Myeloperoxidase-Dependent Oxidation of Etoposide in Human
Myeloid Progenitor CD34?Cells
Irina I. Vlasova, Wei-Hong Feng, Julie P. Goff, Angela Giorgianni, Duc Do,
Susanne M. Gollin, Dale W. Lewis, Valerian E. Kagan, and Jack C. Yalowich1
Department of Pharmacology and Chemical Biology (J.C.Y., A.G., D.D.) and Department of Radiation Oncology (J.P.G.),
University of Pittsburgh School of Medicine and Cancer Institute, Pittsburgh, Pennsylvania; Department of Environmental and
Occupational Health, Center for Free Radical and Antioxidant Health (I.I.V., W.-H.F., V.E.K.) and Department of Human
Genetics (S.M.G., D.W.L.), University of Pittsburgh Graduate School of Public Health, Pittsburgh, Pennsylvania; and Research
Institute of Physico-Chemical Medicine (I.I.V.), Moscow, Russia
Received September 7, 2010; accepted November 19, 2010
Etoposide is a widely used anticancer drug successfully used
for the treatment of many types of cancer in children and
adults. Its use, however, is associated with an increased risk of
development of secondary acute myelogenous leukemia in-
volving the mixed-lineage leukemia (MLL) gene (11q23) trans-
locations. Previous studies demonstrated that the phenoxyl
radical of etoposide can be produced by action of myeloper-
oxidase (MPO), an enzyme found in developing myeloid pro-
genitor cells, the likely origin for myeloid leukemias. We hypoth-
esized, therefore, that one-electron oxidation of etoposide by
MPO to its phenoxyl radical is important for converting this
anticancer drug to genotoxic and carcinogenic species in hu-
man CD34?myeloid progenitor cells. In the present study,
using electron paramagnetic resonance spectroscopy, we pro-
vide conclusive evidence for MPO-dependent formation of eto-
poside phenoxyl radicals in growth factor-mobilized CD34?
cells isolated from human umbilical cord blood and demon-
strate that MPO-induced oxidation of etoposide is amplified in
the presence of phenol. Formation of etoposide radicals re-
sulted in the oxidation of endogenous thiols, thus providing
evidence for etoposide-mediated MPO-catalyzed redox cycling
that may play a role in enhanced etoposide genotoxicity. In
separate studies, etoposide-induced DNA damage and MLL
gene rearrangements were demonstrated to be dependent in
part on MPO activity in CD34?cells. Together, our results are
consistent with the idea that MPO-dependent oxidation of eto-
poside in human hematopoietic CD34?cells makes these cells
especially prone to the induction of etoposide-related acute
Etoposide [VP-16, 4?-demethyl-epipodophyllotoxin-9-(4,6-
O-ethylidene-?-D-gluco-pyranoside)] is a DNA topoisomerase
II-targeting agent that has been used extensively as an an-
ticancer agent to treat a variety of malignancies in adults
and in children (Hande, 1998). However, the use of etoposide
has been associated with an increased risk of developing
secondary leukemias, especially acute myelogenous leukemia
(t-AML), bearing translocations of the MLL gene at human
chromosomal band 11q23 (Libura et al., 2005; Felix et al.,
2006). Etoposide, which contains a hindered ring phenol, can
be converted to phenoxyl radical forms by the action of per-
oxidases (Haim et al., 1987; Kagan et al., 1999). Because
myeloid progenitor CD34?cells in early stages of maturation
contain the enzyme myeloperoxidase (MPO) (Strobl et al.,
1993), we hypothesized that oxidative activation of the eto-
poside phenolic group by MPO may lead to MPO-catalyzed
oxidative stress, including carcinogenic oxidative modifica-
tion of DNA (Kagan et al., 2001). Hence, MPO expressed in
CD34?cells may make these myeloid progenitors especially
sensitive to the leukemogenic action of etoposide.
MPO-induced oxidative stress is triggered by this enzyme’s
reactive intermediates, which have very high (1.35 V) oxidiz-
This research was supported in part by the National Institutes of Health
National Cancer Institute [Grant R01-CA090787].
1Current affiliation: The Ohio State University College of Pharmacy, Co-
Article, publication date, and citation information can be found at
ABBREVIATIONS: VP-16, 4?-demethyl-epipodophyllotoxin-9-(4,6-O-ethylidene-?-D-gluco-pyranoside); MPO, myeloperoxidase; SA, succinylac-
etone; 3-AT, 3-amino-1,2,4-triazole; DTPA, diethylentriaminepentaacetic acid; CB, human umbilical cord blood; MLL, mixed-lineage leukemia;
MLLR, MLL gene rearrangements; t-AML, treatment-related acute myelogenous leukemia; EPR, electron paramagnetic resonance; DMSO,
dimethyl sulfoxide; PBS, phosphate-buffered saline; FBS, fetal bovine serum; FISH, fluorescence in situ hybridization; ACD-A, anticoagulant citrate
Copyright © 2011 The American Society for Pharmacology and Experimental Therapeutics
Mol Pharmacol 79:479–487, 2011
Vol. 79, No. 3
Printed in U.S.A.
ing potential (Jantschko et al., 2005; Davies et al., 2008). In
the presence of reducing substrates, particularly phenolic
compounds such as etoposide, the one-electron oxidation cat-
alyzed by MPO to yield phenoxyl radicals can in turn lead to
interaction with a variety of cellular targets including lipids,
thiols, ascorbate, proteins, and DNA (Zhang et al., 2002;
Borisenko et al., 2004). Depending on the reactivity of the
MPO-generated phenoxyl radicals, the oxidation of these cel-
lular constituents may be directly or indirectly involved in
MPO-driven oxidations and/or carcinogenesis (Goldman et
al., 1999; Kagan et al., 1999). In effect, the reactivity of
phenoxyl radicals determines, to a large extent, their overall
cytotoxicity and genotoxicity in MPO-expressing CD34?
cells, the likely precursors from which t-AML arises. Hence,
characterizing the interactions of etoposide phenoxyl radicals
with major cellular components is essential for a better un-
derstanding of this drug’s effects on cells (Kagan et al., 1999,
The most direct way to detect and monitor the free radical
MPO-initiated reaction is via EPR spectroscopy. We reported
previously that EPR detection of a phenoxyl radical of etopo-
side is feasible in MPO-rich human myeloid leukemia HL60
cells (Kagan et al., 2001). EPR detection of the radicals be-
came possible after depletion of GSH and other thiols, sug-
gesting that etoposide radicals (etoposide-O?) displayed reac-
(Kagan et al., 1999). Furthermore, possible involvement of
secondary reactions of thiol radicals leading to the production
of superoxide radicals and other reactive oxygen species were
considered as important cytotoxic and genotoxic events (Ka-
gan et al., 1999, 2001).
