In this study, human IVD tissues were acquired from 19 patients (10 males and 9 females; 18–35 years old) with IVDD (all with grade II IVDD) who underwent lumbar spine surgery at Qilu Hospital of Shandong University. The degree of IVDD was assessed using the modified Pfirrmann grading system (Pfirrmann et al., 2001). Collection of lumbar disc tissue samples adhered to medical ethics regulations and was approved by the Medical Ethical Committee of Qilu Hospital of Shandong University with the ID of KYll-2021 (ZM)-058. A written informed consent document was requested and received from all study participants. All animal experimental procedures were performed according to the International Guiding Principles for Animal Research and were approved by the Laboratory Animal Centre of Shandong University.

NP cells were isolated and cultured as previously reported (Zhao et al., 2020; Yu et al., 2021). After the human IVD samples were washed 3 times with phosphate-buffered saline (PBS), the endplate cartilage and annulus fibrosus were removed. Then, the IVD tissues were carefully cut into fragments of approximately 1 mm³ in volume and digested with 0.25% trypsin and collagenase II (Sigma, United States). Then, the isolated cells were cultured in Dulbecco’s modified Eagle medium (DMEM)/F12 (HyClone, United States) supplemented with 10% fetal bovine serum (FBS) (Gibco, United States) and 1% penicillin and streptomycin (P1400, Solarbio, China) under standard incubation conditions (37°C, 5% CO₂, 95% air). The cell culture medium was replaced every 3 days, and the cells were passaged when they reached 80–90% density. HNPCs from passages 2–5 were used for further experiments. HNPCs were harvested and cultured in 6-well, 12-well, and 24-well plates and treated with or without TNF-α (HY-P7058, MCE, United States) (10 ng/ml) in the absence or presence of scutellarin (HY-N0751, MCE) (1 μM) for 15 min, 30 min, 1 h, 6 h, 24 h or 48 h before use in various experiments.

We purchased eight-week-old male Sprague Dawley rats from Beijing Vital River Laboratory Animal Technology Co., Ltd. The rats were housed under controlled identical specific pathogen-free (SPF) standard environmental conditions (23 ± 2°C, 12-h light/dark cycle) with free access to food and allowed to move freely. The IVDD model was established as previously reported (Zhao et al., 2020; Wu et al., 2021; Yu et al., 2021). Briefly, 15 rats were anesthetized by intraperitoneal injection with 2% (w/v) pentobarbital (40 mg/kg). As shown in Figure 3 of Supplementary Material, after being located by the X-ray and micro CT, the rat 6/7, 7/8, and 8/9 coccygeal discs were punctured using a 21 G needle. The depth of puncture was controlled at 5 mm. Later, the needles were rotated 360° and kept in position for 1 min. Then, 2.5 μl of PBS or scutellarin (100 ng/μl) was injected into the rat coccygeal discs using a microinjector (Hamilton, United States) with a 30 G needle (n = 5). After the aforementioned surgical procedure, the rats were monitored daily to observe their health condition.

To evaluate the structural differences and signal intensity changes in sagittal T2-weighted images of NPs, the rats in each group underwent X-ray and MRI scanning before execution 14 days after the initial puncture. The rats were placed in the prone position with their tails outstretched on the molybdenum target radiographic-image unit (GE, Boston, MA, United States). Radiographs were captured at a collimator-to-film distance of 66 cm, an exposure of 63 mAs, and a penetration power of 35 kV. T2-weighted images (repetition time: 3,000 ms; echo time: 80 ms; field of view: 200 mm²; slice thickness: 1.4 mm) were obtained by magnetic resonance imaging (MRI) using a 1.5 T system (GE) in the coronal plane. All radiographic images were saved in a Neusoft PACS/RIS DICOM 3.0 medical imaging system (Neusoft, Jinan, China). The IVD height and the adjacent upper and lower vertebral body heights were measured using ImageJ software, and the disc height index (DHI) was calculated from these values. The MRI grade of NPs was evaluated as previously reported (Xu et al., 2019; Zhao et al., 2020).

