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. 2026 Jun 30;34(4):891–900. doi: 10.4062/biomolther.2026.038

Erastin-Induced Ferroptosis Overcomes Oxaliplatin Resistance in Colorectal Cancer Cells

Na Yeong Lee 1,†, Sang Hoon Joo 2,†, Si Yeong Seo 1, Seung-On Lee 1, Goo Yoon 3, Jin Woo Park 1,3,*, Jung-Hyun Shim 1,3,4,*
PMCID: PMC13324481  PMID: 42375092

Abstract

Chemotherapy resistance remains a significant obstacle in the treatment of colorectal cancer (CRC) patients. Here, we examined the ferroptosis susceptibility of the chemotherapy-resistant CRC cell line HCT116-OxR relative to its parental counterpart. The increased basal levels of reactive oxygen species (ROS) and ferroptosis-associated proteins observed in HCT116-OxR cells rendered them more sensitive to erastin-induced cell death, as determined by the MTT and soft agar assays. Mechanistically, erastin elicited a reduction in mitochondrial membrane potential (MMP). Notably, this loss of MMP was not accompanied by detectable apoptotic changes, suggesting that a non-apoptotic cell death pathway may be involved. Rather, erastin elevated intracellular iron and ROS concentrations, resulting in lipid peroxidation consistent with ferroptosis. The pivotal involvement of ROS was supported by N-Acetyl-L-cysteine pretreatment, and ferrostatin-1 pretreatment further verified that erastin-induced cell death was mediated by ferroptosis. The heightened erastin sensitivity of HCT116-OxR cells suggests a therapeutic vulnerability that could be exploited to induce ferroptosis in chemotherapy-resistant CRC. This ferroptosis susceptibility may offer a promising strategy for overcoming treatment resistance and improving therapeutic outcomes.

Keywords: Erastin, Oxaliplatin, Chemotherapy-resistance, Colorectal cancer, Reactive oxygen species, Ferroptosis

INTRODUCTION

Chemotherapy resistance poses a significant obstacle in the management of colorectal cancer (CRC) (Tharin et al., 2021), which remains among the most common malignancies and is responsible for nearly a million deaths annually (Siegel et al., 2023). Although traditional chemotherapeutic regimens utilizing oxaliplatin (Ox) demonstrate efficacy during the initial phases of treatment, they often drive the emergence of aggressive, drug-resistant cancer cell clones, resulting in relapse with a more malignant phenotype (Cayrefourcq et al., 2021). Multiple mechanisms contribute to Ox-resistance, including the increased activity of drug efflux pumps, alterations in apoptotic pathways (Yang et al., 2006), and enhanced secretion of growth factors by the Ox-resistant variant of HCT116 CRC cells (HCT116-OxR), representing a notable resistance mechanism (Bose et al., 2011).

Reactive oxygen species (ROS) are essential in modulating the balance between cell survival and cell death (Schumacker, 2015). Cancer cells may utilize the pro-tumorigenic activities of ROS to promote malignancy and progression (Cui et al., 2018). As a result, moderate elevation of ROS levels is commonly observed in cancer cells, although ROS concentrations exceeding a cytotoxic threshold induce apoptosis (Chun and Joo, 2022). To counter the harmful effects of excessive ROS, cancer cells increasingly depend on potent antioxidant defense systems (Marengo et al., 2016). In contrast, disrupting these antioxidant systems or artificially raising ROS concentrations to cytotoxic levels has emerged as a potential therapeutic approach for targeting cancer cells, given the cytotoxicity of ROS. Substantial increases in ROS can lead to oxidative damage of essential biomolecules, including DNA, proteins, and lipids, and may also result in a collapse of mitochondrial membrane potential (MMP) (Idelchik et al., 2017). Disruption of MMP can initiate apoptotic signaling (Gottlieb et al., 2003) or other forms of regulated cell death (Kroemer and Reed, 2000).

Erastin is recognized for its role in inducing ferroptosis, a specific type of regulated cell death mediated by iron (Dixon et al., 2012). Although studies have demonstrated that erastin can trigger ferroptosis in various cancer cell types (Joo et al., 2026), the use of ferroptosis induction as a method to sensitize cancer cells that are resistant to chemotherapy has not been extensively investigated. In our study, we evaluated the response of the Ox-resistant CRC cell line HCT116-OxR to erastin. The findings reveal that HCT116-OxR exhibited greater sensitivity to erastin-induced cytotoxicity compared to the parental HCT116 cell line.

