ARCHIVES

Vol.

OF BIOCHEMISTRY

283, No. 2, December,

AND

BIOPHYSICS

pp. 537-541,

1990

COMMUNICATION Nitric Oxide Mediates David

W. Reif’

Biology Department,

Received

July

Iron Release from Ferritin

and Roy D. Simmons Fisons Pharmaceuticals,

16, 1990, and in revised

form

755 Jefferson Road, Rochester, New York 14603

September

16, 1990

Nitric oxide (NO) synthesis by cytotoxic activated macrophages has been postulated to result in a progressive loss of iron from tumor target cells as well as inhibition of mitochondrial respiration and DNA synthesis. In the present study, the addition of an NO-generating agent, sodium nitroprusside, to the iron storage protein ferritin resulted in the release of iron from ferritin and the released iron-catalyzed lipid peroxidation. Hemoglobin, which binds NO, and superoxide anion, which reacts with NO, inhibited nitroprusside-dependent iron release from ferritin, thereby providing evidence that NO can mobilize iron from ferritin. These results suggest that NO generation in uivo could lead to the mobilization of iron from ferritin disrupting intracellular iron homeostasis and increasing the level of reactive oxygen species. 0 1990

Academic

Press,

Inc.

It has been known for many years that macrophages are cytotoxic to tumor cells in culture when the macrophages are activated by tumor necrosis factor or interferon-y, in combination with a second signal such as Escherichia coli lipopolysaccharide (l-3). One aspect of this cytotoxicity is a selective inhibition of certain metabolic functions, an effect which is also observed in the activated macrophage effector cells (4). The mechanism of this metabolic inhibition is unknown, but recent evidence suggests that nitric oxide (NO) production by the activated macrophages may contribute to the cytotoxic effects. Stuehr and Marletta (5-7) first reported that activated macrophages produced nitrite and nitrate and subsequently demonstrated that NO was an intermediate in their production (8). L-Arginine is required for NO production as well as the biochemical changes associated with activated macrophage cytotoxicity (9,lO). These effects can be blocked by NG-monomethylL-arginine (g-11), which inhibits the NO synthetase enzyme. Target cells previously loaded with radiolabeled iron lose a major fraction (64% of labeled iron) of their cellular (12, 13)

1 To whom

correspondence

should

0003-9861/90 $3.00 Copyright 0 1990 by Academic Press, All rights of reproduction in any form

be addressed.

and mitochondrial (13) iron as a result of cocultivation with cytotoxic activated macrophages. Granger and Lehninger (14) demonstrated that activated macrophages also cause a selective inhibition of the iron-dependent, mitochondrial enzymes NADH coenzyme Q reductase and succinate coenzyme Q reductase. Additionally, activated macrophages inhibit aconitase in the target cells as a result of the loss of an iron atom from the 4Fe-4S cluster within the aconitase active site (15). Aconitase activity could be recovered if the damaged target cells were subsequently incubated in the presence of ferrous, suggesting that the cells had lost the ability to utilize intracellular stores of iron. The ability of activated macrophages to inhibit non-heme iron-dependent enzymes as well as to synthesize NO, led Lancaster and Hibbs (4) to speculate that mitochondrial inhibition may be a result of a direct interaction between NO and the FeS centers of these enzymes. Indeed, using ESR spectroscopy, they observed iron-nitrosyl complexes in cytotoxic activated macrophage effector cells. The source of this iron is unknown, but these results provide a direct link between NO generation and a disruption in intracellular iron utilization. The relatively large amount of iron that is lost from target cells led Wharton et al. (13) to suggest that some of this iron may arise from the iron storage protein ferritin. Ferritin stores iron as ferric and can store up to 4500 atoms of Fe/molecule (16). The physiological reductant(s) is unknown, but in recent years the ability of numerous toxicological reductants to mobilize ferritin iron has been reported (17, 18). Once released, the iron can react with cellular oxidants and result in the oxidation of critical cellular constituents (19). Since NO interacts with nonheme iron and the majority of intracellular nonheme iron is stored within ferritin, we investigated the ability of NO to mobilize iron from ferritin using sodium nitroprusside as the source of NO. We report here that NO releases iron from ferritin in uitro and the released iron catalyzes lipid peroxidation. These results suggest an additional effect of NO generation in uiuo and may provide a partial mechanism that explains the loss of intracellular iron reported by Hibbs et al. (12). MATERIALS