To further evaluate whether MPO is a cellular determi-
nant of etoposide oxidation, genotoxicity, and leukemogene-
sis, we evaluated MPO-catalyzed production of etoposide
phenoxyl radicals in growth factor-mobilized human CD34?
cells, a proximal progenitor model for t-AML. We report for
the first time the detection of the EPR signal of etoposide
phenoxyl radicals in intact CD34?cells and demonstrate
that this process is MPO-dependent and leads to the deple-
tion of intracellular thiols. In addition, our results demon-
strate an MPO-dependent component of etoposide-induced
DNA damage and MLL gene rearrangements, providing
“proof-of-principle” evidence for MPO as a determinant of
Materials and Methods
Materials. Etoposide (VP-16), phenol, hydrogen peroxide, succi-
nylacetone (SA), guaiacol, 3-amino-1,2,4-triazole (3-AT), phenyl-
methylsulfonyl fluoride, glucose, cetylmethylammonium bromide,
glucose, HEPES, dimethyl sulfoxide (DMSO), sodium chloride, so-
dium phosphate, diethylenetriaminepentaacetic acid (DTPA), and
myeloperoxidase (from human leukocytes, EC 220.127.116.11) were pur-
chased from Sigma-Aldrich (St. Louis, MO). Triton X-100 (t-octylphe-
noxy polyethoxyethanol) was from Bio-Rad Laboratories (Hercules,
CA). ThioGlo-1 was from Covalent Associates, Inc. (Woburn, MA).
Cell Culture and CD34?Cell Isolation. Human umbilical cord
blood (CB) samples were obtained immediately after delivery in
accordance with institutional guidelines and placed in 50-ml tubes
containing anticoagulant citrate dextrose solution (ACD-A; Cytosol
Labs, Braintree, MA). The CB was diluted with calcium- and mag-
nesium-free phosphate-buffered saline (PBS) (?) 0.6% ACD-A, and
low-density mononuclear cells were isolated by Ficoll-Paque density
gradient centrifugation for 30 min at 400g (GE Healthcare, Chalfont
St. Giles, Buckinghamshire, UK). CB mononuclear cells were
washed twice in PBS and resuspended in PBS plus 0.6% ACD-A for
magnetic labeling and separation. CD34?progenitor cells were iso-
lated using immunomagnetic selection techniques. In brief, cells
were incubated with blocking reagent (human IgG) and QBEND/10
CD34 antibody for 15 min at 4°C and washed in PBS/ACD-A followed
by incubation with a secondary antibody-magnetic microbead conju-
gate for an additional 15 min at 4°C. The unlabeled fraction of
CD34(?) cells were separated from the labeled CD34?fraction on a
high-gradient magnetic separation column (Miltenyi Biotec, Sunny-
vale, CA). Isolated CD34?cells were grown in 95% humidity under
5% CO2in air at 37°C for up to 2 weeks in Iscove’s modified Dulbec-
co’s minimal essential medium, supplemented with 10% fetal bovine
serum (FBS), 2 mM L-glutamine, and 100 ng/ml each of interleu-
kin-3, stem cell factor, and granulocyte colony-stimulating factor.
Human myeloid HL-60 cells (from the American Type Culture Col-
lection, Manassas, VA) were grown continuously in RPMI 1640
medium supplemented with 15% FBS and 2 mM L-glutamine.
CD34?and HL-60 cells were collected for experiments in mid-log
phase growth (5–8 ? 105cell/ml).
MPO Peroxidase Activity in Cell Homogenates. CD34?cells
were harvested by centrifugation at 500g for 5 min. Pellets were
washed twice with buffer A containing 25 mM HEPES and 5 mM
NaH2PO4, pH 7.4, 10 mM glucose, 115 mM NaCl, 5 mM KCl, and 1
mM MgCl2. Cell homogenates were prepared by freezing at ?80°C
and then thawing cells. Measurements of MPO activity were made at
room temperature using guaiacol as substrate. Guaiacol oxidation
was monitored by changes of absorbance at 470 nm (? ? 26.6 mM?1?
cm?1) using a spectrophotometer (Shimadzu, Kyoto, Japan). Cell
homogenates (0.1–0.4 ? 106cells) were added to 50 mM disodium
phosphate buffer containing 0.1% Triton X-100, 0.1 mM phenylmeth-
ylsulfonyl fluoride, 10 mM guaiacol, 0.02% cetylmethylammonium
bromide, and 3.75 mM 3-AT, pH 7.0. The reaction was started by the
addition of 500 ?M H2O2. Activity of MPO was calculated as nano-
moles of tetraguaiacol formed per minute per 106cells.
In Vitro MPO Peroxidase Activity Using Guaiacol and Eto-
poside as Substrates. Peroxidase activity of purified myeloperoxi-
dase was assayed both with guaiacol and etoposide in 50 mM sodium
phosphate buffer, pH 7.4, and 100 ?M DTPA at 25°C. Kinetics of
guaiacol oxidation was monitored at 470 nM (? ? 26.6 mM?1? cm?1)
after the addition of 100 ?M H2O2to the solution of 10 nM MPO and
500 ?M guaiacol. Oxidation of etoposide (200 ?M) by MPO (100 nM)
was detected by measurement of the EPR spectra of etoposide phe-
noxyl radicals 1 min after the addition of 100 ?M H2O2to the
solution. The concentration of etoposide radicals produced during the
course of the MPO reaction was estimated by the use of stable
nitroxide 4-amino-2,2,6,6-tetramethylpiperidine-1-oxyl as a stan-
dard. Double integration of the spectra was performed using the
WinSim Inc. Process simulation program (Laboratory of Molecular
Biophysics, National Institute of Environmental Health Sciences,
Bethesda, MD). The relative activity of guaiacol and etoposide as
MPO substrates was characterized by calculating the number of
substrate molecules converted to product per unit time [kcat(micro-
moles of substrate per minute per micromole of enzyme].
Immunoblotting for MPO. Whole-cell lysates were prepared
from 1 million pelleted CD34?cells by the addition of SDS-polyacryl-
amide gel electrophoresis sample buffer [50 mM Tris-HCl, pH 6.8,
1% (w/v) SDS, 10% (v/v) glycerol, and 0.5% (v/v) 2-mercaptoethanol],
followed by boiling for 5 min and brief sonication. Protein samples of
CD34?cell lysate (15 ?g) were resolved using 10% (w/v) SDS-poly-
acrylamide gel electrophoresis and then transferred to nitrocellulose.
Visual inspection of Ponceau S-stained nitrocellulose membranes
was used to ensure equivalent loading/transfer of the lysates. Mem-
branes were blocked with nonfat dry milk [3% (w/v)] in PBS
containing 0.05% (w/v) Tween 20 and then incubated with
1:40,000 dilutions of primary rabbit anti-MPO antibodies kindly
provided by Dr. William Nauseef (University of Iowa School of
Vlasova et al.
