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Introduction
Nitrofen (2, 4-dichloro-40-nitrodiphenyl ether) is a selective contact herbicide used on a variety of food crops for pre- and
post-emergence control of annual grasses and
weeds[1]. It is a member of the chlorophenoxy class of
herbicides and is also called nitrophene, TOK, TOK E-25, and
Nip[1]. Nitrofen is a potent teratogen in rats that produces abnormal development of the
heart, kidneys, diaphragm, and lung when administered during
organogenesis[2]. It has been demonstrated in pregnant
rodents that nitrofen induces congenital diaphragmatic hernia
(CDH)[3], including pulmonary
hypoplasia[4] with abnormal pulmonary arterioles and biochemical lung
immaturity[5] similar to those observed in human CDH. The reliable induction of
CDH and pulmonary hypoplasia-immaturity in fetal rodents exposed to nitrofen has made this teratogenic model the main tool
for understanding the pathogenesis of CDH.
Previous evidence indicates that the lung is malformed
independently, and perhaps is the primary cause of
diaphragmatic anomalies in CDH[6,7]. It has been clearly shown that
nitrofen on its own can, to varying degrees, affect aspects of
lung development[8]. Nitrofen levels in the embryo are likely
persist for several days after administration, allowing
multiple stages of lung embryogenesis to be targeted by this
teratogen throughout the final third of rat
gestation[8]. Recent data have suggested that nitrofen may interfere with
retinoid signaling during lung development, as measured
with transgenic mice containing the Lac Z reporter gene linked
to a retinoic acid response element[9]. Inadequate retinoic
acid signaling resulted in incomplete differentiation cells and
ultimately cell death[9]. However, the mechanisms of
nitrofen-mediated pulmonary hypoplasia are still incompletely
understood.
In developing and adult lungs, type II pneumocytes are
not only the source of alveolar surfactant, but also the
progenitor cells for alveolar
epithelium[10]. Type II pneumocytes
produce and secrete pulmonary surfactant, which is critical
for effective gas exchange[10]. Type II pneumocytes also
proliferate and differentiate into type I pneumocytes to
restore the alveolar epithelium after lung injury and participate
in the innate immune response to exogenous materials and
organisms[10]. Li et al demonstrated that in the lungs of
nitrofen-induced CDH, type II pneumocytes were filled with
only a few lamellar bodies and become metabolically
inactive[11]. The sparse type II pneumocytes also showed
cytoplasmic degenerative changes, such as vacuolization of the
cytoplasm[11]. In cultured type II
pneumocytes, nitrofen decreased synthesis of surfactant components like surfactant
protein B, and downregulated thyroid transcription factor-1,
a homeotic protein that acts as a transcription factor during
pulmonary morphogenesis[12]. These findings show that
nitrofen-induced pulmonary hypoplasia is exerted, at least
in part, by direct action on type II
pneumocytes[12]. However, it still remains largely unknown whether nitrofen exerts
effects on proliferation and apoptosis of type II pneumocytes.
In the present study, we demonstrated that nitrofen
suppressed proliferation of cultured A549 cells accompanied by
the downregulation of proliferating cell nuclear antigen
(PCNA), and induced mitochondria-mediated apoptosis
involving the activation of the p38 mitogen-activated protein
kinase (p38-MAPK) signaling pathway.
Materials and methods
Cell culture and nitrofen treatment A549, a type II
pneumocyte cell line with phenotypic features including
surfactant protein synthesis[13], was obtained from ATCC
(CCL-185, Manassas, VA, USA) and maintained in Dulbecco's minimal Eagle's medium (DMEM, Life
Technologies Inc, Gaithersburg, MD, USA)
supplemented with 10% fetal bovine serum (FBS, Life Technologies, USA), penicillin
(100 U/mL), and streptomycin (100 µg/mL). The cells were
maintained at 37 °C in a humidified atmosphere of 5%
CO2. Confluent monolayers of cells were starved with a medium
supplemented with 0.1% FBS and 2 mmol/L L-glutamine.
