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• W2005703352 abstract "Glucocorticoids regulate the expression of the G1 progression factor, cyclin D3. Cyclin D3 messenger RNA (CcnD3 mRNA) stability decreases rapidly when murine T lymphoma cells are treated with the synthetic glucocorticoid dexamethasone. Basal stability of CcnD3 mRNA is regulated by sequences within the 3′-untranslated region (3′-UTR). RNA-protein interactions occurring within the CcnD3 3′-UTR have been analyzed by RNA electrophoretic mobility shift assay. Three sites of RNA-protein interaction have been mapped using this approach. These elements include three pyrimidine-rich domains of 25, 26, and 37 nucleotides. When the cyclin D3 3′-UTR was stably overexpressed, the endogenous CcnD3 mRNA was no longer regulated by dexamethasone. Likewise, overexpression of a 215-nucleotide transgene that contains the 26- and 37-nucleotide elements blocks glucocorticoid inhibition of CcnD3 mRNA expression. These observations suggest that the 215-nucleotide 3′-UTR element may act as a molecular decoy, competing for proteins that bind to the endogenous transcript and thereby attenuating glucocorticoid responsiveness. UV-cross-linking experiments showed that two proteins of approximate molecular weight 37,000 and 52,000 bind to this 3′-UTR element. Glucocorticoids regulate the expression of the G1 progression factor, cyclin D3. Cyclin D3 messenger RNA (CcnD3 mRNA) stability decreases rapidly when murine T lymphoma cells are treated with the synthetic glucocorticoid dexamethasone. Basal stability of CcnD3 mRNA is regulated by sequences within the 3′-untranslated region (3′-UTR). RNA-protein interactions occurring within the CcnD3 3′-UTR have been analyzed by RNA electrophoretic mobility shift assay. Three sites of RNA-protein interaction have been mapped using this approach. These elements include three pyrimidine-rich domains of 25, 26, and 37 nucleotides. When the cyclin D3 3′-UTR was stably overexpressed, the endogenous CcnD3 mRNA was no longer regulated by dexamethasone. Likewise, overexpression of a 215-nucleotide transgene that contains the 26- and 37-nucleotide elements blocks glucocorticoid inhibition of CcnD3 mRNA expression. These observations suggest that the 215-nucleotide 3′-UTR element may act as a molecular decoy, competing for proteins that bind to the endogenous transcript and thereby attenuating glucocorticoid responsiveness. UV-cross-linking experiments showed that two proteins of approximate molecular weight 37,000 and 52,000 bind to this 3′-UTR element. heterogeneous nuclear ribonucleoprotein untranslated region β-galactosidase Malignant lymphoid cells of thymic origin cease to proliferate and often die when exposed to glucocorticoids (1.Harmon J.M. Norman M.R. Fowlkes B.J. Thompson E.B. J. Cell Physiol. 1979; 98: 267-278Crossref PubMed Scopus (181) Google Scholar, 2.Schwartzman R.A. Cidlowski J.A. Int. Arch. Allergy Immunol. 1994; 105: 347-354Crossref PubMed Scopus (114) Google Scholar), and glucocorticoids are an important tool for treatment of leukemias and other malignant and nonmalignant lymphoproliferative diseases. Glucocorticoids have also been proposed to be responsible for triggering apoptosis of CD4+/CD8+ cells in the thymus (reviewed in Ref. 3.Cohen J.J. Semin. Immunol. 1992; 4: 363-369PubMed Google Scholar). We have analyzed glucocorticoid effects on murine T lymphoma P1798 cells in an attempt to elucidate the molecular mechanisms that account for glucocorticoid inhibition of cell proliferation and induction of cell death. The data indicate that glucocorticoids induce G0/G1arrest by decreasing the expression of two critical cell G1progression factors: c-Myc (4.Forsthoefel A.M. Thompson E.A. Mol. Endocrinol. 1987; 12: 899-907Crossref Scopus (54) Google Scholar) and cyclin D3 (5.Reisman D. Thompson E.A. Mol. Endocrinol. 