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. 2026 Jan 20;21:21. doi: 10.1186/s13062-025-00721-z

Synergistic anticancer effects of cabozantinib and cuproptosis induction driven by DHRS2-KAT2A-H3K27ac axis in clear cell renal cell carcinoma

Yilong Cao 1,#, Chao Xu 1,#, Bowei Zhang 1,#, Changbao Qu 1, Yuepeng Liu 1, Bei Shi 1, Xiaoling Li 1, Jiehan Li 1, Zeyuan Zhang 1, Shengtao Dai 1, Qingyun Sun 1, Yaxuan Wang 1,✉, Junfei Gu 1,✉
PMCID: PMC12895997  PMID: 41559745

Abstract

Background

Clear cell renal cell carcinoma (ccRCC) is the most prevalent subtype of kidney cancer, representing about 70–80% of all renal cell carcinomas. Cuproptosis, a recently identified mode of cell death, has increasingly been linked to tumor initiation, progression, and drug resistance. Our study provides the first evidence that the histone acetyltransferase KAT2A regulates cuproptosis in ccRCC by modulating histone H3 lysine 27 (H3K27) acetylation.

Methods

Our study commenced by establishing, through in vitro and in vivo experiments including CCK8 assays, angiogenesis assays, and Western Blot analysis, that cabozantinib and cuproptosis inducers exhibit synergistic anticancer effects. Subsequently, whole-transcriptome sequencing identified DHRS2 as a key regulatory gene. A series of functional experiments, including colony formation assays, cell proliferation assays, EdU staining, mitochondrial membrane potential staining, and ROS detection, were then conducted to systematically validate the tumor-suppressive role of DHRS2 and its pro-cuproptosis activity.Further investigation, by analyzing co-expressed genes, led us to identify KAT2A as a downstream effector molecule. We employed RNA immunoprecipitation (RIP) and rescue experiments to confirm the interaction between DHRS2 and KAT2A. To elucidate the molecular mechanism, CHIP-seq (chromatin immunoprecipitation sequencing) revealed that H3K27 acetylation promotes the transcription of GLS, a negative regulator of cuproptosis. This finding was subsequently validated through CHIP-qPCR and luciferase reporter gene assays.

Results

Our study reveals that in ccRCC, the combined treatment of cabozantinib and cuproptosis inducers leads to a significant upregulation of DHRS2. Functional experiments demonstrate that DHRS2 exerts a tumor-suppressive effect on ccRCC cells, effectively inhibiting their proliferation and growth. Mechanistically, DHRS2 post-transcriptionally represses KAT2A. Downregulation of KAT2A results in reduced H3K27 acetylation, which, as identified by CHIP-seq, leads to decreased transcriptional activation of its downstream target, GLS. As GLS acts as a negative regulator of cuproptosis, its diminished expression ultimately increases cuproptosis, thereby synergistically suppressing tumor growth.

Conclusions

Our findings demonstrate that DHRS2 suppresses the transcription of KAT2A, leading to reduced H3K27 acetylation levels. This, in turn, diminishes the transcriptional activation of its downstream target, GLS, ultimately inhibiting tumor growth. These results suggest that the combination of cabozantinib and cuproptosis inducers holds promise as a novel therapeutic strategy for ccRCC.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13062-025-00721-z.

Keywords: Clear cell renal cell carcinoma (ccRCC), Cabozantinib, Cuproptosis, DHRS2, KAT2A

Introduction

Clear cell renal cell carcinoma (ccRCC) is the most prevalent subtype of kidney cancer, representing about 70–80% of all renal cell carcinomas [1]. It is a highly malignant tumor with a strong tendency to metastasize. The main treatment approach currently is surgical resection of the tumor when possible [2]. However, roughly 30% of patients face recurrence or distant spread following surgery. The emergence of resistance to existing therapies presents a significant obstacle, underscoring the necessity for ongoing research to discover new therapeutic targets and develop more effective treatment strategies to enhance patient prognosis [3].

Renal cell carcinoma (RCC) is the most common type of kidney cancer, and while historically challenging to treat, the advent of targeted therapies, particularly Tyrosine Kinase Inhibitors (TKIs), has significantly improved patient outcomes [4]. TKIs target key signaling pathways that drive tumor growth and angiogenesis in RCC. Cabozantinib is a potent, orally administered small molecule inhibitor that targets multiple receptor tyrosine kinases. Its primary targets include VEGFRs, MET (mesenchymal-epithelial transition factor), RET (rearranged during transfection), AXL (a tyrosine kinase receptor), and FLT3 (Fms-like tyrosine kinase 3) [5, 6]. This multi-targeted profile allows Cabozantinib to exert broad anti-tumor effects by simultaneously disrupting tumor angiogenesis, inhibiting tumor cell proliferation, decreasing invasiveness, and overcoming resistance mechanisms. Cabozantinib is also utilized in the treatment of various other advanced malignancies, including neuroendocrine tumors, hepatocellular carcinoma, and medullary thyroid carcinoma [7–11]. Consequently, elucidating its precise mechanism of action holds significant importance for therapeutic advancement.

Cuproptosis, a recently identified form of programmed cell death, is triggered by the abnormal accumulation of copper ions, leading to mitochondrial dysfunction and subsequent cell demise [12]. Copper homeostasis plays a critical role in cancer and its recurrence. This process is intimately linked to tumor initiation, progression, and regulation. Cuproptosis significantly impacts the survival and death of tumor cells, making its study crucial for understanding cancer mechanisms [13]. Furthermore, it presents a promising target for developing novel anticancer therapies, potentially offering new avenues for the prevention and treatment of cancer recurrence.

