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NEOPLASIA
Resistance of leukemic cells to 2-chlorodeoxyadenosine is due to a lack
of calcium-dependent cytochrome c release
Joya Chandra, Emma Mansson, Vladimir Gogvadze, Scott H. Kaufmann, Freidoun Albertioni, and Sten Orrenius
The purine nucleoside 2-chlorodeoxyadenosine (CdA) is often used in leukemia
therapy. Its efficacy, however, is compromised by the emergence of resistant cells.
In the present study, 3 CdA-resistant cell
lines were generated and characterized.
Their ability to accumulate 2-chloroadenosine triphosphate (CdATP) varied, reflecting differences in activities of deoxycytidine kinase (dCK) and deoxyguanosine
kinase (dGK). Nonetheless, the selected
lines were uniformly resistant to CdAinduced apoptosis, as assessed by
caspase activation and DNA fragmentation. In contrast, cytosols from resistant
cells were capable of robust caspase
activation when incubated in the presence of cytochrome c and dATP. Moreover, replacement of dATP with CdATP
also resulted in caspase activation in the
parental and some of the resistant cell
lines. Strikingly, CdA-induced decreases
in mitochondrial transmembrane potential and release of cytochrome c from
mitochondria were observed in the parental cells but not in any resistant lines. The
lack of cytochrome c release correlated
with an increased ability of mitochondria
from resistant cells to sequester free Ca2ⴙ.
Consistent with this enhanced Ca2ⴙ buffering capacity, an early increase in cytosolic Ca2ⴙ after CdA treatment of parental
cells but not resistant cells was detected.
Furthermore, CdA-resistant cells were selectively cross-resistant to thapsigargin
but not to staurosporine- or Fas-induced
apoptosis. In addition, CdA-induced
caspase-3 activation and DNA fragmentation were inhibited by the Ca2ⴙ chelator
BAPTA-AM in sensitive cells. Taken together, the data indicate that the mechanism of resistance to CdA may be dictated by changes in Ca2ⴙ-sensitive
mitochondrial events. (Blood. 2002;99:
655-663)
© 2002 by The American Society of Hematology
Introduction
The nucleoside analog, 2-chlorodeoxyadenosine (CdA, cladribine)
is used for the treatment of lymphoproliferative disorders such as
hairy cell leukemia and chronic lymphocytic leukemia (CLL) as
well as for therapy of multiple sclerosis and autoimmune disorders.1 CdA was initially described as a novel chemotherapeutic
agent2 that demonstrated greater activity than previously used
analogs. Subsequent experience, however, has shown that its
efficacy can be limited by the emergence of cells refractory to
treatment. Knowledge about the mechanism of CdA cytotoxicity is
limited, and most studies have focused on the activation and/or
deactivation of the drug.3 Among other proposed mechanisms of
action of CdA are incorporation of the analog into DNA, inhibition
of ribonucleotide reductase, inhibition of DNA repair, DNA strand
break accumulation, and activation of poly (ADP-ribose) polymerase.4 However, these possibilities require progression of cells
through cell cycle, which does not seem to occur in indolent leukemias.
CdA has also been shown to induce apoptosis in leukemic cells
in vitro and in vivo.5 Early reports described endonuclease
activation in lymphocytes isolated from patients with CLL treated
with CdA, which could be blocked by Ca2⫹ chelators such as
BAPTA (1,2-bis(o-aminophenoxy)ethane-N, N, N⬘, N⬘-tetraacetic
acid tetra(acetoxymethyl) ester).5 More recent studies of nucleoside analog–induced apoptosis have centered around protease
activation. The best characterized of the apoptotic proteases,
caspases, are cysteine proteases that cleave their substrates (more
than 100 identified to date) at a defined peptide sequence with an
aspartate residue in the P1 position.6 Two major pathways of
caspase activation have been identified. Ligation of death receptors, such as Fas or tumor necrosis factor receptor-1, can activate a
caspase cascade, beginning with an apical receptor-associated
caspase, such as caspase-8, and culminating in DNA fragmentation
mediated by a caspase-3–regulated deoxyribonuclease. Another
proposed pathway involves mitochondrial events. Cytochrome c,
released from the intermembrane space of mitochondria into the
cytosol, binds to Apaf-1, which is a homolog of the proapoptotic
nematode protein Ced-4.7 Apaf-1 also contains a binding site for
deoxyadenosine triphosphate (dATP); however, this requirement for
dATP is not completely stringent. Adenosine triphosphate (ATP) as
well as a host of intracellularly phosphorylated nuceloside analogs,
including CdATP (2-chloro-2⬘-deoxyadenosine-5⬘-triphosphate),
can also cooperate with cytochrome c to promote Apaf-1–mediated
caspase activation.8 The discovery that CdATP could induce
apoptosis via this mechanism was especially provocative because it explained how this drug might function in nonproliferating
cells in the absence of incorporation into DNA. However, this
proposed mechanism still requires the presence of cytochrome c in
the cytosol.
Despite all the reports indicating that cytochrome c release is a
critical event during apoptosis, the precise mechanism by which
cytochrome c exits mitochondria is poorly understood. One model
From the Institute for Environmental Medicine, Division of Toxicology, and
Department of Medicine, Division of Clinical Pharmacology, Karolinska
Institutet, Stockholm, Sweden.
Reprints: Joya Chandra, Guggenheim 1394, Mayo Clinic, 200 First St SW,
Rochester, MN 55905; e-mail: [email protected].
Submitted December 17, 2000; accepted September 21, 2001.
Supported by grants from the Swedish Medical Research Council, the Swedish
Children Cancer Foundation, and the Swedish Cancer Foundation. J.C. is the
recipient of a Wenner-Gren Foundation Fellowship for Visiting Scientists.
BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.
© 2002 by The American Society of Hematology
655
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656
CHANDRA et al
proposes that mitochondrial permeability transition (MPT) may be
responsible for cytochrome c release.9 Induction of MPT is
characterized by a sudden increase in the permeability of the
mitochondrial inner membrane to small ions and solutes, leading to
collapse of the membrane potential and swelling of the mitochondrial matrix. Calcium is a well-known inducer of MPT. Mitochondria protect the cell against damage due to high cytosolic Ca2⫹
concentrations by readily absorbing Ca2⫹ up to amounts nearing
100 ␮M. A further increase in mitochondrial calcium concentration
can trigger permeability transition.9 Normal lymphocytes and CLL
cells treated with nucleoside analogs or glucocorticoid display
drops in mitochondrial transmembrane potential and cytochrome c
release.10,11 Also in CLL cells and thymocytes, glucocorticoidinduced apoptosis is associated with an early sustained increase in
the cytosolic Ca2⫹ concentration, and Ca2⫹ buffering agents block
DNA fragmentation and delay cell death.12,13 These early studies
focused on the concept of a Ca2⫹-dependent endonuclease and did
not address the possibility that mitochondria might be a more
immediate target for the observed Ca2⫹ changes.
In the present study, we have generated 3 cell lines resistant to
CdA. These lines have differing levels of deoxycytidine kinase
(dCK) and deoxyguanosine kinase (dGK), the 2 enzymes that
purportedly confer activity to the drug. Consequently, the resistant
cell lines accumulate different levels of the phosphorylated drug,
CdATP, intracellularly. However, our data show that resistance
does not correlate solely with CdATP accumulation but also with
changes at the mitochondrial level that confer selective resistance
to Ca2⫹-mediated processes.
BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
were purchased from the Peptide Institute (Osaka, Japan). Thapsigargin and
BAPTA-AM were purchased from Calbiochem (San Diego, CA). Agonistic
anti-Fas monoclonal antibodies (clone CH-11) were purchased from
Medical & Biological Laboratories (Nagoya, Japan). Kaleidoscope protein
standards for Western blot analysis were purchased from Bio-Rad
(Hercules, CA).
Quantitation of dCK and dGK enzyme activities
Cells were suspended at 106/100 ␮L in an extraction buffer containing 50
mM Tris (pH 7.6), 2 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl
fluoride, 20% glycerol, and 0.5% Nonidet P-40. The suspended cells were
then frozen and thawed repeatedly 3 times and centrifuged at 14 000g in an
Eppendorf centrifuge for 5 minutes at 4°C to remove cell debris. The
supernatant was collected and used as a source of protein for the enzyme
assays. The substrates used were deoxycytidine and CdA. The concentration was chosen to be 10 times higher than the Km value for the principle
kinase phosphorylating the substrate. The assays were initiated by addition
of 2 to 3 ␮g protein to a reaction mixture containing 50 mM Tris (pH 7.6), 5
mM MgCl2, 5 mM ATP, 4 mM dithiothreitol, 10 mM sodium fluoride, and
substrate in a total volume of 25 ␮L. After incubation at 37°C for 30
minutes, 10-␮L aliquots were withdrawn and spotted on Whatman DE81
papers. Filters were then washed as described by Spasokoukotskaja et al,14
eluted by 100 ␮L 0.4 M perchloric acid, and counted in 3 mL Ecoscint
scintillation fluid in a liquid scintillation counter (RackBeta, LKB Wallac,
Turku, Finland). Conditions to maintain the linear reaction rate were
determined in preliminary experiments. The specific activity of the
enzymes was expressed in picomoles per milligram of protein per minute.
The activity of dGK was determined by using CdA in the presence of 1 mM
deoxycytidine to block the dCK enzyme.
High-performance liquid chromatography analysis
of CdA nucleotides
Materials and methods
Cell lines
The CCRF-CEM cell line was obtained from American Type Culture
Collection (Rockville, MD). CdA-resistant subclones were selected from
the original parent CCRF-CEM cell line by exposure to increasing
concentrations of CdA until final concentrations of 25 nM, 100 nM, and 1
␮M were reached in the respective cell lines. Cells were maintained in
RPMI 1640 medium containing fetal calf serum (10%), penicillin (100
U/mL), streptomycin (100 ␮g/mL), and L-glutamine (2 mM) and were kept
in humidified air, 5% CO2 at 37°C. The cells were subcultured twice
weekly, and the CdA resistance was maintained by adding relevant
concentrations of respective drugs. Cells were also routinely tested for
mycoplasma contamination. The cells were cultured in drug-free medium
for 3 passages before being used for experiments.
Antibodies and chemicals
All antibodies employed are commercially available. A murine cytochrome
c–specific antibody was purchased from Pharmingen (San Diego, CA). A
murine anti-Bax antibody was purchased from Trevigen (Gaithersburg,
MD). Rabbit anti–Bcl-xL, murine anti-Bad, and murine anti-Bid antibodies
were from Transduction Labs (Lexington, KY). Rabbit anti-Bak was
purchased from Santa Cruz Biotechnology (Santa Cruz, CA). A murine
Bcl-2 antibody was from Dako (Glostrup, Denmark). CdA was synthesized
by Dr Zygmunt Kazimierczuk at the Foundation for the Development of
Diagnostics and Therapy (Warsaw, Poland). 8-3H-CdA (4 Ci/mM
[1.48 ⫻ 1011 Bq]) and [5-3H]-deoxycytidine (16.7 Ci/mM [6.18 ⫻ 1011
Bq]) were purchased from Moravek Biochemicals (Brea, CA). Dithiothreitol, staurosporine, phenylmethylsulfonyl fluoride, Nonidet P-40, and deoxycytidine were purchased from Sigma Aldrich (Stockholm, Sweden). RPMI
1640 medium, heat-inactivated fetal calf serum, L-glutamine, and penicillinstreptomycin were from Gibco (Life Technologies, Paisley, United Kingdom). DEVD-AMC (acetyl-Asp-Glu-Val-Asp-7-amido-4-methylcoumarin)
and LEHD-AMC (acetyl-Leu-Glu-His-Asp-7-amido-4-methylcoumarin)
The accumulation of CdA nucleotides in cells was measured according to a
previously described method.15 Briefly, 10 ⫻ 106 to 20 ⫻ 106 exponentially
growing cells were exposed to 10 ␮M CdA for 12 hours. The cells were
centrifuged at 1200g for 5 minutes, washed 2 times with cold phosphatebuffered saline (PBS), and CdA nucleotides were extracted by addition of
200 ␮L ice-cold 0.4 M perchloric acid containing 0.08 M triethylammonium phosphate. The suspension was mixed and brought to pH 6.2 by
addition of 100 ␮L ice-cold 1.2 M KOH/0.5 M NH4H2PO4. After
centrifugation at 13 100g for 5 minutes at 4°C, a 100-␮L aliquot of the
supernatant was injected into the Ultrasphere ODS (250 ⫻ 4.6 mm, 5 ␮m;
Beckman Instruments, Fullerton, CA) column with the Guard Pak precolumn (Bondapak C18; Millipore, Milford, MA). The mobile phase consisted
of triethylammonium phosphate buffer (0.08 M, pH 6.1) and methanol
(89:11, vol/vol). The elution was carried out at flow rates of 1.5 mL/min at
the ambient temperature (22°C). The temperature of the autosampler was
maintained at 8°C. The concentration of the CdA nucleotides was determined at 265 nm by comparing the peak area of the CdA nucleotides in the
cells to those of the standard substances.
Quantitation of DNA fragmentation
Quantification of apoptosis by propidium iodide staining and fluorescenceactivated cell sorting (FACS) analysis was performed as described previously.16 Following incubation with various doses of CdA, cells were
pelleted by centrifugation and resuspended in PBS containing 50 ␮g/mL
propidium iodide, 0.1% Triton X-100, and 0.1% sodium citrate. Samples
were read on the FL-3 channel (FACScan, Becton Dickinson, Mountain
View, CA). A total of 10 000 events were quantitated.
Measurement of caspase-3 and caspase-9 activity
The measurement of DEVD-AMC and LEHD-AMC cleavage was performed in a fluorometric assay modified from Nicholson et al.17 Cell lysates
(generated from 1.0 ⫻ 106 cells) and substrate were combined in a standard
reaction buffer: 100 mM HEPES (DEVD-AMC) or 100 mM 2-(Nmorpholino)-ethanesulfonic acid for LEHD-AMC; 10% sucrose, 5 mM
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BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
RESISTANCE TO CdA MEDIATED BY MITOCHONDRIAL EVENTS
dithiothreitol, 0.0001% Nonidet P-40, and 0.1% 3-[(3-cholamidopropyl)
dimethylammonio] propane-1-sulphonic acid, pH 7.25 (for DEVD-AMC)
or pH 6.8 (for LEHD-AMC). This mixture was added in duplicate to a
microtiter plate. The cleavage of the fluorogenic peptide substrate DEVDAMC or LEHD-AMC (50 ␮M) was monitored by AMC liberation in a
Fluoroscan II plate reader (LabSystems, Stockholm, Sweden) using 355-nm
excitation and 460-nm emission wavelengths. Fluorescence was measured
every 70 seconds during a 30-minute period, and fluorescence units were
converted to picomoles of AMC using a standard curve generated with free
AMC. Data from duplicate samples were then analyzed by linear regression.
