Growth Hormone- and Prolactin-Induced Proliferation of Insulinoma Cells, INS-1, Depends on Activation of STAT5 (Signal Transducer and Activator of Transcription 5)

Birgitte Nissen Friedrichsen, Elisabeth Douglas Galsgaard, Jens Høiriis Nielsen and Annette Møldrup

The Hagedorn Research Institute Department of Cell Biology 2820 Gentofte, Denmark


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
GH and PRL stimulate proliferation and insulin production of pancreatic ß-cells. Whereas GH- and PRL-regulated transcription of the insulin gene in insulinoma cells has been shown to depend on STAT5 (signal transducer and activator of transcription 5), the signaling pathways involved in GH/PRL-induced ß-cell replication are unknown. The roles of various signaling pathways in human GH (hGH)-induced DNA synthesis were studied by analysis of the effect of specific inhibitors in both the insulin-producing cell line, INS-1, and in primary ß-cells. The mitogen-activated protein kinase kinase (MEK)-inhibitor, PD98059, as well as the mitogen-activated protein kinase p38 (MAPKp38) inhibitor, SB203580, partially inhibited hGH- induced proliferation in INS-1 cells but had no significant effect in primary ß-cells. Staurosporine, a protein kinase C (PKC) and protein kinase A (PKA) inhibitor, blocked both basal and hGH-induced proliferation in INS-1 cells, but had no inhibitory effect in primary ß-cells. Wortmannin, a phosphatidylinositol 3-kinase (PI3K) inhibitor, inhibited hGH-induced proliferation neither in INS-1 cells nor in primary ß-cells, whereas the tyrosine kinase inhibitor, genistein, completely inhibited hGH- induced proliferation in both primary ß-cells and INS-1 cells. To analyze the possible role of STAT5 in hGH-induced proliferation, a dominant negative STAT5 mutant, STAT5{Delta}749, was expressed in INS-1 cells under the control of a doxycycline- inducible promoter by stable transfection. Two clones were found to exhibit dose-dependent, doxycycline-inducible expression of STAT5{Delta}749 and suppression of hGH-stimulated transcriptional activation of a STAT5-regulated PRL receptor (PRLR) promoter-reporter construct. Furthermore, induction of STAT5{Delta}749 expression completely inhibited hGH-induced DNA synthesis. Analysis of endogenous gene expression revealed a doxycycline-dependent inhibition of hGH-stimulated PRLR and cyclin D2 mRNA levels. Our results suggest that GH/PRL-induced ß-cell proliferation is dependent on the Janus Kinase2 (JAK2)/STAT5 signaling pathway but not the MAPK, PI3K, and PKC signaling pathways. Furthermore, the cell cycle regulator cyclin D2 may be a crucial target gene for STAT5 in this process.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The hormones of the GH family, GH , PRL, and placental lactogen (PL), are the most potent growth factors that have been identified for pancreatic ß-cells to date (for review see Refs. 1, 2). In addition, these hormones stimulate insulin production, partially via direct transcriptional activation of the insulin promoter. These effects are mediated through the GH-receptor (GHR) and the PRL-receptor (PRLR) (3), which are both expressed in rat islets as well as in various insulinoma cell lines (4, 5, 6). The GHR and PRLR are transmembrane proteins belonging to the cytokine receptor superfamily (7). These receptors are characterized by the ability to activate STAT (signal transducers and activators of transcription) proteins, which are latent transcription factors that become activated by phosphorylation of a single tyrosine residue by receptor-associated Janus kinases (JAKs). The phosphorylated STAT proteins dimerize and translocate to the nucleus where they bind to specific DNA elements and activate transcription (8). Recently, we identified STAT5 binding elements in the promoters of the rat insulin 1 (9) and PRLR genes (10) that are required for transcriptional activation by GH and PRL. Thus, GH and PRL were found to stimulate DNA binding of STAT5a and STAT5b and, to a lesser extent, STAT1 and STAT3 in the insulinoma cell lines, RIN-5AH (9) and INS-1 (10). Apart from the JAK/STAT pathway, several other signaling pathways are activated in various cell types in response to GH and PRL stimulation (for review see Refs. 8, 11, 12). Thus, the ras/raf/MAPK (mitogen-activated protein kinase) cascade is activated in some cell types, presumably involving the adaptor protein complex, SHC/Grb2/SOS (SH2-containing protein/growth factor receptor-bound protein/Son-of-Sevenless-1), which may bind to either tyrosine-phosphorylated JAK2 or insulin receptor substrates (IRS) 1 and/or 2. Phosphatidylinositol-3'- kinase (PI3K) may also be activated via recruitment by tyrosine-phosphorylated IRS, leading to downstream activation of the 70-kDa-S6-kinase (p70S6K). Furthermore, activation of protein kinase C (PKC), which may involve phospholipase C and increased diacylglycerol (DAG) levels, or PI3K, has been reported for both GH and PRL. The signaling pathways involved in GH/PRL-induced ß-cell mitogenesis have not been identified. Activation of MAPK and p70S6K are known to be required for replication of many mammalian cell types. However, in INS-1 cells, MAPK activity was not induced by GH/PRL, but potently by nerve growth factor (NGF) and glucose (13). Furthermore, whereas insulin-like growth factor-I (IGF-I) in a recent study was found to activate both MAPK and PI3K activities in INS-1 cells, GH had no detectable effect on these enzymes (14). Tyrosine residues in the intracellular domain of the GHR and PRLR have been found to be required for STAT5 activation and transcriptional signaling (15, 16). However, studies in the mouse promyeloid interleukin-3-dependent cell line, FDC-P1, showed that the intracellular tyrosine residues of the GHR were not essential for mitogenic signaling in these cells (17, 18). Similar results have been reported for the PRLR (19), and it has been hypothesized that STAT5 activation is mainly involved in the regulation of differentiated gene expression whereas mitogenic signaling may primarily depend on activation of STAT1 and 3, which are recruited to and activated by JAK2 and/or JAK2-mediated MAPK activation (20). Recently however, STAT5 has been implicated in regulation of cell cycle progression of peripheral lymphocytes (21, 22, 23) and lymphoma cell lines (24, 25, 26, 27). In the present study we have examined the role of the MAPK, PI3K, PKC, and JAK2/STAT5 signaling pathways in GH/PRL-induced cell proliferation using metabolic inhibitors of these enzymes in INS-1 cells and in primary ß-cells from neonatal rat islets. The role of STAT5 was evaluated by inducible overexpression of a dominant negative mutant of STAT5 in INS-1 cells.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Analysis of Signaling Pathways That May Lead to Human GH (hGH)-Induced Proliferation in INS-1 Cells and Primary ß-Cells
To determine whether the MAPK, the PI3K, or PKC signaling pathways are involved in hGH-induced ß-cell proliferation, the effect of specific protein kinase inhibitors on hGH-stimulated DNA synthesis was examined in INS-1 cells and in primary ß-cells in monolayer culture by 3H-thymidine and bromodeoxy uridine (BrdU) incorporation assays, respectively, as previously described (1, 28). After a 24-h stimulation period with 0.5 µg/ml of hGH, 3H-thymidine incorporation into INS-1 cells was increased by 2.0 ± 0.1-fold over the basal level (Fig. 1AGo), and BrdU incorporation into primary ß-cells was increased by 2.7 ± 0.4-fold over the basal level (Fig. 1BGo). Human GH-induced DNA synthesis was completely inhibited in both INS-1 and primary ß-cells (1.1 ± 0.1-fold and 0.9 ± 0.1-fold, respectively, over the basal levels) in the presence of 25 µM genistein, an inhibitor of protein tyrosine kinase activity, as would be expected from inhibition of JAK2. In the presence of the MEK inhibitor, PD98059 (20 µM), and the p38 MAPK inhibitor, SB203580 (10 µM), hGH-induced DNA synthesis was partially reduced in INS-1 cells (1.4 ± 0.1-fold and 1.4 ± 0.1-fold, respectively, over basal levels), whereas these two inhibitors had no significant effect in primary ß-cells. Wortmannin (10 nM), a specific PI3K inhibitor, had no effect on hGH-induced DNA synthesis in either INS-1 or in primary ß-cells, whereas staurosporine (20 nM), an inhibitor of PKC, protein kinase A (PKA), and protein kinase G (PKG), blocked both basal (not shown) and hGH- induced proliferation of INS-1 cells (0.2 ± 0.1-fold of basal) but did not inhibit proliferation of primary ß-cells. On the contrary, staurosporine was found to potentiate DNA synthesis in primary cells. Addition of the various inhibitors in the absence of hGH (data not shown) had no effect on the basal proliferation level in any of the cell types with the exception of staurosporine, as described above.