Recent studies have revealed the potential applications of cuproptosis in various cancers [14–16]. Evidence suggests that combining ferroptosis inducers with cuproptosis inducers can enhance the sensitivity of liver cancer cells to intracellular death [17]. Additionally, research has indicated that targeting cuproptosis may provide a strategy to overcome radiotherapy resistance in cancer patients [18, 19]. Multiple studies also highlight the potential of cuproptosis as a novel therapeutic approach to overcome tumor drug resistance [20].

Histone acetylation, a key epigenetic modification, dynamically regulates gene expression by adding acetyl groups to lysine residues on histone tails, altering chromatin structure [21]. Catalyzed by histone acetyltransferases (HATs), this typically leads to chromatin relaxation and transcriptional activation [22]. KAT2A (also known as GCN5) is a prominent HAT family member that acetylates not only histones (e.g., H3K9, H3K14, H3K27) but also numerous non-histone proteins, participating in vital physiological processes such as cell proliferation, differentiation, immune response, and DNA damage repair [23–26]. Given its dysregulation and pro-tumorigenic role in various cancers, KAT2A has emerged as a prime target in cancer research and potential anti-cancer therapies.

DHRS2 (Dehydrogenase/Reductase (SDR Family) Member 2) is a ubiquitously expressed, pivotal enzyme belonging to the short-chain dehydrogenase/reductase (SDR) superfamily. It performs multiple crucial roles in cellular metabolism, primarily recognized for its reductase activity, catalyzing the conversion of various carbonyl compounds to their corresponding alcohols [27]. Emerging research additionally implicates DHRS2 in modulating cancer cell proliferation, migration, and chemoresistance. Specifically, DHRS2 has been shown to post-transcriptionally downregulate choline kinase alpha (CHKα), thereby suppressing ovarian cancer (OC) cell growth and metastasis both in vitro and in vivo [28]. Furthermore, DHRS2 regulates reactive oxygen species (ROS) and key tumor suppressors like p53 and p-Rb, demonstrating significant involvement in esophageal squamous cell carcinoma [29]. Its regulatory role has also been observed in other malignancies, including lung adenocarcinoma, prostate cancer, and renal cell carcinoma [30–32].

This study demonstrates that DHRS2 suppresses the growth and progression of ccRCC cells through its induction of cuproptosis. Mechanistically, DHRS2 post-transcriptionally downregulates the expression of KAT2A. Subsequently, KAT2A catalyzes histone H3 lysine 27 (H3K27) acetylation, thereby promoting the transcription of the cuproptosis-negative regulator gene, GLS. This cascade ultimately leads to the inhibition of ccRCC proliferation. Notably, our research establishes a novel link between histone acetylation and cuproptosis, offering a new therapeutic avenue for ccRCC.

Materials and methods

Cell culture

Cell lines 786-O, 769-P, A498, and OSRC2 were obtained from Procell LifeScience & Technology Co., Ltd. 786-O, 769-P, and OSRC2 cells were maintained in RPMI-1640 medium (Gibco, USA) with the same concentrations of 10% fetal bovine serum (FBS, Gibco, USA) and 1% streptomycin (Solarbio, China). A498 cells were cultured in minimum essential medium (MEM, Gibco, USA) with the same concentrations of FBS and streptomycin. All cell lines were incubated at 37 °C in a humidified atmosphere containing 5% CO2.

Western blot assays

Cells were initially harvested using ice-cold 1X Phosphate-Buffered Saline (PBS). Following collection, specimens underwent centrifugation at 5000 rpm. The resultant cell pellet was then re-suspended in pre-chilled RIPA lysis buffer and incubated on ice for a duration of thirty minutes to facilitate protein extraction. Subsequently, the lysates were subjected to centrifugation at 13,000 rpm for 10 min at 4 °C. Protein concentration was then quantified via the Bicinchoninic Acid (BCA) assay. Prepared samples, precisely quantified to 30 µg of protein, were denatured by heating at 95 °C for 10 min. These samples were then loaded onto an SDS-PAGE gel for electrophoretic separation. Upon completion of electrophoresis, the resolved proteins were transferred to a PVDF membrane. To mitigate non-specific antibody binding, the PVDF membrane was blocked using a 5% non-fat milk solution at room temperature for 2 h. Following the blocking step, the membrane was incubated with the primary antibody, diluted in blocking buffer, at 4 °C overnight. On the subsequent day, after washing away unbound primary antibody, the membrane was incubated with the correspondingly diluted secondary antibody for a period of 2 h at room temperature. The brands and dilutions of the antibodies used are detailed in the Supplementary Table 1.

mRNA isolation and qRT-PCR

Messenger RNA was meticulously isolated utilizing the RNA Extraction Kit (SM130, SEVEN, China). Following isolation, complementary DNA (cDNA) was synthesized from the extracted RNA via Superscript III reverse transcriptase (Invitrogen). The relative abundance of target gene mRNA was subsequently determined using fluorescent quantitative reverse transcription PCR (qRT–PCR) on the Bio-Rad CFX96 system, employing SYBR Green as the detection chemistry. To account for variations in sample loading and RNA extraction efficiency, all gene expression data were normalized against the expression levels of the housekeeping gene GAPDH. The established qRT–PCR amplification protocol involved an initial denaturation step at 50 °C for 2 min, followed by a high-temperature denaturation at 95 °C for 8 min and 30 s. This was succeeded by 40 cycles of thermal amplification, each comprising a 15-second denaturation at 95 °C and a 1-minute annealing/extension phase at 60 °C. The cycling protocol also included a melting curve analysis, with a preliminary denaturing step at 95 °C for 1 min, followed by a gradual cooling to 55 °C for 1 min, and concluding with a final plate reading at 55 °C for 10 s to assess product specificity. The sequences used in the experiment are listed in Supplementary Table 2.

