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• W2023561018 abstract "Two general pathways of mRNA decay have been characterized in yeast. Both start with deadenylation. The major pathway then proceeds via cap hydrolysis and 5′-exonucleolytic degradation whereas the minor pathway consists of 3′-exonucleolytic decay followed by hydrolysis of the remaining cap structure. In higher eukaryotes, these pathways of mRNA decay are believed to be conserved but have not been well characterized. We have investigated the decay of the hsp70 mRNA in Drosophila Schneider cells. As shown by the use of reporter constructs, rapid deadenylation of this mRNA is directed by its 3′-untranslated region. The main deadenylase is the CCR4·NOT complex; the PAN nuclease makes a lesser contribution. Heat shock prevents deadenylation not only of the hsp70 but also of bulk mRNA. A completely deadenylated capped hsp70 mRNA decay intermediate accumulates transiently and is degraded via cap hydrolysis and 5′-decay. Thus, decapping is a slow step in the degradation pathway. Cap hydrolysis is also inhibited during heat shock. Degradation of reporter RNAs from the 3′-end became detectable only upon inhibition of 5′-decay and thus represents a minor decay pathway. Because two reporter RNAs and at least two endogenous mRNAs were degraded primarily from the 5′-end with cap hydrolysis as a slow step, this pathway appears to be of general importance for mRNA decay in Drosophila. Two general pathways of mRNA decay have been characterized in yeast. Both start with deadenylation. The major pathway then proceeds via cap hydrolysis and 5′-exonucleolytic degradation whereas the minor pathway consists of 3′-exonucleolytic decay followed by hydrolysis of the remaining cap structure. In higher eukaryotes, these pathways of mRNA decay are believed to be conserved but have not been well characterized. We have investigated the decay of the hsp70 mRNA in Drosophila Schneider cells. As shown by the use of reporter constructs, rapid deadenylation of this mRNA is directed by its 3′-untranslated region. The main deadenylase is the CCR4·NOT complex; the PAN nuclease makes a lesser contribution. Heat shock prevents deadenylation not only of the hsp70 but also of bulk mRNA. A completely deadenylated capped hsp70 mRNA decay intermediate accumulates transiently and is degraded via cap hydrolysis and 5′-decay. Thus, decapping is a slow step in the degradation pathway. Cap hydrolysis is also inhibited during heat shock. Degradation of reporter RNAs from the 3′-end became detectable only upon inhibition of 5′-decay and thus represents a minor decay pathway. Because two reporter RNAs and at least two endogenous mRNAs were degraded primarily from the 5′-end with cap hydrolysis as a slow step, this pathway appears to be of general importance for mRNA decay in Drosophila. A characteristic feature of mRNA is its rapid turnover, permitting a continuous qualitative and quantitative adjustment of protein synthesis according to physiological needs. The pathways of eukaryotic mRNA degradation have been characterized mostly in Saccharomyces cerevisiae (1Parker R. Song H. Nat. Struct. Mol. Biol. 2004; 11: 121-127Crossref PubMed Scopus (644) Google Scholar, 2Meyer S. Temme C. Wahle E. Crit. Rev. Biochem. Mol. Biol. 2004; 39: 197-216Crossref PubMed Scopus (287) Google Scholar). The decay of all mRNAs examined in these cells starts with deadenylation, i.e. exonucleolytic shortening of the poly(A) tail. The second step of decay does not occur until the poly(A) tail has been shortened to about a dozen nucleotides (3Decker C.J. Parker R. Genes Dev. 1993; 7: 1632-1643Crossref PubMed Scopus (517) Google Scholar). In the major pathway, this second step is hydrolysis of the 5′-cap (4Hsu C.L. Stevens