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A tale of non-canonical tails: gene regulation by post-transcriptional RNA tailing

Nature Reviews Molecular Cell Biology volume 21, pages 542–556 (2020)Cite this article

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Abstract

RNA tailing, or the addition of non-templated nucleotides to the 3′ end of RNA, is the most frequent and conserved type of RNA modification. The addition of tails and their composition reflect RNA maturation stages and have important roles in determining the fate of the modified RNAs. Apart from canonical poly(A) polymerases, which add poly(A) tails to mRNAs in a transcription-coupled manner, a family of terminal nucleotidyltransferases (TENTs), including terminal uridylyltransferases (TUTs), modify RNAs post-transcriptionally to control RNA stability and activity. The human genome encodes 11 different TENTs with distinct substrate specificity, intracellular localization and tissue distribution. In this Review, we discuss recent advances in our understanding of non-canonical RNA tails, with a focus on the functions of human TENTs, which include uridylation, mixed tailing and post-transcriptional polyadenylation of mRNAs, microRNAs and other types of non-coding RNA.

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Fig. 1: Human terminal nucleotidyltransferases.
Fig. 2: Polyadenylation and mixed tailing of mRNAs by terminal nucleotidyl transferases.
Fig. 3: Uridylation of mRNAs by TUT4 and TUT7.
Fig. 4: Pre-miRNA uridylation by TUT4 and TUT7.

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References

  1. Norbury, C. J. Cytoplasmic RNA: a case of the tail wagging the dog. Nat. Rev. Mol. Cell Biol. 14, 643–653 (2013).

    Article  CAS  PubMed  Google Scholar 

  2. Colgan, D. F. & Manley, J. L. Mechanism and regulation of mRNA polyadenylation. Genes Dev. 11, 2755–2766 (1997).

    Article  CAS  PubMed  Google Scholar 

  3. 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 

  4. Stewart, M. Polyadenylation and nuclear export of mRNAs. J. Biol. Chem. 294, 2977–2987 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  5. Weill, L., Belloc, E., Bava, F. A. & Mendez, R. Translational control by changes in poly(A) tail length: recycling mRNAs. Nat. Struct. Mol. Biol. 19, 577–585 (2012).

    Article  CAS  PubMed  Google Scholar 

  6. 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 

  7. 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 

  8. 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 

  9. Villalba, A., Coll, O. & Gebauer, F. Cytoplasmic polyadenylation and translational control. Curr. Opin. Genet. Dev. 21, 452–457 (2011).

    Article  CAS  PubMed  Google Scholar 

  10. Munoz-Tello, P., Rajappa, L., Coquille, S. & Thore, S. Polyuridylation in eukaryotes: a 3′-end modification regulating RNA life. Biomed. Res. Int. 2015, 968127 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  11. Lim, J. et al. Mixed tailing by TENT4A and TENT4B shields mRNA from rapid deadenylation. Science 361, 701–704 (2018). This article shows that TENT4A and TENT4B incorporate intermittent non-adenosine nucleotides with a preference for guanosine, which results in the formation of heterogeneous poly(A) tails.

    Article  CAS  PubMed  Google Scholar 

  12. Dupasquier, M., Kim, S., Halkidis, K., Gamper, H. & Hou, Y. M. tRNA integrity is a prerequisite for rapid CCA addition: implication for quality control. J. Mol. Biol. 379, 579–588 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  13. Tazi, J., Forne, T., Jeanteur, P., Cathala, G. & Brunel, C. Mammalian U6 small nuclear RNA undergoes 3′ end modifications within the spliceosome. Mol. Cell Biol. 13, 1641–1650 (1993).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. 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 

  15. Kim, Y. K., Heo, I. & Kim, V. N. Modifications of small RNAs and their associated proteins. Cell 143, 703–709 (2010).

    Article  CAS  PubMed  Google Scholar 

  16. Egecioglu, D. E., Henras, A. K. & Chanfreau, G. F. Contributions of Trf4p- and Trf5p-dependent polyadenylation to the processing and degradative functions of the yeast nuclear exosome. RNA 12, 26–32 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  17. LaCava, J. et al. RNA degradation by the exosome is promoted by a nuclear polyadenylation complex. Cell 121, 713–724 (2005). This work identifies the nuclear TRAMP complex in S. cerevisiae, which shows distributive RNA polyadenylation activity on rRNA and snoRNA precursors and promotes their degradation by the exosome.

    Article  CAS  PubMed  Google Scholar 

  18. Shcherbik, N., Wang, M., Lapik, Y. R., Srivastava, L. & Pestov, D. G. Polyadenylation and degradation of incomplete RNA polymerase I transcripts in mammalian cells. EMBO Rep. 11, 106–111 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Labno, A. et al. Perlman syndrome nuclease DIS3L2 controls cytoplasmic non-coding RNAs and provides surveillance pathway for maturing snRNAs. Nucleic Acids Res. 44, 10437–10453 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  20. Pirouz, M., Du, P., Munafo, M. & Gregory, R. I. Dis3l2-mediated decay is a quality control pathway for noncoding RNAs. Cell Rep. 16, 1861–1873 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. Ustianenko, D. et al. TUT-DIS3L2 is a mammalian surveillance pathway for aberrant structured non-coding RNAs. EMBO J. 35, 2179–2191 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. Pirouz, M., Munafo, M., Ebrahimi, A. G., Choe, J. & Gregory, R. I. Exonuclease requirements for mammalian ribosomal RNA biogenesis and surveillance. Nat. Struct. Mol. Biol. 26, 490–500 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Warkocki, Z., Liudkovska, V., Gewartowska, O., Mroczek, S. & Dziembowski, A. Terminal nucleotidyl transferases (TENTs) in mammalian RNA metabolism. Philos. Trans. R. Soc. Lond. B Biol. Sci. 373, 20180162 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  24. Aravind, L. & Koonin, E. V. DNA polymerase beta-like nucleotidyltransferase superfamily: identification of three new families, classification and evolutionary history. Nucleic Acids Res. 27, 1609–1618 (1999).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Martin, G. & Keller, W. RNA-specific ribonucleotidyl transferases. RNA 13, 1834–1849 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Schmidt, M. J. & Norbury, C. J. Polyadenylation and beyond: emerging roles for noncanonical poly(A) polymerases. Wiley Interdiscip. Rev. RNA 1, 142–151 (2010).

