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The importance of prolactin (PRL) in physiological proliferation and differentiation of the mammary gland, together with high levels of PRL receptors in breast tumors, the association of circulating PRL with incidence of breast cancer, and the recognition of locally produced PRL, point to the need for greater understanding of PRL actions in mammary disease. Although PRL has been shown to activate multiple kinase cascades in various target cells, relatively little is known of its signaling pathways in the mammary gland apart from the Janus kinase 2/ signal transducer and activator of transcription 5 pathway, particularly in tumor cells. Another potential effector is activating protein-1 (AP-1), a transcription complex that regulates processes essential for neoplastic progression, including proliferation, survival and invasion. We demonstrate that PRL activates AP-1 in MCF-7 cells, detectable at 4 h and sustained for at least 24 h. Although Janus kinase 2 and ERK1/2 are the primary mediators of PRL-induced signals, c-Src, phosphatidylinositol 3′-kinase, protein kinase C, and other MAPKs contribute to maximal activity. PRL activation of these pathways leads to increased c-Jun protein and phosphorylation, JunB protein, and phosphorylation of c-Fos, elevating the levels of AP-1 complexes able to bind DNA. These active AP-1 dimers may direct expression of multiple target genes, mediating some of PRL’s actions in mammary disease.
In normal mammary development, the peptide hormone prolactin (PRL) is critical for alveolar proliferation and differentiation. In vitro, it also enhances cell survival and motility, and together, these activities suggest a role in human breast cancer. However, this has been controversial; attempts to correlate circulating PRL levels and breast cancer were conflicting, and inhibition of pituitary PRL synthesis with bromocryptine did not alter the disease course (reviewed in Refs. 1–3). More recently, however, new findings have strongly supported a role for PRL in mammary carcinogenesis. A large prospective study within the Nurses’ Health Study demonstrated a correlation between circulating PRL and breast cancer incidence (4), and higher PRL levels in another study were associated with increased mammographic density, a risk factor for breast cancer (5). Moreover, the discovery of local PRL production within mammary epithelial cells indicates autocrine/paracrine signaling, independent of pituitary PRL (reviewed in Ref. 1).
PRL signals through a complex web of kinases including Janus kinase 2 (Jak2), Src kinase, phosphatidylinositol 3′-kinase (PI3K), protein kinase C (PKC), and MAPKs (reviewed in Refs. 1 and 6). The Jak2/ signal transducer and activator of transcription 5 (Stat5) pathway is critical for PRL actions in alveolar development and has been the best characterized signaling pathway employed by PRL (reviewed in Ref. 7). However, little is known about the role of other pathways that may mediate PRL actions in the mammary gland. Furthermore, it is likely that PRL pathways and targets may shift as disease progresses and neoplastic changes accumulate.
Another potential PRL target is activating protein-1 (AP-1), a transcription factor complex that plays fundamental roles in multiple cellular processes, including cell proliferation, survival and differentiation (reviewed in Refs. 8–13). Several families of AP-1 components, including Jun and Fos proteins, form homo- or heterodimers that regulate transcription by binding to closely related DNA sequences originally termed 12-O-tetradecanoylphorbol-13-acetate (TPA)-responsive elements, or AP-1 sites. Depending on the components of the AP-1 complex, different genes can be targeted, or opposite transcriptional effects elicited. Induced expression of some AP-1 proteins in vitro can result in cell transformation and proliferation, and overexpression in transgenic models has been shown to result in tumor formation, including osteosarcoma, lung, skin, and liver tumors. Many genes important in carcinogenesis and tumor progression are regulated by AP-1 enhancer sequences, including collagenase, matrix metalloproteinases, and proteases of the urokinase plasminogen-activator system, TGFβ, epidermal growth factor receptor, and the cell cycle regulators p53, cyclin D1 and A, and p16 and p21CIP/WAF (reviewed in Refs. 8, 9, 12, and 14). AP-1 activity and expression of individual AP-1 proteins have been examined in human breast tumors, and DNA binding activity and Jun/Fos family member expression have correlated with tumor grade (15, 16), cell cycle-regulatory protein expression (17), estrogen receptor expression and/or tamoxifen resistance (18, 19), and metastases (15). These studies support a role for AP-1 in breast cancer and underscore the need to study AP-1 as a possible target for PRL in mammary pathogenesis.
