Introduction

Intervertebral discs (IVDs) are a major contributor to neck and back pain in adults, significantly impairing quality of life and imposing a considerable socioeconomic burden1,2,3. These fibrocartilaginous structures, situated between vertebral bodies, provide spinal flexibility while absorbing mechanical stress4. Each disc is composed of three main components: the outer annulus fibrosus (AF), the central gelatinous nucleus pulposus (NP), and two hyaline cartilaginous endplates that connect to adjacent vertebrae5,6,7. IVDs are the largest avascular tissues in the human body, making them highly dependent on diffusion for nutrient supply. Nutrients originate from capillaries in the vertebral body, traverse the subchondral plate, and terminate in loops near the cartilaginous endplates8. From there, oxygen and glucose must diffuse through the endplate matrix to sustain cells within the AF and NP9. This delicate supply chain is easily disrupted. Aging, mechanical overload, injury, smoking, and subchondral bone sclerosis promote endplate calcification and impair vascular flow, thereby restricting nutrient transport10. A reduction in oxygen and glucose availability creates a hypoxic, nutrient-deprived microenvironment that elicits strong cellular stress responses11,12. Under such conditions, NP cells employ adaptive mechanisms, such as autophagy, a lysosome-dependent process that recycles damaged proteins and organelles to maintain cellular homeostasis13,14. Yoshinori Ohsumi’s Nobel Prize-winning discovery of ATG genes firmly established autophagy as a central mechanism of survival under nutrient deprivation15. Extending this concept to disc biology, nutrient scarcity has been shown to activate autophagy in NP cells, where HMGB1 emerges as a key modulator of this process.

Under normal physiological states, HMGB1 functions as a non-histone DNA-binding protein in the nucleus. During stress, however, it translocates to the cytoplasm or is secreted extracellularly, where it regulates stress responses16,17. In the cytoplasm, HMGB1 interacts with Beclin-1, disrupting its association with Bcl-2 and thereby promoting autophagosome formation18,19,20. While autophagy initially protects NP cells, sustained or excessive activation can shift the balance toward apoptosis, marked by caspase activation, chromatin condensation, and membrane blebbing, ultimately contributing to disc cell loss in IVDD2,21. HMGB1 thus acts as a potential molecular switch between protective autophagy and apoptotic cell death. Reactive oxygen species (ROS), which accumulate under nutrient-deficient conditions, further enhance HMGB1 translocation and autophagic flux, while simultaneously sensitizing cells to apoptosis22,23,24. In line with this, Lee et al. (2010) reported that ROS buildup during serum deprivation is a potent trigger of apoptosis in various cell types25,26. Despite these insights, the precise mechanisms by which HMGB1 dictates NP cell fate under nutrient deprivation remain incompletely understood (Fig. 1).

Moreover, recent investigations have further explored the therapeutic modulation of autophagy to influence cell survival or death in degenerative tissues. CQ, an FDA-approved antimalarial drug, has gained attention for its role as an autophagy inhibitor. It interferes with lysosomal acidification, thereby preventing the degradation of LC3-II and autophagosome-lysosome fusion. In various models, CQ has been shown to enhance apoptosis when autophagy is insufficient to mitigate cellular stress, making it a valuable tool for dissecting autophagy-apoptosis dynamics. Additionally, CQ, a well-known autophagy inhibitor, impairs autophagosome-lysosome fusion by disrupting lysosomal pH, leading to the accumulation of damaged proteins and endoplasmic reticulum (ER) stress, which can culminate in cell death (Fig. 1)27,28,29. This process may be linked to apoptosis and could involve potential off-target effects related to autophagy inhibition under different cellular states. Moreover, the off-target effects of CQ on serum-deprived HNP cells and their autophagic responses remain to be fully characterized.

Given the critical role of nutrient availability in IVD health, this study aimed to investigate how serum deprivation affects HNP cells, with a particular focus on the regulatory role of HMGB1 in autophagy and apoptosis. Specifically, we investigated the time-dependent activation of autophagy by assessing key markers such as LC3-II and P62, as well as the cytoplasmic translocation of HMGB1 and its association with cell stress responses. We further examined the progression of apoptosis through the activation of cleaved caspase-3 and evaluated the effects of chloroquine (CQ)-mediated autophagy inhibition, including its potential off-target impacts on cellular processes. This study highlighted the HMGB1-driven interplay between autophagy and apoptosis under nutrient stress, and the potential off-target effect of CQ, offering new insights into potential therapeutic strategies for IVDD.