Tube formation assay

Under stringent aseptic conditions, prepared or commercially sourced Matrigel® (Corning, USA) was dispensed into a 96-well plate at a volume of 50–80 µL per well. The 96-well plate was then incubated at 37 °C in a humidified incubator with a 5% CO₂ atmosphere for 30–60 min to ensure complete gel solidification. Subsequently, cells were harvested, enzymatically dissociated, and precisely counted. The cell suspension was adjusted to a final concentration of 1.0–1.5 × 10⁵ cells/mL. A volume of 100 µL of this cell suspension in appropriate culture medium was then seeded onto the surface of the solidified Matrigel®. Cells were further cultured under the same conditions (37 °C, 5% CO₂) for a period of 4–6 h. Following the incubation period, the formation of tubular structures was assessed and documented using an inverted microscope at magnifications ranging from 100x to 200x.

CCK-8 assay

Human renal cell carcinoma cells were seeded in 96-well plates at a density of 5 × 103 cells per well and allowed to adhere overnight. The cells were then treated with varying concentrations of the drug for 24 h. Following incubation, the medium was removed and 10 µL of CCK-8 reagent (Solarbio, China) was added to each well. The plates were incubated at 37 °C in a 5% CO2 incubator for 2 h in the dark. Absorbance was then measured at 450 nm using a microplate reader. Data are presented as mean ± standard deviation, and statistical analysis was performed using Student’s t-test to determine significant differences between treatment groups.

Clonogenic assay

Cells were seeded at a low density (500–1000 cells/well in a 6-well plate) and allowed to adhere for 24 h. The cells were incubated for 10–14 days, allowing colonies of 50 or more cells to form. Following incubation, the cells were fixed with methanol and stained with 0.1% crystal violet. Colonies were counted manually or using an automated colony counter, and the colony-forming efficiency was calculated as: Colony-Forming Efficiency (%) = (Number of colonies counted / Number of cells seeded) × 100.

Edu staining

Cells were seeded into 6-well plates and incubated with the EdU reagent for 2 h, following the manufacturer’s instructions (EdU Cell Proliferation Kit, Beyotime). Following incubation, cells were fixed and washed with PBS. Subsequently, they were stained with an EdU detection reagent and counterstained with DAPI. EdU-positive cells, indicating cell proliferation, were quantified using fluorescence microscopy.

Reactive oxygen species assay

Cells were seeded into 6-well plates and incubated with the EdU reagent for 2 h, following the manufacturer’s instructions (EdU Cell Proliferation Kit, Beyotime). After incubation, cells were fixed and washed with PBS. Subsequently, they were stained with an EdU detection reagent. The number of EdU-positive cells, indicative of cell proliferation, was then quantified by counting under a fluorescence microscope.

Assessment of mitochondrial membrane potential with JC-1

To assess the mitochondrial membrane potential (ΔΨm), cells were stained with JC-1 (5 µg/mL) for 20 min at 37 °C. After washing with PBS, cells were visualized using a fluorescence microscope. The presence of red fluorescence indicated healthy mitochondria with high ΔΨm, while green fluorescence indicated mitochondria with disrupted membrane potential.

Cell copper content assay

Following the guidelines of a commercial cellular copper (Cu) quantification assay kit (Solarbio, China), the procedure commences with cellular harvest. A predetermined cellular yield of 5 × 106 cells will be suspended in 400 µL of distilled water, followed by thorough homogenization. Subsequently, cells will undergo ultrasonic lysis under ice-cold conditions at a power output of 200 W, with a pulse sequence of 3 s on and 7 s off, for a total duration of 3 min. The resulting lysate will then be subjected to centrifugation at 10,000 g at 4 °C for 10 min to pellet cellular debris. The supernatant, containing solubilized intracellular copper, will be carefully collected. Finally, in accordance with the kit’s instructions, the absorbance will be measured at a wavelength of 580 nm using a spectrophotometer, enabling the calculation of intracellular copper concentrations.

RNA immunoprecipitation assay

In a strictly RNase-free environment, cell samples were lysed with a lysis buffer pre-supplemented with protease inhibitors and RNase inhibitors to maximize the integrity of RNA-protein complexes (RNPs). Subsequently, the lysate was incubated with magnetic beads conjugated to a specific antibody at 4 °C overnight. Following incubation, the magnetic beads, carrying the antibody-protein-RNA complexes, were effectively isolated from the lysate using magnetic separation. The isolated complexes were then washed 3–5 times to remove non-specifically bound molecules, thereby purifying the target complexes. Finally, the purified magnetic beads were resuspended in an RNA extraction buffer, to which dissociating agents such as proteinase K were added. Incubation at a specific temperature range of 50–70 °C was performed to release the bound RNA molecules. The extracted RNA was obtained using a standard RNA extraction kit and subsequently subjected to RT-qPCR analysis to detect the RNA bound to the target protein.

CHIP-qPCR assay

Cells are treated with formaldehyde for fixation, followed by sonication to fragment the chromatin into DNA fragments of a defined length. Subsequently, under gentle lysis and washing buffer conditions, the sample is incubated with antibodies conjugated to magnetic beads, with a negative control set up concurrently. After immunoprecipitation, the beads are separated from the solution via magnetic separation. This is followed by thorough washing with washing buffer (containing salt, detergent, and inhibitors) to remove non-specific binding. Once washing is complete, the material is incubated at 65 °C with reagents such as proteinase K to reverse cross-linking and release DNA. The resulting DNA precipitate is collected, and finally, real-time quantitative PCR (RT-qPCR) analysis is performed using specific primers to quantify the enrichment of the target protein in that region.