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Figure 1. The Effect of Protein Kinase Inhibitors on hGH-Stimulated DNA Synthesis

A, INS-1 cells were cultured for 24 h in the absence or presence of 0.5 µg/ml hGH. Before addition of hGH, respective cultures were supplemented with 25 µM genistein, 20 µM PD98059, 10 µM SB203580, 10 nM wortmannin, or 20 nM staurosporine. The cells were labeled 4 h with 3H-thymidine before harvesting and counting. The results are expressed as fold induction compared with control levels (mean ± SEM, n = 4). B, Precultured monolayers of primary ß-cells were incubated for 24 h with 10 µM BrdU and also in the absence or presence of hGH and inhibitors as described above. The cells were fixed and stained for BrdU and insulin. The results are expressed as the ratio between BrdU-labeled ß-cells and the total amount of ß-cells (mean ± SEM, n = 5). A total of 1,000 cells were counted for each preparation. *, P <= 0.05; **, P <= 0.001; ***, P <= 0.0001, compared with the group treated with hGH alone.

 
hGH Induces Prolonged Activation of STAT5 in INS-1 Cells and in Cultured Newborn Rat Islets
The kinetics of hGH-induced STAT5 activation in INS-1 cells and primary newborn rat islets in suspension culture were determined by electrophoretic mobility shift analysis (EMSA) (Fig. 2Go). INS-1 cells and islets cultured in 0.5% serum were either left untreated or stimulated with 0.5 µg/ml hGH for 15 min to 24 h. Nuclear extracts were prepared and incubated with radiolabeled, double-stranded oligonucleotide (1A-GLE) representing the previously identified STAT5-binding element of the PRLR 1A promoter (10). Free and bound probe was separated by nondenaturing PAGE and visualized by autoradiography. Nuclear extracts from INS-1 cells that had been stimulated with hGH for 15 min (upper panel, lane 2) produced a strong shifted band, which was also present in incubates with nuclear extracts from INS-1 cells treated for 1, 4, and 24 h with hGH (upper panel, lanes 3, 4, and 5, respectively). In contrast, this complex was absent in incubates with nuclear extracts from untreated INS-1 cells (upper panel, lane 1). A similar pattern was observed with nuclear extracts from newborn rat islets (lower panel, lanes 1–5), although the intensity of the complex was somewhat reduced after prolonged hGH treatment (~30–50% after 4 and 24 h).



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Figure 2. hGH-Induced STAT5 Activation in INS-1 Cells and Newborn Rat Islets

Analysis of hGH-induced activation of STAT5 was performed by EMSA. The radiolabeled probe, 1A-GLE, was incubated with 10 µg nuclear extracts prepared from either INS-1 cells (upper panel) or newborn rat islets (lower panel) that were either unstimulated (lane 1) or stimulated with 0.5 µg/ml hGH for 15 min, 1 h, 4 h, and 24 h (lanes 2–5, respectively). Free and bound probe were separated by nondenaturing gel electrophoresis and visualized by autoradiography. The autoradiograph shown is representative of three independent experiments. The arrows indicate the migration of the 1A-GLE*STAT5 complex.

 
Inducible Overexpression of a Truncated STAT5a Mutant (STAT5a{Delta}749) in INS-1 Cells with Dominant Negative Activity
To address whether GH/PRL-stimulated activation of STAT5 is involved in the mitogenic response of ß-cells to these hormones, the Tet-On gene expression system was employed to generate stably transfected INS-1 cells expressing, in an inducible manner, a STAT5 mutant, STAT5a{Delta}749, which has been reported to exert dominant negative activity (29). An INS-1 clone, INS-r3, stably transfected with an expression plasmid encoding the reverse tetracycline/doxycycline-dependent transactivator (30), was transfected with either the expression vector, pTRE, without insert or the pTRE-vector containing a cDNA encoding STAT5a{Delta}749 under the control of the tetracycline operator. Cotransfection of an expression vector containing a hygromycin resistance gene allowed for selection of stably transfected clones, which were tested for DNA integration of the pTRE plasmids by PCR analysis. Twenty-five of 38 clones examined were found to contain integrated pTRE vector without insert, and 2 of 69 clones examined contained a full-length insert of STAT5a{Delta}749.

Expression of STAT5a{Delta}749 in the two transfected clones (termed BB32 and EB03) was determined by Western blot analysis (Fig. 3AGo). The cells were cultured for 20 h in the absence or presence of 0.05–1.0 µg/ml doxycycline, and total cell lysates were examined for STAT5 expression using an antibody raised against the amino acids (aa) 451–649 of STAT5. In the absence of doxycycline (lanes 1 and 6) only wild-type STAT5 protein was detectable, which in this experiment probably represents STAT5b, since Western blot analysis, using a more potent antibody recognizing the C terminus part of STAT5, indicates that STAT5b is approximately 4–5 times more abundant than STAT5a in these cells (data not shown). In both cell lines, addition of 0.05 µg/ml doxycycline induced the expression of a protein of faster mobility than wild-type STAT5, likely to represent STAT5a{Delta}749 (lanes 2 and 7). The effect of doxycycline was found to be dose dependent with maximal expression observed at a concentration of 0.1 µg/ml doxycycline for both BB32 and EB03 (lanes 2–5 and 7–10, respectively). Doxycycline had no effect on STAT5 protein expression in a control clone (termed BA15) stably transfected with pTRE without insert (data not shown).



