Growth Hormone- and Prolactin-Induced Proliferation of Insulinoma Cells, INS-1, Depends on Activation of STAT5 (Signal Transducer and Activator of Transcription 5)
Birgitte Nissen Friedrichsen,
Elisabeth Douglas Galsgaard,
Jens Høiriis Nielsen and
Annette Møldrup
The Hagedorn Research Institute Department of Cell Biology
2820 Gentofte, Denmark
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ABSTRACT
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GH and PRL stimulate proliferation and insulin
production of pancreatic ß-cells. Whereas GH- and PRL-regulated
transcription of the insulin gene in insulinoma cells has been shown to
depend on STAT5 (signal transducer and activator of transcription 5),
the signaling pathways involved in GH/PRL-induced ß-cell replication
are unknown. The roles of various signaling pathways in human GH
(hGH)-induced DNA synthesis were studied by analysis of the effect of
specific inhibitors in both the insulin-producing cell line, INS-1, and
in primary ß-cells. The mitogen-activated protein kinase kinase
(MEK)-inhibitor, PD98059, as well as the mitogen-activated protein
kinase p38 (MAPKp38) inhibitor, SB203580, partially inhibited hGH-
induced proliferation in INS-1 cells but had no significant effect
in primary ß-cells. Staurosporine, a protein kinase C (PKC) and
protein kinase A (PKA) inhibitor, blocked both basal and hGH-induced
proliferation in INS-1 cells, but had no inhibitory effect in primary
ß-cells. Wortmannin, a phosphatidylinositol 3-kinase (PI3K)
inhibitor, inhibited hGH-induced proliferation neither in INS-1 cells
nor in primary ß-cells, whereas the tyrosine kinase inhibitor,
genistein, completely inhibited hGH- induced proliferation in both
primary ß-cells and INS-1 cells. To analyze the possible role of
STAT5 in hGH-induced proliferation, a dominant negative STAT5 mutant,
STAT5
749, was expressed in INS-1 cells under the control of a
doxycycline- inducible promoter by stable transfection. Two clones
were found to exhibit dose-dependent, doxycycline-inducible expression
of STAT5
749 and suppression of hGH-stimulated transcriptional
activation of a STAT5-regulated PRL receptor (PRLR) promoter-reporter
construct. Furthermore, induction of STAT5
749 expression completely
inhibited hGH-induced DNA synthesis. Analysis of endogenous gene
expression revealed a doxycycline-dependent inhibition of
hGH-stimulated PRLR and cyclin D2 mRNA levels. Our results suggest that
GH/PRL-induced ß-cell proliferation is dependent on the Janus Kinase2
(JAK2)/STAT5 signaling pathway but not the MAPK, PI3K, and PKC
signaling pathways. Furthermore, the cell cycle regulator cyclin D2 may
be a crucial target gene for STAT5 in this process.
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INTRODUCTION
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The hormones of the GH family, GH , PRL, and placental lactogen
(PL), are the most potent growth factors that have been identified for
pancreatic ß-cells to date (for review see Refs. 1, 2). In
addition, these hormones stimulate insulin production, partially via
direct transcriptional activation of the insulin promoter. These
effects are mediated through the GH-receptor (GHR) and the PRL-receptor
(PRLR) (3), which are both expressed in rat islets as well as in
various insulinoma cell lines (4, 5, 6). The GHR and PRLR are
transmembrane proteins belonging to the cytokine receptor superfamily
(7). These receptors are characterized by the ability to activate STAT
(signal transducers and activators of transcription) proteins, which
are latent transcription factors that become activated by
phosphorylation of a single tyrosine residue by receptor-associated
Janus kinases (JAKs). The phosphorylated STAT proteins dimerize and
translocate to the nucleus where they bind to specific DNA elements and
activate transcription (8). Recently, we identified STAT5 binding
elements in the promoters of the rat insulin 1 (9) and PRLR genes (10)
that are required for transcriptional activation by GH and PRL. Thus,
GH and PRL were found to stimulate DNA binding of STAT5a and STAT5b
and, to a lesser extent, STAT1 and STAT3 in the insulinoma
cell lines, RIN-5AH (9) and INS-1 (10). Apart from the JAK/STAT
pathway, several other signaling pathways are activated in various cell
types in response to GH and PRL stimulation (for review see Refs. 8, 11, 12). Thus, the ras/raf/MAPK (mitogen-activated protein kinase)
cascade is activated in some cell types, presumably involving the
adaptor protein complex, SHC/Grb2/SOS (SH2-containing protein/growth
factor receptor-bound protein/Son-of-Sevenless-1), which may bind to
either tyrosine-phosphorylated JAK2 or insulin receptor substrates
(IRS) 1 and/or 2. Phosphatidylinositol-3'- kinase (PI3K) may also
be activated via recruitment by tyrosine-phosphorylated IRS, leading to
downstream activation of the 70-kDa-S6-kinase
(p70S6K). Furthermore, activation of protein
kinase C (PKC), which may involve phospholipase C and increased
diacylglycerol (DAG) levels, or PI3K, has been reported for both GH and
PRL. The signaling pathways involved in GH/PRL-induced ß-cell
mitogenesis have not been identified. Activation of MAPK and
p70S6K are known to be required for replication
of many mammalian cell types. However, in INS-1 cells, MAPK activity
was not induced by GH/PRL, but potently by nerve growth factor (NGF)
and glucose (13). Furthermore, whereas insulin-like growth factor-I
(IGF-I) in a recent study was found to activate both MAPK and PI3K
activities in INS-1 cells, GH had no detectable effect on these enzymes
(14). Tyrosine residues in the intracellular domain of the GHR and PRLR
have been found to be required for STAT5 activation and transcriptional
signaling (15, 16). However, studies in the mouse promyeloid
interleukin-3-dependent cell line, FDC-P1, showed that the
intracellular tyrosine residues of the GHR were not essential for
mitogenic signaling in these cells (17, 18). Similar results have been
reported for the PRLR (19), and it has been hypothesized that STAT5
activation is mainly involved in the regulation of differentiated gene
expression whereas mitogenic signaling may primarily depend on
activation of STAT1 and 3, which are recruited to and activated by JAK2
and/or JAK2-mediated MAPK activation (20). Recently however, STAT5 has
been implicated in regulation of cell cycle progression of peripheral
lymphocytes (21, 22, 23) and lymphoma cell lines (24, 25, 26, 27). In the present
study we have examined the role of the MAPK, PI3K, PKC, and JAK2/STAT5
signaling pathways in GH/PRL-induced cell proliferation using metabolic
inhibitors of these enzymes in INS-1 cells and in primary ß-cells
from neonatal rat islets. The role of STAT5 was evaluated by inducible
overexpression of a dominant negative mutant of STAT5 in INS-1
cells.
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RESULTS
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Analysis of Signaling Pathways That May Lead to Human GH
(hGH)-Induced Proliferation in INS-1 Cells and Primary ß-Cells
To determine whether the MAPK, the PI3K, or PKC signaling pathways
are involved in hGH-induced ß-cell proliferation, the effect of
specific protein kinase inhibitors on hGH-stimulated DNA synthesis was
examined in INS-1 cells and in primary ß-cells in monolayer culture
by 3H-thymidine and bromodeoxy uridine (BrdU)
incorporation assays, respectively, as previously described (1, 28).
