Review

Functional 5′ UTR mRNA structures in eukaryotic translation regulation and how to find them

Published online:

Abstract

RNA molecules can fold into intricate shapes that can provide an additional layer of control of gene expression beyond that of their sequence. In this Review, we discuss the current mechanistic understanding of structures in 5′ untranslated regions (UTRs) of eukaryotic mRNAs and the emerging methodologies used to explore them. These structures may regulate cap-dependent translation initiation through helicase-mediated remodelling of RNA structures and higher-order RNA interactions, as well as cap-independent translation initiation through internal ribosome entry sites (IRESs), mRNA modifications and other specialized translation pathways. We discuss known 5′ UTR RNA structures and how new structure probing technologies coupled with prospective validation, particularly compensatory mutagenesis, are likely to identify classes of structured RNA elements that shape post-transcriptional control of gene expression and the development of multicellular organisms.

  • Subscribe to Nature Reviews Molecular Cell Biology for full access:

    $59

    Subscribe

Additional access options:

Already a subscriber?  Log in  now or  Register  for online access.

References

  1. 1.

    Riboswitches and the RNA world. Cold Spring Harb. Perspect. Biol. 4, 1–15 (2012).

  2. 2.

    , & The ribosome challenge to the RNA world. J. Mol. Evol. 80, 143–161 (2015).

  3. 3.

    et al. Genetic control by a metabolite binding mRNA. Chem. Biol. 9, 1043–1049 (2002).

  4. 4.

    Gene regulation by structured mRNA elements. Trends Genet. 30, 172–181 (2014).

  5. 5.

    & Introns and the origin of nucleus-cytosol compartmentalization. Nature 440, 41–45 (2006).

  6. 6.

    & The rise of regulatory RNA. Nat. Rev. Genet. 15, 423–437 (2014).

  7. 7.

    & Regulation of translation initiation in eukaryotes: mechanisms and biological targets. Cell 136, 731–745 (2009).

  8. 8.

    , & The mechanism of eukaryotic translation initiation and principles of its regulation. Nat. Rev. Mol. Cell Biol. 11, 113–127 (2010).

  9. 9.

    et al. The mammalian ribo-interactome reveals ribosome functional diversity and heterogeneity. Cell 169, 1051–1057 (2017).

  10. 10.

    et al. Heterogeneous ribosomes preferentially translate distinct subpools of mRNAs genome-wide. Mol. Cell 67, 71–83 (2017).

  11. 11.

    , , & Cap and cap-binding proteins in the control of gene expression. Wiley Interdiscip. Rev. RNA 2, 277–298 (2011).

  12. 12.

    et al. RNA binding protein/RNA element interactions and the control of translation. Curr. Protein Pept. Sci. 13, 294–304 (2012).

  13. 13.

    & The roles of RNA processing in translating genotype to phenotype. Nat. Rev. Mol. Cell Biol. 18, 102–114 (2016).

  14. 14.

    Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res. 31, 3406–3415 (2003).

  15. 15.

    et al. ViennaRNA package 2.0. Algorithms Mol. Biol. 6, 26 (2011).

  16. 16.

    Regulation by 3′-untranslated regions. Annu. Rev. Genet. 51, 171–194 (2017).

  17. 17.

    et al. Structural and functional features of eukaryotic mRNA untranslated regions. Gene 276, 73–81 (2001).

  18. 18.

    , & Translational control by the 3′-UTR: the ends specify the means. Trends Biochem. Sci. 28, 91–98 (2003).

  19. 19.

    , & The evolution of transcription-initiation sites. Mol. Biol. Evol. 22, 1137–1146 (2005).

  20. 20.

    The scanning mechanism of eukaryotic translation initiation. Annu. Rev. Biochem. 83, 779–812 (2014).

  21. 21.

    , & Translational control by 5′-untranslated regions of eukaryotic mRNAs. Science 352, 1413–1416 (2016).

  22. 22.

    , & Upstream open reading frames cause widespread reduction of protein expression and are polymorphic among humans. Proc. Natl Acad. Sci. USA 106, 7507–7512 (2009).

  23. 23.

    , & Tuning gene expression with synthetic upstream open reading frames. Proc. Natl Acad. Sci. USA 110, 11284–11289 (2013).

  24. 24.

    , & A perspective on mammalian upstream open reading frame function. Int. J. Biochem. Cell Biol. 45, 1690–1700 (2013).

  25. 25.

    et al. Before it gets started: regulating translation at the 5′ UTR. Comp. Funct. Genomics 2012, 1–8 (2012).

  26. 26.

    Point mutations define a sequence flanking the AUG initiator codon that modulates translation by eukaryotic ribosomes. Cell 44, 283–292 (1986).

  27. 27.

    et al. Quantitative analysis of mammalian translation initiation sites by FACS-seq. Mol. Syst. Biol. 10, 1–14 (2014).

  28. 28.

    , & Origins and principles of translational control. Cold Spring Harbor Monogr. Ser. 48, 1–40 (2007).

  29. 29.

    et al. Translational regulation of specific mRNAs controls feedback inhibition and survival during macrophage activation. PLOS Genet. 10, e1004368 (2014).

  30. 30.

    , & Origins and evolution of the mechanisms regulating translation initiation in eukaryotes. Trends Biochem. Sci. 35, 63–73 (2010).