Twenty-four hours later, the cells were incubated in
serum-free DMEM for 4 h and pretreated with
Z-Val-Ala-Asp(OCH3)-fluoromethylketone (zVAD-fmk, Biomol, Plymouth Meeting,
PA, USA) or SB203580 (Calbiochem, La Jolla, CA, USA) for
1h, then stimulated with different concentrations of
2,4-dichloro-40-nitrodiphenyl ether (nitrofen, Sigma, St Louis,
MO, USA) or DMSO as indicated.
Measurement of cell viability For direct cell counting,
1×104 A549 cells were seeded in 24-well plates, and treated
with nitrofen or DMSO as indicated. At the end of the
incubation period, the cells were washed with
phosphate-buffered saline (PBS), trypsinized, and counted using a Casy
1-model TT cell counter (Schaefer System GmbH, Reutlingen,
Germany). Cell proliferation was monitored by the
3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT,
Sigma, USA) colorimetric assay[14]. Briefly, 20 µL MTT (5
mg/mL) was added to each well. After 4 h incubation at
37 ºC, the cell supernatant was discarded, the MTT crystals
were dissolved with DMSO, and the absorbance was
measured at 570 nm. Percent viability was defined as the relative
absorbance of treated versus untreated control cells.
Cellular DNA synthesis was determined by
[3H]-thymidine incorporation assay. Briefly, the cells
(2×105/well) were grown on 24-well plates. A pulse of 1 µCi/mL
[3H]-thymidine (DuPont, Boston, MA, USA; sp act 80_90 Ci/mmol) was added to the
cells for 1 h. DNA synthesis was determined as
trichloroacetic acid (TPA)-precipitable incorporation of
[3H]-thymidine as described by
Sheffield[15]. All experiments were done with
6_8 wells per experiment and repeated at least 3 times.
Colony formation assay The cells were seeded at a
density of 500 mL on 35 mm dishes[16]. After an overnight
incubation to allow cell attachment, the cells were incubated with
nitrofen or DMSO as indicated. After being incubated for 24
h, the medium was replaced with fresh medium containing
10% FBS. The colonies were allowed to grow for 10_14 d.
The medium was discarded and each well was washed twice
with PBS carefully. The cells were fixed in methanol for 15
min and then stained with crystal violet for 20 min. Finally,
positive colony formation (more than 50 cells/colony) was
counted. The survival fraction for the cells was expressed
as the ratio of plating efficiency of treated cells to that of
untreated control cells.
Cell cycle assay According to the
literature[17], cell cycles in untreated, DMSO- or nitrofen-treated cells were examined
by flow cytometry. Briefly, 2×105
cells were collected, washed twice with PBS, and fixed in 70% ethanol overnight at 4
°C. Then the cells were washed once with PBS, digested with
200 µL RNase (1 mg/mL) at 37 °C for 30 min, and stained with
800 µL propidium iodide (PI, 50 µg/mL, Sigma, USA) at room
temperature for 30 min. The DNA histograms were assayed
with a flow cytometer (Becton-Dickinson , San Jose, CA,
USA), using the CELLQUEST software (Becton-Dickinson,
USA).
Cellular morphological observation To observe the
changes in cellular morphology, the in situ terminal
deoxynucleotidyl transferase-mediated dUTP
nick-end-labeling (TUNEL, Roche, Indianapolis, IN, USA) method was
performed according to the manufacturer's instructions.
Briefly, the cells were fixed immediately in 4%
paraformaldehyde for 20 min, washed with PBS, and permeabilized with
0.1% Triton X-100 in 0.1% sodium citrate. Each of the sample
slides received 50 µL TUNEL reaction mixture and was
incubated for 60 min at 37 ºC. After washing with PBS, the
sections were analyzed under a Leitz fluorescent microscope
(Leitz, Wetzlar, Germany). Negative control slides were
performed without the TUNEL mixture. The TUNEL
positively-stained cells were counted in 10 randomly selected
high-power (×200) fields. The rate of the positively
stained cells was determined by calculating the average percentage.
Morphological evidence of apoptosis was further
obtained using acridine orange (AO, Sigma, USA) and
ethidium bromide (EB, Sigma, USA)
staining[18]. Briefly, the cells were harvested with 0.125% trypsin and 0.01% EDTA,
resuspended in 95 µL DMEM medium, and incubated with 5
µL AO/EB staining solution (100 mg/L PBS of each dye) at
room temperature for 15 min. The cells were examined using
fluorescence microscopy and photographed (Olympus,
Tokyo, Japan). Viable cells were colored green with intact
nuclei. Nonviable cells had bright orange chromatin.
