| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Biochemistry and Biophysics |
Departments of Environmental and Occupational Health [V. E. K., A. I. K., Y. Y. T., T. M.] and Pharmacology [V. E. K., A. I. K., T. M., J. C. Y.], University of Pittsburgh, Pittsburgh, Pennsylvania 15238; Health Effects Laboratory Division, Pathology and Physiology Research Branch, National Institute for Occupational Safety and Health, Morgantown, West Virginia 26505 [A. A. S.]; and A.V. Palladin Institute of Biochemistry, Ukrainian National Academy of Sciences, Kiev, 252030, Ukraine [A. I. K.]
 |
ABSTRACT |
|---|
 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
The DNA topo3
II poison etoposide (VP-16), a widely used anticancer agent, contains a hindered phenolic ring, a critical structural prerequisite for its antitumor activity (6
, 7)
. As a hindered phenol, etoposide can act as an effective donor of electrons for scavenging reactive (peroxyl) radicals, i.e., it can act as an antioxidant as indicated below.
 |
The protective antioxidant effects of etoposide against lipid peroxidation have been demonstrated in model biochemical systems (8 , 9) . In contrast, etoposide-induced formation of lipid peroxyl radicals and increased lipid peroxidation have been reported in different clones derived from Chinese hamster ovary cells (10) .
Reactivity of the phenoxyl radical formed from etoposide (etoposide-O·) in the course of Reaction 1
toward different biomolecules (Reaction 2) is essential for determining whether net antioxidant or pro-oxidant effects of etoposide will be realized in cells.
 |
Thus, etoposide may act as either a pro- or antioxidant toward different intracellular constituents. In particular, MPO-catalyzed one-electron oxidation of etoposide and the secondary reactions of the resultant phenoxyl radical cause oxidative stress (thiol oxidation) in cell-free model systems, cell homogenates (9) , and myeloblastic leukemia cells (11) .
It has been suggested that oxidative activation of etoposide by cytochrome P450 monooxygenases, MPO, prostaglandin synthetase, and tyrosinase may contribute to its cytotoxicity (12, 13, 14, 15) . Whereas etoposide is highly efficacious as an antitumor drug, its use is also associated with an increased risk of secondary AML (16) . This has prompted its withdrawal from some treatment regimens, potentially compromising efficacy against the original tumor (17) . Based on the results of our experiments in cell-free systems and cell homogenates, we have hypothesized that etoposide genotoxicity and carcinogenicity are due to MPO-catalyzed production of etoposide phenoxyl radicals (etoposide-O·) in myeloid progenitors and their subsequent pro-oxidant effects (9) .
In the present work, we present direct EPR evidence for the MPO-catalyzed generation of etoposide-O· radicals in intact HL-60 cells, their interactions with intracellular thiols, and the resulting modulation of etoposide-induced covalent complex formation between DNA and topo II. In addition, using a sensitive procedure for assay of oxidative stress in different classes of membrane phospholipids, we demonstrate that etoposide-O· phenoxyl radicals are not involved in pro-oxidant reactions toward lipids but rather act as antioxidants against H2O2-induced phospholipid peroxidation.
|
MATERIALS AND METHODS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Cell Culture.
Human promyelocytic HL-60 cells (from the American Type Culture Collection) were grown in Iscove’s medium supplemented with 15% FBS in 95% humidity atmosphere under 5% CO2 in air at 37°C. Cells from passages 25–40 were used for the experiments. The density of cells at collection time was 0.5 x 106 cells/ml. Etoposide was dissolved in DMSO and added to cells (DMSO concentration did not exceed 1%). HL-60 cells (1.5 x 105 cell/ml) were incubated for 48 h with SA (500 µM) dissolved in Iscove’s medium. Cell viability was determined microscopically by trypan blue dye exclusion.
MPO Activity.
