Glutaminolysis and Transferrin Regulate Ferroptosis

Abstract

Ferroptosis has emerged as a new form of regulated necrosis that is implicated in various human diseases. However, the mechanisms of ferroptosis are not well defined. This study reports the discovery of multiple molecular components of ferroptosis and its intimate interplay with cellular metabolism and redox machinery. Nutrient starvation often leads to sporadic apoptosis. Strikingly, we found that upon deprivation of amino acids, a more rapid and potent necrosis process can be induced in a serum-dependent manner, which was subsequently determined to be ferroptosis. Two serum factors, the iron-carrier protein transferrin and amino acid glutamine, were identified as the inducers of ferroptosis. We further found that the cell surface transferrin receptor and the glutamine-fueled intracellular metabolic pathway, glutaminolysis, played crucial roles in the death process. Inhibition of glutaminolysis, the essential component of ferroptosis, can reduce heart injury triggered by ischemia/reperfusion, suggesting a potential therapeutic approach for treating related diseases.
Copyright © 2015 Elsevier Inc. All rights reserved.

Full text

Article
Glutaminolysis and Transferrin Regulate Ferroptosis
Graphical Abstract
Highlights
dTransferrin and glutamine are essential for the induction of
ferroptotic cell death
dTransferrin import is required for ferroptosis
dThe cellular metabolic process glutaminolysis is essential for
ferroptosis
dInhibiting glutaminolysis reduces ischemia/reperfusion-
induced heart injury ex vivo
Authors
Minghui Gao, Prashant Monian,
Nosirudeen Quadri, Ravichandran
Ramasamy, Xuejun Jiang
Correspondence
jiangx@mskcc.org
In Brief
In this article, Gao et al. show that the
extracellular iron carrier protein
transferrin and the intracellular metabolic
process glutaminolysis are required for
the execution of a form of programmed
necrosis known as ferroptosis; targeting
glutaminolysis, presumably via inhibiting
ferroptosis, represents a potential
therapy for treating ischemia/
reperfusion-induced organ damage.
Gao et al., 2015, Molecular Cell 59, 298–308
July 16, 2015 ª2015 Elsevier Inc.
http://dx.doi.org/10.1016/j.molcel.2015.06.011

Molecular Cell
Article
Glutaminolysis and Transferrin
Regulate Ferroptosis
Minghui Gao,
1
Prashant Monian,
1
Nosirudeen Quadri,
2
Ravichandran Ramasamy,
2
and Xuejun Jiang
1,
*
1
Cell Biology Program, Memorial Sloan-Kettering Cancer Center, 1275 York Avenue, New York, NY 10065, USA
2
Department of Medicine, Langone Medical Center, New York University, 522 1
st
Avenue, New York, NY 10016, USA
*Correspondence: jiangx@mskcc.org
http://dx.doi.org/10.1016/j.molcel.2015.06.011
SUMMARY
Ferroptosis has emerged as a new form of regulated
necrosis that is implicated in various human dis-
eases. However, the mechanisms of ferroptosis are
not well defined. This study reports the discovery of
multiple molecular components of ferroptosis and
its intimate interplay with cellular metabolism and
redox machinery. Nutrient starvation often leads to
sporadic apoptosis. Strikingly, we found that upon
deprivation of amino acids, a more rapid and potent
necrosis process can be induced in a serum-depen-
dent manner, which was subsequently determined
to be ferroptosis. Two serum factors, the iron-carrier
protein transferrin and amino acid glutamine,
were identified as the inducers of ferroptosis. We
further found that the cell surface transferrin receptor
and the glutamine-fueled intracellular metabolic
pathway, glutaminolysis, played crucial roles in the
death process. Inhibition of glutaminolysis, the
essential component of ferroptosis, can reduce heart
injury triggered by ischemia/reperfusion, suggesting
a potential therapeutic approach for treating related
diseases.
INTRODUCTION
In multicellular organisms, programmed cell death, particularly
apoptosis, is frequently activated in a highly orchestrated
manner to fulfill specific physiological functions (Budihardjo
et al., 1999; Danial and Korsmeyer, 2004; Fuchs and Steller,
2011; Green and Kroemer, 2004; Thompson, 1995). Defects in
apoptosis contribute to the development of numerous human
diseases.
Apoptosis is not the only mechanism for programmed cell
death. Recent studies have led to the identification of several
other cell death processes that appear to be programmed but
distinctive from apoptosis (Bergsbaken et al., 2009; Blum
et al., 2012; Vanden Berghe et al., 2014; Yuan and Kroemer,
2010). The RIP3-dependent necrosis pathway is one such pro-
cess (Moriwaki and Chan, 2013; Vandenabeele et al., 2010).
RIP3-dependent necrosis can be triggered by tumor necrosis
factor-a(TNF-a) and is mediated by a signaling cascade
involving protein kinases RIP1 (Degterev et al., 2008) and RIP3
(Cho et al., 2009; He et al., 2009; Kaiser et al., 2011; Newton
et al., 2014; Oberst et al., 2011; Zhang et al., 2009), leading to
activation of the downstream necrotic response. To date, the
precise physiological function of RIP3-dependent necrosis has
not been unambiguously established. However, mounting evi-
dence suggests that it may benefit the organism under various
infectious or inflammatory conditions (Cho et al., 2009; He
et al., 2009; Murphy et al., 2013; Sun et al., 2012).
Recently, another form of regulated necrosis, known as fer-
roptosis, has been identified. It was shown that a synthetic com-
pound, erastin, can induce a form of non-apoptotic cell death
that requires iron (thus the name ferroptosis) (Dixon et al.,
2012; Yagoda et al., 2007). Subsequent studies demonstrate
that erastin inhibits cystine import and downstream glutathione
synthesis, leading to deregulated cellular redox homeostasis
and ultimately cell death (Dixon et al., 2012; Yang et al., 2014).
