Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

Augmented genetic decoding: global, local and temporal alterations of decoding processes and codon meaning

Key Points

  • Alternative genetic decoding can be either a global phenomenon that affects the decoding of most or all mRNAs, or a local recoding that affects particular sites in a few mRNAs.

  • Global alterations in the genetic code are usually caused by changes in the translation machinery.

  • Recoding is observed at specific mRNA locations and can be modulated by a variety of cis-elements and trans-factors.

  • Diverse recoding phenomena include the incorporation of non-standard amino acids, ribosomes that shift between reading frames or that bypass large untranslated gaps, and the trans-translation of two mRNA molecules for the synthesis of a single polypeptide.

  • Although only a small number of genes require recoding mechanisms for their expression, these genes probably occur in all organisms and are usually evolutionary conserved.

  • For genes that do not require altered decoding for expression, recoding may still have regulatory importance.

Abstract

The non-universality of the genetic code is now widely appreciated. Codes differ between organisms, and certain genes are known to alter the decoding rules in a site-specific manner. Recently discovered examples of decoding plasticity are particularly spectacular. These examples include organisms and organelles with disruptions of triplet continuity during the translation of many genes, viruses that alter the entire genetic code of their hosts and organisms that adjust their genetic code in response to changing environments. In this Review, we outline various modes of alternative genetic decoding and expand existing terminology to accommodate recently discovered manifestations of this seemingly sophisticated phenomenon.

This is a preview of subscription content, access via your institution

Access options

Buy this article

Prices may be subject to local taxes which are calculated during checkout

Figure 1: Freezing and melting of genetic decoding.
Figure 2: Components that shape alternative genetic decoding.
Figure 3: Relationship between nucleotide sequences and alternatively decoded proteins.

Similar content being viewed by others

References

  1. Crick, F. H. The origin of the genetic code. J. Mol. Biol. 38, 367–379 (1968). This is a seminal paper in which the frozen accident hypothesis of the origin of the genetic code was formulated.

    CAS  PubMed  Google Scholar 

  2. Riyasaty, S. & Atkins, J. F. External suppression of a frameshift mutant in salmonella. J. Mol. Biol. 34, 541–557 (1968).

    CAS  PubMed  Google Scholar 

  3. Weiner, A. M. & Weber, K. Natural read-through at the UGA termination signal of Q-β coat protein cistron. Nat. New Biol. 234, 206–209 (1971).

    CAS  PubMed  Google Scholar 

  4. Barrell, B. G., Bankier, A. T. & Drouin, J. A different genetic code in human mitochondria. Nature 282, 189–194 (1979).

    CAS  PubMed  Google Scholar 

  5. Yamao, F. et al. UGA is read as tryptophan in Mycoplasma capricolum. Proc. Natl Acad. Sci. USA 82, 2306–2309 (1985).

    CAS  PubMed  PubMed Central  Google Scholar 

  6. Horowitz, S. & Gorovsky, M. A. An unusual genetic code in nuclear genes of Tetrahymena. Proc. Natl Acad. Sci. USA 82, 2452–2455 (1985).

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Preer, J. R. Jr., Preer, L. B., Rudman, B. M. & Barnett, A. J. Deviation from the universal code shown by the gene for surface protein 51A in Paramecium. Nature 314, 188–190 (1985).

    CAS  PubMed  Google Scholar 

  8. Clare, J. & Farabaugh, P. Nucleotide sequence of a yeast Ty element: evidence for an unusual mechanism of gene expression. Proc. Natl Acad. Sci. USA 82, 2829–2833 (1985).

    CAS  PubMed  PubMed Central  Google Scholar 

  9. Jacks, T. & Varmus, H. E. Expression of the Rous sarcoma virus pol gene by ribosomal frameshifting. Science 230, 1237–1242 (1985).

    CAS  PubMed  Google Scholar 

  10. Mellor, J. et al. A retrovirus-like strategy for expression of a fusion protein encoded by yeast transposon Ty1. Nature 313, 243–246 (1985).

    CAS  PubMed  Google Scholar 

  11. Chambers, I. et al. The structure of the mouse glutathione peroxidase gene: the selenocysteine in the active site is encoded by the 'termination' codon, TGA. EMBO J. 5, 1221–1227 (1986).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. Craigen, W. J. & Caskey, C. T. Expression of peptide chain release factor 2 requires high-efficiency frameshift. Nature 322, 273–275 (1986).

    CAS  PubMed  Google Scholar 

  13. Huang, W. M. et al. A persistent untranslated sequence within bacteriophage T4 DNA topoisomerase gene 60. Science 239, 1005–1012 (1988).

    CAS  PubMed  Google Scholar 

  14. Matsufuji, S. et al. Autoregulatory frameshifting in decoding mammalian ornithine decarboxylase antizyme. Cell 80, 51–60 (1995).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. Keiler, K. C., Waller, P. R. & Sauer, R. T. Role of a peptide tagging system in degradation of proteins synthesized from damaged messenger RNA. Science 271, 990–993 (1996). This work reveals the function of tmRNA that rescues ribosomes from truncated mRNA lacking stop codons through the process of trans -translation.

    CAS  PubMed  Google Scholar 

  16. Srinivasan, G., James, C. M. & Krzycki, J. A. Pyrrolysine encoded by UAG in Archaea: charging of a UAG-decoding specialized tRNA. Science 296, 1459–1462 (2002). This work describes the discovery of twenty-second proteinogenic amino acid.

    CAS  PubMed  Google Scholar 

  17. Jungreis, I. et al. Evidence of abundant stop codon readthrough in Drosophila and other metazoa. Genome Res. 21, 2096–2113 (2011). This work reveals signatures of protein-coding evolution in the sequences downstream of stop codons in many genes in Drosophila species. This is strong evidence that stop codon readthrough has important functional roles in these organisms.

    CAS  PubMed  PubMed Central  Google Scholar 

  18. Prat, L. et al. Carbon source-dependent expansion of the genetic code in bacteria. Proc. Natl Acad. Sci. USA 109, 21070–21075 (2012). This paper describes a microorganism that uses alternative genetic codes depending on its environment.

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Lang, B. F. et al. Massive programmed translational jumping in mitochondria. Proc. Natl Acad. Sci. USA 111, 5926–5931 (2014). From 1988 until the publication of this paper in 2014, only a single naturally occurring case of translational bypassing was known. In this work the authors describe numerous cases of such bypassing that occur during the translation of mRNA in the mitochondria of the yeast Magnusiomyces capitatus.