MATERIALS AND METHODS

Chemicals and reagents

Ox was acquired from MedKoo Biosciences (Durham, NC, USA) and prepared in N,N-dimethylformamide (DMF), while erastin was sourced from GLPBIO Technology Inc. (Montclair, CA, USA) and dissolved in dimethyl sulfoxide (DMSO; Thermo Fisher Scientific, Waltham, MA, USA). N-Acetyl-L-cysteine (NAC) was sourced from Sigma-Aldrich (St. Louis, MO, USA), and ferrostatin-1 (Fer-1) was obtained from Selleck Chemicals (Houston, TX, USA). Dulbecco’s modified Eagle’s medium (DMEM), Roswell Park Memorial Institute 1640 medium (RPMI-1640), and Minimum Essential Medium (MEM) were procured from Welgene (Gyeongsan, Korea). MEM vitamin solution, MEM non-essential amino acids (NEAA), and penicillin-streptomycin (P/S) were purchased from Gibco (Carlsbad, CA, USA). Furthermore, 0.05% trypsin/0.53 mM EDTA solution was sourced from Corning (NY, USA).

Cell culture and treatment

The human keratinocyte cell line HEKa and the Ox-sensitive colorectal cancer cell line HCT116 were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA). The Ox-resistant derivative HCT116-OxR was generously provided by MD Anderson Cancer Center (Houston, TX, USA). HEKa cells were maintained in DMEM, while HCT116 cells were cultured in RPMI-1640 medium, both supplemented with 10% FBS and 1% P/S. For HCT116-OxR cells, MEM was used along with 10% FBS, 1% P/S, 1% MEM vitamin solution, 1% MEM NEAA, and 1% sodium pyruvate. Incubation was performed at 37°C in a humidified 5% CO₂ environment. To ensure maintenance of Ox resistance, HCT116-OxR cells were continuously cultured with 2 µM Ox added to the medium, and the drug was removed before cell treatment. For experimental treatments, cells were exposed to designated compounds for either 24 h or 48 h. In inhibitor assays, NAC (4 mM, 3 h) or Fer-1 (1 µM, 12 h) was applied as a pre-treatment prior to administering erastin for an additional 48 h.

Western blot analysis

After harvesting, the collected cell pellets were lysed on ice, and total cellular proteins were extracted using the PRO-PREP Protein Extraction Solution (iNtRON Biotechnology, Seongnam, Korea) supplemented with protease inhibitors. The lysates were subsequently sonicated and centrifuged at 13,000 rpm for 15 min at 4°C, after which the supernatants were recovered for analysis. Protein concentrations were quantified using the DC Protein Assay Kit (Bio-Rad, Hercules, CA, USA). Equivalent amounts of protein (20–30 µg) were resolved on 8%, 10%, or 15% SDS–polyacrylamide gels, with the gel percentage selected based on the molecular weight of the target proteins, and separated by electrophoresis at 100–120 V for 90–120 min. The proteins were transferred onto PVDF membranes at 100 V for 90–120 min in Tris–glycine transfer buffer (pH 8.3). Following transfer, membranes were blocked using 3%–5% skim milk for 2 h at room temperature and then incubated overnight at 4°C with the following primary antibodies: β-actin (1:1000, sc-47778), cyclin B1 (1:500, sc-7393), Cdc2 (1:1000, sc-8395), p21 (1:1000, sc-6246), ACSL4 (1:1000, sc-365230) (Santa Cruz Biotechnology), and xCT/SLC7A11 (1:1000, 12691S) and GPX4 (1:1000, 59735S) (Cell Signaling Technology). After thorough washing, the membranes were incubated with HRP-conjugated secondary antibodies (Thermo Fisher Scientific, Waltham, MA, USA) for 2 h at room temperature. Protein bands were detected using an enhanced chemiluminescence detection system and visualized with an iBright CL1500 system (Thermo Fisher Scientific).

Determination of intracellular ROS levels

Intracellular ROS levels were quantified using the CellROX Green Reagent (C10444, Invitrogen, Carlsbad, CA, USA). Both control and treated cells were collected and resuspended in 1× PBS containing 5 µM CellROX Green and incubated at 37°C for 30 min in the dark. Cells were then pelleted by centrifugation at 3,000 rpm for 5 min at 24°C, washed once with 1× PBS, and resuspended in 1× PBS. Fluorescence was analyzed using a MACSQuant Analyzer 16 flow cytometer (Miltenyi Biotec, Bergisch Gladbach, Germany) with the B1 channel (525/50 nm). Data were acquired and processed with the MACSQuantify software.