    Article  CAS  PubMed  Google Scholar 

  27. Lohr, J. G. et al. Widespread genetic heterogeneity in multiple myeloma: implications for targeted therapy. Cancer Cell 25, 91–101 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  28. Martin, N. T. et al. Homozygous mutation of MTPAP causes cellular radiosensitivity and persistent DNA double-strand breaks. Cell Death Dis. 5, e1130 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Doyard, M. et al. FAM46A mutations are responsible for autosomal recessive osteogenesis imperfecta. J. Med. Genet. 55, 278–284 (2018).

    Article  CAS  PubMed  Google Scholar 

  30. Le Pen, J. et al. Terminal uridylyltransferases target RNA viruses as part of the innate immune system. Nat. Struct. Mol. Biol. 25, 778–786 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  31. Mueller, H. et al. PAPD5/7 are host factors that are required for hepatitis B virus RNA stabilization. Hepatology 69, 1398–1411 (2019).

    Article  CAS  PubMed  Google Scholar 

  32. Martin, G., Keller, W. & Doublie, S. Crystal structure of mammalian poly(A) polymerase in complex with an analog of ATP. EMBO J. 19, 4193–4203 (2000).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  33. Trippe, R. et al. Identification, cloning, and functional analysis of the human U6 snRNA-specific terminal uridylyl transferase. RNA 12, 1494–1504 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Laishram, R. S. & Anderson, R. A. The poly A polymerase Star-PAP controls 3′-end cleavage by promoting CPSF interaction and specificity toward the pre-mRNA. EMBO J. 29, 4132–4145 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  35. Mellman, D. L. et al. A PtdIns4,5P2-regulated nuclear poly(A) polymerase controls expression of select mRNAs. Nature 451, 1013–1017 (2008).

    Article  CAS  PubMed  Google Scholar 

  36. Yamashita, S., Takagi, Y., Nagaike, T. & Tomita, K. Crystal structures of U6 snRNA-specific terminal uridylyltransferase. Nat. Commun. 8, 15788 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Kadyk, L. C. & Kimble, J. Genetic regulation of entry into meiosis in Caenorhabditis elegans. Development 125, 1803–1813 (1998).

    CAS  PubMed  Google Scholar 

  38. Wang, L., Eckmann, C. R., Kadyk, L. C., Wickens, M. & Kimble, J. A regulatory cytoplasmic poly(A) polymerase in Caenorhabditis elegans. Nature 419, 312–316 (2002). This work discovrs that C. elegans GLD-2 is a cytoplasmic poly(A) polymerase whose nucleotidyltransferase activity is stimulated by GLD-3 in vitro.

    Article  CAS  PubMed  Google Scholar 

  39. Barnard, D. C., Ryan, K., Manley, J. L. & Richter, J. D. Symplekin and xGLD-2 are required for CPEB-mediated cytoplasmic polyadenylation. Cell 119, 641–651 (2004).

    Article  CAS  PubMed  Google Scholar 

  40. Suh, N., Jedamzik, B., Eckmann, C. R., Wickens, M. & Kimble, J. The GLD-2 poly(A) polymerase activates gld-1 mRNA in the Caenorhabditis elegans germ line. Proc. Natl Acad. Sci. USA 103, 15108–15112 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Benoit, P., Papin, C., Kwak, J. E., Wickens, M. & Simonelig, M. PAP- and GLD-2-type poly(A) polymerases are required sequentially in cytoplasmic polyadenylation and oogenesis in Drosophila. Development 135, 1969–1979 (2008).

    Article  CAS  PubMed  Google Scholar 

  42. Kwak, J. E. et al. GLD2 poly(A) polymerase is required for long-term memory. Proc. Natl Acad. Sci. USA 105, 14644–14649 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Kim, K. W., Wilson, T. L. & Kimble, J. GLD-2/RNP-8 cytoplasmic poly(A) polymerase is a broad-spectrum regulator of the oogenesis program. Proc. Natl Acad. Sci. USA 107, 17445–17450 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Sartain, C. V., Cui, J., Meisel, R. P. & Wolfner, M. F. The poly(A) polymerase GLD2 is required for spermatogenesis in Drosophila melanogaster. Development 138, 1619–1629 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  45. Cui, J., Sartain, C. V., Pleiss, J. A. & Wolfner, M. F. Cytoplasmic polyadenylation is a major mRNA regulator during oogenesis and egg activation in Drosophila. Dev. Biol. 383, 121–131 (2013).

    Article  CAS  PubMed  Google Scholar 

  46. Kwak, J. E., Wang, L., Ballantyne, S., Kimble, J. & Wickens, M. Mammalian GLD-2 homologs are poly(A) polymerases. Proc. Natl Acad. Sci. USA 101, 4407–4412 (2004). This article describes the first in vivo tethering assay, which identifies human and mouse TENT2 as poly(A) polymerases.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  47. Nakanishi, T. et al. Possible role of mouse poly(A) polymerase mGLD-2 during oocyte maturation. Dev. Biol. 289, 115–126 (2006).

    Article  CAS  PubMed  Google Scholar 

  48. Faehnle, C. R., Walleshauser, J. & Joshua-Tor, L. Multi-domain utilization by TUT4 and TUT7 in control of let-7 biogenesis. Nat. Struct. Mol. Biol. 24, 658–665 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  49. Yamashita, S., Nagaike, T. & Tomita, K. Crystal structure of the Lin28-interacting module of human terminal uridylyltransferase that regulates let-7 expression. Nat. Commun. 10, 1960 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  50. Lim, J. et al. Uridylation by TUT4 and TUT7 marks mRNA for degradation. Cell 159, 1365–1376 (2014). This article shows that TUT4 and TUT7 selectively uridylate deadenylated mRNAs with a short poly(A) tail and facilitate mRNA decay.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. Liu, X. et al. A microRNA precursor surveillance system in quality control of microRNA synthesis. Mol. Cell 55, 868–879 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  52. Hubstenberger, A. et al. P-Body purification reveals the condensation of repressed mRNA regulons. Mol. Cell 68, 144–157 e5 (2017).

    Article  CAS  PubMed  Google Scholar 

  53. Vanacova, S. et al. A new yeast poly(A) polymerase complex involved in RNA quality control. PLoS Biol. 3, e189 (2005).

    Article  PubMed  CAS  Google Scholar 

  54. Wyers, F. et al. Cryptic pol II transcripts are degraded by a nuclear quality control pathway involving a new poly(A) polymerase. Cell 121, 725–737 (2005).

    Article  CAS  PubMed  Google Scholar 

  55. Kadaba, S., Wang, X. & Anderson, J. T. Nuclear RNA surveillance in Saccharomyces cerevisiae: Trf4p-dependent polyadenylation of nascent hypomethylated tRNA and an aberrant form of 5S rRNA. RNA 12, 508–521 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  56. Houseley, J., Kotovic, K., El Hage, A. & Tollervey, D. Trf4 targets ncRNAs from telomeric and rDNA spacer regions and functions in rDNA copy number control. EMBO J. 26, 4996–5006 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  57. Callahan, K. P. & Butler, J. S. TRAMP complex enhances RNA degradation by the nuclear exosome component Rrp6. J. Biol. Chem. 285, 3540–3547 (2010).

    Article  CAS  PubMed  Google Scholar 

  58. Dez, C., Houseley, J. & Tollervey, D. Surveillance of nuclear-restricted pre-ribosomes within a subnucleolar region of Saccharomyces cerevisiae. EMBO J. 25, 1534–1546 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  59. Lubas, M. et al. Interaction profiling identifies the human nuclear exosome targeting complex. Mol. Cell 43, 624–637 (2011).

    Article  CAS  PubMed  Google Scholar 

  60. Rammelt, C., Bilen, B., Zavolan, M. & Keller, W. PAPD5, a noncanonical poly(A) polymerase with an unusual RNA-binding motif. RNA 17, 1737–1746 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  61. Sudo, H., Nozaki, A., Uno, H., Ishida, Y. & Nagahama, M. Interaction properties of human TRAMP-like proteins and their role in pre-rRNA 5′ETS turnover. FEBS Lett. 590, 2963–2972 (2016).

    Article  CAS  PubMed  Google Scholar 

  62. Ogami, K., Cho, R. & Hoshino, S. Molecular cloning and characterization of a novel isoform of the non-canonical poly(A) polymerase PAPD7. Biochem. Biophys. Res. Commun. 432, 135–140 (2013).

    Article  CAS  PubMed  Google Scholar 

  63. Kim, D. et al. Viral hijacking of the TENT4-ZCCHC14 complex protects viral RNAs via mixed tailing. Nat. Struct. Mol. Biol. https://doi.org/10.1038/s41594-020-0427-3 (2020).

    Article  PubMed  Google Scholar 

  64. Fasken, M. B. et al. Air1 zinc knuckles 4 and 5 and a conserved IWRXY motif are critical for the function and integrity of the Trf4/5-Air1/2-Mtr4 polyadenylation (TRAMP) RNA quality control complex. J. Biol. Chem. 286, 37429–37445 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  65. Kuchta, K., Knizewski, L., Wyrwicz, L. S., Rychlewski, L. & Ginalski, K. Comprehensive classification of nucleotidyltransferase fold proteins: identification of novel families and their representatives in human. Nucleic Acids Res. 37, 7701–7714 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  66. Kuchta, K. et al. FAM46 proteins are novel eukaryotic non-canonical poly(A) polymerases. Nucleic Acids Res. 44, 3534–3548 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  67. Mroczek, S. et al. The non-canonical poly(A) polymerase FAM46C acts as an onco-suppressor in multiple myeloma. Nat. Commun. 8, 619 (2017). This article shows for the first time that TENT5C is an active poly(A) polymerase, which stabilizes mRNAs and enhances gene expression.

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  68. Tomecki, R., Dmochowska, A., Gewartowski, K., Dziembowski, A. & Stepien, P. P. Identification of a novel human nuclear-encoded mitochondrial poly(A) polymerase. Nucleic Acids Res. 32, 6001–6014 (2004).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  69. Nagaike, T., Suzuki, T., Katoh, T. & Ueda, T. Human mitochondrial mRNAs are stabilized with polyadenylation regulated by mitochondria-specific poly(A) polymerase and polynucleotide phosphorylase. J. Biol. Chem. 280, 19721–19727 (2005).

    Article  CAS  PubMed  Google Scholar 

  70. Bratic, A. et al. Mitochondrial polyadenylation is a one-step process required for mRNA integrity and tRNA maturation. PLoS Genet. 12, e1006028 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  71. Borowski, L. S., Szczesny, R. J., Brzezniak, L. K. & Stepien, P. P. RNA turnover in human mitochondria: more questions than answers? Biochim. Biophys. Acta 1797, 1066–1070 (2010).

    Article  CAS  PubMed  Google Scholar 

  72. Wilson, W. C. et al. A human mitochondrial poly(A) polymerase mutation reveals the complexities of post-transcriptional mitochondrial gene expression. Hum. Mol. Genet. 23, 6345–6355 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  73. 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). This work develops a sequencing technique that allows genomic-scale investigation of poly (A) tail lengths and 3′-end modifications of mRNAs.

    Article  CAS  PubMed  Google Scholar 

  74. Radford, H. E., Meijer, H. A. & de Moor, C. H. Translational control by cytoplasmic polyadenylation in Xenopus oocytes. Biochim. Biophys. Acta 1779, 217–229 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  75. Charlesworth, A., Meijer, H. A. & de Moor, C. H. Specificity factors in cytoplasmic polyadenylation. Wiley Interdiscip. Rev. RNA 4, 437–461 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  76. Ivshina, M., Lasko, P. & Richter, J. D. Cytoplasmic polyadenylation element binding proteins in development, health, and disease. Annu. Rev. Cell Dev. Biol. 30, 393–415 (2014).

    Article  CAS  PubMed  Google Scholar 

  77. Nakanishi, T. et al. Disruption of mouse poly(A) polymerase mGLD-2 does not alter polyadenylation status in oocytes and somatic cells. Biochem. Biophys. Res. Commun. 364, 14–19 (2007).

    Article  CAS  PubMed  Google Scholar 

  78. Rouhana, L. et al. Vertebrate GLD2 poly(A) polymerases in the germline and the brain. RNA 11, 1117–1130 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  79. Udagawa, T. et al. Bidirectional control of mRNA translation and synaptic plasticity by the cytoplasmic polyadenylation complex. Mol. Cell 47, 253–266 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  80. Swanger, S. A., He, Y. A., Richter, J. D. & Bassell, G. J. Dendritic GluN2A synthesis mediates activity-induced NMDA receptor insertion. J. Neurosci. 33, 8898–8908 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  81. Mansur, F. et al. Gld2-catalyzed 3′ monoadenylation of miRNAs in the hippocampus has no detectable effect on their stability or on animal behavior. RNA 22, 1492–1499 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  82. Yamagishi, R., Tsusaka, T., Mitsunaga, H., Maehata, T. & Hoshino, S. The STAR protein QKI-7 recruits PAPD4 to regulate post-transcriptional polyadenylation of target mRNAs. Nucleic Acids Res. 44, 2475–2490 (2016).

    Article  PubMed  PubMed Central  Google Scholar 

  83. Burns, D. M. & Richter, J. D. CPEB regulation of human cellular senescence, energy metabolism, and p53 mRNA translation. Genes. Dev. 22, 3449–3460 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  84. Burns, D. M., D’Ambrogio, A., Nottrott, S. & Richter, J. D. CPEB and two poly(A) polymerases control miR-122 stability and p53 mRNA translation. Nature 473, 105–108 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  85. Shin, J., Paek, K. Y., Ivshina, M., Stackpole, E. E. & Richter, J. D. Essential role for non-canonical poly(A) polymerase GLD4 in cytoplasmic polyadenylation and carbohydrate metabolism. Nucleic Acids Res. 45, 6793–6804 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  86. Zhu, Y. X. et al. Loss of FAM46C promotes cell survival in myeloma. Cancer Res. 77, 4317–4327 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  87. Bilska, A. et al. B cell humoral response and differentiation is regulated by the non-canonical poly(A) polymerase TENT5C. bioRxiv https://doi.org/10.1101/686683 (2019).

    Article  Google Scholar 

  88. Lagali, P. S., Kakuk, L. E., Griesinger, I. B., Wong, P. W. & Ayyagari, R. Identification and characterization of C6orf37, a novel candidate human retinal disease gene on chromosome 6q14. Biochem. Biophys. Res. Commun. 293, 356–365 (2002).

    Article  CAS  PubMed  Google Scholar 

  89. Barragan, I. et al. Genetic analysis of FAM46A in Spanish families with autosomal recessive retinitis pigmentosa: characterisation of novel VNTRs. Ann. Hum. Genet. 72, 26–34 (2008).

    Article  CAS  PubMed  Google Scholar 

  90. Etokebe, G. E. et al. Family-with-sequence-similarity-46, member A (Fam46a) gene is expressed in developing tooth buds. Arch. Oral. Biol. 54, 1002–1007 (2009).

    Article  CAS  PubMed  Google Scholar 

  91. Diener, S. et al. Exome sequencing identifies a nonsense mutation in Fam46a associated with bone abnormalities in a new mouse model for skeletal dysplasia. Mamm. Genome 27, 111–121 (2016).

    Article  CAS  PubMed  Google Scholar 

  92. Etokebe, G. E. et al. Susceptibility to large-joint osteoarthritis (hip and knee) is associated with BAG6 rs3117582 SNP and the VNTR polymorphism in the second exon of the FAM46A gene on chromosome 6. J. Orthop. Res. 33, 56–62 (2015).

    Article  CAS  PubMed  Google Scholar 

  93. Benjachat, T. et al. Biomarkers for refractory lupus nephritis: a microarray study of kidney tissue. Int. J. Mol. Sci. 16, 14276–14290 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  94. Bettoni, F. et al. Identification of FAM46D as a novel cancer/testis antigen using EST data and serological analysis. Genomics 94, 153–160 (2009).

    Article  CAS  PubMed  Google Scholar 

  95. Hamilton, S. M. et al. Multiple autism-like behaviors in a novel transgenic mouse model. Behav. Brain Res. 218, 29–41 (2011).

    Article  CAS  PubMed  Google Scholar 

  96. Kwak, J. E. & Wickens, M. A family of poly(U) polymerases. RNA 13, 860–867 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  97. Rissland, O. S., Mikulasova, A. & Norbury, C. J. Efficient RNA polyuridylation by noncanonical poly(A) polymerases. Mol. Cell Biol. 27, 3612–3624 (2007). This is the first description of the in vitro activities of the poly(U) polymerase Cid1 in Schizosaccharomyces pombe and of TUT4 in humans.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  98. Rissland, O. S. & Norbury, C. J. The Cid1 poly(U) polymerase. Biochim. Biophys. Acta 1779, 286–294 (2008).

    Article  CAS  PubMed  Google Scholar 

  99. 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 

  100. Chung, C. Z. et al. RNA surveillance by uridylation-dependent RNA decay in Schizosaccharomyces pombe. Nucleic Acids Res. 47, 3045–3057 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  101. Lunde, B. M., Magler, I. & Meinhart, A. Crystal structures of the Cid1 poly (U) polymerase reveal the mechanism for UTP selectivity. Nucleic Acids Res. 40, 9815–9824 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  102. Munoz-Tello, P., Gabus, C. & Thore, S. Functional implications from the Cid1 poly(U) polymerase crystal structure. Structure 20, 977–986 (2012).

    Article  CAS  PubMed  Google Scholar 

  103. Yates, L. A. et al. Structural basis for the activity of a cytoplasmic RNA terminal uridylyl transferase. Nat. Struct. Mol. Biol. 19, 782–787 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  104. Kuhn, 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  CAS  PubMed  Google Scholar 

  105. 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 

  106. Sement, F. M. et al. Uridylation prevents 3′ trimming of oligoadenylated mRNAs. Nucleic Acids Res. 41, 7115–7127 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  107. Thomas, M. P. et al. Apoptosis triggers specific, rapid, and global mRNA decay with 3′ uridylated intermediates degraded by DIS3L2. Cell Rep. 11, 1079–1089 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  108. Morgan, M. et al. mRNA 3′ uridylation and poly(A) tail length sculpt the mammalian maternal transcriptome. Nature 548, 347–351 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  109. Chang, H. et al. Terminal uridylyltransferases execute programmed clearance of maternal transcriptome in vertebrate embryos. Mol. Cell 70, 72–82 (2018).

    Article  CAS  PubMed  Google Scholar 

  110. Morgan, M. et al. A programmed wave of uridylation-primed mRNA degradation is essential for meiotic progression and mammalian spermatogenesis. Cell Res. 29, 221–232 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  111. Marzluff, W. F., Wagner, E. J. & Duronio, R. J. Metabolism and regulation of canonical histone mRNAs: life without a poly(A) tail. Nat. Rev. Genet. 9, 843–854 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  112. Osley, M. A. The regulation of histone synthesis in the cell cycle. Annu. Rev. Biochem. 60, 827–861 (1991).

    Article  CAS  PubMed  Google Scholar 

  113. Mullen, T. E. & Marzluff, W. F. Degradation of histone mRNA requires oligouridylation followed by decapping and simultaneous degradation of the mRNA both 5′ to 3′ and 3′ to 5′. Genes. Dev. 22, 50–65 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  114. Schmidt, M. J., West, S. & Norbury, C. J. The human cytoplasmic RNA terminal U-transferase ZCCHC11 targets histone mRNAs for degradation. RNA 17, 39–44 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  115. Su, W. et al. mRNAs containing the histone 3′ stem-loop are degraded primarily by decapping mediated by oligouridylation of the 3′ end. RNA 19, 1–16 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  116. Lackey, P. E., Welch, J. D. & Marzluff, W. F. TUT7 catalyzes the uridylation of the 3′ end for rapid degradation of histone mRNA. RNA 22, 1673–1688 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  117. Slevin, M. K. et al. Deep sequencing shows multiple oligouridylations are required for 3′ to 5′ degradation of histone mRNAs on polyribosomes. Mol. Cell 53, 1020–1030 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  118. Welch, J. D. et al. EnD-Seq and AppEnD: sequencing 3′ ends to identify nontemplated tails and degradation intermediates. RNA 21, 1375–1389 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  119. Marzluff, W. F. & Koreski, K. P. Birth and death of histone mRNAs. Trends Genet. 33, 745–759 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  120. Warkocki, Z. et al. Uridylation by TUT4/7 restricts retrotransposition of human LINE-1s. Cell 174, 1537–1548 e29 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  121. Yeo, J. & Kim, V. N. U-tail as a guardian against invading RNAs. Nat. Struct. Mol. Biol. 25, 903–905 (2018).

    Article  CAS  PubMed  Google Scholar 

  122. Burroughs, A. M. et al. A comprehensive survey of 3′ animal miRNA modification events and a possible role for 3′ adenylation in modulating miRNA targeting effectiveness. Genome Res. 20, 1398–1410 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  123. Wyman, S. K. et al. Post-transcriptional generation of miRNA variants by multiple nucleotidyl transferases contributes to miRNA transcriptome complexity. Genome Res. 21, 1450–1461 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  124. Wang, F., Johnson, N. R., Coruh, C. & Axtell, M. J. Genome-wide analysis of single non-templated nucleotides in plant endogenous siRNAs and miRNAs. Nucleic Acids Res. 44, 7395–7405 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  125. Chiang, H. R. et al. Mammalian microRNAs: experimental evaluation of novel and previously annotated genes. Genes. Dev. 24, 992–1009 (2010). This article provides a comprehensive analysis of non-templated nucleotide addition on murine miRNAs.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  126. Newman, M. A., Mani, V. & Hammond, S. M. Deep sequencing of microRNA precursors reveals extensive 3′ end modification. RNA 17, 1795–1803 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  127. Lee, Y. et al. The nuclear RNase III Drosha initiates microRNA processing. Nature 425, 415–419 (2003).

    Article  CAS  PubMed  Google Scholar 

  128. Ha, M. & Kim, V. N. Regulation of microRNA biogenesis. Nat. Rev. Mol. Cell Biol. 15, 509–524 (2014).

    Article  CAS  PubMed  Google Scholar 

  129. Khvorova, A., Reynolds, A. & Jayasena, S. D. Functional siRNAs and miRNAs exhibit strand bias. Cell 115, 209–216 (2003).

    Article  CAS  PubMed  Google Scholar 

  130. Frank, F., Sonenberg, N. & Nagar, B. Structural basis for 5′-nucleotide base-specific recognition of guide RNA by human AGO2. Nature 465, 818–822 (2010).

    Article  CAS  PubMed  Google Scholar 

  131. Suzuki, H. I. et al. Small-RNA asymmetry is directly driven by mammalian Argonautes. Nat. Struct. Mol. Biol. 22, 512–521 (2015).

    Article  CAS  PubMed  Google Scholar 

  132. Bartel, D. P. Metazoan microRNAs. Cell 173, 20–51 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  133. Bussing, I., Slack, F. J. & Grosshans, H. let-7 microRNAs in development, stem cells and cancer. Trends Mol. Med. 14, 400–409 (2008).

    Article  PubMed  CAS  Google Scholar 

  134. Heo, I. et al. Lin28 mediates the terminal uridylation of let-7 precursor MicroRNA. Mol. Cell 32, 276–284 (2008).

    Article  CAS  PubMed  Google Scholar 

  135. Hagan, J. P., Piskounova, E. & Gregory, R. I. Lin28 recruits the TUTase Zcchc11 to inhibit let-7 maturation in mouse embryonic stem cells. Nat. Struct. Mol. Biol. 16, 1021–1025 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  136. Heo, I. et al. TUT4 in concert with Lin28 suppresses microRNA biogenesis through pre-microRNA uridylation. Cell 138, 696–708 (2009).

    Article  CAS  PubMed  Google Scholar 

  137. Yeom, K. H. et al. Single-molecule approach to immunoprecipitated protein complexes: insights into miRNA uridylation. EMBO Rep. 12, 690–696 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  138. Thornton, J. E., Chang, H. M., Piskounova, E. & Gregory, R. I. Lin28-mediated control of let-7 microRNA expression by alternative TUTases Zcchc11 (TUT4) and Zcchc6 (TUT7). RNA 18, 1875–1885 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  139. Ustianenko, D. et al. LIN28 selectively modulates a subclass of Let-7 microRNAs. Mol. Cell 71, 271–283 e5 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  140. Chang, H. M., Triboulet, R., Thornton, J. E. & Gregory, R. I. A role for the Perlman syndrome exonuclease Dis3l2 in the Lin28-let-7 pathway. Nature 497, 244–248 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  141. Ustianenko, D. et al. Mammalian DIS3L2 exoribonuclease targets the uridylated precursors of let-7 miRNAs. RNA 19, 1632–1638 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  142. Faehnle, C. R., Walleshauser, J. & Joshua-Tor, L. Mechanism of Dis3l2 substrate recognition in the Lin28-let-7 pathway. Nature 514, 252–256 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  143. Roush, S. & Slack, F. J. The let-7 family of microRNAs. Trends Cell Biol. 18, 505–516 (2008).

    Article  CAS  PubMed  Google Scholar 

  144. Heo, I. et al. Mono-uridylation of pre-microRNA as a key step in the biogenesis of group II let-7 microRNAs. Cell 151, 521–532 (2012).

    Article  CAS  PubMed  Google Scholar 

  145. Park, J. E. et al. Dicer recognizes the 5′ end of RNA for efficient and accurate processing. Nature 475, 201–205 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  146. Kim, B. et al. TUT7 controls the fate of precursor microRNAs by using three different uridylation mechanisms. EMBO J. 34, 1801–1815 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  147. Kim, H. et al. Bias-minimized quantification of microRNA reveals widespread alternative processing and 3′ end modification. Nucleic Acids Res. 47, 2630–2640 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  148. Kim, H. et al. A mechanism for microRNA arm switching regulated by uridylation. Mol. Cell https://doi.org/10.1016/j.molcel.2020.04.030 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  149. Chung, C. Z., Jo, D. H. & Heinemann, I. U. Nucleotide specificity of the human terminal nucleotidyltransferase Gld2 (TUT2). RNA 22, 1239–1249 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  150. Berezikov, E., Chung, W. J., Willis, J., Cuppen, E. & Lai, E. C. Mammalian mirtron genes. Mol. Cell 28, 328–336 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  151. Okamura, K., Hagen, J. W., Duan, H., Tyler, D. M. & Lai, E. C. The mirtron pathway generates microRNA-class regulatory RNAs in Drosophila. Cell 130, 89–100 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  152. Ruby, J. G., Jan, C. H. & Bartel, D. P. Intronic microRNA precursors that bypass Drosha processing. Nature 448, 83–86 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  153. Westholm, J. O., Ladewig, E., Okamura, K., Robine, N. & Lai, E. C. Common and distinct patterns of terminal modifications to mirtrons and canonical microRNAs. RNA 18, 177–192 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  154. Bortolamiol-Becet, D. et al. Selective suppression of the splicing-mediated microRNA pathway by the terminal uridyltransferase tailor. Mol. Cell 59, 217–228 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  155. Reimao-Pinto, M. M. et al. Uridylation of RNA hairpins by tailor confines the emergence of microRNAs in Drosophila. Mol. Cell 59, 203–216 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  156. Reimao-Pinto, M. M. et al. Molecular basis for cytoplasmic RNA surveillance by uridylation-triggered decay in Drosophila. EMBO J. 35, 2417–2434 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  157. Chang, J. et al. miR-122, a mammalian liver-specific microRNA, is processed from hcr mRNA and may downregulate the high affinity cationic amino acid transporter CAT-1. RNA Biol. 1, 106–113 (2004).

    Article  CAS  PubMed  Google Scholar 

  158. Katoh, T. et al. Selective stabilization of mammalian microRNAs by 3′ adenylation mediated by the cytoplasmic poly(A) polymerase GLD-2. Genes. Dev. 23, 433–438 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  159. D’Ambrogio, A., Gu, W., Udagawa, T., Mello, C. C. & Richter, J. D. Specific miRNA stabilization by Gld2-catalyzed monoadenylation. Cell Rep. 2, 1537–1545 (2012).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  160. Hojo, H. et al. The RNA-binding protein QKI-7 recruits the poly(A) polymerase GLD-2 for 3′ adenylation and selective stabilization of microRNA-122. J. Biol. Chem. 295, 390–402 (2020).

    Article  PubMed  Google Scholar 

  161. Katoh, T., Hojo, H. & Suzuki, T. Destabilization of microRNAs in human cells by 3′ deadenylation mediated by PARN and CUGBP1. Nucleic Acids Res. 43, 7521–7534 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  162. Lee, M. et al. Adenylation of maternally inherited microRNAs by Wispy. Mol. Cell 56, 696–707 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  163. Boele, J. et al. PAPD5-mediated 3′ adenylation and subsequent degradation of miR-21 is disrupted in proliferative disease. Proc. Natl Acad. Sci. USA 111, 11467–11472 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  164. Feng, Y. H. & Tsao, C. J. Emerging role of microRNA-21 in cancer. Biomed. Rep. 5, 395–402 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  165. Shukla, S., Bjerke, G. A., Muhlrad, D., Yi, R. & Parker, R. The RNase PARN controls the levels of specific miRNAs that contribute to p53 regulation. Mol. Cell 73, 1204–1216 e4 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  166. Li, J., Yang, Z., Yu, B., Liu, J. & Chen, X. Methylation protects miRNAs and siRNAs from a 3′-end uridylation activity in Arabidopsis. Curr. Biol. 15, 1501–1507 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  167. Yu, B. et al. Methylation as a crucial step in plant microRNA biogenesis. Science 307, 932–935 (2005).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  168. Ren, G., Chen, X. & Yu, B. Uridylation of miRNAs by hen1 suppressor1 in Arabidopsis. Curr. Biol. 22, 695–700 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  169. Zhao, Y. et al. The Arabidopsis nucleotidyl transferase HESO1 uridylates unmethylated small RNAs to trigger their degradation. Curr. Biol. 22, 689–694 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  170. Jones, M. R. et al. Zcchc11-dependent uridylation of microRNA directs cytokine expression. Nat. Cell Biol. 11, 1157–1163 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  171. Jones, M. R. et al. Zcchc11 uridylates mature miRNAs to enhance neonatal IGF-1 expression, growth, and survival. PLoS Genet. 8, e1003105 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  172. Yang, A. et al. 3′ Uridylation confers miRNAs with non-canonical target repertoires. Mol. Cell 75, 511–522 e4 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  173. Gutierrez-Vazquez, C. et al. 3′ Uridylation controls mature microRNA turnover during CD4 T-cell activation. RNA 23, 882–891 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  174. Fuchs Wightman, F., Giono, L. E., Fededa, J. P. & de la Mata, M. Target RNAs strike back on microRNAs. Front. Genet. 9, 435 (2018).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  175. Choi, Y. S., Patena, W., Leavitt, A. D. & McManus, M. T. Widespread RNA 3′-end oligouridylation in mammals. RNA 18, 394–401 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  176. Slomovic, S., Fremder, E., Staals, R. H., Pruijn, G. J. & Schuster, G. Addition of poly(A) and poly(A)-rich tails during RNA degradation in the cytoplasm of human cells. Proc. Natl Acad. Sci. USA 107, 7407–7412 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  177. Slomovic, S., Laufer, D., Geiger, D. & Schuster, G. Polyadenylation of ribosomal RNA in human cells. Nucleic Acids Res. 34, 2966–2975 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  178. Sinturel, F. et al. Diurnal Oscillations in liver mass and cell size accompany ribosome assembly cycles. Cell 169, 651–663 e14 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  179. Son, A., Park, J. E. & Kim, V. N. PARN and TOE1 constitute a 3′ end maturation module for nuclear non-coding RNAs. Cell Rep. 23, 888–898 (2018).

    Article  CAS  PubMed  Google Scholar 

  180. Shukla, S. & Parker, R. PARN modulates Y RNA stability and Its 3′-End formation. Mol Cell Biol https://doi.org/10.1128/MCB.00264-17 (2017).

    Article  PubMed  PubMed Central  Google Scholar 

  181. Nguyen, D. et al. A polyadenylation-dependent 3′ end maturation pathway is required for the synthesis of the human telomerase RNA. Cell Rep. 13, 2244–2257 (2015).

    Article  CAS  PubMed  Google Scholar 

  182. Shukla, S., Schmidt, J. C., Goldfarb, K. C., Cech, T. R. & Parker, R. Inhibition of telomerase RNA decay rescues telomerase deficiency caused by dyskerin or PARN defects. Nat. Struct. Mol. Biol. 23, 286–292 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  183. Stuart, B. D. et al. Exome sequencing links mutations in PARN and RTEL1 with familial pulmonary fibrosis and telomere shortening. Nat. Genet. 47, 512–517 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  184. Hajnsdorf, E. & Kaberdin, V. R. RNA polyadenylation and its consequences in prokaryotes. Philos. Trans. R. Soc. Lond. B. Biol. Sci. https://doi.org/10.1098/rstb.2018.0166 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

  185. Schmid, M., Kuchler, B. & Eckmann, C. R. Two conserved regulatory cytoplasmic poly(A) polymerases, GLD-4 and GLD-2, regulate meiotic progression in C. elegans. Genes. Dev. 23, 824–836 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  186. Nousch, M., Yeroslaviz, A., Habermann, B. & Eckmann, C. R. The cytoplasmic poly(A) polymerases GLD-2 and GLD-4 promote general gene expression via distinct mechanisms. Nucleic Acids Res. 42, 11622–11633 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  187. Hyrina, A. et al. A genome-wide CRISPR screen identifies ZCCHC14 as a host factor required for hepatitis B surface antigen production. Cell Rep. 29, 2970–2978 e6 (2019).

    Article  CAS  PubMed  Google Scholar 

  188. Bai, Y., Srivastava, S. K., Chang, J. H., Manley, J. L. & Tong, L. Structural basis for dimerization and activity of human PAPD1, a noncanonical poly(A) polymerase. Mol. Cell 41, 311–320 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  189. Hamill, S., Wolin, S. L. & Reinisch, K. M. Structure and function of the polymerase core of TRAMP, a RNA surveillance complex. Proc. Natl Acad. Sci. USA 107, 15045–15050 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  190. Nakel, K., Bonneau, F., Eckmann, C. R. & Conti, E. Structural basis for the activation of the C. elegans noncanonical cytoplasmic poly(A)-polymerase GLD-2 by GLD-3. Proc. Natl Acad. Sci. USA 112, 8614–8619 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  191. 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 

  192. Eichhorn, S. W. et al. mRNA poly(A)-tail changes specified by deadenylation broadly reshape translation in Drosophila oocytes and early embryos. Elife 5, e16955 (2016).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  193. 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 

  194. Liu, Y., Nie, H., Liu, H. & Lu, F. Poly(A) inclusive RNA isoform sequencing (PAIso-seq) reveals wide-spread non-adenosine residues within RNA poly(A) tails. Nat. Commun. 10, 5292 (2019).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  195. Preston, M. A. et al. Unbiased screen of RNA tailing activities reveals a poly(UG) polymerase. Nat. Methods 16, 437–445 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  196. Shukla, A. et al. Poly(UG)-tailed RNAs in genome protection and epigenetic inheritance. Nature https://doi.org/10.1038/s41586-020-2323-8 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  197. Huo, Y. et al. Widespread 3′-end uridylation in eukaryotic RNA viruses. Sci. Rep. 6, 25454 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  198. Han, X. et al. Discovery of RG7834: the first-in-class selective and orally available small molecule hepatitis B virus expression inhibitor with novel mechanism of action. J. Med. Chem. 61, 10619–10634 (2018).

    Article  CAS  PubMed  Google Scholar 

  199. Mueller, H. et al. A novel orally available small molecule that inhibits hepatitis B virus expression. J. Hepatol. 68, 412–420 (2018).

    Article  CAS  PubMed  Google Scholar 

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Acknowledgements

The authors are grateful to the members of their laboratory for helpful discussions and comments, especially to H. Kim, D. Kim, Y.-s. Lee and H. Jang for their insights and suggestions. Work in the laboratory of V.N.K. was supported by the Institute for Basic Science from the Ministry of Science and ICT of Korea (IBS-R008-D1 to S.Y. and V.N.K.).

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  1. Center for RNA Research, Institute for Basic Science, Seoul, Korea

    Sha Yu & V. Narry Kim

  2. School of Biological Sciences, Seoul National University, Seoul, Korea

    V. Narry Kim

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HUGO Gene Nomenclature Committee: https://www.genenames.org/

Glossary

mRNA decay

Post-transcriptional mRNA degradation through decapping, 5′–3′ exonucleolytic decay, deadenylation, 3′–5′ exonucleolytic decay and endonucleolytic cleavage.

Small Cajal body-specific RNAs

(scaRNAs). A class of small nucleolar RNAs that localize to nuclear Cajal bodies and guide 2′-O-methylation and pseudouridylation of certain RNA polymerase II-dependent transcripts.

RNA exosome

A ribonuclease complex with both endoribonucleolytic function and 3′–5′ exoribonucleolytic activity that degrades various types of RNA, including mRNAs, ribosomal RNAs and many species of small RNA.

P granules

Membraneless ribonucleoprotein organelles that form in the germline of Caenorhabditis elegans. P granules are associated with RNA metabolism and their formation is driven by liquid–liquid phase separation.

Zinc finger domain

A small, independently folded domain that coordinates with one or more zinc ions to stabilize its structure; binds DNA, RNA, proteins and/or lipids.

Zinc knuckle domain

A zinc-binding motif; in this context a Cys2HisCys (CCHC)-type zinc finger domain.

Cryptic transcripts

A class of non-coding RNAs that are produced by relaxation of transcriptional control and are rapidly degraded by RNA surveillance pathways.

Ameloblasts

A group of cells that exist only during tooth development and deposit tooth enamel.

External transcribed spacer

A non-functional part of precursor ribosomal RNAs, positioned between the different ribosomal RNA species.

Y RNAs

Highly structured small non-coding RNAs essential for the initiation of chromosomal DNA replication in vertebrates. Humans encode four Y RNAs, which localize to both the nucleus and the cytoplasm.

Vault RNAs

Small non-coding RNA components of the large vault ribonucleoprotein. Vault RNAs are predicted to fold into stem–loop structures, which are processed into small vault RNAs through a Dicer-dependent mechanism.

7SL RNAs

Abundant cytoplasmic non-coding RNAs that function in protein secretion as the scaffold that makes possible the assembly of the signal recognition particle complex.

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Yu, S., Kim, V.N. A tale of non-canonical tails: gene regulation by post-transcriptional RNA tailing. Nat Rev Mol Cell Biol 21, 542–556 (2020). https://doi.org/10.1038/s41580-020-0246-8

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