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Introduction
Ovarian cancer is the third most common gynecological cancer in women, but it has the highest mortality rate of all
gynecological cancers. Ovarian cancer is so deadly because it is infrequently diagnosed at an early stage. There are often
no symptoms in its earlier stages, and the cancer may grow for some time before it causes pressure, pain, or other problems.
However, even when symptoms appear, it is difficult to tell whether they are caused by ovarian cancer, or by some other less
threatening disease. Approximately two-thirds of
patients present with advanced-stage disease (stage III or IV) at diagnosis.
The majority of cases initially respond to standard therapies
with different anticancer drugs, including
platinum-containing compounds and taxol, but long-term survival and cure
rates are still disappointingly low because of the
development of drug resistance[1]. As a result, while women
diagnosed with localized cancer, or one that has not spread
beyond the ovary, have a 93.5% 5 year survival rate, only about
19% of all cases are detected at this
stage[2]. When the cancer is already regional or distant by the time it is first
diagnosed, the 5 year survival rate drops to 68.8% and
28.5%, respectively[3]. Ovarian cancer remains a difficult cancer
to treat in that the mortality rate has not
improved in the past few decades. Therefore, while it is
challenging, finding a new drug capable of killing these
resistant ovarian cancer lines would drastically reduce the
disease's mortality rate.
Artemisinin (Qinghaosu), the active constituent of the
herb Artemisia annua (sweet wormwood), has been used as
early as the Han Dynasty in ancient China to treat malaria
and is still widely used as an effective antimalaria drug in
clinics. More recently, some laboratory studies have
suggested that artemisinin and its derivatives have anticancer
properties, such as in breast cancer and oral squamous cell
carcinomas[4_22], although the underlying mechanisms is not
clearly known[11,23,24]. In addition, artemisinin and its
derivatives can increase sensitivity of human glioma cells to
ionizing radiation[25] and inhibit tumor
angiogenesis[5,26]. In the present studies, we examined the anticancer activity of
dihydroartemisinin (DHA), an artemisinin derivative, in
human ovarian cancer cells and demonstrated that DHA is a
new potent drug for the inhibition of ovarian cancer cell
growth via the induction of apoptosis (programmed cell death)
and blocking of the cell cycle progression. These studies
will provide important preclinical information and a
mechanism for the clinical application of DHA in ovarian cancer
therapy in the near future.
Materials and methods
Cell culture and DHA Ten ovarian cancer cell lines, as
listed in Table 1, were purchased from American Type
Culture Collection and maintained in Dulbecco's modified Eagle
medium ( DMEM, Invitrogen, Carlsbad, CA, USA)
supplemented with 10% fetal bovine serum, 100 unit/mL penicillin,
100 µg/mL streptomycin, and a mixture of nonessential amino
acids (Sigma-Aldrich, St Louis, MO, USA). Seven normal
human ovarian surface epithelial (HOSE) cell lines, as listed
in Table 1, were cultured in a mixture of medium 199 and
MMCDB105 medium supplemented with 10% fetal bovine
serum. Yellow DHA was purchased from Sigma-Aldrich
(USA) and dissolved in sterile double distilled water (DD
water) with a stock solution of 20 mmol/L and stored at
-20 oC. DHA was directly diluted in medium to appropriate
concentrations just before the experiments. Artemisinin,
artesunate, arteether, artemether, and arteannuin were kindly
provided by the Laboratory of Molecular Pharmacology,
Division of Basic Science, National Cancer Institute, National
Institutes of Health, Bethesda, Maryland, USA.
Transfection of siRNA siRNA for human Bcl-2 and Bax
were designed and synthesized by Ambion (Austin, TX,
USA). The specific sequences of these siRNA were as
follows: Bcl-2, 5'-CAGGACCUCGCCGCUGCAGACC-3'; Bax,
5'-GACGAACUGGACAGUAA-CA-3'; and luciferase, 5'-CGUACGCGGAAUACUUCGA-3' (as a control).
