Involvement of STAT5 (Signal Transducer and Activator of Transcription 5) and HNF-4 (Hepatocyte Nuclear Factor 4) in the Transcriptional Control of the hnf6 Gene by Growth Hormone
Olivier Lahuna1,2,
Mojgan Rastegar2,
Dominique Maiter,
Jean-Paul Thissen,
Frédéric P. Lemaigre and
Guy G. Rousseau
Hormone and Metabolic Research Unit (O.L., M.R., F.P.L.,
G.G.R.) Christian de Duve Institute of Cellular
Pathology Unité de Diabétologie (D.M., J.-P.T.)
Université catholique de Louvain B-1200 Brussels, Belgium
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ABSTRACT
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HNF-6 is a tissue-restricted transcription factor
that participates in the regulation of several genes in liver. We
reported earlier that in adult rats, HNF-6 mRNA concentration in liver
drops to almost undetectable levels after hypophysectomy and returns to
normal after 1 week of GH treatment. We now show that this results from
a rapid effect of GH, and we characterize its molecular mechanism. In
hypophysectomized rats, HNF-6 mRNAs increased within 1 h after a
single injection of GH. The same GH-dependent induction was reproduced
on isolated hepatocytes. To determine whether GH regulates
hnf6 expression at the gene level, we studied its promoter.
DNA binding experiments showed that 1) the transcription factors STAT5
(signal transducer and activator of transcription 5) and HNF-4
(hepatocyte nuclear factor 4) bind to sites located around -110 and
-650, respectively; and 2) STAT5 binding is induced and HNF-4 binding
affinity is increased in liver within 1 h after GH injection to
hypophysectomized rats. Using transfection experiments and
site-directed mutagenesis, we found that STAT5 and HNF-4 stimulated
transcription of an hnf6 gene promoter-reporter construct.
Furthermore, GH stimulated transcription of this construct in cells
that express GH receptors. Consistent with our earlier finding that
HNF-6 stimulates the hnf4 and hnf3ß gene
promoters, GH treatment of hypophysectomized rats increased the liver
concentration of HNF-4 and HNF-3ß mRNAs. Together, these data
demonstrate that GH stimulates transcription of the hnf6
gene by a mechanism involving STAT5 and HNF-4. They show that HNF-6
participates not only as an effector, but also as a target, to the
regulatory network of liver transcription factors, and that several
members of this network are GH regulated.
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INTRODUCTION
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Hepatocyte nuclear factor (HNF)-6 is the prototype of a new class
of cut-homeoproteins conserved from Caenorhabditis elegans
to humans (1, 2, 3). These proteins are transcription factors that contain
a single cut domain and a divergent homeodomain. Two HNF-6 isoforms,
(465 residues) and ß (491 residues), have been cloned in the rat.
They differ only by the length (27 or 53 amino acids) of the linker
between the cut domain and the homeodomain (2). These two isoforms
originate from the same gene by differential splicing (4). They differ
in affinity for DNA target sequences, but both behave as
transcriptional activators in transient transfection assays (2). The
hnf6 gene is strongly expressed in the liver (1).
Transfection experiments performed with HNF-6
showed that it
stimulates the transcription of liver-expressed genes that code for
proteins such as 6-phosphofructo-2-kinase, an enzyme involved in
glucose metabolism (1), CYP2C12, an enzyme of steroid metabolism (5),
transthyretin, a plasma transport protein (2), protein C, which
controls coagulation (6), and the transcription factors HNF-4 and
HNF-3ß (7, 8). These two transcription factors are involved in the
differentiation of hepatocytes and the maintenance of liver-specific
functions (9). Thus, the HNF-6 family participates to the regulatory
network of factors that controls liver development and
differentiation.
How the hnf6 gene is regulated is therefore an important
issue. One candidate is GH. Indeed, we found in adult rats that the
liver concentration of HNF-6 mRNAs drops dramatically after
hypophysectomy and returns to normal after administration of GH (5).
The aim of the present work was to determine whether this effect
results from a direct action of GH on the hepatocyte and on the
hnf6 gene. We show here that this is the case. We also
identify transcription factors that mediate the effect of GH on the
hnf6 gene promoter. Our data show that GH rapidly induces
the binding of signal transducer and activator of transcription (STAT)5
and increases the binding of HNF-4 to the hnf6 gene
promoter. This results in a stimulation of the promoter. Finally, we
provide evidence that GH controls the network of liver-enriched
transcription factors and that HNF-6 participates not only as an
effector, but also as a target, to this regulatory network.
