4′-Epidoxorubicin To Re-explore Anthracycline Degradation in

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Communication 4′-Epidoxorubicin To Re-explore Anthracycline Degradation in Cardiomyocytes Pierantonio Menna,† Emanuela Salvatorelli,† and Giorgio Minotti* Integrated Research Center and Laboratory of Drug Sciences, UniVersity Campus Bio-Medico, and Fondazione Alberto Sordi-Research Institute on Aging, 00128 Rome, Italy ReceiVed February 3, 2009

Cardiotoxicity limits the clinical use of doxorubicin (DOX) and other quinone-hydroquinone antitumor anthracyclines. One-electron reduction of the quinone moiety is followed by the formation of reactive oxygen species (ROS) that have been proposed to induce cardiotoxicity through an oxidative stress; conversely, one-electron oxidation of the hydroquinone moiety by hydrogen peroxide (H2O2) and oxyferrous myoglobin (MbIIO2) is followed by an anthracycline degradation process that has been proposed to limit cardiotoxicity. We previously reported that tert-butoxycarbonyl-alanine (t-BA) impeded DOX oxidation/degradation by H2O2/MbIIO2 in a cell-free system; accordingly, t-BA increased the levels of DOX, its conversion to ROS, and its concentration-related toxicity in cardiomyocytes. To re-explore methodological and toxicological aspects of anthracycline degradation, we used 4′-epidoxorubicin (EPI), an anthracycline analogue that is very similar to DOX but undergoes protonation-sequestration in cytoplasmic acidic organelles. t-BA lacked an effect on H9c2 cardiomyocytes exposed to EPI; however, blocking the protonation-sequestration mechanism with the vacuolar H+-ATPase inhibitor, bafilomycin A1 (BFL), enabled t-BA to increase the cellular levels of EPI, its conversion to ROS, and its concentrationrelated toxicity. This suggested that t-BA was specific enough to increase the cellular levels and toxicity of only those anthracyclines that were liable to oxidation/degradation by H2O2/MbIIO2. By exposing cardiomyocytes to nontoxic concentrations of DOX or EPI and by increasing their cellular levels by means of appropriate combinations with t-BA, BFL, or t-BA+BFL, we nonetheless found that the loss of cardiomyocyte viability correlated with the accumulation of undegraded anthrayclines but not with their ability to form ROS or to induce lipid peroxidation. This suggested that an accumulation of undegraded anthracyclines might induce cardiotoxicity also by mechanisms independent of ROS and oxidative stress. Thus, EPI proved useful to refine the value of t-BA in the studies of anthracycline degradation and to reappraise the role of anthracycline degradation in cardiotoxicity. Introduction 1

The clinical value of doxorubicin (DOX) and other antitumor anthracyclines is limited by cardiotoxicity (1). DOX is composed of a methoxy-substituted tetracyclic ring with adjacent quinonehydroquinone moieties, a short side chain, and a sugar with a protonatable -NH2 residue (Figure 1A). One-electron reduction of the quinone moiety, catalyzed primarily by mitochondrial reductases, causes formation of a semiquinone free radical that regenerates its parent quinone by reducing oxygen to reactive oxygen species (ROS) such as superoxide anion and its dismutation product, hydrogen peroxide (H2O2). There is * To whom correspondence should be addressed. Tel: 011-39-06225419109. Fax: 011-39-06-22541456. E-mail: [email protected]. † P.M. and E.S. contributed equally to this paper. 1 Abbreviations: DOX, doxorubicin [7-(3-amino-2,3,6-trideoxy-R-L-lyxohexopyranosyl)doxorubicinone]; ROS, reactive oxygen species; H2O2, hydrogen peroxide; MbIIO2, oxyferrous myoglobin; MbIVdO, ferrylmyoglobin; t-BA, tert-butoxycarbonyl-alanine; EPI, 4′-epidoxorubicin (epirubicin) [7-(3-amino-2,3,6-trideoxy-R-L-arabino-hexopyranosyl)doxorubicinone]; DCFH-(DA), dichlorofluorescin-(diacetate); DCF, dichlorofluorescein; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; MbIII, metmyoglobin; BFL, bafilomycin (type A1); HPLC, high-performance liquid chromatography; MDA, malondialdehyde.

