Differentiation-Dependent Prolactin Responsiveness and Stat (Signal Transducers and Activators of Transcription) Signaling in Rat Ovarian Cells

Darryl L. Russell and JoAnne S. Richards

Baylor College of Medicine Department of Cell Biology Houston, Texas 77030


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
PRL activates an important cytokine signaling cascade that is obligatory for maintaining luteal cell function in the rat ovary. To determine when specific components of this cascade are expressed and can be activated by PRL, we analyzed the expression of receptor subtypes (short, PRL-RS, and long, PRL-RL), the presence and kinetics of Stat (signal transducer and activator of transcription) activation using the PRL-response element (PRL-RE) of the {alpha}2M ({alpha}2-macroglobulin) gene, and the content and hormonal regulation of three specific modulators of cytokine signaling; the tyrosine phosphatases (SHP-1 and SHP-2), and the protein inhibitor of activated Stat3 (PIAS-3). These components were analyzed in differentiating granulosa/luteal cells of hypophysectomized (H) rats and in corpora lutea of pregnant rats. Levels of PRL-R mRNAs increased as granulosa cells differentiated and reached maximal levels in luteal cells of pregnant rats where levels of PRL-RS approached those of PRL-RL. The relative concentrations shifted from a 27-fold excess of PRL-RL in preovulatory granulosa cells to a 3.7-fold difference in luteal cells during midgestation. Despite the increased PRL-RL expression in differentiated granulosa cells, PRL did not stimulate detectable activation of Stats. Rather PRL activation of Stat5, principally Stat5b, occurred in association with luteinization. In contrast, granulosa cells of untreated immature and H rats contained a high level of DNA binding activity, which was shown to be comprised entirely of activated, phosphorylated Stat3. Treatment with estrogen and FSH reduced the amount of phosphorylated Stat3 and abolished its ability to bind DNA, an effect temporally related to increased PIAS-3. Expression of SHP-1 (but not SHP-2) was also hormonally regulated; SHP-1 mRNA and protein were high in granulosa cells of H rats, decreased by estrogen and FSH, and subsequently increased dramatically with luteinization. Of particular note, SHP-1 was localized in cytoplasm of granulosa cells in atretic follicles but was distinctly nuclear in luteal cells, indicative of different functional roles. Collectively, these results indicate that Stat3 and Stat5 are activated by distinct cytokine-signaling pathways modulated through differentiation-dependent transcriptional regulation of signaling pathway components and mediate distinct functional processes in the rat ovary: early follicle growth and atresia vs. luteinization.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The mechanisms controlling the progression of granulosa cell fate (growth vs. atresia) and terminal differentiation (luteinization) are not well understood. During the ovarian cycle numerous cytokines modulate gene expression and function (Ref. 1 for review), while luteinization involves the acquisition of PRL-induced Stat5 (signal transducer and activator of transcription 5) responsiveness, a process requisite for maintenance of luteal cell function. The specific modulation of cytokine-signaling mechanisms through progression of follicle differentiation and acquisition of PRL-responsive Stat5-mediated gene induction during luteinization comprise the primary focus of the studies described herein. PRL is a pleiotropic pituitary hormone long known for its luteotropic actions in the rodent (2). PRL receptors (PRL-R) are expressed in the ovary (3, 4, 5) and have been shown to regulate multiple genes and events in ovarian follicular development and function, as evidenced by the recently reported PRL-R null mutant mice (6). Female PRL-R-/- mice never become pregnant and fail to establish pseudopregnancy, indicating impaired function of corpora lutea (CL). These PRL-R-/- mice also had fewer primary follicles, fewer ovulations, delayed or mistimed oocyte release, and impaired oocyte maturation, all signs of disruption in follicular development and possibly atresia. Additionally, PRL has been shown to increase vascular endothelial cell proliferation (7) and leukocyte infiltration in luteinizing ovaries as well as ovaries undergoing luteal regression (8). The pleiotropic effects of PRL in ovarian function are related to the different patterns of pituitary PRL secretion and release of placental lactogens (2), as well as the involvement of diverse components of the PRL-signaling cascade including: PRL-R subtypes (9), their associated Janus kinases (Jaks; Ref. 10), and substrates for Jak2 such as tyrosine phosphatases (11, 12), and Stat1, Stat3, and primarily Stat5 (Refs. 13, 14 for review).

Briefly, cytokine signaling occurs through ligand-induced dimerization of receptors which associate with and are phosphorylated by Jak2. Targets for Jak2 include Stats (13) and phosphotyrosine phosphatases (PTPs) (11, 12). In the rat, PRL-R are expressed in two variant forms, long (PRL-RL) and short (PRL-RS). As products of differential mRNA splicing from a single gene, the ligand binding and transmembrane regions as well as 44 amino acids of the cytoplasmic domains are identical, but these receptor isoforms diverge over the majority of the intracellular sequences (9, 14). Importantly, although both forms associate with Jak2, the signaling events mediated by PRL-R remain controversial and may be more complicated than originally proposed (14, 15, 16, 17, 18, 19). In particular, PRL-RS appears not to transduce Stat5-dependent induction of milk protein genes in mammary cells in culture (15, 16, 20, 21). Thus, PRL-RL is thought to be the only endogenous PRL-R form capable of transducing a Stat5-dependent signal (22), while PRL-RS may act in a dominant-negative fashion to impair the activity of PRL-RL (23, 24).

Negative regulation of cytokine signaling downstream of Stat activation can also occur through several families of modulators. Two SH-2 domain containing PTPs have been shown to modulate PRL/GH-induced Stat responses. The PTP SHP-2 is a substrate for Jak2 and is obligatory for PRL/Stat5 induction of ß-casein (11). SHP-2 may bind P-Tyr residues on PRL-R, Jak-2, and/or Stat5 via dual SH-2 domains which, along with catalytic activity, are required to maintain the intact signaling pathway (11). The highly homologous phosphatase SHP-1 binds activated Stat5b catalyzing rapid dephosphorylation resulting in the transient activation pattern characteristic of this family of transcription factors (25). In addition, a recently described class of proteins can bind and neutralize activated Stats. One of these, PIAS-3 (protein inhibitor of activated Stat3), is a specific inhibitor of Stat3 DNA binding and transactivation (26). Thus, regulated expression of SHP-1, SHP-2, and PIAS-3 in the ovary and the manner in which they interact with PRL-R/Stat signaling complexes can modulate cell-specific responses to PRL and other cytokines.

