The antibodies used in this study included NRF2 (16396-1-AP), Histone H3 (17168-1-AP), SLC7A11 (26864-1-AP), HMOX-1 (10701-1-AP), and GAPDH (16396-1-AP) (ProteinTech Group, IL, United States). 4-Hydroperoxy cyclophosphamide (4HC) was purchased from TCR (39800-16-3, Santa Cruz Biotechnology, TA, United States). Cyclophosphamide (CTX), Hemin, zinc protoporphyrin (ZNPP), ferrostatin-1 (Fer1), and deferoxamine mesylate salt (DFO) were purchased from Sigma–Aldrich (Sigma–Aldrich, MO, United States).

The murine glioblastoma cell lines (GL261, CT-2A, and KR-158) and murine breast cancer cell line (4T1) were kindly provided by Cell Bank, Chinese Academy of Sciences (Shanghai, China). Cells were maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin (Gibco, Massachusetts, United States) in a humidified incubator at 37 °C with 5% CO₂. Cells grown on 60 mm culture dishes (Corning, New York, NY, United States) were stimulated with or without different concentrations of 4HC (from 5 to 160 μM)for various times (from 0.5 to 48 h). Fer1 or DFO were added 1 h before 4HC supplementation. After treatment, cells were collected and used for subsequent experiments.

RNA sequencing was performed by LC Sciences (Hangzhou, China). Briefly, total RNA was extracted from samples using TRIzol (Thermo Fisher Scientific, MA, United States) according to the manufacturer’s protocol. High-quality RNA was used to construct sequencing libraries and analyze the enrichment of differentially expressed genes (DEGs). The number of gene counts in each sample was normalized to the base mean value, and the p value and fold change (FC) of the difference in each comparison were calculated. The DEGs were screened based on p < 0.05 and FC > 2. Kyoto Encyclopedia of Genes and Genomes (KEGG) was used to perform the pathway enrichment analysis of the differentially expressed genes.

The CCK-8 assay was performed using a Cell Counting Kit-8 (CCK-8) (Bimake, TX, United States) according to the manufacturer’s protocol to assess cell proliferation. Cells were seeded in 96-well plates at a density of 5,000 cells per well. Cells were treated with a series of 4HC concentrations (from 0 to 160 μM) for various times (from 0 to 48 h) in the assay. Then, 100 μL of CCK-8 solution were added to each well and incubated at 37 °C for 1 h. Next, the absorbance of each well was measured at 450 nm.

The amount of LDH released in cell culture supernatants was measured using the LDH Cytotoxicity Assay Kit (Beyotime Biotech, Nanjing, China) according to the manufacturer’s instructions. A total of 5,000 cells/well were treated with 4HC as described above in medium containing 1% FBS (Gibco, MA, United States). Sixty microliters of the reaction solution were added to each well and incubated in the dark at room temperature for 30 min. Then, the amount of LDH released was measured by detecting the absorbance at a wavelength of 490 nm, and cytotoxicity was calculated using the following equation: cytotoxicity (%) = (LDH released in the sample–background)/(total LDH release - background)  × 100.

GSH levels were measured using a GSH assay kit (Beyotime Biotech, Nanjing, China) according to the manufacturer’s instructions. Cells or lysed tissue were incubated with Protein Removal Buffer. The GSH concentration was measured in the supernatant collected from each well by detecting the absorbance at 412 nm. A standard curve was generated and used to determine GSH levels in the specimens.

Cells or lysed tissue were washed twice and digested with a 3:1 ratio of trace element grade nitric acid:HCl for 20 min at 200 °C. Iron concentrations were analyzed using ICP–MS (inductively coupled plasma–mass spectrometry, PerkinElmer, MA, United States) according to the manufacturer’s instructions.

Intracellular ROS levels were measured with the fluorescent probe 2′,7′-dichlorofluorescin diacetate (DCFH) (Solarbio, Beijing, China). As a cell-permeable nonfluorescent probe, DCFH is deesterified intracellularly and transformed into the highly fluorescent molecule 2′,7′-dichlorofluorescin (DCF) in the presence of intracellular ROS. Cells were incubated with 5 μM DCFH for 60 min at 37 °C. After the incubation, cells were observed using a Leica microscope system (Leica, Wetzlar, Germany) at an excitation wavelength of 488 nm and an emission wavelength of 525 nm or directly collected before the immediate detection of the fluorescence intensity of DCF using flow cytometry (BD Biosciences, CA, United States).

