IRP1 Activation by Extracellular Oxidative Stress in the Perfused Rat Liver*

Sebastian MuellerDagger §, Kostas Pantopoulos||, Christian A. HübnerDagger , Wolfgang StremmelDagger , and Matthias W. Hentze

From the Dagger  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

    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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.

    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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&cjs1138;2, are inevitable byproducts of aerobiosis, cells have to tightly control intracellular iron levels to minimize iron toxicity and satisfy its metabolic needs. Pathologic conditions of iron overload are associated with tissue injury, neurodegeneration, and malignancy (3). For example, hereditary hemochromatosis, one of the most prevalent human genetic disorders, is caused by a defect in the regulation of iron absorption (4). As there is no physiological mechanism for iron secretion, this defect leads to iron accumulation and gradual deposition in liver (and heart) parenchymal cells. Because of extensive liver iron overload with progressing age, hemochromatosis patients eventually develop liver fibrosis and cirrhosis or hepatocellular carcinoma (5).

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.

    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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 (epsilon 290 = 350 mol liter-1 cm-1 at pH 12 (14) and epsilon 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).

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 (10-4 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).

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 × 10-7 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.

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.

    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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-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.

Generation of H2O2 by Stimulated Neutrophils-- During the oxidative burst of neutrophils, a plasma membrane-bound NADPH oxidase generates O&cjs1138;2, which, at physiological pH, is spontaneously disproportionated to H2O2 and water (25). Superoxide dismutase increases the rate of this reaction almost 10000-fold (26). Compared with O&cjs1138;2, H2O2 constitutes a rather stable oxygen metabolite. To trigger an oxidative burst, we treated a suspension of neutrophils with 100 ng/ml PMA and determined the actual concentration of H2O2 in suspension at different time points (Fig. 2a). Under these conditions, a maximum concentration of H2O2 in the micromolar range (~2 µM H2O2) was reached, confirming earlier observations (17, 27). Interestingly, levels of H2O2 cannot be further elevated by increasing the cell number. At higher cell numbers, H2O2 is also more rapidly degraded upon stimulation with PMA. This degradation is completely inhibited by sodium azide, a known inhibitor of catalase and myeloperoxidase (17, 28, 29), suggesting that these heme-containing enzymes are the main H2O2 consumers (data not shown). Because the degradation rate of H2O2 by catalase depends on the H2O2 concentration, this enzyme seems to restrict the maximum H2O2 concentration in stimulated leukocytes. The inhibition of H2O2 degradation by sodium azide can be utilized to determine the generation rate of H2O2 (in M/s). When 106 neutrophils per ml were stimulated with PMA, we determined the H2O2 generation rate to be 3.2 × 10-8 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 (black-triangle) and 105 cells/ml (black-square) 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.

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.


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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.

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%.


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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
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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&cjs1138;2) and H2O2, has been reported to cause a reversible inhibition of IRP activity (36, 37). In an ischemia-reperfusion model, a pathologic condition that is also thought to involve oxidative stress, the same investigators observed an initial reduction of IRP activity (38).

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.

    ACKNOWLEDGEMENTS

We thank Dr. Lucas K<A><AC>&ogr;</AC><AC>´</AC></A>hn for plasmids and Angelika Weber for technical assistance.

    FOOTNOTES

* 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

    ABBREVIATIONS

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.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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