From the Department of Internal Medicine IV,
University of Heidelberg, Bergheimer Str. 58, 69115 and the ¶ Gene
Expression Programme, European Molecular Biology Laboratory,
Meyerhofstr. 1, 699117 Heidelberg, Germany
Received for publication, January 24, 2001
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ABSTRACT |
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The expression of several proteins with critical
functions in iron metabolism is regulated post-transcriptionally by the
binding of iron regulatory proteins, IRP1 and IRP2, to mRNA iron
responsive elements (IREs). In iron-deficient tissues and
cultured cells, both IRP1 and IRP2 are activated for high affinity IRE
binding. Previous work showed that IRP1 is also activated when
cultured cells are exposed to H2O2. The
well established role of iron and H2O2 in
tissue injury (based on Fenton chemistry) suggests that this response
may have important pathophysiological implications. This is
particularly relevant in inflammation, where cytotoxic immune cells
release large amounts of reactive oxygen species. Here, we describe a
rat liver perfusion model to study IRP1 activation under
H2O2 generation conditions that mimic a
physiological inflammatory response, using steady-state concentrations
of H2O2 produced by a glucose/ glucose
oxidase/catalase system. We show first that stimulated neutrophils are
able to increase serum levels of H2O2 by a
factor of 10, even in the presence of
H2O2-degrading erythrocytes. We further show
that perfusion of rat liver with glucose oxidase leads to a rapid
activation of IRE binding activity in the intact organ. Mobility shift
assays with liver extracts and IRP1 or IRP2-specific probes indicate
that only IRP1 responds to H2O2. Our
study demonstrates a principal existence of iron regulation by
oxidative stress at the intact organ level. It also provides a link
between iron metabolism and the inflammatory response, as
H2O2 is a major product of the oxidative burst
of neutrophils and macrophages.
Because of the flexible coordination chemistry of iron
and its redox potential, cells and organisms utilize iron-containing proteins for vital metabolic functions, such as oxygen transport, electron transfer, and catalysis (1). While these properties explain
why iron is an essential constituent for a multitude of biochemical
activities, they also render it potentially toxic for cells and
tissues. In the presence of reactive oxygen intermediates (ROIs),1 iron catalyzes the
generation of hydroxyl radicals (Fenton/Haber-Weiss reactions) that
damage membrane lipids, proteins, and nucleic acids (2). Considering
that ROIs, including H2O2 and O Cellular iron uptake involves binding of the soluble plasma iron
carrier transferrin to the cell-surface transferrin receptor (TfR),
followed by endocytosis and iron release into the cytoplasm. A fraction
of iron is utilized for the synthesis of iron-containing proteins, and
excess iron is stored in ferritin, a multisubunit protein of H- and
L-chains. The expression of TfR and ferritin are controlled
post-transcriptionally by binding of two cytoplasmic iron regulatory
proteins, IRP1 and IRP2, to iron-responsive elements (IREs) in their
mRNAs. Both IRPs are activated by iron deficiency to bind to IREs
as monomers, resulting in the stabilization of TfR mRNA, which
contains multiple IREs in the 3'-untranslated region, and translational
inhibition of ferritin mRNA, that harbors a single IRE in the
5'-untranslated region (6, 7).
IRP1 is a bifunctional protein that assembles a cubane
4Fe-4S cluster and functions as a cytosolic aconitase in iron-repleted cells. Conversely, the cluster dissociates in iron-starved cells, and
IRP1 binds to IREs in its apo form. IRP2 is homologous to IRP1;
however, IRP2 activity rises in iron-deficient cells by stabilization
of the protein. Both IRPs respond to additional, iron-independent
signals such as NO and oxidative stress (8, 9). By employing various
cell lines, we and others have previously shown that IRP1 is rapidly
activated when cells are exposed to low micromolar concentrations of
H2O2 (10, 11). Mechanistically, IRP1 activation
by H2O2 is not the simple consequence of a
direct attack of H2O2 on the 4Fe-4S cluster but
rather appears to involve stress-response signaling (12). Moreover,
this finding has established a novel regulatory connection between iron
metabolism and oxidative stress, which is striking considering the role
of H2O2 in iron toxicity (Fenton chemistry).