controversy about whether ROS formation and oxidative stress were the only or prevailing mechanisms of anthracycline-related cardiotoxicity (1-5); in fact, anthracyclines might cause toxicity by direct mechanisms if they accumulate above a threshold (6-9). DOX is also liable to an one-electron oxidation of the hydroquinone in juxtaposition to the quinone. This process is followed by the opening and degradation of the tetracyclic ring system of DOX, as evidenced by a dissipation of the optical, fluorescent, and chromatographic properties of its quinonehydroquinone chromophore (10-12). The precise chemistry of anthracycline degradation is unkown, as is the nature of the terminal product(s) of such a process. An in vitro oxidation of DOX with the compound I of many authentic or pseudoperoxidases was accompanied by the formation of 3-methoxyphthalic acid, a product of oxidative modifications of the methoxy-substituted D ring of DOX (11-14). Unfortunately, neither 3-methoxyphthalic acid nor other degradation products could be identified when DOX oxidized with H2O2-activated oxyferrous myoglobin (MbIIO2), which is the prevailing catalyst

10.1021/tx900039p CCC: $40.75  2009 American Chemical Society Published on Web 04/27/2009

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Figure 1. Molecules of interest in this study: DOX (A), t-BA (B), and EPI (C) (the arrow indicates the axial-to-equatorial epimerization of the hydroxyl group at C-4′ in the aminosugar).

of anthracycline degradation in cardiomyocytes, and only forms a ferrylmyoglobin (MbIVdO) formally similar to compound II (12). Absent biochemical indices of DOX degradation induced by MbIVdO, the role of this process in cardiomyocytes could only be explored by indirect approaches. Whole organic extracts of MbIVdO-degraded DOX were shown to lack toxicity to isolated cardiomyocytes under conditions in which DOX per se induced ROS formation and cellular damage (12). This suggested that the products of DOX degradation might be considered nontoxic, but uncertainties about the structure of such products did not allow us to establish whether they diffused in cardiomyocytes and reached the same sites as those reached by DOX under conditions of cellular damage. Thus, direct comparisons of parent DOX with MbIVdO-degraded DOX were not possible. We considered the opportunity to stimulate or inhibit DOX oxidation by means of oxidants or antioxidants, heme poisons, or genetic manipulations of myoglobin or ROS-detoxifying enzymes, but all such approaches were felt to introduce too many changes in the redox balance of cardiomyocytes and anthracycline pharmacokinetics (12, 15, 16). For example, vitamin E and other antioxidants might well protect cardiomyocytes by diminishing the levels of ROS and oxidative stress IVdO (5); however, antioxidants might also reduce Mb and protect anthracyclines from degradation (10, 17, 18), possibly making anthracyclines accumulate undegraded and cause toxicity independent of ROS and oxidative stress. After screening for numerous compounds, we found that DOX degradation could be blocked with tert-butoxycarbonylalanine (t-BA) (Figure 1B). Being a redox-inactive molecule, t-BA neither decomposed H2O2 before it oxidized MbIIO2 to IVdO IVdO nor reduced Mb before it oxidized DOX in vitro; Mb however, t-BA blocked DOX degradation by introducing sterical IVdO and DOX (12). Of note, t-BA did not barriers between Mb

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inhibit DOX degradation induced by many other (pseudo)peroxidases (12). In isolated cardiomyocytes, t-BA did not cause toxicity by its own, did not make cells more vulnerable by endogenous or exogenous sources of ROS, and did not alter DOX uptake or efflux; nevertheless, t-BA augmented the cellular levels of DOX, its conversion to ROS, and its concentration-related toxicity. These results suggested that blocking DOX degradation with t-BA eliminated a salvage pathway, which diminished the levels of DOX, its redox activation to ROS, and its toxicity to cardiomyocytes (12). 4′-Epidoxorubicin (epirubicin, EPI) differs from DOX in an axial-to-equatorial epimerization of the hydroxyl group at C-4′ in daunosamine (Figure 1C). EPI is appreciably more lipophilic than DOX (octanol:water partitioning coefficients: 2.3 vs 0.8) (19), which makes it liable to diffuse in cytoplasmic acidic organelles such as recycling endosomes, lysosomes, and vesicles of the trans-Golgi network. Protonation of the aminosugar in these organelles then makes EPI too polar to diffuse back to cytosol, causing the so-called vesicular sequestration of EPI (20). Protonation-sequestration diverted EPI from the mitochondrial sites of ROS formation, possibly explaining how EPI proved less cardiotoxic than DOX in many clinical conditions (1, 21). In principle, protonation-sequestration might also divert EPI from oxidation with MbIVdO in the cytosol. Here, we characterized the effects of t-BA in cardiomyocytes treated with EPI. By performing the experiments in the absence or presence of an inhibitor of vesicular acidification, we probed the specificity with which t-BA modulated the cellular fate of EPI that underwent protonation-sequestration in the vesicles or oxidation/degradation with MbIVdO in the cytosol. By the same approach, we could also reappraise the different toxicity of EPI vs DOX and the relative importance of anthracycline per se or ROS formation and oxidative stress in these settings.