Luciferase reporter assay

First, the GLS promoter was subcloned into the pGL3.0-basic vector. After successfully transfecting the constructed plasmids into cells, continue culturing the cells until they reach an optimal density. Subsequently, collect the cells and add lysis buffer for cell lysis. Following this, proceed with the subsequent steps according to the detailed instructions of the luciferase assay kit. Finally, place the samples in a microplate reader to detect luminescence intensity and analyze the obtained data.

Animals

This study utilized twelve male BALB/c nude mice, aged 3–5 weeks, sourced from SPF (Beijing) Biotechnology Co., Ltd. All animal procedures were approved by the Animal Care and Use Committee of the Second Hospital of Hebei Medical University and adhered strictly to ethical guidelines for animal welfare. A subcutaneous renal cancer xenograft model was established by inoculating each mouse intraperitoneally with 1 × 107 cells. Beginning five days post-inoculation, mice received intraperitoneal drug administration for three weeks. Following the treatment period, mice were euthanized via cervical dislocation, and tumor volumes were quantified using the formula: Volume (mm³) = Width (mm) × Width (mm) × Length (mm) / 2. Animal allocation into control and experimental groups was achieved through randomization and blinded assessment.

Statistical analysis

Statistical analyses were performed using R version 4.1.3, and data visualization was generated with GraphPad Prism version 8.0.0. Correlations between variables were evaluated using Pearson and Spearman correlation coefficients, with statistical significance defined as a p-value < 0.05.

Results

Synergistic effect of cuproptosis inducers and cabozantinib

To investigate whether cuproptosis inducers can exhibit synergistic effects with anti-renal cell carcinoma drugs, we performed CCK8 assays on various renal cell carcinoma (RCC) cell lines. These experiments aimed to determine the impact of different concentrations of cuproptosis inducers, cabozantinib, and axitinib on the viability of RCC cells after varying treatment durations. Upon co-administration of cabozantinib or axitinib with the cuproptosis inducer into RCC cells, we observed that the combination of the cuproptosis inducer and cabozantinib more readily produced synergistic effects. Specifically, the synergy index reached as low as 0.19 (Fig. 1A, D), indicating that the combined application of the cuproptosis inducer and cabozantinib elicited a potent synergistic anti-tumor effect in RCC cell lines.

Fig. 1.

Fig. 1

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Synergistic effects of cabozantinib and cuproptosis inducers in ccRCC. (A) Cell viability of 786-O and 769-P cells treated with various concentrations of cuproptosis inducers. (B) Cell viability of 786-O and 769-P cells treated with various concentrations of cabozantinib. (C) Cell viability of 786-O and 769-P cells treated with various concentrations of axitinib. (D) Effect of combination treatment (cabozantinib and cuproptosis inducers) on ccRCC cell viability. (E) Inhibition of angiogenesis in ccRCC cells by the combination of cabozantinib and cuproptosis inducers. (F) Western Blot analysis of Ve-Cadherin expression under combination treatment. (G) Schematic diagram of the RNA sequencing workflow. (All data were shown as the mean ± SD, * p < 0.05; ** p < 0.01; *** p < 0.001, ns = non-significant)

For subsequent experiments, we ultimately selected 2.5 nM of the cuproptosis inducer and 5 µM of cabozantinib as a combination for further investigation. To further validate the synergistic effect, we performed angiogenesis assays (Fig. 1E). The results indicated that the cuproptosis inducer alone had a minor impact on angiogenesis. While cabozantinib could inhibit angiogenesis, its inhibitory effect was significantly enhanced when used in combination with the cuproptosis inducer. This observed potentiation demonstrates the synergistic anti-angiogenic effect of the combined treatment. To further validate the anti-angiogenic effect of the combined treatment, we performed Western blot analysis to examine the expression of Vascular Endothelial (VE)-Cadherin (Fig. 1F). The results demonstrated that Ve-Cadherin expression was lowest when the two drugs were administered in combination, compared to single-agent treatments. In summary, these experimental results demonstrated a significant synergistic effect between the cuproptosis inducer and cabozantinib.

Transcriptomics reveals DHRS2’s role in synergy

To investigate the mechanism of drug synergy, we performed RNA sequencing and analysis on four treatment groups (Fig. 1G). Differential gene expression analysis, based on pairwise comparisons, revealed the number of differentially expressed genes (Fig. 2A). By taking the intersection, we identified 9 key genes (Fig. 2B), which were then highlighted in a volcano plot (Fig. 2C). Furthermore, GO and KEGG enrichment analyses, performed on the top 30 differentially expressed genes, revealed biological functions related to extracellular matrix and cell adhesion, as well as molecular mechanisms linked to reactive oxygen species (ROS) metabolism and various signaling pathways (Fig. 2D, E). GSEA enrichment analysis suggested potential impact on functions like cell migration (Fig. 2F).

Fig. 2.

Fig. 2

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Transcriptome sequencing data analysis. (A) Lollipop chart and bar chart displaying differential gene comparisons across different groups. (B) Venn diagram illustrating the intersection of gene sets. (C) Volcano plot identifying significantly differentially expressed genes. (D) Gene Ontology (GO) enrichment analysis. (E) Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis. (F) Gene Set Enrichment Analysis (GSEA) revealing pathways related to cell migration and angiogenesis

In vivo synergism of cuproptosis inducers and cabozantinib

To validate the synergistic effect of two drugs in vivo, we utilized a xenograft tumor model. Following tumor establishment in nude mice, mice were administered drugs intraperitoneally once every five days for a total of 20 days. During the experimental period, mouse body weight and tumor volume were continuously monitored and recorded. The results demonstrated that both the cuproptosis inducer and cabozantinib exhibited tumor growth inhibitory effects individually. However, the combined administration of both drugs yielded the most pronounced inhibition (Figs. 3A, C).

Fig. 3.