  31. 31.

    , , & Evolution of the Protein Synthesis Machinery and its Regulation (eds Hernández, G. & Jagus, R.) 81–107 (Springer Int. Pub. 2016).

  32. 32.

    , , & Untranslated regions of mRNAs. Genome Biol. 3, 4.1–4.10 (2002).

  33. 33.

    , , & Formation, regulation and evolution of Caenorhabditis elegans 3′UTRs. Nature 469, 97–101 (2011).

  34. 34.

    et al. The transcriptional landscape of the yeast genome defined by RNA sequencing. Science 320, 1344–1349 (2008).

  35. 35.

    & Insertion mutagenesis to increase secondary structure within the 5′ noncoding region of a eukaryotic mRNA reduces translational efficiency. Cell 40, 515–526 (1985). This is one of the first studies to show a role for a 5′ UTR RNA secondary structure in influencing mRNA translation.

  36. 36.

    & Regulation of rat ornithine decarboxylase mRNA translation by its 5′-untranslated region. J. Biol. Chem. 265, 11817–11822 (1990).

  37. 37.

    et al. RNA sequence context effects measured in vitro predict in vivo protein binding and regulation. Mol. Cell 64, 294–306 (2016).

  38. 38.

    , & Distinctive features of the 5′-terminal sequences of the human mitochondrial mRNAs. Nature 290, 465–470 (1981).

  39. 39.

    , & Cap-dependent, scanning-free translation initiation mechanisms. Biochim. Biophys. Acta 1849, 1313–1318 (2015).

  40. 40.

    et al. Identification of the iron responsive element for the translational regulation of human ferritin mRNA. Science 238, 1570–1573 (1987).

  41. 41.

    , , & A red carpet for iron metabolism. Cell 3, 1–18 (2017).

  42. 42.

    & Iron regulatory protein prevents binding of the 43S translation pre-initiation complex to ferritin and eALAS mRNAs. EMBO J. 13, 3882–3891 (1994).

  43. 43.

    , & IRP-1 binding to ferritin mRNA prevents the recruitment of the small ribosomal subunit by the cap-binding complex eIF4F. Mol. Cell 2, 383–388 (1998).

  44. 44.

    , , & Control of mammalian translation by mRNA structure near caps. RNA 12, 851–861 (2006).

  45. 45.

    Influences of mRNA secondary structure on initiation by eukaryotic ribosomes. Proc. Natl Acad. Sci. USA 83, 2850–2854 (1986).

  46. 46.

    et al. mRNA helicases: the tacticians of translational control. Nat. Rev. Mol. Cell Biol. 12, 235–245 (2011).

  47. 47.

    et al. RNA G-quadruplexes cause eIF4A-dependent oncogene translation in cancer. Nature 513, 65–70 (2014). Ribosome profiling in cancer cells following eIF4A perturbation identifies stable RG4 structures in the 5′ UTR in eIF4A-sensitive target mRNAs.

  48. 48.

    et al. Transcriptome-wide characterization of the eIF4A signature highlights plasticity in translation regulation. Genome Biol. 15, 1–19 (2014).

  49. 49.

    , & Rocaglates convert DEAD-box protein eIF4A into a sequence-selective translational repressor. Nature 534, 558–561 (2016).

  50. 50.

    , , & Duplex unwinding and ATPase activities of the DEAD-box helicase eIF4A are coupled by eIF4G and eIF4B. J. Mol. Biol. 412, 674–687 (2011).

  51. 51.

    et al. Efficient translation initiation directed by the 900-nucleotide-long and GC-rich 5′ untranslated region of the human retrotransposon LINE-1 mRNA is strictly cap dependent rather than internal ribosome entry site mediated. Mol. Cell. Biol. 27, 4685–4697 (2007).

  52. 52.

    et al. Therapeutic suppression of translation initiation modulates chemosensitivity in a mouse lymphoma model. J. Clin. Invest. 118, 2651–2660 (2008).

  53. 53.

    et al. Antitumor activity and mechanism of action of the cyclopenta[b]benzofuran, silvestrol. PLoS ONE 4, e5223 (2009).

  54. 54.

    et al. Evidence for a functionally relevant rocaglamide binding site on the eIF4A-RNA complex. ACS Chem. Biol. 8, 1519–1527 (2013).

  55. 55.

    & Targeting the eIF4A RNA helicase as an anti-neoplastic approach. Biochim. Biophys. Acta 1849, 781–791 (2014).

  56. 56.

    et al. NanoCAGE reveals 5′ UTR features that define specific modes of translation of functionally related MTOR-sensitive mRNAs. Genome Res. 26, 636–648 (2016).

  57. 57.

    et al. The malignant phenotype in breast cancer is driven by eIF4A1-mediated changes in the translational landscape. Cell Death Dis. 6, e1603 (2015).

  58. 58.

    , , & Genome-wide analysis of translational efficiency reveals distinct but overlapping functions of yeast DEAD-box RNA helicases Ded1 and eIF4A. Genome Res. 25, 1196–1205 (2015).

  59. 59.

    , , , & Translation initiation on mammalian mRNAs with structured 5′UTRs requires DexH-box protein DHX29. Cell 135, 1237–1250 (2008).