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Figure 3. Inducible Expression and Dominant Negative Activity of STAT5a{Delta}749 in INS-1 Cells

A, Western blot analysis was performed as described in Materials and Methods on total protein extracts prepared from stably STAT5a{Delta}749-transfected clones, BB32 and EB03 (STAT5a{Delta}749 transfected) that had been cultured for 24 h in the absence (lanes 1 and 6) or presence of the indicated amounts of doxycycline (lanes 2–5 and 7–10). Forty microliters of each extract were separated on a 7.5% SDS polyacrylamide gel, and proteins were transferred to nitrocellulose. STAT5 protein was visualized using STAT5 antibody raised against the aa 451–649 of STAT5 in a 1:1000 dilution. B, The radiolabeled oligonucleotide probe, 1A-GLE, was incubated with nuclear extracts from either the BA15 (lanes 1–4), the BB32 (lanes 5–8), or the EB03 cells (lanes 9–16) that had been cultured for 24 h in the absence or presence of the indicated amounts of doxycycline and in the absence or presence of 0.5 µg/ml hGH 15 min before harvesting. Free and bound probe were separated by nondenaturing gel electrophoresis and visualized by autoradiopgraphy. The autoradiograph shown is representative of two independent experiments. The arrow indicates the migration of the 1A-GLE*STAT5/STAT5a{Delta}749 complex. C, INS-1 cells, BA15, BB32, and EB03 were transiently transfected with the firefly luciferase reporter gene construct containing the PRLR 1A promoter and the internal control construct, pRL-SV40, which contain the Renilla luciferase gene. The cells were cultured for 24 h in the absence or presence of the indicated amounts of doxycycline. Seven hours before harvest the cells were cultured in the absence or presence of 0.5 µg/ml hGH. The results are expressed as ratios between the firefly and the Renilla luciferase activities (mean ± SEM, n = 3). Significant differences between the groups treated with hGH+Dox and the group treated with hGH alone are indicated: *, P <= 0.05; **, P <= 0.001; ***, P <= 0.0001. A significant difference (P <= 0.0001–0.05) between the hGH-treated groups and the control groups ± Dox was found in all four cell lines.

 
To determine the hGH-induced DNA-binding activity of STAT5a{Delta}749, EMSA was performed as described above. The stably transfected clones, BA15, BB32, and EB03, were cultured for 24 h in the absence or presence of doxycycline and stimulated the last 15 min of the culture period with 0.5 µg/ml hGH. Nuclear extracts of the cells were prepared and incubated with radiolabeled 1A-GLE oligonucleotide probe, the mobility of which was examined by gel electrophoresis. hGH treatment alone was found to induce 1A GLE binding of nuclear protein in all the cell lines (Fig. 3BGo, lanes 2, 6, and 10), and this binding was unaffected by the addition of doxycycline in BA15 cells (Fig. 3BGo, lane 4). In BB32 and EB03 cells, a doxycycline-dependent increase of DNA complex formation was observed (Fig. 3BGo, lanes 8 and 12–16) likely to represent binding of STAT5a{Delta}749. A promoter-reporter gene assay (10) was used to address whether overexpression of STAT5a{Delta}749 would suppress signaling via endogenously expressed STAT5 (Fig. 3CGo). The parental and the three transfected lines were transiently transfected with the firefly luciferase reporter gene under the transcriptional control of the 5'-flanking region of the PRLR gene containing a STAT5-responsive element. hGH stimulation induced transcriptional activity of the PRLR promoter in INS-1 cells by 2.9 ± 0.1-fold and in BA15 cells by 5.5 ± 0.8-fold (upper panel), in BB32 cells by 4.6 ± 0.4-fold (middle panel) and in EB03 cells by 17.7 ± 0.3-fold (lower panel) over the basal levels. The addition of 0.5 µg/ml doxycycline affected neither the basal nor the hGH-induced transcriptional activity in INS-1 and BA15 cells. However, addition of doxycycline to BB32 and EB03 cells increased the basal transcriptional level by 1.8 ± 0.3-fold and a 4.3 ± 1.3-fold, respectively (middle and lower panel). The reason for this stimulatory activity is not known. However, hGH-induced transcriptional activity in both BB32 and EB03 was inhibited by the induction of STAT5a{Delta}749 in a doxycycline dose-dependent manner, which was significant in BB32 cells at a concentration of 0.25 µg/ml and in EB03 cells at a concentration of 0.1 µg/ml.

Induction of STAT5a{Delta}749 Results in Inhibition of hGH-Stimulated DNA Synthesis in INS-1 Cells
To determine the influence of dominant negative STAT5 activity on cell proliferation, 3H-thymidine incorporation assay was carried out, as described above, on the transfected clones (Fig. 4Go). hGH stimulated 3H-thymidine incorporation by approximately 2-fold in the two transfected clones (1.9 ± 0.1 and 2.4 ± 0.4 fold over the basal levels in BB32 and EB03 cells, respectively) similar to what was found in INS-1 cells and BA15 cells (data not shown). The addition of 1 µg/ml doxycycline had no effect on either the basal or the hGH-induced DNA synthesis in INS-1 and BA15 cells (data not shown). Neither was the basal level in BB32 (upper panel) and EB03 (lower panel) affected by doxycycline addition alone. However, induction of STAT5a{Delta}749 by doxycycline in BB32 and EB03 blocked hGH-induced 3H-thymidine incorporation in a dose-dependent manner, whereas no effect of doxycycline was observed on the growth response to 10% FCS. The inhibition of 3H-thymidine incorporation was maximal at a doxycycline concentration of 0.5 µg/ml for both the BB32 cells and the EB03 cells, at which close to basal levels of DNA synthesis were detected (1.25 ± 0.03-fold and 1.1 ± 0.2-fold over basal in BB32 and EB03 cells, respectively).



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Figure 4. The Effect of STAT5a{Delta}749 Expression on hGH-Stimulated 3H-Thymidine Incorporation

BB32 and EB03 cells seeded in 96-well dishes were precultured in RPMI 1640 containing 0.25% BSA in the absence or presence of the indicated amounts of doxycycline for 24 h and then stimulated with either 0.5 µg/ml hGH or 10% FCS for a further 24-h period, the last 4 h in the presence of 3H-thymidine. Incorporated 3H-thymidine was measured as described in Materials and Methods. All experiments were done in quadruplicate in four independent experiments, and the results are expressed as fold induction compared with the untreated control cells (mean ± SEM, n = 3–4). Significant differences between the groups treated with hGH+Dox and the group treated with hGH alone are indicated: *, P <= 0.05; **, P <= 0.001, ***, P <= 0.0001. No significant differences were observed between the groups treated with 10% FCS+Dox and the group treated with 10% FCS alone. In both cell lines a significant difference (P <= 0.0001–0.01) between the hGH- or FCS-treated groups and the control groups ± Dox was detected.