Measurement of lipid peroxidation

Lipid peroxidation was evaluated using BODIPY 581/591 C11 (D3861, Thermo Fisher Scientific). A stock solution of 5 mM was prepared in DMSO, and cells were incubated with 2.5 µM BODIPY C11 for 15 min at 37°C in darkness. Following staining, cells were washed once with 1× HBSS, detached, collected, and centrifuged at 3,000 rpm for 5 min at 24°C. The pellets were then washed again, resuspended in 1× HBSS, and maintained on ice before measurement. Fluorescence was detected using a MACSQuant VYB flow cytometer (Miltenyi Biotec) with the B1 channel (525/50 nm), and the data were analyzed via MACSQuantify software.

Cell viability assay

Cell viability was assessed using the MTT assay. MTT powder (M6494, Invitrogen) was dissolved to prepare a working solution, which was then added to each well at a final concentration of 1 mg/mL. Following incubation at 37°C to facilitate formazan formation, the supernatant was carefully removed, and the resulting crystals were dissolved in DMSO. Absorbance was recorded at 570 nm using a Varioskan LUX Multimode Microplate Reader (Thermo Fisher Scientific).

Anchorage-independent growth assay

Anchorage-independent growth was quantified by conducting a soft agar colony formation assay as previously described (Lee et al., 2025). After cells were seeded, the plates were incubated for 10 days and colonies were examined under a light microscope (Leica, Wetzlar, Germany). Images were analyzed with i-Solution software (Vancouver, BC, Canada). Colonies measuring ≥40 µm in diameter were counted for quantitative evaluation.

Cell cycle analysis

Cell cycle distribution was evaluated by propidium iodide (PI) staining. After experimental treatment, cells were fixed in 70% ethanol at −20°C overnight. Subsequently, cells were washed with 1× PBS to eliminate residual ethanol and then incubated with 50 µg/mL RNase A (EN0531, Thermo Fisher Scientific) for 15 min at room temperature. PI solution (130-093-233, Miltenyi Biotec) was added directly before flow cytometry. DNA content was analyzed using a MACSQuant VYB flow cytometer with PI fluorescence detected at the Y2 channel (615/20 nm). Data interpretation was performed using MACSQuantify software.

MMP analysis

MMP was determined by employing JC-1 Dye (T3168, Invitrogen). After harvesting, cells were suspended in serum- and supplement-free RPMI-1640 or MEM supplemented with 5 µg/mL JC-1, and incubated at 37°C for 15 min in the dark. Following centrifugation (3,000 rpm, 5 min, 24°C) and removal of the supernatant, cells were resuspended in 1× PBS. JC-1 monomer (green) and aggregate (red) signals were detected using a MACSQuant VYB flow cytometer in the B1 (525/50 nm) and Y1 (586/15 nm) channels, respectively. Data analysis was performed with MACSQuantify software.

Apoptosis analysis

Apoptosis was evaluated using the Annexin V–FITC apoptosis detection kit (130-092-052, Miltenyi Biotec). Cells were collected, washed in cold 1× PBS, and then resuspended in 1× binding buffer. Annexin V–FITC (10 µL) was added and samples were incubated for 15 min at room temperature in the dark. After an additional wash, cells were resuspended in 500 µL 1× binding buffer, and PI solution was administered immediately before analysis. Fluorescence was acquired with a MACSQuant Analyzer 16 flow cytometer using the B1 (525/50 nm) and B3 (615/20 nm) channels. Data were interpreted using MACSQuantify software.

Intracellular Iron Measurement

Intracellular iron levels were quantified using FerroOrange (F374, Dojindo Laboratories). Following cell collection, cells were resuspended in 300 µL serum- and supplement-free RPMI-1640 or MEM and incubated with 1 µM FerroOrange at 37°C for 30 min in the dark. After centrifugation (3,000 rpm, 5 min, 24°C) and a subsequent wash, cells were again resuspended in 1× HBSS. Fluorescence was detected on a MACSQuant VYB flow cytometer using the Y1 channel (586/15 nm), and results were processed with MACSQuantify software.

Statistical analysis

Data are shown as means ± standard deviation (SD). All statistical analyses were conducted using KaleidaGraph 4.5.2 (Synergy Software, Reading, PA, USA). Differences between groups were assessed using one-way analysis of variance, with Tukey’s post hoc test applied where appropriate. A p value of <0.05 was regarded as indicating statistical significance.