  60. 60.

    , , , & eIF4B stimulates translation of long mRNAs with structured 5′ UTRs and low closed-loop potential but weak dependence on eIF4G. Proc. Natl Acad. Sci. USA 113, 10464–10472 (2016).

  61. 61.

    et al. A novel translational control mechanism involving RNA structures within coding sequences. Genome Res. 27, 95–106 (2017).

  62. 62.

    , & DNA G-quadruplexes in the human genome: detection, functions and therapeutic potential. Nat. Rev. Mol. Cell Biol. 5, 279–284 (2017).

  63. 63.

    & RNA G-quadruplexes: emerging mechanisms in disease. Nucleic Acids Res. 45, 1584–1595 (2016).

  64. 64.

    , & RNA G-quadruplexes in biology: principles and molecular mechanisms. J. Mol. Biol. 429, 2127–2147 (2017).

  65. 65.

    & 5′-UTR RNA G-quadruplexes: translation regulation and targeting. Nucleic Acids Res. 40, 4727–4741 (2012).

  66. 66.

    , , & RNA G-quadruplexes and their potential regulatory roles in translation. Translation (Austin) 4, e1244031 (2016).

  67. 67.

    & 5′-UTR G-quadruplex structures acting as translational repressors. Nucleic Acids Res. 38, 7022–7036 (2010).

  68. 68.

    , , & Irregular G-quadruplexes found in the untranslated regions of human mRNAs influence translation. J. Biol. Chem. 291, 21751–21760 (2016).

  69. 69.

    , & Position and stability are determining factors for translation repression by an RNA G-quadruplex-forming sequence within the 5′ UTR of the NRAS proto-oncogene. Biochemistry 47, 12664–12669 (2008).

  70. 70.

    , & Predictable suppression of gene expression by 5′-UTR-based RNA quadruplexes. Nucleic Acids Res. 37, 6811–6817 (2009).

  71. 71.

    , , & An RNA G-quadruplex in the 5′ UTR of the NRAS proto-oncogene modulates translation. Nat. Chem. Biol. 3, 218–221 (2007).

  72. 72.

    et al. Inhibition of translation in living eukaryotic cells by an RNA G-quadruplex motif. RNA 14, 1290–1296 (2008).

  73. 73.

    & The role of G-quadruplex in RNA metabolism: involvement of FMRP and FMR2P. Biochimie 92, 919–926 (2010).

  74. 74.

    et al. Fragile X mental retardation protein targets G quartet mRNAs important for neuronal function. Cell 107, 489–499 (2001).

  75. 75.

    et al. Identification of consensus binding sites clarifies FMRP binding determinants. Nucleic Acids Res. 44, 6649–6659 (2016).

  76. 76.

    et al. The fragile X mental retardation protein binds specifically to its mRNA via a purine quartet motif. EMBO J. 20, 4803–4813 (2001).

  77. 77.

    et al. The G-quartet containing FMRP binding site in FMR1 mRNA is a potent exonic splicing enhancer. Nucleic Acids Res. 36, 4902–4912 (2008).

  78. 78.

    et al. FMRP interferes with the Rac1 pathway and controls actin cytoskeleton dynamics in murine fibroblasts. Hum. Mol. Genet. 14, 835–844 (2005).

  79. 79.

    et al. FMRP stalls ribosomal translocation on mRNAs linked to synaptic function and autism. Cell 146, 247–261 (2011).

  80. 80.

    , , , & Fragile X mental retardation protein regulates translation by binding directly to the ribosome. Mol. Cell 54, 407–417 (2014).

  81. 81.

    et al. The human CCHC-type zinc finger nucleic acid-binding protein binds G-rich elements in target mRNA coding sequences and promotes translation. Cell Rep. 18, 2979–2990 (2017).

  82. 82.

    , , , & Human interferon-γ mRNA autoregulates its translation through a pseudoknot that activates the interferon-inducible protein kinase PKR. Cell 108, 221–232 (2002).

  83. 83.

    et al. Dynamic refolding of IFN-γ mRNA enables it to function as PKR activator and translation template. Nat. Chem. Biol. 5, 896–903 (2009). The studies in Refs 82 and 83 show how short helical segments assemble into a pseudoknot structure in a 5′ UTR, which is bound by PKR and activates it, lead to inhibition of translation initiation.

  84. 84.

    , & Regulation of innate immunity through RNA structure and the protein kinase PKR. Curr. Opin. Struct. Biol. 21, 119–127 (2011).

  85. 85.

    , & Posttranscriptional gene regulation by long noncoding RNA. J. Mol. Biol. 425, 3723–3730 (2013).

  86. 86.

    et al. Long non-coding antisense RNA controls Uchl1 translation through an embedded SINEB2 repeat. Nature 491, 454–457 (2012).

  87. 87.

    , & RNA pseudoknots and the regulation of protein synthesis. Biochem. Soc. Trans. 36, 684–689 (2008).

  88. 88.

    , , & A cis-acting element in the 3′-untranslated region of human TNF-α mRNA renders splicing dependent on the activation of protein kinase PKR. Genes Dev. 13, 3280–3293 (1999).