AND

METHODS

Chemicals. Xanthine, glucose, (2-pyridylj-5,6-bis(4-phenylsulfonic (2.hydroxyethyl)-1-piperazineethanesulfonic

sodium hydrosulfite (dithionite), 3acid)-1,2,4-triazine (ferrozine), 4acid (Hepes), and Sephadex

537 Inc. reserved.

538

REIF

AND

G-25 and G-100 were from Sigma. Sodium nitroferricyanide (nitroprusside) was from Baker, Inc. (Phillipsburg, NJ). Chelex 100 was obtained from Bio-Rad (Richmond, CA). Distilled deionized water was purified with a Milli Q Water Purification System (resistivity 2 16 Mohm/cm). All reagents were prepared in 50 mM Hepes, pH 7, which had been chromatographed over Chelex 100 to remove trace metal impurities. Proteins. Glucose oxidase (EC 1.1.3.4), xanthine oxidase (EC 1.1.3.22) grade III from buttermilk, bovine erythrocyte superoxide dismutase (EC 1.15.1.1), catalase (EC 1.11.1.6), human hemoglobin, and horse spleen ferritin were purchased from Sigma. Glucose oxidase and ferritin were incubated with 10 mM EDTA for 1 h on ice followed by chromatography over a Sephadex G-25 column equilibrated in 50 mM Hepes, pH 7, prior to use (17). Human hemoglobin was treated with dithionite to obtain reduced hemoglobin as described (20) and quantified as described (21). Xanthine oxidase was purified by chromatography over a G-100 column equilibrated in 50 mM Hepes, pH 7, and assayed according to McCord and Fridovich (22). Superoxide dismutase was assayed according to McCord and Fridovich (22) and catalase according to Beers and Sizer (23). Iron release assay. Iron release from ferritin was measured utilizing the ferrous chelator ferrozine and the amount of iron released calculated using cr,62 = 27.9 mM-‘cm-’ (24). Reaction mixtures (1.0 ml final volume) in 50 mM Hepes, pH 7.0, contained 500 ~.LM ferrozine and other reactants as indicated in the figure and table legends.. Sodium nitroprusside (SNP)’ was prepared in argon-purged 50 mM Hepes, pH 7.0, stored on ice, and used within 4 h. All reactions were performed at 37”C, utilized a reference cuvette that contained all reactants except ferritin, and were initiated by the addition of SNP. The increase in absorbance at 562 nm was continuously monitored using a Beckman Model 35 spectrophotometer. For the anaerobic experiments, all solutions were purged with argon and glucose and glucose oxidase were included to scavenge any remaining Op. Lipid peronidation assay. Rat liver microsomal phospholipid was isolated and liposomes were prepared anaerobically as described (19). Reaction mixtures (2.5 ml final volume) in 50 mM Hepes, pH 7.0, contained phospholipid liposomes (1 pmol/ml) and other reactants as described in the legend to Table III. The extent of lipid peroxidation following a 20-min incubation at 37°C was determined as the amount of malondialdehyde (MDA) formed as assayed by the thiobarbituric acid assay (25). Statistical analysis. The data in Tables II and III and Figs. 1 and 2 were analyzed by ANOVA and the data in Table I by Analysis of Covariance. Significance was designated at the P < 0.01 level.

RESULTS Sodium nitroprusside releases NO spontaneously and provides a continuous flux of NO (26). The addition of increasing concentrations of SNP (0.1-10 mM) to horse spleen ferritin in the presence of ferrozine (an iron chelator) resulted in a corresponding increase in the formation of the Fe(II):(ferrozine), complex, indicating that iron is being released from ferritin (Fig. 1). The rate of iron release from ferritin is dependent upon the concentration of SNP and is linear over a 30-min incubation period (P < 0.01). At 1 mM SNP, NO is released at a rate of 110 nM min-’ (26). Thus, assuming that a mole of NO could mobilize a mole of iron from ferritin, then NO exhibits an efficiency of approximately 32% in this test system.

