Prolactin (PRL)-PRL Receptor System Increases Cell Proliferation Involving JNK (c-Jun Amino Terminal Kinase) and AP-1 Activation: Inhibition by Glucocorticoids
Isabel Olazabal,
Jaime Muñoz,
Samuel Ogueta,
Eva Obregón and
Josefa P. García-Ruiz
Departamento de Biología Molecular-Centro de
Biología Molecular "Severo Ochoa" Facultad de
Ciencias Universidad Autónoma de Madrid 28049 Madrid
Spain
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ABSTRACT
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PRL receptor (PRLR) signal transduction
supports PRL-induced growth/differentiation processes. While PRL is
known to activate Jak2-Stat5 (signal transducer and activator of
transcription 5) signaling pathway, the mechanism by which cell
proliferation is stimulated is less known. We show that PRL induces
proliferation of bovine mammary gland epithelial cells and AP-1 site
activation. Using PRLR mutants and the PRLR short form, we have found
that both homodimerization of PRLR wild type and the integrity of box-1
and C-distal tyrosine of PRLR intracellular domain are needed in
PRL-induced proliferation and AP-1 activation. The effect of PRL has
been assayed in the presence of dexamethasone (Dex), insulin, and
alone. We found that Dex negatively regulates PRL-induced proliferation
and AP-1 site activation. We demonstrate that PRL exerts activation of
AP-1 transcriptional complex, and the mechanism by which this
activation is produced is also studied. We show that PRL induces an
increase in the c-Jun content of AP-1 transcriptional complexes. The
PRL-induced c-Jun of AP-1 transcriptional complex diminishes in the
presence of Dex in a dose-dependent manner. Dex inhibition was reversed
by the higher concentration of PRL added to cells. Despite the fact
that the regulation of the AP-1 site is complex, we found that PRL
activates the c-Jun amino terminal kinase (JNK), while glucocorticoid
prevents this JNK activation. These data support a regulation of
cellular growth by PRL-PRLR system by increasing AP-1 transcriptional
complex activity via JNK activation. JNK activation can be repressed by
glucocorticoid in a DNA-binding-independent manner.
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INTRODUCTION
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PRL is a peptide hormone that stimulates cellular proliferation
and differentiation processes (1, 2). PRL signals through the PRL
receptor (PRLR), a member of the cytokine receptor superfamily (3, 4).
The PRLR is expressed in rat tissues in two forms by alternative
splicing of a single gene, so that they only diverge at the
intracellular domain (5). PRL induces dimerization of cell surface
receptors leading to phosphorylation and activation of the
receptor-associated protein tyrosine kinase JAK2 and the PRLR (6, 7).
These phosphorylations create sites for SH2 type interactions with
intracellular signaling molecules including protein tyrosine
phosphatase (8), phosphatidylinositol 3-kinase (9), and STAT-5
(signal transducer and activator of transcription) (10). The activated
STAT-5 translocates from the cytoplasm to the nucleus where it binds to
specific DNA motifs of ß-casein promoter and activates the
transcription of ß-casein gene (11). The mechanisms by which PRL
exerts cell growth are less documented. PRL induction of cell
proliferation has been determined in different cell lines (12, 13).
Furthermore, PRLR immune-complexes have associated signaling molecules
used by growth factors. Tyrosine kinases of the Src family have been
described associated with and activated by PRL (14, 15). Activation of
Grb2/Sos/Ras (16, 17), Raf (18), Vav (19), and IRS1/PI3-K (9) has also
been reported. These experiments support pathways by which the PRL-PRLR
system may induce activation of MAPKs (mitogen-activated protein
kinases) (16, 17). However, the genes activated in these pathways
remain to be elucidated.
Growth factors, proinflammatory cytokines, oxidative stress, and
UV irradiation initiate cellular signals by different mechanisms that
converge in the activation of one of the three families of MAP kinases,
ERK, JNK, or p38, leading to AP-1 complex activation. The AP-1 family
of transcriptional factors consists of homodimers and heterodimers of
Jun, Fos, ATF, and Maf family members (20). It is becoming clear that
different AP-1 factors may regulate different target genes and, thus,
have distinct biological functions. c-Jun plays a role in cell
proliferation in response to external growth factor ligands forming
Fos-Jun and Jun-Jun dimers and activating AP-1 sites (21). This was
suggested after cell progression from G1 into S phases failed when
c-Jun was neutralized with antibodies (22). Moreover, it has been
recently demonstrated that c-Jun regulation is critical in two
different cellular processes, proliferation and survival, which involve
distinct biochemical mechanisms. In fibroblasts derived from c-Jun null
embryos, c-Jun is required for progression through the G1 phase of the
cell cycle. c-Jun-mediated G1 progression occurs by a mechanism that
involves direct transcriptional control of the cyclin D1 gene. This
establishes a molecular link between growth factor signaling and cell
cycle regulators. In addition, c-Jun protects cells from UV-induced
apoptosis when it is phosphorylated in serines 63/73 by JNK activity
(23). Thus, c-Jun is involved in two different cellular processes,
proliferation and antiapoptosis, that can be independently modulated by
extracellular stimuli.
