Lessons from non-canonical splicing

Journal name:
Nature Reviews Genetics
Year published:
Published online


Recent improvements in experimental and computational techniques that are used to study the transcriptome have enabled an unprecedented view of RNA processing, revealing many previously unknown non-canonical splicing events. This includes cryptic events located far from the currently annotated exons and unconventional splicing mechanisms that have important roles in regulating gene expression. These non-canonical splicing events are a major source of newly emerging transcripts during evolution, especially when they involve sequences derived from transposable elements. They are therefore under precise regulation and quality control, which minimizes their potential to disrupt gene expression. We explain how non-canonical splicing can lead to aberrant transcripts that cause many diseases, and also how it can be exploited for new therapeutic strategies.

At a glance


  1. Cryptic exons and microexons.
    Figure 1: Cryptic exons and microexons.

    a | Many introns contain proximally spaced sequences that resemble splice sites (such as GU for 5′ splice sites and YAG for 3′ splice sites), which can in some cases lead to splicing of 'cryptic' exons. Cryptic exons often introduce premature termination codons, which may target the resulting transcripts for nonsense-mediated decay (NMD). Such NMD exons are common within transcripts that encode splicing activators and function as part of autoregulatory mechanisms33, 34, 35. In this example, a serine/arginine-rich (SR) protein enhances the inclusion of an NMD exon within its own mRNA as part of a negative autoregulatory feedback that maintains appropriate steady-state abundance. b | An Alu element is normally composed of two arms, which contain an adenine-rich linker (A-linker; left arm) and a poly(A) tail (right arm). The Alu element can become retrotransposed into the antisense strand relative to the gene, so that transcription of the gene produces an antisense Alu sequence that contains two poly(U) tracts (U tracts) at the beginning of each arm. Many such antisense Alu elements are capable of forming cryptic exons owing to the presence of splice-site-like motifs37. However, they are normally repressed by a heterogeneous nuclear ribonucleoprotein C (HNRNPC) tetramer (C, green circle), possibly because each U tract can bind the two RNA recognition motif domains that are present on the opposite surfaces of the tetramer (as indicated by the double arrow)8, 190. The example provided here shows the U tracts around the Alu exon from the CD55 gene (encoding CD55 molecule). Mutations in the U tracts can decrease the binding of HNRNPC, allowing the binding of U2 small nuclear RNA auxiliary factor (U2AF2) and T cell-restricted intracellular antigen 1 (TIA1), which initiate splicing of a cryptic Alu exon8, 37, 39. c | Microexons (denoted here by μ) can be detected from gapped regions in sequencing reads11, 13, 44. After mapping of multiple parts of the sequence read to flanking exons, unmapped intervening sequences are aligned to the intronic sequence present between the two exons, with preference given to those that are flanked by conserved splice site motifs. Inclusion of microexons can be enhanced by RNA-binding proteins (RBPs), such as serine/arginine repetitive matrix 4 (SRRM4), an SR protein that binds upstream of microexons and promotes microexon splicing. Inclusion of microexons typically leads to the modulation of overlapping or adjacent protein domains and a change in protein activity. SRRM4 is reduced in patients with autism spectrum disorder (ASD), leading to decreased inclusion of microexons12.

  2. Recursive splicing of long introns.
    Figure 2: Recursive splicing of long introns.

    a | Total RNA sequencing (RNA-seq) read counts display a characteristic pattern of depletion from the start to the end of long introns, which can be used to infer exon positions and splicing events47, 48. The 'saw-tooth' patterns that overlap novel junction reads indicate splicing at deep intronic loci and are candidates for recursive splicing14, 15. Here, the upstream exon first uses a 3′ splice site (3ss) to remove the first part of the intron. This process reconstitutes a 5′ splice site (5ss) that can then be used to remove the next section of the intron. This special type of splice site is referred to as a recursive splice site (RS site). b | Recursive splicing in vertebrates requires the RS site to overlap a cryptic 'RS exon', which initiates the exon definition mechanism that is required for the recognition of the 3′ splice site (YAG) of the RS site14. After the first splicing step, the 5′ splice site (GURAG) of the RS site competes with the 5′ splice site of the RS exon. In the second step, the outcome of this competition determines whether the RS exon is skipped, owing to recursive splicing, or included as a nonsense-mediated decay (NMD) exon. Whereas the preceding exons from major isoforms end in sequences that favour RS exon skipping, the minor isoforms and cryptic elements end in sequences that favour RS exon inclusion.