Medicine, Iowa City, IA). The secondary donkey anti-rabbit anti-
body used at 1:20,000 dilution was purchased from Jackson Im-
munoResearch Laboratories (West Grove, PA). Bound antibody
was detected using enhanced chemiluminescence (PerkinElmer
Life and Analytical Sciences, Waltham, MA).
Samples for EPR Measurements. In experiments with purified
MPO, etoposide (200 ?M) was added to the solution of MPO (25 nM)
in phosphate buffer (50 mM and 100 ?M DTPA, pH 7.4 at 25°C).
Etoposide radical formation was monitored at room temperature 1
min after the addition of 100 ?M H2O2. Generation of these radicals
in suspensions of HL-60 cells (3 ? 106cell/ml) or CD34?cells (8–10 ?
106cell/ml) in buffer A were recorded by EPR spectroscopy at room
temperature. 3-AT (6 mM) was added to the cell suspension and
incubated for 6 min, after which etoposide (200 ?M) was added.
Incubation for 2 min with etoposide was followed by the addition of
H2O2(100 ?M), and EPR spectra were recorded beginning 1 min
thereafter. For thiol depletion, the cells were preincubated in buffer
A for 10 min with ThioGlo-1 (30 ?M) followed by washout of this
maleimide reagent and additions sequentially of 3-AT, etoposide,
and H2O2as indicated above. In experiments with phenol, the ex-
perimental procedure was identical except that phenol (at various
concentrations) was added 2 min after 3-AT.
EPR Measurements. EPR spectra were recorded on a JEOL-
REIX spectrometer with 100-kHz modulation (JEOL, Kyoto, Japan)
in gas-permeable Teflon tubing (0.8 mm internal diameter, 0.013
mm thickness) obtained from Alpha Wire Corporation (Elizabeth,
NJ). The tubing was filled with 60 ?l of sample, folded doubly, and
placed in an open 3.0 mm internal diameter EPR quartz tube. Eto-
poside phenoxyl radical spectra were recorded at 3350 G, center
field; 50 G, sweep width; 10 mW, microwave power; 0.5 G, field
modulation; 103, receiver gain; 0.1 s, time constant; and 2 min, scan
time. The time course of etoposide radical EPR signals was obtained
by repeated scanning (25 s) of part of the spectrum (3350 G, centered
field; 5 G, sweep width; and other instrumental conditions were the
same). A computer simulation of the experimental spectrum was
made by the use of WinSim software package, and the numbers of
hyperfine couplings for etoposide phenoxyl radicals were published
previously (Kalyanaraman et al., 1989).
Flow Cytometry Assay for Intracellular Thiols in Native
Cells. After treatment, the cells were collected and resuspended in
PBS at a density of 0.1 ? 106cells/ml followed by incubation with
ThioGlo-1 (10 ?M) at room temperature for 10 min after washing
once with PBS. The fluorescence of ThioGlo-1 inside cells was
measured using a FACScan (BD Biosciences, San Jose, CA) flow
cytometer, equipped with a 488-nm argon ion laser and supplied
with the Cell Quest software. Mean fluorescence intensity from
104cells was acquired using a 530-nm filter (FL-1 channel). Al-
though excitation of ThioGlo-1 at 488-nm is at the tailing end of
the absorption spectrum, absorbance was sufficient for the record-
ing of emission spectra.
Fluorescence Assay for Low Molecular Weight Thiols
(GSH). Low molecular weight thiols (predominantly GSH) in cells
were determined using ThioGlo-1, which produces a highly fluores-
cent adduct upon its reaction with SH-groups. Cells (0.2 ? 105) were
suspended in PBS and lysed by repeated freeze-thaw. GSH content
was estimated by an immediate fluorescence response registered
upon the addition of ThioGlo-1 to the cell homogenate using excita-
tion at 388 nm and emission at 500 nm. The total amount of protein
was determined using the Bradford assay.
Cytogenetic Analysis for MLL Gene Rearrangements.
Growth factor-mobilized CD34?cells were treated for 60 min with
etoposide (50 ?M) or vehicle (DMSO). Cells were then washed free of
drug and allowed to grow for an additional 7 days, after which they
were treated with demecolcine (Colcemid, 0.1 mg/ml; Invitrogen,
Carlsbad, CA) for 1 h before standard cytogenetic harvesting. After
mitotic arrest, the cells were incubated in 0.075 M KCl, fixed with
Carnoy’s fixative (3:1 methanol/glacial acetic acid), and slides were
prepared for cytogenetic analysis. A dual-color break-apart DNA
probe for the detection of human MLL gene rearrangements was
obtained from Abbott Molecular Inc. (Des Plaines, IL). This probe
consists of a 350-kilobase segment centromeric to the MLL break-
point cluster region (bcr) labeled in SpectrumGreen and a 190-
kilobase segment, mostly telomeric to the MLL bcr, labeled in Spec-
trumOrange. The probe was diluted 1:5 in tdenhyb (Insitus
Biotechnologies, Albuquerque, NM). Fluorescence in situ hybridiza-
tion (FISH) assays were carried out to detect and quantify MLL gene
rearrangements. Slides were pretreated with RNase, dehydrated in
an ethanol series, denatured in 70% formamide, and hybridized with
probe overnight at 37°C in a humidified chamber. Posthybridization
washes were carried out according to the Abbott Molecular protocol.
The slides were stained with 4?,6-diamidino-2-phenylindole and
mounted with antifade composed of 1 mg/ml 1,4-phenyene-diamine
(Sigma-Aldrich) in 86% glycerol/PBS at pH 8.0. FISH signals on
metaphase spreads and in interphase nuclei were analyzed. Hybrid-
izations enabled analysis of between 124 and 274 cells for DMSO and
between 213 and 220 cells for the etoposide-treated samples. A
yellow (orange ? green) fluorochrome fusion signal suggested the
presence of an intact MLL gene, whereas separation of the two
signals indicated the presence of an MLL gene rearrangement. All
FISH analyses were carried out using an Olympus BX61 epifluores-
cence microscope (Olympus Microscopes, Melville, KY). The Genus
software platform on the Cytovision System was used for image
capture and analysis (Applied Imaging, San Jose, CA).
Comet Assays. The alkaline single-cell gel electrophoresis assay
for DNA damage (Comet assay) was performed essentially according
to instructions provided in the Trevigen Comet Assay kit (Trevigen,
Gaithersburg, MD). CD34?cells growing in complete Iscove’s
DMEM supplemented with FBS, and growth factors were incubated
in the absence or presence of succinylacetone for 63 h, after which
cells were washed and resuspended in buffer A at 37°C to a density
of approximately 500,000 cells/ml. Cells were incubated for 30 min
with etoposide (5 ?M) dissolved in DMSO or with DMSO vehicle
alone [0.2% (v/v)] followed by centrifugal collection of cell pellets.