After a 24-h stimulation period with 0.5 µg/ml of hGH,
3H-thymidine incorporation into INS-1 cells was
increased by 2.0 ± 0.1-fold over the basal level (Fig. 1A
), and BrdU incorporation into primary
ß-cells was increased by 2.7 ± 0.4-fold over the basal level
(Fig. 1B
). Human GH-induced DNA synthesis was completely inhibited in
both INS-1 and primary ß-cells (1.1 ± 0.1-fold and 0.9 ±
0.1-fold, respectively, over the basal levels) in the presence of 25
µM genistein, an inhibitor of protein tyrosine kinase
activity, as would be expected from inhibition of JAK2. In the presence
of the MEK inhibitor, PD98059 (20 µM), and the p38 MAPK
inhibitor, SB203580 (10 µM), hGH-induced DNA synthesis
was partially reduced in INS-1 cells (1.4 ± 0.1-fold and 1.4
± 0.1-fold, respectively, over basal levels), whereas these two
inhibitors had no significant effect in primary ß-cells. Wortmannin
(10 nM), a specific PI3K inhibitor, had no effect on
hGH-induced DNA synthesis in either INS-1 or in primary ß-cells,
whereas staurosporine (20 nM), an inhibitor of PKC, protein
kinase A (PKA), and protein kinase G (PKG), blocked both basal (not
shown) and hGH- induced proliferation of INS-1 cells (0.2 ±
0.1-fold of basal) but did not inhibit proliferation of primary
ß-cells. On the contrary, staurosporine was found to potentiate DNA
synthesis in primary cells. Addition of the various inhibitors in the
absence of hGH (data not shown) had no effect on the basal
proliferation level in any of the cell types with the exception of
staurosporine, as described above.

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Figure 1. The Effect of Protein Kinase Inhibitors on
hGH-Stimulated DNA Synthesis
A, INS-1 cells were cultured for 24 h in the absence or presence
of 0.5 µg/ml hGH. Before addition of hGH, respective cultures
were supplemented with 25 µM genistein, 20
µM PD98059, 10 µM SB203580, 10
nM wortmannin, or 20 nM staurosporine. The
cells were labeled 4 h with 3H-thymidine
before harvesting and counting. The results are expressed as fold
induction compared with control levels (mean ± SEM,
n = 4). B, Precultured monolayers of primary ß-cells were
incubated for 24 h with 10 µM BrdU and also in the
absence or presence of hGH and inhibitors as described above. The cells
were fixed and stained for BrdU and insulin. The results are expressed
as the ratio between BrdU-labeled ß-cells and the total amount of
ß-cells (mean ± SEM, n = 5). A total of 1,000
cells were counted for each preparation. *, P 0.05;
**, P 0.001; ***, P 0.0001,
compared with the group treated with hGH alone.
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hGH Induces Prolonged Activation of STAT5 in INS-1 Cells and in
Cultured Newborn Rat Islets
The kinetics of hGH-induced STAT5 activation in INS-1 cells and
primary newborn rat islets in suspension culture were determined by
electrophoretic mobility shift analysis (EMSA) (Fig. 2
). INS-1 cells and islets cultured in
0.5% serum were either left untreated or stimulated with 0.5 µg/ml
hGH for 15 min to 24 h. Nuclear extracts were prepared and
incubated with radiolabeled, double-stranded oligonucleotide (1A-GLE)
representing the previously identified STAT5-binding element of the
PRLR 1A promoter (10). Free and bound probe was separated by
nondenaturing PAGE and visualized by autoradiography. Nuclear extracts
from INS-1 cells that had been stimulated with hGH for 15 min
(upper panel, lane 2) produced a strong shifted band, which
was also present in incubates with nuclear extracts from INS-1 cells
treated for 1, 4, and 24 h with hGH (upper panel, lanes
3, 4, and 5, respectively). In contrast, this complex was absent in
incubates with nuclear extracts from untreated INS-1 cells (upper
panel, lane 1). A similar pattern was observed with nuclear
extracts from newborn rat islets (lower panel, lanes 15),
although the intensity of the complex was somewhat reduced after
prolonged hGH treatment (
3050% after 4 and 24 h).

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Figure 2. hGH-Induced STAT5 Activation in INS-1 Cells and
Newborn Rat Islets
Analysis of hGH-induced activation of STAT5 was performed by EMSA. The
radiolabeled probe, 1A-GLE, was incubated with 10 µg nuclear extracts
prepared from either INS-1 cells (upper panel) or
newborn rat islets (lower panel) that were either
unstimulated (lane 1) or stimulated with 0.5 µg/ml hGH for 15 min,
1 h, 4 h, and 24 h (lanes 25, respectively). Free and
bound probe were separated by nondenaturing gel electrophoresis and
visualized by autoradiography. The autoradiograph shown is
representative of three independent experiments. The
arrows indicate the migration of the 1A-GLE*STAT5
complex.
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Inducible Overexpression of a Truncated STAT5a Mutant
(STAT5a
749) in INS-1 Cells with Dominant Negative Activity
To address whether GH/PRL-stimulated activation of STAT5 is
involved in the mitogenic response of ß-cells to these hormones, the
Tet-On gene expression system was employed to generate stably
transfected INS-1 cells expressing, in an inducible manner, a STAT5
mutant, STAT5a
749, which has been reported to exert dominant
negative activity (29). An INS-1 clone, INS-r3, stably transfected with
an expression plasmid encoding the reverse
tetracycline/doxycycline-dependent transactivator (30), was transfected
with either the expression vector, pTRE, without insert or the
pTRE-vector containing a cDNA encoding STAT5a
749 under the control
of the tetracycline operator. Cotransfection of an expression vector
containing a hygromycin resistance gene allowed for selection of stably
transfected clones, which were tested for DNA integration of the pTRE
plasmids by PCR analysis. Twenty-five of 38 clones examined were found
to contain integrated pTRE vector without insert, and 2 of 69 clones
examined contained a full-length insert of STAT5a
749.
Expression of STAT5a
749 in the two transfected clones (termed BB32
and EB03) was determined by Western blot analysis (Fig. 3A
). The cells were cultured for 20
h in the absence or presence of 0.051.0 µg/ml doxycycline, and
total cell lysates were examined for STAT5 expression using an antibody
raised against the amino acids (aa) 451649 of STAT5. In the absence
of doxycycline (lanes 1 and 6) only wild-type STAT5 protein was
detectable, which in this experiment probably represents STAT5b, since
Western blot analysis, using a more potent antibody recognizing the C
terminus part of STAT5, indicates that STAT5b is approximately 45
times more abundant than STAT5a in these cells (data not shown). In
both cell lines, addition of 0.05 µg/ml doxycycline induced the
expression of a protein of faster mobility than wild-type STAT5, likely
to represent STAT5a
749 (lanes 2 and 7). The effect of doxycycline
was found to be dose dependent with maximal expression observed at a
concentration of 0.1 µg/ml doxycycline for both BB32 and EB03 (lanes
25 and 710, respectively). Doxycycline had no effect on STAT5
protein expression in a control clone (termed BA15) stably transfected
with pTRE without insert (data not shown).