HL-60 cells were harvested by centrifugation at 1000 rpm for 5 min. Pellets were washed twice with buffer A containing 25 mM HEPES, 10 mM glucose, 115 mM NaCl, 5 mM KCl, 1 mM MgCl2, and 5 mM NaH2PO4 (pH 7.4). The homogenate was prepared by freezing at -77°C and thawing the cells. A spectrophotometric assay (Shimadzu UV 160U spectrophotometer; Kyoto, Japan) of MPO activity was used in which guaiacol oxidation was monitored by changes of absorbance at 470 nm (
= 26.6 mM-1/cm-1; Ref. 1
). Cell homogenate (0.5 x 106 cells) was added to 100 mM disodium phosphate buffer containing 0.1% Triton X-100, 0.1 mM phenylmethylsulfonyl fluoride, 13 mM guaiacol, 0.02% cetylmethylammonium bromide, and 3.75 mM 3-AT (pH 7.0). The reaction was started by the addition of 670 µM H2O2. Activity of MPO was calculated in nmol tetraguaiacol formed/min/106 cells. The data were acquired using Shimadzu PC 160 software version 1.2.
EPR Spectroscopy of Etoposide Phenoxyl Radicals.
Measurements of etoposide phenoxyl radicals were performed on a JEOL-RE1X EPR spectrometer (Tokyo, Japan) at 25°C as described previously (9)
. Samples (50 µl) contained viable HL-60 cells (8 x 104), etoposide (50–400 µM), 3-AT (5 mM), and H2O2 (5–200 µM). Measurements were done in gas-permeable Teflon tubing (internal diameter, 0.8 mm; thickness, 0.013 mm) from Alpha Wire Corp. (Elizabeth, NJ). The tubing filled with mixed sample was folded twice and placed into a 3.0-mm EPR quartz tube. The EPR conditions were as follows: 335.7 mT center field; 2 mT sweep width; 0.04 mT field modulation; 10 mW microwave power; 0.1 s time constant; 4000 receiver gain; and 1 min time scan. The kinetics of etoposide phenoxyl radical formation was measured by repeated scanning of its EPR signal.
Assay of GSH and Protein SH Groups in HL-60 Cells.
GSH and total protein SH concentration in HL-60 cells was determined using ThioGlo-1, a maleimide reagent that produces a highly fluorescent adduct on its reaction with SH groups (18)
. GSH content was estimated by an immediate fluorescence response registered on the addition of ThioGlo-1 to the cell homogenate. Protein SHs were determined as an additional increase in fluorescence response after the addition of SDS (4.0 mM) to the same cell homogenate. To demonstrate the initial interaction of ThioGlo-1 mainly with GSH, HL-60 cell homogenates were treated with glutathione peroxidase to specifically deplete GSH, followed by analysis of fluorescence responses before and after the addition of SDS (i.e., GSH and other low molecular weight thiols or protein thiols, respectively). We found that glutathione peroxidase treatment almost completely eliminated that part of the fluorescence response to ThioGlo-1 obtained in the absence of detergent. For non-detergent-treated cell homogenates, glutathione peroxidase treatment decreased the measured SH level from 4.98 ± 1.00 to 0.48 ± 0.14 nmol/mg protein. At the same time, no effect of glutathione peroxidase treatment on the fluorescence response to ThioGlo-1 was revealed in the presence of detergent (19.94 ± 4.53 versus 17.92 ± 3.32 nmol SH/mg protein). In HL-60 cells, therefore, the initial fluorescence response elicited by ThioGlo-1 in the absence of detergent was due almost entirely to GSH, whereas the response evoked by the addition of detergent was due mainly to protein SH groups.
For standard assays, homogenate (8 x 104 cells), ThioGlo-1 (10 µM in DMSO), and disodium phosphate buffer [100 mM (pH 7.4)] in a final volume of 1 ml were used. A standard curve was established by the addition of 0.04–4.0 µM GSH to 100 mM phosphate buffer (pH 7.4) containing 10 µM ThioGlo-1 (DMSO solution). A model RF-5301PC spectrofluorophotometer (Shimadzu) was used for the assay of fluorescence using excitation at 388 nm and emission at 500 nm. The data obtained were exported and treated using RF-5301PC Personal Fluorescence Software (Shimadzu). To titrate endogenous GSH, intact HL-60 cells (8 x 104) were incubated with different concentrations of ThioGlo-1 (2–10 µM, 10 min) in buffer A (see above). HL-60 cells were then washed twice with buffer A to remove excess non-reacted ThioGlo-1.
