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ABSORPTION, DISTRIBUTION, METABOLISM, AND EXCRETION
Research Service, Veterans Administration Medical Center, Iowa City, Iowa (K.J.R., L.H.B.); Free Radical and Radiation Biology Program of the Department of Radiation Oncology, University of Iowa Roy J. and Lucille A. Carver College of Medicine, Iowa City, Iowa (K.J.R.); Research Service and Department of Internal Medicine, Veterans Administration Medical Center, Cincinnati, Ohio (B.E.B., K.J.R.); and Departments of Internal Medicine (B.E.B., K.J.R.) and Biochemistry, Molecular Genetics and Microbiology (B.E.B.), University of Cincinnati, Cincinnati, Ohio
Received May 13, 2005; accepted June 22, 2005.
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Abstract |
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; Andrews et al., 2002; Gwyn and Sinicrope, 2002; Thun et al., 2002). These functions are partly due to their inhibition of COX-2, an inducible form of COX that is overexpressed in cancer cells (Hussain et al., 2003).
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). These reactions seemed to be so specific that dihydroxybenzoic acids have been used as an index of generation of ·OH in vitro and in vivo (Floyd et al., 1986; Sagone and Husney, 1987; Davis et al., 1989; Ramos et al., 1992). SA reacts with peroxynitrite (Kaur et al., 1997), quenches singlet oxygen (Kalyanaraman et al., 1993), and inhibits superoxide/NO-dependent low density lipoprotein oxidation (Hermann et al., 1999a).
Similar to other phenolic compounds, SA is a substrate for peroxidases. HRP/H2O2 and methemoglobin/H2O2 oxidize SA to the corresponding phenoxyl radical, as demonstrated using EPR (Shiga and Imaizumi, 1973, 1975). Incubation of SA with metmyoglobin/H2O2 affords 2,3- and 2,5-dihydroxybenzoic acids (Galaris et al., 1988). SA stimulates low density lipoprotein oxidation by MPO/H2O2 through the intermediacy of SA-derived phenoxyl radicals (Hermann et al., 1999b). Ascorbate peroxidase and lactoperoxidase metabolize SA at acidic pH (Kvaratskhelia et al., 1997; Muraoka and Miura, 2005). Stimulated granulocytes induce decarboxylation of SA; however, no major role for MPO in this process was envisaged (Sagone and Husney, 1987).
It has also been reported that NSAIDs modulate the cytotoxic action of anticancer agents (Inchiosa and Smith, 1990; Duffy et al., 1998). This aspect of the biochemistry of NSAIDs is of particular interest given that during chemotherapy, cancer patients may also be administered NSAIDs. Earlier, we have reported that acetaminophen, a phenolic compound and the active ingredient of the popular analgesic drug Tylenol, stimulates oxidation of the anticancer anthracyclines doxorubicin (DXR) and daunorubicin (DNR) by peroxidases (Reszka et al., 2004). Because the reaction leads to degradation of anthracyclines and loss of their anticancer and cytotoxic activities, better understanding of this process and mechanisms involved may be important for clinical oncology. It was of interest to find out whether other phenolic compounds also stimulate oxidative degradation of DNR(DXR). We were particularly interested in SA since it may be used by cancer patients undergoing anthracycline chemotherapy. We report that at pharmacologically relevant concentrations (<2 mM; Stead and Moffat, 1983), SA efficiently stimulates oxidative degradation of DNR(DXR) by LPO(MPO)/H2O2 systems especially at acidic pH. We also show that the peroxidative metabolism of SA gives rise to a redox-active product, presumably of a biphenol type, which also mediates oxidation of anthracyclines.
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240 = 39.4 M-1 cm-1 for H2O2 (Nelson and Kiesow, 1972), 412 = 1.12 x 105 M-1 cm-1 for LPO (Jenzer et al., 1986), and 480 = 1.15 x 104 M-1 cm-1 for DNR(DXR) (Chaires et al., 1982). Because DNR and DXR tend to form dimers in aqueous solutions, in the present study, they were used in low micromolar concentrations (<10 µM) to assure that they were present predominantly as monomers.