The composition of AP-1 dimers depends on the relative expression of AP-1 components, which varies with cell type as well as environment. Levels of AP-1 proteins are tightly regulated at many levels, including transcription, mRNA stability, and protein stability (reviewed in Refs. 10, 20, and 21). Expression of c-Jun and c-Fos, in particular, is dramatically increased after exposure to many stimuli, resulting in proliferation and/or transformation in a variety of cell types. Multiple MAPK family members, including c-Jun N-terminal kinases (JNKs), ERKs, and p38 MAPK, have been implicated in transcriptional regulation. These kinases also can phosphorylate AP-1 components, enhancing DNA binding affinity, transactivating potential, and stability (reviewed in Refs. 9 and 22).
Activation of JNK was implicated in PRL-induced proliferation of bovine mammary epithelial cells (23), the rat lymphoma Nb2 cell line (24), and the pheochromocytoma PC12 cell line (25). This was linked to c-Jun and AP-1 activity in some studies (23, 25). However, upstream mediators and other MAPKs converging on this transcription factor complex, as well as the role of other AP-1 components, have not been explored.
The study of PRL effects on human breast cancer cells has been complicated by the production of PRL within the mammary epithelial cells themselves. We have derived cells from the well-characterized, hormonally responsive MCF-7 cell line that do not express endogenous PRL, but retain the ability to respond to exogenous PRL (26). In this PRL-deficient MCF-7 cell model, we have shown that PRL alters levels of cell cycle regulators and increases cell proliferation through many signaling pathways (26, 27). Overexpression of c-Jun in the parental cells increased tumorigenicity, invasiveness, and motility (28, 29), and adriamycin-resistant cells displayed increased AP-1 activity (30), demonstrating that this AP-1 protein regulates clinically relevant target genes in this breast cancer cell line. To investigate the mechanism whereby PRL regulates AP-1 activity in the PRL-deficient MCF-7 cell line, we used an AP-1 reporter construct, which preferentially binds Jun and Fos AP-1 family members. We found that PRL employs multiple proximal signaling pathways, as well as multiple MAPKs, particularly ERK1/2, to maximally activate AP-1. Activation of these kinases increases protein levels of c-Jun and JunB, as well as phosphorylation of both c-Jun and c-Fos. Together, these data indicate that PRL signals to AP-1 through multiple pathways that may modulate cell proliferation and aggressive tumor behavior in breast cancer cells.
To examine the ability of PRL to activate AP-1 in breast cancer cells, we transiently transfected PRL-deficient MCF-7 cells (26) with an AP-1 reporter plasmid containing four AP-1 tandem repeats (4XAP-1-luc) (31) and treated cells with 4 nM PRL. PRL rapidly activated AP-1 activity, which peaked at 4 h and was sustained for at least 24 h (Fig. 1A). Based on these results, we further investigated PRL activation of AP-1 after 6 h exposure to PRL.
Although the predominant PRL receptor (PRLR) iso-form in MCF-7 cells is the long PRLR (lPRLR), transcripts for the intermediate PRLR (iPRLR) isoform are also present in MCF-7 cells (Brockman, J. L., and L. A. Schuler, unpublished observations). The differing cytoplasmic domains result in different signaling capabilities; unlike the lPRLR, iPRLR is able to activate only Jak2 and not the Src family member, Fyn (32). To determine the capacity of the lPRLR and iPRLR to transduce signals to AP-1, we examined PRL-induced AP-1 activity in the presence of endogenous PRLR isoforms only or supplemental lPRLR or iPRLR after cotransfection (Fig. 1B). In the presence of the endogenous complement of receptors, PRL activated AP-1 activity about 2-fold. Transfected iPRLR did not further increase PRL stimulation. However, cells transfected with lPRLR demonstrated an increase in unstimulated activity as well as a greater PRL response (> 3-fold). PRL, in the presence of the lPRLR, did not activate the enhancer-less parent vector, pXP2. These results suggest that PRL signals to AP-1 in these cells primarily via the lPRLR isoform. For subsequent experiments, the lPRLR was transfected along with the AP-1-responsive reporter to maximize signal.