An RNAfast200 Kit (220011, Fastagen, China) was used to extract total RNA from the HNPCs according to the recommended procedure. Complementary cDNA was synthesized using a cDNA synthesis kit (FSQ-101, Toyobo, Japan). Real-time PCR was carried out with SYBR Green PCR Master Mix (OPK-201, Toyobo). For each sample, the relative mRNA levels were normalized to the group treated with PBS. The expression levels of target genes were normalized to GAPDH. The specific PCR products for each gene were confirmed by performing a melting curve analysis. The experiment was repeated three times for each group and each target gene. The sequence-specific primers used for real-time PCR are listed in Table 1. The results were calculated using the 2^−ΔΔCt method and are expressed as the fold change.

HNPCs were cultured using the methods described above to detect different indicators by Western blotting. Total proteins were extracted from HNPCs from each treated group with precooled radioimmunoprecipitation assay (RIPA) lysis buffer (P0013C, Beyotime Biotechnology, China) containing 1 μM phenylmethylsulfonyl fluoride (PMSF) with or without phosphatase inhibitors. The protein concentration was detected with a bicinchoninic acid (BCA) protein assay kit (PC0020, Solarbio). Equal amounts of protein from each sample were separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) on 8%, 10%, or 12% SDS-polyacrylamide gels and then transferred to a polyvinylidene difluoride (PVDF) membrane (Millipore, United States). After blocking with QuickBlockTM Blocking Buffer (P0252, Beyotime Biotechnology) for 20 min at room temperature, the membranes were incubated with rabbit primary antibodies against iNOS(1:1000, 18985-1-AP, Proteintech, United States), COX-2 (1:1000, ab15191, Abcam, United States), GAPDH (1:5000, 10494-1-AP, Proteintech), Aggrecan (1:1000, 13880-1-AP, Proteintech), Col-2 (1:1000, 28459-1-AP, Proteintech), ADAMTs-5 (1:1000, ab41037, Abcam), MMP-13 (1:1000, sc-515284, Santa Cruz Biotechnology), Bcl-2 (1:1000, ab196495, Abcam), Bax (1:1000, BM3964, Boster, China), Cleaved caspase-3 (1:1000, AF7022, Proteintech), OPA1(1:1000, PB0773, Boster), Drp1 (1:1000, 12957-1-AP, Proteintech), Mfn1 (1:1000, 13798-1-AP, Affinity, China), Mfn2 (1:1000 12186-1-AP, Proteintech), NLRP3 (1:1000, ab214185, Abcam), p-p65 (1:1000, 8242T, Cell Signaling Technology, United States), p65 (1:1000, 3033S, Cell Signaling Technology), p-p38 (1:1000, 4511T, Cell Signaling Technology), p38 (1:1000, 8690T, Cell Signaling Technology), p-JNK (4688S, Cell Signaling Technology), JNK (1:1000, 9252T, Cell Signaling Technology), p-Erk (1:1000, AF1015, Affinity), and Erk (1:1000, 4695T, Cell Signaling Technology) overnight at 4°C. Then, the membranes were incubated for 90 min at room temperature with a secondary antibody. The bands were visualized using an Amersham Imager 600, and the density of protein bands was quantified using ImageJ software.

The rats were sacrificed 2 weeks after the indicated surgery, and IVD tissues were collected. After fixation in 4% paraformaldehyde, the IVD tissues were decalcified, dehydrated, cleared using dimethylbenzene, and embedded in paraffin. Then, they were cut into 5-μm sections. Immunohistochemistry was performed with an Immunohistochemistry Kit (SP-9000, ZSGB-BIO, China) according to the instructions. After dewaxing and hydration, the sections were treated with Tris-ethylenediaminetetraacetic acid (EDTA) antigen retrieval buffer (C1038, Solarbio) for 10 min at 95°C and endogenous peroxidase blocker for 10 min. After blocking in goat serum for 30 min at room temperature, the cells were incubated with primary antibodies against iNOS (1:200, 18985-1-AP, Proteintech), COX-2 (1:200, ab15191, Abcam), ADAMTS-5 (1:200, ab41037, Abcam), MMP-13 (1:200, sc-515284, Santa Cruz Biotechnology, China), NLRP3 (1:200, Affinity, DF7438) at 4°C overnight. Then, the sections were incubated with goat antirabbit immunoglobulin (IgG) secondary antibodies for 1 h at room temperature and horseradish peroxidase (HRP)-labeled Streptomyces ovalbumin for 15 min. Detection was performed by using the DAB Substrate kit (ZLI-9018, ZSGB). Then, the slides were counterstained with 1% hematoxylin. Images were captured by a microscope (Leica DMI3000B). The positive areas were quantified using ImageJ.

