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Introduction
Constriction and remodeling of the pulmonary arteries
induced by hypoxia are very important physiological
phenomena in order to maintain ventilation_perfusion matching.
However, a sustained hypoxic pulmonary vasoconstriction
and artery remodeling also contribute to hypoxic pulmonary
hypertension. Until now, the mechanism of hypoxic
pulmonary hypertension has still not been fully elucidated. The
proliferation of pulmonary artery smooth muscle cells plays
an important role in hypoxic pulmonary artery remodeling.
The imbalance between apoptosis and proliferation may
contribute to this cell
proliferation[1,2]. Mitochondria play a great
part in regulating cell apoptosis and
proliferation[3]. When mitochondrial membrane potential
(Δψm) collapses, cytochrome c, which normally exists in the mitochondrial
intermembrane space, is released from the mitochondrial
intermembrane space to the cytosol. An increase in cytosolic
cytochrome c is a trigger for the activation of caspase-9,
which then activates a set of cysteine proteases that are
central executioners of the cell apoptotic pathway. The
mitochondrial membrane ATP-sensitive potassium channel
(MitoKATP) is a key factor which
determinesΔψm[4].
MitoKATP is very sensitive to hypoxia. In a normoxic environment,
MitoKATP always is closed and
Δψm collapses; while in a hypoxic environment,
MitoKATP is opened and Dym increases.
Recently, some studies have demonstrated that
MitoKATP contributes to apoptosis and proliferation of myocardial
cells[5,6]. So far, there has not been any reports as to whether
MitoKATP plays any important role in human hypoxic pulmonary artery
remodeling. Diazoxide, an opener of
MitoKATP, and 5-hydroxydecanoate (5-HD), an antagonist of
MitoKATP, were used to influence
MitoKATP. Both diazoxide and 5-HD, which
are specific for KATP on the mitochondrial membrane, have
no effect on cell membrance. This study was aimed to
investigate the roles of MitoKATP and
Δψm in the proliferation of hypoxic human pulmonary artery smooth muscle cells and
the pharmacological effect of 5-HD on this process.
Materials and methods
Ethics of experimentation This study was approved by
the ethical committee in our hospital and by the patients
involved in this study. The experiments were conducted in
accordance with the Declaration of Helsinki.
Culture and identification of human pulmonary artery
smooth muscle cells Normal human lung tissues were
obtained from 6 patients receiving partial lung resection.
Pulmonary arteries (diameter: 2_3 mm) were dissected from the
surrounding parenchyma and washed twice in precooled
D-Hanks' solution containing 100 U/mL penicillin and 100
U/mL streptomycin. Under a dissecting microscope, smooth
muscle bundles were dissected from the artery. The minced
muscle fragments (about 1 mm3 in size) were digest-ed in
D-Hanks' solution containing 3 mg/mL collagenase type I
(Sigma, St Louis, MO, USA) at 37 °C for 50_60 min. Then
1_2 mL of 0.25% trypsin (Sigma, St Louis, MO, USA) was added
and incubated for about 10 min for digestion. The digestion
was finally stopped with Dulbecco's modified Eagle's
medium (DMEM) containing 20% fetal bovine serum (FBS,
Invitrogen, Carlsbad, CA, USA). The dissociated cells were
resuspended in DMEM containing 20% FBS, 100 U/mL penicillin, and 100 U/mL streptomycin, seeded onto 50 mL
culture flasks and maintained at 37 °C in an humidified
atmosphere with 5% CO2. The culture medium was replaced
every 2_3 d until conflu-ence was achieved (at d 15_20).
The cells were digested with 0.25% trypsin and subcultured
following the same procedure. Smooth muscle cells were
confirmed by immunohistochemistry staining against
α-actin using a mouse antihuman smooth muscle
α-actin polyclonal antibody was purchased from Santa Cruz
Biotechnology (Santa Cruz, CA, USA), and by the typical "hill
and valley" morphological pattern of confluent smooth
muscle cells under phase contrast microscope.