In this study, we analyzed the potential PRL-regulated proliferation of
bovine mammary gland epithelium cells (BMGE). These cells conserve the
pathway to induce ß-casein gene promoter. We considered it of
interest to study both PRL-modulated cell proliferation in comparison
with ß-casein induction and the influence of glucocorticoids and
insulin (Ins) in both processes. To this end, BMGE cells were
transiently transfected with PRLR, wild type and mutated, expression
vectors. Since AP-1 transcriptional elements mediate cell
proliferation, we assessed the PRL regulation of the AP-1 site of
collagenase gene promoter in comparison with ß-casein gene promoter.
The regulation of AP-1 transcriptional factors varies with the specific
family member and with cell type (21). For this reason we assessed
PRL-regulated AP-1 transcriptional complexes via c-Jun modulation. Our
findings indicate that PRL activates the AP-1 element of collagenase
gene promoter mediated by JNK activation, which is inhibited by
glucocorticoids.
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RESULTS
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PRL-PRLR System Induces Proliferation of BMGE Cells
BMGE cells have been proven to be a useful system to study
PRL-induced ß-casein gene promoter (23). We explored whether, in
addition to the PRLR signaling pathway to ß-casein gene promoter, PRL
could induce BMGE cells to grow by analyzing the rate of BrdU or
thymidine incorporation into cell nuclei. Transient transfections of
BMGE cells were performed using wild-type (PRLR-L) and mutated PRLR
(PRLR-L4PA, mutated in the proline box motif and PRLR-LY580F, mutated
in the distal tyrosine of the intracellular domain) expression vectors.
Since PRLR signaling to ß-casein is often studied in the presence of
glucocorticoids and Ins, we considered it of interest to analyze the
role of both hormones in PRL-induced proliferation experiments. Cells
stimulated with 10% FCS have been used as control and referred to as
100% response. Results are summarized in Fig. 1
. In cells incubated with
dexamethasone (Dex) and Ins, only the expression of PRLR-L caused a
significant increase in the rate of cell proliferation (Fig. 1A
). The
rate was 2.5-fold higher than that of cells transfected with control
vector (P < 0.01) and double the rate observed in
cells stimulated with FCS (P < 0.01). However, only a
nonsignificant increase in proliferation was observed by PRL
stimulation. We explored whether these cells synthesized PRL by RT-PCR,
and the results were negative (data not shown). Is there a constitutive
activation of PRLR in these cells? Interestingly, PRLR expression of
nonstimulated 32Dc13 cells was found to increase the pattern of
phosphotyrosine-containing proteins that was further increased by PRL
or interleukin-3 addition to cells (25). When the PRL effect was
analyzed in cells cultured in the presence of Ins (Fig. 1B
), the rate
of proliferation of cells expressing PRLR-L was similar to cells
stimulated with 10% FCS and increased by 2-fold after PRL treatment
(P < 0.05). Upon expression of PRLR mutant forms:
PRLR-L4PA and PRLR-LY580F, no PRL-induced proliferation was detected.
In the absence of Dex and Ins, PRLR expression was observed to cause an
increase in the rate of proliferation (P < 0.01) and a
2-fold increase upon PRL stimulation (P < 0.01) (Fig. 1C
). As occurred in cells incubated in the presence of Ins, PRL-induced
proliferation was not observed in cells expressing PRLR mutant forms.
Thus, the results show that PRL stimulates the rate of proliferation in
BMGE cells and suggests that Ins and glucocorticoids may contribute to
a constitutive activation of PRLR in these cells.

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Figure 1. PRL Activates the Proliferation of BMGE Cells
Cells were transfected with the expression vectors encoding for PRLR-L
and mutant forms PRLR-L4PA and PRLR-LY580F and pCMV empty vector as a
control. PRL-induced proliferation was determined after 48 h in
cells incubated in: A, 2.5 nM Dex and 3 µg/ml Ins; B,
Ins; and C, vehicle. In each experiment, cells transfected with pCMV
vector and stimulated with 10% FCS were used as proliferation-positive
controls (hatched bars). Cells were labeled with BrdU,
fixed, and processed following Amersham Pharmacia Biotech
instructions. Results, expressed as percentage of positive controls,
are the mean ± SEM of three to five
independent experiments.
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PRL-PRLR System Activates AP-1 Site of Collagenase Promoter, Which
Is Inhibited by Glucocorticoids
The results shown above prompted us to study the mechanism
of PRL-induced cell proliferation. Thus, we assessed the ability of the
PRL-PRLR system to stimulate AP-1 transcriptional activity by using an
AP-1-dependent expression vector in which a deletion derivative of the
collagenase promoter is fused to Luc reporter gene, -73-Col-Luc (26).
In addition, we explored the influence of Dex and Ins on PRL regulation
of both ß-casein and AP-1 promoters. To this end, ß-Cas-Luc or
-73-Col-Luc expression vectors were cotransfected into BMGE cells with
the PRLR-L expression vector. As shown in Fig. 2
, the expression of PRLR-L in cells that
were incubated in a hormone-free medium caused a 2-fold increase
(P < 0.01) of ß-casein gene promoter and AP-1
elements. The presence of PRL significantly stimulated the activity of
both promoters (P < 0.01). When cells were incubated
with Ins, the expression of PRLR-L and the stimulation with PRL of both
promoters did not induce any significant alteration with respect to
data observed in the absence of hormones. However, in the presence of
Dex, PRL significantly stimulated ß-casein promoter
(P < 0.01) (Fig. 2A
), as already reported (27), while
Luc activity driven by AP-1 elements was diminished (P
< 0.05) (Fig. 2B
). In the presence of Dex and Ins, the data observed
with the expression of PRLR and the PRL stimulation of both promoters
were similar to those obtained in the presence of Dex
alone.