Experimental Procedures Chemicals. DOX [7-(3-amino-2,3,6-trideoxy-R-L-lyxo-hexopyranosyl)doxorubicinone] and EPI [7-(3-amino-2,3,6-trideoxy-R-Larabino-hexopyranosyl)doxorubicinone] were obtained through the courtesy of Nerviano Medical Sciences (Milan, Italy). Dichlorofluorescin-diacetate (DCFH-DA) and dichlorofluorescein (DCF) were from Molecular Probes (Invitrogen, Carlsbad, CA), while t-BA and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) were from Fluka (Milan, Italy). Horse heart metmyoglobin (MbIII), 1,1,3,3-tetramethoxypropane, 2-thiobarbituric acid, and all other chemicals were from Sigma Aldrich (Milan, Italy). MbIIO2 was prepared by ascorbate reduction of MbIII and quantitated by assuming ε581 nm ) 11 mM-1 cm-1 (22). Spectrophotometric Assays for MbIVdO-Dependent Anthracycline Degradation. We adopted previously validated procedures (12). The incubations (1 mL final volume) were prepared to contain 10 µM DOX or EPI and equimolar MbIIO2. The spectrum and the net absorbance of the anthracycline chromophore (ε477 nm ) 12.11 mM cm-1) were recorded against a reference cuvette that contained MbIIO2 only. Anthracycline degradation was started by adding 20 µM H2O2 to convert MbIIO2 to MbIVdO, and the decay of the anthracycline chromophore was recorded every 2-3 min for 20 min. H2O2 was also added to the reference cuvette that contained MbIIO2 but not DOX or EPI; this was done to constantly correct the spectrum of the anthracycline for the two peaks of MbIVdO at 546 and 586 nm (12). Where indicated, the incubations contained 25-50 µM t-BA. All of the experiments were carried out at 37 °C in 0.3 M NaCl, pH 7.0, so as to avoid interferences of the most common buffers with anthracycline redox reactions (10, 12). Cell Cultures and Treatments. We used H9c2 cardiomyocytes (American Type Culture Collection-CRL 1446), which contained 1.4 ( 0.2 nmol of myoglobin/mg of cytosolic proteins (n ) 4).