2 Abbreviations used: SNP, sodium of variance; MDA, malondialdehyde; laxation factor.

nitroprusside; ANOVA, EDRF, endothelium-derived

analysis re-

SIMMONS

1.6 2

10

20 TIME

30

(minutes)

FIG. 1. Effect of sodium nitroprusside on iron release from ferritin. Reaction mixtures (1 ml final volume) in 50 mM Hepes, pH 7.0, contained 500 pM ferrozine, 200 pM Fe as horse spleen ferritin iron (0.1 pM protein), and sodium nitroprusside (0,O.l mM; n , 1.0 mM; 0,lO mM). The reaction was initiated with nitroprusside and the amount of iron released determined as described under Material and Methods. Values presented are the means + SE of five replicate incubations from two experiments.

Since the reaction of NO with O2 proceeds rapidly, the ability of NO to release iron from ferritin should be attenuated by the O2 present in the system. Accordingly, when the reactions were performed in an anaerobic environment, an enhanced (P < 0.01) rate of SNP-dependent iron release from ferritin was detected (Table I). The rate of iron release from ferritin anaerobically was consistently between 20 and 40% greater than the rate aerobically (P < 0.01). Increasing the amount of ferritin iron present (25-200 FM) also resulted in an increase in the amount of iron released from ferritin (Table I). Hemoglobin binds NO and has been employed as a functional antagonist to confirm the involvement of NO (27). Thus, the addition of hemoglobin (l-25 FM) to incubations containing 1 mM SNP and 200 PM ferritin iron resulted in the inhibition (P < 0.01) of iron release from ferritin (Fig. 2). In contrast to NO, neither sodium nitrite (1 mM) nor sodium nitrate (1 mM) was capable of releasing iron from ferritin (data not shown). These results strongly suggest that either NO or some species derived from NO directly mobilizes iron from ferritin. Another criterion for establishing the involvement of NO is inactivation by superoxide anion (27). As shown in Table II, the combination of SNP and xanthine/xanthine oxidase decreased the iron released as compared to either the SNP-dependent system or the superoxide-dependent system (P < 0.01). Since superoxide also mobilizes iron from ferritin (19, 28), then the products of the reaction of NO and superoxide, N204 and Nz03, as well as their hydrolysis products NO*- and NOBe (as mentioned above), do not appear to release iron from ferritin. SNP (1 mM) did not affect the activity of xanthine oxidase as assayed by urate production (data not shown). The inclusion of superoxide dismutase (100 U/ml) to scavenge superoxide increased iron release in the presence of both SNP and xanthine/xanthine oxidase to a level approximating the iron release observed with SNP alone. Superoxide dismutase completely inhibited iron release in the xanthine/xanthine oxidase system (Table II), but

NITRIC TABLE

OXIDE

MEDIATES

IRON

25 50

System” Anaerobic

0.18 + 0.02 0.30 + 0.01

100

0.49

200

0.75 I!z 0.03

0.29 0.44 0.58 0.90

f 0.01

f 0.02 * 0.02 + 0.02 +_ 0.04

Note. The aerobic system was identical to those described in the legend for Fig. 1 and utilized 1 mM sodium nitroprusside with the indicated concentrations of ferritin iron. Anaerobiosis was achieved by purging the solutions in the cuvette with argon for 5 min. The anaerobic reaction mixture also contained catalase (100 U/ml) to prevent iron oxidation by hydrogen peroxide, and glucose (5 mM) and glucose oxidase (15 U/ ml) to scavenge traces of Or. Iron release was determined for 20 min as described under Materials and Methods and the values presented are the means f SE of replicate (n = 5) incubations of two experiments.