Spectrophotometric Measurements. Oxidation of anthracyclines was studied by measuring their absorption spectra at designated time points. The spectra were measured using an Agilent diode array spectrophotometer model 8453 (Agilent Technologies, Palo Alto, CA). Samples were prepared in phosphate buffers (50 mM) for pH 6.0 to 8.0 and acetate buffers (50 mM) for pH < 6. All measurements were performed at room temperature. Typically the reaction was initiated by addition of a small aliquot H2O2 (5 or 10 µl) as the last component to a sample consisting of DNR(DXR), SA, and LPO (or MPO) in buffer solution. Time course measurements were carried out following changes in absorbance at 480 nm (
max for DNR and DXR). Data were collected in 2-, 5-, or 10-s intervals during continuous stirring of the sample in a spectrophotometric cuvette (1-cm light path). All experiments were repeated at least twice.
The initial rate of DNR(DXR) oxidation by MPO/H2O2/SA (Vi) was determined from the initial linear portion of the A480 versus time traces using the method of linear regression. When DNR(DXR) was oxidized by LPO/H2O2 in the presence of SA, curves of a Z-shape were recorded. They were characterized by the maximum rate (Vmax) determined by linear fitting to the portion of the curve with the largest slope.
Oxidation of SA by LPO/H2O2 and MPO/H2O2 was determined by recording absorption spectra of its metabolite showing maximum absorption at 412 nm and by measuring time course of its formation in buffers of various pHs and at various anthracycline concentrations. Concentrations of the neutral form of SA, HOOC-SA-OH, were calculated using the known total concentration of the salicylate, the pKa of the SA carboxylic group of 2.98 (Lide, 2004), and the actual pH of the sample solution.
EPR Measurements. EPR spectra were recorded using a Bruker EMX EPR spectrometer (Bruker Biospin Co., Billerica, MA), operating in X band and equipped with a high-sensitivity resonator ER 4119HS. Samples were prepared in pH 7.1 or 5.1 buffers (total volume, 250 µl), and the reaction was initiated by addition of H2O2 as the last component. To facilitate detection of radicals, 400 µM DNR was used in these experiments. Similar experiments were carried out with DXR. The sample was transferred to a flat aqueous EPR cell, and recording was started 1 min after initiation of the reaction (H2O2 addition). The spectra were recorded using microwave power (40 mW), modulation amplitude (2 G), receiver gain (2 x 106), conversion time (40.96 ms), time constant (81.92 ms), and scan rate (80 G/41.92 s). Spectra shown (see Fig. 10) are average of seven scans and represent results of typical experiments.
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). However, when DXR in pH 7.0 buffer was exposed to MPO/H2O2 in the presence of micromolar concentrations of SA, no changes in the absorption spectrum of DXR were observed, suggesting that under these conditions, DXR was not oxidized.2 In contrast, oxidation of DXR became evident when the reaction was carried out at acidic pH. Figure 2 shows spectra recorded at selected time points following the addition of H2O2 to DXR/MPO/SA at pH 5.46. The decrease in intensity of the drug's characteristic absorption band at 480 nm indicates that the drug undergoes oxidation. When A480 reached a near zero level (indicating that almost all DXR was consumed), a new absorption band with maximum at 412 nm began to emerge. As will be shown later, this new band originates not from DXR, but from SA, and the corresponding metabolite is assigned the symbol X.
Dependence of the reaction on pH was studied next by recording the time course of A480 changes in buffers of various pHs at constant initial concentration of SA of 0.5 mM. It may be seen that the rate of the reaction increases as the pH decreases (Fig. 3, inset A). Because the SA carboxylic group (pKa = 2.98) (Lide, 2004) is the only group that can be affected by changes in pH in the studied pH range of approximately 7 to 5, the observed stimulatory effects are attributed to the higher concentration of the neutral (nonionized) form of salicylic acid, HO-SA-COOH, at acidic pH. Indeed, the initial rate of DXR oxidation (Vi) depends linearly on [HO-SA-COOH], with the latter being calculated for a given pH (Fig. 3, main panel). Similar results were obtained for DNR (data not shown). This dependence of DXR(DNR) oxidation on pH is completely opposite to that observed in the presence of acetaminophen, in which the maximum stimulation was observed at near neutral pH, and no effect was observed at pH 
5 (Reszka et al., 2004). We emphasize that ionization of the DXR(DNR) hydroquinone group does not change in this pH range (pKa of first ionization of the drugs' hydroquinone moiety is 9.5, Razzano et al., 1990); accordingly, its redox potential should remain invariant at these pHs. These results further support the idea that the observed dependence on pH should be linked to ionization status of the cofactor and not the anthracyclines.