  3. Intron retention and exitrons.
    Figure 3: Intron retention and exitrons.

    a | Intron retention events are detected as an accumulation of reads across intronic regions or increases in the ratio of exon–intron reads to exon–exon reads21, 22, 23, 24, 25. Intron retention events are characterized by numerous features including weak splice sites, high GC content and short intron lengths. Trans-acting factors such as RNA-binding proteins (RBPs), the spliceosome and the exon junction complex (EJC) can also regulate specific intron retention events. The resulting transcripts are typically either retained in the nucleus or targeted for nonsense-mediated decay (NMD) in the cytoplasm or may result in truncated proteins21, 53, 55. b | Exonic introns (exitrons) are introns within annotated protein-coding exons that can be removed owing to the presence of internal splice site motifs within the exon25, 26. Exitron-containing exons are longer than typical exons, and removal of the exitron can lead to changes in protein structure or degradation through NMD.

  4. Formation of circularRNAs and chimeric transcripts.
    Figure 4: Formation of circularRNAs and chimeric transcripts.

    a | Circular RNAs (circRNAs) are produced by head-to-tail splicing and can be either mono- or multi-exonic. In this multi-exonic example, the 3′ splice site of an upstream exon becomes spliced to the 5′ splice site of a downstream exon to generate a circular transcript in which the intervening intron is either removed (exonic circRNA) or retained between the two circularized exons (exon–intron circRNA)20. circRNA formation is promoted when the pre-mRNA regions flanking the exon termini are brought into close proximity. This can be due to the action of RNA-binding proteins (RBPs) such as quaking (QKI) or muscleblind-like (MBNL), which bind to the flanking regions74, 75. Alternatively, this can be due to RNA hybridization of the flanking regions, which can be caused by Alu elements in primates70. b | CircRNAs are resistant to RNase R exoribonuclease activity, which can be used for their enrichment during preparation of cDNA libraries. They can then be detected in RNA sequencing (RNA-seq) data by junction reads that are in a head-to-tail orientation16, 17, 18, 19. c | Chimeric RNA products can also be produced by cis-splicing when transcript termination is deficient76. This process results in read-through of one gene into its neighbouring gene, before splicing occurs between the penultimate exon of gene 1 and the second exon of gene 2, which is seen in the chimeric cathepsin C (CTSC)–RAB38 gene in some cancers. d |Trans-splicing occurs when exons of two different transcripts become spliced together80, 81, 82, 83, 84, 85, 86, 87. Alternatively, the same chimeric transcripts can be produced when genes become fused, such as in JAZF1–SUZ12 gene fusion in some cancers, which leads to the same chimeric transcript being produced by a linear splicing reaction.

  5. A summary of human splice site DNA consensus motifs.
    Figure 5: A summary of human splice site DNA consensus motifs.