Ferroptosis inhibition has been shown to be effective in treating
diseases such as ischemia/reperfusion-induced organ damage
in experimental models (Friedmann Angeli et al., 2014; Linker-
mann et al., 2014). Further, because cancer cells harboring
oncogenic Ras appear to be more sensitive to ferroptosis induc-
tion, this form of cell death has also been explored for cancer
treatment (Yagoda et al., 2007; Yang et al., 2014). Although fer-
roptosis is strongly implicated in human diseases, currently the
precise molecular mechanisms and biological functions of fer-
roptosis remain poorly understood.
This study reports the discovery of essential components and
mechanisms for ferroptosis regulation, as well as an intimate
functional interplay between ferroptosis and cellular metabolism.
We identified transferrin and L-glutamine as extracellular regula-
tors of ferroptosis. We also demonstrated that both transferrin
transport and the cellular metabolic process glutaminolysis are
essential for ferroptosis triggered by deprivation of full amino
acids or of cystine alone. Further, we present evidence to sup-
port that glutaminolysis is a potential therapeutic target for treat-
ing heart injury caused by ischemia/reperfusion, likely due to the
essential role of glutaminolysis in ferroptosis.
RESULTS
Serum Can Induce RIP3-Independent Necrosis upon
Amino Acid Starvation
Nutrient availability is one of the key parameters for cells to make
life-or-death decisions. It has been documented that long-term
298 Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc.

deprivation of growth factors, amino acids, or glucose causes
gradual cell death (Wei et al., 2001). Although apoptotic machin-
ery is often elicited in such starvation-induced death, this never-
theless can be considered a passive death process due to failure
of the cell to survive the stressful conditions of nutrient/growth
factor deprivation.
To recapitulate cell death under nutrient/growth factor depri-
vation, we incubated mouse embryonic fibroblasts (MEFs) in
growth medium containing glucose but lacking amino acids
and serum. Modest cell death was observed after 12 hr (Fig-
ure 1A). We then incubated MEFs in amino acid-free medium
containing full serum, expecting that growth factors in serum
would mitigate cell death. Surprisingly, we observed much
more potent cell death (Figure 1A), which was further confirmed
by propidium iodide (PI) staining (Figures 1A and 1B) and mea-
surement of cellular ATP levels (Figure 1C).
We subsequently investigated the molecular nature of this
serum-induced cell death process. Cell death induced by
combined deprivation of amino acids and serum was typical
apoptosis, associated with caspase activation (Figure 1D) and
characteristic morphological changes, such as chromosomal
condensation, membrane blebbing, and formation of apoptotic
bodies (Movie S1). However, in the presence of serum, although
cell death was significantly more potent, there was no caspase
activation (Figure 1D). Consistently, use of the pan-caspase
inhibitor zVAD-FMK or deletion of bax and bak, genes essential
for mitochondria-mediated apoptosis, failed to block such
serum-induced cell death (Figures S1A and S1B). Further, the
morphological changes associated with this cell death process
were distinct from apoptosis (Figure 1E; Movies S2 and S3).
These results demonstrate that upon amino acid starvation,
serum can induce non-apoptotic cell death in MEFs.
Through live-cell time-lapse imaging, we observed that this
cell death process shared key morphological features with ne-
crosis, including cell rounding, swelling, and plasma membrane
rupture (Figure 1E; Movies S2 and S3). Recent studies have
established a programmed necrosis pathway dependent on
the protein kinase RIP3 (Cho et al., 2009; He et al., 2009; Zhang
et al., 2009). However, RIP3 gene deletion did not prevent
serum-induced cell death in MEFs, although it completely
blocked TNF-a-induced necrosis as predicted (Figures 1F, 1G,
and S1C). These results indicate that the RIP3-dependent necro-
sis pathway is not responsible for this serum-induced cell death.
Importantly, induction of potent cell death by serum under
the condition of amino acid starvation can be observed in various
types of cancerous and noncancerous cells. While amino
acid/serum double starvation could induce different levels of
death in a time- and cell type-dependent manner, addition of
serum unanimously further potentiated cell death in these cells
(Figure S2).
Figure 1. Serum Induces Potent Non-
apoptotic, RIP3-Independent Cell Death
upon Amino Acid Starvation
(A) Microscopy showing cell death induced by
serum upon amino acid starvation. MEFs were
treated as indicated for 12 hr. Upper panel: phase-
contrast, lower panel: propidium iodide (PI) stain-
ing. AA: amino acids, FBS: 10% (v/v) fetal bovine
serum.
(B) Quantitation of cell death by PI staining
coupled with flow cytometry. Cells were subjected
to the same treatment as in (A). Quantitative data
here and thereafter are presented as mean ± SEM
from three independent experiments (*p < 0.05,
**p < 0.01, ***p < 0.001 by unpaired Student’s
t test).
(C) Determination of cell viability by measuring
cellular ATP levels. Cells were subjected to the
same treatment as in (A).
(D) Serum-induced cell death is independent of
caspase activation. MEFs were treated as indi-
cated for 12 hr and caspase-3 activation was as-
sessed by immunoblotting.
(E) Serum-induced cell death shows necrotic
morphology. Representative still images from
confocal time-lapse imaging of MEFs treated with
amino acid starvation in the presence of FBS. Time
(hr) after treatment is indicated.
(F and G) Serum-induced cell death is indepen-
dent of RIP3. RIP3+/+ MEFs and RIP3/MEFs
were treated as indicated. Cell death was
monitored by phase-contract microscopy and PI
staining (F), and cell viability was determined by
measuring cellular ATP levels (G). CTRL: control.
See also Figure S1 and Movies S1,S2, and S3.
Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc. 299

Multiple Serum Factors Are Required for the Induction
of Serum-Dependent Necrosis
Results in Figure 1 indicate that certain serum factor(s) are
responsible for the activation of this type of RIP3-independent
necrosis. Biologically active components in serum include both
macromolecules (such as proteins) and small molecules. We
removed small molecules in fetal bovine serum (FBS) by dialysis
and then tested whether the dialyzed FBS (diFBS) can induce
cell death upon amino acid starvation. Unlike full FBS, diFBS
failed to induce potent cell death, and it even protected against
the modest apoptotic cell death induced by amino acid/serum
double starvation in MEFs (Figure 2;Movie S4), presumably
due to pro-survival growth factors present in serum. The small
molecule fraction of FBS (smFBS) prepared by filtering FBS
through a size-exclusion membrane also failed to mimic the
death-inducing activity of FBS. Only a combination of diFBS
with smFBS fully restored the potent killing (Figure 2). Therefore,
multiple serum factors, of both macromolecule and small mole-
cule natures, are required to induce this form of necrosis upon
amino acid starvation.