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Figure 2. PRL Stimulation of ß-Casein Promoter and AP-1
Elements
BMGE cells were cotransfected with pCMV vector or with expression
vectors encoding PRLR-L and ß-casein-Luc (A) or AP-1-Luc (B)
constructs. PRL effect was determined after 48 h of treatment in
the presence or absence of Dex (D) and Ins (I). Cells were lysed and
luciferase and ß-galactosidase activities were determined. Results
are the mean ± SEM of four experiments expressed as
percentage of the maximum stimulus detected in each type of experiment.
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Since the PRL-PRLR system stimulates AP-1 elements, we assessed which
structural domain of the PRLR-L was involved in this function. To this
end, PRLR-L or its mutant forms, PRLR-L4PA and PRLR-LY580F, were
cotransfected in BMGE cells with the -73-Col-Luc construct. After
transfection, the cells were either untreated or stimulated with PRL
for 24 h and Luc activity was determined. Cells treated with
phorbol ester [phorbol myristol acetate (PMA)] (28) during
4 h were used as a positive control in all the experiments. As
shown in Fig. 3A
, BMGE cells in the
absence of PRLR expression presented basal reporter activity and
increased by 2-fold when stimulated with PMA. The expression of PRLR-L
increased the reporter Luc activity by 2.5- to 3-fold
(P < 0.01), and PRL stimulation caused an additional
rise (P < 0.01). The expression of PRLR mutant forms
did not stimulate Col-Luc, and PRL treatment did not cause a
significant stimulation.

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Figure 3. PRLR Mutants and Short Isoform Silence AP-1 Element
Activation
A, BMGE cells were cotransfected with vectors expressing PRLR-L
or its mutant forms and with -73-Col-Luc construct. They were incubated
in the presence of Dex and Ins and stimulated (+) or not (-) with PRL
for 48 h. As a positive control, cells were stimulated with PMA
for 4 h. B, Cells were transfected with different amounts of
vectors specifying the PRLR long or short form separately (B.1) and
cotransfected with different proportions of PRLR-L and PRLR-S (B.2), as
indicated above. Cells were lysed, and luciferase and ß-galactosidase
activity were determined. Results are the mean ± SEM
of four experiments expressed as percentage of the PRLR stimulus.
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The PRLR short (PRLR-S) form has been proved to have a silencing
function in the activation of ß-casein promoter (24). We analyzed
whether PRLR-S could act in the same way in the stimulation of AP-1
elements. Cells were transfected with different amounts of PRLR-L or
PRLR-S constructs separately as controls (Fig. 3B
.1) or with a mixture
of PRLR-L and PRLR-S constructs (Fig. 3B
.2). The amount of DNA was
maintained at 3.33 µg by adding pCMV plasmid when necessary. As can
be observed in Fig. 3B
.1, the Col-Luc was efficiently stimulated by the
PRLR-L at the different concentrations used (P <
0.01). In contrast, PRLR-S did not induce Col-Luc at any of the
concentrations assayed. Interestingly, cotransfection of PRLR-L: PRLR-S
constructs in a 1:1 ratio (Fig. 3B
.2) abolished the stimulation of
Col-Luc compared with the PRLR-L construct alone and decreased by 50%
the ability of PRL stimulation of the Col-Luc (P <
0.01). Thus, PRLR-S blocks PRL signaling to both ß-casein promoter
and AP-1 elements.
PRL Increases AP-1 DNA Binding Activity in BMGE Cells
The ability of PRL to induce AP-1 DNA binding activity was
analyzed in BMGE cells transfected with the PRLR-L expression vector.
To this end, cells were transfected and incubated for 24 h in GC-3
medium and then stimulated with PRL for different periods of time.
Nuclear extracts were prepared, normalized for protein concentration,
and incubated with radiolabeled oligonucleotide probe containing the
consensus AP-1-binding site or a mutant oligonucleotide (Fig. 4A
, lane 1). A representative result of
electrophoretic mobility shift assays (EMSAs) is shown in Fig. 4A
. PRL
was able to increase binding to the AP-1 site after 5 min (lane 4) that
may reflect activation of preexisting AP-1 factors, followed by a
significant increase after 6 h (lane 5) and 24 h (lane 6)
that could reflect a sustained PRL induction.

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Figure 4. PRL Induction of AP-1 and c-Jun DNA Binding
Activity
Nuclear extracts were prepared from PRLR-L-transfected BMGE cells
treated with PRL for the specified times. A, Each nuclear extract (5
µg) was analyzed by EMSA using [32P]-AP-1 probe (lanes
26). In lane 1, a [32P]-mutated AP-1 probe is used. The
arrows indicate the retarded complex and the free probe.