Differential patterns of Stat activation and gene expression are also determined by the manner in which PRL is secreted in female rats. Pulsatile release of PRL from the pituitary occurs during early pregnancy (days 1–9) but is replaced at midpregnancy by elevated and chronic secretion of placental lactogenic hormones (rPL) until luteal regression and parturition (2). Several genes are regulated by these lactogenic hormones including the estrogen receptor (ER) subtype, ER{alpha}, which confers autocrine effects of estrogen to synergize with PRL and maintain luteal function and the progression of pregnancy to term (27). This luteotropic complex maintains expression of several genes in CL including P450arom (CYP19; Ref. 28) and LH receptor (29). Inhibitory effects of PRL are also exerted at this time on 20{alpha}-hydroxysteroid dehydrogenase expression, thereby preventing the metabolism of progesterone to its inactive metabolite 20{alpha}OH-progesterone (30). In addition, PRL and rat placental lactogen (rPL) induce and regulate {alpha}2-macroglobulin ({alpha}2M) expression by luteinized cells (31, 32). The initial increase in {alpha}2M occurring soon after luteinization is followed by a secondary increase in pregnant CL around day 10 stimulated by high, constitutive rPL release. We subsequently demonstrated that in luteal cells, but not granulosa cells, PRL activates Stat5b and Stat5a (33), which bind the {alpha}2M promoter to induce expression (34, 35).

In this study, we have addressed the question of how a pleiotropic factor such as PRL elicits specific effects in ovarian tissues at different stages of differentiation. Specifically we addressed three questions. How do the levels of PRL-R subtypes relate to the activation of Stat5 or Stat3? Is the activation of Stat3 or Stat5 related to specific stages of granulosa and luteal cell differentiation? Is the activation of Stat5 or Stat3 modulated by specific Stat regulators SHP-1, SHP-2, or PIAS? For this we have used two in vivo models. Hypophysectomized (H) rats were treated with estradiol (E) and FSH (HEF) to stimulate the growth of healthy preovulatory follicles that would otherwise become atretic. Luteal cell responses to PRL were analyzed in HEF rats treated with an ovulatory dose of hCG (HEF/hCG) and in pregnant rats when fully functional CL are maintained by endogenous pulsatile (day 1–9) or chronic (day 10–21) exposure to lactogenic hormones or are undergoing regression (day 21–post partum). Results of these studies provide the novel observation that expression of PRL-RS is increased markedly in luteal cells, suggesting a role other than to exert a dominant negative effect on Stat5 activation by PRL. Likewise, high levels of SHP-1 are present in luteal cell nuclei, suggesting it may exert positive regulatory effects on Stat activation/turnover. Lastly, high Stat3 DNA binding activity, as well as SHP-1 expression, are present in granulosa cells of unstimulated H rat ovaries. Thus, Stat3 and Stat5 are activated by different signaling pathways in the rat ovary, modulated by changes in positive and negative effectors, and associated with distinct stages of granulosa cell differentiation and function.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
PRL Receptor Long (PRL-RL) and Short (PRL-RS) Isoforms
To determine whether changes in the response of ovarian cells to PRL might be related to expression of specific forms of the PRL-R, the acute regulation of mRNA levels encoding the short and long forms of PRL-R was analyzed by specific semiquantitative, as well as quantitative, RT-PCR assays (3). The quantitative assay specifically amplifies each rat PRL-R isoform yielding single products of predicted molecular size; 422-bp PRL-RL and 332-bp PRL-RS (Fig. 1Go). Product identities were further confirmed by restriction digest analysis (data not shown). Truncated cDNA clones of each PCR amplicon were introduced into the RT-PCR reactions at titrated concentrations from 0.1–30 x 105 copies per tube. Competition of the cloned DNA with the RNA-derived cDNA from reverse transcriptase reactions permitted quantiative comparisons of the two receptor mRNAs (see Materials and Methods).



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Figure 1. Regulated Expression of PRL-R Long and Short Isoforms during Follicular Development and Luteinization

Competitive RT-PCR analysis of PRL-R isoforms using RNA from hormonally stimulated H rats or pregnant rats. Specific primer sets were used to amplify PRL-RL or PRL-RS mRNA-derived cDNA or truncated competitor introduced at indicated concentrations. A, Representative autoradiographs of amplified competitive RT-PCR products resolved on nonreducing acrylamide gels. mRNA was analyzed from immature/atretic (H), or preovulatory (HEF) granulosa cells or luteal cells of newly formed (HEF/hCG) or midgestational CL (day 15). B, Combined mean ± SEM data from three competitive RT-PCR analyses performed on RNA from granulosa or luteal cells. Quantitated results were calculated from phosphorimager analysis of target vs. competitor amplification as described in Materials and Methods. C, Semiquantitative RT-PCR analysis of specific PRL-R isoforms in immature/atretic granulosa cells (H), preovulatory granulosa cells (HEF), newly formed CL (HEF/hCG), or mature functional CL at early (day 7), midgestation (day 15), or regressing CL (day 22). PRL was administered as described in Materials and Methods for 1 h, 24 h, or 24 h followed by a 1-h pulse as indicated. Results are expressed as the ratio of each specific PRL-R isoform to the L-19 internal control; mean ± SEM values are shown for three repeated analyses.

 
By quantitative RT-PCR analysis we found that granulosa cells isolated from H rat ovaries contained near-undetectable levels of PRL-RL (0.4 x 106 copies/µgRNA) and PRL-RS (0.16 x 106 copies/µg RNA, Fig. 1Go, A and B). In preovulatory granulosa cells of HEF rats, each PRL-R isoform had increased 20-fold and 2-fold, respectively such that expression of PRL-RL (7.7 x 106) was 27-fold greater than PRL-RS (0.28 x 106). PRL treatment at this stage of differentiation resulted in a time-dependent reduction of both PRL-R isoforms, making their expression after 24 h similar to that in H granulosa cells (not shown). However, 24 h of hCG treatment induced luteinization in which PRL-RL increased 4-fold (30.5 x 105 copies/µg RNA), whereas PRL-RS increased 10-fold (2.8 x 106 copies/µg RNA) above that in HEF granulosa cells. In CL on day 15 of pregnancy, PRL-RS mRNA had further increased, reaching 10.2 x 106 copies/µg, 3.7-fold lower than mRNA encoding PRL-RL, which changed little compared with HEF/hCG CL. Similar quantities of both isoforms were found on day 7 of gestation, but each had declined dramatically on day 22 (not shown).

Semiquantitative RT-PCR analyses comparing specific PRL-R amplification against that of the ribosomal proteins L-19 or S-16 (as internal controls for equal RNA loading and reaction efficiency) produced highly similar patterns of regulation to competitive PCR studies; however, higher reaction efficiency using PRL-RS primer resulted in overestimation of the relative PRL-RS/PRL-RL ratio. Thus, comparison between the two different assays by semiquantitative methods was impossible. Consistent with the fully quantitative results, however, we found the most dramatic increase in PRL-RL after E+FSH treatment of granulosa cells, which decreased time dependently with PRL treatment (Fig. 1CGo) but was increased only 2-fold further after hCG-induced luteinization. The transcript for PRL-RS, conversely, increased most dramatically after luteinization and further increased in early and midgestational CL, while both transcripts declined upon onset of luteolysis (day 22, Fig. 1CGo).

Collectively, these results demonstrate that the amount of PRL-R as well as the ratio of PRL-RL/PRL-RS is altered during ovarian cell differentiation. PRL-RL is most predominant in granulosa cells of preovulatory ovaries. In contrast, PRL-RS increases most during luteinization and attains levels quantitatively similar to that of PRL-RL in CL of pregnant rats. These changes are remarkably similar and ascribe additional isoform-specific information to the observed changes in 125I-PRL binding in HEF, HEF/hCG, and pregnant rat ovaries (36).