Transferrin Is Required for Serum-Dependent Necrosis
To identify the active component(s) in diFBS, we fractionated
diFBS by ammonium sulfate precipitation in combination with
various chromatographic columns. The killing activity of each
fraction was monitored by incubating the fraction with cultured
cells freshly switched to amino acid/serum-free medium, sup-
plemented with the smFBS fraction. Cell death was measured
after incubation for 12 hr. In this assay, we used bax/bak dou-
ble-knockout (DKO) MEFs to avoid amino acid/serum double
starvation-induced apoptosis. After a four-step fractionation
procedure (Figure 3A), we purified a single protein (Figure 3B)
that correlated with killing activity (Figure 3C). Mass spectrom-
etry analysis revealed the identity of the protein to be bovine
transferrin. To validate this activity of transferrin, we immuno-
depleted transferrin from FBS and found that the death-inducing
activity of FBS was indeed dramatically decreased (Figure 3D).
Further, upon amino acid starvation, addition of commercial
bovine holo-transferrin induced robust cell death in the presence
of smFBS (Figure S3). The effective concentration of transferrin
in these assays was well within the range of serum transferrin
concentration (0.49–2.63 mg/ml) (Valaitis and Theil, 1984). To
rule out the possibility that the killing activity came from certain
serum molecules co-purified with transferrin instead of trans-
ferrin per se, we tested recombinant human holo-transferrin
that was expressed in rice (thus serum was not involved in the
expression and purification). Again, upon amino acid starvation
and in the presence of smFBS, recombinant human holo-trans-
ferrin could induce cell death in a dose-dependent manner
(Figure 3E).
Transferrin is an iron carrier protein in serum that can be
transported into the cell via receptor-mediated endocytosis
(Andrews and Schmidt, 2007). To determine whether trans-
ferrin needs to be imported into the cell to exert its death-
inducing function, we first tested the requirement of transferrin
receptor (TfR) for serum-induced cell death. Indeed, TfR RNAi
significantly inhibited cell death (Figure 3F). Transferrin can
only interact with TfR and be transported into the cell when it
is loaded with iron. Consistently, iron-free bovine apo-trans-
ferrin does not possess killing activity (Figure 3G), and multiple
iron chelators can inhibit cell death (Figure 3H), in agreement
with the requirement of transferrin import for this mode of
cell death.
Glutamine and Glutaminolysis Are Required
for Serum-Dependent Necrosis
To identify the small molecule component of FBS required for
death induction, the smFBS fraction was subjected to multiple
steps of fractionation (Figure 4A). The resulting fractions of
each step were assessed for the death-inducing activity in
MEFs in combination with diFBS. The active fractions obtained
from the last step of purification (reverse phase-C18 HPLC)
were analyzed by mass spectrometry. Three major mass peaks
showed mass equal to that of two abundant components of
serum: L-glutamine (146 Da, with Na
+
or H
+
) and glucose
(180 Da, with Na
+
)(Figure 4B). Glucose failed to recapitulate
the killing activity in combination with diFBS or transferrin, which
was expected, because the cells were cultured in high-glucose
medium and diFBS alone could not induce cell death upon
amino acid starvation.
To determine whether glutamine (Gln) plays a role in serum-
dependent necrosis, we compared L-Gln with several related
amino acids. In a dose-dependent manner and upon amino
acid starvation, L-Gln but not D-Gln or other tested amino acids
induced potent cell death in the presence of diFBS (Figure 4C).
Figure 2. Multiple Serum Components Are Required for Serum-
Induced Cell Death
(A) MEFs were treated as indicated for 12 hr. Upper panel, phase-contrast;
lower panel, PI staining. diFBS: 10% (v/v) dialyzed FBS, smFBS: 10% (v/v).
(B and C) MEFs were treated as indicated for 12 hr. Cell death was determined
by PI staining coupled with flow cytometry (B), and cell viability was deter-
mined by measuring cellular ATP levels (C).
See also Figure S2 and Movie S4.
300 Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc.

To rule out the possibility that the killing activity is due to a
degraded product rather than L-Gln itself, we tested a chemically
stable L-Gln replacement, L-alanine-L-glutamine (A-Q). A-Q in
combination with diFBS was also able to induce cell death
upon amino acid starvation (Figure S4). Further, as expected,
the requirement of diFBS in this assay can be replaced with
transferrin (Figure 4D). In this later experiment, bax/bak-DKO
MEFs were used to avoid apoptosis induced by amino acid/
serum double starvation. It should be noted that unlike in wild-
type (WT) MEFs, L-Gln alone induced modest but measurable
cell death in bax/bak-DKO MEFs, and addition of transferrin
further enhanced cell death (Figure 4D).
L-Gln is the most abundant amino acid in the body. Through
glutaminolysis, proliferating cells use L-Gln both as a nitrogen
source for the biosynthesis of nucleotides, amino acids, and hex-
amine and as an important carbon source for the tricarboxylic
acid (TCA) cycle (DeBerardinis et al., 2008). We sought to identify
the molecular basis underlying the role of L-Gln and glutaminol-
ysis in serum-dependent necrosis. L-Gln uptake is mainly
dependent on receptors SLC38A1and SLC1A5 (McGivan and
Bungard, 2007)(Figure 5A). We found that pharmacological
inhibition of SLC1A5 by L-g-glutamyl-p-nitroanilide (GPNA)
(Esslinger et al., 2005) or RNAi knockdown of these receptors
markedly blocked serum-dependent necrosis (Figures 5B, 5C,
and S5A), indicating that efficient import of L-Gln is essential
for this type of cell death. In cells, Gln is converted into glutamate
(Glu) by glutaminases (GLS) (Curthoys and Watford, 1995).