To confirm that PRL induced AP-1 activity was not a result of PRL-induced increases in cell cycle progression (26), we pretreated cells with cell cycle inhibitors for 24 h (1 mM hydroxyurea or 100 ng/ml nocodazole to block the cells in G0/G1 or G2/M, respectively). PRL-induced AP-1 activity was slightly, but not significantly, lower in the presence of both hydroxyurea and nocodazole (Fig. 1C), demonstrating that PRL can activate AP-1 independently of effects on the cell cycle.
PRL binding to the lPRLR initiates multiple signaling cascades in mammary epithelial cells, although the Jak2/Stat5 pathway has been the most studied (reviewed in Refs. 1, 6, and 7). Moreover, several pathways have been shown to be involved in PRL-induced proliferation in our cell model (27). To determine the proximal pathways important for PRL signaling to AP-1 in this system, we used dominant negative (DN) constructs or selective inhibitors for Jak2, c-Src, PI3K, and PKC. As shown in Fig. 2, many pathways also mediate PRL signals to AP-1. Transfection with DN Jak2 (as well as treatment with AG490; data not shown) completely blocked PRL activation of AP-1 without altering unstimulated levels, indicating a critical role for this kinase. DN Src (as well as PP1, data not shown) decreased, but did not eliminate, PRL-induced activity, as did the selective inhibitors for PI3K and PKC, LY294002, and Bisindolylmaleimide II (BisII), respectively. The failure of these agents to completely block PRL-induced activity suggested that c-Src, PI3K, and PKC are all important for maximal PRL induction of AP-1, but are not required. DN Src, LY294002, and BisII also reduced unstimulated AP-1 activity, perhaps due to inhibition of signals initiated by factors secreted during the preincubation period. In contrast, treatment with other inhibitors such as ICI 182,780 had no effect on PRL-induced AP-1 activity (data not shown). This array of proximal kinases that mediate PRL activation of AP-1 is consistent with the ability of PRL to stimulate these pathways in various cell types (reviewed in Refs. 6 and 14), and the demonstrated roles of these kinases in initiation of MAPK cascades, critical for AP-1 activation (reviewed in Refs. 18 and 26).
AP-1 family members have been shown to be activated by phosphorylation cascades, especially those involving the MAPKs, to enhance their transcriptional activity and/or increase their synthesis (reviewed in Refs. 9 and 22). To investigate the ability of PRL to activate MAPKs in this cell line, cells were treated over a short time course with PRL, and cell lysates were analyzed for MAPK phosphorylation, which is associated with activation of these proteins. As shown in Fig. 3, PRL strongly and transiently induced ERK1/2 and ERK5 phosphorylation within 15 min and 5 min, respectively. Activation of JNK1/2 and p38 was apparent at 5 min and remained elevated for at least 135 min. These results indicated that many possible mechanisms are available to PRL to activate AP-1.
To further investigate the role of ERKs in PRL-induced AP-1 activity, we used the selective ERK inhibitors, PD98059 and U0126. Although, historically, these compounds have been used as specific inhibitors for the MEK1/2-ERK1/2 cascade, it has been shown recently that PD98059 and U0126 blocked EGF-stimulated ERK5 phosphorylation in HeLa cells (33) and may also inhibit other unknown signaling cascades. As shown in Fig. 4A, PD98059 decreased PRL-induced ERK1/2 phosphorylation only moderately and had no effect on PRL-induced ERK5 or JNK1/2 phosphorylation. However, U0126 blocked the PRL-induced ERK1/2 phosphorylation, without affecting ERK5 phosphorylation. Although it slightly decreased levels of phosphorylated JNK1/2 in unstimulated cells, U0126 did not alter the magnitude of the PRL response. We therefore used U0126 to further investigate PRL activation of AP-1. This compound blocked PRL induction of AP-1 (Fig. 4B), suggesting that ERK1/2 are involved.