Activation of IRP1 by H2O2 results in enhanced
expression of TfR and increased cellular iron uptake from transferrin,
whereas it also leads to a significant reduction of the iron storage
protein ferritin (13).
Although these data may help to understand iron-mediated tissue damage,
as yet, studies have only been performed in cultured cell lines. Here
we investigate the response of IRP1 to H2O2 in the intact liver, a key organ for iron homeostasis, iron pathogenesis, and oxygen turnover. We demonstrate the ability of
H2O2 to modulate hepatic iron metabolism at the
intact organ level. This finding is particularly relevant in the light
of the capacity of the liver to efficiently degrade
H2O2.
Reagents--
Luminol, phosphate-buffered saline, HANKS buffer,
H2O2, catalase, sodium hypochlorite, glucose
oxidase, and sodium azide were from Sigma (Deisenhofen, Germany). Human
HepG2 cells were grown in Dulbecco's modified Eagle's medium
supplemented with 2 mM glutamine, 4.5 g/l glucose, 100 units/ml penicillin, 0.1 ng/ml streptomycin, and 10% fetal calf serum.
Solutions--
Stock solutions of luminol were prepared in 10 mM phosphate-buffered saline and adjusted to pH 7.4. Stock
solutions of NaOCl and H2O2 were prepared in
water. Their concentrations were determined spectrophotometrically
( Isolation of Leukocytes and Erythrocytes--
Unfractionated
leukocytes (neutrophils and lymphocytes/monocytes) or isolated
erythrocytes, neutrophils, monocytes, and lymphocytes were obtained as
described by Boyum et al. (16) with slight modifications.
Briefly, heparinized blood from healthy volunteers was underlayed with
50% (by volume) histopaque (Sigma) and centrifuged at 400 × g for 30 min to separate all cells from the blood plasma. The interphase (lymphocytes and monocytes) was further washed with NaCl
0.9%, and erythrocytes were removed by hypotonic lysis using distilled
water. The pellet (neutrophils and erythrocytes) was resuspended in
0.9% NaCl, centrifuged at 600 × g for 7 min. The
pellet was resuspended in Hanks I buffer and incubated with 5% dextran
sulfate (Sigma) at a volume ratio of 1:10 for 40 min at room
temperature. The pellet (erythrocytes) was further washed with 0.9%
NaCl. The supernatant (neutrophils) was further washed with 0.9% NaCl,
and erythrocytes were removed by hypotonic lysis using distilled water.
For the determination of cell number, a Celldyn 1700 (Abbott, Great
Britain) was used.
Determination of H2O2 Generation by
Glucose Oxidase and Leukocytes--
A sensitive nonenzymatic
chemiluminescence assay was used for the determination of
H2O2 (17-19), and a flow technique was
employed to determine glucose oxidase activity (20, 21). Briefly, a solution of glucose and glucose oxidase aspirated by a peristaltic pump
(4 ml/min) was mixed with luminol (10 Liver Perfusion--
Liver perfusion was established according
to standard procedures (22). Rat livers were perfused at 20 ml/min via
a peristaltic pump in a single path system with Krebs-Henseleit buffer
(0.3 mM pyruvate, 2 mM lactate). The
temperature was kept at 37 °C, and the buffer was saturated with
oxygen. The pH was maintained at 7.4 by exposure to carbon dioxide. As
determined with an oxygen electrode, the oxygen concentration was only
reduced to 80% after liver passage under our perfusion conditions
indicating a sufficient oxygen supply of the organ. Lactate
dehydrogenase, potassium, and sodium were determined in a routine
clinical laboratory to determine common parameters of cell toxicity. In
the experiments with GOX, GOX was added to the buffer reservoir
with an final activity of 3 × 10 Electrophoretic Mobility Shift Assay (EMSA)--
EMSAs were
performed as described earlier using a radiolabeled human ferritin
H-chain IRE probe (9). IRP1- or IRP2-specific probes were generated
from plasmids CG 125 (IRP2) (23) and no. 34 (IRP1) (24). RNA-protein
complex formation was quantified by densitometric scanning of the
depicted autoradiographs.