Stat-DNA Binding Activity during Folliculogenesis and Luteinization
Changes in the expression of PRL-R were then compared with the ability of PRL to induce specific DNA binding complexes. This was analyzed in whole cell extracts (WCE) of hormonally treated granulosa or luteal cells by electrophoretic mobility shift assays (EMSA) using the {alpha}2M PRL-RE as a probe (33, 34). Extracts of unstimulated H rat granulosa cells formed an intense band (Fig. 2AGo, lane 1), which was specifically competed by 10-fold excess unlabeled probe (Fig. 2BGo, lane 2). Treatment with E and FSH separately (not shown) or in sequential combination (Fig. 2AGo, lane 2) reduced the granulosa cell DNA binding activity without inducing the formation of any new complexes either before or after PRL stimulation (lanes 2–4). Extracts prepared from CL 1 day after induction of luteinization of HEF/hCG-treated rats also showed little specific DNA binding; however, 1 h after PRL treatment a protein/DNA complex was induced (Fig. 2AGo, lanes 5 and 6). This activity was absent 24 h after PRL treatment (lane 7). However, intense DNA binding activity was stimulated by acute injection of PRL to HEF/hCG-PRL (24 h) rats (Fig. 2AGo, lane 8). Supershift analysis demonstrated that the DNA binding complex in H granulosa cells contained exclusively Stat3, not Stat5a or Stat5b (Fig. 2BGo left panel), while the complex in luteal cells is composed predominantly of Stat5b and some Stat5a but not Stat3 (Fig. 2BGo, right panel). Both the Stat3 and Stat5 complexes were competed by incubation with 10-fold excess unlabeled probe DNA (Fig. 2BGo). Supershift analysis for Stat1 has repeatedly failed to detect any Stat1 binding activity in any ovarian extract (not shown; Refs. 33, 34).



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Figure 2. Stat Binding Activity in Granulosa Cells and CL: Shift from Stat3 to Stat5 with Luteinization

A, DNA binding activities in isolated immature/atretic (H) and preovulatory (HEF) granulosa cells or CL (HEF/hCG). WCE prepared from granulosa cells or luteal tissue of hypophysectomized (H) rats with or without E, FSH, and PRL stimulation were incubated with labeled {alpha}2M PRL-RE probe and analyzed by EMSA as described in Materials and Methods. Animals were treated with PRL either for 24 h by ip injection or with a 1-h pulse iv injection, or for 24 h followed by a 1-h pulse as indicated. B, Supershift analysis of protein complexes binding the {alpha}2M PRL-RE probe in extracts of H granulosa cells and HEF/hCG+PRL 24+1 h-treated CL. Antisera to Stat3, Stat5a, or Stat5b were incubated with protein extracts for 15 min before incubation with PRL-RE probe and EMSA as for panel A.

 
Extracts of CL isolated on days 4–9 of gestation contained no constitutive PRL-RE binding complexes. However, within 5 min of exogenous PRL administration on days 4, 7, and 9, an intense but transient DNA binding activity was induced (Fig. 3AGo). A more complete time course of PRL treatment on day 7 of gestation demonstrated maximal binding after 5 min with decreasing amounts at 15 and 30 min and none by 60 min. These rapid kinetics of Stat5 activation/deactivation differ from those observed in the H rat model (data herein and Ref. 33) where maximal activation of Stat5b occurred 1 h after PRL injection. On days 11–15 of gestation a constitutive protein/DNA complex was formed, and exogenous PRL administration had little or no influence on the magnitude or duration of activity (Fig. 3AGo). Supershift analysis of the induced complex present on day 7 and the constitutive complex from day 15 CL confirmed that the activity observed was Stat5b (Fig. 3BGo). Stat3-specific antibody did not supershift any of either complex.



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Figure 3. Stat5b Is Transiently Activated in Response to PRL in CL of Early Gestation but Constitutively Active in Midluteotropic Phase of Gestation

A, EMSA of protein complexes binding labeled PRL-RE probe in extracts of isolated CL from pregnant rats before or 5 min after iv injection of 10 µg PRL on days 4, 7, 9, 11, 12, and 15 of gestation. During early (day 7) and mid (day 15) luteotropic phases of pregnancy, more complete time courses of response to exogenous PRL were examined. B, Supershift analysis of PRL inducible (day 7, left panel) and constitutive (day 15, right panel) DNA binding complexes from early or midgestation, respectively. Note exposure of the right panel was 3 times longer than that in the left panel, enabling supershifted bands to be more easily visualized.

 
Thus, four distinct phases of Stat-DNA binding activity were identified during differentiation of ovarian cells. Constitutive Stat3 activity is present in immature/atretic (H) granulosa cells, while no Stat activity is detectable in preovulatory granulosa cells. In CL of nonpregnant (HEF/hCG) rats or rats at early stages of gestation (days 4–9), Stat5a/b are inducibly activated by PRL while after day 9 of gestation Stat5b retains a low constitutive activity.

Regulation of Cytokine Signaling Pathways: Phosphorylation and Expression of Stat3 and Stat5
To investigate whether Stat3 activation is unique to granulosa cells of H rats or whether activated Stat3 is prevented from binding DNA through additional changes in the hormonally differentiated granulosa cells, we analyzed the levels of phospho-Stat3 (P-Stat3) in extracts of granulosa cells at each stage of differentiation. As shown by Western blot analyses using a specific P-Stat3 antibody, levels of P-Stat3 were high in granulosa cells of H rats (Fig. 4AGo, lane 1), affirming the Stat3 DNA binding activity seen at this time. Despite the absence of DNA binding activity in HEF extracts (Fig. 2AGo), P-Stat3 remained present in E+FSH-treated granulosa cells and hCG stimulated luteal cells (Fig. 4AGo, lanes 2–6), albeit at 50–60% reduced levels. PRL treatment had no influence on the phosphorylation of Stat3 in granulosa cells at any stage of differentiation (Fig. 4AGo, lanes 3, 5, and 6). Protein extracts of CL isolated from pregnant rats on days 7 and 15 of gestation had lower amounts (10–20%) of P-Stat3, which appeared slightly induced 1 h after iv PRL administration (Fig. 4AGo, right panel). Total Stat3 content in these extracts remained consistent except for a 60% reduction in HEF granulosa cells (Fig. 4AGo, lower panel).



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Figure 4. Stat3 Phosphorylation and Stat Expression during Stages of Follicle Differentiation and Luteinization

A, Immunoblot analysis using antibodies specific for Stat3 phosphorylated on tyrosine-705 (upper panel); after stripping the same blot was reprobed for total Stat3 content of cells (lower panel). Protein extracts from isolated granulosa cells of immature/atretic (H) or preovulatory granulosa cells (HEF) or luteal cells (HEF/hCG) were isolated before or after PRL treatment for 1 h or 24+1 h as indicated or from CL of pregnant rats before or after 1 h PRL treatment on the indicated days of gestation. Different blots were probed using specific antibodies for Stat5a (B, upper panel), or Stat5b (B, lower panel); a slower migrating Stat5b immunoreactive band was evident 1 h after PRL treatment in CL extracts (arrow) corresponding to phosphorylated Stat5b. Rats were treated with PRL for 1 h, 24 h, or 24+1 h as indicated.