Determination of cytochrome c release from mitochondria
Release of cytochrome c from mitochondria was measured by immunoblotting. Cells were incubated in the absence or presence of 100 nM CdA or 1
␮M CdA for 12 hours, harvested by centrifugation, and gently lysed in an
ice-cold STE buffer containing 250 mM sucrose, 25 mM Tris, and 1 mM
ethylenediaminetetraacetic acid, pH 6.8. Lysates were immediately centifuged for 15 minutes at 15 000g. Supernatants were mixed with 2 ⫻
Laemmli reducing sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer, and extracts from equal numbers of
cells (10 ⫻ 106) were resolved by 12% SDS-PAGE. Polypeptides were
transferred to nitrocellulose membranes (0.2 ␮M, Schleicher & Scheull,
Keene, NH), and cytochrome c was detected with a monoclonal antibody (Pharmingen).
657
2-oxoethyl]amino]-3-[2-[2-[bis[2-[(acetyloxy)methoxy]-2-oxoetyl]amino]5-methylphenoxy]ethoxy]phenyl] (acetyloxy)methyl ester) (Molecular
Probes). Cells were then harvested and washed in 1 mL of Ca2⫹-free PBS.
Cells were resuspended in 1 mL Ca2⫹-free PBS and read on a Becton
Dickinson FACS Vantage SE using a UV laser on the FL4 channel
(excitation 351 nm, emission 530 nm).
Immunoblotting for Bcl-2 family proteins
For detection of Bax, Bcl-2, Bcl-xL, Bad, Bak, and Bid, cells were lysed for
1 hour at 4°C in a buffer containing 150 mM NaCl, 1% Triton X-100, a
cocktail of protease inhibitors (Complete Mini tablets; BoehringerMannheim, Indianapolis, IN), and 25 mM Tris (pH 7.5). Insoluble material
was sedimented by centrifugation for 5 minutes at 12 000g, and the
supernatants were solubilized for 5 minutes at 100°C in Laemmli SDSPAGE sample buffer containing 100 mM dithiothreitol. Polypeptides were
resolved at 130 V on a 12% gel and electrophoretically transferred to 0.2␮m nitrocellulose membranes for 2 hours at 100 V. Membranes were
blocked overnight in a TBS-T buffer (25 mM Tris [pH 8.0], 150 mM NaCl,
and 0.5% Tween 20) containing 5% (wt/vol) nonfat dried milk. Blots were
then probed for 4 hours with primary antibody and developed using a
horseradish peroxidase–coupled antimouse (Bax, Bcl-2, Bad) or antirabbit
(Bcl-xL, Bid, Bak) secondary antibody by enhanced chemiluminescence
(ECL, Amersham, Uppsala, Sweden) according to the supplier’s instructions.
Quantitation of changes in ⌬⌿mito
The potential-sensitive fluorochrome tetramethylrhodamine ethyl ester
(TMRE; Molecular Probes, Eugene, OR) was used to measure ⌬⌿mito. Cells
were obtained by centrifugation and resuspended in HEPES buffer (10 mM
HEPES-NaOH, 150 mM NaCl, 5 mM KCl, 1 mM MgCl2, and 1.8 mM
CaCl2, pH 7.4) containing 25 nM TMRE for 30 minutes at 37°C in the dark.
Cells were then analyzed on a Becton Dickinson FACScan flow cytometer
on the FL2 channel.
Isolation of cytosol and reconstitution of a cell-free model
for caspase activation
Cytosols were prepared by lysing cells in STE buffer as described above.
Cytosols were then incubated in the absence or presence of 1 mM dATP or 1
mM CdATP and 40 nM cytochrome c (Sigma, St Louis, MO) for 1 hour at
30°C. Samples were then analyzed for caspase-3–like activity as
described above.
Induction of MPT in permeabilized cells
Cells were washed in Ca2⫹-free PBS and resuspended in a buffer containing
0.15 M KCl, 5 mM KH2PO4, 1 mM MgSO4, 5 mM succinate, 5 mM Tris,
pH 7.4. After 2 minutes, cells were permeabilized with 0.005% digitonin,
and 5 ␮M rotenone was added. MPT was induced by sequential additions of
Ca2⫹ as previously described.18 Ca2⫹ fluxes were monitored with a
Ca2⫹-selective electrode.
Quantitation of intracellular Ca2ⴙ
Cells were incubated for 30 minutes at 37°C in RPMI containing 1 ␮M
Indo-1 AM (1H-Indole-6-carboxylic acid, 2-[4-[bis[2-[(acetyloxy)methoxy]-
Results
To investigate molecular mechanisms underlying CdA resistance,
cell lines that were able to survive in the presence of CdA were
generated and characterized. In brief, CEM cells were incubated in
the presence of increasing concentrations of CdA to confer
resistance. Thus, to generate the CEM CdA25 cell line, parental
CEM 0 cells were treated with 5-nM increments of CdA, increasing
every 2 weeks until the cells were in a final concentration of 25 nM
CdA. To generate the CEM CdA100 cell line, some of the CEM
CdA25 cells were further treated with 5-nM increments of CdA
every 2 weeks until the cells were finally cultured in 100 nM CdA.
The CEM CdA1000 cell line was generated in a similar manner,
except we began with CEM CdA100 cells and treated them with 50
nM increasing increments every 2 weeks. Finally, a cell line resistant to
9-␤-D-arabinofuranosyl-2-fluoroadenine (FaraA, fludarabine) that demonstrated cross-resistance to CdA was used for some of the studies
described below. These cells were designated CEM FaraA1000 and
were generated in an identical manner as the CEM CdA1000 cells,
except they were treated with FaraA instead of CdA.
The conversion of CdA to its phosphorylated form is initiated
by the generation of CdAMP from CdA. This reaction can be
catalyzed by either dCK or dGK. We measured the activity of these
enzymes in our cell lines using radioactive CdA as a substrate
(Table 1). In both CEM CdA100 and CdA1000 cells, dCK enzyme
activity was significantly lower than levels seen in CEM 0 cells.
However, in CEM CdA25 cells, dCK activity was only slightly less
Table 1. Intracellular drug activity and bioactivation enzymes vary among resistant cell lines
CEM/wt
CEM/CdA25
CEM/CdA100
CEM/CdA1000
CdATP formation (␮M) after 12-h incubation with 1 ␮M CdA
7.6 ⫾ 0.6
4.5 ⫾ 0.4
3.7 ⫾ 0.6
ND
dCK activity (pmole/mg/min)
245 ⫾ 18
204 ⫾ 5
142 ⫾ 14
4⫾1
dGK activity (pmole/mg/min)
313 ⫾ 24
428 ⫾ 48
502 ⫾ 4
426 ⫾ 15
Levels of CdATP were quantitated by high-performance liquid chromatography as described in “Materials and methods” after cells were incubated in the presence of 1 ␮M
CdA for 12 hours. Enzyme activity of dCK and dGK was measured in extracts made from sensitive and resistant cells. Radioactive CdA was used as a substrate to measure
both dCK and dGK activities. In the case of dGK enzyme activity, measurements were conducted in the presence of excess (1 mM) deoxycytidine to block dCK enzyme activity.
The specific activities of dCK and dGK are expressed in picomoles per milligram of protein per minute. Data shown are from 1 typical experiment performed 3 times in duplicate.
ND indicates not detectable.
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658
CHANDRA et al
BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
than levels in sensitive cells. We also measured dGK enzyme
activity and found that all the resistant cell lines had varying
degrees of heightened dGK activity (Table 1). Despite the increased dGK activity, intracellular CdATP levels were lower in
resistant cell lines than in parental CEM cells (Table 1). In fact, in
the case of the CdA1000 cells, levels of CdATP were extremely
low and below the detection level of our UV-based highperformance liquid chromatography assay.