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Figure 3. Inducible Expression and Dominant Negative Activity
of STAT5a 749 in INS-1 Cells
A, Western blot analysis was performed as described in Materials
and Methods on total protein extracts prepared from stably
STAT5a 749-transfected clones, BB32 and EB03 (STAT5a 749
transfected) that had been cultured for 24 h in the absence (lanes
1 and 6) or presence of the indicated amounts of doxycycline (lanes
25 and 710). Forty microliters of each extract were separated on a
7.5% SDS polyacrylamide gel, and proteins were transferred to
nitrocellulose. STAT5 protein was visualized using STAT5 antibody
raised against the aa 451649 of STAT5 in a 1:1000 dilution. B, The
radiolabeled oligonucleotide probe, 1A-GLE, was incubated with nuclear
extracts from either the BA15 (lanes 14), the BB32 (lanes 58), or
the EB03 cells (lanes 916) that had been cultured for 24 h in
the absence or presence of the indicated amounts of doxycycline and in
the absence or presence of 0.5 µg/ml hGH 15 min before harvesting. Free and
bound probe were separated by nondenaturing gel electrophoresis and
visualized by autoradiopgraphy. The autoradiograph shown is
representative of two independent experiments. The arrow
indicates the migration of the 1A-GLE*STAT5/STAT5a 749 complex. C,
INS-1 cells, BA15, BB32, and EB03 were transiently transfected with the
firefly luciferase reporter gene construct containing the PRLR 1A
promoter and the internal control construct, pRL-SV40, which contain
the Renilla luciferase gene. The cells were cultured for
24 h in the absence or presence of the indicated amounts of
doxycycline. Seven hours before harvest the cells were cultured in the
absence or presence of 0.5 µg/ml hGH. The results are expressed as
ratios between the firefly and the Renilla luciferase
activities (mean ± SEM, n = 3). Significant
differences between the groups treated with hGH+Dox and the group
treated with hGH alone are indicated: *, P 0.05;
**, P 0.001; ***, P
0.0001. A significant difference (P
0.00010.05) between the hGH-treated groups and the control
groups ± Dox was found in all four cell lines.
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To determine the hGH-induced DNA-binding activity of STAT5a
749, EMSA
was performed as described above. The stably transfected clones, BA15,
BB32, and EB03, were cultured for 24 h in the absence or presence
of doxycycline and stimulated the last 15 min of the culture period
with 0.5 µg/ml hGH. Nuclear extracts of the cells were prepared and
incubated with radiolabeled 1A-GLE oligonucleotide probe, the mobility
of which was examined by gel electrophoresis. hGH treatment alone was
found to induce 1A GLE binding of nuclear protein in all the cell lines
(Fig. 3B
, lanes 2, 6, and 10), and this binding was unaffected by the
addition of doxycycline in BA15 cells (Fig. 3B
, lane
4). In BB32 and EB03 cells, a doxycycline-dependent
increase of DNA complex formation was observed (Fig. 3B
, lanes 8 and
1216) likely to represent binding of STAT5a
749. A
promoter-reporter gene assay (10) was used to address whether
overexpression of STAT5a
749 would suppress signaling via
endogenously expressed STAT5 (Fig. 3C
). The parental and the three
transfected lines were transiently transfected with the firefly
luciferase reporter gene under the transcriptional control of the
5'-flanking region of the PRLR gene containing a STAT5-responsive
element. hGH stimulation induced transcriptional activity of the PRLR
promoter in INS-1 cells by 2.9 ± 0.1-fold and in BA15 cells by
5.5 ± 0.8-fold (upper panel), in BB32 cells by
4.6 ± 0.4-fold (middle panel) and in EB03 cells by
17.7 ± 0.3-fold (lower panel) over the basal levels.
The addition of 0.5 µg/ml doxycycline affected neither the basal nor
the hGH-induced transcriptional activity in INS-1 and BA15 cells.
However, addition of doxycycline to BB32 and EB03 cells increased the
basal transcriptional level by 1.8 ± 0.3-fold and a 4.3 ±
1.3-fold, respectively (middle and lower panel).
The reason for this stimulatory activity is not known. However,
hGH-induced transcriptional activity in both BB32 and EB03 was
inhibited by the induction of STAT5a
749 in a doxycycline
dose-dependent manner, which was significant in BB32 cells at a
concentration of 0.25 µg/ml and in EB03 cells at a concentration of
0.1 µg/ml.
Induction of STAT5a
749 Results in Inhibition of hGH-Stimulated
DNA Synthesis in INS-1 Cells
To determine the influence of dominant negative STAT5 activity on
cell proliferation, 3H-thymidine incorporation
assay was carried out, as described above, on the transfected clones
(Fig. 4
). hGH stimulated
3H-thymidine incorporation by approximately
2-fold in the two transfected clones (1.9 ± 0.1 and 2.4 ±
0.4 fold over the basal levels in BB32 and EB03 cells, respectively)
similar to what was found in INS-1 cells and BA15 cells (data not
shown). The addition of 1 µg/ml doxycycline had no effect on either
the basal or the hGH-induced DNA synthesis in INS-1 and BA15 cells
(data not shown). Neither was the basal level in BB32 (upper
panel) and EB03 (lower panel) affected by doxycycline
addition alone. However, induction of STAT5a
749 by doxycycline in
BB32 and EB03 blocked hGH-induced 3H-thymidine
incorporation in a dose-dependent manner, whereas no effect of
doxycycline was observed on the growth response to 10% FCS. The
inhibition of 3H-thymidine incorporation was
maximal at a doxycycline concentration of 0.5 µg/ml for both the BB32
cells and the EB03 cells, at which close to basal levels of DNA
synthesis were detected (1.25 ± 0.03-fold and 1.1 ±
0.2-fold over basal in BB32 and EB03 cells, respectively).
To be able to determine whether the suppression of
3H-thymidine incorporation by dominant negative
STAT5 expression was due to an effect on S-phase entry, the cell cycle
profiles of hGH-stimulated parental INS-1 cells and transfected cells
were examined by fluorescence-activated cell sorting (FACS) analysis
(Table 1
) using BrdU incorporation and
DNA staining with propidium iodide. Stimulation with hGH for 24 h
was found to increase the percentage of cells in S-phase by
approximately 2-fold in both INS-1 cells and the transfected clones
(INS-1, 2.5 ± 0.2; BA15, 1.9 ± 0.3; BB32, 1.8 ± 0.1;
and EB03, 2.5 ± 0.3) (Table 1
, middle column). The
percentage of cells in the G2/M phase was not
affected by hGH stimulation (Table 1
, third column), whereas
the percentage of cells in the
G0/G1 phase was found to
decrease upon hGH stimulation, indicating that hGH primarily in these
experiments increases the
G0/G1-to-S phase transition
frequency. Neither the basal nor the hGH-induced cell cycle profiles in
INS-1 and BA15 cells were affected by the addition of 0.5 µg/ml
doxycycline. Furthermore, addition of doxycycline alone did not affect
the cell cycle profiles in either BB32 or EB03 cells. However, the
hGH-induced increase of cells in S-phase was totally blocked by the
induction of STAT5a
749 expression in both BB32 and EB03 cells.