Apoptosis was demonstrated by the appearance of cell
shrinkage with condensation and fragmentation
of the nuclei. Necrotic cells appeared orange with a normal
nuclear structure. The numbers of viable cells with normal nuclei
(VN), viable cells with apoptotic nuclei (VA), nonviable cells
with apoptotic nuclei(NVA), and nonviable cells with normal
nuclei cells (NVN) were determined by counting 5 randomly
selected high-power (×200) fields. Apoptosis rates were
calculated as (%)=(VA+ NVA)/(VN+ NVN+VA+NVA)×100%.
Apoptosis rate detection The apoptotic ratios of cells
were determined by annexin V- fluorescein isothiocyanate
(FITC) (BD Pharmingen, San Diego, CA, USA) and PI
staining flow cytometry[18]. Briefly, the cells from the above
groups were collected, washed twice with cold PBS,
resuspended with 100 µL binding buffer [10 mmol/L
4-(2-Hydroxy-ethyl)-1-piperazineethanesulfonic acid , 140 mmol/L
NaCl, and 2.5 mmol/L CaCl2, pH 7.4] into
2×105_5×105 cells/mL density,
and incubated with annexin V-FITC at room temperature for
10 min. After washing with the binding buffer, the cells were
resuspended in 400 µL binding buffer containing 10 µL PI
(20 µg/mL, Sigma, USA), and incubated on ice for 15 min.
Apoptosis was analyzed by flow cytometry (Becton-Dickinson, USA) at a wavelength of 488 nm. This method
can be used to distinguish between living cells (annexin
V_/PI_), early apoptotic/primary apoptotic cells (annexin
V+/PI_), late apoptotic/secondary necrotic cells (annexin
V+/PI+), and necrotic cells (annexin
V_/PI+)[19].
Measurement of mitochondrial membrane potential
(Dy m) The mitochondrial membrane potential was analyzed
using 5,5',6,6'- tetrachloro-1,1',3,3'-tetraethylbenzimidazolyl-
carbocyanine iodide (JC-1, Sigma, USA), a lipophilic cationic
fluorescence dye. JC-1 exists as a green fluorescent
monomer (emission wavelength is 527 nm) at low mitochondrial
membrane potential. Mitochondrial depolarization is
indicated by an increase in green fluorescence
(FL-1)[20]. The cells
(1×106) were incubated with 5 mg/mL JC-1 for 15 min at
room temperature in dark conditions. After centrifugation at
200×g for 5 min, the cells were washed twice with PBS at 4 °C,
resuspended in 0.5 mL PBS, and analyzed on a flow
cytometer (Becton-Dickinson, USA).
Immunofluorescence for apoptosis-inducing factor
(AIF) The cells were grown on sterile coverslips in a 6-well
plate and treated with nitrofen as indicated. The cells were
fixed with fresh 4% formaldehyde in PBS at 4 °C for 20 min
and permeabilized with pre-chilled PBS with 0.2% Triton
X-100. The cells were then incubated with a rabbit antibody
against AIF (1:100, Santa Cruz Biotechnology, Santa Cruz,
CA, USA) in 3% bovine serum albumin/PBS overnight at
4 °C followed by incubation with fluorescein
isothiocyanate-conjugated secondary antibodies. After PBS washes, nuclei
were counterstained with PI. Images were captured on a
fluorescent microscope.
Real-time RT-PCR for PCNA Total RNA was isolated
with a RNeasy mini kit (Qiagen, Valencia, CA, USA). The RT
reactions were conducted with a Transcriptor First Strand
cDNA Synthesis kit (Roche, USA). Real-time PCR with SYBR
Green PCR Master Mix (Applied Biosystems, Foster City,
CA, USA) was performed using the ABI Prism 7700 Sequence
Detector (Applied Biosystems, USA). The PCR primers for
PCNA were as follows: 5'-AAACTAGCTAGACTTTCCTC-3' and 5'-TCACGCCCATGGCC AGGTTG-3', amplifying a 274
bp fragment.