 
To be able to determine whether the suppression of 3H-thymidine incorporation by dominant negative STAT5 expression was due to an effect on S-phase entry, the cell cycle profiles of hGH-stimulated parental INS-1 cells and transfected cells were examined by fluorescence-activated cell sorting (FACS) analysis (Table 1Go) using BrdU incorporation and DNA staining with propidium iodide. Stimulation with hGH for 24 h was found to increase the percentage of cells in S-phase by approximately 2-fold in both INS-1 cells and the transfected clones (INS-1, 2.5 ± 0.2; BA15, 1.9 ± 0.3; BB32, 1.8 ± 0.1; and EB03, 2.5 ± 0.3) (Table 1Go, middle column). The percentage of cells in the G2/M phase was not affected by hGH stimulation (Table 1Go, third column), whereas the percentage of cells in the G0/G1 phase was found to decrease upon hGH stimulation, indicating that hGH primarily in these experiments increases the G0/G1-to-S phase transition frequency. Neither the basal nor the hGH-induced cell cycle profiles in INS-1 and BA15 cells were affected by the addition of 0.5 µg/ml doxycycline. Furthermore, addition of doxycycline alone did not affect the cell cycle profiles in either BB32 or EB03 cells. However, the hGH-induced increase of cells in S-phase was totally blocked by the induction of STAT5a{Delta}749 expression in both BB32 and EB03 cells.


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Table 1. FACS Analysis of the Effect of STAT5a{Delta}749 Expression on hGH-Stimulated S-Phase Entry

 
Suppression of hGH-Induced PRLR 1A and Cyclin D2 mRNA Levels by Overexpression of STAT5a{Delta}749
The effects of STAT5a{Delta}749 overexpression on the endogenous mRNA levels of the PRLR 1A and the cell cycle regulators, cyclin D1, D2, and D3, were investigated by quantitative RT-PCR in BA15, BB32, and EB03 cells (Fig. 5Go). The PRLR mRNA levels were increased by hGH in BA15 cells by 7.4 ± 0.4-fold (upper panel) in BB32 by 4.1 ± 0.5-fold (middle panel), and in EB03 cells by 4.4 ± 0.4-fold (lower panel), respectively, in accordance with previous results in INS-1 cells (10). Addition of doxycycline alone had no effect on either the basal or the hGH-induced PRLR mRNA level in BA15 cells. Neither was the basal level affected by doxycycline in BB32 and EB03 cells. However, doxycyline-induced STAT5a{Delta}749 expression in BB32 and EB03 significantly suppressed the hGH-induced PRLR mRNA levels in a dose-dependent manner at a concentration of 0.05 µg/ml for both BB32 and EB03. At 0.5 µg/ml doxycycline, the induction by hGH was reduced to 0.8 ± 0.03-fold in BB32 cells and to 1.6 ± 0.2-fold in EB03 cells, supporting the previous finding of a primary role of STAT5 in the regulation of the PRLR 1A promoter.



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Figure 5. Effect of STAT5a{Delta}749 Expression on hGH-Stimulated mRNA Levels of the PRLR and Cyclin D1, D2, and D3

The three clones, BA15, BB32, and EB03, were cultured in the absence or presence of the indicated amounts of doxycycline for 48 h. Twenty-four hours before harvesting, 0.5 µg/ml hGH was added to the respective wells. cDNA was synthesized from total RNA, and RT-PCR was performed as described in Materials and Methods with primer sets specific for the PRLR exon 1A, and cyclin D1, D2, and D3. Primer sets specific for G6PDH or TBP were included as internal controls. The PCR products were separated by denaturing polyacrylamide gel electrophoresis and visualized and quantified by phosphoimager analysis. The PRLR 1A and the cyclin D1 and D3 PCR products were quantified relative to the internal standard TBP. The basal relative expression range in the three different clones were for PRLR1A, 25–88% of TBP; for cyclin D1, 99–281%; and for cyclin D3, 95–207%. The cyclin D2 products were quantified relative to G6PDH and ranged from 370–942% of G6PDH in the three untreated clones. The results are expressed as fold induction (mean ± SEM, n = 3) compared with control levels and represents the mean of three independent cDNA preparations. Significant differences between the groups treated with hGH+Dox and the group treated with hGH alone are indicated: * P <= 0.05; **, P <= 0.001; ***, P <= 0.0001. In all the cell lines, a significant difference (P <= 0.0001–0.001) was detected between the hGH-treated groups and the control groups ± Dox for the PRLR, cyclin D1, and cyclin D2 mRNA levels, respectively.

 
Examination of cyclin D mRNA levels revealed that cyclin D2 was positively regulated by hGH in all the cell lines (4.5 ± 0.5-fold in BA15, 3.7 ± 0.2-fold in BB32, and 4.2 ± 0.4-fold in EB03) over the basal expression levels which were 513% ± 64, 513% ± 35, and 565% ± 154 of G6PDH (glucose-6-phosphate-dehydrogenase), respectively (mean ± SEM, n = 3). Whereas doxycycline treatment had no effect on hGH-induced cyclin D2 mRNA levels in BA15 cells, a dose-dependent decrease was observed in both BB32 and EB03 cells, which was significant at 0.05 µg/ml in both BB32 and in EB03 cells. At 0.5 µg/ml doxycycline, the hGH-stimulated cyclin D2 mRNA levels were reduced to approximately 50% and 20%, in BB32 and EB03, respectively. The cyclin D3 mRNA levels (142% ± 14, 126% ± 14, and 163% ± 20 of TBP (TATA-binding-protein) in unstimulated BA15, BB32, and EB03 cells, respectively; means ± SEM, n = 3) were affected by neither hGH stimulation nor doxycycline treatment in all three clones. However, hGH significantly decreased the expression of cyclin D1 (0.42 ± 0.01-fold in BA15, 0.55 ± 0.06 in BB32, and 0.86 ± 0.05 in EB03) below basal levels which were 207% ± 39, 153% ± 25, and 230% ± 21, respectively (mean ± SEM, n = 3) of TBP mRNA levels, and treatment with doxycycline seemed to potentiate this effect in both BB32 cells (reduced to 0.1 ± 0.005) and EB03 cells (reduced to 0.4 ± 0.01) in a dose-dependent manner, whereas doxycycline treatment had no effect on cyclin D1 mRNA levels in BA15 cells. The basal mRNA expression levels of the three cyclin Ds were markedly different. Based on the estimation that G6PDH mRNA is approximately 25 times more abundant than TBP mRNA, the cyclin D2 mRNA level is 50- to 100-fold higher than that of cyclin D1 and D3.

Immunocytochemical Detection of Cyclin D2 in INS-1 Cells and Primary ß-Cells
Cyclin D2 protein expression was examined by peroxidase staining in INS-1 cells (Fig. 6Go, panel A) and primary ß-cells cultured in monolayer (Fig. 6Go, panel B). The INS-1 cells were cultured 4 days in the presence of 10% FCS, and the primary ß-cells were stimulated with hGH for 24 h before fixation and staining using a monoclonal anticyclin D2 antibody. Cyclin D2 protein was detected in the nucleus of a subpopulation of INS-1 cells and primary ß-cells.