Gentle resuspension in 1.5 ml of ice-cold 1? PBS (calcium- and
magnesium-free) was followed by a second centrifugation and resus-
pension of pellets in 0.5 ml of the same ice-cold PBS. Thirty micro-
liters of cell suspension was carefully mixed with 300 ?l of low
melting temperature agarose (Trevigen LMAgarose) at 39°C. A cell/
agarose mixture (75 ?l) was then transferred evenly to Trevigen
COMET glass slides, which were immediately placed in a desiccator
jar at 4°C in the dark for 30 min to allow the agarose to set. Slides
were then transferred into prechilled Trevigen lysis solution for 2 h
in the dark. Next, the slides were immersed in an alkaline solution
(pH ? 13, 300 mM NaOH, and 1 mM EDTA) for 1 h in the dark at
room temperature. Slides were transferred to the center of a hori-
zontal gel electrophoresis apparatus (with electrodes 34 cm apart)
containing just enough ice-cold alkaline solution (pH ?13, 300 mM
NaOH) to cover the slides. Electrophoresis proceeded in a cold room
at 1 V/cm for 45 min. After electrophoresis, the slides were drained
of excess alkaline solution, immersed in 70% ethanol for 5 min, and
air-dried overnight. Slides were stained with SYBR green, and im-
ages were visualized under a fluorescence microscope and captured
with a charge-coupled device camera. Images were imported and
analyzed using a version of the Comet Assay Software Project
(CASP) the public domain program specifically designed for the
Comet assay (Konca et al., 2003). From each slide, at least 150 cells
were analyzed. DNA damage is presented as the Olive tail moment
in etoposide-treated minus DMSO-treated cells. The Olive tail mo-
ment is defined as the product of the comet tail length and the
fraction of total DNA in the comet tail (Olive, 2002).
Statistical Analysis. The results are presented as the mean
values ? S.D. values for n ? 3, and statistical analysis was per-
formed by one-way analysis of variance. The statistical significance
of differences was set at P ? 0.05.
MPO-Dependent Oxidation of Etoposide in Human CD34?Cells
Detection of the Etoposide-O?in Intact CD34?Cells.
Our previous work demonstrated MPO-dependent one-elec-
tron oxidation of etoposide to its phenoxyl radical, etoposide-
O?, in intact HL-60 leukemia cells (Kagan et al., 2001). In
those studies etoposide-O?was directly detectable by EPR
spectroscopy after depletion of nonprotein thiols with the
maleimide reagent ThioGlo-1. Using similar experimental
conditions, we now detect the characteristic EPR signal for
etoposide-O?with g ? 2.004 in growth factor-mobilized hu-
man CD34?myeloid progenitor cells isolated from umbilical
cord blood (Fig. 1A, b). Computer simulation of the experi-
mental spectrum provided additional proof that the radical
was etoposide-O?(Fig. 1A, c). For our experimental conditions
the best simulation of the spectrum was achieved with hy-
perfine couplings aOCH3
? 1.4 G (6), aring
G (1), and a?
denote the number of equivalent protons. These are in good
agreement with hyperfine couplings published earlier for the
etoposide phenoxyl radical (Kalyanaraman et al., 1989).
Growth factor mobilization of the CD34?progenitors leads to
a progressive expression of MPO (Morabito et al., 2005).
MPO activity in these CD34?cells cultured for 1 week in the
presence of 100 ?g/ml stem cell factor, interleukin-3, and
granulocyte colony-stimulating factor was found to be 9.0 ?
2.5 nmol tetraguaiacol formed/min/106cells. In contrast,
freshly isolated CD34?cells contained no detectable MPO
activity and did not convert etoposide to its phenoxyl radical
(Fig. 1A, a). Together our results indicate that etoposide-O?is
formed in myeloid CD34?cells and that this oxidation of
etoposide is dependent on the level of MPO.
? 1.4 G (2), a?
H? 0.64 G (1). The numbers in parentheses
Because both etoposide and the added MPO cosubstrate
H2O2are cytotoxic, thereby releasing cellular contents (in-
cluding MPO), we next established that etoposide-O?radicals
were generated exclusively inside cells. CD34?cells pre-
treated with ThioGlo-1 were incubated with 200 ?M etopo-
side and 100 ?M H2O2for 45 min (H2O2was added at time 0
and every 15 min thereafter). Viability of cells treated with
etoposide and H2O2was 91 ? 5% (by trypan blue dye exclu-
sion assay) after 45-min incubation. Peroxidase activity in
collected supernatants during this incubation period was
virtually undetectable. In addition, the magnitude of the
EPR signal for etoposide-O?in supernatants collected at 15
and 45 min (after fresh addition of etoposide and H2O2)was 5
to 10% of that observed in intact cells (results not shown).
Together, these results indicate that etoposide-O?formation
is an intracellular event and that CD34?cells remained
intact during the incubation period with H2O2and etoposide.
Formation of Etoposide-O?in CD34?Cells Is MPO-
Dependent. To confirm the involvement of endogenous
MPO in one-electron oxidation of etoposide in CD34?cells,
we compared the main characteristics of the peroxidase re-
action and etoposide-O?radical generation. First, as indi-
cated above, the magnitude of the EPR signal of etoposide
radicals correlated with the MPO activity in freshly obtained
CD34?cells compared with cells grown for 1 week in growth
factor containing media (Fig. 1A, a compared with b). In
addition, the EPR signal of etoposide-O?in CD34?cells was
not detected in the absence of the MPO cosubstrate, H2O2
(Fig. 1A, d).
We next demonstrated that the formation of etoposide-O?
radicals in CD34?cells is a heme-mediated process. To this
end, we used SA, an inhibitor of heme synthesis (Pinnix et
al., 1994) and studied its effect on the EPR signal of etopo-
side-O?radicals in mobilized CD34?cells. Immunoblot anal-
ysis (Fig. 1B) indicated that mobilized CD34?cells contain
mature (heavy subunit) MPO. A 48-h incubation with 200
?M SA dramatically decreased levels of MPO in CD34?cells
(Fig. 1B). Quantitation of replicate immunoblots indicated a
reduction of MPO levels to 33.0 ? 5.5% of control cells. Under
these conditions (?SA), MPO activity was diminished to
30.5 ? 7.1% of that seen in control cells (data not shown).
