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Introduction
The epidermal growth factor receptor (EGFR) is a
receptor tyrosine kinase involved in multiple physiological
processes including cell proliferation, survival and
migration[1]. EGRF is highly expressed in solid tumor cells such as the
breast, lung, colon, ovaries and
brain[2]. According to previous studies, the constitutive expression of EGFR activates
intracellular signal transduction cascades involving the
Ras-Raf-mitogen-activated protein kinase (MAPK),
phosphatidyl inositol 3-kinase (PI3K)/Akt, and signal transducer and
activator of transcription (STAT)
pathways[3]. All of these pathways have been implicated in the inhibition of apoptosis
and the promotion of tumor cell
motility[4,5]. Therefore, drugs and monoclonal antibodies designed to inhibit EGFR tyrosine
kinase activity are actively being developed for cancer
therapy[6_8].
ZD1839 (Iressa) is a quinazoline derivative that inhibits
EGFR tyrosine kinase activity by binding to the adenosine
triphosphate pocket within EGFR's catalytic
domain[9]. Although ZD1839 has been proposed as a biologically targeted
agent for the treatment of
cancer[10_14], Govindan et
al[15] proved that ZD1839 was not active in malignant
mesothe-lioma, regardless of EGFR expression. The clustered
incidence of acute promyelocytic leukemia was also detected during
ZD1839 treatment of non-small-cell lung
cancer[16]. On the other hand, Stegmaier
et al[17] recently reported that ZD1839
induced myeloid differentiation of acute myeloid leukemia
without EGFR at clinically achievable doses. However, the
mechanisms responsible for its effects on leukemia cells
without EGFR have yet to be fully elucidated.
In the present study we examined the effect of ZD1839
on antiproliferation and apoptosis in human leukemic U937
cells without EGFR. The results provide new insight about
the function of Bcl-2 and caspase-3 during apoptosis in
response to ZD1839 exposure in leukemic U937 cells. Our
results also demonstrate that ZD1839 can initiate a substantial
apoptotic response through the downregulation of
extracellular signal-regulated kinase (ERK) and PI3K/Akt.
Materials and methods
Reagents Propidium iodide (PI),
4,6-diamidino-2-phenylindole (DAPI) and
3-(4,5-dimethyl-2-thiazolyl)-2,5-diphnyl-2H-tetrazolim bromide (MTT) were purchased
from Sigma (St Louis, MO, USA). Caspase activity assay
kits were purchased from R&D systems (Minneapolis, MN,
USA). An enhanced chemiluminescence (ECL) kit was
purchased from Amersham (Arlington Heights, IL, USA).
Caspase-3 inhibitor 1 z-DEVD-fmk, PD98059, SP600125,
SB203580, and LY294002 were obtained from Calbiochem
(San Diego, CA). RPMI-1640 medium and fetal bovine
serum (FBS) were purchased from Invitrogen Co (Carlsbad,
CA, USA) and GIBCO-BRL (Gaithersburg, MD, USA),
respec-tively. All other chemicals not specifically cited here were
purchased from Sigma (USA).
Antibodies Anti-inhibitor of apoptosis protein (cIAP)-1,
anti-cIAP-2, anti-X-linked inhibitor of apoptosis (XIAP),
anti-Bcl-2, anti-Bax, anti-Bad, anti-poly (ADP-ribose) polymerase
(PARP), anti-phospholipase (PLC)-γ1, anti-caspase-3,
anti-caspase-8, anti-caspase-9, antibodies against PI3K and
phosphor (p)-PI3K, respectively, were purchased form
Santa Cruz Biotechnology (Santa Cruz, CA, USA).
Antibodies against ERK, p-ERK, p38, p-p38, c-Jun-N-terminal
kinase (JNK), p-JNK, Akt and p-Akt were purchased from
PharMingen (San Diego, CA, USA), and the antibody against
b-actin was from Sigma (USA). Peroxidase-labeled
donkey anti-rabbit and sheep anti-mouse immunoglobulin were
purchased from Amersham (USA).
Cell culture Human leukemic U937 cells were obtained
from the American Type Culture Collection (Manassas, VA,
USA). Bcl-2-overexpressing U937 (U937/Bcl-2) cells were
kindly provided by Professor TK KWON (Department of
Immunology, School of Medicine, Keimyung University,
Daegu, South Korea) in South Korea. The cells were
cultured at 37 °C in a humidified incubator with 5%
CO2 and maintained in RPMI-1640 medium containing 10%
heat-inactivated FBS.