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Figure 6. Immunocytochemical Detection of Cyclin D2 in INS-1 Cells and in Primary ß-Cells

A, INS-1 cells were seeded and cultured 4 days in complete medium, after which they were fixed and cyclin D2 expression was detected by peroxidase staining. B, Primary newborn rat islet-cells were cultured in monolayer for 5–7 days in the presence of 0.5 µg/ml hGH. The cells were deprived of hGH for 24 h and then restimulated for a further 24-h period before fixation and peroxidase staining. The staining shown is representative of two independent experiments.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Little is known about the postreceptor signaling of GH and PRL in proliferating insulin-producing cells. In primary neonatal ß-cells, stimulation of either GHRs or PRLRs using rat GH and rat PRL, respectively, leads to the same maximal effect on proliferation as when both receptors are activated using hGH, indicating that GHRs and PRLRs share downstream signaling molecules involved in proliferation (3). In INS-1 cells, cAMP was found to play a permissive role in hGH-induced proliferation, whereas hGH-stimulated increases of intracellular Ca2+ levels are probably not involved (28). In a recent study, rat GH (rGH)-stimulated INS-1 cell proliferation was found to be glucose dependent, and inhibitor experiments indicated involvement of the PI3K pathway in this effect whereas MAPK activation did not appear to play a role, in accordance with previous findings (13, 14). The INS-1 cell line has proven useful for the study of the effects of secretagogues, IGF-I and GH/PRL (13, 14, 31), and the role of tissue-specific transcription factors (32). Although it exhibits a mitogenic response comparable to that of primary rat ß-cells in culture, it is not known to what extent the mitogenic signaling apparatus in these cells is perturbed.

In the present study we examined the signaling pathways that may play a role in hGH-induced mitogenic signaling in INS-1 cells as well as in primary neonatal rat ß-cells. At 11 mM glucose, hGH was found to increase DNA synthesis by 2- to 3-fold over a 24-h stimulation period in both cell types in accordance with previous findings (1, 28). Inhibition of tyrosine protein kinase activity by coincubation with genistein resulted in a complete inhibition of hGH-induced proliferation in both cell types, as would be expected from inhibition of JAK2 and, thereby, most downstream signaling from both the GHR and the PRLR. PKC is involved in cell cycle progression in mammalian cells during the G1-phase and G2/M phase, and both positive and negative regulatory effects of PKC on cell growth have been reported (33). Staurosporine, an inhibitor of PKC, PKA, and protein kinase G, was found to inhibit basal as well as hGH-induced proliferation in INS-1 cells, but not in the primary ß-cells in which, on the contrary, a tendency for a growth-potentiating effect was observed. This paradoxical effect may reflect an important difference in the growth-signaling pathway between the primary ß-cells and the tumor cell line, INS-1. Inhibition of MAPK signaling pathways using the MEK inhibitor PD98059 and the p38 MAPK inhibitor SB203580 was only partially inhibitory to hGH-induced mitogenic signaling in INS-1 cells but did not affect primary ß-cells. Furthermore, we found no indications of hGH-stimulated activity of these enzymes by Western blot analysis of protein extracts using antiphosphothreonine-phosphotyrosine (anti-pTpY)-specific antibodies against these proteins (data not shown). These data confirm previous negative effects of this pathway in GH/PRL-stimulated INS-1 cell proliferation as mentioned above. The partial effect of the MAPK inhibitors may relate to a role of p42 and p38 MAPK in the glucose-induced proliferation of these cells (13, 34) and/or the glucose dependence of GH-induced INS-1 cell proliferation reported by Cousin et al. (14). The PI3K pathway seems to play an essential role in glucose- and IGF-I-stimulated INS-1 proliferation (31). However, inhibition of this pathway using coincubation with wortmannin did not influence hGH-induced proliferation in either INS-1 cells or primary ß-cells cultured in 11 mM glucose. This correlates well with the finding that rGH is unable to mediate the association of IRS-1, IRS-2, Grb2, or mSOS with PI3K (14) and the previous report that the mitogenic effects of IGF-I and hGH are additive, indicating distinct signaling mechanisms of these growth factors. Intriguingly, the study by Cousin et al. found an inhibitory effect of wortmannin on both glucose- and rGH-stimulated INS-1 cell proliferation in assays in which the cells had been precultured in the absence of glucose (14). Using these culture conditions, we have also seen a partial wortmannin-sensitive component of the hGH response (~50% inhibition, data not shown). However, in our hands considerable cell death occurs in the absence of glucose, which may obscure the interpretation of the data. A cautious conclusion from our inhibitor experiments is that the MAPK, PI3K, and the PKC signaling pathways, in contrast to findings in other cell types, are probably not essential for hGH-stimulated ß-cell replication, and we therefore turned our focus to the STAT5 pathway.

We have previously shown that activation of either GHRs or PRLRs in INS-1 cells induces binding of both STAT5a and STAT5b to the GLE of the PRLR 1A promoter (10). Stimulation of both GHRs and PRLRs using hGH led to a similar STAT5 activation. In the present study the kinetics of STAT5 activation in hGH-stimulated INS-1 and primary islet cells was examined. Pronounced STAT5 binding was observed after 15 min that persisted up to 24 h in both cell types. In INS-1 cells, the STAT5 activity after 24 h was comparable to that of 15 min hGH stimulation, whereas in islets a partial reduction was observed, indicating an influence of counterregulatory mechanisms operating in the primary cells, e.g. phosphatases or members of the suppressors of cytokine signaling (SOCS) family (35, 36). The long-term activation of STAT5 DNA binding in hGH-stimulated INS-1 cells correlates with the prolonged nuclear translocation of STAT5 in these cells in response to PRL as reported by Sorenson and co-workers (37). Interestingly, hGH-induced STAT5 DNA binding in the insulin-producing RIN-5AH cells was found to be only transient (9) with no detectable changes in STAT5 localization (37). The effects of GH and PRL on proliferation and PRLR gene expression in RIN-5AH cells are small compared with the effect of these hormones in INS-1 cells and cultured newborn rat islets (5, 10, 37, 38). Thus, the potency of GH and PRL action in insulin-producing cells may be correlated to the kinetics of GH- and PRL-induced STAT5 activation.

To address the role of STAT5 in GH/PRL-stimulated mitogenic signaling in ß-cells, we took advantage of a mutant of STAT5a that is deleted in its C-terminal transactivation domain and reportedly exerts dominant negative activity by its ability to inhibit the effect of both wild-type STAT5a and STAT5b (29). This mutant, STAT5a{Delta}749, was stably expressed in INS-1 cells using the inducible Tet-On gene expression system, and we showed in two different clones (BB32 and EB03) that STAT5a{Delta}749 expression and activation were doxycycline inducible. Furthermore, dominant negative activity of STAT5a{Delta}749 in the two clones was verified by analysis of the transcriptional effect on STAT5-regulated PRLR promoter, as previously described (10). The expression of STAT5a{Delta}749 was highly controllable, and there were no signs of leakage, as determined by Western blotting, EMSA, and reporter gene assay. Although the STAT5a{Delta}749 expressing cells do not have increased expression of PRLRs, they should still express the initial amount of GHRs (these are not regulated by STAT5) and the initial amount of PRLRs, as only one of the three characterized PRLR promoters contain a STAT5 response element (10). The expression of functional receptors in the presence of doxycycline was confirmed by the hGH-induced activation and binding of STAT5a{Delta}749 to DNA. Analysis of the effect of STAT5a{Delta}749 expression on DNA synthesis measured by 3H-thymidine incorporation and FACS analysis revealed a close correlation between STAT5 activity and hGH-stimulated S-phase entry, and we conclude that STAT5 activation is essential for GH/PRL-induced proliferation of INS-1 cells. Since previous experiments show that hGH, rat GH, and rat PRL have the same maximal mitogenic effect in neonatal rat ß-cells (3), the STAT5-mediated up-regulation of PRLRs is probably not rate limiting in short-term experiments. Whether this up-regulation after prolonged stimulation could be important for the extent of the ß-cell response remains to be addressed.