Treated HNPCs on coverslips were fixed with 4% paraformaldehyde for 30 min. After incubation with 0.1% Triton X-100 for 5 min, the samples were blocked with 5% bovine serum albumin (BSA) at 37°C for 1 h. Next, the samples were incubated with primary antibodies against iNOS (1:1000, 18985-1-AP, Proteintech), COX-2 (1:1000, ab15191, Abcam), Col-2 (1:1000, 28459-1-AP, Proteintech), MMP-13 (1:1000, sc-515284, Santa Cruz Biotechnology), and p65 (1:1000, AF5008, Affinity) at 4°C overnight and then with fluorescently labeled goat anti-IgG (A23220, Amyjet Scientific or A23320, Amyjet Scientific) for 1 h at 37°C. The nuclei were stained with diamidino-2-phenylindole (DAPI).

The IVD sections were dewaxed and dehydrated, and the sections were treated with Tris-EDTA antigen retrieval buffer (C1038, Solarbio) for 10 min at 95°C. Next, the sections were incubated with 0.1% Triton X-100 for 5 min and 5% BSA at 37°C for 1 h. Then, the tissue sections were incubated with primary antibodies against cleaved caspase-3 (1:100, AF7022, Affinity) at 4°C overnight. The sections were incubated with fluorescently labeled goat anti-IgG (A23220, Amyjet Scientific or A23320, Amyjet Scientific) for 1 h at 37°C, and the nuclei were stained with DAPI.

The coverslips and sections were then observed by a fluorescence microscope (ZEISS Vert. A1), and the immunofluorescence signal intensities were quantified with ImageJ software.

The cell culture supernatants of each group were collected and preserved at −80°C until detection. Then, the IL-1β secretory expression levels were detected by an ELISA kit (E-EL-H0149c, Elabscience, China) according to the manufacturer’s instructions with a microplate reader (Infinite M NANO, TECAN).

MitoTracker Red CMXRos (C1049B, Beyotime Biotechnology) was used to detect mitochondrial activity and membrane potential. According to the manufacturer’s instructions, the cells were incubated with a culture medium containing 20 nM MitoTracker Red CMXRos for 30 min at 37°C in the dark and then observed and captured with a fluorescence microscope (ZEISS Vert. A1) after changing the fresh culture medium.

The mitochondrial membrane potential was detected using a JC-1 assay kit (C2003S, Beyotime Biotechnology). Treated HNPCs in 6-well plates were stained with a JC-1 staining solution at 37°C for 20 min in the dark according to the manufacturer’s instructions. Then, the cells were washed twice with 1× JC-1 staining buffer, and the images were observed and captured using a fluorescence microscope (ZEISS Vert. A1).

The ATP Bioluminescence Assay kit (S0026, Beyotime Biotechnology) was used to quantify intracellular ATP production. ATPs were extracted from cells treated with ATP for lysis. All procedures were performed according to the manufacturer’s instructions with a luminometer (Centro XS3 LB 960, Berthold Technologies).

Human NP cells on coverslips were stained with the TUNEL assay Kit (E-CK-A331, Elabscience) to examine the cellular apoptosis of each experimental group. All procedures were performed in accordance with the manufacturer’s instructions. The images were captured using a fluorescence microscope (ZEISS Vert. A1).