Cell viability and proliferation The cells were treated
with the indicated concentrations of ZD1839 for 48 h, and
the cell number and viability were determined by trypan blue
exclusion staining and MTT assays, respectively. The
control cells were supplemented with complete media
containing 0.1% DMSO as a vehicle control for 48 h.
Nuclear staining After treatment with ZD1839, the cells
were harvested, washed in ice-cold phosphate-buffered
solution (PBS), fixed with 3.7% paraformaldehyde, and then
permeabilized in PBS containing Triton X-100. The fixed
cells were washed with PBS and the nuclei were stained with
DAPI solution. Nuclear morphology was evaluated by
fluorescence microscopy.
Cell cycle analysis The cells were serum-starved for
24 h in order to synchronize them in the
G0 phase of the cell cycle. Synchronous populations of cells were treated with
ZD1839 for 48 h. The cells were fixed in 75%
(v/v) ethanol for 1 h at 4 °C and resuspended in cold PI solution (50 µg/mL)
containing RNase A (0.1 mg/mL) in PBS (pH 7.4) for 30 min in
the dark. Flow cytometry was performed using FACSCalibur
(Becton Dickinson, San Jose, CA, USA). Forward light
scatter characteristics were used to exclude the cell debris from
the analysis. The sub-G1 population was calculated in order
to estimate the apoptotic cell population.
Protein extraction and Western blot
analysis The cells were harvested and gently lysed for 2 min in ice-cold lysis
buffer [20 mmol/L sucrose, 1 mmol/L EDTA, 20 µmol/L
Tris-Cl (pH 7.2), 1 mmol/L dithiothreitol, 10 mmol/L KCl,
1.5 mmol/L MgCl2, 5 µg/mL pepstatin A, 10 µg/mL leupeptin
and 2 µg/mL aprotinin]. The lysates were centrifuged at
14 000×g at 4 °C for 10 min. The supernatant was collected
and the protein concentrations were determined using a
Bio-Rad protein assay kit (Bio-Rad, Hercules, CA, USA). The
samples were either stored at -80 °C or used immediately
for the Western blot analysis. Aliquots containing 30 µg
total protein were separated on SDS-PAGE and transferred
to nitrocellulose membranes (Schleicher & Schuell,
Keene, NH, USA). The proteins were detected using an
ECL detection system (Amersham, USA).
Determination of caspase activity Caspase activities were
determined by a colorimetric assay according to the
manu-facturer's instructions. Briefly, the cells were lysed in the
supplied lysis buffer. The supernatant was collected and
then incubated with the supplied reaction buffer containing
dithiothreitol and substrate. The reaction was measured by
changes in absorbance at 405 nm using the
VERSA-max microplate reader (Molecular Devices, Palo Alto, CA, USA).
DNA fragmentation assay U937 and U937/Bcl-2 cells
were treated with different concentrations of ZD1839 for
48 h and were lysed on ice in a buffer containing 10
mmol/L
Tris-HCl (pH 7.4), 150 mmol/L NaCl, 5 mmol/L EDTA, and
0.5% Triton X-100 for 30 min. The lysates were vortexed and
cleared by centrifugation at 10 000×g for 20 min. Fragmented
DNA in the supernatant was extracted with an equal volume
of neutral phenol:chloroform:isoamyl alcohol (25:24:1,
v/v/v) and analyzed electrophoretically on 1.5% agarose gel
containing ethidium bromide.
Determination of cytotoxicity For the determination of
plasma membrane integrity loss, lactate dehydrogenase
(LDH) release into the extracellular medium was measured
using the cyto-tox96 non-radioactive assay from Promega
(Madison, WI, USA) in order to determine cytotoxicity. This
assay measures the formation of a red formazan product
after the conversion of lactate and nicotinamide adenine
dinucleotide (NAD+) to pyruvate and
NAD+ hydrogen. The assay was used according to the manufacturer's instructions.
Briefly, the maximum release of LDH was obtained by adding
100 µL of 2% Triton X-100 to the untreated cells. One
hundred microliters of each sample were incubated with 100
µL of LDH assay reagents for 10 min and the absorbance of
the samples was measured at 490 nm. The percentage of
LDH release was determined by dividing the amount of LDH
released by the cells under each condition by the maximum
amount of LDH release, and then multiplying the fraction by
100.