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Cells were grown, plated subconfluently (50 × 103) on uncoated 35 mm polystyrene dishes, and then incubated for 16 h in Dulbecco’s modified minimum essential medium added with 0.01-50 µM anthracyclines, precisely as described (12). Where indicated, anthracycline treatment was preceded by preloading of cardiomyocytes with t-BA or bafilomycin (BFL), an inhibitor of the H+-ATPase that acidifies cytoplasmic organelles (23). On the basis of earlier characterizations, 10 mM or 50 nM concentrations of t-BA and BFL were used, respectively; 10 nM BFL was also included during 16 h of exposure of cardiomyocytes to anthracyclines to ensure a long-lasting inhibition of the vacuolar H+-ATPase (12, 20). To measure basal or anthracycline-augmented ROS formation, separate sets of cells were loaded for 40 min with 10 µM DCFH-DA; next, the medium was removed and replaced with fresh medium added with anthracyclines. Where indicated, the loading with DCFH-DA was followed by a 2 h loading with t-BA and/or BFL prior to the final exposure to anthracyclines. High-Performance Liquid Chromatography (HPLC) Assays for Anthracyclines, DCF-Detectable H2O2, and Malondialdehyde (MDA). At the end of experiments, cells from 10 identical dishes were scraped with H2O, homogenized, and extracted with a 4-fold volume of (1:1) CHCl3/CH3OH. The organic phases were combined, and 100 µL was analyzed by reversed-phase HPLC in a HewlettPackard 1100 system (Palo Alto, CA). The extracts were loaded onto a Macrosphere RP 300 C-18 column (250 mm × 4.6 mm, 5 µm/Alltech Associates, Inc., Deerfield, IL), operated at 25 °C and eluted at a flow rate of 1 mL/min for a total 25 min run time (15 min linear gradient from 100% 50 mM NaH2PO4, pH 4.0, to 50-50% CH3CN-50 mM NaH2PO4 followed by a 10 min isocratic elution with 50-50% CH3CN-50 mM NaH2PO4). DOX (retention time ) 14.3 min) and EPI (retention time ) 14.9 min) were detected fluorimetrically (excitation at 477 nm/emission at 560 nm) and quantified against appropriate standard curves (detection limit ) 0.5 × 10-3 µM). DCF, the product of the oxidation of DCFH with H2O2 and cellular peroxidases or iron traces, was identified by cochromatography with a DCF standard (excitation at 488 nm/ emission at 525 nm, retention time ) 18.3 min, detection limit ) 1 × 10-3 µM). DCF formation was expressed as H2O2 equivalents by assuming that 1 mol of H2O2 oxidized 0.43 mol of DCFH (12, 20). MDA, the product of lipid peroxidation, was assayed by mixing 500 µL of cell homogenates or incubation medium with 15 µL of 2% butylhydroxytoluene and 1 mL of 0.75% thiobarbituric acid. After they were heated at 100 °C for 15 min, the mixtures were cooled on ice and extracted with 1.5 mL of n-butanol. One hundred microliters of the organic phase was loaded onto a HxSil C-18 column (250 mm × 4.6 mm, 5 µm/Hamilton Co., Reno, NV) operated under the same conditions as those described above. The adduct of MDA with thiobarbituric acid (retention time ) 11.65 min) was detected fluorimetrically (excitation at 515 nm/emission at 550 nm). The MDA content of cells or medium was quantified against a standard curve obtained by reacting thiobarbituric acid with known amounts of MDA, prepared by acid hydrolysis of 1,1,3,3-tetramethoxypropane (detection limit ) 1 × 10-3 µM) (24). Other Conditions and Assays. The cell viability was measured by the antimycin A-inhibitable reduction of MTT to formazan, an indicator of mitochondrial function. The assay was performed directly on the cell cultures at the end of their exposure to test compounds, and viability was expressed as the percentage of antimycin A-inhibitable MTT reduction in anthracycline-treated cells as compared with control cells (12). Proteins were measured by the bicinchoninic acid method. Myoglobin was determined by an electrochemiluminescence immunoassay with an Elecsys 2010 Analyzer (Roche Diagnostics), according to the manufacturer’s instructions. Unless otherwise indicated, all of the values were means ( standard errors (SEs) of at least three experiments. Data were analyzed by one-way analysis of variance (ANOVA) followed by Bonferroni’s test for multiple comparisons. The Student’s test was also applied as appropriate. Differences were considered to be significant when P was 10 times more H2O2 than it was measured in these experiments (see also Figure 4A). It follows that the ratio of ∆H2O2 to anthracycline, a concentration-adjusted index of anthracycline redox cycling, plateaued and decreased under conditions when (DOX + t-BA) or (EPI + BFL + t-BA) caused the highest loss of viability (Figure 4B). These results suggested that anthracycline toxicity was not tightly coupled with ROS formation and oxidative stress. To probe this hypothesis while not introducing spurious effects by

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Figure 4. Anthracycline toxicity in H9c2 cardiomocytes. Experimental conditions were as described in the legend to Figure 3. In panel A, cardiomyocytes viability was measured by the antimycin-inhibitable MTT reduction assay and plotted against free anthracycline levels and anthracycline-dependent ∆H2O2. In panel B, cardiomyocytes viability was plotted against the ratio of ∆H2O2 to anthracycline (percent) and ∆MDA in the cells. The values were means ( SE of 3-5 experiments; those without vertical bars had their SE within symbol. ∆H2O2, net increase over H2O2 levels in control cells (0.016 ( 0.004 nmol/mg prot); ∆MDA, net increase over MDA levels in control cardiomyocytes (0.022 ( 0.004 nmol/mg prot).

vitamin E or other antioxidants, we measured the lipid peroxidation product, MDA. Lipid peroxidation has been proposed to mediate cardiotoxicity induced by anthracyclines and oxidative stress (5). In comparison with control incubations, DOX or the combinations of (EPI + BFL), (DOX + t-BA), or (EPI + BFL + t-BA) did not induce a release and net increase of MDA (∆MDA) in the medium (not shown); in contrast, cardiomyocytes showed ∆MDA values that increased with DOX and (EPI + BFL) but decreased when (DOX + t-BA) or (EPI + BFL + t-BA) caused the highest levels of free anthracycline and loss of viability (see also Figure 4B). Thus, lipid peroxidation too was poorly coupled with cardiomyocyte toxicity induced by free DOX or EPI.