Fig. 3

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In vivo studies demonstrate synergistic inhibition of renal cancer growth by cabozantinib and cuproptosis inducers. (A) In vivo xenograft model illustrating the suppression of tumor size by combination therapy. (B, C) Tumor weight and volume following monotherapy with cabozantinib or cuproptosis inducers, and combination therapy. (D) Immunohistochemical (IHC) staining depicting the expression of Ki67 and DLAT across different treatment groups. (E) Immunofluorescence (IF) staining showing the expression of Ki67 in different treatment groups. (All data were shown as the mean ± SD, * p < 0.05; ** p < 0.01; *** p < 0.001, ns = non-significant)

To further investigate the underlying mechanisms, immunohistochemistry was performed to compare the expression levels of Ki67 and DLAT in tumor tissues from different treatment groups. Immunohistochemical analysis revealed a decreasing trend in Ki67 expression and an increasing trend in DLAT expression in the combination therapy group. This suggests that the combined treatment not only suppressed tumor growth but also promoted the progression of cuproptosis (Fig. 3D). Subsequently, immunofluorescence experiments further confirmed that the combined administration group exhibited the lowest Ki67 expression, reinforcing the most potent tumor growth inhibition efficacy (Fig. 3E).

DHRS2-mediated inhibition of proliferation in ccRCC

To validate the nine intersecting genes identified in previous sequencing analysis, we performed qPCR assays on the target genes in two renal cancer cell lines. The results showed that the expression of DHRS2, NUDT3, and ERFE exhibited the most significant changes in the combination treatment group, with DHRS2 showing the most prominent changes (Fig. 4A). Similar results were observed at the protein level, where DHRS2 expression was highest in the combination treatment group (Fig. 4B).

Fig. 4.

Fig. 4

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DHRS2 suppresses ccRCC cell proliferation. (A) RT-qPCR analysis of downstream gene expression following treatment with cabozantinib and a cuproptosis inducer. (B) Western blot analysis of DHRS2 expression after drug treatment of cells. (C) WB analysis to confirm the efficiency of DHRS2 knockdown and overexpression. (D) RT-qPCR analysis to confirm the efficiency of DHRS2 knockdown and overexpression. (E) Colony formation assay to assess the effect of DHRS2 knockdown and overexpression on cell proliferation. (F) EdU staining assay to assess the effect of DHRS2 knockdown and overexpression on cell proliferation. (G) CCK8 assay to assess the proliferative capacity of cells after DHRS2 knockdown and overexpression. (H) Intracellular copper ion concentration after knockdown and overexpression of DHRS2. (All data were shown as the mean ± SD, * p < 0.05; ** p < 0.01; *** p < 0.001, ns = non-significant)

To further investigate the function of DHRS2, we conducted both DHRS2 knockdown and overexpression experiments. For the knockdown experiments, we screened three shRNA sequences and selected the most effective, sequence #3, for subsequent experiments. We also constructed a DHRS2 overexpression plasmid (Fig. 4C, D). Based on our focus on cell proliferation, we performed CCK-8, colony formation, and EdU staining assays. The CCK-8 assay results indicated that DHRS2 knockdown promoted cell proliferation, while DHRS2 overexpression inhibited cell proliferation. Colony formation assays further validated this conclusion, with a significant increase in the number of colonies formed in the DHRS2 knockdown group and a significant decrease in the DHRS2 overexpression group. EdU staining experiments also showed similar results, with an increase in EdU-positive cells in the DHRS2 knockdown group and a decrease in EdU-positive cells in the DHRS2 overexpression group (Fig. 4E, G). Collectively, these experimental results consistently demonstrate that DHRS2 inhibits cell proliferation in renal cancer cells.

The role of DHRS2 in regulating cuproptosis progression in ccRCC

To explore the relationship between DHRS2 and cuproptosis, we first employed Western Blotting (WB) to validate the changes in key cuproptosis-related genes, DLAT, FDX1, and GLS, in cells with DHRS2 knockdown and overexpression. The results revealed that DHRS2 knockdown led to a decrease in DLAT and FDX1 protein expression and an increase in GLS protein expression, suggesting that DHRS2 may promote the progression of cuproptosis. Subsequent qPCR experiments confirmed that these changes observed at the protein level were consistent at the transcriptional level, with similar trends in mRNA expression (Fig. 5A, B). Cuproptosis is generally associated with the accumulation of reactive oxygen species (ROS). Therefore, we investigated intracellular ROS levels in DHRS2 knockdown cells. Our findings indicated that after DHRS2 knockdown, intracellular ROS levels also significantly decreased (Fig. 5C).

Fig. 5.

Fig. 5

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DHRS2 promotes intracellular cuproptosis. (A, B) Expression levels of DLAT, FDX1, and GLS following DHRS2 knockdown and overexpression. (C) Changes in intracellular ROS levels after DHRS2 knockdown. (D) JC-1 staining to assess mitochondrial membrane potential following DHRS2 knockdown and overexpression. (E) Changes in downstream mRNA expression following DHRS2 knockdown and overexpression. (F, G) WB and RT-qPCR analyses to confirm the efficiency of KAT2A knockdown and overexpression. (All data were shown as the mean ± SD, * p < 0.05; ** p < 0.01; *** p < 0.001, ns = non-significant)

Furthermore, cuproptosis is known to induce a decline in mitochondrial membrane potential. To assess this effect, we measured the changes in JC-1 fluorescence upon DHRS2 knockdown and overexpression. Compared to the control group, DHRS2 knockdown resulted in an increase in red fluorescence (indicating normal mitochondrial membrane potential) and a decrease in green fluorescence (indicating decreased mitochondrial membrane potential). Conversely, DHRS2 overexpression led to a reduction in red fluorescence and an increase in green fluorescence (Fig. 5D). Finally, we examined the changes in intracellular copper ion concentration. The results showed that DHRS2 knockdown led to a decrease in intracellular copper ion levels, while DHRS2 overexpression significantly increased intracellular copper ion concentration (Fig. 4H). Collectively, these experimental results, including Western Blotting, qPCR, ROS detection, JC-1 mitochondrial membrane potential assessment, and intracellular copper ion concentration measurements, strongly support the conclusion that DHRS2 promotes the progression of cuproptosis.