Grouping of human pulmonary artery smooth muscle
cells HPASMC (passages 4_8) were cultured in DMEM with
20% FBS and divided into 6 groups: (i) control group, where
cells were cultured in normoxia for 24 h (20%
O2, 5% CO2); (ii) diazoxide group, where cells were pretreated with a
MitoKATP specific
opener-diazoxide[8] (100 µmol/L; Sigma, USA) and
cultured in normoxia for 24 h. Diazoxide was dissolved in
dimethylsulfoxide(DMSO) before being added to the
experimental solution (DMEM without FBS). The final
concentration of DMSO in DMEM was
<0.1%[8]; (iii) 5-HD group, where cells were pretreated with a
MitoKATP specific antagonist_5-HD
[8] (500 μmol/L Sigma, USA) and cultured in normoxia for
24 h. 5-HD was dissolved in the experimental solution (DMEM
without FBS); (iv) 24 h hypoxia group, where cells were
cultured at 37 °C in an atmosphere containing 5%
O2, 5% CO2, and 90% N2
for 24 h; (v) 24 h hypoxia+diazoxide group, which
involved 24 h hypoxia treatment in the concomitant
stimulation of diazoxide; and (vi) 24 h hypoxia+5-HD group, which
involved 24 h hypoxia treatment in the concomitant
stimulation of 5-HD.
Measurement of rhodamine-123
fluorescence[9] The cells seeded on 25 mm coverslips were stained with rhodamine
123 (R-123; Sigma, St Louis, MO, USA) by incubating with
10 µg/mL R-123 for 30 min at 37 °C. The cells were perfused
with physiological salt solution to establish a baseline of
fluorescence. R-123 fluorescence was excited at 488 nm and
measured at 530 nm with Leica SP-1 confocal microscopy
(Leica, wiesbaden, Germany). The fluorescence signals were
stored in a computer and analyzed with Leica TCS NT
software.
R-123 is taken up selectively by
mitochondria[10,11] and its uptake depends on
Δψm. In isolated mitochondria, the relationship between the intensity of R-123 fluorescence
and Δψm is linear. The uptake of R-123 fluorescence, which is
quenched at resting Δψm, increases with the depolarization
of the mitochondrial membrane[11].
Measurement of HPASMC proliferation by
3-(4,5-dimethyl-2-thiazol-yl)-2,5-
diphenyl-2H-tetrazolium bromide (MTT) assay
HPASMC were seeded at a density of
1×104 cells/well into 96-well plates, cultivated, and divided into
different groups as described above. MTT (5 mg/mL) was
added to the wells (20 µL/well) at the end of the experimental
period. After 4 h incubation at 37 °C, media were removed
from the wells, and DMSO was added (150 µL/well). The
plates were agitated at room temperature for 30 min.
Absorbance (value A) of every well at a 490 nm wavelength was
read on an ELISA reader.
Detection of expression of proliferative cell nuclear
antigen in HPASMC by immunohistochemistry staining
HPASMC were seeded at a density of
1×105 cells/mL onto
6-well plates, cultivated, and divided into the 6 different
groups as described earlier. The HPASMC were fixed with
cool acetone for 10 min. The expression of proliferative cell
nuclear antigen (PCNA) in the HPASMC was detected by
immunohistochemistry staining using a polyclonal antibody
against PCNA (Santa Cruz, USA) at 4 °C overnight. The
same process without the primary antibody was used as a
control. After incubation with the secondary antibody
(Beijing Zhongshan Biotechnology, Beijing, China) at 37
°C for 1 h, the HPASMC were observed under light microscopy
and recorded photographically. The criteria for judgement
were as follows: the cells with brown staining in nuclei were
PCNA positive. On each slide, 500 cells (with the detection
of 6 slides) were randomly counted under a light microscope
and then the positive percentage per 100 cells was calculated.
Terminal deoxynucleotidyl transferase-mediated
dUTP-biotin nick end-labeling technique For the detection of
apoptotic cells, the apoptotic index was examined by the
terminal deoxynucleotidyl transferase-mediated dUTP-biotin
nick end-labeling (TUNEL) method. An in
situ cell death detection kit (ISCDD, Boehringer Mannheim, Germany) was
used to detect the apoptotic cells. The procedures were
carried out according to the protocol of the kit and other
references. The HPASMC were observed under microscopy
at once and recorded photographically. The cells with deep
blue staining in nuclei were TUNEL positive. In each slide (6
slides), 500 cells were randomly counted under a light
microscope and then the positive percentage per 100 cells was
calculated.