To confirm the results with U0126, we examined the role of ERK1/2 using dominant negative (DN) ERK1 and ERK2 constructs (Fig. 4C). Cotransfection with DN ERK1/2 dramatically decreased PRL-induced AP-1 activity from about 4-fold to 1.5-fold, and slightly decreased unstimulated activity. However, the response was not completely blocked, and transfection of additional DN ERK1/2 did not further reduce the response (data not shown), suggesting that additional kinases were involved. To investigate other MAPKs, we employed DN JNK1/2, DN ERK5, kinase-dead p38α (p38αKD) constructs, and the p38-selective inhibitor, SB202190. As shown in Fig. 4D, JNK1/2, ERK5, and p38α were less important than ERK1/2, but all played roles in maximal activation of AP-1 by PRL. SB202190 (10 μM) had an effect similar to the p38α KD construct (data not shown). JNK1/2 were also important for maintenance of unstimulated AP-1 activity. Overexpression of the JNK-interacting protein-1 (JIP-1), which binds JNKs in the cytoplasm and inhibits their action (34), confirmed the findings with DN JNK1/2. As shown in Fig. 4E, JIP-1 overexpression resulted in a dose-dependent decrease in both un-stimulated activity as well as PRL-induced activity. Together, these data indicate that ERK1/2 are the most critical MAPKs involved in PRL-induced AP-1 activity in this model system, whereas JNK1/2 are critical determinants of unstimulated and therefore potential maximal activities. ERK5 and p38 play minor roles in PRL-induced AP-1 activity.
In MCF-7 cells, JunB and c-Fos are the predominant AP-1 components, although cycling MCF-7 cells also express c-Jun, JunD, and FosB, but not Fra-1 (35). To determine whether PRL treatment increased the ability of AP-1 complexes to bind DNA, we performed EMSAs. As expected, we observed an AP-1-bound complex in the absence of hormone. PRL increased AP-1 DNA binding, apparent after 30 min of treatment, which remained elevated for at least 24 h (Fig. 5A). To determine the composition of the DNA-bound complex, we incubated nuclear extracts from cells treated with PRL for 3 h with specific antibodies for Jun/Fos family members (Fig. 5B). Antibodies to c-Jun, JunB, and c-Fos all partially inhibited formation of the un-stimulated as well as the PRL-induced complex, possibly due to disruption of DNA binding sites. Antibodies to JunD and FosB had no effect on the AP-1 complex. Together, these results are consistent with increases in c-Jun, JunB, and c-Fos activity mediating the PRL-induced increase in AP-1 activity.
Based on the established role for c-Jun in the growth of breast cancer cells in vitro (28, 36), we examined the importance of c-Jun in our system, employing the dominant negative c-Jun, TAM-67. TAM-67 is a truncated c-Jun mutant that lacks the transactivation domain, but retains the ability to dimerize with other AP-1 proteins and to bind DNA (37). Increasing concentrations of TAM-67 resulted in dose-dependent decreases in both unstimulated, as well as PRL-induced activity (Fig. 5C). However, the ability of PRL to activate AP-1 above the corresponding unstimulated control remained constant. These data suggest that c-Jun is critical for both unstimulated AP-1 activity, as well as determining the potential total AP-1 activity after exposure to PRL. Interestingly, the effect of TAM-67 was very similar to the results with the JIP-1 construct (Fig. 4E), consistent with the importance of JNK1/2 in control of c-Jun activity.