Fig. 1
figure 1

Nutrient deprivation inhibits mTOR signaling, thereby activating ATG proteins, which interact with Beclin-1 and ULK1 to initiate phagophore formation. Under stress conditions, nuclear proteins such as HMGB1 translocate to the cytoplasm, further promoting autophagy initiation. As the process progresses, LC3-II is recruited to facilitate autophagosome maturation, while the ubiquitin-binding protein p62 tethers autophagosomes to lysosomes. Upon fusion, autophagosomes are degraded within lysosomes, completing the autophagic process. Enhanced autophagy results in increased autophagic flux. However,15 μm CQ inhibits autophagosome-lysosome fusion, leading to the accumulation of damaged proteins and the induction of ER stress. Additionally, CQ can trigger Tumor necrosis factor (TNF)-related apoptosis-inducing ligand (TRAIL)-mediated apoptosis. Furthermore, nutrient deprivation increases ROS production due to mitochondrial dysfunction, ultimately leading to apoptosis.

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Materials and methods

Isolation and culture of NP cells

HNP samples were obtained from the human biobank of Daejeon St. Mary’s Hospital, with exemption from IRB screening (Task number DC25SASI0012) for experimentation. The tissue was washed three times with PBS and then minced into small fragments, approximately 2–3 mm in size, using a sterile scalpel or scissors. The dissected NP tissue was digested at 37 °C with 0.25% trypsin–EDTA for 25–30 min. Following trypsinization, the tissue was washed with PBS and neutralized with medium containing serum. It was then subjected to digestion with collagenase type II (0.1–0.2%) for 10 h to achieve complete dissociation. The resulting cell suspension was filtered through a 70 μm strainer to remove large debris and undigested material. The cells were subsequently washed with DMEM, and the isolated cells were cultured in Dulbecco’s Modified Eagle Medium/Nutrient Mixture F-12 (DMEM/F-12) supplemented with 10% fetal bovine serum (FBS) under 5% O₂ conditions. Cells at approximately 80% confluence were used for subsequent experiments. In the second passage, monolayer HNP cells were cultured under varying concentrations of nutrients and serum. Hank’s Balanced Salt Solution (HBSS) or DMEM-F12 with the addition of 0 to 20% FBS was used. Experiments were repeated to test the influence of variability in serum factors, and results were confirmed by using FBS from multiple lots. Cells were pre-cultured in DMEM-F12 with 1% FBS for 24 h, followed by 10% FBS for 72 h (~ 80% confluence), and cultured for up to 48 h. All experiments were carried out under 5% O2 to simulate the physiological environment of disc NP cells.

Cell morphology, dehydrogenase activity, DNA amount, and cell metabolic activity assays

Cell morphology, total dehydrogenase activity, and deoxyribonucleic acid (DNA) amount were assessed using an electron microscope, CCK-8, and NanoDropOne. The CCK-8 absorbance (450 nm) and DNA amount were measured using the NanoDropOne. Cell metabolic activity was calculated as dehydrogenase activity normalized to DNA amount.

Western blotting

The expression levels of proteins were determined by WB analysis of the total protein extracts from NP cells. Cell samples were lysed using Radioimmunoprecipitation Assay (RIPA) buffer and sonicated, and the protein concentrations were calculated using the Bradford assay. Proteins were loaded onto 10–15% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels and transferred to nitrocellulose membranes (Amersham Western Blotting Membranes, GE Healthcare UK Limited). After blocking for 1 h, the membranes were incubated with primary antibodies overnight at 4 °C. The following primary antibodies were used: LC3B (1:1000, NB100-2220, Novus Biologicals USA), P62 (1:1000, ab56416, Abcam), HMGB1 (1:1000, 6893, Cell Signaling Technology, Massachusetts, USA), Cleaved caspase-3 (1:1000, NB100-56113, Novus Biologicals USA), and beta-actin (1:5000, 5125, Cell Signaling Technology, Massachusetts, USA). Negative controls were performed with normal rabbit IgG-HRP (Cell Signaling Technology, Massachusetts, USA) and goat anti-mouse IgG-HRP (sc-2005, Santa Cruz Biotechnology, Inc., USA) under the same conditions. After washing tris tris-buffered saline with tween (TBST) three times, the membranes were incubated with the respective secondary antibodies. The bands were then detected with ECL Plus reagent (Millipore) using the ChemiDoc™ XRS + System (BIO-RAD, USA). Relative expression levels of proteins were determined by quantitative densitometric analysis using image analysis software (Image Lab, Bio-Rad, USA).