A. Mol. Cell Biol. 1993; 13: 4826-4835Crossref PubMed Scopus (322) Google Scholar, 5Muhlrad D. Decker C.J. Parker R. Genes Dev. 1994; 8: 855-866Crossref PubMed Scopus (408) Google Scholar, 6Muhlrad D. Decker C.J. Parker R. Mol. Cell Biol. 1995; 15: 2145-2156Crossref PubMed Scopus (262) Google Scholar). Cap hydrolysis results in free m7GDP and a 5′-monophosphate left on the mRNA and is catalyzed by the Dcp2p subunit of the Dcp1p·Dcp2p heterodimer (7Cougot N. van Dijk E. Babajko S. Séraphin B. Trends Biochem. Sci. 2004; 29: 436-444Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar, 8Coller J. Parker R. Annu. Rev. Biochem. 2004; 73: 861-890Crossref PubMed Scopus (392) Google Scholar). In vivo, the activity of the decapping enzyme is enhanced by several other proteins (8Coller J. Parker R. Annu. Rev. Biochem. 2004; 73: 861-890Crossref PubMed Scopus (392) Google Scholar). The decapped RNA is degraded by the 5′-exonuclease Xrn1p (4Hsu C.L. Stevens A. Mol. Cell Biol. 1993; 13: 4826-4835Crossref PubMed Scopus (322) Google Scholar, 5Muhlrad D. Decker C.J. Parker R. Genes Dev. 1994; 8: 855-866Crossref PubMed Scopus (408) Google Scholar). In a minor pathway, the deadenylated RNA is degraded from the 3′-end by the exosome (9Jacobs Anderson J.S. Parker R. EMBO J. 1998; 17: 1497-1506Crossref PubMed Scopus (518) Google Scholar) before the remaining oligonucleotide is decapped by the enzyme Dcs1p, liberating m7GMP (10Wang Z. Kiledjian M. Cell. 2001; 107: 751-752Abstract Full Text Full Text PDF PubMed Scopus (206) Google Scholar, 11Liu H. Rodgers N.D. Jiao X. Kiledjian M. EMBO J. 2002; 21: 4699-4708Crossref PubMed Scopus (203) Google Scholar).The deadenylation-dependent decapping pathway of mRNA decay is in a fundamental competition with translation (8Coller J. Parker R. Annu. Rev. Biochem. 2004; 73: 861-890Crossref PubMed Scopus (392) Google Scholar). Two proteins required for cap hydrolysis in vivo, Pat1p and the RNA helicase Dhh1p, act by shifting the balance in favor of cap hydrolysis at the expense of translation (12Coller J. Parker R. Cell. 2005; 122: 875-886Abstract Full Text Full Text PDF PubMed Scopus (477) Google Scholar). Because the poly(A) tail is involved in translation initiation via the poly(A)-binding protein, deadenylation can be seen as promoting decapping and decay by removing the mRNA from the translated pool (13Tharun S. Parker R. Mol. Cell. 2001; 8: 1075-1083Abstract Full Text Full Text PDF PubMed Scopus (185) Google Scholar). In fact, the inhibitor of decapping is the poly(A)-binding protein, not the poly(A) tail per se (8Coller J. Parker R. Annu. Rev. Biochem. 2004; 73: 861-890Crossref PubMed Scopus (392) Google Scholar, 14Caponigro G. Parker R. Genes Dev. 1995; 9: 2421-2432Crossref PubMed Scopus (230) Google Scholar).In animal cells, the decay of most mRNAs analyzed is also initiated by deadenylation (15Wilson T. Treisman R. Nature. 1988; 336: 396-399Crossref PubMed Scopus (504) Google Scholar, 16Shyu A.-B. Belasco J.G. Greenberg M.E. Genes Dev. 1991; 5: 221-231Crossref PubMed Scopus (400) Google Scholar, 17Yamashita A. Chang T.-C. Yamashita Y. Zhu W. Zhong Z. Chen C.-Y. A. Shyu A.-B. Nat. Struct. Mol. Biol. 2005; 12: 1054-1063Crossref PubMed Scopus (334) Google Scholar). Subsequent decay of the deadenylated message is thought to proceed via the two pathways analyzed in yeast, mostly because the relevant proteins are conserved, but experimental data on the later steps of mRNA decay are relatively scarce. Based mainly on in vitro evidence, several groups have argued that the exosomal 3′-decay pathway may be the predominant mode of decay of unstable mammalian mRNAs (10Wang Z. Kiledjian M. Cell. 2001; 107: 751-752Abstract Full Text Full Text PDF PubMed Scopus (206) Google Scholar, 18Chen C.-Y. Gherzi R. Ong S.