1995; 9: 1500-1509PubMed Google Scholar). P1798 cells utilize cyclin D3 as the principal D-type cyclin in G1/S phase transition (6.Ajchenbaum F. Ando K. De Caprio J.A. Griffin J.D. J. Biol. Chem. 1993; 268: 4113-4119Abstract Full Text PDF PubMed Google Scholar). Cyclin D2 is barely detectable and cyclin D1 is undetectable in P1798 cells (5.Reisman D. Thompson E.A. Mol. Endocrinol. 1995; 9: 1500-1509PubMed Google Scholar). Consequently, inhibition of cyclin D3 blocks activation of Cdk4/Cdk6 and precludes progression through G1 phase (5.Reisman D. Thompson E.A. Mol. Endocrinol. 1995; 9: 1500-1509PubMed Google Scholar). Simultaneous overexpression of c-Myc plus cyclin D3 prevents cell cycle arrest and apoptosis of glucocorticoid-treated cells, although neither c-Myc nor cyclin D3 alone will suffice to protect cells (7.Rhee K. Bresnahan W. Hirai M. Thompson E.A. Cancer Res. 1995; 55: 4188-4195PubMed Google Scholar). Glucocorticoids control cyclin D3 expression in T-lymphoid cells by decreasing the stability of CcnD3 mRNA (5.Reisman D. Thompson E.A. Mol. Endocrinol. 1995; 9: 1500-1509PubMed Google Scholar). CcnD3 mRNA is quite stable in mid-log phase P1798 cells, as determined by measuring mRNA abundance after treatment with actinomycin D. However, the rate of degradation of CcnD3 mRNA increases within 2 h after addition of glucocorticoids, to the extent that a 50% decrease in mRNA abundance occurs within 60–90 min after addition of actinomycin D to glucocorticoid-treated cells (5.Reisman D. Thompson E.A. Mol. Endocrinol. 1995; 9: 1500-1509PubMed Google Scholar). This posttranscriptional mechanism of gene expression, coupled with the short half-life of the cyclin D3 protein, ensures a rapid response to glucocorticoids. There are several examples of glucocorticoid effects on mRNA stability (8.Petersen D.D. Koch S.R. Granner D.K. Proc. Natl. Acad. Sci. U. S. A. 1989; 86: 7800-7804Crossref PubMed Scopus (88) Google Scholar, 9.Gadson Jr., P. McCoy J. Biochim. Biophys. Acta. 1993; 1173: 22-31Crossref PubMed Scopus (4) Google Scholar, 10.Peppel K. Vinci J.M. Baglioni C. J. Exp. Med. 1991; 173: 349-355Crossref PubMed Scopus (122) Google Scholar), but the molecular mechanisms that underlie such events are largely unknown. During the last decade, several RNA motifs have been identified that control the degradation rate of specific mRNAs. Iron-responsive elements (reviewed in Ref. 11.Hentze M.W. Kuhn L.C. Proc. Natl. Acad. Sci. U. S. A. 1996; 16: 8175-8182Crossref Scopus (1137) Google Scholar) influence the stability of the transcripts of several genes that are involved in iron metabolism. AU-rich sequences (10.Peppel K. Vinci J.M. Baglioni C. J. Exp. Med. 1991; 173: 349-355Crossref PubMed Scopus (122) Google Scholar, 12.Bohjanen P.R. Petryniac B. June C.H. Thompson C.B. Lindsten T. Mol. Cell. Biol. 1991; 11: 3288-3295Crossref PubMed Scopus (233) Google Scholar, 13.Winstall E. Gamache M. Raymond V. Mol. Cell. Biol. 1995; 15: 3796-3804Crossref PubMed Scopus (48) Google Scholar), and polypyrimidine tracts (14.Kohn D.T. Tsai K. Cansino V. Neve R.L. Perrone-Bizzozero N.I. Brain Res. 1996; 36: 240-250Google Scholar, 15.Irwin N. Baekelandt V. Goritchenko L. Benowitz L.I. Nucleic Acids Res. 1997; 25: 1281-1288Crossref PubMed Scopus (76) Google Scholar, 16.Czyzyk-Krzeska M.F. Beresh J.E. J. Biol. Chem. 1996; 271: 3293-3299Abstract Full Text Full Text PDF PubMed Scopus (95) Google Scholar) are known to control the stability of some mRNAs. Several families of RNA-binding proteins have been implicated in controlling the stability of mRNAs. The heterogeneous nuclear ribonucleoprotein (hnRNP)1 complexes involved in maturation of mRNAs contain more than 20 protein species, many of which are involved in mRNA stability control (17.Matunis M.J. Michael W.M. Dreyfuss G. Mol. Cell. Biol. 1992; 12: 164-171Crossref PubMed Scopus (237) Google Scholar, 18.Zaidi S.H.E. Malter J.S. J. Biol. Chem. 1995; 270: 17292-17298Abstract Full Text Full Text PDF PubMed Scopus (101) Google Scholar). Vertebrate homologues of the Drosophila embryonic lethal abnormal vision protein family regulate the stability of mRNAs encoding tumor necrosis factor-α (19.Sakai K. Kitagawa Y. Hirose G. FEBS Lett. 1999; 446: 157-162Crossref PubMed Scopus (32) Google Scholar), N-Myc (20.Changnovich D. Fayos B.E. Cohn S. J. Biol. Chem. 1996; 271: 33587-33591Abstract Full Text Full Text PDF PubMed Scopus (35) Google Scholar, 21.Changnovich D. Cohn S.L. J. Biol. Chem. 1996; 271: 33580-33586Abstract Full Text Full Text PDF PubMed Scopus (38) Google Scholar), and the cyclin-dependent kinase inhibitor p21Cip1 (22.Joseph B. Orlian M. Furneaux H. J. Biol. Chem. 1998; 273: 20511-20516Abstract Full Text Full Text PDF PubMed Scopus (112) Google Scholar). Regulation of replication-dependent histone mRNAs involves specific RNA-protein interactions within the 3′-UTR. Processing and degradation of those mRNAs are mediated by a 26-base stem-loop structure and a M r 31,000 protein (23.Wang Z.F. Whitfield M.L. Ingledue III, T.C. Diminski Z. Marzluff W.F. Genes Dev. 1996; 10: 3028-3040Crossref PubMed Scopus (218) Google Scholar). The general conclusions that one may advance are that 1) mRNA stability is frequently determined by RNA-protein interactions, and 2) those interactions frequently occur within the 3′-UTR. The experiments described in this paper were designed to test the hypothesis that the 3′-UTR of CcnD3 mRNA contains specific protein-binding elements that are involved in glucocorticoid regulation of degradation of the mature transcript. We have undertaken to analyze RNA-protein interactions within the 3′-UTR, to map the nucleotide components that are required for such interactions, and to characterize the RNA-binding proteins with which these elements interact. Our results indicate that there is a protein-binding element within the 3′-UTR of CcnD3 mRNA. This element interacts with at least two proteins, and this interaction is necessary for glucocorticoid-mediated destabilization of CcnD3 mRNA. Murine T-lymphoma P1798-S20 cells were maintained in suspension culture in RPMI 1640 medium supplemented with 3 mm glutamine, 25 mm HEPES, 20 mm 2-mercaptoethanol, and 2% fetal bovine serum, at 37 °C in 5% CO2. Mid-log phase cultures containing 5–8 × 105 cell/ml were used for all experiments. Dexamethasone was dissolved in 70% ethanol as a 0.1 mm stock, and this was diluted into medium to a final concentration of 0.1 μm dexamethasone and 0.07% ethanol in all experiments. Ethanol has no effect on P1798 cultures at this concentration. RNA isolation was performed using TRIzol reagent from Life Technologies Inc. Northern blotting was carried out under standard conditions, and all Northern blotting data were normalized to 18 S RNA. Autoradiographic data were quantified using a Lynx 5000 digital workstation. Cytosolic protein extracts were prepared using the method described by Sun and Antony (24.Sun X.L. Antony A.C. J. Biol. Chem. 1996; 271: 25539-25547Abstract Full Text Full Text PDF PubMed Google Scholar). Cells were washed with 5 packed cell volumes of phosphate-buffered saline by centrifugation at 2000 rpm in a Beckman JA-14 rotor at 4 °C. Cells (5 × 107) were incubated at 4 °C in Buffer A (10 mm HEPES, pH 7.9, containing 1.5 mm MgCl2, 10 mm KCl, 0.5 mm dithiothreitol) for 10 min at a ratio of 5 ml of Buffer A per ml of packed cell volume. The cells were centrifuged (5 min at 2000 rpm in a Beckman JA-14 rotor), suspended in 2 ml of Buffer A per ml of packed cells, and then lysed by 10 strokes of a glass Dounce homogenizer (B pestle). Cell lysis was confirmed by microscopy. The lysate was centrifuged at 2000 rpm for 10 min at 4 °C. The resultant supernatant fraction was mixed with 0.11 supernatant volumes of Buffer B (300 mm HEPES, pH 7.0, containing 10 mm MgCl2, 1.4M KCl), and this solution was centrifuged for 60 min at 100,000 × g. The supernatant fraction was recovered and dialyzed for 8 h against 20 volumes of Buffer C (20 mm HEPES, pH 7.9, containing 100 mm KCl, 0.2 mm EDTA 20% (v/v) glycerol, 0.5 mm phenylmethylsulfonyl fluoride, and 0.5 mmdithiothreitol). Aliquots (100 μl) were frozen at –80 °C, and once thawed, they were never reused. The MAXIscript kit from AMBION Inc. was used for the synthesis of cyclin D3 3′-UTR probes. [α-32P]CTP (800Ci/mmol, NEN Life Science Products, Inc.) was used to label the probes. The full-length 3′-UTR of murine CcnD3 mRNA from nucleotide 933 to 1871 was amplified by polymerase chain reaction and cloned into InVitrogen pCRII to yield pCRII/D3UTR. This plasmid was digested with BamHI, and run-off transcription was performed from the T7 promoter to generate the D3 3′-UTR probe (specific activity 217.8Ci/μm). Nucleotides 933–1574 of the cyclin D3 cDNA were eliminated by digesting pCRII/D3UTR with StuI and EcoRV and ligating the blunt ends to generate a plasmid named pCRII/D3UTRΔ1–1574. pCRII/D3UTRΔ1–1574 was digested with AvrII (which cuts at 1789), and run-off transcription was performed with T7 RNA polymerase to yield a probe corresponding to nucleotides 1575–1789. This probe was named BC (specific activity, 54.4Ci/μm). The probes were purified by electrophoresis in 8 m urea on 5% polyacrylamide gels. RNA-protein binding reactions and electrophoresis of the complexes formed were carried out using a variation of the method described by Leibold and Munro (25.Leibold E.A. Munro H.N. Proc. Natl. Acad. Sci. U. S. A. 1989; 85: 2171-2175Crossref Scopus (560) Google Scholar). Binding reactions were carried out with various amounts of cytosolic extract and 15 pg of 32P-labeled RNA probe in 30 μl of Buffer D (containing 10 mm HEPES, pH 7.6, 3 mmMgCl2, 40 mm KCl, 2% glycerol, 1 mm dithiothreitol, and 5 mg/ml heparin). Cytosolic protein extracts were diluted in Buffer C (described above) to equalize the volumes among different reactions in each experiment. Reactions performed with the CcnD3 3′-UTR probe were incubated for 20 min at 30 °C, and then 20 units of ribonuclease T1 were added and the reaction was incubated for another 20 min at 30 °C. Under these conditions, the ribonuclease T1 digestion is incomplete. In one experiment, proteinase K was added to a final concentration of 2 mg/ml for 20 min at 30 °C. Reactions performed with the BC probe (1575–1789) were incubated for 30 min at 30 °C and contained 3.3 ng/μl of yeast tRNA as a nonspecific competitor, and there was no ribonuclease T1 digestion of the BC-protein complex prior to electrophoresis. Binding conditions were otherwise identical to those described for the full-length 3′-UTR. In competition experiments, labeled probe was mixed with the unlabeled probes before addition of the cytosolic extracts. Electrophoresis of RNA-protein complexes was carried out in 5% nondenaturing polyacrylamide gels. Autoradiography was performed at –80 °C. RNA-protein complexes were resolved by electrophoretic mobility shift. These complexes were excised from polyacrylamide gels, and the labeled RNA was isolated by phenol-chloroform extraction. The isolated RNA fragments were digested in 10 mm Tris-HCl, pH 7.4, containing 1 mm EDTA and 20 units of ribonuclease T1 for 60 min at 37 °C. Ribonuclease T1-resistant fingerprint fragments were separated by electrophoresis in 8 m urea and 0.1%SDS on 20% polyacrylamide gels. The method used to cross-link the BC probe to the RNA-binding proteins was performed as described by Chang et al. (26.Chang K.H. Brown E.A. Lemon S.M. J. Virol. 1993; 67: 6716-6725Crossref PubMed Google Scholar) with some modifications. Binding reactions were carried out with various amounts of cytosolic extract and 32P-labeled RNA probe in Buffer D containing 0.1 μg/μl of yeast tRNA. After 20 min at 30 °C the samples were transferred to ice and irradiated for 30 min with a UV light source (254 nm, 15 W) at a distance of approximately 4 cm. After irradiation, RNA was digested with 1 mg/ml of ribonuclease A for 1 h at 37 °C. The UV cross-linked products were separated on a 12% SDS-polyacrylamide gel and detected by autoradiography. pUHD10–4 was constructed by replacing the polylinker of pUHD10–3 (26.Chang K.H. Brown E.A. Lemon S.M. J. Virol. 1993; 67: 6716-6725Crossref PubMed Google Scholar) with a multiple cloning site that contains unique SacII (442), EcoRI (449), EcoRV (457), SalI (462), AccI (463), PstI (473), BglII (475), XbaI (481), andBamHI (487) sites. Numbers in parentheses are positions relative to the XhoI site of the parental pUHD10–3.pUHD10–5 was made by replacing the polylinker of pUHD10–3 with a multiple cloning site that contains unique sites forSacII (446), NotI (446), EcoRV (455),SphI (469), AvrII (470), and PacI (480). pUHDZeo4 was made by ligating theXhoI/PvuII fragment of pZeo (InVitrogen) with theXhoI/PvuII fragment of pUHD10–4.pUHDZeo5 was made by ligating theXhoI/NaeI fragment of pUHD10–5 into theXhoI/PvuII fragment of pZeo. These pUHDZeo plasmid conveys stable resistance to the antibiotic zeocin. Modified tetracycline-repressible expression vectors and their nucleotide sequences are available upon request. The 1878-base pair EcoRI fragment containing full-length CcnD3 cDNA was cloned into pUHDZeo4 to create p4ZD3FL (full-length), which we have designated tetD3FL. p4ZD3FL was digested with XbaI to excise the 3′-UTR and then ligated to form p4ZD3D3P (Δ3′-UTR), which is designated tetD3Δ-UTR. The 215-nucleotide BC element was excised as a StuI/AvrII fragment from pCRIID3FL and cloned into theEcoRV/XbaI sites of pUHD4Zeo to form p4zBC. The p4zBC transcript has a predicted size of about 250 nucleotides and is terminated by SV40 polyadenylylation signals. Transcribed from the tetO/human cytomegalovirus major immediate early TATA box (HCMV) chimeric promoter, the p4zBC transcript is presumed to be capped. We have not been able to quantify this transcript, and we have not ascertained that the transcript is either capped or polyadenylylated. p5zβGal was made by ligating theNotI/AhaIII fragment of cytomegalovirus/βGal (Promega) into the NotI/EcoRV sites of pUHDZeo5. This expression vector is identified herein as β-gal.p5BGD3UTR was made by inserting theSpeI/XbaI fragment from pCRII/D33-UTR (containing the 3′-UTR) into the AvrII site ofp5zβGal. This transgene is called β-galD3UTR herein. P1798 cells were initially transfected to stably express the tetracycline transactivator from pUHD15–1neo (27.Gossen M. Bujard H. Proc. Natl. Acad. Sci. U. S. A. 1992; 89: 5547-5551Crossref PubMed Scopus (4268) Google Scholar). G418-resistant clones were screened for transient expression of pUHD13–3 (tet/luciferase). Secondary transfections with appropriate expression vectors were carried out. Some of the expression vector harbor linked zeocin resistance genes; others were co-transfected with pZeo. Stable, zeocin-resistant clones were selected and analyzed for expression of the appropriate transgenes. Cyclin D3 mRNA is very stable in P1798 cells treated with actinomycin D. As shown in Fig.1 A, there was little or no decrease in CcnD3 mRNA within the first 4 h after addition of actinomycin D to mid-log phase cells. In other experiments, we observed no decrease in CcnD3 mRNA after 24 h in the presence of actinomycin D (5.Reisman D. Thompson E.A. Mol. Endocrinol. 1995; 9: 1500-1509PubMed Google Scholar), which leads us to suspect that actinomycin D affects the turnover of CcnD3 mRNA. Data that will be presented below are consistent with this supposition. Addition of dexamethasone accelerated CcnD3 mRNA degradation, to the extent that 50% decrease in CcnD3 mRNA abundance was observed within about 2 h after addition of actinomycin D to cells that had been exposed to dexamethasone for 2 h (Fig. 1 A). These data suggest that glucocorticoids regulate the rate of degradation of CcnD3 mRNA. The great majority of mRNA stability control mechanisms reported to date have been found in the 3′-UTRs of the respective transcripts. Consequently, our first objective was to analyze the turnover of cyclin D3 mRNA derivatives with and without the 3′-UTR. Tetracycline-repressible expression systems were developed to permit analysis of mRNA abundance without resort to nonspecific inhibitors of transcription, such as actinomycin D. The development of the appropriate cell lines is described under “Materials and Methods,” and Fig. 1 B shows the results obtained upon analysis of two cell lines that stably express full-length cyclin D3 mRNA (tetD3FL) or cyclin D3 mRNA from which the 3′-UTR had been deleted (tetD3Δ-UTR). The properties of the two clonal cell lines used in these experiments are representative of at least four independently isolated clones that express either full-length or truncated D3 transgenes from the tet promoter. A ribonuclease protection assay was developed to discriminate between the transgenic tetD3 transcripts and the endogenous CcnD3 mRNA. Expression of both full-length and 3′-truncated tetD3 transcripts was repressed upon addition of tetracycline (Fig. 1 B, lanes 2 and 5), whereas expression of the endogenous CcnD3 mRNA was not affected. The cell lines illustrated in Fig. 1 B were used to measure the stability of full-length and truncated tetD3 transcripts, as shown in Fig. 1 C. The abundance of full-length tetD3 transcripts decreased 50% within about 2 h after addition of tetracycline (filled circles). In contrast, the tetD3 derivative that was deleted of the 3′-UTR exhibited a slower rate of degradation (filled squares). We estimate a T1/2 of degradation of about 12 h for the 3′-deleted transcript. These data suggest that there are elements within the 3′-UTR of cyclin D3 mRNA that control basal stability of the transcript. The rapid turnover of full-length transgenic tetD3 mRNA, (t 12 of about 2 h) is inconsistent with the data shown in Fig. 1 A, in which the abundance of endogenous CcnD3 mRNA remained relatively constant for several hours after addition of actinomycin D. These observations suggested that actinomycin D might interfere with degradation of cyclin D3 mRNA. The rate of degradation of transgenic tetD3FL mRNA was measured in cells that had been treated with actinomycin D for 2 h prior to addition of tetracycline. As shown in Fig. 1 C(diamonds), no significant decrease in the abundance of the full-length tetD3 transcript was observed within 4 h after addition of tetracycline to actinomycin d-treated cells. This apparent stability should be contrasted with the rapid rate of degradation that occurs upon addition of tetracycline to tetD3FL-expressing cultures in the absence of actinomycin D (Fig.1 C, circles). We conclude that actinomycin D has an effect on the basal stability of cyclin D3 mRNA, but this phenomenon has not been pursued. Cells that express tetD3 derivatives were treated for 24 h with dexamethasone, as shown in Fig. 1 B. Neither the full-length tetD3 transcript (lane 3) nor the 3′-truncated tetD3Δ-UTR derivative (lane 6) was regulated by glucocorticoids. The stability of both full-length and tetD3Δ-UTR transcripts was determined by measuring the abundance of the RNAs in dexamethasone-treated cells after addition of tetracycline. As shown in Fig. 1 C, dexamethasone did not affect the stability of transgenic tetD3 transcripts (open circles and open squares). This result was unanticipated, because we had previously shown that glucocorticoids do not inhibit transcription of the cyclin D3 gene (5.Reisman D. Thompson E.A. Mol. Endocrinol. 1995; 9: 1500-1509PubMed Google Scholar) and that endogenous CcnD3 mRNA is degraded more rapidly in dexamethasone-treated cells (Fig. 1 A). We were also surprised to note that dexamethasone failed to inhibit the expression of the endogenous CcnD3 mRNA in cells that were transfected with the full-length tetD3 cDNA expression vectors (Fig. 1 B, lane 3), whereas CcnD3 mRNA was inhibited when dexamethasone was added to cells that express the 3′-truncated derivative (lane 6). The data shown in Fig. 1 B suggest that overexpression of the cyclin D3 3′-UTR from a transgene (tetD3FL) interfered with glucocorticoid regulation of the endogenous CcnD3 mRNA expression. The abundance of the tetD3FL transcript, when maximally derepressed by withdrawal of tetracycline, never exceeded five times that of the endogenous CcnD3 mRNA (Fig. 1 B), suggesting that the effect prevailed at relatively low concentrations of the transgenic 3′-UTR. We concluded that the transgenic D3 3′-UTR was acting in trans to block glucocorticoid-mediated destabilization of CcnD3 mRNA. To test this prediction, a number of P1798 cell lines were generated that stably expressed the β-galactosidase gene, with or without the cyclin D3 3′-UTR, under the control of a tetracycline-repressible promoter. All of the clones that were subsequently analyzed for cyclin D3 expression (six clones) exhibited essentially the same properties as the two clones for which β-galactosidase activity data are shown in Fig. 2 A. Four such clones were analyzed, and all showed essentially the same properties. Theopen bars in Fig. 2 A illustrate the properties of those clones that express β-gal, whereas the filled barsillustrate the properties of those cells that express β-galactosidase fuse to the D3 3′-UTR chimera (β-galD3UTR). Initially, we noted that the level of expression of the β-galD3UTR chimeric gene was consistently lower than that of authentic β-gal. (Note that the scales are different in Fig. 2 A.) We have shown that the D3 3′-UTR increases the turnover of tetD3 transgenic RNAs, and we suspect that this reproducible difference in β-galactosidase activity may be due to an increased turnover of β-galD3UTR transcripts; however, the rates of degradation of the β-galactosidase mRNAs have not been measured. We also observed that dexamethasone had no effect on β-galactosidase expression, irrespective of the presence or absence of the D3 3′-UTR. This observation is consistent with the data shown in Fig. 1 B, which indicate that the presence or absence of the D3 3′-UTR did not affect glucocorticoid regulation of the tetD3 transgenes. Finally, tetracycline inhibition of β-galactosidase activity was >95% for the cell line illustrated in Fig. 2 Aand for all cell lines used in the experiments described below. Because tetracycline regulates the transcription of the transgene, it is reasonable to assume that the decrease in β-galactosidase activity that can be observed upon addition of tetracycline is associated with and attributable to a corresponding decrease in β-galactosidase mRNA. Glucocorticoid regulation of CcnD3 mRNA was measured in several clones that express β-gal and β-galD3UTR transgenes, as shown in Fig. 2. Fig. 2, B and C, illustrates the properties of two clones that express authentic β-gal mRNA. The abundance of CcnD3 mRNA decreased rapidly upon addition of dexamethasone, indicating that expression of β-galactosidase had no effect on glucocorticoid-mediated inhibition of CcnD3 mRNA abundance (filled circles). Conversely, little or no decrease in CcnD3 mRNA was observed when dexamethasone was added to βgalD3UTR clones, as shown in Fig. 2, D–G (filled circles). These results indicate that cells that express the β-galactosidase/cyclin D3 3′-UTR chimera were resistant to glucocorticoid inhibition of cyclin D3 expression. The abundance of CcnD3 mRNA decreased rapidly when dexamethasone was added to βgalD3UTR cells that had been treated with tetracycline (Fig. 2,D–G, open circles), indicating that glucocorticoid inhibition of CcnD3 mRNA expression was restored when transcription of the β-galD3UTR chimeric transgene was repressed. Tetracycline had no effect upon glucocorticoid inhibition of cyclin D3 expression in cells that express β-galactosidase without the CcnD3 3′-UTR (Fig. 2,B and C, open circles). The data shown in Figs. 1 and 2 suggest that there are sequences within the D3 3′-UTR that influence glucocorticoid regulation of the stability of CcnD3 mRNA. These elements appear to act in trans, to the extent that overexpression of the 3′-UTR from a transgene (tetD3FL or β-galD3UTR) interferes with regulation of the endogenous CcnD3 mRNA. The data are consistent with the hypothesis that sequences within the transgenic 3′-UTR are acting as molecular decoys, titrating cellular proteins that would otherwise interact with the endogenous transcript to affect glucocorticoid regulation of CcnD3 mRNA stability. RNA mobility shift assays were used to identify potential RNA-protein interactions within the CcnD3 3′-UTR. The full-length CcnD3 3′-UTR was transcribed in vitro in the presence of α-32P-labeled nucleoside triphosphates. The full-length, labeled 3′-UTR transcript, ∼1000 bases long, was incubated with S-100 cytoplasmic extracts, which were prepared from exponentially growing P1798 cells. Ribonuclease T1 was added to degrade those regions of the transcript that were not protected by stable RNA-protein complexes. The T1 resistance complexes were then resolved by electrophoresis on nondenaturing polyacrylamide gels, as shown in Fig.3. Lane 1 of Fig.3 A contains a ribonuclease T1-resistant RNA-protein complex (arrow). Formation of this complex was precluded by addition of 120 ng of unlabeled CcnD3 3′-UTR (lane 2) However, addition of 120 ng of unlabeled E. coli β-galactosidase mRNA failed to compete for binding of S100 proteins to the CcnD3 3′-UTR (lane 3). Fig. 3 B shows the results of an experiment in which labeled CcnD3 3′-UTR was incubated with increasing amounts of S100 protein. The results indicate that formation of the gel shift entity requires the presence of S100 protein (lane 1) and is not simply a ribonuclease T1-resistant fragment. The intensity of the shifted band increased as a function of the protein concentration in the binding reaction (lanes 2 and 3). Furthermore, the shifted band was abolished when the ribonuclease T1-resistant complex was treated with proteinase K before electrophoresis (lane 4), indicating that the labeled band that we observed results from formation of an RNA-protein complex. An RNA mobility shift assay was performed, and the RNA-protein complex was excised from the gel. The labeled RNA was isolated and digested to completion with ribonuclease T1. The products of ribonuclease T1 digestion were s" @default.
• W2005703352 created "2016-06-24" @default.
• W2005703352 creator A5012215691 @default.
• W2005703352 creator A5051842779 @default.
• W2005703352 creator A5053729027 @default.
• W2005703352 date "2000-07-01" @default.
• W2005703352 modified "2023-10-16" @default.
• W2005703352 title "Glucocorticoid-mediated Destabilization of Cyclin D3 mRNA Involves RNA-Protein Interactions in the 3′-Untranslated Region of the mRNA" @default.
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