| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
(PPAR) Messenger RNA Expression and Stimulates Adipogenic Conversion of NIH-3T3 Cells
Helix Research Institute (R.N.-W., Y.M., M.-a.M., H.W.) Chiba,
292-0812, Japan
The First Department of Internal Medicine
(Y.F.) Osaka University Medical School Osaka, 565 Japan
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
(PPAR) play crucial
roles in this process. Although ectopic expression of these factors in
NIH-3T3 cells, a multipotential mesenchymal stem cell line, results in
adipogenic conversion, little is known as to hormonal factors that
regulate adipogenesis in these cells. In this report we demonstrate
that PRL, a lactogenic hormone, enhances C/EBPß and PPAR mRNA
expression and augments adipogenic conversion of NIH-3T3 cells.
Moreover, we show that ectopic expression of the PRL receptor in
NIH-3T3 cells results in efficient adipocyte conversion when stimulated
with PRL and a PPAR ligand, as evidenced by expression of the
adipocyte differentiation-specific genes as well as the presence of
fat-laden cells. We further demonstrate that signal transducer and
activator of transcription 5 (Stat5), a PRL signal transducer,
activates aP2 promoter in a PRL-dependent manner. These results suggest
that PRL acts as an adipogenesis-enhancing hormone in NIH-3T3 cells. |
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
(PPAR) are potential regulators of this process, relaying external
signals to gene expressions that ultimately lead to adipocyte
differentiation.
C/EBP are characterized by a common structure, the presence of a
C-terminal leucine zipper for dimerization and basic residues
responsible for DNA binding. Among the family of C/EBPs, C/EBP
,
-ß, -
, and CHOP (Gadd153) have been shown to be involved in
adipogenesis (3, 4, 5, 6, 7). Chronologically, the expression of C/EBPß and
C/EBP precedes that of C/EBP during differentiation of 3T3-L1
cells (8). Evidence that these C/EBPs are crucial regulators of
adipogenesis stems in part from the observation that ectopic expression
of C/EBPß and, to a lesser extent, C/EBP results in the conversion
of preadipocytes or multipotential mesenchymal stem cells into
adipocytes (6). As well, overexpression of a dominant-negative form of
C/EBPß inhibits 3T3-L1 cell differentiation (6). C/EBP has also
been shown, from both antisense and overexpression studies, to play a
crucial role in adipogenesis (3, 4, 5). Recent studies using gene-targeted
mice support these findings. For example, mice ablated of C/EBP
suffer significant decreases in both brown adipose tissue (BAT) and in
white adipose tissue (WAT) (9). Also, while depletion of either the
C/EBPß or gene results in only a mild perturbation of adipogenic
differentiation of primary embryonic fibroblasts and a slight volume
loss in epidydimal WAT, the C/EBPß and - double knockout mice
display an almost complete abrogation of adipocyte differentiation as
well as severely reduced WAT weight due to a greatly diminished number
of adipocytes (10).
PPAR is a member of the ligand-stimulated nuclear hormone
receptor superfamily (11, 12). PPAR plays a central role in
adipocyte differentiation. Enforced expression of PPAR in
multipotential mesenchymal stem cells results in adipogenic conversion
in the presence of its ligands/agonists, thiazolidinedione, or
prostaglandin (13, 14, 15, 16). In adipocytes, PPAR is responsible for the
expression of adipose differentiation-related genes such as 422/aP2,
phosphoenol pyruvate carboxykinase, and lipoprotein lipase. Indeed,
promoters of these genes contain PPAR binding sites (17, 18, 19).
Although these families of transcription factors have been shown to
play important roles in adipogenesis, little is known about which
extracellular stimuli drive the mesenchymal cell to commit toward the
adipogenic lineage. In vitro studies have established that
hormones such as dexamethasone (DEX), methylisobutylxanthine (MIX),
insulin, and those present in FBS trigger the adipogenic conversion of
3T3-L1 cells as well as NIH-3T3 cells ectopically expressing C/EBPs or
PPAR. MIX and DEX are direct inducers of C/EBPß and ,
respectively (8). This induction, in turn, promotes PPAR expression
and ultimately stimulates adipogenesis (20, 21). In contrast, simple
treatment of NIH-3T3 cells with the above mentioned adipogenic hormones
does not lead to adipogenic conversion. These observations indicate
that some signal transducers are absent or present at low levels in
NIH-3T3 cells, and the addition of some hormones or factors that
up-regulate the expression of C/EBPs or PPAR may have a potential to
convert NIH-3T3 cells into adipocytes. Recently, it has been shown that
the PRL receptor mRNA is up-regulated during preadipocyte
differentiation (22). PRL is best known as a lactogenic hormone
responsible for the development of the mammary gland, and it plays a
crucial role in reproduction, although many other functions are also
reported (23). With regards to diseases, PRL-secreting pituitary
adenoma is occasionally associated with obesity (24). In some cases of
prolactinoma and obesity, normalization of serum PRL level results in
reduction of body weight (25). These observations prompted us to
investigate a possible role of PRL and its cognate receptor in
adipogenesis. We herein demonstrate that PRL enhances C/EBPß and
PPAR mRNA production in conjunction with MIX and DEX in NIH-3T3
cells and provide evidence that ectopic expression of the PRL receptor
results in adipogenic conversion of NIH-3T3 cells in the presence of a
PPAR stimulator. We also show that PRL contributes to the activity
of aP2 gene promoter via signal transducer and activator of
transcription 5 (Stat5).