    Summarized splice site sequences are classified using the nucleotides marked by the grey boxes. All borders of human exons within Ensembl v83 multi-exon transcripts that overlap with Reference Sequence (RefSeq) database mRNA IDs were used. Identical coordinates from overlapping transcripts were collapsed into a single occurrence so that junctions were not counted multiple times. The first exons had only their exon–intron junction evaluated, whereas terminal exons had only their intron–exon junction evaluated. This led to a total of 189,255 5′ splice sites (left panels, with the black line marking the exon–intron border) and 187,091 3′ splice sites (right panels, with the black line marking the intron–exon border). Splice site sequences of U12-type introns were obtained from the U12 Intron Database191. After identifying the 5′ and 3′ sites that overlap with the respective U11-type and U12-type splice sites, the remaining U2-type intron splice site sequences were examined. 5′ and 3′ splice sites were classified independently and sequentially based on the indicated nucleotides. For example, 53.58% of unique U1-type exon–intron junctions contain GTRAG, and the remaining U1-type junctions were classified on the basis of the first two intronic nucleotides, GT. The percentage of unique junctions containing each motif is indicated. Weblogo 3 was used to show the relative frequency of nucleotides at each position192. a | Consensus motifs of the U1-type 5′ splice sites with GT at the border and the U2-type 3′ splice sites with AG at the border. b | Consensus motifs of the U11-type 5′ splice sites and U12-type 3′ splice sites. c | Consensus motifs of U1-type 5′ splice sites with GC at the border, or with TN or VN at the border (in which N stands for any nucleotide, and V stands for any nucleotide except T), and U2-type 3′ splice sites with BG or W at the border (in which B stands for any nucleotide except A, and W stands for T or A).

  6. Cryptic splicing in disease and therapeutic strategies.
    Figure 6: Cryptic splicing in disease and therapeutic strategies.

    a | Cryptic exons are normally repressed by RNA-binding proteins (RBPs) such as heterogeneous nuclear ribonucleoprotein C (HNRNPC) or by U1 small nuclear ribonucleoproteins (snRNP). b | Examples of mutations (α, β, γ and δ) in deep intronic regions that can activate cryptic splicing events in disease-associated genes. Mutation α: HNRNPC binding to a poly(U) tract (U tract) upstream of an antisense Alu element represses recognition of the cryptic 3′ splice site within the element. Intronic deletions or point mutations that shorten the U tract can impede HNRNPC recruitment but allow U2 small nuclear RNA auxiliary factor (U2AF2) binding, leading to Alu exonization. A deletion within an Alu in the PTS gene (encoding 6-pyruvoyltetrahydropterin synthase) leads to splicing of an Alu exon that introduces a frameshift, thereby causing the neurologic disease hyperphenylalaninaemia8, 141. Mutation β: in ataxia telangiectasia mutated (ATM), U1 snRNP binding to an intronic element within a cryptic exon inhibits its recognition as a splicing competent exon. Patients with ataxia telangiectasia present a 4 nt deletion that abolishes U1 snRNP interaction, causing cryptic exon activation110. Mutation γ: a point mutation within a deep intronic sequence of cystic fibrosis transmembrane conductance regulator (CFTR) generates an active 5′ splice site that allows insertion of a cryptic exon within the CFTR transcripts, which causes cystic fibrosis135. Mutation δ: in breast cancer 2 (BRCA2), a point mutation that disrupts a canonical 3′ splice site activates an upstream cryptic exon (grey arrow)136. Disrupted BRCA2 expression causes breast, ovarian and other cancer types. c | New therapeutic strategies in cancer involve spliceosome targeting156, 161, 162. In MYC-driven tumours, oncogenic MYC causes transcriptional amplification, which overloads the splicing machinery and makes these cells more sensitive to alterations in splicing fidelity. Genetic knockdown or pharmacological inhibition of spliceosomal components leads to the accumulation of retained introns, which results in increased apoptosis and reduced tumorigenic and metastatic potential of MYC-driven tumours.