B, Nuclear extracts were analyzed by supershift assay using an anti
c-Jun (H-79) antibody (lanes 3, 6, 8, 10, and 12). In lane 1, a
[32P]-mutated AP-1 probe is used. After 15 min of nuclear
extract-AP-1 probe incubation, 0.2 µg of the anti c-Jun or of a
nonspecific antibody (N) (lane 5) was added. Then, reactions were left
for an additional 30 min as indicated in Materials and
Methods. Figure shows a representative experiment out of the
four performed. The arrows indicate the overexposed
retarded AP-1 complex, and the supershifted complex.
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We then assayed whether or not the c-Jun concentration in PRL-induced
AP-1 complexes was altered. For this purpose, BMGE cells were
transfected with the expression vector carrying the PRLR-L coding
sequence or with pCMV vector and treated as above. c-Jun factor present
in AP-1 complexes was analyzed by supershift assay. Films needed longer
exposure time to detect c-Jun-antibody complexes than to detect AP-1
complexes. As can be observed in a representative experiment shown in
Fig. 4B
, PRL stimulates an enrichment of c-Jun factor in AP-1
complexes. The c-Jun content was increased after 5 min (lane 8) and was
sustained after 6 or 24 h (lanes 10, 12) of PRL addition to cells.
Thus, there is a correlation between PRL-induced c-Jun content in AP-1
and the activation of AP-1 complex.
Glucocorticoids and PRL Modulation of AP-1 Complexes
Inhibition of PRL-induced Col-Luc promoter activity by
glucocorticoids prompted us to study whether glucocorticoids
transmodulate PRL-induced AP-1 complexes. To this end, cells were
transfected with PRLR-L expression vector and incubated for 24 h
in GC-3 medium. Then, cells were treated for 6 h either with 40
nM PRL and increasing amounts of Dex, or with 2.5
nM Dex and increasing amounts of PRL. In both types of
experiments, the c-Jun content of AP-1 complexes was assayed by
supershift analysis. As can be observed in a representative experiment
(Fig. 5A
), PRL-induced c-Jun factor of
AP-1 complexes (lane 3) was diminished by Dex treatment in a
concentration-dependent manner (lanes 5, 7, and 9). In addition, Dex
inhibitory effect was reversed by increasing amounts of PRL added to
cells (Fig. 5B
). Films were overexposed to detect the supershifted
complex. These results were consistently observed in four independent
experiments, as shown in the densitometric quantification (Fig. 5C
).

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Figure 5. PRL and Dex Modulation of AP-1 Complexes
Nuclear extracts were prepared from PRLR-L-transfected BMGE cells. A
and B, Cells were treated with the indicated amounts of PRL and Dex for
6 h. After 15 min of nuclear extract-AP-1 probe incubation, 0.2
µg of the anti c-Jun (H-79) antibody (lanes 3, 5, 7, and 9), or a
nonspecific antibody (N) (lane 2) was added. Then, reactions were left
for an additional 30 min as indicated in Materials and
Methods. The figure shows a representative experiment. The
films were overexposed to visualize the supershifted complexes. C,
Densitometric evaluation of supershifted c-Jun, in four independent
experiments as performed in panels A and B, is expressed in relative
percentage.
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PRL Activates JNK
Since AP-1 elements can be activated by at least three distinct
cascades of protein kinases (20), we assessed whether PRL exerts JNK
and p38 kinase activation in BMGE cells. To this end, we determined
kinase activity in immune complexes obtained from PRLR-L-transfected
cells using purified GST-c-Jun and GST-ATF-2 as substrates. Cells were
transfected with PRLR-L expression vector or with pCMV vector as
control. In each experiment, aliquots of cells were treated with
anisomycin for 30 min, as a positive control of JNK activity (29). The
effect of PRL was analyzed after 15 min and 24 h of addition, as
representative short and long lasting duration. The result of a
representative experiment is shown in Fig. 6A
. Cells transfected with PRLR (lane 2)
or with pCMV (lane 1) had basal levels of JNK activity. Within 15 min,
PRL addition to cells doubles JNK activity (P <
0.01)(lane 3) and after 24 h a 50% increase was detected,
although this was not a significant stimulation (P >
0.05)(lane 4). As expected, anisomysin treatment caused a significant
increase of JNK (P < 0.001) (lane 5). When p38 kinase
was assessed in identical experiments, no alteration was observed after
PRL treatment (results not shown). In addition, we explored whether JNK
activity was the place for glucocorticoids inhibition of the PRLR
signaling to c-Jun. To this end, cells transfected as above were
stimulated or not with PRL for 15 min and incubated with increasing
amounts of Dex for 6 h. As can be observed in a representative
experiment (Fig. 6B
), Dex addition at 2.5 nM
caused a significant decrease (P < 0.01) (lane 4) of
PRL-stimulated JNK activity (lane 3 vs. lane 2) that was
sustained when increasing amounts of Dex were added (lanes 5 and 6).
Results of JNK activity normalized by the amount of JNK
immunoprecipitated of three independent experiments, A and B type, are
together shown in Fig. 6C
. In accordance with these results, the PRLR
signaling to AP-1 is mediated by the activation of JNK, which is
inhibited by glucocorticoids.