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
are the initial transcription factors
required for adipogenesis and their mRNAs are induced by a variety of
stimuli, including MIX, DEX, insulin, lipopolysaccharide,
interleukin (IL)-1, IL-6, and GH (8, 26, 27), we first asked whether
PRL enhances C/EBPß mRNA expression in multipotential mesenchymal
stem cells (NIH-3T3) and in 3T3-L1 preadipocyte cells. Confluent
NIH-3T3 or 3T3-L1 cells were treated with or without increasing
concentrations of PRL or with various effectors, and Northern blot
analysis was performed (Fig. 1
). While
nontreated NIH-3T3 cells expressed a low level of C/EBPß mRNA, PRL
increased its transcript in a dose-dependent manner (Fig. 1A, lanes
1–5). At 333 ng/ml of PRL, the induction was saturated (Fig. 1A, lane
4). This dose of PRL was as efficacious as MIX, DEX, and insulin (Fig. 1A, lanes 4 and 6–8). FBS was not as potent as PRL (Fig. 1A, lane 9).
The effect of PRL was also examined in 3T3-L1 cells. PRL failed to
enhance C/EBPß mRNA (Fig. 1B, lanes 1–5), while MIX, DEX, and
insulin increased C/EBPß mRNA expression as previously reported (Fig. 1B, lanes 6–8) (8). FBS moderately enhanced its expression but less
efficiently than the other stimulators (Fig. 1B, lane 9). No
PRL-dependent expression of C/EBP mRNA was detected in either cell
type (data not shown). These results suggest that PRL has an inductive
effect on C/EBPß transcription, at least in NIH-3T3 cells.
|
mRNA in NIH-3T3 Cells as Well as in
Preadipocyte 3T3-L1 Cells expression
(20), we next asked whether PRL might influence PPAR expression. As
optimal PPAR mRNA production requires MIX, DEX, and insulin (21),
PRL was challenged together with these adipogenic hormones. Figure 2 shows Northern blot analysis for
NIH-3T3 (Fig. 2A) and 3T3-L1 cells (Fig. 2B). While PPAR mRNA was
efficiently induced by MIX, DEX, and insulin in 3T3-L1 cells, it was
barely detected in NIH-3T3 cells in the absence of PRL (Fig. 2, A and
B, lane 1). In both cell lines, however, addition of PRL resulted in a
dose-dependent transcriptional augmentation (Fig. 2, A and B, lanes
2–5). PRL at the concentration of 333 ng/ml was as potent as FBS in
both cell lines (Fig. 2, A and B, lanes 4 and 6). To further determine
which adipogenic effector(s) synergize with PRL for PPAR mRNA
expression, different combinations were tested in NIH-3T3 (Fig. 2C) and
in 3T3-L1 cells (Fig. 2D). MIX, DEX, and insulin efficiently induced
PPAR mRNA in 3T3-L1 cells but much less efficiently in NIH-3T3 cells
(Fig. 2, D and C, lane 1). In NIH-3T3 cells PRL alone or in combination
with MIX, insulin, or DEX induced little mRNA (Fig. 2C, lanes 2–5).
Combination of MIX and DEX, but not MIX and insulin, nor DEX and
insulin, led to strong PPAR mRNA expression together with PRL (Fig. 2C, lanes 6–8). Insulin plus MIX and DEX resulted in a maximized
induction, and FBS was as efficacious as PRL (Fig. 2C, lanes 9 and 10).
Similar results were obtained in 3T3-L1 cells (Fig. 2D). It is
noteworthy that DEX and PRL had an inductive effect, which was slightly
enhanced by insulin (Fig. 2D, lanes 5 and 8). These data demonstrate
that PRL enhances MIX- and DEX-induced PPAR expression in both cell
lines.
|
.