  89. 89.

    et al. An ancient pseudoknot in TNF-α pre-mRNA activates PKR, inducing eIF2α phosphorylation that potently enhances splicing. Cell Rep. 20, 188–200 (2017).

  90. 90.

    Molecular determinants and guided evolution of species-specific RNA editing. Nature 434, 409–413 (2005).

  91. 91.

    Programmed translational frameshifting. Microbiol. Rev. 60, 103–134 (1996).

  92. 92.

    Mechanisms and implications of programmed translational frameshifting. Wiley Interdiscip. Rev. RNA 3, 661–673 (2012).

  93. 93.

    & Reprogramming the genetic code: the emerging role of ribosomal frameshifting in regulating cellular gene expression. Bioessays 38, 21–26 (2016).

  94. 94.

    , & Characterization of an efficient coronavirus ribosomal frameshifting signal: requirement for an RNA pseudoknot. Cell 57, 537–547 (1989).

  95. 95.

    , , & The frameshift stimulatory signal of human immunodeficiency virus type 1 group O is a pseudoknot. J. Mol. Biol. 331, 571–583 (2003).

  96. 96.

    , , , & A mechanical explanation of RNA pseudoknot function in programmed ribosomal frameshifting. Nature 441, 244–247 (2006).

  97. 97.

    , & Endogenous ribosomal frameshift signals operate as mRNA destabilizing elements through at least two molecular pathways in yeast. Nucleic Acids Res. 39, 2799–2808 (2011).

  98. 98.

    et al. Ribosomal frameshifting in the CCR5 mRNA is regulated by miRNAs and the NMD pathway. Nature 512, 265–269 (2014).

  99. 99.

    & Translational control in stress and apoptosis. Nat. Rev. Mol. Cell Biol. 6, 318–327 (2005).

  100. 100.

    & Preferential translation of internal ribosome entry site-containing mRNAs during the mitotic cycle in mammalian cells. J. Biol. Chem. 279, 13721–13728 (2004).

  101. 101.

    The current status of vertebrate cellular mRNA IRESs. Cold Spring Harb. Perspect. Biol. 5, a011569 (2013).

  102. 102.

    et al. Molecular mechanisms of translation initiation in eukaryotes. Proc. Natl Acad. Sci. USA 98, 7029–7036 (2001).

  103. 103.

    & Cellular internal ribosome entry segments: structures, trans-acting factors and regulation of gene expression. Oncogene 23, 3200–3207 (2004).

  104. 104.

    , , & Re-programming of translation following cell stress allows IRES-mediated translation to predominate. Biol. Cell 100, 27–38 (2008).

  105. 105.

    , & More than just scanning: the importance of cap-independent mRNA translation initiation for cellular stress response and cancer. Cell. Mol. Life Sci. 74, 1659–1680 (2016).

  106. 106.

    & Internal initiation of translation mediated by the 5′ leader of a cellular mRNA. Nature 353, 90–94 (1991). This study presents the first description of IRES activity in the 5′ UTR of a cellular mRNA.

  107. 107.

    , , , & Enhancement of IRES-mediated translation of the c-myc and BiP mRNAs by the poly(A) tail is independent of intact eIF4G and PABP. Mol. Cell 15, 925–935 (2004).

  108. 108.

    , , & Human cytomegalovirus induces the endoplasmic reticulum chaperone BiP through increased transcription and activation of translation by using the BiP internal ribosome entry site. J. Virol. 84, 11479–11486 (2010).

  109. 109.

    et al. IRESite — a tool for the examination of viral and cellular internal ribosome entry sites. Nucleic Acids Res. 38, D131–D136 (2010).

  110. 110.

    et al. Splicing mediates the activity of four putative cellular internal ribosome entry sites. Proc. Natl Acad. Sci. USA 105, 4733–4738 (2008).

  111. 111.

    & Internal ribosome entry sites in cellular mRNAs: mystery of their existence. J. Biol. Chem. 280, 23425–23428 (2005).

  112. 112.

    & Cellular IRES-mediated translation: the war of ITAFs in pathophysiological states. Cell Cycle 10, 229–240 (2011).

  113. 113.

    , , & A search for structurally similar cellular internal ribosome entry sites. Nucleic Acids Res. 35, 4664–4677 (2007).

  114. 114.

    et al. Identification of a motif that mediates polypyrimidine tract-binding protein-dependent internal ribosome entry. Genes Dev. 19, 1556–1571 (2005).

  115. 115.

    , & The role of IRES trans-acting factors in regulating translation initiation. Biochem. Soc. Trans. 38, 1581–1586 (2010).

  116. 116.

    & The role of IRES trans-acting factors in carcinogenesis. Biochim. Biophys. Acta 1849, 887–897 (2014).

  117. 117.

    & For IRES trans-acting factors, it is all about location. Oncogene 27, 1033–1035 (2008).

  118. 118.

    et al. Upregulated c-myc expression in multiple myeloma by internal ribosome entry results from increased interactions with and expression of PTB-1 and YB-1. Oncogene 29, 2884–2891 (2010).

  119. 119.

    , , , & The Apaf-1 internal ribosome entry segment attains the correct structural conformation for function via interactions with PTB and unr. Mol. Cell 11, 757–771 (2003).

  120. 120.