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Simultaneously with measurements of drug oxidation, we measured the formation of the species X versus pH. Inset B in Fig. 3 shows the time course of absorption changes at 412 nm at various pHs. It is apparent that the appearance of the species X is well correlated with the complete oxidation of the anthracycline. Figure 4 shows that the initial rate of DXR oxidation measured at various total [SA] but at one pH (5.25) changes linearly with [HO-SA-COOH], additionally supporting the idea that the neutral form of SA is involved in the reaction.
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85.8% (n = 2; [LPO] = 24 nM, [H2O2] = 35 µM). This suggests that the SA metabolite had to redox cycle several times to accomplish this level of degradation. When ASA was used instead of SA, oxidation of DXR(DNR) was observed neither at neutral nor acidic pHs.
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). Figure 5, inset A, shows that the rate of DNR oxidation by LPO/H2O2/SA increases as pH decreases; however, in contrast to the system with MPO, the A480 versus time traces assume Z-shape, indicating that the process is autocatalytic. The time course of DNR oxidation recorded at constant pH of 5.0 but at various [SA] also assumes Z-shape (Fig. 5, inset B). Figure 5, main panel, shows that the maximal rate of the reaction (Vmax) increases linearly with [HO-SA-COOH]. Similar results were obtained for DXR (data not shown). Based on these observations, we infer that the stimulatory action by SA may also involve its secondary metabolite that is readily formed at acidic pH. This species is presumably formed by recombination of the primary metabolites of HOOC-SA-OH, the SA-derived phenoxyl radicals (HOOC-SA-O·), to corresponding biphenols (SA-BPH), which then are oxidized to biphenol quinone (SA-BPQ). It is known that oxidation of various phenolic compounds, e.g., phenol (hydroxybenzene), tyrosine, or p-cresol, leads to formation of the corresponding dimers, which are also substrates for peroxidases (Bayse et al., 1972; Sawahata and Neal, 1982; Marquez and Dunford, 1995; Monzani et al., 1997).
Measurements of the position of the enzyme's Soret band during turnover were conducted next. Addition of H2O2 (5 µM) to DXR, SA, and LPO (0.46 µM) in pH 5.0 buffer caused the peak at 412 nm (ferric LPO) to shift to 430 nm (LPO compound II).3 In this form, the enzyme lived for 45 s, after which it returned to native LPO. The corresponding spectral lines intersect at 421 nm, consistent with conversion of LPO-II to native LPO (Jenzer et al., 1986). During the LPO-II lifetime, the A480 decreased by 
A480 = 0.041, which corresponds to the loss of 3.6 µM DXR (Fig. 6A). The presence of LPO in the form of compound II during the reaction suggests that reduction of LPO-II by HOOC-SA-OH is the rate-limiting step. When the same experiment was repeated at pH 7.0, the decrease at 480 nm was very small (Fig. 6B) ([DXR] = 0.59 µM in 75 s), and the peak at 435 nm was observed for at least 20 min. This result confirms that reaction of ionized SA (HO-SA-COO-) with LPO-II at pH 7 is very slow. When SA was omitted, the amount of DXR degraded at pH 7.0 was nearly the same as when SA was present, further confirming that SA is inactive at this pH (data not shown).
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Oxidation of Salicylic Acid by MPO and LPO Systems. We observed that oxidation of SA/DNR(DXR) at acidic pH by either LPO/H2O2 or MPO/H2O2 generated species X, but only when the anthracycline was depleted. This suggested to us that X could be derived from SA and not from DXR or DNR. Therefore, we next studied the formation of this metabolite in the absence of the anthracyclines.
Reaction of SA with MPO/H2O2 in pH 5.1 buffer generated spectrum with max at 412 nm (Fig. 7), which is attributed to the species X. The absorbance at 412 nm, after reaching maximum starts to decrease, suggesting that X is unstable (Fig. 7, inset A). The formation of this metabolite is pH-dependent as the rate of its formation increases sharply upon changing pH from 6.4 to 5.0 (Fig. 7, inset A). No peak at 412 nm was formed at pH 7.0 and above, during prolonged observation.4 Thus, efficient metabolism of SA by MPO occurs only at acidic pH, with maximum efficiency at pH 5.0, the lowest pH used in our experiments. The initial rate of the formation of X changes linearly with [HOOC-SA-OH] (Fig. 7, inset B). Similar observations were made when LPO was used instead of MPO. We tentatively assign the product X to SA-BPQ and analog of 4,4'-biphenol quinone generated during enzymatic oxidation of phenol (hydroxybenzene) (Sawahata and Neal, 1982). We note that the Vi versus pH relationship determined here for SA is opposite to that found for other phenolics, for which decrease in pH was associated with decrease in Vi (Marquez and Dunford, 1995; Monzani et al., 1997).