Statistical analysis All data are presented as mean±SD.
Significant differences between the groups were determined
using the unpaired Student's t-test. A value of
P<0.05 was accepted as an indication of statistical significance. All the
figures shown were obtained from at least 4 independent
experiments with a similar pattern.
Results
ZD1839-induced apoptosis in a concentration-dependent
manner in U937 cells We first evaluated the ability of
ZD1839 to attenuate the proliferation and viability of U937
cells by cell-counting and MTT assay. As shown in Figure
1A and 1B, ZD1839 treatment significantly decreased cell
proliferation and viability in a concentration-dependent
manner. Cellular death was not observed when treated with
ZD1839 at a concentration of 10 µmol/L, however, a marked
decrease in cell numbers
[(465±21)×103 cells/mL)] and
viability (69%±4%) was observed at 15 µLmol/L. Exposure to 25
µmol/L of ZD1839 sharply decreased cell numbers and
viability to (332±35)×103 cells/mL and
46%±7%, respectively. Additionally, the U937 cells showed a marked change in
morphology including shrinkage, irregular shape, and
condensed chromatin in their nuclei (Figure 1C) following 48 h
of exposure to 20 or 25 µmol/L ZD1839. The cell cycle
analysis also revealed that 25 µmol/L ZD1839 significantly
increased the sub-G1 DNA content (33%±4%), indicating
apoptosis (Figure 1D). These data indicate that ZD1839
inhibits cell proliferation and induces cell death in U937 cells.
Caspase-3 is a potential target of ZD1839-induced
apoptosis In order to examine whether caspases were likely
involved in the apoptotic response, the expression and the
activation of caspases-3, -8, and -9 were evaluated
following treatment with ZD1839 (Figure 1). The cells were
treated with ZD1839 at the indicated concentrations for
48 h, and the caspase activity from the cell lysates was
determined. As shown in Figure 2A, ZD1839 triggered
caspase-3 activation in a dose-dependent manner; however
caspases-8 and -9 were activated only at ZD1839
concentrations greater than 20 µmol/L. The activation of caspases
was also confirmed by the decrease of proform caspase by
Western blot analysis (Figure 2B). In a parallel experiment,
treatment with ZD1839 at concentrations of more than 20
µmol/L decreased procaspases-3, -8, and -9, followed by
increases in PARP cleavage and PLC-γ degradation. We next
attempted to determine whether caspase-3 was likely to play
an important role in ZD1839-induced apoptosis by treating
cells with the specific caspase-3 inhibitor z-DEVD-fmk. The
inhibition of caspase-3 activity by pretreatment with 50
µmol/L z-DEVD-fmk significantly decreased the appearance of
sub-G1 DNA content and apoptotic bodies following ZD1839
treatment (Figure 2C). Additionally, the DNA fragmentation
(Figure 2D) and LDH release (Figure 2E) associated with
ZD1839 were significantly
attenuated 1 h after pretreatment with z-DEVD-fmk.
Further-more, ZD1839 treatment was associated with the appearance
of caspase-3 and PARP cleavage products; however, a
significant decrease in these products was observed following
pretreatment with z-DEVD-fmk (Figure 2F). These results
suggest that ZD1839-induced apoptosis may be executed
by activating the caspase-3 pathway.
Ectopic expression of Bcl-2 significantly attenuates
ZD1839-induced apoptosis Bcl-2 and IAP family members
could ultimately inhibit or promote apoptosis. Therefore, we
also investigated whether ZD1839 treatment can modulate
the expression of these pro- and anti-apoptotic proteins.