    , , & Polypyrimidine tract binding protein and poly r(C) binding protein 1 interact with the BAG-1 IRES and stimulates its activity in vitro and in vivo. Nucleic Acids Res. 31, 639–646 (2003).

  121. 121.

    , , , & Bag-1 internal ribosome entry segment activity is promoted by structural changes mediated by poly(rC) binding protein 1 and recruitment of polypyrimidine tract binding protein 1. Mol. Cell. Biol. 24, 5595–5605 (2004).

  122. 122.

    , , & Derivation of a structural model for the c-myc IRES. J. Mol. Biol. 310, 111–126 (2001).

  123. 123.

    et al. A mutation in the c-myc-IRES leads to enhanced internal ribosome entry in multiple myeloma: a novel mechanism of oncogene de-regulation. Oncogene 19, 4437–4440 (2000).

  124. 124.

    et al. The zipper model of translational control: a small upstream ORF is the switch that controls structural remodeling of an mRNA leader. Cell 113, 519–531 (2003).

  125. 125.

    et al. Ribosome stalling regulates ires-mediated translation in eukaryotes, a parallel to prokaryotic attenuation. Mol. Cell 17, 405–416 (2005).

  126. 126.

    et al. Overexpression of FGF9 in colon cancer cells is mediated by hypoxia-induced translational activation. Nucleic Acids Res. 42, 2932–2944 (2014).

  127. 127.

    et al. An upstream open reading frame within an IRES controls expression of a specific VEGF-A isoform. Nucleic Acids Res. 36, 2434–2445 (2008).

  128. 128.

    et al. Regulation of internal ribosome entry site-mediated translation by eukaryotic initiation factor-2a phosphorylation and translation of a small upstream open reading frame. J. Biol. Chem. 277, 2050–2058 (2002).

  129. 129.

    , , , & An RNA G-quadruplex is essential for cap-independent translation initiation in human VEGF IRES. J. Am. Chem. Soc. 132, 17831–17839 (2010).

  130. 130.

    et al. Stabilization of the G-quadruplex at the VEGF IRES represses cap-independent translation. RNA Biol. 12, 320–329 (2015).

  131. 131.

    et al. A single internal ribosome entry site containing a G quartet RNA structure drives fibroblast growth factor 2 gene expression at four alternative translation initiation codons. J. Biol. Chem. 278, 39330–39336 (2003).

  132. 132.

    , & An independently folding RNA G-quadruplex domain directly recruits the 40S ribosomal subunit. Biochemistry 54, 1879–1885 (2015).

  133. 133.

    et al. RNA regulons in Hox 5′ UTRs confer ribosome specificity to gene regulation. Nature 517, 33–38 (2015). The first targeted knockout of a cellular IRES in mice reveals a key role for IRES-driven translation of Hox mRNAs in vivo and identifies a new cis-regulatory element, the TIE, which blocks cap-dependent translation.

  134. 134.

    , & RPS25 is essential for translation initiation by the Dicistroviridae and hepatitis C viral IRESs. Genes Dev. 23, 2753–2764 (2009).

  135. 135.

    et al. rRNA pseudouridylation defects affect ribosomal ligand binding and translational fidelity from yeast to human cells. Mol. Cell 44, 660–666 (2011).

  136. 136.

    et al. Ribosome-mediated specificity in Hox mRNA translation and vertebrate tissue patterning. Cell 145, 383–397 (2011).

  137. 137.

    et al. Internal ribosome entry site structural motifs conserved among mammalian fibroblast growth factor 1 alternatively spliced mRNAs. Mol. Cell. Biol. 24, 7622–7635 (2004).

  138. 138.

    et al. Consistent global structures of complex RNA states through multidimensional chemical mapping. eLife 4, 1–38 (2015).

  139. 139.

    , , & Fibroblast growth factor 2 internal ribosome entry site (IRES) activity ex vivo and in transgenic mice reveals a stringent tissue-specific regulation. J. Cell Biol. 150, 275–281 (2000).

  140. 140.

    , , , & c-Myc internal ribosome entry site activity is developmentally controlled and subjected to a strong translational repression in adult transgenic mice. Mol. Cell. Biol. 21, 1833–1840 (2001).

  141. 141.

    et al. Hypoxia induces VEGF-C expression in metastatic tumor cells via a HIF-1a-independent translation-mediated mechanism. Cell Rep. 6, 155–167 (2014).

  142. 142.

    et al. Potent activation of FGF-2 IRES-dependent mechanism of translation during brain development. RNA 14, 1852–1864 (2008).

  143. 143.

    et al. FGF2 translationally induced by hypoxia is involved in negative and positive feedback loops with HIF-1α. PLOS ONE 3, e3078 (2008).

  144. 144.

    et al. Systematic discovery of cap-independent translation sequences in human and viral genomes. Science 351, 1–24 (2016). This study reports the first genome-wide approach to identify novel IRES-like elements in vivo and finds short poly(U) sequences and small structured elements that harbour IRES activity.

  145. 145.

    , & eIF3 targets cell-proliferation messenger RNAs for translational activation or repression. Nature 522, 111–114 (2015). This study identifies a novel role for eIF3 in interacting with RNA stem–loop structures in the 5′ UTR of mRNAs to directly recruit the ribosome.