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Figure 6. PRL Activates Jun N-terminal Kinase: Inhibition by
Glucocorticoids
A, BMGE cells were transfected with pCMV vector (lane 1) or with
PRLR-L expression construct (lane 25). After maintenance for 24
h, cells were left untreated (lane 2) or treated with PRL for 30 min
(lane 3) for 24 h (lane 4) or with 0.2 µM anisomycin
(lane 5). The cells were lysed, extracts were normalized for protein
concentration, and JNK was immunoprecipitated with 2 µg of anti-JNK
antibody. Kinase assays were determined in immune complexes using 2
µg of GST-c-Jun peptide as substrate and 5 µCi of
( -32P)-ATP. B, BMGE cells were transfected with pCMV
control vector (lanes 1 and 7) or with PRLR-L expression vector (lanes
26). After maintenance for 24 h, cells were untreated (lane 2)
or treated with 0.2 µg anisomycin (lane 7) or with PRL for 15 min
(lane 36). After PRL stimulation, Dex at the specified concentrations
was added to cells for 6 h (lanes 46). JNK assays were performed
as above. C, The results of JNK assays, after normalization for the
amount of JNK by immunoblotting with anti-JNK antibody, in three
independent experiments as performed in panels A and B. JNK activity is
expressed as -fold induction and is presented as mean ±
SE (error bars).
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DISCUSSION
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The binding of PRL to its receptor initiates a specific cascade of
signaling events. After a ligand-induced homodimerization of the
receptor, it activates both Jak2 tyrosine kinase (6, 7) and src-related
tyrosine kinase (14, 15). Jak2 mediates the activation of the
transcriptional factor STAT-5 that increases the transcription of
ß-casein (10). The mechanism of PRL-induced cellular proliferation is
less well known. PRL-induced proliferation has been observed in
different cell lines, Nb2 rat T-lymphoma (12), factor-dependent myeloid
cells (30) and human breast cancer cells (13). This correlates well
with PRL-induced activation of SHC, RAS, RAF, PI3-K, and IRS-1 (9, 16, 17, 18), similar to that described for cell growth factors and
growth-promoting cytokines. However, it is not clear whether PRL alone
can stimulate the proliferation of mammary epithelial cells or the role
of PRL and PRLR in the growth/differentiation processes of the mammary
gland. Despite this fact, PRL and PRLR must play a critical role in
mammary gland growth and development since knockout female mice of
either PRL, PRLR, or STAT-5 genes present failures in mammary gland
development (31, 32, 33).
We show that BMGE cells increase their rate of proliferation by
boththe only expression of PRLR-L form and subsequent PRL stimulation
of this receptor. As we have found no synthesis of PRL in these cells,
this effect needs to be mediated by the expression of PRLR. It is known
that mammary epithelial cells are adherent, and phenomena such as
migration, proliferation, survival, and differentiation are strongly
influenced by cell interactions with the extracellular matrix (ECM). In
fact, both laminin-1 and ß1 integrin are required for differentiating
mammary cells and for PRL signaling pathways to milk protein synthesis
(34, 35, 36). In this sense, we suggest that BMGE cell-ECM
interactions must organize PRLR signaling components into a
functionally active complex to increase cell proliferation upon PRLR
expression. PRL exerts increased cell
proliferation mediated by PRLR-L while mutant PRLR forms at proline-box
or distal tyrosine do not transmit PRL signals. However, the results
show that these mutants increase cell proliferation, suggesting that
cell-ECM interactions can be modulated by the expression of the PRLR.
It is well documented that growth factor responses synergize with
integrin-mediated signaling. However, the mechanisms underlying the
interplay of signaling pathways are not well established (37). Our
results are in accordance with those showing that Ins potently
activates a specific type of integrin in CHO cells and, in turn, this
integrin activates insulin receptor kinase (38). Moreover, Ins and
platelet-derived growth factor receptors associate with integrin
since both receptors have been detected in integrin immunoprecipitates
(39). Interestingly, the constitutive activation of the deleted
178
PRLR mutant at the extracellular ligand-binding domain argues that
disruption of the WSXWS motif plays an important role for the
ligand-independent proliferation induced by this PRLR mutant (25). In
this complex situation, the increased proliferation detected by
the PRL-PRLR system in BMGE cells incubated with Ins can be a
consequence of Ins and PRL synergistic actions. In addition, the
PRLR-induced proliferation seems to be important in epithelial cells
since it was higher in the presence of Ins and Dex. In fact, Dex
regulates epithelial cell phenotype, including ECM composition and
ECM receptors (40, 41). Although future work is needed, our
results are consistent with PRL-PRLR system stimulation of BMGE
cell proliferation mediated by both PRLR interactions with ECM or ECM
receptors and PRLR intracellular interactions with signaling
proteins.
The results show that, in BMGE cells, PRL-PRLR system activates both
ß-casein gene promoter and AP-1 site containing collagenase gene
promoter. The PRL modulation of both gene promoter elements is
transregulated by glucocorticoids, which provides a functional
antagonism between both signaling pathways. Both glucocorticoid actions
can be considered as DNA-binding-independent activities of
glucocorticoid receptor. In relation to ß-casein regulation, our
results expand on the results of previous studies on mechanisms of
cooperation of glucocorticoid receptor and STAT-5. Both proteins
synergistically cooperate in the transcription of the ß-casein gene
by protein-protein type interaction between glucocorticoid receptor and
STAT-5 at STAT-5-DNA binding domain (27, 42).
