the model group. This apparent reduction was less evident when dezocine was combined with SBI-797812, as indicated by increased tumor cell infiltration in the images (Figure 3(d)). Mice treated with SBI-797812 alone exhibited pain behaviors, bone destruction, and tumor infiltration similar to those of the model group.

Figure 3.

Dezocine alleviates pain and mitigates bone destruction in the CIBP model. Mice received intraperitoneal injections once daily from day 3 to day 28 post-inoculation: dezocine (30 mg/kg), SBI-797812 (10 mg/kg), or their combination, starting from day 3 post‑inoculation. (a) Statistical analysis of thermal withdrawal latency in mice from different groups measured by the hot plate test. (b) Statistical analysis of the mechanical withdrawal threshold in mice from different groups assessed by the von Frey filament test. (c) Representative radiographic (X-ray) images of the affected tibiae from mice in each group. Arrows indicate osteolytic lesions. (d) Representative H&E-stained images of tibial tissue sections from mice in each group; scale bars: 25 μm and 100 μm. Data are presented as mean ± SEM (n = 6). **p < 0.01, ***p < 0.001 versus Control; ##p < 0.01, ###p < 0.001 versus Model.

Dezocine exerts its effects in vivo via modulation of the NAMPT/NAD⁺ axis

Immunohistochemical staining of tibial lesion sections showed that representative images suggested increased Ki67 and NAMPT immunoreactivity in the model group compared with saline-injected controls. Dezocine monotherapy visibly reduced the immunostaining of both markers, whereas co-administration of SBI-797812 partially restored their expression (Figure 4(a)). Western blot analysis further demonstrated that dezocine treatment downregulated NAMPT protein expression and upregulated cleaved Caspase-3, an apoptosis marker. Furthermore, the expression level of phosphorylated AMPK (p-AMPK) was significantly enhanced without a change in total AMPK protein, indicating an increased level of AMPK phosphorylation. Co-treatment with SBI-797812 partially reversed the dezocine-induced changes in NAMPT, cleaved caspase-3, and p-AMPK/AMPK expression (Figure 4(b)–(e)) and partially restored the NAD+/NADH ratio (Figure 4(f)). In contrast, SBI-797812 alone increased NAMPT expression and the NAD⁺/NADH ratio but did not elevate cleaved Caspase-3 or p-AMPK levels (Figure 4(a)–(f)). Consistent with these findings, the NAD⁺/NADH ratio was elevated in tibial lesion tissues from the model group compared to saline-injected controls. Dezocine monotherapy significantly lowered this ratio, whereas co-administration of SBI-797812 partially restored it (Figure 4(f)). Collectively, these in vivo results suggest that the antitumor and analgesic effects of dezocine may be associated with its suppression of the NAMPT/NAD⁺ axis, accompanied by enhanced AMPK phosphorylation and promotion of tumor cell apoptosis.

Figure 4.

Dezocine exerts antitumor effects in vivo by regulating the NAMPT/NAD⁺ axis. Mice received intraperitoneal injections once daily from day 3 to day 28 post-inoculation: dezocine (30 mg/kg), SBI-797812 (10 mg/kg), or their combination, starting from day 3 post‑inoculation. (a) Representative images of Ki67 and NAMPT expression detected by IHC staining; scale bar: 25 μm. (b) Representative Western blot bands showing the expression of NAMPT, cleaved Caspase-3, p-AMPK, AMPK, and the loading control (GAPDH) in tibial lesion tissues. (c–e) Quantitative densitometric analysis of the protein bands in (b) was performed using ImageJ. (f) Statistical analysis of the NAD⁺/NADH ratio in tibial lesion tissues from mice of different groups. Data are presented as mean ± SEM (n = 6). **p < 0.01, ***p < 0.001 versus Control; ##p < 0.01, ###p < 0.001 versus Model, bracketed comparisons indicate Dezocine vs Dezocine+SBI.