D-type cyclins, which regulate the activity of the cyclin-dependent kinases and S-phase entry, exhibit both cell type-specific and differential regulation in their expression, suggesting that the biological functions of these cyclins are not fully redundant (39). In this study we found that cyclin D2 mRNA was abundant in the INS-1 cells, whereas cyclin D3 and cyclin D1 mRNA were detectable at lower levels. The role of STAT5 in the growth of lymphoma cells has been discovered quite recently and, importantly, revealed a direct transcriptional effect of STAT5 on the human cyclin D1 gene (27). Furthermore, STAT5-deficient mice lacked peripheral T cell expression of cyclin D2, cyclin D3, cyclin A, and the cyclin-dependent kinase (cdk)-6 in response to interleukin-2 (IL-2) (21). In mouse and rat, no STAT5 binding element is present in the cyclin D1 promoter, whereas the cyclin D2 and D3 promoters contain PRL-responsive elements (40, 41), and a potential STAT5 binding sequence is present in the cyclin D2 promoter (Ref. 21 and our unpublished observations). In the present study we found that hGH increased the mRNA level of cyclin D2 and PRLR in the stably transfected cell lines, and the doxycycline- inducible inhibition of this effect indicates that STAT5 is involved in this regulation. However, whereas expression of the PRLR 1A mRNA was totally inhibited at the maximal doxycycline dose, only partial inhibition was observed for cyclin D2, indicating that other hGH-stimulated signaling pathways play a role in the transcriptional regulation of cyclin D2. Interestingly, an inhibitory effect of hGH on cyclin D1 mRNA levels was observed, which was augmented dose-dependently by doxycycline treatment. Our finding may indicate that STAT5 is involved in negative regulation of this cyclin through a mechanism that may involve competition of cofactors in transcription factor complexes (42). In a recent study a reciprocal regulation of cyclin D1 and D2 was identified in mouse myeloid leukemia cells in response to granulocyte-colony-stimulating-factor (G-CSF), indicating a correlation between G-CSF-stimulated up-regulation of cyclin D1 mRNA levels and G-CSF-induced apoptosis (43). One may speculate that the inverse effect of hGH on these cyclins in insulin-producing cells is related to a combined effect on antiapoptosis and replication.

In conclusion, our study has shown a requirement for STAT5 signaling in hGH-stimulated proliferation of INS-1 cells, which may involve a direct transcriptional effect on the cyclin D2 promoter. Future studies using adenoviral transfer of STAT5 mutants and establishment of transgenic animals expressing STAT5 mutants under the control of the insulin promoter will hopefully reveal whether these findings are relevant to primary ß-cells.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Cells, Hormones, and Chemicals
INS-1 cells, kindly provided by Dr. C. B. Wollheim, Geneva, Switzerland, were cultured in RPMI 1640 with glutamax supplemented with 10% heat-inactivated FCS, 100 U/ml penicillin, 100 µg/ml streptomycin, and 50 µM ß-mercaptoethanol (complete medium) at 37 C in a humidified atmosphere containing 5% CO2. INS-r3 cells were cultured in the media described above plus 100 µg/ml G418. For stably transfected clones, 100 µg/ml hygromycin B (Calbiochem, La Jolla, CA) was additionally added. Islets were isolated from 3- to 5-day-old Wistar rats by the collagenase digestion method (44) and cultured until further processing in RPMI 1640 supplemented with 10% newborn calf serum (NCS), 100 U/ml penicillin, 100 µg/ml streptomycin, 2 mM L-glutamine, 0.0375% NaHCO3, and 20 mM HEPES at 37 C in a humidified atmosphere. Recombinant hGH was obtained from Novo Nordisk A/S (Gentofte, Denmark). Doxycycline was from Sigma (St. Louis, MO). Wortmannin, PD98059, SB203580, genistein, and staurosporine were purchased from Calbiochem. All inhibitors were dissolved in dimethylsulfoxide (DMSO) (Sigma).

3H-Thymidine Incorporation
Cells were seeded in 96-wells plates (50,000 cells per well) and cultured for 2 days in 200 µl/well complete medium. The medium was changed to RPMI 1640 containing 0.25% BSA (Fraction V, Sigma) and culture proceeded for 24 h. For the inhibitor assay, various inhibitors (see cells, hormones, and chemicals) were added followed by stimulation with 0.5 µg/ml hGH for 24 h. For the stable clones, the cells were cultured in the absence or presence of doxycycline for 20–24 h followed by stimulation with either 0.5 µg/ml hGH or 10% FCS containing medium for an additional 24-h culture period. The last 4 h before harvesting 0.5 µCi [methyl, 1',2'-3H]Thymidine (no. TRK.565, Amersham Pharmacia Biotech, Buckinghamshire, UK) was additionally added per well. Cells were harvested onto a filter paper (Filtermate 196, Packard Instruments, Meriden, CT) by a cell harvester (Wallac, Inc., Gaithersburg, MD) using H2O for lysis. The filter paper was dried for 1 h at 37 C and transferred to a bag containing 5 ml of Optiscent scintillation fluid (Wallac, Inc.). The filter was counted in a 1450 Microbeta Plus counter (Wallac, Inc.).

Preparation of Monolayers of ß-Cells and 5-BrdU Labeling
Monolayer cultures of islet cells were prepared essentially as previously described (1). Briefly, islets were precultured for 5–7 days in RPMI 1640, containing 0.5% human serum (HS) and were then dispersed into single cells by trypsin-EDTA treatment. The cells (50–100,000) were plated in plastic cell culture 9-cm2 slideflasks (Nunc, Roskilde, Denmark) in RPMI 1640 medium containing 2% HS and 0.5 µg/ml hGH. The cells were allowed to attach and establish a monolayer for 5–7 days, after which they were washed twice in medium without hGH and then cultured for 24 h in RPMI 1640 containing 2% HS. The medium was changed, and the cells were cultured further for 24 h in the presence of 10 µM BrdU, and in the absence or presence of the various protein kinase inhibitors, which were added before the addition of 0.5 µg/ml of hGH. The cells were washed twice in RPMI 1640 without serum before fixation in 1% paraformaldehyde in 0.1 M phosphate buffer, pH 7.4. The cells were double stained for BrdU and insulin as described previously (1). Briefly, the cells were exposed to 1.5 M HCl for 30 min and washed. They were stained with a monoclonal mouse antibody to BrdU (no. M 0744, DAKO Corp., Glostrup, Denmark) diluted 1:50 and with guinea pig antiinsulin antibody (Novo Nordisk A/S) diluted 1:500. The antibodies were visualized by a Texas red-conjugated goat antimouse-IgG (no. 115–075-100, Jackson ImmunoResearch Laboratories, Inc. West Grove, PA) and fluorescein isothiocyanate (FITC)-conjugated goat antiguinea pig-IgG (no. 106–095-003, Jackson ImmunoResearch Laboratories, Inc.) both diluted 1:100. Dilution of the antibodies was performed in PBS with 0.3% Triton X-100 and 0.1% human serum albumin (HSA). The slides were mounted in 20% glycerol/0.05 M Trisma base adjusted to pH 8.4 and stored at 4 C.