  146. 146.

    et al. Aberrant expression of c-Jun in glioblastoma by internal ribosome entry site (IRES)-mediated translational activation. Proc. Natl Acad. Sci. USA 109, E2875–E2884 (2012).

  147. 147.

    , , & eIF3d is an mRNA cap-binding protein that is required for specialized translation initiation. Nature 536, 96–99 (2016).

  148. 148.

    , & Post-transcriptional gene regulation by mRNA modifications. Nat. Rev. Mol. Cell Biol. 18, 31–42 (2016).

  149. 149.

    , & Chemical and structural effects of base modifications in messenger RNA. Nature 541, 339–346 (2017).

  150. 150.

    et al. Topology of the human and mouse m6A RNA methylomes revealed by m6A-seq. Nature 485, 201–206 (2012).

  151. 151.

    et al. 5′ UTR m6A promotes cap-independent translation. Cell 163, 999–1010 (2015).

  152. 152.

    et al. Dynamic m(6)A mRNA methylation directs translational control of heat shock response. Nature 526, 591–594 (2015). Refs 151 and 152 demonstrate that m6A promotes cap-independent mRNA translation initiation, which is mediated by direct eIF3 recruitment. They also show that m6A levels in 5′ UTRs are increased during stress.

  153. 153.

    et al. N(6)-methyladenosine-dependent RNA structural switches regulate RNA-protein interactions. Nature 518, 560–564 (2015). This study reports that the incorporation of m6A into a hairpin modulates the local secondary structure of RNA ('m6A switch'), thereby facilitating the binding of the indirect m6A reader hnRNPC.

  154. 154.

    et al. Structural imprints in vivo decode RNA regulatory mechanisms. Nature 519, 486–490 (2015). This paper presents the first global in vivo RNA structure probing method, icSHAPE, and connects m6A modifications with destabilization of local RNA structure across the transcriptome.

  155. 155.

    et al. High-Resolution mapping reveals a conserved, widespread, dynamic mRNA methylation program in yeast meiosis. Cell 155, 1409–1421 (2013).

  156. 156.

    et al. Structure and thermodynamics of N6-methyladenosine in RNA: a spring-loaded base modification. J. Am. Chem. Soc. 137, 2107–2115 (2015).

  157. 157.

    et al. N6-methyladenosine-dependent regulation of messenger RNA stability. Nature 505, 117–120 (2014).

  158. 158.

    et al. N6-methyladenosine modulates messenger RNA translation efficiency. Cell 161, 1388–1399 (2015).

  159. 159.

    , , , & The m6A methyltransferase METTL3 promotes translation in human cancer cells. Mol. Cell 62, 335–345 (2015).

  160. 160.

    et al. The dynamic N1-methyladenosine methylome in eukaryotic messenger RNA. Nature 530, 441–446 (2016).

  161. 161.

    et al. Transcriptome-wide distribution and function of RNA hydroxymethylcytosine. Science 351, 282–285 (2016).

  162. 162.

    et al. Transcription impacts the efficiency of mRNA translation via co-transcriptional N6-adenosine methylation. Cell 169, 326–337 (2017).

  163. 163.

    et al. N(6)-methyladenosine in mRNA disrupts tRNA selection and translation-elongation dynamics. Nat. Struct. Mol. Biol. 23, 110–115 (2016).

  164. 164.

    et al. Circ-ZNF609 is a circular RNA that can be translated and functions in myogenesis. Mol. Cell 66, 22–37 (2017).

  165. 165.

    et al. Translation of CircRNAs. Mol. Cell 66, 9–21 (2017).

  166. 166.

    et al. Genome-wide maps of m6A circRNAs identify widespread and cell-type-specific methylation patterns that are distinct from mRNAs. Cell Rep. 20, 2262–2276 (2017).

  167. 167.

    , & Insights into RNA structure and function from genome-wide studies. Nat. Rev. Genet. 15, 469–479 (2014).

  168. 168.

    , , & The RNA structurome: transcriptome-wide structure probing with next-generation sequencing. Trends Biochem. Sci. 40, 221–232 (2015).

  169. 169.

    , & Progress and challenges for chemical probing of RNA structure inside living cells. Nat. Chem. Biol. 11, 933–941 (2015).

  170. 170.

    , , & Genome-wide analysis of RNA secondary structure. Annu. Rev. Genet. 50, 235–266 (2016).

  171. 171.

    & The RNA epistructurome: uncovering RNA function by studying structure and post-transcriptional modifications. Trends Biotechnol. 35, 318–333 (2017).

  172. 172.

    , , , & Genome-wide probing of RNA structure reveals active unfolding of mRNA structures in vivo. Nature 505, 701–705 (2014).

  173. 173.

    et al. In vivo genome-wide profiling of RNA secondary structure reveals novel regulatory features. Nature 505, 696–700 (2014). Refs 172 and 173 introduce the first methods for in vivo DMS-based chemical modification of accessible RNA nucleotides to globally probe RNA secondary structure.

  174. 174.

    et al. Genome-wide measurement of RNA secondary structure in yeast. Nature 467, 103–107 (2010).

  175. 175.

    , , , & RNA motif discovery by SHAPE and mutational profiling (SHAPE-MaP). Nat. Methods 11, 959–965 (2014).

  176. 176.