 
Western blot analyses using specific antibodies for the detection of Stat5a and Stat5b were also performed. Stat5a protein was present and showed little regulation throughout granulosa and luteal cell differentiation (Fig. 4BGo, upper panel). Stat5b protein was also present in granulosa cell extracts and increased approximately 2-fold in luteal cell extracts of HEF/hCG rats. After 1 h PRL treatment in extracts prepared from luteal cells of HEF/hCG rats, a slower migrating band was observed (Fig. 4BGo, lower panel, arrow), representing a phosphorylated form of Stat5b and supporting the observed Stat5b activation in these samples in EMSA experiments (Fig. 2Go). Concentrations of Stat5a/b in extracts of pregnant CL were comparable to those in CL of HEF/hCG-treated rats and remained constant throughout gestation (Fig. 4BGo, right panels). The phosphorylated form of Stat5b was evident in PRL-treated extracts of day 7 CL and was constitutively present on day 15.

Stat Signaling Regulators
Downstream of ligand-receptor interaction several mechanisms modulate cytokine signaling. One such regulator is PIAS-3. To determine whether this inhibitor might be involved in the regulation of ovarian Stat activity, specific primers for PIAS-3 were designed and used to amplify a specific band of the predicted size (308 bp). This band was cloned into the PCRII-topo vector and sequenced confirming the identity of the amplicon as PIAS-3. RT-PCR analyses showed that PIAS-3 mRNA expression was low in H rat granulosa cells but increased 4-fold in response to treatment with E+FSH (Fig. 5Go, lanes 1 and 2, respectively). In CL of HEF/hCG- treated rats, PIAS-3 expression declined slightly (lanes 5–8), was low in RNA from CL of pregnant rats on days 7, and approached nondetectable levels on days 15 and 22 of gestation. Thus, the highest levels of PIAS-3 mRNA were observed in the HEF granulosa cells that contained phosphorylated Stat3 (Fig. 4AGo) but no DNA binding activity (Fig. 2Go).



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Figure 5. PIAS-3 Expression Is Induced during Granulosa Cell Differentiation

Semiquantitative RT-PCR analysis using specific primer sets for PIAS-3 and L-19 as internal control. Total RNA was isolated from immature/atretic (H), or preovulatory granulosa cells (HEF) or luteal cells (HEF/hCG) before or after PRL treatment for 1 h, 24 h, or 24+1 h as indicated or from pregnant rat CL on the indicated days of gestation (right panels). Lower panels are mean ± SEM of phosphorimage analysis from three separate RT-PCR assays.

 
To investigate the importance of the SH-2 domain-containing phosphatases, SHP-1 and SHP-2, as potential regulators of Stat activation, specific RT-PCR assays for each of these phosphatases were designed and used to analyze their expression in granulosa and luteal cells. Single bands of predicted sizes were obtained in RT-PCR of ovarian cell RNA for SHP-1 (443 bp) and SHP-2 (223 bp). Products from all RT-PCRs showed the predicted restriction digest patterns, confirming their specific identities (data not shown). SHP-2 was highly expressed at relatively constant levels in granulosa and luteal cells of H, HEF-, and HEF/hCG-treated rats with or without PRL treatment (Fig. 6AGo, left panel). Levels of SHP-2 mRNA remained high through both luteotropic (days 7 and 15) and luteolytic (d22) phases of pregnancy (Fig. 6AGo, right panel). Western blot also showed SHP-2 present and not regulated throughout stages of follicle differentiation and luteal function (Fig. 6BGo).



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Figure 6. Expression of SHP-2 Is Not Regulated during Follicle Differentiation and Luteinization

A, Semiquantitative RT-PCR analysis using specific primer sets for SHP-2 and L-19 and RNA from granulosa cells or CL of hormone-treated H rats isolated either before or after PRL stimulation for 1 h, 24 h, or 24 h followed by 1 h pulse as indicated. CL of pregnant rats were isolated on day 7, 15, and 22 of gestation as indicated. Upper panel shows autoradiography from one representative experiment; lower panel shows combined mean ± SEM data from three repeated experiments quantitated by phosphorimage analysis. B, Immunoblot analysis of SHP-2 protein in extracts from isolated granulosa cells or CL of H rats that received the same treatments as in panel A, or pregnant rats on days 7, 15, or 22 of gestation before or after 1 h PRL treatment.

 
In contrast, expression of SHP-1 showed dramatic changes with hormonal treatments. Granulosa cells of H rats contained moderately detectable SHP-1 mRNA, which declined to nondetectable concentrations in HEF granulosa cells both with or without PRL treatment (Fig. 7AGo). Expression increased 4-fold after hCG treatment compared with HEF samples and increased 2-fold further 24 h after PRL treatment of HEF/hCG rats (Fig. 7AGo). SHP-1 was also expressed at elevated levels in CL of pregnant rats (Fig. 7AGo, right panel) where the ratio of SHP-1/L-19 was 2-fold higher than in HEF/hCG samples (Fig. 7AGo, right panel; note that the autoradiogram was exposed for less time than for HEF samples) and remained elevated through the luteolytic phase (Fig. 7Go). Western blot analysis of whole cell protein extracts from granulosa and luteal cells confirmed observations of mRNA regulation of SHP-1. Immunoreactive SHP-1 was low but clearly detectable in H extracts and declined to undetectable levels in preovulatory granulosa cells. After luteinization, SHP-1 protein concentration increased and remained high in luteal extracts of HEF/hCG-treated rats and throughout gestation (Fig. 7BGo).



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Figure 7. Expression of SHP-1 Is Hormonally Regulated during Follicle Differentiation and Luteinization

A, Semiquantitative RT-PCR analysis using specific primer sets for SHP-1 and L-19 internal control. Total RNA was from granulosa cells or CL of hormone- treated H rats isolated either before or after PRL stimulation for 1 h, 24 h, or 24 h followed by 1 h pulse as indicated. CL of pregnant rats were isolated on day 7, 15, and 22 of gestation as indicated. Upper panel shows autoradiography from one representative experiment; lower panel shows combined mean ± SEM data from three repeated experiments quantitated by phosphorimage analysis. B, Immunoblot analysis of SHP-1 protein in extracts from isolated granulosa cells or CL of H rats that recieved the same treatments as in panel A or pregnant rats on days 7, 15, or 22 of gestation before or after 1 h PRL treatment.