Intracellular ROS levels were detected with a ROS assay kit (S0033S, Beyotime Biotechnology). Briefly, after washing twice with sterile PBS, HNPCs were stained with 10 μM 2′,7′-dichlorofluorescein diacetate (DCFDA) at 37°C for 20 min in the dark and mixed every 5 min. Then, the HNPCs were washed three times with a basal culture medium before being observed and captured using a fluorescence microscope (ZEISS Vert. A1).

The rats were sacrificed 2 weeks after surgery. The IVD tissues were collected and fixed in 4% paraformaldehyde. After decalcification, dehydration, and embedding in paraffin, the IVDs were cut into 5-μm sections. Hematoxylin and eosin (H & E) staining was performed to evaluate the morphological changes of NPs with an H & E staining Kit (EE0012, Sparkjade), while Safranin O-Fast green staining was also performed to examine the cellularity and morphology of NPs and AF with a Modified Safranine O-Fast Green FCF Cartilage Stain Kit (G1371, Solarbio). The images were captured by a microscope (Leica DMI3000B).

For H&E staining, we evaluated histological grading of the IVD samples in accordance with the grading scale, which is based on the morphology of the AF and the cellularity of the NP (Keorochana et al., 2010). For safranin O-fast green staining, we conducted histological grading of the IVD samples according to the grading scale previously published (Han et al., 2008). The grading scale is based mainly on the cellularity and morphology of the AF and NP as well as the border between the two structures. The grades range from 5–15. The higher the score is, the more severe the degeneration.

HNPCs were collected by trypsinization, transferred into 2-ml centrifuge tubes, and fixed with fixation solution (G1102, Servicebio, China) for 2 h at 4°C in the dark. Then, the cells were proembedded with the agarose solution and postfixed in 1% osmium tetroxide in 0.1 M phosphate buffer (pH 7.4). Then, the cells were dehydrated in a graded series of ethanol solutions (30%, 50%, 70%, 80%, 95%, and 100%) for 15 min for each solution and infiltrated with a propylene oxide embedding medium overnight. Ultrathin sections (50 nm) were obtained using a Leica UC7 ultramicrotome, poststained with uranyl acetate and lead citrate, and visualized using a TEM (HT7800/HT7700).

The apoptosis of HNPCs from each experimental group was detected by flow cytometry. Cells were collected into flow cytometry tubes and stained with propidium iodide (PI) and Annexin V-FITC for 15 min at room temperature away from light with a fluorescein isothiocyanate (FITC) Annexin V Apoptosis Detection Kit (E-CK-A211, Elabscience) according to the manufacturer’s instructions. Cell apoptosis was detected with a CytoFLEX S flow cytometer (Beckman Coulter, United States), and the data obtained were analyzed with CtyExpert software.

The blinded method was conducted in all data acquisitions. Data are presented as the means ± SD of results derived from three independent experiments performed in triplicate. Analysis of data was performed with GraphPad Prism software (version 7.0; GraphPad Inc., La Jolla, CA, United States). Comparisons of different treatment groups were performed using analysis of variance (ANOVA) with Tukey’s post-hoc test. Statistical significance was considered when p < 0.05.

Previous studies have shown that elevated expression of inflammatory cytokines is a critical factor leading to IVDD (Liu K. et al., 2021). To investigate the role of scutellarin in the progression of the inflammatory response during IVDD, primary human nucleus pulposus cells (HNPCs) were isolated and then cultured with TNF-α in the presence or absence of scutellarin for 24 h to detect the mRNA level and 48 h to detect the protein level of inflammatory biomarkers. Following the indicated treatments, mRNA was extracted from the HNPCs, and real-time PCR was performed. Figures 1A,B show that the expression levels of COX-2 and iNOS were increased after TNF-α stimulation, while administration of scutellarin decreased the production of these cytokines induced by TNF-α. Next, total protein was extracted from each group of HNPCs, and Western blotting was performed to detect the expression levels of COX-2 and iNOS. As shown in Figures 1C–E, TNF-α-mediated enhancement of inflammatory cytokines was repressed by additional treatment with scutellarin. In the present study, immunofluorescence staining was performed for COX-2 and iNOS, indicating that the TNF-α-induced inflammatory reaction of HNPCs was markedly attenuated by scutellarin treatment (Figures 1F–I).