Troglitazone is a member of thiazolidinediones, which has
been shown to elevate the potential of multipotential mesenchymal cells
to differentiate (14, 28). Under these conditions, we set out to study
the function of PRL in adipogenic conversion of NIH-3T3 cells. The PRL
receptor gene was stably transfected into NIH-3T3 cells along with the
neomycin resistance gene. As a control, cells harboring only the
neomycin resistance gene were also selected. G418-resistant colonies
were cultured for 2 weeks and were induced to differentiate in
situ. Before differentiation stimulation, no fat-laden
differentiated cells were observed (data not shown). However, upon
exposure to the strong permissive regimen for 10 days, some terminally
differentiated adipocyte colonies, as evidenced by Oil-Red-O staining,
appeared in the plates transfected with the PRL receptor (Fig. 3B, PRLR). From three independent
experiments, 11% of the G418-resistant colonies cotransfected with the
PRL receptor gave rise to differentiated adipocytes. In
contrast, 2% of differentiated colonies were present in the control
cells (Fig. 3A, Neo). The omission of PRL from the medium resulted in a
significant decrease of differentiated colony number in both cases
(data not shown). These data further reinforced our hypothesis that PRL
and the PRL receptor play an important role in adipogenic
differentiation of NIH-3T3 cells.
|
mRNA expression. Cells transfected only with
the neomycin resistance gene exhibited similar dose-dependent C/EBPß
mRNA expression as parental cells (Figs. 1A
and Fig. 4A, lanes 1–5). In contrast, the PRL
receptor-expressing cells showed an enhanced sensitivity to a given
dose of PRL (Fig. 5A, lanes 6–10). Even
at the highest concentration of PRL (1 µg/ml), the C/EBPß mRNA
level in cells expressing the neomycin resistance gene was lower than
that induced with 111 ng/ml of PRL in the PRL receptor-expressing cells
(Fig. 4A, compare lanes 5 and 8). As for PPAR mRNA, the same PRL
dose dependency was observed in parental and in the control cells
(Figs. 2A and Fig. 4B, lanes 1–5). Interestingly, ectopic expression
of the PRL receptor made these cells much more sensitive to PRL as in
the case for C/EBPß mRNA (Fig. 4B, lanes 1–5 and 6–10). Addition of
111 ng/ml of PRL to the PRL receptor-expressing cells resulted in the
similar induction of PPAR mRNA as seen with 1 µg/ml of PRL to the
control cells (Fig. 4B, compare lanes 5 and 8). We next examined the
differentiation program of these two cells with or without PRL (Fig. 5). PRL slightly elevated C/EBPß mRNA in the control cells after 3
days (Fig. 5A). In the PRL receptor-overexpressing cells, further
up-regulation was detected especially at day 3 (compare lanes 4 and 9
in Fig. 5A and those in Fig. 5B), and thereafter PRL sustained C/EBPß
mRNA at a slightly higher level (Fig. 5, A and B, compare lanes 9–11).
As for C/EBP mRNA, addition of PRL had no effect in either cell
throughout the entire differentiation process (data not shown).
Expression of PPAR mRNA peaked on day 2 in the control cells, and
then sharply declined in the absence of PRL (Fig. 5A, lanes 1–6).
Ectopic expression of the PRL receptor resulted in enhanced expression,
even in the absence of exogenous PRL (Fig. 5B, lane 3). Addition of PRL
kept the level of PPAR mRNA slightly high during the later period of
the differentiation program in both control and PRL
receptor-overexpressing cells (Fig. 5, A and B, lanes 9–11). Adipsin
mRNA was detected on day 5 in both control and PRL
receptor-overexpressing cells, and PRL significantly enhanced the mRNA
in the PRL receptor-overexpressing cells (Fig. 5, A and B, lanes 5 and
10). Quite interestingly, the control cells failed to accumulate
adipsin mRNA by day 8, while the PRL receptor-overexpressing cells kept
adipsin mRNA high until day 8 (Fig. 5, A and B, lanes 5, 6, 10, and
11). aP2 mRNA was detected as early as day 3 in both control and PRL
receptor-overexpressing cells (lanes 4 in both panels A and B).