Correlating with a decrease in the level and activity of MPO,
formation of etoposide-O?was dramatically decreased in
these SA-treated heme-depleted cells. The magnitudes of the
EPR signal for etoposide-O?were 45 ? 8 AU and 13 ? 4 AU
for control CD34?cells and for SA-treated CD34?cells, re-
spectively (n ? 3). The representative spectra are shown in
Fig. 1A, b and e. Cyanide (0.5 mM), an inhibitor of heme-
containing enzymes, inhibited etoposide-O?formation by ap-
proximately 90% (Fig. 1A, f). Although cyanide and SA are
not highly specific inhibitors of MPO but rather block heme-
containing enzymes and total heme synthesis, respectively,
together, these results strongly suggest that H2O2-dependent
MPO catalysis is involved in the generation of etoposide-O?
radicals, especially because MPO is a major heme-containing
protein in growth factor-mobilized CD34?cells and there is
little expression of cytochromes P450 in CD34?cells capable
of etoposide oxidation (Soucek et al., 2005).
Effect of Phenol on MPO-Dependent Etoposide-O?
Formation. The high-oxidizing potential of the reactive in-
termediate MPO compound I (1.35 V) allows for one-electron
oxidation of reducing substrates such as phenolic compounds
Fig. 1. EPR signal of etoposide phenoxyl radicals (A) and MPO levels (B)
in CD34?cells. A, CD34?cells (8–10 ? 106cells/ml) pretreated with
ThioGlo-1 were incubated with 6 mM 3-AT for 6 min. Etoposide (200 ?M)
was then added, and cells were incubated 2 min longer. EPR signal of
etoposide-O?radical was recorded beginning 2 min after the addition of
100 ?M H2O2. a, freshly obtained CD34?cells; b, growth factor-mobilized
CD34?cells; c, computer simulation of experimental EPR spectrum of
etoposide phenoxyl radical. The computer simulation was obtained by
using a line width of 0.2 G and 50% Lorentzian/50% Gaussian line
shapes. d, mobilized CD34?cells without addition of H2O2; e, mobilized
CD34?cells incubated for 63 h in the presence of succinylacetone (200
?M), an inhibitor of heme synthesis; f, mobilized CD34?cells in the
presence of cyanide (0.5 mM). B, Western blot analysis of MPO levels in
CD34?cells without (?SA) or with (?SA) a 63-h exposure to 200 ?M
succinylacetone, a heme synthesis inhibitor. The level of mature, enzy-
matically active MPO (59 kDa) was decreased in the SA-treated com-
pared with untreated CD34?cells.
Vlasova et al.
to form phenoxyl radicals (Jantschko et al., 2005; Davies et
al., 2008). Despite its low redox potential (E0? 0.56 V),
etoposide is expected to be a very poor substrate for MPO
because access of this large phenolic compound (molecular
weight ? 589) to the MPO active site is highly constrained.
The MPO active site is located at the base of a narrow and
deep heme pocket (Day et al., 1999; Zhang et al., 2002). We
measured MPO activity using both etoposide and guaiacol as
substrates. MPO activity toward etoposide was 2 orders of
magnitude less than that toward guaiacol (kcat? 8.8 ? 2.4
versus 1050 ? 150/min, respectively). This result is consis-
tent with our observation that high concentrations of etopo-
side are required to observe MPO-catalyzed production of
Compared with etoposide, small phenolic molecules (phe-
nol, tyrosine) are good substrates of MPO (Goldman et al.,
1999). These small molecules may act as cosubstrates for
oxidation of large, bulky compounds such as etoposide be-
cause they have free access to the MPO active site and
because of the relatively high oxidizing potential (approxi-
mately 0.7–0.9 V) of their phenoxyl radicals compared with
etoposide. Hence, to further indicate that MPO is responsible
for H2O2-dependent etoposide oxidation in CD34?cells, we
examined a specific feature of MPO catalysis, namely the
amplification of MPO-induced oxidation of the bulky etopo-
side molecule in the presence of the smaller phenol molecule.
Phenol-derived phenoxyl radicals were not detected under
our experimental conditions. These phenoxyl radicals are
short-lived because of their high oxidizing potential and
chemical reactivity. Phenol-derived phenoxyl radicals recom-
bine with a second order rate constant of approximately 108
M?1?s?1(Goldman et al., 1997) compared with the recombi-
nation rate constant for the relatively long-lived etoposide
phenoxyl radicals; 3 ? 103M?1?s?1(Tyurina et al., 2006).
Figure 2A shows the effect of phenol on the H2O2-depen-
dent formation of etoposide-O?by purified MPO. Given the
second order recombination rate constant for etoposide rad-
icals of 3 ? 103M?1? s?1(Tyurina et al., 2006), their lifetime
is expected to exceed the time of measurements (approxi-
mately 250 s) under our experimental conditions. Hence,
progressive accumulation of etoposide radicals can be de-
tected. We easily measured the time-dependent increase of
EPR signals of etoposide-O?when etoposide and H2O2were
added to purified MPO (Fig. 2Aa). Under the same experi-
mental conditions but in the presence of 100 ?M phenol, the
production of etoposide-O?after 1 min increased dramatically
The concentration-dependent effects of phenol on purified
MPO enzyme oxidation of etoposide are demonstrated in Fig.
2B, top. If cellular etoposide oxidation is similarly catalyzed
by MPO, then phenol-induced enhancement of etoposide ox-
idation should be observed in intact CD34?cells. Indeed, we
next demonstrate the phenol-dependent amplification of the
EPR signal of etoposide phenoxyl radicals in MPO-rich hu-
man leukemia HL-60 cells and in growth factor-mobilized
CD34?cells (Fig. 2B). In cells, there are many potential
intracellular targets beside etoposide and thiols for the
highly reactive phenol radicals. For example, in contrast to
etoposide phenoxyl radicals, phenol-derived phenoxyl radi-
cals react with lipids and proteins and oxidize them very well
(Goldman et al., 1999; Tyurina et al., 2006). This may ac-
count for the less pronounced phenol potentiation of etopo-
side-O?production in cells compared with an in vitro isolated
enzyme system (Fig. 2B). Using this in vitro model system,
phenol at 10 ?M was sufficient to increase the etoposide-O?
signal more than 2-fold. In cells, by comparison, reliable
increases of the etoposide radical signal (30%) were observed
at much higher added phenol concentrations (50 ?M) and
only after depletion of intracellular thiols (using ThioGlo-1),
one of the main targets of phenoxyl radicals (Fig. 2B). On the
other hand, it is also possible that the increase of etoposide
radical signals under these conditions may be mediated via
secondary phenoxyl radical reactions such as oxidation of
tyrosines, tryptophans, and lipids, resulting in intermediates
with high oxidizing redox potentials. Overall, in cells, the
phenol-dependent enhancement of etoposide-O?production
indicates that MPO is involved in etoposide oxidation.