This study shows that PRL stimulates AP-1 transcriptional activity by
which PRL may regulate different target genes and thus may execute
distinct biological functions such as cell proliferation and cell
survival. The mechanism for PRL-induced AP-1 transcriptional complex
activity involves at least an increase in the c-Jun concentration in
the AP-1 complex. The fact that PRL-induced activation of AP-1 complex
was observed in minutes is consistent with the activation of
preexisting c-Jun proteins, while in a longer time it may correspond
with synthesis of c-Jun. c-Jun protein is a central component of AP-1
transcriptional factors and mediates the control of genes that regulate
cell growth. Studies performed in fibroblasts derived from c-Jun null
embryos demonstrate that c-Jun is required for progression through the
G1 phase of the cell cycle. c-Jun-mediated
G1 progression occurs by a mechanism that
involves direct transcriptional control of the cyclin D1 gene (23).
Interestingly, an early work showed, in Nb2 cells, that PRL stimulates
the transcription of cyclin D2 (43). Thus, our results support that PRL
stimulates cell growth, and reveal an end point of several
intracellular signals induced by PRL, including Grb2 and SOS to
SHC/RAS/RAF (16, 17). The studies performed to determine the
functional/structural relationship of PRLR signaling to AP-1 showed no
difference with those of the signaling to ß-casein gene promoter.
PRL-induced cell proliferation and AP-1 activation need
homodimerization of PRLR long form, since the coexpression of PRLR
short and long isoforms decreased PRL-induced proliferation and AP-1
activation. In addition, results derived from PRLR mutants show that
proline-box and the distal residue of tyrosine of the PRLR are needed
in PRL-induced proliferation and AP-1 activation. Thus, molecules that
interact with PRLR by means of SH3 and SH2 type of interactions are
needed for growth signals. In contrast, erythropoietin receptor that
belongs to the cytokine family of receptors, and has signaling pathways
through Jak2-STAT-5 and AP-1 activations, signals to AP-1 depending on
multiple intracellular receptor tyrosines, but not depending on Jak2
activation (44).
In this study we also show that PRL alone stimulates BMGE cell
proliferation, an effect that is transmodulated negatively by
glucocorticoids. These results correlate well with both glucocorticoid
inhibition of PRL-induced AP-1 complex transcriptional activity and the
parallel decrease in c-Jun content of AP-1 complex. Thus, both PRL and
glucocorticoids have the c-Jun factor as a target with antagonistic
functions. These results expand on evidence of glucocorticoid
transrepression of AP-1-driven genes by a glucocorticoid receptor
DNA-independent mechanism (45). Indeed, glucocorticoid receptor
represses AP-1 transcriptional factor by interacting with activated
c-Jun (46, 47). Thus, our observations that PRL stimulates JNK
activity, which is prevented by glucocorticoids, provide evidence for
the mechanism of glucocorticoid transmodulation of PRLR signaling and
gives a tempting explanation for the antiapoptotic role attributed to
PRL (48). As it has been shown that c-Jun phosphorylated by JNK at
serines 63/73 protects from apoptosis in response to UV (23), further
experimental work is needed to establish whether PRL is involved in the
antiapoptotic function in this way.
In summary, our results show that the PRL-PRLR system stimulates cell
proliferation by regulating c-Jun content of AP-1 transcriptional
complex, and that glucocorticoids play a key role in the cellular
signaling of PRL-PRLR system in BMGE cells. This cross-talk between PRL
and glucocorticoids may arbitrate the PRL-induced differentiation and
growth processes.
 |
MATERIALS AND METHODS
|
---|
Plasmids
The -73 collagenase gene promoter-luciferase (-73-Col-Luc)
construct was a gift from J. M. Redondo (Centro de
Biología Molecular, Madrid, Spain) and constitutes an AP-
1-dependent expression vector in which a deletion derivative of the
collagenase promoter is fused to the Luc reporter gene (26). The (-344
to -1) ß-casein gene promoter luciferase (ß-Cas-Luc) reporter
plasmid and expression vectors for PRLR forms were provided by P.
A. Kelly (10, 49): pECE-PRLR long and short forms, pECE-4PA, and
pCMV-Y580F. The PRLR intracellular domain mutant forms, 4PA and Y580F,
present a deletion of the proline box, and the distal tyrosine mutated
to phenylalanine, respectively. pRc/CMV vector was used as control of
an empty vector in different experiments (pCMV)
(Invitrogen, San Diego, CA). The pECE-PRLR long and short
forms were subcloned, respectively, into EcoRI or
HindIII-XbaI of the pRc/CMV vector. To generate
the pCMV-4PA mutant form, pECE-4PA and pRc/CMV vectors were digested
with EcoRI and HindIII, respectively, and then
blunt-ended. One-end products were XbaI digested, and the
2.35- and 5.15-kb fragments generated were ligated.