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RESULTS AND DISCUSSION
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GH Induces Liver HNF-6 mRNAs by a Direct Effect on Hepatocytes
In rat liver, HNF-6 mRNAs almost disappear after hypophysectomy.
When these rats are treated with GH and their liver RNA is analyzed
after 1 week of continuous treatment, HNF-6 mRNAs have returned to
normal levels (5). We therefore determined the time-course of liver
HNF-6 mRNAs induction by GH in vivo. Total RNA was extracted
from the liver of hypophysectomized rats before, or at several time
intervals after, a single injection of GH. HNF-6 mRNAs were quantified
by a ribonuclease (RNase) protection assay using
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA as a reference.
In untreated hypophysectomized animals, HNF-6
mRNA was barely
detectable and HNF-6ß mRNA was below the threshold of sensitivity of
the assay (Fig. 1A
), consistent with the
fact that the concentration of HNF-6ß mRNA in the liver of intact
rats is lower than that of HNF-6
mRNA (1). Within 1 h after the
injection of GH, the concentrations of HNF-6
and -ß mRNAs
increased 6-fold, to reach 50-fold after 3 h and return to basal
levels by 9 h after the injection (Fig. 1
, A and B).

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Figure 1. Effect of GH on HNF-6 mRNAs Concentration in Rat
Liver and in Isolated Hepatocytes
A, RNase protection assays were performed on RNA from the liver of
hypophysectomized rats, before (0 h) or after (at the times indicated)
a single injection of GH, with the HNF-6 and GAPDH riboprobes and size
markers (first three lanes). The other lanes each
correspond to a sample from a different rat (four animals per
experimental condition). B, Quantitation of relative HNF-6 mRNA
concentration determined in the experiments described in panel A. **,
P < 0.01 and ***, P < 0.001
vs. the untreated (0 h) group. C, RNase protection
assays were performed (see panel A) on RNA from hepatocytes (four 60-mm
plates) incubated with GH (500 ng/ml) for the times indicated. RNA from
the liver of an intact male rat was included as a positive control
(lane 1). D, Relative concentration of HNF-6 mRNA determined by
RNase protection assay (see panel C) in hepatocytes incubated for
24 h with different concentrations of GH. Data are means ±
SEM for three separate experiments.
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We next investigated whether the GH-dependent regulation of liver HNF-6
mRNAs observed in vivo is a direct effect of the hormone on
hepatocytes. To do so, hepatocytes were isolated from intact rats and
cultured on a matrix to maintain the expression of liver-specific
functions (10). After 48 h, GH (500 ng/ml) was added and the cells
were harvested at different time points for quantifying HNF-6 mRNAs as
above. As shown in Fig. 1C
, HNF-6
and ß mRNAs concentration
increased between 30 min and 1 h after addition of GH, to reach a
maximum after 2 h. The experiment was repeated with different
concentrations of GH. This yielded a typical dose-response curve (Fig. 1D
), with a near-maximal response at a concentration of 50 ng/ml (2
nM) GH, which is physiological. We concluded that
the hnf6 gene is expressed at a low level in the absence of
GH and that GH increases the concentration of liver HNF-6 mRNAs by a
direct effect on the hepatocyte.
Involvement of STAT5 in the Transcriptional Stimulation of the
hnf6 Gene by GH
The experiments reported above suggested that GH directly
stimulates the transcription of the hnf6 gene in liver. This
was consistent with the observed coordinate effect of GH on HNF-6
and HNF-6ß mRNA (Fig. 1
), which both originate from the same gene
(4). Upon binding to its receptor at the surface of the hepatocyte, GH
induces receptor dimerization and association with the tyrosine kinase
Jak-2. The latter phosphorylates the receptor, which then serves as a
docking site for STAT factors. These become phosphorylated, dimerize,
and bind to regulatory regions of GH-responsive genes (11). We
therefore searched for such regions in the hnf6 gene
promoter. Nuclear extracts were prepared from the rat livers that were
used to demonstrate an effect of GH on HNF-6 mRNAs (see Fig. 1A
). These
extracts were incubated with labeled fragments of the hnf6
gene promoter to conduct deoxyribonuclease I (DNase I) footprinting
assays. These experiments showed GH-dependent protein binding to the
region from -105 to -124 of the promoter (Fig. 2A
). This footprint was not seen with
liver extracts from hypophysectomized rats. The footprint appeared
within 1 h of GH treatment and had disappeared after 6 h
(Fig. 2A
). The underlying sequence, TTCTAAGAA (from -116 to -108), is
compatible with the binding consensus for STAT factors (12), among
which STAT1, STAT3, and STAT5 are known to mediate several actions of
GH in the liver (13).