Perfusion of Rat Liver with H2O2 Fails to
Activate IRP1--
Treatment of cultured cells with a bolus of 100 µM H2O2 results in rapid
activation of IRP1 within 30-60 min. Previous studies have provided
evidence that this response is elicited by extracellular H2O2 and that a mere increase in intracellular
H2O2 levels is not sufficient to activate IRP1
(10); inhibition of peroxisomal catalase by aminotriazole caused a
significant and rapid increase in intracellular
H2O2, but no IRP1 activation could be detected under these conditions. By contrast, even a modest increase in the
extracellular H2O2 concentration was sufficient
to rapidly induce IRP-1 in the absence of an observable increase in
intracellular H2O2 concentration (10).
Stimulated neutrophils and macrophages are considered to represent a
major physiological source of extracellular
H2O2 in the organism, although other cell types
also release smaller quantities of H2O2 (3). To
study the effects of extracellular H2O2 on hepatic IRP1 at the intact organ level, we established a
non-recirculating rat liver perfusion model, which allows the
application of H2O2 from the
sinusoid/extracellular compartment of the liver. Contrary to
experiments with cultured cell lines, perfusion of the rat liver with
an oxygenated Krebs-Henseleit buffer containing 100 µM
H2O2 for 60 min failed to activate IRP1 (Fig.
1, upper panel), although the
presence of activable IRP1 could be demonstrated by the addition of 2%
2-mercaptoethanol to all lysates (Fig. 1, lower panel).
H2O2 was stable in perfusion buffer that was
not used for liver perfusion (data not shown). A possible
interpretation of this unexpected result might be that
H2O2 does not activate IRP1 at the intact organ
level, challenging the physiological significance of previous studies.
Alternatively, the perfusion of the liver with
H2O2 may not accurately mimic in
vivo conditions, as hepatic catalase and glutathione peroxidase
are very efficient in H2O2 degradation. For
instance, catalase activity in liver homogenate is as high as
~kcat = 200 s Generation of H2O2 by Stimulated
Neutrophils--
During the oxidative burst of neutrophils, a plasma
membrane-bound NADPH oxidase generates O
To study whether erythrocytic catalase and glutathione peroxidase would
efficiently remove oxygen-burst-derived H2O2,
we determined H2O2 concentrations in highly
diluted blood. Under these conditions, the steady-state concentration
of H2O2 still increased by a factor of ten upon
stimulation with 100 ng/ml PMA (Fig. 2b). Because erythrocytes and leukocytes were used at their original ratio, the
results are expected to reflect the in vivo conditions. In conclusion, H2O2 concentrations may drastically
change during the inflammatory response of leukocytes, potentially
affecting redox-sensitive signaling cascades such as the IRE/IRP
regulatory network.
Perfusion of Rat Liver with H2O2 at
Steady-State Concentrations That Mimic the Oxidative Burst of
Neutrophils Activates IRP1--
The isolated liver of a male Wistar
rat was perfused in situ with Krebs-Henseleit buffer
containing glucose/GOX. This procedure leads to the generation of a
continuous flux of H2O2 (30). The amount of GOX
was calculated to yield 0.2 µM/s
H2O2, comparable with that released by
stimulated neutrophils. To assess the response of liver IRP1 to the
exposure to H2O2, liver lobules were removed every 10 min, and IRE binding activity in liver cytoplasmic extracts was monitored by an electrophoretic mobility shift assay (Fig. 3). Under these conditions, IRP
activation is apparent within 20 min of liver perfusion, increasing IRP
activity within 1 h by a factor of 25. To confirm the equal
loading of all lanes, cell extracts were treated with 2%
2-mercaptoethanol, which activates IRP1 in vitro (31). In
kinetic terms, this result corroborates previous observations in
cultured fibroblasts (10). It should be noted that no significant
release of lactate dehydrogenase, change in electrolyte composition
(sodium and potassium), or visible morphological alteration of the
liver were observed during perfusion (data not shown), indicating that
H2O2 was, as expected, not toxic under the
conditions of the experiment.