Resistant cell lines do not undergo DNA fragmentation
when treated with CdA and are defective in caspase-3–
and caspase-9–like activities
Several reports indicate that CdA induces apoptosis in parental
CEM cells. To ascertain the degree of resistance in the various cell
lines, we quantitated a classical biochemical event associated with
apoptosis: the cleavage of chromatin into oligonucleosomal fragments. By staining cells with propidium iodide to visualize cell
cycle distribution, cells with subdiploid DNA content were designated to have undergone cleavage of chromatin into oligonucleosomal fragments.16 From Figure 1A it is apparent that none of the
resistant cell lines underwent DNA fragmentation after treatment
with CdA. Notably, all of the resistant cell lines, including the
FaraA1000 cells, demonstrated similarly low levels of DNA
fragmentation despite the earlier-mentioned differences in
CdATP formation.
Activation of caspases is generally considered to be a requisite
event during apoptosis. Fourteen mammalian caspases have been
described to date and can be grouped by their substrate specificity.
Fluorescently tagged versions of these preferred amino acid substrates
can be used to investigate the presence of these enzyme activities.
We conducted such assays for caspase-3–like (DEVDase) (Figure
1B) and caspase-9–like (LEHDase) (Figure 1C) activity. The results
showed that upon drug treatment, resistant cell lines displayed uniformly low levels of DEVDase and LEHDase activities.
Cytosols isolated from resistant cell lines are capable
of activating caspases
Because our data indicated that CdA treatment was not accompanied by
activation of caspase-3 and -9 in resistant cells, we explored whether
caspases could be activated in a cell-free system in these cells. Cytosols
were prepared from all cell lines and incubated with dATP and
cytochrome c. DEVDase activity was then monitored spectrofluorometrically. In the absence of added cytochrome c and/or dATP, none of the
cytosols displayed any caspase-3–like activity. This indicates that
mitochondrial release of cytochrome c did not occur during preparation
of the cytosols. When cytosols incubated in the presence of added
cytochrome c and dATP were tested for caspase-3–like activity, we
observed robust responses in cytosols from both sensitive and resistant
cells (Figure 2). In fact, in 2 of the resistant cell lines (CdA25 and
CdA100), DEVDase activity was higher than that seen in parental cells.
We included cytosols from the FaraA1000 cells in these experiments
and noted caspase activity comparable to that seen in the CdA1000 cells.
Others have shown that ATP as well as a number of purine
deoxyribonucleotide analogs can cooperate with cytochrome c to
activate caspases.8 In fact, this has been put forth as a potential
explanation for the activity of these nucleoside analogs in indolent
lymphoproliferative diseases. We tested the validity of this hypothetical mechanism in the context of our resistant cells. Incubation
of cytosols from parental CEM cells with CdATP and cytochrome c
did result in caspase-3–like activity, although the response was not
as high as that seen with dATP and cytochrome c. In the cytosols
isolated from resistant cell lines incubated with cytochrome c and
Figure 1. DNA fragmentation, caspase-3–like activity, and caspase-9–like
activity in sensitive and resistant cell lines treated with CdA. All cells were
treated in the absence or presence of 100 nM or 1 ␮M CdA. (A) DNA fragmentation.
Cells were incubated in the presence or absence of drug for 48 hours. Subsequent
propidium iodide staining and FACS analysis (as described in “Materials and
methods”) was conducted to quantitate the number of cells with subdiploid amounts
of DNA. This subdiploid population was indicative of cells with fragmented DNA and is
depicted graphically. Open bars depict cells treated with diluent alone, gray bars
represent cells treated with 100 nM CdA, and black bars show cells treated with 1 ␮M
CdA. (B) Caspase-3–like activity. Cells were incubated in the presence or absence of
CdA for 12 hours. DEVDase activity was measured using a commercially available
labeled peptide, DEVD-AMC. Cleavage of this fluorescently labeled peptide and
consequent release of free AMC, which was measurable with the use of a
spectrofluorimeter, indicated the presence of caspase-3–like proteases in cell
extracts. Color designation of bars is identical to that for panel A. (C) Caspase-9–like
activity. Cells were incubated in the presence or absence of CdA for 12 hours.
LEHDase activity was quantitated using the caspase-9 substrate, LEHD-AMC.
Cleavage of this fluorescently labeled peptide and consequent release of free AMC,
which was measurable with the use of a spectrofluorimeter, indicated the presence of
caspase-9–like proteases in cell extracts. Results shown for panels A-C are
representative of 4 separate experiments.
CdATP, DEVDase activity varied greatly (Figure 2). The CEM
CdA100 cells exhibited a response similar to that seen in the
parental CEM cells. However, none of the other cell lines showed
appreciable levels of caspase-3–like activity. These diverse responses suggest that an inability of CdATP to activate caspases via
binding to Apaf-1 does not account for resistance to CdA.
Lack of mitochondrial perturbations in all resistant cell lines
A drop in mitochondrial transmembrane potential (⌬⌿mito), which
can be quantitated with commercially available lipophilic cationic
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BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
RESISTANCE TO CdA MEDIATED BY MITOCHONDRIAL EVENTS
659
lines, including FaraA1000 cells, treated with CdA indicates that
cytochrome c release did not occur.
Protein levels of antiapoptotic Bcl-2 family members are similar
in sensitive and resistant cells
Figure 2. Caspase activity in a cell-free system. Cytosols were isolated using an
STE buffer as described in “Materials and methods.” Cytosols were then incubated at
30°C for 1 hour in the absence or presence of 1 mM dATP or 1 mM CdATP and in the
absence or presence of 40 nM cytochrome c. Extracts were then plated onto a
96-well plate, and caspase-3–like activity was quantitated with the DEVD-AMC
reagent on a spectofluorometer as described previously. Results are characteristic of
3 independent experiments.
dyes, appears to be a frequently occurring event during apoptosis.19
Another, possibly independent mitochondrial perturbation associated with apoptosis is the release of cytochrome c, which is
normally sequestered in the intermembrane space of mitochondria,
into the cytosol. This event is considered essential for caspase
activation mediated by the apoptosome pathway.
We used TMRE, a dye that loses its fluorescence upon
dissipation of mitochondrial inner membrane potential, to quantitate the effect of CdA treatment on ⌬⌿mito. In parental CEM 0 cells,
1 ␮M CdA caused a marked decrease in fluorescence, suggesting a
drop in ⌬⌿mito (Figure 3A). However, in resistant cell lines, CdA
treatment had no effect on ⌬⌿mito.
To determine whether cytochrome c was released from mitochondria upon CdA treatment, we prepared cytosols from all the cell
lines and conducted Western blot analysis using a specific antibody
to cytochrome c. This experiment revealed that in sensitive cells,
CdA treatment resulted in the appearance of a 14-kd band
corresponding to cytochrome c in the cytosolic fraction (Figure
3B). The band was stronger in cytosol from cells treated with 1 ␮M
CdA (lane 3) than in cytosol from cells treated with 100 nM CdA
(lane 2). The absence of a similar band in any of the resistant cell
Overexpression of Bcl-2 abrogates apoptosis in numerous settings
and also protects against apoptosis induced by microinjection of
low doses of cytochrome c into cytosol.20 Bcl-2 family members
have been ascribed pore-forming abilities, and several studies have
shown localization of these proteins to mitochondrial membranes.21 In fact, it has been suggested that proapoptotic Bcl-2
homologs may potentiate cytochrome c release by inserting into
mitochondria and forming a channel either alone or in combination
with constituents of the MPT. Alternatively, antiapoptotic family
members may attenuate apoptosis by binding and inactivating these
pores. We measured levels of Bcl-2 in parental and resistant cell
lines and found no difference in expression (Figure 4). Bcl-xL
levels also did not differ in sensitive or resistant cells (Figure 4).