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Figure 2. GH Induction of STAT5 Binding to the
hnf6 Gene Promoter
A, DNase I footprinting on the rat hnf6 gene with
liver nuclear extracts obtained from hypophysectomized rats at the
times indicated after a single GH injection and incubated without or
with 50 ng of a STAT-binding oligonucleotide (GRR). B, EMSA with the
labeled fragment from -98 to -126 of the hnf6 gene
promoter as a probe (H6STAT) without or with 50 ng of the competing
cold oligonucleotides or 1.5 µl of the antisera indicated. The
nuclear extracts were the same as in panel A.
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To confirm that the protein involved in the GH-induced footprint was a
member of the STAT family, a labeled oligonucleotide corresponding to
the footprinted region and containing the hnf6 gene fragment
-126 to -98 was used as a probe in electrophoretic mobility shift
assay (EMSA) with the nuclear extracts that were used in the
footprinting experiments. As shown in Fig. 2B
, the protein-DNA complex
was seen exclusively with the liver extracts obtained from rats that
had been killed 1 or 3 h after GH injection (lanes 3 and 4),
consistent with the footprinting data. This complex disappeared with
excess of cold probe (lane 8) or cold STAT-binding oligonucleotide
(lane 9), but not with cold Sp1-binding oligonucleotide (lane 10).
Moreover, the migration of the complex was retarded by an anti-STAT5,
but not by an anti-STAT3, antibody (lanes 11 and 12). We concluded from
these experiments that GH induces the binding of STAT5 to the
hnf6 gene promoter within 1 h.
To test the role of STAT5 in the control of the hnf6 gene
promoter, we overexpressed a constitutively active form of STAT5
(STAT5
750VP16Jak2) in transfected cells. This chimeric protein
includes the receptor-binding (SH2) and DNA-binding domains of STAT5
and the transactivation domain of VP16 fused to the kinase domain of
Jak2, which ensures the phosphorylation-dependent dimerization and
nuclear translocation required for gene targeting of the chimera (14).
The cells were cotransfected with a luciferase reporter gene linked to
0.75 kb of the hnf6 promoter. As shown in Fig. 3A
, constitutively active STAT5
stimulated transcription from the hnf6 gene promoter. This
did not occur when the cells were transfected with a transcriptionally
inactive form of STAT5 (STAT5
750Jak2), which lacks the
transactivation domain of VP16, or with a reporter construct in which
the STAT5-binding site in the hnf6 promoter had been
destroyed by mutation (Fig. 3A
).
Since GH induces STAT5 binding to the hnf6 gene promoter and
STAT5 stimulates the expression of a reporter gene linked to this
promoter, then GH should stimulate this reporter construct in
transfection experiments. To verify this prediction, we used BRL-4
hepatoma cells stably transfected with the rat GH receptor (15). When
these cells were transiently transfected with the hnf6
promoter-reporter construct, luciferase activity increased 1.8-fold
(0.001 < P < 0.01) after exposure to rat GH
(Fig. 3B
). The amplitude of this GH effect was comparable to that
reported in the same cells for another GH-inducible gene (15). Our
experiments with the hnf-6 promoter-reporter construct also
showed that the STAT-binding site in this promoter is absolutely
required for the stimulation by GH, as GH had no effect when this
binding site had been destroyed (Fig. 3B
).
The demonstration that GH can transactivate the hnf6 gene
promoter and that this depends on the integrity of the STAT-binding
site allowed us to provide evidence for the role of STAT5 in the GH
effect. BRL-4 cells were now transiently cotransfected with the
hnf6 reporter construct and an expression vector for
wild-type STAT5. As shown in Fig. 3B
, overexpression of STAT5 clearly
amplified the stimulation of hnf6 promoter activity by GH.
Overexpressed STAT5 had little effect in the absence of GH, which is
consistent with the notion that the transcriptional action of STAT
factors requires their ligand (i.e. GH)-dependent
phosphorylation by Jak proteins. The amplification of the GH effect by
exogenous STAT5 did not occur with an expression vector lacking the
transactivation domain of STAT5 (Fig. 3B
). Taken together, our data
demonstrate that GH can stimulate transcription of the hnf6
gene through an induction of STAT5 binding to its cognate site in the
hnf6 promoter.