To further address the mechanism whereby PRL activates AP-1, we investigated the effect of PRL treatment on expression and phosphorylation of AP-1 components. As shown in Fig. 6A, PRL induced a transient increase in c-Jun phosphorylation at Ser 63, detectable at 15 min. PRL also raised levels of total c-Jun and JunB protein, peaking after 1 h. Total c-Jun and JunB were increased 4.9 ± 0.36 and 3.77 ± 0.10 fold, respectively, at this time (mean ± SD of three independent experiments). PRL did not affect the protein expression of c-Fos, JunD, or FosB during this period. However, a longer exposure revealed PRL-induced modifications of c-Fos that altered mobility, including a prominent signal of reduced mobility (solid arrowhead), and a more variable faint band of increased mobility (open arrowhead). Due to the lack of antibodies to detect phosphorylation of c-Fos, we employed λ-protein phosphatase (λ-PPase) as done previously (38), to determine whether the upper band indicated a phosphorylation event. As shown in Fig. 6B, in the presence of λ-PPase, this signal was not detectable, supporting this likelihood. In contrast, the PRL-induced lower band was not affected, suggesting some other modification.
The observations that PRL-induced AP-1 activity is blocked by U0126 (Fig. 4B), and that c-Jun, JunB, and c-Fos are targets for PRL, and participate in PRL-induced complexes (Figs. 5 and and6),6), suggested that U0126 should inhibit the PRL-induced changes in c-Jun, JunB, and c-Fos. As expected from the effect on basal levels of total cellular phosphorylated JNK1/2, pretreatment with U0126 decreased levels of phosphorylated c-Jun in unstimulated cells, but not the fold induction by PRL (1.9 ± 0.16; n = 3, mean ± SE). It also partially inhibited the PRL-induced increases in c-Jun protein, completely blocked increases in JunB protein, as well as greatly reduced the PRL-induced phosphor-ylation of c-Fos (Fig. 6C). Together, these results indicate that PRL-induced AP-1 activity in these breast cancer cells is mediated primarily by a U0126-sensitive pathway involving ERK1/2, and that c-Jun, JunB, and c-Fos are the important AP-1 components that mediate this response.
The importance of PRL in physiological proliferation and differentiation of the mammary gland, together with the high level of PRLR in breast tumors, the association of circulating PRL with incidence of breast cancer, and the recognition of locally produced PRL, call for greater understanding of its actions in mammary disease. Little is known about the breadth of pathways whereby this cytokine/hormone may directly modulate or synergize with other oncogenes in the disease process. Here we have demonstrated that PRL is a potent inducer of AP-1 activity in breast cancer cells, which may alter expression of many target genes that are critical for neoplasia, including cell survival, proliferation, differentiation, angiogenesis, and invasion. As depicted in Fig. 7, PRL activates this transcription factor by complex signaling cascades, involving multiple proximal pathways, including Jak2, c-Src, PI3K, and PKC, although only Jak2 is required for a PRL response. ERK1/2 are the primary downstream activators of c-Jun, JunB, and c-Fos in this system, resulting in increased DNA binding of these components. Although JNK1/2 activity is a major determinant of the magnitude of AP-1 activity in this system, it is not necessary for the response to PRL, in contrast to other reports. ERK5 and p38 are minor effectors of the PRL response.