Cell Culture and Transient Transfection Assays
Bovine mammary gland epithelium (BMGE) cells (50) were grown in
DMEM supplemented with 10% (vol/vol) FCS (Life Technologies, Inc., Gaithersburg, MD), 2 mM glutamine,
nonessential amino acids, 0,01% penicillin-streptomycin, 50 µg/ml
gentamycin, and antimicotics. For transfection, 20 x
104 cells were grown until they reached 7080%
of confluence. Then, they were starved overnight in GC3 medium composed
of 1:1 DMEM and Hams F12 (Life Technologies, Inc.)
supplemented with 10 µg/ml transferrin (Sigma,
Madrid, Spain) and 3 µg/ml Ins (Humulina, Lilly Ltd., Madrid, Spain).
Cells were transfected with 3.33 µg of total DNA by the calcium
phosphate precipitation procedure (24). To study the inducibility of
different promoters, 0.5 µg of -73-Col-Luc or ß-Cas-Luc plasmids
were used. In every assay, 1 µg of the PRLR expression vectors or the
empty vector pCMV and 0.33 µg of CMV-ßgalactosidase plasmid were
cotransfected. After glycerol shock, cells were incubated during 4548
h in GC3 medium in the presence or absence of hormones. When added, the
amounts were, 3 µg/ml Ins, 2.5 nM Dex (Decadran,
Merck & Co., Inc., St. Louis, MO) and 40 nM
ovine PRL (National Hormone and Pituitary Program, Rockville, MD).
Cells were washed twice with cold PBS and then lysed with 0.15 ml of
lysis buffer (25 mM Tris-phosphate, pH 7.8, 2
mM dithiothreitol (DTT), 2 mM EDTA, 10%
glycerol, and 1% Triton X-100). Luciferase activity was measured in
arbitrary light units and normalized with ß-galactosidase activity.
Results are expressed as percentage and represent the mean ±
SEM of at least four different experiments.
Proliferation Assay
BMGE cells (2.5 x 104) were grown in
M24 dishes and transfected with 2 µg of total DNA containing 0.6 µg
of PRLR expression vectors or the pCMV empty vector. After stimulation
for 40 h with 40 nM PRL or 10% FCS, as a positive
control, thymidine-deficient RPMI medium was added to cells for 1
h. Then, cells were incubated for 3 h in the same medium
containing 5-bromo-2'-deoxyuridine (BrdU). Cells were washed twice with
cold PBS and processed following Amersham Pharmacia Biotech (Piscataway, NJ) kit instructions. Positive cells were
counted under microscopy light. Alternatively, proliferation rate was
assessed measuring the incorporation of
[3H]-thymidine to cells. In this case,
triplicates of 0.5 x 104 cells were grown
in M24 dishes as indicated above and were pulsed with 1 µCi of
[3H]-thymidine for 5 h. Cells were
harvested with an automatic collector (Skatron), and the radioactivity
incorporated was determined in a Rack Beta scintillation counter
(LKB Wallac, Inc., Turku, Finland).
Results are expressed as percentages.
Nuclear Extracts
Nuclear extracts were prepared as described (51). Briefly,
3 x 106 cells were washed with cold PBS and
swelled in 400 µl of lysis mixture. This contained 10 mM
HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.2
mM phenylmethylsulfonyl fluoride (PMSF), 1 mM
DTT, 2 µg/ml leupeptin, and 4 µg/ml pepstatin A. After 15 min at 4
C, samples were adjusted to 0.6% Nonidet P-40 (NP-40) and vigorously
shaken for 10 sec. Nuclei were pelleted by centrifugation during 1 min
at 12,000 x g. Nuclear proteins were extracted with a
high-salt solution containing 20 mM HEPES, pH
7.9, 400 mM KCl, 0.2 mM
EDTA, 2 mM PMSF, 1 mM DTT,
2 µg/ml leupeptin, and 4 µg/ml pepstatin A. The volume added
was equal to that of the nuclear pellets. Tubes were vigorously shaken
for 20 min and then centrifuged at 12,000 x g for 5
min. Supernatants were harvested as the nuclear protein extracts and
stored at -70 C. Protein concentration was determined in triplicate by
the Bradford method.
EMSAs
EMSAs were developed as described (38). DNA-protein binding
reactions were conducted in 20 µl volume. Reaction mixtures were
composed of 1 µg poly (dI-dC) (Sigma, St. Louis, MO), 5
µg nuclear protein extracts, 0.1 µg denatured salmon sperm DNA, 10
µg BSA, 0.15 ng [32P]-labeled double-stranded
oligonucleotide (100,000150,000 cpm), and 10 µl (2x) binding
reaction buffer. Binding buffer was composed of 20 mM
HEPES, pH 7.9, 60 mM KCl, 5 mM
MgCl2, 0.2 mM EDTA, 8% glycerol, 0.2
mM PMSF, and 1% NP-40. Reactions were incubated at room
temperature for 15 min and resolved on 5% nondenaturing polyacrylamide
gel and Tris-glycine buffer (prerun at 110 V for 2 h). The loaded
gel was run at 30 mA for 90 min, dried, and exposed on X Omat film
(Eastman Kodak, Rochester, NY). The AP-1 probe and its
mutant were purchased from Santa Cruz Biotechnology, Inc.