Immunofluorescence (IF)

To detect LC3, P62, and Cleaved Caspase-3 proteins in HNP cells, cells were prepared at a density of 50,000 cells per confocal plate. Cells cultured on the confocal plates were washed three times with phosphate-buffered saline (PBS) and fixed with 4% paraformaldehyde in PBS (pH 8.0) for 20 min, followed by three additional washes with PBS. The cells were then permeabilized with 0.1% Triton-X 100 in PBS for 15 min and washed three times with PBS. Antigenic sites were blocked with 5% bovine serum albumin in PBS for 30 min. The cells were incubated overnight at 4 °C with primary antibodies: LC3 (1:100), P62 (1:200), or cleaved Caspase-3 (1:200). After incubation, the cells were washed three times with PBS and then incubated with a fluorescein-labeled secondary antibody for one hour at room temperature. The cells were subsequently washed three times with PBS for five minutes each. Immunopositive cells were visualized using a fluorescent microscope (Olympus IX73, Tokyo, Japan).

Fluorescence-activated cell sorting(FACS)

Apoptotic cells were detected using the annexin V-FITC apoptosis Kit I (Invitrogen, Thermo Fisher Scientific, Bender Med Systems GmbH, Vienna, Austria). Briefly, both adherent cells and those present in the supernatant were collected and resuspended in 1× binding buffer at a concentration of 1 × 10⁶ cells per mL. A 100 µL sample containing 1 × 10⁵ cells was incubated with 5 µL of Annexin V-FITC and 5 µL of propidium iodide for 15 min at room temperature in the dark. After incubation, 400 µL of 1× binding buffer was added. Samples were then analyzed using a fluorescence-activated cell sorter (BD FACSCanto II Flow Cytometer, USA) within one hour. Apoptotic cells, including those staining positive for annexin V-FITC and negative for propidium iodide and double-positive cells, were counted and represented as a percentage of the total cell count.

Real-Time-quantitative polymerase chain reaction (RT-qPCR)

After the second passage, NP cells were treated with different concentrations of FBS (10%, 1%, and 0%), and total RNA was isolated using the RNeasy Mini Kit (Qiagen, Hilden, Germany). One microgram of RNA was reverse transcribed using the Revertra Ace RT Master Mix Kit (TOYOBO, Osaka, Japan). Relative mRNA expression of autophagy and apoptosis-related genes (LC3, P62, and Caspase-3) was evaluated by real-time (RT) polymerase chain reaction (PCR). The Applied Biosystems 7500 Fast real-time PCR System (Thermo Fisher Scientific, USA) and Thunderbird SYBR Green RT-PCR Mix (TOYOBO, Osaka, Japan) were used for this analysis. Primer sequences are listed in Table 1. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) mRNA expression was measured as an endogenous control. The mRNA expression of each gene in the experimental groups was normalized to the control group using the 2−ΔΔCt method. The formula ΔCt = Cttarget gene – CtGAPDH was used to calculate the difference in threshold cycles between the target gene and the reference gene. Finally, the fold change in mRNA expression of the target gene in the treated groups was calculated as 2−ΔΔCt.

Table 1 List of primers.
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Statistical analysis

Results were expressed as mean ± standard deviation. One-way ANOVA was used to investigate the effects of 0%, 1%, and 10% FBS on NP cells. The 10% FBS condition at 48 h was considered the normal condition for comparison with stress conditions, such as 1% and 0% FBS. To examine time-dependent autophagy activation, the 12-hour time point was considered the baseline (normal condition) for 0%, 1%, and 10% FBS across 12, 24, and 48 h. Statistical significance was assessed using GraphPad Prism 10.0 (GraphPad Software, Inc., USA). The levels of significance were indicated as follows: *P < 0.05,**P < 0.01,***P < 0.001, and ****P < 0.0001.

Results

Diminished serum and nutrient levels: impacts on NP cell proliferation and metabolic function

To understand how the NP cells respond to the alteration of nutrient supply associated with serum, we investigated cell morphology using HBSS or DMEM-F12 with varying serum concentrations, ranging from 0%,1%, and 10% FBS. We observed that cellular morphology underwent gradual changes as the serum and nutrient concentrations decreased. At 10% FBS, the cells retained their normal shape after 48 h. However, with decreasing serum and nutrient concentrations, the cell shape gradually shrank, and the cell number reduced. In HBSS, after 48 h, the cell morphology changed completely, significantly altering and approaching a state of reduced cell numbers (Fig. 2a).