    , , & Probing xist RNA structure in cells using targeted structure-seq. PLoS Genet. 11, e1005668 (2015).

  177. 177.

    et al. DMS-MaPseq for genome-wide or targeted RNA structure probing in vivo. Nat. Methods 14, 75–82 (2016). At the current vanguard of in vivo chemical mapping techniques, DMS–MaPseq leverages several methodological advances to pilot applications to probe low-abundance mRNAs and pre-mRNAs and to derive structural models that inform compensatory mutagenesis experiments.

  178. 178.

    et al. Single-molecule correlated chemical probing of RNA. Proc. Natl Acad. Sci. USA 111, 13858–13863 (2014).

  179. 179.

    , & Chemical methods for the structural study of the ribosome: judgment day. Mol. Biol. 35, 472–495 (2001).

  180. 180.

    et al. RNA-puzzles round III: 3D RNA structure prediction of five riboswitches and one ribozyme. RNA 23, 655–672 (2017).

  181. 181.

    , , & A two-dimensional mutate-and-map strategy for non-coding RNA structure. Nat. Chem. 3, 954–962 (2011).

  182. 182.

    , , & High-throughput mutate-map-rescue evaluates SHAPE-directed RNA structure and uncovers excited states. RNA 20, 1815–1826 (2014).

  183. 183.

    & RNA structure through multidimensional chemical mapping. Q. Rev. Biophys. 49, 1–30 (2016).

  184. 184.

    , & Detection of RNA-protein interactions in living cells with SHAPE. Biochemistry 54, 6867–6875 (2015).

  185. 185.

    , , & RNA structure inference through chemical mapping after accidental or intentional mutations. Proc. Natl Acad. Sci. USA 114, 9876–9881 (2017).

  186. 186.

    , , & Variable chromatin structure revealed by in situ spatially correlated DNA cleavage mapping. Nature 541, 237–241 (2016).

  187. 187.

    et al. Operon mRNAs are organized into ORF-centric structures that predict translation efficiency. eLife 6, e22037 (2017).

  188. 188.

    , , & Messenger-RNA-binding proteins and the messages they carry. Nat. Rev. Mol. Cell Biol. 3, 195–205 (2002).

  189. 189.

    et al. A compendium of RNA-binding motifs for decoding gene regulation. Nature 499, 172–177 (2013).

  190. 190.

    , , & A sequence-independent, unstructured IRES is responsible for internal expression of the coat protein of Turnip Crinkle Virus. J. Virol. 91, e02421–e02416 (2017).

  191. 191.

    , & A statistical test for conserved RNA structure shows lack of evidence for structure in lncRNAs. Nat. Methods 14, 45–48 (2017).

  192. 192.

    et al. Identification and classification of conserved RNA secondary structures in the human genome. PLoS Comput. Biol. 2, 251–262 (2006).

  193. 193.

    et al. New families of human regulatory RNA structures identified by comparative analysis of vertebrate genomes. Genome Res. 21, 1929–1943 (2011).

  194. 194.

    et al. Systematic discovery of structural elements governing stability of mammalian messenger RNAs. Nature 485, 264–268 (2012).

  195. 195.

    , , , & Translation initiation of ornithine decarboxylase and nucleocytoplasmic transport of cyclin D1 mRNA are increased in cells overexpressing eukaryotic initiation factor 4E. Proc. Natl Acad. Sci. USA 93, 1065–1070 (1996).

  196. 196.

    Structural insights into the mechanism of scanning and start codon recognition in eukaryotic translation initiation. Trends Biochem. Sci. 1348, 1–23 (2017).

  197. 197.

    et al. Ribosomal 18S rRNA base pairs with mRNA during eukaryotic translation initiation. Nat. Commun. 7, 1–7 (2016).

  198. 198.

    et al. Visualizing the molecular sociology at the HeLa cell nuclear periphery. Science 351, 969–972 (2016).

  199. 199.

    Computational analysis of conserved RNA secondary structure in transcriptomes and genomes. Annu. Rev. Biophys. 43, 433–456 (2014).

  200. 200.

    et al. The hnRNA-binding proteins hnRNP L and PTB are required for efficient translation of the Cat-1 arginine/lysine transporter mRNA during amino acid starvation. Mol. Cell. Biol. 29, 2899–2912 (2009).

  201. 201.

    et al. A majority of m6A residues are in the last exons, allowing the potential for 3′ UTR regulation. Genes Dev. 29, 2037–2053 (2015).

Download references

Acknowledgements

The authors apologize to those researchers whose work was not cited owing to space limitations and thank G. W. Byeon in the Barna laboratory for data analysis and figure preparation of eukaryotic 5′ UTR lengths, the Barna laboratory members for helpful comments on the manuscript, and H.-G. Wendel, J. Kieft, P. Sarnow and M. Hentze for thoughtful suggestions. RNA research in the authors' laboratories is supported by the New York Stem Cell Foundation (M.B.), Mallinckrodt Foundation Award (M.B.), Pew Scholars Award (M.B.) and NIH grants R01HD086634 (M.B.), R21HD086730 (M.B.), R01 GM102519 (R.D.) and R35 GM122579 (R.D.). M.B. is a New York Stem Cell Foundation Robertson Investigator. K.L. is supported by an EMBO Long-Term Fellowship (ALTF 539–2015) and is the Layton Family Fellow of the Damon Runyon Cancer Research Foundation (DRG-2237-15).