Compound 968, an inhibitor of GLS (Wang et al., 2010),
significantly inhibited serum-dependent necrosis (Figure 5B).
There are two isoforms of mammalian GLS, GLS1 and GLS2
(Curthoys and Watford, 1995). We found that knockdown of
GLS2 but not GLS1 inhibited serum-dependent necrosis in
Figure 3. Transferrin and Transferrin Re-
ceptor Are Required for Serum-Dependent
Necrosis
(A) Purification scheme for the death-inducing
component in diFBS. See Experimental Pro-
cedures for detailed description.
(B) The final heparin column fractions were
resolved by SDS-PAGE and stained with Coo-
massie blue. Arrow indicates the protein band
correlating with killing activity.
(C) In amino acid-free medium, bax/bak-DKO
MEFs were incubated with the heparin fractions in
combination with smFBS as indicated, and cell
death was determined by PI staining coupled with
flow cytometry.
(D) Immuno-depletion of transferrin (TF) abrogated
the killing activity of serum. Serum was immuno-
depleted with control IgG or anti-transferrin anti-
body (a-TF) as indicated and subsequently used to
induce cell death in bax/bak-DKO MEFs under
amino acid-free conditions for 12 hr. Western blot
showed the efficiency of transferrin depletion from
FBS, with BSA as the loading control.
(E) Recombinant human holo-transferrin (rhTF)
induced cell death in bax/bak-DKO MEFs under
amino acid-free conditions in a smFBS-depen-
dent manner.
(F) RNAi of transferrin receptor (TfR) inhibited
serum-dependent necrosis. MEFs expressing
control shRNA (NT) or two independent shRNAs
targeting TfR were treated as indicated for 12 hr
and cell viability was measured. Western blot
confirmed knockdown of TfR expression.
(G) Iron-free bovine apo-transferrin (apo-bTF) did
not have death-inducing activity.
(H) Iron chelators blocked serum-dependent
necrosis. MEFs were treated with three different
iron chelators as indicated, and cell viability was
determined by measuring cellular ATP levels. DFO
(Deferoxamine), 80 mM; CPX (ciclopirox olamine),
10 mM; BIP (2, 2-bipyridyl), 100 mM.
See also Figure S3.
Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc. 301

MEFs (Figures 5D and S5B). Consistently, Bis-2-(5-phenylaceta-
mido-1,3,4-thiadiazol-2-yl)ethyl sulfide (BETPS), a GLS1-spe-
cific inhibitor (Robinson et al., 2007), failed to block serum-
dependent necrosis in MEFs (Figure S5C). Whether the specific
requirement of GLS2 but not GLS1 is due to differential regula-
tion of these two enzymes or due to predominant expression
of GLS2 in MEFs requires further investigation.
Downstream of glutaminolysis, glutamate can be further
converted into a-ketoglutarate (a-KG) either by glutamate
dehydrogenase (GLUD1)-mediated glutamate deamination or
by transaminase-mediated transamination (Hensley et al.,
2013)(Figure 5A). We found that amino-oxyacetate (AOA), a
pan inhibitor of transaminases (Wise et al., 2008), could inhibit
serum-dependent necrosis but not TNF-a-induced apoptosis
in MEFs (Figures 5B and S5E). Consistently, RNAi of the trans-
aminase GOT1 inhibited serum-dependent necrosis in MEFs
(Figure 5E). However, GLUD1 RNAi failed to do so (Figure S5D).
We then explored whether downstream metabolites of gluta-
minolysis can mimic the killing activity of L-Gln. Indeed, upon
amino acid starvation, a-KG in combination with diFBS can
Figure 4. Glutamine Is the Death-Inducing
Small Molecule Component in Serum
(A) Purification scheme for the death-inducing
small molecule component in FBS. S, superna-
tant; P, pellet. See Experimental Procedures for
detailed description.
(B) LC/MS spectra of the active fraction from the
XDB-C18 column.
(C) L-Gln (left panel) but not D-Gln (right panel)
induces cell death in a diFBS-dependent manner
under AA starvation. MEFs were treated as indi-
cated for 12 hr, and cell viability was subsequently
measured.
(D) L-Gln in combination with transferrin recapit-
ulated the cell death-inducing activity of serum.
bax/bak-DKO MEFs were treated as indicated
for 12 hr, and cell viability was subsequently
measured. L-Gln concentration, 0.1 mM.
See also Figure S4.
induce potent cell death even in the pres-
ence of transaminase inhibitor AOA (Fig-
ure 5F). This result is consistent with the
fact that in the glutaminolysis pathway,
a-KG is a downstream metabolite of
transaminases, the target of AOA.
Serum-Dependent Necrosis
Requires Cystine Starvation and
Subsequent Depletion of Cellular
Glutathione
Does serum-dependent necrosis require
deprivation of all amino acids or certain
specific amino acid(s)? To address this
question, we examined which amino
acid(s) can rescue cells from necrosis
triggered by serum, or transferrin in com-
bination with L-Gln, upon full amino acid
starvation. Adding back a single amino acid, cysteine or cystine,
but no other amino acids, rescued cells from death (Figures 6A,
S6A, and S6B). Conversely, cystine starvation alone, in the
presence of all other amino acids and serum, is sufficient to
induce cell death (Figure 6B). Further, even a partial starvation
of cystine can induce necrosis in a glutamine dose-dependent
manner (Figure S6C).
As a building block of the cellular reducing agent glutathione,
cysteine is required for maintaining cellular redox homeostasis.
Therefore, it is likely that cysteine starvation induces cell death
via depleting cellular glutathione (GSH) and consequently
increasing reactive oxygen species (ROS). For this reason, we
determined the cellular GSH and ROS levels under conditions
that trigger serum-dependent necrosis. Indeed, these conditions
caused dramatic decrease of GSH level and increase of ROS
level in cells (Figures 6C and 6D). Supplement of GSH or
the biosynthetic precursor of GSH, N-acetylcysteine (NAC), or
ROS scavenger Trolox (6-hydroxy-2,5,7,8-tetramethylchro-
man-2-carboxylic acid) all effectively blocked cell death under
these conditions (Figures 6E and 6F). Further, RNAi knockdown
302 Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc.

