MDA-MB-231, T47D, BT549 and MCF7 human breast cancer cells were purchased from the American Type Culture Collection (ATCC) and grown in medium supplemented with 10% heat-inactivated fetal bovine serum (HI-FBS), 100 µg/ml streptomycin, and 100 units/ml penicillin (Invitrogen; Thermo Fisher Scientific, Inc.) in a humidified incubator with 5% CO₂ at 37°C.

The intracellular ROS level in the form of cellular peroxides was assessed using a Reactive Oxygen Species Assay Kit (Beyotime Institute of Biotechnology) after treatment with or without SAS (Abcam) and liproxstatin-1 (MedChemExpress). In the liproxstatin-1-treatment group, the cells were treated with 300 nM liproxstatin-1 for 24 h before the addition of SAS. The cells were collected, exposed to 10 µM 2′,7′-dichlorofluorescein diacetate (DCFH-DA), and incubated at 37°C for 20 min. The cells were washed three times with PBS, and fluorescence intensity was analyzed by flow cytometry (excitation at 488 nm; emission at 525 nm). ROS levels were expressed as the mean fluorescence intensity of 20,000 cells. Three independent experiments were performed.

Total RNA from MDA-MB-231 and T47D cells was extracted with a total RNA extraction kit and reverse-transcribed using the PrimeScript RT reagent kit (Promega Corporation). Quantitative RT-PCR was performed using SYBR Premix Ex Taq™ II (Takara Bio, Inc.) in a 10-µl PCR mixture on a Bio-Rad CFX96 Real-Time PCR system (Bio-Rad Laboratories, Inc.) according to the manufacturer's standard protocols. An initial cycling for 2 min at 95°C, followed by 39 cycles at 95°C for 30 sec, 30 sec at 58°C and 20 sec at 72°C. The primer sequences were as follows: Divalent metaltransporter 1 (DMT1) forward, 5′-AGCTCCACCATGACAGGAACCT-3′ and reverse, 5′-TGGCAATAGAGCGAGTCAGAACC-3′; TFRC forward, 5′-ATCGGTTGGTGCCACTGAATGG-3′ and reverse, 5′-ACAACAGTGGGCTGGCAGAAAC-3′; glyceraldehyde 3-phosphate dehydrogenase (GAPDH) forward, 5′-TGACTTCAACAGCGACACCCA-3′ and reverse, 5′-CACCCTGTTGCTGTAGCCAAA-3′; and estrogen receptor 1 (ESR1) forward, 5′-GGTCAGTGCCTTGTTGGATG-3′ and reverse, 5′-CAGGTTGGTGAGTAAGC-3′. Three independent experiments were performed for each group. Relative gene expression was normalized to GAPDH and calculated using the 2^−ΔΔCq method (13).

Cell growth and SAS-mediated inhibition were detected using the Cell Counting Kit-8 (CCK-8; Beyotime Institute of Biotechnology). According to the manufacturer's instructions, cells were plated at a density of 2.5×10⁴/well in 96-well plates with or without different concentrations of SAS, and cell viability was assessed at 24 h. The absorbance of each well was measured at a wavelength of 450 nm using a Synergy H1 microplate reader (BioTek Instruments, Inc.). Empty wells served as blank controls. The test was performed three times under the same operating conditions. The cells were treated with SAS at concentrations of 0.1, 0.5, 1.0, 1.5, 2.0, and 3.0 mM, and cell viability was determined using the CCK-8 assay at 24 h. The half maximal inhibitory concentration (IC₅₀) of SAS for different cells was calculated according to the standard curve.