Imaging of GSH in HL-60 Cells Using ThioGlo-1.
HL-60 cells (106 cells/75-cm2 flask) were incubated with 200 µM H2O2, 400 µM etoposide, and 5 mM 3-AT for 15 min. After incubation, the cells were pelleted, washed twice, and resuspended in PBS. Cells were labeled with 40 µM ThioGlo-1. Cell slides were prepared by spinning cell suspensions at 800 rpm for 5 min using Cytospin 3 (Shandon; Life Sciences International Ltd., Cheshire, United Kingdom). Cells on slides were fixed with 2% paraformaldehyde for 30 min. The glass coverslips (Fisher Scientific) were mounted over slides using Prolong Mounting Medium (Molecular Probes, Eugene, OR). Cells were visualized using an Olympus AX70 photomicroscope (Tokyo, Japan) with a xenon epifluorescence attachment (filter cubes 
= 360–390 nm) and barrier filter ( = 515 nm) to record fluorescence images. Images were captured with a Sony 3CCD color video camera, DXC9000 (Kyoto, Japan) and saved as tif. files using Simple32 software (Compix Inc., Cranberry, PA). Magnification 1000x.
Determination of Phospholipid Peroxidation in HL-60 Cells.
PnA was incorporated into HL-60 cells by the addition of complex PnA-human serum albumin to yield a final concentration of 2 µg PnA/106 cells to serum-free RPMI 1640 without phenol red as described previously (19)
. PnA-labeled HL-60 cells were treated with 100 µM H2O2 in the presence or absence of 50 µM etoposide for 2 h at 37°C in buffer A (pH 7.4) containing 5 mM 3-AT. H2O2 (25 µM) was added every 30 min. Etoposide was added to cell suspensions 35 min before the addition of H2O2. At the end of the incubation period, total lipids were extracted using Folch’s procedure. The lipid extract was dried under N2, dissolved in 0.2 ml of 2-propanol:hexane:water (4:3:0.16, by volume), and separated by normal phase high-performance liquid chromatography using a 5-µm microsorb MV column (4.6 x 250 mm; Rainin Instrument Co. Inc.) and an ammonium acetate gradient as described previously (19)
. The separations were performed using a Shimadzu LC-600 high-performance liquid chromatography system equipped with an in-line configuration of RF-551 fluorescence detector. Fluorescence of PnA was measured at 420 nm emission after excitation at 324 nm. Data were processed and stored in digital form with Shimadzu EZChrom software. Lipid phosphorus was determined using a micromethod (20)
.
topo II Covalent Complex Formation.
Mid-log HL-60 cells (2.5–3.0 x 105 cells/ml) were labeled overnight with 0.5 µCi/ml [methyl-3H]thymidine (0.5 Ci/mmol) and 0.1 µCi/ml [U-14C]leucine (318 mCi/mmol) in Iscove’s media containing 15% FBS. Cells were then pelleted by centrifugation, resuspended in fresh DMEM/7.5% calf serum, and incubated for 1 h at 37°C. Cells were pelleted and resuspended in buffer A at 37°C at a final density of 1.0 x 106 cells/ml. Cells were then incubated with etoposide (50 µM) for 30 min in the presence or absence of ThioGlo-1 (0.1–10 µM). Cells (1.0 x 106) were then pelleted (10 min at 1700 x g) after being plunged into 10 ml of ice-cold 10 mM PBS, washed with ice-cold buffer A, pelleted, and lysed. Cellular DNA was sheared, and protein-DNA complexes were precipitated with SDS and KCl as described previously (21)
. topo II-DNA complexes were quantified by scintillation counting (Beckman LS 6500 multipurpose scintillation counter), and [3H]DNA was normalized to cell number using coprecipitated 14C-labeled protein as an internal control.
Statistical Analysis.
The results are presented as the mean ± SE values from at least three experiments, and statistical analysis was performed by either paired/unpaired Student’s t test or one-way ANOVA. The statistical significance of differences was set at P < 0.05.
|
RESULTS |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Effects of Etoposide on Phospholipid Peroxidation.