 
Cellular and Subcellular Localization of SHP-1
Immunohistochemical localization of SHP-1 in ovaries from H, HEF, and HEF/hCG-treated rats as well as pregnant rats confirmed the results from RT-PCR and Western analysis and further revealed dynamic changes in the cellular and subcellular localization of this protein in granulosa/luteal cells at different stages of differentiation. In H and HEF-treated ovaries, SHP-1 protein was localized in the cytoplasm of granulosa cells in small follicles that had the appearance of being atretic. Healthy follicles present in H and HEF ovaries had no detectable SHP-1 (Fig. 8Go, A, B, E, and F). SHP-1 was detected in nuclei of a subpopulation of cells throughout the ovarian stroma surrounding healthy follicles of H and HEF ovaries (Fig. 8FGo). The highest amount of immunoreactive SHP-1 was observed in CL of HEF/hCG-treated rats (data not shown) and pregnant rats in early (day 7) and middle (day 15) stages of gestation (Fig. 7Go, C, D, G, and H). In luteal cells, SHP-1 was selectively localized in nuclei irrespective of PRL treatment or levels of endogenous PRL/rPL. The distinct cellular and subcellular compartmentalization of SHP-1 in atretic granulosa cells compared with luteal tissue suggests differing functional roles possibly related to the specific activation of Stat3 vs. Stat5 and atresia vs. luteinization, respectively.



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Figure 8. Changes in Cellular and Subcellular Immunolocalization of SHP-1 Protein in Immature, Preovulatory, or Luteinized Ovaries

Ovaries from H rats (A and E), rats treated with E and FSH (B and F), and pregnant rats on day 7 (C and G) or 15 (D and H) of gestation were subjected to immunohistochemical detection of SHP-1 using nickel chloride diaminobenzidine color enhancement (black coloration) as described in Materials and Methods. Shown at 50x (upper panels) and 150x (lower panels) magnification. SHP-1 immunoreactivity was present in cytoplasm of granulosa cells in atretic follicles (arrowheads) but was not detected in healthy follicle granulosa cells (gc). In CL, strong staining was detected in nuclei of luteal cells on days 7 (C and G) and 15 (D and H) of gestation. Atretic follicles in pregnant rat sections also showed intense cytoplasmic staining (not shown).

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
In the ovary, follicles are predestined to undergo atresia and degenerate by apoptotic processes unless rescued and induced to grow, ovulate, and luteinize by hormonal stimuli. The transition from immature follicles to terminally differentiated CL involves specific reprogramming of cellular responsiveness to external signals (such as gonadotropins and cytokines), through changing interactions of positive and negative intracellular regulatory elements. Results of the studies described herein indicate that the actions of the gonadotropins and steroids are linked to specific changes in cytokine-signaling pathways as ovarian cells differentiate. We document that increases in PRL-R mRNAs, specifically a marked increase in PRL-RS, are associated with luteinization and related to enhanced activation of Stat5b during luteinization and in functional CL of pregnancy. Thus, PRL-RS may not exert a dominant negative regulatory mechanism in luteal cells. We also show for the first time that Stat3 is selectively activated in granulosa cells of small follicles by a factor other than PRL, and that the DNA binding capacity of Stat3 is reduced in growing follicles, possibly by the up-regulation of the specific inhibitory protein PIAS-3. Lastly, the expression as well as the cellular and subcellular localization of SHP-1 are selectively regulated. SHP-1 was elevated and cytoplasmic in granulosa cells of small atretic follicles, absent in healthy growing follicles, and then expressed at elevated levels in luteal cells where it is specifically nuclear.

The differentiation of granulosa cells by E+FSH to preovulatory stage is associated with several changes in the PRL signaling cascade, most notably the pronounced increase in the amount of PRL-RL (but not PRL-RS) mRNA. Despite an approximately 27-fold increase in preovulatory granulosa cells compared with small follicles of H rats, no Stat activation occurred after PRL administration. The increased PRL-RL expression in preovulatory (compared with immature) granulosa cells suggests a role for PRL in mature follicles, one of which is the time-dependent decrease in expression of its own receptor. Whether such a negative feedback mechanism serves a physiological function in preovulatory follicles is not yet clear, but similar observations have been noted in other tissues (37, 38, 39, 40). Since PRL does not activate Stat5b in granulosa cells, alternative signaling pathways appear to be involved at this time (41, 42, 43), while clearly factors other than PRL-RL are requisite for PRL-Stat5 signaling in ovarian cells.

PRL-RS prevalently increased (10-fold vs. 4-fold) after luteinization in HEF/hCG rats in which PRL-responsive Stat5 activation was observed. PRL-RS further increased in pregnant rat CL, attaining levels only 3.7-fold less than the long form on day 15, which contains rPL-mediated constitutively active Stat5. The up-regulation and relative abundance of PRL-RS in lactogen-responsive luteal cells suggests that it may not exert potent inhibitory effects on ovarian gene expression as demonstrated for milk protein gene activation in cultured cells of mammary epithelial origin cotransfected with PRL-RS and PRL-RL (23, 24). Rather, the appearance of high PRL-RS is associated with PRL-induced activation of Stat5b and induction of {alpha}2M gene expression as well as other genes in the CL, including LH receptor (43, 44), aromatase (30), and ER{alpha} (27). In support of our observations, identical patterns of PRL binding were reported in similarly treated granulosa and luteal cells (36). Highly quantitative RT-PCR analysis of whole ovary mRNA (3) indicated that PRL-RL transcripts are 1 order of magnitude more abundant than PRL-RS in proestrous ovaries, whereas PRL-RL and PRL-RS mRNAs were essentially equal in diestrus-I ovaries containing newly formed CL. Furthermore, in studies employing a different semiquantitative RT-PCR analysis, both isoforms showed equal expression in RT-PCR of RNA of rat CL day 15 of pregnancy (45), a time when PRL is critical to functional CL maintenance. Thus, the acquisition and maintenance of Stat5b activation and expression of PRL-responsive luteal genes such as {alpha}2M correlates with a high PRL-RS/PRL-RL ratio compared with that in Stat5 quiescent granulosa cells. That PRL-RS is directly involved with acquisition of Stat5 responsiveness cannot be concluded, but its relative abundance renders it unlikely that PRL-RS exerts dominant negative influence on PRL signaling in luteal cells.

One potentially important difference between ovarian and mammary cells is that the milk protein genes transcriptionally repressed by PRL-RS are selectively Stat5a-induced genes (46, 47), whereas we have found that ovarian {alpha}2M expression is Stat5b dependent (34, 35) and PRL activated predominantly Stat5b in luteinized cells despite the presence of high levels of Stat5a (present study). Additionally, PRL-RS may antagonize milk protein gene induction by mechanisms other than or in addition to changing Stat phosphorylation, such as by modulating nuclear translocation (19). Furthermore, in the ovary, PRL-RS has been shown to specifically interact with the CL-specific PRL-RS-associated phosphoprotein (48, 49), which has recently been identified as 17ß-hydroxysteroid dehydrogenase-7 (50). These associations link the PRL-signaling pathway not only to the regulation of estrogen receptor (27) but also to the endogenous production of estradiol within the CL from exogenous testosterone (51). Estradiol is a potent inducer of protein kinase C{delta} (52), which has recently been shown to enhance PRL- regulated expression of relaxin (53), a mediator of CL formation and function (54). Thus, luteal PRL-RS may serve significant luteotropic functions that are independent of PRL-RL and Stat5b.