    CAS  PubMed  PubMed Central  Google Scholar 

  20. Knight, R. D., Freeland, S. J. & Landweber, L. F. Rewiring the keyboard: evolvability of the genetic code. Nat. Rev. Genet. 2, 49–58 (2001).

    CAS  PubMed  Google Scholar 

  21. Ambrogelly, A., Palioura, S. & Soll, D. Natural expansion of the genetic code. Nat. Chem. Biol. 3, 29–35 (2007).

    CAS  PubMed  Google Scholar 

  22. Atkins, J. F. & Gesteland, R. F. Recoding: Expansion of Decoding Rules Enriches Gene Expression (Springer, 2010).

    Google Scholar 

  23. Sharma, V. et al. A pilot study of bacterial genes with disrupted ORFs reveals a surprising profusion of protein sequence recoding mediated by ribosomal frameshifting and transcriptional realignment. Mol. Biol. Evol. 28, 3195–3211 (2011). This work introduces a scheme for the large-scale identification of evolutionary conserved recoded genes in bacterial genomes.

    CAS  PubMed  PubMed Central  Google Scholar 

  24. Antonov, I., Coakley, A., Atkins, J. F., Baranov, P. V. & Borodovsky, M. Identification of the nature of reading frame transitions observed in prokaryotic genomes. Nucleic Acids Res. 41, 6514–6530 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  25. Antonov, I., Baranov, P. & Borodovsky, M. GeneTack database: genes with frameshifts in prokaryotic genomes and eukaryotic mRNA sequences. Nucleic Acids Res. 41, D152–D156 (2013).

    CAS  PubMed  Google Scholar 

  26. Ivanova, N. N. et al. Stop codon reassignments in the wild. Science 344, 909–913 (2014). In this work the authors used metagenome sequences to analyse the occurrence of stop codon reassignment in organisms from different environments. Among several intriguing findings was a bacteriophage that modifies the genetic code of its host during infection.

    CAS  PubMed  Google Scholar 

  27. Ohama, T. et al. Non-universal decoding of the leucine codon CUG in several Candida species. Nucleic Acids Res. 21, 4039–4045 (1993).

    CAS  PubMed  PubMed Central  Google Scholar 

  28. Duarte, I., Nabuurs, S. B., Magno, R. & Huynen, M. Evolution and diversification of the organellar release factor family. Mol. Biol. Evol. 29, 3497–3512 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. Bove, J. M. Molecular features of mollicutes. Clin. Infect. Dis. 17 (Suppl. 1), S10–S31 (1993).

    CAS  PubMed  Google Scholar 

  30. McCutcheon, J. P., McDonald, B. R. & Moran, N. A. Origin of an alternative genetic code in the extremely small and GC-rich genome of a bacterial symbiont. PLoS Genet. 5, e1000565 (2009).

    PubMed  PubMed Central  Google Scholar 

  31. Campbell, J. H. et al. UGA is an additional glycine codon in uncultured SR1 bacteria from the human microbiota. Proc. Natl Acad. Sci. USA 110, 5540–5545 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. Osawa, S. & Jukes, T. H. Codon reassignment (codon capture) in evolution. J. Mol. Evol. 28, 271–278 (1989).

    CAS  PubMed  Google Scholar 

  33. Chater, K. F. & Chandra, G. The use of the rare UUA codon to define “expression space” for genes involved in secondary metabolism, development and environmental adaptation in streptomyces. J. Microbiol. 46, 1–11 (2008).

    CAS  PubMed  Google Scholar 

  34. Nakabachi, A. et al. The 160-kilobase genome of the bacterial endosymbiont Carsonella. Science 314, 267 (2006).

    CAS  PubMed  Google Scholar 

  35. Schultz, D. W. & Yarus, M. On malleability in the genetic code. J. Mol. Evol. 42, 597–601 (1996).

    CAS  PubMed  Google Scholar 

  36. Suzuki, T., Ueda, T. & Watanabe, K. The 'polysemous' codon — a codon with multiple amino acid assignment caused by dual specificity of tRNA identity. EMBO J. 16, 1122–1134 (1997).

    CAS  PubMed  PubMed Central  Google Scholar 

  37. Santos, M. A., Ueda, T., Watanabe, K. & Tuite, M. F. The non-standard genetic code of Candida spp.: an evolving genetic code or a novel mechanism for adaptation? Mol. Microbiol. 26, 423–431 (1997).

    CAS  PubMed  Google Scholar 

  38. Netzer, N. et al. Innate immune and chemically triggered oxidative stress modifies translational fidelity. Nature 462, 522–526 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  39. Lee, J. Y. et al. Promiscuous methionyl-tRNA synthetase mediates adaptive mistranslation to protect cells against oxidative stress. J. Cell Sci. 127, 4234–4245 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  40. Namy, O. et al. Adding pyrrolysine to the Escherichia coli genetic code. FEBS Lett. 581, 5282–5288 (2007).

    CAS  PubMed  Google Scholar 

  41. Longstaff, D. G., Blight, S. K., Zhang, L., Green-Church, K. B. & Krzycki, J. A. In vivo contextual requirements for UAG translation as pyrrolysine. Mol. Microbiol. 63, 229–241 (2007).

    CAS  PubMed  Google Scholar 

  42. Zhang, Y., Baranov, P. V., Atkins, J. F. & Gladyshev, V. N. Pyrrolysine and selenocysteine use dissimilar decoding strategies. J. Biol. Chem. 280, 20740–20751 (2005).

    CAS  PubMed  Google Scholar 

  43. King, G. M. Methanogenesis from methylated amines in a hypersaline algal mat. Appl. Environ. Microbiol. 54, 130–136 (1988).

    CAS  PubMed  PubMed Central  Google Scholar 

  44. Atkins, J. F. & Baranov, P. V. The distinction between recoding and codon reassignment. Genetics 185, 1535–1536 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Firth, A. E. & Brierley, I. Non-canonical translation in RNA viruses. J. Gen. Virol. 93, 1385–1409 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  46. Dreher, T. W. & Miller, W. A. Translational control in positive strand RNA plant viruses. Virology 344, 185–197 (2006).

    CAS  PubMed  Google Scholar 

  47. Steneberg, P. & Samakovlis, C. A novel stop codon readthrough mechanism produces functional Headcase protein in Drosophila trachea. EMBO Rep. 2, 593–597 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  48. Namy, O. et al. Identification of stop codon readthrough genes in Saccharomyces cerevisiae. Nucleic Acids Res. 31, 2289–2296 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  49. Namy, O., Duchateau-Nguyen, G. & Rousset, J. P. Translational readthrough of the PDE2 stop codon modulates cAMP levels in Saccharomyces cerevisiae. Mol. Microbiol. 43, 641–652 (2002).