Transfections were performed at approximately 70%
confluency in 6-well plates using oligofectamine (Invitrogen, USA)
according to the manufacturer's instructions. Briefly,
2.5×105 cells were seeded in complete growth
medium the day before transfection. For each transfection
reaction, siRNA-oligo-fectamine complexes were prepared by mixing
oligonucleotides with oligofectamine diluted at 1:5
(v/v) with medium directly
before addition to the cells. Final concentrations of
the siRNA were between 1 and 100 nmol/L. Transfections
were performed in 1 mL serum-free medium for 4 h. Thereafter,
0.5 mL medium containing 30% Fetal Calf Serum (FCS,
v/v) was added to achieve complete
growth conditions (10% FCS). In each experiment, control cells receiving only
oligofecta-mine without siRNA and luciferase siRNA-transfected cells
(as a negative control), were included. Cells were assayed
48 h after transfection.
Cell viability analysis Cell viability was performed by a
MTT (3-[4,5-methylthiazol-2-yl]-2,5-diphenyl-tetrazolium
bromide) viability assay, the most commonly used assay to
determine cell growth and death. The MTT survival assay is
described in detail in previous
studies[27]. Briefly, exponentially growing cells were recultured (5000 cells/well)
overnight in 96-well tissue culture plates. Different doses of
DHA diluted in medium were added to the respective wells,
and the cells were then postincubated for either 24 or 48 h.
20 µL MTT (Sigma-Aldrich, USA) was directly added to the
media in each well with a final concentration of 2
mg/mL. After 4 h incubation, the medium containing MTT was
discarded and 120 µL DMSO (Sigma-Aldrich, USA) was added
for 10 min. Absorbance was measured in an ELISA reader at
570 nm, with the absorbance at 690 nm to correct for
back-ground, and viability was expressed as the percentage of
untreated controls. As an alternative method to analyze cell
viability, trypan blue exclusion assays were performed after
adherent and non-adherent cells were harvested. To
validate the MTT assay as a measure of cell viability, trypan
blue exclusion was performed in some experiments in parallel.
All experiments were performed at least 3 times.
Flow cytometry assay Cell cycle analysis was performed
by flow cytometry. After the cells were untreated and treated
with DHA, the medium was collected into centrifuge tubes.
The removed cells by trypsin were poured into the same
tubes. The cells were centrifuged for 5 min at
1800×g. The supernatant were poured out, washed once with
1×phos-phate-buffered saline (PBS), and centrifuged for 5 min again.
The cells were finally fixed by 5 mL of precooled 70% ethanol
for at least 4 h. The fixed cells were centrifuged and washed
with 1×PBS. After centrifugation, the cell pellets were
resuspended in 500 µL propidium iodine (10 µg/mL) containing
300 µg/mL RNase (Sigma-Aldrich, USA). Then the cells were
incubated on ice for 30 min and filtered with a 53 µm nylon
mesh. Cell cycle distribution was calculated from
10 000 cells with ModFit LT software (Becton Dickinson, CA, USA)
using FACScaliber (Becton Dickinson, San Jose, CA, USA).
DNA fragmentation gel electrophoresis Apoptosis
induction was determined by a DNA fragmentation gel
electrophoresis, as described in previous
studies[28]. The cells untreated and treated with DHA were collected by
trypsin and centrifuged for 5 min at
13000×g. The cells were washed with 1×PBS and centrifuged again. The cell pellets
were then lysated with a 200 µL lysis buffer (10 mmol/L
Tris-HCl at pH 7.6, 100 mmol/L EDTA, 20 mmol/L NaCl) and
centrifuged. The supernatant was transferred into new tubes.
20 µL SDS plus 200 µL RNase A (10 mg/mL, Sigma-Aldrich,
USA) was added to each tube. After 2 h incubation at 56 °C,
30 µL proteinase K (50 mg/mL, Sigma-Aldrich, USA) was
added to each tube. The cell lysates were incubated at 37 °C
for another 2 h. The DNA was finally precipitated with the
addition of 10 µL of 10 mol/L potassium acetate and 1 mL
100% ethanol at -80 °C for 30 min. The extracted DNA samples
were centrifuged and washed with 70% ethanol. Pure DNA
were finally loaded and run on a 1% agarose gel at 80 V in
running buffer (89 mmol/L Tris-acetate, 2 mmol/L
Na2EDTA, and 89 mmol/L boric acid),
stained with ethidium bromide, and photographed. The samples were run in
tandem with a DNA molecular weight ladder (Life Technologies,
Gaithers-burg, MD, USA) providing molecular size markers of 0.5_12
kilobase pairs. Gel photographs were evaluated for typical
ladder patterns of low molecular weight DNA fragments in
multiples of 180_200 base pairs, a hallmark of apoptosis.