The presence of distinct, active Stat/DNA complexes at defined stages of granulosa cell differentiation indicates that Stat3 and Stat5b control different cellular functions and are regulated by distinct pathways. That activated Stat3 was observed in the granulosa cells of H rats, while E or FSH treatment caused dramatic reductions in its DNA binding activity, suggests that E and FSH may regulate expression of the ligand or the receptor involved in activating Stat3. By analyzing the phosphorylation of Stat3 using a specific phospho-Stat3 antibody, we were able to demonstrate that although the amount of phospho-Stat3 in healthy growing follicles decreased, significant amounts of phosphorylated Stat3 remained present in differentiated granulosa cells. The absence of Stat3 binding to DNA in granulosa cells treated with E+FSH was temporally associated with a 4-fold increase in PIAS-3 expression, a negative regulator of Stat3 DNA binding activity. These results provide evidence that suppression of Stat3 function may be requisite for steroid- and gonadotropin-supported stages of growth and differentiation of follicles as opposed growth of primary follicles or apoptosis. Since many of the follicles in immature and H rats are undergoing atresia, it is possible that activated Stat3, as well as SHP-1, are associated with the apoptosis of granulosa cells. SHP-1 may mediate the atretogenic effect of angiotensin-II (55) through suppression of FSH-mediated growth signals involving the inhibition of extracellular regulated kinase activity (56). Thus, down-regulation of SHP-1 and induction of PIAS-3 may be important events in rescue of follicles from atresia. Although a selective role for Stat3 in small follicles is not yet known, activated Stat3 has been linked in other systems to both proapoptotic (57) and antiapoptotic cell survival pathways (58). When the ligand activating Stat3 becomes known and its targets identified, the role of Stat3 in early follicular cell function will be clearer.

In contrast to Stat3, an essential role for Stat5 in ovarian cell function is evidenced by the phenotypes of a line of Stat5b-specific null mutant mice that prematurely abort litters between days 8 and 17 of gestation unless exogenous progesterone is administered to substitute for CL function (59). Interestingly Stat5a/b-/- mice fail to produce CL at all (47), suggesting that Stat5a can act in a redundant or alternative but requisite fashion at least in early stages of luteal function in the absence of Stat5b, and that acquisition of PRL-inducible Stat5 activation and the resulting changes in gene expression outlined above are essential steps in the transition from granulosa to luteal cells. In this study we found that Stat5b is specifically induced in response to an exogenous pulse of PRL administered in a transient fashion in early gestation, while from day 11 constitutive Stat5b activation is unresponsive to the same exogenous PRL dose. These two patterns of activation probably result from the change in lactogen secretion in the latter stages of gestation when the placenta replaces the pituitary as the major source of lactogenic hormone secretion. Whether these different patterns of Stat5b activation mediate changes in luteal gene expression in early vs. late CL, as has been demonstrated for signaling through Stat5 by interleukin vs. erythropoietin (60), will be interesting to determine. It is known that {alpha}2M at this time undergoes a secondary phase increase in expression (31, 32, 33), while progesterone production and luteal cell hypertrophy also undergo incremental increases (2).

Activation of Stats is complex and involves many factors, and in some situations gene regulation may be dependent on signal duration (25, 60, 61, 62), indicating that the deactivation of signals is as important as the ligand- dependent activation in establishing and maintaining tissue responsiveness. In this regard, protein tyrosine phosphatases have been shown to play key roles in cytokine signaling and cell function. The tyrosine phosphatase SHP-2 plays an essential mediatory role in PRL-regulated milk protein gene induction (11) as well as in signal transduction by several growth factors and interferon-{alpha}/ß involving activation of Stats1 and 2 (63). Therefore, we initially hypothesized that the acquisition of Stat5 responsiveness and/or changes in rapid and transient vs. chronic Stat5b activation in luteal cells at specific times during pregnancy might be related to changes in the expression of SHP-2 mRNA and protein. However, SHP-2 was present with little change in abundance throughout granulosa cell differentiation, luteinization, and through to the onset of luteal regression. Thus, although this phosphatase may be a necessary mediator of PRL-regulated gene expression, its presence is not a limiting factor in luteal cells.

In contrast, we did observe a positive correlation between the presence of activated Stats and the expression of SHP-1, a tyrosine phosphatase thought to be required for recycling of activated Stat5b and maintaining responsiveness to pulsatile GH in hepatocytes (25, 61). In the ovary, SHP-1 expression was detected in H granulosa cells when Stat3-DNA binding is detected. As discussed above, it is suggested that SHP-1 may be involved with the process of follicular atresia. SHP-1 expression (mRNA and protein) was undetectable in preovulatory follicles but maximal in luteal cells, in a temporal pattern that mimics the acquisition of PRL-responsive activation of Stat5b. Immunoreactivity for SHP-1 was intensely localized in the nuclei of luteal cells. Transient (<1 h) Stat5b activation in PRL-treated CL during early gestation implicates the activity of a phosphatase such as SHP-1 in rapid dephosphorylation of Stat5b in CL. SHP-1 was expressed at its highest level and remained specifically nuclear in CL at day 15 of gestation, a time when Stat5b-DNA binding was constitutively maintained. We propose that nuclear SHP-1 may continuously dephosphorylate activated Stat5b, recycling it to the cytoplasm and enabling reactivation through continuous rPL/PRL-R stimulation in midpregnancy. Thus, nuclear SHP-1 may be requisite for mediating the rapid turnover of activated Stat5b, resulting in a transient activation profile during pulsatile PRL exposure and maintaining a pool of inactive Stat5b able to respond to constant circulating lactogens in midterm pregnant rats. While this manuscript was under review, it was reported that a vanadate-sensitive factor, probably SHP-1, is involved in the constant turnover of activated Stat5b in GH-treated liver cells. Moreover, enhanced dephosphorylation of Stat5b (as well as Jak2) in constant GH-exposed cells results in a lower continuous activation similar to that seen in midpregnant luteal cells (64). In contrast to GH signaling in female liver, constant rPL exposure in luteal cells and the resulting low constitutive Stat5b activation have enhanced tropic effects on midgestational CL function.

Attractive as this hypothesis for the role of SHP-1 might be, a functional role in the corpus luteum remains to be verified. A naturally occurring mutation in SHP-1 gene has been described in the motheaten strain of mice, which are infertile and fail to form CL (65). However, when ovaries of the mutant mice were transplanted to wild-type recipients, fertility (normal ovarian function?) was rescued. These results indicate that the defect may not be intrinsic to the ovary, that factors present in the recipient regulate other SHP-1-like activities in the mutant ovaries, or that SHP-1 is present and plays a role in extraovarian (immune?) cells to regulate fertility and CL formation. The presence of SHP-1 in the nuclei of luteal cells suggests that there is an intrinsic role for this phosphatase in ovarian cells; however, it should be noted that SHP-1 expression was present in additional cells surrounding follicles (Fig. 6Go, B and F). This expression pattern mimicked that seen with macrophage-specific antibodies (Ref. 66 and R. Robker and J. S. Richards, unpublished). Since SHP-1 is a known modulator of immune cell function and leukocytes have numerous important actions in the ovary (66), immune defects, the predominant phenotype of motheaten mice, may explain their ovarian failure.