Addition of PRL strongly augmented the expression of aP2 mRNA,
particularly in cells overexpressing the PRL receptor (lanes 9–11 in
both panels A and B). Glycerol-3-phosphate dehydrogenase (GPD)
mRNA was detected on day 8 in the control cells; however, the effect of
PRL on the mRNA was not clear (panel A, lanes 6 and 11). On the other
hand, PRL increased GPD mRNA in cells overexpressing the PRL receptor
(Fig. 5B, lanes 5, 6, 10, and 11). We have repeated these experiments
four times and obtained essentially identical results.
|
|
, Neo, FBS+I+T), 4% of control cells exhibited adipogenic conversion in
the presence of PRL as judged by Oil-Red-O-positive cell number (Fig. 6A, Neo, FBS+I+T+PRL). In contrast, 18% of the PRL
receptor-transfected cells underwent adipogenic conversion in the
presence of PRL, as assessed by the formation of fat droplets in cells
(Fig. 6B, PRLR, FBS+I+T+PRL). On the other hand, omission of PRL
resulted in a significant decrease of differentiated cell population to
5% (Fig. 6B, PRLR, FBS+I+T). Four independent experiments were
performed and 13–23% of G-418-resistant cells overexpressing the PRL
receptor converted into lipid-laden adipocytes in the presence of PRL.
In light of these results, we concluded that PRL and its cognate
receptor up-regulate the adipogenic conversion of NIH-3T3
multipotential mesenchymal stem cells.
|
2, Stat5A plus PPAR2, or an
ovine dominant negative form of Stat5(Y694F) (31). After transfection
cells were treated with different combinations of PRL and
troglitazone (a PPAR ligand). When the aP2 promoter
alone was transfected, any combination of PRL/troglitazone
failed to enhance the promoter activity (Fig. 7, lanes 1–4). In contrast, transfection
with Stat5A cDNA resulted in a PRL-dependent 2-fold increase in
promoter activity (Fig. 7, lanes 7 and 8). Troglitazone
did not affect the promoter activity (Fig. 7, lane 6). When PPAR2
cDNA was cotransfected, troglitazone-dependent enhancement
of the promoter activity was observed. Challenge with
troglitazone resulted in 60% augmentation, while
combination with PRL gave 100% increase in promoter activity (Fig. 7, lanes 9, 10, and 12). PRL alone had no effect in this case (Fig. 7, lane 11). Cotransfection with both Stat5A and PPAR2 cDNA showed
additive effects. Troglitazone or PRL challenge doubled
the promoter activity, whereas the combination of both ligands resulted
in a 2.5-fold increase (Fig. 7, lanes 13–16). These data strongly
suggested that Stat5 regulates aP2 promoter activity. To further
strengthen this hypothesis, ovine dominant negative Stat5 (Y694) was
transfected into cells together with aP2 promoter. When challenged with
PRL, no ligand-dependent promoter activity was observed (Fig. 7, lanes
17, 18, and 19). Addition of troglitazone had no effect on
promoter activity (Fig. 7, lanes 18 and 20). Use of Stat5B and the
corresponding dominant negative Stat5B (Y699F) gave essentially
identical results (data not shown). These data indicate that the aP2
gene is regulated by PRL, at least partially, via Stat5.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
mRNA in conjunction with MIX and DEX
in NIH-3T3 cells. 2) PRL enhances the adipogenic conversion of NIH-3T3
cells under strong permissive conditions (i.e. in the
presence of a PPAR ligand/agonist). 3) ectopic expression of the PRL
receptor efficiently converts NIH-3T3 cells into fat-laden adipocytes.
4) PRL regulates aP2 promoter via Stat5.
3T3-L1 cells are of preadipocyte lineage and differentiate into
adipocytes when stimulated appropriately (1). Growth factors such as
insulin, insulin-like growth factor-1, epidermal growth factor, and
platelet-derived growth factor have been shown to promote
adipogenesis in 3T3-L1 cells, and, in fact, FBS is a potent stimulus
(32, 33, 34). On the other hand, NIH-3T3 cells are considered to be a
multipotential mesenchymal cell line that does not readily
differentiate into adipocytes as 3T3-L1 cells do. Nevertheless, NIH-3T3
cells do convert into adipocytes when transcription factors such as
C/EBPs, PPAR, and ADD-1 are overexpressed (4, 6, 13, 35). These data
imply that NIH-3T3 cells possess a potential to undergo adipogenesis.
To our knowledge, there has been no report of any growth factor or
cytokine that enhances adipogenesis in NIH-3T3 cells. Our present data
clearly show that PRL and its cognate receptor are required but not
sufficient for the activation of adipogenic program in NIH-3T3 cells.
While PRL increased C/EBPß mRNA in a concentration-dependent fashion
in NIH-3T3 cells, no such effect was observed in 3T3-L1 preadipocytes.