had no effect on the SNP-dependent system (data not shown). These results provide additional evidence that No is capable of mobilizing iron from ferritin and support the finding that No does not appear to complex with the Type 2 copper of SOD (29). To evaluate the potential biological significance of NO-dependent iron release from ferritin, the effect of SNP-dependent iron release from ferritin in a model lipid peroxidation system was investigated. As shown in Table III, SNP-dependent iron release is sufficient to catalyze lipid peroxidation. The extent of lipid peroxidation detected increased more than 16-fold in the presence of both ferritin and SNP (P < 0.01). Parallel experi-

Iron

Nitroprusside Xanthine oxidase Nitroprusside and xanthine oxidase Nitroprusside, xanthine oxidase, and SOD Xanthine oxidase plus SOD

z 2 Ei

0.6

2 Y 2

0.4

1

L

8 fi

0.2

-

0

0.67 0.83 0.44 0.60

I1 I

1

HEMOGLOBIN

10

1

25

(PM)

FIG. 2. Inhibition of sodium nitroprusside-dependent iron release from ferritin by hemoglobin. The reaction mixtures were identical to those described in the legend to Fig. 1 and utilized 1 mM sodium nitroprusside. Oxhemoglobin (tetramer) was added to the reaction at the indicated concentrations. Iron release was determined for 20 min as described under Materials and Methods and the values presented are the means i SE of five replicate incubations from two experiments.

f f f + ND*

(PM)

0.03 0.02 0.03 0.02

ments utilizing xanthine oxidase and xanthine to generate superoxide at a rate resulting in an equivalent rate of iron release from ferritin as the SNP system resulted in similar extents of lipid peroxidation (Table III) (P < 0.01). The strikingly similar results between SNP-dependent and superoxide-dependent lipid peroxidation support the premise that NO-dependent iron release from ferritin could be significantly damaging to biological membranes. DISCUSSION Cytotoxic activated macrophages produce NO, which results in a loss of iron from tumor target cells (12, 13) and inhibition

Sodium

III

Nitroprussideand Ferritin-Dependent Lipid Peroxidation MDA

1,

released

’ The reactions in 50 mM Hepes, pH 7.0, contained 200 PM ferritin iron, 0.5 mM ferrozine, and where indicated, 1 mM sodium nitroprusside, 0.33 mM xanthine, 1 mU xanthine oxidase, and 100 U/ml superoxide dismutase (SOD). Iron release from ferritin was determined as described under Materials and Methods following a 20-min incubation and the values presented are the means + SE of five replicate incubations of two experiments. * Rate not detectable.

TABLE

0.8

II

Effect of Superoxide on Sodium Nitroprusside-Dependent Iron Release from Ferritin

released (PM)

Aerobic

(PM)

539

FERRITIN TABLE

Iron iron

FROM

I

Effect of Ferritin Iron Concentration and O2 on Sodium Nitroprusside-Dependent Iron Release

Ferritin

RELEASE

Additions None Complete -Ferritin -Nitroprusside or superoxide

Nitroprusside

b

(PM)”

Superoxide’

0.08 + 0.01 1.34 f 0.05 0.19 Zk 0.03

0.07 k 0.01 1.50 f 0.05 0.21 2 0.02

0.09 + 0.03

0.13 2 0.01

o All values presented are the means + SE of five replicate incubations from two experiments. * Reaction mixtures (2.5 ml final volume) in 50 mM Hepes, pH 7.0, contained 1 pmol/ml rat liver microsomal phospholipid liposomes, horse spleen ferritin (200 PM Fe), and sodium nitroprusside (1 mM) as indicated. MDA formation at 20 min was assayed as described under Materials and Methods. ’ These incubations were identical to the nitroprusside system with the omission of the nitroprusside and the inclusion of 0.33 mM xanthine and 1 mu/ml xanthine oxidase to generate superoxide. Catalase (100 U/ml) was also included to eliminate iron oxidation by hydrogen peroxide.