The cells were digested and resuspended in PBS, and the fluorescence intensity was measured using flow cytometry (BD Biosciences, CA, United States). The MFI (mean fluorescence intensity) was calculated and displayed as a histogram using the FlowJo program (Tree Star, OR, United States).

The mitochondrial membrane potential was assayed using JC-1 according to the manufacturer’s instructions (Beyotime Biotech, Nanjing, China). Briefly, cells were cultured, treated with staining buffer for JC-1 at 37 °C for 20 min, and monitored using flow cytometry (BD Biosciences, CA, United States). The excitation wavelength of JC-1 is 488 nm, and the approximate emission wavelengths of the monomeric and J-aggregate forms are 529 and 590 nm, respectively.

Total RNA was extracted from cultured cells or tissues with TRIzol (Thermo Fisher Scientific, MA, United States) according to the manufacturer’s instructions. After quantifying the concentrations and purities of RNA, 1 μg of RNA was reverse transcribed into cDNAs with an RT reagent Kit with gDNA Eraser (Takara, Dalian, China). Subsequently, cDNA templates from the samples were amplified with TB Green (Takara, Dalian, China). Quantitative PCR was performed using a C1000 Touch Thermocycler CFX96 Real-Time System (Bio-Rad, CA, United States). The relative expression of the target genes was standardized and determined using the 2^−ΔΔ CT method. All primers used for RT–qPCR are listed as follows:

18s-F: CGC​CGC​TAG​AGG​TGA​AAT​TC

Cells were washed twice, fixed with 1% glutaraldehyde and postfixed with 2% osmium tetroxide. Following dehydration with a graded series of ethanol solutions, the samples were embedded in epon resin and cut into ultrathin sections using a microtome. The sections were then counterstained with saturated uranyl acetate and lead citrate before being examined with a transmission electron microscope (HITACHI, Japan). The morphological characteristics of mitochondria were observed in randomly selected cytoplasmic fields of view from each group.

After treatment, cells were washed with PBS, scraped for collection, gently suspended in extraction buffer with PMSF and incubated on ice for 10 min. The cell suspension was vigorously vortexed for 5 s to lyse the cells and release the cytoplasmic contents. The cellular mixtures were then centrifuged at 12,000-16,000 g for 5 min at 4 °C to separate the cytoplasmic components (supernatant) from the nuclear fraction (pellet). For the purification of nuclear proteins, the nuclear pellets were washed with hypotonic buffer three times and suspended in PBS containing proteinase inhibitors and phosphatase inhibitors. The suspension was further sonicated and centrifuged at 12,000 g for 10 min at 4 °C. The isolated fractions were stored at −80 °C until further analysis. The concentration of the protein lysate was measured using a BCA protein assay reagent kit (Thermo Pierce, MA, United States) and used for western blotting. The following primary antibodies were used at the indicated dilutions: 1:1,000 for NRF2, Histone H3, SLC7A11, HMOX-1, and GAPDH.

The siRNAs targeting Hmox-1 were synthesized by EgyptBio (Egypt Biotechnology, Guangzhou, China). The shRNA segments for Nrf2 were donated by Professor Xuesong Yang from the Department of Histology Embryology, Jinan University. After reaching 80–90% confluence, cells were transfected with siRNAs or the specific shRNA using Lipofectamine 2000 Transfection Reagent (Thermo Fisher, MA, United States) in Opti-MEM (Thermo Fisher, MA, United States) according to the manufacturer’s instructions. After 6–8 h of incubation at 37 °C, the medium was replaced with complete medium. Cells were prepared for tests and harvested after 48 h. The sequences of siRNAs/shRNAs used for RNA interference are listed as follows:

Hmox-1-F: GGG​UCA​GGU​GUC​CAG​AGA​A

Cells on glass slides were fixed with 4% paraformaldehyde for 15 min at room temperature. After two rinses with PBS, cells were permeabilized with 0.1% Triton (Sinopharm, Shanghai, China) for 5 min, incubated with a primary antibody against NRF2 at 4 °C overnight and secondary antibody at room temperature for 1 h, and counterstained with DAPI. Images were captured using a Leica microscope system (Leica, Wetzlar, Germany).

