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Role of Mitochondria in Ferroptosis
Article
Role of Mitochondria in Ferroptosis Graphical Abstract
Authors Minghui Gao, Junmei Yi, Jiajun Zhu, Alexander M. Minikes, Prashant Monian, Craig B. Thompson, Xuejun Jiang
Correspondence [email protected] (M.G.), [email protected] (X.J.)
In Brief
Highlights d
Mitochondria play a pivotal role in cysteine-deprivationinduced (CDI) ferroptosis
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Mitochondrial membrane potential hyperpolarization is associated with CDI ferroptosis
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TCA cycle, electron transport chain, and glutaminolysis function in CDI ferroptosis
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Mutation of tumor suppressor fumarate hydratase confers resistance to CDI ferroptosis
Gao et al., 2019, Molecular Cell 73, 354–363 January 17, 2019 ª 2018 Elsevier Inc. https://doi.org/10.1016/j.molcel.2018.10.042
Gao et al. show that mitochondria play a crucial and proactive role in cysteinedeprivation-induced ferroptosis but not in GPX4 inhibition-induced ferroptosis. Mechanistically, the mitochondrial TCA cycle and electron transport chain promote cysteine-deprivation-induced ferroptosis by serving as the major source for cellular lipid peroxide production. The anaplerotic role of glutaminolysis in replenishing the TCA cycle intermediates explains its involvement in cysteinedeprivation-induced ferroptosis. Importantly, mitochondria-mediated ferroptosis might contribute to the antitumor function of fumarate hydratase, a component of the TCA cycle and a tumor suppressor in renal cancer.
Molecular Cell
Article Role of Mitochondria in Ferroptosis Minghui Gao,1,2,5,* Junmei Yi,2,5 Jiajun Zhu,3 Alexander M. Minikes,2,4 Prashant Monian,2 Craig B. Thompson,3 and Xuejun Jiang2,6,* 1The
HIT Center for Life Sciences, School of Life Science and Technology, Harbin Institute of Technology, Harbin 150080, China Biology Program, Memorial Sloan Kettering Cancer Center, New York City, NY 10065, USA 3Department of Cancer Biology and Genetics, Memorial Sloan Kettering Cancer Center, New York City, NY 10065, USA 4BCMB Allied Program, Weill Cornell Graduate School of Medical Sciences, New York City, NY 10065, USA 5These authors contribute equally 6Lead Contact *Correspondence: [email protected] (M.G.), [email protected] (X.J.) https://doi.org/10.1016/j.molcel.2018.10.042 2Cell
SUMMARY
Ferroptosis is a regulated necrosis process driven by iron-dependent lipid peroxidation. Although ferroptosis and cellular metabolism interplay with one another, whether mitochondria are involved in ferroptosis is under debate. Here, we demonstrate that mitochondria play a crucial role in cysteinedeprivation-induced ferroptosis but not in that induced by inhibiting glutathione peroxidase-4 (GPX4), the most downstream component of the ferroptosis pathway. Mechanistically, cysteine deprivation leads to mitochondrial membrane potential hyperpolarization and lipid peroxide accumulation. Inhibition of mitochondrial TCA cycle or electron transfer chain (ETC) mitigated mitochondrial membrane potential hyperpolarization, lipid peroxide accumulation, and ferroptosis. Blockage of glutaminolysis had the same inhibitory effect, which was counteracted by supplying downstream TCA cycle intermediates. Importantly, loss of function of fumarate hydratase, a tumor suppressor and TCA cycle component, confers resistance to cysteine-deprivation-induced ferroptosis. Collectively, this work demonstrates the crucial role of mitochondria in cysteine-deprivation-induced ferroptosis and implicates ferroptosis in tumor suppression. INTRODUCTION Ferroptosis, an iron-dependent form of regulated necrosis, has emerged as a new cell death modality highly relevant to disease (Angeli et al., 2017; Gao and Jiang, 2018; Stockwell et al., 2017; Yang and Stockwell, 2016). Ferroptosis results from the accumulation of cellular reactive oxygen species (ROS) that exceed the redox contents maintained by glutathione (GSH) and the phospholipid hydroperoxidases that use GSH as a substrate. The synthetic, small molecule compound erastin can trigger ferroptosis by inhibiting the activity of cystine-glutamate antiporter (system Xc ), leading to the depletion of cellular cysteine and
GSH, thus the collapse of cellular redox homeostasis (Dixon et al., 2012). Importantly, it is the lipid ROS/peroxides, rather than cytosolic ROS, that unleash ferroptosis. As such, inactivation of glutathione peroxidase 4 (GPX4), an enzyme required for the clearance of lipid ROS, can induce ferroptosis even when cellular cysteine and GSH contents are normal (Friedmann Angeli et al., 2014; Ingold et al., 2018; Yang et al., 2014). Although the physiological function of ferroptosis is still elusive, its involvement in multiple human diseases has been established. Ferroptosis is a major mechanism for cell death associated with ischemic organ injury, including ischemic heart diseases, brain damage, and kidney failure (Friedmann Angeli et al., 2014; Gao et al., 2015a; Linkermann et al., 2014). A role of ferroptosis in neurodegeneration has also been implicated (Chen et al., 2015; Do Van et al., 2016; Skouta et al., 2014). In cancer, ferroptosis has been shown to contribute to the tumor suppressive function of p53 (Galluzzi et al., 2015; Jennis et al., 2016; Jiang et al., 2015; Wang et al., 2016). Many types of cancer cells that are resistant to chemotherapy and certain targeted therapies appear to be sensitive to ferroptosis induced by GPX4 inhibition (Hangauer et al., 2017; Viswanathan et al., 2017). These findings suggest that modulating ferroptosis might be potential therapeutic approaches in treating cancer or other diseases. Cellular metabolism is essential for ferroptosis, presumably because lipid ROS is mainly generated from various steps of cellular metabolism. Mounting evidence has demonstrated that diverse cellular metabolic processes, including lipid metabolism and amino acid metabolism (particularly that involves cysteine and glutamine), contribute to ferroptosis (Gao and Jiang, 2018; Stockwell et al., 2017). Interestingly, the stress-responsive catabolic pathway, autophagy, can also promote ferroptosis by degrading iron-storage protein ferritin and thus increase cellular iron concentration (Gao et al., 2016; Hou et al., 2016). Iron, as an essential cofactor of a plethora of metabolic enzymes and as a catalyst of lipid peroxide-generating Fenton reaction, drives oxygen and redox-based metabolism and cellular ROS production. Despite the central role of mitochondria in oxidative metabolism, it remains unclear whether mitochondria play a central role in ferroptosis. Supporting this possibility, ferroptosis is associated with dramatic morphological changes of mitochondria, including mitochondrial fragmentation and cristae enlargement
354 Molecular Cell 73, 354–363, January 17, 2019 ª 2018 Elsevier Inc.
(Dixon et al., 2012; Doll et al., 2017), and some potent ferroptosis inhibitors appear to be exquisitely targeted to mitochondria (Krainz et al., 2016). Furthermore, glutamine metabolism, known as glutaminolysis, is required for cysteine-deprivation-induced (CDI) ferroptosis (Gao et al., 2015a, 2015b), and one of the major functions of glutaminolysis is to fuel the mitochondrial tricarboxylic acid (TCA) cycle. Intriguingly, mitochondrial glutaminase (GLS), GLS2, instead of cytosolic GLS1, has been shown to be required for ferroptosis, although both enzymes catalyze glutaminolysis (Cassago et al., 2012; Gao et al., 2015a; Jennis et al., 2016). All these observations are consistent with the potential involvement of mitochondria in ferroptosis. However, there is also strong evidence arguing against a major role of mitochondria in ferroptosis. For example, a comparison between a mitochondrial DNA-depleted (r0) cancer cell line and its parental line did not show a significant difference in ferroptosis sensitivity (Dixon et al., 2012), and a mechanistic investigation into a cohort of ferrostatin analogs (inhibitors of ferroptosis) failed to establish a correlation between mitochondrial localization of ferrostatins with their anti-death potency (Gaschler et al., 2018). Taken together, the functional relevance of mitochondria in ferroptosis is still highly debatable. In this study, through a series of cellular, molecular, pharmacological, and metabolomic analyses, we demonstrated that the mitochondrion is a crucial player in ferroptosis induced by cysteine deprivation (inhibition of system Xc by erastin or using culture medium free of cystine). Mechanistically, the canonical metabolic activity of mitochondria, including both the TCA cycle and mitochondrial electron transport chain (ETC) activity, are required for the generation of sufficient lipid ROS to initiate ferroptosis. Furthermore, cancer cells deficient of the mitochondrial tumor suppressor fumarate hydratase (FH) are resistant to CDI ferroptosis—this result supports the notion that ferroptosis might be a physiologically relevant tumor suppressive mechanism and provides insights into potential ferroptosis-inducing cancer therapeutic approaches. Importantly, the role of mitochondria in ferroptosis is context dependent; if the activity of the glutathione-dependent peroxidase (GPX4) is inhibited, cells undergo ferroptosis independent of mitochondrial function. RESULTS Mitochondria Regulate Cysteine-Deprivation-Induced Ferroptosis To determine unambiguously whether mitochondria play a role in ferroptosis, we engineered human fibrosarcoma HT1080 cells depleted of mitochondria. Mitochondrial depletion was achieved by activating mitochondria-targeted autophagy, known as mitophagy (Youle and Narendra, 2011). First, we treated control and Parkin (tagged with mCherry)-overexpressing HT1080 cells with the mitochondrial uncoupler carbonyl cyanide m-chlorophenyl hydrazine (CCCP). Under this condition, mitochondria are depolarized by CCCP and consequently subjected to clearance via Parkin-mediated mitophagy (Narendra et al., 2008). As shown in Figures 1A and 1B, upon CCCP treatment for 24 or 48 hr followed by recovery in CCCP-free culture medium for 24 hr, mitochondria were significantly eliminated in Parkinoverexpressed cells (see the disappearance of MitoTracker
signal and mitochondrial proteins TOM20 and TIM23). Subsequently, cells were incubated with cystine (CC)-free medium or a pharmacological inducer of ferroptosis, erastin. Compared to control cells, Parkin-overexpressing, mitochondria-depleted cells were less sensitive to ferroptosis triggered by CC starvation or erastin (Figure 1C), confirming that mitochondria promote CDI ferroptosis. Lipid ROS accumulation is a hallmark of ferroptosis, but where and how it is generated during ferroptosis have not been defined. As the mitochondrion is a major organelle for cellular ROS production, does this organelle contribute to lipid ROS generation upon cysteine deprivation? To test this possibility, we monitored lipid ROS accumulation using C11 BODIPY 581/591, a fluorescent probe for membrane-localized ROS. Indeed, upon CC starvation or erastin treatment, mitochondria-depleted cells had a decreased level of lipid ROS accumulation (Figure 1D). Further, when we examined subcellular localization of the lipid ROS probe by confocal imaging (the probe changes fluorescence from red to green upon oxidation), we found that, in both HT1080 cells and mouse embryonic fibroblasts (MEFs) treated with erastin, the oxidized probe first appeared in a distribution that significantly colocalized with mitochondria, then colocalized with the plasma membrane at later time points (Figures 1E and S1). Mitochondrial TCA Cycle Participates in CysteineDeprivation-Induced Ferroptosis Glutaminolysis has been demonstrated to be required for CDI ferroptosis (Gao et al., 2015a). In the absence of glutamine, neither CC starvation nor pharmacological inhibition of CC transporter subunit SLC7A11 by erastin can induce ferroptosis. As a major function of glutaminolysis is to maintain the mitochondrial TCA cycle through anaplerosis (Figure 2A), is the TCA cycle required for ferroptosis? To test this possibility, we examined whether TCA cycle metabolites downstream of glutaminolysis can recapitulate the function of glutamine in ferroptosis. Indeed, alphaKetoglutaric acid (aKG), which is the TCA cycle metabolite immediately downstream of glutaminolysis, can replace glutamine for CDI lipid ROS accumulation and ferroptosis (Figures 2B and 2C). Further, TCA metabolites downstream of aKG, including succinate (Suc), fumarate (Fum), and malate (Mal), can all replace the role of glutamine in lipid ROS accumulation and ferroptosis induced by CC starvation or system Xc inhibition in both MEFs and HT1080 cells (Figures 2D–2F and S2). To further confirm that glutamine supports ferroptosis via the TCA cycle, we measured TCA metabolites under different conditions by gas chromatography–mass spectrometry (GC/MS) metabolomic analysis. The results (Figure 2G) showed that levels of TCA cycle metabolites were not changed by CC starvation alone, within the time range of our analysis. In the absence of glutamine, levels of multiple TCA cycle metabolites, including aKG, fumarate, and malate, were significantly decreased. In contrast, TCA cycle metabolites upstream of aKG, such as citrate, were affected more by glucose status than by glutamine status. Mitochondrial Electron Transport Chain Regulates Cysteine-Deprivation-Induced Ferroptosis One of the most important functions of the TCA cycle is to support electron transport activity of protein complexes embedded Molecular Cell 73, 354–363, January 17, 2019 355
Figure 1. Mitochondria Regulate CysteineDeprivation-Induced Ferroptosis (A–D) Mitochondrial depletion by Parkin-mediated mitophagy inhibits cysteine-deprivation-induced ferroptosis. WT (control) or mCherry-Parkin overexpressing (Parkin OE) HT1080 cells were treated with 10 mM Carbonyl cyanide m-chlorophenyl hydrazine (CCCP) for 48 hr, unless otherwise indicated (as in B, 24 or 48 hr), to induce mitochondrial depletion. After recovery from CCCP treatment by incubating cells in CCCP-free normal culture medium for 24 hr, the following experiments were performed: the presence of mitochondria was assessed by MitoTracker staining (scale bar, 10 mM) (A) or by western blot for endogenous TOM20 and TIM23 (B); cells were treated with erastin or CC starvation (10% (V/V) dialyzed FBS was used for CC starvation assay medium hereafter) for 12 hr for cell death measurement (C) or 8 hr for lipid ROS measurement (D). Cell death was measured with Sytox Green staining coupled with flow cytometry. The accumulation of lipid ROS was assessed by BODIPY C11 staining coupled with flow cytometry analysis. All quantitative data are presented as mean ± SD from three independent experiments, and p values were calculated with unpaired t test. **p < 0.01, ***p < 0.001, ****p < 0.0001. (E) Colocalization of oxidized lipid and mitochondria. MEFs were treated as indicated for 4 hr, then cells were co-stained with BODIPY C11 and MitoTracker. Oxidized BODIPY C11 (green) indicating lipid ROS and mitochondrial signals (red) were imaged by fluorescent microscope. Scale bar, 10 mm. See also Figure S1.
in the inner-membrane of mitochondria (Figure 3A). Since the TCA cycle is required for ferroptosis induced by cellular cysteine deprivation, are components of the ETC also required? To address this question, we utilized an array of ETC inhibitors. As shown in Figures 3B–3E, inhibitors of mitochondrial complex I (rotenone), complex II (DBM), complex III (antimycin), and complex IV (NaN3) all suppressed lipid ROS accumulation and ferroptosis induced by CC starvation or erastin. Taken together, these results indicate that ETC contributes to ferroptosis induced by cellular cysteine deprivation. Notably, although all tested ETC inhibitors significantly blocked ferroptosis at earlier time points (12 hr), some of them (antimycin and NaN3) maintained their inhibitory effect at later time points, but others (inhibitors for complex I and complex II) gradually lost such effect (Figure S3). This might be due to different stability of these compounds; alternatively, it may be that inhibition of complex I or complex II alone is not sufficient to block electron transfer to the downstream complex III. Mitochondrial Membrane Potential Hyperpolarization Is Associated with Cysteine-Deprivation-Induced Lipid ROS Accumulation and Ferroptosis Mitochondria produce ATP by utilizing the proton electrochemical gradient potential across the mitochondrial membrane, 356 Molecular Cell 73, 354–363, January 17, 2019
which is the result of the action of the TCA cycle and ETC. As the TCA cycle and ETC promote CDI ferroptosis, is ferroptosis associated with a change of mitochondrial membrane potential (MMP)? To test this possibility, we used MitoTracker (tetramethylrhodamine ethyl ester, or TMRE) staining to monitor MMP, upon induction of ferroptosis and as a control, of apoptosis. As expected, during mitochondria-mediated apoptosis, a decrease of MMP was observed prior to eventual cell death (Figure S4A; Video S1). This decrease of MMP is indicative of mitochondrial outer membrane permeabilization (MOMP), a crucial characteristic of mitochondria-mediated apoptosis (Green and Reed, 1998; Kroemer and Reed, 2000). In striking contrast, ferroptosis inducers, such as CC starvation (Figures 4A and 4B; and Video S2), amino acid-free medium plus full serum (Gao et al., 2015a) (Figure S4B; Video S3), erastin (Figure S4C), or glutamate (Figure S4D), can all induce MMP hyperpolarization. To further investigate the role of MMP in CDI ferroptosis, we examined the effect of the mitochondrial uncoupler CCCP on ferroptosis. As shown in Figures 4C, 4E, S4E, and S4F, CCCP disrupted MMP and completely blocked CDI lipid ROS accumulation and ferroptosis. Moreover, proliferation can be recovered after removing both erastin and CCCP, as assessed by cell colony formation assay (Figure 4F), indicating that temporarily disrupting MMP by CCCP can prevent ferroptotic cell death without
Figure 2. Mitochondrial TCA Cycle Promotes Cysteine-Deprivation-Induced Ferroptosis (A) An overview of mammalian tricarboxylic acid cycle. (B and C) a-ketoglutarate can mimic the death-inducing activity of L-Glutamine (L-Gln). (B) Microscopy showing cell death. MEFs were treated as indicated for 10 hr. Upper panel: phase-contrast; lower panel: propidium iodide (PI) staining for dead cells (Scale bar, 100 mM). (C) For cell death measurement, MEFs were subjected to the same treatment as in (B), and cell death was determined by propidium iodide (PI) staining coupled with flow cytometry; for lipid ROS measurement, MEFs were treated as in (B) for 8 hr and then lipid ROS was determined by BODIPY C11 staining coupled with flow cytometry. (D–F) TCA metabolites succinate (D), fumarate (E), and malate (F) can mimic the death inducing activity of L-Gln. (G) Glutamine is required for the maintenance of the normal level of multiple TCA cycle metabolites during ferroptosis. MEFs were treated as indicated for 8 hr, and samples were collected and measured by GC/MS metabolomic analysis as detailed in STAR Methods. For all experiments, 10% (V/V) dialyzed FBS was used in the assay medium. Q, L-glutamine (1 mM); CC, L-cystine (0.2 mM); aKG, Dimethyl-a-2-oxoglutarate (mM); Suc, Dimethyl succinate (mM); Fum, Dimethyl fumarate (mM); Mal, Dimethyl (S)-( )-malate (mM); and Fer-1, ferrostatin-1 (1 mM). All quantitative data are presented as mean ± SD from three independent experiments. **p < 0.01, ***p < 0.001, ****p < 0.0001 by unpaired Student’s t test. See also Figure S2.
compromising long-term cell viability. This conclusion is consistent with the effect of pharmacological inhibition of ETC on ferroptosis (Figures 3 and S4G–S4J). Glutaminolysis and TCA Cycle Drive FerroptosisAssociated MMP Hyperpolarization We subsequently determined the role of glutaminolysis and the TCA cycle in ferroptosis-associated MMP hyperpolarization. In the absence of glutamine, CC starvation failed to induce MMP hyperpolarization (Figure 4G). Similarly, inhibition of glutaminolysis by aminooxyacetic acid (AOA) blocked CC starvation-induced MMP hyperpolarization (Figure 4H). Therefore, glutamine metabolism is essential for CC starvation-induced MMP hyperpolarization. Furthermore, TCA cycle intermediates downstream of glutaminolysis, including aKG, succinate, fumarate, and malate, were all able to substitute glutamine for CC starvation-induced MMP hyperpolarization (Figures 4I–4L). Therefore, glutaminolysis and TCA cycle are essential for ferroptosis-associated MMP hyperpolarization. Presumably, MMP hy-
perpolarization reflects an increase of mitochondrial ETC activity and subsequent lipid ROS generation, which explains its role in ferroptosis. However, the mechanisms by which cysteine deprivation triggers MMP hyperpolarization are not clear and warrant further investigation. Mitochondrial Function Is Dispensable for GPX4 Inhibition-Induced Ferroptosis Importantly, although glutaminolysis and mitochondrial TCA cycle/ETC activity contribute to CDI ferroptosis, ferroptosis can still be triggered by pharmacological inhibition or genetic elimination of GPX4 when these mitochondrial activities are disrupted. Mitochondria-depleted HT1080 cells did not show significant resistance when ferroptosis was triggered by the GPX4 inhibitor RSL3 (Figures 5A and 5B). ETC inhibitors also failed to impinge on lipid ROS accumulation and ferroptosis upon RSL3 treatment (Figures 5C, 5D, and S5A). Further, we generated GPX4-knockout (KO) HT1080 cells by CRIPSR/Cas9 technology and found that ETC inhibitors could not block ferroptosis Molecular Cell 73, 354–363, January 17, 2019 357
MMP hyperpolarization (Figure S5B) without inhibiting RSL3induced lipid ROS accumulation or ferroptosis in both MEFs and HT1080 cells (Figures 5C, 5D, S5A, and S5B). Moreover, we found that GPX4 inhibition-induced ferroptosis can occur even in the absence of glutamine (Figure 5G). Therefore, glutamine and mitochondria are dispensable for GPX4 inhibitioninduced ferroptosis.
Figure 3. ETC Activity Promotes Cysteine-Deprivation-Induced Ferroptosis (A) Scheme shows mitochondrial electron transport chain (ETC) complexes and their inhibitors. (B) Inhibitors of ETC can inhibit CC starvation-induced ferroptosis. MEFs were treated with CC starvation for 12 hr in the absence or presence of ETC inhibitors as indicated. Cell death was determined by PI staining coupled with flow cytometry. Complex I inhibitor rotenone (Rot), 10 mM; Complex II inhibitor diethyl butylmalonate (DBM), 2 mM; Complex III inhibitor antimycin A (Anti A), 50 mM; and Complex IV inhibitor NaN3, 15 mM. (C) ETC inhibitors can inhibit CC starvation-induced lipid ROS accumulation. MEFs were treated with CC for 6 hr in the absence or presence of ETC inhibitors as indicated. Accumulation of lipid ROS was determined by BODIPY C11 staining coupled with flow cytometry. (D) ETC inhibitors can inhibit erastin-induced ferroptosis. MEFs were treated with 1 mM erastin for 12 hr in the absence or presence of ETC inhibitors as indicated. Cell death was determined by PI staining coupled with flow cytometry. (E) ETC inhibitors can inhibit erastin-induced lipid ROS accumulation. MEFs were treated with 1 mM erastin for 6 hr in the absence or presence of ETC inhibitors as indicated. Accumulation of lipid ROS was determined by BODIPY C11 coupled with flow cytometry. All quantitative data are presented as mean ± SD from three independent experiments. ***p < 0.001, ****p < 0.0001 by unpaired Student’s t test. See also Figure S3.
induced by GPX4-KO (Figures 5E–5G). Consistently, although RSL3 treatment can induce MMP hyperpolarization, the mitochondrial uncoupler CCCP can completely ablate RSL3-induced 358 Molecular Cell 73, 354–363, January 17, 2019
Loss of Function of Mitochondrial Tumor Suppressor Fumarate Hydratase Confers Resistance to Ferroptosis in Renal Cancer Cells Genetic mutation of the gene encoding for the TCA cycle enzyme fumarate hydratase (FH, also known as fumarase) has been identified in benign and malignant renal cancer lesions and has been demonstrated to be a bona fide tumor suppressor in renal cancer (Alam et al., 2005; Chuang et al., 2005; Gottlieb and Tomlinson, 2005; King et al., 2006; Tomlinson et al., 2002). In light of our finding that mitochondrial TCA cycle and respiration drive CDI ferroptosis, we examined whether loss of FH function renders cells more resistant to ferroptosis, thus contributing to its tumor suppressive function. We tested this possibility in two pairs of patient-derived isogenic renal cancer cell lines, FH mutant UOK262 and UOK262 reconstituted with wild-type (WT) FH and FH mutant UOK268 and UOK268 reconstituted with WT FH (Tong et al., 2011). Compared to cells expressing WT FH, FH mutant renal cancer cells were less sensitive to CC starvationinduced ferroptosis (Figures 6A–6C), and cell death in these mutant cells could be restored by supplying exogenous malate, which is the metabolite downstream of FH in the TCA cycle (Figures 6A and 6B). Importantly, in a cell colony formation experiment, FH mutant renal cancer cells remained viable and proliferative upon CC starvation, whereas the isogenic cells expressing WT FH failed to proliferate under this condition (Figure 6D). These results suggest that under oxidative stress (common for growth environment of solid tumors), loss of FH function can make cancer cells more resistant to ferroptotic death, thus conferring tumorigenic advantage. DISCUSSION Cellular metabolism and ferroptosis closely interact with one another, and lipid ROS, mainly a product of oxygen/iron-driven metabolism, is essential for the execution of ferroptosis. For these reasons, it appears logical that mitochondria should play a central role in ferroptosis. Nonetheless, whether the mitochondria are an important component in ferroptosis remains highly controversial. In this study, we present evidence to demonstrate that the mitochondrion is indeed a crucial player in ferroptotic cell death induced by cysteine deprivation. When mitochondria are depleted via Parkin-dependent mitophagy, cells become more resistant to ferroptosis upon CC starvation or pharmacological inhibition of CC import (Figure 1). Mechanistically, we showed that the role of the mitochondrion in ferroptosis is due to its metabolic function, and both mitochondrial TCA cycle and the action of ETC are required for a potent ferroptosis (Figures 2, 3, S2, and S3). Ferroptotic function of TCA cycle is the reason why glutaminolysis, a major source of anaplerosis, is required for ferroptosis (Figures 2 and S2).
Figure 4. Mitochondrial Membrane Potential Hyperpolarization Is Associated with Cysteine-Deprivation-Induced Lipid ROS Accumulation and Ferroptosis (A) Ferroptosis is accompanied by mitochondrial membrane potential (MMP) hyperpolarization. Representative still images from time-lapse imaging of MMP in MEFs undergoing CC starvationinduced ferroptosis. Cells were incubated with TMRE (200 nM) and subjected to CC starvation for indicated time (scale bar, 5 mM). (B) Quantification of MMP during CC starvationinduced ferroptosis. MEFs were treated with CC starvation, and 500 nM TMRE was added 30 min before each indicated time point. MMP was measured by flow cytometry and mean fluorescence was calculated. (C) CCCP can inhibit ferroptosis-associated MMP hyperpolarization. MEFs were treated with CC starvation plus or minus 10 mM CCCP for indicated time, and 500 nM TMRE was added 30 min before each indicated time point. MMP was measured by flow cytometry and mean fluorescence was calculated. (D) CCCP can block ferroptosis induced by CC starvation or erastin. MEFs were treated as indicated for 12 hr and cell death was determined by PI staining coupled with flow cytometry. (E) CCCP can block lipid ROS accumulation induced by CC starvation or erastin. MEFs were treated as indicated for 8 hr, and lipid ROS was measured by BODIPY C11 staining coupled with flow cytometry. (F) Cells protected by CCCP from erastininduced ferroptosis maintain viability. MEFs and HT1080 cells were seeded at low density in 12-well dishes and treated with erastin for 16 hr in the absence or presence of CCCP as indicated. Cells were then allowed to recover in drug-free DMEM medium for 3 days and subsequently fixed and stained with crystal violet. (G and H) Glutaminolysis and TCA cycle drive ferroptosis-associated MMP hyperpolarization in MEFs. (G) Glutamine is required for CC starvation-induced MMP hyperpolarization. (H) Transaminase inhibitor AOA (Aminooxyacetic acid, 0.5 mM) can block CC starvation-induced MMP hyperpolarization. (I–L) TCA intermediates a-ketoglutarate (I) succinate (J), fumarate (K), and malate (L) can replace the requirement of glutamine for CC starvation-induced MMP hyperpolarization. aKG, Dimethyl-a-2-oxoglutarate (8 mM); Suc, Dimethyl succinate (8 mM); Fum, Dimethyl fumarate (5 mM); and Mal, Dimethyl (S)-(-)-malate (32 mM). All quantitative data are presented as mean ± SD from three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by unpaired Student’s t test. See also Figure S4 and Videos S1, S2, and S3.
Why, then, is the role of mitochondria in ferroptosis so controversial? The following three reasons can reconcile this discrepancy. First, our study indicates that the role of mitochondria in ferroptosis is context dependent: blockage of mitochondrial function potently inhibits CDI ferroptosis; however, upon elimination or pharmacological inhibition of GPX4, the very downstream component of the ferroptosis pathway responsible for lipid ROS clearance, cells can commit ferroptosis independently
of mitochondria (Figures 5 and S5). When cysteine is scarce, mitochondrial metabolism contributes significantly to rapid glutathione depletion and subsequent lipid ROS generation and ferroptosis. On the other hand, upon GPX4 inhibition, although one would predict that inhibition of mitochondria should also attenuate lipid ROS generation and thus may at least delay ferroptosis, we did not observe a measurable effect of mitochondrial ETC inhibitors on slowing down ferroptosis in Molecular Cell 73, 354–363, January 17, 2019 359
Figure 5. Mitochondria Are Dispensable for GPX4 InhibitionInduced Ferroptosis (A and B) Mitochondrial depletion by Parkin-mediated mitophagy cannot inhibit RSL3-induced ferroptosis. Mitochondria-depleted HT1080 cells were created as described in Figure 1. Cells were treated with RSL3 for 6 hr for cell death measurement (A) or 4 hr for lipid ROS measurement (B). Cell death was measured with DAPI staining coupled with flow cytometry. The accumulation of lipid ROS was assessed by BODIPY C11 staining coupled with flow cytometry analysis. (C and D) Mitochondrial ETC inhibitors cannot inhibit RSL3-induced ferroptosis (C) and lipid ROS accumulation (D). (E) Western blotting confirmed CRISPR-Cas9-mediated knockout of GPX4 in HT1080 cells (see STAR Methods for detail). (F) Mitochondrial ETC inhibitors cannot inhibit GPX4 knockout-induced ferroptosis. GPX4 KO HT1080 cells were seeded and subsequently grew in DMEM medium containing 0.2 mM Trolox. Ferroptosis induction was initiated by switching cells to Trolox-free medium in the presence or absence of indicated ETC complex inhibitors for 8 hr. Complex I inhibitor rotenone (Rot), 10 mM; Complex II inhibitor Diethyl butylmalonate (DBM), 2 mM; Complex III inhibitor Myxothiazol (Myxo), 15 mM; and Complex IV inhibitor NaN3, 15 mM. (G) Glutamine is dispensable for GPX4 knockout-induced ferroptosis. GPX4 KO HT1080 cells were seeded and subsequently grew in DMEM medium containing 0.2 mM Trolox. Ferroptosis induction was initiated by switching cells to Trolox-free medium in the presence or absence of 1 mM glutamine for 8 hr. Scale bar, 100 mM. All quantitative data are presented as mean ± SD from three independent experiments. ns, non-significant by unpaired Student’s t test. See also Figure S5.
360 Molecular Cell 73, 354–363, January 17, 2019
GPX4-KO HT1080 cells (Figure S5D). It is likely that once GPX4 is eliminated, the low amount of lipid ROS produced by other mechanisms will be rapidly amplified through the Fenton chain reaction, leading to full-blown ferroptotic cell death even when mitochondrial activity is diminished. It is also possible that GPX4 inactivation may transduce a signal to, and thus hyperactivate, certain enzymes responsible for lipid ROS generation. Future investigation is needed to clarify this issue. Second, since ferroptosis is dictated by cellular metabolism, long-term alteration of cellular metabolism may rewire the molecular processes underlying ferroptosis. For example, in a type of experimental model cells known as r0 cells, mitochondrial DNA is depleted permanently (King and Attardi, 1989, 1996). To compensate the long-term lack of mitochondria, these cells need to be cultured with nutrients different from mitochondria-containing cells (King and Attardi, 1989, 1996). Therefore, metabolism in r0 cells, including lipid ROS generation (mainly a consequence of metabolism), is fundamentally rewired. For this reason, the mechanism governing the mitochondria-independent ferroptosis in r0 cells may differ significantly from that in normal, mitochondria-containing cells. Third, the methods for cell death measurement need to be carefully considered. For example, although cell viability assays are often used to infer cell death, they are not suitable for the study of the role of mitochondria in ferroptosis. This is because that manipulation of mitochondrial function may impact the outcome of these assays (many of them are ATP content based or metabolic activity based), regardless of cell death status. Assays directly measuring cell death are preferable for this purpose. Mitochondria are also involved in other modalities of programmed cell death, particularly apoptosis. Intriguingly, the roles of mitochondria in apoptosis and ferroptosis are fundamentally different. In mitochondria-mediated apoptosis, the mitochondrion serves as a reservoir, storing apoptosis-regulatory proteins such as cytochrome c and Smac, which, upon release to the cytoplasm, promote caspase activation (Jiang and Wang, 2004; Wang, 2001). Notably, the cytosolic apoptotic activities of these mitochondrial proteins are independent of, and can be molecularly dissected from, their mitochondrial functions. As such, a compromise of mitochondrial integrity and loss of MMP, known as MOMP, precedes the release of these proteins and subsequent caspase-9 activation. Interestingly, in CDI ferroptosis, it is the canonical metabolic function of mitochondria that actively contributes to lipid ROS generation, a prerequisite of ferroptosis, and inhibition of TCA cycle or ETC function, or disruption of MMP, can all abrogate ferroptosis under this condition. Strikingly, a transient hyperpolarization of MMP can be observed during CDI ferroptosis, followed by the eventual collapse of MMP (Figures 4 and S4; Videos S1, S2, and S3). This unique feature is indicative of the crucial role of mitochondrial metabolic activity in CDI ferroptosis. Taken together, while mitochondria play a passive role in apoptosis by storing various pro-apoptotic proteins, they play a crucial and proactive role in CDI ferroptosis by fueling metabolism and lipid ROS production. Further, the findings presented in this work provide novel mechanistic insights into some enigmatic observations in cancer biology. FH, a metabolic enzyme in TCA cycle, has
Figure 6. Loss of Function of Mitochondrial Tumor Suppressor Fumarate Hydratase Confers Resistance to Ferroptosis (A–C) Fumarase (FH) mutant cancer cells are more resistant to CC starvation-induced ferroptosis than isogenic cells expressing WT FH. (A) Microscopy showing cell death. Two pairs of FH mutant cancer cells and their isogenic cells expressing WT FH were treated as indicated for 36 hr (UOK262 and UOK262+WT FH) or 60 hr (UOK268 and UOK268+WT FH). Upper panel: phase-contrast; lower panel: PI staining for dead cells (scale bar, 100 mM). Western blot shows the expression level of FH in these cell lines. (B) Cell death was determined by PI staining coupled with flow cytometry. (C) Lipid ROS was determined by BODIPY C11 staining coupled with flow cytometry. (D) FH mutant cancer cells recovered and proliferated after CC starvation. Cells were treated with CC starvation for 24 hr (UOK262 and UOK262+WT FH) or 60 hr (UOK268 and UOK268+WT FH), then cells were incubated in full DMEM medium for 3 days. Colony formation was measured by crystal violet staining. All quantitative data are presented as mean ± SD from three independent experiments. ****p < 0.0001 by unpaired Student’s t test. See also Figure S6.
been demonstrated to be a tumor suppressor. This is counterintuitive because one would assume that defects in the TCA cycle or ETC would be disadvantageous for cancer cell growth. Indeed, under normal culture condition, reconstitution of WT FH into UOK262/268 cancer cells rendered them proliferating better (Figure S6). So, what is the molecular basis underlying the tumor suppressive activity of FH? A pseudo-hypoxic pathway was proposed to be the link between FH mutation and tumorigenesis. FH defect leads to aberrant accumulation of fumarate, which, in turn, inactivates prolyl hydroxylases and thus stabilizes hypoxia-inducible factor (HIF)-1a (Isaacs et al., 2005; Selak et al., 2005). However, both HIF-1a and HIF-2a are
dispensable for the formation of hyperplastic renal cysts in mice lacking expression of FH in the kidney (Adam et al., 2011), suggesting additional tumorigenic consequences caused by FH mutation. Our finding that loss of FH function renders cells more resistant to CDI ferroptosis (Figure 6) provides a novel mechanistic explanation of how dysfunction of FH contributes to malignancy independent of hypoxic signaling—we hypothesize that stronger resistance to ferroptosis caused by FH dysfunction contributes to tumorigenesis, although this is most likely not sufficient on its own. It remains to be determined why FH mutation preferentially drives tumorigenesis only in certain specific tissue types. Conceptually, our finding that ferroptosis contributes to the tumor suppressive function of FH is important. It has been reported recently that p53 can suppress cancer development via its ferroptosis-promoting activity even when it loses its function to promote apoptosis, senescence, and growth arrest (Jiang et al., 2015; Wang et al., 2016). Based on these novel findings about FH and p53, we hypothesize that ferroptosis is a natural tumor suppressive mechanism under diverse biological conditions. The significance of this hypothesis in cancer biology is obvious. As for the biology of ferroptosis, tumor suppression might represent an authentic physiological role of this unique modality of cell death, which, until now, remains to be defined. Molecular Cell 73, 354–363, January 17, 2019 361
STAR+METHODS
Angeli, J.P.F., Shah, R., Pratt, D.A., and Conrad, M. (2017). Ferroptosis inhibition: mechanisms and opportunities. Trends Pharmacol. Sci. 38, 489–498.
Detailed methods are provided in the online version of this paper and include the following:
Carey, B.W., Finley, L.W., Cross, J.R., Allis, C.D., and Thompson, C.B. (2015). Intracellular a-ketoglutarate maintains the pluripotency of embryonic stem cells. Nature 518, 413–416.
KEY RESOURCES TABLE CONTACT FOR REAGENT AND RESOURCE SHARING METHOD DETAILS B Cell culture B Measurement of mitochondrial membrane potential B Measurement of lipid ROS B CRISPR-mediated GPX4 deletion B Fluorescence microscopy B Metabolite analysis by gas chromatography-mass spectrometry (GC-MS) QUANTIFICATION AND STATISTICAL ANALYSIS DATA AND SOFTWARE AVAILABILITY
Cassago, A., Ferreira, A.P., Ferreira, I.M., Fornezari, C., Gomes, E.R., Greene, K.S., Pereira, H.M., Garratt, R.C., Dias, S.M., and Ambrosio, A.L. (2012). Mitochondrial localization and structure-based phosphate activation mechanism of Glutaminase C with implications for cancer metabolism. Proc. Natl. Acad. Sci. USA 109, 1092–1097.
d d d
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SUPPLEMENTAL INFORMATION Supplemental Information includes six figures and three videos and can be found with this article online at https://doi.org/10.1016/j.molcel.2018.10.042. ACKNOWLEDGMENTS The authors thank Dr. W. Marston Linehan for kindly providing the FH-mutant UOK262 and UOK268 cell lines, as well as their wild-type FH reconstituted counterparts. The authors thank members of the Jiang lab and the Thompson lab for critical reading and suggestions. This work is supported by NIH R01CA204232 (to X.J.), NIH R01CA201318 (to C.B.T.), a Geoffrey Beene Cancer Research fund (to X.J.), National Natural Science Foundation of China 31871388 (to M.G.), a Leukemia and Lymphoma Society Fellowship (to J.Z.), and NIH T32 fellowship 5T32GM008539-23 (to A.M.M.). This work is also supported by NCI cancer center core grant P30CA008748 to Memorial Sloan Kettering Cancer Center. AUTHOR CONTRIBUTIONS M.G. and X.J. conceived and supervised the study. M.G. and J.Y. performed most of the experiments in collaboration with J.Z., A.M.M., and P.M. M.G., J.Y., C.B.T., and X.J. analyzed the data and wrote the paper with suggestions from other authors. All authors reviewed the paper.
Chen, L., Hambright, W.S., Na, R., and Ran, Q. (2015). Ablation of the ferroptosis inhibitor glutathione peroxidase 4 in neurons results in rapid motor neuron degeneration and paralysis. J. Biol. Chem. 290, 28097–28106. Chuang, G.S., Martinez-Mir, A., Geyer, A., Engler, D.E., Glaser, B., CserhalmiFriedman, P.B., Gordon, D., Horev, L., Lukash, B., Herman, E., et al. (2005). Germline fumarate hydratase mutations and evidence for a founder mutation underlying multiple cutaneous and uterine leiomyomata. J. Am. Acad. Dermatol. 52, 410–416. Dixon, S.J., Lemberg, K.M., Lamprecht, M.R., Skouta, R., Zaitsev, E.M., Gleason, C.E., Patel, D.N., Bauer, A.J., Cantley, A.M., Yang, W.S., et al. (2012). Ferroptosis: an iron-dependent form of nonapoptotic cell death. Cell 149, 1060–1072. Do Van, B., Gouel, F., Jonneaux, A., Timmerman, K., Gele´, P., Pe´trault, M., Bastide, M., Laloux, C., Moreau, C., Bordet, R., et al. (2016). Ferroptosis, a newly characterized form of cell death in Parkinson’s disease that is regulated by PKC. Neurobiol. Dis. 94, 169–178. Doll, S., Proneth, B., Tyurina, Y.Y., Panzilius, E., Kobayashi, S., Ingold, I., Irmler, M., Beckers, J., Aichler, M., Walch, A., et al. (2017). ACSL4 dictates ferroptosis sensitivity by shaping cellular lipid composition. Nat. Chem. Biol. 13, 91–98. Friedmann Angeli, J.P., Schneider, M., Proneth, B., Tyurina, Y.Y., Tyurin, V.A., Hammond, V.J., Herbach, N., Aichler, M., Walch, A., Eggenhofer, E., et al. (2014). Inactivation of the ferroptosis regulator Gpx4 triggers acute renal failure in mice. Nat. Cell Biol. 16, 1180–1191. Galluzzi, L., Bravo-San Pedro, J.M., and Kroemer, G. (2015). Ferroptosis in p53-dependent oncosuppression and organismal homeostasis. Cell Death Differ. 22, 1237–1238. Gao, M., and Jiang, X. (2018). To eat or not to eat-the metabolic flavor of ferroptosis. Curr. Opin. Cell Biol. 51, 58–64. Gao, M., Monian, P., Quadri, N., Ramasamy, R., and Jiang, X. (2015a). glutaminolysis and transferrin regulate ferroptosis. Mol. Cell 59, 298–308. Gao, M., Monian, P., and Jiang, X. (2015b). Metabolism and iron signaling in ferroptotic cell death. Oncotarget 6, 35145–35146.
DECLARATION OF INTERESTS
Gao, M., Monian, P., Pan, Q., Zhang, W., Xiang, J., and Jiang, X. (2016). Ferroptosis is an autophagic cell death process. Cell Res. 26, 1021–1032.
C.B.T. is a founder of Agio Pharmaceuticals and a member of its scientific advisory board. Agio does not hold financial interests in the work reported in this paper.
Gaschler, M.M., Hu, F., Feng, H., Linkermann, A., Min, W., and Stockwell, B.R. (2018). Determination of the subcellular localization and mechanism of action of ferrostatins in suppressing ferroptosis. ACS Chem. Biol. 13, 1013–1020.
Received: July 6, 2018 Revised: September 16, 2018 Accepted: October 29, 2018 Published: December 20, 2018 REFERENCES Adam, J., Hatipoglu, E., O’Flaherty, L., Ternette, N., Sahgal, N., Lockstone, H., Baban, D., Nye, E., Stamp, G.W., Wolhuter, K., et al. (2011). Renal cyst formation in Fh1-deficient mice is independent of the Hif/Phd pathway: roles for fumarate in KEAP1 succination and Nrf2 signaling. Cancer Cell 20, 524–537. Alam, N.A., Olpin, S., Rowan, A., Kelsell, D., Leigh, I.M., Tomlinson, I.P., and Weaver, T. (2005). Missense mutations in fumarate hydratase in multiple cutaneous and uterine leiomyomatosis and renal cell cancer. J. Mol. Diagn. 7, 437–443.
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Tomlinson, I.P., Alam, N.A., Rowan, A.J., Barclay, E., Jaeger, E.E., Kelsell, D., Leigh, I., Gorman, P., Lamlum, H., Rahman, S., et al.; Multiple Leiomyoma Consortium (2002). Germline mutations in FH predispose to dominantly inherited uterine fibroids, skin leiomyomata and papillary renal cell cancer. Nat. Genet. 30, 406–410.
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Molecular Cell 73, 354–363, January 17, 2019 363
STAR+METHODS KEY RESOURCES TABLE
REAGENT or RESOURCE
SOURCE
IDENTIFIER
g-tubulin
Sigma
Cat# T6557; RRID:AB_477584
Tom20
Santa Cruz
Cat#SC11415; RRID:AB_2207533
Tim23
Cell Signal
Cat# 3130S; RRID:AB_10693298
Fumarase
Cell Signal
Cat#4567; RRID:AB_11178522
Vinculin
Cell Signal
Cat#13901; RRID:AB_2728768
Erastin
Sigma
Cat#E7781
RSL3
Selleckchem
Cat# S8155
Dimethyl 2-oxoglutarete
Sigma
Cat#349631
Dimethyl succinate
Sigma
Cat#W239607
Dimethyl fumarate
Sigma
Cat#242926
Dimethyl (S)-(-)-malate
Sigma
Cat#374318
Carbonyl cyanide m-chlorophenyl hydrazone
Sigma
Cat#C2759
Rotenone
Sigma
Cat#R8875
Antibodies
Chemicals and Reagents
Diethyl butylmalonate
Sigma
Cat#112038
Antimycin A
Sigma
Cat#A8674
Sodium azide
Sigma
Cat# S2002
Oligomycin A
Sigma
Cat#75351
Myxothiazol
Sigma
Cat# T5580
Ferrostatin-1
Sigma
Cat# M60042
TMRE
Enzo Life Sciences
Cat#ENZ-52309
BODIPY C11
Thermo Fisher Scientific
Cat# D3861
Lipofectamine 2000
Life Technologies
Cat# 11668027
Propidium Iodide Staining Solution
BD PharMingen
Cat# 556463
SYTOX Green Nucleic Acid Stain
Thermo Fisher Scientific
S7020
Mendeley
https://dx.doi.org/10.17632/95zbjbj95x.1
HT1080
ATCC
N/A
HT1080 mCherry Parkin
Dr. Xuejun Jiang Lab
N/A
MEFs
ATCC
N/A
UOK262
Dr. W. Marston Linehan Lab
N/A
UOK262 WT
Dr. W. Marston Linehan Lab
N/A
UOK268
Dr. W. Marston Linehan Lab
N/A
UOK268 WT
Dr. W. Marston Linehan Lab
N/A
pLx-gRNA
Addgene
#50662
GPX4-CACGCCCGATACGCTGAGTG
NA
N/A
Deposited Data Raw Data Experimental Models: Cell Lines
Plasmid and sgRNA sequence
CONTACT FOR REAGENT AND RESOURCE SHARING Requests for resources, reagents and further information should be directed to, and will be fulfilled by Xuejun Jiang (jiangx@ mskcc.org). e1 Molecular Cell 73, 354–363.e1–e3, January 17, 2019
METHOD DETAILS Cell culture Unless specified otherwise, all mammalian cells are maintained in DMEM with high glucose, sodium pyruvate (1 mM), glutamine (2 mM), penicillin (100 U/ml), streptomycin (0.1 mg/ml) and 10% (v/v) FBS at 37 C and 5% CO2. Cell death was analyzed by PI (100 ng/ml), Sytox (5 nM) or DAPI (1 mg/ml) staining coupled with microscopy or flow cytometry. Measurement of mitochondrial membrane potential Time-lapse imaging: cells were pre-loaded with 200 nm TMRE (Enzo Life Sciences, Cat#ENZ-52309) for 1 h. Subsequently, cells were washed with PBS twice to remove traces of TMRE from the media and treated as indicated. Fluorescence and differential interference contrast (DIC) images were acquired every 5-7 min, and images were analyzed using NIS elements software (Nikon) and ImageJ software (NIH). Flow cytometry: Cells were treated as indicated, and then 0.5 mM TMRE was added and incubated for 30 min. Excess TMRE was removed by washing the cells with PBS. Labeled cells were trypsinized and resuspended in PBS plus 2% FBS. Fluorescence at Ex/ Em = 549/575 nm was analyzed using a flow cytometer. Measurement of lipid ROS Lipid ROS was analyzed by flow cytometry: Cells were seeded at a density of 2.5x105 per well in a 6-well dish and grown overnight in DMEM. 5 mM BODIPY C11 (Thermo Fisher, Cat# D3861) was added into cell culture medium and incubated for 30 min after indicated treatment. Excess BODIPY C11 was then removed by washing the cells with PBS twice. Labeled cells were trypsinized and resuspended in PBS plus 2% FBS. Oxidation of BODIPY C11 resulted in a shift of the fluorescence emission peak from 590 nm to 510 nm proportional to lipid ROS generation and was analyzed using a flow cytometer. Lipid ROS imaging: Cells were seeded at a density of 2.5x105 per well on coverslips placed in a 6-well dish and grown overnight in DMEM. Cells were washed twice in HBSS and treated as indicated. Coverslips were washed in HBSS and incubated in HBSS containing 2 mM BODIPY 581/591 C11 (Invitrogen) and 200 nM MitoTracker Deep Red FM (Invitrogen) for 20 minutes. Coverslips were then inverted onto microscope slides. Slides were imaged using a Nikon Eclipse Ti-U inverted microscope and images were processed in Photoshop (Adobe). CRISPR-mediated GPX4 deletion Guide RNA sequence CACGCCCGATACGCTGAGTG targeting human GPX4 was used. pLx-gRNA (Addgene#50662) expression plasmid encoding guide RNA was generated and co-transfected with Streptococcus pyogenes Cas9 expression plasmid and transfected into HT1080 cells in the presence of 0.2 mM Trolox using Lipofectamine 2000 (Life Technologies). 24 h after the transfection, the cells were trypsinized and about 200 cells were seeded into a 10-cm plate and cultured in the presence of 0.2 mM Trolox. After cell clones were formed and expanded, western Blot was performed to screening for GPX4 knockout clones. Fluorescence microscopy MEFs stably expressing SU9-GFP or HT1080 cells stably expressing mCherry-Parkin were grown on glass coverslips in a 6-well plate. 24 h later, cells were treated as indicated. Coverslips were then fixed with 3.7% (vol/vol) paraformaldehyde (PFA) in 20 mM HEPES pH 7.5 for 30 min at room temperature. Coverslips were then mounted on microscope slides for visualization using Nikon Eclipse Ti-U Microscope with a 60 3 magnification objective. Metabolite analysis by gas chromatography-mass spectrometry (GC-MS) Metabolite analysis by GC-MS was performed as previously described (Carey et al., 2015). In brief, cells were seeded in standard culture medium in 6-well plates and the next day were changed into experimental medium. Metabolites were extracted with icecold 80% methanol supplemented with 20 mM deuterated 2-hydroxyglutarate (D-2-hydroxyglutaric2,3,3,4,4-d5 acid (d5-2HG)) as an internal standard. After overnight incubation at 80 C, lysates were harvested and centrifuged at 21,000 g for 20 min to remove protein. Extracts were dried in an evaporator (Genevac EZ-2 Elite) and resuspended by incubation at 30 C for 90 min in 50 ml of 40 mg/ml methoxyamine hydrochloride in pyridine. Metabolites were further derivatized by addition of 80 ml of MSTFA plus 1% TMCS (Thermo Scientific) and 70 ml ethyl acetate (Sigma) and incubated at 37 C for 30 min. Samples were analyzed using an Agilent 7890A GC coupled to Agilent 5975C mass selective detector. The GC was operated in splitless mode with constant helium gas flow at 1 ml/min. One microliter of derivatized metabolites was injected onto an HP-5MS column and the GC oven temperature ramped from 60 C to 290 C in 25 min. Peaks representing compounds of interest were extracted and integrated using MassHunter software (Agilent Technologies) and then normalized to both the internal standard (d5-2HG) peak area and the relative total biomass as determined by total cell number 3 mean cell volume. Ions used for quantification of metabolite levels are as follows: d5-2HG m/z 354; Citrate, m/z 465; aKG, m/z 304; fumarate, m/z 245 and malate, m/z 335. All peaks were manually inspected and verified relative to known spectra for each metabolite.
Molecular Cell 73, 354–363.e1–e3, January 17, 2019 e2
QUANTIFICATION AND STATISTICAL ANALYSIS All statistical analyses were performed using Prism 7.0c GraphPad Software. Data are presented as mean ± SD from 3 independent experiments. p values were calculated with Student’s unpaired t test, except for the measurement of mitochondrial membrane potential for which a two-way ANOVA test was used (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, see figure legend for further detail). DATA AND SOFTWARE AVAILABILITY Raw data for this paper has been deposited to Mendeley: https://dx.doi.org/10.17632/95zbjbj95x.1.
e3 Molecular Cell 73, 354–363.e1–e3, January 17, 2019
Role of Mitochondria in Ferroptosis Graphical Abstract
Authors Minghui Gao, Junmei Yi, Jiajun Zhu, Alexander M. Minikes, Prashant Monian, Craig B. Thompson, Xuejun Jiang
Correspondence [email protected] (M.G.), [email protected] (X.J.)
In Brief
Highlights d
Mitochondria play a pivotal role in cysteine-deprivationinduced (CDI) ferroptosis
d
Mitochondrial membrane potential hyperpolarization is associated with CDI ferroptosis
d
TCA cycle, electron transport chain, and glutaminolysis function in CDI ferroptosis
d
Mutation of tumor suppressor fumarate hydratase confers resistance to CDI ferroptosis
Gao et al., 2019, Molecular Cell 73, 354–363 January 17, 2019 ª 2018 Elsevier Inc. https://doi.org/10.1016/j.molcel.2018.10.042
Gao et al. show that mitochondria play a crucial and proactive role in cysteinedeprivation-induced ferroptosis but not in GPX4 inhibition-induced ferroptosis. Mechanistically, the mitochondrial TCA cycle and electron transport chain promote cysteine-deprivation-induced ferroptosis by serving as the major source for cellular lipid peroxide production. The anaplerotic role of glutaminolysis in replenishing the TCA cycle intermediates explains its involvement in cysteinedeprivation-induced ferroptosis. Importantly, mitochondria-mediated ferroptosis might contribute to the antitumor function of fumarate hydratase, a component of the TCA cycle and a tumor suppressor in renal cancer.
Molecular Cell
Article Role of Mitochondria in Ferroptosis Minghui Gao,1,2,5,* Junmei Yi,2,5 Jiajun Zhu,3 Alexander M. Minikes,2,4 Prashant Monian,2 Craig B. Thompson,3 and Xuejun Jiang2,6,* 1The
HIT Center for Life Sciences, School of Life Science and Technology, Harbin Institute of Technology, Harbin 150080, China Biology Program, Memorial Sloan Kettering Cancer Center, New York City, NY 10065, USA 3Department of Cancer Biology and Genetics, Memorial Sloan Kettering Cancer Center, New York City, NY 10065, USA 4BCMB Allied Program, Weill Cornell Graduate School of Medical Sciences, New York City, NY 10065, USA 5These authors contribute equally 6Lead Contact *Correspondence: [email protected] (M.G.), [email protected] (X.J.) https://doi.org/10.1016/j.molcel.2018.10.042 2Cell
SUMMARY
Ferroptosis is a regulated necrosis process driven by iron-dependent lipid peroxidation. Although ferroptosis and cellular metabolism interplay with one another, whether mitochondria are involved in ferroptosis is under debate. Here, we demonstrate that mitochondria play a crucial role in cysteinedeprivation-induced ferroptosis but not in that induced by inhibiting glutathione peroxidase-4 (GPX4), the most downstream component of the ferroptosis pathway. Mechanistically, cysteine deprivation leads to mitochondrial membrane potential hyperpolarization and lipid peroxide accumulation. Inhibition of mitochondrial TCA cycle or electron transfer chain (ETC) mitigated mitochondrial membrane potential hyperpolarization, lipid peroxide accumulation, and ferroptosis. Blockage of glutaminolysis had the same inhibitory effect, which was counteracted by supplying downstream TCA cycle intermediates. Importantly, loss of function of fumarate hydratase, a tumor suppressor and TCA cycle component, confers resistance to cysteine-deprivation-induced ferroptosis. Collectively, this work demonstrates the crucial role of mitochondria in cysteine-deprivation-induced ferroptosis and implicates ferroptosis in tumor suppression. INTRODUCTION Ferroptosis, an iron-dependent form of regulated necrosis, has emerged as a new cell death modality highly relevant to disease (Angeli et al., 2017; Gao and Jiang, 2018; Stockwell et al., 2017; Yang and Stockwell, 2016). Ferroptosis results from the accumulation of cellular reactive oxygen species (ROS) that exceed the redox contents maintained by glutathione (GSH) and the phospholipid hydroperoxidases that use GSH as a substrate. The synthetic, small molecule compound erastin can trigger ferroptosis by inhibiting the activity of cystine-glutamate antiporter (system Xc ), leading to the depletion of cellular cysteine and
GSH, thus the collapse of cellular redox homeostasis (Dixon et al., 2012). Importantly, it is the lipid ROS/peroxides, rather than cytosolic ROS, that unleash ferroptosis. As such, inactivation of glutathione peroxidase 4 (GPX4), an enzyme required for the clearance of lipid ROS, can induce ferroptosis even when cellular cysteine and GSH contents are normal (Friedmann Angeli et al., 2014; Ingold et al., 2018; Yang et al., 2014). Although the physiological function of ferroptosis is still elusive, its involvement in multiple human diseases has been established. Ferroptosis is a major mechanism for cell death associated with ischemic organ injury, including ischemic heart diseases, brain damage, and kidney failure (Friedmann Angeli et al., 2014; Gao et al., 2015a; Linkermann et al., 2014). A role of ferroptosis in neurodegeneration has also been implicated (Chen et al., 2015; Do Van et al., 2016; Skouta et al., 2014). In cancer, ferroptosis has been shown to contribute to the tumor suppressive function of p53 (Galluzzi et al., 2015; Jennis et al., 2016; Jiang et al., 2015; Wang et al., 2016). Many types of cancer cells that are resistant to chemotherapy and certain targeted therapies appear to be sensitive to ferroptosis induced by GPX4 inhibition (Hangauer et al., 2017; Viswanathan et al., 2017). These findings suggest that modulating ferroptosis might be potential therapeutic approaches in treating cancer or other diseases. Cellular metabolism is essential for ferroptosis, presumably because lipid ROS is mainly generated from various steps of cellular metabolism. Mounting evidence has demonstrated that diverse cellular metabolic processes, including lipid metabolism and amino acid metabolism (particularly that involves cysteine and glutamine), contribute to ferroptosis (Gao and Jiang, 2018; Stockwell et al., 2017). Interestingly, the stress-responsive catabolic pathway, autophagy, can also promote ferroptosis by degrading iron-storage protein ferritin and thus increase cellular iron concentration (Gao et al., 2016; Hou et al., 2016). Iron, as an essential cofactor of a plethora of metabolic enzymes and as a catalyst of lipid peroxide-generating Fenton reaction, drives oxygen and redox-based metabolism and cellular ROS production. Despite the central role of mitochondria in oxidative metabolism, it remains unclear whether mitochondria play a central role in ferroptosis. Supporting this possibility, ferroptosis is associated with dramatic morphological changes of mitochondria, including mitochondrial fragmentation and cristae enlargement
354 Molecular Cell 73, 354–363, January 17, 2019 ª 2018 Elsevier Inc.
(Dixon et al., 2012; Doll et al., 2017), and some potent ferroptosis inhibitors appear to be exquisitely targeted to mitochondria (Krainz et al., 2016). Furthermore, glutamine metabolism, known as glutaminolysis, is required for cysteine-deprivation-induced (CDI) ferroptosis (Gao et al., 2015a, 2015b), and one of the major functions of glutaminolysis is to fuel the mitochondrial tricarboxylic acid (TCA) cycle. Intriguingly, mitochondrial glutaminase (GLS), GLS2, instead of cytosolic GLS1, has been shown to be required for ferroptosis, although both enzymes catalyze glutaminolysis (Cassago et al., 2012; Gao et al., 2015a; Jennis et al., 2016). All these observations are consistent with the potential involvement of mitochondria in ferroptosis. However, there is also strong evidence arguing against a major role of mitochondria in ferroptosis. For example, a comparison between a mitochondrial DNA-depleted (r0) cancer cell line and its parental line did not show a significant difference in ferroptosis sensitivity (Dixon et al., 2012), and a mechanistic investigation into a cohort of ferrostatin analogs (inhibitors of ferroptosis) failed to establish a correlation between mitochondrial localization of ferrostatins with their anti-death potency (Gaschler et al., 2018). Taken together, the functional relevance of mitochondria in ferroptosis is still highly debatable. In this study, through a series of cellular, molecular, pharmacological, and metabolomic analyses, we demonstrated that the mitochondrion is a crucial player in ferroptosis induced by cysteine deprivation (inhibition of system Xc by erastin or using culture medium free of cystine). Mechanistically, the canonical metabolic activity of mitochondria, including both the TCA cycle and mitochondrial electron transport chain (ETC) activity, are required for the generation of sufficient lipid ROS to initiate ferroptosis. Furthermore, cancer cells deficient of the mitochondrial tumor suppressor fumarate hydratase (FH) are resistant to CDI ferroptosis—this result supports the notion that ferroptosis might be a physiologically relevant tumor suppressive mechanism and provides insights into potential ferroptosis-inducing cancer therapeutic approaches. Importantly, the role of mitochondria in ferroptosis is context dependent; if the activity of the glutathione-dependent peroxidase (GPX4) is inhibited, cells undergo ferroptosis independent of mitochondrial function. RESULTS Mitochondria Regulate Cysteine-Deprivation-Induced Ferroptosis To determine unambiguously whether mitochondria play a role in ferroptosis, we engineered human fibrosarcoma HT1080 cells depleted of mitochondria. Mitochondrial depletion was achieved by activating mitochondria-targeted autophagy, known as mitophagy (Youle and Narendra, 2011). First, we treated control and Parkin (tagged with mCherry)-overexpressing HT1080 cells with the mitochondrial uncoupler carbonyl cyanide m-chlorophenyl hydrazine (CCCP). Under this condition, mitochondria are depolarized by CCCP and consequently subjected to clearance via Parkin-mediated mitophagy (Narendra et al., 2008). As shown in Figures 1A and 1B, upon CCCP treatment for 24 or 48 hr followed by recovery in CCCP-free culture medium for 24 hr, mitochondria were significantly eliminated in Parkinoverexpressed cells (see the disappearance of MitoTracker
signal and mitochondrial proteins TOM20 and TIM23). Subsequently, cells were incubated with cystine (CC)-free medium or a pharmacological inducer of ferroptosis, erastin. Compared to control cells, Parkin-overexpressing, mitochondria-depleted cells were less sensitive to ferroptosis triggered by CC starvation or erastin (Figure 1C), confirming that mitochondria promote CDI ferroptosis. Lipid ROS accumulation is a hallmark of ferroptosis, but where and how it is generated during ferroptosis have not been defined. As the mitochondrion is a major organelle for cellular ROS production, does this organelle contribute to lipid ROS generation upon cysteine deprivation? To test this possibility, we monitored lipid ROS accumulation using C11 BODIPY 581/591, a fluorescent probe for membrane-localized ROS. Indeed, upon CC starvation or erastin treatment, mitochondria-depleted cells had a decreased level of lipid ROS accumulation (Figure 1D). Further, when we examined subcellular localization of the lipid ROS probe by confocal imaging (the probe changes fluorescence from red to green upon oxidation), we found that, in both HT1080 cells and mouse embryonic fibroblasts (MEFs) treated with erastin, the oxidized probe first appeared in a distribution that significantly colocalized with mitochondria, then colocalized with the plasma membrane at later time points (Figures 1E and S1). Mitochondrial TCA Cycle Participates in CysteineDeprivation-Induced Ferroptosis Glutaminolysis has been demonstrated to be required for CDI ferroptosis (Gao et al., 2015a). In the absence of glutamine, neither CC starvation nor pharmacological inhibition of CC transporter subunit SLC7A11 by erastin can induce ferroptosis. As a major function of glutaminolysis is to maintain the mitochondrial TCA cycle through anaplerosis (Figure 2A), is the TCA cycle required for ferroptosis? To test this possibility, we examined whether TCA cycle metabolites downstream of glutaminolysis can recapitulate the function of glutamine in ferroptosis. Indeed, alphaKetoglutaric acid (aKG), which is the TCA cycle metabolite immediately downstream of glutaminolysis, can replace glutamine for CDI lipid ROS accumulation and ferroptosis (Figures 2B and 2C). Further, TCA metabolites downstream of aKG, including succinate (Suc), fumarate (Fum), and malate (Mal), can all replace the role of glutamine in lipid ROS accumulation and ferroptosis induced by CC starvation or system Xc inhibition in both MEFs and HT1080 cells (Figures 2D–2F and S2). To further confirm that glutamine supports ferroptosis via the TCA cycle, we measured TCA metabolites under different conditions by gas chromatography–mass spectrometry (GC/MS) metabolomic analysis. The results (Figure 2G) showed that levels of TCA cycle metabolites were not changed by CC starvation alone, within the time range of our analysis. In the absence of glutamine, levels of multiple TCA cycle metabolites, including aKG, fumarate, and malate, were significantly decreased. In contrast, TCA cycle metabolites upstream of aKG, such as citrate, were affected more by glucose status than by glutamine status. Mitochondrial Electron Transport Chain Regulates Cysteine-Deprivation-Induced Ferroptosis One of the most important functions of the TCA cycle is to support electron transport activity of protein complexes embedded Molecular Cell 73, 354–363, January 17, 2019 355
Figure 1. Mitochondria Regulate CysteineDeprivation-Induced Ferroptosis (A–D) Mitochondrial depletion by Parkin-mediated mitophagy inhibits cysteine-deprivation-induced ferroptosis. WT (control) or mCherry-Parkin overexpressing (Parkin OE) HT1080 cells were treated with 10 mM Carbonyl cyanide m-chlorophenyl hydrazine (CCCP) for 48 hr, unless otherwise indicated (as in B, 24 or 48 hr), to induce mitochondrial depletion. After recovery from CCCP treatment by incubating cells in CCCP-free normal culture medium for 24 hr, the following experiments were performed: the presence of mitochondria was assessed by MitoTracker staining (scale bar, 10 mM) (A) or by western blot for endogenous TOM20 and TIM23 (B); cells were treated with erastin or CC starvation (10% (V/V) dialyzed FBS was used for CC starvation assay medium hereafter) for 12 hr for cell death measurement (C) or 8 hr for lipid ROS measurement (D). Cell death was measured with Sytox Green staining coupled with flow cytometry. The accumulation of lipid ROS was assessed by BODIPY C11 staining coupled with flow cytometry analysis. All quantitative data are presented as mean ± SD from three independent experiments, and p values were calculated with unpaired t test. **p < 0.01, ***p < 0.001, ****p < 0.0001. (E) Colocalization of oxidized lipid and mitochondria. MEFs were treated as indicated for 4 hr, then cells were co-stained with BODIPY C11 and MitoTracker. Oxidized BODIPY C11 (green) indicating lipid ROS and mitochondrial signals (red) were imaged by fluorescent microscope. Scale bar, 10 mm. See also Figure S1.
in the inner-membrane of mitochondria (Figure 3A). Since the TCA cycle is required for ferroptosis induced by cellular cysteine deprivation, are components of the ETC also required? To address this question, we utilized an array of ETC inhibitors. As shown in Figures 3B–3E, inhibitors of mitochondrial complex I (rotenone), complex II (DBM), complex III (antimycin), and complex IV (NaN3) all suppressed lipid ROS accumulation and ferroptosis induced by CC starvation or erastin. Taken together, these results indicate that ETC contributes to ferroptosis induced by cellular cysteine deprivation. Notably, although all tested ETC inhibitors significantly blocked ferroptosis at earlier time points (12 hr), some of them (antimycin and NaN3) maintained their inhibitory effect at later time points, but others (inhibitors for complex I and complex II) gradually lost such effect (Figure S3). This might be due to different stability of these compounds; alternatively, it may be that inhibition of complex I or complex II alone is not sufficient to block electron transfer to the downstream complex III. Mitochondrial Membrane Potential Hyperpolarization Is Associated with Cysteine-Deprivation-Induced Lipid ROS Accumulation and Ferroptosis Mitochondria produce ATP by utilizing the proton electrochemical gradient potential across the mitochondrial membrane, 356 Molecular Cell 73, 354–363, January 17, 2019
which is the result of the action of the TCA cycle and ETC. As the TCA cycle and ETC promote CDI ferroptosis, is ferroptosis associated with a change of mitochondrial membrane potential (MMP)? To test this possibility, we used MitoTracker (tetramethylrhodamine ethyl ester, or TMRE) staining to monitor MMP, upon induction of ferroptosis and as a control, of apoptosis. As expected, during mitochondria-mediated apoptosis, a decrease of MMP was observed prior to eventual cell death (Figure S4A; Video S1). This decrease of MMP is indicative of mitochondrial outer membrane permeabilization (MOMP), a crucial characteristic of mitochondria-mediated apoptosis (Green and Reed, 1998; Kroemer and Reed, 2000). In striking contrast, ferroptosis inducers, such as CC starvation (Figures 4A and 4B; and Video S2), amino acid-free medium plus full serum (Gao et al., 2015a) (Figure S4B; Video S3), erastin (Figure S4C), or glutamate (Figure S4D), can all induce MMP hyperpolarization. To further investigate the role of MMP in CDI ferroptosis, we examined the effect of the mitochondrial uncoupler CCCP on ferroptosis. As shown in Figures 4C, 4E, S4E, and S4F, CCCP disrupted MMP and completely blocked CDI lipid ROS accumulation and ferroptosis. Moreover, proliferation can be recovered after removing both erastin and CCCP, as assessed by cell colony formation assay (Figure 4F), indicating that temporarily disrupting MMP by CCCP can prevent ferroptotic cell death without
Figure 2. Mitochondrial TCA Cycle Promotes Cysteine-Deprivation-Induced Ferroptosis (A) An overview of mammalian tricarboxylic acid cycle. (B and C) a-ketoglutarate can mimic the death-inducing activity of L-Glutamine (L-Gln). (B) Microscopy showing cell death. MEFs were treated as indicated for 10 hr. Upper panel: phase-contrast; lower panel: propidium iodide (PI) staining for dead cells (Scale bar, 100 mM). (C) For cell death measurement, MEFs were subjected to the same treatment as in (B), and cell death was determined by propidium iodide (PI) staining coupled with flow cytometry; for lipid ROS measurement, MEFs were treated as in (B) for 8 hr and then lipid ROS was determined by BODIPY C11 staining coupled with flow cytometry. (D–F) TCA metabolites succinate (D), fumarate (E), and malate (F) can mimic the death inducing activity of L-Gln. (G) Glutamine is required for the maintenance of the normal level of multiple TCA cycle metabolites during ferroptosis. MEFs were treated as indicated for 8 hr, and samples were collected and measured by GC/MS metabolomic analysis as detailed in STAR Methods. For all experiments, 10% (V/V) dialyzed FBS was used in the assay medium. Q, L-glutamine (1 mM); CC, L-cystine (0.2 mM); aKG, Dimethyl-a-2-oxoglutarate (mM); Suc, Dimethyl succinate (mM); Fum, Dimethyl fumarate (mM); Mal, Dimethyl (S)-( )-malate (mM); and Fer-1, ferrostatin-1 (1 mM). All quantitative data are presented as mean ± SD from three independent experiments. **p < 0.01, ***p < 0.001, ****p < 0.0001 by unpaired Student’s t test. See also Figure S2.
compromising long-term cell viability. This conclusion is consistent with the effect of pharmacological inhibition of ETC on ferroptosis (Figures 3 and S4G–S4J). Glutaminolysis and TCA Cycle Drive FerroptosisAssociated MMP Hyperpolarization We subsequently determined the role of glutaminolysis and the TCA cycle in ferroptosis-associated MMP hyperpolarization. In the absence of glutamine, CC starvation failed to induce MMP hyperpolarization (Figure 4G). Similarly, inhibition of glutaminolysis by aminooxyacetic acid (AOA) blocked CC starvation-induced MMP hyperpolarization (Figure 4H). Therefore, glutamine metabolism is essential for CC starvation-induced MMP hyperpolarization. Furthermore, TCA cycle intermediates downstream of glutaminolysis, including aKG, succinate, fumarate, and malate, were all able to substitute glutamine for CC starvation-induced MMP hyperpolarization (Figures 4I–4L). Therefore, glutaminolysis and TCA cycle are essential for ferroptosis-associated MMP hyperpolarization. Presumably, MMP hy-
perpolarization reflects an increase of mitochondrial ETC activity and subsequent lipid ROS generation, which explains its role in ferroptosis. However, the mechanisms by which cysteine deprivation triggers MMP hyperpolarization are not clear and warrant further investigation. Mitochondrial Function Is Dispensable for GPX4 Inhibition-Induced Ferroptosis Importantly, although glutaminolysis and mitochondrial TCA cycle/ETC activity contribute to CDI ferroptosis, ferroptosis can still be triggered by pharmacological inhibition or genetic elimination of GPX4 when these mitochondrial activities are disrupted. Mitochondria-depleted HT1080 cells did not show significant resistance when ferroptosis was triggered by the GPX4 inhibitor RSL3 (Figures 5A and 5B). ETC inhibitors also failed to impinge on lipid ROS accumulation and ferroptosis upon RSL3 treatment (Figures 5C, 5D, and S5A). Further, we generated GPX4-knockout (KO) HT1080 cells by CRIPSR/Cas9 technology and found that ETC inhibitors could not block ferroptosis Molecular Cell 73, 354–363, January 17, 2019 357
MMP hyperpolarization (Figure S5B) without inhibiting RSL3induced lipid ROS accumulation or ferroptosis in both MEFs and HT1080 cells (Figures 5C, 5D, S5A, and S5B). Moreover, we found that GPX4 inhibition-induced ferroptosis can occur even in the absence of glutamine (Figure 5G). Therefore, glutamine and mitochondria are dispensable for GPX4 inhibitioninduced ferroptosis.
Figure 3. ETC Activity Promotes Cysteine-Deprivation-Induced Ferroptosis (A) Scheme shows mitochondrial electron transport chain (ETC) complexes and their inhibitors. (B) Inhibitors of ETC can inhibit CC starvation-induced ferroptosis. MEFs were treated with CC starvation for 12 hr in the absence or presence of ETC inhibitors as indicated. Cell death was determined by PI staining coupled with flow cytometry. Complex I inhibitor rotenone (Rot), 10 mM; Complex II inhibitor diethyl butylmalonate (DBM), 2 mM; Complex III inhibitor antimycin A (Anti A), 50 mM; and Complex IV inhibitor NaN3, 15 mM. (C) ETC inhibitors can inhibit CC starvation-induced lipid ROS accumulation. MEFs were treated with CC for 6 hr in the absence or presence of ETC inhibitors as indicated. Accumulation of lipid ROS was determined by BODIPY C11 staining coupled with flow cytometry. (D) ETC inhibitors can inhibit erastin-induced ferroptosis. MEFs were treated with 1 mM erastin for 12 hr in the absence or presence of ETC inhibitors as indicated. Cell death was determined by PI staining coupled with flow cytometry. (E) ETC inhibitors can inhibit erastin-induced lipid ROS accumulation. MEFs were treated with 1 mM erastin for 6 hr in the absence or presence of ETC inhibitors as indicated. Accumulation of lipid ROS was determined by BODIPY C11 coupled with flow cytometry. All quantitative data are presented as mean ± SD from three independent experiments. ***p < 0.001, ****p < 0.0001 by unpaired Student’s t test. See also Figure S3.
induced by GPX4-KO (Figures 5E–5G). Consistently, although RSL3 treatment can induce MMP hyperpolarization, the mitochondrial uncoupler CCCP can completely ablate RSL3-induced 358 Molecular Cell 73, 354–363, January 17, 2019
Loss of Function of Mitochondrial Tumor Suppressor Fumarate Hydratase Confers Resistance to Ferroptosis in Renal Cancer Cells Genetic mutation of the gene encoding for the TCA cycle enzyme fumarate hydratase (FH, also known as fumarase) has been identified in benign and malignant renal cancer lesions and has been demonstrated to be a bona fide tumor suppressor in renal cancer (Alam et al., 2005; Chuang et al., 2005; Gottlieb and Tomlinson, 2005; King et al., 2006; Tomlinson et al., 2002). In light of our finding that mitochondrial TCA cycle and respiration drive CDI ferroptosis, we examined whether loss of FH function renders cells more resistant to ferroptosis, thus contributing to its tumor suppressive function. We tested this possibility in two pairs of patient-derived isogenic renal cancer cell lines, FH mutant UOK262 and UOK262 reconstituted with wild-type (WT) FH and FH mutant UOK268 and UOK268 reconstituted with WT FH (Tong et al., 2011). Compared to cells expressing WT FH, FH mutant renal cancer cells were less sensitive to CC starvationinduced ferroptosis (Figures 6A–6C), and cell death in these mutant cells could be restored by supplying exogenous malate, which is the metabolite downstream of FH in the TCA cycle (Figures 6A and 6B). Importantly, in a cell colony formation experiment, FH mutant renal cancer cells remained viable and proliferative upon CC starvation, whereas the isogenic cells expressing WT FH failed to proliferate under this condition (Figure 6D). These results suggest that under oxidative stress (common for growth environment of solid tumors), loss of FH function can make cancer cells more resistant to ferroptotic death, thus conferring tumorigenic advantage. DISCUSSION Cellular metabolism and ferroptosis closely interact with one another, and lipid ROS, mainly a product of oxygen/iron-driven metabolism, is essential for the execution of ferroptosis. For these reasons, it appears logical that mitochondria should play a central role in ferroptosis. Nonetheless, whether the mitochondria are an important component in ferroptosis remains highly controversial. In this study, we present evidence to demonstrate that the mitochondrion is indeed a crucial player in ferroptotic cell death induced by cysteine deprivation. When mitochondria are depleted via Parkin-dependent mitophagy, cells become more resistant to ferroptosis upon CC starvation or pharmacological inhibition of CC import (Figure 1). Mechanistically, we showed that the role of the mitochondrion in ferroptosis is due to its metabolic function, and both mitochondrial TCA cycle and the action of ETC are required for a potent ferroptosis (Figures 2, 3, S2, and S3). Ferroptotic function of TCA cycle is the reason why glutaminolysis, a major source of anaplerosis, is required for ferroptosis (Figures 2 and S2).
Figure 4. Mitochondrial Membrane Potential Hyperpolarization Is Associated with Cysteine-Deprivation-Induced Lipid ROS Accumulation and Ferroptosis (A) Ferroptosis is accompanied by mitochondrial membrane potential (MMP) hyperpolarization. Representative still images from time-lapse imaging of MMP in MEFs undergoing CC starvationinduced ferroptosis. Cells were incubated with TMRE (200 nM) and subjected to CC starvation for indicated time (scale bar, 5 mM). (B) Quantification of MMP during CC starvationinduced ferroptosis. MEFs were treated with CC starvation, and 500 nM TMRE was added 30 min before each indicated time point. MMP was measured by flow cytometry and mean fluorescence was calculated. (C) CCCP can inhibit ferroptosis-associated MMP hyperpolarization. MEFs were treated with CC starvation plus or minus 10 mM CCCP for indicated time, and 500 nM TMRE was added 30 min before each indicated time point. MMP was measured by flow cytometry and mean fluorescence was calculated. (D) CCCP can block ferroptosis induced by CC starvation or erastin. MEFs were treated as indicated for 12 hr and cell death was determined by PI staining coupled with flow cytometry. (E) CCCP can block lipid ROS accumulation induced by CC starvation or erastin. MEFs were treated as indicated for 8 hr, and lipid ROS was measured by BODIPY C11 staining coupled with flow cytometry. (F) Cells protected by CCCP from erastininduced ferroptosis maintain viability. MEFs and HT1080 cells were seeded at low density in 12-well dishes and treated with erastin for 16 hr in the absence or presence of CCCP as indicated. Cells were then allowed to recover in drug-free DMEM medium for 3 days and subsequently fixed and stained with crystal violet. (G and H) Glutaminolysis and TCA cycle drive ferroptosis-associated MMP hyperpolarization in MEFs. (G) Glutamine is required for CC starvation-induced MMP hyperpolarization. (H) Transaminase inhibitor AOA (Aminooxyacetic acid, 0.5 mM) can block CC starvation-induced MMP hyperpolarization. (I–L) TCA intermediates a-ketoglutarate (I) succinate (J), fumarate (K), and malate (L) can replace the requirement of glutamine for CC starvation-induced MMP hyperpolarization. aKG, Dimethyl-a-2-oxoglutarate (8 mM); Suc, Dimethyl succinate (8 mM); Fum, Dimethyl fumarate (5 mM); and Mal, Dimethyl (S)-(-)-malate (32 mM). All quantitative data are presented as mean ± SD from three independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 by unpaired Student’s t test. See also Figure S4 and Videos S1, S2, and S3.
Why, then, is the role of mitochondria in ferroptosis so controversial? The following three reasons can reconcile this discrepancy. First, our study indicates that the role of mitochondria in ferroptosis is context dependent: blockage of mitochondrial function potently inhibits CDI ferroptosis; however, upon elimination or pharmacological inhibition of GPX4, the very downstream component of the ferroptosis pathway responsible for lipid ROS clearance, cells can commit ferroptosis independently
of mitochondria (Figures 5 and S5). When cysteine is scarce, mitochondrial metabolism contributes significantly to rapid glutathione depletion and subsequent lipid ROS generation and ferroptosis. On the other hand, upon GPX4 inhibition, although one would predict that inhibition of mitochondria should also attenuate lipid ROS generation and thus may at least delay ferroptosis, we did not observe a measurable effect of mitochondrial ETC inhibitors on slowing down ferroptosis in Molecular Cell 73, 354–363, January 17, 2019 359
Figure 5. Mitochondria Are Dispensable for GPX4 InhibitionInduced Ferroptosis (A and B) Mitochondrial depletion by Parkin-mediated mitophagy cannot inhibit RSL3-induced ferroptosis. Mitochondria-depleted HT1080 cells were created as described in Figure 1. Cells were treated with RSL3 for 6 hr for cell death measurement (A) or 4 hr for lipid ROS measurement (B). Cell death was measured with DAPI staining coupled with flow cytometry. The accumulation of lipid ROS was assessed by BODIPY C11 staining coupled with flow cytometry analysis. (C and D) Mitochondrial ETC inhibitors cannot inhibit RSL3-induced ferroptosis (C) and lipid ROS accumulation (D). (E) Western blotting confirmed CRISPR-Cas9-mediated knockout of GPX4 in HT1080 cells (see STAR Methods for detail). (F) Mitochondrial ETC inhibitors cannot inhibit GPX4 knockout-induced ferroptosis. GPX4 KO HT1080 cells were seeded and subsequently grew in DMEM medium containing 0.2 mM Trolox. Ferroptosis induction was initiated by switching cells to Trolox-free medium in the presence or absence of indicated ETC complex inhibitors for 8 hr. Complex I inhibitor rotenone (Rot), 10 mM; Complex II inhibitor Diethyl butylmalonate (DBM), 2 mM; Complex III inhibitor Myxothiazol (Myxo), 15 mM; and Complex IV inhibitor NaN3, 15 mM. (G) Glutamine is dispensable for GPX4 knockout-induced ferroptosis. GPX4 KO HT1080 cells were seeded and subsequently grew in DMEM medium containing 0.2 mM Trolox. Ferroptosis induction was initiated by switching cells to Trolox-free medium in the presence or absence of 1 mM glutamine for 8 hr. Scale bar, 100 mM. All quantitative data are presented as mean ± SD from three independent experiments. ns, non-significant by unpaired Student’s t test. See also Figure S5.
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GPX4-KO HT1080 cells (Figure S5D). It is likely that once GPX4 is eliminated, the low amount of lipid ROS produced by other mechanisms will be rapidly amplified through the Fenton chain reaction, leading to full-blown ferroptotic cell death even when mitochondrial activity is diminished. It is also possible that GPX4 inactivation may transduce a signal to, and thus hyperactivate, certain enzymes responsible for lipid ROS generation. Future investigation is needed to clarify this issue. Second, since ferroptosis is dictated by cellular metabolism, long-term alteration of cellular metabolism may rewire the molecular processes underlying ferroptosis. For example, in a type of experimental model cells known as r0 cells, mitochondrial DNA is depleted permanently (King and Attardi, 1989, 1996). To compensate the long-term lack of mitochondria, these cells need to be cultured with nutrients different from mitochondria-containing cells (King and Attardi, 1989, 1996). Therefore, metabolism in r0 cells, including lipid ROS generation (mainly a consequence of metabolism), is fundamentally rewired. For this reason, the mechanism governing the mitochondria-independent ferroptosis in r0 cells may differ significantly from that in normal, mitochondria-containing cells. Third, the methods for cell death measurement need to be carefully considered. For example, although cell viability assays are often used to infer cell death, they are not suitable for the study of the role of mitochondria in ferroptosis. This is because that manipulation of mitochondrial function may impact the outcome of these assays (many of them are ATP content based or metabolic activity based), regardless of cell death status. Assays directly measuring cell death are preferable for this purpose. Mitochondria are also involved in other modalities of programmed cell death, particularly apoptosis. Intriguingly, the roles of mitochondria in apoptosis and ferroptosis are fundamentally different. In mitochondria-mediated apoptosis, the mitochondrion serves as a reservoir, storing apoptosis-regulatory proteins such as cytochrome c and Smac, which, upon release to the cytoplasm, promote caspase activation (Jiang and Wang, 2004; Wang, 2001). Notably, the cytosolic apoptotic activities of these mitochondrial proteins are independent of, and can be molecularly dissected from, their mitochondrial functions. As such, a compromise of mitochondrial integrity and loss of MMP, known as MOMP, precedes the release of these proteins and subsequent caspase-9 activation. Interestingly, in CDI ferroptosis, it is the canonical metabolic function of mitochondria that actively contributes to lipid ROS generation, a prerequisite of ferroptosis, and inhibition of TCA cycle or ETC function, or disruption of MMP, can all abrogate ferroptosis under this condition. Strikingly, a transient hyperpolarization of MMP can be observed during CDI ferroptosis, followed by the eventual collapse of MMP (Figures 4 and S4; Videos S1, S2, and S3). This unique feature is indicative of the crucial role of mitochondrial metabolic activity in CDI ferroptosis. Taken together, while mitochondria play a passive role in apoptosis by storing various pro-apoptotic proteins, they play a crucial and proactive role in CDI ferroptosis by fueling metabolism and lipid ROS production. Further, the findings presented in this work provide novel mechanistic insights into some enigmatic observations in cancer biology. FH, a metabolic enzyme in TCA cycle, has
Figure 6. Loss of Function of Mitochondrial Tumor Suppressor Fumarate Hydratase Confers Resistance to Ferroptosis (A–C) Fumarase (FH) mutant cancer cells are more resistant to CC starvation-induced ferroptosis than isogenic cells expressing WT FH. (A) Microscopy showing cell death. Two pairs of FH mutant cancer cells and their isogenic cells expressing WT FH were treated as indicated for 36 hr (UOK262 and UOK262+WT FH) or 60 hr (UOK268 and UOK268+WT FH). Upper panel: phase-contrast; lower panel: PI staining for dead cells (scale bar, 100 mM). Western blot shows the expression level of FH in these cell lines. (B) Cell death was determined by PI staining coupled with flow cytometry. (C) Lipid ROS was determined by BODIPY C11 staining coupled with flow cytometry. (D) FH mutant cancer cells recovered and proliferated after CC starvation. Cells were treated with CC starvation for 24 hr (UOK262 and UOK262+WT FH) or 60 hr (UOK268 and UOK268+WT FH), then cells were incubated in full DMEM medium for 3 days. Colony formation was measured by crystal violet staining. All quantitative data are presented as mean ± SD from three independent experiments. ****p < 0.0001 by unpaired Student’s t test. See also Figure S6.
been demonstrated to be a tumor suppressor. This is counterintuitive because one would assume that defects in the TCA cycle or ETC would be disadvantageous for cancer cell growth. Indeed, under normal culture condition, reconstitution of WT FH into UOK262/268 cancer cells rendered them proliferating better (Figure S6). So, what is the molecular basis underlying the tumor suppressive activity of FH? A pseudo-hypoxic pathway was proposed to be the link between FH mutation and tumorigenesis. FH defect leads to aberrant accumulation of fumarate, which, in turn, inactivates prolyl hydroxylases and thus stabilizes hypoxia-inducible factor (HIF)-1a (Isaacs et al., 2005; Selak et al., 2005). However, both HIF-1a and HIF-2a are
dispensable for the formation of hyperplastic renal cysts in mice lacking expression of FH in the kidney (Adam et al., 2011), suggesting additional tumorigenic consequences caused by FH mutation. Our finding that loss of FH function renders cells more resistant to CDI ferroptosis (Figure 6) provides a novel mechanistic explanation of how dysfunction of FH contributes to malignancy independent of hypoxic signaling—we hypothesize that stronger resistance to ferroptosis caused by FH dysfunction contributes to tumorigenesis, although this is most likely not sufficient on its own. It remains to be determined why FH mutation preferentially drives tumorigenesis only in certain specific tissue types. Conceptually, our finding that ferroptosis contributes to the tumor suppressive function of FH is important. It has been reported recently that p53 can suppress cancer development via its ferroptosis-promoting activity even when it loses its function to promote apoptosis, senescence, and growth arrest (Jiang et al., 2015; Wang et al., 2016). Based on these novel findings about FH and p53, we hypothesize that ferroptosis is a natural tumor suppressive mechanism under diverse biological conditions. The significance of this hypothesis in cancer biology is obvious. As for the biology of ferroptosis, tumor suppression might represent an authentic physiological role of this unique modality of cell death, which, until now, remains to be defined. Molecular Cell 73, 354–363, January 17, 2019 361
STAR+METHODS
Angeli, J.P.F., Shah, R., Pratt, D.A., and Conrad, M. (2017). Ferroptosis inhibition: mechanisms and opportunities. Trends Pharmacol. Sci. 38, 489–498.
Detailed methods are provided in the online version of this paper and include the following:
Carey, B.W., Finley, L.W., Cross, J.R., Allis, C.D., and Thompson, C.B. (2015). Intracellular a-ketoglutarate maintains the pluripotency of embryonic stem cells. Nature 518, 413–416.
KEY RESOURCES TABLE CONTACT FOR REAGENT AND RESOURCE SHARING METHOD DETAILS B Cell culture B Measurement of mitochondrial membrane potential B Measurement of lipid ROS B CRISPR-mediated GPX4 deletion B Fluorescence microscopy B Metabolite analysis by gas chromatography-mass spectrometry (GC-MS) QUANTIFICATION AND STATISTICAL ANALYSIS DATA AND SOFTWARE AVAILABILITY
Cassago, A., Ferreira, A.P., Ferreira, I.M., Fornezari, C., Gomes, E.R., Greene, K.S., Pereira, H.M., Garratt, R.C., Dias, S.M., and Ambrosio, A.L. (2012). Mitochondrial localization and structure-based phosphate activation mechanism of Glutaminase C with implications for cancer metabolism. Proc. Natl. Acad. Sci. USA 109, 1092–1097.
d d d
d d
SUPPLEMENTAL INFORMATION Supplemental Information includes six figures and three videos and can be found with this article online at https://doi.org/10.1016/j.molcel.2018.10.042. ACKNOWLEDGMENTS The authors thank Dr. W. Marston Linehan for kindly providing the FH-mutant UOK262 and UOK268 cell lines, as well as their wild-type FH reconstituted counterparts. The authors thank members of the Jiang lab and the Thompson lab for critical reading and suggestions. This work is supported by NIH R01CA204232 (to X.J.), NIH R01CA201318 (to C.B.T.), a Geoffrey Beene Cancer Research fund (to X.J.), National Natural Science Foundation of China 31871388 (to M.G.), a Leukemia and Lymphoma Society Fellowship (to J.Z.), and NIH T32 fellowship 5T32GM008539-23 (to A.M.M.). This work is also supported by NCI cancer center core grant P30CA008748 to Memorial Sloan Kettering Cancer Center. AUTHOR CONTRIBUTIONS M.G. and X.J. conceived and supervised the study. M.G. and J.Y. performed most of the experiments in collaboration with J.Z., A.M.M., and P.M. M.G., J.Y., C.B.T., and X.J. analyzed the data and wrote the paper with suggestions from other authors. All authors reviewed the paper.
Chen, L., Hambright, W.S., Na, R., and Ran, Q. (2015). Ablation of the ferroptosis inhibitor glutathione peroxidase 4 in neurons results in rapid motor neuron degeneration and paralysis. J. Biol. Chem. 290, 28097–28106. Chuang, G.S., Martinez-Mir, A., Geyer, A., Engler, D.E., Glaser, B., CserhalmiFriedman, P.B., Gordon, D., Horev, L., Lukash, B., Herman, E., et al. (2005). Germline fumarate hydratase mutations and evidence for a founder mutation underlying multiple cutaneous and uterine leiomyomata. J. Am. Acad. Dermatol. 52, 410–416. Dixon, S.J., Lemberg, K.M., Lamprecht, M.R., Skouta, R., Zaitsev, E.M., Gleason, C.E., Patel, D.N., Bauer, A.J., Cantley, A.M., Yang, W.S., et al. (2012). Ferroptosis: an iron-dependent form of nonapoptotic cell death. Cell 149, 1060–1072. Do Van, B., Gouel, F., Jonneaux, A., Timmerman, K., Gele´, P., Pe´trault, M., Bastide, M., Laloux, C., Moreau, C., Bordet, R., et al. (2016). Ferroptosis, a newly characterized form of cell death in Parkinson’s disease that is regulated by PKC. Neurobiol. Dis. 94, 169–178. Doll, S., Proneth, B., Tyurina, Y.Y., Panzilius, E., Kobayashi, S., Ingold, I., Irmler, M., Beckers, J., Aichler, M., Walch, A., et al. (2017). ACSL4 dictates ferroptosis sensitivity by shaping cellular lipid composition. Nat. Chem. Biol. 13, 91–98. Friedmann Angeli, J.P., Schneider, M., Proneth, B., Tyurina, Y.Y., Tyurin, V.A., Hammond, V.J., Herbach, N., Aichler, M., Walch, A., Eggenhofer, E., et al. (2014). Inactivation of the ferroptosis regulator Gpx4 triggers acute renal failure in mice. Nat. Cell Biol. 16, 1180–1191. Galluzzi, L., Bravo-San Pedro, J.M., and Kroemer, G. (2015). Ferroptosis in p53-dependent oncosuppression and organismal homeostasis. Cell Death Differ. 22, 1237–1238. Gao, M., and Jiang, X. (2018). To eat or not to eat-the metabolic flavor of ferroptosis. Curr. Opin. Cell Biol. 51, 58–64. Gao, M., Monian, P., Quadri, N., Ramasamy, R., and Jiang, X. (2015a). glutaminolysis and transferrin regulate ferroptosis. Mol. Cell 59, 298–308. Gao, M., Monian, P., and Jiang, X. (2015b). Metabolism and iron signaling in ferroptotic cell death. Oncotarget 6, 35145–35146.
DECLARATION OF INTERESTS
Gao, M., Monian, P., Pan, Q., Zhang, W., Xiang, J., and Jiang, X. (2016). Ferroptosis is an autophagic cell death process. Cell Res. 26, 1021–1032.
C.B.T. is a founder of Agio Pharmaceuticals and a member of its scientific advisory board. Agio does not hold financial interests in the work reported in this paper.
Gaschler, M.M., Hu, F., Feng, H., Linkermann, A., Min, W., and Stockwell, B.R. (2018). Determination of the subcellular localization and mechanism of action of ferrostatins in suppressing ferroptosis. ACS Chem. Biol. 13, 1013–1020.
Received: July 6, 2018 Revised: September 16, 2018 Accepted: October 29, 2018 Published: December 20, 2018 REFERENCES Adam, J., Hatipoglu, E., O’Flaherty, L., Ternette, N., Sahgal, N., Lockstone, H., Baban, D., Nye, E., Stamp, G.W., Wolhuter, K., et al. (2011). Renal cyst formation in Fh1-deficient mice is independent of the Hif/Phd pathway: roles for fumarate in KEAP1 succination and Nrf2 signaling. Cancer Cell 20, 524–537. Alam, N.A., Olpin, S., Rowan, A., Kelsell, D., Leigh, I.M., Tomlinson, I.P., and Weaver, T. (2005). Missense mutations in fumarate hydratase in multiple cutaneous and uterine leiomyomatosis and renal cell cancer. J. Mol. Diagn. 7, 437–443.
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Molecular Cell 73, 354–363, January 17, 2019 363
STAR+METHODS KEY RESOURCES TABLE
REAGENT or RESOURCE
SOURCE
IDENTIFIER
g-tubulin
Sigma
Cat# T6557; RRID:AB_477584
Tom20
Santa Cruz
Cat#SC11415; RRID:AB_2207533
Tim23
Cell Signal
Cat# 3130S; RRID:AB_10693298
Fumarase
Cell Signal
Cat#4567; RRID:AB_11178522
Vinculin
Cell Signal
Cat#13901; RRID:AB_2728768
Erastin
Sigma
Cat#E7781
RSL3
Selleckchem
Cat# S8155
Dimethyl 2-oxoglutarete
Sigma
Cat#349631
Dimethyl succinate
Sigma
Cat#W239607
Dimethyl fumarate
Sigma
Cat#242926
Dimethyl (S)-(-)-malate
Sigma
Cat#374318
Carbonyl cyanide m-chlorophenyl hydrazone
Sigma
Cat#C2759
Rotenone
Sigma
Cat#R8875
Antibodies
Chemicals and Reagents
Diethyl butylmalonate
Sigma
Cat#112038
Antimycin A
Sigma
Cat#A8674
Sodium azide
Sigma
Cat# S2002
Oligomycin A
Sigma
Cat#75351
Myxothiazol
Sigma
Cat# T5580
Ferrostatin-1
Sigma
Cat# M60042
TMRE
Enzo Life Sciences
Cat#ENZ-52309
BODIPY C11
Thermo Fisher Scientific
Cat# D3861
Lipofectamine 2000
Life Technologies
Cat# 11668027
Propidium Iodide Staining Solution
BD PharMingen
Cat# 556463
SYTOX Green Nucleic Acid Stain
Thermo Fisher Scientific
S7020
Mendeley
https://dx.doi.org/10.17632/95zbjbj95x.1
HT1080
ATCC
N/A
HT1080 mCherry Parkin
Dr. Xuejun Jiang Lab
N/A
MEFs
ATCC
N/A
UOK262
Dr. W. Marston Linehan Lab
N/A
UOK262 WT
Dr. W. Marston Linehan Lab
N/A
UOK268
Dr. W. Marston Linehan Lab
N/A
UOK268 WT
Dr. W. Marston Linehan Lab
N/A
pLx-gRNA
Addgene
#50662
GPX4-CACGCCCGATACGCTGAGTG
NA
N/A
Deposited Data Raw Data Experimental Models: Cell Lines
Plasmid and sgRNA sequence
CONTACT FOR REAGENT AND RESOURCE SHARING Requests for resources, reagents and further information should be directed to, and will be fulfilled by Xuejun Jiang (jiangx@ mskcc.org). e1 Molecular Cell 73, 354–363.e1–e3, January 17, 2019
METHOD DETAILS Cell culture Unless specified otherwise, all mammalian cells are maintained in DMEM with high glucose, sodium pyruvate (1 mM), glutamine (2 mM), penicillin (100 U/ml), streptomycin (0.1 mg/ml) and 10% (v/v) FBS at 37 C and 5% CO2. Cell death was analyzed by PI (100 ng/ml), Sytox (5 nM) or DAPI (1 mg/ml) staining coupled with microscopy or flow cytometry. Measurement of mitochondrial membrane potential Time-lapse imaging: cells were pre-loaded with 200 nm TMRE (Enzo Life Sciences, Cat#ENZ-52309) for 1 h. Subsequently, cells were washed with PBS twice to remove traces of TMRE from the media and treated as indicated. Fluorescence and differential interference contrast (DIC) images were acquired every 5-7 min, and images were analyzed using NIS elements software (Nikon) and ImageJ software (NIH). Flow cytometry: Cells were treated as indicated, and then 0.5 mM TMRE was added and incubated for 30 min. Excess TMRE was removed by washing the cells with PBS. Labeled cells were trypsinized and resuspended in PBS plus 2% FBS. Fluorescence at Ex/ Em = 549/575 nm was analyzed using a flow cytometer. Measurement of lipid ROS Lipid ROS was analyzed by flow cytometry: Cells were seeded at a density of 2.5x105 per well in a 6-well dish and grown overnight in DMEM. 5 mM BODIPY C11 (Thermo Fisher, Cat# D3861) was added into cell culture medium and incubated for 30 min after indicated treatment. Excess BODIPY C11 was then removed by washing the cells with PBS twice. Labeled cells were trypsinized and resuspended in PBS plus 2% FBS. Oxidation of BODIPY C11 resulted in a shift of the fluorescence emission peak from 590 nm to 510 nm proportional to lipid ROS generation and was analyzed using a flow cytometer. Lipid ROS imaging: Cells were seeded at a density of 2.5x105 per well on coverslips placed in a 6-well dish and grown overnight in DMEM. Cells were washed twice in HBSS and treated as indicated. Coverslips were washed in HBSS and incubated in HBSS containing 2 mM BODIPY 581/591 C11 (Invitrogen) and 200 nM MitoTracker Deep Red FM (Invitrogen) for 20 minutes. Coverslips were then inverted onto microscope slides. Slides were imaged using a Nikon Eclipse Ti-U inverted microscope and images were processed in Photoshop (Adobe). CRISPR-mediated GPX4 deletion Guide RNA sequence CACGCCCGATACGCTGAGTG targeting human GPX4 was used. pLx-gRNA (Addgene#50662) expression plasmid encoding guide RNA was generated and co-transfected with Streptococcus pyogenes Cas9 expression plasmid and transfected into HT1080 cells in the presence of 0.2 mM Trolox using Lipofectamine 2000 (Life Technologies). 24 h after the transfection, the cells were trypsinized and about 200 cells were seeded into a 10-cm plate and cultured in the presence of 0.2 mM Trolox. After cell clones were formed and expanded, western Blot was performed to screening for GPX4 knockout clones. Fluorescence microscopy MEFs stably expressing SU9-GFP or HT1080 cells stably expressing mCherry-Parkin were grown on glass coverslips in a 6-well plate. 24 h later, cells were treated as indicated. Coverslips were then fixed with 3.7% (vol/vol) paraformaldehyde (PFA) in 20 mM HEPES pH 7.5 for 30 min at room temperature. Coverslips were then mounted on microscope slides for visualization using Nikon Eclipse Ti-U Microscope with a 60 3 magnification objective. Metabolite analysis by gas chromatography-mass spectrometry (GC-MS) Metabolite analysis by GC-MS was performed as previously described (Carey et al., 2015). In brief, cells were seeded in standard culture medium in 6-well plates and the next day were changed into experimental medium. Metabolites were extracted with icecold 80% methanol supplemented with 20 mM deuterated 2-hydroxyglutarate (D-2-hydroxyglutaric2,3,3,4,4-d5 acid (d5-2HG)) as an internal standard. After overnight incubation at 80 C, lysates were harvested and centrifuged at 21,000 g for 20 min to remove protein. Extracts were dried in an evaporator (Genevac EZ-2 Elite) and resuspended by incubation at 30 C for 90 min in 50 ml of 40 mg/ml methoxyamine hydrochloride in pyridine. Metabolites were further derivatized by addition of 80 ml of MSTFA plus 1% TMCS (Thermo Scientific) and 70 ml ethyl acetate (Sigma) and incubated at 37 C for 30 min. Samples were analyzed using an Agilent 7890A GC coupled to Agilent 5975C mass selective detector. The GC was operated in splitless mode with constant helium gas flow at 1 ml/min. One microliter of derivatized metabolites was injected onto an HP-5MS column and the GC oven temperature ramped from 60 C to 290 C in 25 min. Peaks representing compounds of interest were extracted and integrated using MassHunter software (Agilent Technologies) and then normalized to both the internal standard (d5-2HG) peak area and the relative total biomass as determined by total cell number 3 mean cell volume. Ions used for quantification of metabolite levels are as follows: d5-2HG m/z 354; Citrate, m/z 465; aKG, m/z 304; fumarate, m/z 245 and malate, m/z 335. All peaks were manually inspected and verified relative to known spectra for each metabolite.
Molecular Cell 73, 354–363.e1–e3, January 17, 2019 e2
QUANTIFICATION AND STATISTICAL ANALYSIS All statistical analyses were performed using Prism 7.0c GraphPad Software. Data are presented as mean ± SD from 3 independent experiments. p values were calculated with Student’s unpaired t test, except for the measurement of mitochondrial membrane potential for which a two-way ANOVA test was used (*p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001, see figure legend for further detail). DATA AND SOFTWARE AVAILABILITY Raw data for this paper has been deposited to Mendeley: https://dx.doi.org/10.17632/95zbjbj95x.1.
e3 Molecular Cell 73, 354–363.e1–e3, January 17, 2019