Determination of apoptosis by terminal deoxynucleotidyl
transferase-mediated nick end labeling assay
Terminal deoxynucleotidyl transferase-mediated nick end
labeling (TUNEL) was performed using the
Apo-BrdUTM apoptosis detection kit (PharMingen,
San Diego, CA, USA) to quantify induction of apoptosis following the
manufacturer's instructions. Briefly, the untreated or treated cells were fixed
in 1% (w/v) paraformaldehyde
in PBS and incubated on ice for 15 min. Then the cells
were washed 3 times with PBS and preserved in 70%
(v/v) ethanol overnight. About
1×106 cells/treatment in duplicate, along with
positive and negative controls, were counted, pelletized, washed
twice with wash buffer, and subjected to labeling reaction using
terminal deoxynucleotidyl transferase overnight at RT. At the end
of the reaction, the cells were rinsed twice before treatment
with fluorescein-labeled anti-BrdU antibody solution in the
dark for 30 min at RT. The cells were stained with propidium
iodide/RNase solution for 30 min in the dark and analyzed by
flow cytometry (Epics XL-MCL, Beckman Coulter, Miami,
FL, USA).
Western blot assay The protein expression was detected
by using Western blot assays, as previously
described[27,28]. The cells untreated and treated with DHA were harvested by
trypsin and centrifugation. The cells were washed twice
with ice-cold PBS and lysed with a protein lysis buffer
containing Tris-HCl (50 mol/L, pH 7.4), Nonidet P-40 lysis buffer
(1%), Na-deoxycholate (0.25%), NaCl (150 mmol/L), EDTA (1
mmol/L), phenylmethanesulfonyl fluoride (1 mmol/L),
Na3VO4 (1 mmol/L), NaF (1 mmol/L), and an inhibitor cocktail
(Sigma-Aldrich, USA). After centrifugation, the supernatant was
transferred into new tubes. The protein concentration was
determined by the Bio-Rad protein assay (Bio-Rad, USA).
50 µg total whole cell lysates were separated on SDS-PAGE
and electroblotted onto nitrocellulose membranes (Millipore,
Billerica, MA, USA). The membranes were then incubated
in blocking solution [5% non-fat milk in 20 mmol/L Tris-HCl,
150 mmol/L NaCl, 0.1% Tween-20 (TBS-T)], followed by
overnight incubation with the appropriate primary antibodies
(Santa Cruz Biotechnology, Santa Cruz, CA, USA). After
being completely washed with TBS-T, the membranes were
incubated with horseradish peroxidase-conjugated
secondary antibodies (Santa Cruz Biotechnology, USA) for 1 h.
Finally, immunocomplexes were developed with an enhanced
horseradish peroxidase/luminol chemiluminescence reagent
(Sigma-Aldrich, USA) according to the
manufacturer's instruction. Mouse monoclonal antibodies against Bax (2D2,
1:500 dilution), Bad (C-7, 1:1000 dilution), Bcl-2 (100, 1:2000
dilution), Bcl-xL (H-5, 1:1000 dilution), and a goat antibody
against α-actin (I-19, 1:2000 dilution) were purchased from
Santa Cruz Biotechnology (USA). Each protein band was
then quantitatively analyzed using Imagemaster (Amersham,
Pittsburg, PA, USA) and Software Image Analyzer
(Amer-sham, Pittsburg, PA, USA).
RT-PCR assay The relative changes of mRNA transcripts
were performed by semi-quantitative RT-PCR assays, as
describe in previous studies[27]. Briefly,
1.5×106 cells untreated and treated with DHA were lysated with 1 mL TRIZOL
method (according to Invitrogen specifications). The
concentration of RNA was determined by absorbance at 260 nm,
and its integrity was confirmed by means of electrophoresis
on 1% agarose gels, and then stained with 0.1 mg/L ethidium
bromide (EB). A total of 1 µg RNA was converted to cDNA
using 15 U reverse transcriptase in 20 µL buffer, that
contained 1 mmol/L deoxynucleoside triphosphate (dNTP), 1
U RNase inhibitor, and an 0.5 µg oligo (deoxythymidine) primer.