Effects of Etoposide on H2O2-Induced Oxidation of
Endogenous Thiols in Human CD34?Cells. Our previ-
ous studies demonstrated that etoposide-O?radicals are re-
active toward intracellular reductants such as ascorbate (Ka-
gan et al., 1994, 1999) (suggesting a future chemoprevention
strategy to diminish the leukemogenic effects of etoposide)
and endogenous thiols such as GSH (Tyurina et al., 1995;
Kagan et al., 1999, 2001). The addition of ThioGlo-1, a ma-
Fig. 2. Phenol-induced amplification of etoposide radical formation by
MPO. A, EPR spectrum (inset) and kinetics of etoposide-O?generated by
the addition of purified MPO. MPO (25 nM) was added together with
etoposide (200 ?M) (a) or with etoposide and phenol (100 ?M) (b). The
time course of the EPR signal was monitored beginning 1 min after the
addition of H2O2(100 ?M). B, magnitude of EPR signal of etoposide-O?1
min after the addition of H2O2to a solution containing purified MPO (E,
top curve) or suspensions of cells in buffer A [Œ, HL60 cells; F, growth
factor-mobilized CD34?cells] in the presence of various concentrations of
phenol. HL60 cells treated with ThioGlo-1 were at a concentration of 3 ?
0.5 ? 106cells/ml. All other experimental conditions were the same as
detailed in the legend to Fig. 1.
MPO-Dependent Oxidation of Etoposide in Human CD34?Cells
leimide reagent capable of titrating out thiols, was essential
for observation of intracellular etoposide-O?in human my-
eloid leukemia HL-60 cells (Kagan et al., 2001). To determine
whether titrating out thiols is similarly required to observe
etoposide-O?in CD34?cells, we compared the time course of
etoposide-O?formation in this cell population before and after
treatment with ThioGlo-1. We found that the maximal EPR
signal for etoposide-O?was several times greater after Thio-
Glo-1 treatment (Fig. 3A). When cells were pretreated with
Thioglo-1, the magnitude of the signal increased over time
and then decreased (Fig. 3B), probably because of the deple-
tion of H2O2. In the absence of ThioGlo-1, there was a less
pronounced increase in the EPR signal and a more dramatic
decrease of the signal within 10 min of the addition of H2O2
(Fig. 3B). Another addition of H2O2(100 ?M) after 10 min
reconstituted the EPR signal for etoposide-O?observed 1 min
later with a greater signal recorded in cells pretreated with
ThioGlo-1 (Fig. 3B).
Repeated additions of H2O2and etoposide/H2O2in cell
suspensions not pretreated with Thioglo-1 resulted in a pro-
gressive increase in the peak magnitude of etoposide-O?rad-
ical formation followed by a decrease in EPR signal (Fig. 4A).
These results suggest that etoposide-O?reactivity toward
endogenous thiols caused oxidation and depletion of the thiol
pool, allowing for an initial increase of the signal upon fur-
ther addition of H2O2and etoposide/H2O2(Fig. 4A). Hence,
reactivity of etoposide-O?toward intracellular thiols should
be observable directly through oxidation and depletion of
SH-groups. Therefore, we measured low molecular weight
thiols and protein thiols in growth factor-mobilized CD34?
cells after incubation with etoposide and H2O2(Fig. 4B). At
various time points, aliquots of CD34?cell suspension were
taken for measurements of low molecular weight thiol con-
tent by flow cytometry after addition of ThioGlo-1. In addi-
tion, for estimation of low molecular weight thiol content in
CD34?cells, the immediate fluorescence response to the
addition of ThioGlo-1 in cell homogenates was measured in a
spectrofluorometer. Protein SH-groups were determined by
the additional increase in fluorescence response after the
addition of SDS (4 mM) to the same cell homogenates. The
validity of this technique for assessment of low molecular
weight thiols and protein-SH groups has been demonstrated
previously (Kagan et al., 2001).
We observed a time-dependent decrease in low molecular
weight thiol content both in intact cells by flow cytometry
(Fig. 4Ba) and in cell homogenates by spectrofluorometry
(Fig. 4Bb). In contrast, there was no statistically significant
decrease in free total protein SH content after 45-min incu-
bation with etoposide and H2O2(Fig. 4Bc).
These results are consistent with the idea that the abun-
dant low molecular weight thiols such as GSH are the imme-
diate reductants of the etoposide-O?, whereas protein SH-
groups are oxidized only after depletion of endogenous GSH
or when GSH levels are relatively low. In support of this
concept, we have demonstrated previously in MPO-rich
HL-60 cells that pretreatment with ThioGlo-1 to deplete cells
of GSH resulted in enhanced oxidation of protein thiols (Ka-
gan et al., 2001). When GSH levels are low, etoposide-medi-
ated protein SH oxidation in CD34?cells may be significant
at the level of DNA topoisomerase II cysteines as a determi-
nant of cytotoxicity, genotoxicity, and the known leukemoge-
nicity of this agent.
Effects of Etoposide on DNA Damage and MLL Gene
Rearrangement in CD34?Cells: Role of Myeloperoxi-
dase. When CD34?cells were pretreated with 200 ?M SA for
63 h to decrease MPO levels, etoposide (5 ?M)-induced DNA
damage assessed by Comet assay (after 30 min) was signifi-
cantly reduced compared with CD34?cells with replete MPO
(Fig. 5). DNA damage was quantified in etoposide-treated
compared with vehicle alone (DMSO) controls in SA pre-
treated or untreated cells. SA did not perturb vehicle alone
control levels of DNA damage (data not shown). These results
indicate an MPO-dependent component of etoposide-induced
DNA damage in CD34?cells consistent with the idea that
MPO catalyzed oxidation of etoposide can result in increased
genotoxicity and potentially carcinogenicity. To further es-
tablish the role of MPO in etoposide-mediated carcinogenic-
ity, we examined the effects of SA pretreatment on etoposide-
induced MLL gene rearrangements in CD34?cells. Growth
factor-mobilized human CD34?cells were pretreated for 48 h
with or without SA (200 ?M) to deplete cells of mature MPO.
Fig. 3. Time course of etoposide-O?formation in CD34?cells. A, EPR
spectra of etoposide radical formation in growth factor mobilized CD34?
cells treated with (top spectra) or without (bottom spectra) ThioGlo-1.
CD34?cells (9–11 ? 106cells/ml) in buffer A were incubated with 6 mM
3-AT, etoposide (200 ?M), and H2O2(100 ?M) was added sequentially as
in Fig. 1. Complete spectra were subsequently recorded 5 min after the
addition of 100 ?M H2O2. After 10 min, an additional aliquot of H2O2was
added, and spectra were recorded 1 min later. B, magnitude of EPR signal
of etoposide-O?in suspensions of CD34?cells treated with (?) or without
(f) ThioGlo-1 at different incubation times with 200 ?M etoposide and
100 ?M H2O2.
Vlasova et al.