Author information

Affiliations

  1. Department of Developmental Biology, Stanford University, Stanford, California 94305, USA.

    • Kathrin Leppek
    •  & Maria Barna
  2. Department of Genetics, Stanford University, Stanford, California 94305, USA.

    • Kathrin Leppek
    •  & Maria Barna
  3. Departments of Biochemistry and Physics, Stanford University, Stanford, California 94305, USA.

    • Rhiju Das

Authors

  1. Search for Kathrin Leppek in:

  2. Search for Rhiju Das in:

  3. Search for Maria Barna in:

Contributions

All authors substantially contributed to researching data for the article, discussion of the content, writing and editing and/or reviewing the manuscript before submission.

Competing interests

The authors declare no competing financial interests.

Corresponding authors

Correspondence to Rhiju Das or Maria Barna.

Supplementary information

PDF files

  1. 1.

    Supplementary information S1 (box)

    Genome-wide RNA-structure probing technologies

  2. 2.

    Supplementary information S2 (box)

    Viral IRESs and systematic IRES discovery

  3. 3.

    Supplementary information S3 (table)

    Experimental probing of cellular IRES RNA structures

Glossary

Secondary structures

Patterns of Watson–Crick base pairs (G–C, A–U and G–U) that define the double helices of an RNA.

Tertiary structures

Interactions that orient RNA double helices into specific three-dimensional arrangements, often involving non-Watson–Crick base pairs.

Ribozymes

RNA molecules that catalyse specific biochemical reactions.

Riboswitches

Non-coding mRNA structures that sense the environmental status of a cell by directly binding to small molecule ligands, such as a metabolite or an ion. This interaction changes the RNA conformation and controls gene expression through alternative splicing, transcription or translation.

Peptidyl transferase centre

The site in the large ribosomal subunit that catalyses peptide bond formation and peptide release; it is composed entirely of RNA.

Pseudoknots

RNA tertiary structures formed by base pairing of a single-stranded loop of a hairpin with a complementary sequence outside that hairpin.

Upstream open reading frames

(uORFs). Small ORFs located in the 5′ UTR of some mRNAs. Translation of uORFs can regulate the translation of the downstream ORF.

Kozak sequence

A favourable sequence (GCCRCCAUGG in mammals, where R is a purine) surrounding the translation start codon (AUG); also called the Kozak consensus or Kozak context.

GC content

Percentage of guanine and cytosine nucleotides in an RNA molecule. G–C base pairs are more stable than A–U base pairs and can form more stable RNA structures.

Free energy

G). The stability of an RNA secondary structure, estimated as the sum of the free energies assigned to all loops and base pair stacks of a folded RNA, based on a computational folding algorithm.

5′ cap

A modified guanine nucleotide, m7GpppN (where m7G is 7-methylguanosine, p is a phosphate group and N is any base), located at the 5′ end of eukaryotic mRNAs.

Ribonucleoprotein

(RNP). A complex of proteins in association with an RNA (RNP) or mRNA (mRNP).

Ribosome profiling

Sequencing of the RNA fragments protected by ribosomes, providing a quantitative signature of the translated mRNAs at a given time.

RNA G-quadruplex

(RG4). Extremely stable RNA structure formed in G-rich regions by the stacking of at least two G-tetrads, each of them forming a square-shaped structure by non-Watson–Crick interactions between two or more layers of paired G-quartets.

Toeprinting

Nucleotide-resolution assay that uses primer extension inhibition induced by the complex of a protein, or the ribosome, bound to an mRNA, to report its position on the mRNA.

In-line probing

Nucleotide-resolution structure probing method that uses the natural instability of RNA: the 2′ hydroxyl of each nucleotide can attack its phosphodiester backbone at a rate dependent on local structure.

No-go decay

An mRNA and translation quality-control mechanism that recognizes and degrades mRNAs following prolonged ribosome stalling during translation.

Nonsense-mediated decay

An RNA surveillance mechanism that recognizes and degrades mRNAs containing premature termination codons to prevent their translation.

Selective 2′-hydroxyl acylation analysed by primer extension

(SHAPE). SHAPE and in vivo click SHAPE (icSHAPE) use cell-permeable reagents to acylate the 2′-OH of accessible, single-stranded RNA at all four nucleotide bases.

Mutate-and-map

Two-dimensional expansion of chemical probing that infers RNA base pairing by mutating each nucleotide and detecting increased chemical accessibility at other nucleotides.

m6A switches

mRNA sequences that adopt a different secondary structure depending on N6-adenosine methylation.

Dimethyl sulfate

(DMS). A cell-permeable reagent that methylates adenine and cytosine nucleotides that are not protected by base pairing. Modification sites stall or induce a mutation during primer extension by reverse transcriptase, and sequencing the resulting complementary DNA indicates the folding state of a nucleotide in an RNA.

Psoralen

A natural plant product that intercalates into DNA and RNA and reversibly crosslinks interacting RNA duplex molecules at UpA motifs upon irradiation with ultraviolet light.

Sequence covariation

Nucleotide substitutions that differ between two or more homologous genes but retain the potential for RNA base pairing in each sequence.