Suppression of hGH-Induced PRLR 1A and Cyclin D2 mRNA Levels by
Overexpression of STAT5a
749
The effects of STAT5a
749 overexpression on the endogenous mRNA
levels of the PRLR 1A and the cell cycle regulators, cyclin D1, D2, and
D3, were investigated by quantitative RT-PCR in BA15, BB32, and EB03
cells (Fig. 5
). The PRLR mRNA levels were
increased by hGH in BA15 cells by 7.4 ± 0.4-fold (upper
panel) in BB32 by 4.1 ± 0.5-fold (middle panel),
and in EB03 cells by 4.4 ± 0.4-fold (lower panel),
respectively, in accordance with previous results in INS-1 cells (10).
Addition of doxycycline alone had no effect on either the basal or the
hGH-induced PRLR mRNA level in BA15 cells. Neither was the basal level
affected by doxycycline in BB32 and EB03 cells. However,
doxycyline-induced STAT5a
749 expression in BB32 and EB03
significantly suppressed the hGH-induced PRLR mRNA levels in a
dose-dependent manner at a concentration of 0.05 µg/ml for both BB32
and EB03. At 0.5 µg/ml doxycycline, the induction by hGH was reduced
to 0.8 ± 0.03-fold in BB32 cells and to 1.6 ± 0.2-fold in
EB03 cells, supporting the previous finding of a primary role of STAT5
in the regulation of the PRLR 1A promoter.

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Figure 5. Effect of STAT5a 749 Expression on hGH-Stimulated
mRNA Levels of the PRLR and Cyclin D1, D2, and D3
The three clones, BA15, BB32, and EB03, were cultured in the absence or
presence of the indicated amounts of doxycycline for 48 h.
Twenty-four hours before harvesting, 0.5 µg/ml hGH was added to the
respective wells. cDNA was synthesized from total RNA, and RT-PCR was
performed as described in Materials and Methods with
primer sets specific for the PRLR exon 1A, and cyclin D1, D2, and D3.
Primer sets specific for G6PDH or TBP were included as internal
controls. The PCR products were separated by denaturing polyacrylamide
gel electrophoresis and visualized and quantified by phosphoimager
analysis. The PRLR 1A and the cyclin D1 and D3 PCR products were
quantified relative to the internal standard TBP. The basal relative
expression range in the three different clones were for PRLR1A,
2588% of TBP; for cyclin D1, 99281%; and for cyclin D3,
95207%. The cyclin D2 products were quantified relative to G6PDH and
ranged from 370942% of G6PDH in the three untreated clones. The
results are expressed as fold induction (mean ±
SEM, n = 3) compared with control levels and
represents the mean of three independent cDNA preparations. Significant
differences between the groups treated with hGH+Dox and the group
treated with hGH alone are indicated: * P 0.05;
**, P 0.001; ***, P
0.0001. In all the cell lines, a significant difference (P 0.00010.001) was
detected between the hGH-treated groups and the control groups ±
Dox for the PRLR, cyclin D1, and cyclin D2 mRNA levels, respectively.
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Examination of cyclin D mRNA levels revealed that cyclin D2 was
positively regulated by hGH in all the cell lines (4.5 ± 0.5-fold
in BA15, 3.7 ± 0.2-fold in BB32, and 4.2 ± 0.4-fold in
EB03) over the basal expression levels which were 513% ± 64, 513% ±
35, and 565% ± 154 of G6PDH (glucose-6-phosphate-dehydrogenase),
respectively (mean ± SEM, n = 3). Whereas
doxycycline treatment had no effect on hGH-induced cyclin D2 mRNA
levels in BA15 cells, a dose-dependent decrease was observed in both
BB32 and EB03 cells, which was significant at 0.05 µg/ml in both BB32
and in EB03 cells. At 0.5 µg/ml doxycycline, the hGH-stimulated
cyclin D2 mRNA levels were reduced to approximately 50% and 20%, in
BB32 and EB03, respectively. The cyclin D3 mRNA levels (142% ± 14,
126% ± 14, and 163% ± 20 of TBP (TATA-binding-protein) in
unstimulated BA15, BB32, and EB03 cells, respectively; means ±
SEM, n = 3) were affected by neither hGH stimulation
nor doxycycline treatment in all three clones. However, hGH
significantly decreased the expression of cyclin D1 (0.42 ±
0.01-fold in BA15, 0.55 ± 0.06 in BB32, and 0.86 ± 0.05 in
EB03) below basal levels which were 207% ± 39, 153% ± 25, and 230%
± 21, respectively (mean ± SEM, n = 3) of TBP
mRNA levels, and treatment with doxycycline seemed to potentiate this
effect in both BB32 cells (reduced to 0.1 ± 0.005) and EB03 cells
(reduced to 0.4 ± 0.01) in a dose-dependent manner, whereas
doxycycline treatment had no effect on cyclin D1 mRNA levels in BA15
cells. The basal mRNA expression levels of the three cyclin Ds were
markedly different. Based on the estimation that G6PDH mRNA is
approximately 25 times more abundant than TBP mRNA, the cyclin D2 mRNA
level is 50- to 100-fold higher than that of cyclin D1 and D3.
Immunocytochemical Detection of Cyclin D2 in INS-1 Cells and
Primary ß-Cells
Cyclin D2 protein expression was examined by peroxidase staining
in INS-1 cells (Fig. 6
, panel A) and
primary ß-cells cultured in monolayer (Fig. 6
, panel B). The INS-1
cells were cultured 4 days in the presence of 10% FCS, and the primary
ß-cells were stimulated with hGH for 24 h before fixation and
staining using a monoclonal anticyclin D2 antibody. Cyclin D2 protein
was detected in the nucleus of a subpopulation of INS-1 cells and
primary ß-cells.

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Figure 6. Immunocytochemical Detection of Cyclin D2 in INS-1
Cells and in Primary ß-Cells
A, INS-1 cells were seeded and cultured 4 days in complete medium,
after which they were fixed and cyclin D2 expression was detected by
peroxidase staining. B, Primary newborn rat islet-cells were cultured
in monolayer for 57 days in the presence of 0.5 µg/ml hGH. The
cells were deprived of hGH for 24 h and then restimulated for a
further 24-h period before fixation and peroxidase staining. The
staining shown is representative of two independent experiments.
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DISCUSSION
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Little is known about the postreceptor signaling of GH and PRL in
proliferating insulin-producing cells. In primary neonatal ß-cells,
stimulation of either GHRs or PRLRs using rat GH
and rat PRL, respectively, leads to the same maximal effect on
proliferation as when both receptors are activated using hGH,
indicating that GHRs and PRLRs share downstream signaling molecules
involved in proliferation (3). In INS-1 cells, cAMP was found to play a
permissive role in hGH-induced proliferation, whereas hGH-stimulated
increases of intracellular Ca2+ levels are
probably not involved (28). In a recent study, rat GH (rGH)-stimulated
INS-1 cell proliferation was found to be glucose dependent, and
inhibitor experiments indicated involvement of the PI3K pathway in this
effect whereas MAPK activation did not appear to play a role, in
accordance with previous findings (13, 14). The INS-1 cell line has
proven useful for the study of the effects of secretagogues, IGF-I and
GH/PRL (13, 14, 31), and the role of tissue-specific transcription
factors (32). Although it exhibits a mitogenic response comparable to
that of primary rat ß-cells in culture, it is not known to what
extent the mitogenic signaling apparatus in these cells is
perturbed.
