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Introduction
Peroxisome proliferator-activated receptor α
(PPARα) is a member of the nuclear receptor superfamily of transcription
factors and binds cognate response elements as an obligate heterodimer with the retinoid X receptor.
PPARα plays a key role in the transcriptional regulation of genes encoding mitochondrial fatty acid
β-oxidation (FAO) enzymes during cardiac development and in response to physiological and pathophysiological stimuli, including fasting, cardiac hypertrophy, and
cellular hypoxia[1-4]. FAO enzymes, including medium chain acyl co-enzyme A, dehydrogenase, and
muscle carnitine palmitoyltransferase I (M-CPT I or CPT
Iβ)[5_7], are the principal sources of energy production in adult mammalian
cardiomyocytes. PPARα is rich in tissues that have high
energy demand, such as heart, skeletal muscle, brown fat,
kidney, liver, and brain[8], demonstrating a close association
between PPARα and energy turnover.
However, little is known about how physiological or
pathophysiological signals are transduced for the
modulation of the transcription of PPARα in cardiomyocytes. The
p38 mitogen-activated protein kinase (MAPK) signaling
pathway could be activated by cellular stressors in the heart,
including ischemia, hypoxia, and hypertrophic growth
stimuli[9]. As mentioned earlier, the expression of
PPARα is modulated to adapt to different demands in these processes.
Therefore, we hypothesize that p38 MAPK may act as the
upstream event that regulates the transcription of
PPARα.
PPARα, together with PPARδ and PPARγ, form a
subgroup within the nuclear receptor
superfamily[8,10]. PPAR isoforms have been demonstrated to be involved in the
differentiation of several cell types, including nerve cells,
adipose cells, and some tumor cell
types[11_13]. Prominent increased mRNA of
PPARα has been observed in the heart during embryonic development starting on d
7 in vivo[14,15]. This prominent expression abates before birth, suggesting
that a period of PPARα exposure may be critical to the
normal development of the heart.
It is increasingly clear that the p38 MAPK pathway plays
an important role in a large number of cellular processes,
such as cell growth, cell differentiation, cell cycle arrest, and
apoptosis[9,16_20]. In mice, p38 MAPK activity was recently
demonstrated to be required for the development of the 8_16
cell stage embryos[17]. In the mouse embryonic carcinoma
cell model, the activation of p38 MAPK has been shown to
be necessary for cardiac
differentiation[16,21].
Although multiple roles for PPARα have been proposed,
little is known of the significance of PPARα in early cardiac
development, especially during the differentiation of
cardiomyocytes. Using embryonic stem (ES) cells as a
model system faithfully recapitulates cardiomyocyte
differentiation[22_24]. The present study was designated to
determine the possible function of PPARα in early cardiomyocyte differentiation of ES cells
in vitro. In addition, the p38 MAPK pathway was also detected to
evaluate the possible pathway regulating the transcription of
PPARα.
Materials and methods
Cell culture and differentiation The permanent ES cell
line D3 (American Type Culture Collection, CRL-1934,
Manassas, VA, USA) was cultivated in undifferentiated
states on primary cultures of mouse embryonic fibroblasts
in Dulbecco's modified Eagle's minimal essential medium
(DMEM, Gibco BRL, Life Technologies, Germany),
supplemented with 10% fetal calf serum (FCS, Gibco, Germany),
1×10-4 mol/L beta-mercaptoethanol (Sigma, St Louis, MO,
USA), non-essential amino acids (NEAA, Hyclone, Logan,
UT, USA, stock solution dilution 1:100), and
1×106 units/L recombinant mouse leukemia inhibitory factor (Chemicon,
Temecula, California, USA). For the differentiation of ES
cells, embryoid bodies (EB) were generated using the
hanging drop method[25,26]. On d 0, 30 microlitres of drops
containing approximately 600 ES cells were placed on the lids of
Petri dishes filled with D-Hanks' solution, and cultivated in
hanging drops for 3 d followed by another 2 d in the Petri
dishes. On d 5, the EB were plated separately onto
gelatin-coated 24-well culture plates in differentiation medium that
consisted of DMEM, 20% FCS, 1 µmol/L mercaptoethanol,
and 1% NEAA. GW6471 and SB203580 were added, respectively, at the time points indicated in the text.
Reagents SB203580 was purchased from Biomol
(Plymouth Meeting, PA, USA). GW6471 was obtained from
Sigma-Aldrich (USA). Each inhibitor was dissolved in DMSO
at 1000×immediately prior to use. Unused inhibitor was
aliquoted into Eppendorf tubes and stored at -20 °C.