(Santa Cruz, CA). The double-stranded probe was labeled with
[
-32P] ATP and Polynucleotide Kinase
(Promega Corp.). For supershift analysis, the c-Jun
antibody (H-79 from Santa Cruz Biotechnology, Inc.) or rabbit antimouse
[RAM, IgG + IgM (H + L), Jackson ImmunoResearch Laboratories, Inc., West Grove, PA] to detect nonspecific binding were used.
They were added to binding reactions after 15 min of incubation time
and left for an additional 30 min.
Isolation and Quantification of mRNA
Total RNA was isolated from 5 x 106
BMGE transfected cells and bovine pituitary using the standard
guanidinium-thiocyanate-phenol procedure (52). RT-PCR determinations of
PRL messenger were performed using the primers:
5'-GCTGCTTGTTTTGTTCCTCAATCTC-3' and 5'-CTCTCCGAGAGCTGTTTGACCG-3'
corresponding to the mouse PRL gene. These primers were used since
bovine PRL corresponding sequences present, at the sense and antisense
primers, three and five mismatches, respectively. These mismatches are
located beyond positions 11 and 12, respectively, from the 3'-expanding
ends of both primers. RT reaction was carried at 42 C for 1 h with
5 U of AMV retrotransferase (Promega Corp.). PCR
conditions were three cycles: 94 C/30 sec, 45 C/30 sec, 72 C/30 sec and
35 cycles: 94 C/30 sec, 45 C/30 sec, and 72 C/30 sec. Southern
hybridization was carried out using an internal oligomer
5'-TCCTGGAATGAGCCTCTGTATCA-3'corresponding to bovine and mouse PRL
sequences following the protocol described (24).
Solid-Phase JNK Assays
Cells (6 x 106) were lysed in 1 ml
of lysis buffer containing 50 mM Tris-HCl (pH 7.5), 100
mM NaCl, 5 mM EDTA, 30 mM sodium
pyrophosphate, 50 mM sodium fluoride, 0.8% Triton X-100,
8% glycerol, 2 mM sodium orthovanadate, 1 mM
PMSF, 5 µg/ml apronitrin, 4 µg/ml pepstatin A, and 2 µg/ml
leupeptin. Lysates were left for 30 min and microcentrifuged for 15 min
at 4 C. Supernatants were normalized by their protein concentration,
and p38 and JNK-1 were reacted with anti-p38 and anti-JNK-1 antibodies
(Santa Cruz Biotechnology, Inc.) at 2 µg/ml. Immune
complexes were precipitated with rabbit antimouse (RAM), bound to
protein A-Sepharose. Immune complexes were washed twice with buffer A
(20 mM Tris-HCl (pH 7.4), 140 mM NaCl, 5
mM EDTA, 1% Triton X-100) and twice with kinase buffer (50
mM HEPES (pH 7.1), 0.1 mM EDTA, 0.1 mg/ml BSA,
0.1% ß-mercaptoethanol, 20 mM
MgCl2). Fusion proteins,
Glutathione-S-transferase GST-c-Jun and GST-ATF-2 were
purified on glutathione-agarose, as described previously (53, 54). The
activity of the immune complex was assayed at 30 C for 20 min in a
volume of 30 µl of kinase buffer containing 5 µCi
[
-32P] ATP, and 2 µg of GST-c-Jun or
GST-ATF-2 as substrates. Reactions were stopped by the addition of
SDS-nonreducing-gel loading buffer with 18.5 mg/ml of iodoacetamide and
heating at 80 C for 3 min. Proteins were resolved by SDS-9% PAGE,
transferred onto nitrocellulose (Schleicher & Schuell, Inc., Dassel, Germany) and exposed to X-Omat autoradiography
films (Eastman Kodak Co.). Autoradiograms were scanned
using a Molecular Dynamics scanner (Sunnyvale, CA).
Immunoblotting
Nitrocellulose membranes were blocked at room temperature for
2 h with Tris-buffered saline containing 5% milk proteins and
0.1% Tween 20 (TTBS). The blotted proteins were probed overnight at 4
C with anti-p38 and anti-JNK-1 antibodies diluted 1:5000 in TTBS
containing 2% BSA. The secondary antibody used for detection was
labeled with antirabbit horseradish peroxidase. The blots were washed
and developed using the ECL chemiluminescence system (Amersham Pharmacia Biotech), according to manufacturers
instructions.
Statistical Analysis
Data were compared by Students t test with a
significant level of 95% (P < 0.05) or 99%
(P < 0.01).
 |
ACKNOWLEDGMENTS
|
---|
We would like to thank Paul A. Kelly and Juan M. Redondo for
providing PRLR-vectors and GST-c-Jun vector. We also thank L. Alvarez
and J. L. Castrillo for their help in reviewing the
manuscript.
 |
FOOTNOTES
|
---|
Address requests for reprints to: Josefa P. García-Ruiz, Departamento de Biología Molecular Facultad de Ciencias, Universidad Autónoma de Madrid, Cantoblanco, 28049 Madrid Spain.
This work has been supported by Grants 08.6/0015 (Com-unidad
Autónoma de Madrid), PM980023 (Programa Sectorial de
Promoción General del Conocímíento) and Ramón
Areces Foundation.
Received for publication July 8, 1999.
Revision received December 1, 1999.
Accepted for publication January 5, 2000.
 |
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