SREBP2 confers ferroptosis resistance by targeting GPX4 in colorectal cancer

Abstract

Background

The upregulated expression of sterol regulatory element-binding protein 2 (SREBP2) has been observed in multiple types of malignant cancers. Ferroptosis is a form of cell death that is iron-dependent and driven by the accumulation of lipid peroxides. It has recently garnered considerable attention in Colorectal cancer (CRC) research. This study aims to investigate the role of SREBP2 in CRC ferroptosis resistance and the underlying molecular mechanisms.

Methods

SREBP2 expression was assessed in CRC. Functional assays were conducted in HT29 and RKO cells following SREBP2 knockdown or treatment with Betulin, the SREBP2 inhibitor. RNA-seq was used to screen the potential downstream targets of SREBP2. Mechanistically, ferroptosis-related markers and rescue assays revealed the relation between SREBP2 and CRC ferroptosis resistance mediated by GPX4 expression regulation. Furthermore, ChIP and luciferase assays were used to confirm the upstream that regulates SREBP2 expression. Finally, subcutaneous tumorigenesis model was employed to evaluate the therapeutic potential of targeting SREBP2 in CRC.

Results

The upregulated SREBP2 expression in CRC drives the proliferation, migration, and invasion of CRC cells. Mechanistically, SREBP2 directly increases GPX4 transcription, thereby reducing the sensitivity of CRC to ferroptosis and facilitating CRC progression. Additionally, β-catenin was identified as an upstream regulator of SREBP2. Inhibition of SREBP2 sensitizes CRC cells to ferroptosis and suppresses tumor growth.

Conclusion

SREBP2 enhances ferroptosis resistance in CRC by upregulating GPX4, thereby contributing to tumor progression. Our findings highlight SREBP2 as a potential therapeutic target and provide a rationale for the development of SREBP2-targeted strategies in colorectal cancer.

Supplementary Information

The online version contains supplementary material available at 10.1007/s10495-025-02209-7.

Background

Colorectal cancer (CRC) is a malignant neoplasm of the digestive system that significantly endangers human health, with high incidence and mortality rates. Currently, colorectal cancer accounts for 9.6% of global cancer cases and 9.3% of cancer-related deaths, ranking as the second leading cause of cancer mortality worldwide and remaining a major global health concern [1]. Although surgery, radiotherapy, chemotherapy, or their combination has reduced the mortality of colorectal cancer, advanced CRC patient prognosis remains suboptimal [2]. Therefore, a deeper understanding of the molecular mechanisms underlying colorectal cancer development is crucial for advancing its diagnosis and treatment.

Recently, ferroptosis has garnered considerable attention in cancer including CRC. Ferroptosis is a form of cell death that is iron-dependent and driven by the accumulation of lipid peroxides [3–5]. When levels of lipid reactive oxygen species (ROS) exceed the antioxidant capacity of glutathione peroxidase 4 (GPX4), cellular redox homeostasis collapses, ultimately leading to oxidative damage and ferroptosis [6, 7]. However, cancer cells have ferroptosis defense mechanisms for survival under oxidative stress [8–10]. Disruption of these defense systems can be fatal to cancer cells, making ferroptosis a promising target for cancer therapy. GPX4 plays a critical role in catalyzing the reduction of lipid peroxides to lipid alcohols, thereby serving as a pivotal regulator in the suppression of ferroptosis. Study has shown that targeting GPX4 can induce ferroptosis in CRC cells [11]. However, the precise molecular mechanisms of ferroptosis in colorectal cancer and strategies for its targeted regulation remain unclear. Consequently, elucidating the molecular regulation of ferroptosis in colorectal cancer is essential for advancing ferroptosis-based therapeutic strategies.

Sterol regulatory element-binding protein 2 (SREBP2), a transcription factor, requires translocation of its precursor (Pre-SREBP2) from the endoplasmic reticulum to the Golgi apparatus, where proteolytic cleavage releases the N-terminal domain (n-SREBP2) [12]. This domain subsequently translocates to the nucleus and interacts with the promoters of rate-limiting enzymes in cholesterol biosynthesis, thereby regulating cellular lipid metabolism [12]. Recent studies have demonstrated that SREBP2 plays a pivotal role in the growth and malignant progression of various solid tumors, including breast cancer and liver cancer [13–16]. In breast cancer, SREBP2 upregulates the expression of key factors essential for mature osteoclast formation, thereby promoting tumor invasion and bone metastasis [14]. In hepatocellular carcinoma (HCC), targeting SREBP2 effectively suppresses tumor growth and metastasis [15]. Furthermore, SREBP2 has been identified to be involved in cancer cell ferroptosis. SREBP2 contributes to circulating melanoma cell resistance to inducers of ferroptosis by regulating the expression of TF (Transferrin) [17]. Inhibition of SREBP2 nuclear localization has been shown to induce ferroptosis in myeloma cells [18]. However, the specific regulatory role of SREBP2 in ferroptosis in CRC remains to be fully elucidated.

Betulin is a natural pentacyclic triterpene abundantly [19]. Mechanistically, betulin reinforces the interaction between SCAP and INSIG, thereby blocking the association of SCAP with COPII vesicles and preventing the release of SCAP–SREBP2 complex from the endoplasmic reticulum (ER); which suppresses SREBP2 activation [20]. Whether Betulin exerts anti-cancer effects in colorectal cancer through inhibiting SREBP2 needs to be further studied.

APC mutation, leading to the Wnt/β-catenin signaling pathway activation, has been recognized as a driver of colorectal cancer [21]. It is noteworthy that emerging evidence suggests β-catenin may directly regulate SREBP2 expression. A breast cancer study demonstrated that phosphorylated β-catenin upregulates SREBP2 expression, thereby promoting colony formation and recurrence of circulating tumor cells [22]. These findings suggest a potential direct regulatory role of β-catenin in SREBP2 expression. In this study, we identified β-catenin as a direct regulator of SREBP2 transcription which causes the upregulation of SREBP2 in CRC. Functionally, inhibition of SREBP2 by genetic knockdown or pharmacological inhibition suppressed CRC cell proliferation, migration and invasion. Mechanistically, SREBP2 promoted CRC ferroptosis resistance and tumor progression by transcriptionally upregulating GPX4. A subcutaneous tumorigenesis model further proved that inhibition of SREBP2 in vivo promoted CRC ferroptosis while suppressing CRC progression. These findings reveal a novel mechanism of ferroptosis regulation in CRC and highlight the role of SREBP2 in CRC progression.

Materials and methods

Patient samples

Tumor and adjacent non-tumor tissues were collected from 47 patients diagnosed with colorectal cancer who underwent radical surgical intervention at the Affiliated Cancer Hospital of Guangxi Medical University between January 2018 and August 2019. Inclusion criteria: (1) patients who underwent surgical treatment for colorectal cancer with postoperative pathological diagnosis of primary adenocarcinoma; (2) patients who had not received any systemic or local treatment before surgery. Exclusion criteria: patients diagnosed with other concomitant malignancies. The fresh tissue specimens were immediately frozen in liquid nitrogen and subsequently stored at -80 °C in a medical refrigerator (Haier, China). A subset of these fresh samples was fixed in 4% paraformaldehyde (Biosharp, China), followed by paraffin embedding, sectioning, and preparation for immunohistochemical (IHC) analysis. The research protocol received approval from the Science and Technology Ethics Committee of the Affiliated Cancer Hospital of Guangxi Medical University (Approval Number: KY20251099).

IHC analysis

Perform IHC analysis on paraffin-embedded tissue sections. The sections were subjected to dewaxing, hydration, and antigen retrieval utilizing sodium citrate buffer. Subsequently, endogenous peroxidase activity was inhibited with a 3% hydrogen peroxide aqueous solution. The sections were then blocked with 5% bovine serum albumin (BSA, neofroxx, Germany) prepared in PBS (servicebio, China), prior to incubation with the primary antibody, followed by three 10-minute washes with PBS and incubation with secondary antibody (Maxim, China). Cell nuclei were stained with hematoxylin. Staining results were visualized by incubation with diaminobenzidine (DAB, Maxim, China). Observe and take photos using a Axio scope5 microscope (ZESIS, Germany). The staining intensity was quantitatively assessed using ImageJ software, with scores assigned as follows: negative (0), weak positive (1), positive (2), and strong positive (3).

Cell culture

The human colon epithelial cell line NCM460 and six colorectal cancer cell lines—SW480, RKO, Lovo, HT29, HCT116 and Caco2—were obtained from Procell Life Science & Technology Co (Wuhan, China). 293T cell were purchased from Icell bioscience company (Shanghai, China). The Caco2 cell line was cultured in Dulbecco’s Modified Eagle Medium (DMEM, Gibco, USA) supplemented with 20% fetal bovine serum (FBS, Vazyme, China), 1% penicillin-streptomycin (P/S, Gibco, USA), and 100 ng/mL of recombinant human non-essential amino acids (NEAA, MCE, USA). Other cell lines including 293T were cultured in DMEM (10% FBS and 1% P/S). All cell cultures were incubated at 37 °C in an atmosphere containing 5% CO₂.

Virus packaging and cell transfection

To establish a stable SREBP2 knockout cell line, we designed the SREBP2 sgDNA sequence utilizing the CHOPCHOP web platform, which was subsequently synthesized by Sangon Biotech (Shanghai, China). The sgRNA was constructed in the LentiCRISPR v2 vector (Promega, USA). To produce lentivirus, the LentiCRISPR v2 sgRNA construct, VSVG, psPAX2, TAT3, and HEPACAM2 plasmids were transfected into 293T cells. Following this, HT29 cells were infected with the harvested supernatant containing viral particles, and the infected cells were subjected to selection using 2 µg/mL puromycin ((Solarbio, China). The GPX4 promoter region (-3000 to 0 bp) was amplified via polymerase chain reaction (PCR) from human genomic DNA and subsequently cloned into the pGL4.11[luc2P] vector (Promega, USA). PCR was conducted on the Biometra TAdvanced (analytik-jena, Germany). The plasmids for SREBP2, n-SREBP2 (1-484 amino acids) and the SREBP2 promoter (-2000 to 0 bp) were purchased from the MiaoLing Plasmid Platform (Wuhan, China). The mutant promoter plasmids were produced by site-directed mutagenesis system (Vazyme, China; Cat# C214). All small interfering RNAs (siRNAs) were synthesized by Sangon Biotech (Shanghai, China). The transfection of all plasmids and siRNAs into cell lines was conducted in accordance with the standard operating procedure for the Lipomaster 3000 Transfection Reagent (Vazyme, China). Comprehensive information regarding the sequences utilized in this study is listed in Supplementary Tables 1–3.

Cell proliferation assay

Cell proliferation was evaluated through cell counting and colony formation assays. In the cell counting assay, 1 × 10³ pretreated cells were inoculated into a 96-well plate and 10 µL of Cell Counting Kit-8 was added later (Vazyme, China). After incubation for 2 h in a 37 °C incubator, the each well absorbance was measured at a wavelength of 450 nm. Absorbance measurements were performed on a SpectraMax iD5e Multi-Mode Microplate Reader (Molecular Devices, USA). Cell counting experiments were conducted every 24 h. In the colony formation assay, 1 × 10³ pretreated cells were plated in a 6-well plate and cultured in medium (10% serum) for two weeks. Subsequently, the plates were fixed with 4% paraformaldehyde, stained with crystal violet, and the number of stained colonies was quantified using ImageJ software.

Cell migration and invasion assays

In the Transwell migration assay, a total of 3 × 10⁴ pre-treated cells in 200 µL of serum-free medium were seeded onto the upper chamber of the Transwell insert (8.0 μm pore size, Corning, USA). The lower chamber was supplemented with 700 µL of medium containing 10% FBS to serve as an attractant, thereby facilitating cell migration toward the lower chamber. For the Transwell invasion assay, the upper chamber was pre-coated with Matrigel (Corning, USA), while all other procedural steps remained consistent with those of the migration assay. Following a 48-hour incubation period, the cells were subjected to staining with crystal violet, and those that had not migrated into the lower chamber were subsequently removed. Images were then captured using a microscope (Axio observer3, ZESIS, Germany), and the cell count was counted utilizing ImageJ.

Western blotting

Cells were lysed on ice by RIPA buffer (CWBIO, China). After lysis, the resultant lysate underwent sonication. Subsequently, the lysate was centrifuged at 12,000 × g for 5 min to remove cellular debris. Loading buffer was added to the sample and mixed well; subsequently the sample was denatured at a temperature of 100 °C using a metal bath. The sample was separated using SDS-PAGE gels and transferred to a nitrocellulose (NC) membrane (PALL, USA). Protein electrophoresis was performed using the Mini-PROTEAN Tetra Cell (BIO RAD, USA), and transfering protein to membrane was carried out using the Trans-Blot Turbo Blotting System (BIO RAD, USA). The membrane was then blocked with 5% non-fat milk for one hour and incubated with the primary antibody, including anti-SREBP2 (1:2000; 28212-1-AP, Proteintech), anti-GPX4 (1:1000; sc-166570, santa), anti-ACSL4 (1:5000; 22401-1-AP, Proteintech), anti-SLC7A11 (1:2000; 26864-1-AP, Proteintech), anti-β-catenin (1:1000, #8480, CST), anti-active-β-catenin (1:1000, #19807, CST), anti-active-β-catenin (1:1000, #19807, CST), anti-β-ACTIN antibody (1:2000, sc-376421, Santa Cruz), anti-TCF4 antibody (1:2000, sc-166699, Santa Cruz), overnight at 4 °C. The following day, the membrane was treated with an IRDye^® secondary antibody (LI-COR, USA) at room temperature in the dark for one hour. Ultimately, the fluorescent signal was detected using the Odyssey CLX Western Blot imaging system. For tissue samples, grinding was performed using a grinder, and subsequent steps remained consistent with those for cell samples.

RNA extraction and real-time PCR

RNA was isolated from cellular and tissue samples utilizing TRIzol^® RNA Isolation Reagent (Invitrogen, USA). Subsequently, reverse transcription to complementary DNA (cDNA) was conducted using HiScript III qRT SuperMix (Vazyme, China), followed by quantitative polymerase chain reaction (qPCR) utilizing ChamQ Universal SYBR qPCR Master Mix (Vazyme, China). Reverse transcription was performed using the Biometra TAdvanced (analytik-jena, Germany), and qPCR was conducted on the qTOWER3G (analytik-jena, Germany). β-Actin served as an internal control. The relative expression levels of the target genes were determined using the 2^−ΔΔCt method. The primers utilized in this investigation are listed in Supplementary Table 4.

RNA-sequencing and analysis

RNA sequencing was conducted by Shanghai majorbio Biotechnology Co., Ltd. Briefly, RNA was extracted and reverse transcribed from si-SREBP2 and control transfected HT29 cells. The RNA-seq transcriptome library was prepared following Illumina^® Stranded mRNA Prep Ligation from Illumina. The reads were separately aligned to reference genome using HISAT2 software. The mapped reads of each sample were assembled by StringTie in a reference- based approach. RSEM was used to quantify gene abundances. Essentially, differential expression analysis was performed using the DESeq2. Additionally, the DAVID database (https://davidbioinformatics.nih.gov) was used for KEGG pathway enrichment analysis for the differentially expressed genes. Enrichment bubble plots and heatmaps were plotted by https://www.bioinformatics.com.cn, an online platform for data analysis and visualization.

MDA level measurement

The MDA Colorimetric Assay Kit (Lot: WW056V6 × 8360, Elabscience, China) was used to quantify MDA levels in cellular samples. Cells were seeded into 6-well cell culture plates (Vazyme, China) at a density of approximately 2 × 10⁶ cells per well. After cell collection, 300 µL of Extracting Solution was added, and the cells were disrupted by ultrasonication at 90 W for 4 s per cycle with 2 s intervals for a total duration of 10 min. Protein concentrations in the samples were determined using a BCA assay (Thermo Fisher Scientific, USA). Reagents were added and standards were prepared according to the manufacturer’s instructions. Samples were incubated in a 100 °C water bath for 40 min, followed by centrifugation at 1078 × g for 10 min. Then, 0.25 mL of the supernatant was transferred to a microplate, and the OD value of each well was measured at 532 nm using a microplate reader. Absorbance measurements were performed on a SpectraMax iD5e Multi-Mode Microplate Reader (Molecular Devices, USA).

Fe²⁺ level measurement

The concentration of Fe²⁺ in cells was determined using the Cell Ferrous Iron Colorimetric Assay Kit (Lot: WU03JZN62369, Elabscience, China) according to the manufacturer’s instructions. Cells were seeded into 6-well cell culture plates (Vazyme, China) at a density of approximately 2 × 10⁶ cells per well. After cell collection, 400 µL of Buffer Solution was added, and the samples were lysed on ice for 10 min, followed by centrifugation at 15,000 × g for 10 min. The supernatant was collected for further analysis. Reagents and standards were prepared according to the instructions. After mixing, the samples were incubated at 37 °C for 10 min, and the OD value of each well was measured at 593 nm using a microplate reader. Absorbance measurements were performed on a SpectraMax iD5e Multi-Mode Microplate Reader (Molecular Devices, USA).

GSH/GSSG level measurement

The GSH and GSSG Assay Kit (Lot: A060A241227, Beyotime, China) was used to measure the levels of GSH and GSSG in cells. Cells were seeded into 6-well cell culture plates (Vazyme, China). After digestion with 0.25% trypsin, the cells were collected by centrifugation at 1000 rpm for 5 min. The samples were subjected to two rapid freeze–thaw cycles using liquid nitrogen and a 37 °C water bath, followed by centrifugation at 10,000 × g for 10 min at 4 °C. The supernatant was collected for total glutathione measurement. Reagents and standards were prepared according to the manufacturer’s instructions. A portion of the prepared total glutathione sample was used for GSH removal, as instructed, and then processed for GSSG measurement following the same procedure. After a 25 min reaction between the samples and the reagents, the OD value of each well was measured at 412 nm using a microplate reader. Absorbance measurements were performed on a SpectraMax iD5e Multi-Mode Microplate Reader (Molecular Devices, USA).

Measurement of reactive oxygen species (ROS)

The DCFDA/H2DCFDA - Cellular ROS Assay Kit (Abcam, UK) was used to detect reactive oxygen species (ROS). Briefly, 2.5 × 10⁴ pre-treated cells were inoculated into a dark, clear bottom 96-well microplate and were permitted the cells to adhere overnight. Subsequently, 100 µL of a 20 µM DCFDA solution was added to each well and incubated for 45 min in a light-protected environment. Following this incubation period, the fluorescence of the plate was assessed promptly using a fluorescence plate reader (SpectraMax iD5e Multi-Mode Microplate Reader, Molecular Devices, USA) set to an Ex/Em = 485/535 nm.

Lipid peroxidation assay

The pretreated cells were collected and subsequently resuspended in PBS containing 2 µM of 581/591 C-11 BODIPY dye (Thermo Fisher Scientific, USA). After incubation at 37 °C for 30 min, the cells were centrifuged at 400 × g for 5 min, and the supernatant was discarded. The cells were then washed twice with PBS and resuspended in PBS. Lipid peroxidation was evaluated by flow cytometry (CytoFLEX, Beckman Coulter, USA).

Intracellular cholesterol detection

The pretreated cells were harvested and resuspended in 300 µL of saline. Cell suspension was performed with an ultrasonic processor on ice (300 W, 5 s on / 30 s off, 5 cycles). Protein concentration was quantified by the BCA assay, and total cellular cholesterol was determined using the Total Cholesterol Assay Kit (A111-1-1, Nanning jiancheng, China) according to the manufacturer’s instructions.

Luciferase reporter assay

The gene luciferase reporter plasmid, siRNA, and the pRL-TK (Renilla luciferase plasmid) were co-transfected into HT29 cells using the Lipofectamine 3000 transfection reagent (Vazyme, China). Subsequent quantitative analysis was measured utilizing the Dual-Luciferase^® Reporter Assay System kit (Promega, USA).

Chromatin Immunoprecipitation (ChIP)

Cells were crosslinked with 0.75% formaldehyde for 10 min. Subsequently, glycine was added to attain a final concentration of 125 mM to terminate the reaction. The cells were then collected and lysed in ChIP lysis buffer, while being kept on ice for 10 min, followed by sonication. Then, the cells were incubated with 10 µg of antibody at 4 °C overnight. The resulting immunocomplex was incubated with 20 µl of Protein A/G PLUS-Agarose (Santa Cruz Biotechnology, USA) on a rotator for 2 h to facilitate binding. Following centrifugation of the beads, the complex was washed three times with Low Salt Wash Buffer and once with High Salt Wash Buffer. Finally, the DNA was eluted using Elution Buffer, purified utilizing a PCR purification kit (TIANGWN, China), and subsequently analyzed through qPCR. The primers for ChIP-qPCR are listed in Supplementary Table 5.

Coimmunoprecipitation (Co-IP)

In the co-immunoprecipitation (Co-IP) assay, proteins were extracted using RIPA lysis buffer (CWBIO, China) supplemented with protease inhibitors. Protein A/G magnetic beads (MCE, USA) were pretreated with 0.5% PBST buffer. The beads were then incubated with the indicated primary antibody or IgG control at 4 °C for 2 h. After three washes with 0.5% PBST buffer, an equal volume of protein lysate supernatant was added, followed by incubation at 4 °C for another 2 h. The bead–antibody–protein complexes were washed three times with 0.5% PBST buffer and then eluted with loading buffer. The eluted complexes were heated at 100 °C for 10 min. Finally, the immunoprecipitated protein complexes were subjected to western blot analysis.

Animal experiments

APC^min/+ mice were purchased from Cyagen Biosciences Inc. Male 4-week-old BALB/C nude mice were purchased from the Guangxi Medical University Laboratory Animal Center. Mice were housed under SPF conditions at 24 °C with 50% humidity and a 12-hour light–dark cycle. 5 × 10⁶ control or HT29 KO-SREBP2 cells were subcutaneously injected into each nude mouse (male BALB/c nu/nu nude mice, 4-week-old), with a total of six mice per group. Before the animal experiments, body weights were measured. The body weight of mice in the control group was 16.26 ± 1.954 g, while that of mice in the experimental group was 16.89 ± 1.767 g, with no significant difference observed between the two groups. The mice were euthanized 14 days later. To investigate the inhibitory effects of Betulin on CRC progression, we injected 5 × 10⁶ HT29 wild-type cells into a total of six mice per group. Before the animal experiments, body weights were measured. The body weight of mice in the control group was 16.57 ± 2.324 g, while that of mice in the experimental group was 17.39 ± 2.355 g, with no significant difference observed between the two groups. One group was treated with 20 mg/kg/day Betulin (MCE, USA), while the control group received PEG300 (MCE, USA) as a control. The administration of the drugs was conducted via oral gavage on a daily basis, commencing from the third day post-cell injection, with euthanasia performed after 14 days. Following euthanasia, the tumors were excised, weighed, photographed, and utilized for a variety of experimental analyses. Statistical analyses were performed to evaluate tumor weight and volume. These animal experiments were approved by Animal Trial Ethics Committee of the Guangxi Medical University (Approval Number: 202306024).

Statistical analysis

The data are presented as mean ± SEM derived from at least three independent experiments. Paired t-tests were used to analyze the RNA levels, protein expression, and IHC staining scores of SREBP2 in clinical colorectal cancer tissue samples. Comparisons between two treatment groups were conducted using two-tailed Student’s t-tests. One-way ANOVA was conducted to compare differences among the multiple groups. GraphPad Prism 8 software was used for the assessment of statistical significance and the visualization of data. P-value < 0.05 was considered statistically significant.

Results

SREBP2 was upregulated in human colorectal cancer

Considering the abnormal expression of SREBP2 in various malignant tumors, we first detected the mRNA levels of SREBP2 in 47 CRC and paired adjacent non-tumor tissue samples. As shown in Fig. 1A, SREBP2 was remarkably upregulated in cancerous tissues compared with paired adjacent tissue samples. In addition, the elevated SREBP2 expression positively correlated with CRC clinical stage (Fig. S1). We further detected the protein level of SREBP2 in CRC tissues and the paired adjacent tissues by Western blotting and immunohistochemistry (Fig. 1B-E). The results showed a significant increase in the protein levels of SREBP2 in CRC tissues compared to the paired adjacent tissues. The active SREBP2, namely n-SREBP2, increased obviously in CRC tissues (Fig. 1B). Furthermore, elevated levels of both SREBP2 mRNA and protein were identified in various human CRC cell lines (Fig. 1F-H). Consequently, we found that SREBP2 was markedly upregulated in both colorectal cancer tissues and cell lines.

Fig. 1.

SREBP2 was upregulated in human colorectal cancer. A. Real-time PCR analysis of SREBP2 expression levels in colorectal cancer and adjacent tissues. B, C. Western blotting analysis of SREBP2 in CRC tissues and adjacent tissues. The quantitative results of SREBP2 expression levels were evaluated. D, E. Immunohistochemistry of SREBP2 in tumor sections generated from CRC samples, utilizing adjacent tissues as a control. Scale bar, 50 μm. Analysis of SREBP2 IHC scores in two groups of tissue samples. F. Expression levels of SREBP2 in different CRC cell lines and normal NCM460 cells were determined by real-time PCR. G, H. Western blotting analysis of SREBP2 in different CRC cell lines and normal NCM460 cells. The quantitative results of SREBP2 expression levels were evaluated. The data are presented as the mean ± SEM of at least three independent experiments. ns, not significant, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

SREBP2 knockdown suppressed CRC cell proliferation, migration and invasion in vitro

Previously, we found that the expression of SREBP2 increased in colorectal cancer (CRC). To elucidate the role of SREBP2 in CRC, we employed siRNA-mediated knockdown of SREBP2 in HT29 and RKO cell lines—both of which exhibit higher levels of SREBP2 mRNA and protein. The mRNA and protein levels of SREBP2 was markedly decreased in SREBP2 knockdown group indicating by qPCR and Western blotting experiments, respectively (Fig. 2A–D). Subsequently, colony formation and CCK-8 assays demonstrated that knockdown of SREBP2 suppressed the proliferation of CRC cells, reflected by the colony formation ability and growth rate in si-SREBP2 cells (Fig. 2E–H). Furthermore, wound healing assay revealed that SREBP2 knockdown markedly impaired the migratory abilities of CRC cells. In the wound healing assay, a decrease in the migration rate of si-SREBP2 cells was observed, indicating that SREBP2 was essential for cellular migratory function (Fig. 2I, J). Subsequently, alterations in the migratory and invasive capacities of CRC cells were evaluated through transwell assays (Fig. 2K, L). As anticipated, the depletion of SREBP2 significantly suppressed the migratory and invasive abilities of colorectal cancer cells. Collectively, knockdown SREBP2 in CRC cell lines by siRNA suppressed cell proliferation, migration and invasion.

Fig. 2.

SREBP2 knockdown suppressed CRC cell proliferation, migration and invasion in vitro. The following assays were performed in HT29 or RKO cells treated with si-SREBP2 or si-NC (as control). A–D. Knockdown of SREBP2 was confirmed by real-time PCR (A, C) and western blotting (B, D) in HT29 and RKO cells. E, F. Representative images of colony formation in HT29 and RKO cells. Quantification of colonies was presented. G, H. CCK-8 assay was performed to evaluate cell viabilities of CRC cells. I, J. Wound healing assay was conducted to measure cell migratory abilities. Scale bar, 200 μm. Representative images and quantification were presented. K, L. Transwell migration and invasion assays were performed to determine the effects of SREBP2 on migratory and invasive abilities of HT29 and RKO cells. Scale bar, 200 μm. The number of migrated and invaded cells were quantified. Error bars represent mean ± SEM from at least three independent experiments. ns not significant, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Betulin suppressed CRC cell proliferation, migration, and invasion in vitro

Acting as a specific small-molecule inhibitor of SREBP2, Betulin prevents the proteolytic cleavage of pre-SREBP2 into its active nuclear form (n-SREBP2). It has been reported in the literature that 6 µg/mL Betulin can significantly inhibit the cleavage of Pre-SREBP2 in HT29 cells [23]. To further investigate the function of SREBP2 in CRC cells, HT29 and RKO cells were treated with 6 µg/mL Betulin for 24 h. Western blotting results showed that Betulin significantly decreased n-SREBP2 levels in CRC cells (Fig. 3A, B). Consistent with the observations in si-SREBP2 cells, colony formation assays showed that Betulin significantly suppressed the colony-forming capacity of CRC cells (Fig. 3C, D). Furthermore, CCK-8 assays revealed a marked reduction in the proliferation rate of CRC cells following Betulin treatment (Fig. 3E, F). Subsequently, wound healing and Transwell assays were used to evaluate the effects of Betulin on the migratory and invasive capacities of CRC cells. The results found that Betulin significantly inhibited the migration and invasion of CRC cells (Fig. 3G–J). In summary, these findings demonstrate that Betulin exhibits potent anti-CRC activity through suppressing CRC cell proliferation, migration, and invasion.

Fig. 3.

Betulin suppressed CRC cell proliferation, migration, and invasion in vitro. The following assays were performed in HT29 or RKO cells treated with Betulin (6 µg/mL) or DMSO (as control). A, B. SREBP2 inhibition was confirmed by western blotting in HT29 and RKO cells. The levels of precursor of SREBP2 (pre-SREBP2) and nuclear form of SREBP2 (n-SREBP2) were presented. C, D. Colony formation assays were used to assess treated HT29 and RKO cells proliferation abilities. Representative images and quantification of colonies were presented. E, F. HT29 and RKO cell viabilities were evaluated by CCK-8 assay. G, H. The CRC cell migratory abilities were measured by wound-healing assays. Scale bar, 200 μm. Representative images and quantification were presented. I, J. Transwell migration and invasion assays were performed to determine the effects of Betulin on migratory and invasive abilities of CRC cells. Scale bar, 200 μm. All results represent mean ± SEM from at least three independent experiments. ns, not significant, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

SREBP2 transcriptionally upregulated GPX4 expression

As a transcription factor, SREBP2 activates the transcription of downstream target genes. To elucidate the potential downstream target gene of SREBP2 in CRC, we performed RNA sequencing (RNA-seq) in HT29 cells transfected with si-NC or si-SREBP2. Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis revealed that ferroptosis was among the top 50 pathways most affected by SREBP2 knockdown (Fig. 4A). Consistently, Gene Set Enrichment Analysis (GSEA) of DEGs showed that differentially expressed genes were significantly enriched on ferroptosis in (Fig. S2A). Heatmap visualization revealed that SREBP2 knockdown modulated the expression of key genes involved in the SREBP2-regulated mevalonate (MVA) pathway as well as three critical ferroptosis-associated pathways: polyunsaturated fatty acid-phospholipid (PUFA-PL) synthesis and peroxidation, iron metabolism, and reactive oxygen species (ROS) generation (Fig. 4B). Subsequent qPCR validation in si-SREBP2 HT29 cells showed a significant downregulation of GPX4 and SLC7A11 mRNA levels, whereas ACSL4 expression remained unchanged (Fig. 4C, Fig. S2B). Western blotting analysis further demonstrated a marked decrease in GPX4 protein expression, an upregulation of ACSL4 protein, and no significant alteration in SLC7A11 protein levels in si-SREBP2 cells (Fig. 4D, Fig. S2C).

Fig. 4.

SREBP2 transcriptionally upregulated GPX4 expression. A–D. HT29 cells transfected with si-SREBP2 or si-NC (as control). A, B. RNA-sequence was used to study the function of SREBP2. A. Kyoto Encyclopaedia of Genes and Genomes (KEGG) analysis was used to investigate the signaling pathways that the differentially expressed genes enriched on. B. Heatmap was conducted to show the DEGs. C. The mRNA levels of SREBP2 and GPX4 were examined by real-time PCR. D. The protein levels of SREBP2 and GPX4 were determined by western blotting. E. The sequence logo of a potential SREBP2 binding site (TBS) predicted by JASPAR database. The schematic diagram of the predicted binding site on the GPX4 promoter sequence (from − 431 to − 422 bp) and the corresponding site-directed mutation were showed. F, G. ChIP-qPCR assay was performed using anti-SREBP2 or IgG (as control) antibody in HT29 cells. Agarose gel electrophoresis analysis of ChIP-qPCR products (F). Analysis and visualization of ChIP-qPCR results (G). H. HT29 cells were transfected with luciferase reporter plasmids containing GPX4 promoter (wild/mute type) and further transfected with si-SREBP2 and si-NC (as control). The luciferase activity was measured and normalized. All the results were obtained from at least three independent replicate experiments. The results represent mean ± SEM from at least three independent experiments. ns, not significant, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Regarding the decrease of both mRNA and protein levels of GPX4 in si-SREBP2 HT29 cells, we further investigated whether SREBP2 transcriptionally increases GPX4 expression. Using JASPAR website (https://jaspar.elixir.no/), we predicted eight SREBP2 binding sites on the GPX4 promoter (Fig. S3). Chromatin immunoprecipitation (ChIP) experiment confirmed that SREBP2 can bind to -431~ -422 bp upstream of the GPX4 transcription start site (GTGCCCTGAT) (Fig. 4E-G and Fig. S4). To further verify whether SREBP2 directly regulates GPX4 gene expression, we analyzed the effect of SREBP2 on the transcriptional activity of the GPX4 promoter-luciferase reporter vector. In addition, we constructed a mutant GPX4 promoter luciferase reporter plasmid (wild type, GTGCCCTGAT; mutant type, TGTAAAGTCG) and performed a dual-luciferase reporter assay to identify the SREBP2 binding site (Fig. 4E). The results revealed a decrease in GPX4 promoter transcriptional activity following SREBP2 knockdown. Moreover, a marked reduction in the activity of the mutant promoter was observed, with no significant difference in luciferase intensity between the control and si-SREBP2 groups (Fig. 4H). In conclusion, these results proved that SREBP2 directly increased GPX4 transcription in CRC.

SREBP2 promoted ferroptosis resistance through GPX4

To elucidate the role of SREBP2 in CRC ferroptosis, SREBP2 expression was suppressed in HT29 cells via si-SREBP2, followed by induction of ferroptosis by Erastin. During Erastin induction of CRC ferroptosis, SREBP2 knockdown resulted in elevated levels of malondialdehyde (MDA), reactive oxygen species (ROS), Fe²⁺, and lipid ROS, concomitant with a reduction in the glutathione disulfide/glutathione (GSSG/GSH) ratio (Fig. 5A–E). Like Erastin, SREBP2 knockdown alone promoted CRC ferroptosis indicating by the increased levels of MDA, ROS, Fe²⁺, and lipid ROS, as well as a decrease in the GSSG/GSH ratio (Fig. 5A–E). Conversely, overexpression of the nuclear form of SREBP2 (n-SREBP2) in HT29 cells led to decreased concentrations of MDA, ROS, Fe²⁺, and lipid ROS, alongside an increased GSSG/GSH ratio, indicating that n-SREBP2 conferred ferroptosis resistance in CRC (Fig. 5F–K). Notably, GPX4 knockdown blocked the ferroptosis resistance conferred by n-SREBP2 overexpression (Fig. 5F–K). To further confirm that GPX4 serves as a key downstream effector of SREBP2 in regulating ferroptosis, we knocked down SREBP2 by siRNA and simultaneously overexpressed GPX4, then measured MDA and Fe²⁺ levels in HT29 cells. The results showed that GPX4 overexpression reversed the increased ferroptosis sensitivity induced by SREBP2 knockdown in colorectal cancer cells (Fig. S5A, B). As the key transcription factor in cholesterol synthesis and metabolism, SREBP2 increases intracellular cholesterol levels [12]. We found that SREBP2 knockdown decreased the expression of key cholesterol synthesis enzymes (HMGCR and LDLR) as well as total cholesterol levels in HT29, whereas SREBP2 overexpression showed the opposite effects (Fig. S6A–D). To determine whether SREBP2 regulates ferroptosis through cholesterol synthesis, we supplemented exogenously cholesterol during SREBP2 knockdown. SREBP2 knockdown increased ferroptosis sensitivity, as indicated by elevated levels of MDA and Fe²⁺ while supplementation with exogenous cholesterol cannot reverse the increased ferroptosis sensitivity induced by SREBP2 knockdown (Fig. S6E, F). Collectively, these findings suggested that SREBP2 suppressed ferroptosis in CRC through a GPX4-dependent mechanism.

Fig. 5.

SREBP2 promoted ferroptosis resistance through GPX4. A–E. HT29 cells transfected with si-SREBP2 or si-NC (as control for siRNA) were treated with Erastin (a ferroptosis inducer) or DMSO (as control for erastin). The experiment was divided into four groups: (1) DMSO + si-NC, (2) DMSO + si-SREBP2, (3) Erastin + si-NC, (4) Erastin + si-SREBP2. DMSO + si-NC is the control group for DMSO + si-SREBP2 and Erastin + si-NC. Erastin + si-NC is the control group for Erastin + si-SREBP2. MDA A, ROS (B), Fe²⁺ C levels, and the relative ratio of GSH/GSSG D were detected by assay kits, respectively. E. Flow cytometry was used to analyze lipid peroxidation stained with C11-BODIPY in treated HT29 cells. F–K. HT29 cells transfected with n-SREBP2 or pLEX (as control for n-SREBP2) as well as si-GPX4 or si-NC (as control for siRNA). The experiment was divided into four groups: (1) pLEX + si-NC, (2) n-SREBP2 + si-NC, (3) pLEX + si-GPX4, (4) n-SREBP2 + si-GPX4. pLEX + si-NC is the control group for n-SREBP2 + si-NC. n-SREBP2 + si-NC is the control group for n-SREBP2 + si-GPX4. pLEX + si-GPX4 is the control group for n-SREBP2 + si-GPX4. F. Western blotting of n-SREBP2 and GPX4 in HT29 cells. G-J. MDA G, ROS H, Fe²⁺ I levels, and the relative ratio of GSH/GSSG J were detected by assay kits, respectively. K. Flow cytometry was used to analyze lipid peroxidation stained with C11-BODIPY. All the results represent mean ± SEM from at least three independent experiments. ns, not significant; *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

β-catenin transcriptionally upregulated the expression of SREBP2

Abnormal activation of the Wnt/β-catenin signaling pathway has been observed in the majority of CRC patients and is recognized as a critical factor in the initiation and progression of CRC [24, 25]. β-catenin functions as a transcriptional coactivator by interacting with members of the TCFs/LEF family to modulate gene expression [26]. To study the mechanism involved in SREBP2 increase in CRC, we examined whether β-catenin promoted SREBP2 transcription. qPCR and Western blotting experiments demonstrated that β-catenin knockdown by siRNA decreased the mRNA and protein levels of both SREBP2 and GPX4 (Fig. 6A, B). Furthermore, JASPAR database analysis identified a predicted TCF4 binding site on the SREBP2 promoter region, which was subsequently validated by ChIP assays confirming β-catenin binding at this site (Fig. 6C-E). Dual luciferase reporter assays revealed that the transcriptional activity of the SREBP2 promoter was decreased in cells with β-catenin knockdown (Fig. 6F). To determine whether β-catenin exerts its regulatory effects through interaction with TCF4, we constructed a mutant SREBP2 promoter luciferase reporter plasmid targeting the TCF4 binding site within the SREBP2 promoter region (Fig. 6C). The results revealed a marked reduction in the activity of the mutant promoter, with no significant difference in luciferase intensity between the control and si-β-catenin groups (Fig. 6F). ChIP and Co-IP assays further confirmed that TCF4 binds to the SREBP2 promoter region and interacts with β-catenin (Fig. 6G–I). Collectively, these findings indicate that β-catenin promotes the transcriptional activation of SREBP2 by forming a complex with the transcription factor TCF4.

Fig. 6.

β-catenin transcriptionally upregulated the expression of SREBP2. A, B. HT29 cells transfected with si-β-catenin or si-NC (as control). The levels of β-catenin, SREBP2 and GPX4 were examined by real-time PCR. B. The protein levels of total β-catenin, active-β-catenin, SREBP2 and GPX4 were determined by western blotting. C. The sequence logo of a potential TCF4 binding site (TBS) predicted by JASPAR database. The schematic diagram of the predicted binding site in the SREBP2 promoter sequence (from − 1628 to − 1621 bp) and the corresponding site-directed mutation were showed. D, E. ChIP-qPCR assay was performed using anti-β-catenin or control IgG antibody in HT29 cells. Agarose gel electrophoresis analysis of ChIP-qPCR products D. Analysis and visualization of ChIP-qPCR result (E). F. HT29 cells were transfected with luciferase reporter plasmids containing SREBP2 promoter (wild/mute type) and further transfected with si-β-catenin and si-NC (as control). The luciferase activity was measured and normalized. G, H. ChIP-qPCR assay was performed using anti-TCF4 or control IgG antibody in HT29 cells. Agarose gel electrophoresis analysis of ChIP-qPCR products G. Analysis and visualization of ChIP-qPCR result H. I. The interaction between β-catenin and TCF4 was examined by Co-IP. All the results represent mean ± SEM from at least three independent experiments. ns not significant, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

SREBP2 Inhibition suppressed CRC progression through promoting ferroptosis in vivo

Previously, we have demonstrated that SREBP2 suppressed ferroptosis by upregulating the expression of GPX4, thereby facilitating the progression of CRC in vitro. To further elucidate the role of SREBP2 in tumor growth, we established an SREBP2 knockout (SREBP2-KO) cell model and subcutaneously injected HT29-SREBP2-KO cells or control cells into nude mice to evaluate the effect of SREBP2 on CRC tumorigenicity (Fig. S7A, B). Consistent with the findings in vitro, SREBP2 knockout resulted in the inhibition of CRC progression, reflected by a marked decrease in both tumor volume and tumor weight (Fig. 7A–D). Furthermore, IHC analysis revealed a marked decrease in the SREBP2 IHC score and Ki-67 positivity within xenograft tumors formed by SREBP2-KO cells (Fig. S8A, Fig. 7E). Additionally, in the xenograft tumors, GPX4 expression was significantly diminished, while the expression of 4-hydroxynonenal (4-HNE), a lipid peroxidation product, was notably elevated (Fig. 7E), indicating that SREBP2 inhibition promoted ferroptosis sensitivity in vivo. To further assess the anti-tumor potential of targeting SREBP2, wild-type HT29 cells were subcutaneously implanted into nude mice, and the mice were subsequently treated with vehicle control or the SREBP2 inhibitor Betulin. Expectedly, Betulin treatment induced ferroptosis and exerted antitumor effects, reflected by reductions in tumor volume and weight, decreased Ki-67 positivity, reduced GPX4 expression, and increased 4-HNE levels (Fig. 7F–J). We further performed IHC analyses of β-catenin, ACSL4, and SLC7A11 in tumor tissues. Si-NCe only SREBP2 was targeted, there was no significant change in the expression of its upstream molecule β-catenin. In contrast, ACSL4 expression increased in tumors with SREBP2 knockout or inhibition, whereas SLC7A11 protein levels remained unchanged, consistent with our in vitro findings (Fig. S8A–B). In addition, analyses of orthotopic intestinal tumors in APC^min/+ mice, as well as CRC liver and lung metastatic tumors in nude mice, revealed a positive correlation between SREBP2 and GPX4 expression (Fig. S9). This positive association was further validated in clinical CRC specimens (Fig. S10). Collectively, these results demonstrate that GPX4 functions as a critical downstream effector of SREBP2. Targeting SREBP2 enhances ferroptosis in CRC, thereby exerting antitumor effects and underscoring its therapeutic potential in CRC treatment.

Fig. 7.

SREBP2 inhibition suppressed proliferation and promoted ferroptosis of CRC in vivo. A–E. Xenograft tumors formed by HT29-SREBP2-KO or HT29-v2 (LentiCRISPR v2 vector cells as control) in nude mice. Tumor volume of xenografts C and quantitative analysis of tumor weight (D) formed by HT29-v2 or HT29-SREBP2-KO cells. E. Ki-67, GPX4, and 4-HNE were examined by IHC. Scale bar, 50 μm. Analysis of Ki-67 positive rate, GPX4, and 4-HNE IHC score in xenograft tumor tissue from v2 and SREBP2-KO groups. F–J. Xenograft tumors formed by wild-type HT29 in nude mice treated with Betulin or PEG300 (as control). H, I. Tumor volume of xenografts H)and quantitative analysis of tumor weight (I) from nude mice treated with Betulin or PEG300 as control. J. Ki-67, GPX4, and 4-HNE were examined by IHC. Scale bar, 50 μm. Analysis of Ki-67 positive rate, GPX4, and 4-HNE IHC score in xenograft tumor tissue from PEG300 and Betulin groups. All the results represent mean ± SEM from at least three independent experiments. ns not significant, *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001

Discussion

Numerous research studies have established that SREBP2 plays a key role in cancer development. Elevated SREBP2 has been shown to participate in the advancement of various malignancies, including hepatocellular carcinoma, breast carcinoma, and ovarian cancer [27–29]. In this study, we found that the expression of SREBP2 increased significantly in clinical colorectal cancer (CRC) specimens and CRC cell lines. Functionally, SREBP2 knockdown suppressed the in vitro proliferation, migration, and invasion of CRC cells, as well as tumor growth in vivo. Furthermore, both in vitro and in vivo studies demonstrated that treatment with the SREBP2 inhibitor Betulin exhibited significant anti-CRC effects. Mechanistically, SREBP2 promoted CRC ferroptosis resistance through transcriptionally increasing GPX4 expression. Furthermore, β-catenin activation contributed to increased SREBP2 expression in CRC. Collectively, these results highlight the pivotal role of SREBP2 in CRC progression and ferroptosis resistance, indicating its significant potential for clinical application in CRC treatment.

Previous studies have reported that SREBP2 overcomes KRAS mutation-induced senescence in pancreatic cancer [30]. In hepatocellular carcinoma, SREBP2 promotes epithelial–mesenchymal transition [15]. Additionally, SREBP2 contributes to chemotherapy resistance in non-small cell lung cancer [31]. Although these studies have highlighted the role of SREBP2 in various cancers, its specific mechanisms in colorectal cancer progression remain to be fully elucidated. To study the function of SREBP2 in CRC progression, we used RNA-seq and identified that SREBP2 participates in the regulation of the ferroptosis pathway. Typically, precursor SREBP2 undergoes proteolytic processing within the Golgi apparatus to generate the nuclear form (n-SREBP2), which translocates to the nucleus to bind promoter regions of downstream target genes, thereby modulating their transcription [12]. However, little is known about the transcriptional regulation of key ferroptosis-related genes by SREBP2. We found a significant downregulation of GPX4 expression, indicated by the decrease in both mRNA and protein levels following SREBP2 knockdown. Mechanistically, our study indicated that SREBP2 directly binds to the promoter region of GPX4, enhancing its transcriptional activity. In our study, we observed that knockdown SREBP2 led to a decrease in SLC7A11 mRNA levels, while the protein level remained unchanged. For ACSL4, we found that SREBP2 knockdown had no significant effect on its mRNA while increased its protein level. To analyze this inconsistency, we conducted a further analysis. miR-33 is located within an intron of the SREBP2 gene [32–34]. Using the TargetScan database, we predicted the potential targets of miR-33 and found that miR-33 may bind to the 3′ UTR of SLC7A11 and ACSL4 mRNA through which suppress their translation (Fig. S11, S12). SREBP2 knockdown caused a significantly decrease of SLC7A11 mRNA and a slightly decrease of ACSL4 mRNA (with no significance). When SREBP2 is knocked down, the expression of miR-33 is also reduced, thereby relieving its inhibitory effect on SLC7A11 and ACSL4 translation. As a result, no significant change in SLC7A11 protein levels and an increase in ACSL4 protein was observed. However, this assumption requires further experimental validation. Acting as the key transcription factor in cholesterol synthesis and metabolism, SREBP2 has been reported to drive tumor EMT and drug resistance via cholesterol regulation. Whether SREBP2 participates in CRC ferroptosis through cholesterol regulation needs further study. Controversially, a study demonstrated that SREBP2 mediated cholesterol metabolism promoted ferroptosis [35]. We further found that exogenous cholesterol supplementation fails to reverse the increased ferroptosis sensitivity induced by SREBP2 knockdown indicating that SREBP2 regulates ferroptosis primarily through a GPX4-dependent pathway. Collectively, this study uncovers the role of SREBP2 as a negative regulator of ferroptosis in colorectal cancer, further expanding its known biological roles in malignant tumors.

Mutations in APC lead to the activation of the Wnt/β-catenin signaling pathway that results in β-catenin accumulation and its translocation to the nucleus, where it associates with TCF/LEF1 to stimulate the transcription of Wnt target genes [36, 37]. This mechanism is a critical contributor to the initiation and progression of CRC. Recent studies have shown that the β-catenin/TCF4 complex confers resistance to cuproptosis in cancer cells by transcriptionally activating ATP7B [38]. We therefore investigated whether the β-catenin/TCF4 complex regulates SREBP2 expression to modulate ferroptosis in CRC. Our results demonstrate that β-catenin acts as a transcriptional coactivator with TCF4 to enhance SREBP2 transcription. Knockdown of β-catenin in CRC cells led to a significant decrease in the expression of both SREBP2 and GPX4. In summary, our findings indicate that the upregulation of SREBP2 in CRC is mediated by β-catenin and suggest that β-catenin may influence ferroptosis in CRC through SREBP2.

Betulin, a pentacyclic triterpene abundantly found in birch bark, inhibits SREBP2 activation by preventing the release of the SCAP–SREBP2 complex from the ER [19, 20]. Numerous studies in breast and liver cancers have found tumor-suppressive effects of Betulin [39, 40]. In this study, to further investigate the therapeutic potential of Betulin targeting SREBP2 in CRC, we evaluated its effects in both cellular and animal models. Consistent with the findings in other cancers, Betulin exhibited significant antitumor activity in vitro and in vivo, supporting SREBP2 as a promising anticancer target. Nonetheless, whether Betulin can be developed into a clinical anticancer drug requires further investigation.

In conclusion, our study reveals that SREBP2 promotes the progression of CRC through ferroptosis resistance. Mechanistic investigations demonstrate that SREBP2 suppresses ferroptosis in CRC by transcriptionally activating GPX4 and upregulating its expression (Fig. 8). These findings identify SREBP2 as a novel molecular marker and a promising therapeutic target for CRC treatment, while also providing foundational support for the potential clinical application of Betulin.

Fig. 8.

The mechanism of SREBP2-suppressed ferroptosis in CRC. In colorectal cancer, the activation of β-catenin contributes to SREBP2 upregulation. Acting as a transcription factor, the active SREBP2 promotes GPX4 transcription. And then, the elevated GPX4 catalyzes the conversion of GSH to GSSG, leading to a reduction in lipid ROS level, thereby enhancing CRC cell resistance to ferroptosis and promoting colorectal cancer progression. Created in BioRender. Chen, R. (2025) https://BioRender.com/dsllbwn

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

HZ: Writing—original draft, Writing—review & editing, Validation, Data curation. RC: Writing—original draft, Writing—review & editing, Investigation, Visualization. TC: Writing—original draft, Investigation, Visualization. ML: Visualization, Data curation. SW: Visualization, Data curation. MZ: Visualization, Data curation. DL: Visualization, Data curation. JY: Visualization, Data curation. JC: Visualization, Data curation. BY: Visualization, Data curation. RL: Visualization, Data curation. WJ: Visualization, Data curation. ZL: Writing—review & editing, Supervision, Project administration, Funding acquisition, Conceptualization. All authors reviewed the manuscript and approved the final version for publication.

Funding

This work is supported by the Youth Project of Guangxi Natural Science Foundation (2023GXNSFBA026004), Bagui youth top talent training program, National Natural Science Foundation of China (82170421 and 82360464), Young leading talents training program of Guangxi Medical University, and Outstanding Young Talent Training Program of Guangxi Medical University.

Data availability

The data that support the findings of this study are available from the corresponding author upon reasonable request.

Declarations

Conflict of interest

The authors declare no competing interests.