  1. Raj, B. & Blencowe, B. J. Alternative splicing in the mammalian nervous system: recent insights into mechanisms and functional roles. Neuron 87, 1427 (2015).
  2. Fu, X. D. & Ares, M. Jr. Context-dependent control of alternative splicing by RNA-binding proteins. Nat. Rev. Genet. 15, 689701 (2014).
  3. Derrien, T. et al. The GENCODE v7 catalog of human long noncoding RNAs: analysis of their gene structure, evolution, and expression. Genome Res. 22, 17751789 (2012).
  4. Matera, A. G. & Wang, Z. A day in the life of the spliceosome. Nat. Rev. Mol. Cell Biol. 15, 108121 (2014).
  5. Scotti, M. M. & Swanson, M. S. RNA mis-splicing in disease. Nat. Rev. Genet. 17, 1932 (2015).
  6. Jangi, M., Boutz, P. L., Paul, P. & Sharp, P. A. Rbfox2 controls autoregulation in RNA-binding protein networks. Genes Dev. 28, 637651 (2014).
    In this study, RBFOX2 is found to cross-regulate conserved NMD exons within transcripts that encode nearly 70 RBPs, thus forming a broad auto- and cross-regulatory splicing network for fine-tuning the expression levels of RBPs.
  7. Eom, T. et al. NOVA-dependent regulation of cryptic NMD exons controls synaptic protein levels after seizure. eLife 2, e00178 (2013).
    In this study, NOVA proteins are found to regulate the splicing of many cryptic NMD exons, which mediates the regulation of transcripts encoding synaptic proteins in response to excitation, indicating its role in the homeostasis of synaptic activity.
  8. Zarnack, K. et al. Direct competition between hnRNP C and U2AF65 protects the transcriptome from the exonization of Alu elements. Cell 152, 453466 (2013).
    In this paper, U tracts within thousands of antisense Alu elements are found to act as a platform for competition between HNRNPC and U2AF2, and therefore the length of the U tract affects splicing efficiency of cryptic Alu exons.
  9. Ling, J. P., Pletnikova, O., Troncoso, J. C. & Wong, P. C. TDP-43 repression of nonconserved cryptic exons is compromised in ALS-FTD. Science 349, 650655 (2015).
  10. Yan, Q. et al. Systematic discovery of regulated and conserved alternative exons in the mammalian brain reveals NMD modulating chromatin regulators. Proc. Natl Acad. Sci. USA 112, 34453450 (2015).
  11. Wu, J., Anczukow, O., Krainer, A. R., Zhang, M. Q. & Zhang, C. OLego: fast and sensitive mapping of spliced mRNA-Seq reads using small seeds. Nucleic Acids Res. 41, 51495163 (2013).
    This paper describes a sequence mapping algorithm that incorporates short seed mapping and identified >500,000 non-canonical splicing events, including microexons, in humans and mice.
  12. Irimia, M. et al. A highly conserved program of neuronal microexons is misregulated in autistic brains. Cell 159, 15111523 (2014).
    In this study, a new strategy to map RNA-seq data identified more than 400 ORF-modifying microexons, many of which are regulated by SRMM4 and are differentially spliced in patients with ASD.
  13. Li, Y. I., Sanchez-Pulido, L., Haerty, W. & Ponting, C. P. RBFOX and PTBP1 proteins regulate the alternative splicing of micro-exons in human brain transcripts. Genome Res. 25, 113 (2015).
  14. Sibley, C. R. et al. Recursive splicing in long vertebrate genes. Nature 521, 371375 (2015).
    This paper describes the identification of highly conserved vertebrate RS sites within extremely long introns that require an unusual exon definition mechanism for their splicing. Genes with such long introns are found to be more highly expressed in the brain.