Using our sensitive and specific procedure to quantitate peroxidation of different classes of membrane phospholipids in intact cells (19)
, we determined whether etoposide, as a hindered phenolic compound, could act as a radical scavenger. In addition, this technique allowed us to determine whether etoposide-O· radicals were reactive enough to cause direct oxidation of intracellular phospholipids. We prepared HL-60 cells in which the major classes of membrane phospholipids were metabolically labeled with the oxidation-sensitive fluorescent fatty acid PnA (19)
, and then we exposed cells to etoposide, H2O2, or a combination of etoposide plus H2O2.
Our data on oxidation of two major phospholipids in HL-60 cells, PC and PE, are shown in Table 1
. We found that etoposide alone caused no oxidation of either PC or PE. Consistent with our previous observations (22)
, H2O2 produced significant oxidative damage to both PC and PE. When intact cells were exposed to H2O2 in the presence of etoposide, there was significant protection against PC and PE oxidation, suggesting etoposide-mediated antioxidant effects. We found that neither etoposide nor H2O2 alone or in combination caused any significant cytotoxicity (trypan blue-positive cells were <5%) during the same 2-h incubation time used in phospholipid peroxidation experiments. Thus, etoposide-induced loss of cell viability is not responsible for the antioxidant effects of etoposide.
|
. Analysis of other minor phospholipid classes (phosphatidylserine, phosphatidylinositol, and sphingomyelin) demonstrated that none were oxidized by etoposide in SA-pretreated or non-SA-pretreated HL-60 cells. On the contrary, etoposide protected these phospholipids against H2O2-induced oxidation in both naive and SA-pretreated cells (data not shown). Together, these results indicate that etoposide exerted a pronounced antioxidant effect against H2O2-induced phospholipid peroxidation in HL-60 cells. In addition, results show that MPO-catalyzed etoposide-O· radicals do not oxidize membrane phospholipids in HL-60 cells.
Detection of Etoposide-O· Radicals in Intact HL-60 Cells.
Our previous work has established that in the presence of H2O2, MPO in HL-60 cell homogenates catalyzed one-electron oxidation of etoposide, yielding etoposide-O· radicals that were directly detectable by EPR after depletion of thiols (9)
. We sought to determine whether intact cells were able to metabolize etoposide via the same pathway and whether intracellular thiols were involved in further reactions involving these radical species. We used EPR spectroscopy to monitor the production of etoposide-O· radicals in HL-60 cells. We found that typical EPR spectra of etoposide-O· radicals could be directly recorded from the cells in the presence of etoposide and H2O2 (Fig. 1A)
. Importantly, the signal could be recorded only after a lag period that lasted for 16 min under the conditions used. The presence of the MPO cosubstrate, H2O2, was essential because no signals were detectable in the absence of the catalase inhibitor, 3-AT.
|
. In the absence of ThioGlo-1, there was a lag period (Fig. 2B)
before typical EPR spectra of etoposide-O· radicals could be observed. Titration out of HL-60 cell thiols with different concentrations of ThioGlo-1 showed that at 1 µM, the cells retained 
30% of endogenous GSH (Fig. 2A)
. Increasing ThioGlo-1 concentrations up to 5–10 µM essentially depleted all intracellular GSH. Importantly, at concentrations 
10 µM, ThioGlo-1 did not affect the viability of HL-60 cells within 30 min of incubation (cell viability was 93.3 ± 0.8% and 86.1 ± 3.6% in control samples and after incubation with 10 µM ThioGlo-1, respectively). As ThioGlo-1 concentrations were increased from 0.1–10 µM, the lag period before observation of EPR signals of etoposide-O· radicals was shortened (Fig. 2B)
, corresponding to the titration out of thiol reductant (Fig. 2A)
. In fact, at ThioGlo-1 concentrations of >1 µM, the signal was immediately detectable (Fig. 2B)
. Both the magnitude (Fig. 2B)
and the rate of etoposide-O· formation (Fig. 3)
were proportional to the concentration of added ThioGlo-1. Repeated scanning of a portion of the etoposide-O· EPR spectrum (Fig. 1, A and B)
over time demonstrated that the steady-state concentrations of the radicals were sufficiently high to readily allow its monitoring over 16 min (Fig. 2B)
. The magnitude of the signal was dependent on the concentrations of both etoposide and H2O2 (Fig. 4)
, suggesting the involvement of MPO in etoposide-O· radical production.