    CAS  PubMed  Google Scholar 

  50. Robinson, D. N. & Cooley, L. Examination of the function of two kelch proteins generated by stop codon suppression. Development 124, 1405–1417 (1997).

    CAS  PubMed  Google Scholar 

  51. Klagges, B. R. et al. Invertebrate synapsins: a single gene codes for several isoforms in Drosophila. J. Neurosci. 16, 3154–3165 (1996).

    CAS  PubMed  PubMed Central  Google Scholar 

  52. Dunn, J. G., Foo, C. K., Belletier, N. G., Gavis, E. R. & Weissman, J. S. Ribosome profiling reveals pervasive and regulated stop codon readthrough in Drosophila melanogaster. eLife 2, e01179 (2013). In this work the authors used ribosome profiling to analyse the protein-coding potential of the Drosophila melanogaster genome and confirmed the widespread occurrence of stop codon readthrough in this organism.

    PubMed  PubMed Central  Google Scholar 

  53. Loughran, G. et al. Evidence of efficient stop codon readthrough in four mammalian genes. Nucleic Acids Res. 42, 8928–8938 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  54. Eswarappa, S. M. et al. Programmed translational readthrough generates antiangiogenic VEGF-Ax. Cell 157, 1605–1618 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  55. Schueren, F. et al. Peroxisomal lactate dehydrogenase is generated by translational readthrough in mammals. eLife 3, e03640 (2014).

    PubMed  PubMed Central  Google Scholar 

  56. Stiebler, A. C. et al. Ribosomal readthrough at a short UGA stop codon context triggers dual localization of metabolic enzymes in fungi and animals. PLoS Genet. 10, e1004685 (2014).

    PubMed  PubMed Central  Google Scholar 

  57. Pavlov, M. Y. et al. A direct estimation of the context effect on the efficiency of termination. J. Mol. Biol. 284, 579–590 (1998).

    CAS  PubMed  Google Scholar 

  58. Bonetti, B., Fu, L., Moon, J. & Bedwell, D. M. The efficiency of translation termination is determined by a synergistic interplay between upstream and downstream sequences in Saccharomyces cerevisiae. J. Mol. Biol. 251, 334–345 (1995).

    CAS  PubMed  Google Scholar 

  59. Namy, O., Hatin, I. & Rousset, J. P. Impact of the six nucleotides downstream of the stop codon on translation termination. EMBO Rep. 2, 787–793 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  60. Skuzeski, J. M., Nichols, L. M., Gesteland, R. F. & Atkins, J. F. The signal for a leaky UAG stop codon in several plant viruses includes the two downstream codons. J. Mol. Biol. 218, 365–373 (1991).

    CAS  PubMed  Google Scholar 

  61. Firth, A. E., Wills, N. M., Gesteland, R. F. & Atkins, J. F. Stimulation of stop codon readthrough: frequent presence of an extended 3′ RNA structural element. Nucleic Acids Res. 39, 6679–6691 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  62. Lee, S. R. et al. Mammalian thioredoxin reductase: oxidation of the C-terminal cysteine/selenocysteine active site forms a thioselenide, and replacement of selenium with sulfur markedly reduces catalytic activity. Proc. Natl Acad. Sci. USA 97, 2521–2526 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  63. Zhong, L., Arner, E. S. & Holmgren, A. Structure and mechanism of mammalian thioredoxin reductase: the active site is a redox-active selenolthiol/selenenylsulfide formed from the conserved cysteine–selenocysteine sequence. Proc. Natl Acad. Sci. USA 97, 5854–5859 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  64. Heider, J., Baron, C. & Bock, A. Coding from a distance: dissection of the mRNA determinants required for the incorporation of selenocysteine into protein. EMBO J. 11, 3759–3766 (1992).

    CAS  PubMed  PubMed Central  Google Scholar 

  65. Berry, M. J., Banu, L., Harney, J. W. & Larsen, P. R. Functional characterization of the eukaryotic SECIS elements which direct selenocysteine insertion at UGA codons. EMBO J. 12, 3315–3322 (1993). This work describes discovery of the SECIS element, an RNA structure in 3′ UTRs of eukaryotic mRNAs that is required for Sec incorporation.

    CAS  PubMed  PubMed Central  Google Scholar 

  66. Howard, M. T. et al. Recoding elements located adjacent to a subset of eukaryal selenocysteine-specifying UGA codons. EMBO J. 24, 1596–1607 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  67. Howard, M. T., Moyle, M. W., Aggarwal, G., Carlson, B. A. & Anderson, C. B. A recoding element that stimulates decoding of UGA codons by Sec tRNA[Ser]Sec. RNA 13, 912–920 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  68. Squires, J. E. & Berry, M. J. Eukaryotic selenoprotein synthesis: mechanistic insight incorporating new factors and new functions for old factors. IUBMB Life 60, 232–235 (2008).

    CAS  PubMed  Google Scholar 

  69. Driscoll, D. M. & Copeland, P. R. Mechanism and regulation of selenoprotein synthesis. Annu. Rev. Nutr. 23, 17–40 (2003).

    CAS  PubMed  Google Scholar 

  70. Labunskyy, V. M., Hatfield, D. L. & Gladyshev, V. N. Selenoproteins: molecular pathways and physiological roles. Physiol. Rev. 94, 739–777 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  71. Yoshizawa, S. & Bock, A. The many levels of control on bacterial selenoprotein synthesis. Biochim. Biophys. Acta 1790, 1404–1414 (2009).

    CAS  PubMed  Google Scholar 

  72. Rother, M., Resch, A., Wilting, R. & Bock, A. Selenoprotein synthesis in archaea. Biofactors 14, 75–83 (2001).

    CAS  PubMed  Google Scholar 

  73. Gursinsky, T., Jager, J., Andreesen, J. R. & Sohling, B. A selDABC cluster for selenocysteine incorporation in Eubacterium acidaminophilum. Arch. Microbiol. 174, 200–212 (2000).

    CAS  PubMed  Google Scholar 

  74. Otero, L. et al. Adjustments, extinction, and remains of selenocysteine incorporation machinery in the nematode lineage. RNA 20, 1023–1034 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  75. Chapple, C. E. & Guigo, R. Relaxation of selective constraints causes independent selenoprotein extinction in insect genomes. PLoS ONE 3, e2968 (2008).