TNF-α enhances the expression of ADAMTSs and MMPs, leading to a reduction in ECM synthesis and the exaggeration of ECM degradation, among which MMP-13 and ADAMTS-5 play a critical role in Col-2 and Aggrecan levels (Fu et al., 2021). In this study, primary HNPCs were cultured and stimulated with TNF-α with or without scutellarin treatment. At the 24 h time point, mRNA was collected from each group, and real-time PCR was performed. The mRNA levels of the anabolic biomarkers Aggrecan and Col-2 were diminished, while the catabolic factors MMP-13 and ADAMTS-5 were elevated following stimulation with TNF-α. Nevertheless, additional treatment with scutellarin displayed a protective effect in the presence of TNF-α (Figures 2A–D). Moreover, total protein was isolated from HNPCs after the indicated treatment for 48 h, and the levels of Aggrecan, Col-2, MMP-13, and ADAMTS-5 were detected through Western blotting. As shown in Figures 2E–I, TNF-α stimulation decreased Aggrecan and Col-2 and promoted MMP-13 and ADAMTS-5 expression, while scutellarin antagonized the impairment of metabolism in HNPCs induced by TNF-α. These results were further verified by immunofluorescence staining, as shown in Figures 2J–M. TNF-α-induced reduction of Col-2 and elevation of MMP-13 signals were reversed by scutellarin administration.

Apoptosis of nucleus pulposus cells has been reported to be a critical factor associated with IVDD (Li BL. et al., 2021). To demonstrate the function of scutellarin in apoptosis of HNPCs, the cells were cultured with TNF-α in the presence or absence of scutellarin for 24 h to detect the mRNA levels of apoptosis-associated molecules. As shown in Figures 3A,B, Bcl-2 levels were reduced, and Bax levels were increased after stimulation with TNF-α. Scutellarin reversed the alteration of these biomarkers induced by TNF-α. Moreover, the cells were subjected to the indicated treatments for 48 h, and total protein was extracted from each group of HNPCs. Western blotting results indicated that TNF-α-mediated disorganization of these apoptosis biomarkers, including reduction of Bcl-2 and elevation of Bax as well as cleaved caspase-3, was attenuated by scutellarin treatment (Figures 3C–F). To further determine the role of scutellarin in apoptosis of HNPCs, TUNEL staining (Figures 3G,H) and flow cytometry (Figures 3I–L) were performed, which revealed that TNF-α promoted cell apoptosis, while scutellarin repressed this disorganization. Additionally, a live/dead assay was performed to verify the apoptosis rate of HNPCs, which indicated that TNF-α exaggerated the death of HNPCs, which was markedly diminished by scutellarin treatment (Figures 3M,N).

Exacerbation of oxidative stress and dysfunction of mitochondria are known to be harmful to inflammation and metabolism (Zhao et al., 2020); together with the finding of this study that scutellarin protects against inflammation and metabolism in NP cells, we were prompted to study whether the role of scutellarin is related to these detrimental factors. Herein, HNPCs were cultured, and transmission electron microscopy (TEM) was performed to observe the morphology of mitochondria. As shown in Figure 4A, the mitochondria were swollen and exhibited disrupted mitochondrial cristae following stimulation with TNF-α. However, treatment with scutellarin reversed this disordered alteration in mitochondrial morphology, which proved that scutellarin might be protective against mitochondrial damage in HNPCs. OPA1, Drp1, Mfn1, and Mfn2 are biomarkers of mitochondrial function (Zhao et al., 2020), all of which displayed a disorganized expression pattern after TNF-α stimulation, as assayed by Western blotting. Intriguingly, scutellarin treatment remarkably reversed the production of these molecules, suggesting the beneficial effect of scutellarin on mitochondrial function (Figures 4B–F) and ATP production. (Figure 4G). To further investigate the potential connection between scutellarin and mitochondrial ROS in HNPCs during IVDD, DCFDA, JC-1, and MitoTracker staining assays were performed to detect the production of ROS, mitochondrial membrane potential, and distribution pattern of mitochondria. As revealed in Figures 4H–M, our results further proved that scutellarin reversed TNF-α-induced mitochondrial dysfunction and oxidative stress in HNPCs.