-E. Chan E.L. Raijmakers R. Pruijn G.J.M. Stoecklin G. Moroni C. Mann M. Karin M. Cell. 2001; 107: 451-464Abstract Full Text Full Text PDF PubMed Scopus (728) Google Scholar, 19Mukherjee D. Gao M. O'Connor J.P. Raijmakers R. Pruijn G. Lutz C.S. Wilusz J. EMBO J. 2002; 21: 165-174Crossref PubMed Scopus (307) Google Scholar). In vivo studies have supported a role of the 3′–5′ pathway (20Brewer G. J. Biol. Chem. 1999; 274: 16174-16179Abstract Full Text Full Text PDF PubMed Scopus (35) Google Scholar, 21Chou C.-F. Mulky A. Maitra S. Lin W.-J. Gherzi R. Kappes J. Chen C.-Y. Mol. Cell. Biol. 2006; 26: 3695-3706Crossref PubMed Scopus (101) Google Scholar, 22Stoecklin G. Mayo T. Anderson P. EMBO Rep. 2006; 7: 72-77Crossref PubMed Scopus (189) Google Scholar, 23Franks T.M. Lykke-Andersen J. Genes Dev. 2007; 21: 719-735Crossref PubMed Scopus (194) Google Scholar), but evidence for the existence of the 5′–3′ pathway, sometimes acting on the same RNAs, has also been provided (17Yamashita A. Chang T.-C. Yamashita Y. Zhu W. Zhong Z. Chen C.-Y. A. Shyu A.-B. Nat. Struct. Mol. Biol. 2005; 12: 1054-1063Crossref PubMed Scopus (334) Google Scholar, 21Chou C.-F. Mulky A. Maitra S. Lin W.-J. Gherzi R. Kappes J. Chen C.-Y. Mol. Cell. Biol. 2006; 26: 3695-3706Crossref PubMed Scopus (101) Google Scholar, 22Stoecklin G. Mayo T. Anderson P. EMBO Rep. 2006; 7: 72-77Crossref PubMed Scopus (189) Google Scholar, 23Franks T.M. Lykke-Andersen J. Genes Dev. 2007; 21: 719-735Crossref PubMed Scopus (194) Google Scholar, 24Couttet P. Fromont-Racine M. Steel D. Pictet R. Grange T. Proc. Natl. Acad. Sci. U. S. A. 1997; 94: 5628-5633Crossref PubMed Scopus (186) Google Scholar, 25Fenger-Gron M. Fillman C. Norrild B. Lykke-Andersen J. Mol. Cell. 2005; 20: 905-915Abstract Full Text Full Text PDF PubMed Scopus (371) Google Scholar). Conservation of the 5′–3′ decay pathway is also supported by the co-localization of the responsible proteins in distinct cytoplasmic structures, P bodies or GW bodies, in both yeast and mammalian cells (26Eulalio A. Behm-Ansmant I. Izaurralde E. Nat Rev. Mol. Cell Biol. 2007; 8: 9-22Crossref PubMed Scopus (741) Google Scholar). Furthermore, microRNA (miRNA) 4The abbreviations used are: miRNA, microRNA; UTR, untranslated region; ARE, AU-rich element; dsRNA, double-stranded RNA; Ab, antibody; nt, nucleotide; RNP, ribonucleoprotein.4The abbreviations used are: miRNA, microRNA; UTR, untranslated region; ARE, AU-rich element; dsRNA, double-stranded RNA; Ab, antibody; nt, nucleotide; RNP, ribonucleoprotein.-induced mRNA destabilization in Drosophila Schneider cells requires Dcp1·Dcp2-dependent decapping (27Behm-Ansmant I. Rehwinkel J. Doerks T. Stark A. Bork P. Izaurralde E. Genes Dev. 2006; 20: 1885-1898Crossref PubMed Scopus (721) Google Scholar). However, the relative contributions of the two decay pathways to overall mRNA decay remain unknown, and decay intermediates have been analyzed only to a limited extent.Deadenylation is the rate-limiting step in the decay of most mRNAs. Several arguments support this view. First, deadenylation occurs by continuous shortening of the poly(A) tail throughout the lifetime of the RNA; the process can be easily followed in vivo even for unstable RNAs. However, once deadenylation has proceeded beyond a critical limit, the RNA usually disappears rapidly without detectable intermediates, showing that all subsequent steps are fast (28Chen C.-Y. Shyu A.-B. Trends Biochem. Sci. 1995; 20: 465-470Abstract Full Text PDF PubMed Scopus (1666) Google Scholar). Second, unstable RNAs are deadenylated faster than stable RNAs, and sequence manipulations affecting the overall rate of decay change the rate of deadenylation in a similar manner (8Coller J. Parker R. Annu. Rev. Biochem. 