    PubMed  PubMed Central  Google Scholar 

  76. Hill, K. E., Lloyd, R. S., Yang, J. G., Read, R. & Burk, R. F. The cDNA for rat selenoprotein P contains 10 TGA codons in the open reading frame. J. Biol. Chem. 266, 10050–10053 (1991).

    CAS  PubMed  Google Scholar 

  77. Lobanov, A. V., Hatfield, D. L. & Gladyshev, V. N. Reduced reliance on the trace element selenium during evolution of mammals. Genome Biol. 9, R62 (2008).

    PubMed  PubMed Central  Google Scholar 

  78. Berry, M. & Howard, M. in Recoding: Expansion of Decoding Rules Enriches Gene Expression (eds Atkins, J. F. & Gesteland, R. F.) 29–52 (Springer New York, 2010).

    Google Scholar 

  79. Xu, X. M. et al. Targeted insertion of cysteine by decoding UGA codons with mammalian selenocysteine machinery. Proc. Natl Acad. Sci. USA 107, 21430–21434 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  80. Hoffman, D. C., Anderson, R. C., DuBois, M. L. & Prescott, D. M. Macronuclear gene-sized molecules of hypotrichs. Nucleic Acids Res. 23, 1279–1283 (1995).

    CAS  PubMed  PubMed Central  Google Scholar 

  81. Turanov, A. A. et al. Genetic code supports targeted insertion of two amino acids by one codon. Science 323, 259–261 (2009). This work describes a unique case of codon redefinition that involves a sense codon.

    CAS  PubMed  PubMed Central  Google Scholar 

  82. Lobanov, A. V., Kryukov, G. V., Hatfield, D. L. & Gladyshev, V. N. Is there a twenty third amino acid in the genetic code? Trends Genet. 22, 357–360 (2006).

    CAS  PubMed  Google Scholar 

  83. Fujita, M., Mihara, H., Goto, S., Esaki, N. & Kanehisa, M. Mining prokaryotic genomes for unknown amino acids: a stop-codon-based approach. BMC Bioinformatics 8, 225 (2007).

    PubMed  PubMed Central  Google Scholar 

  84. Baranov, P. V., Gesteland, R. F. & Atkins, J. F. Release factor 2 frameshifting sites in different bacteria. EMBO Rep. 3, 373–377 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. Bekaert, M., Atkins, J. F. & Baranov, P. V. ARFA: a program for annotating bacterial release factor genes, including prediction of programmed ribosomal frameshifting. Bioinformatics 22, 2463–2465 (2006).

    CAS  PubMed  Google Scholar 

  86. Ivanov, I. P. & Atkins, J. F. Ribosomal frameshifting in decoding antizyme mRNAs from yeast and protists to humans: close to 300 cases reveal remarkable diversity despite underlying conservation. Nucleic Acids Res. 35, 1842–1858 (2007). This is a comprehensive survey of ribosomal frameshifting sites and stimulatory elements in antizyme mRNAs occurring in species from a large phylogenetic spectrum of organisms.

    CAS  PubMed  PubMed Central  Google Scholar 

  87. Gupta, P., Kannan, K., Mankin, A. S. & Vazquez-Laslop, N. Regulation of gene expression by macrolide-induced ribosomal frameshifting. Mol. Cell 52, 629–642 (2013).

    CAS  PubMed  Google Scholar 

  88. Cobucci-Ponzano, B., Rossi, M. & Moracci, M. Translational recoding in archaea. Extremophiles 16, 793–803 (2012).

    CAS  PubMed  Google Scholar 

  89. Dinman, J. D. Control of gene expression by translational recoding. Adv. Protein Chem. Struct. Biol. 86, 129–149 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  90. Baranov, P. V., Gesteland, R. F. & Atkins, J. F. Recoding: translational bifurcations in gene expression. Gene 286, 187–201 (2002).

    CAS  PubMed  Google Scholar 

  91. Baranov, P. V., Fayet, O., Hendrix, R. W. & Atkins, J. F. Recoding in bacteriophages and bacterial IS elements. Trends Genet. 22, 174–181 (2006).

    CAS  PubMed  Google Scholar 

  92. Farabaugh, P. J. Programmed Alternative Reading of the Genetic Code (R. G. Landes, 1997).

    Google Scholar 

  93. Namy, O., Rousset, J. P., Napthine, S. & Brierley, I. Reprogrammed genetic decoding in cellular gene expression. Mol. Cell 13, 157–168 (2004).

    CAS  PubMed  Google Scholar 

  94. Firth, A. E., Bekaert, M. & Baranov, P. V. in Recoding: Expansion of Decoding Rules Enriches Gene Expression 435–461 (Springer New York, 2010).

    Google Scholar 

  95. Jiang, H. et al. Orsay virus utilizes ribosomal frameshifting to express a novel protein that is incorporated into virions. Virology 450–451, 213–221 (2014).

    PubMed  Google Scholar 

  96. Fang, Y. et al. Efficient -2 frameshifting by mammalian ribosomes to synthesize an additional arterivirus protein. Proc. Natl Acad. Sci. USA 109, E2920–E2928 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  97. Loughran, G., Firth, A. E. & Atkins, J. F. Ribosomal frameshifting into an overlapping gene in the 2B-encoding region of the cardiovirus genome. Proc. Natl Acad. Sci. USA 108, E1111–E1119 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  98. Firth, A. E., Blitvich, B. J., Wills, N. M., Miller, C. L. & Atkins, J. F. Evidence for ribosomal frameshifting and a novel overlapping gene in the genomes of insect-specific flaviviruses. Virology 399, 153–166 (2010).

    CAS  PubMed  Google Scholar 

  99. Melian, E. B. et al. NS1' of flaviviruses in the Japanese encephalitis virus serogroup is a product of ribosomal frameshifting and plays a role in viral neuroinvasiveness. J. Virol. 84, 1641–1647 (2010).

    CAS  PubMed  Google Scholar 

  100. Firth, A. E. & Atkins, J. F. A conserved predicted pseudoknot in the NS2A-encoding sequence of West Nile and Japanese encephalitis flaviviruses suggests NS1' may derive from ribosomal frameshifting. Virol. J. 6, 14 (2009).

    PubMed  PubMed Central  Google Scholar 

  101. Firth, A. E., Chung, B. Y., Fleeton, M. N. & Atkins, J. F. Discovery of frameshifting in Alphavirus 6K resolves a 20-year enigma. Virol. J. 5, 108 (2008).