KAT2A acts as a promoter of ccRCC cell proliferation

To investigate how DHRS2 promotes cuproptosis progression, we first identified the intersection of DHRS2-related genes and cuproptosis-related genes. we performed a Spearman correlation analysis between DHRS2 and all differentially expressed genes (DEGs) in KIRC (Kidney Clear Cell Carcinoma). The DEGs associated with DHRS2 were ranked based on their p-values, and the top 100 genes were selected (∣R∣>0.3). Subsequently, these 100 genes underwent a prognostic analysis in KIRC, requiring a p-value less than 0.05. Subsequently, combining this with genes that impact the prognosis of clear cell renal cell carcinoma, we finally screened out four genes: ARID5A, CSNK1E, KAT2A, and ZNF692. Through qPCR validation in renal cancer cell lines with DHRS2 knockdown and overexpression, we observed that upon DHRS2 knockdown, KAT2A and ZNF692 levels increased, while ARID5A levels decreased (Fig. 5D). We paid particular attention to KAT2A, a member of the histone acetyltransferase family. Based on this, we hypothesized that DHRS2 might influence cuproptosis progression by affecting histone acetylation through KAT2A. To explore this, we first performed knockdown and overexpression of KAT2A (Fig. 5F, G). To further investigate its function, we conducted CCK8 assays. The results showed that after KAT2A knockdown, the cell proliferation ability was enhanced, whereas after KAT2A overexpression, cell proliferation ability was attenuated (Fig. 6A). Subsequently, colony formation assays were performed to further validate these findings, revealing that KAT2A knockdown resulted in an increased number of colonies, while KAT2A overexpression led to a reduced number of colonies (Fig. 6B). Finally, we also performed EdU staining experiments, which demonstrated that EdU staining was increased after KAT2A knockdown and decreased after KAT2A overexpression (Fig. 6C). All these experiments collectively indicate that KAT2A promotes cell proliferation.

Fig. 6.

Fig. 6

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KAT2A promotes ccRCC cell proliferation. (A) CCK8 assay to assess the proliferative capacity of cells following KAT2A knockdown and overexpression. (B) Colony formation assay to evaluate the effect of KAT2A knockdown and overexpression on cell proliferation. (C) EdU staining assay to evaluate the effect of KAT2A knockdown and overexpression on cell proliferation. (D, E) Expression levels of DLAT, FDX1, and GLS following KAT2A knockdown and overexpression. (F) Changes in intracellular ROS levels after KAT2A knockdown. (G) Intracellular copper ion concentration following knockdown and overexpression of KAT2A. (H) JC-1 staining to assess changes in mitochondrial membrane potential following KAT2A knockdown and overexpression. (All data were shown as the mean ± SD, * p < 0.05; ** p < 0.01; *** p < 0.001, ns = non-significant)

The role of KAT2A in inhibiting cuproptosis progression in ccRCC

To investigate the relationship between KAT2A and cuproptosis, we first examined the changes in cuproptosis-related genes in cell lines with KAT2A knockdown and overexpression using Western blot. The results showed that, contrary to DHRS2, knockdown of KAT2A led to increased levels of DLAT and FDX1, and decreased levels of GLS. Conversely, overexpression of KAT2A resulted in decreased DLAT and FDX1, and increased GLS. Subsequent qPCR analysis confirmed that mRNA expression changes mirrored protein level alterations (Fig. 6D, E). We then measured intracellular ROS content and found that KAT2A knockdown led to increased ROS (Fig. 6F). Mitochondrial membrane potential was also assessed, revealing that KAT2A knockdown resulted in weaker red fluorescence and stronger green fluorescence, while KAT2A overexpression led to enhanced red fluorescence and weakened green fluorescence (Fig. 6H). Finally, we measured intracellular copper ion concentration, observing that KAT2A knockdown increased intracellular copper levels, while KAT2A overexpression reduced them (Fig. 6G). Collectively, these experiments indicate that KAT2A inhibits the progression of cuproptosis.

DHRS2 suppresses ccRCC growth by inhibiting KAT2A

DHRS2, a member of the SDR family, is known to regulate lipid and cholesterol metabolism. However, recent research has revealed that DHRS2 can also suppress RNA expression via post-transcriptional mechanisms [28]. Building on this, we hypothesized whether DHRS2 might inhibit KAT2A through similar post-transcriptional regulation. To investigate this, we performed RNA immunoprecipitation (RIP) assays, which demonstrated that DHRS2 indeed binds to KAT2A mRNA in various cell types (Fig. 7A). To assess DHRS2’s impact on KAT2A mRNA stability, we treated cells with actinomycin D to inhibit RNA synthesis for 24 h. RT-qPCR analysis revealed that DHRS2 overexpression shortened KAT2A mRNA half-life, whereas DHRS2 depletion increased it (Fig. 7B). To further confirm the inhibitory effect of DHRS2 on KAT2A, we conducted rescue experiments. Initially, we overexpressed DHRS2, observing an upregulation of DLAT and downregulation of GLS, indicating DHRS2’s influence on downstream signaling pathways (Fig. 7C). Subsequently, when KAT2A was co-overexpressed in DHRS2-overexpressing cells, these DHRS2-induced changes in DLAT and GLS were abrogated. Furthermore, we validated this mechanism using colony formation assays. The results showed that DHRS2 overexpression significantly reduced cell colony numbers, while co-overexpression of DHRS2 and KAT2A attenuated this inhibitory effect (Fig. 7D). CCK8 assays yielded similar findings. DHRS2 overexpression markedly inhibited cell proliferation, but co-overexpression of DHRS2 and KAT2A weakened this inhibitory capacity (Fig. 7E). To assess the impact on cuproptosis, we performed JC-1 staining. The experiment revealed a significant enhancement of green fluorescence upon DHRS2 overexpression, suggesting increased copper-induced cell death (Fig. 7F). Crucially, this cuproptosis-enhancing effect was substantially attenuated when KAT2A was co-overexpressed with DHRS2. In conclusion, our experiments provide compelling evidence that DHRS2 suppresses ccRCC through post-transcriptional regulation of KAT2A.