The proapoptotic Bcl-2 family members Bax, Bak, Bid, and
Bad have been reported to promote cytochrome c release. A recent
study using cells from mice deficient in both Bax and Bak reported
that murine embryonic fibroblasts from these animals are resistant
to several diverse triggers of apoptosis.22 We sought to determine
the status of these 2 proteins in our cell lines by Western blot.
Interestingly, the proapoptotic family member Bax was virtually
undetectable in parental cells, while in all resistant cells Bax levels
were significantly higher (Figure 4). Bak, however, was present in
equivalent amounts in both sensitive and resistant cells. Bid and
Bad are 2 proapoptotic Bcl-2 family members that contain only the
BH3 domain. Expression of neither Bid nor Bad differed between
sensitive and resistant cells (Figure 4).
Lack of cytochrome c release correlates with decreased
susceptibility of mitochondria to undergo MPT
in resistant cells
Apart from the contribution of Bcl-2 family members, an alternative mechanism of cytochrome c release is due to MPT with
subsequent swelling and rupture of the outer membrane.9 As
described above, resistant cells did not release cytochrome c or lose
Figure 3. Effect of CdA treatment on mitochondria in sensitive and resistant cell lines. (A) Effect of CdA treatment on ⌬⌿mito. Cells were incubated in the absence (solid
line) or presence (dotted line) of 1 ␮M CdA for 12 hours and then stained with the cationic, lipophilic, membrane potential-sensitive fluorochrome TMRE (25 nM) for 30 minutes.
Samples were read on the FL2 channel of a Becton Dickinson FACScan. Data shown are representative of 3 independent experiments. (B) Effect of CdA treatment on release
of cytochrome c from mitochondria into cytosol. Cells were incubated in the absence or presence of 100 nM or 1 ␮M CdA for 12 hours. Cytosols were isolated as described in
“Materials and methods,” and the presence of cytochrome c was determined by Western blotting. Jurkat subcellular fractions were included as a positive control to illustrate that
although there was no cytochrome c release in resistant cells, the protein was detectable in mitochondrial fractions of untreated Jurkat cells.
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660
CHANDRA et al
Figure 4. Protein levels of Bcl-2 family members in sensitive and resistant cells.
Protein levels were detected by Western blotting as described in “Materials and
methods.” The antiapoptotic Bcl-2 family members, Bcl-2 and Bcl-xL, and the
proapoptotic members, Bax, Bak, Bid, and Bad, were investigated because of their
relevance to the process of cytochrome c release. Equal loading is evident from the
Bcl-2, Bcl-xL, Bak, and Bid blots. Protein levels for Bcl-2, Bcl-xL, Bax, Bak, and Bid
were also measured in a cell line developed with resistance to FaraA (CEM
FaraA1000) but displaying cross-resistance to CdA. Data shown are representative
of that obtained in 3 independent experiments.
⌬⌿mito upon CdA treatment. Thus, we hypothesized that this might
be due to a decreased susceptibility to MPT induction in
resistant cells.
It is well known that Ca2⫹ is an obligatory component for MPT
induction.9 Accumulation of Ca2⫹ by mitochondria results in
mitochondrial swelling, dissipation of membrane potential, and
induction of MPT, which is accompanied by release of accumulated cations and low molecular weight components (such as
adenine and pyridine nucleotides).9 In Figure 5B, MPT was
induced in digitonin-permeabilized cells by sequential additions of
Ca2⫹. Permeabilization of plasma membrane by digitonin allows
energized mitochondria to accumulate Ca2⫹ when added to the cell
suspension.18 As can be seen in Figure 5A, addition of Ca2⫹ to
permeabilized cell suspensions resulted in a rapid elevation of the
Ca2⫹ concentration in the buffer followed by the restoration of the
initial level due to accumulation of Ca2⫹ in mitochondria. Aliquots
of Ca2⫹ were added to the permeabilized cells until spontaneous
Ca2⫹ release occurred (Figure 5A). In Figure 5B, the amount of
Ca2⫹ required to trigger spontaneous Ca2⫹ release (indicative of
MPT induction) was tallied and is represented graphically. The
amount of Ca2⫹ necessary for MPT induction in resistant untreated
cells was higher than in untreated sensitive cells (Figure 5B),
indicating that the mitochondria of CdA-resistant cells are able to
buffer higher amounts of Ca2⫹ before undergoing MPT induction.
Interestingly, CdA treatment sensitized control cells, making them
even more vulnerable toward MPT, whereas in CdA25 and
CdA100 cells, more Ca2⫹ was necessary to induce MPT after
CdA treatment.
CdA-induced apoptosis requires an increase in intracellular
Ca2ⴙ and is inhibited by the intracellular Ca2ⴙ
chelator BAPTA-AM
In further experiments, we explored the potential importance of
these differences in Ca2⫹ buffering capacity. We first examined the
contribution of Ca2⫹ to the effects of CdA in sensitive parental
BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
CEM cells. If CdA-induced apoptosis requires an increase in
intracellular Ca2⫹ levels, agents that bind Ca2⫹ should reverse the
effects of CdA in these cells. To determine whether this was the
case, we pretreated parental cells with a cell-permeant Ca2⫹
chelator, BAPTA-AM, for 15 minutes prior to CdA treatment. We
found that CdA-induced caspase-3–like activity (Figure 6A) and
DNA fragmentation (data not shown) were completely abrogated
by 50 ␮M BAPTA-AM in CEM 0 cells.
The preceding results suggest that changes in intracellular Ca2⫹
should be detectable in sensitive cells after CdA treatment.
Moreover, if the altered Ca2⫹ buffering capacity of mitochondria
contributes to resistance, the Ca2⫹ fluxes should be blunted in the
resistant cells. To address these possibilities, Indo-1 AM, a
cell-permeant UV light–excitable Ca2⫹ indicator that can be
detected using a flow cytometer, was used to assess changes in
cytosolic-free Ca2⫹. Within 30 minutes after addition of CdA, an
increase in Indo-1 AM fluorescence was apparent. (Figure 7). At
later time points, this increase was not detectable. In contrast,
Figure 5. Quantitation of Ca2ⴙ-induced MPT induction in sensitive and resistant
cells. (A) Illustration of method of detection. Curves were generated using a
Ca2⫹-sensitive electrode. Two million cells were resuspended in a buffer containing
0.15 mM KCl, 5 mM KH2PO4, 1 mM MgSO4, 5 mM succinate, and 5 mM Tris, pH 7.4
(see “Materials and methods”). After 2 minutes, cells were permeabilized with 0.005%
digitonin and 5 ␮M rotenone was added to keep mitochondrial pyridine nucleotides in
a reduced state. Ca2⫹ addition, in 20 nM increments, resulted in a steep increase in
Ca2⫹, followed by restoration of the initial level due to Ca2⫹ accumulation in
mitochondria, at which point 20 nM Ca2⫹ was again added. The Ca2⫹ capacity was
calculated by adding the total amount of Ca2⫹ that could be taken up in mitochondria
before MPT induction occurred, and mitochondrial Ca2⫹ was released. (B) Ca2⫹
capacity in sensitive and resistant cells treated with diluent (open bars) or 1 ␮M CdA
(gray bars) for 12 hours. MPT was measured as described in panel A, and the amount
of Ca2⫹ required for MPT induction is expressed graphically. Data shown are
representative of 3 separate experiments.