An aliquot (5%) of cDNA was amplified using the following
primers:
Bcl-2:
5'-CTC AGT CAT CCA CAG GGC GA-3' (forward) 540 bp
5'-AGA GGG GCT ACG AGT GGG AT-3' (reverse)
Bax:
5'-ACA AAG ATG GTC ACG GTC TGC C-3' (forward) 429 bp
5'-GGT TTC ATC CAG GAT CGA GAC GG-3' (reverse)
Bad:
5'-GTT TGA GCC GAG TGA GCA GG-3' (forward) 315 bp
5'-ATA GCG CTG TGC TGC CCA GA-3' (reverse)
Bcl-xL:
5'-CAG TGA GTG AGC AGG TGT TTT GG-3' (forward) 264 bp
5'-GTT CCA CAA AAG TAT CCC AGC CG-3' (reverse) GAPDH:
5'-CAA CTA CAT GGT CTA CAT GTT CC-3' (forward) 724 bp
5'-CAA CCT GGT CCT CAG TGT AG-3' (reverse)
The PCR conditions were as follows: predenaturation at
94 ºC for 5 min, denaturation at 94 ºC for 30 s, annealing at
60 ºC (Bcl-2 and β-actin) or 64 ºC
(Bcl-xL, Bax and Bad) for
40 s, and polymerization at 72 ºC for 40 s with
Taq DNA polymerase. After 28 cycles, PCR products, together with
the β-actin PCR product of the same template, were
separated by electrophoresis and revealessd by EB staining. Then
each band of the electrophoresis gel was semiquantitatively
analyzed using Imagemaster and Software Image Analyzer.
Intensities of the mRNA levels were
normalized to those of the β-actin products as ratios to
produce arbitrary units of relative abundance.
Statistics Assays were performed in at least triplicate
for each sample exposure, and an average value was
deter-mined. Results were expressed as mean±SD. Statistical
analysis was performed using ANOVA and
Student-Newman-Keuls comparison for parametric data sets.
P-values of <0.05 were considered significant.
Results
Artemisinin and its derivatives inhibits growth of
human ovarian cancer cells We first compared the effects of
artemisinin and its 5 derivatives, artesunate, arteether,
artemether, DHA, and arteannuin (10 µmol/L, 24 h exposure),
on cell survival of 2 ovarian cancer cell lines, Human ovarian
adenoma cells SK-OV-3 and OVCAR-432, by using MTT
viability assays. As shown in Figure 1, 24 or 48 h exposure to
20 µmol/L artemisinin and its derivatives caused a loss of
cell viability to different extents. Similar results were
observed with trypan blue dye exclusion assays (data not
shown) and in other human ovarian cancer cell lines (data
not shown). It is notable that among them, DHA was the
most effective and potent in inhibiting cell growth of ovarian
cancer cells following 24 or 48 h exposure with a single dose
of 20 µmol/L. For example, 24 h treatment with 20 µmol/L
DHA reduced cell viability to approximately 50%, while 48 h
exposure caused about a 90% loss of cell viability compared
to the untreated control. Thus, DHA was used for
subsequent studies. These findings indicate that artemisinin and
it's derivatives examined can indeed be used as a new class
of compounds with possible therapeutic potential in ovarian
cancer.
DHA inhibits cell growth in a dose- and time-dependent
manner To further investigate the antitumor activity of DHA
in ovarian cancer cells, we next examined the DHA inhibition
of cell growth in 7 normal human ovarian epithelial cell lines
and 10 human ovarian cancer cell lines. The cells were
exposed to different doses of DHA for 48 h and then subjected
to MTT assays. In order to easily see the different effects of
DHA in a panel of the cell lines examined, Inhibitory
Concentration 50% (IC50)or Inhibitory Concentration 90%
(IC90), a DHA dose required to cause 50% or 90% of cell viability
loss, in each cell line were calculated. As summarized in
Table 1 and Figure 2, although DHA showed an inhibitive
activity of cell growth in all examined cell lines, normal
ovarian epithelial cell lines were much more resistant to DHA
treatment than ovarian cancer cell lines. Among them,
ovarian cancer cell line HEY is the most sensitive to DHA, while
normal ovarian cell line HOSE642 was the most resistant.