To evaluate whether the H2O2-mediated increase
in IRE binding activity in the liver corresponds to IRP1 and/or IRP2
activation, we utilized IRP1 and IRP2-specific IRE probes and performed
electrophoretic mobility shift assays in liver extracts. The results
shown in Fig. 4 reveal that IRP1 not only
accounts for the H2O2-mediated increase in IRP
activity but also demonstrates that IRP1 is the predominant iron
regulatory protein in rat liver. In contrast to this, HepG2 cells, a
human hepatoma cell line, are rich in IRP2. Whereas the analysis with
IRP2-specific probes showed that any IRE binding activity of IRP2 in
rat liver extracts is below detection levels, the respective fraction
of IRE binding activity in HepG2 extracts is about 40%.
The initial finding that H2O2 activates
IRP1 in tissue culture cells raised intriguing mechanistic and
physiological questions. A series of experiments focused on the
elucidation of the underlying mechanism. The results obtained so far
have provided evidence that the activation of IRP1 requires signaling
activity by H2O2, rather than a direct chemical
modification of its 4Fe-4S cluster by H2O2.
First, IRP1 activation takes place in intact cells but not when cell
extracts are exposed to H2O2 (32). Moreover,
treatment of highly purified human recombinant 4Fe-4S IRP1 with
H2O2 yields 3Fe-4S IRP1, which suffices to
destroy the aconitase of the 4Fe-4S IRP1 activity but does not activate
IRE binding (33). Second, in cells treated with
H2O2, activation of IRP1 follows biphasic kinetics and can be completed in the absence of
H2O2 (8). In fact, the presence of a threshold
concentration of H2O2 (estimated ~10
µM) is only required for 10-15 min (10, 30). Third, a mere increase in intracellular levels of reactive oxygen intermediates (including H2O2) is not sufficient to elicit
IRP1 activation (10). Finally, reconstitution of
H2O2-mediated induction of IRP1 in an in
vitro system of permeabilized cells suggests the involvement of
membrane-associated factor(s) and phosphorylation-dephosphorylation steps in the pathway (34).
Whereas IRP1 activation by H2O2 has been
extensively studied in cultured cells and in vitro, the
physiological implications of IRP1 activation by
H2O2 in intact organs have not yet received comparable attention. The responses of the IRP/IRE regulatory network
toward oxidative stress warrant particular consideration in the context
of inflammation, where activated immune effector cells, including
neutrophils and macrophages, undergo a respiratory burst resulting in
the generation and release of reactive oxygen species to combat infection.
To gain insight into pathophysiological aspects of the IRP1 response to
H2O2, we perfused rat liver with
H2O2 under conditions that mimic the oxidative
burst of phagocytes. We show that perfusion of the liver with 100 µM H2O2 is not sufficient to
activate liver IRP1, although an analogous treatment of cultured cells
is effective. The high H2O2-degrading capacity
of the liver plausibly explains this result, since the half-life of
H2O2 in the liver was estimated to lie within
the millisecond range (21). These considerations suggested that the
perfusion of rat liver with H2O2 might not faithfully mimic in vivo conditions. Following the
determination of the rate of H2O2 generation in
PMA-stimulated neutrophils, we calibrated an
H2O2-generating system based on the oxidation of glucose by glucose oxidase. This setting allowed to mimic
H2O2 generation during the oxidative burst.
Moreover, it provided a model system to study the effects of
H2O2 on IRP1 in an intact organ, uncoupled from
the pleiotropic effects of neutrophil stimulation. Perfusion of rat
liver with GOX leads to the rapid, although incomplete, activation of
IRP1. This result validates earlier observations made in cultured cells
at the intact organ level. The increase in IRE binding activity by
H2O2 is based on IRP1 activation, as no IRP2
activity was detectable even with IRP2-specific probes. This is in
agreement with the previous observation that the liver contains
predominantly IRP1 (35). Our study not only demonstrates a principal
existence of iron regulation by oxidative stress in the liver, but it
also links it to the inflammatory response, as
H2O2 is a main product of the oxidative burst
cascade of leukocytes.