Cells were then incubated for 60 min with 50 ?M etoposide
(VP-16). Cells were washed free of drug and allowed to grow
for an additional 7 days, after which metaphase chromo-
somes were analyzed for MLL gene rearrangements (MLLR)
by FISH using a dual-color break-apart probe for MLL. For
each etoposide treatment, more than 200 cells were analyzed.
No MLLR were observed in controls. For etoposide-treated
CD34?cells 3.8% of cells exhibited MLLR, and this was
reduced to 0.5% when cells were depleted of MPO. Hence, the
depletion of MPO resulted not only in decreased etoposide-O?
formation (Fig. 1A, e) but also in a reduction in both etopo-
side-induced DNA damage (Fig. 5) and MLLR.
This report presents, for the first time, direct detection and
monitoring of etoposide radicals and their enhancement by
small phenolic molecules in CD34?myeloid progenitor cells.
Detection of free radicals in cells is not trivial. Few reports
have been published demonstrating endogenous production
of free radicals in cells or in animals. An ESR spin-trapping
system was successfully implemented to detect free radicals
produced in rodents after treatment with tert-butyl hydroper-
oxide (Hix et al., 2000). A spin-trapping technique was used
as well for the detection of radicals inside macrophages
(Lopes de Menezes and Augusto, 2001). We presented evi-
dence previously for the formation of etoposide-O?in HL-60
human myeloid leukemia cells (Kagan et al., 2001) using
clinically relevant concentrations of etoposide. To date, no
reports have been published demonstrating the direct detec-
tion of MPO-derived radicals in normal cells.
Detection of etoposide phenoxyl radicals in bone marrow
CD34?progenitor cells, the likely precursors from which
myeloid tumors arise, is critical to understand the genotoxic
effects of etoposide. These myeloid progenitor cells contain
MPO even in the early stages of maturation (Strobl et al.,
1993). In our experiments, after the addition of etoposide to
human CD34?cells, we were able to detect and monitor the
EPR signal of etoposide phenoxyl radicals. Detection of eto-
poside phenoxyl radicals was dependent on the depletion of
nonprotein thiols such as GSH by preincubation of cells with
the maleimide reagent ThioGlo-1. At the same time, we were
able to obtain EPR spectra with distinguishable EPR signals
even without ThioGlo-1. The radical signal can be measured
reliably after repeated additions of H2O2and (etoposide ?
H2O2), indicating that intracellular reactions of etoposide
radicals lead to the depletion of endogenous GSH (Figs. 3 and
4A). Because the sensitivity of EPR spectroscopy is relatively
low, we used higher concentrations of etoposide. Neverthe-
less these concentrations were clinically relevant because
etoposide plasma levels of 100 ?g/ml (180 ?M) can be
achieved in patients receiving high doses of this drug
(Stremetzne et al., 1997). Under our experimental conditions,
the concentration of etoposide phenoxyl radicals accumu-
lated inside CD34?cells was as high as 0.2 ? 0.05 ?M
We provide several lines of evidence associating the ob-
served etoposide-O?radical signals in CD34?cells with the
activity of the heme protein, MPO. First, peroxidase activity
correlated with the detection of the EPR signal of etoposide-
O?. Second, H2O2, an MPO cosubstrate, dramatically stimu-
lated formation of the etoposide phenoxyl radical. Third,
generation of etoposide phenoxyl radicals was demonstrated
to be a heme-mediated process because phenoxyl radical for-
mation was inhibited by the addition of cyanide or by prein-
cubation of cells with the heme-synthesis inhibitor, succiny-
lacetone. Finally, the established MPO substrate, phenol,
amplified production of etoposide phenoxyl radicals within
cells, strongly suggesting that MPO-catalyzed one-electron
oxidation is responsible for etoposide “activation.”
As a bulky molecule, etoposide is a relatively poor sub-
strate for MPO compared with the small molecule phenol,
which has greater access to the relatively restricted active
site of the enzyme (Day et al., 1999; Zhang et al., 2002).
MPO-dependent production of nonspecific phenol radicals
results in subsequent production of the etoposide-O?based on
its specific property as a phenoxyl radical with low redox
potential compared with phenol-derived phenoxyl radicals
(Goldman et al., 1997; Goldman et al., 1999). Hence, incuba-
tion of MPO-rich cells with both phenol and etoposide may be
Fig. 4. Oxidation of SH-groups and radical formation in the
presence of etoposide in CD34?cells. A, time course of
etoposide radical formation in the suspension of growth
factor mobilized CD34?cells (without ThioGlo-1 treat-
ment) with repeated addition of H2O2. Etoposide (100 ?M)
and H2O2(100 ?M) were added to cell suspensions at time
0. As indicated, more H2O2(100 ?M) and etoposide (100
?M) plus H2O2(100 ?M)) were added to cell suspensions 10
and 30 min later, respectively. Samples for the measure-
ments of thiols were taken before any additions (control)
and then 30 and 45 min after first addition of H2O2. B, free
thiol group content in CD34?cells was measured by addi-
tion of ThioGlo-1 after cell incubation with etoposide and
H2O2either to cell suspensions (a) or to cell lysates (b and
c). a, the fluorescence of ThioGlo-1 within intact CD34?
cells assessed by flow cytometry. (Inset indicates the de-
crease in total free thiols groups in cells 45 min after
several additions of etoposide and H2O2as indicated in A);
b, time-dependent change in the level of low molecular
weight thiols (predominantly GSH) in cell lysates; c, time-
dependent change in the level of free protein thiols in cell
lysates. ?, P ? 0.05 compared with 0 time control.
MPO-Dependent Oxidation of Etoposide in Human CD34?Cells
responsible for converting nonspecific phenol radicals, which
are broadly reactive toward different biomolecules, to specific
interactions of etoposide-O?radicals with GSH and pro-
tein-SH groups. This specificity may be responsible for mod-
ified and/or inhibited protein activities.
Because the etoposide phenoxyl radical is relatively long-
lived compared with the phenol radical (Goldman et al., 1997;
Tyurina et al., 2006), the oxidative potential of MPO can be
more readily transferred to the nucleus when cells are rela-
tively depleted of intracellular reductants. In addition, it has
been demonstrated that MPO can be found in the nucleus of
normal and leukemic human myeloid cells (Murao et al.,
1988). Hence, one suggestion from our results is that expo-
sure of MPO-containing myeloid progenitors to etoposide or
concurrently to etoposide and phenols increases the risk of
etoposide-induced secondary acute myeloid leukemias based
on increased oxidative stress and potential oxidative DNA
damage, resulting in abasic DNA sites. These abasic sites can
act to enhance DNA topoisomerase II poisoning (Kingma et
al., 1997) and may thereby increase the recombinogenic ac-
tivity of etoposide.