RT-PCR The total RNA was isolated from the ES cells
and EB using Trizol reagent (Gibco BRL, Germany) in
accordance with the manufacturer's instructions. To synthesize
first strand cDNA, 1 µg total RNA was incubated with 0.5 µg
of oligo (dT) 6 primer (Sangon, Shanghai, China) and 5 µL
deionized water at 65 °C for 15 min. Reverse transcription
reactions of 20 µL were performed with 200 units of M-MuLV
reverse transcriptase (Gibco BRL, Germany), 4 µL of
5×reaction buffer, and 1 mmol/L deoxynucleoside
triphosphate (dNTP) mixture for 1 h at 42 °C. Polymerase chain
reactions of 50 µL contained 1 µL of the RT reaction product,
5 µL of 10×PCR buffer, 25 units Taq polymerase (Sangon,
China), 1 µL of 10 mmol/L dNTP mixture, and 30 pmol of each
primer.
Primers, annealing temperature, product size, and the
number of PCR cycles are depicted in Table 1. Products were
analyzed in 1.5% agarose, ethidium bromide staining gels.
β-actin was used as an internal standard. No PCR products
were found without RT reaction and in water controls. RNA
from 3 independent experiments were analyzed.
Western blot analysis The cells were washed with
phosphate-buffered saline PBS, collected in
radioimmuno-precipitation assay RIPA buffer (containing 0.2% Triton X-100, 5
mmol/L EDTA, 1 mmol/L phenylmethanesulfonyl fluoride
PMSF, 0.01 g/L leupeptin, and 0.01 g/L aprotinin) and lysed
for 30 min on ice. Aliquots were assayed for protein
concentration using the Bio-Rad protein assay kit (Hercules, CA,
USA); equal amounts of proteins were loaded per well on a
12% SDS-PAGE. Subsequently, the proteins were transferred
onto 0.45 µm pore size nitrocellulose membranes and blocked
with 5% dry milk in PBS (pH 7.4, with 0.1% Tween 20) at room
temperature. The blots were probed with either rabbit
polyclonal anti-PPARα, mouse monoclonal anti-troponin-T,
goat polyclonal anti-actin (dilution 1:500, Santa Cruz
Biotechnology, Santa Cruz, CA, USA), mouse monoclonal
anti-α-actinin (dilution 1:500, Sigma-Aldrich, USA), rabbit
polyclonal anti-phospho-p38 MAPK or rabbit polyclonal
anti-p38 MAPK (dilution 1:1000, Cell Signaling Technology, USA)
antibodies overnight at 4 °C, followed by washing 3 times
with PBS-Tween (0.1% Tween 20) at room temperature and
challenged with horseradish peroxidase HRP-conjugated goat
anti-rabbit, rabbit anti-goat, or mouse anti-mouse antibodies
(dilution 1:4000, Affinity Bioreagents, Golden, CO, USA),
respectively. The proteins were visualized
autoradiographi-cally with an enhanced chemiluminescent substrate (Pierce,
Rockford, IL, USA) and scanned using a bio-imaging
analyzer (Bio-Rad, USA).
Immunofluorescence analysis Differentiated EB that had
been grown on coverslips were fixed for 20 min in methanol
at -20 °C, followed by permeabilization in 0.1% Tween 20 in
PBS. After washing in PBS three times, the EB on the
coverslips were transferred to PBS containing 10% goat serum for
30 min at room temperature. The EB were then placed into
blocking buffer containing mouse monoclonal anti-α-actinin
antibodies (dilution 1:100) and incubated overnight at 4 °C,
followed by incubation in blocking buffer containing
fluorescein isothiocyanate (FITC)-labeled mouse anti-mouse
antibody (1:500, Sigma-Aldrich, USA). For double staining,
rabbit polyclonal anti-PPARα and mouse monoclonal
anti-troponin-T were added together with the last dilution of
1:100, followed by incubation in blocking buffer containing
FITC-labeled mouse anti-mouse (1:250, Sigma-Aldrich, USA)
and Texa red-labeled goat anti-rabbit (1:250, Santa Cruz
Biotechnology, USA) antibodies. For the morphometric
analysis for the expression of sarcomeric proteins, all
aspects of cell processing, immunostaining, and imaging were
rigorously standardized. Digital images were obtained from
randomly selected fields using 10×objective lens and
analyzed by the image analysis software of the confocal setup
(Leica TCS SP2, Bensheim, Germany). For fluorescence
excitation, the 488 nm band of the argon ion laser of the
confocal setup was used. Emission was recorded using a
longpass LP505-nm filter set (Leica, Germany). Data are
expressed as mean±SD calculated as a percentage of the cells
in the controls. At least 5 different fields were measured for
each dish.