In conclusion, we have demonstrated a dynamic transition in Stat activation during the maturation of ovarian follicle cells. In undifferentiated/atretic follicles deprived of gonadotropin, Stat3-DNA binding is uniquely active possibly because a permissive milieu is maintained through low PIAS-3 expression. Increased PIAS-3 mRNA in granulosa cells of growing follicles suggests that suppression of Stat3-DNA binding may be an important requirement for progression of follicle growth and differentiation. Specific expression of SHP-1 and localization to the cytoplasm in atretic granulosa cells may also regulate Stat signaling and mediate apoptotic events, directing cells toward atresia rather than growth and differentiation. Acquisition of Stat5b responsiveness to PRL is initiated after luteinization and associated with increased PRL-R, most strikingly increased PRL-RS, providing evidence that the transition from granulosa cells to luteal cells is not related to a dominant negative effect of PRL-RS on Stat5b activation. Additionally, high PRL-RL expression alone in granulosa cells appears insufficient to activate Stat5 signaling. Lastly, our results indicate that SHP-1 is temporally expressed in a pattern that mimics the cellular activation of Stats. Atresia in granulosa cells involves moderate expression of cytoplasmic SHP-1. Luteal cells express higher levels of SHP-1 where it is localized to nuclei and may be an important requirement for maintaining PRL responsiveness through Stat5 deactivation and recycling.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Materials
17ß-Estradiol was purchased from Sigma (St Louis MO), ovine FSH and PRL from National Hormone and Pituitary Program (Baltimore MD), hCG from Organon special chemicals (West Orange NJ) and [{alpha}-32P] from ICN Biochemicals, Inc. (Cleveland OH). BenchMark molecular size markers were from Life Technologies, Inc.(Gaithersburg, MD), and the enhanced chemiluminescence (ECL) detection system was from Amersham Pharmacia Biotech (Arlington Heights IL). Kodak X-Omat AR film was from Eastman Kodak Co. (Rochester, NY). AMV-Reverse Transcriptase and Taq Polymerase were from Promega Corp. (Madison WI). Anti-Stat5a and Stat5b-specific antibodies, catalog nos. sc-1081 and sc-835, respectively, were from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA ). Anti-Stat3 antiserum used for supershift analysis was generously provided by Dr. David Levy (New York School of Medicine, New York, NY). Stat3 Western blots employed an antibody purchased from Santa Cruz Biotechnology, Inc. (catalog no. 7179), P-Stat3 antiserum was from New England Biolabs, Inc. (Beverly, MA). Antibodies to SHP-1 and SHP-2 were from Transduction Laboratories, Inc. (Lexington KY).

Animals
Pregnant, hypophysectomized, and immature (day 26) rats were purchased from Harlan Bioproducts for Science, Inc. (Indianapolis, IN), provided food and water ad libitum, and housed under a 16-h light, 8-h dark schedule. Animals were treated in accordance with the NIH Guide for Care and Use of Laboratory Animals. Protocols were approved by the Institutional Animal Care and Use Committee, Baylor College of Medicine (Houston TX).

H Rats
Proliferation and differentiation of ovarian follicles were stimulated in H rats by hormonal treatment as described previously (33). Follicles of H rats inevitably become atretic and undergo apoptotic degeneration unless rescued by E and FSH administration (67). Commencing 3–4 days after hypophysectomy, H rats received sc injections of 17ß-estradiol (1.5 mg) for 3 consecutive days, followed on the next 2 days with two injections of FSH (1 µg) each day (HEF rats). Luteinization was induced in HEF rats by ip injection of 10 IU hCG (HEF/hCG). At each stage of follicular development, animals were killed either before or 1 h after tail vein injection of 10 µg PRL. Additional groups of HEF- and HEF/hCG-treated rats were treated ip with PRL 24 h before receiving a second iv PRL injection 1 h before ovaries were collected. Each treatment group included 6 rats except for the H group for which 16 animals were used. After removal, ovaries were divided at random into two groups for RNA or WCE preparation.

Pregnant Rats
Day 1 of pregnancy was assigned as the day a sperm-positive vaginal swab was observed. On selected days of pregnancy, ovaries were collected from two rats either before or 5 min after iv PRL administration. On days 7 and 15, representing the early (pulsatile endogenous PRL) and mid (continuous rPL) luteotropic stages of gestation, a time course of response to exogenous PRL was examined by collecting ovaries 5, 15, 30, and 60 min after PRL injection. All PRL treatments (10 µg) were given at approximately 1100 h.

WCE and RNA Isolation
Ovaries were extirpated and granulosa cells were isolated from preovulatory ovaries by puncture with a 26-gauge needle, or CL were dissected from luteinized ovaries. Granulosa cells isolated from preovulatory ovaries were resuspended in 150–200 µl of 10 mM Tris buffer containing 1 mM EDTA, 1 mM dithiothreitol, 10% glycerol, 400 mM potassium chloride, 1 mM vanadate, and protease inhibitors (WCE buffer; Ref. 68). Cells and nuclei were lysed by three rapid freeze-thaw cycles and centrifuged at 12,000 x g, and protein concentrations of soluble extracts were measured (Bradford method, Bio-Rad Laboratories, Inc., Richmond CA). Isolated CL were homogenized at 4 C in WCE buffer and then treated as for granulosa cell extracts.

For RNA extraction, granulosa cells or CL were homogenized in 25 mM Tris buffer containing 1% Nonidet P-40. RNA was extracted in phenol-chloroform, ethanol precipitated, and resuspended in RNase free water.

EMSA
EMSA were performed as described previously (33). Briefly, protein extracts (15 µg per lane) were incubated for 30 min at room temperature with 50,000 cpm of end-labeled double-stranded oligonucleotide probe and poly(deoxyinosinic-deoxycytidylic)acid in a final buffer volume of 20 µl containing 15 mM Tris-HCl (pH 7.5) 100 mM KCl, 5 mM dithiothreitol, 1 mM EDTA, 5 mM MgCl2, and 12% glycerol. For supershift and competition experiments, antibodies or 10-fold excess of unlabeled competitor DNA were incubated with extracts for 30 min on ice before labeled probe DNA was added. Bound probe-DNA complexes were resolved on 5% acrylamide gel electrophoresis before autoradiography. The sequences of oligonucleotides used for {alpha}2M PRL-RE probe were as follows:: 5'-TGGATCATCCTTCTGGGAATTCTGATATCCTTC-TGGGAATTCTG-3' annealed to the reverse complimentary strand.