Stable Transfection
The establishment of the stable clone, INS-r3, expressing the reverse tetracycline-dependent transactivator, has been described previously (30) and was kindly provided by Dr. P. B. Iynedjian (Geneva, Switzerland). A vector containing the cDNA encoding the STAT5a mutant, STAT5a{Delta}749, which lacks the C-terminal transactivation domain, was provided by Dr. B. Groner (Freiburg, Germany). This mutant cDNA was subcloned into the Tet-On gene expression vector, pTRE (CLONTECH Laboratories, Inc. Palo Alto, CA) using the EcoRI restriction site of the polylinker. INS-r3 cells were seeded (1 x 107 cells per 100-mm dish) and cultured overnight in complete medium containing 100 µg/ml G418. The following day the medium was changed to Opti-MEM, and transfection was carried out using the LipofectAMINE PLUS Reagent (Life Technologies, Inc., Paisley, UK) essentially as described by the manufacturer. The cells were transfected overnight with 3.7 µg pTK-Hyg Vector (CLONTECH Laboratories, Inc.) and 18.1 µg pTRE vector with or with out STAT5a{Delta}749 insert. The cells were cultured for 2 days in complete medium and were then trypsinized and reseeded in 5 x 100 mm dishes using complete medium containing 100 µg/ml G418 and hygromycin. Medium was changed every fourth day, and antibiotic-resistant colonies of cells were isolated after 2–4 weeks, transferred to 24-well plates, and split to single cells by trypsination. Hygromycin-resistant clones were tested for integration of pTRE by PCR amplification on purified DNA. The DNeasy Tissue kit (QIAGEN, Valencia, CA) was used for DNA purification. The PCR reaction was run with 250 ng of total DNA as a template, and 50 pmol per specific pTRE primers (CLONTECH Laboratories, Inc.), 1.25 U Taq polymerase (Promega Corp., Madison, WI), thermophilic DNA 10x buffer (Promega Corp.), 0.2 mM deoxynucleoside triphosphate (dNTP) (Amersham Pharmacia Biotech), 1.5 mM MgCl2 (Promega Corp.), and H2O to 50 µl. A single denaturing step at 94 C/1 min was followed by 25 cycles as given: 94 C/15 sec; 63 C/1 min; 68 C/3 min. The products were detected on 1% agarose gel.

Western Blot Analysis
Cells were seeded in 100-mm dishes (4 x 106 cells per dish) and cultured for 2 days in 10 ml complete medium per dish. The medium was changed to medium containing 0.25% BSA, and cells were cultured approximately 20 h in the presence and absence of doxycyline as indicated. The cells were washed once in cold PBS, scraped off in 1 ml PBS, transferred to microfuge tubes, and pelleted. The cells were resuspended in 500 µl PBS, containing 1% NP40, 0.1% SDS, 1 µg/ml leupeptin, 1 µg/ml aprotinin, and 0.5 mM AEBSF (Calbiochem) and allowed to lyse on ice for 30 min. Cell debris was removed by centrifugation at 15,000 x g for 20 min, and the supernatants were stored at -80 C after addition of 125 µl 5x sample buffer. Proteins were denatured by boiling for 2 min and separated by electrophoresis on a 7.5% SDS polyacrylamide gel. For protein size determination, High-Range Rainbow Marker (Amersham Pharmacia Biotech) was used. The proteins were transferred to a nitrocellulose membrane by Western blotting for 2 h at 200 mA. The membrane was blocked for 15 min and washed once in PBS before probing with antibodies. The primary antibody was either no. S21520 (Transduction Laboratories, Inc., Lexington, KY) raised against aa 451–649 of STAT5 or (C-17)-G no. SC835 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA) raised against the carboxy terminus of STAT5b diluted 1:1000. The secondary antibody was antimouse IgG horseradish peroxidase-linked whole antibody (Amersham Pharmacia Biotech) diluted 1:5,000. Proteins were visualized using the ECL Western blotting detection reagents (Amersham Pharmacia Biotech).

Nuclear Extracts
Isolated newborn rat islets (5,000 islets per dish), which had been precultured for 1 week in 15 ml of RPMI 1640 supplemented with 0.5% normal HS, were stimulated with 0.5 µg/ml hGH for 15 min before harvest. INS-1 cells and the three INS-r3 clones (BA15, BB32, and EB03) were seeded in 100-mm dishes (4 x 106 cells per dish) and cultured for 2 days in 10 ml complete medium per dish. The medium was changed to medium containing 0.25% BSA, and cells were cultured for 20 h in the presence or absence of doxycyline. When indicated, the cells were stimulated with 0.5 µg/ml hGH for 15 min. Nuclear extracts were prepared essentially as described previously (9, 10). Briefly, cells were lysed in hypotonic buffer containing 1% Triton X-100. Nuclei were collected by centrifugation, and nuclear proteins were extracted in hypertonic buffer containing 400 mM NaCl. After centrifugation, aliquots of the supernatants were frozen in liquid nitrogen and stored at -80 C. Protein concentrations were measured using the Bio-Rad protein assay (Bio-Rad Laboratories, Inc. Hercules, CA).

EMSA
EMSA was performed essentially as described previously (9, 10). Briefly, the double-stranded oligonucleotide 1A-GLE (5'-agctAGTTCTAGGAATAagct) containing a STAT5 binding element derived from the rat PRLR 1A promoter (45) was radiolabeled in a fill-in reaction using [{alpha}-32P] dATP (Amersham Pharmacia Biotech) and DNA polymerase (Klenow fragment) and used as probe (10). Nuclear extracts (10 µg) were incubated at 30 C with 20 fmol of probe in a 20 µl reaction. Free and bound probe were separated by nondenaturing PAGE and visualized by autoradiography.

Transient Transfection and Dual Luciferase Reporter (DLR) Assay
Cells were seeded in 24-well plates (300,000 cells per well) in 500 µl/well complete medium. The cells were transfected as described previously (10) with 1.5 µl LipofectAMINE PLUS Reagent (Life Technologies, Inc.) and 0.5 µg DNA [250 ng of pGL2–1A, 10 ng of pRL-SV40 plasmid (internal control) and 240 ng of pUC18 plasmid]. The cells were transfected overnight in Opti-Mem (240 µl/well). The medium was changed to RPMI 1640 containing 0.5% FCS (500 µl/well) and incubated for 24 h in the presence or absence of doxycycline. Seven hours before harvesting, 0.5 µg/ml hGH was added to the respective wells. The cells were lysed by adding 100 µl/well of 1x passive lysis buffer (supplied with DLR Assay System no. E1910, Promega Corp.) followed by shaking the plate for 15 min at room temperature. The cell extracts were stored in the plate at -80 C until measuring was performed as described previously (10). The pRL-SV40 vector contains the coding region of the Renilla luciferase gene under the transcriptional control of the SV40 early enhancer/promoter (Promega Corp.). The pGL2–1A is generated by the insertion of the 5'-flanking region of PRLR exon 1A (-462/+81) into the pGL2-basic vector that contains the coding region of the firefly luciferase gene (45). The pUC18 vector was used as carrier plasmid.