|
|
|
. As expected, etoposide phenoxyl radical formation increased as ThioGlo-1 concentration increased (Figs. 2B
and 5
). In SA-treated MPO-depleted cells, etoposide-O· formation was dramatically decreased even in the presence of ThioGlo-1 concentrations as high as 10 µM (Fig. 5)
. In separate experiments, we found that short-term (2–6-h) incubation of HL-60 cells with 500 µM SA did not affect either MPO activity or etoposide-O· radical generation (data not shown).
|
. ThioGlo-1 (1 µM) significantly increased etoposide-induced topo II-DNA covalent complexes (>25%). This result is consistent with previous reports that partial depletion of GSH resulted in increased etoposide-induced DNA damage and cytotoxicity (28)
. At higher concentrations, ThioGlo-1 progressively inhibited etoposide activity, with essentially complete inhibition observed in the range of 5–10 µM ThioGlo-1.
|
, the ThioGlo-1 (1 µM)-mediated enhancement of etoposide-induced topo II-DNA covalent complexes was prevented in cells pretreated for 48 h with 500 µM SA. Similarly, control etoposide activity was reduced by 22% in SA (500 µM)-pretreated HL-60 cells (data not shown).
|
We speculate that when concentrations of endogenous GSH are low (either intrinsically or as a result of depletion), MPO-catalyzed etoposide-O· formation may allow for direct oxidation of cysteines on topo II that are essential for its function (29
, 30)
. Consistent with this idea, we found that the protein SH groups in HL-60 cells pretreated with 10 µM ThioGlo-1 (a concentration that titrated out all intracellular GSH; Fig. 8A
) were significantly decreased during a 15-min incubation with etoposide + H2O2 (Fig. 8B)
. This etoposide/H2O2-induced oxidation of protein SH groups was not detectable in cells with substantially higher levels of GSH [i.e., cells not pretreated with ThioGlo-1 or cells pretreated with lower concentrations of ThioGlo-1 (0.1–1.0 µM; Fig. 8A
)].
|
|
DISCUSSION |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
The cytotoxicity of different antitumor drugs toward neoplastic cells often leads to apoptosis and phagocytosis without induction of an inflammatory response. In contrast, necrotic cells lyse and release their contents into the extracellular space, thus inducing inflammation. Recent results indicate that oxidative stress and antioxidants can modulate chemotherapy-induced cell death pathways (1 , 2) . Antioxidants may protect tumor cells from apoptosis and phagocytosis, whereas pro-oxidants may induce apoptosis, thus affecting the overall effectiveness of antitumor therapy. A significant decline in antioxidant status and elevation of biomarkers of oxidative stress are typical of tissues and plasma of patients in the course of commonly used chemotherapy/radiotherapy protocols. In particular, this relates to treatment regimens that include etoposide (3 , 5) . Because the toxicity of etoposide and its homologues to tumor cells is mainly dependent on its ability to induce apoptosis (32, 33, 34) , its anti-/pro-oxidant effects may modulate apoptotic pathways and hence modulate the overall antitumor activity of the drug.
Over the past 10 years, numerous groups have reported that the same treatment schedules associated with the impressive efficacy of etoposide are also associated with an increased risk of secondary AML (35, 36, 37) . This has prompted the withdrawal of etoposide from some treatment regimens, potentially compromising efficacy against the original tumor (17) . Whereas the causative link between treatment of cancer with etoposide and the development of secondary AML in children and adults has been firmly established, the biochemical mechanisms explaining the extremely high susceptibility of myeloid stem cells to the leukemogenic effects of etoposide have not been elucidated. Recently, we hypothesized that MPO-catalyzed activation of etoposide to a reactive intermediate, etoposide-O·, may trigger oxidative stress and cause cytotoxic and genotoxic effects responsible for leukemogenesis (9) . As a direct consequence of this mechanism, we further proposed nutritional antioxidant strategies for preventing etoposide-induced leukemias (9) .
Thus, both the clinical efficacy and the leukemogenic action of etoposide may be dependent, at least in part, on the ability of etoposide to modulate the anti-/pro-oxidant status of cells. This prompted us to conduct the present study to investigate both the radical scavenging antioxidant effects of etoposide and its potential pro-oxidant role(s) in MPO-rich HL-60 cells.