In support of the idea that hematopoietic cell cytotoxicity
(and presumably genotoxicity) can be enhanced by MPO-
catalyzed oxidation of phenol, it was reported that phenol
stimulated hydroquinone oxidation by peroxidases (Smith et
al., 1989). In addition, repeated coadministration of phenol
and hydroquinone to B6C3F1mice resulted in a dramatic
decrease in bone marrow cellularity (Eastmond et al., 1987).
Together with the results presented here, these reports raise
a cautionary note concerning environmental phenol exposure
in patients receiving etoposide therapy. In contrast, dietary
flavonoids, relatively weak oxidants, can be viewed as
potentially important competitive inhibitors of MPO-cata-
lyzed etoposide oxidation useful for the prevention of eto-
Peroxidase-dependent formation of phenoxyl radicals in
the presence of glutathione is known to derive thiyl radicals
and to provide an additional important source of reactive
oxygen species, thus propagating oxidative stress in MPO-
rich cells (Borisenko et al., 2004). The reaction of thiyl radi-
cals with GSH in the presence of oxygen leads to disulfide
anion-radical formation, a reducing radical that can readily
donate an electron to molecular oxygen to yield O2.. Dispro-
portionation of superoxide radicals leads to the accumulation
of H2O2, a source of oxidizing equivalents for the MPO-
catalyzed reactions. Hence, preferable oxidation of SH-
groups by etoposide phenoxyl radicals may cause formation
of oxygen radicals and etoposide redox-cycling implying the
amplification of oxidative damage (and potentially recombi-
nogenic oxidative DNA damage) in MPO-rich cells, including
myeloid progenitor cells.
Rearrangements involving the MLL gene on chromosome
band 11q23 are a hallmark of therapy-related acute myeloid
leukemias after treatment with DNA topoisomerase II poi-
sons including etoposide (Felix et al., 2006). Acute myeloid
leukemia-like MLL rearrangements are induced by etoposide
in primary human CD34?cells and remain stable after
clonal expansion (Libura et al., 2005). Etoposide promotes
specific rearrangements of MLL in CD34?consistent with
the full spectrum of oncogenic events identified in leukemic
samples. However, the mechanisms underlying etoposide-
induced genotoxicity and leukemogenesis remain controver-
sial. One possibility, as suggested above, is that MPO-cata-
lyzed oxidation of etoposide leads to oxidative DNA damage,
abasic sites, and resultant enhancement of DNA topoisomer-
ase II-mediated MLL gene rearrangements. Others have sug-
gested illegitimate recombination events in response to for-
mation of DNA topoisomerase II covalent complexes may
initiate MLL rearrangements (Sung et al., 2006). Any dys-
function or variation of DNA damage sensor and repair pro-
teins might be expected to influence both the frequency and
spectrum of repair products. In addition, when concentra-
tions of endogenous GSH are low (either intrinsically or as a
result of an oxidative process), MPO-catalyzed etoposide-O?
formation may allow for direct oxidation of cysteines on to-
poisomerase II or other DNA-repair proteins that are essen-
tial for their functions. In particular, topoisomerase II is a
strongly SH-dependent endonuclease. Its inhibition is asso-
ciated with genotoxicity/mutagenicity and development of a
procarcinogenic phenotype (Hutt and Kalf, 1996). Regardless
of the mechanism(s) responsible for etoposide-induced leuke-
mogenesis, our results demonstrate MPO dependence for
both etoposide-induced DNA damage (Fig. 5) and for MLL
MPO is not the only enzyme that oxidizes etoposide. It has
been suggested that oxidative activation of etoposide by cy-
tochrome P450 monooxygenases, prostaglandin synthetase,
and tyrosinase may contribute to its cytotoxicity (Haim et al.,
1987; Usui and Sinha, 1990; Relling et al., 1994). However,
these enzymes are not expressed appreciably in CD34?cells
(Fan et al., 2006). We hypothesize, therefore, that it is the
oxidizing enzyme MPO in highly proliferative CD34?cells that
increases the risk of etoposide-induced secondary leukemias.
Oxidation of etoposide results in the formation of several
metabolites (Fan et al., 2006; Zheng et al., 2006). Patients
receiving etoposide accumulate appreciable levels of etopo-
side catechol catalyzed by the action of CYP3A4 and CYP3A5
(Zheng et al., 2004; Zhuo et al., 2004). Etoposide catechol can
be converted to semiquinone radicals by one-electron oxida-
tion catalyzed by MPO. MPO may also catalyze (in cells and
in vitro) the formation of a two-electron oxidation product of
Fig. 5. MPO-dependent etoposide-induced DNA damage in CD34?cells.
Growth factor-mobilized CD34?cells were incubated for 63 h in the
absence or presence of SA (200 ?M) followed by a 30-min incubation with
etoposide (5 ?M) or vehicle control [DMSO 0.2% (v/v)]. Cells were then
processed and evaluated for DNA damage by Comet assay as described
under Materials and Methods. Results shown are the mean ? S.E.M. for
triplicate measurements during each of three experiments run on sepa-
rate days. ?, P ? 0.05 comparing etoposide effects in SA-treated and
Vlasova et al.
etoposide, etoposide ortho-quinone that forms conjugates
with GSH (Mans et al., 1992; Fan et al., 2006; Zheng et al.,
2006) and may be responsible in part for the cyto- and geno-
toxic effects of etoposide because of its ability to stabilize
topoisomerase II/DNA covalent complexes as studied in vitro
(Lovett et al., 2001).
In conclusion, our study demonstrates MPO-dependent
generation of etoposide-O?radicals in human myeloid progen-
itor CD34?cells relevant to etoposide genotoxicity and car-
cinogenesis. EPR signals of etoposide phenoxyl radicals can
be directly measured in these cells and can be used as a
model to study MPO-induced oxidation of phenolic com-
pounds. The data obtained should facilitate future studies of
the mechanisms of etoposide-associated secondary leukemias
and potentially lead to the development of nutritional anti-
oxidant strategies to limit and/or prevent MPO-catalyzed
formation of etoposide metabolites causative for therapy-
related myeloid leukemias.
We thank Dr. Ching-Shih Chen and Dr. Mary K. Ritke for critical
evaluation of this manuscript.
Participated in research design: Vaslova, Feng, Gollin, Lewis, Ka-
gan, and Yalowich.
Conducted experiments: Vaslova, Feng, Giorgianni, Do, Lewis, and
Contributed new reagents or analytic tools: Goff.
Performed data analysis: Vaslova, Feng, Giorgianni, Do, Gollin,
Lewis, and Yalowich.
Wrote or contributed to the writing of the manuscript: Vaslova,
Goff, Giorgianni, Do, Gollin, Lewis, Kagan, and Yalowich.
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MPO-Dependent Oxidation of Etoposide in Human CD34?Cells