To assess cellular metabolism, we quantified both cell dehydrogenase activity and DNA content in HBSS or DMEM-F12 containing (0–20)% FBS. Total dehydrogenase activity was determined through the CCK-8 assay, while DNA quantity was measured using NanoDropOne. Both metrics exhibited a decline under conditions of reduced serum and nutrient availability (Fig. 2b), indicating alterations not only in cell proliferation but also in metabolic activity. Interestingly, when evaluating cell metabolic activity by normalizing dehydrogenase activity to DNA content, the values remained relatively consistent in DMEM-F12 with 5–20% FBS. Notably, 10% FBS exhibited the highest metabolic activity, which is crucial for the survival of NP cells, with a declined metabolic activity observed only at the lowest FBS concentrations (0–1%) (Fig. 2c). Concurrently, visual observation of cell morphology revealed a flattened and enlarged appearance in DMEM-F12 with 0–1% FBS, while most cells became detached in HBSS (Fig. 2a).

Fig. 2
figure 2

(a) Cell responses to varying serum levels, observe the morphology using HBSS or DMEM-F12 with 0%, 1%, and 10% FBS. At 10% FBS, cells maintained their normal shape after 48 h. As serum levels decreased, cells shrank, and their numbers declined. In HBSS, after 48 h, cells exhibited significant morphological changes and a further reduction in cell numbers. (b) Measuring dehydrogenase activity (n = 5 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001) (2c) DNA content in HBSS or DMEM with 0–20% FBS showed that both metrics declined with reduced serum, indicating decreased cell proliferation and metabolic activity (n = 5 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001 and ****P < 0.0001). (2d) In DMEM with 5–20% FBS, metabolic activity remained almost stable, with 10% FBS showing the maximum metabolic activity, but it decreased at 0–1% FBS. Cells in 0–1% FBS appeared flattened and enlarged, while most cells detached in HBSS (n = 5 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001,and ****P < 0.0001).

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Reduced serum and nutrients increase NP cellular autophagy

We performed WB blot analysis to evaluate the time-dependent autophagic activity of LC3-II, P62, and HMGB1 under varying serum concentrations (10%, 1%, and 0% FBS) over 12, 24, and 48 h, aiming to identify the peak autophagic response to nutrient deprivation. Our results demonstrated that LC3-II levels were highest at 48 h under 0% and 1% FBS, indicating maximal autophagic response under these conditions. In contrast, under 10% FBS, LC3-II expression peaked at 12 h and gradually declined over time, reflecting reduced autophagic activity with sustained nutrient availability. P62 expression exhibited a decreasing trend across all serum conditions, further supporting autophagy activation through its degradation. HMGB1 expression peaked at 12 h under both 0% and 10% FBS, followed by a decline, while under 1% FBS, HMGB1 levels showed an unexpected increase over time. Despite this discrepancy, the overall expression patterns of LC3-II, P62, and HMGB1 supported the activation of autophagy in response to serum and nutrient deprivation. Based on these findings, we selected the 48-hour time point as the optimal duration to assess autophagic activity under nutrient-deprived conditions (Fig. 3).

Fig. 3
figure 3figure 3

WB analysis of autophagic activity in response to nutrient deprivation.(Fig: 3a to 3e) LC3, P62, and HMGB1 expression for 0% FBS at 12, 24, and 48 h (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001). (Fig: 3f to 3j) LC3, P62, and HMGB1 expression for 1% FBS at 12, 24, and 48 h (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001). (Fig: 3k to 3o) LC3, P62, and HMGB1 expression for 10% FBS at 12, 24, and 48 h (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001). The ratio of LCII/LC3I was calculated at 48 h (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001 and ****P < 0.0001). β-actin was used as a loading control. LC3-II accumulation was highest at 48 h in 0% and 1% FBS, indicating enhanced autophagy under nutrient deprivation. P62 showed a decreasing trend across all conditions, consistent with autophagic degradation. HMGB1 peaked at 12 h in 0% and 10% FBS, but increased over time under 1% FBS.

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We checked the HMGB1-mediated autophagy and late phase of autophagic flux, specifically autophagosome degradation, by analyzing LC3, HMGB1, and p62/SQSTM1, an autophagy-mediated degradation substrate. WB demonstrated both LC3-I (16 kDa) and LC3-II (14 kDa) in HNP cells. Our results showed a decrease in LC3-I and a corresponding increase in LC3-II expression, indicating the conversion of soluble LC3-I to lipid-bound LC3-II, which is a key marker of autophagic activity. The calculated LC3-II/LC3-I ratio was statistically significant, further confirming enhanced autophagy.