At present, the reason for this difference is unclear. In both NIH-3T3
and 3T3-L1 cells, PRL enhanced PPAR mRNA expression in conjunction
with other adipogenic hormones such as MIX and DEX (Fig. 2, A–D). As
MIX and DEX induce C/EBPß and -, respectively (8), it is
conceivable that there is a cooperation between PRL-emanated signals
and those of C/EBPß and/or - for the maximum production of PPAR
mRNA. Further study is required to fully understand how PRL
receptor-triggered signals act together with these transcription
factors. The fact that FBS was as potent as PRL in inducing C/EBPß
and PPAR mRNA suggests that FBS contains PRL, or that some factors
present in FBS enhance PPAR mRNA (Figs. 1A and 2C and 2D). The
report that FBS is rich in PRL rather supports the former possibility
(36). The presence of PRL in FBS may explain the relatively high level
of PPAR mRNA expression in the PRL receptor-overexpressing cells in
the absence of exogenous PRL (Fig. 5B, lane 3). This, in turn, may
contribute the slight increase in the number of differentiated cells in
cells overexpressing PRL receptors (Fig. 6B, PRLR, FBS+I+T).
PRL has been shown to induce calcium influx and trigger
activation of signaling molecules such as Ras, PI3 kinase, and the
STATs (23, 30, 37, 38). It remains to be determined which of these
signaling pathways are responsible for C/EBPß and PPAR expression.
As for the transcriptional regulation of PPAR mRNA, it has been
shown that granulocyte-colony stimulating factor (G-CSF),
12-O-tetradecanoylphorbol-13-acetate, 1,25-dihydroxyvitamin
D3, and oxidized low density lipoprotein promote
its transcription in macrophages (39, 40). Since both G-CSF and PRL
activate JAK-STAT pathway, it is tempting to speculate that the maximum
induction of PPAR mRNA may be dependent on this pathway.
PRL has a potential in enhancing PPAR transcript in parental NIH-3T3
cells as well as the PRL receptor-transfected cells (Fig. 5). This
enhancement may, in part, explain why NIH-3T3 cells receiving PRL and
troglitazone exhibit morphological changes and express
adipocyte-specific marker genes such as adipsin, aP2, and GPD.
Intriguingly, PPAR mRNA level peaked at day 2 and gradually declined
during the adipogenic conversion (Fig. 5). This is in contrast to the
observation during the differentiation process of 3T3-L1 cells or
NIH-3T3 cells ectopically expressing C/EBPß (17, 20). In these cells
a high level of PPAR mRNA is sustained at later steps in
differentiation. Our data suggest that transient PPAR mRNA
production is sufficient for the induction of some terminal marker
genes, i.e. adipsin, aP2, and GPD and for the accumulation
of fat droplets (Figs. 5B and 6). Why do cells treated with PRL promote
adipogenic conversion to a certain extent even though the increase of
PPAR mRNA is rather transient? A possible explanation is that
PRL-elicited signals and those emanated from the activated PPAR
might coordinate to stimulate adipogenic program. In agreement with
this hypothesis, a significant drop of adipsin, aP2, and GPD mRNAs as
well as of Oil-Red-O positive cell number was observed in the absence
of PRL, while the PPAR mRNA expression profile remained nearly
identical in the PRL receptor-expressing cells (Figs. 5B and 6). Our
data from aP2 promoter analysis further indicate that aP2 gene is
controlled, at least in part, by Stat5, which is activated by PRL (Fig. 7). These results also suggest that the effect of PPAR and Stat5 on
the aP2 promoter is ligand dependent and additive. However, it has yet
to be elucidated how Stat5 regulates the aP2 promoter activity.
Recently, a cross-talk between PPAR, another member of PPAR family,
and STAT5B in GH signaling has been shown in COS cells (41). It will be
interesting to examine whether there are other adipocyte-specific genes
regulated by both PPAR and STAT5 as shown here for the aP2 gene.
It is noteworthy that GH shows opposite effects on the expression of
adipocyte-specific genes such as PPAR, aP2, and fatty acid synthase
and on adipogenesis in primary preadipocytes (42). Whether PRL exhibits
stimulatory or inhibitory effects on primary preadipocytes remains to
be addressed. GH deficiency is often associated with obesity (43),
while some prolactinoma patients are obese (24). In both cases,
treatment with either GH or with reagents lowing serum PRL results in
improvement of obesity (25, 43). These data imply that the action of GH
and PRL is opposite in vivo. Given the fact that GH and PRL
share many signaling molecules such as Jak2 and Stat5A and B, it is
necessary to delineate molecular mechanisms underling these opposing
effects.