Western blotting for PCNA, caspase-3, Bcl-2,
Bcl-xL, Bax, Bak, AIF, and phosphorylation of the p38 kinase
The cells were collected and extracted with 1×cell lysis buffer
(Promega, Madison, WI, USA). Nuclei and mitochondria
were isolated according to reported
protocols[21]. Protein (50 µg) from each sample was subjected to 4%_20% pre-cast
polyacrylamide gel (Bio-Rad, Hercules, CA, USA)
electrophoresis and transferred to nitrocellulose membranes
(Bio-Rad, USA). For PCNA, caspase-3, Bcl-2,
Bcl-xL, Bax, Bak, and AIF (Santa Cruz Biotechnology, USA) detections, the
specific primary antibody dilution was 1:1000, 1:500, 1:500,
1:500, 1:500, 1:500, and 1:500, respectively. For the
phosphorylation assay of the p38 kinase, the membranes were probed
with the specific primary antibodies against the
phospho-specific and non-phosphorylated p38 kinase at a dilution of
1:500 (Santa Cruz Biotechnology, USA). The second
antibody used in this assay was the goat anti-rabbit
horseradish peroxidase -labeled antibody at a dilution of 1:3000
(Bio-Rad, USA). An enhanced chemiluminescence substrate kit
(Amersham, Piscataway, NJ, USA) was used for the
chemiluminescent detection of the signals with autoradiography
film (Amersham, USA).
Statistical analysis Unless otherwise stated, all data
were shown as mean±SEM. Statistical significance
(P<0.05) was determined by t-test or ANOVA followed by an
assessment of differences using SigmaStat 2.03 software (Jandel,
Erkrath, Germany).
Results
Nitrofen suppressed the proliferation of A549 cells
Precisely controlled cell proliferation and apoptosis are
prerequisites for normal development and homeostasis of lungs.
Abnormalities in these processes are emerging to explain
features of disorders, including pulmonary
hypoplasia[22]. To explore the effects of nitrofen on type II pneumocytes,
we first observed cell proliferation changes in the cultured
A549 cell line. As shown in Figure 1A, nitrofen
administration resulted in distinct decreases of cell viability in a dose-
and time-dependent manner. However, DMSO treatment led
to no significant change in cell viability of A549 cells. MTT
colorimetric assay indicated that nitrofen inhibited the
proliferation of A549 cells, which was lower than those induced
by mitomycin as the positive controls (Figure 1B). The colony
formation assay further demonstrated the cell proliferation
inhibition effects of nitrofen in cultured A549 pneumocytes
(Figure 1C). These findings indicated that nitrofen
suppressed the in vitro cell proliferation of cultured type II
pneumocytes.
Downregulation of PCNA might be associated with
nitrofen-induced decreases in the proliferation of A549 cells
PCNA is an intranuclear polypeptide maximally synthesized
during the S-phase of cell cycles and participates in cell
proliferation[23]. Since the above evidence indicated that nitrofen
suppressed cell proliferation of type II pneumocytes, we
hypothesized that nitrofen may modulate PCNA expression in
these cells. To meet this end, A549 cells were treated with
various concentrations of nitrofen for different periods. As
shown in Figure 2A, Western blotting indicated that nitrofen
downregulated the expression level of PCNA in A549 cells in
a dose- and time-dependent manner. However, DMSO
administration had no influence on the expression of PCNA
(Figure 2A). In addition, the results of real-time RT-PCR
indicated that nitrofen can induce decreases of PCNA mRNA
in A549 cells (Figure 2B). Consistent with these findings,
the DNA synthesis of nitrofen-treated A549 cells was
attenuated (Figure 2C). Moreover, nitrofen treatment reduced
the ratios of the S-phase, and induced
G0/G1 arrest in cultured A549 cells (Table 1). These findings indicated that
nitrofen-induced decreases in cell proliferation might be
associated, at least in part, with the downregulation of PCNA.