FACS
Cells were seeded in six-well plates (500,000 cells per well) and cultured for 2 days in 3 ml complete medium per well. The medium was changed to medium containing 0.25% BSA, and cells were cultured approximately 20 h in the presence and absence of doxycyline as indicated. Respective wells were stimulated 24 h with 0.5 µg/ml hGH and 2 h before harvesting, 10 µM BrdU was added per well. The cells were harvested by adding 100 µl 0.5% trypsin-EDTA per well. Trypsination was stopped by adding 900 µl serum-containing medium, and cells were transferred to Eppendorf (Madison, WI) tubes, pelleted, and resuspended in 500 µl -20 C 70% ethanol. Cells were stored at 4 C up to 1 week. For denaturation of DNA, 500 µl of 3 M HCl were added and the cells were incubated at room temperature for 30 min. The cells were washed once in PBS containing 0.1% HSA and 0.3% Triton X-100 and were resuspended in 1 ml of this buffer containing mouse anti-BrdU antibody (DAKO Corp.) in a 1:100 dilution. Incubation was carried out overnight at 4 C with rotation. Cells were washed twice, resuspended in 1 ml buffer containing FITC-conjugated goat-antimouse IgG no. 115–095-003 (Jackson ImmunoResearch Laboratories, Inc.), and incubated for 45 min at 4 C in the dark. The cells were washed twice, resuspended in 500 µl PBS containing 5 µg/ml propidium iodide, and placed in the dark. FACS analysis was carried out using Cell Quest (Becton Dickinson and Co., San Jose, CA) as software.

RT-PCR
Cells were seeded in 60-mm dishes (2 x 106 cells per dish). The cells were cultured for 2 days in 4 ml complete medium per dish. The medium was changed to medium containing 0.25% BSA, and cells were cultured approximately 20 h in the presence and absence of doxycyline as indicated. Respective wells were stimulated 24 h with 0.5 µg/ml hGH. Total RNA was extracted using the RNeasy method from QIAGEN (Chatsworth, CA). cDNA was synthesized from 1 µg RNA using AMV Reverse Transcriptase and dNTP mix from Stratagene (La Jolla, CA) and random primers from Life Technologies, Inc. The reaction was run at 42 C for 1 h, and the sample was diluted in 40 µl of 0.1% Triton-X-100 and stored at -20 C. PCR was carried out in 12.5 µl reactions using 0.75 µl of cDNA as template. The primer sequences were: PRLR, 5'-TTG TGG ATC TCA GGT TTC CCT GGT G (forward); 5'-AGC GAG CTG GAT TCT AGG GAA ACA T (reverse); cyclin D1, 5'-TCT ACA CTG ACA ACT CTA TCC G (forward); 5'-TAG CAG GAG AGG AAG TTG TTG G (reverse); cyclin D2, 5'- AGA CCT TCA TCG CTC TGT GT (forward); 5'- TAG CAGATG ACG AAC ACG CC (reverse); cyclin D3, 5'-CTG CTG GCG GGA ATC ACA (forward); 5'-GGC CCC TCC TCT GCT TGG T (reverse); G6PDH, 5'-GAC CTG CAG AGC TCC AAT CAA C (forward); 5'-CAC GAC CCT CAG TAC CAA AGG G (reverse); TBP, 5'-ACC CTT CAC CAA TGA CTC CTA TG (forward); 5'-ATG ATG ACT GCA GCA AAT CGC (reverse). The expected lengths of the various PCR products were as follows: PRLR, 329 bp; cyclin D1, 304 bp; cyclin D2, 372 bp; cyclin D3, 246 bp; G6PDH, 214 bp; TBP, 192 bp. The PCR incubates contained 50 mM KCl; 10 mM Tris-HCl; 1.5 mM MgCl2; 40 µM dATP; dGTP; and dTTP; 20 µM dCTP; 2.5 mCi of 3,000 Ci/mmol [{alpha}-33P]dCTP; 10 pmol of each primer and 2.5 U Ampli Tag Gold polymerase. A single denaturing step at 94 C/10 min was followed by either 20 cycles (cyclin D2) or 25 cycles (PRLR, cyclin D1/D3) as given: 94 C/30 sec; 55 C/1 min; 72 C/1.5 min. PCR products were separated on 6% denaturing polyacrylamide gels (GEL-MIX 6, Life Technologies, Inc.), dried, and exposed to Phosphorimage storage screens that were scanned by Phosphorimager series 400 (Molecular Dynamics, Inc., Sunnyvale, CA), and band intensities were calculated using the program Image Quant (Molecular Dynamics, Inc.). The PRLR and the cyclin D1 and D3 were quantified relative to the internal standard TBP and cyclin D2 was quantified relative to the internal standard G6PDH.

Immunocytochemistry
INS-1 cells were seeded in slide flasks (100,000 cells per flask) and cultured 4 days in complete medium. Newborn rat islets were dispersed into single ß-cells and cultured as described above. The cells were fixed in 1% paraformaldehyde and stained with peroxidase using the HISTOSTAIN-PLUS KIT (Zymed Laboratories, Inc. South San Francisco, CA) according to the instructions provided by the manufacturer. The primary antibody monoclonal cyclin D2 antibody no. MS-221 (NeoMarkers, Union City, CA) diluted 1:100 in PBS + 0.1% HSA + 0.3% Triton-X-100 was incubated for 1 h.

Statistical Analysis
Statistical analysis was performed using SAS 6.12 software (SAS Institute, Cary, NC). Two-way ANOVA with Dunnett’s method for adjustment of multiple comparisons was carried out.


    ACKNOWLEDGMENTS
 
We want to thank Dagny Jensen and Hanne Richter Olesen for excellent technical assistance. We are indebted to Dr. B. Groner, Tumor Biology Center, Freiburg, Germany, for kindly supplying STAT5a{Delta}749 cDNA; to Dr. P. B. Iynedjian, University of Geneva School of Medicine, Geneva, Switzerland, for supplying INS-r3 cells; and Dr. C. B. Wollheim, Centre Medical Universitaire, Geneva, Switzerland, for the supply of INS-1 cells. We thank Dr. S. N. Jakobsen and Dr. N. Billestrup, Novo Nordisk, Bagsværd, Denmark, and Dr. K. Seedorf, Lilly Research Laboratories, Hamburg, Germany, for helpful advice and discussion.


    FOOTNOTES
 
Address requests for reprints to: Annette Møldrup, Department of Islet Discovery Research, Novo Nordisk A/S, Novo Alle 1, 1KMS03, 2880 Bagsværd, Denmark. E-mail amp{at}novo.dk

Received for publication March 30, 2000. Revision received September 11, 2000. Accepted for publication September 20, 2000.


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

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