PRL gene knockout mice grow normally but fail to develop mammary glands (44). PRL receptor gene ablation leads to a similar but somewhat different phenotype, resulting in reproductive defects (45). In both cases, gene-targeted mice grow as wild type. While mice lacking STAT5A grow normally, STAT5B-deficient male mice manifest a significant decrease in weight (46, 47). The STAT5A/STAT5B double-knockout mice exhibit more severe growth impairment, and the size of the fat pads is decreased to one-fifth of the wild type (48). These data suggest that STAT5A and B, both of which are activated by PRL, play a key role in the development of the fat pad. The results presented herein appear to be in line with these observations. The reason why mice lacking PRL or the PRL receptor do not manifest weight loss, as observed in STAT5B or STAT5A/STAT5B knockout mice, can be explained as follows. In the absence of PRL or its cognate receptors, other factor(s) or receptor(s) that activate STAT5A and/or B compensate for their functions. The redundancy in cytokine signaling may mask all of the PRL/PRL receptor functions from gene targeting experiments.
In summary, we have shown here that PRL enhances adipocyte differentiation of NIH-3T3 fibroblasts. Our results may have important implications for defining hormonal stimulation along the adipocyte differentiation pathway. Understanding the interactions between PRL-triggered signals and those emanated from other adipogenic inducers should provide valuable insight into the mechanisms that control adipogenic differentiation.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
Plasmids
The cDNAs for C/EBP, C/EBPß, C/EBP, PPAR, aP2, and
GPD used in this study were isolated from a 3T3-L1 preadipocyte library
constructed 9 days after induction of differentiation, using
SuperScript 
system (Life Technologies, Inc.,
Gaithersburg, MD). The sequences of the cDNAs were verified by DNA
sequencing (ABI 377 DNA sequencer). aP2 promoter comprising a -1 to
-5.4 kbp fragment was inserted into the SmaI site of
pGL2-basic vector (Promega Corp., Madison, WI).
Cell Culture and Induction of Differentiation
3T3-L1 preadipocytes (NIHS cell bank, Tokyo, Japan, catalogue
number JCRB9014) and multipotential mesenchymal stem cells (NIH-3T3;
Riken cell bank, Tsukuba, Japan; catalogue no. RCB0150) were
maintained in growth medium consisting of DMEM (Nisseiken, Kyoto,
Japan) containing 10% normal calf serum (CS; Life Technologies, Inc.). To establish stable cell lines, NIH-3T3
cells were transfected with the rat PRL receptor (PRLR) in pME18S and
pSV2neo with lipofectAMINE-PLUS reagent following the protocol provided
by the supplier (Life Technologies, Inc.). Briefly, a
mixture of 8 µg of PRLR plasmids, 0.4 µg of pSV2neo plasmids, 20
µl of PLUS reagent, and 30 µl of lipofectAMINE in 1.5 ml of
OPTI-MEM I (Life Technologies, Inc.) was incubated at room
temperature for 30 min and then diluted to 8 ml with OPTI-MEM I. Cells
plated at a density of 8 x105 cells per 10-cm
dish the day before transfection were washed with OPTI-MEM I, and the
diluted DNA-lipid mixture was added. After 3 h of incubation at 37
C, cells were refed with DMEM supplemented with 10% normal calf serum
and were cultured for an additional 24 h before selection with 0.4
mg/ml of G418. The transfectants were selected for 14 days and
subjected to in situ colony differentiation assay. In some
experiments, more than 20,000 G418-resistant clones were pooled and
assayed for their ability to differentiate. To convert NIH-3T3 cells
and the transfectants into adipocytes, cells were grown to confluence
(considered as day 0). At this stage, cells were exposed to fresh DMEM
containing 10% FBS (Life Technologies, Inc.), 1
µM DEX, 0.5 mM MIX, and
10 µg/ml of insulin for 48 h. After this treatment, the medium
was replaced with DMEM containing 10% FBS, and 2.5 µg/ml of insulin,
and cells were refed every other day (normal permissive conditions).
For strong permissive conditions, 5 µM
of troglitazone was included. To distinguish the effect of
PRL, 10% CS instead of 10% FBS was used in some experiments, as
indicated in the text and figure legends. Adipocyte conversion was
assessed by the presence of accumulated fat droplets in the cytoplasts.
Cells were fixed with 2% formaldehyde, 0.2% glutaraldehyde in PBS and
stained with Oil-Red-O (49).