The response of the IRE/IRP regulatory system to different
conditions of oxidative stress may be more complex. Different from the
observed activation of IRP1 by H2O2 in the
intact liver, incubation of rat liver homogenates with xanthine
oxidase, which generates both superoxide anions (O The liver is an important iron storage organ that is primarily
affected in disorders of iron overload. As activation of IRP1 directly
leads to up-regulation of TfR and down-regulation of ferritin
synthesis, increased H2O2 levels would trigger
a shift of extracellular iron toward the free cytosolic compartment. A shift of iron into various intracellular compartments is exactly the
situation observed in humans under conditions of inflammation. In
contrast to chronic iron deficiency, chronic inflammation decreases blood iron levels i.e. extracellular iron concentration.
This inflammation-induced decrease in extracellular iron concentration is not a global net deficiency in iron but reflects a shift from the
extracellular phase into the intracellular compartment (39, 40).
Interestingly, also more localized chronic inflammations like chronic
hepatitis C seems to be associated with an increased intracellular iron
level. Almost half of all cases with hepatitis C show
nonhemochromatosis iron overload (41). Our findings hence resemble some
clinical observations and suggest a study of the involvement of
H2O2 in determining the distribution of iron in inflammation.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
290 = 350 mol liter
1 cm
1 at
pH 12 (14) and
230 = 74 mol liter
1
cm
1 (15) for NaOCl and H2O2,
respectively). Solutions of NaOCl and H2O2 were
freshly prepared. Phosphate-buffered saline/Dulbecco (137 mM NaCl, 2.7 mM KCl, 8.1 mM
Na2HPO4, 1.48 mM
KH2PO4, 0.49 mM
MgCl2·6H2O, 0.9 mM
CaCl2) was used as phosphate buffer (phosphate-buffered saline). For cell preparations, a HANKS buffer was used (137 mM NaCl, 5 mM KCl, 5 mM glucose, 2 mM Na2HPO4, 2 mM
KH2PO4, 1.47 mM
MgCl2·, 0.9 mM
CaCl2).
4 mol/liter) and
hypochlorite (10
4 mol/liter), continuously added by a
perfusion pump (6 ml/min). This procedure allows monitoring of the
actual H2O2 concentration in real time by
measuring the luminescence emitted. H2O2
generation by leukocytes was determined using an injection technique as
described previously (17). Briefly, 50 µl of 5 × 10
6 M NaOCl (final concentration) were
injected into 950 µl of a cell suspension containing 5 × 10
5 M luminol. Chemiluminescence was
determined immediately after injection over 2 s. Samples with
known concentrations of H2O2 were used for
calibration. All luminescence measurements were performed using a
AutoLumat LB 953 luminometer (Fa. Berthold, Wildbad, Germany).
7 M/s
H2O2 corresponding to the
H2O2 release by human leukocytes at cell
concentrations found in vivo (see "Results"). Catalase was further added (final activity: k = 0.003 s
1) to prevent accumulation of
H2O2.
RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
1 (with
kcat as the exponential rate constant of
catalase activity), reducing the half-time of
H2O2 to less than 3.5 ms (21). As a direct
consequence, only a few liver cells would be exposed to concentrations
of H2O2 sufficient to activate IRP1 in the
perfusion experiment. This is certainly not the case during the
oxidative burst of neutrophils and macrophages during inflammation,
where H2O2 can be generated continuously over
hours and days and within a more extensive area. To simulate exposure
of the liver to H2O2 in vivo more
closely, the generation of H2O2 in activated
neutrophils was investigated.
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Fig. 1.