540

REIF

AND

of iron-dependent enzymes (14, 15). The mechanism by which iron is lost from target cells is presently unknown. However, the amount of iron lost from these target cells is not accounted for simply by losses from iron-dependent enzymes and may result from iron release from ferritin (13). Supporting a role for NO in this process, Lancaster and Hibbs (4), utilizing ESR spectroscopy, recently demonstrated an iron:nitrosyl complex in cytotoxic activated macrophages, but the source of the iron was not identified. The results of the present study demonstrate that NO is capable of releasing iron from ferritin in uitro, which suggests that this may occur in uiuo. In support of the hypothesis that NO can mobilize iron from ferritin, it was demonstrated that both hemoglobin and superoxide anion inhibited SNP-dependent iron release from ferritin. In the latter case, superoxide dismutase reversed the inhibition by superoxide, but did not affect NOdependent iron release from ferritin. Cytotoxic activated macrophages also produce superoxide and it has been shown that superoxide is capable of releasing iron from ferritin (19, 28). Thus, the overall effects of NO production by cytotoxic activated macrophages are complex and will be influenced by the rate of production of NO and the extent of scavenging of superoxide by endogenous superoxide dismutase. The production of NO is not limited to macrophages and neutrophils, but has also been reported to occur in an array of cell types. For instance, nitric oxide generation has been detected in Kiippfer cells and hepatocytes, and Kiippfer cells release another factor that activates hepatocytes to produce NO (30). The production of NO correlated with a decrease in protein synthesis, but not necessarily cell death. NO is also synthesized by the endothelium and has been postulated to be responsible for endothelium-dependent vascular smooth muscle relaxation through the stimulation of guanylate cyclase. NO has been proposed to be endothelium-derived relaxation factor (EDRF) (27). Recently, it has been reported that EDRF may not be NO, but may be a nitrosyl:thiol complex (31). In addition, NO has been shown to inhibit platelet aggregation (20,32), an activity which may be related to the ability of NO to activate a cytosolic ADPribosyltransferase (20). In cerebellar cells, glutamate stimulates NO production, which then stimulates cGMP synthesis (33,34) through the activation of guanylate cyclase. The results of this study suggest an additional novel link between NO generation and the loss of iron from cells exposed to cytotoxic activated macrophages. Thus, NO generation could lead to cellular injury and cytotoxicity by disrupting the normal cellular iron homeostasis in three ways. First, NO inactivates Fe-requiring enzymes important for mitochondrial respiration (14), the citric acid cycle (aconitase) (15), and DNA synthesis (35,36). Second, NO may decrease the intracellular storage pool of iron (within ferritin) hindering the cell’s ability to recover the lost iron-dependent enzymatic activities. This could explain why aconitase activity in damaged cells can be recovered by incubating the cells in the presence of exogenous ferrous as reported by Drapier and Hibbs (15). Finally, NO-dependent mobilization of iron from utilization (iron-dependent enzymes) and from storage sites (ferritin) could increase the generation of reactive oxygen species resulting in uncontrolled and irreversible damage to DNA, proteins, and membrane lipids (37). It is apparent that NO generation serves critical physiological functions and can have beneficial effects, but it is also apparent

SIMMONS

that NO has the capacity to catalyze deleterious to the affinity of NO for transition metals (29). biological effect(s) of NO will depend upon the the rate of NO generation. This may explain why like intracellular calcium concentration, appears regulated (27).

reactions due Therefore, the site as well as NO generation, to be so tightly

REFERENCES 1. Hibbs, J. B., Jr., Lambert, New Biol. 235,48-W. 2. Ruco, 2042.