of glutamate cysteine ligase catalytic subunit (GCLC), an essen-
tial enzyme for GSH synthesis, sensitized cell death induced by
cystine starvation (Figure 6G). In addition, when mitochondrial
oxidative phosphorylation, an important resource for ROS pro-
duction, was inhibited by oligomycin, serum-induced necrosis
was greatly reduced (Figure S6D).
Serum-Dependent Necrosis Is Ferroptosis
Requirement of cystine starvation, cellular glutathione depletion,
and iron-carrier transferrin led us to test whether serum-depen-
dent necrosis is the same as ferroptosis, a recently discovered
regulated necrosis process triggered by the synthetic chemical
compound erastin and dependent on iron (Dixon et al., 2012;
Dolma et al., 2003; Yagoda et al., 2007). Erastin triggers ferrop-
tosis by inhibiting cystine uptake and thus depleting cellular
glutathione (Dixon et al., 2012; Yang et al., 2014). The following
experiments confirmed that serum-dependent necrosis is
indeed ferroptosis, or at least these two modes of cell death
share central mechanisms. First, ferrastatin-1 (Fer-1), a specific
ferroptosis inhibitor (Dixon et al., 2012), can block serum-depen-
dent cell death triggered by either full amino acid starvation
or cystine starvation (Figure 7A). Second, both transferrin and
glutamine were required for erastin-induced ferroptosis (Fig-
ure 7B). Further, iron chelators (DFO and CPX), glutaminolysis in-
hibitor Compound 968, and AOA can also inhibit erastin-induced
ferroptosis (Figures 7C and 7D), demonstrating that ferroptosis
requires the cellular metabolic process glutaminolysis.
Previous studies suggest that ferroptosis depends on Ras-
ERK signaling and can be completely blocked by MEK inhibi-
tion (Yagoda et al., 2007). We found that this conclusion is
inaccurate, likely due to the use of MEK inhibitor U0126 in
the previous studies. We compared U0126 with a more selec-
tive and potent MEK1/2 inhibitor, PD0325901, and found that
U0126 but not PD0325901 is able to block cell death induced
by either erastin or amino acid starvation in the presence of
serum (Figure S7A). Therefore, MEK activity is not essential
Figure 5. Glutaminolysis Mediates Serum-
Dependent Necrosis
(A) Schematic overview of the glutaminolysis
pathway.
(B) Pharmacological inhibition of multiple compo-
nents in the glutaminolysis pathway abrogated
serum-dependent necrosis. The following in-
hibitors were used as indicated: L-Gln transporter
inhibitor GPNA (5 mM); GLS inhibitor Compound
968 (968, 20 mM); Pan-transaminases inhibitor
AOA (0.5 mM).
(C) RNAi knockdown of SLC38A1 inhibited serum-
dependent necrosis. Left: MEFs expressing
non-targeting (NT) shRNA or shRNA targeting
SLC38A1 were treated as indicated for 12 hr, and
cell viability was subsequently measured. Right:
qPCR measurement of SLC38A1 mRNA levels in
MEFs infected with NT shRNA or shRNA targeting
SLC38A1.
(D) Knockdown of GLS2 blocked serum-depen-
dent necrosis. MEFs expressing non-targeting
shRNA (NT) or two independent shRNAs targeting
GLS2 were treated as indicated for 12 hr, and cell
viability was subsequently measured. FBS: 5%
(v/v). Western blotting confirmed the knockdown
of GLS2 expression.
(E) GOT1 RNAi reduced serum-dependent
necrosis. Left: MEFs expressing non-targeting
(NT) shRNA or shRNA targeting GOT1 were
treated as indicated for 12 hr, and cell viability was
subsequently measured. Right: qPCR measure-
ment of GOT1 mRNA levels in MEFs infected with
NT shRNA or shRNA targeting GOT1.
(F) a-ketoglutarate can mimic the death-inducing
activity of L-Gln but in a manner insensitive to the
transaminase inhibitor AOA. MEFs were treated
as indicated for 12 hr, and cell viability was sub-
sequently measured. a-KG (Dimethyl-a-Ketoglu-
tarate), 4 mM; AOA, 0.5 mM.
See also Figure S5.
Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc. 303

for ferroptosis. The unintended effect of U0126 may be due
to off-target inhibition of certain unknown enzymes required
for ferroptosis; it might also be due to the anti-oxidative prop-
erty of this compound as measured by its ability to reduce 2,2-
Diphenyl-1-picrylhydrazyl (DPPH) (Figure S7B). It should also
be noted that PD0325901 and Compound 968 showed no
anti-oxidant function (Figure S7B).
Inhibitors of Glutaminolysis Prevent Heart Injury
Induced by Ischemia/Reperfusion
Recent studies indicate that ferroptosis is associated with
ischemia/reperfusion injury of organs such as liver and kidney
and is thus a potential therapeutic target (Friedmann Angeli
et al., 2014; Linkermann et al., 2014). Since we identified glutami-
nolysis as an essential factor for ferroptosis, we sought to
test if the glutaminolysis inhibitor Compound 968 might also
be able to reduce ischemia/reperfusion injuries in an ex vivo
heart model. We subjected the hearts isolated from WT mice
to ischemia/reperfusion stress (Figure 7E). At the end of reperfu-
sion, the hearts treated with either iron chelator DFO, a docu-
mented ferroptosis inhibitor (Dixon et al., 2012), or GLS inhibitor
Compound 968 showed significant improved function compared
to hearts treated with vehicle (DMSO) as assessed by measuring
the left ventricular developed pressure (LVDP) (Figure 7F; vehicle
43.20% ± 1.66% of Baseline, DFO 64.75% ± 4.09%, and
Compound 968 75.25% ± 5.75%). This functional improvement
was consistent with the reduction in myocardial infarcts size
(Figure 7G; vehicle treatment 30.82% ± 0.57% of myocardium,
DFO treatment 22.60% ± 0.97%, and Compound 968 treatment
17.36% ± 0.47%). Similarly, DFO or Compound 968 significantly
inhibited the release of lactate dehydrogenase (LDH) during
reperfusion, another indicator of myocardial injury (Figure 7H).