Flow cytometry analysis To determine the apoptosis,
the treated cells (1×109 cells/L) were washed in
phosphate-buffered saline (PBS) and fixed in 70% ethanol at 4 °C for 12
h. After incubation, 195 µL of the solution was transferred to
a 5 mL culture tube with 5 µL Annexin V-fluorescein
isothio-cyanate (FITC) added. The tube was then incubated for 30
min at room temperature in the dark. The cells were washed
with PBS and resuspended in 190 µL binding buffer with 10
µL propidium iodide (PI) added. Finally, the tube was gently
vortexed and incubated for another 30 min in the dark and
then the cells were analyzed immediately by flow
cytometry. Samples were acquired on a FACScan flow cytometer (Becton
Dickinson, Franklin Lakes, New Jersey, USA) and analyzed
with CellQuest software (Becton Dickinson, Franklin
Lakes, New Jersey, USA). Early apoptotic cells were characterized
by high Annexin binding and low PI staining, whereas late
apoptotic and necrotic cells were stained strongly with both
Annexin and PI.
Statistical analysis All data were expressed as mean±SD.
ANOVA was used for the comparison of variance among
several groups. The significance of difference between 2
groups was tested by t-test. P<0.05 was considered
statistically significant.
Results
Cell identification As shown in Figure
1, under
inverted phase contrast microscope, the HPASMC showed
spindle-shaped features, central oval nuclei with prominent
nucleoli, as well as the characteristic "hill and valley"
appearance. Immunohistochemistry staining of smooth
muscle-specific α-actin was positive. These results
indicated that the cells in culture are smooth muscle cells.
Changes of HPASMC mitochondrial membrane
potential in HPASMC As shown in Figure 2, diazoxide (100
µmol/L), a MitoKATP specific opener, increased the intensity
of R-123 fluorescence in the HPASMC as compared with the
control group (increased by 87%±39.86%,
n=6, P<0.05; Figure 2), indicating an increase of mitochondrial membrane
potential. However, 5-HD (500 µmol/L) did not markedly
change the intensity of R-123 fluorescence in the HPASMC
(n=6, P>0.05; Figure 2). Both 24 h hypoxia treatment and 24
h hypoxia treatment in the concomitant stimulation of
diazoxide increased R-123 fluorescent intensity (increased
by 52%±32.42%, and 117%±40.17%, respectively; Figure 2).
The changes of R-123 fluorescence intensity were obviously
enhanced under 24 h hypoxia treatment in the concomitant
stimulation of diazoxide as compared to 24 h hypoxia
treatment or diazoxide stimulation alone (217%±40.17%
vs 152%±32.42% and 217%±40.17%
vs 187%±39.86%, respec-tively,
n=6, P<0.05; Figure 2). 5-HD attenuated a 24 h
hypoxia-induced increase in R-123 fluorescence intensity
(114%±12.57% vs 152%±32.42%,
n=6, P<0.05; Figure 2). These results imply that hypoxia opens
MitoKATP and
induces an increase in Δψm in HPASMC.
Measurement of HPASMC proliferation by MTT
colorimetric assay Diazoxide increased the value A of HPASMC
as compared with the control group
(0.457%±0.041 vs 0.318%± 0.032%,
n=6, P<0.05; Figure 3). However, 5-HD
did not markedly change the value A of HPASMC compared
with the control group (0.323%±0.039%
vs 0.318%±0.032%, n=6, P>0.05; Figure 3). Both 24 h hypoxia treatment and 24 h
hypoxia treatment in the concomitant stimulation of diazoxide
increased the value A of HPASMC (0.478%±0.058%
vs 0.318%±0.032% and 0.621%±0.067%
vs 0.318%±0.032%, respectively,
n=6, P<0.05; Figure 3). Diazoxide significantly
enhanced the effects of 24 h hypoxia on the value A of
HPASMC (0.621%±0.067% vs 0.478%±0.058%,
n=6, P<0.05; Figure 3). 5-HD could weaken the increasing effect of 24 h
hypoxia on the value A of HPASMC (0.328%±0.044%
vs 0.478%±0.058%, n=6, P<0.05; Figure 3). These results
demonstrate that hypoxia and diazoxide induced cell proliferation;
5-HD almost completely inhibited the increase of value A
induced by hypoxia treatment.