Fig. 7.

Fig. 7

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DHRS2 promotes cuproptosis through KAT2A. (A) RIP assay demonstrating mRNA binding between DHRS2 and KAT2A in 786-O and 769-P cells. (B) After treated with actinomycin D (5 µg/ml), KAT2A mRNA levels were measured by RT-qPCR, and the percentage of remaining mRNAs in the designated group were plotted. (C) Western blot to assess the rescue effect of KAT2A on DHRS2-regulated cuproptosis. (D) Colony formation assay to evaluate the rescue effect of KAT2A on DHRS2-regulated cell proliferation. (E) CCK8 assay to assess the rescue effect of KAT2A on DHRS2-regulated cell proliferation. (F) JC-1 staining to investigate the rescue effect of KAT2A on DHRS2-regulated cuproptosis. (G) KEGG enrichment analysis following ChIP-seq. (H) Annotation of ChIP-seq results. (I) Detection of H3K27ac binding peaks at the GLS promoter region. (J) Segmentation of the GLS promoter region. (K) ChIP-qPCR validation of H3K27ac binding at different segments of the GLS promoter. (L) Impact of KAT2A knockdown and overexpression on H3K27ac binding at the GLS promoter. (M) Dual luciferase activity assays to analyze the fluorescence intensity of KAT2A-overexpressing and KAT2A-depleted with the GLS promoter region. (All data were shown as the mean ± SD, * p < 0.05; ** p < 0.01; *** p < 0.001, ns = non-significant)

KAT2A promotes GLS transcription through histone acetylation to inhibit cuproptosis

To investigate whether KAT2A promotes cuproptosis inhibition by upregulating the transcription of the cuproptosis-suppressing gene GLS via histone acetylation, we performed histone acetylation level sequencing for KAT2A. Bioinformatic analysis, including KEGG and GO enrichment, revealed that genes associated with KAT2A binding were significantly involved in pathways such as cell cycle, renal cell carcinoma, and autophagy (Fig. 7G). Annotation of the enriched regions indicated that the majority of binding fragments were located within introns (45.84%) and promoter regions (22.47%) (Fig. 7H). Subsequent analysis of the sequencing-obtained binding peaks revealed their presence at GLS (Fig. 7I). Given that KAT2A negatively regulates cuproptosis and histone acetylation generally promotes downstream transcription, we hypothesized that KAT2A influences histone acetylation levels to facilitate the transcription of GLS, ultimately inhibiting cuproptosis progression. To pinpoint the exact binding site to GLS, we designed ten DNA fragments, and based on the identified binding peak locations, selected four fragments of varying sizes for primer design (Fig. 7J). ChIP-qPCR experiments demonstrated the most significant binding to the second fragment (Fig. 7K), thus indicating a primary binding location within the GLS region spanning from position 190,880,471 to 190,880,634. Further validation in cell lines with KAT2A knockdown and overexpression confirmed these findings: KAT2A knockdown led to a significant decrease in binding at the second GLS fragment, while KAT2A overexpression enhanced binding (Fig. 7L). Luciferase activity was elevated in the p-GL3-promoter group compared to the p-GL3-Basic group. KAT2A overexpression potentiated GLS promoter activity, whereas KAT2A knockdown attenuated it (Fig. 7M). Collectively, these results demonstrate that KAT2A promotes the transcription of GLS, a negative regulator of cuproptosis, likely by influencing histone H3K27 acetylation levels, thereby suppressing cuproptosis progression.

Discussion

As a newly discovered mode of cell death in recent years, the role of cuproptosis in suppressing tumorigenesis requires further exploration and discussion [12]. Nevertheless, recent studies have already unveiled its capacity to inhibit tumor growth and its significant involvement in chemotherapy resistance. In the study by Weikai Wang et al., it was found that ferroptosis inducers and cuproptosis regulators exhibit a synergistic effect, enhancing cuproptosis levels and suppressing the progression of hepatocellular carcinoma [17]. Research by Lei et al. found that downregulation of BACH1 suppresses the expression of copper-transporting metallothioneins (MT) 1E/X. This consequently reduces Cuproptosis, thereby promoting radioresistance in esophageal cancer cells and demonstrating the potential of targeting Cuproptosis to overcome tumor radioresistance [18]. Liao et al.‘s research discovered that p53 enhances cancer cell sensitivity to Cuproptosis through glycolytic metabolism. Furthermore, in vivo experiments confirmed the synergistic tumor-inhibitory effect of p53 agonists combined with Cuproptosis inducers [14]. Wen et al. also found that Cuproptosis influences the sensitivity of prostate cancer cells to docetaxel through autophagy [33]. These observations collectively suggest that Cuproptosis holds substantial significance in the context of both oncogenesis suppression and the resolution of chemoresistance, warranting further investigation.

The central aim of this study was to elucidate the synergistic effects and underlying mechanisms of cabozantinib and cuproptosis inducers in renal cell carcinoma (RCC) cells. Cuproptosis, a novel programmed cell death modality dependent on intracellular copper overload, presents a promising avenue for oncotherapy [34–36]. Our findings reveal that DHRS2, KAT2A, and GLS collaboratively form a complex regulatory network that profoundly influences the fate of RCC cells, providing a theoretical basis for targeted therapeutic interventions.