Statistical analysis Student's t-test and ANOVA were
used to determine the statistical significance of differences
between the values for the various experimental and control
groups. P<0.05 was considered statistically significant.
Results
In vitro cardiomyocyte differentiation of ES cells
The study protocol of cardiac differentiation of murine ES cells
in vitro are shown in Figure 1, indicating the time points for
RT-PCR, Western blotting, and immunofluorescence. The
attached culture was established by plating a single, d 5 EB
(Figure 2A) onto a 24-well plate and allowing continued
cellular proliferation and differentiation. Within this
multicellular arrangement in EB outgrowth, cardiomyocytes appeared
as spontaneously contracting, round cell clusters. Each EB
contained 1 or more beating areas (Figure 2B, 5 d after
plating). An increase in size, strength of contraction, and
beat frequency was observed during further differentiation.
The synchronously contracting cardiomyocytes were
positive for the anti-troponin-T antibody (Figure 2C). At higher
magnification, cross striations were demonstrated with
α-actinin immunolabeling (Figure 2E).
PPARα increased during cardiomyocyte
differentiation Differentiation of ES cells into cardiomyocytes first
required their forming into EB by growing them in hanging
drop cultures. After 3 d in hanging drops, the EB were placed
into suspension for 2 d, after which they were plated into
individual wells of a 24-well plate. Beating commenced in
distinct areas within EB on average 2 d later (culture d 7), the
area of contracting cells expanded multifocally thereafter and
reached a plateau around d 12. Therefore, these 2 time points
were chosen for future analysis. As determined by Western
blotting, the expression of PPARα was at low levels in early
differentiation, but increased obviously in parallel with the
appearance of contracting cardiomyocytes (Figure 3A).
To confirm the result of the protein expression found by
Western blotting, cultures were fixed at d 10 and doubly
stained with antibodies against PPARα and cardiac
troponin-T. In each case, we selected fields which were full of
cells so that a comparison could be made between the level
of staining in the cells positive for the marker and those that
were negative. There was a strong correlation between
PPARα and troponin-T during differentiation; a high level of
PPARα was observed in cardiomyocytes positive for
troponin-T (Figure 3B). Negative areas were also positive for
PPARα, but the staining was at considerably low levels.
Specific inhibition of PPARα prevented cardiomyocyte
differentiation of ES cells in a time-dependent manner
GW6471, the specific inhibitor of
PPARα[27], was used in our experiment to address whether these changes in the
PPARα expression was a causative or a simple effect resulting from
beating area formation. To closely define the exact time point
during the course of the developmental program at which
PPARα activity was essential, GW6471 was applied at
different time courses of differentiation. The data showed that
only the application of GW6471 before d 9 could efficiently
prevent EB from cardiomyocyte differentiation, indicated by
the reduced formation of the beating area (Figure 4) and
expression of α-actinin and troponin-T (Figure 5B). When
GW6471 was applied after d 9, cardiomyocyte differentiation
and formation was slightly affected.
The expressions of α-MHC and MLC2v were analyzed
by semiquantitative RT-PCR as they were specific
transcription factors in cardiomyocyte differentiation. As a result,
GW6471 tended to reduce the concentration of α-MHC and
MLC2v compared with the appropriate controls (Figure 5A).
Inhibition of p38 MAPK prevented cardiomyocyte
differentiation and reduced the expression of
PPARα To explore the function of p38 MAPK in cardiomyocyte
differentiation and the expression of PPARα, SB203580, the specific
inhibitor of p38 MAPK, was employed in our experiment.
P-p38 MAPK activity was maintained at a high level from d 3
and was followed by a decrease on d 10 (Figure 6A). The
specific inhibition of p38 MAPK from d 3 to d 7 greatly
reduced the formation of beating cardiomyocytes in the EB
(Figure 6B) which was well matched with expression of
PPARα (Figure 6C). While treated with SB203580 from d 5, cardiac
differentiation and PPARα expression were just slightly
affected. The expression of troponin-T was analyzed to
confirm the inhibition of differentiation which was completely
absent from the undifferentiated cells and the cells treated
with SB203580 from d 3 to d 7. These results implied that the
upregulation of PPARα was restrained to cardiac
differentiation of ES cells in vitro, and p38 MAPK might be the signal
pathway regulating the expression of PPARα.