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BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
RESISTANCE TO CdA MEDIATED BY MITOCHONDRIAL EVENTS
661
Discussion
Figure 6. Inhibition of CdA-induced apoptosis by BAPTA-AM and selective
cross-resistance to thapsigargin. (A) Cells were treated with 1 ␮M thapsigargin or
1 ␮M CdA for 12 hours, and DEVDase activity was quantitated as described in
“Materials and methods.” CdA-induced caspase-3 activity was abrogated by preincubating the cells for 15 minutes in the presence of 50 ␮M BAPTA-AM, a cell-permeant
Ca2⫹ chelator. (B) Cells were treated with diluent alone, staurosporine (100 nM) for 6
hours, or anti-Fas mAb (250 ng/mL) for 3 hours. DNA fragmentation was assessed by
propidium iodide staining and subsequent FACS analysis on the FL3 channel of a
Becton Dickinson FACScan.
CdA25 cells did not display an increase in Indo-1 fluorescence after
CdA treatment, indicating that the rise in intracellular Ca2⫹ levels
was, as predicted, blunted in these cells (Figure 7). CdA100 and
CdA1000 cells likewise failed to exhibit increases in cytosolic
Ca2⫹ (data not shown).
CdA-resistant cells are cross-resistant to thapsigargin
Because CdA sensitivity appears to rely on intracellular Ca2⫹
increases that are blocked in the CdA-resistant lines, we
postulated that the CdA-resistant cells would also be resistant to
other apoptosis inducers that rely on perturbations of intracellular Ca2⫹ homeostasis. Thapsigargin is a cell-permeable lactone
that releases endoplasmic reticular (ER) stores of Ca2⫹ by
inhibiting ER Ca2⫹-ATPases. In thymocytes and a number of
other cells, thapsigargin is known to induce apoptosis.23 In
response to thapsigargin, parental CEM cells readily underwent
apoptosis as measured by caspase-3 activation (Figure 6A) and
DNA fragmentation (data not shown). In contrast, CdA-resistant
cells were resistant to the apoptotic effects of this agent (Figure
6A). However, CdA-resistant cells were not pan-resistant to
diverse triggers of apoptosis and in fact appeared to be more
sensitive to Fas or staurosporine treatment than sensitive cells
(Figure 6B).
Although the development of nucleoside analogs has led to better
management of indolent lymphoproliferative disorders, the emergence of refractory disease remains an obstacle. A poor understanding of the mechanism of CdA action has compounded this problem.
In the present study, we have developed a series of CdA-resistant
cell lines and examined the possibility that the resistance was
related to changes in metabolic activation. Because there was not a
good correlation between CdATP formation and sensitivity, we
examined the action of CdA in the sensitive and resistant cells in
greater detail. Results of this analysis demonstrated that caspase-3
could be activated by addition of cytochrome c and dATP to cytosol
from sensitive or resistant cells. However, cytochrome c release,
MPT, and caspase-3 activation were impaired in the resistant cells.
Further experiments indicated that the resistant cells had increased
mitochondrial Ca2⫹ buffering capacity and were cross-resistant to
thapsigargin, an agent that triggers apoptosis by increasing cytosolic free Ca2⫹ concentrations. These results have important implications for the current understanding of the action of CdA in both
sensitive and resistant cells.
In previous studies of CdA resistance, enzymes controlling the
activity of nucleoside analogs have been extensively studied, but
contradictory results have been reported. In some studies dCK
levels correlate with patient responses to CdA,24 whereas in other
studies no such correlations were found.25 In fact, leukemia
patients have been shown to exhibit varying and heterogenous
levels of dCK that are not always indicative of in vivo responses.25
Consistent with this clinical situation, our resistant cell lines
displayed varying levels of dCK that did not correlate with CdA
sensitivity. Other enzymatic factors important in CdA biotransformation include dGK and 5⬘ nucleotidase (5⬘NT). The ability of
dGK to efficiently phosphorylate CdA and the mitochondrial
localization of this enzyme26 prompted us to examine levels of this
enzyme in our cell lines. However, we observed that CdA-resistant
cells display various degrees of heightened dGK activity, which did
not correlate with CdATP formation (Table 1). A recent study
illustrated that overexpression of mitochondrial dGK caused increased sensitivity to several purine nucleoside analogs, including
CdA, in solid tumor cell lines.27 Based on this observation, it was
proposed that incorporation of CdA into mitochondrial DNA was a
potential mechanism of drug action and dGK facilitated this
process. However, a separate study concluded that mitochondrial
DNA content was not affected by CdA.28 Moreover, in indolent
leukemias, incorporation of analogs into mitochondrial DNA is
Figure 7. Quantitation of intracellular Ca2ⴙ changes in sensitive and resistant
cells. Cells were treated with diluent alone (solid line) or 1 ␮M CdA (dotted line) for 30
minutes. Five minutes into the incubation, 1 ␮M Indo-1 AM was added to the culture
dishes. At the end of the incubation, cells were washed with and then resuspended in
Ca2⫹-free PBS and read on a Becton Dickinson FACS Vantage using a UV laser.
Histograms depict Indo-1 fluorescence, which indicates intracellular Ca2⫹ levels.
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662
CHANDRA et al
unlikely. Other activities that participate in CdA biotransformation
include a 5⬘NT enzyme, which opposes dCK and dGK phosphorylation of CdA. Although heightened 5⬘NT activity has been linked
to drug resistance, overexpression of 5⬘NT in a human kidney cell
line did not increase resistance to CdA.29 Because the sensitivity of
our cell lines did not correlate with levels of CdATP formed (Table
1), we considered it unlikely that alterations in CdA biotransformation provided the sole explanation for the resistance.
The recent suggestion that CdATP may directly participate in
caspase activation by binding to Apaf-130 prompted us to examine
the possibility that CdATP-induced caspase activation might be
specifically lost in the resistant cells due to metabolic inactivation
of CdATP or altered sensitivity of Apaf-1. We found that CdATP
was a poor trigger of caspase activation in sensitive cells and that
cytosols made from some CdA-resistant cells could also elicit
caspase activation when incubated with CdATP and cytochrome c
(Figure 2). Although we could not rule out the possibility that
altered sensitivity of the apoptotic apparatus to CdATP contributes
to the inhibition of apoptosis in some of the resistant lines, it cannot
contribute in all of them.
In contrast, all of the resistant lines displayed diminished
CdA-induced release of cytochrome c, the other requirement for
caspase activation by the Apaf-1–mediated pathway. Other mitochondrial perturbations associated with apoptosis—notably, loss
of ⌬⌿mito—also failed to occur in the resistant cells. Direct
estimation of MPT induction in permeabilized cells revealed that
resistant cells demonstrated greater resistance toward MPT induction by Ca2⫹.
One model for cytochrome c release cites the ability of Bcl-2
family proteins to insert into mitochondrial membranes, thus
creating a pore capable of shuttling cytochrome c from the
intermembrane space of mitochondria into the cytosol.21 Bax, Bid,
Bik, and Bak have all been implicated in cytochrome c release by
this mechanism, and their effects are antagonized by antiapoptotic
members such as Bcl-2 and Bcl-xL. In the current study, there was
no difference in either Bcl-2 or Bcl-xL expression between
sensitive and resistant lines (Figure 4). Unexpectedly, we saw that
expression of the proapoptotic family member Bax was virtually
undetectable in parental cells. Because these cells release cytochrome c without appreciable levels of Bax, this polypeptide clearly is
dispensable for CdA-induced cytochrome c release. Bak, Bid, and Bad,
all of which were detected in the parental cells, may contribute to
CdA-induced cytochrome c release in the absence of Bax. Surprisingly, we observed Bax levels in all of the resistant lines. Although
the up-regulation of Bax is unlikely to contribute to the observed
CdA and thapsigargin resistance (Figure 6A), it might explain the
slightly enhanced apoptotic responses observed when the CdAselected cells were treated with staurosporine and agonistic Fas
antibody (Figure 6B). In any case, it does not appear that changes in
the Bcl-2 family members examined explained the CdA resistance.