Most studies have focused on Jak2 as a proximal mediator of PRL action. However, PRL also activates other pathways in various target cells, including c-Src, PI3K, and PKC (reviewed in Refs. 1 and 6), although the hierarchical relationships between Jak2 and these other kinases are not well understood. Because these proteins also activate MAPKs, it is not surprising that all these pathways play roles in PRL signaling to AP-1. We have previously shown the importance of Jak2, PI3K, and c-Src in PRL induction of the critical cell cycle regulator, cyclin D1, and in PRL-induced proliferation of this MCF-7 derived cell line (27). PRL induction of c-Src has been also reported to be a critical mediator of proliferation and activation of PI3K and ERK1/2 in MCF-7 parent cells as well as T47D breast cancer cells (39). In contrast, although PRL has been shown to activate PKC-linked signaling cascades (40), the role of these kinases in PRL action has received relatively little attention in any cell type, especially in breast cancer cells. Whereas Jak2/Stat5 mediates most PRL actions in alveolar development (reviewed in Ref. 7), and Jak2 is required for PRL activation of AP-1 in PRL-deficient MCF-7 cells, Stat5 is not required for the AP-1 response, and indeed inhibits this pathway (Gutzman, J. H., L. M. Arendt, D. E. Rugowski, S. E. Nikolai, H. Rai, and L. A. Schuler, manuscript in preparation). Our findings underscore the wide range of signaling pathways activated by PRL and the need for further investigation into this kinase network.
PRL also activated many MAPK family members in these breast cancer cells. Whereas ERK1/2 and JNK1/2 have been reported to be downstream of the PRLR in several in vitro models, our data indicate that PRL also is able to initiate phosphorylation of ERK5 and p38, suggesting other unexplored distal targets. Whereas various p38 isoforms have been reported to differentially affect AP-1 activity (41), p38α accounted for the modest effect observed with a p38-selective inhibitor in the current study. All of these MAPKs have been shown to modulate AP-1 activity in response to other stimuli (reviewed in Refs. 9 and 22); the ability of DN constructs specific for each family member to partially inhibit PRL-induced activation of AP-1 here suggests that they play specific roles in mediating PRL action and that they are not redundant pathways.
AP-1 enhancer activation requires increased Jun/ Fos expression, DNA binding affinity, and/or transactivational potential of one or more of these proteins. Our studies showed that PRL employed all of these mechanisms to increase the activity of dimers composed of c-Jun, JunB, and/or c-Fos. Phosphorylation of c-Jun at Ser-63 was the earliest detectable modification after exposure to PRL; this modification by JNK1/2 is a well-characterized event that potentiates its transactivation properties by enhancing recruitment of coactivators (reviewed in Ref. 22). Activated c-Jun, in combination with other transcription factors stimulated by a complex network of other MAPKs, increases c-jun transcription (11, 42). Indeed, in our studies, increased c-Jun protein was detectable after 1 h exposure to PRL. U0126 also partially blocked the PRL-induced rise in c-Jun levels; this is consistent with roles for both JNK1/2 as well as the U0126-sensitive ERKs in this event. In addition, PRL elevated levels of JunB, apparent after 1 h. Like the other AP-1 components, amounts of this protein can be regulated at many levels. However, two Serum Response Elements in the promoter can be activated by MAPK-activated Ternary Complex Factor, providing at least one potential mechanism. In contrast, net levels of c-Fos were not altered over the time examined here. Although MAPKs can also activate multiple cis-inducible elements in the c-fos promoter (reviewed in Ref. 22), it is possible that increased protein turnover may have stabilized steady-state levels in our studies (38, 43). However, PRL-induced, U0126-sensitive phosphorylation of c-Fos was clearly evident. This is consistent with the well-characterized phosphorylation of c-Fos at multiple sites in the C-terminal transactivation domain primarily by ERK1/2 and downstream kinases, which enhances its transactivation potential (38, 44). Together, our data support a model whereby PRL directs activation of a complex web of MAPKs to alter AP-1 activity at multiple levels. Interestingly, the effects observed here on AP-1 proteins were transient; the dynamic mechanism(s) sustaining the PRL-induced activity are under investigation.