2004; 73: 861-890Crossref PubMed Scopus (392) Google Scholar, 15Wilson T. Treisman R. Nature. 1988; 336: 396-399Crossref PubMed Scopus (504) Google Scholar, 16Shyu A.-B. Belasco J.G. Greenberg M.E. Genes Dev. 1991; 5: 221-231Crossref PubMed Scopus (400) Google Scholar, 29Muhlrad D. Parker R. Genes Dev. 1992; 6: 2100-2111Crossref PubMed Scopus (160) Google Scholar, 30Chen C.-Y. A. Chen T.-M. Shyu A.-B. Mol. Cell. Biol. 1994; 14: 416-426Crossref PubMed Google Scholar, 31Lagnado C.A. Brown C.Y. Goodall G.J. Mol. Cell. Biol. 1994; 14: 7984-7995Crossref PubMed Scopus (310) Google Scholar, 32Zubiaga A.M. Belasco J.G. Greenberg M.E. Mol. Cell. Biol. 1995; 15: 2219-2230Crossref PubMed Scopus (471) Google Scholar). Third, when a normally unstable RNA is stabilized because of a physiological stimulus, the rate of its deadenylation is reduced (33Dean J.L.E. Sarsfield S.J. Tsounakou E. Saklatvala J. J. Biol. Chem. 2003; 278: 39470-39476Abstract Full Text Full Text PDF PubMed Scopus (87) Google Scholar, 34Winzen R. Gowrishankar G. Bollig F. Redich N. Resch K. Holtmann H. Mol. Cell. Biol. 2004; 24: 4835-4847Crossref PubMed Scopus (114) Google Scholar). Fourth, kinetic modeling suggests that changes in the rate of deadenylation have the greatest influence on the overall stability of the RNA (35Cao D. Parker R. RNA (N. Y.). 2001; 7: 1192-1212Crossref PubMed Scopus (103) Google Scholar). For some RNAs cap hydrolysis is a slow step, leading to the accumulation of a fully deadenylated, capped decay intermediate (3Decker C.J. Parker R. Genes Dev. 1993; 7: 1632-1643Crossref PubMed Scopus (517) Google Scholar). Slow decapping is often associated with slow deadenylation (8Coller J. Parker R. Annu. Rev. Biochem. 2004; 73: 861-890Crossref PubMed Scopus (392) Google Scholar).The main enzyme responsible for deadenylation in yeast as well as in mammals and in Drosophila is the CCR4·NOT complex (17Yamashita A. Chang T.-C. Yamashita Y. Zhu W. Zhong Z. Chen C.-Y. A. Shyu A.-B. Nat. Struct. Mol. Biol. 2005; 12: 1054-1063Crossref PubMed Scopus (334) Google Scholar, 36Tucker M. Valencia-Sanchez M.A. Staples R.R. Chen J. Denis C.L. Parker R. Cell. 2001; 104: 377-386Abstract Full Text Full Text PDF PubMed Scopus (466) Google Scholar, 37Daugeron M.-C. Mauxion F. Seraphin B. Nucleic Acids Res. 2001; 29: 2448-2455Crossref PubMed Scopus (164) Google Scholar, 38Temme C. Zaessinger S. Simonelig M. Wahle E. EMBO J. 2004; 23: 2862-2871Crossref PubMed Scopus (167) Google Scholar). A second universally conserved poly(A)-degrading 3′-exonuclease is the PAN2·PAN3 heterodimer, PAN for short (39Brown C.E. Tarun S.Z. Boeck R. Sachs A.B. Mol. Cell. Biol. 1996; 16: 5744-5753Crossref PubMed Scopus (129) Google Scholar, 40Boeck R. Tarun S. Rieger M. Deardorff J.A. Müller-Auer S. Sachs A.B. J. Biol. Chem. 1996; 271: 432-438Abstract Full Text Full Text PDF PubMed Scopus (146) Google Scholar, 41Uchida N. Hoshino S. Katada T. J. Biol. Chem. 2004; 279: 1383-1391Abstract Full Text Full Text PDF PubMed Scopus (106) Google Scholar). In yeast, this enzyme catalyzes the residual deadenylation observed upon inactivation of the CCR4·NOT complex (36Tucker M. Valencia-Sanchez M.A. Staples R.R. Chen J. Denis C.L. Parker R. Cell. 2001; 104: 377-386Abstract Full Text Full Text PDF PubMed Scopus (466) Google Scholar). Under normal conditions, PAN is thought to act before the CCR4·NOT complex, catalyzing a rapid initial shortening of the poly(A) tail (17Yamashita A. Chang T.-C. Yamashita Y. Zhu W. Zhong Z. Chen C.-Y. A. Shyu A.-B. Nat. Struct. Mol. Biol. 2005; 12: 1054-1063Crossref PubMed Scopus (334) Google Scholar, 42Brown C.E. Sachs A.B. Mol. Cell. Biol. 1998; 18: 6548-6559Crossref PubMed Scopus (180) Google Scholar). A third poly(A)-degrading enzyme, the homodimeric PARN (43Körner C.G. Wormington M. Muckenthaler M. Schneider S. Dehlin E. Wahle E. EMBO J. 1998; 17: 5427-5437Crossref PubMed Scopus (202) Google Scholar, 44Martinez J. Ren Y.-G. Thuresson A.-C. Hellman U. Astrom J. Virtanen A. J. Biol. Chem. 2000; 275: 24222-24230Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar, 45Wu M. Reuter M. Lilie H. Liu Y. Wahle E. Song H. EMBO J. 2005; 24: 4082-4093Crossref PubMed Scopus (91) Google Scholar), is conserved in most eukaryotes, but not in yeast and in Drosophila. PARN has been implicated in the decay of several unstable RNAs (21Chou C.-F. Mulky A. Maitra S. Lin W.-J. Gherzi R. Kappes J. Chen C.-Y. Mol. Cell. Biol. 2006; 26: 3695-3706Crossref PubMed Scopus (101) Google Scholar, 46Lai W.S. Kennington E.A. Blackshear P.J. Mol. Cell. Biol. 2003; 23: 3798-3812Crossref PubMed Scopus (194) Google Scholar), but its general role in mRNA turnover remains poorly defined (17Yamashita A. Chang T.-C. Yamashita Y. Zhu W. Zhong Z. Chen C.-Y. A. Shyu A.-B. Nat. Struct. Mol. Biol. 2005; 12: 1054-1063Crossref PubMed Scopus (334) Google Scholar).RNA elements promoting rapid deadenylation and decay are often located in the 3′-UTR, and the so-called AU-rich elements (AREs) are the best-studied class among them (28Chen C.-Y. Shyu A.-B. Trends Biochem. Sci. 1995; 20: 465-470Abstract Full Text PDF PubMed Scopus (1666) Google Scholar). Destabilizing sequences are bound by specific proteins, which can directly recruit deadenylases and other decay enzymes (21Chou C.-F. Mulky A. Maitra S. Lin W.-J. Gherzi R. Kappes J. Chen C.-Y. Mol. Cell. Biol. 2006; 26: 3695-3706Crossref PubMed Scopus (101) Google Scholar, 25Fenger-Gron M. Fillman C. Norrild B. Lykke-Andersen J. Mol. Cell. 2005; 20: 905-915Abstract Full Text Full Text PDF PubMed Scopus (371) Google Scholar, 47Lykke-Andersen J. Wagner E. Genes Dev. 2005; 19: 351-361Crossref PubMed Scopus (364) Google Scholar, 48Semotok J.L. Cooperstock R.L. Pinder B.D. Vari H.K. Lipshitz H.D. Smibert C.A. Curr. Biol. 2005; 15: 284-294Abstract Full Text Full Text PDF PubMed Scopus (183) Google Scholar, 49Goldstrohm A.C. Hook B.A. Seay D.J. Wickens M. Nat. Struct. Mol. Biol. 2006; 13: 533-539Crossref PubMed Scopus (243) Google Scholar, 50Zaessinger S. Busseau I. Simonelig M. Development. 2006; 133: 4573-4583Crossref PubMed Scopus (137) Google Scholar). Recently, miRNAs have also been found to be able to induce mRNA deadenylation (27Behm-Ansmant I. Rehwinkel J. Doerks T. Stark A. Bork P. Izaurralde E. Genes Dev. 2006; 20: 1885-1898Crossref PubMed Scopus (721) Google Scholar, 51Jing Q. Huang S. Guth S. Zarubin T. Motoyama A. Chen J. Di Padova F. Lin S.-C. Gram H. Han J. Cell. 2005; 120: 623-634Abstract Full Text Full Text PDF PubMed Scopus (702) Google Scholar, 52Wu L. Fan J. Belasco J.G. Proc. Natl. Acad. Sci. U. S. A. 2006; 103: 4034-4039Crossref PubMed Scopus (902) Google Scholar, 53Giraldez A.J. Mishima Y. Rihel J. Grocock R.J. Van Dongen S. Inoue K. Enright A.J. Schier A.F. Science. 2006; 312: 75-79Crossref PubMed Scopus (1217) Google Scholar).The hsp70 mRNA of Drosophila is very unstable at normal growth temperatures with a functional half-life of 15–30 min suggested by indirect measurements (54Petersen R. Lindquist S. Gene (Amst.). 1988; 72: 161-168Crossref PubMed Scopus (79) Google Scholar). Rapid degradation of the hsp70 message requires its 3′-UTR (55Petersen R.B. Lindquist S. Cell Regulation. 1989; 1: 135-149Crossref PubMed Scopus (101) Google Scholar). Upon heat shock, the RNA is stabilized at least 10-fold (54Petersen R. Lindquist S. Gene (Amst.). 1988; 72: 161-168Crossref PubMed Scopus (79) Google Scholar). This contributes to the rapid and massive (more than 1000-fold (56Lindquist S. Petersen R. Enzyme. 1990; 44: 147-166Crossref PubMed Scopus (67) Google Scholar)) induction of HSP70 protein synthesis. During recovery from heat shock, the normal instability of the message is restored, and production of HSP70 protein ceases (54Petersen R. Lindquist S. Gene (Amst.). 1988; 72: 161-168Crossref PubMed Scopus (79) Google Scholar, 55Petersen R.B. Lindquist S. Cell Regulation. 1989; 1: 135-149Crossref PubMed Scopus (101) Google Scholar). The first step in hsp70 mRNA degradation during recovery is its deadenylation catalyzed by the CCR4·NOT complex (38Temme C. Zaessinger S. Simonelig M. Wahle E. EMBO J. 2004; 23: 2862-2871Crossref PubMed Scopus (167) Google Scholar, 57Dellavalle R.P. Petersen R. Lindquist S. Mol. Cell. Biol. 1994; 14: 3646-3659Crossref PubMed Google Scholar). An unusual, transient but pronounced accumulation of a completely deadenylated RNA species is also observed both during heat shock and recovery. Two observations suggest that deadenylation is inhibited by heat shock: First, during heat shock, the fraction of polyadenylated hsp70 RNA increases with increasing temperature. Second, deadenylation is faster after a mild heat shock than after a severe heat shock, and deadenylation during recovery from a severe heat shock is faster if the cells have been preconditioned by a mild heat shock (57Dellavalle R.P. Petersen R. Lindquist S. Mol. Cell. Biol. 1994; 14: 3646-3659Crossref PubMed Google Scholar). However, the temperature dependence of deadenylation and its dependence on 3′-UTR sequences have not been tested directly.We have examined the decay pathway of the hsp70 mRNA in Drosophila Schneider cells. We find that this RNA is degraded via rapid deadenylation controlled by its 3′-UTR, followed by slow cap hydrolysis and 5′-decay. Exosome-dependent 3′-decay plays a minor role in the degradation of the hsp70 mRNA. Both deadenylation and decapping are inhibited during heat shock. The 5′-decay pathway is also important for the degradation of several other mRNAs examined in Schneider cells.EXPERIMENTAL PROCEDURESCell Culture—Drosophila melanogaster Schneider 2 cells were grown as semi-adherent layers at 25 °C in Schneider′s Insect Medium (Sigma) supplemented with 10% heat-inactivated fetal calf serum (Biochrom) and 1% antibiotic/antimycotic mix (Invitrogen). Stably transformed cell lines were generated by calcium phosphate transfection and hygromycin B selection according to procedures suggested by Invitrogen (Drosophila Expression System Version C handbook) and grown in medium containing 300 μg/ml hygromycin B. Expression of reporter gene constructs was induced at 25 °C by addition of 500 μm CuSO4. Transcription was stopped by addition of 5 μg/ml actinomycin D (Sigma).Heat shock was performed by incubation of cells in a temperature-controlled water bath at the temperature indicated. For induction of endogenous hsp70 or hsp83 RNA and analysis of their decay during subsequent recovery, heat shock was at 35.5 or 36 °C, followed by recovery at 25 °C. For the analysis of RNAs during heat shock, 36 or 37 °C were used. No significant differences were observed between these conditions. It should be noted that the metallothionein promoter driving the reporter constructs (see below) is nearly inactive under heat shock conditions. Thus, when actinomycin D was added some time after heat shock, the distribution of RNA decay intermediates at the time of actinomycin D addition reflected a period of net decay preceding the addition of the drug. Apart from that, no difference in RNA decay was seen when actinomycin D was added at the time of heat shock or 30 min later.Plasmids—The reporter plasmids were constructed as follows: The mutated RNP domain of bovine PABPN1 (Met161–Thr258 with the mutations Y175A, K213Q, and F215A) (58Kühn U. Nemeth A. Meyer S. Wahle E. J. Biol. Chem. 2003; 278: 16916-16925Abstract Full Text Full Text PDF PubMed Scopus (63) Google Scholar) was amplified by PCR with the primers GGTACCATGTCCATTGAGGAGAAGAT introducing a KpnI restriction site (underlined) at the beginning of the open reading frame and GTCGACTGTTGTGCTGATGCCTGGT introducing a SalI site (underlined) at the end. The 3′-UTR and additional downstream sequences of the distal hsp70 gene of the 87A7 locus (gene Ab, GenBank™ accession number CG18743 nucleotides 1–243, 1 being the first nucleotide behind the stop codon) were amplified from genomic D. melanogaster DNA (strain Oregon 1) with primers HSP70 Sal (5′-GTCGACTAAGGCCAAAGA-3′), binding at the border of the open reading frame and the 3′-UTR and containing the naturally occuring SalI site immediately 5′ of the stop codon, and HSP70 DSE (5′-GAATTCAGACTTTCAAAAGTCTACAA-3′), introducing a new EcoRI site at the 3′-end. The 3′-UTR plus downstream sequences of the Drosophila adh gene (GenBank™ accession number X78384, nucleotides 2926–3226) were amplified from a plasmid clone (a gift of Saverio Brogna) with the primers ADH sense Sal (5′-GTCGACTAAGAAGTGATAATCCCA-3′), which introduces a SalI restriction site followed by a stop codon at the 5′-end, and ADH DSE (5′-GAATTCTTCTTTCTAACCGCTTTCA-3′), which introduces an EcoRI restriction site at the 3′-end of the PCR product. All fragments were cloned into a pBS-SK vector (Stratagene) opened with SmaI. The RNP fragment was cut out with KpnI/SalI, purified, and ligated into the KpnI/SalI-opened plasmids already containing the HSP70 and ADH 3′-UTRs. The SalI site introduces the additional sequence Val-Thr at the C terminus of the RNP domain. The complete RNP-UTR fragments were then cut out with KpnI and EcoRI and ligated into the KpnI/EcoRI-opened pMT/V5 expression vector (version A, Invitrogen), from which a BclI fragment containing the SV40 poly(A) signal had been deleted. Point mutations in the ARE sequences of the HSP70 reporter plasmid were introduced in two sequential reactions via the QuikChange site-directed mutagenesis method (Stratagene). The sequence was verified after each round of mutagenesis.RNA Interference—dsRNA was prepared by enzymatic synthesis (Ambion MEGAscript® T7 Kit) from DNA templates obtained by PCR. PCR templates were either plasmid clones (Pan2 and Xrn1, the latter a gift of Sarah Newbury) or oligo(dT)-primed cDNA derived from S2 cell RNA (all others). Primer pairs are listed in supplemental Table S1. 45 μg of dsRNA was placed into a 60-mm cell culture dish. Logarithmically growing cells were diluted with serum-free medium to 1 × 106/ml (final serum concentration 1.5% or less), 3 ml were added per dish, incubated for 30 min at 25 °C and then mixed with 6 ml of medium containing 10% serum. After 4 days, cells were split into 1-ml aliquots. When the dsRNA treatment reduced the growth rate, two batches of knock-down cells were combined and split into 2-ml aliquots. If knock-down efficiency was to be confirmed by Western blotting, protein was isolated from one aliquot: Cells were pelleted, resuspended in 75 μl of lysis buffer (50 mm Tris-HCl, pH 8.0, 150 mm NaCl, 1% Triton X-100, 8% glycerol) and frozen in liquid nitrogen and thawed on ice 3 times. RNA was isolated from all other aliquots according to the TRIzol® protocol (Invitrogen). Pelleted RNA was dried at room temperature and dissolved in water by incubation at 65 °C for 20–30 min. Concentrations were determined by spectrophotometry. If no antibody was available to confirm the knock-down, semi-quantitative RT-PCR was done as follows: 1.5 μg of total RNA from knock-down and control cells was reverse-transcribed with 20 pmol dT12 primer and 150 units of murine leukemia virus reverse transcriptase (Promega) in a 25-μl reaction as recommended by the supplier. For the subsequent PCR, 1 μl of this