For example, 5.51 and 11.13 µmol/L DHA were respectively
required for IC50 and IC90 in ovarian cancer cell line HEY
compared to 54.9 µmol/L and 99.8 µmol/L, for
IC50 and IC90 in normal ovarian cell line HOSE642 (a 10-fold difference,
P<0.005). Typical survival curves in 2 ovarian cancer cell
lines, OVCA-432 and SK-OV-3, following 24 or 48 hr
exposure to DHA, are shown in Figure 3. Moreover, a significant
morphology change in DHA-treated cells was able to be seen
in a dose-dependent manner (Figure 4).
It was noted that among the human ovarian cancer cells
examined, the cell lines having wild-type (functional)
p53 gene status were more sensitive to DHA compared to the
cell lines containing mutations of p53 or null p53 status. For
instance, 3.83 and 10.13 µmol/L were required for
IC50 and IC90 in wild-type p53 cell line OVCA-439, while the
IC50 and IC90 were 15.2 µmol/L and 23.15 µmol/L in mutant p53 cell line
ALST. The means of IC50 and
IC90 in the 5 cell lines with wild-type p53 respectively were 5.04±0.52 and 11.52±0.4, but the
mean of IC50 and IC90 in the 5 cell lines with mutant p53 were
14.5±0.39 and 22.36±0.49 (about 2_3-fold,
P<0.01). It is obvious that p53 mediates cell susceptibility of ovarian cancer to
DHA treatment. In other words, p53 gene status is an
important determinant of DHA chemosensitivity and p53
mutations are often associated with decreased sensitivity to DHA
in ovarian cancer.
DHA induces apoptosis and cell cycle arrest
We investigated whether DHA, like many anticancer drugs in ovarian
cancer therapy, induced apoptosis. Four ovarian cancer cell
lines, OVCA-420, OVCA-432, SK-OV-3, and OVCAR-3, were
untreated and treated with 10 µmol/L DHA for 48 h, and the
cells were collected for DNA fragmentation determination
by using DNA fragmentation gel electrophoresis assay. As
shown in Figure 5, DHA treatment induced a significant
apoptosis (DNA ladders) in all 4 cell lines containing either
wild-type or mutant p53. Similar results with apoptosis
induction were observed in other cell lines treated with DHA,
including cell lines containing p53 mutation (data not shown).
To further determine the apoptosis induction, flow
cytometry assay was performed in OVCA-420 cells treated
with DHA at various doses (0_15 µmol/L) for 48 h as
described in Materials and methods. Apoptotic cells
contain less than 2 N DNA content and appear as a
sub-G1 cell population. As shown in Figure 6, a significant
accumulation of sub-G1 cells was detected in the cells treated with
increased doses of DHA, which was consistent with our
apoptosis data using DNA fragmentation gel
electrophoresis assays (Figure 5). For example, only 10% of
sub-G1 cells were shown with 2.5 µmol/L DHA, however, 47% of
sub-G1 cells were seen following treatment with 15 µmol/L DHA.
DHA also caused an arrest of cell cycle progression. DHA
treatment resulted in a gradual G1 and
G2/M arrest with increased doses, that is, an increase of both
G1 and G2 peaks (Figure 6). For example, 56% of cells were arrested at the
G2/M phase following 15 µmol/L exposure of DHA, while only 26%
of cells were at the G2/M phase after treatment with 2.5
µmol/L DHA. In contrast, the population of the
G0/G1 phase decreased with increasing doses of DHA. For example, 32% of
cells were arrested at the
G0/G1 phase following 15 µmol/L
exposure of DHA, while 67% of cells were at the
G0/G1 phase after treatment with 2.5 µmol/L DHA. Similar results were
obtained with other human ovarian cancer cell lines (data
not shown). These findings indicate that cell cycle arrest
and apoptosis induction may be important mechanisms for
antitumor cell growth activity of DHA in ovarian cancer.