Expression of PCNA in HPASMC Diazoxide increased
the expression of PCNA compared with the control group
(P<0.05; Figure 4). However, 5-HD did not markedly change
the expression of PCNA compared with the control group
(n=6, P>0.05; Figure 4). The 24 h exposure of the HPASMC
to hypoxia increased the PCNA expression (n=6,
P<0.05; Figure 4) and this was significantly augmented by a
concomitant stimulation of diazoxide (n=6,
P<0.05; Figure 4). 5-HD obviously attenuated a 24 h hypoxia-increased expression
of PCNA (n=6, P<0.05; Figure 4). These results demonstrated
that both hypoxia and diazoxide induced the proliferation of
the HPASMC, a process that can be inhibited by 5-HD.
TUNEL staining Diazoxide decreased the rate of
positive TUNEL-staining cells in the HPASMC compared with
the control group (n=6, P<0.05; Figure 5). However, 5-HD
did not markedly alter the TUNEL staining result in the
HPASMC compared with the control group (n=6,
P>0.05; Figure 5). The exposure of the HPASMC to hypoxia for
24 h prominently decreased the rate of positive
TUNEL-staining cells (n=6, P<0.05; Figure 5).These effects of hypoxia on
TUNEL staining is obviously augmented by the
concomitant treatment of diazoxide (n=6,
P<0.05; Figure 5), whereas 5-HD significantly reversed the hypoxia-induced decline in
TUNEL staining in the HPASMC (n=6, P<0.05; Figure 5).
These results demonstrate that both hypoxia and diazoxide
can inhibit apoptosis in HPASMC, which can be reversed by
5-HD.
Flow cytometric analysis To determine whether 5-HD
and diazoxide affect the apoptotic rate in HPASMC, we also
examined the expression of Annexin V and the exclusion of
PI by flow cytometry analysis. Diazoxide decreased the
percentage of apoptosis in HPASMC compared with the
control group (7.41%±1.32% vs 17.23%±4.16%,
n=6, P<0.05; Figure 6). However, 5-HD did not markedly alter the positive
percentage of apoptosis compared with the control group
(20.37%±4.89% vs 17.23%±4.16%,
n=6, P>0.05). The exposure of HPASMC to hypoxia for 24 h prominently decreased
the positive percentage of apoptosis (8.27%±1.82%
vs 17.23%± 4.16%, n=6, P<0.05). These effects of hypoxia on the
positive percentage of apoptosis are obviously decreased by
the concomitant treatment of diazoxide (2.52%±0.22%
vs
8.27%±1.82%, n=6, P<0.05), whereas 5-HD significantly
reversed the hypoxia-induced decline in the positive
percentage of apoptosis in HPASMC
(12.48%±3.07% vs 8.27%±1.82%, n=6, P<0.05). These results also demonstrate that
both hypoxia and diazoxide inhibit apoptosis in HPASMC,
which can be reversed by 5-HD.
Discussion
The precise control of the balance between pulmonary
artery smooth muscle cell (PASMC) proliferation and
apoptosis plays an important role in maintaining normal
pulmonary vascular structure and function. The reinforcement
of proliferation or the weakness of apoptosis in PASMC leads
to pulmonary hypertension and pulmonary vascular
structure remodeling. Apoptosis is a physiological mode of cell
death that is triggered by diverse external or internal signals.
Apoptosis in pulmonary artery smooth muscle cells allows
the pulmonary vasculature to tightly control the vascular
wall tissue mass (or thickness). Dysfunction of this process
has been widely linked to the pathogenesis of pulmonary
vascular diseases[12]. Some data have demonstrated that
apoptosis in PASMC and endothelial cells is involved in the
regression of pulmonary arterial
hypertrophy[13]. Most pulmonary hypertension was caused by hypoxia induced by
various reasons[14]; this pulmonary hypertension was named
hypoxic pulmonary hypertension. Some studies have shown
that hypoxia can promote PASMC proliferation, which can
subsequently trigger hypoxic pulmonary vascular structure
remodeling[14]. The mechanism of hypoxic pulmonary
vascular structure remodeling has not yet been fully elucidated,
and studies about the mechanism on human PASMC are
very scarce. It is a complex network that relates to many
cytokines and transcriptional factors.