Nitrofen induced apoptosis in A549 cells in a
caspase-independent manner Previous studies demonstrated
enhanced apoptosis in the cervical somites of nitrofen-exposed
rat embryos as a mechanism for diaphragmatic
maldevelopment in CDH[24]. We hypothesized in this study that
disturbed apoptosis may participate in nitrofen-induced cell
death in cultured type II pneumocytes. As shown in Figure
3A, TUNEL indicated that treatment of A549 cells with
nitrofen resulted in an obvious DNA strand break, a
characteristic change of apoptosis. To further investigate the type
of cell death induced by nitrofen treatment, the cells were
stained with AO/EB, which allows the
identification of viable, apoptotic, and necrotic cells based on color
and appearance[18]. As shown in Figure 3A, the treatment of A549 cells with
nitrofen for 24 h resulted in nuclei-shrunk and orange-stained
cells. However, after treatment with DMSO for 24 h, there
were no such morphological changes in the A549 cells (Figure
3A). This procedure and annexin V-FITC/PI staining flow
cytometry were also used to quantify the number
of apoptotic cells induced by nitrofen treatment. As shown in Table 2
and Figure 3B, 20_80 µmol/L nitrofen exerted strong
apoptosis-inducing effects on A549 cells. Additionally,
pretreatment of A549 cells with zVAD-fmk (50 µmol/L), a
pan-caspase inhibitor[25], did not abolish nitrofen-induced
apoptosis (Table 2; Figure 3B). Furthermore, Western
blotting indicated that nitrofen administration would not
influence the expression level and cleavage of procaspase-3 within
A549 cells (Figure 3C). These findings indicated that a
caspase-independent mechanism was involved in
nitrofen-induced apoptosis.
Participation of the mitochondria-mediated pathway in
nitrofen-induced apoptosis in A549 cells To determine the
involvement of the mitochondria-mediated pathway in
nitrofen-induced apoptotic cell death, we first performed
Western blotting to measure the expression of Bcl-2,
Bcl-xL, Bax, and Bak, which are members of the Bcl-2 family proteins
critical to maintaining the integrity of the mitochondrial
membrane (Figure 4A). There was a decrease in the expression of
anti-apoptotic Bcl-xL in a dose- and time-dependent manner
(Figure 4A). The decrease was significant after 6 h, and at 24
h, expression decreased to 15.2% compared to the control
(Figure 4A). However, there was no alteration in the
expression of Bcl-2, Bax, and Bak (Figure 4A). Then we measured
changes in mitochondrial membrane potential
(Dym). As shown in Figure 4B, nitrofen treatment of A549 cells resulted
in a rapid dissipation of Dym in a time-dependent manner
with an increase in green fluorescence emission. Previous
studies demonstrated that a loss of permeability of the
mitochondrial membrane leads to the release of apoptogenic
proteins normally confined to mitochondrial intermembrane
space[21]. AIF is synthesized as a 67 kDa preprotein and
localizes in mitochondrial intermembrane
space[26]. Upon
induction of apoptosis, AIF is processed to a 57 kDa form,
translocated into the nucleus, and plays important roles in
caspase-independent apoptosis[26]. We hypothesize that AIF
may participate in nitrofen-induced apoptosis. Western
blotting and immunofluorescence studies confirmed that nitrofen
administration lead to the translocation of AIF from the
mitochondria to the nucleus in A549 cells (Figure 4C). Moreover,
pretreatment with the general caspase inhibitor zVAD-fmk
(50 µmol/L) did not prevent the AIF translocation (Figure
4C). These results indicated that the mitochondria-mediated
pathway participated in nitrofen-induced apoptosis.
Involvement of the p38-MAPK signaling pathway in
nitrofen-induced apoptosis in A549 cells Apoptosis has
been demonstrated to be associated with changes in MAPK
activity in a number of different cell
systems[27,28]. To determine whether MAPK were involved in nitrofen-induced
apoptosis, its activation was evaluated by Western blot
analysis with antibodies specific for the phosphorylated
(activated) forms of MAPK. As shown in Figure 5A, nitrofen
strongly induced the phosphorylation of p38-MAPK. The
activation of extracellular signal-regulated protein kinase
(ERK)1/2 and the c-Jun N-terminal kinase (JNK) were
examined using the same extracts, but did not show changes in
phosphorylation, suggesting that ERK-1/2 and JNK were
not affected by nitrofen (Figure 5A). To clarify whether
mitochondria-mediated apoptosis induced by nitrofen is
dependent upon p38-MAPK activation, the p38-MAPK
inhibitor SB203580 was used to treat A549 cells prior to nitrofen
exposure. As shown in Figure 5B, the incubation of cells
with SB203580 (5 µmmol/L) blocked the nitrofen-induced
phosphorylation of p38-MAPK and downregulation of
Bcl-xL, resulting in rescued nuclear translocation of AIF. In
addition, pretreatment of A549 cells with SB203580 (5
µmol/L) abolished nitrofen-induced apoptosis (Figure 5C). These
findings indicate that activated p38-MAPK is involved in
nitrofen-induced apoptosis in A549 cells.