RESULTS

The Ox-resistant HCT116-OxR CRC cells exhibit sustained increases in ferroptosis-related proteins and ROS

To assess possible variations in ferroptosis-related protein expression between HCT116 and HCT116-OxR cells, western blot analysis was conducted. The analysis revealed that proteins involved in antioxidant defense—namely SLC7A11 and GPX4—along with ACSL4, which enhances lipid peroxidation, were found at higher levels in HCT116-OxR cells compared to HCT116 cells (Fig. 1A). The normalized expression levels of SLC7A11, GPX4, and ACSL4, relative to β-actin, were measured to be 3.43-fold, 1.74-fold, and 2.09-fold greater, respectively, in HCT116-OxR cells (Fig. 1B-1D). To further characterize baseline oxidative stress, flow cytometry was performed with CellROX Green staining. These results demonstrated that HCT116-OxR cells had an ROS level of 74.00%, corresponding to a 2.49-fold elevation over the 29.77% found in HCT116 cells (Fig. 1E). Additionally, flow cytometry using BODIPY-C11 revealed the lipid peroxidation level in HCT116-OxR cells to be 66.56%, which is 2.76-fold higher than in HCT116 cells (24.09%) (Fig. 1F).

Fig. 1.

Fig. 1

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Expression of ferroptosis-associated proteins and levels of intracellular ROS/lipid peroxidation in CRC cells. (A) Western blot results of SLC7A11, GPX4, and ACSL4 in HCT116 and HCT116-OxR cells. (B-D) Quantitative assessment of SLC7A11, GPX4, and ACSL4 expression normalized to β-actin. (E) Representative flow cytometry images of CellROX Green staining (left) and its quantification (right). (F) Representative flow cytometry images of BODIPY 581/591 C11 staining (left) and corresponding quantification (right). Data are presented as mean ± SD (n=3). **p<0.01, ***p<0.001 versus HCT116.

HCT116-OxR cells demonstrate greater susceptibility to erastin-induced cytotoxicity

To assess the viability of HCT116, HCT116-OxR, and human keratinocyte HEKa cells exposed to erastin, an MTT assay was conducted. Erastin (1-16 μM) reduced cell viability in both a time- and concentration-dependent fashion. The IC50 values after 48 h incubation were 14.57 μM for HCT116 and 2.85 μM for HCT116-OxR cells (Fig. 2A). For HCT116 cells treated with Ox (2 μM), cell viability decreased to 76.17% at 24 h and 48.83% at 48 h relative to the vehicle control. In contrast, HCT116-OxR cells maintained high viability, at 95.67% (24 h) and 100.66% (48 h), confirming the Ox-resistance of the HCT116-OxR cell line (Fig. 2B). In comparison, treatment with erastin (1-16 μM) for 24 h did not significantly alter HEKa cell viability (Fig. 2C). After 48 h at the highest concentration of erastin (16 μM), a slight reduction in HEKa cell viability to 90.26% was observed. However, after Ox treatment (2 μM) for 48 h, HEKa cell viability declined to 75.80% compared to the vehicle control. The soft-agar assay indicated that erastin (1, 2, 4, and 16 µM) administration for 10 days significantly suppressed colony formation (Fig. 2D). The percentage of remaining colonies in HCT116 cells was 87.94%, 45.73%, 18.59%, and 0.00% after erastin treatment at 1, 2, 4, and 16 µM, respectively. The corresponding values for HCT116-OxR cells were lower, at 33.95%, 1.86%, 0.00%, and 0.00%, respectively. Moreover, following Ox treatment (2 μM), the percentage of remaining colonies was 13.07% in HCT116 and 74.88% in HCT116-OxR cells compared to the vehicle control (Fig. 2E). The relative colony size in HCT116 cells was 100.16%, 104.41%, 75.06%, and 0.00% after erastin treatment at 1, 2, 4, and 16 µM, respectively, and the respective values in HCT116-OxR cells were 121.88%, 16.65%, 0.00%, and 0.00%. The relative colony size after Ox treatment (2 μM) was 41.69% in HCT116 cells and 111.62% in HCT116-OxR cells (Fig. 2F).

Fig. 2.

Fig. 2

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Impact of erastin on cell viability and anchorage-independent growth of CRC cells. (A, B) HCT116 and HCT116-OxR cells were exposed to escalating doses of erastin (1–16 µM) or oxaliplatin (Ox, 2 µM) for 24 h and 48 h. Cell viability was assessed using the MTT assay. Data are presented as mean ± SD (n=4). **p<0.01, ***p<0.001 compared to control. (C) HEKa cells were treated with erastin (1–16 µM) or Ox (2 µM) under matching conditions to evaluate drug selectivity. Data are presented as mean ± SD (n=4). ***p<0.001 compared to control. (D) Anchorage-independent cell proliferation was evaluated through a soft agar colony formation assay after administering erastin (1, 2, 4, 16 µM) or Ox (2 µM) to HCT116 and HCT116-OxR cells for 10 days. Representative images are displayed. (E, F) Colony count and colony size were quantitatively measured using i-Solution software. Data are presented as mean ± SD (three independent areas in each triplicate experiment). *p<0.05, ***p<0.001 compared to control.