In the present study we examined the signaling pathways that may play a
role in hGH-induced mitogenic signaling in INS-1 cells as well as in
primary neonatal rat ß-cells. At 11 mM glucose, hGH was
found to increase DNA synthesis by 2- to 3-fold over a 24-h stimulation
period in both cell types in accordance with previous findings (1, 28).
Inhibition of tyrosine protein kinase activity by coincubation with
genistein resulted in a complete inhibition of hGH-induced
proliferation in both cell types, as would be expected from inhibition
of JAK2 and, thereby, most downstream signaling from both the GHR and
the PRLR. PKC is involved in cell cycle progression in mammalian cells
during the G1-phase and
G2/M phase, and both positive and negative
regulatory effects of PKC on cell growth have been reported (33).
Staurosporine, an inhibitor of PKC, PKA, and protein kinase G, was
found to inhibit basal as well as hGH-induced proliferation in INS-1
cells, but not in the primary ß-cells in which, on the contrary, a
tendency for a growth-potentiating effect was observed. This
paradoxical effect may reflect an important difference in the
growth-signaling pathway between the primary ß-cells and the tumor
cell line, INS-1. Inhibition of MAPK signaling pathways using the MEK
inhibitor PD98059 and the p38 MAPK inhibitor SB203580 was only
partially inhibitory to hGH-induced mitogenic signaling in INS-1 cells
but did not affect primary ß-cells. Furthermore, we found no
indications of hGH-stimulated activity of these enzymes by Western blot
analysis of protein extracts using antiphosphothreonine-phosphotyrosine
(anti-pTpY)-specific antibodies against these proteins (data not
shown). These data confirm previous negative effects of this pathway in
GH/PRL-stimulated INS-1 cell proliferation as mentioned above. The
partial effect of the MAPK inhibitors may relate to a role of p42 and
p38 MAPK in the glucose-induced proliferation of these cells (13, 34)
and/or the glucose dependence of GH-induced INS-1 cell proliferation
reported by Cousin et al. (14). The PI3K pathway seems to
play an essential role in glucose- and IGF-I-stimulated INS-1
proliferation (31). However, inhibition of this pathway using
coincubation with wortmannin did not influence hGH-induced
proliferation in either INS-1 cells or primary ß-cells cultured in 11
mM glucose. This correlates well with the finding
that rGH is unable to mediate the association of IRS-1, IRS-2, Grb2, or
mSOS with PI3K (14) and the previous report that the mitogenic effects
of IGF-I and hGH are additive, indicating distinct signaling mechanisms
of these growth factors. Intriguingly, the study by Cousin et
al. found an inhibitory effect of wortmannin on both glucose- and
rGH-stimulated INS-1 cell proliferation in assays in which the cells
had been precultured in the absence of glucose (14). Using these
culture conditions, we have also seen a partial wortmannin-sensitive
component of the hGH response (
50% inhibition, data not shown).
However, in our hands considerable cell death occurs in the absence of
glucose, which may obscure the interpretation of the data. A cautious
conclusion from our inhibitor experiments is that the MAPK, PI3K, and
the PKC signaling pathways, in contrast to findings in other cell
types, are probably not essential for hGH-stimulated ß-cell
replication, and we therefore turned our focus to the STAT5
pathway.
We have previously shown that activation of either GHRs or PRLRs in
INS-1 cells induces binding of both STAT5a and STAT5b to the GLE of the
PRLR 1A promoter (10). Stimulation of both GHRs and PRLRs using hGH led
to a similar STAT5 activation. In the present study the kinetics of
STAT5 activation in hGH-stimulated INS-1 and primary islet cells was
examined. Pronounced STAT5 binding was observed after 15 min that
persisted up to 24 h in both cell types. In INS-1 cells, the STAT5
activity after 24 h was comparable to that of 15 min hGH
stimulation, whereas in islets a partial reduction was observed,
indicating an influence of counterregulatory mechanisms operating in
the primary cells, e.g. phosphatases or members of the
suppressors of cytokine signaling (SOCS) family (35, 36). The long-term
activation of STAT5 DNA binding in hGH-stimulated INS-1 cells
correlates with the prolonged nuclear translocation of STAT5 in these
cells in response to PRL as reported by Sorenson and co-workers
(37). Interestingly, hGH-induced STAT5 DNA binding in the
insulin-producing RIN-5AH cells was found to be only transient (9) with
no detectable changes in STAT5 localization (37). The effects of GH and
PRL on proliferation and PRLR gene expression in RIN-5AH cells are
small compared with the effect of these hormones in INS-1 cells and
cultured newborn rat islets (5, 10, 37, 38). Thus, the potency of GH
and PRL action in insulin-producing cells may be correlated to the
kinetics of GH- and PRL-induced STAT5 activation.
To address the role of STAT5 in GH/PRL-stimulated mitogenic signaling
in ß-cells, we took advantage of a mutant of STAT5a that is deleted
in its C-terminal transactivation domain and reportedly exerts dominant
negative activity by its ability to inhibit the effect of both
wild-type STAT5a and STAT5b (29). This mutant, STAT5a
749, was stably
expressed in INS-1 cells using the inducible Tet-On gene expression
system, and we showed in two different clones (BB32 and EB03) that
STAT5a
749 expression and activation were doxycycline inducible.
Furthermore, dominant negative activity of STAT5a
749 in the two
clones was verified by analysis of the transcriptional effect on
STAT5-regulated PRLR promoter, as previously described (10). The
expression of STAT5a
749 was highly controllable, and there were no
signs of leakage, as determined by Western blotting, EMSA, and reporter
gene assay. Although the STAT5a
749 expressing cells do not have
increased expression of PRLRs, they should still express the initial
amount of GHRs (these are not regulated by STAT5) and the initial
amount of PRLRs, as only one of the three characterized PRLR promoters
contain a STAT5 response element (10). The expression of functional
receptors in the presence of doxycycline was confirmed by the
hGH-induced activation and binding of STAT5a
749 to DNA. Analysis of
the effect of STAT5a
749 expression on DNA synthesis measured by
3H-thymidine incorporation and FACS analysis
revealed a close correlation between STAT5 activity and hGH-stimulated
S-phase entry, and we conclude that STAT5 activation is essential for
GH/PRL-induced proliferation of INS-1 cells. Since previous experiments
show that hGH, rat GH, and rat PRL have the same maximal mitogenic
effect in neonatal rat ß-cells (3), the STAT5-mediated up-regulation
of PRLRs is probably not rate limiting in short-term experiments.
Whether this up-regulation after prolonged stimulation could be
important for the extent of the ß-cell response remains to be
addressed.