These studies document the ability of PRL to augment signaling through AP-1 components but do not address the likely plethora of target genes. The clear evidence that DNA binding of AP-1 proteins is dependent on flanking sequences and promoter context (reviewed in Refs. 3 and 45–47) suggest that the interactions detected with our reporter construct and EMSAs may be a subset of the total activity. Analysis of gene expression in diverse cell types in response to overexpression of individual components and tethered AP-1 dimers has confirmed the multiplicity of factors determining these targets and is beginning to reveal the diverse AP-1 repertoire (12, 21, 29, 48, 49). Cell context, dictating levels of potential dimerization partners and activity of cooperating pathways, is paramount in determining the response. JunB, for example, is generally considered antiproliferative; however, it increases cyclin A transcription permitting entry into S phase (50), up-regulates angiogenic factors during progression of fibrosarcomas (51), transforms Rat1a fibroblasts (48), and can replace c-Jun during development (reviewed in Refs. 8 and 9). In addition to the direct DNA binding examined here, PRL-activated AP-1 components may also modify gene expression via interaction with a multitude of other transcription factors, such as nuclear factor-κ B, NFAT, Smad, Ets, Stats, and various nuclear receptors (reviewed in Refs. 13 and 21). This well-documented underlying complexity suggests that targets of PRL-activated AP-1 may differ in breast tumors of various etiologies or as tumorigenesis progresses.
Our studies demonstrated the ability of PRL to direct a complex signaling network resulting in activation of AP-1 components in breast cancer cells, which may modify numerous aspects of tumor cell behavior. Many other growth factors and hormones that play central roles in the normal mammary gland or carcinogenesis can also activate AP-1, providing opportunities for cross-talk. Investigation of PRL interactions with these factors in different cell contexts and responsive AP-1 target genes will increase our understanding of the role of PRL in mammary neoplasia and aid in the development of new therapeutic approaches for breast cancer.
The following antibodies were used for Western analyses: c-Jun, phospho-c-Jun (Ser-63), ERK1/2, phospho-ERK1/2, ERK5, phospho-ERK5, p38, and phospho-p38 from Cell Signaling Technology (Beverly, MA); c-Jun (sc-45X), c-Fos (sc-52X), JunB (sc-46X), JunD (sc-74X), and FosB (sc-48X) from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). Recombinant hPRL (lot AFP795) was obtained through the National Hormone and Pituitary Program, NIDDK, and Dr. Parlow. The following inhibitors were used in some experiments: U0126 from Promega Corp. (Madison, WI); PD98059 and BisII from Calbiochem (San Diego, CA); and LY294002 from Sigma-Aldrich Corp. (St. Louis, MO) and PP1 (4-amino-1-tert-butyl-3-(1′-naphtyl)pyrazolo[3,4α] pyrimidine from Biomol (Ply-mouth Meeting, PA). λ-PPase (no. P0753S) was obtained from New England Biolabs, Inc. (Beverly, MA). All other reagents were obtained from Sigma-Aldrich Corp. unless otherwise noted.
The AP-1-luciferase construct (4XAP-1-luc) contains four GCN4 consensus AP-1-responsive elements located upstream of a luciferase reporter (31). The control parent lucif-erase vector (pXP2-luc) does not contain the AP-1 elements. Expression constructs employed were: DN Jak2 -Δ 829 (DN Jak2) from D. M. Wojchowski (52); DN c-Src K296R/Y528F (DN Src) from Upstate Biotechnology, Inc. (Lake Placid, NY); DN ERK1 K71R (DN ERK1), and DN ERK2 K52R (DN ERK2) from M. Cobb (53); DN ERK5-AEF (DN ERK5) from J. D. Lee (54); JIP-1 from M. Dickens (34); DN c-Jun (TAM-67) from M. J. Birrer (37); KD p38α from P. Lobie (55); DN JNK1-APF (DN JNK1) and DN JNK2-APF (DN JNK2) from L. Heasley (56); iPRLR and lPRLR isoforms from C. Clevenger (32).
