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


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
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.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
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.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
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. 1Go. 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. 1AGo). 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. 1BGo), 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. 1CGo). 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.

 
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. 2Go, 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. 2AGo), as already reported (27), while Luc activity driven by AP-1 elements was diminished (P < 0.05) (Fig. 2BGo). 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.

 
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. 3AGo, 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.

 
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. 3BGo.1) or with a mixture of PRLR-L and PRLR-S constructs (Fig. 3BGo.2). The amount of DNA was maintained at 3.33 µg by adding pCMV plasmid when necessary. As can be observed in Fig. 3BGo.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. 3BGo.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. 4AGo, lane 1). A representative result of electrophoretic mobility shift assays (EMSAs) is shown in Fig. 4AGo. 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 2–6). 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.

 
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. 4BGo, 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. 5AGo), 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. 5BGo). Films were overexposed to detect the supershifted complex. These results were consistently observed in four independent experiments, as shown in the densitometric quantification (Fig. 5CGo).



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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.

 
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. 6AGo. 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. 6BGo), 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. 6CGo. 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 2–5). 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 ({gamma}-32P)-ATP. B, BMGE cells were transfected with pCMV control vector (lanes 1 and 7) or with PRLR-L expression vector (lanes 2–6). 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 3–6). After PRL stimulation, Dex at the specified concentrations was added to cells for 6 h (lanes 4–6). 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).

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
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 both—the 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 {Delta}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
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
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 70–80% of confluence. Then, they were starved overnight in GC3 medium composed of 1:1 DMEM and Ham’s 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 45–48 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 (100,000–150,000 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 [{gamma}-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 [{gamma}-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 manufacturer’s instructions.

Statistical Analysis
Data were compared by Student’s 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), PM98–0023 (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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 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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