D-type cyclins, which regulate the activity of the cyclin-dependent
kinases and S-phase entry, exhibit both cell type-specific and
differential regulation in their expression, suggesting that the
biological functions of these cyclins are not fully redundant (39). In
this study we found that cyclin D2 mRNA was abundant in the INS-1
cells, whereas cyclin D3 and cyclin D1 mRNA were detectable at lower
levels. The role of STAT5 in the growth of lymphoma cells has been
discovered quite recently and, importantly, revealed a direct
transcriptional effect of STAT5 on the human cyclin D1 gene (27).
Furthermore, STAT5-deficient mice lacked peripheral T cell expression
of cyclin D2, cyclin D3, cyclin A, and the cyclin-dependent kinase
(cdk)-6 in response to interleukin-2 (IL-2) (21). In mouse and rat, no
STAT5 binding element is present in the cyclin D1 promoter, whereas the
cyclin D2 and D3 promoters contain PRL-responsive elements (40, 41),
and a potential STAT5 binding sequence is present in the cyclin D2
promoter (Ref. 21 and our unpublished observations). In the present
study we found that hGH increased the mRNA level of cyclin D2 and PRLR
in the stably transfected cell lines, and the doxycycline-
inducible inhibition of this effect indicates that STAT5 is
involved in this regulation. However, whereas expression of the PRLR 1A
mRNA was totally inhibited at the maximal doxycycline dose, only
partial inhibition was observed for cyclin D2, indicating that other
hGH-stimulated signaling pathways play a role in the transcriptional
regulation of cyclin D2. Interestingly, an inhibitory effect of hGH on
cyclin D1 mRNA levels was observed, which was augmented
dose-dependently by doxycycline treatment. Our finding may indicate
that STAT5 is involved in negative regulation of this cyclin through a
mechanism that may involve competition of cofactors in transcription
factor complexes (42). In a recent study a reciprocal regulation of
cyclin D1 and D2 was identified in mouse myeloid leukemia cells in
response to granulocyte-colony-stimulating-factor (G-CSF), indicating a
correlation between G-CSF-stimulated up-regulation of cyclin D1 mRNA
levels and G-CSF-induced apoptosis (43). One may speculate that the
inverse effect of hGH on these cyclins in insulin-producing cells is
related to a combined effect on antiapoptosis and replication.
In conclusion, our study has shown a requirement for STAT5 signaling in
hGH-stimulated proliferation of INS-1 cells, which may involve a direct
transcriptional effect on the cyclin D2 promoter. Future studies using
adenoviral transfer of STAT5 mutants and establishment of transgenic
animals expressing STAT5 mutants under the control of the insulin
promoter will hopefully reveal whether these findings are relevant to
primary ß-cells.
 |
MATERIALS AND METHODS
|
---|
Cells, Hormones, and Chemicals
INS-1 cells, kindly provided by Dr. C. B. Wollheim, Geneva,
Switzerland, were cultured in RPMI 1640 with glutamax supplemented with
10% heat-inactivated FCS, 100 U/ml penicillin, 100 µg/ml
streptomycin, and 50 µM ß-mercaptoethanol (complete
medium) at 37 C in a humidified atmosphere containing 5%
CO2. INS-r3 cells were cultured in the media
described above plus 100 µg/ml G418. For stably transfected clones,
100 µg/ml hygromycin B (Calbiochem, La Jolla, CA) was
additionally added. Islets were isolated from 3- to 5-day-old Wistar
rats by the collagenase digestion method (44) and cultured until
further processing in RPMI 1640 supplemented with 10% newborn calf
serum (NCS), 100 U/ml penicillin, 100 µg/ml streptomycin, 2
mM L-glutamine, 0.0375%
NaHCO3, and 20 mM HEPES at 37 C in a
humidified atmosphere. Recombinant hGH was obtained from Novo Nordisk A/S (Gentofte, Denmark). Doxycycline was from
Sigma (St. Louis, MO). Wortmannin, PD98059, SB203580,
genistein, and staurosporine were purchased from
Calbiochem. All inhibitors were dissolved in
dimethylsulfoxide (DMSO) (Sigma).
3H-Thymidine Incorporation
Cells were seeded in 96-wells plates (50,000 cells per well) and
cultured for 2 days in 200 µl/well complete medium. The medium was
changed to RPMI 1640 containing 0.25% BSA (Fraction V,
Sigma) and culture proceeded for 24 h. For the
inhibitor assay, various inhibitors (see cells, hormones, and
chemicals) were added followed by stimulation with 0.5 µg/ml hGH for
24 h. For the stable clones, the cells were cultured in the
absence or presence of doxycycline for 2024 h followed by stimulation
with either 0.5 µg/ml hGH or 10% FCS containing medium for an
additional 24-h culture period. The last 4 h before harvesting 0.5
µCi [methyl, 1',2'-3H]Thymidine (no. TRK.565,
Amersham Pharmacia Biotech, Buckinghamshire, UK) was
additionally added per well. Cells were harvested onto a filter paper
(Filtermate 196, Packard Instruments, Meriden, CT) by a cell harvester
(Wallac, Inc., Gaithersburg, MD) using
H2O for lysis. The filter paper was dried for
1 h at 37 C and transferred to a bag containing 5 ml of Optiscent
scintillation fluid (Wallac, Inc.). The filter was counted
in a 1450 Microbeta Plus counter (Wallac, Inc.).
Preparation of Monolayers of ß-Cells and 5-BrdU
Labeling
Monolayer cultures of islet cells were prepared essentially as
previously described (1). Briefly, islets were precultured for 57
days in RPMI 1640, containing 0.5% human serum (HS) and were then
dispersed into single cells by trypsin-EDTA treatment. The cells
(50100,000) were plated in plastic cell culture
9-cm2 slideflasks (Nunc, Roskilde, Denmark) in
RPMI 1640 medium containing 2% HS and 0.5 µg/ml hGH. The cells were
allowed to attach and establish a monolayer for 57 days, after which
they were washed twice in medium without hGH and then cultured for
24 h in RPMI 1640 containing 2% HS. The medium was changed, and
the cells were cultured further for 24 h in the presence of 10
µM BrdU, and in the absence or presence of the various
protein kinase inhibitors, which were added before the addition of 0.5
µg/ml of hGH. The cells were washed twice in RPMI 1640 without serum
before fixation in 1% paraformaldehyde in 0.1 M phosphate
buffer, pH 7.4. The cells were double stained for BrdU and insulin as
described previously (1). Briefly, the cells were exposed to 1.5
M HCl for 30 min and washed. They were stained with a
monoclonal mouse antibody to BrdU (no. M 0744, DAKO Corp.,
Glostrup, Denmark) diluted 1:50 and with guinea pig antiinsulin
antibody (Novo Nordisk A/S) diluted 1:500. The antibodies
were visualized by a Texas red-conjugated goat antimouse-IgG (no.
115075-100, Jackson ImmunoResearch Laboratories, Inc.
West Grove, PA) and fluorescein isothiocyanate
(FITC)-conjugated goat antiguinea pig-IgG (no. 106095-003,
Jackson ImmunoResearch Laboratories, Inc.) both diluted
1:100. Dilution of the antibodies was performed in PBS with 0.3%
Triton X-100 and 0.1% human serum albumin (HSA). The slides were
mounted in 20% glycerol/0.05 M Trisma base adjusted to pH
8.4 and stored at 4 C.