PRL-deficient MCF-7 cells were maintained as previously described (26). Cells were grown 4–5 d before all experiments in phenol red-free RPMI-1640 containing 5% fetal bovine serum stripped three times with charcoal (3× charcoal-stripped serum), penicillin, and streptomycin. Cells were plated into 12-well tissue culture plates at 3 × 105 cells per well and allowed to adhere overnight. The next day, cells were serum starved for 24 h and then transiently transfected using SuperFect (QIAGEN, Valencia, CA) as previously described (57). Within each experiment, total amount of transfected DNA was equalized with vector DNA. After 4 h, the transfection complex was replaced with serum-free media ± 4 nM PRL for 6 h, or allowed to recover from transfection in serum-free media overnight and then treated ± 4 nM PRL for 6 h, depending on the experiment. Cell lysates were harvested and analyzed for luciferase and β-galactosidase activity, and luciferase values were corrected for transfection efficiency as described (57). Relative activity is the mean of at least three independent experiments represented as fold change relative to the vehicle control.
PRL-deficient MCF-7 cells were maintained as above and plated at 1 × 106 cells per 100-mm dish, serum starved for 24 h, and treated ± 4 nM PRL for different times. For Western analyses, cell lysates were harvested and analyzed as described previously (26). Primary antibody concentrations were as follows: phospho-ERK1/2, 1:5,000; ERK1/2, 1:1,000; phospho-JNK, 1:1,000; JNK, 1:1,000; phospho-ERK5, 1:1,000; ERK5, 1:1,000; p38, 1:1,000; phospho-p38, 1:1,000; c-Jun, 1:1,000; phospho-c-Jun, 1:1,000; c-Fos, 1:10,000; JunB, 1:10,000; JunD, 1:10,000; and FosB, 1:10,000. Signals were visualized by enhanced chemiluminescence, followed by autoradiography. For some experiments, the signals were quantified by densitometry (ImageQuant software, version 4.2a; Molecular Dynamics, Inc., Sunnyvale, CA).
PRL-deficient MCF-7 cells were plated at 1.5 × 106 cells per 100-mm dish, serum starved for 24 h, and treated ± 4 nM PRL for different times. Nuclear extracts were prepared and EMSAs were performed as previously described (57). The AP-1 probe consisted of a synthetic double-stranded DNA oligonucleotide (Integrated DNA Technologies, Coralville, IA) containing a single AP-1 consensus site with the following sequence (5′-CGC TTG ATG ACT CAG CCG GAA-3′). The oligonucleotide was labeled with [γ-32P]ATP (PerkinElmer Life Sciences, Boston, MA) and 5 μ g of nuclear protein were incubated with the labeled probe for 20 min at room temperature. To assay for the specificity of binding to the AP-1 DNA sequence, unlabeled oligonucleotide was incubated with nuclear protein for 20 min before addition of labeled oligonucleotide. To assay for antibody competition, nuclear protein was incubated with 2 μg of antibody (c-Jun sc-45X, c-Fos sc-52X, JunB sc-46X, JunD sc-74X, or FosB sc-48X) for 40 min at 25 C before the addition of labeled oligonucleotide. These antibodies have been successfully used previously for EMSAs (58, 59). Samples were fractionated on a nondenaturing 4% polyacrylamide gel, and the complexes were visualized using a Storm Phospho-Imaging System (Molecular Dynamics, Piscataway, NJ).
Cells were grown and plated as described above for immunoblotting. After treatment with or without 4 nM PRL for 1 h, cells were lysed in 25 mM Tris, 2 mM EDTA, 10% glycerol, 1% Triton-X 100, 2 mM Na3VO4, and 20 mM NaF. Samples were diluted and supplemented with 10 μg/ml leupeptin, 10 μg/ml aprotinin, and 0.5 mM PMSF. Each cell extract (30 μg) was incubated in the presence or absence of λ-PPase at 600 U/50μl total volume in the supplied buffer for 1 h at 30 C. The reaction was stopped by the addition of 50 μl of 2× SDS-PAGE loading buffer, and 10 μg of protein were analyzed by immunoblotting as described above.
We are grateful to the many generous investigators who made constructs available for these studies.
This work was supported by National Institutes of Health Grant R01 CA 78312 and Department of Defense Breast Cancer Research Grant DAMD17-01-1-0460.