Erastin induces cell cycle arrest at the G2/M phase in HCT116-OxR cells

Flow cytometric analysis using PI was performed on CRC cells exposed to erastin (1, 2, and 4 µM) or Ox (2 µM) for 48 h (Fig. 3A). In HCT116 cells, the proportion of cells in the G2/M phase remained relatively stable, changing from 28.23% to 28.09%, 27.56%, and 28.17% after erastin treatment at 1, 2, and 4 µM, respectively (Fig. 3B). Conversely, in HCT116-OxR cells, the G2/M population increased from 33.95% to 36.45%, 37.37%, and 38.58%, respectively (Fig. 3C). Western blot analysis demonstrated that the protein expression of cyclin B1, Cdc2, and p21 was largely unchanged in HCT116 cells after erastin exposure. In contrast, HCT116-OxR cells showed a substantial increase in p21 and decreases in cyclin B1 and Cdc2 (Fig. 3D).

Fig. 3.

Fig. 3

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Erastin-induced alterations in cell cycle progression in CRC cells. (A) Representative flow cytometry plots stained with PI for HCT116 and HCT116-OxR cells after 48 h exposure to erastin (1, 2, 4 µM) or oxaliplatin (Ox, 2 µM). (B, C) Quantitative analysis of cell cycle phase distribution (G0/G1, S, and G2/M phases) derived from the flow cytometry data shown in (A) for HCT116 and HCT116-OxR cells. Data are presented as mean ± SD (n=3). *p<0.05, **p<0.01, ***p<0.001 compared to control. (D) Immunoblotting of cell cycle-related proteins Cyclin B1, Cdc2, and p21 in HCT116 and HCT116-OxR cells following erastin (1, 2, 4 µM) treatment for 48 h.

Erastin induces a non-apoptotic type of mitochondrial dysfunction

To evaluate whether erastin triggers mitochondrial dysfunction in CRC cells, we used flow cytometry to measure the percentage of JC-1 green–positive cells following treatment with erastin (1, 2, and 4 µM) or Ox (2 µM) for 48 h (Fig. 4A). The proportion of JC-1 green HCT116 cells remained largely unchanged following erastin exposure. 5.69% in the vehicle control versus 6.09%, 7.28%, and 8.25% after exposure to 1, 2, and 4 µM erastin, respectively. Conversely, in HCT116-OxR cells, erastin significantly elevated the percentage of JC-1 green cells from 3.08% in the vehicle control to 6.81%, 16.43%, and 32.37% upon treatment with 1, 2, and 4 µM erastin, respectively. Following Ox treatment, 28.95% and 6.09% of JC-1 green cells were detected in HCT116 and HCT116-OxR cells, respectively. To investigate if the loss of MMP was linked to apoptosis, we conducted flow cytometry with annexin V/PI staining (Fig. 4B). There were no significant erastin-induced shifts in annexin V–positive cells in HCT116 cells: 4.91% in the vehicle control compared to 5.62%, 4.86%, and 4.52% after administration of 1, 2, and 4 µM erastin, respectively. Similarly, in HCT116-OxR cells, the percentage of annexin V–positive cells remained stable, from 4.28% in the vehicle control to 3.95%, 3.94%, and 4.22% after 1, 2, and 4 µM erastin, respectively. As anticipated, Ox treatment led to an increase in annexin V–positive cells to 17.66% in HCT116 cells only while the corresponding value in HCT116-OxR cells was 4.14%.

Fig. 4.

Fig. 4

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Erastin-mediated effects on MMP and induction of apoptosis in CRC cells. (A) Representative JC-1-stained flow cytometry plots (left) and the corresponding quantitative analysis (right) for HCT116 and HCT116-OxR cells after a 48 h exposure to erastin (1, 2, 4 µM) or oxaliplatin (Ox, 2 µM). (B) Flow cytometry data of Annexin V/PI staining (left) and related quantitative analysis (right) in HCT116 and HCT116-OxR cells following 48 h treatment with erastin (1, 2, 4 µM) or Ox (2 µM). Results are shown as mean ± SD (n=3). ***p<0.001 compared to control.