Our results demonstrated for the first time that etoposide can act as both an antioxidant and a pro-oxidant toward different intracellular constituents. Using our sensitive and specific assay of phospholipid peroxidation in intact cells (19) , we established that etoposide does not cause peroxidation of any of major phospholipids (PC, PE, PI, and PS) in HL-60 cells. In fact, it acts as a potent antioxidant against H2O2-induced phospholipid peroxidation. This antioxidant effect of etoposide was completely independent of MPO activity in HL-60 cells. In a recent report, etoposide-induced formation of lipid peroxyl radicals and accumulation of lipid peroxidation products were found in Chinese hamster ovary cells with compromised genomic integrity (10) . However, the authors failed to determine whether these pro-oxidant effects of etoposide in lipids were typical of live cells, and they did not establish that the elevated levels of free radical damage to lipids were derived from the fraction of damaged cells dying by apoptotic or necrotic processes. In contrast, our results provide unequivocal direct evidence for the lack of pro-oxidant effects toward lipids and the potent antioxidant effect of etoposide in membrane phospholipids of intact HL-60 cells. Thus, the reactivity of etoposide-O· is not sufficient to directly attack intracellular phospholipids and cause their oxidative damage.
Our studies on thiol oxidation, however, established that etoposide can directly oxidize GSH and protein SH groups (after prior depletion of GSH by pretreatment with ThioGlo-1) in HL-60 cells. This oxidizing activity can be attributed entirely to MPO-catalyzed generation of etoposide-O· radicals and their subsequent interactions with thiols. These conclusions are supported by several results observed in the present study. First, our EPR measurements directly detected formation of etoposide-O· radicals in HL-60 cells. The EPR signals were observable in cells incubated in the presence of etoposide and H2O2 only after a lag period, the duration of which was strictly dependent on the intracellular GSH levels. When intracellular GSH was titrated out with ThioGlo-1, a GSH-specific maleimide reagent, the lag period was no longer observed. This suggests that reduction of etoposide-O· by GSH was occurring during the lag period. Second, manipulations of MPO activity in HL-60 cells using a heme synthesis inhibitor, SA, demonstrated that the enzyme-catalyzed reaction was absolutely required for both the production of etoposide-O· radicals and their interactions with GSH. Finally, etoposide-O· radicals were able to attack protein SH groups only after essentially complete depletion of intracellular GSH by ThioGlo-1. Thus, GSH acts as a potent protector of protein SHs against etoposide-O· radical oxidation. Our demonstration of etoposide-O· radical-induced oxidation of thiols explains previous findings on GSH depletion in cells treated with etoposide (11 , 28 , 38) .
Throughout these experiments, we used several reagents and inhibitors such as 3-AT, SA, and ThioGlo-1 to readily observe etoposide phenoxyl radicals in HL-60 cells. However, in cell-based assays, neither exogenous H2O2 nor 3-AT was required to observe MPO-dependent activity of etoposide where SA pretreatment alone reduced etoposide activity by 22%. Hence, we have demonstrated that a portion of etoposide activity is correlated with both MPO activity and the production of etoposide phenoxyl radicals. Overall, our results indicate that formation of etoposide phenoxyl radicals occurs in MPO-containing cells and that there are potentially important biological consequences related to etoposide activation.
We have previously put forth the idea (9) that etoposide phenoxyl radicals can result in oxidative stress, causing both genotoxicity and cytotoxicity. Because etoposide phenoxyl radicals can be produced in cells that contain oxidizing enzymes capable of one-electron transfer reactions such as peroxidases and tyrosinase, these metabolites may contribute to (a) the antitumor activity of etoposide in malignant cells capable of producing these phenoxy radicals and (b) the carcinogenic action of this agent in myeloid stem cells that contain MPO.