Cell were washed twice with cold PBS and harvested. Protein lysates were prepared from MDA-MB-231 and T47D cells using a protein extraction kit (Nanjing KeyGen Biotech. Co. Ltd.). Protein concentrations were assessed using a BCA Protein Assay kit (Beyotime Institute of Biotechnology). Equal amounts of proteins (40 µg/lane) from each group were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis on 12.5% gels (Bio-Rad Laboratories, Inc.) and transferred to polyvinylidene difluoride membranes (EMD Millipore). After blocking with 5% skim milk for 1 h at room temperature, the membranes were incubated with specific primary antibodies at 4°C overnight. Then, after washing three times with Tris-buffered saline containing Tween-20, the membranes were incubated with horseradish peroxidase-conjugated secondary antibodies for 1 h at room temperature. Proteins were visualized by chemiluminescence using enhanced chemiluminescent substrate (Beyotime Institute of Biotechnology). Immunoreactive bands were examined using the ChemiDoc Imaging System (Bio-Rad Laboratories, Inc.). The following antibodies were used: Rabbit anti-xCT antibody (cat. no. 12691S; dilution 1:800; Cell Signaling Technology, Inc.); rabbit anti-glutathione peroxidase 4 (GPX4) antibody (cat. no. SAB2108670; dilution 1:1,000; Sigma-Aldrich; Merck KGaA); rabbit anti-ESR1 (cat. no. 13258; dilution 1:1,000) and rabbit anti-TFRC (cat. no. 13113; dilution 1:1,000; both from Cell Signaling Technology, Inc.); and rabbit anti-GAPDH (cat. no. ATGA0181; dilution 1:1,000; 4A Biotech Co., Ltd.). Horseradish peroxidase-conjugated goat anti-rabbit antibodies were used as secondary antibodies (cat. no. 7074S; dilution 1:1,000; Cell Signaling Technology, Inc.). GAPDH was used as an internal control for each membrane.

After the indicated treatment, cells on a 100-mm dish were fixed in 2.5% glutaraldehyde in 0.1 M PBS (pH 7.4) at room temperature for 1 h, post-fixed in 1% OsO₄ in 0.1 M PBS (pH 7.4) at room temperature for 1 h, dehydrated using a graded series of ethanol and embedded with epoxy resin and sectioned. Ultrathin (60-nm) sections were collected on grids and stained with uranyl acetate and lead citrate. Images were obtained using an H-7100 transmission electron microscope (Hitachi, Ltd.).

Cells were plated in six-well plates and transfected with specific siRNAs using Lipofectamine^™ 2000 transfection reagent (Invitrogen; Thermo Fisher Scientific, Inc.) according to the manufacturer's instructions. Two siRNA sequences were used to ensure the accuracy of the experiment. The sequences of siRNA were as follows: ESR1-siRNA1 (sense 5′-GGAGGAUGUUGAAACACAATT-3′, antisense 5′-UUGUGUUUCAACAUUCUCCTT-3′) and ESR1-siRNA2 (sense 5′-GGAUUUGACCCUCCAUGAUTT-3′, antisense 5′-AUCAUGGAGGGUCAAAUCCTT-3′). All siRNA oligomers were purchased from Shanghai GenePharma Co., Ltd.

The human tissues were fixed with 4% formaldehyde buffer, the fixed time of the specimens was 12–24 h at room temperature 15–28°C. Deparaffinized specimens were then sectioned at 4-µm-thick slices. The slices were autoclaved at 115°C for 5 min for antigen retrieval in citric acid buffer (pH 6.0) and quenched for endogenous peroxidase activity with 0.3% H₂O₂ solution for 10–15 min. Then, the samples were blocked for nonspecific binding with normal goat serum for 10–15 min and incubated with the specific rabbit primary antibody against TFRC (cat. no. PB9233; dilution 1:700; Boster Biological Technology Co., Ltd.) overnight at 4°C. Subsequently, the sections were treated with the goat anti-rabbit antibody for 30 min at room temperature. After staining with diaminobenzidine (DAB) (cat. no. PV-9000; Beijing Zhongshan Golden Bridge Biotechnology Co., Ltd.; OriGene Technologies), images were captured using a Nikon Eclipse 80i microscope (Nikon Corporation).

The mitochondrial membrane potential (MMP) was assayed using tetramethylrhodamine methyl ester (TMRM) according to the manufacturer's instructions (AAT Bioquest, Inc.). TMRM is readily sequestered by healthy mitochondria, but its fluorescence is rapidly lost when the MMP dissipates. Cells were seeded in confocal dishes before treatment with or without SAS. The cells were washed twice with PBS and incubated for 20 min at 37°C with 100 nM TMRM for staining mitochondria and then carefully washed three times with PBS. To ensure the accuracy of MMP detection, cell activity should be maintained when staining for TMRM; the cells should not be killed by reagents such as formaldehyde. Finally, the cells were analyzed by confocal fluorescence microscopy (Nikon N-SIM; Nikon Corporation) to assess the intensity of red fluorescence (excitation at 549 nm; emission at 573 nm).