Discussion
In this investigation, we gain insight into the expression
and possible function of PPARα during the cardiomyocyte
differentiation of ES cells in vitro.
PPARα was observed to increase immediately after beating area formation.
Interestingly, this phenomenon was recently replicated as
the mRNA levels of PPARα increased almost 10-fold during
myogenesis[28].
Cardiomyocyte differentiation can be divided into two
processes: cardiogenesis and cardiac myofibrillogenesis. At
d 3, cardiac transcription factors (ie GATA4, NKX2.5, MEF2C)
are not expressed yet, while on d 5, cardiac transcription
factors start to be fully expressed, but those encoding
sarcomeric proteins are not. Since d 5 is a critical window at
which the cardiac differentiation program becomes fully
activated, but no cardiac cells identifiable by organized
cardiac sarcomeric proteins (ie α-MHC, MLC2V) are yet
present[29], PPARα appeared to mainly play a role in
myofibrillogenesis. Our data showed that the inhibition of
PPARα before d 9 significantly disrupted differentiation and
reduced mRNA levels of α-MHC and MLC2v.
Mitochondrial number and functional capacity are
dynamically regulated in accordance with cardiac energy
demands during developmental stages and in response to
diverse physiological conditions[30]. Since the cardiomyocyte
differentiation of ES cells in vitro faithfully replicates the
process in vivo, and these cardiomyocytes display
properties similar to those observed in those in
vivo or in primary cultures, there must be an increase in mitochondrial number
with the emergence of beating clusters. In addition to the
activation of suites of genes encoding contractile proteins,
cardiac differentiation is accompanied by an organization of
metabolism. Therefore, the most obvious explanation for
our findings was that the inhibition of PPARα impaired
mitochondria ATP production or function of mitochondria, thus
hampering normal development. That blockade of
mitochondrial activity by the inhibition of mitochondrial protein
synthesis, uncoupling of the inner membrane potential from
ATP synthesis, and the inhibition of mitochondrial ATP
production inhibits differentiation of skeletal muscle cells from
myoblast precursors[31]. The other explanation for our result
may be that PPARα was involved in the regulation of the
nitric oxid (NO)/nitric oxid synthase (NOS) system and thus
affected cardiac differentiation. The beneficial effects of the
activation of PPARα in the improvement of cardiovascular
function have long been reported and these beneficial
effects in cardiovascular diseases have been suggested to
involve the NO/NOS system[32,33]. Interestingly, NO
signaling plays an important role in cardiac differentiation and NO
treatment can accelerate mouse ES cells to differentiate into
cardiomyocyte by both inducing a switch toward a cardiac
phenotype and inducing apoptosis in cells not committed to
cardiac differentiation[34].
p38 MAPK was involved in cardiomyocyte
differentiation of mouse embryonic carcinoma cells. Recently, p38
MAPK was further demonstrated to commit ES cells to
cardiomyogenesis[18]. Consistent with these previous
studies, p-p38 MAPK was found to maintain at a high level
from d 3 to d 10, which overlaps with the increase of
PPARα. Furthermore, the inhibition of p38 MAPK from d 3 to d 7
significantly prevented cardiac differentiation and reduced
the expression PPARα, which implied a relationship
between PPARα and p38 MAPK.
A previous study revealed that ligand-activated
PPARα would drive its own
transcription[35], and this effect would
be increased if PPARα was phosphorylated by p38
MAPK[36]. There would have been abundant natural ligands (eg fatty
acids) for the activation of PPARα since EB were grown in
the presence of 20% fetal bovine serum during differentiation.
p38 MAPK-mediated phosphorylation activates PPARα in a
ligand-influenced manner and results in enhanced functional
cooperation with the transcriptional co-activator peroxisome
proliferator-activated receptor γ coactivator
(PGC)-1α. Although we have no detailed experimental data on the
mechanism by which p38 MAPK mediates the expression of
PPARα, two possibilities can be considered. First, p38 MAPK may
enhance the binding between PPARa and PGC-1α and thus
stimulates the transcription of PPARα. Second, the
activation of p38 MAPK can improve the stability of
PPARα during differentiation and thus contributes to the increase of
PPARα. However, we could not exclude the possibility that
other upstream proteins might be responsible for the
mediation of PPARα during cardiac differentiation
in vitro.
In conclusion, PPARα increased in cardiomyocyte
differentiation in vitro, and p38 MAPK partly acted as the
upstream event regulating the expression of PPARα. The
inhibition of PPARα in the early course markedly prevented
cardiac differentiation, as indicated by the reduced
expression of cardiac sarcomeric proteins and specific genes.
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