DHA targets the Bcl-2 family for apoptosis induction
To study the possible contribution of the
Bcl-2 gene family to DHA-induced apoptosis, the effects of DHA on several
members of the Bcl-2 gene family were examined, including 2
apoptotic inhibitors Bcl-2 and Bcl-xL and 2 apoptotic
promoters Bax and Bad. OVCA-420 and OVCA-432 cells were
exposed to 24 or 48 h of 10 µmol/L DHA exposure, collected,
and subjected to protein expression analysis by Western
blot assays and mRNA expression analysis by RT-PCR
assays. As shown in Figure 7, an increase of 2 promoter
proteins, Bax and Bad, was detectable in both OVCA-420
and OVCA-432 cells after DHA treatment. However, DHA
significantly reduced 2 antiapoptotic proteins, Bcl-2 and
Bcl-xL, in both cell lines. Moreover, the alterations of these
proteins were time-dependent. For example, a 54% decrease
of Bcl-xL protein expression appeared in OVCA-420 cells
following 24 h exposure to DHA, while a 27% decrease of
Bcl-xL protein expression occurred in response to 48 h
treatment of DHA.
The alterations of mRNA expression in OVCA-420 cells
treated with DHA are shown in Figure 8, as determined with
RT-PCR assays. 2.5 µmol/L DHA caused a significant
decrease of Bcl-xL and Bcl-2 mRNA. Moreover, both
Bcl-xL and Bcl-2 mRNA became undetectable after 5_7.5 µmol/L
treatment (Figure 8A). Both Bcl-2 and
Bcl-xL mRNA expression started to decline 2 h after DHA addition to the medium and
became undetectable 16_24 h following 10 µmol/L DHA
treatment (Figure 8B). Both Bax and Bad mRNA expressions were
elevated in response to higher doses of DHA (Figure 8A)
and later time points following DHA exposure (Figure 8B).
These results with protein expression results indicate that
DHA regulates members of the Bcl-2 gene family at both the
transcription and post-transcription levels.
To further determine the role of the Bcl-2 family in DHA
antitumor cancer activity, Bcl-2 siRNA and Bax siRNA were
employed. RNAi is the process where the introduction of
double stranded RNA into a cell inhibits gene expression in
a sequence-dependent fashion. RNAi is usually described
as a post-transcriptional, gene-silencing mechanism in which
dsRNA triggers the degradation of homologous messenger
RNA. The mediators of RNAi are 21- and 23-nucleotide
siRNA that bind to a ribonuclease complex called
RNA-induced silencing complex (RISC), which guides the small
dsRNA to its homologous mRNA target. Consequently, RISC
cuts the mRNA approximately in the middle of the region
paired with the antisense siRNA, after which the mRNA is
further degraded. As shown in Figure 9A, transfection of
Bcl-2 siRNA or Bax siRNA significantly reduced the Bcl-2 or
Bax protein level. The cells transfected with Bcl-2 siRNA
exhibited a significantly increased sensitivity to the
DHA-induced apoptosis compared to the untransfected control
cells (Figure 9B), that is, more apoptotic cells were observed
in the Bcl-2 siRNA-transfected cells than in the control cells
(49% vs 28%, P<0.01). In contrast, the decreased
sensitivity obtained in the cells transfected with Bax siRNA in response
to DHA caused apoptosis in comparison with the control
cells, that is, 8% of apoptotic cells vs 28% of apoptotic cells
(P<0.01). However, transfection of luciferase siRNA did not
produce any effect on DHA-triggered apoptosis. These
results suggest that the Bcl-2 gene family, at least Bcl-2 and
Bax, is an important target in DHA induction of apoptosis in
ovarian cancer cells.
Discussion
Although it has been reported that artemisinin inhibits
cell proliferation in breast cancer and other types of
cancers[4_22], it is still not known whether artemisinin and its
derivatives work as growth inhibitors in ovarian cancer cells.
Our results in this study suggest that artemisinin and its 5
derivatives examined, including artesunate, arteether,
artemether, arteannuin, and DHA, differentially inhibit cell
growth in human ovarian cancer. DHA is the most effective
one in the inhibition of ovarian cancer cell division among
them. Moreover, DHA inhibits growth of human ovarian
cancer cells by doses at micromolar levels (Table 1 and
Figures 1_3), but these doses of DHA exhibit little or no
significant cytotoxicity in normal ovarian cells (Table 1).