Constant IRP activity following the perfusion
of rat liver with H2O2. Rat liver was
perfused in a single path rat liver model with 100 µM
H2O2 for 50 min. Liver samples were taken every
10 min and lysed. Cytoplasmic extracts (25 µg) were analyzed by EMSA
with 25,000 cpm of 32P-labeled IRE probe in the absence
(top panel) or presence of 2% 2-mercaptoethanol
(ME) (bottom panel). The position of the
IRE·IRP complexes and excess free IRE probe is indicated by
arrows.
8 M/s corresponding to 19 nmol
H2O2 (107 cells)
1
min
1 (data not shown). Assuming normal leukocyte
concentrations in the whole blood of about 6 million cells/ml, a
maximum generation rate of 0.2 µM
H2O2/s is calculated. This experimentally
determined H2O2 generation rate was used to
study the activation of IRP1 by H2O2 in the
perfused liver.
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Fig. 2.
Time course of H2O2
concentration in unfractionated leukocytes (a) and in highly diluted
whole blood samples upon stimulation with PMA (b).
a, 20,000 cells/ml ( ) and 105 cells/ml
(
) were stimulated at 0 s with 100 ng/ml PMA, and the
H2O2 concentration was determined at different
time points using the luminol/hypochlorite assay (see "Materials and
Methods"). b, in this representative experiment, blood
from healthy donors was diluted 1:1000 with buffer, treated with 100 ng/ml PMA, and the H2O2 concentration was
determined at different time points using the luminol/hypochlorite
assay (see "Materials and Methods"). The depicted experiment is
representative of three independent determinations.
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Fig. 3.
Activation of IRP by a continuous flux of
H2O2 in the perfused rat liver. Liver of
male Wistar rats was perfused in a single pass rat liver model with GOX
for 50 min using Krebs-Henseleit buffer containing 5 mM
glucose. At this glucose concentration, H2O2
generation rate was determined to be 1.9 × 10 7
M/s corresponding to quantities found in vivo.
Liver samples were taken every 10 min and lysed. Cytoplasmic extracts
(25 µg) were analyzed by EMSA with 25,000 cpm of
32P-labeled IRE probe in the absence (top panel)
or presence of 2% 2-mercaptoethanol (ME, bottom
panel). Arrows indicate the position of the IRE·IRP
complexes and of excess free IRE probe. The depicted experiment is
representative of five independent measurements.
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Fig. 4.
H2O2 activates IRP1
but not IRP2 in the perfused rat liver. The liver of a male Wistar
rat was perfused in a single path rat liver model with GOX for 50 min
as described in Fig. 3. Cytoplasmic extracts (25 µg) were analyzed by
EMSA with 25,000 cpm of 32P-labeled probes specific for
IRP1/2, IRP1, or IRP2 in the absence (top panel) or presence
of 2% 2-mercaptoethanol (ME, bottom panel). The
position of the respective IRE·IRP1 complexes and of excess free IRE
probe is indicated by arrows. A positive control (HepG2
lysate) in the right panel indicates the position of
IRE·IRP2 complexes and demonstrates the function of the IRP2-specific
probe.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES
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ACKNOWLEDGEMENTS |
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We thank Dr. Lucas K
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FOOTNOTES |
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* This work was supported by Grant SFB 601 from the Deutsche Forschungsgemeinschaft.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ To whom correspondence should be addressed. Tel.: 49 6221 56 8611/12; Fax: 49/6221 40 83 66; E-mail: sebastian. mueller@urz.uni-heidelberg.de.
Present address: Lady Davis Inst. for Medical Research, Sir
Mortimer B. Davis Jewish General Hospital, 3755 Cote-Ste-Catherine Road, Montreal, Quebec H3T 1E2 Canada.
Published, JBC Papers in Press, April 10, 2001, DOI 10.1074/jbc.M100654200
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ABBREVIATIONS |
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The abbreviations used are: ROI, reactive oxygen intermediates; IRE, iron responsive element; IRP, iron regulatory protein; EMSA, electrophoretic mobility shift assay; PMA, phorbol myristate acetate; TfR, transferrin receptor; GOX, glucose oxidase.
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