  15. Duff, M. O. et al. Genome-wide identification of zero nucleotide recursive splicing in Drosophila. Nature 521, 376379 (2015).
    This paper identifies197 RS sites in Drosophila, one of which is also found in an orthologous human gene.
  16. Hansen, T. B. et al. Natural RNA circles function as efficient microRNA sponges. Nature 495, 384388 (2013).
    This study identifies a brain-enriched circRNA that acts as a miR-7 sponge and a testes-specific circRNA acting as a miR-138 sponge.
  17. Memczak, S. et al. Circular RNAs are a large class of animal RNAs with regulatory potency. Nature 495, 333338 (2013).
    This paper reports the discovery of thousands of new circRNAs across multiple tissues and species, including a circRNA that acts as a miR-7 sponge.
  18. Salzman, J., Gawad, C., Wang, P. L., Lacayo, N. & Brown, P. O. Circular RNAs are the predominant transcript isoform from hundreds of human genes in diverse cell types. PLoS ONE 7, e30733 (2012).
  19. Danan, M., Schwartz, S., Edelheit, S. & Sorek, R. Transcriptome-wide discovery of circular RNAs in Archaea. Nucleic Acids Res. 40, 31313142 (2012).
  20. Chen, L. L. The biogenesis and emerging roles of circular RNAs. Nat. Rev. Mol. Cell Biol. 17, 205211 (2016).
  21. Braunschweig, U. et al. Widespread intron retention in mammals functionally tunes transcriptomes. Genome Res. 24, 17741786 (2014).
    This paper reports that intron retention can be detected in three-quarters of human and mouse multiexonic genes; it is often coupled to RNA polymerase II stalling to suppress inappropriately expressed transcripts.
  22. Yap, K., Lim, Z. Q., Khandelia, P., Friedman, B. & Makeyev, E. V. Coordinated regulation of neuronal mRNA steady-state levels through developmentally controlled intron retention. Genes Dev. 26, 12091223 (2012).
    This study shows that PTBP1 regulates the expression of at least four neuron-specific genes by inhibiting splicing of 3′ terminal introns in non-neuronal cells, which promotes nuclear retention and degradation of the resulting transcripts.
  23. Boutz, P. L., Bhutkar, A. & Sharp, P. A. Detained introns are a novel, widespread class of post-transcriptionally spliced introns. Genes Dev. 29, 6380 (2015).
  24. Wong, J. J. et al. Orchestrated intron retention regulates normal granulocyte differentiation. Cell 154, 583595 (2013).
    This paper reports that coordinated intron retention in 86 functionally related genes is used to regulate gene expression during granulopoiesis.
  25. Marquez, Y., Brown, J. W., Simpson, C., Barta, A. & Kalyna, M. Transcriptome survey reveals increased complexity of the alternative splicing landscape in Arabidopsis. Genome Res. 22, 11841195 (2012).
  26. Marquez, Y., Hopfler, M., Ayatollahi, Z., Barta, A. & Kalyna, M. Unmasking alternative splicing inside protein-coding exons defines exitrons and their role in proteome plasticity. Genome Res. 25, 9951007 (2015).
    This paper reports the discovery of 923 alternative introns inside annotated human protein-coding exons, which are referred to as exitrons.
  27. De Conti, L., Baralle, M. & Buratti, E. Exon and intron definition in pre-mRNA splicing. Wiley Interdiscip. Rev. RNA 4, 4960 (2013).
  28. Robberson, B. L., Cote, G. J. & Berget, S. M. Exon definition may facilitate splice site selection in RNAs with multiple exons. Mol. Cell. Biol. 10, 8494 (1990).
  29. Trapnell, C., Pachter, L. & Salzberg, S. L. TopHat: discovering splice junctions with RNA-Seq. Bioinformatics 25, 11051111 (2009).
  30. Dobin, A. et al. STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29, 1521 (2013).