Involvement of HNF-4 in the Transcriptional Stimulation of the
hnf6 Gene by GH
Another DNase I footprint, from -633 to -670, was detected in
the hnf6 gene promoter with liver nuclear extracts (Fig. 4A
). This region encompasses a sequence,
CGGGCAAAGGCCA (-652 to -640), compatible with the binding consensus
for HNF-4 (4, 16). To identify the protein involved in this footprint,
EMSA were performed with the corresponding oligonucleotide probe and
with the liver extracts used to demonstrate GH-dependent STAT5 binding
to the hnf6 promoter. The data (Fig. 4B
) indeed showed
specific binding of HNF-4, as demonstrated by competition with the cold
probe (lane 8) and with an HNF-4-binding oligonucleotide (lane 9), but
not with an Sp1-binding oligonucleotide (lane 10). A supershift was
observed with an anti-HNF-4 antibody (lane 11). A complex exhibiting
the same properties was seen in EMSA with this hnf6 promoter
probe when using, instead of liver extracts, extracts from Cos-7 cells
that had been transfected with an HNF-4 expression vector (data not
shown). Consistent with the footprinting data (Fig. 4A
, lanes 2 and 3),
HNF-4 binding to the hnf-6 promoter did not depend on GH
(Fig. 4B
, lane 2). However, HNF-4 binding increased strongly within the
hour after the injection of GH and returned to uninduced levels between
6 and 12 h (Fig. 4B
, lanes 37).

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Figure 4. Binding of the HNF-4 Protein to the
hnf6 Gene Promoter and Effect of GH
A, DNase I footprinting on the rat hnf6 gene with liver
nuclear extracts obtained from hypophysectomized rats before or 3
h after a single GH injection. B, EMSA with the labeled fragment from
-633 to -657 of the hnf6 gene promoter as a probe (PH4)
without or with 50 ng of the competing cold oligonucleotides or 1 µl
of a 1:5 dilution of the antiserum indicated. The nuclear extracts were
the same as for the experiments shown in Fig. 2 . C, Immunoblot, with an
anti-HNF-4 antibody, of the extracts used in panel B.
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In earlier studies we showed that GH treatment of
hypophysectomized rats increases the concentration of HNF-6 protein in
liver (5) and that HNF-6 binds to, and stimulates the activity of, the
hnf4 gene promoter in transiently transfected cells (8).
Thus, it was likely that GH increases the expression of the
hnf4 gene. To test this, we measured the concentration of
HNF-4 mRNA in rat liver after a single injection of GH to
hypophysectomized animals. The data in Fig. 5
, A and B, show that liver HNF-4 mRNA
increased only slightly. This took place 6 h after the GH
injection and returned to uninduced levels by 9 h. Obviously, this
late and minor change in HNF-4 mRNA concentration could not account for
the rapid and intense GH-induced increase in HNF-4 binding. Therefore,
the signaling cascade triggered by GH in rat liver most probably
targets the HNF-4 protein itself, thereby increasing its stability or
its affinity for the hnf6 gene promoter. To investigate
these possibilities, we determined by immunoblotting the concentration
of HNF-4 in the nuclear extracts used for testing HNF-4 binding after
GH treatment. As shown in Fig. 4C
, the concentration of HNF-4 did not
change over the 6 h that followed the GH injection, a period
during which HNF-4 binding increased dramatically. Thus, GH treatment
confers to HNF-4 a higher affinity for its DNA target in the
hnf-6 promoter. The fact that HNF-4 mRNA had returned to
normal levels between 6 and 9 h after the injection of GH (Fig. 5B
) is consistent with the uninduced levels of HNF-4 protein seen by
immunoblotting after 12 and 24 h (Fig. 4C
). We concluded from
these experiments that HNF-4 binds to the hnf6 promoter and
that this interaction is increased by GH treatment.

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Figure 5. Effect of GH on HNF-4 and HNF-3ß mRNA
Concentration in Rat Liver
A, Northern blotting of the liver RNA samples obtained as described in
Fig. 1A . The two lanes per timepoint correspond to samples from
different rats. B and C, Relative concentration of HNF-4 mRNA (panel B)
and HNF-3ß mRNA (panel C) determined by Northern blotting. Values
obtained by densitometry were corrected for variations in RNA loading
with reference to the 18 S rRNA values. Data shown in panels B and C
are means ± SEM for four rats. *,
P < 0.05 and **, P < 0.01
vs. the untreated (0 h) group.
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We therefore determined by transfection the functionality of this HNF-4
binding site in the context of the intact hnf6 gene
promoter. As shown in Fig. 6A
, the
activity of the hnf6 promoter-luciferase reporter construct
was stimulated by the cotransfection of an HNF-4 expression vector.