Six-week-old female Balb/c mice were purchased from Huafukang Co. (Hfkbio, Beijing, China) and maintained under specific pathogen-free (SPF) conditions. The animal experiment was performed under the supervision of the Jinan University Animal Ethics Committee in accordance with the Animal Research: Reporting of In Vivo Experiments (ARRIVE) guidelines (approval no. 20180604–02). A total of 1 × 10⁶ 4T1 cells was injected into the mammary fat pad of each mouse. Once the tumor size reached ∼200 mm³, mice were randomly allocated to six different groups as follows (6-7 mice per group): 1) untreated (UT) group (PBS); 2) Fer1 treatment group (4 mg/kg); 3) Hemin treatment group; 4) CTX treatment group; 5) CTX + Fer1 treatment group; and 6) CTX + Hemin treatment group. CTX (140 mg/kg, i. p.) was administered every 6 days (Days 0, 6, 12 and 18) after Fer1 (4 mg/kg, i. p.), Hemin (20 mg/kg, i. p.) or the equivalent vehicle was administered 1 day in advance.

During the treatment period, the tumor size and body weights of the 4T1-bearing mice were monitored every other day. The tumor volume of each mouse was measured using the following equation: tumor volume (mm³) = 1/2 length (mm)  × width (mm)². Then, the mice were sacrificed by cervical dislocation on Days 12 and 18 after the first administration of CTX. Tumor masses were harvested for further investigations.

The experiments were all independently repeated at least three times. Statistical analysis were conducted using GraphPad Prism software (GraphPad, CA, United States). Significant differences between the groups were analyzed using one-way ANOVA, followed by Student-Newman-Keuls test or Student’s t-test; p < 0.05 was considered to indicate a statistically significant difference.

We first performed RNA sequencing in GL261 murine glioblastoma cells treated with 4-hydroxycyclophosphamide (4HC), the pharmacologically active moiety of CTX, to systematically investigate the biological processes underlying the therapeutic effect of CTX. Differentially expressed genes were identified based on a p value <0.05 and fold change >2. Among differentially expressed genes, Hmox-1 was one of the most significantly upregulated genes, and the cystine/glutamate antiporter solute carrier family 7 membrane 11 (Slc7a11) gene was slightly upregulated (Figure 1B). The KEGG analysis based on RNA-seq data also revealed enriched pathways related to ferroptosis, suggesting that iron homeostasis was among the pathways that were remarkably altered in 4HC-treated cells (Figure 1A). In addition, some other enriched pathways such as protein processing in endoplasmic reticulum or the mitogen-activated protein kinases (MAPKs) were also shown. Consistently, the real-time PCR analysis confirmed that 4HC treatment led to a significant increase in Hmox-1 and Slc7a11 mRNA levels at an early stage (within 6 h) in the murine glioblastoma cell lines GL261, CT-2A, and KR-158 and the 4T1 murine breast cancer (Figure 1C). Therefore, the ferroptosis pathway tended to be altered by CTX in tumor cells, which at least partially involved HMOX-1 regulation.

Based on the findings described above, we further confirmed the validity of ferroptosis as one of the main causes of CTX-induced cell death by applying various inhibitors of ferroptosis, such as ferrostatin-1 (Fer1) and deferoxamine (DFO). The viability of the GL261 and 4T1 cell lines treated with 4HC was significantly reduced in a dose- and time-dependent manner compared with that of the control groups, accompanied by increased cytotoxicity (Figures 2A,B). As shown in Figures 2C,D, 4HC treatment also altered the cell morphology under an inverted microscope, which was characterized by a reduced neurite-like shape and broadened intercellular gaps. Notably, 4HC-induced cell death and morphological changes were significantly attenuated in the presence of Fer1 or DFO (Figures 2C–E). Since ferroptosis depends on ROS production for its cytotoxicity (Dixon et al., 2012), we also observed prominent intracellular ROS accumulation in both GL261 and 4T1 cells (Figures 3A,B) after 4HC treatment, as visualized by the strong fluorescence intensity of the DCF probe. The quantitative flow cytometry analysis clearly showed the induction of oxidative stress in a concentration-dependent manner, which was almost completely offset by ferroptosis inhibitors (Figures 3C,D).

Furthermore, mitochondrial physiological function was evaluated by assessing the mitochondrial membrane potential (MMP) with JC-1 staining reagent in live cells. As shown in Figures 4A,B, 4HC reduced the MMP, as evidenced by a pronounced decrease in JC-1 aggregate formation (with specific red fluorescence), and more free green puncta indicating JC-1 monomers. However, pre-treatment with ferroptosis inhibitors (Fer1 or DFO) significantly restored the loss of the MMP induced by 4HC. Moreover, the quantitative flow cytometry analysis further indicated a concentration-dependent decrease in the MMP of 4HC-stimulated tumor cells (Figures 4C,D).