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Effect of Anthracyclines on the Formation of Species X. Based on the observations that MPO(LPO)/H2O2/SA oxidizes DNR(DXR) that MPO(LPO)/H2O2 oxidizes SA and also that the SA metabolite (species X) appears only when DNR(DXR) is depleted, we asked how formation of X depends on anthracyclines. Therefore, we studied formation of this product as a function of [DNR]. It was expected that if X, and/or its precursors, reacts with DNR, the appearance of the 412-nm band should depend on [DNR]. Figure 8A shows A412 versus time traces observed at [DNR] of 0, 3.9, 7.2, and 14.8 µM. The figure shows that there is a distinctive [DNR]-dependent lag period, preceding the formation of X. Concomitantly measured changes in absorbance at 480 nm indicate that in contrast to the formation of X, oxidation of DNR starts immediately after the H2O2 addition (Fig. 8B). These results suggest that a precursor of X, or the X itself, may react with DNR. The latter possibility was studied next.
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EPR Study. EPR measurements were carried out to find whether oxidation of the anthracyclines by peroxidases in the presence of SA generates free radicals. When DNR(DXR) was incubated with MPO/H2O2 in the presence of SA in pH 5.1 buffer, the EPR signal shown in Fig. 10, trace A, was observed. The signal line width of 0.195 mT and g of 2.00479 are close to those reported previously for DNR(DXR) radicals in other peroxidizing systems (Reszka et al., 2004, 2005a). No signal was detected when SA was omitted (Fig. 10, trace B) or when the reaction was carried out at pH 7.0 (data not shown). The dependence of the signal on pH corroborates our results of spectrophotometric measurements. These results are consistent with the mechanism whereby the anthracycline hydroquinone moiety undergoes oxidation to the corresponding semiquinone by an SA-derived metabolite(s). We did not observe any EPR signals from control samples consisting of SA/peroxidase/H2O2 in acidic buffer. Although oxidation of SA by HRP/H2O2 or methemoglobin/H2O2 generates phenoxyl radicals (Shiga and Imaizumi, 1973, 1975), they cannot be detected using stationary EPR.
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). Because oxidation of anthracyclines leads to their inactivation (Cartoni et al., 2004; Reszka et al., 2005b), this reaction may be of clinical importance.
Our study shows that SA stimulates oxidation of DNR(DXR) by peroxidases but does this in a pH-dependent fashion. The stimulatory effect increases as the pH decreases from 7 to 5, which parallels the dependence on pH of the peroxidative metabolism of SA itself. These observations suggest that oxidation of the anthracyclines is mediated by an SA metabolite and that the protonated (neutral) form of SA (HOOC-SA-OH) is the preferred substrate for peroxidases. Oxidation of phenols yields the respective phenoxyl radicals (Shiga and Imaizumi, 1973, 1975; Marquez and Dunford, 1995; Monzani et al., 1997; Hermann et al., 1999b); accordingly, oxidation of SA should yield the corresponding phenoxyl radical, HOOC-SA-O·, as described by eq. 1, using MPO as a typical peroxidase. The resulting radicals may dimerize, forming a corresponding biphenol (SA-BPH) (eq. 2), or react with other substrates. We propose that in the presence of DXR or DNR, the SA-derived phenoxyl radicals react with the quinone-hydroquinone group (Q-QH2) of the anthracyclines, causing its oxidation to a semiquinone radical (Q-QH·) (eq. 3). During this reaction, the phenoxyl radical is reduced back to HOOC-SA-OH.
 | (1) |
 | (2) |
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HOOC-SA-O· HOOC-SA-OH to uphold the oxidation of the much higher concentrations of the drug. Thus, the cycling of the HOOC-SA-OH/HOOC-SA-O· couple sustains continuous oxidation of the drug. The scheme in Fig. 11 illustrates the proposed mechanism of this pro-oxidant action of SA. In contrast to SA, ASA appeared to be inactive. The most likely reason behind this is that in ASA, the phenolic group has been blocked by acetylation (Fig. 1); accordingly, the compound is not metabolized by peroxidases and does not generate phenoxyl radicals.