To investigate the role of scutellarin in IVDD in vivo, a rat IVD needle puncture model was established, and local delivery of scutellarin was performed through intradiscal injection (Figure 5A) as we have previously reported (Zhao et al., 2020; Wu et al., 2021; Yu et al., 2021). To detect the IVD signal, MRI-T2WI analysis was performed, which revealed higher signal intensity in the rat IVD following scutellarin treatment than in the PBS group following needle puncture, suggesting the improvement of IVDD (Figures 5B,C). In addition, an X-ray was performed for each group of IVD samples, the IVD height was diminished after needle puncture, and the IVD height was largely retained following scutellarin treatment (Figures 5D,E). Thereafter, the IVD samples were collected, followed by histological analysis. Figures 5F–I indicate that the morphological degeneration score of the IVD based on HE and Safranin O staining in this IVDD model was alleviated by treatment with scutellarin. Moreover, Safranin O staining showed that scutellarin reduced proteoglycan loss during the process of IVDD. To further study the degeneration-associated biomarkers in each group, IHC for the inflammatory factors iNOS and COX-2 (Figures 6A–D) and the metabolism molecules ADAMTS-5 and MMP-13 (Figures 6E–H) was performed, which showed that scutellarin greatly reduced the expression of these biomarkers in the rat IVDD model. Moreover, an IHC analysis of NLRP3 expression levels was performed, and Figures 6I,J indicate that scutellarin alleviated the production of NLRP3 in the IVDD rat model. Furthermore, immunofluorescence was performed, and as shown in Figures 6K,L, scutellarin treatment reduced the expression of cleaved caspase-3 in IVDD.

To date, the NF-κB and MAPK signaling pathways are known to facilitate the ageing process under several conditions and have been extensively studied in the development of IVDD. In this study, HNPCs were cultured and stimulated with TNF-α with or without scutellarin treatment. Thereafter, phosphorylation of critical parameters in the NF-κB signaling pathway p65 (Chen et al., 2019) and components of the MAPK signaling pathway (Li L. et al., 2021), including JNK, p38, and ERK1/2, were assayed through Western blotting. As shown in Figures 7A–E, phosphorylated p65 was enhanced by TNF-α, which was largely abolished by additional treatment with scutellarin. Moreover, Western blotting for p-JNK, p-p38, and p-ERK1/2 indicated that TNF-α activated all the mentioned MAPK signaling pathways, while scutellarin retained the activity of the p38, JNK, and ERK1/2 signaling pathways in cultured HNPCs. To further test the involvement of the NF-κB signaling pathway in the effect of scutellarin, IF staining was performed, and nuclear translocation p65 was assayed. Figure 7F indicates that TNF-α induced the nuclear translocation of p65, while scutellarin suppressed this change in HNPCs mediated by TNF-α.

The NLRP3 inflammasome is deeply associated with IVDD through the production of IL-1β as its downstream molecule and the induction of exaggerated cell death, including apoptosis (Chao-Yang et al., 2021). To determine whether NLRP3 is related to the function of scutellarin in HNPCs, mRNA and protein were isolated from HNPCs. Real-time PCR was performed for NLRP3 and IL-1β (Figures 7G,H), and Western blotting was performed for NLRP3 (Figures 7I,J), indicating that scutellarin administration attenuated the activation of NLRP3 inflammasome induced by TNF-α. Furthermore, the supernatant was collected from each group, and an enzyme-limled immunoassay (ELISA) was performed for IL-1β, which showed that TNF-α-mediated secretion of IL-1β in HNPCs was inhibited by scutellarin (Figure 7K). In addition, the cells of each group were collected, and IF staining for NLRP3 and its downstream molecule IL-1β was performed. As revealed in Figures 7L–O TNF-α enhanced the expression of NLRP3 and IL-1β in HNPCs, and scutellarin treatment significantly suppressed this trend.