Our investigation demonstrates that DHRS2 promotes the cuproptosis process, a finding consistent with its known roles in other cellular metabolic pathways [37–39]. The pro-cuprodeath activity of DHRS2 is likely multifaceted. Firstly, this aligns with our prior observations of DHRS2 inhibiting RCC cell proliferation, suggesting that DHRS2 expression can suppress tumor activity [29, 30, 40, 41]. Secondly, beyond its metabolic contributions, DHRS2 may exert its effects by modulating downstream gene transcription [28, 42]. The mechanism by which DHRS2 regulates copper toxicity may be multifaceted. On one hand, DHRS2 could indirectly influence intracellular copper ion concentrations by modulating metabolic pathways such as lipid and cholesterol metabolism [33]. On the other hand, it may exert its effects through post-transcriptional regulation of key gene expression. Our investigation focused on DHRS2’s suppression of KAT2A expression via post-transcriptional control. Whether DHRS2 also impacts copper toxicity through metabolic pathways warrants further experimental investigation. Nevertheless, DHRS2 plays a significant role in suppressing renal cancer progression.

In contrast to DHRS2, KAT2A in our study exhibited an inhibitory effect on cuproptosis and promoted cell proliferation. As a histone acetyltransferase, KAT2A modulates gene expression by altering levels [43–45]. Our research highlights KAT2A’s critical role in regulating GLS. We found that KAT2A binds to specific regions of the GLS gene and upregulates its transcription through histone acetylation. Since GLS is a negative regulator of cuproptosis, KAT2A is capable of inhibiting cuproptosis. This is achieved by promoting the transcription of GLS, which in turn is facilitated by KAT2A-mediated enhancement of H3K27 histone acetylation levels. Nevertheless, our research is subject to limitations. While we focused on H3K27 acetylation, it is possible that acetylation at other loci, including H3K9 and H3K14, also plays a role. Additionally, KAT2A has the potential to regulate other histone modifications, such as succinylation and lactylation, which could be explored in subsequent research [43, 46].

In summary, our study made several key discoveries. Firstly, through in vivo and in vitro experiments, we demonstrated the synergistic anti-tumor effects of cabozantinib and cuproptosis inducers. Subsequently, RNA-sequencing identified the key gene DHRS2. Further investigations using RIP, CHIP-seq, and CHIP-qPCR experiments unequivocally showed that DHRS2 downregulates KAT2A via post-transcriptional regulation. This reduction in KAT2A led to decreased histone H3K27 acetylation levels, consequently downregulating the transcription of GLS, a negative regulator of cuproptosis. The enhanced cuproptosis levels in renal clear cell carcinoma cells ultimately achieved the inhibition of tumor growth (Fig. 8).

Fig. 8.

Fig. 8

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Proposed mechanism of DHRS2 in suppressing renal clear cell carcinoma growth. The combination of Cabozantinib and a cuproptosis inducer upregulates DHRS2 and enhances its transcriptional repression of KAT2A. KAT2A, in turn, promotes H3K27 acetylation, which facilitates the transcription of GLS. GLS negatively regulates the cuproptosis pathway, ultimately leading to the inhibition of cancer growth

Overall, our study indicates potential avenues for treating renal clear cell carcinoma, highlighting the synergistic effects of cabozantinib and cuproptosis inducers and suggesting their therapeutic promise.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary Material 1 (509.5KB, pdf)
Supplementary Material 2 (787.9KB, docx)
Supplementary Material 3 (15.2KB, docx)
Supplementary Material 4 (18.4KB, docx)

Acknowledgements

We would like to express our gratitude for the support from the Urology Laboratory at the Second Hospital of Hebei Medical University.

Abbreviations

ccRCC

Clear cell renal cell carcinoma

TKIs

Tyrosine Kinase Inhibitors

DHRS2

Dehydrogenase/Reductase (SDR Family) Member 2

HATs

Histone acetyltransferases

ROS

Reactive oxygen species

H3K27Ac

Histone H3 lysine 27 acetylation

Author contributions

J.F. Gu and Y.X Wang provided Data curation. Y.L. Cao, C. Xu and B.W. Zhang provided Writing-original draft and review& editing. C.B Qu and Y.P Liu provided Visualization and Methodology. B. Shi and X.L. Li and J.H. Li provided Formal analysis. Z.Y. Zhang, S.T. Dai, and Q.Y. Sun provided Project administration. All authors have reviewed and approved the final manuscript.

Funding

This work was supported by the National Natural Science Foundation of China (grant number 82072842), the Yanzhao Golden Talent Program of Hebei Province for 2024 (Returnee Platform, grant number B2024027) and Post-graduate’s Innovation Fund Project of Hebei Province (grant number CXZZBS2024122).

Data availability

The data underlying this article are available in the article and in its online supplementary material.

Declarations

Ethics approval and consent to participate

This study was approved by the research ethics committee of the second hospital of Hebei Medical University(2024-R244). Clinical trial number: not applicable.

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Yilong Cao, Chao Xu and Bowei Zhang equal contributions to this study and shared the first authorship.

Contributor Information

Yaxuan Wang, Email: wangyaxuan87@126.com.

Junfei Gu, Email: Junfei_Gu2020@hebmu.edu.cn.

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Supplementary Materials

Supplementary Material 1 (509.5KB, pdf)
Supplementary Material 2 (787.9KB, docx)
Supplementary Material 3 (15.2KB, docx)
Supplementary Material 4 (18.4KB, docx)

Data Availability Statement

The data underlying this article are available in the article and in its online supplementary material.

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