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The dependence of the reaction on pH is not unexpected since there is precedence with nitrite and acetaminophen (Reszka et al., 2001, 2004). What was unusual was the direction of these changes because they do not follow the trend of oxidation of phenols versus pH. It has been reported that the rate of oxidation of p-cresol decreases on changing the pH from neutral to acidic values, and this effect was correlated with protonation of an amino acid with pKa 5.8 near the enzyme active site, presumably histidine (Monzani et al., 1997). p-Cresol does not have a carboxylic group. Therefore, its ionization status is not affected by changes in pH. That the dependence on pH observed in our study is primarily due to protonation of the SA anion, and much less due to changes in the enzyme's reactivity, is demonstrated by the very good linear correlation between the rate of SA oxidation at various pHs and the actual content of the neutral form of the compound (HOOC-SA-OH). Similarly good correlation was found between the rate of DXR and DNR oxidation and the concentration of HOOC-SA-OH at various pHs. Thus, we conclude that oxidation of SA to phenoxyl radicals (eq. 1) involves mostly the neutral form of the compound. The observation that the SA-dependent oxidation of anthracyclines occurs at acid pHs is relevant to the situation in vivo since the extracellular pH of solid tumors, against which anthracyclines are frequently used, is acidic. pH as low as 6.1 occurs with some types of tumors (Gillies et al., 2002).
Oxidation of SA in acid solutions gives rise to a product X, which shows maximum absorption at 412 nm. Based on the known chemistry of phenoxyl radicals and using the simplest phenol (hydroxybenzene) as a reference, we tentatively identify this product as the respective biphenol quinone (SA-BPQ). This compound could be formed by enzymatic oxidation of a biphenol, which is the product of recombination of phenoxyl radicals. Although oxidation of phenol can produce 2,2'- and 4,4'-biphenol quinones, only the latter one shows the characteristic intense absorption at 398 nm (Sawahata and Neal, 1982). Therefore, by analogy to oxidation of phenol to 4,4'-biphenol quinone, the species X could be assigned to 5,5'-SA-BPQ, in which the >C = O functions are in para position. The structure of this compound and the proposed mechanism of its formation are shown in Fig. 11. The probable reason why this dimeric product is efficiently formed in acid solutions is the low pKa value of the -COOH group in SA phenoxyl radicals, 3 (Neta and Fessenden, 1974). Dimerization of these radicals is facile when their carboxyl group is protonated. In contrast, recombination of SA-derived phenoxyl radical anions (·O-SA-COO-) may be hindered due to repulsion of their negative charges. Thus, as the pH increases from 5 to 7, the proportion of neutral radicals become so low that any dimers formed are below the detection limit (but see footnote 4).
We found that the SA derived biphenol quinone can be reduced by DNR (Fig. 9A). The resulting biphenol was reoxidized with H2O2 and reduced again by a second dose of the drug. The reduction of 5,5'-SA-BPQ was concomitant with DNR oxidation. Thus, in the peroxidase/H2O2/SA system, oxidation of anthracyclines can be carried out by both the SA-derived phenoxyl radical and the SA-derived biphenol quinone (Fig. 11, steps 1 and 2, respectively). The former reaction seems to play a more important role since presence of anthracyclines inhibits formation of biphenols (Fig. 11, step 1).
We have previously shown that oxidation of anthracyclines can be inhibited by ascorbate or reduced glutathione (Reszka et al., 2004). Therefore, the efficacy of this reaction in vivo will certainly depend on the presence of endogenous antioxidants and may become evident under conditions of oxidative stress, when these antioxidants are depleted. We emphasize that redox cycling of anthracyclines might promote development of oxidative stress.
