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Differential accumulation of mRNA and interspersed RNA during Xenopus oogenesis and embrypgenesis

Published online by Cambridge University Press:  26 September 2008

Chengyu Liu  and
L. Dennis Smith
Chengyu Liu*
Affiliation:
Depratment of Developmental and Cell Biology and Developmental Biology Center, University of California at Irvina, Irvine, Califonira, USA.
L. Dennis Smith
Affiliation:
Depratment of Developmental and Cell Biology and Developmental Biology Center, University of California at Irvina, Irvine, Califonira, USA.
*
Dr Chengyu Liu, Roche Institue of molecular Biology, 340 Kingsland street, Nutley, NJ 07110, USA. Telephone: (201)-235-4574. (201)-235-2839.
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Summary

Xenopus ooctye cytoplasmic poly(A)+ RNA has been shown to include two major complex classes: mRNA and interspersed RNA. the former is defined by its translatalility, the latter consists of non–translatable repeat–containing transcripts with unknown functions. In this study we compared the accumulation patterns of total mRNA and a subfamily of interspersed RNA, the XR family (McGrew&Richter, 1989, Dev. Biot. 134, 267–70)

Keywords

Interspersed RNAmRNA accumulationOogenesisPolyadenylationXenopus
Type
Article
Information
Zygote , Volume 2 , Issue 4 , November 1994 , pp. 307 - 316
Copyright
Copyright © Cambridge University Press 1994