Mitochondrial is a very important oxygen sensor in
mammalian cells. However, the studies about the modulation
mechanism of mitochondrial on hypoxic pulmonary vascular
structure remodeling are rarely documented. Recently, some
studies have found that Δψm was markedly linked to cell
proliferation in myocardial cells[15], but studies about
Δψm are seldom on PASMC. It is known that
MitoKATP is
extremely sensitive to hypoxia[16] and is a major factor in the
control of Δψm. Furthermore, some studies have identified a
mechanistic link between MitoKATP
channels and the mitochondrial apoptotic pathway. The
principal findings are as follows: (i) in isolated cardiac myocytes,
the MitoKATP channel opener diazoxide inhibits the activation of
the mitochondrial apoptotic pathway induced by oxidative
stress in a concentration-dependent
manner[17]; And (ii) the channel
blocker 5-HD (as well as glibenclamide) abolishes the
anti-apoptotic effect of
diazoxide[18]. These observations support the
hypothesis that the activation of
MitoKATP channels inhibits
apoptosis[19]. In a normoxic condition, the production of ATP
is at a natural level and the MitoKATP is always closed.
Mean-while, Δψm collapses and cytochrome
c is released from the mitochondrial intermembrane space to the cytosol.
An increase in cytosolic cytochrome
c is a trigger for the activation of caspase-9, which then activates a set of cysteine
proteases that are central executioners of the cell apoptotic
pathway[20]. In a hypoxic environment, the production of
ATP decreases[21], which triggers the activation of
MitoKATP and the Dym
increase, subsequently inhibits cytochrome
c releasing from the mitochondrial intermembrane space to the
cytosol and triggers cell proliferation. So far, it has not been
reported whether MitoKATP plays an important role in human
hypoxic pulmonary vascular structure remodeling.
In the present study, hypoxia for 24 h increased the value
of MTT and PCNA staining, decreased TUNEL staining, as
well as the percentage of apoptotic cells in HPASMC. These
results showed that hypoxia promoted HPASMC
proliferation and inhibited HPASMC apoptosis. Diazoxide opened
MitoKATP, increased Δψm and the proliferative status of
HPASMC in a similar way as that of HPASMC exposed to
hypoxia. These results also showed that an increase in
Δψm promoted HPASMC proliferation. In the presence of 5-HD,
MitoKATP was closed and the proliferative status of HPASMC
showed no any remarkable change compared with the
control group. This study has shown that an exposure to
hypoxia for 24 h increases Δψm and promotes HPASMC
proliferation. 5-HD can inhibit
MitoKATP and therefore attenuate hypoxia- and diazoxide-induced proliferation in
HPASMC. These results prove that
MitoKATP plays an important role in HPASMC proliferation induced by hypoxia.
In addition, many other factors may be involved in this
signaling pathway, for instance, hypoxia-inducible factor
1 [22], protein kinase C[23],
fas/fasL[23], c-jun,
Bcl-2[24],
[Ca2+]i[25].
Other factors that might be involved in the mechanism of
hypoxic pulmonary vascular structure remodeling remain to
be investigated.
In the present study, we found that 5-HD can prevent
hypoxic pulmonary artery smooth muscle cell proliferation.
We speculate that the mechanisms of 5-HD on hypoxic
pulmonary vascular structure remodeling may involve several
routes as follows: (i) to change mitochondrial morphology
and structure via changing mitochondrial membrane potential; (ii) to change the transmission and metabolism of
cell energy; (iii) to impact other cellular factors involved in
pulmonary artery smooth muscle cell proliferation; and (iv)
other potential routes. In future, we will keep focusing on
the effects of 5-HD in vivo (animal model) and further
investigate the pharmacological mechanisms of 5-HD as an
inhibitor of mitochondrial KATP channel activation on hypoxic
pulmonary vascular structure remodeling.
In summary, our study suggests that
MitoKATP and Dym play an important role in hypoxic pulmonary vascular
structure remodeling in humans, providing theoretical evidence
for the further study of the mechanism and prevention of
human pulmonary hypertension.
Acknowledgement
We thank all of the surgeons in the Department of
Thoracic Surgery, Tongji Hospital, Tongji Medical College,
Huazhong University of Science and Technology for their
assistance.
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