L. P., and Meltzer,

3. Weinberg, J. B., Chapman, J. Immunol. 121,72-80.

L. H., and Remington, M.

S. (1978)

J. S. (1972)

J. Zmmurwl.

H. A., Jr., and Hibbs,

Nature

121, 2035-

J. B., Jr. (1978)

4. Lancaster, J. R., Jr., and Hibbs, J. B., Jr. (1990) Proc. Natl. Acad. Sci. USA 87, 122331227. 5. Stuehr, D. J., and Marletta, M. A. (1985) Proc. N&l. Acad. Sci. USA 82, 7738-7742. 6. Stuehr, D. J., and Marletta, M. A. (1987) J. Zmmud. 139, 518 525. 7. Stuehr, D. J., and Marletta, M. S. (1987) Cancer Res. 47, 55905594. 8. Marletta, M. A., Yoon, P. S., Iyengar, R., Leaf, C. D., and Wishnok, J. S. (1988) Biochemistry 27,8706-8711. 9. Hibbs, J. B., Jr., Vavrin, Z., and Taintor, R. R. (1987) J. Zmmunol. 38,550-565. 10. Hibbs, J. B., Jr., Taintor, R. R., and Vavrin, Z. (1987) Science 236, 473-476. 11. Drapier, J.-C., and Hibbs, J. B., Jr. (1988) J. Zmmunol. 140,28292838. 12. Hibbs, J. B., Jr., Taintor, R. R., and Vavrin, Z. (1984) Biochem. Biophys. Res. Commun. 123, 716-723. 13. Wharton, M., Granger, D. L., and Durack, D. T. (1988) J. Zmmud. 141,1311-1317. 14. Granger, D. L., and Lehninger, A. L. (1982) J. Cell Biol. 95, 527535. 15. Drapier, J.-C., and Hibbs, J. B., Jr. (1986) J. Clin. Invest. 78, 790797. 16. Harrison, P. M. (1977) Semin. Hematol. 14, 55-70. 17. Thomas, C. E., and Aust, S. D. (1986) J. Bid. Chem. 261, 1306413070. 18. Samokyszyn, V. M., Thomas, C. E., Reif, D. W., Saito, M., and Aust, S. D. (1988) Drug Metab. Reu. 19, 283-303. 19. Thomas, C. E., Morehouse, L. A., and Aust, S. D. (1985) J. Biol. Chem. 260,3275-3280. 20. Salvemini, D., deNucci, G., Gryglewski, R. J., and Vane, J. R. (1989) Proc. Natl. Acad. Sci. USA 86,6328-6332. 21. Winterbourn, C. C. (1985) in Handbook of Methods for Oxygen Radical Research (Greenwald, R. A., Ed.), pp. 137-141, CRC Press, Boca Raton, FL. 22. McCord, J. M., and Fridovich, I. (1969) J. Biol. Chem. 244, 60496055. 23. Beers, R. F., Jr., and Sizer, T. W. (1952) J. Biol. Chem. 195, 133140. 24. Stookey, L. L. (1970) Anal. Chem. 42, 779-781. 25. Buege, J. A., and Aust, S. D. (1977) in Methods in Enzymology (Fleischer, S., and Packer, L., Eds.), Vol. 52, pp, 302-310, Academic Press, New York. 26. Feelisch, M., and Noack, E. (1987) Eur. J. Pharmacol. 142, 465-

469.

NITRIC 27. Moncada, Pharmacol.

OXIDE

S., Radomski, M. W., and Palmer, 37, 24952501.

28. Reif, D. W., Schubert, Biophys. 264, 238-243. 29. Gorren, A. C. F., deBoer, Acta 916,38-47. 30. Billiar, T. R., Curran, Simmons, R. L. (1990)

J., and Aust, E., and Wever,

MEDIATES

R. M. J. (1988)

S. D. (1989) R. (1987)

Arch. B&him.

R. D., Ferrari, F. K., Williams, J. Surgical Res. 48, 349-353.

31. Myers, P. R., Minor, R. L., Jr., Guerra, Harrison, D. G. (1990) Nature (London)

R., Jr., Bates, 345, 161-163.

IRON

Bio&m. Biochem. Biophys. D. L., and J. N., and

RELEASE

FROM

FERRITIN

541

32. Briine, B., and Lapetina, E. G. (1986) J. Biol. Chem. 264, 84558458. 33. Knowles, R. G., Palacios, M., Palmer, R. M. J., and Moncada, S. (1989) Proc. Natl. Acad. Sci. USA 86, 5159-5162. 34. Bredt, D. S., and Snyder, S. H. (1989) Proc. Natl. Acad. Sci. USA 86,9030-9033. 35. Keller, R. (1973) J. Exp. Med. 138, 625-644. 36. Krahenbuhl, J. L., and Remington, J. S. (1974) J. Zmmunol. 113, 507-516. 37. Aust, S. D., Morehouse, L. A., and Thomas, C. E. (1985) J. Free Rad. Biol. Med. 1, 3-25.