Discussion
PCNA is a protein that acts in conjunction with DNA
polymerase delta during mitosis[29]. It also plays a central
role in DNA replication, DNA repair, and cell cycle
progression[29]. Until now, PCNA expression has been an
established operational marker for proliferating cells. In the
current study, we demonstrated that nitrofen treatment
suppressed the cell proliferation of cultured pneumocytes by
cell counting, MTT colorimetry, and colony formation assay
(Figure 1). In addition, we found that nitrofen downregulated
the transcription and expression level of PCNA in cultured
pneumocytes in a dose- and time-dependent manner (Figure
2A,B), which was consistent with the findings of Keijzer
et al that when lung explants were treated with nitrofen, PCNA
immunoreactivity was clearly reduced in the mesenchymal
component[30]. In addition, PCNA reactivity was only
observed perinuclear and in the cytoplasm of epithelial cells of
the explants exposed to nitrofen[30]. In the present study, we
also demonstrated that nitrofen treatment could induce
G0/G1 arrest and reduce S-phase cells by flow cytometry (Table
1), which was consistent with previous findings that PCNA
has several roles in progression through the S
phase[29]. Moreover, DNA synthesis of nitrofen-treated A549 cells
decreased compared to those of the controls (Figure 2C).
These results indicated that proliferation is disturbed in
cultured type II pneumocytes exposed to nitrofen, which might
be associated with the downregulation of PCNA. Impaired
regulation of cell proliferation may therefore underpin
nitrofen-induced pulmonary hypoplasia in CDH.
Apoptosis is a controlled type of cell death
characterized by cell shrinkage, membrane blebbing, and DNA
fragmentation. Apoptosis is the result of coordinated
signaling pathways, which can be triggered by a variety of
extracellular stimuli, such as radiation, cytokines, and growth
factor withdrawal[31]. We hypothesized in this study that the
disturbed apoptosis may participate in nitrofen-induced
pulmonary hypoplasia. Although a previous study examined
the possibility of nitrofen-mediated apoptosis
in type II H441 pneumocytes, it found no observable increases in
apoptosis by TUNEL assay[12]. Our present study found that
nitrofen, with high concentrations (20_80 µmol/L) rather than low
concentrations (1.5 µmol/L), can induce apoptosis of A549
cells as demonstrated not only by TUNEL, but also by
AO/EB assay and flow cytometry (Figure 3A,B; Table 2). In a
recent report, Kling et al demonstrated that administration
of nitrofen to teratocarcinoma P19 cells resulted in
caspase-3 cleavage and caspase-dependent
apoptosis[32]. However, in the present study we observed no cleavage of caspase-3,
the key executor of apoptotic cell death, within
nitrofen-treated type II A549 pneumocytes (Figure 3C). In addition,
pretreatment of A549 cells with zVAD-fmk, a pan-caspase
inhibitor[25], did not abolish nitrofen-induced apoptosis
(Figure 3B), which indicated that a caspase-independent
mechanism is involved in nitrofen-induced apoptosis of type
II pneumocytes. We analyzed that the different findings
between the current study and Kling's
report[32] may be due to the characteristic of different cell types. Recent evidence
indicates that differentiated cells appear to be more resistant
to caspase-dependent apoptosis than the undifferentiated
ones[33]. Teratocarcinoma cells, including the P19 cell line,
are pluripotent and exhibit similar characteristics to
undifferentiated embryonic and fetal cell
types[34]. However, the A549 pneumocytes are likely more
differentiated[13] and consequently, might be resistant to nitrofen-induced caspase
machinery, in which case the caspase-independent pathway
is triggered and enrolled in apoptosis.