reaction was used as template in a total volume of 80 μl of 1× green GoTaq® Flexi buffer (Promega) containing 1.5 mm MgCl2, 200 μm dNTPs, 1 μm primer (each), and 0.5 units of GoTaq® DNA polymerase (Promega). Primers are listed in supplemental Table S2. After 2 min at 94 °C, cycles consisted of 15 s at 94 °C, 30 s at 55 °C, 60 s at 72 °C. After 29, 31 and 33 cycles, aliquots were taken and analyzed on a 1% agarose gel. Cytochrome c cDNA was amplified as a control.Northern Blots—Probes directed against the PABPN1 RNP domain and the hsp70 3′-UTR were synthesized by transcription of linearized plasmids similar to those described above. All other probes were generated by transcription of templates obtained by RT-PCR (supplemental Table S3). T7 RNA polymerase (Promega) was used in the presence of 10–40 μCi of [α-32P]UTP (Amersham Biosciences) as recommended by the supplier. Total RNA (between 1 and 2.5 μg depending on the experiment) was electrophoretically separated on a 5% polyacrylamide-8.3 m urea gel and transferred to a nylon membrane (Hybond-N, Amersham Biosciences) by semidry electro-blotting in 0.5× Tris borate EDTA. After UV irradiation, hybridization and washing were carried out as described (59Sambrook J. Russell D.W. Molecular Cloning.Vol. I. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York2001: 7.42-7.44Google Scholar), and the blots were exposed to a phosphorimager screen.Cap Analysis—Total RNA (10 or 15 μg depending on the experiment) was denatured at 95 °C for 3 min and placed on ice. Depending on the amount of RNA used, 1 or 1.5 units of Terminator™ 5′-phosphate-dependent exonuclease (Epicentre) was added, the volume was adjusted to 20 or 30 μl, and the mixture was incubated at 30 °C for 2 h under conditions specified by the supplier. The volume was adjusted to 200 μl with water, 10 μg of glycogen was added, the RNA was purified by phenol/chloroform extraction and ethanol precipitation and dissolved either in 10 μl of 1 mm EDTA (for subsequent RNase H digestion) or in formamide-loading buffer.RNase H Digestion—RNase H digestion was performed as described (57Dellavalle R.P. Petersen R. Lindquist S. Mol. Cell. Biol. 1994; 14: 3646-3659Crossref PubMed Google Scholar) with the following modifications: 10 μg of RNA was used for each digestion. Oligonucleotide 1 (5′-GTCCAGAGTAGCCGCCAAATCCTCCGGCC-3′) was used for the hsp70 mRNA, oligonucleotide 2 (5′-TTAATCGACCTCCTCCATGTGGGAAGC-3′) for the hsp83 mRNA, and dT12 instead of dT15 for removal of the poly(A) tail. RNase H was from Promega, and digestion was for 45 min at 37 °C. 10 μg of glycogen was added, RNA was pelleted by ammonium acetate/ethanol precipitation, washed with 70% ethanol, air-dried, and dissolved in formamide-loading buffer.Mapping of the hsp70 Ab Polyadenylation Site—3 μg of total RNA from stable cell lines in which the transcription of the RNP-HSP70 3′-UTR or the RNP-HSP70 3′-UTR ARE- constructs had been induced was used for RT-PCR amplification of the 3′-UTR and poly(A) tail as described earlier (38Temme C. Zaessinger S. Simonelig M. Wahle E. EMBO J. 2004; 23: 2862-2871Crossref PubMed Scopus (167) Google Scholar). Amplification was carried out with the HSP70 Sal primer (see above) and the oligo(dT) anchor primer. The PCR products were purified and sequenced with the Hsp70 Sal primer.Bulk Poly(A) Analysis—The length of bulk poly(A) was analyzed essentially as described previously (38Temme C. Zaessinger S. Simonelig M. Wahle E. EMBO J. 2004; 23: 2862-2871Crossref PubMed Scopus (167) Google Schol" @default.
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• W2023561018 title "Degradation of hsp70 and Other mRNAs in Drosophila via the 5′–3′ Pathway and Its Regulation by Heat Shock" @default.
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