Discussion EPI proved useful to refine the value of t-BA in studies of anthracycline degradation. In fact, t-BA inhibited DOX degradation in both cell-free systems and cardiomyocytes, as evidenced by an increase of the cellular levels of DOX, its conversion to ROS, and its concentration-related toxicity; by contrast, t-BA inhibited the degradation of EPI in cell-free systems but not in cardiomyocytes, unless these cells had been treated with BFL to prevent the protonation-sequestration of EPI in cytoplasmic acidic organelles. These findings show that t-BA is specific enough to modulate the cellular fate of only those anthracyclines that were freely moveable in the cell and liable to degradation by MbIVdO in the cytoplasmic milieu. On a different note, these studies suggest that t-BA would be specific enough to probe the cardiac distribution and degradation of anthracyclines also in more elaborate animal models. When considering future studies in that direction, one may want to clarify that micromolar

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t-BA was high enough to inhibit anthracycline degradation in cell-free systems, whereas an effect of t-BA in cardiomyocytes usually occurred over a millimolar range (cf. ref 12 and this study). This discrepancy is explained by the limited partitioning (e2%) of t-BA from extracellular fluids to cardiomyocytes (12). Pharmacokinetic and safety studies of high dose t-BA should therefore precede toxicologic evaluation of t-BA/anthracycline schedules in laboratory animals. Improving the intracellular diffusion of t-BA through esterification of its carboxylate might also be considered. The experiments described in this communication offered an opportunity to also re-explore some aspects of cardiotoxicity induced by EPI or by anthracyclines in general. Under conditions of vesicular acidification, EPI was much less toxic than DOX; however, inhibition of vesicular acidification enabled EPI to induce toxicity that was further aggravated by a t-BA blockade of its degradation, similar to what observed with DOX. These results identify vesicular sequestration as a major determinant of the reduced cardiotoxicity of EPI vs DOX. It is noteworthy that vesicular sequestration would diminish the toxicity of EPI in cardiomyocytes but not in cancer cells, which usually exhibit a defective acidification of cytoplasmic organelles (29); anthracycline protonation-sequestration only occurred in cancer cells that regained vesicular acidification following acquisition of a multidrug resistant phenotype (30). These notions should be kept in mind when considering strategies for improving antitumor therapy. For example, proton pump inhibitors currently used in the antiacid treatment of peptic disease have been considered as potential tools to increase the sensitiveness of drug-resistant cells that sequestered anthracyclines in acidic organelles (31). Our experiments with 0.5 µM anthracyclines show that BFL actually made EPI reach higher levels, induce more ROS formation and lipid peroxidation, and cause more toxicity than we observed with DOX (cf. Figures 3 and 4). This may have been caused by the higher lipophilicity of EPI and its better partitioning toward the mitochondrial sites of ROS formation under conditions of inhibited vesicular sequestration. These observations caution that pharmacological inhibition of the vesicular sequestration of EPI in tumors might paradoxically render EPI more cardiotoxic than DOX. Aggravation of DOX or EPI toxicity by t-BA or (t-BA + BFL), respectively, shows that anthracycline degradation serves a salvage pathway for diminishing the levels and noxious effects of these drugs in cardiomyocytes; therefore, the toxicokinetics of anthracycline-based therapies are more complex than commonly believed. By having said of the possible unfavorable interactions of EPI with BFL-like drugs, one should consider that drugs administered in combination with anthracyclines might also induce t-BA-like effects. It is not known which particular drug(s) would be able to induce t-BA-like effects, but the higher than expected cardiotoxicity that often complicates multiagent therapies may well fit in this scenario. Cardiomyocyte toxicity induced by modulating the degradation and cellular levels of DOX or EPI correlated linearly with an accumulation of either anthracycline; however, conditions that favored the highest cellular levels of anthracyclines and toxicity were accompanied by a decrease of anthracycline redox cycling and lipid peroxidation (cf. Figure 4A,B). This might suggest that the contribution of ROS and oxidative stress to the mechanisms of cardiotoxicity is only marginal or confined to a narrow range of anthracycline levels in cardiomyocytes. Redox-independent mechanisms of anthracycline cardiotoxicity have been described and shown to include inactivation of calcium-handling proteins and energy metabolism enzymes,

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chaotropic effects in mitochondria, suppression of gene expression programs, and others (5-9, 32). It is in keeping with these notions that antioxidants often failed to protect against cardiotoxicity in preclinical studies (2, 3) as well as in controlled clinical trials (4, 5). The multifactorial nature of anthracycline cardiotoxicity cautions against a definitive interpretation of its mechanisms. The results of this communication support a possible role for anthracyclines per se, or metabolites other than ROS, and suggest that modulation of anthracycline vesicular sequestration or degradation may serve a valuable approach to reappraise this subject matter in future studies. Acknowledgment. This work was supported by the Associazione Italiana Ricerca sul Cancro and University Campus Bio-Medico (Special Project “CardioOncology”).

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