Western blot analysis revealed that ZD1839 significantly
downregulated anti-apoptotic proteins such as Bcl-2,
cIAP-1 and XIAP, and upregulated the pro-apoptotic protein Bax in
a dose-dependent manner (Figure 3A). However, ZD1839
had no effect on the levels of Bad or cIAP-2. These results
indicate that ZD1839 is likely to induce apoptosis in U937
cells by downregulating the expression levels of Bcl-2,
cIAP-1, XIAP, and upregulating Bax. In order to further elucidate
the role of Bcl-2 in ZD1839-induced apoptosis in U937 cells,
we next investigated the suggestive apoptotic features in
U937/Bcl-2 cells. Western blot analysis confirmed that
U937/Bcl-2 cells had significantly higher levels of Bcl-2 than the
control U937 cells (data not shown). The ectopic expression
of Bcl-2 also completely protected U937 cells from the
formation of sub-G1 and apoptotic bodies induced by ZD1839
(Figure 3B). Although ZD1839 clearly induced a potent DNA
fragmentation and LDH release in U937 cells after 48 h of
treatment, these features were not observed or were
significantly reduced in U937/Bcl-2 cells (Figures 3C, D).
Consistent with the inhibition of apoptosis, the cleavage of
caspase-3 and PARP in U937/Bcl-2 cells was significantly reduced.
These results indicate that the downregulation of Bcl-2
proteins might be associated with ZD1839-induced apoptosis
in U937 cells, and might act as an upstream regulator of
caspase-3 and PARP.
Inhibition of the ERK pathway increases
ZD1839-induced apoptosis In order to investigate the role of the
MAPK signaling pathway in ZD1839-induced apoptosis, we
investigated the effect of ZD1839 on the expression and
function of MAPK. As shown in Figure 4A, the
phosphorylation of p38 MAPK increased significantly from 24 to
48 h after ZD1839 treatment. However, ERK and JNK
activation gradually decreased after 24 h of ZD1839 treatment.
We next evaluated the possible roles of MAPK in
ZD1839-induced apoptosis. As shown in Figures 4B and 4C,
treatment with SB203580, a specific p38 MAPK inhibitor, slightly
increased cellular viability (55%±4%), although this increase
was not statistically significant. However, pretreating cells
with PD98059, a potent inhibitor of ERK, modestly increased
the sub-G1 phases (32%±4%) and decreased cellular
viability (35%±6%) in the presence of ZD1839. Interestingly,
treating U937 cells with 5 µmol/L of SP600125, a potent inhibitor
of JNK, reduced the number of cells with
sub-G1 DNA content from 24%±4% to 10%±3% and increased cellular
viability from 47%±5% to 92%±4%, but increased arrest to the
G2 phase. Treatment with 5 or 10 µmol/L SP600125 could
significantly attenuate the appearance of ZD1839-induced DNA
fragmentation and LDH release through an increase in
G2 levels, however, a high dose (20 µmol/L) of SP600125 did not
inhibit the appearance of apoptotic features (Figures 4D, E).
It is possible that SP600125 itself increased apoptosis or
G2 arrest. These results led us to believe that ZD1839 is likely
to downregulate the ERK pathway, suggesting that this
pathway is likely involved in ZD1839-induced apoptosis.
PI3K inhibitor LY294002 sensitizes ZD1839-induced
apoptosis We further questioned whether ZD1839 alters PI3K
and Akt activation in U937 cells. In order to investigate the
activation of these cascades, we determine the expression
and phosphorylation levels of PI3K and Akt after treatment
with 25 µmol/L of ZD1839. The levels of phosphorylated
PI3K decreased significantly in response to ZD1839 after
treatment for 6 h. Consistent with this, the levels of
phosphorylated Akt also decreased from 24 to 48 h (Figure 5A).
The total PI3K and Akt protein levels remained constant at
various times points during ZD1839 treatment. We further
investigated whether the activation of the PI3K/Akt
pathways is necessary for ZD1839-induced apoptosis. The PI3K
inhibitor LY294002 was used to determine whether the Akt
phosphorylation inhibitor was responsible for the induction
of apoptosis. As shown in Figure 5B, the inhibitor (25
µmol/L) alone modestly increased in apoptosis and
G1 arrest, and cotreatment with ZD1839 markedly increased apoptosis, as
determined by morphological change, DNA condensation,
and sub-G1 DNA content. We next analyzed cellular viability,
DNA fragmentation, and LDH release in order to further
elucidate the relationship between the Akt pathway and
ZD1839-induced apoptosis in U937 cells. Treatment with 25
µmol/L of LY294002 significantly decreased ZD1839-induced
cell viability from 53%±4% to 36%±7% (Figure 5C). We also
found that co-treatment with LY294002 and ZD1839
markedly increased the level of DNA fragmentation (Figure 5D)
and LDH release (Figure 5E). These results indicate that
ZD1839-induced apoptosis may be associated with the
down-regulation of the Akt signal pathway in this system.