Discussion

RCC bone metastasis causes severe pain, skeletal complications, and a poor prognosis. Current treatments often fail to balance analgesia with tumor control. ¹⁷ Dezocine, an analgesic reported to exhibit a favorable safety profile in certain clinical settings, has recently shown antitumor activity in various cancer models. ¹⁸ Thus, clarifying dezocine’s role in RCC bone metastasis improves our understanding of pain-tumor interplay and supports the development of dual-action therapies.

We hypothesized that the mechanism by which dezocine alleviates cancer-induced bone pain involves not only analgesic signaling but also the inhibition of intraosseous tumor growth and bone destruction via the NAMPT/NAD⁺ axis. In vitro, dezocine significantly inhibited the proliferation, clonogenicity, migration, and invasion of human clear cell RCC 786-O cells in a concentration- and time-dependent manner and induced apoptosis. Mechanistic exploration revealed that dezocine treatment downregulated the protein expression of the key metabolic enzyme NAMPT and reduced the level of its product, NAD⁺. Reverse rescue experiments using the NAMPT activator SBI-797812 further confirmed the critical role of the NAMPT/NAD⁺ axis in dezocine-mediated proliferation inhibition and apoptosis induction.

In the in vivo segment, we successfully established a CIBP model via intra-tibial inoculation of RCC cells. We found that dezocine monotherapy effectively alleviated mechanical allodynia and thermal hyperalgesia in the model animals, but also significantly suppressed tumor-associated osteolytic destruction and intraosseous tumor progression. The bone microenvironment plays a pivotal role in the colonization, growth, and progression of metastatic tumors, and targeting the tumor–bone interaction has emerged as a promising therapeutic strategy for bone metastasis.^19,20 Our findings extend this concept to renal cell carcinoma, suggesting that dezocine may confer dual benefits by simultaneously inhibiting intraosseous tumor growth and mitigating osteolytic destruction. Importantly, the co-administration of SBI-797812 partially counteracted the analgesic and bone-protective effects of dezocine, thereby providing in vivo pharmacological evidence supporting the central role of the NAMPT/NAD⁺ axis. Further molecular analyses indicated that dezocine downregulated the expression of the proliferation marker Ki67 and NAMPT, reduced NAD⁺ biosynthesis, and concurrently activated the apoptotic signaling (upregulation of cleaved Caspase-3) and the AMPK phosphorylation pathway in tumor tissues in vivo. SBI-797812 similarly reversed these effects. In summary, dezocine may inhibit the NAMPT/NAD⁺ axis, disrupt tumor energy metabolism, activate AMPK, and promote apoptosis, thereby achieving both pain relief and tumor suppression.

Conclusion

This study preliminarily demonstrates the potential of dezocine to exert dual analgesic and antitumor effects in RCC bone tumors by suppressing the NAMPT/NAD⁺ axis, providing experimental evidence for expanding its clinical applications. However, this work has certain limitations. The research focused on a single metabolic axis and did not explore the possible connection between its κ-opioid receptor-mediated analgesic pathway and its antitumor mechanism. The in vivo model was based on a single cell line, and the intra-tibial implantation approach does not fully recapitulate the complete metastatic cascade from a primary renal tumor to bone. Furthermore, analgesic behavioral assessments might be influenced by sedative effects. Future studies could validate these effects across additional model systems, delve deeper into the potential of combining dezocine with existing standard therapies, and employ techniques such as gene editing to definitively establish the critical role of the NAMPT/NAD⁺ axis. Additionally, a recent review highlights that NAMPT can negatively regulate PD-L1 expression in bladder cancer. ²¹ Whether dezocine-mediated NAMPT suppression influences PD-L1 levels and thereby modulates the tumor immune microenvironment in RCC bone metastases remains an intriguing question for future investigation. Another limitation is the cross-species design (human 786-O cells in vitro versus murine Renca cells in vivo), which warrants validation in additional RCC models to confirm the translational relevance of the findings. In conclusion, this work provides a new perspective and preclinical evidence for the comprehensive treatment of RCC bone tumors.