As one of the distinct morphological features of ferroptosis, mitochondria appear smaller than normal, with an increased membrane density and reduced cristae (Yagoda et al., 2007; Dixon et al., 2012). In our study, the transmission electron microscopy (TEM) analysis displayed similar mitochondrial deformation in 4HC-treated cells, which was blocked by Fer1 (Figures 5A,B).

Iron, a basic component within the mitochondrial respiratory chain, is required for ferroptosis (Dixon et al., 2012), (Gkouvatsos et al., 2012). We measured the intracellular iron concentrations and found that iron accumulated after treatment with 4HC and reached its highest level at the 2-h time point (Figure 5C). Glutathione (GSH) exhaustion is another critical event leading to excess oxidative stress during ferroptosis-mediated cell death (Gao et al., 2015; Yan et al., 2021); hence, we observed reduced GSH levels after 4HC treatment, and these changes were also inhibited by Fer1 or DFO (Figure 5D). All the results collectively demonstrate that ferroptosis is the relevant mechanism underlying the effects of CTX on inducing cell death.

HMOX-1 expression is regulated by nuclear factor E2 related factor 2 (NRF2), a main regulator of detoxifying/antioxidant phase II enzymes (Biswas et al., 2014; Chiang et al., 2018). Moreover, according to a previous study, increasing HMOX-1 expression increases the labile iron pool (LIP) and triggers ferroptosis (Chang et al., 2018). In this study, we logically hypothesized that the NRF2/HMOX-1 pathway plays a role in CTX-initiated ferroptosis and performed experiments to test our hypothesis. The expression of the Nrf2 mRNA was obviously upregulated after 4HC treatment for 1 h but returned to normal levels at 6 h (Figures 6A,B). Meanwhile, the nuclear translocation of NRF2 was also increased in the 4HC group, with stronger nuclear NRF2 expression observed using immunofluorescence staining (Figures 6B,C). To verify the up- and downstream regulatory mechanisms, cells were transfected with a small interfering RNA against Hmox-1 or short hairpin RNA against Nrf2. As expected, Nrf2 knockdown attenuated the levels of SLC7A11 and HMOX-1, but Hmox-1 silencing had no significant effect on the expression of NRF2 and SLC7A11 (Figure 6E), indicating that the NRF2/HMOX-1 signaling pathway was activated by the CTX treatment. Furthermore, we used a specific inhibitor or inducer of HMOX-1 to determine whether HMOX-1 is a key protein involved in CTX-induced ferroptosis. Compared with the 4HC group, treatment with the HMOX-1 inhibitor zinc protoporphyrin (ZNPP) noticeably reduced the cell viability and intracellular GSH level, as well as the MMP; in contrast, these ferroptosis-related features were more significantly increased in the HMOX-1 agonist (Hemin) group (Figures 6D,F). Therefore, HMOX-1 plays a pivotal role in promoting CTX-induced cancer cell ferroptosis.

Based on the in vitro findings described above, a murine xenograft model of 4T1 breast cancer was established to examine the antitumor mechanism of CTX in BALB/c mice in vivo. 4T1 tumor-bearing mice were intraperitoneally injected with CTX (140 mg/kg) with or without Fer1 (4 mg/kg), Hemin (25 mg/mg) or the control (PBS) every 6 days (Figure 7A). CTX treatment significantly inhibited tumor growth in mice (Figure 7B), with more iron accumulation and decreased GSH level in tumor tissues (Figure 7C). Co-treatment with Fer1 abrogated these effects while treatment combined with HMOX-1 agonist Hemin enhanced these effects (Figures 7B,C). Consistently, the mRNA expression of Nrf-2, Hmox-1 and Slc7a11 were obviously upregulated in CTX group, as well as Fth1 and Ptgs2, two downstream key genes involved in ferroptosis (Figure 7D). However, compared with the CTX group, the CTX + Fer1 and the CTX + Hemin group did not show significant changes in Nrf2, Hmox-1 and Slc7a11 levels (Figure 7D). These results implied that CTX promoted tumor cell ferroptosis and combination treatment with HMOX-1 agonist could enhanced its chemotherapy effect in animals, which involved NRF2/HMOX-1 system.