Discussion
In the present study, we first demonstrated that ZD1839
significantly inhibited the proliferation of, and promoted
apoptosis in, human leukemic U937 cells without EGFR.
Thus, this study indicates that ZD1839-induced apoptosis
in U937 cells is mediated by Bcl-2 downregulation, which
subsequently activates caspase-3 followed by the cleavage
of PARP and PLC-γ for apoptosis. These pathways occurred
from unknown causes, because most leukemic cells have
little EGFR and are insensitive to low doses.
Bcl-2 and IAP proteins have been investigated as
potential therapeutic targets on the basis of their ability to disrupt
apoptosis and to confer resistance to chemotherapy in
cancer cells[18,19]. Bcl-2 can form ion channels in biological
membranes[20], and those channels may control apoptosis
by influencing intracellular membranes' permeability and
cytochrome c release from
mitochondria[21]. In this study, ectopic expression of Bcl-2 and inhibition of caspase-3
significantly promoted cell viability. In contrast, Chang
et al[22] found that ZD1839 did not change Bcl-2 or Bax
expression, and that ectopic expression of Bcl-2 could not
prevent ZD1839-induced apoptosis, thus suggesting that a
Bcl-2 protein-independent pathway was associated with
ZD1839-mediated apoptosis in human lung adenocarcinoma
A549 cells. Conversely, some researchers have reported
that ZD1839-induced apoptosis involves a Bcl-2-dependent
mechanism[11] and that combined targeted inhibition of
Bcl-2 and other anti-apoptotic proteins resulted in potent
apoptotic activity[23]. These disparities may be due to the
inherent biological differences between the investigated cell
types. Current insight also suggested that the IAP family,
including cIAP-1, cIAP-2, and XIAP, inhibit apoptosis by
directly inhibiting activation of effector
caspases[24,25]. A shift toward death signals is the activation of effector
caspases such as caspases-3 and -9, and caspase signaling
is initiated and propagated by substrates such as
PARP[26]. Nevertheless, it is still not known whether ZD1839-induced
apoptosis is related to the downregulation of IAP family
proteins. Our results suggest that ZD1839-induced
apopto-sis is associated with decreased expression levels of cIAP-1
and XIAP, but not cIAP-2. These results indicate that the
downregulation of Bcl-2 and the IAP family might also
activate caspase-3 and induce apoptosis in U937 cells in
response to ZD1839.
The MAPK family proteins and PI3K/Akt pathways play
critical roles in cell survival and death in many physiological
and pathological settings. It is well known that the
activation of the p38 MAPK and JNK pathways leads to the
phosphorylation of a variety of pro-apoptotic downstream
effectors, whereas the ERK and PI3K/Akt pathways are more
often associated with cell
survival[27,28]. In the present study we found that ZD1839 modulated the downregulation of ERK
and Akt for apoptotic death. In contrast to our data, another
recent study reported that combined treatment with ZD1839
and SP600125 had no effect on colony formation in bile duct
carcinoma[29]. More experiments are required in order to find
the relationship between ZD1839 and the JNK pathway in
leukemic cells. Sumitomo et al also suggested that ERK, but
not Akt, is involved in the mechanism by which ZD1839
exerts its chemosensitizing effect[30]. On the other hand, the
inhibition of ERK or PI3K has been shown in preclinical and
clinical studies to provide a potential antitumor effect in some
cancer types[11]. These disparate observations regarding
the nature of the signaling pathways represent the different
apoptotic mechanisms associated with ZD1839 in different
cellular systems. Although exceptions occur, the bulk of the
evidence indicates that constitutive activation of PI3K/Akt
signaling increases the survival threshold of cancer cells.
In conclusion, our findings may provide insight into one
of the mechanisms to improve the efficacy of anticancer drugs
and to support the idea that ZD1839 could induce apoptosis
in human leukemic U937 cells through the
EGFR-independent pathway. However, further studies are necessary in
order to investigate what kinds of molecular pathways are
related to EGFR-independent apoptosis.
Acknowledgments
We also thank Prof TK KWON for his critical comments
on the manuscript and providing Bcl-2 overexpressing U937
cells. We are also grateful to AstraZeneca for kindly
providing us with ZD1839 for experimental studies.
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