  31. Kelly, S. et al. Splicing of many human genes involves sites embedded within introns. Nucleic Acids Res. 43, 47214732 (2015).
  32. Kapustin, Y. et al. Cryptic splice sites and split genes. Nucleic Acids Res. 39, 58375844 (2011).
  33. Ni, J. Z. et al. Ultraconserved elements are associated with homeostatic control of splicing regulators by alternative splicing and nonsense-mediated decay. Genes Dev. 21, 708718 (2007).
  34. Lareau, L. F., Inada, M., Green, R. E., Wengrod, J. C. & Brenner, S. E. Unproductive splicing of SR genes associated with highly conserved and ultraconserved DNA elements. Nature 446, 926929 (2007).
  35. Jangi, M. & Sharp, P. A. Building robust transcriptomes with master splicing factors. Cell 159, 487498 (2014).
  36. Vaz-Drago, R. et al. Transcription-coupled RNA surveillance in human genetic diseases caused by splice site mutations. Hum. Mol. Genet. 24, 27842795 (2015).
  37. Keren, H., Lev-Maor, G. & Ast, G. Alternative splicing and evolution: diversification, exon definition and function. Nat. Rev. Genet. 11, 345355 (2010).
  38. Quentin, Y. Origin of the Alu family: a family of Alu-like monomers gave birth to the left and the right arms of the Alu elements. Nucleic Acids Res. 20, 33973401 (1992).
  39. Gal-Mark, N., Schwartz, S., Ram, O., Eyras, E. & Ast, G. The pivotal roles of TIA proteins in 5′ splice-site selection of Alu exons and across evolution. PLoS Genet. 5, e1000717 (2009).
  40. Konig, J., Zarnack, K., Luscombe, N. M. & Ule, J. Protein–RNA interactions: new genomic technologies and perspectives. Nat. Rev. Genet. 13, 7783 (2011).
  41. Corvelo, A. & Eyras, E. Exon creation and establishment in human genes. Genome Biol. 9, R141 (2008).
  42. Dominski, Z. & Kole, R. Selection of splice sites in pre-mRNAs with short internal exons. Mol. Cell. Biol. 11, 60756083 (1991).
  43. Black, D. L. Does steric interference between splice sites block the splicing of a short c-src neuron-specific exon in non-neuronal cells? Genes Dev. 5, 389402 (1991).
  44. Volfovsky, N., Haas, B. J. & Salzberg, S. L. Computational discovery of internal micro-exons. Genome Res. 13, 12161221 (2003).
  45. Burnette, J. M., Miyamoto-Sato, E., Schaub, M. A., Conklin, J. & Lopez, A. J. Subdivision of large introns in Drosophila by recursive splicing at nonexonic elements. Genetics 170, 661674 (2005).
  46. Hatton, A. R., Subramaniam, V. & Lopez, A. J. Generation of alternative Ultrabithorax isoforms and stepwise removal of a large intron by resplicing at exon–exon junctions. Mol. Cell 2, 787796 (1998).
  47. Herzel, L. & Neugebauer, K. M. Quantification of co-transcriptional splicing from RNA-Seq data. Methods, 85, 3643 (2015).
  48. Ameur, A. et al. Total RNA sequencing reveals nascent transcription and widespread co-transcriptional splicing in the human brain. Nat. Struct. Mol. Biol. 18, 14351440 (2011).
  49. Parra, M. K., Tan, J. S., Mohandas, N. & Conboy, J. G. Intrasplicing coordinates alternative first exons with alternative splicing in the protein 4.1R gene. EMBO J. 27, 122131 (2008).
  50. Ner-Gaon, H. et al. Intron retention is a major phenomenon in alternative splicing in Arabidopsis. Plant J. 39, 877885 (2004).
  51. Galante, P. A., Sakabe, N. J., Kirschbaum-Slager, N. & de Souza, S. J. Detection and evaluation of intron retention events in the human transcriptome. RNA 10, 757765 (2004).
  52. Kan, Z., States, D. & Gish, W. Selecting for functional alternative splices in ESTs. Genome Res. 12, 18371845 (2002).