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References

Andéol, Y., Gusse, M. & Méchali, M.. (1990). Characterization and expression of a Xeno pus ras during oogenesis and development. Dev. Biol. 139, 24–34.CrossRefGoogle Scholar
Anderson, D.M., Richter, J.D., Chamberlin, M.E., Price, D.H., Britten, R.J., Smith, L.D. & Davidson, E.H.. (1982). Sequence organization of poly(A) RNA synthesized and accumulated in lampbrush chromosome stage Xeno pus laevis. J. Mol. Biol. 155, 281–309.CrossRefGoogle Scholar
Bagni, C., Mariottini, P., Annesi, F. & Amaldi, F.. (1990). Structure of Xeno pus laevis ribosomal protein 132 and its expression during development. Nucleic Acids Res. 18, 442–36.CrossRefGoogle Scholar
Baum, E.Z. & Wormington, W.J.. (1985). Coordinate expression of protein genes during Xenopus development. Dev Biol. 111, 488–98.CrossRefGoogle ScholarPubMed
Cabada, M.O., Darnbrough, C., Ford, P.J. & Turner, P.C.. (1977). Differential accumulation of two size classes of poly(A) associated with messenger RNA during oogenesis in Xenopus laevis. Dev. Biol. 57, 427–39.CrossRefGoogle ScholarPubMed
Caizone, F.J., Angerer, R.C. & Gorovsky, M.A.. (1982). Regulation of protein synthesis in Tetrahyma: isolation and characterization of polysomes by gel filtration and precipitation at pH 5.3. Nucleic Acids Res. 10, 2145–61.CrossRefGoogle Scholar
Calzone, F.J., Jacobs, H.T., Flytzanis, C.N., Posakony, J.W. & Davidson, E.H.. (1985). Interspersed maternal RNA of sea urchin and amphibian eggs. In Biology of Fertilization: The Fertilization Response of the Eggs, ed. Mertz, CB & Monroy, A, 3, 347–66. Orlando, Florida: Academic Press.Google Scholar
Caizone, F.J., Lee, J.J., Le, N., Britten, R.J. & Davidson, E.H.. (1988). A long, nontranslatable poly(A) RNA stored in the egg of the sea urchin Strongylocentrotus purpuratus. Genes Dev. 2, 305–18.CrossRefGoogle Scholar
Costantini, F.D., Britten, R.J. & Davidson, E.H.. (1980). Message sequences and short repetitive sequences are interspersed in sea urchin egg poly(A) + RNAs. Nature 287, 111–17.CrossRefGoogle ScholarPubMed
Dale, L., Matthews, C., Tabe, L.. & Colman, A.. (1989). Developmental expression of the protein product of Vgl, a localized maternal mRNA in the frog Xeno pus laevis. EMBO J. 8, 1057–65.CrossRefGoogle Scholar
Darnbrough, C.H. & Ford, P.J.. (1979). Turnover and processing of poly(A) in full-grown oocytes and during progesterone-induced oocyte maturation in Xeno pus laevis. Dev. Biol. 71, 323–40.CrossRefGoogle Scholar
Davidson, E.H.. (1986). Gene Activity in Early Development, 3rd edn. Orlando, Florida: Academic Press.Google Scholar
Deschamps, S., Viel, A., Garrigos, M., Denis, H. & Maire, M.. (1992). mRNP4, a major mRNA-binding protein from Xenopus oocytes is identical to transcription factor FRG Y2. J. Biol. Chem. 267, 13799–802.CrossRefGoogle Scholar
Dolecki, G.J. & Smith, L.D. (1979). Poly(A)+ RNA metabolism during oogenesis in Xenopus laevis. Dev. Biot. 69, 217–36.CrossRefGoogle ScholarPubMed
Dumont, J.N.. (1972). Oogenesis in Xeno pus laevis (Daudin). I. Stages of oocyte development in laboratory maintained animals. I. Morphol. 136, 153–80.Google Scholar
Ford, P.J., Mathieson, T. & Rosbash, M.. (1977). Very long–lived messenger RNA in ovaries of Xeno pus taevis. Dev. Biol. 57, 417–26.CrossRefGoogle Scholar
Fox, C.A. & Wickens, M.. (1990). Poly(A) removal during oocyte maturation: a default reaction selectively prevented by specific sequences in the 3' UTR of certain maternal mRNAs. Genes Dev. 4, 2287–98.CrossRefGoogle ScholarPubMed
Fox, C.A., Sheets, M.D. & Wickens, M.P.. (1989). Poly(A) addition during maturation of frog oocytes distinct nuclear and cytoplasmic activities and regulation by the sequence. UA. Genes Dev. 3, 2151–62.CrossRefGoogle Scholar
Fox, C.A., Sheets, M.D., Wahle, E. & Wickens, M.. (1992). Polyadenylation of maternal mRNA during oocyte maturation: poly(A) addition in vitro requires a regulated RNA binding activity and a poly(A) polymerase. EMBO. 11, 5021–32.CrossRefGoogle Scholar
Golden, L., Schafer, U. & Rosbash, M.. (1980). Accumulation of individual pA + RNAs during oogenesis of Xeno pus laevis. Cell 22 835–44.CrossRefGoogle Scholar
Hyman, L.E. & Wormington, M.W.. (1988). Translational inactivation of ribosomal protein mRNA during Xenopus oocyte maturation, Genes Dev. 2, 598–605.CrossRefGoogle ScholarPubMed
Ibanez, C.F., Hallbook, F. & Persson, H.. (1992). Expression of neurotrophin–4 mRNA during oogenesis in Xenopus laevis. Int J Dev Biol. 36, 239–45.Google ScholarPubMed
Jackson, R.J.. (1993). Cytoplasmic regulation of mRNA function: the importance of the 3 untranslated region. Cell 74, 9–14.CrossRefGoogle ScholarPubMed
Liu, C.. (1992). Accumulation, polyadenylation, storage, and the possible function of interspersed RNA during Xenopus oogenesis and embryogenesis. PhD thesis, University of Califonia Irvine.Google Scholar
Liu, C. & Smith, L.D.. (1994). Evidence that XR family interspersed RNA may regulate translation in Xenopus oocytes. Mol. Reprod. Dev. In press.Google Scholar
McGrew, L.L. & Richter, J.D.. (1989). Xenopus oocyte poly(A) RNAs that hybridize to a cloned interspersed repeat sequence are not translatable. Dev. Biol. 134, 267–70.CrossRefGoogle Scholar