In the present study, clinical tissue samples were collected from 87 patients with breast cancer, and RNA was extracted. The human breast cancer specimens used in this study were collected in the operating room of the Department of Endocrine and Breast Surgery, First Affiliated Hospital of Chongqing Medical University, from July 2015 to June 2017. All clinical specimens were diagnosed by clinical and pathological examinations, and all patient information data were true and valid. The First Affiliated Hospital of Chongqing Medical University Ethics Committee reviewed and approved the use of human tissue specimens and all patients provided written informed consent. Additional information for these patients is provided in Table I. RNA from the tissue samples from 87 patients was extracted with a total RNA extraction kit and reverse-transcribed using the PrimeScript RT reagent kit (Promega Corp.).

Data was downloaded from The Cancer Genome Atlas (TCGA) database (https://cancergenome.nih.gov) for 1,208 breast cancer samples. TFRC expression was analyzed in different subtypes of breast cancer cells and the correlation between TFRC and ESR1 with ggstatspot (a package in R, version 3.5.1). TFRC expression in cancer tissues and normal tissues was determined using Gene Expression Profiling Interactive Analysis (GEPIA; http://gepia.cancer-pku.cn/index.html).

All experiments were independently performed at least three times. The mean ± SD was determined for each group. Statistical analyses were performed using one-way analysis of variance (ANOVA) for multiple group comparisons or Student's t-test for individual comparisons. Multiple comparisons between the groups were performed using the Student-Newman-Keuls method. Statistical significance was considered at P<0.05.

The effects of SAS on breast cancer were investigated. Different human breast cancer cell lines (MDA-MB-231 and T47D, representing TNBC and ER⁺ breast cancer, respectively) were used. Previous studies have reported that higher levels of SAS can reduce glioma cell growth (14). Both breast cancer cell lines were treated with two different concentrations of SAS for 24 h. Microscopic observation revealed that cell growth and viability were significantly inhibited. The two cell lines displayed a significant reduction in cell viability in response to 1.0 and 2.0 mM SAS (Fig. S1). To better understand the effect of SAS in breast cancer cells, the inhibition rate for MDA-MB-231 cells was determined by treating the cells with 1.0 or 2.0 mM SAS at 3, 6, 12 and 24 h. In fact, 2.0 mM SAS induced death in >70% of the breast cancer cells in a time-dependent manner, peaking at 24 h. With an increase in SAS treatment time, the cell growth inhibition rate also increased, and the inhibition rate in the 2.0 mM group was higher than that in the 1.0 mM group (Fig. 1A). Similar results were obtained in T47D cells (Fig. 1B). To confirm the effects of SAS in breast cancer, two other breast cancer lines (BT549 and MCF7, representing TNBC and ER⁺ breast cancer, respectively) were included. The four breast cancer cell lines were treated with different concentrations of SAS for 24 h. The inhibition rates were dependent on the SAS concentration (Fig. 1C). From the trend indicated in the figure, ER⁺ breast cancer cells (T47D and MCF7) were less sensitive to SAS than TNBC cells (MDA-MB-231 and BT549) at the same concentrations. To confirm our hypothesis, the half maximal inhibitory concentration (IC₅₀) of the four cell lines for SAS, was examined. The results revealed that the IC₅₀ values of MDA-MB-231 and BT549 cells were significantly lower than those of T47D and MCF7 cells (Fig. 1D). Therefore, SAS reduced the growth of breast cancer cells; however, the sensitivity varied for different breast cancer cells.

It was demonstrated that SAS could reduce cell growth, however, whether this effect was due to ferroptosis required more evidence. A large and growing body of literature has demonstrated that xCT and GPX4 play important roles in ferroptosis (15). Previous studies have indicated that inhibition of xCT triggers ferroptosis and that GPX4 plays a role in preventing ferroptosis (5). Traditionally, western blotting has been used to assess the expression of xCT and GPX4. A clear decreasing trend in xCT and GPX4 expression with an increasing concentration of SAS were observed. Similar results were obtained in MDA-MB-231 and T47D cells (Fig. 2A). The accumulation of ROS in cells is a direct cause of ferroptosis (16). Flow cytometry is currently the most popular method for detecting ROS. MDA-MB-231 cells treated with 1.0 or 2.0 mM SAS displayed increased ROS production than untreated and negative control cells (Fig. 2B). Similar results were obtained in T47D cells (Fig. 2C). Detailed flow cytometric data is revealed in Fig. S1.

Many studies have revealed that iron is closely related to oxidative stress. If iron ions cannot bind to proteins or other ligands in an appropriate manner, they can catalyze the formation of metabolically toxic ROS through the H:O: Dependent Fenton reaction. Iron chelators (DFO) and ferroptosis-specific inhibitors (liproxstatin-1) have no significant effects on other forms of cell death (7). Therefore, these reagents can be used to confirm that the death mode caused by SAS is ferroptosis. Liproxstatin-1 prevents ROS accumulation and cell death in cells. Moreover, liproxstatin-1 inhibits ferroptosis induced by ferroptosis-inducing agents (FINs; a series of small molecule inducers, e.g., erastin) in vitro (17). Liproxstatin-1 has also been used to determine whether SAS-induced cell death is ferroptosis (18). The results, as revealed in Fig. 2D, indicated that SAS-induced ROS generation can be inhibited by liproxstatin-1 in MDA-MB-231 cells; similar results were obtained in T47D cells (Fig. 2E).

Change in mitochondrial morphology is also a characteristic of ferroptosis (19). Transmission electron microscopy revealed that compared with untreated cells (Fig. 3A), T47D cells treated with 1.0 mM SAS for 24 h had shrunken mitochondria with increased membrane density (Fig. 3B). The white arrow in the figure indicates the mitochondria of T47D cells. This phenomenon was more pronounced in cells treated with 2.0 mM SAS for 24 h (Fig. 3C). Notably, numerous autophagosomes were found in cells treated with 2.0 mM SAS (Fig. 3D), indicating that SAS may induce autophagy. Literature studies have indicated that ferroptosis may be related to autophagy (20). However, further research is required to demonstrate this hypothesis. The same positive result was not obtained in MDA-MB-231 cells. Next, MMP was examined using the probe TMRM, which accumulates in the mitochondria and produces bright red fluorescence in living cells. Confocal microscopy analysis of red fluorescence intensity revealed that compared with untreated cells, SAS-treated MDA-MB-231 cells displayed a reduction in red fluorescence; similar results were obtained for T47D cells (Fig. 3E). These data indicated that SAS induced mitochondrial depolarization, consistent with the morphological characteristics of ferroptosis observed by transmission electron microscopy. In summary, the mode of death caused by SAS was in fact ferroptosis.

Iron metabolism and lipid peroxidation signaling are increasingly being recognized as central mediators of ferroptosis (21). Fe³⁺ is imported into cells through the membrane protein TFRC and then sequestered in endosomes. In the endosome, Fe³⁺ is reduced to ferrous iron (Fe²⁺). DMT1, also known as natural resistance-associated macrophage protein 2, belongs to the soluble carrier family member (solute carrier family 11 member 2, SLC11A2) and is a proton-dependent metal ion transporter. DMT1 can mediate transmembrane transport, including Cu²⁺, Fe²⁺, Zn²⁺, and Mn²⁺ ions and is considered to be a transporter of various divalent metal ions in mammalian cells. When the concentration of metal ions in the body changes, DMT1 expression may change accordingly, and it plays an important physiological role in the metabolism and balance of divalent metal ions in cells. The literature indicates that DMT1 mediates the release of Fe²⁺ from the endosome into a labile iron pool in the cytoplasm (22). Therefore, the accumulation of iron in cells is closely related to TFRC and DMT1. However, the role of SAS in iron metabolic pathways is unknown. To explore the function of SAS in iron metabolic pathways, RT-PCR was used to detect the expression of TFRC and DMT1 in cells treated with different concentrations of SAS. As revealed in Fig. 4A and B, as the SAS concentration increased, the expression of TFRC and DMT1 increased in MDA-MB-231 cells. Consistent with the results in MDA-MB-231 cells, TFRC and DMT1 expression also increased in T47D cells in response to increasing SAS concentrations (Fig. 4C and D), indicating that SAS may activate TFRC and DMT1 to induce ferroptosis.

TFRC is a key factor in iron metabolism, and after determining the role of TFRC in ferroptosis in breast cancer cells, its expression in breast cancer tissues was also investigated. First, TFRC expression was determined in cancer tissues and normal tissues using GEPIA. As revealed in Fig. 5A, TFRC expression in tumor tissues was significantly higher than that in normal tissues. TFRC expression was detected in different breast cancer tissues using immunohistochemistry. Next, bioinformatics was used to analyze TFRC expression in breast cancer subtypes. As revealed in Fig. 5B, TFRC expression in basal (TNBC usually has a basal molecular phenotype) and Her2 subtypes was higher than that in luminal and normal subtypes. The expression of ER is negative in TNBC and Her2⁺ breast cancer. Therefore, the relationship between ER and TFRC was determined. Bioinformatics was used again to analyze the correlation between TFRC and ER. Notably, a negative correlation between TFRC and ER (Fig. 5C) was revealed. This result may explain why T47D cells are less sensitive than MDA-MB-231 cells to SAS-induced ferroptosis (Fig. 1D). To obtain more convincing results, clinical samples of different tissue types were selected for IHC. The results revealed that TFRC expression was significantly higher in breast cancer tissues of basal and Her2⁺ cells than in luminal type cancer tissues. This finding was consistent with the results of our previous bioinformatics analysis (Fig. 5D). Similar results were obtained in different breast cancer cells. TFRC expression was lower in ER⁺ breast cancer cell lines (T47D and MCF7) than in TNBC cells (BT549 and MB231) (Fig. 5E). Due to the different TFRC expression levels, the sensitivity to SAS-induced ferroptosis varies. The difference in TFRC expression may be due to differences in ER expression in different breast cancer cells. To verify our speculation, siRNAs were used to knockdown ESR1 (the gene encodes the ER) in T47D cells. As revealed in Fig. 5F and H, the expression of ER was successfully knocked down at both the RNA and protein levels. In response to ER knockdown, an increase in TFRC expression (Fig. 5G and H) was observed. These results demonstrated that ER may inhibit TFRC. As revealed in Fig. 1C, MDA-MB-231 and T47D cells had the greatest difference in sensitivity to SAS at concentrations of 0.1 and 0.5 mM. Therefore, these two concentrations were selected for the following experiments. Low concentrations of SAS (0.1 and 0.5 mM) were used to treat ER-knockdown T47D cells. Subsequently, CCK-8 assays were used to assess the SAS-mediated inhibition of T47D cells. The inhibition rate was significantly increased after knocking down ER (Fig. 5I). Collectively, our findings indicated that ER has a protective effect on SAS-induced ferroptosis, and the inhibition of TFRC expression may be one of the underlying mechanisms. To determine TFRC expression in breast cancer tissues, 87 clinical tissue samples from patients with breast cancer were collected and total RNA was extracted. RT-PCR was used to detect TFRC expression in these different breast cancer tissues. Concurrently, to identify new relationships between TFRC and clinical characteristics, the clinical data was analyzed. As revealed in Fig. 6A, TFRC was expressed at lower levels in ER⁺ breast cancer tissues than in ER⁻ breast cancer tissues, indicating that ER may inhibit TFRC. This result was consistent with our findings in the in vitro cell experiments. TFRC expression in tissues representing different breast cancer subtypes was also consistent with TCGA database analysis results. Compared with other subtypes, the Her2 subtype had the highest expression of TFRC (Fig. 6B). Indicative of the relationship between the Her2 gene and TFRC, a high expression of TFRC in Her2⁺ breast cancer tissues (Fig. 6C) was observed. Estrogen is a messenger that must bind to cellular receptors to function. Once combined with estrogen, the ER is activated and transported into the nucleus. After gene transcription leads to PR synthesis, PR synthesis must be achieved by the action of ER synthesis. PR synthesis occurs in conjunction with ER synthesis. Therefore, if the PR is expressed, the ER must also be expressed. Therefore, the expression patterns of the two receptors are similar. The expression of TFRC in PR⁻ breast cancer tissues was higher than that in PR⁺ breast cancer tissues (Fig. 6D). The relationship between TFRC expression and TNM stage, histological grade and Ki-67 expression was also explored, however, no significant associations were revealed (Fig. S1).