    PubMed  PubMed Central  Google Scholar 

  102. Firth, A. E. et al. Ribosomal frameshifting used in influenza A virus expression occurs within the sequence UCC_UUU_CGU and is in the +1 direction. Open Biol. 2, 120109 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  103. Jagger, B. W. et al. An overlapping protein-coding region in influenza A virus segment 3 modulates the host response. Science 337, 199–204 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  104. Gao, X., Havecker, E. R., Baranov, P. V., Atkins, J. F. & Voytas, D. F. Translational recoding signals between gag and pol in diverse LTR retrotransposons. RNA 9, 1422–1430 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. Shigemoto, K. et al. Identification and characterisation of a developmentally regulated mammalian gene that utilises -1 programmed ribosomal frameshifting. Nucleic Acids Res. 29, 4079–4088 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  106. Wills, N. M., Moore, B., Hammer, A., Gesteland, R. F. & Atkins, J. F. A functional -1 ribosomal frameshift signal in the human paraneoplastic Ma3 gene. J. Biol. Chem. 281, 7082–7088 (2006).

    CAS  PubMed  Google Scholar 

  107. Lin, M. F. et al. Revisiting the protein-coding gene catalog of Drosophila melanogaster using 12 fly genomes. Genome Res. 17, 1823–1836 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  108. Baranov, P. V. et al. Programmed ribosomal frameshifting in the expression of the regulator of intestinal stem cell proliferation, adenomatous polyposis coli (APC). RNA Biol. 8, 637–647 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  109. Ingolia, N. T. Ribosome profiling: new views of translation, from single codons to genome scale. Nat. Rev. Genet. 15, 205–213 (2014).

    CAS  PubMed  Google Scholar 

  110. Michel, A. M. et al. Observation of dually decoded regions of the human genome using ribosome profiling data. Genome Res. 22, 2219–2229 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  111. Shah, A. A. et al. Computational identification of putative programmed translational frameshift sites. Bioinformatics 18, 1046–1053 (2002).

    CAS  PubMed  Google Scholar 

  112. Gurvich, O. L. et al. Sequences that direct significant levels of frameshifting are frequent in coding regions of Escherichia coli. EMBO J. 22, 5941–5950 (2003).

    CAS  PubMed  PubMed Central  Google Scholar 

  113. Jacobs, J. L., Belew, A. T., Rakauskaite, R. & Dinman, J. D. Identification of functional, endogenous programmed -1 ribosomal frameshift signals in the genome of Saccharomyces cerevisiae. Nucleic Acids Res. 35, 165–174 (2007).

    CAS  PubMed  Google Scholar 

  114. Sharma, V. et al. Analysis of tetra- and hepta-nucleotides motifs promoting -1 ribosomal frameshifting in Escherichia coli. Nucleic Acids Res. 42, 7210–7225 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  115. Belew, A. T., Hepler, N. L., Jacobs, J. L. & Dinman, J. D. PRFdb: a database of computationally predicted eukaryotic programmed -1 ribosomal frameshift signals. BMC Genomics 9, 339 (2008).

    PubMed  PubMed Central  Google Scholar 

  116. Belew, A. T., Advani, V. M. & Dinman, J. D. Endogenous ribosomal frameshift signals operate as mRNA destabilizing elements through at least two molecular pathways in yeast. Nucleic Acids Res. 39, 2799–2808 (2011).

    CAS  PubMed  Google Scholar 

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

    CAS  PubMed  PubMed Central  Google Scholar 

  118. Weiss, R. B., Dunn, D. M., Dahlberg, A. E., Atkins, J. F. & Gesteland, R. F. Reading frame switch caused by base-pair formation between the 3′ end of 16S rRNA and the mRNA during elongation of protein synthesis in Escherichia coli. EMBO J. 7, 1503–1507 (1988).

    CAS  PubMed  PubMed Central  Google Scholar 

  119. Larsen, B., Wills, N. M., Gesteland, R. F. & Atkins, J. F. rRNA-mRNA base pairing stimulates a programmed -1 ribosomal frameshift. J. Bacteriol. 176, 6842–6851 (1994).

    CAS  PubMed  PubMed Central  Google Scholar 

  120. Prere, M. F., Canal, I., Wills, N. M., Atkins, J. F. & Fayet, O. The interplay of mRNA stimulatory signals required for AUU-mediated initiation and programmed -1 ribosomal frameshifting in decoding of transposable element IS911. J. Bacteriol. 193, 2735–2744 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. Gurvich, O. L., Nasvall, S. J., Baranov, P. V., Bjork, G. R. & Atkins, J. F. Two groups of phenylalanine biosynthetic operon leader peptides genes: a high level of apparently incidental frameshifting in decoding Escherichia coli pheL. Nucleic Acids Res. 39, 3079–3092 (2011).

    CAS  PubMed  Google Scholar 

  122. Yordanova, M. M., Wu, C., Andreev, D. E., Sachs, M. S. & Atkins, J. F. A nascent peptide signal responsive to endogenous levels of polyamines acts to stimulate regulatory frameshifting on antizyme mRNA. J. Biol. Chem. http://dx.doi.org/10.1074/jbc.M115.647065 (2015).

  123. Kim, H. K. et al. A frameshifting stimulatory stem loop destabilizes the hybrid state and impedes ribosomal translocation. Proc. Natl Acad. Sci. USA 111, 5538–5543 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  124. Yu, C. H., Noteborn, M. H., Pleij, C. W. & Olsthoorn, R. C. Stem–loop structures can effectively substitute for an RNA pseudoknot in -1 ribosomal frameshifting. Nucleic Acids Res. 39, 8952–8959 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  125. Mazauric, M. H., Licznar, P., Prere, M. F., Canal, I. & Fayet, O. Apical loop–internal loop RNA pseudoknots: a new type of stimulator of -1 translational frameshifting in bacteria. J. Biol. Chem. 283, 20421–20432 (2008).

    CAS  PubMed  Google Scholar 

  126. Brierley, I., Digard, P. & Inglis, S. C. Characterization of an efficient coronavirus ribosomal frameshifting signal: requirement for an RNA pseudoknot. Cell 57, 537–547 (1989). This is the first work that demonstrated the ability of RNA pseudoknot to promote −1 ribosomal frameshifting at 'slippery' patterns.