Northern Blot Analysis
Total RNA was isolated according to the method of Chomczynski
and Sacchi (50). The indicated amount of the total RNA was fractionated
using 1% agarose/2.2 M formaldehyde gel electrophoresis
and was transferred to a nylon membrane (51). rRNA was stained on the
filters with methylene blue to assess RNA loading and transfer
efficiency (52). The DIG-labeled RNA probes were transcribed from
EcoRI-linearized cDNA plasmid in pZL1 according to
the Roche Molecular Biochemicals protocol. Hybridization
was performed with DIG-labeled antisense RNA probes according to the
protocol provided by the supplier (Roche Molecular Biochemicals, Indianapolis, IN).
Transcriptional Activation Assay
The expression vectors for murine Stat5A and ovine Stat5(Y694F)
were constructed by ligating the EcoRI-NotI
fragment containing Stat5A or Stat5(Y694F) into the
EcoRI-NotI site of pME18S. The
SalI-NotI fragment comprising PPAR2 was
inserted into the XhoI-NotI site of pME18S. aP2
promoter-luciferase vector (pGL2-aP2) was transiently transfected into
NIH-3T3 cells without or with expression vectors. NIH-3T3 cells were
maintained in DMEM containing 10% CS and transfected at 70%
confluency with LipofectAMINE-PLUS reagent (Life Technologies, Inc.). A renilla luciferase expression vector (pRL-CMV) was
cotransfected as a control for the transfection efficiency
(Promega Corp.). After DNA removal by washing, cells were
serum starved for 16 h in OPTI-MEM I, and then left untreated or
challenged with PRL, troglitazone, or PRL plus
troglitazone for 6 h. After cell lysis, both firefly
and renilla luciferase activities were measured. Transfections were
performed in duplicate and repeated two to four times.
|
ACKNOWLEDGMENTS |
|---|
Helix Research Institute is supported by the Ministry of International Trade and Industry (MITI), Chugai Pharmaceutical Co., Fujisawa Pharmaceutical Co., Hitachi Co., Mitsubishi Chemical Co., Nippon Godou Finance Co., Kyowa Hakko Co., Sumitomo Chemical Co., Taisho Pharmaceutical Co., Yamanouchi Pharmaceutical Co., and Yoshitomi Pharmaceutical Co.
|
FOOTNOTES |
|---|
Received for publication July 21, 1999. Revision received September 30, 1999. Accepted for publication November 11, 1999.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
is
sufficient to initiate the 3T3–L1 adipocyte differentiation program.
Proc Natl Acad Sci USA 91:8757–87612, a lipid-activated
transcription factor. Cell 79:1147–1156[CrossRef][Medline]
12,14-prostaglandin J2 is a ligand for the
adipocyte determination factor PPAR. Cell 83:803–812[CrossRef][Medline]
and promotes adipocyte
differentiation. Cell 83:813–819[CrossRef][Medline]
and C/EBP. Proc
Natl Acad Sci USA. 92:9856–9860
2: tissue-specific regulator of an adipocyte enhancer. Genes
Dev 8:1224–1234 2 regulates adipose expression of the phosphoenolpyruvate
carboxykinase gene. Mol Cell Biol 15:351–357[Abstract]
and PPAR
activators direct a distinct tissue-specific transcriptional response
via a PPRE in the lipoprotein lipase gene. EMBO J 15:5336–5348[Medline]
and stimulates
adipogenesis. Genes Dev 9:2350–2363 during the conversion of 3T3
fibroblasts into adipocytes is mediated by C/EBPß, C/EBP, and
glucocorticoids. Mol Cell Biol 16:4128–4136[Abstract]
. Mol Endocrinol 9:108–120). J Biol Chem 270:12953–12956, growth
hormones, and a tyrosine phosphatase inhibitor in rabbit primary
mammary epithelial cells. J Biol Chem 270:20952–20961 is a negative regulator
of macrophage activation. Nature 391:79–82[CrossRef][Medline]
promotes monocyte/macrophage differentiation and uptake of
oxidized LDL. Cell 93:241–252[CrossRef][Medline]
(PPAR) signaling pathways.