These data suggest that inhibition of ferroptosis via ablating glu-
taminolysis can protect heart tissue from ischemia/reperfusion
injury.
DISCUSSION
Collectively, our study uncovered several essential components
and regulatory mechanisms for ferroptosis, a RIP3-independent
programmed necrosis process. Extracellularly, serum compo-
nents L-glutamine and transferrin were identified here as crucial
regulators of ferroptosis. Intracellularly, the specific metabolic
pathway glutaminolysis and components mediating transferrin
Figure 6. Cystine Starvation and Subse-
quent Cellular Redox Homeostasis Unbal-
ance Trigger Serum-Dependent Necrosis
(A) Addition of cysteine (C, 0.2 mM) or cystine (CC,
0.2 mM) inhibited serum-dependent necrosis.
MEFs were treated as indicated for 12 hr. Cell
death was subsequently measured by PI staining
followed by flow cytometry.
(B) Cystine starvation alone is sufficient to induce
cell death. MEFs were treated as indicated for
12 hr, and cell death was determined by PI staining
coupled with flow cytometry.
(C) Glutathione (GSH) was depleted in the condi-
tion of serum-induced necrosis or cystine starva-
tion. MEFs were treated as indicated for 6 hr and
harvested for total glutathione measurement.
(D) Accumulation of ROS in MEFs induced by
serum under AA starvation condition (left) or
cystine starvation (right). ROS levels in MEFs were
determined by H
2
DCFDA staining. H
2
DCFDA
assay was performed at 8 hr (left) or 6 hr (right).
(E) Supplement of GSH blocked cell death induced
by serum upon total AA starvation or cystine
starvation. MEFs were treated as indicated for
12 hr, and cell death was subsequently measured.
(F) Cell death induced by serum upon AA starva-
tion or cystine starvation can be prevented by
antioxidant reagents NAC (0.2 mM) and Trolox
(0.2 mM). MEFs were treated as indicated for
12 hr, and cell death was subsequently measured.
(G) GCLC RNAi sensitized serum-dependent
cell death upon cystine starvation. Left: MEFs
expressing non-targeting (NT) shRNA or shRNA
targeting GCLC were treated as indicated for
10 hr, and cell death was subsequently measured.
Right: qPCR measurement of GCLC mRNA levels
in MEFs infected with NT shRNA or shRNA tar-
geting GCLC.
See also Figure S6.
304 Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc.

import are required for ferroptosis. Several intriguing and seem-
ingly counter-intuitive observations are associated with these
findings: under normal conditions, both transferrin and glutamine
are required for cell survival and growth. However, upon amino
acid starvation, these growth/survival factors function to unleash
ferroptotic cell death that is more rapid and potent than that
caused by amino acid starvation alone. Such unexpected func-
tion of transferrin and glutamine is reminiscent of the pro-death
function of the life-essential protein cytochrome cin the intrinsic
apoptotic pathway. Similarly, glutaminolysis is crucial for cellular
biosynthesis and proliferation, but here it is actively involved in
and essential for this intriguing metabolic cell death process.
Under what biological conditions can ferroptosis, a necrotic
process induced experimentally by cystine starvation or
blockage of cystine uptake, occur? Because both transferrin
and glutamine are highly abundant in blood, whereas cystine
amount is much lower, and a partial deprivation of cystine is
sufficient to induce necrosis (Figure S6C), it is possible that, un-
der certain pathological conditions such as ischemia, cells might
be susceptible to ferroptosis. Alternatively, blockage of cystine
uptake or deficiency in glutathione synthesis can also lead to
ferroptosis. Indeed, there are hemolytic anemia patients carrying
hereditary deficiency of the glutathione synthetase (GSS) gene
(Nja
˚lsson, 2005) or that of gamma-glutamyl cysteine synthetase
(GCS) gene (Beutler et al., 1999; Dolma et al., 2003; Hirono et al.,
1996; Man
˜u
´Pereira et al., 2007; Ristoff et al., 2000). These
patients might also be subject to higher risk of ferroptosis.
This study raised many important mechanistic questions for
further understanding of ferroptosis. How do transferrin, cellular
glutaminolysis, glutathione synthesis, and cellular ROS genera-
tion process communicate with each other to lead to cell death?
Further, are the extracellular cues of ferroptosis regulated?
There are a variety of potential mechanisms for such extracellular
regulation. For example, we found that the killing activity of
transferrin is dictated by its iron-loading status; thus mecha-
nisms controlling iron-loading of transferrin may impact ferropto-
sis. Additionally, blood glutamine concentration can be variable.
Indeed, when we used different FBS preparations, we found
that death-inducing activity ranged from very potent, modest,
to almost undetectable (Figures S7C and S7D). Strikingly, we
found a positive correlation between the killing activity of indi-
vidual FBS preparations and their L-Gln concentrations, and
addition of L-Gln or the smFBS fraction prepared from death-
competent FBS can restore the killing activity of those inactive
FBS preparations (Figures S7D–S7F).
Relevant to human disease, this study provided further evi-
dence to support that ferroptosis is responsible, at least partially,
for organ injury triggered by ischemia/reperfusion. This study
also demonstrated that enzymes involved in glutaminolysis are
potential therapeutic targets because of their crucial role in
ferroptosis. Furthermore, the cancer therapeutic potential of
ferroptosis should also be explored. Previous studies have
linked ferroptosis with oncogenic Ras, and most recently it has
been demonstrated that p53 tumor suppressor positively regu-
lates ferroptosis by transcriptionally inhibiting the expression of
the cystine/glutamate antiporter SLC7A11 (Dixon et al., 2012;
Jiang et al., 2015; Yagoda et al., 2007). In light of our finding
that ferroptosis requires glutaminolysis, a metabolic pathway
Figure 7. Serum-Dependent Necrosis Is Ferroptosis and Is Involved
in Ischemia/Reperfusion Heart Injury
(A) Ferroptosis inhibitor Ferrastatin-1 (Fer-1) can inhibit serum-induced ne-
crosis upon total AA starvation or cystine (CC) starvation. MEFs were treated
as indicated for 12 hr, and cell viability was subsequently measured by PI
staining followed by flow cytometry. Erastin: 1 mM.