    CAS  PubMed  PubMed Central  Google Scholar 

  127. Baranov, P. V. et al. Programmed ribosomal frameshifting in decoding the SARS-CoV genome. Virology 332, 498–510 (2005).

    CAS  PubMed  Google Scholar 

  128. Plant, E. P. et al. A three-stemmed mRNA pseudoknot in the SARS coronavirus frameshift signal. PLoS Biol. 3, e172 (2005).

    PubMed  PubMed Central  Google Scholar 

  129. Su, M. C., Chang, C. T., Chu, C. H., Tsai, C. H. & Chang, K. Y. An atypical RNA pseudoknot stimulator and an upstream attenuation signal for -1 ribosomal frameshifting of SARS coronavirus. Nucleic Acids Res. 33, 4265–4275 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  130. Chou, M. Y. & Chang, K. Y. An intermolecular RNA triplex provides insight into structural determinants for the pseudoknot stimulator of -1 ribosomal frameshifting. Nucleic Acids Res. 38, 1676–1685 (2010).

    CAS  PubMed  Google Scholar 

  131. Chen, G., Chang, K. Y., Chou, M. Y., Bustamante, C. & Tinoco, I. Jr. Triplex structures in an RNA pseudoknot enhance mechanical stability and increase efficiency of -1 ribosomal frameshifting. Proc. Natl Acad. Sci. USA 106, 12706–12711 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  132. Herold, J. & Siddell, S. G. An 'elaborated' pseudoknot is required for high frequency frameshifting during translation of HCV 229E polymerase mRNA. Nucleic Acids Res. 21, 5838–5842 (1993).

    CAS  PubMed  PubMed Central  Google Scholar 

  133. Endoh, T. & Sugimoto, N. Unusual -1 ribosomal frameshift caused by stable RNA G-quadruplex in open reading frame. Anal. Chem. 85, 11435–11439 (2013).

    CAS  PubMed  Google Scholar 

  134. Yu, C. H., Teulade-Fichou, M. P. & Olsthoorn, R. C. Stimulation of ribosomal frameshifting by RNA G-quadruplex structures. Nucleic Acids Res. 42, 1887–1892 (2014).

    CAS  PubMed  Google Scholar 

  135. Tajima, Y., Iwakawa, H. O., Kaido, M., Mise, K. & Okuno, T. A long-distance RNA–RNA interaction plays an important role in programmed -1 ribosomal frameshifting in the translation of p88 replicase protein of Red clover necrotic mosaic virus. Virology 417, 169–178 (2011).

    CAS  PubMed  Google Scholar 

  136. Barry, J. K. & Miller, W. A. A -1 ribosomal frameshift element that requires base pairing across four kilobases suggests a mechanism of regulating ribosome and replicase traffic on a viral RNA. Proc. Natl Acad. Sci. USA 99, 11133–11138 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  137. Giedroc, D. P. & Cornish, P. V. Frameshifting RNA pseudoknots: structure and mechanism. Virus Res. 139, 193–208 (2009).

    CAS  PubMed  Google Scholar 

  138. Chung, B. Y., Firth, A. E. & Atkins, J. F. Frameshifting in alphaviruses: a diversity of 3′ stimulatory structures. J. Mol. Biol. 397, 448–456 (2010).

    CAS  PubMed  Google Scholar 

  139. Brierley, I., Gilbert, R. J. & Pennell, S. RNA pseudoknots and the regulation of protein synthesis. Biochem. Soc. Trans. 36, 684–689 (2008).

    CAS  PubMed  Google Scholar 

  140. Li, Y. et al. Transactivation of programmed ribosomal frameshifting by a viral protein. Proc. Natl Acad. Sci. USA 111, E2172–E2181 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  141. Howard, M. T., Gesteland, R. F. & Atkins, J. F. Efficient stimulation of site-specific ribosome frameshifting by antisense oligonucleotides. RNA 10, 1653–1661 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  142. Olsthoorn, R. C. et al. Novel application of sRNA: stimulation of ribosomal frameshifting. RNA 10, 1702–1703 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  143. Kurian, L., Palanimurugan, R., Godderz, D. & Dohmen, R. J. Polyamine sensing by nascent ornithine decarboxylase antizyme stimulates decoding of its mRNA. Nature 477, 490–494 (2011).

    CAS  PubMed  Google Scholar 

  144. Plant, E. P., Rakauskaite, R., Taylor, D. R. & Dinman, J. D. Achieving a golden mean: mechanisms by which coronaviruses ensure synthesis of the correct stoichiometric ratios of viral proteins. J. Virol. 84, 4330–4340 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  145. Stochmanski, S. J. et al. Expanded ATXN3 frameshifting events are toxic in Drosophila and mammalian neuron models. Hum. Mol. Genet. 21, 2211–2218 (2012).

    CAS  PubMed  Google Scholar 

  146. Wills, N. M. & Atkins, J. F. The potential role of ribosomal frameshifting in generating aberrant proteins implicated in neurodegenerative diseases. RNA 12, 1149–1153 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  147. Toulouse, A. et al. Ribosomal frameshifting on MJD-1 transcripts with long CAG tracts. Hum. Mol. Genet. 14, 2649–2660 (2005).

    CAS  PubMed  Google Scholar 

  148. Girstmair, H. et al. Depletion of cognate charged transfer RNA causes translational frameshifting within the expanded CAG stretch in huntingtin. Cell Rep. 3, 148–159 (2013).

    CAS  PubMed  Google Scholar 

  149. Belcourt, M. F. & Farabaugh, P. J. Ribosomal frameshifting in the yeast retrotransposon Ty: tRNAs induce slippage on a 7 nucleotide minimal site. Cell 62, 339–352 (1990). This work demonstrated that ribosomal frameshifting in yeast requires a specific sequence of only seven nucleotides to occur at a high efficiency (~40%).

    CAS  PubMed  PubMed Central  Google Scholar 

  150. Baranov, P. V., Gesteland, R. F. & Atkins, J. F. P-site tRNA is a crucial initiator of ribosomal frameshifting. RNA 10, 221–230 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  151. Clare, J. J., Belcourt, M. & Farabaugh, P. J. Efficient translational frameshifting occurs within a conserved sequence of the overlap between the two genes of a yeast Ty1 transposon. Proc. Natl Acad. Sci. USA 85, 6816–6820 (1988).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Asakura, T. et al. Isolation and characterization of a novel actin filament-binding protein from Saccharomyces cerevisiae. Oncogene 16, 121–130 (1998).