Growth hormone inhibition of PPAR transcriptional activity mediated
by stat5b. J Biol Chem 274:2672–2681This article has been cited by other articles:
 |
|
 |
|
  F. Fang, M. A. Rycyzyn, and C. V. Clevenger Role of c-Myb during Prolactin-Induced Signal Transducer and Activator of Transcription 5a Signaling in Breast Cancer Cells Endocrinology, April 1, 2009; 150(4): 1597 - 1606. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 E. R. Hugo, D. C. Borcherding, K. S. Gersin, J. Loftus, and N. Ben-Jonathan Prolactin Release by Adipose Explants, Primary Adipocytes, and LS14 Adipocytes J. Clin. Endocrinol. Metab., October 1, 2008; 93(10): 4006 - 4012. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 N. Ben-Jonathan, C. R. LaPensee, and E. W. LaPensee What Can We Learn from Rodents about Prolactin in Humans? Endocr. Rev., February 1, 2008; 29(1): 1 - 41. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. C. Hogan and J. M. Stephens The Regulation of Fatty Acid Synthase by STAT5A Diabetes, July 1, 2005; 54(7): 1968 - 1975. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. Fleenor, J. Oden, P. A. Kelly, S. Mohan, S. Alliouachene, M. Pende, S. Wentz, J. Kerr, and M. Freemark Roles of the Lactogens and Somatogens in Perinatal and Postnatal Metabolism and Growth: Studies of a Novel Mouse Model Combining Lactogen Resistance and Growth Hormone Deficiency Endocrinology, January 1, 2005; 146(1): 103 - 112. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S Viengchareun, H Bouzinba-Segard, J-P Laigneau, M-C Zennaro, P A Kelly, A Bado, M Lombes, and N Binart Prolactin potentiates insulin-stimulated leptin expression and release from differentiated brown adipocytes J. Mol. Endocrinol., December 1, 2004; 33(3): 679 - 691. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. Yamasaki, T. Nakashima, A. Kawakami, T. Miyashita, F. Tanaka, H. Ida, K. Migita, T. Origuchi, and K. Eguchi Cytokines regulate fibroblast-like synovial cell differentiation to adipocyte-like cells Rheumatology, April 1, 2004; 43(4): 448 - 452. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. A. Shang and M. J. Waters Constitutively Active Signal Transducer and Activator of Transcription 5 Can Replace the Requirement for Growth Hormone in Adipogenesis of 3T3-F442A Preadipocytes Mol. Endocrinol., December 1, 2003; 17(12): 2494 - 2508. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. Sun, A. C. Nicholson, D. P. Hajjar, A. M. Gotto Jr., and J. Han Adipogenic differentiating agents regulate expression of fatty acid binding protein and CD36 in the J744 macrophage cell line J. Lipid Res., October 1, 2003; 44(10): 1877 - 1886. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. R. Kappeler, Z. Farah, and Z. Puhan 5'-Flanking Regions of Camel Milk Genes Are Highly Similar to Homologue Regions of Other Species and Can be Divided into Two Distinct Groups J Dairy Sci, February 1, 2003; 86(2): 498 - 508. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Meirhaeghe, L. Fajas, F. Gouilleux, D. Cottel, N. Helbecque, J. Auwerx, and P. Amouyel A Functional Polymorphism in a STAT5B Site of the Human PPAR{gamma}3 Gene Promoter Affects Height and Lipid Metabolism in a French Population Arterioscler Thromb Vasc Biol, February 1, 2003; 23(2): 289 - 294. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. Nanbu-Wakao, Y. Morikawa, I. Matsumura, Y. Masuho, M.-a. Muramatsu, E. Senba, and H. Wakao Stimulation of 3T3-L1 Adipogenesis by Signal Transducer and Activator of Transcription 5 Mol. Endocrinol., July 1, 2002; 16(7): 1565 - 1576. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L.-Y. Yu-Lee Prolactin Modulation of Immune and Inflammatory Responses Recent Prog. Horm. Res., January 1, 2002; 57(1): 435 - 455. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Frasor, K. Park, M. Byers, C. Telleria, T. Kitamura, L.-y. Yu-Lee, J. Djiane, O.-K. Park-Sarge, and G. Gibori Differential Roles for Signal Transducers and Activators of Transcription 5a and 5b in PRL Stimulation of ER{alpha} and ER{beta} Transcription Mol. Endocrinol., December 1, 2001; 15(12): 2172 - 2181. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L.-y. Yu-Lee Stimulation of interferon regulatory factor-1 by prolactin Lupus, October 1, 2001; 10(10): 691 - 699. [Abstract] [PDF]
|
|
|
|
|
|
M. Freemark, D. Fleenor, P. Driscoll, N. Binart, and P. A. Kelly Body Weight and Fat Deposition in Prolactin Receptor-Deficient Mice Endocrinology, February 1, 2001; 142(2): 532 - 537. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
| Molecular Endocrinology | Recent Prog. Horm. Res. | All Endocrine Journals |