Erastin promotes lipid peroxidation, leading to ferroptosis in CRC cells

Flow cytometry using FerroOrange was conducted to assess intracellular iron accumulation in CRC cells following erastin treatment (Fig. 5A). In HCT116 cells, there were no significant alterations in the FerroOrange signal, which reflects intracellular iron levels: 30.19% in the vehicle control compared to 31.16%, 27.86%, and 29.22% after exposure to 1, 2, and 4 µM erastin, respectively. In contrast, HCT116-OxR cells demonstrated an increase in the FerroOrange signal from 16.04% in the vehicle control to 46.79%, 68.54%, and 91.79% upon administration of 1, 2, and 4 µM erastin, respectively. Treatment with Ox elevated the FerroOrange signal to 84.48% in HCT116 cells, while the signal in HCT116-OxR cells remained constant (16.93%), consistent with expectations. Next, flow cytometry using BODIPY 581/591 C11 was conducted to assess lipid peroxidation following erastin treatment (1, 2, and 4 µM; Fig. 5B). The BODIPY C11 signal demonstrated minimal change in HCT116 cells: 25.89 % in the vehicle control compared to 24.83%, 25.62%, and 25.57% after exposure to 1, 2, and 4 µM erastin, respectively. In contrast, the BODIPY C11 signal was elevated in HCT116-OxR cells, increasing from 32.49% in the vehicle control to 44.31%, 60.69%, and 86.90% following treatment with 1, 2, and 4 µM erastin, respectively. Ox treatment led to an increase in the signal to 86.77% in HCT116 cells, while the signal was not altered in HCT116-OxR cells (41.06%), consistent with expectations. Western blotting indicated that erastin treatment did not significantly modify SLC7A11, GPX4, or ACSL4 expression in HCT116 cells. Conversely, in HCT116-OxR cells, erastin resulted in decreased SLC7A11 and GPX4 levels, alongside upregulation of ACSL4 (Fig. 5C). Finally, MTT assay results demonstrated that cell viability, which was reduced to 43.05% and 26.93% in HCT116 and HCT116-OxR cells, respectively, following erastin exposure, was increased to 70.31% and 60.94% by pretreatment with Fer-1 for 12 h. Administration of Fer-1 alone did not significantly impact cell viability (Fig. 5D).

Fig. 5.

Fig. 5

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Effect of erastin on intracellular iron and lipid peroxidation in CRC cells. (A) Representative flow cytometry plots of FerroOrange staining (left) and associated quantification (right) in HCT116 and HCT116-OxR cells after 48 h of treatment with erastin (1, 2, 4 µM) or oxaliplatin (Ox, 2 µM). Data are expressed as mean ± SD (n=3). ***p<0.001 versus control. (B) Representative flow cytometry plots of BODIPY 581/591 C11 staining (left) and associated quantification (right) in HCT116 and HCT116-OxR cells after 48 h of treatment with erastin (1, 2, 4 µM) or oxaliplatin (Ox, 2 µM). Data are presented as mean ± SD (n=3). *p<0.05, ***p<0.001 versus control. (C) Western blot analysis of ferroptosis-regulatory proteins SLC7A11, GPX4 and ACSL4. (D) Cell viability was measured by MTT assay following erastin treatment preceded by pre-treatment with ferrostatin-1 (Fer-1). Data are shown as mean ± SD (n=4). ***p<0.001 versus control, ###p<0.001 versus erastin alone.

Erastin leads to ROS generation in HCT116-OxR cells

Flow cytometry with CellROX Green was utilized to measure erastin-induced ROS production (Fig. 6A). ROS levels in HCT116 cells showed minimal change: 21.68% in the vehicle control versus 21.81%, 24.52%, and 29.40% following 1, 2, and 4 µM erastin treatment, respectively. However, a substantial elevation in ROS levels was observed in HCT116-OxR cells, increasing from 29.35% in the vehicle control to 59.43%, 79.85%, and 92.20% in response to 1, 2, and 4 µM erastin, respectively. Ox treatment raised ROS levels to 87.58% in HCT116 cells, whereas ROS in HCT116-OxR cells remained stable (31.61%), as anticipated. To investigate whether ROS production mediates erastin-induced cytotoxicity, CRC cells underwent a 3 h pretreatment with NAC prior to erastin exposure (Fig. 6B). The MTT assay indicated that the reduction in cell viability to 40.96% and 30.66% in HCT116 and HCT116-OxR cells, respectively, after erastin administration, was reversed to 72.90% and 84.38% following NAC pretreatment. NAC alone slightly decreased cell viability. Furthermore, western blot analysis demonstrated that NAC pretreatment restored the expression levels of SLC7A11, GPX4, and ACSL4 (Fig. 6C).