Although the present study has focused on caspase activation
via a mitochondrial route, death receptor ligation represents a
second way in which a caspase cascade may be initiated. Upregulation of the Fas-FasL system has been proposed to contribute
to the cytotoxic effects of certain drugs.31 Whether CdA–induced
apoptosis is triggered in this manner has been a matter of some
controversy. One recent study contends that up-regulation of Fas
and FasL occurs during CdA-induced apoptosis in MOLT-4 cells, a
lymphoma cell line.32 Other studies using CEM cells state that the
Fas pathway is not required for CdA-induced apoptosis, because
preincubation with blocking antibodies to Fas (ZB4) or FasL
(NOK-1) does not inhibit CdA-induced apoptosis.33,34 Our results
BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
complement the results of these latter studies by showing that
CdA-resistant cells retain sensitivity to Fas ligation (Figure 6B).
Also, our data corroborate and extend a recent study showing that
CdA directly affects mitochondria.35
Key experiments in the present study have included a cell line
developed specifically with resistance to FaraA but exhibiting
cross-resistance to CdA (FaraA1000). Like the CdA-resistant cells,
these cells also did not undergo cytochrome c release (Figure 3B)
or Ca2⫹-induced MPT (Figure 5B) while expressing high levels of
Bax protein (Figure 4). This suggests that mitochondrial events
may not only mediate CdA sensitivity but also control responses to
other nucleoside analogs. Interestingly, our observations mirror a
clinical study in which CdA treatment had no effect in patients with
FaraA-resistant disease.36
Mitochondria from CdA-resistant cells not only display diminished release of cytochrome c upon CdA treatment but are also less
susceptible to Ca2⫹-induced MPT. Several additional observations
indicate that an enhanced Ca2⫹ binding capacity might contribute
to the CdA resistance. First, we observed increased cytosolic free
Ca2⫹ within minutes of CdA treatment in sensitive cells (Figure 7).
Second, treatment of the sensitive cells with the Ca2⫹ chelator
BAPTA prevented CdA-induced apoptosis (Figure 6A), providing
evidence that the transient Ca2⫹ increase normally plays a role in
the cytotoxicity of CdA. Third, the increase in cytosolic free Ca2⫹
is blunted in the resistant cells, as would be predicted if they had
increased Ca2⫹ uptake into mitochondria (Figure 7). Finally, the
CdA-resistant cells are selectively cross-resistant to thapsigargin,
which triggers apoptosis by elevating cytosolic free Ca2⫹, but not
to staurosporine or anti-Fas antibody (Figure 6B). Collectively,
these observations support a model (Figure 8) in which transient
CdA-induced elevations in Ca2⫹ lead to MPT, cytochrome c
release, and caspase activation, whereas enhanced Ca2⫹ buffering
capacity leads to resistance.
The model depicted in Figure 8 explains not only the data
presented above but also the previous demonstration that Ca2⫹
chelators can prevent CdA-induced apoptosis in lymphocytes
Figure 8. Proposed scheme for CdA-induced apoptosis in CEM cells. Resistance to CdA is characterized by a lack of cytochrome c release, which is preceded by
an abrogation of cytosolic Ca2⫹ elevations and subsequent induction of MPT. CdA
treatment causes a decrease in mitochondrial Ca2⫹ buffering capacity in sensitive
CEM 0 cells but not in any of the resistant cell lines. Consequently, there is no
caspase-3–like or caspase-9–like activity in CdA-resistant cells. The cell-permeable
Ca2⫹ chelator, BAPTA-AM (50 ␮M), blocks CdA-induced caspase activation and
DNA fragmentation in parental cells, suggesting that mitochondria may be a target for
Ca2⫹-dependent apoptotic events initiated by CdA.
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BLOOD, 15 JANUARY 2002 䡠 VOLUME 99, NUMBER 2
RESISTANCE TO CdA MEDIATED BY MITOCHONDRIAL EVENTS
isolated from CLL patients.5 On the other hand, it also raises
several important questions. In attempting to understand how
transient elevations in cytosolic-free Ca2⫹ lead to MPT, we
performed additional experiments that rule out a number of
possibilities. Nur77 is a Ca2⫹-dependent transcription factor that
has recently been reported to translocate from nuclei to mitochondria, where it promotes cytochrome c release.37 In preliminary
experiments we have not observed any differences between Nur77
levels in mitochondria prepared from sensitive versus resistant
cells after CdA treatment (J.C., unpublished observations, April
2001). Calcineurin is a Ca2⫹-sensitive phosphatase that is capable
of dephosphorylating the proapoptotic protein, Bad—thus allowing
it to translocate to mitochondria, where it binds to and neutralizes
Bcl-xL.38 Preliminary observations have failed to demonstrate any
clear-cut differences between levels of Bad present in mitochondria
of sensitive versus resistant cells after CdA treatment (J.C.,
663
unpublished observations, April 2001). Calpains are a family of
Ca2⫹-dependent cysteine proteases that have been reported to
cleave Bax and other apoptotic regulators.39 In experiments designed to
examine the role of calpains in CdA-induced apoptosis, we observed
that the calpain inhibitor calpeptin did not affect CdA-induced DNA
fragmentation or cytochrome c release (data not shown). Further
experiments are therefore required to understand the manner in which
elevated cytosolic-free Ca2⫹ contributes to CdA-induced apoptosis.
Additional studies are likewise required to determine the molecular
nature of the change that renders mitochondria from CdA-resistant
cells better able to withstand Ca2⫹-induced MPT. Nonetheless, the
present results help frame these questions by providing the
unexpected finding that Bcl-2–independent alterations in mitochondrial function contribute to CdA resistance. If this resistance can be
better understood, it is possible that strategies for modulating this
resistance might ultimately be developed.
References
1. Beutler E, Sipe JC, Romine JS, Koziol JA, Mcmillan R, Zyroff J. The treatment of chronic progressive multiple sclerosis with cladribine. Proc Natl
Acad Sci U S A. 1996;93:1716-1720.
2. Keating MJ, O’Brien S, Kantarjian H, et al.
Nucleoside analogs in treatment of chronic lymphocytic leukemia. Leuk Lymphoma. 1993;
10(suppl):139-145.
3. Carson DA, Wasson DB, Taetle R, Yu A. Specific
toxicity of 2-chlorodeoxyadenosine toward resting
and proliferating human lymphocytes. Blood.
1983;62:737-743.
4. Plunkett W, Gandhi V. Pharmacology of purine
nucleoside analogues. Hematol Cell Ther. 1996;
38:S67–S74.
5. Robertson LE, Chubb S, Meyn RE, et al. Induction of apoptotic cell death in chronic lymphocytic
leukemia by 2-chloro-2⬘-deoxyadenosine and
9-beta-D-arabinosyl-2-fluoroadenine. Blood.
1993;81:143-150.
6. Thornberry NA, Lazebnik Y. Caspases: enemies
within. Science. 1998;281:1312-1316.
7. Budihardjo I, Oliver H, Lutter M, Luo X, Wang X.
Biochemical pathways of caspase activation during apoptosis. Annu Rev Cell Dev Biol. 1999;15:
269-290.