In recent years, mitochondria are thought to act as key
coordinators of cell death. Numerous studies have
demonstrated that most cells manifest a collapse of mitochondrial
membrane potential as a prelude to nuclear DNA
degradation and apoptosis[35]. Several pro-apoptotic signaling and
damage pathways converge on mitochondria to induce
mitochondrial membrane permeabilization, and the
responsible molecules are Bcl-2 family
proteins[36]. In healthy cells, Bax and Bak are in an inactive conformation in which their
NH2 and COOH termini are folded into a hydrophobic
pocket[34]. In response to apoptotic stimuli, these proteins
unfold and form multimers in the mitochondrial membrane.
The lethality of this multimerization is blocked through the
formation of heterodimers with Bcl-2 or
Bcl-xL[37]. In the present study, we found that nitrofen did not affect the
expression of Bcl-2, Bax, and Bak (Figure 4A). However, the
expression of Bcl-xL within nitrofen-treated cells was
downregulated (Figure 4A), which was predicted to favor
Bak and Bax homodimerization and apoptosis induction. Our
findings also demonstrate a decline in the mitochondrial
membrane potential after exposure to nitrofen (Figure 4B),
indicating leakage of proteins from mitochondrial
intermembrane space. AIF is a flavoprotein with activities of both
oxidoreductase and DNA-binding domains, but no intrinsic
DNase activity[38]. In the mitochondria, AIF is involved in
cellular respiration[37] and is essential for cell
survival[39]. Emerging evidence suggests that translocation of
mitochondrial AIF into the nucleus is a hallmark of
caspase-independent apoptosis[40]. When translocated to the nucleus, AIF
binds the DNase Endo G, resulting in DNA fragmentation
and cell death[41]. In this report, we demonstrate the nuclear
translocation of AIF after treatment with nitrofen (Figure 4C),
which provides obvious support for the contention that
nitrofen is able to induce mitochondria-mediated
caspase-independent apoptosis in A549 cells.
MAPK are phosphorylated in response to a series of
extracellular stimuli. There are at least 3 subfamilies of the
MAPK superfamily: ERK, JNK, and
p38-MAPK[42], which have been implicated in both apoptosis and survival
signaling. ERK are activated and play a critical role in
transmitting signals initiated by growth factors, such as
epidermal growth factor and platelet-derived growth
factor[43]. JNK and p38-MAPK are potently activated by various forms of
inflammatory signals or stress[44]. Previous studies
demonstrated that the activation of MAPK was involved in
ceramide-activated apoptotic signaling upstream of the
mitochondria[45]. The p38-MAPK and ERK1/2 inhibitors
attenuated the mitochondrial release of
AIF[45]. In this study, we found that nitrofen administration resulted in the
activation of p38-MAPK, but not of ERK1/2 or JNK (Figure 5A).
This finding was similar to a recent report that found that
nitrofen induced p38-MAPK activity in P19 teratocarcinoma
cells, which was associated with reactive oxygen
species[32]. Moreover, pharmacological inhibition of p38-MAPK in A549
cells was shown to abolish the nitrofen-induced
downregula-tion of Bcl-xL, nuclear translocation of AIF, and apoptosis
(Figure 5B,C), demonstrating that the activation of
p38-MAPK is involved in nitrofen-induced apoptosis in A549
cells. However, at present, the involvement of other
signaling cascades can not be ruled out, which warrants our
further study.
In summary, in the present study we demonstrated that
nitrofen treatment would decrease the proliferation of
cultured A549 pneumocytes accompanied with the
downregula-tion of PCNA. The inhibition of cell proliferation would
relate to mitochondria-mediated caspase-independent
apoptosis involving the activation of the p38-MAPK
signaling pathway. These findings lay the groundwork for further
investigation into the mechanisms of nitrofen-mediated
pulmonary hypoplasia and for the characterization of the
pathways involved in nitrofen-mediated apoptosis in type II
pneumocytes.
Acknowledgements
We are grateful to Dr Dechun LI and Mr John LANGER
(Johns Hopkins University School of Medicine, Baltimore,
MD, USA) for their help in the preparation of this manuscript.
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