  53. Sakabe, N. J. & de Souza, S. J. Sequence features responsible for intron retention in human. BMC Genom. 8, 59 (2007).
  54. Martinez-Contreras, R. et al. Intronic binding sites for hnRNP A/B and hnRNP F/H proteins stimulate pre-mRNA splicing. PLoS Biol. 4, e21 (2006).
  55. Wickramasinghe, V. O. et al. Regulation of constitutive and alternative mRNA splicing across the human transcriptome by PRPF8 is determined by 5′ splice site strength. Genome Biol. 16, 201 (2015).
  56. Marinescu, V., Loomis, P. A., Ehmann, S., Beales, M. & Potashkin, J. A. Regulation of retention of FosB intron 4 by PTB. PLoS ONE 2, e828 (2007).
  57. Bergeron, D., Pal, G., Beaulieu, Y. B., Chabot, B. & Bachand, F. Regulated intron retention and nuclear pre-mRNA decay contribute to PABPN1 autoregulation. Mol. Cell. Biol. 35, 25032517 (2015).
  58. Malone, C. D. et al. The exon junction complex controls transposable element activity by ensuring faithful splicing of the piwi transcript. Genes Dev. 28, 17861799 (2014).
  59. Hayashi, R., Handler, D., Ish-Horowicz, D. & Brennecke, J. The exon junction complex is required for definition and excision of neighboring introns in Drosophila. Genes Dev. 28, 17721785 (2014).
  60. Wang, Z., Murigneux, V. & Le Hir, H. Transcriptome-wide modulation of splicing by the exon junction complex. Genome Biol. 15, 551 (2014).
  61. Nigro, J. M. et al. Scrambled exons. Cell 64, 607613 (1991).
  62. Schindewolf, C., Braun, S. & Domdey, H. In vitro generation of a circular exon from a linear pre-mRNA transcript. Nucleic Acids Res. 24, 12601266 (1996).
  63. Pasman, Z., Been, M. D. & Garcia-Blanco, M. A. Exon circularization in mammalian nuclear extracts. RNA 2, 603610 (1996).
  64. Braun, S., Domdey, H. & Wiebauer, K. Inverse splicing of a discontinuous pre-mRNA intron generates a circular exon in a HeLa cell nuclear extract. Nucleic Acids Res. 24, 41524157 (1996).
  65. Suzuki, H. et al. Characterization of RNase R-digested cellular RNA source that consists of lariat and circular RNAs from pre-mRNA splicing. Nucleic Acids Res. 34, e63 (2006).
  66. Guo, J. U., Agarwal, V., Guo, H. & Bartel, D. P. Expanded identification and characterization of mammalian circular RNAs. Genome Biol. 15, 409 (2014).
  67. Jeck, W. R. & Sharpless, N. E. Detecting and characterizing circular RNAs. Nat. Biotechnol. 32, 453461 (2014).
  68. You, X. et al. Neural circular RNAs are derived from synaptic genes and regulated by development and plasticity. Nat. Neurosci. 18, 603610 (2015).
  69. Liang, D. & Wilusz, J. E. Short intronic repeat sequences facilitate circular RNA production. Genes Dev. 28, 22332247 (2014).
  70. Jeck, W. R. et al. Circular RNAs are abundant, conserved, and associated with ALU repeats. RNA 19, 141157 (2013).
  71. Kramer, M. C. et al. Combinatorial control of Drosophila circular RNA expression by intronic repeats, hnRNPs, and SR proteins. Genes Dev. 29, 21682182 (2015).
  72. Zhang, X. O. et al. Complementary sequence-mediated exon circularization. Cell 159, 134147 (2014).
  73. Ivanov, A. et al. Analysis of intron sequences reveals hallmarks of circular RNA biogenesis in animals. Cell Rep. 10, 170177 (2015).
  74. Ashwal-Fluss, R. et al. circRNA biogenesis competes with pre-mRNA splicing. Mol. Cell 56, 5566 (2014).
  75. Conn, S. J. et al. The RNA binding protein quaking regulates formation of circRNAs. Cell 160, 11251134 (2015).
    This study shows that QKI facilitates certain circRNA back-splicing events across developmental pathways, possibly through its dimerization when bound to flanking intronic elements.