Our results show that oxidation of anthracyclines leads to their irreversible bleaching, suggesting a significant modification of their chromophores. In agreement with this, recent studies revealed that oxidation of anthracyclines leads to their degradation to low-molecular weight products, 3-methoxyphthalic acid and 3-methoxysalicylic acid (Bomgaars et al., 1997; Cartoni et al., 2004; Reszka et al., 2005b). This degradation could be mediated by the drug-derived semiquinone radicals (Q-QH·), which decay, presumably, by disproportion to the parent drug and the electron-deficient diquinone, Q-Q (2 Q-QH· 
Q-QH2 + Q-Q). It seems likely that subsequent reactions of this diquinone species could give rise to the ultimate colorless stable products. We emphasize that semiquinone radicals generated by oxidation of anthracyclines (Q-QH·) differ from the better known radicals formed by metabolic reduction (·QH-QH2). The latter can reduce O2 to superoxide, which restores the drug to its original form (Kalyanaraman et al., 1980). In contrast, semiquinones formed by oxidation undergo structural modifications.
It has been reported that products of anthracycline oxidation are virtually nontoxic to human leukemia HL-60 cells, human prostate cancer PC3 cells, and rat heart cardiomyocytes (H9c2) (Reszka et al., 2005b). These results agree with lower toxicity of 3-methoxyphthalic acid in H9c2 cells reported by another group (Cartoni et al., 2004) and with the lower toxicity of photochemically degraded DXR in P388 murine leukemia cell line (Bomgaars et al., 1997). Altogether, these observations rise the possibility that oxidation of anthracyclines in vivo may suppress their therapeutic activity. Because cancer patients undergoing anthracycline chemotherapy may be administered salicylates to control pain and inflammation, possible complications and decreased anticancer activity of the drugs should be considered. One possible beneficial effect of the drugs' degradation could be reduced cardiotoxicity as suggested by results of in vitro studies on toxicity of anthracycline degradation products in mouse cardiomyocytes (Cartoni et al., 2004; Reszka et al., 2005b). Together, these results suggest that it should be possible to modulate inactivation of the anthracyclines in vivo by pharmacological interventions, using stimulants, or inhibitors of peroxidative processes.
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Acknowledgements |
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Footnotes |
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Results of preliminary studies were presented at: Second International Conference on Prostate Cancer Research, Iowa City, IA, October 12-15, 2002. Abstract: Reszka KJ, Britigan LH, McCormick ML, Britigan BE, and Spitz DR (2002) Do pain killers counteract the anticancer action of anthracyclines? Book of Conference Abstracts, p 50.
Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: ASA, acetylsalicylic acid (aspirin); COX, cyclooxygenase; SA, salicylic acid; NSAID, nonsteroidal anti-inflammatory drug; MPO, myeloperoxidase; DXR, doxorubicin (Adriamycin); DNR, daunorubicin; LPO, lactoperoxidase; HOOC-SA-OH, -OOC-SA-OH, HOOC-SA-O·, neutral and anionic forms of salicylic acid and the respective phenoxyl radical; SA-BPH, biphenol form of SA; SA-BPQ, biphenol quinone form of SA; Q-QH2, Q-QH·, and Q-Q, the quinone-hydroquinone moiety of anthracyclines, and the corresponding semiquinone and di-quinone forms; EPR, electron paramagnetic resonance.
1 Current address: Student at College of Liberal Arts, University of Iowa, Iowa City, Iowa. 
2 This refers only to low, pharmacological concentrations of SA because oxidation of anthracyclines was readily accomplished at pH 7.0 when using high, cytotoxic concentrations of SA (10 mM).
3 The actually measured max was 414 nm for native LPO and 435 nm after H2O2 addition (assigned to LPO-II). These apparent red shifts in peaks positions must be due to the fact that these peaks are on the uphill slope of the DXR absorption spectrum. When DXR was omitted the corresponding spectra, measured before and after H2O2, addition showed max at 412 and 430 nm, respectively, as expected for ferric and compound II forms of LPO (Jenzer et al., 1986).
4 The absence of this metabolite at pH 7 refers to low, pharmacologically relevant concentrations of SA. When the concentration of SA was increased to 10 mM, formation of this metabolite was apparent even at pH 7 (data not shown). Note that at pH 7.0 and 10 mM SA, the concentration of the neutral form of SA is nearly the same as from 0.1 mM SA at pH 5.0.
Address correspondence to: Dr. Krzysztof J. Reszka, Department of Internal Medicine, University of Cincinnati, 231 Albert Sabin Way, ML 0557, Cincinnati, OH 45267-0557. E-mail: reszkakj{at}ucmail.uc.edu
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