PRL-R Competitive PCR
Quantitative PCR assays for PRL-RL and PRL-RS were developed using modifications of the method of Nagano et al. (3). A sense PRL-R oligonucleotide primer targeted to the conserved extracellular domain present in all rat PRL-R isoforms used for amplification of both PRL-R isoforms was 5'-ATACTGGAGTAGATGGAGCCAGGAGAGTTC-3'. Specific antisense primer to the divergent cytoplasmic domains of the long isoform sequence was 5'-CTTCCGTGACCAGAGTCACTGTCGGGATCT-3', and the short isoform was 5'-TCCTATTTGAGTCTGCAGCTTCAGTAGTCA-3'. These primer pairs gave predicted amplification products of 422 bp and 332 bp from the long and short PRL-R, respectively. Amplicons from long and short PRL-R PCR with 100-bp deletions were cloned into the pCRII-topo vector (Invitrogen, Carlsbad CA). Total RNA (350 ng) from each sample was reverse transcribed in a 25 µl total volume containing 500 ng oligo-dT12–18 and 1 IU AMV-RT at 42 C for 90 min; 4 µl of RT reaction mix was aliquoted into each of five tubes containing titrated concentrations of competitor cDNA from 0.01–10 x 106 cDNA copies and 15 µl PCR reaction cocktail [20 mM Tris-HCl (pH 8.0), 2.5 mM MgCl2 and 100 mM NaCl, 2.5 IU Taq DNA polymerase, 1 µCi 32P-dCTP, and 40 pmol forward and isoform-specific reverse primer]. PCR reactions were performed for 30 cycles at 95 C (2 min), 65 C (2 min), and 72 C (3 min). To confirm equivalent amounts of each competitor construct and their efficiency in PCR reactions, PCR was also performed for each competitor dilution using T7 and SP6 oligonucleotide primers that flank the cloned inserts. Reaction products were separated on 5% polyacrylamide gels, and intensity of the competitor and target bands was analyzed as for semiquantitative PCR and expressed as the log10 of the target to competitor ratio. When plotted on a log scale against input competitor concentration, parallel straight lines were obtained for the titration of competitor against each RNA sample. The Y axis origin represents the concentration at which competitor and target amplification are equivalent, and thus the number of cDNA copies present in the original RT-cDNA mix. This procedure was repeated a minimum of three times for each RNA sample with highly reproducible results, and PRL-RL and PRL-RS were analyzed in parallel in each assay.

Semiquantitative RT-PCR Analysis
Primer pairs based on rat SHP-1 sequence were 5'-AGCCGTGTCATCGTCATGACCACCCGAGAG-3' and 5'-CATC-TGGATGGTCTTCTGGATGTCAATGTC-3'; for rat SHP-2 primer pairs were 5'-AGCCAGAGCCACCCTGGGGACTTCGTCCTC-3' and 5'-AATACGAGTTGTGTTGAGGGGCTGTTTGAG-3'. Predicted products from SHP-1 and SHP-2 amplification were 443 bp and 248 bp, respectively. Primers for PIAS-3 were 5'-CAGATGAATGAGAAGAAGCCGACATGG-3' and 5'-TCTGATGAGCTTTCGATGGTCAAG-3' and generated a product of predicted size 308 bp. Primer pairs for the internal control ribosomal protein L-19 were as described previously (69); predicted product size for L-19 PCR amplification was 194 bp. Total RNA (350 ng) was reverse transcribed using 500 ng oligo-dT12–18 primer (Pharmacia Biotech, Piscataway NJ) at 42 C for 90 min in a 20 µl reaction volume. To the RT reactions were added 80 µl of buffer containing 20 mM Tris-HCl (pH 8.0), 2.5 mM MgCl2, and 100 mM NaCl, 2.5 IU Taq DNA polymerase, [32P]dCTP (5 µCi of 3000 Ci/mmol), and specific oligonucleotide primer pairs (50–80 pmol) for each individual gene along with L-19 internal control. Twenty-cycle PCR reactions (within the linear amplification range for input RNA) were performed in a DNA Engine thermocycler (MJ Research, Inc., Watertown, MA) using conditions of 95 C (2 min) for denaturing, 65 C (2 min) annealing, and 72 C (3 min) extension. Reaction products were separated on 5% polyacrylamide gels and exposed to Kodak X-Omat AR x-ray film followed by quantitation of products using a Storm860 PhosphorImager and ImageQuant version 2.1 software (Molecular Dynamics, Inc., Sunnyvale, CA). Intensity of signal for each sample was normalized to the L-19 internal control. All PCR assays were performed three separate times from the same RNA samples, and the mean ± SE for normalized results was calculated. All genes were also analyzed in comparison to ribosomal protein S-16 internal controls in separate PCR analyses with identical results to L-19 obtained.

Immunoblot Analyses
Whole-cell protein extracts (50 µg) were resolved on 10% acrylamide gels by reducing SDS-PAGE, followed by electrophoretic transfer to polyvinylidine fluoride membrane (Immobilon-P, Millipore Corp., Bedford, MA). Membranes were blocked by incubation for 1 h at room temperature with 3% nonfat milk, followed by 1 h incubation with specific primary antibodies in 3% milk and washing in TBST [10 mM Tris (pH 7.5), 150 mM NaCl, and 0.05% Tween-20]. Blots were then incubated with 1:10,000 of horseradish peroxidase-linked antirabbit or antimouse IgG (Amersham Pharmacia Biotech) followed by six 5-min washes with TBST. ECL detection was performed according to the manufacturer’s specifications. Blots were stripped for subsequent reanalysis by washing at 50 C for 30 min in 62.5 mM Tris-HCl, pH 6.7, 100 mM 2-mercaptoethanol, and 2% SDS. Quantitation from Western blots was performed using a Molecular Dynamics, Inc. densitometer and ImageQuant software.

Immunohistochemistry
SHP-1 tissue and cellular localization was analyzed by immunostaining of 4% paraformaldehyde-fixed paraffin sections of ovaries from each of the indicated treatment groups. Rehydrated sections were boiled in 20 mM sodium citrate for 10 min, and endogenous peroxidase activity was quenched by 10 min treatment with 0.1% H2O2 followed by PBS wash. Nonspecific antibody binding was blocked by 30 min incubation with 10% nonimmune goat serum after which anti-SHP-1 IgG (Santa Cruz Biotechnology, Inc.), 0.5 µg/ml in 10% goat serum, was incubated with sections overnight at room temperature. After washing with PBS, biotinylated antirabbit antiserum (Vector Laboratories, Inc., Burlingame, CA) was added for 30 min, slides were washed, and streptavidin-conjugated horseradish peroxidase was applied for 30 min. After washing, sections were incubated with diaminobenzidine substrate containing nickel chloride color enhancement (Vector Laboratories, Inc.) for 2 min and then dehydrated and mounted without counterstaining.


    FOOTNOTES
 
Address requests for reprints to: Dr. Darryl L. Russell, Department of Cell Biology, Baylor College of Medicine, One Baylor Plaza, Houston, Texas 77030.

Received for publication October 27, 1998. Revision received July 9, 1999. Accepted for publication August 24, 1999.


    REFERENCES
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 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

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