(B) Erastin-induced ferroptosis required both transferrin and glutamine. bax/
bak DKO MEFs were treated as indicated for 12 hr, and cell death was sub-
sequently measured by PI staining followed by flow cytometry. Erastin: 1 mM.
(C) Iron chelators inhibited erastin-induced ferroptosis. MEFs were treated as
indicated for 12 hr, and cell death was subsequently measured. DFO, 80 mM;
CPX, 10 mM; Erastin: 1 mM.
(D) Inhibition of GLS by Compound 968 (20 mM) or transaminases by pan-
transaminases inhibitor AOA (0.5 mM) blocked erastin-induced ferroptosis.
MEFs were treated as indicated for 12 hr, and cell death was subsequently
measured. Erastin, 1 mM.
(E) Schematic protocol for the ischemia/reperfusion study involving 30 min of
global ischemia followed by a 60 min reperfusion period.
(F) Determination of myocardial ischemic injury and function by left ventricular
developed pressure (LVDP) recovery, showing improved functional recovery
upon treatment with DFO or Compound 968 (968). DFO, 80 mM; 968, 25 mM.
(G) Representative cross-sections of hearts stained with TTC, demonstrating
reduced infarct injury (pale region) upon DFO or 968 treatment. The plot (lower
panel) shows quantification of infarct size of each group.
(H) Determination of myocardial ischemic injury by lactate dehydrogenase
(LDH) release, demonstrating reduced infarct injury in the hearts treated with
DFO or 968.
See also Figure S7.
Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc. 305

highly active in cancer tissues and essential for cancer growth,
can ferroptosis be preferentially triggered in cancer cells as a
therapeutic option?
EXPERIMENTAL PROCEDURES
For full experimental procedures, see the Supplemental Information.
Induction and Measurement of Cell Death
To induce cell death, 80%-confluent cells were washed with PBS twice
and then cultured in amino acid-free medium, with specific factors added as
indicated in individual experiments. Cell death was analyzed by PI staining
coupled with microscopy or flow cytometry. Alternatively, cell viability was
determined using the CellTiter-Glo luminescent Cell Viability Assay (Promega).
In assays using WT MEFs, viability was calculated by normalizing ATP levels to
cells treated with amino acid starvation in the presence of 10% (v/v) diFBS,
while in assays using bax/bak-DKO MEFs, ATP levels were normalized to cells
treated with amino acid and FBS double starvation.
Purification and Identification of Transferrin from FBS
All purification steps were carried out at 4C, and chromatography was
performed with an Amersham FPLC system. For the purification, 20 ml FBS
(664 mg protein) was applied to 50%–70% (saturation) ammonium sulfate pre-
cipitation. The protein pellet (242 mg) that contained the activity was dissolved
in 4 ml Buffer A (20 mM HEPES [pH 7.5], 10 mM NaCl) and dialyzed overnight.
The activity was applied to HiTrap SP Sepharose (GE Healthcare). The
flowthrough containing the activity was subjected to HiTrap Q Sepharose
(GE Healthcare). After washing the column with Buffer A, the fraction was
eluted by a gradient of 10–300 mM NaCl in Buffer A. Activity-containing frac-
tions were further fractionated with HiTrap Heparin Sepharose (GE Healthcare)
by using a gradient 10–300 mM NaCl in Buffer A. Fractions of 1 ml was
collected. After dialysis, filtered with 0.22 mM filter, the fractions was assayed
for activity. SDS-PAGE and Coomassie staining (Bio-Rad) was preformed, and
a single band correlated with the killing activity was subject to protein identity
determination by mass spectrometry analysis (MALDI-TOF-MS/MS). The ac-
tivity was identified as bovine transferrin.
Immuno-Depletion of Transferrin
To deplete transferrin from serum, amino acid-free DMEM containing 10% FBS
was incubated with control IgG or anti-bovine transferrin antibody bound to
Protein G Agarose (GE Healthcare) overnight at 4C. Protein G Agarose was
removed by centrifugation, and the supernatant was assayed for killing activity.
Purification and Identification of L-Glutamine from FBS
FBS was filtered through Centrifugal Filter Units (MWCO 10 kDa) (Millipore) to
obtain smFBS. One milliliter of smFBS was dried under vacuum and dissolved
in 1 ml methanol, and insoluble material was removed by centrifugation. The
supernatant was dried and dissolved in 750 ml methanol first and then mixed
with 614 ml acetonitrile (final ratio of methanol:acetonitrile is 55:45). After
incubating the mix for 30 min at 4C, precipitated material was removed by
centrifugation, and the supernatant was dried and dissolved in 750 ml meth-
anol. Aliquot of 150 ml was mixed with 1,350 ml acetonitrile (final ratio of
methanol:acetonitrile is 10:90) and incubated for 30 min at 4C. The insoluble
material was obtained by centrifugation and then dissolved in 75 ml ddH
2
O.
An aliquot of 50 ml as input was applied to a reversed-phase XDB-C18
(4.6 3250 mm) HPLC analytical column (Agilent). Separation was achieved
by use of step elution consisting of A (ddH
2
O) and B (methanol) as follows:
0.00–11.00 min: 100% A, flow rate 1 ml/min; 11.00–21.00 min: 100% B, flow
rate 1 ml/min. All fractions were dried and dissolved in 100 ml ddH
2
O, and
25 ml of each fraction was subjected to activity assay. The fraction with the
highest killing activity was subjected to mass spectrometry (PE SCIEX API-
100 LC/MS system, mass range: 30.0–500.0 by amu).
Time-Lapse Microscopy
Live-cell imaging of H2b-mCherry-expressing MEFs was performed on glass-
bottom six-well plates (MatTek) using a Nikon Ti-E inverted microscope
attached to a CoolSNAP CCD camera (Photometrics). Fluorescence and dif-
ferential interference contrast (DIC) images were acquired ever y 7 min, and im-
ages were analyzed using NIS elements software (Nikon) and ImageJ software
(NIH). For confocal imaging, MEFs were grown on 35-mm glass-bottom plates
(MatTek), and DIC images were acquired every 5 min with the Ultraview Vox
spinning disc confocal system (PerkinElmer) equipped with a Yokogawa
CSU-X1 spinning disc head and EMCCD camera (Hamamatsu C9100-13)
and coupled with a Nikon Ti-E microscope. Image analysis was performed
with Volocity software (PerkinElmer). All imaging was carried out in incubation
chambers at 5% CO
2
and 37C.
GSH Measurement
2310
5
MEFs were seeded in six-well plates. One day later, cells were treated
as indicated for 6 hr. Cells were harvested and cell numbers were determined.
Total glutathione was measured as described previously (Rahman et al., 2 006).
Measurement of ROS
MEFs were treated as indicated, and then 10 mM2
0,7’-dichlorodihydrofluores-
cein diacetate (H
2
DCFDA, Life Technologies Cat. #D-399) was added and
incubated for 1 hr. Excess H
2
DCFDA was removed by washing the cells twice
with PBS. Labeled cells were trypsinized and resuspended in PBS plus 5%
FBS. Oxidation of H
2
DCFDA to the highly fluorescent 20,7’-dichlorofluorescein
(DCF) is proportional to ROS generation and was analyzed using a flow cytom-
eter (Fortessa, BD Biosciences). A minimum of 10,000 cells was analyzed per
condition.
2,2-Diphenyl-1-Picrylhydrazyl Assay for Antioxidant Activity
The experiment was performed as described previously (Blois, 1958; Dixon
et al., 2012). 2,2-Diphenyl-1-picrylhydrazyl (DPPH) (Sigma Cat#D9132) was
dissolved in methanol to a final concentration of 50 mM. The tested compounds
were added to 1 ml of DPPH solution with a final concentration of 50 mM.
Samples were mixed well and incubated at room temperature for 1 hr. The
absorbance at 517 nm (indicating the concentration of non-reduced DPPH)
was measured using methanol as control. Results were normalized to
DMSO (which has no antioxidant activity; set as 100%).
Ischemia/Reperfusion Analysis Using Isolated Hearts
Male C57BL/6J mice weighing 25–30 g at age 12–14 weeks were used in all
experiments and maintained in a temperature-controlled room with alter-
nating 12:12-hr light-dark cycles. Experiments were performed using an
isovolumic isolated heart preparation as published and modified for the
use in mice hearts (Ananthakrishnan et al., 2009; Hwang et al., 2004). Hearts
from 12- to 14-week-old WT mice were isolated and retrograde perfused at
37C in a non-recirculating mode through the aorta at a rate of 2.5 ml/min.
Hearts were perfused with modified Krebs-Henseleit (KH) buffer (118 mM
NaCl, 4.7 mM KCl, 2.5 mM CaCl
2
, 1.2 mM MgCl
2
, 25 mM NaHCO
3
,5mM
glucose, 0.4 mM palmitate, 0.2 mM glutamine, 10 mg/ml human recombinant
transferrin, 0.4 mM BSA, and 70 mU/l insulin). Left ventricular developed
pressure (LVDP) was measured using a latex balloon in the left ventricle.
LVDP and coronary perfusion pressure were monitored continuously on a
four-channel Gould recorder. Hearts were perfused either with KH buffer
containing vehicle (DMSO) or the Compounds throughout the ischemia/re-
perfusion protocol. After an equilibration period of 30 min, global ischemia
was performed for 30 min followed by 60 min of reperfusion. Cardiac
injury due to ischemia/reperfusion stress was assessed by measuring LDH
release in the perfusates that were collected during 60 min of reperfusion.
Infarct area was measured using 2, 3, 5-triphenyltetrazolium chloride (TTC)
staining. After 60 min of reperfusion, the heart is perfused with Evans blue
in situ and then removed. Hearts were sliced into cross-sections at approx-
imately 1-mm intervals. The sections were embedded in the TTC solution at
37C for 10 min, and area of infarct as a percent of the whole heart was
quantified as described. Functional recovery of LVDP was expressed by
comparing to the initial LVDP before ischemia. All animal experiments
were approved by the Institutional Animal Care and Use Committees of
New York University School of Medicine and conformed to the guidelines
outlined in the NIH Guide for Care and Use of Laboratory Animals (NIH
Pub. No. 85–23, 1996).
306 Molecular Cell 59, 298–308, July 16, 2015 ª2015 Elsevier Inc.

Statistical Analyses
All statistical analyses were performed using Prism 5.0c GraphPad Software.
p values were calculated with unpaired Student’s t test. Data are presented
as mean ± SEM from three independent experiments (*p < 0.05, **p < 0.01,
***p < 0.001 by unpaired Student’s t test).
SUPPLEMENTAL INFORMATION
Supplemental Information includes Supplemental Experimental Procedures,
seven figures, and four movies and can be found with this article online at
http://dx.doi.org/10.1016/j.molcel.2015.06.011.
ACKNOWLEDGMENTS
We thank Memorial Sloan Kettering Cancer Center (MSKCC) colleagues Drs.
Ron Hendrickson and Hediye Erdjument-Bromage for identification of bovine
transferrin by mass spectrometry, Drs. George Sukenick and Hui Fang for
analyzing small molecule activity by mass spectrometry, and Dr. Michael Over-
holtzer and his lab members for assistance with live-cell imaging. We also
thank Drs. Junru Wang (MSKCC) and Qing Li (New York University) for tech-
nical support. This work is supported by NIH grants R01CA166413 (to X.J.),
R01GM113013 (to X.J.), R01HL102022 (to R.R.), P01HL60901 (to R.R.),
P01AG026467 (to R.R.), and Mr. William H. and Mrs. Alice Goodwin and the
Commonwealth Foundation for Cancer Research of the Experimental Thera-
peutics Center of MSKCC (to X.J.).
Received: February 2, 2015
Revised: April 30, 2015
Accepted: June 4, 2015
Published: July 9, 2015
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