Our result also showed that the highest expression of LC3-II occurred under 0% FBS at 48 h, and this expression gradually decreased as the FBS concentration increased, reaching its lowest level at 10% FBS. Conversely, p62/SQSTM1 exhibited the opposite expression pattern, validating our hypothesis that serum deprivation enhances autophagic flux (Fig. 4).

HMGB1, a nuclear protein, translocates to the cytoplasm under conditions of increasing serum deprivation or serum stress. Our findings showed significantly increased HMGB1 protein expression with decreasing serum concentration, supporting our hypothesis. LC3 protein exists in two forms: LC3-I and the phosphatidyl-ethanolamine-conjugated LC3-II, with LC3-II serving as a marker of ongoing autophagy when recruited to the autophagosomal membrane. Under stress, HMGB1 translocates from the nucleus to the cytoplasm and is eventually released extracellularly, indicating the activation of beclin-1, which is essential for autophagosome formation. Our findings demonstrated that cytoplasmic HMGB1 levels continuously increased over 48 h in DMEM with 0–1% FBS. Thus, changes in LC3 and HMGB1 levels correlate with the severity of nutrient withdrawal (Fig. 4).

Fig. 4
figure 4

WB analysis of autophagic flux under nutrient-deprived conditions. HNP cells were cultured in DMEM supplemented with 0%, 1%, and 10% FBS for 48 h to assess late-phase autophagic flux by analyzing LC3, HMGB1, and p62/SQSTM1 expression. Western blot results revealed a decrease in LC3-I and an increase in LC3-II under serum-deprived conditions, indicating active autophagosome formation. LC3-II expression peaked at 0% FBS and progressively declined as serum concentration increased, while p62/SQSTM1 exhibited an inverse expression pattern, confirming enhanced autophagic degradation. Additionally, HMGB1, a nuclear protein that translocates to the cytoplasm during autophagy, showed increased expression with decreasing serum levels, reinforcing its role in Beclin-1 activation and autophagosome formation. These findings highlight the dynamic regulation of autophagy in response to nutrient withdrawal, with LC3-II and HMGB1 serving as key indicators of autophagic activity. The ratio of LCII/LC3I was calculated at 48 h (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001). β-actin was used as a loading control.(n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001).

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Furthermore, we conducted an LC3 turnover assay using CQ, a lysosomotropic compound known to induce the accumulation of autophagosomes containing LC3-II by inhibiting lysosomal acidification, thereby blocking autophagic flux. Our observation showed, CQ treatment led to a marked elevation of LC3-II expression across all serum conditions, including the nutrient-rich 10% FBS environment, whereas untreated cells showed a huge declined the LC3-II expression (Fig. 5).

Fig. 5
figure 5

CQ treatment led to a marked elevation of LC3-II expression across all serum conditions, including the nutrient-rich 10% FBS environment, whereas untreated cells showed a huge declined the LC3-II expression. Two-way ANOVA was used to investigate the effect of CQ treatment (n = 3, with level of significance *P < 0.05, **P < 0.01, ***P < 0.001).

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Additionally, to assess autophagic expression in NP cells, we conducted IF staining for LC3 and SQSTM1/P62 after 48 h of incubation with varying concentrations of FBS ranging from 0%,1%, and 10%. The expression of LC3-immunopositive cells exhibited an inverse relationship with serum concentration, reaching its peak at 0% FBS and diminishing progressively with increasing FBS concentration, being lowest at 10% FBS. Conversely, SQSTM1/P62 immunopositivity demonstrated a positive correlation with serum concentration. The highest number of immunopositive cells was observed at 10% FBS, gradually decreasing as the serum concentration decreased, with the lowest count noted at 0% FBS (Fig. 6a and b).

Fig. 6
figure 6

(6a) and (6b) Autophagic expression in NP cells via IF staining for LC3 and SQSTM1/P62 after 48 h with 0%, 1%, and 10% FBS. LC3 expression peaked at 0% FBS and decreased with higher serum levels. Conversely, SQSTM1/P62 expression was highest at 10% FBS and declined with lower serum concentrations.(6c) Elevated the nuclear-cleaved caspase-3-positive cells with decreasing FBS, indicating higher apoptosis at 0% FBS and lower at 10% FBS after 48 h.