Stable Transfection
The establishment of the stable clone, INS-r3, expressing the
reverse tetracycline-dependent transactivator, has been described
previously (30) and was kindly provided by Dr. P. B. Iynedjian
(Geneva, Switzerland). A vector containing the cDNA encoding the STAT5a
mutant, STAT5a
749, which lacks the C-terminal transactivation
domain, was provided by Dr. B. Groner (Freiburg, Germany). This mutant
cDNA was subcloned into the Tet-On gene expression vector, pTRE
(CLONTECH Laboratories, Inc. Palo Alto, CA) using the
EcoRI restriction site of the polylinker. INS-r3 cells were
seeded (1 x 107 cells per 100-mm dish) and
cultured overnight in complete medium containing 100 µg/ml G418. The
following day the medium was changed to Opti-MEM, and transfection was
carried out using the LipofectAMINE PLUS Reagent (Life Technologies, Inc., Paisley, UK) essentially as described
by the manufacturer. The cells were transfected overnight with 3.7 µg
pTK-Hyg Vector (CLONTECH Laboratories, Inc.) and 18.1 µg
pTRE vector with or with out STAT5a
749 insert. The cells were
cultured for 2 days in complete medium and were then trypsinized and
reseeded in 5 x 100 mm dishes using complete medium containing
100 µg/ml G418 and hygromycin. Medium was changed every fourth day,
and antibiotic-resistant colonies of cells were isolated after 24
weeks, transferred to 24-well plates, and split to single cells by
trypsination. Hygromycin-resistant clones were tested for integration
of pTRE by PCR amplification on purified DNA. The DNeasy Tissue kit
(QIAGEN, Valencia, CA) was used for DNA purification. The
PCR reaction was run with 250 ng of total DNA as a template, and 50
pmol per specific pTRE primers (CLONTECH Laboratories, Inc.), 1.25 U Taq polymerase (Promega Corp., Madison, WI), thermophilic DNA 10x buffer (Promega Corp.), 0.2 mM deoxynucleoside
triphosphate (dNTP) (Amersham Pharmacia Biotech),
1.5 mM MgCl2 (Promega Corp.), and H2O to 50 µl. A single
denaturing step at 94 C/1 min was followed by 25 cycles as given: 94
C/15 sec; 63 C/1 min; 68 C/3 min. The products were detected on 1%
agarose gel.
Western Blot Analysis
Cells were seeded in 100-mm dishes (4 x
106 cells per dish) and cultured for 2 days in 10
ml complete medium per dish. The medium was changed to medium
containing 0.25% BSA, and cells were cultured approximately 20 h
in the presence and absence of doxycyline as indicated. The cells were
washed once in cold PBS, scraped off in 1 ml PBS, transferred to
microfuge tubes, and pelleted. The cells were resuspended in 500 µl
PBS, containing 1% NP40, 0.1% SDS, 1 µg/ml leupeptin, 1 µg/ml
aprotinin, and 0.5 mM AEBSF (Calbiochem) and
allowed to lyse on ice for 30 min. Cell debris was removed by
centrifugation at 15,000 x g for 20 min, and the
supernatants were stored at -80 C after addition of 125 µl 5x
sample buffer. Proteins were denatured by boiling for 2 min and
separated by electrophoresis on a 7.5% SDS polyacrylamide gel. For
protein size determination, High-Range Rainbow Marker (Amersham Pharmacia Biotech) was used. The proteins were transferred to a
nitrocellulose membrane by Western blotting for 2 h at 200 mA. The
membrane was blocked for 15 min and washed once in PBS before probing
with antibodies. The primary antibody was either no. S21520
(Transduction Laboratories, Inc., Lexington, KY) raised
against aa 451649 of STAT5 or (C-17)-G no. SC835 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) raised against the carboxy
terminus of STAT5b diluted 1:1000. The secondary antibody was antimouse
IgG horseradish peroxidase-linked whole antibody (Amersham Pharmacia Biotech) diluted 1:5,000. Proteins were visualized
using the ECL Western blotting detection reagents (Amersham Pharmacia Biotech).
Nuclear Extracts
Isolated newborn rat islets (5,000 islets per dish), which had
been precultured for 1 week in 15 ml of RPMI 1640 supplemented with
0.5% normal HS, were stimulated with 0.5 µg/ml hGH for 15 min before
harvest. INS-1 cells and the three INS-r3 clones (BA15, BB32, and EB03)
were seeded in 100-mm dishes (4 x 106 cells
per dish) and cultured for 2 days in 10 ml complete medium per dish.
The medium was changed to medium containing 0.25% BSA, and cells were
cultured for 20 h in the presence or absence of doxycyline. When
indicated, the cells were stimulated with 0.5 µg/ml hGH for 15 min.
Nuclear extracts were prepared essentially as described previously (9, 10). Briefly, cells were lysed in hypotonic buffer containing 1%
Triton X-100. Nuclei were collected by centrifugation, and nuclear
proteins were extracted in hypertonic buffer containing 400
mM NaCl. After centrifugation, aliquots of the supernatants
were frozen in liquid nitrogen and stored at -80 C. Protein
concentrations were measured using the Bio-Rad protein assay
(Bio-Rad Laboratories, Inc. Hercules, CA).
EMSA
EMSA was performed essentially as described previously (9, 10).
Briefly, the double-stranded oligonucleotide 1A-GLE
(5'-agctAGTTCTAGGAATAagct) containing a STAT5 binding element derived
from the rat PRLR 1A promoter (45) was radiolabeled in a fill-in
reaction using [
-32P] dATP (Amersham Pharmacia Biotech) and DNA polymerase (Klenow fragment) and used
as probe (10). Nuclear extracts (10 µg) were incubated at 30 C with
20 fmol of probe in a 20 µl reaction. Free and bound probe were
separated by nondenaturing PAGE and visualized by autoradiography.
Transient Transfection and Dual Luciferase Reporter (DLR)
Assay
Cells were seeded in 24-well plates (300,000 cells per well) in
500 µl/well complete medium. The cells were transfected as described
previously (10) with 1.5 µl LipofectAMINE PLUS Reagent (Life Technologies, Inc.) and 0.5 µg DNA [250 ng of pGL21A, 10 ng
of pRL-SV40 plasmid (internal control) and 240 ng of pUC18 plasmid].
The cells were transfected overnight in Opti-Mem (240 µl/well). The
medium was changed to RPMI 1640 containing 0.5% FCS (500 µl/well)
and incubated for 24 h in the presence or absence of doxycycline.
Seven hours before harvesting, 0.5 µg/ml hGH was added to the
respective wells. The cells were lysed by adding 100 µl/well of 1x
passive lysis buffer (supplied with DLR Assay System no. E1910,
Promega Corp.) followed by shaking the plate for 15 min at
room temperature. The cell extracts were stored in the plate at -80 C
until measuring was performed as described previously (10). The
pRL-SV40 vector contains the coding region of the Renilla
luciferase gene under the transcriptional control of the SV40 early
enhancer/promoter (Promega Corp.). The pGL21A is
generated by the insertion of the 5'-flanking region of PRLR exon 1A
(-462/+81) into the pGL2-basic vector that contains the coding region
of the firefly luciferase gene (45). The pUC18 vector was used as
carrier plasmid.