Another important result from this study is that p53 gene
status may be an important regulator in ovarian cancer cell
susceptibility to DHA. Ovarian cancer cells with wild-type
(functional) p53 gene status exhibit a higher sensitivity to
DHA than ovarian cancer cells with mutant or null
p53 gene status (Table 1 and Figure 2). It has been well known that the
tumor suppressor protein p53 plays a central role in the
regulation of cell cycle arrest and cell
death[29] and also plays an important role in modulating radiosensitivity and
chemosensitivity of tumor cells, including ovarian
cancer[30_32]. Thus, the different response to DRAT in ovarian cancer cells with
different p53 gene status will be valuable in determining the
actual application of DHA to ovarian cancer patients in
clinics, since the p53 gene is the most frequently mutated
gene in human cancers (including ovarian cancer),
specifically at later stages for invasive
cancers[33,34]. The role of p53 in DHA sensitivity of ovarian cancer cells will be needed
for further study.
Cell death has been shown to occur by 2 major
mechanisms, necrosis and apoptosis (a programmed cell death). Classical
necrotic cell death occurs due to noxious injury or trauma,
while apoptosis takes place during normal cell development,
regulating cellular differentiation and number. While
necrotic cell death results in cell lysis, cellular apoptosis is
characterized morphologically by cell shrinkage, nuclear
pyknosis, chromatin condensation, and blebbing of the
plasma membrane. It seems that all known anticancer drugs
kill cancer cells predominantly through
apoptosis[35,36]. A cascade of molecular and biochemical events has been
identified, including the activation of an endogenous
endonuclease that cleaves DNA into oligonucleosomes
detectable as a ladder of DNA fragments in agarose gels. Typically,
the DNA of apoptotic cells is cleaved into a population of
fragments composed of multimers that are 180_200 bp in
length. These multimeric fragments are readily observed on
agarose gels as a "ladder", thus, observation of
oligonucleo-somal DNA fragments by DNA laddering has long been the
most acceptable assay used for the detection of apoptosis.
DHA triggers significant "DNA fragments" as shown by
agarose gels (Figure 5) and induces the
sub-G1 phase as determined by flow cytometry (Figure 6). DHA also cause
dose-dependent G2 arrest, although the involved mechanisms
is unclear. Thus, cell cycle arrest and apoptosis might be
responsible, at least in part, to DHA inhibition of cell growth
in ovarian cancer as a working model presented in Figure 9.
It is now well established that members of the Bcl-2
family are the most prominent regulators of apoptosis in a
variety of cell types, including cancer
cells[37_39]. A large number of antitumor drugs cause apoptosis via the regulation of the
Bcl-2 gene family[40]. The Bcl-2 family consists of a growing
number of proteins, which contain 4 conserved Bcl-2
homology domains (Bcl-2 Homology (BH)1, BH2, BH3, and BH4),
together with a transmembrane domain, all being identified
as crucial for the regulation of apoptosis. Based on
functional studies and the retention of BH
domains, the Bcl-2 family can be divided into 3
subgroups[39]. The Bcl-2 subgroup includes all anti-apoptotic proteins, such as Bcl-2 and
Bcl-xL. The Bax subgroup consists
of pro-apoptosis members, such as Bax and Bad. Both
groups contain more than 1 BH domain. The third subgroup contains only BH3 proteins,
such as Bid and Bim, which can interact with
either anti-apoptotic proteins or proapoptosis members and promote
apoptosis. Our observations have demonstrated that
during the induction of apoptosis, DHA downregulates Bcl-2
and Bcl-xL and upregulates Bax and Bad at both the mRNA
and protein levels (Figures 7 and 8). Furthermore, as expected,
silencing Bcl-2 or Bax increases or reduces cell sensitivity to
DHA-induced apoptosis (Figure 9). Therefore, these results
with gain of function or a loss of function demonstrate that
targeting the Bcl-2 family-mediated apoptosis components
is an important mechanism for DHA antitumor activity in
ovarian cancer cells. Identification of mechanisms that
regulate DHA-induced apoptosis might provide a new target for
further therapeutic intervention of DHA to therapy of
ovarian cancer.
In summary, these results show for the first time that
DHA inhibits cell growth via inhibiting cell cycle
progression and inducing apoptosis in ovarian cancer(Figure 10)
and provides evidence of potential implications for the
rational application of DHA as a novel anticancer drug against
ovarian cancer, although lots of work need to be done
before DHA will really be utilized in clinical treatments of
ovarian cancer.
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