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Besides, RT-qPCR was conducted to assess the expression of autophagy-related genes (P62 and LC3). The mRNA expression of LC3 showed a gradual upregulation with decreasing serum concentration, reaching its peak at 0% FBS and its lowest point at 10% FBS. The expression of P62 was detected during autophagosome degradation. Consequently, the downregulation of P62 expression, as anticipated, was observed to coincide with activated autophagy, being highest at 10% FBS and lowest at 0% FBS. This pattern suggests that serum deprivation activates autophagy (Fig. 7).

Fig. 7
figure 7

RT-qPCR assessed autophagy-related gene expression (P62 and LC3). LC3 mRNA expression increased with decreasing serum, peaking at 0% FBS and lowest at 10% FBS. P62 expression, indicative of autophagosome degradation, was highest at 10% FBS and lowest at 0% FBS. The downregulation of P62 coincided with activated autophagy (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001). Expression of apoptosis-related gene of caspase-3 increased with serum withdrawal, peaking at 0% FBS and lowest at 10% FBS, indicating that serum deprivation induces apoptosis (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001).

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Reduced serum concentration increases HNP cellular caspase-dependent apoptosis

We conducted WB to check the cleaved caspase-3 expression under varying serum concentrations (0%, 1%, and 10% FBS) to evaluate the expression of apoptosis levels. Our findings revealed a marked increase in cleaved caspase-3 expression at 48 h under 0% FBS, indicating significant apoptosis activation. This expression declined progressively with increasing serum concentration, reaching its lowest level under 10% FBS (Fig. 4a and f). To further investigate the effect of autophagy inhibition on apoptosis, we examined the impact of CQ treatment under serum-deprived conditions. Notably, CQ treatment led to a dramatic increase in cleaved caspase-3 levels, exhibiting a pronounced elevation compared to serum deprivation alone, providing compelling evidence of enhanced apoptosis upon autophagy inhibition (Fig. 8).

Fig. 8
figure 8

Nutrient deprivation increased cleaved caspase-3 expression in a manner proportional to the level of deprivation, with a peak observed at 48 h under 0% FBS conditions. While CQ treatment combined with serum deprivation elevated cleaved caspase-3 expression by 2- to 3-fold compared to serum deprivation alone, also peaking at 48 h under 0% FBS (n = 4 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001).

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Additionally, IF staining was performed to visualize cleaved caspase 3 positive cells following serum deprivation. The results demonstrated a significant increase in nuclear cleaved caspase 3 positive cells with decreasing FBS concentrations (Fig. 6c). The highest percentage was observed under 0% FBS, and the lowest under 10% FBS after 48 h, further supporting the conclusion that serum deprivation induces apoptosis in a dose-dependent manner.

Furthermore, we performed FACS analysis to assess the presence of apoptotic cells after treatment with 10%, 1%, and 0% FBS for 48 h. The lowest percentage of apoptotic cells was found in 10% FBS, while the highest was in 0% FBS, indicating that serum deprivation induces apoptosis. Although we used human tissues collected from defective discs, which may carry apoptosis-induced genes, the increase in apoptotic cells in 0% and 1% FBS compared to 10% FBS strongly indicates that serum deprivation induces apoptosis (Fig. 9).

Fig. 9
figure 9

FACS analysis revealed the highest percentage of apoptotic cells in 0% FBS and the lowest in 10% FBS, indicating that serum deprivation induces apoptosis (n = 3 with level of significance *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001).

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Moreover, RT-qPCR was performed to assess apoptotic-related gene expression, specifically caspase-3. The expression of caspase-3 increased gradually with serum withdrawal, reaching its highest level at 0% FBS and its lowest at 10% FBS after 48 h, indicating that serum deprivation induces apoptosis. (Fig:7)

Discussion

Our study investigated the effects of serum deprivation on autophagy in NP cells, providing substantial evidence that reduced nutrient availability triggers significant autophagic activity and apoptosis. The IVD, characterized by its avascular environment, relies on nutrient diffusion from the vascularized cartilage endplate6. When external stimuli, such as compression, exert pressure on the disc, nutrient supply from the endplate may be obstructed, leading to NP cell degeneration. This degeneration can occur through various pathways, including autophagy, apoptosis, and occasionally, necroptosis30. Clinically, such degeneration is observed in disc-related pathologies.

Our findings reveal that serum and nutrient availability play a crucial role in maintaining NP cell morphology, proliferation, and metabolic activity. At optimal conditions (10% FBS), cells retained their normal shape and metabolic function, whereas serum deprivation led to progressive cell shrinkage and detachment, especially in HBSS. The decline in dehydrogenase activity and DNA content under reduced serum conditions suggests impaired cellular metabolism and proliferation. Interestingly, metabolic activity remained stable in DMEM-F12 with moderate serum levels (5–20% FBS), emphasizing the importance of serum-derived factors for cell survival. The highest metabolic activity observed at 10% FBS underscores its essential role in maintaining NP cell homeostasis. In contrast, severe serum deprivation (0–1% FBS) led to noticeable morphological alterations and cell detachment, highlighting the vulnerability of NP cells to nutrient stress. These findings suggest that maintaining an optimal serum concentration is critical for preserving NP cell function and viability, consistent with the results reported by T. Yurube et al.20197.

Our findings showed that HMGB1 is a significant regulator of autophagy in HNP cells under nutrient stress. WB data revealed that HMGB1 expression peaked at 48 h in 0% and 1% FBS, coinciding with elevated LC3-II and reduced SQSTM1/P62 levels, hallmarks of active autophagic flux. This suggests that HMGB1 may facilitate sustained autophagy during prolonged serum deprivation. In contrast, under 10% FBS, HMGB1 showed an early peak at 12 h, followed by a decline, indicating a transient autophagic response in nutrient-rich conditions. These dynamics mirror the time-dependent role of HMGB1 in modulating autophagy, aligning with prior findings by T. Yurube et al., 2019. The data positioned HMGB1 as a responsive mediator that helps NP cells adapt to metabolic stress, especially during severe nutrient scarcity.

We investigated the expression of key regulatory proteins involved in autophagy and apoptosis, LC3, HMGB1, p62/SQSTM1, and cleaved caspase-3, using WB under varying serum conditions. Our findings demonstrated a significant association between serum deprivation and elevated autophagic flux, accompanied by increased apoptotic activity, consistent with the observations reported by Yurube et al. (2019) and Zhao et al. (2019). Notably, HMGB1 expression significantly increased at 48 h under 0% FBS, accompanied by elevated LC3-II and cleaved caspase-3 levels. This suggests that HMGB1 functions as a dual modulator, promoting autophagy while concurrently sensitizing cells to apoptosis during prolonged stress31.

The marked reduction in p62/SQSTM1 levels under serum deprivation further confirmed active autophagic degradation, consistent with observations by Klionsky et al. (2021). Interestingly, the highest p62/SQSTM1 expression was observed in 10% FBS conditions, showing an inverse relationship with LC3-II levels, supporting its role as a substrate degraded during autophagy32. Our findings also suggested HMGB1 translocation from the nucleus to the cytoplasm under nutrient-deprived conditions, which is indicative of its role in activating the beclin-1-dependent autophagy pathway, consistent with Yurube et al. (2019). This mechanism aligns with the work of Tang et al. (2010), who identified HMGB1 as a critical regulator of autophagy under stress. Collectively, our study highlights HMGB1 as a central switch modulating autophagy and apoptosis, facilitating cellular adaptation to metabolic stress21.

Furthermore, IF staining for LC3 and SQSTM1/P62 further validated these observations. LC3-positive cells peaked at 0% FBS and decreased with increasing serum concentrations, whereas SQSTM1/P62-positive cells followed the opposite trend, highest at 10% FBS and lowest at 0% FBS. This pattern confirms that serum deprivation robustly induces autophagy, leading to enhanced degradation of SQSTM1/P62. These results are in line with previous studies, such as those by M. Mauthe et al. (2018), who demonstrated similar staining patterns under autophagic induction33,34. RT-qPCR analysis of autophagy-related genes revealed a significant upregulation of LC3 and a downregulation of p62 mRNA levels with decreasing serum concentrations, peaking at 0% FBS and reaching the lowest levels at 10% FBS. This transcriptional response suggests that serum deprivation not only activates autophagic flux but also modulates gene expression to sustain autophagy. Similar findings have been reported by Shin, D. W. (2020), who observed that nutrient deprivation upregulates autophagy-related genes in various cell types33,35.

Moreover, our IF observation showed that the significant rise in cleaved caspase-3-positive cells with decreasing FBS concentration indicates the apoptosis induction in response to NP cells under serum deprivation. The highest levels of apoptosis were observed at 0% FBS, while the lowest were at 10% FBS after 48 h. These observations are consistent with those of H. E. Gruber et al. (2015), who reported that serum deprivation induces apoptosis in disc cells, contributing to cellular degeneration and tissue breakdown11,12. Further confirmation of apoptosis through FACS analysis demonstrated an increased percentage of apoptotic cells in conditions of reduced serum, with the highest percentage at 0% FBS. This trend mirrors findings from C. Q. Zhao et al. (2006), who showed that NP cells undergo significant apoptosis under nutrient-poor conditions, exacerbating IVDD36,37.