Fig. 6.

Fig. 6

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Impact of erastin on ROS levels in CRC cells. (A) Representative flow cytometry plots of CellROX Green staining (left) and respective quantification (right) in HCT116 and HCT116-OxR cells following 48 h treatment with erastin (1, 2, 4 µM) or Ox (2 µM). Results are presented as mean ± SD (n=3). *p<0.05, ***p<0.001 versus control. (B) Cell viability was determined using the MTT assay after treatment with erastin and pre-treatment with NAC. Results are shown as mean ± SD (n=4). **p<0.01, ***p<0.001 versus control, ###p<0.001 versus erastin alone. (C) Western blot analysis of ferroptosis-regulatory proteins SLC7A11, GPX4 and ACSL4 was performed under the same experimental conditions as described in (B).

DISCUSSION

In this study, we aimed to utilize the distinct susceptibility of drug-resistant cancer cells to devise approaches for managing cancers prone to therapy resistance. The Ox-resistant variant of HCT116 CRC cells displays significantly higher expression of the proteins SLC7A11, GPX4, and ACSL4 (Fig. 1A-1D). Both SLC7A11 and GPX4 are involved in maintaining cellular glutathione levels, and a decrease in their expression can result in enhanced lipid peroxidation (Feng et al., 2021). In contrast, ACSL4 has an established role in facilitating ferroptosis (Doll et al., 2017). The observed upregulation of these proteins most likely acts as a protective adaptation against elevated ROS levels commonly linked with drug resistance. Notably, this protein upregulation was paralleled by an increase in basal ROS in HCT116-OxR cells, with levels approximately 2.49-fold higher than those in the parental HCT116 strain (Fig. 1E). In addition, basal lipid peroxidation, measured via BODIPY-C11 staining, was 2.76-fold greater in HCT116-OxR cells, in line with the increased ROS (Fig.1F). This distinction may explain the heightened vulnerability of HCT116-OxR cells to erastin-induced cytotoxicity in comparison to HCT116 cells (Fig. 2). This increased sensitivity was clearly demonstrated using the MTT cell viability assay. The IC50 for erastin following 48 h exposure in HCT116 (14.57 μM) was roughly 5.11-fold greater than in HCT116-OxR cells (2.85 μM). Therefore, all subsequent experiments used ≤16 µM erastin for HCT116 cells and ≤4 µM erastin for HCT116-OxR cells to induce cell death. Importantly, this range of erastin concentrations used on both cell lines was non-toxic to HEKa human keratinocytes, supporting the conclusion that erastin exerts a cancer cell-specific cytotoxic effect. Lastly, the respective sensitivities of HCT116 and HCT116-OxR cells to Ox were further validated independently through MTT viability and soft-agar assays.

The progression of the cell cycle is frequently altered by the administration of anticancer therapeutics (Shapiro and Harper, 1999). Analysis of the cell cycle using PI staining demonstrated a modest increase in the proportion of cells in the G2/M phase (Fig. 3). Supporting this observation, western blot analysis revealed that erastin resulted in the accumulation of p21, a CDK inhibitor (Besson et al., 2008), while concurrently decreasing the expression of cyclin B1 and Cdc2 (Dash and El-Deiry, 2005). However, the degree of cell cycle dysregulation induced by erastin likely does not play a major role in its cytotoxic effect. Nonetheless, these results demonstrate that erastin impacts cell cycle control in CRC cells.

The mitochondrial membrane integrity and the associated MMP are essential for cell viability (Marchi et al., 2012). The influence of erastin on MMP was evaluated using JC-1 staining. The data showed a significant rise in the proportion of HCT116-OxR cells exhibiting depolarized mitochondrial membranes following exposure to erastin at 1, 2, and 4 µM (Fig. 4A). In contrast, parental HCT116 cells did not display an increased proportion of depolarized mitochondrial membranes under the same concentrations of erastin. Furthermore, erastin-induced mitochondrial dysfunction in HCT116-OxR cells did not promote apoptosis. Annexin V/PI double staining demonstrated that neither HCT116 nor HCT116-OxR cells underwent apoptosis after erastin exposure. This observation differs from the response of HCT116 cells to Ox, which reliably induces apoptosis (Fig. 4B). Collectively, these findings suggest that erastin generates a non-apoptotic mitochondrial dysfunction in CRC cells.

Increases in intracellular iron levels and lipid peroxidation represent critical hallmarks of ferroptosis (Dixon et al., 2012). Initially, intracellular iron was evaluated using FerroOrange staining. A significant elevation in intracellular iron was detected in HCT116-OxR cells treated with erastin (1, 2, and 4 µM), but not in parental HCT116 cells (Fig. 5A). Next, lipid peroxidation was measured by BODIPY-C11 staining. A marked increase in lipid peroxidation was noted in HCT116-OxR cells following exposure to erastin at 1, 2, and 4 µM, while it remained unchanged in parental HCT116 cells (Fig. 5B). Western blot findings showed decreased levels of SLC7A11 and GPX4, and increased ACSL4 in HCT116-OxR cells (Fig. 5C). These molecular changes are in agreement with the observed rise in lipid peroxidation. Pretreatment with Fer-1, a ferroptosis inhibitor, abrogated erastin-induced cell death (Fig. 5D). This elevation corresponded with a parallel increase in cellular ROS, determined through CellROX Green staining (Fig. 6A). Importantly, NAC pretreatment (4 mM) markedly attenuated erastin-induced cell death (Fig. 6B), indicating that ROS generation mediated the cytotoxicity induced by erastin. Furthermore, western blot analysis revealed the mechanistic basis for this rescue: erastin treatment decreased SLC7A11 and GPX4 levels while increasing ACSL4 levels—changes that were reversed by NAC pretreatment (Fig. 6C). These results suggest that the initial accumulation of ROS upon erastin treatment drove these protein expression changes, whereas NAC pretreatment prevented ROS accumulation. Collectively, these findings indicate that erastin-induced cell death in HCT116-OxR cells is mediated through ferroptosis.

In summary, we show that HCT116-OxR cells exhibit elevated basal expression of ferroptosis-related proteins and ROS, rendering them more sensitive to erastin-induced cytotoxicity than the parental HCT116 cells. Erastin triggers G2/M cell cycle arrest, mitochondrial dysfunction of a non-apoptotic phenotype, increased intracellular iron, enhanced lipid peroxidation, ROS accumulation, and ultimately ferroptosis (Fig. 7). The heightened vulnerability of Ox-resistant HCT116-OxR cells to ferroptosis induction offers a potential therapeutic strategy for targeting drug-resistant cancer cells. These findings merit further investigation into the use of this approach as a novel therapeutic strategy.

Fig. 7.

Fig. 7

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Schematic diagram of erastin-induced ferroptosis in HCT116-OxR cells. Erastin induces ferroptosis in HCT116-OxR cells by inhibiting system Xc⁻, leading to cystine/GSH depletion and impaired GPX4 activity. In addition, the accumulation of intracellular iron (Fe²⁺) and ACSL4 expression trigger ROS accumulation and lipid peroxidation, leading to cell death. These effects are mitigated by NAC and Fer-1, which suppresses ROS production and lipid peroxidation, respectively. The model is presented as a simplified representation of the principal mechanisms identified in this study. System Xc⁻, cystine/glutamate antiporter; SLC3A2, solute carrier family 3 member 2; SLC7A11, solute carrier family 7 member 11; NAC, N-Acetyl-L-cysteine; ROS, reactive oxygen species; GSH, reduced glutathione; GSSG, oxidized glutathione; ACSL4, acyl-CoA synthetase long-chain family member 4; GPX4, glutathione peroxidase 4; PUFA, polyunsaturated fatty acid; PL-PUFA, phospholipid-containing polyunsaturated fatty acid; Fer-1, ferrostatin-1.

ACKNOWLEDGMENTS

This work was supported by the National Research Foundation of Korea (NRF) grants funded by the Korea government (MSIT) (RS-2022-NR070862 and RS-2024-00336900).

Footnotes

CONFLICT OF INTEREST

The authors confirm that there are no conflicts of interest related to this study.

AUTHOR CONTRIBUTIONS

Na Yeong Lee: Conceptualization, Investigation, Methodology, and Writing – original draft & review & editing. Sang Hoon Joo: Methodology, Validation, and Writing – review & editing. Si Yeong Seo: Methodology, Investigation, and Writing – review & editing. Seung-On Lee: Validation and Writing – review & editing. Goo Yoon: Resources, Validation, and Writing – review & editing. Jin Woo Park: Resources, Supervision, and Project administration. Jung-Hyun Shim: Data Curation, Writing – review & editing, Supervision, Project administration, and Funding acquisition.

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