8. Genini D, Budihardjo I, Plunkett W, et al. Nucleotide requirements for the in vitro activation of the
apoptosis protein-activating factor-1-mediated
caspase pathway. J Biol Chem. 2000;275:29-34.
9. Bernardi P, Colonna R, Costantini P, et al. The
mitochondrial permeability transition. Biofactors.
1998:8:273-281.
10. Hirsch T, Dallaporta B, Zamzami N, et al. Proteasome activation occurs at an early, premitochondrial step of thymocyte apoptosis. J Immunol.
1998;161:35-40.
11. Chandra J, Niemer I, Gilbreath J, et al. Proteasome inhibitors induce apoptosis in glucocorticoid-resistant chronic lymphocytic leukemic lymphocytes. Blood. 1998;92:4220-4229.
12. McConkey DJ, Aguilar-Santelises M, Hartzell P,
et al. Induction of DNA fragmentation in chronic
B-lymphocytic leukemia cells. J Immunol. 1991;
146:1072-1076.
13. Neamati N, Fernandez A, Wright S, Kiefer J, McConkey DJ. Degradation of lamin B1 precedes
oligonucleosomal DNA fragmentation in apoptotic
thymocytes and isolated thymocyte nuclei. J Immunol. 1995;154:3788-3795.
14. Spasokoukotskaja T, Arner ES, Brosjo O, et al.
Expression of deoxycytidine kinase and phosphorylation of 2-chlorodeoxyadenosine in human
normal and tumour cells and tissues. Eur J Cancer. 1995;31A:202-208.
15. Reichelova V, Albertioni F, Liliemark JJ. Liquid
chromatographic study of acid stability of
2-chloro-2⬘-arabino-fluoro-2⬘-deoxyadenosine,
2-chloro-2⬘-deoxyadenosine and related analogues. J Chromatogr B Biomed Appl. 1996;682:
115-123.
16. Nicoletti I, Migliorati G, Pagliacci MC, Grignani F,
Riccardi CJ. A rapid and simple method for measuring thymocyte apoptosis by propidium iodide
staining and flow cytometry. Immunol Methods.
1991;139:271-279.
17. Nicholson DW, Ali A, Thornberry NA, et al. Identification and inhibition of the ICE/CED-3 protease
necessary for mammalian apoptosis. Nature.1995;376:37-43.
18. Robertson JD, Gogvadze V, Zhivotovsky B, Orrenius S. Distinct pathways for stimulation of cytochrome c release by etoposide. J Biol Chem.
2000;275:32438-32443.
19. Marchetti P, Castedo M, Susin SA, et al. Mitochondrial permeability transition is a central coordinating event of apoptosis. J Exp Med. 1996;
184:1155-1160.
20. Zhivotovsky B, Orrenius S, Brustugun OT, Doskeland SO. Injected cytochrome c induces apoptosis. Nature. 1998;391:449-450.
21. Schendel SL, Montal M, Reed JC. Bcl-2 family
proteins as ion-channels. Cell Death Differ. 1998;
5:372-380.
22. Wei MC, Zong WX, Cheng EH, et al. Proapoptotic
BAX and BAK: a requisite gateway to mitochondrial dysfunction and death. Science. 2001;292:
727-730.
overexpression of mitochondrial deoxyguanosine
kinase in cancer cell lines. J Biol Chem. 1998;
273:14707-14711.
28. Hentosh P, Tibudan M. 2-Chloro-2⬘-deoxyadenosine, an antileukemic drug, has an early effect
on cellular mitochondrial function. Mol Pharmacol. 1997;51:613-619.
29. Gazziola C, Moras M, Ferraro P, et al. Induction
of human high K(M) 5⬘-nucleotidase in cultured
293 cells. Exp Cell Res. 1999;253:474-482.
30. Leoni LM, Chao Q, Cottam HB, et al. Induction of
an apoptotic program in cell-free extracts by
2-chloro-2⬘-deoxyadenosine 5⬘-triphosphate and
cytochrome c. Proc Natl Acad Sci U S A. 1998;95:
9567-9571.
31. Friesen C, Herr I, Krammer PH, Debatin KM. Involvement of the CD95 (APO-1/FAS) receptor/
ligand system in drug-induced apoptosis in leukemia cells. Nat Med. 1996;2:574-577.
32. Nomura Y, Inanami O, Takahashi K, Matsuda A,
Kuwabara M. 2-Chloro-2⬘-deoxyadenosine induces apoptosis through the Fas/Fas ligand pathway in human leukemia cell line MOLT-4. Leukemia. 2000;14:299-306.
33. Richardson DS, Allen PD, Kelsey SM, Newland
AC. Inhibition of FAS/FAS-ligand does not block
chemotherapy-induced apoptosis in drug sensitive and resistant cells. Adv Exp Med Biol. 1999;
457:259-266.
34. McGahon AJ, Costa Pereira AP, Daly L, Cotter
TG. Chemotherapeutic drug-induced apoptosis in
human leukaemic cells is independent of the Fas
(APO-1/CD95) receptor/ligand system. Br J
Haematol. 1998;101:539-547.
23. Jiang S, Chow SC, Nicotera P, Orrenius S. Intracellular Ca2⫹ signals activate apoptosis in thymocytes: studies using the Ca(2⫹)-ATPase inhibitor
thapsigargin. Exp Cell Res. 1994;212:84-92.
35. Genini D, Adachi S, Chao Q, et al. Deoxyadenosine analogs induce programmed cell death in
chronic lymphocytic leukemia cells by damaging
the DNA and by directly affecting the mitochondria. Blood. 2000;96:3537-3543.
24. Kawasaki H, Carrera CJ, Piro LD, Saven A, Kipps
TJ, Carson DA. Relationship of deoxycytidine kinase and cytoplasmic 5⬘-nucleotidase to the chemotherapeutic efficacy of 2-chlorodeoxyadenosine. Blood. 1993;81:597-601.
36. O’Brien S, Kantarjian H, Estey E, et al. Lack of
effect of 2-chlorodeoxyadenosine therapy in patients with chronic lymphocytic leukemia refractory to fludarabine therapy. N Engl J Med. 1994;
330:319-322.
25. Albertioni F, Lindemalm S, Reichelova V, et al.
Pharmacokinetics of cladribine in plasma and its
5⬘-monophosphate and 5⬘-triphosphate in leukemic cells of patients with chronic lymphocytic leukemia. Clin Cancer Res. 1998;4:653-658.
37. Li H, Kolluri SK, Gu J, et al. Cytochrome c release
and apoptosis induced by mitochondrial targeting
of nuclear orphan receptor TR3. Science. 2000;
289:1159-1164.
26. Johansson M, Karlsson A. Cloning and expression of human deoxyguanosine kinase cDNA.
Proc Natl Acad Sci U S A. 1996;93:7258-7262.
27. Zhu C, Johansson M, Permert J, Karlsson A. Enhanced cytotoxicity of nucleoside analogs by
38. Wang HG, Pathan N, Ethell IM, et al. Ca2⫹-induced apoptosis through calcineurin dephosphorylation of BAD. Science. 1999;284:339-343.
39. Wood DE, Thomas A, Devi LA, et al. Bax cleavage is mediated by calpain during drug-induced
apoptosis. Oncogene 1998;17:1069-1078.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
2002 99: 655-663
doi:10.1182/blood.V99.2.655
Resistance of leukemic cells to 2-chlorodeoxyadenosine is due to a lack of
calcium-dependent cytochrome c release
Joya Chandra, Emma Mansson, Vladimir Gogvadze, Scott H. Kaufmann, Freidoun Albertioni and Sten
Orrenius
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