CQ was employed to inhibit lysosomal degradation by blocking LC3-II turnover, thereby enabling the assessment of autophagic flux. While CQ effectively suppressed autophagy, it also markedly enhanced apoptosis, as indicated by increased cleaved caspase-3 expression relative to untreated controls. This effect is likely attributable to CQ-mediated disruption of lysosomal acidification, resulting in the accumulation of LC3-II and impaired clearance of damaged proteins and organelles27,33. The consequent cellular overload contributes to pronounced ER stress, which, together with ROS-driven mitochondrial dysfunction, may activate apoptotic pathways as a protective mechanism against the survival of compromised cells. Importantly, this does not necessarily demonstrate a direct causal relationship between autophagy and apoptosis under nutrient deprivation, where autophagy normally proceeds. To address this, we considered possible off-target actions of CQ and clarified that the observed increase in apoptosis may reflect such effects. Nonetheless, these findings highlight the need for further studies to delineate the combined influence of nutrient deprivation and CQ-induced apoptosis on NP cell homeostasis.

Additionally, RT-qPCR analysis revealed a progressive increase in caspase-3 expression with decreasing serum concentrations, reaching its highest level under complete serum deprivation (0% FBS). This indicates a strong association between serum starvation and apoptosis in NP cells, aligning with previous observations in chondrocytes by Jia et al. (2018)28. Additionally, LC3 turnover assays demonstrated LC3-II accumulation across all serum conditions, suggesting that both basal and stress-induced autophagy are active in NP cells, even in nutrient-rich environments.

Our study emphasized the pivotal role of HMGB1 in orchestrating the balance between autophagy and apoptosis in NP cells under varying serum conditions. Serum deprivation led to increased LC3-II levels, decreased p62/SQSTM1, and elevation (cytoplasmic translocation stated Yurube, T., et al.,0.2019) of HMGB1, all indicative of enhanced autophagic flux as an adaptive response to nutrient stress. Concurrently, elevated cleaved caspase-3 levels and a rise in apoptotic cells revealed that apoptosis was also activated under these conditions. Notably, HMGB1 emerged as a central regulator, promoting autophagy through its interaction with beclin-1 and ATG5 during moderate stress, while its reduction under prolonged stress shifted the balance toward apoptosis, mirroring the model proposed by Chen, Ruochan, et al. These dynamics were further supported by our time-course data, which showed HMGB1 expression and gradual reduction patterns aligning with the shift in cellular responses23. Eventually, while CQ effectively inhibited autophagy by blocking lysosomal degradation, it also significantly promoted apoptosis.

Clinical relevance of this study

Clinically, IVDD develops in a microenvironment marked by chronic nutrient deficiency, hypoxia, and inflammation, which collectively accelerate NP cell death and extracellular matrix breakdown. Strategies that can restore homeostasis and maintain NP cell viability are therefore highly relevant for disease-modifying therapies38. In this context, pharmacological agents such as rapamycin and its analogs have been shown to enhance autophagy, suppress apoptosis, and delay disc degeneration in preclinical models39. In parallel, emerging gene-editing and RNA interference approaches targeting HMGB1 provide promising avenues for fine-tuning the autophagy–apoptosis balance40.

our findings highlight HMGB1 as a key molecular switch regulating NP cell fate under nutrient stress and underscore its potential as a therapeutic target in IVDD. Enhancing protective autophagy or preventing HMGB1-driven apoptotic shifts may help preserve disc cell survival and slow degeneration. Future studies combining HMGB1 modulation with pharmacological autophagy regulators will be essential to advance these strategies toward clinical application. A limitation of our study is the lack of direct mechanistic validation through HMGB1 knockdown, overexpression, or immunofluorescence staining, which should be explored in future research. To address this limitation, we aligned our statement with previously published studies and cited relevant references that support our conclusions.

Conclusion

 Serum deprivation triggers a time-dependent interplay between autophagy and apoptosis in human nucleus pulposus cells, orchestrated by HMGB1. Chloroquine-mediated autophagy inhibition further amplifies apoptosis, underscoring the critical balance between these pathways. Understanding this balance may guide targeted therapies for IVDD.