    CAS  PubMed  Google Scholar 

  153. Vimaladithan, A. & Farabaugh, P. J. Special peptidyl-tRNA molecules can promote translational frameshifting without slippage. Mol. Cell. Biol. 14, 8107–8116 (1994).

    CAS  PubMed  PubMed Central  Google Scholar 

  154. Sundararajan, A., Michaud, W. A., Qian, Q., Stahl, G. & Farabaugh, P. J. Near-cognate peptidyl-tRNAs promote +1 programmed translational frameshifting in yeast. Mol. Cell 4, 1005–1015 (1999).

    CAS  PubMed  Google Scholar 

  155. Temperley, R., Richter, R., Dennerlein, S., Lightowlers, R. N. & Chrzanowska-Lightowlers, Z. M. Hungry codons promote frameshifting in human mitochondrial ribosomes. Science 327, 301 (2010).

    CAS  PubMed  Google Scholar 

  156. Young, D. J. et al. Bioinformatic, structural, and functional analyses support release factor-like MTRF1 as a protein able to decode nonstandard stop codons beginning with adenine in vertebrate mitochondria. RNA 16, 1146–1155 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  157. Akabane, S., Ueda, T., Nierhaus, K. H. & Takeuchi, N. Ribosome rescue and translation termination at non-standard stop codons by ICT1 in mammalian mitochondria. PLoS Genet. 10, e1004616 (2014).

    PubMed  PubMed Central  Google Scholar 

  158. Klobutcher, L. A. & Farabaugh, P. J. Shifty ciliates: frequent programmed translational frameshifting in euplotids. Cell 111, 763–766 (2002). This review discusses evidence supporting the frequent utilization of ribosomal frameshifting during protein synthesis in ciliates of the Euplotes genus.

    CAS  PubMed  Google Scholar 

  159. Vallabhaneni, H., Fan-Minogue, H., Bedwell, D. M. & Farabaugh, P. J. Connection between stop codon reassignment and frequent use of shifty stop frameshifting. RNA 15, 889–897 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  160. Wills, N. M. et al. Translational bypassing without peptidyl-tRNA anticodon scanning of coding gap mRNA. EMBO J. 27, 2533–2544 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  161. Herr, A. J., Gesteland, R. F. & Atkins, J. F. One protein from two open reading frames: mechanism of a 50 nt translational bypass. EMBO J. 19, 2671–2680 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  162. Herr, A. J., Atkins, J. F. & Gesteland, R. F. Coupling of open reading frames by translational bypassing. Annu. Rev. Biochem. 69, 343–372 (2000).

    CAS  PubMed  Google Scholar 

  163. Weiss, R. B., Huang, W. M. & Dunn, D. M. A nascent peptide is required for ribosomal bypass of the coding gap in bacteriophage T4 gene 60. Cell 62, 117–126 (1990).

    CAS  PubMed  PubMed Central  Google Scholar 

  164. Samatova, E., Konevega, A. L., Wills, N. M., Atkins, J. F. & Rodnina, M. V. High-efficiency translational bypassing of non-coding nucleotides specified by mRNA structure and nascent peptide. Nat. Commun. 5, 4459 (2014).

    CAS  PubMed  Google Scholar 

  165. Smith, M. C. et al. Evolutionary relationships among actinophages and a putative adaptation for growth in Streptomyces spp. J. Bacteriol. 195, 4924–4935 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  166. Keiler, K. C. & Ramadoss, N. S. Bifunctional transfer–messenger RNA. Biochimie 93, 1993–1997 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  167. Himeno, H., Kurita, D. & Muto, A. tmRNA-mediated trans-translation as the major ribosome rescue system in a bacterial cell. Front. Genet. 5, 66 (2014).

    PubMed  PubMed Central  Google Scholar 

  168. Hudson, C. M. & Williams, K. P. The tmRNA website. Nucleic Acids Res. 43, D138–D140. (2015).

    CAS  PubMed  Google Scholar 

  169. Hudson, C. M., Lau, B. Y. & Williams, K. P. Ends of the line for tmRNA-SmpB. Front. Microbiol. 5, 421 (2014).

    PubMed  PubMed Central  Google Scholar 

  170. Donnelly, M. L. et al. Analysis of the aphthovirus 2A/2B polyprotein 'cleavage' mechanism indicates not a proteolytic reaction, but a novel translational effect: a putative ribosomal 'skip'. J. Gen. Virol. 82, 1013–1025 (2001). In this work the authors demonstrated that ribosomes can interrupt continuous elongation of polypeptide chains without interruption of mRNA decoding, thus producing two peptide products from a single ORF.

    CAS  PubMed  Google Scholar 

  171. Donnelly, M. L. et al. The 'cleavage' activities of foot-and-mouth disease virus 2A site-directed mutants and naturally occurring '2A-like' sequences. J. Gen. Virol. 82, 1027–1041 (2001).

    CAS  PubMed  Google Scholar 

  172. Chin, J. W. Expanding and reprogramming the genetic code of cells and animals. Annu. Rev. Biochem. 83, 379–408 (2014).

    CAS  PubMed  Google Scholar 

  173. Haimovich, A. D., Muir, P. & Isaacs, F. J. Genomes by design. Nat. Rev. Genet. http://www.dx.doi.org/10.1038/nrg3956 (2015).

  174. Timmis, J. N., Ayliffe, M. A., Huang, C. Y. & Martin, W. Endosymbiotic gene transfer: organelle genomes forge eukaryotic chromosomes. Nat. Rev. Genet. 5, 123–135 (2004).

    CAS  PubMed  Google Scholar 

  175. Kondrashov, A. S. Deleterious mutations and the evolution of sexual reproduction. Nature 336, 435–440 (1988).

    CAS  PubMed  Google Scholar 

  176. Bender, A., Hajieva, P. & Moosmann, B. Adaptive antioxidant methionine accumulation in respiratory chain complexes explains the use of a deviant genetic code in mitochondria. Proc. Natl Acad. Sci. USA 105, 16496–16501 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  177. Korkmaz, G., Holm, M., Wiens, T. & Sanyal, S. Comprehensive analysis of stop codon usage in bacteria and its correlation with release factor abundance. J. Biol. Chem. 289, 30334–30342 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  178. Maas, S. Posttranscriptional recoding by RNA editing. Adv. Protein Chem. Struct. Biol. 86, 193–224 (2012).

    PubMed  Google Scholar 

  179. Kiran, A., Loughran, G., O'Mahony, J. J. & Baranov, P. V. Identification of A-to-I RNA editing: dotting the i's in the human transcriptome. Biochem. (Mosc) 76, 915–923 (2011).