This effect was abolished when the HNF-4-binding site in the
hnf6 promoter was destroyed by mutation (Fig. 6A
). These
experiments were repeated with BRL-4 cells to evaluate the effect of GH
under these conditions (Fig. 6B
). The 1.8-fold increase in wild-type
promoter activity induced by GH was reduced to 1.3-fold (0.001<
P< 0.01) after destruction of the HNF-4 binding site. The
residual activity was probably due to the activation by GH of
endogenous STAT5. We could not test this with a promoter in which both
the STAT5 and the HNF-4 sites had been destroyed, as destruction of the
STAT5 site alone abolishes the response to GH (see Fig. 3B
).
Overexpression of HNF-4 in the BRL-4 cells amplified 2.4-fold (0.001<
P <0.01) promoter activity in the absence of GH, and this
effect was lost after destruction of the HNF-4 binding site (Fig. 6B
).
This GH-independent effect of HNF-4 is consistent with our
demonstration by footprinting and by EMSA (Fig. 4
) that, unlike for
STAT5, the binding of HNF-4 to the hnf6 gene promoter does
not depend on GH. In keeping with this interpretation, GH did not
significantly stimulate the promoter after transfection of HNF-4,
presumably because overexpressed HNF-4 saturated its binding site on
the promoter, without need for the GH-induced increase in binding
affinity that takes place with endogenous HNF-4. These results,
together with the binding data, show that HNF-4 participates in, but is
not essential to, the stimulatory action of GH on the
transcription of the hnf6 gene.

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Figure 6. Stimulation of the hnf6 Gene
Promoter by HNF-4
Rat-1 cells (A) or BRL-4 cells (B) were cotransfected transiently with
the wild-type or mutated hnf6 promoter-reporter
constructs and with an HNF-4 expression vector, as indicated. See
legend of Fig. 3A for other details. Data are means ±
SD for at least three experiments. In panel B the cells
were exposed for 24 h to 100 nM rat GH (solid
bars).
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Implications for the Control of the Hepatic Network of Liver
Transcription Factors
Liver transcription factors are organized in a network that
involves reciprocal as well as autoregulatory positive and negative
feedback loops (9, 17). We had shown earlier that HNF-6 belongs to this
network and that it stimulates transcription of the hnf3ß
gene in transfected cells (1, 8). The data presented here demonstrate
that GH stimulates transcription of the hnf6 gene. In this
way, GH could control the expression of the hnf3ß gene
in vivo. To investigate this possibility, we measured the
concentration of HNF-3ß mRNA in the liver of hypophysectomized rats
treated or not with a single injection of GH. As shown in Fig. 5
, A and
C, the concentration of HNF-3ß mRNA was increased 3 h after the
GH injection, and it decreased later to values below those in untreated
rats. The time course of induction of HNF-3ß mRNA is compatible with
an HNF-6-mediated effect of GH, since hnf6 gene expression
increases 6-fold within the hour after the injection of GH (see Fig. 1B
).
In conclusion, our data demonstrate that GH signaling stimulates the
expression of the hnf6 gene in liver by a direct action on
the hepatocyte. They provide evidence that the effect of GH described
here involves 1) GH-dependent binding of STAT5 to the promoter and a
GH-induced increase in the affinity of HNF-4 for the promoter, and 2)
increased transcription from the hnf6 gene promoter. The
mechanism by which GH triggers STAT5 binding to target genes is well
documented. In contrast, a GH-dependent increase in the affinity of
HNF-4 for a DNA target sequence has not, to our knowledge, been
reported before. Consistent with its rapid kinetics, this effect could
result from GH-induced tyrosine phosphorylation, e.g. by
Jak2, of HNF-4. Indeed, HNF-4 is a tyrosine phosphoprotein, and
dephosphorylation of its tyrosine residues in vivo or
in vitro decreases its affinity for DNA (18). We cannot
exclude the possibility that regions of the hnf6 gene other
than the one studied in this paper are involved in the stimulatory
effect of GH on hnf6 gene expression seen in
vivo. Still, the STAT binding site identified here was absolutely
necessary for the transcriptional stimulation of the hnf6
gene promoter by GH. An identical phenomenon has been described for the
ß-casein gene, whose transcriptional activation by GH depends on
STAT5 binding (19).
The experiments described in this paper show that GH controls the
network of hepatocyte transcription factors in three ways. First, GH
increases the transcription of the hnf6 gene by the
mechanisms discussed above. Second, GH increases the affinity of HNF-4
for DNA. Whether this holds true for HNF-4 target sequences other than
the one studied here remains to be established. Third, GH increases the
amount of HNF-3ß mRNA. Since HNF-6, HNF-4, and HNF-3ß in turn
control the transcription of a number of genes, the latter might in
this way be indirectly regulated by GH. Insofar as these three
transcription factors are tissue-restricted, they should have a key
role in the tissue specificity of the action of GH on gene
expression.
Finally, the demonstration that HNF-4 controls the transcription of the
hnf6 gene adds a new loop to the regulatory network of liver
transcription factors. This, together with the fact that HNF-6
stimulates the activity of the hnf4 gene promoter in
transfected cells (8), would predict the existence of a positive
feedback mechanism involving HNF-4 and HNF-6. However, such an
autoregulatory loop must be kept in check in vivo by
negative control mechanisms. These might explain why, in the animal, GH
treatment leads to a spectacular increase in hnf6 gene
expression, but only to a modest increase in hnf4 gene
expression. The sharp drop in liver HNF-6 mRNA concentration after
3 h was not surprising in view of the existence of mechanisms that
rapidly terminate GH-induced STAT5 signaling (20, 21). Experiments on
transfected cells and on embryoid bodies have shown that HNF-3ß can
induce HNF-3
, which inhibits the HNF-4 gene both directly, and
indirectly via inhibition of the expression of HNF-1, which is an
inducer of the hnf4 gene (17, 22). In these ways, induction
of HNF-3ß by HNF-6 could eventually turn off the hnf4
gene. The kinetics and cell type specificity of such positive and
negative regulatory loops should be taken into account to fully
understand how GH controls gene expression.
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MATERIALS AND METHODS
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Animals
Four-week-old male Wistar rats obtained 7 days after
hypophysectomy (IFFA-Credo, Lyon, France) were maintained under
standardized conditions of light and temperature with free access to
rat chow and to water containing 0.9% (wt/vol) NaCl. Hypophysectomized
rats were pretreated during 7 days with a daily subcutaneous injection
of L-T4 (Aldrich,
Milwaukee, WI) (1 µg/100 g body weight) and cortisol hemisuccinate
(Pharmacia & Upjohn, Inc., Kalamazoo, MI) (50
µg/100 g body weight). On day 8, groups of hypophysectomized rats
(four per group) were killed before (0 h) or after (at the time
indicated) a single subcutaneous injection of purified rat GH (NIDDK
rat pituitary hormone distribution program) (100 µg/100 g body
weight). These experiments were conducted in accordance with the
highest standards of humane animal care.
Cell Cultures and Transfections
For hepatocyte cultures, matrigel was prepared (23) from
Engelbreth-Holm-Swarm sarcoma cells propagated in C57BL/6 female mice.
Hepatocytes were obtained by nonrecirculating collagenase perfusion
through the portal vein of normal male rats anesthetized with
pentobarbital (6 mg/100 g body weight), as described (24). Cells were
seeded at a density of 1.8 x 106 per 60-mm
plates and incubated for 48 h at 37 C in 5%
CO2 in serum-free DMEM/Hams F-12 medium
supplemented with penicillin (100 U/ml) and streptomycin (100 µg/ml),
hydrocortisone (50 nM), insulin (175 nM),
L-ornithine (0.4 mM), L-lactic acid
(17.7 µM), selenium (25 nM), and ethanolamine
(1 µM). After 48 h of culture, cells were incubated
with rat GH as indicated. Cos-7 cells were cultured and transfected as
described (3). Rat-1 cells, kindly provided by J. Wyke, were grown in
DMEM with 10% FCS. For transient transfection, 3 x
105 cells were plated on 60-mm dishes and
incubated with N-[1-(2,
3-dioleoyloxy)propyl]-N,N,N-triethyl-ammonium methylsulfate
(DOTAP, Roche Molecular Biochemicals, Indianapolis,
IN) and 7 µg of reporter construct, 50 ng to 2 µg of
expression vector, and 1 µg of pRL138 as internal control. After
16 h, the cells were washed with PBS and further incubated for
24 h before luciferase activities were measured with the
Dual-Luciferase kit (Promega Corp., Madison, WI) and a
TD20/20 Luminometer (Promega Corp.). The data were
expressed as the ratio of firefly luciferase (reporter activity) to
Renilla luciferase (internal control).
BRL-4-GHR1-638 cells (15), from a rat hepatoma
cell line stably transfected with the rat GH receptor cDNA and kindly
provided by G. Norstedt, were grown in DMEM with 10% FCS and cultured
to confluence in six-well plates. Transient transfection was performed
in serum-free DMEM with DOTAP according to the manufacturers
instructions. One microgram of reporter plasmid and 1 µg of pRL138 as
internal control were transfected per well. After 8 h the medium
was changed to serum-free DMEM with or without 100
nM rat GH, and the cells were further incubated
for 24 h.
Detection of mRNA
RNase protection assays were performed as described (1) with 20
µg of total RNA isolated from individual livers or from cultured
hepatocytes by the guanidine thiocyanate/cesium chloride method (25)
and with an HNF-6 probe (290 b) that allows detection of two
specifically protected fragments (254 and 215 b) originating from
HNF-6
and HNF-6ß mRNA, respectively. A rat GAPDH antisense RNA
probe (Ambion, Inc., Beverly, MA) was cohybridized (4
x 104 cpm) with the HNF-6 probe (300 x
104 cpm) as an internal reference to correct for
variations in RNA concentration. After digestion with RNase and
separation of the protected fragments on a 6% polyacrylamide
denaturing gel, the GAPDH and HNF-6 mRNAs were quantified with a
PhosphorImager (Molecular Dynamics, Inc., Sunnyvale,
CA). For Northern blot analysis, total RNA (20 µg) was size
fractionated on a denaturing 1% agarose gel and transferred to nylon
membranes (Hybond-N, Amersham Pharmacia Biotech,
Buckinghamshire, UK) by overnight vacuum blotting (VacuGeneXL blotting
system, Pharmacia LKB, Uppsala, Sweden). After UV
cross-linking (Stratalinker, Stratagene, La Jolla, CA),
the membranes were hybridized to 32P-labeled
probes specific for rat HNF-3ß or HNF-4. HNF-3ß and HNF-4 mRNAs
were detected with random-primed rat cDNA fragments containing
nucleotides 947-2218 (26) and 614-1424 (27), respectively. Each
Northern blot was rehybridized with a 32P-labeled
oligonucleotide specific for 18 S rRNA to correct for variations in RNA
concentration. The relative concentration of mRNA in each lane was
quantified by scanning autoradiograms (8-day exposure at -80 C with
two intensifying screens) with a LKB Ultroscan XL
laser densitometer (Pharmacia Biotech, Uppsala, Sweden).
Results were expressed by assigning a value of 1 arbitrary
densitometric unit to liver mRNA from hypophysectomized rats killed at
time 0. Statistical analysis was performed with an ANOVA test.
DNase I Footprinting and EMSAs
For the DNase I footprint of STAT5, a
BamHI-ClaI fragment of the rat hnf6
gene promoter was labeled on one strand at the BamHI site
(-44) with 32P [
-dGTP] and cleaved at the
SacI site (-323). For the DNase I footprint of HNF-4, a
XhoI-StuI fragment of the rat hnf6
gene promoter was labeled on one strand at the XhoI site
(-196) with 32P[
-dGTP] and cleaved at the
StuI site (-756). The incubations, which contained 15 µg
of rat liver nuclear protein and the probe (100150 counts per
second), were carried out as described previously (28) and were
followed by analysis on 6% polyacrylamide-8 M
urea sequencing gels. For EMSA, nuclear extracts were prepared from rat
livers as described (29). The following double-stranded
oligonucleotides were used as probes: PH4,
5'-GCGAACGGGCAAAGGCCATGGCATA-3' (from -657 to -633 of the rat
hnf6 gene promoter); HNF-4,
5'-AAGGCTGAAGTCCAAAGTTCAGTCCCTTC-3' (HNF-4-binding oligonucleotide,
-71 to -43 of the rat hnf1 gene promoter); H6STAT,
5'-GGCAGCAGGATTCTAAGAAAGAGAGGGGC-3' (-126 to -98 of the rat
hnf6 gene promoter); GRR, 5'-ATGTATTTCCCAGAAA-3'
(STAT-binding oligonucleotide from the Fc
RI gene
promoter) (30); Sp1, 5'-ATTCGATCGGGGCGGGGCGAGC-3' (Promega Corp.). They were labeled with
[
32P]-ATP (Amersham Pharmacia Biotech) by T4 polynucleotide kinase (United States Biochemical Corp., Cleveland , OH) and EMSAs were performed as
described (5). Antibodies were added to the liver nuclear extracts on
ice 45 min before addition of the labeled probe. Incubation with the
labeled probe was then allowed to proceed for 45 min on ice before
electrophoresis. The anti-STAT3 and anti-STAT5 antibodies were from
Santa Cruz Biotechnology, Inc., (Santa Cruz, CA), and the
anti-HNF-4 antibody was kindly provided by M. Pontoglio.
Immunoblotting
Liver nuclear extracts (20 µg of protein) from
hypophysectomized male rats that had received a single injection of GH
as indicated were loaded on an 8% acrylamide gel. After SDS-PAGE the
proteins were transferred to a polyvinylidene fluoride membrane
(Amersham Pharmacia Biotech) that was incubated overnight
with the anti-HNF-4 antiserum (1:15,000) used for EMSA.
Protein-antibody complexes were visualized using the Enhanced
Chemiluminescence Detection System of Roche Molecular Chemicals
(Indianapolis, IN).
Expression Vectors and Reporter Constructs
The expression vectors STAT5
750JAK2,
STAT5
750VP16-JAK2, pXM-MGF-STAT5, and
pXM-MGF-STAT5
750 were kindly given by B. Groner. The expression
vector HNF-4
1 was kindly given by B. Laine. The reporter construct
pNF/0.75 luc has been described previously (4). pRL138, used as an
internal control, contains the pfk2 gene promoter (-138 to +86) cloned
in pRLnull (Promega Corp.). The pNF/0.75 (HNF-4 mut)luc
was made by PCR amplification with two sets of primers (the
mutated oligonucleotides are underlined): PH6IIIS,
5'-TTGTGAGGGTCATGGATACCAGTTCTA-3' (-803 to -777 of the rat
hnf6 gene promoter, sense strand) and GAH4NAS,
5'-AAAAGTACTCCGCCATTGGGCTTTATTCCC-TGG-3' (the 3'-end
corresponds to -678 of the rat hnfh6 gene promoter,
antisense strand), GAH4S,
5'-AAAAGTACTGTCCTCCGATGGCATAGTCTCCAGCTCC-3' (the 3'-end
corresponds to -621 of the rat hnf6 gene promoter, sense
strand) and SacAS, 5'-CCGCTGCCCACCCTCACGCCC-3' (-273 to - 253
of the rat hnf6 gene promoter, antisense strand). The first
fragment was digested with ScaI and
StuI, while the second fragment was digested with
SacI and ScaI. The digested fragments were gel
purified and ligated with pNF/0.75 luc opened at the StuI
and SacI sites. In this way the HNF-4 site was replaced with
a GAL-4 binding site. To prepare the pNF/0.75 (STATmut)luc construct,
two PCR reactions were carried out. The first PCR was done with the
primers H6STATM,
5'-CTCGCCCCTCTCTTGAATTCAATCCTGCTGCCCCC-3'
(-129 to -95 of the rat hnf6 gene promoter, antisense
strand) and HNF-63'Sac, 5'-CTACCGAATCTCAGCCACAG-3' (-240 to -221 of
the rat hnf6 gene promoter, sense strand). In the second PCR
the amplified fragment from the first PCR was used as a primer with GL
Primer 2, 5'-CTTTATGTTTTTGGCGTCTTCC-3' (standard primer of pGL3 basic
vector of Promega Corp.). The product of this PCR was
digested with XhoI and cloned in the XhoI site of
pNF/0.75 luc.
 |
ACKNOWLEDGMENTS
|
---|
We would like to thank P. Lause and M. A. Gueuning for
their excellent technical assistance, P. Vandoolaeghe for help with
preparation of nuclear extracts, T. Lambert and V. O Connor
for secretarial work, and all the colleagues mentioned in the text for
their generous gifts.
 |
FOOTNOTES
|
---|
Address requests for reprints to: Guy G. Rousseau, HORM 7529, 75 Avenue Hippocrate, B-1200 Brussels, Belgium.
This work was supported by grants from the Belgian State Program on
Interuniversity Poles of Attraction, Prime Ministers Office, Federal
Office for Scientific, Technical and Cultural Affairs; from the
Délégation Générale Higher Education and
Scientific Medical Research, French Community of Belgium; from the Fund
for Scientific Medical Research (Belgium); from the National Fund for
Scientific Research (Belgium); from the Fonds de Développement
Scientifique (Louvain University); and from the Danone Institute
(Belgium). J.-P.T is Research Associate and F.P.L. is Senior Research
Associate of the National Fund for Scientific Research (Belgium).
1 Present address: Unité INSERM 135, Hôpital
de Bicêtre, 78 Rue du Général Leclerc, 94275 Le
Kremlin Bicêtre, France. 
2 Both authors have contributed equally to this work. 
Received for publication July 21, 1999.
Accepted for publication November 16, 1999.
 |
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