FACS
Cells were seeded in six-well plates (500,000 cells per well)
and cultured for 2 days in 3 ml complete medium per well. The medium
was changed to medium containing 0.25% BSA, and cells were cultured
approximately 20 h in the presence and absence of doxycyline as
indicated. Respective wells were stimulated 24 h with 0.5 µg/ml
hGH and 2 h before harvesting, 10 µM BrdU was added
per well. The cells were harvested by adding 100 µl 0.5%
trypsin-EDTA per well. Trypsination was stopped by adding 900 µl
serum-containing medium, and cells were transferred to
Eppendorf (Madison, WI) tubes, pelleted, and resuspended
in 500 µl -20 C 70% ethanol. Cells were stored at 4 C up to 1 week.
For denaturation of DNA, 500 µl of 3 M HCl were added and
the cells were incubated at room temperature for 30 min. The cells were
washed once in PBS containing 0.1% HSA and 0.3% Triton X-100 and were
resuspended in 1 ml of this buffer containing mouse anti-BrdU antibody
(DAKO Corp.) in a 1:100 dilution. Incubation was carried
out overnight at 4 C with rotation. Cells were washed twice,
resuspended in 1 ml buffer containing FITC-conjugated goat-antimouse
IgG no. 115095-003 (Jackson ImmunoResearch Laboratories, Inc.), and incubated for 45 min at 4 C in the dark. The cells
were washed twice, resuspended in 500 µl PBS containing 5 µg/ml
propidium iodide, and placed in the dark. FACS analysis was carried out
using Cell Quest (Becton Dickinson and Co., San Jose, CA)
as software.
RT-PCR
Cells were seeded in 60-mm dishes (2 x
106 cells per dish). The cells were cultured for
2 days in 4 ml complete medium per dish. The medium was changed to
medium containing 0.25% BSA, and cells were cultured approximately
20 h in the presence and absence of doxycyline as indicated.
Respective wells were stimulated 24 h with 0.5 µg/ml hGH. Total
RNA was extracted using the RNeasy method from QIAGEN
(Chatsworth, CA). cDNA was synthesized from 1 µg RNA using AMV
Reverse Transcriptase and dNTP mix from Stratagene (La
Jolla, CA) and random primers from Life Technologies, Inc.
The reaction was run at 42 C for 1 h, and the sample was diluted
in 40 µl of 0.1% Triton-X-100 and stored at -20 C. PCR was carried
out in 12.5 µl reactions using 0.75 µl of cDNA as template. The
primer sequences were: PRLR, 5'-TTG TGG ATC TCA GGT TTC CCT GGT G
(forward); 5'-AGC GAG CTG GAT TCT AGG GAA ACA T (reverse); cyclin D1,
5'-TCT ACA CTG ACA ACT CTA TCC G (forward); 5'-TAG CAG GAG AGG AAG TTG
TTG G (reverse); cyclin D2, 5'- AGA CCT TCA TCG CTC TGT GT (forward);
5'- TAG CAGATG ACG AAC ACG CC (reverse); cyclin D3, 5'-CTG CTG GCG GGA
ATC ACA (forward); 5'-GGC CCC TCC TCT GCT TGG T (reverse); G6PDH,
5'-GAC CTG CAG AGC TCC AAT CAA C (forward); 5'-CAC GAC CCT CAG TAC CAA
AGG G (reverse); TBP, 5'-ACC CTT CAC CAA TGA CTC CTA TG (forward);
5'-ATG ATG ACT GCA GCA AAT CGC (reverse). The expected lengths of the
various PCR products were as follows: PRLR, 329 bp; cyclin D1, 304 bp;
cyclin D2, 372 bp; cyclin D3, 246 bp; G6PDH, 214 bp; TBP, 192 bp. The
PCR incubates contained 50 mM KCl; 10 mM
Tris-HCl; 1.5 mM MgCl2; 40
µM dATP; dGTP; and dTTP; 20 µM dCTP; 2.5
mCi of 3,000 Ci/mmol [
-33P]dCTP; 10 pmol of
each primer and 2.5 U Ampli Tag Gold polymerase. A single denaturing
step at 94 C/10 min was followed by either 20 cycles (cyclin D2) or 25
cycles (PRLR, cyclin D1/D3) as given: 94 C/30 sec; 55 C/1 min; 72 C/1.5
min. PCR products were separated on 6% denaturing polyacrylamide gels
(GEL-MIX 6, Life Technologies, Inc.), dried, and exposed
to Phosphorimage storage screens that were scanned by Phosphorimager
series 400 (Molecular Dynamics, Inc., Sunnyvale, CA), and
band intensities were calculated using the program Image Quant
(Molecular Dynamics, Inc.). The PRLR and the cyclin D1 and
D3 were quantified relative to the internal standard TBP and cyclin D2
was quantified relative to the internal standard G6PDH.
Immunocytochemistry
INS-1 cells were seeded in slide flasks (100,000 cells per
flask) and cultured 4 days in complete medium. Newborn rat islets were
dispersed into single ß-cells and cultured as described above. The
cells were fixed in 1% paraformaldehyde and stained with peroxidase
using the HISTOSTAIN-PLUS KIT (Zymed Laboratories, Inc.
South San Francisco, CA) according to the instructions provided by the
manufacturer. The primary antibody monoclonal cyclin D2 antibody no.
MS-221 (NeoMarkers, Union City, CA) diluted 1:100 in PBS + 0.1% HSA +
0.3% Triton-X-100 was incubated for 1 h.
Statistical Analysis
Statistical analysis was performed using SAS 6.12
software (SAS Institute, Cary, NC). Two-way ANOVA with
Dunnetts method for adjustment of multiple comparisons was carried
out.
 |
ACKNOWLEDGMENTS
|
---|
We want to thank Dagny Jensen and Hanne Richter Olesen for
excellent technical assistance. We are indebted to Dr. B. Groner, Tumor
Biology Center, Freiburg, Germany, for kindly supplying STAT5a
749
cDNA; to Dr. P. B. Iynedjian, University of Geneva School of
Medicine, Geneva, Switzerland, for supplying INS-r3 cells; and Dr.
C. B. Wollheim, Centre Medical Universitaire, Geneva, Switzerland,
for the supply of INS-1 cells. We thank Dr. S. N. Jakobsen and Dr.
N. Billestrup, Novo Nordisk, Bagsværd, Denmark, and Dr. K. Seedorf,
Lilly Research Laboratories, Hamburg, Germany, for helpful advice and
discussion.
 |
FOOTNOTES
|
---|
Address requests for reprints to: Annette Møldrup, Department of Islet Discovery Research, Novo Nordisk A/S, Novo Alle 1, 1KMS03, 2880 Bagsværd, Denmark. E-mail amp{at}novo.dk
Received for publication March 30, 2000.
Revision received September 11, 2000.
Accepted for publication September 20, 2000.
 |
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