McGrew, L.L. & Richter, J.D.. (1990). Translational control by cytoplasmic polyadenylation during xenopus oocyete matution: characterizatiopn of cis and trans elenents and regulation by cyclin/Mpf. EMBO J. 9, 3743–51.CrossRefGoogle ScholarPubMed
McGrew, L.L., Dworkin–Rastl, E., Dworkin, M.B. & Richter, J.D.. (1989). Poly(A) elongation during Xenopus oocyte maturation is required for translational recruitment and is mediated by a short sequence element. Genes Dev. 3,803–15.CrossRefGoogle ScholarPubMed
Melton, D.A.. (1987). Translocation of a localized maternal mRNA to the vegetal pole of Xenopus oocytes. Nature 328, 80–2.CrossRefGoogle Scholar
Miller, T.J., Stephens, D.L. & Mertz, J.E. (1982). Kinetics of accumulation and processing of simian virus 40 rna in Xenopus laeivs oocytes injected with simian virus 40 Dna. Mol Cell Biol 2 1581–94.Google ScholarPubMed
MInshull, J., Blow, J.J. & Hunt, T.. (1989). Translation of cycilin mRNA is necessary for extracts of activated Xenopus eggs to enter mitosis. Cell 56, 947–56.CrossRefGoogle Scholar
Murray, M.T., Schiller, D.L.. & Franke, W.W.. (1992). Sequence analysis of cytoplasmic mRNA-binding proteins of Xenopus oocytes identifies a family of RNA-binding proteins. Proc. Natl. Acad. Sci. USA 89, 11–15.CrossRefGoogle ScholarPubMed
Newport, J. & Kirschner, M.. (1982a). A major developmental transition in early Xeno pus embryos. I. Characterization and timing of cellular changes at midblastula stage. Cell 30, 675–86.CrossRefGoogle Scholar
Newport, J. & Kirschner, M.. (1982b). A major developmental transition in early Xenopus embryos. II. Control of the onset of transcription. Cell 30, 687–96.CrossRefGoogle Scholar
Nieuwkoop, P.D. & Faber, J.. (1967). Normal Table of Xenopus laevis (Daudin). Amsterdam: North-Holland.Google Scholar
Paris, J. & Richter, J.D.. (1990). Maturation–specific polyadenylation and translation control: diversity of cytoplamic polyadenylation elements, influence of poly(A) tail size and formation of ftable polyadenylation complexes. Mol. Cell Biol. 10, 5634–45.Google Scholar
Paris, J., Swenson, K., PiwnicaWorms, H. & Richter, J.D.. (1991). Maturation-specific polyadenylation: in vitro activation by p34C2 and phosphorylation of a 58-kD CPE-binding protein. Genes Dev. 5, 1697–708.CrossRefGoogle ScholarPubMed
Pierandrei–Amaldi, P., Campioni, N., Beccari, E., Bozzoni, I. & Amaldi, F.. (1982). Expression of ribosomal–protein genes in Xenopus laevis development. Cell 30, 163–71.CrossRefGoogle ScholarPubMed
Richter, J.D., Anderson, D.M., Davidson, E.H. & Smith, L.D.. (1984). Interspersed poly(A) RNAs of amphibian oocytes are not translatable, at. J. Mol. Biol. 173, 227–41.CrossRefGoogle ScholarPubMed
Rosbash, M. & Ford, P.J. (1974). Polyadenylic acid-containing RNA in Xenopus laevis oocytes. I. Mol. Biol. 85, 87–101.CrossRefGoogle ScholarPubMed
Sagata, N., Shiokawa, K. & Yamana, K.. (1980). A study of the steady–state population of poly(A) RNA during early development of Xenopus laevis. Dev. Biol. 77, 431–48.CrossRefGoogle ScholarPubMed
Sagata, N., Oskarsson, M., Copeland, T., Brumbaugh, J. & VandeWoude, G.F.. (1988). Function of c-mos protooncogene product in meiotic maturation in Xenopus oocytes. Nature 335, 519–25.CrossRefGoogle ScholarPubMed
Sambrook, J., Fritsch, E.F. & Maniatis, T.. (1989). Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.Google Scholar
Simon, R. & Richter, J.D.. (1992). Translational control by poly(A) elongation during Xenopus development: differential repression and enhancement by a novel cytoplasmic polyadenylation element. Genes Dev. 6, 2580–91.CrossRefGoogle ScholarPubMed
Smith, L.D.. (1992). Translational regulation of maternal messenger RNA. In Advances in Development Biochemistry, ed. Wasserman, PM, 1, 136–67. Greenwich, Connecticut: JAI Press.Google Scholar
Tafuri, R. & Wolffe, A.P.. (1993). Dual roles for transcription and translation factors in the RNA storage particles of Xeno pus oocytes. Trends Cell Biol. 3, 94–8.CrossRefGoogle Scholar
Tannahill, D. & Melton, D.A.. (1989). Localized synthesis of the Vgl protein during early Xenopus development. Development 106, 775–85.CrossRefGoogle Scholar
Taylor, M.A. & Smith, L.D.. (1985). Quantitative changes in protein synthesis during oogenesis in Xeno pus laevis. Dev. Biol. 110, 230–7.CrossRefGoogle Scholar
Varnum, S.M. & Wormington, W.M.. (1990). Deadenylation of mRNAs during Xenopus oocyte maturation does not require specific cis–sequences: a default mechanism for translational control. Genes Dev. 4, 2278–86.CrossRefGoogle Scholar
Wallace, R.A., Jared, D.W., Dumont, J.N. & Sega, M.W.. (1973). Protein incorporation by isolated amphibian oocytes. III. Optimum incubation conditions. I. Exp. Zool. 184, 321–34.CrossRefGoogle ScholarPubMed
Wickens, M.. (1992). Forward, backward, how much, when: mechanisms of poly(A) addition and removal and their role in early development. Semin. Dev. Biol. 3, 399–412.Google Scholar
Wickens, M.P.. & Gurdon, J.B. (1983). Post-transcriptional processing of simian virus 40 late transcripts in injected frog oocytes. I. Mol. Biol. 163, 1–26.CrossRefGoogle ScholarPubMed
Yisraeli, J.K., Sokol, S. & Melton, D.A.. (1990). A two-step model for the locelization of maternal mrna in Xenopus oocytes: invoolvement of microtubes and microfilamrnts in the translocation and anchoring of Vg1 mRna. Development 108, 289–98.CrossRefGoogle Scholar