It is tempting to speculate that oxidation of essential cysteines on topo II by etoposide-O· radicals is responsible for both the enhancement and inhibition of etoposide activity, depending on the extent of oxidation. Indeed, we observed a significant oxidation of protein SH groups in HL-60 cells in which GSH was completely depleted with ThioGlo-1. Interestingly, topo II expression is sensitive to thiol-mediated redox regulation at the posttranscriptional level by modulation of the interaction of the 3'-untranslated region of topo II mRNA with redox-sensitive protein complexes (39) . However, this important regulatory mechanism is likely to differ from the mechanism(s) involving modulation of etoposide activity after a decrease in GSH levels induced by treatment with ThioGlo-1. Additional studies are necessary to establish whether this oxidative modification of protein cysteines does in fact include those on topo II and whether cysteines are oxidized to any significant extent in the presence of physiologically relevant levels of GSH. It is obvious, however, that modulation of the intracellular redox status by etoposide metabolism can affect drug-induced activity at the level of topo II because moderately decreased levels of GSH correlated with potentiation of etoposide-induced topo II-DNA complexes only when MPO-catalyzed metabolism to etoposide-O· radicals was sufficiently high.
In conclusion, etoposide can act as both an antioxidant toward membrane phospholipids and a pro-oxidant toward intracellular thiols in cells. This redox activity of etoposide may contribute to its antitumor effectiveness and to its genotoxic and carcinogenic potential. Understanding the intracellular redox biochemistry of etoposide may be instrumental for the development of nutritional antioxidant strategies for safer and more efficacious clinical use of etoposide.
|
FOOTNOTES |
|---|
1 Supported by NIH Grants CA 90787-01, CA74972, and CA77468; American Institute for Cancer Research Grant 97-B128; and the Leukemia Research Foundation and the International Neurological Science Fellowship Program (F05 NS 10669) administered by NIH/National Institute of Neurological Disorders and Stroke in collaboration with WHO, Unit of Neuroscience, Division of Mental Health and Prevention of Substance Abuse (Y. Y. T.). 
2 To whom requests for reprints should be addressed, at (V. E. K.) Department of Environmental and Occupational Health, University of Pittsburgh, 3343 Forbes Avenue, Pittsburgh, PA 15260. Phone: (412) 383-2136; Fax: (412) 383-2123; E-mail: kagan@pitt.edu, or: (J. C. Y.) Department of Pharmacology, University of Pittsburgh, W1355 Biomedical Science Tower, Pittsburgh, PA 15261. Phone: (412) 648-8136; Fax: (412) 648-1945; E-mail: yalowich{at}pitt.edu
3 The abbreviations used are: topo, topoisomerase; EPR, electron paramagnetic resonance; GSH, reduced glutathione; MPO, myeloperoxidase; AML, acute myeloid leukemia; SA, succinyl acetone (4,6-dioxoheptanoic acid); 3-AT, 3-amino-1,2,4-triazole; FBS, fetal bovine serum; SH, sulfhydryl; PnA, cis-parinaric acid; PC, phosphatidylcholine; PE, phosphatidylethanolamine.
Received 3/ 9/01. Accepted 8/28/01.
|
REFERENCES |
|---|
|
TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
-glutamylcysteine synthetase levels during etoposide-induced oxidative stress. Ann. Oncol., 10: 1361-1367, 1999.