    CAS  Google Scholar 

  180. Mallela, A. & Nishikura, K. A-to-I editing of protein coding and noncoding RNAs. Crit. Rev. Biochem. Mol. Biol. 47, 493–501 (2012).

    CAS  PubMed  Google Scholar 

  181. Carlile, T. M. et al. Pseudouridine profiling reveals regulated mRNA pseudouridylation in yeast and human cells. Nature 515, 143–146 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  182. Karijolich, J. & Yu, Y. T. Converting nonsense codons into sense codons by targeted pseudouridylation. Nature 474, 395–398 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  183. Larsen, B., Wills, N. M., Nelson, C., Atkins, J. F. & Gesteland, R. F. Nonlinearity in genetic decoding: homologous DNA replicase genes use alternatives of transcriptional slippage or translational frameshifting. Proc. Natl Acad. Sci. USA 97, 1683–1688 (2000).

    CAS  PubMed  PubMed Central  Google Scholar 

  184. Ivanov, I. P., Firth, A. E., Michel, A. M., Atkins, J. F. & Baranov, P. V. Identification of evolutionarily conserved non-AUG-initiated N-terminal extensions in human coding sequences. Nucleic Acids Res. 39, 4220–4234 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  185. Wojciechowska, M., Olejniczak, M., Galka-Marciniak, P., Jazurek, M. & Krzyzosiak, W. J. RAN translation and frameshifting as translational challenges at simple repeats of human neurodegenerative disorders. Nucleic Acids Res. 42, 11849–11864 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  186. Kearse, M. G. & Todd, P. K. Repeat-associated non-AUG translation and its impact in neurodegenerative disease. Neurotherapeutics 11, 721–731 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  187. Cleary, J. D. & Ranum, L. P. Repeat-associated non-ATG (RAN) translation in neurological disease. Hum. Mol. Genet. 22, R45–R51 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

This Review was inspired by the discussions during EMBO Workshop “Recoding: Reprogramming genetic decoding” that took place in Killarney, Ireland, on the 13–18 May 2014. Therefore, the authors are grateful to all participants who shared their ideas and experimental data during this meeting. They apologize to their colleagues whose relevant works were not cited as this article is not intended as a comprehensive review of the topic and the authors aimed to keep it concise. The authors wish to acknowledge support by SFI grants [12/IA/1335 to P.V.B. and 08/IN.I/B1889,12/IP/1492 to J.F.A], P.V.B. is also supported by Wellcome Trust grant [094423].

Author information

Authors and Affiliations

Authors

Corresponding author

Correspondence to Pavel V. Baranov.

Ethics declarations

Competing interests

The authors declare no competing financial interests.

Related links

PowerPoint slides

Glossary

Genetic code

A correspondence between 64 triplet combinations of four nucleotides and their standard amino acid or stop meanings.

Variant genetic codes

Genetic codes that differ from the standard genetic code shown in Figure 1a.

Proteinogenic amino acids

Amino acids that are incorporated into proteins co-translationally.

Ribosomal frameshifting

A process in which a ribosome changes its reading frame.

Codon redefinition

A local change of codon meaning that is dependent on the context in which it occurs.

Translational bypassing

A process in which ribosomes skip three or more nucleotides without decoding.

Fixed codon reassignment

A complete unconditional change of the standard meaning of a codon.

Standard meaning

The way the translational machinery interprets a codon (coding for a proteinogenic amino acid or a signal for translation termination) unless it occurs in a specific context.

Codon capture

An evolutionary event in which a codon that disappears from a genome reappears in its descendant and acquires a different standard meaning, thus leading to a variant genetic code.

Ambiguous intermediate

An evolutionary state in the history of an organism evolving a variant genetic code in which a particular codon has two standard meanings.

Regulated codon reassignment

A conditional change of the standard meaning of a codon.

Recoding

A process of context- or condition-specific alteration of genetic decoding.

Stop codon readthrough

A redefinition of a stop codon to a sense codon irrespective of functional implications of the identity of the incorporated amino acid.

Proteoforms

Groups of sequence-related proteins arising from the same mRNA.

SECIS element

(Sec insertion sequence element). An mRNA secondary structure that functions as a stimulatory element for selenocysteine (Sec) incorporation.

Programmed ribosomal frameshifting

(PRF). Ribosomal frameshifting that is programmed (by a sequence context) to occur at a specific mRNA location.

Productive PRF

Programmed ribosomal frameshifting (PRF) that is required for the production of a functional protein product.

Abortive PRF

Programmed ribosomal frameshifting (PRF) that results in the synthesis of dysfunctional protein products or in the downregulation of functional protein synthesis.

Purifying evolutionary selection

The removal of disadvantageous traits. In the case of protein-coding sequences it results in a higher rate of synonymous substitutions relative to non-synonymous substitutions.

Frameshifting site

(Also known as frameshift site and shift site). A sequence in which ribosomal frameshifting takes place. It includes codons in the A- and P-sites of the ribosomes just before and after the frameshifting. It is useful to describe the sequence of the frameshifting site denoting codons in the original and new frames, for example, C.UU_A.GG_C. Such representation unambiguously reflects the direction (minus or plus) as well as the mechanism of frameshifting (+1, +2, and so on)

Stimulatory element

An mRNA element that is required for the efficient local alteration of genetic decoding.

P-site

The ribosomal site that accommodates the peptidyl-tRNA carrying the growing polypeptide chain.

Isoacceptor

One of a group of tRNA species carrying the same amino acids but with different anticodon sequences

A-site

The ribosomal site that accommodates either the aminoacyl-tRNA carrying the next amino acid to be added to the growing polypeptide chain or a release factor.

Byps

Non-coding gaps in mRNAs of mitochondria (in Magnusiomyces capitatus and related species) that escape decoding through frequent translational bypassing.

Trans-translation

A process in which a single protein is translated from two mRNA molecules as templates.

StopGo

(Also known as Stop-Carry on). A process in which the production of a polypeptide chain is interrupted at a specific place while triplet mRNA decoding continues. This results in the production of two protein products from a single open reading frame.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Baranov, P., Atkins, J. & Yordanova, M. Augmented genetic decoding: global, local and temporal alterations of decoding processes and codon meaning. Nat Rev Genet 16, 517–529 (2015). https://doi.org/10.1038/nrg3963

Download citation

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/nrg3963

This article is cited by

Search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing