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  • Review Article
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Roles of mRNA poly(A) tails in regulation of eukaryotic gene expression

Abstract

In eukaryotes, poly(A) tails are present on almost every mRNA. Early experiments led to the hypothesis that poly(A) tails and the cytoplasmic polyadenylate-binding protein (PABPC) promote translation and prevent mRNA degradation, but the details remained unclear. More recent data suggest that the role of poly(A) tails is much more complex: poly(A)-binding protein can stimulate poly(A) tail removal (deadenylation) and the poly(A) tails of stable, highly translated mRNAs at steady state are much shorter than expected. Furthermore, the rate of translation elongation affects deadenylation. Consequently, the interplay between poly(A) tails, PABPC, translation and mRNA decay has a major role in gene regulation. In this Review, we discuss recent work that is revolutionizing our understanding of the roles of poly(A) tails in the cytoplasm. Specifically, we discuss the roles of poly(A) tails in translation and control of mRNA stability and how poly(A) tails are removed by exonucleases (deadenylases), including CCR4–NOT and PAN2–PAN3. We also discuss how deadenylation rate is determined, the integration of deadenylation with other cellular processes and the function of PABPC. We conclude with an outlook for the future of research in this field.

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Fig. 1: Overview of the function of mRNA poly(A) tails as master regulators of gene expression in the cytoplasm.
Fig. 2: mRNA poly(A) tails stimulate translation.
Fig. 3: Eukaryotic mRNA deadenylation and decay.
Fig. 4: Deadenylation by PAN2–PAN3 and CCR4–NOT.
Fig. 5: Factors that influence deadenylation rate.
Fig. 6: Summary of recent insights into gene regulation by poly(A) tails.

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References

  1. Edmonds, M. & Abrams, R. Polynucleotide biosynthesis: formation of a sequence of adenylate units from adenosine triphosphate by an enzyme from thymus nuclei. J. Biol. Chem. 235, 1142–1149 (1960).

    Article  CAS  PubMed  Google Scholar 

  2. Hadjivassiliou, A. & Brawerman, G. Polyadenylic acid in the cytoplasm of rat liver. J. Mol. Biol. 20, 1–7 (1966).

    Article  CAS  PubMed  Google Scholar 

  3. Edmonds, M. & Caramela, M. G. The isolation and characterization of adenosine monophosphate-rich polynucleotides synthesized by Ehrlich ascites cells. J. Biol. Chem. 244, 1314–1324 (1969).

    Article  CAS  PubMed  Google Scholar 

  4. Burr, H. & Lingrel, J. B. Poly A sequences at the 3′ termini of rabbit globin mRNAs. Nat. New Biol. 233, 41–43 (1971).

    Article  CAS  PubMed  Google Scholar 

  5. Lim, L. & Canellakis, E. S. Adenine-rich polymer associated with rabbit reticulocyte messenger RNA. Nature 227, 710–712 (1970).

    Article  CAS  PubMed  Google Scholar 

  6. Kates, J. & Beeson, J. Ribonucleic acid synthesis in vaccinia virus. II. Synthesis of polyriboadenylic acid. J. Mol. Biol. 50, 19–33 (1970).

    Article  CAS  PubMed  Google Scholar 

  7. Edmonds, M., Vaughan, M. H. & Nakazato, H. Polyadenylic acid sequences in the heterogeneous nuclear RNA and rapidly-labeled polyribosomal RNA of HeLa cells: possible evidence for a precursor relationship. Proc. Natl Acad. Sci. USA 68, 1336–1340 (1971).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  8. McLaughlin, C. S., Warner, J. R., Edmonds, M., Nakazato, H. & Vaughan, M. H. Polyadenylic acid sequences in yeast messenger ribonucleic acid. J. Biol. Chem. 248, 1466–1471 (1973).

    Article  CAS  PubMed  Google Scholar 

  9. Gallie, D. R. The cap and poly(A) tail function synergistically to regulate mRNA translational efficiency. Genes Dev. 5, 2108–2116 (1991).

    Article  CAS  PubMed  Google Scholar 

  10. Goldstrohm, A. C. & Wickens, M. Multifunctional deadenylase complexes diversify mRNA control. Nat. Rev. Mol. Cell Biol. 9, 337–344 (2008).

    Article  CAS  PubMed  Google Scholar 

  11. Vende, P., Piron, M., Castagné, N. & Poncet, D. Efficient translation of rotavirus mRNA requires simultaneous interaction of NSP3 with the eukaryotic translation initiation factor eIF4G and the mRNA 3′ end. J. Virol. 74, 7064–7071 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  12. Hsu, M. T., Parvin, J. D., Gupta, S., Krystal, M. & Palese, P. Genomic RNAs of influenza viruses are held in a circular conformation in virions and in infected cells by a terminal panhandle. Proc. Natl Acad. Sci. USA 84, 8140–8144 (1987).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Cakmakci, N. G., Lerner, R. S., Wagner, E. J., Zheng, L. & Marzluff, W. F. SLIP1, a factor required for activation of histone mRNA translation by the stem-loop binding protein. Mol. Cell. Biol. 28, 1182–1194 (2008).

    Article  CAS  PubMed  Google Scholar 

  14. Holstege, F. C. et al. Dissecting the regulatory circuitry of a eukaryotic genome. Cell 95, 717–728 (1998).

    Article  CAS  PubMed  Google Scholar 

  15. Wang, Z.-Y. et al. Transcriptome and translatome co-evolution in mammals. Nature 588, 642–647 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Geisberg, J. V., Moqtaderi, Z., Fan, X., Ozsolak, F. & Struhl, K. Global analysis of mRNA isoform half-lives reveals stabilizing and destabilizing elements in yeast. Cell 156, 812–824 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  17. Eisen, T. J. et al. The dynamics of cytoplasmic mRNA metabolism. Mol. Cell 77, 786–799.e10 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  18. Xiang, K. & Bartel, D. P. The molecular basis of coupling between poly(A)-tail length and translational efficiency. eLife 10, e66493 (2021).

    Article  PubMed  PubMed Central  Google Scholar 

  19. Wilt, F. H. Polyadenylation of maternal RNA of sea urchin eggs after fertilization. Proc. Natl Acad. Sci. USA 70, 2345–2349 (1973).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Slater, D. W., Slater, I. & Gillespie, D. Post-fertilization synthesis of polyadenylic acid in sea urchin embryos. Nature 240, 333–337 (1972).

    Article  CAS  PubMed  Google Scholar 

  21. Rosenthal, E. T., Tansey, T. R. & Ruderman, J. V. Sequence-specific adenylations and deadenylations accompany changes in the translation of maternal messenger RNA after fertilization of Spisula oocytes. J. Mol. Biol. 166, 309–327 (1983).

    Article  CAS  PubMed  Google Scholar 

  22. Dworkin, M. B. & Dworkin-Rastl, E. Changes in RNA titers and polyadenylation during oogenesis and oocyte maturation in Xenopus laevis. Dev. Biol. 112, 451–457 (1985).

    Article  CAS  PubMed  Google Scholar 

  23. Dworkin, M. B., Shrutkowski, A. & Dworkin-Rastl, E. Mobilization of specific maternal RNA species into polysomes after fertilization in Xenopus laevis. Proc. Natl Acad. Sci. USA 82, 7636–7640 (1985).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  24. Fox, C. A. & Wickens, M. 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, 2287–2298 (1990).

    Article  CAS  PubMed  Google Scholar 

  25. McGrew, L. L. & Richter, J. D. Translational control by cytoplasmic polyadenylation during Xenopus oocyte maturation: characterization of cis and trans elements and regulation by cyclin/MPF. EMBO J. 9, 3743–3751 (1990).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. McGrew, L. L., Dworkin-Rastl, E., Dworkin, M. B. & Richter, J. D. Poly(A) elongation during Xenopus oocyte maturation is required for translational recruitment and is mediated by a short sequence element. Genes Dev. 3, 803–815 (1989).

    Article  CAS  PubMed  Google Scholar 

  27. Sheets, M. D., Wu, M. & Wickens, M. Polyadenylation of c-mos mRNA as a control point in Xenopus meiotic maturation. Nature 374, 511–516 (1995).

    Article  CAS  PubMed  Google Scholar 

  28. Barkoff, A., Ballantyne, S. & Wickens, M. Meiotic maturation in Xenopus requires polyadenylation of multiple mRNAs. EMBO J. 17, 3168–3175 (1998).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Wu, L. et al. CPEB-mediated cytoplasmic polyadenylation and the regulation of experience-dependent translation of alpha-CaMKII mRNA at synapses. Neuron 21, 1129–1139 (1998).

    Article  CAS  PubMed  Google Scholar 

  30. Doel, M. T. & Carey, N. H. The translational capacity of deadenylated ovalbumin messenger RNA. Cell 8, 51–58 (1976).

    Article  CAS  PubMed  Google Scholar 

  31. Jacobson, A. & Favreau, M. Possible involvement of poly(A) in protein synthesis. Nucleic Acids Res. 11, 6353–6368 (1983).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Subtelny, A. O., Eichhorn, S. W., Chen, G. R., Sive, H. & Bartel, D. P. Poly(A)-tail profiling reveals an embryonic switch in translational control. Nature 508, 66–71 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  33. Lima, S. A. et al. Short poly(A) tails are a conserved feature of highly expressed genes. Nat. Struct. Mol. Biol. 24, 1057–1063 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Groppo, R. & Richter, J. D. Translational control from head to tail. Curr. Opin. Cell Biol. 21, 444–451 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. Huang, Y.-S. & Richter, J. D. Regulation of local mRNA translation. Curr. Opin. Cell Biol. 16, 308–313 (2004).

    Article  PubMed  Google Scholar 

  36. Vicens, Q., Kieft, J. S. & Rissland, O. S. Revisiting the closed-loop model and the nature of mRNA 5′-3′ communication. Mol. Cell 72, 805–812 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Baer, B. W. & Kornberg, R. D. The protein responsible for the repeating structure of cytoplasmic poly(A)-ribonucleoprotein. J. Cell Biol. 96, 717–721 (1983).

    Article  CAS  PubMed  Google Scholar 

  38. Blobel, G. A protein of molecular weight 78,000 bound to the polyadenylate region of eukaryotic messenger RNAs. Proc. Natl Acad. Sci. USA 70, 924–928 (1973).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Baer, B. W. & Kornberg, R. D. Repeating structure of cytoplasmic poly(A)-ribonucleoprotein. Proc. Natl Acad. Sci. USA 77, 1890–1892 (1980).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  40. Smith, R. W. P., Blee, T. K. P. & Gray, N. K. Poly(A)-binding proteins are required for diverse biological processes in metazoans. Biochem. Soc. Trans. 42, 1229–1237 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Burd, C. G., Matunis, E. L. & Dreyfuss, G. The multiple RNA-binding domains of the mRNA poly(A)-binding protein have different RNA-binding activities. Mol. Cell. Biol. 11, 3419–3424 (1991).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. Kühn, U. & Pieler, T. Xenopus poly(A) binding protein: functional domains in RNA binding and protein-protein interaction. J. Mol. Biol. 256, 20–30 (1996).

    Article  PubMed  Google Scholar 

  43. Deo, R. C., Bonanno, J. B., Sonenberg, N. & Burley, S. K. Recognition of polyadenylate RNA by the poly(A)-binding protein. Cell 98, 835–845 (1999).

    Article  CAS  PubMed  Google Scholar 

  44. Xie, J., Kozlov, G. & Gehring, K. The ‘tale’ of poly(A) binding protein: the MLLE domain and PAM2-containing proteins. Biochim. Biophys. Acta 1839, 1062–1068 (2014).

    Article  CAS  PubMed  Google Scholar 

  45. Wigington, C. P., Williams, K. R., Meers, M. P., Bassell, G. J. & Corbett, A. H. Poly(A) RNA-binding proteins and polyadenosine RNA: new members and novel functions. Wiley Interdiscip. Rev. RNA 5, 601–622 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  46. Schäfer, I. B. et al. Molecular basis for poly(A) RNP architecture and recognition by the Pan2-Pan3 deadenylase. Cell 177, 1619–1631.e21 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  47. Melo, E. O., Dhalia, R., Martins de Sa, C., Standart, N. & de Melo Neto, O. P. Identification of a C-terminal poly(A)-binding protein (PABP)-PABP interaction domain: role in cooperative binding to poly (A) and efficient cap distal translational repression. J. Biol. Chem. 278, 46357–46368 (2003).

    Article  CAS  PubMed  Google Scholar 

  48. Rissland, O. S. et al. The influence of microRNAs and poly(A) tail length on endogenous mRNA-protein complexes. Genome Biol. 18, 211 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  49. Sachs, A. B. & Davis, R. W. The poly(A) binding protein is required for poly(A) shortening and 60S ribosomal subunit-dependent translation initiation. Cell 58, 857–867 (1989).

    Article  CAS  PubMed  Google Scholar 

  50. Brune, C., Munchel, S. E., Fischer, N., Podtelejnikov, A. V. & Weis, K. Yeast poly(A)-binding protein Pab1 shuttles between the nucleus and the cytoplasm and functions in mRNA export. RNA 11, 517–531 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. Caponigro, G. & Parker, R. Multiple functions for the poly(A)-binding protein in mRNA decapping and deadenylation in yeast. Genes Dev. 9, 2421–2432 (1995).

    Article  CAS  PubMed  Google Scholar 

  52. Coller, J. M., Gray, N. K. & Wickens, M. P. mRNA stabilization by poly(A) binding protein is independent of poly(A) and requires translation. Genes Dev. 12, 3226–3235 (1998).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  53. Gray, N. K., Coller, J. M., Dickson, K. S. & Wickens, M. Multiple portions of poly(A)-binding protein stimulate translation in vivo. EMBO J. 19, 4723–4733 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  54. Blagden, S. P. et al. Drosophila Larp associates with poly(A)-binding protein and is required for male fertility and syncytial embryo development. Dev. Biol. 334, 186–197 (2009).

    Article  CAS  PubMed  Google Scholar 

  55. Shirokikh, N. E. & Preiss, T. Translation initiation by cap-dependent ribosome recruitment: recent insights and open questions. Wiley Interdiscip. Rev. RNA 9, e1473 (2018).

    Article  PubMed  Google Scholar 

  56. Tarun, S. Z. & Sachs, A. B. Association of the yeast poly(A) tail binding protein with translation initiation factor eIF-4G. EMBO J. 15, 7168–7177 (1996).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  57. Jacobson, A. & Peltz, S. W. Interrelationships of the pathways of mRNA decay and translation in eukaryotic cells. Annu. Rev. Biochem. 65, 693–739 (1996).

    Article  CAS  PubMed  Google Scholar 

  58. Borman, A. M., Michel, Y. M. & Kean, K. M. Biochemical characterisation of cap-poly(A) synergy in rabbit reticulocyte lysates: the eIF4G-PABP interaction increases the functional affinity of eIF4E for the capped mRNA 5′-end. Nucleic Acids Res. 28, 4068–4075 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  59. Bi, X. & Goss, D. J. Wheat germ poly(A)-binding protein increases the ATPase and the RNA helicase activity of translation initiation factors eIF4A, eIF4B, and eIF-iso4F. J. Biol. Chem. 275, 17740–17746 (2000).

    Article  CAS  PubMed  Google Scholar 

  60. Kapp, L. D. & Lorsch, J. R. The molecular mechanics of eukaryotic translation. Annu. Rev. Biochem. 73, 657–704 (2004).

    Article  CAS  PubMed  Google Scholar 

  61. Munroe, D. & Jacobson, A. mRNA poly(A) tail, a 3′ enhancer of translational initiation. Mol. Cell. Biol. 10, 3441–3455 (1990).

    CAS  PubMed  PubMed Central  Google Scholar 

  62. Preiss, T. & Hentze, M. W. Dual function of the messenger RNA cap structure in poly(A)-tail-promoted translation in yeast. Nature 392, 516–520 (1998).

    Article  CAS  PubMed  Google Scholar 

  63. Wells, S. E., Hillner, P. E., Vale, R. D. & Sachs, A. B. Circularization of mRNA by eukaryotic translation initiation factors. Mol. Cell 2, 135–140 (1998).

    Article  CAS  PubMed  Google Scholar 

  64. Amrani, N., Ghosh, S., Mangus, D. A. & Jacobson, A. Translation factors promote the formation of two states of the closed-loop mRNP. Nature 453, 1276–1280 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  65. Adivarahan, S. et al. Spatial organization of single mRNPs at different stages of the gene expression pathway. Mol. Cell 72, 727–738.e5 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  66. Costello, J. et al. Global mRNA selection mechanisms for translation initiation. Genome Biol. 16, 10 (2015).

    Article  PubMed  PubMed Central  Google Scholar 

  67. Sheiness, D., Puckett, L. & Darnell, J. E. Possible relationship of poly(A) shortening to mRNA turnover. Proc. Natl Acad. Sci. USA 72, 1077–1081 (1975).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Merkel, C. G., Kwan, S. & Lingrel, J. B. Size of the polyadenylic acid region of newly synthesized globin messenger ribonucleic acid. J. Biol. Chem. 250, 3725–3728 (1975).

    Article  CAS  PubMed  Google Scholar 

  69. Gorski, J., Morrison, M. R., Merkel, C. G. & Lingrel, J. B. Poly(A) size class distribution in globin mRNAs as a function of time. Nature 253, 749–751 (1975).

    Article  CAS  PubMed  Google Scholar 

  70. Nudel, U., Soreq, H. & Littauer, U. Z. Globin mRNA species containing poly(A) segments of different lengths. Their functional stability in Xenopus oocytes. Eur. J. Biochem. 64, 115–121 (1976).

    Article  CAS  PubMed  Google Scholar 

  71. Mercer, J. F. & Wake, S. A. An analysis of the rate of metallothionein mRNA poly(A)-shortening using RNA blot hybridization. Nucleic Acids Res. 13, 7929–7943 (1985).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  72. Peng, J. & Schoenberg, D. R. mRNA with a <20-nt poly(A) tail imparted by the poly(A)-limiting element is translated as efficiently in vivo as long poly(A) mRNA. RNA 11, 1131–1140 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  73. Wilson, T. & Treisman, R. Removal of poly(A) and consequent degradation of c-fos mRNA facilitated by 3′ AU-rich sequences. Nature 336, 396–399 (1988).

    Article  CAS  PubMed  Google Scholar 

  74. Swartwout, S. G. & Kinniburgh, A. J. c-myc RNA degradation in growing and differentiating cells: possible alternate pathways. Mol. Cell. Biol. 9, 288–295 (1989).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. Brewer, G. & Ross, J. Poly(A) shortening and degradation of the 3′ A+U-rich sequences of human c-myc mRNA in a cell-free system. Mol. Cell. Biol. 8, 1697–1708 (1988).

    CAS  PubMed  PubMed Central  Google Scholar 

  76. Bernstein, P., Peltz, S. W. & Ross, J. The poly(A)-poly(A)-binding protein complex is a major determinant of mRNA stability in vitro. Mol. Cell. Biol. 9, 659–670 (1989).

    CAS  PubMed  PubMed Central  Google Scholar 

  77. Simón, E. & Séraphin, B. A specific role for the C-terminal region of the poly(A)-binding protein in mRNA decay. Nucleic Acids Res. 35, 6017–6028 (2007).

    Article  PubMed  PubMed Central  Google Scholar 

  78. Tucker, M., Staples, R. R., Valencia-Sanchez, M. A., Muhlrad, D. & Parker, R. Ccr4p is the catalytic subunit of a Ccr4p/Pop2p/Notp mRNA deadenylase complex in Saccharomyces cerevisiae. EMBO J. 21, 1427–1436 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  79. Viswanathan, P., Ohn, T., Chiang, Y.-C., Chen, J. & Denis, C. L. Mouse CAF1 can function as a processive deadenylase/3′-5′-exonuclease in vitro but in yeast the deadenylase function of CAF1 is not required for mRNA poly(A) removal. J. Biol. Chem. 279, 23988–23995 (2004).

    Article  CAS  PubMed  Google Scholar 

  80. Decker, C. J. & Parker, R. A turnover pathway for both stable and unstable mRNAs in yeast: evidence for a requirement for deadenylation. Genes Dev. 7, 1632–1643 (1993).

    Article  CAS  PubMed  Google Scholar 

  81. Muhlrad, D., Decker, C. J. & Parker, R. Deadenylation of the unstable mRNA encoded by the yeast MFA2 gene leads to decapping followed by 5′–>3′ digestion of the transcript. Genes Dev. 8, 855–866 (1994).

    Article  CAS  PubMed  Google Scholar 

  82. Hsu, C. L. & Stevens, A. Yeast cells lacking 5′–>3′ exoribonuclease 1 contain mRNA species that are poly(A) deficient and partially lack the 5′ cap structure. Mol. Cell. Biol. 13, 4826–4835 (1993).

    CAS  PubMed  PubMed Central  Google Scholar 

  83. Garneau, N. L., Wilusz, J. & Wilusz, C. J. The highways and byways of mRNA decay. Nat. Rev. Mol. Cell Biol. 8, 113–126 (2007).

    Article  CAS  PubMed  Google Scholar 

  84. Kurosaki, T., Popp, M. W. & Maquat, L. E. Quality and quantity control of gene expression by nonsense-mediated mRNA decay. Nat. Rev. Mol. Cell Biol. 20, 406–420 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  85. Shyu, A. B., Belasco, J. G. & Greenberg, M. E. Two distinct destabilizing elements in the c-fos message trigger deadenylation as a first step in rapid mRNA decay. Genes Dev. 5, 221–231 (1991).

    Article  CAS  PubMed  Google Scholar 

  86. Muhlrad, D. & Parker, R. Mutations affecting stability and deadenylation of the yeast MFA2 transcript. Genes Dev. 6, 2100–2111 (1992).

    Article  CAS  PubMed  Google Scholar 

  87. Chen, C. Y., Chen, T. M. & Shyu, A. B. Interplay of two functionally and structurally distinct domains of the c-fos AU-rich element specifies its mRNA-destabilizing function. Mol. Cell. Biol. 14, 416–426 (1994).

    Article  PubMed  PubMed Central  Google Scholar 

  88. Schiavi, S. C. et al. Multiple elements in the c-fos protein-coding region facilitate mRNA deadenylation and decay by a mechanism coupled to translation. J. Biol. Chem. 269, 3441–3448 (1994).

    Article  CAS  PubMed  Google Scholar 

  89. Schwartz, D. C. & Parker, R. mRNA decapping in yeast requires dissociation of the cap binding protein, eukaryotic translation initiation factor 4E. Mol. Cell. Biol. 20, 7933–7942 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  90. Schwartz, D. C. & Parker, R. Mutations in translation initiation factors lead to increased rates of deadenylation and decapping of mRNAs in Saccharomyces cerevisiae. Mol. Cell. Biol. 19, 5247–5256 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  91. Khanna, R. & Kiledjian, M. Poly(A)-binding-protein-mediated regulation of hDcp2 decapping in vitro. EMBO J. 23, 1968–1976 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  92. Lobel, J. H. & Gross, J. D. Pdc2/Pat1 increases the range of decay factors and RNA bound by the Lsm1-7 complex. RNA 26, 1380–1388 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  93. Montemayor, E. J. et al. Molecular basis for the distinct cellular functions of the Lsm1-7 and Lsm2-8 complexes. RNA 26, 1400–1413 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. He, W. & Parker, R. Functions of Lsm proteins in mRNA degradation and splicing. Curr. Opin. Cell Biol. 12, 346–350 (2000).

    Article  CAS  PubMed  Google Scholar 

  95. Fabian, M. R. et al. Mammalian miRNA RISC recruits CAF1 and PABP to affect PABP-dependent deadenylation. Mol. Cell 35, 868–880 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  96. Sachs, A. B. & Deardorff, J. A. Translation initiation requires the PAB-dependent poly(A) ribonuclease in yeast. Cell 70, 961–973 (1992).

    Article  CAS  PubMed  Google Scholar 

  97. Boeck, R. et al. The yeast Pan2 protein is required for poly(A)-binding protein-stimulated poly(A)-nuclease activity. J. Biol. Chem. 271, 432–438 (1996).

    Article  CAS  PubMed  Google Scholar 

  98. Brown, C. E., Tarun, S. Z., Boeck, R. & Sachs, A. B. PAN3 encodes a subunit of the Pab1p-dependent poly(A) nuclease in Saccharomyces cerevisiae. Mol. Cell. Biol. 16, 5744–5753 (1996).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  99. Wolf, J. et al. Structural basis for Pan3 binding to Pan2 and its function in mRNA recruitment and deadenylation. EMBO J. 33, 1514–1526 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  100. Schäfer, I. B., Rode, M., Bonneau, F., Schüssler, S. & Conti, E. The structure of the Pan2-Pan3 core complex reveals cross-talk between deadenylase and pseudokinase. Nat. Struct. Mol. Biol. 21, 591–598 (2014).

    Article  PubMed  Google Scholar 

  101. Jonas, S. et al. An asymmetric PAN3 dimer recruits a single PAN2 exonuclease to mediate mRNA deadenylation and decay. Nat. Struct. Mol. Biol. 21, 599–608 (2014).

    Article  CAS  PubMed  Google Scholar 

  102. Siddiqui, N. et al. Poly(A) nuclease interacts with the C-terminal domain of polyadenylate-binding protein domain from poly(A)-binding protein. J. Biol. Chem. 282, 25067–25075 (2007).

    Article  CAS  PubMed  Google Scholar 

  103. Lowell, J. E., Rudner, D. Z. & Sachs, A. B. 3′-UTR-dependent deadenylation by the yeast poly(A) nuclease. Genes Dev. 6, 2088–2099 (1992).

    Article  CAS  PubMed  Google Scholar 

  104. Tang, T. T. L., Stowell, J. A. W., Hill, C. H. & Passmore, L. A. The intrinsic structure of poly(A) RNA determines the specificity of Pan2 and Caf1 deadenylases. Nat. Struct. Mol. Biol. 26, 433–442 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  105. Braun, J. E., Huntzinger, E., Fauser, M. & Izaurralde, E. GW182 proteins directly recruit cytoplasmic deadenylase complexes to miRNA targets. Mol. Cell 44, 120–133 (2011).

    Article  CAS  PubMed  Google Scholar 

  106. Christie, M., Boland, A., Huntzinger, E., Weichenrieder, O. & Izaurralde, E. Structure of the PAN3 pseudokinase reveals the basis for interactions with the PAN2 deadenylase and the GW182 proteins. Mol. Cell 51, 360–373 (2013).

    Article  CAS  PubMed  Google Scholar 

  107. Tucker, M. et al. The transcription factor associated Ccr4 and Caf1 proteins are components of the major cytoplasmic mRNA deadenylase in Saccharomyces cerevisiae. Cell 104, 377–386 (2001).

    Article  CAS  PubMed  Google Scholar 

  108. Chen, J., Chiang, Y.-C. & Denis, C. L. CCR4, a 3′-5′ poly(A) RNA and ssDNA exonuclease, is the catalytic component of the cytoplasmic deadenylase. EMBO J. 21, 1414–1426 (2002).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  109. Finoux, A.-L. & Séraphin, B. In vivo targeting of the yeast Pop2 deadenylase subunit to reporter transcripts induces their rapid degradation and generates new decay intermediates. J. Biol. Chem. 281, 25940–25947 (2006).

    Article  CAS  PubMed  Google Scholar 

  110. Thore, S., Mauxion, F., Séraphin, B. & Suck, D. X-ray structure and activity of the yeast Pop2 protein: a nuclease subunit of the mRNA deadenylase complex. EMBO Rep. 4, 1150–1155 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  111. Daugeron, M. C., Mauxion, F. & Séraphin, B. The yeast POP2 gene encodes a nuclease involved in mRNA deadenylation. Nucleic Acids Res. 29, 2448–2455 (2001).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  112. Maillet, L., Tu, C., Hong, Y. K., Shuster, E. O. & Collart, M. A. The essential function of Not1 lies within the Ccr4-Not complex. J. Mol. Biol. 303, 131–143 (2000).

    Article  CAS  PubMed  Google Scholar 

  113. Bai, Y. et al. The CCR4 and CAF1 proteins of the CCR4-NOT complex are physically and functionally separated from NOT2, NOT4, and NOT5. Mol. Cell. Biol. 19, 6642–6651 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  114. Alhusaini, N. & Coller, J. The deadenylase components Not2p, Not3p, and Not5p promote mRNA decapping. RNA 22, 709–721 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  115. Muhlrad, D. & Parker, R. The yeast EDC1 mRNA undergoes deadenylation-independent decapping stimulated by Not2p, Not4p, and Not5p. EMBO J. 24, 1033–1045 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  116. Sgromo, A. et al. A CAF40-binding motif facilitates recruitment of the CCR4-NOT complex to mRNAs targeted by Drosophila Roquin. Nat. Commun. 8, 14307 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  117. Sgromo, A. et al. Drosophila Bag-of-marbles directly interacts with the CAF40 subunit of the CCR4-NOT complex to elicit repression of mRNA targets. RNA 24, 381–395 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  118. Raisch, T. et al. Reconstitution of recombinant human CCR4-NOT reveals molecular insights into regulated deadenylation. Nat. Commun. 10, 3173 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  119. Albert, T. K. et al. Isolation and characterization of human orthologs of yeast CCR4-NOT complex subunits. Nucleic Acids Res. 28, 809–817 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  120. Jiang, H., Wolgast, M., Beebe, L. M. & Reese, J. C. Ccr4-Not maintains genomic integrity by controlling the ubiquitylation and degradation of arrested RNAPII. Genes. Dev. 33, 705–717 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  121. Panasenko, O. O. The role of the E3 ligase Not4 in cotranslational quality control. Front. Genet. 5, 141 (2014).

    Article  PubMed  PubMed Central  Google Scholar 

  122. Buschauer, R. et al. The Ccr4-Not complex monitors the translating ribosome for codon optimality. Science 368, eaay6912 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  123. Körner, C. G. & Wahle, E. Poly(A) tail shortening by a mammalian poly(A)-specific 3′-exoribonuclease. J. Biol. Chem. 272, 10448–10456 (1997).

    Article  PubMed  Google Scholar 

  124. Körner, C. G. et al. The deadenylating nuclease (DAN) is involved in poly(A) tail removal during the meiotic maturation of Xenopus oocytes. EMBO J. 17, 5427–5437 (1998).

    Article  PubMed  PubMed Central  Google Scholar 

  125. Aström, J., Aström, A. & Virtanen, A. In vitro deadenylation of mammalian mRNA by a HeLa cell 3′ exonuclease. EMBO J. 10, 3067–3071 (1991).

    Article  PubMed  PubMed Central  Google Scholar 

  126. Yi, H. et al. PABP cooperates with the CCR4-NOT complex to promote mRNA deadenylation and block precocious decay. Mol. Cell 70, 1081–1088.e5 (2018).

    Article  CAS  PubMed  Google Scholar 

  127. Berndt, H. et al. Maturation of mammalian H/ACA box snoRNAs: PAPD5-dependent adenylation and PARN-dependent trimming. RNA 18, 958–972 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  128. Tang, W., Tu, S., Lee, H.-C., Weng, Z. & Mello, C. C. The RNase PARN-1 trims piRNA 3′ ends to promote transcriptome surveillance in C. elegans. Cell 164, 974–984 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  129. Shukla, S. & Parker, R. PARN modulates Y RNA stability and its 3′-end formation. Mol. Cell. Biol. 37, e00264-17 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  130. Webster, M. W. et al. mRNA deadenylation is coupled to translation rates by the differential activities of Ccr4-Not nucleases. Mol. Cell 70, 1089–1100.e8 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  131. Yamashita, A. et al. Concerted action of poly(A) nucleases and decapping enzyme in mammalian mRNA turnover. Nat. Struct. Mol. Biol. 12,1054–1063 (2005).

    Article  CAS  PubMed  Google Scholar 

  132. Stowell, J. A. W. et al. Reconstitution of targeted deadenylation by the Ccr4-Not complex and the YTH domain protein Mmi1. Cell Rep. 17, 1978–1989 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  133. Cheng, J., Maier, K. C., Avsec, Ž., Rus, P. & Gagneur, J. Cis-regulatory elements explain most of the mRNA stability variation across genes in yeast. RNA 23, 1648–1659 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  134. Lykke-Andersen, J. & Wagner, E. Recruitment and activation of mRNA decay enzymes by two ARE-mediated decay activation domains in the proteins TTP and BRF-1. Genes Dev. 19, 351–361 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  135. Brooks, S. A. & Blackshear, P. J. Tristetraprolin (TTP): interactions with mRNA and proteins, and current thoughts on mechanisms of action. Biochim. Biophys. Acta 1829, 666–679 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  136. Goldstrohm, A. C., Hall, T. M. T. & McKenney, K. M. Post-transcriptional regulatory functions of mammalian Pumilio proteins. Trends Genet. 34, 972–990 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  137. Wang, X., Zamore, P. D. & Hall, T. M. Crystal structure of a Pumilio homology domain. Mol. Cell 7, 855–865 (2001).

    Article  CAS  PubMed  Google Scholar 

  138. Wickens, M., Bernstein, D. S., Kimble, J. & Parker, R. A PUF family portrait: 3′UTR regulation as a way of life. Trends Genet. 18, 150–157 (2002).

    Article  CAS  PubMed  Google Scholar 

  139. Goldstrohm, A. C., Hook, B. A., Seay, D. J. & Wickens, M. PUF proteins bind Pop2p to regulate messenger RNAs. Nat. Struct. Mol. Biol. 13, 533–539 (2006).

    Article  CAS  PubMed  Google Scholar 

  140. Jonas, S. & Izaurralde, E. Towards a molecular understanding of microRNA-mediated gene silencing. Nat. Rev. Genet. 16, 421–433 (2015).

    Article  CAS  PubMed  Google Scholar 

  141. Du, H. et al. YTHDF2 destabilizes m(6)A-containing RNA through direct recruitment of the CCR4-NOT deadenylase complex. Nat. Commun. 7, 12626 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  142. Webster, M. W., Stowell, J. A. & Passmore, L. A. RNA-binding proteins distinguish between similar sequence motifs to promote targeted deadenylation by Ccr4-Not. eLife 8, e40670 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  143. Weidmann, C. A. & Goldstrohm, A. C. Drosophila pumilio protein contains multiple autonomous repression domains that regulate mRNAs independently of nanos and brain Tumor. Mol. Cell. Biol. 32, 527–540 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  144. Arvola, R. M. et al. Unique repression domains of Pumilio utilize deadenylation and decapping factors to accelerate destruction of target mRNAs. Nucleic Acids Res. 48, 1843–1871 (2020).

    Article  CAS  PubMed  Google Scholar 

  145. Fabian, M. R. et al. Structural basis for the recruitment of the human CCR4-NOT deadenylase complex by tristetraprolin. Nat. Struct. Mol. Biol. 20, 735–739 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  146. Jonas, S. & Izaurralde, E. The role of disordered protein regions in the assembly of decapping complexes and RNP granules. Genes Dev. 27, 2628–2641 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  147. Raisch, T. et al. Distinct modes of recruitment of the CCR4-NOT complex by Drosophila and vertebrate Nanos. EMBO J. 35, 974–990 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  148. Marchese, F. P. et al. MAPKAP kinase 2 blocks tristetraprolin-directed mRNA decay by inhibiting CAF1 deadenylase recruitment. J. Biol. Chem. 285, 27590–27600 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  149. Akira, S. & Maeda, K. Control of RNA stability in immunity. Annu. Rev. Immunol. 39, 481–509 (2021).

    Article  CAS  PubMed  Google Scholar 

  150. Kundu, P., Fabian, M. R., Sonenberg, N., Bhattacharyya, S. N. & Filipowicz, W. HuR protein attenuates miRNA-mediated repression by promoting miRISC dissociation from the target RNA. Nucleic Acids Res. 40, 5088–5100 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  151. Srikantan, S., Tominaga, K. & Gorospe, M. Functional interplay between RNA-binding protein HuR and microRNAs. Curr. Protein Pept. Sci. 13, 372–379 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  152. Beelman, C. A. & Parker, R. Differential effects of translational inhibition in cis and in trans on the decay of the unstable yeast MFA2 mRNA. J. Biol. Chem. 269, 9687–9692 (1994).

    Article  CAS  PubMed  Google Scholar 

  153. Muhlrad, D., Decker, C. J. & Parker, R. Turnover mechanisms of the stable yeast PGK1 mRNA. Mol. Cell. Biol. 15, 2145–2156 (1995).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  154. Coller, J. & Parker, R. General translational repression by activators of mRNA decapping. Cell 122, 875–886 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  155. Bazzini, A. A. et al. Codon identity regulates mRNA stability and translation efficiency during the maternal-to-zygotic transition. EMBO J. 35, 2087–2103 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  156. Presnyak, V. et al. Codon optimality is a major determinant of mRNA stability. Cell 160, 1111–1124 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  157. Radhakrishnan, A. & Green, R. Connections underlying translation and mRNA stability. J. Mol. Biol. 428, 3558–3564 (2016).

    Article  CAS  PubMed  Google Scholar 

  158. Forrest, M. E. et al. Codon and amino acid content are associated with mRNA stability in mammalian cells. PLoS ONE 15, e0228730 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  159. Harigaya, Y. & Parker, R. Analysis of the association between codon optimality and mRNA stability in Schizosaccharomyces pombe. BMC Genomics 17, 895 (2016).

    Article  PubMed  PubMed Central  Google Scholar 

  160. de Freitas Nascimento, J., Kelly, S., Sunter, J. & Carrington, M. Codon choice directs constitutive mRNA levels in trypanosomes. eLife 7, e32467 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  161. Burow, D. A. et al. Attenuated codon optimality contributes to neural-specific mRNA decay in drosophila. Cell Rep. 24, 1704–1712 (2018).

    Article  CAS  PubMed  Google Scholar 

  162. Mishima, Y. & Tomari, Y. Codon usage and 3′ UTR length determine maternal mRNA stability in zebrafish. Mol. Cell 61, 874–885 (2016).

    Article  CAS  PubMed  Google Scholar 

  163. Wu, Q. et al. Translation affects mRNA stability in a codon-dependent manner in human cells. eLife 8, e45396 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  164. Narula, A., Ellis, J., Taliaferro, J. M. & Rissland, O. S. Coding regions affect mRNA stability in human cells. RNA 25, 1751–1764 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  165. Hia, F. et al. Codon bias confers stability to human mRNAs. EMBO Rep. 20, e48220 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  166. Hanson, G. & Coller, J. Codon optimality, bias and usage in translation and mRNA decay. Nat. Rev. Mol. Cell Biol. 19, 20–30 (2018).

    Article  CAS  PubMed  Google Scholar 

  167. Panasenko, O. et al. The yeast Ccr4-Not complex controls ubiquitination of the nascent-associated polypeptide (NAC-EGD) complex. J. Biol. Chem. 281, 31389–31398 (2006).

    Article  CAS  PubMed  Google Scholar 

  168. Ikeuchi, K. et al. Collided ribosomes form a unique structural interface to induce Hel2-driven quality control pathways. EMBO J. 38, e100276 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  169. Panasenko, O. O. & Collart, M. A. Presence of Not5 and ubiquitinated Rps7A in polysome fractions depends upon the Not4 E3 ligase. Mol. Microbiol. 83, 640–653 (2012).

    Article  CAS  PubMed  Google Scholar 

  170. Mauger, D. M. et al. mRNA structure regulates protein expression through changes in functional half-life. Proc. Natl Acad. Sci. USA 116, 24075–24083 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  171. Mauxion, F., Faux, C. & Séraphin, B. The BTG2 protein is a general activator of mRNA deadenylation. EMBO J. 27, 1039–1048 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  172. Chen, C.-Y. A., Strouz, K., Huang, K.-L. & Shyu, A.-B. Tob2 phosphorylation regulates global mRNA turnover to reshape transcriptome and impact cell proliferation. RNA 26, 1143–1159 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  173. Mattijssen, S., Kozlov, G., Fonseca, B. D., Gehring, K. & Maraia, R. J. LARP1 and LARP4: up close with PABP for mRNA 3′ poly(A) protection and stabilization. RNA Biol. 18, 259–274 (2021).

    Article  CAS  PubMed  Google Scholar 

  174. Mattijssen, S., Iben, J. R., Li, T., Coon, S. L. & Maraia, R. J. Single molecule poly(A) tail-seq shows LARP4 opposes deadenylation throughout mRNA lifespan with most impact on short tails. eLife 9, e59186 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  175. Funakoshi, Y. et al. Mechanism of mRNA deadenylation: evidence for a molecular interplay between translation termination factor eRF3 and mRNA deadenylases. Genes Dev. 21, 3135–3148 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  176. Chang, H., Lim, J., Ha, M. & Kim, V. N. TAIL-seq: genome-wide determination of poly(A) tail length and 3′ end modifications. Mol. Cell 53, 1044–1052 (2014).

    Article  CAS  PubMed  Google Scholar 

  177. Lim, J. et al. Mixed tailing by TENT4A and TENT4B shields mRNA from rapid deadenylation. Science 361, 701–704 (2018).

    Article  CAS  PubMed  Google Scholar 

  178. Legnini, I., Alles, J., Karaiskos, N., Ayoub, S. & Rajewsky, N. FLAM-seq: full-length mRNA sequencing reveals principles of poly(A) tail length control. Nat. Methods 16, 879–886 (2019).

    Article  CAS  PubMed  Google Scholar 

  179. Kim, D. et al. Viral hijacking of the TENT4-ZCCHC14 complex protects viral RNAs via mixed tailing. Nat. Struct. Mol. Biol. 27, 581–588 (2020).

    Article  CAS  PubMed  Google Scholar 

  180. Rissland, O. S. & Norbury, C. J. Decapping is preceded by 3′ uridylation in a novel pathway of bulk mRNA turnover. Nat. Struct. Mol. Biol. 16, 616–623 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  181. Lim, J., Lee, M., Son, A., Chang, H. & Kim, V. N. mTAIL-seq reveals dynamic poly(A) tail regulation in oocyte-to-embryo development. Genes Dev. 30, 1671–1682 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  182. Workman, R. E. et al. Nanopore native RNA sequencing of a human poly(A) transcriptome. Nat. Methods 16, 1297–1305 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  183. Shapiro, R. A., Herrick, D., Manrow, R. E., Blinder, D. & Jacobson, A. Determinants of mRNA stability in Dictyostelium discoideum amoebae: differences in poly(A) tail length, ribosome loading, and mRNA size cannot account for the heterogeneity of mRNA decay rates. Mol. Cell. Biol. 8, 1957–1969 (1988).

    CAS  PubMed  PubMed Central  Google Scholar 

  184. Zekri, L., Kuzuoğlu-Öztürk, D. & Izaurralde, E. GW182 proteins cause PABP dissociation from silenced miRNA targets in the absence of deadenylation. EMBO J. 32, 1052–1065 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  185. Tsuboi, T. & Inada, T. Tethering of poly(A)-binding protein interferes with non-translated mRNA decay from the 5′ end in yeast. J. Biol. Chem. 285, 33589–33601 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  186. Wu, C., Roy, B., He, F., Yan, K. & Jacobson, A. Poly(A)-binding protein regulates the efficiency of translation termination. Cell Rep. 33, 108399 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  187. de Melo Neto, O. P., Standart, N. & Martins de Sa, C. Autoregulation of poly(A)-binding protein synthesis in vitro. Nucleic Acids Res. 23, 2198–2205 (1995).

    Article  PubMed  PubMed Central  Google Scholar 

  188. Wan, R., Bai, R. & Shi, Y. Molecular choreography of pre-mRNA splicing by the spliceosome. Curr. Opin. Struct. Biol. 59, 124–133 (2019).

    Article  CAS  PubMed  Google Scholar 

  189. Chen, Y. et al. A DDX6-CNOT1 complex and W-binding pockets in CNOT9 reveal direct links between miRNA target recognition and silencing. Mol. Cell 54, 737–750 (2014).

    Article  CAS  PubMed  Google Scholar 

  190. Mathys, H. et al. Structural and biochemical insights to the role of the CCR4-NOT complex and DDX6 ATPase in microRNA repression. Mol. Cell 54, 751–765 (2014).

    Article  CAS  PubMed  Google Scholar 

  191. Radhakrishnan, A. et al. The DEAD-Box protein Dhh1p couples mRNA decay and translation by monitoring codon optimality. Cell 167, 122–132.e9 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  192. Saenger, W., Riecke, J. & Suck, D. A structural model for the polyadenylic acid single helix. J. Mol. Biol. 93, 529–534 (1975).

    Article  CAS  PubMed  Google Scholar 

  193. Tang, T. T. L. & Passmore, L. A. Recognition of Poly(A) RNA through its intrinsic helical structure. Cold Spring Harb. Symp. Quant. Biol. 84, 21–30 (2019).

    Article  PubMed  Google Scholar 

  194. Chandrasekaran, V. et al. Mechanism of ribosome stalling during translation of a poly(A) tail. Nat. Struct. Mol. Biol. 26, 1132–1140 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  195. Tesina, P. et al. Molecular mechanism of translational stalling by inhibitory codon combinations and poly(A) tracts. EMBO J. 39, e103365 (2020).

    Article  CAS  PubMed  Google Scholar 

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Acknowledgements

We thank A. Jacobson, M. Wickens, F. He, T. Tang, E. Absmeier, J. Leipheimer, S. Martin, R. Kawalerski and members of the Passmore and Coller laboratories for helpful insight and discussion. This Review would not have been possible without contributions from many individuals; this field has been rich in discovery, collaboration and friendship. We thank all of you and apologize for any oversights we may have made. In particular, we dedicate this Review to the memory of Dr Elisa Izaurralde and Dr Richard Jackson; their seminal discoveries and keen intellect have inspired us all. Funding is provided to L.A.P. by the European Union’s Horizon 2020 research and innovation programme (ERC grant No. 725685) and the Medical Research Council as part of United Kingdom Research and Innovation (MRC grant No. MC_U105192715) and to J.C. by the National Institutes of Health (USA; GM118018 and GM125086) and Bloomberg Philanthropies.

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Glossary

mRNA decay

The process of removing mRNA from the cytosolic pool of transcripts. Decay of mRNA occurs in a defined pathway with each mRNA having an intrinsic half-life.

Translation efficiency

The amount of protein output relative to the amount of transcribed mRNA.

Bypass suppressors

A mutation at a distinct locus, which restores viability following mutation of an essential gene. Bypass suppressors often provide insight into the function of essential genes.

Intrinsically disordered regions

(IDRs). Polypeptide segments enriched in polar or charged amino acids and lacking hydrophobic amino acids that would mediate cooperative folding. IDRs generally lack a secondary structure.

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Passmore, L.A., Coller, J. Roles of mRNA poly(A) tails in regulation of eukaryotic gene expression. Nat Rev Mol Cell Biol 23, 93–106 (2022). https://doi.org/10.1038/s41580-021-00417-y

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