mRNA is regulated by the 3'-untranslated region. Possible role of redox-sensitive protein binding in mRNA accumulation. J. Biol. Chem., 275: 38384-38392, 2000.This article has been cited by other articles:
|
|
 |
|
  S. Jitkaew, A. Trebinska, E. Grzybowska, G. Carlsson, A. Nordstrom, J. Lehtio, A.-S. Frojmark, N. Dahl, and B. Fadeel N{alpha}-Tosyl-L-phenylalanine Chloromethyl Ketone Induces Caspase-dependent Apoptosis in Transformed Human B Cell Lines with Transcriptional Down-regulation of Anti-apoptotic HS1-associated Protein X-1 J. Biol. Chem., October 9, 2009; 284(41): 27827 - 27837. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 F. Rombaldi, C. Cassini, M. Salvador, J. Saffi, and B. Erdtmann Occupational risk assessment of genotoxicity and oxidative stress in workers handling anti-neoplastic drugs during a working week Mutagenesis, March 1, 2009; 24(2): 143 - 148. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Watabe, K. Aoyama, and T. Nakaki A Dominant Role of GTRAP3-18 in Neuronal Glutathione Synthesis J. Neurosci., September 17, 2008; 28(38): 9404 - 9413. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. Y. Tyurina, V. Kini, V. A. Tyurin, I. I. Vlasova, J. Jiang, A. A. Kapralov, N. A. Belikova, J. C. Yalowich, I. V. Kurnikov, and V. E. Kagan Mechanisms of Cardiolipin Oxidation by Cytochrome c: Relevance to Pro- and Antiapoptotic Functions of Etoposide Mol. Pharmacol., August 1, 2006; 70(2): 706 - 717. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
B. B. Hasinoff, X. Wu, O. V. Krokhin, W. Ens, K. G. Standing, J. L. Nitiss, T. Sivaram, A. Giorgianni, S. Yang, Y. Jiang, et al. Biochemical and Proteomics Approaches to Characterize Topoisomerase II{alpha} Cysteines and DNA as Targets Responsible for Cisplatin-Induced Inhibition of Topoisomerase II{alpha} Mol. Pharmacol., March 1, 2005; 67(3): 937 - 947. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Darst, M. Al-Hassani, T. Li, Q. Yi, J. M. Travers, D. A. Lewis, and J. B. Travers Augmentation of Chemotherapy-Induced Cytokine Production by Expression of the Platelet-Activating Factor Receptor in a Human Epithelial Carcinoma Cell Line J. Immunol., May 15, 2004; 172(10): 6330 - 6335. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Y. Tyurina, F. B. Serinkan, V. A. Tyurin, V. Kini, J. C. Yalowich, A. J. Schroit, B. Fadeel, and V. E. Kagan Lipid Antioxidant, Etoposide, Inhibits Phosphatidylserine Externalization and Macrophage Clearance of Apoptotic Cells by Preventing Phosphatidylserine Oxidation J. Biol. Chem., February 13, 2004; 279(7): 6056 - 6064. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. Corazzari, P. E. Lovat, S. Oliverio, A. D. J. Pearson, M. Piacentini, and C. P. F. Redfern Growth and DNA Damage-Inducible Transcription Factor 153 Mediates Apoptosis in Response to Fenretinide but Not Synergy between Fenretinide and Chemotherapeutic Drugs in Neuroblastoma Mol. Pharmacol., December 1, 2003; 64(6): 1370 - 1378. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Shimoda, W. E. Achanzar, W. Qu, T. Nagamine, H. Takagi, M. Mori, and M. P. Waalkes Metallothionein Is a Potential Negative Regulator of Apoptosis Toxicol. Sci., June 1, 2003; 73(2): 294 - 300. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Li, M. D. Southall, Q. Yi, Y. Pei, D. Lewis, M. Al-Hassani, D. Spandau, and J. B. Travers The Epidermal Platelet-activating Factor Receptor Augments Chemotherapy-induced Apoptosis in Human Carcinoma Cell Lines J. Biol. Chem., May 2, 2003; 278(19): 16614 - 16621. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Kumar, N. Mishra, D. Raina, S. Saxena, and D. Kufe Abrogation of the Cell Death Response to Oxidative Stress by the c-Abl Tyrosine Kinase Inhibitor STI571 Mol. Pharmacol., February 1, 2003; 63(2): 276 - 282. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Cancer Research | Clinical Cancer Research |
| Cancer Epidemiology Biomarkers & Prevention | Molecular Cancer Therapeutics |
| Molecular Cancer Research | Cancer Prevention Research |
| Cancer Prevention Journals Portal | Cancer Reviews Online |
| Annual Meeting Education Book | Meeting Abstracts Online |
), 1.0 µM (
and 
), and 10 µM (
and 
) ThioGlo-1, and then the cells were washed free of extracellular drug and incubated in buffer A at 25°C in the presence (closed symbols) or absence (open symbols) of catalase inhibitor 3-AT (5 mM) and etoposide (400 µM) and H2O2 (20 µM). The time course of etoposide phenoxyl radical formation was recorded as described in "Materials and Methods." Data points are presented as the mean ± SE from four to six separate experiments. The thick solid line represents the level of the noise signal in the EPR spectra.
), 1.0 (speckled bars), or 10 µM (