  76. Grosso, A. R. et al. Pervasive transcription read-through promotes aberrant expression of oncogenes and RNA chimeras in renal carcinoma. eLife 4, e09214 (2015).
    This paper reports that transcription read-through beyond the termination site in clear cell renal cell carcinoma (ccRCC) leads to chimeric transcripts through cis-splicing that correlates with poor survival rates.
  77. Akiva, P. et al. Transcription-mediated gene fusion in the human genome. Genome Res. 16, 3036 (2006).
  78. Qin, F. et al. Discovery of CTCF-sensitive cis-spliced fusion RNAs between adjacent genes in human prostate cells. PLoS Genet. 11, e1005001 (2015).
  79. Jividen, K. & Li, H. Chimeric RNAs generated by intergenic splicing in normal and cancer cells. Genes Chromosomes Cancer 53, 963971 (2014).
  80. Sutton, R. E. & Boothroyd, J. C. Evidence for trans splicing in trypanosomes. Cell 47, 527535 (1986).
  81. Allen, M. A., Hillier, L. W., Waterston, R. H. & Blumenthal, T. A global analysis of C. elegans trans-splicing. Genome Res. 21, 255264 (2011).
  82. McManus, C. J., Duff, M. O., Eipper-Mains, J. & Graveley, B. R. Global analysis of trans-splicing in Drosophila. Proc. Natl Acad. Sci. USA 107, 1297512979 (2010).
  83. Dorn, R., Reuter, G. & Loewendorf, A. Transgene analysis proves mRNA trans-splicing at the complex mod(mdg4) locus in Drosophila. Proc. Natl Acad. Sci. USA 98, 97249729 (2001).
  84. Gabler, M. et al. Trans-splicing of the mod(mdg4) complex locus is conserved between the distantly related species Drosophila melanogaster and D. virilis. Genetics 169, 723736 (2005).
  85. Kong, Y. et al. The evolutionary landscape of intergenic trans-splicing events in insects. Nat. Commun. 6, 8734 (2015).
  86. Li, H., Wang, J., Mor, G. & Sklar, J. A neoplastic gene fusion mimics trans-splicing of RNAs in normal human cells. Science 321, 13571361 (2008).
    This paper reports that physiologically regulated trans-splicing between precursor mRNAs for JAZF1 and JJAZ1 forms a chimeric transcript and protein with anti-apoptotic activity identical to that produced from chromosomal rearrangements in human tumours.
  87. Wu, C. S. et al. Integrative transcriptome sequencing identifies trans-splicing events with important roles in human embryonic stem cell pluripotency. Genome Res. 24, 2536 (2014).
  88. Dietrich, R. C., Incorvaia, R. & Padgett, R. A. Terminal intron dinucleotide sequences do not distinguish between U2- and U12-dependent introns. Mol. Cell 1, 151160 (1997).
  89. Wu, Q. & Krainer, A. R. Splicing of a divergent subclass of AT-AC introns requires the major spliceosomal snRNAs. RNA 3, 586601 (1997).
  90. Sheth, N. et al. Comprehensive splice-site analysis using comparative genomics. Nucleic Acids Res. 34, 39553967 (2006).
  91. Parada, G. E., Munita, R., Cerda, C. A. & Gysling, K. A comprehensive survey of non-canonical splice sites in the human transcriptome. Nucleic Acids Res. 42, 1056410578 (2014).
  92. Mercer, T. R. et al. Genome-wide discovery of human splicing branchpoints. Genome Res. 25, 290303 (2015).
  93. DeBoever, C. et al. Transcriptome sequencing reveals potential mechanism of cryptic 3′ splice site selection in SF3B1-mutated cancers. PLoS Comput. Biol. 11, e1004105 (2015).
  94. Darman, R. B. et al. Cancer-associated SF3B1 hotspot mutations induce cryptic 3′ splice site selection through use of a different branch point. Cell Rep. 13, 10331045 (2015).
    This paper shows that mutations in SF3B1 lead to tumour-specific splicing changes by using an alternative branch point that induces aberrant 3′ splice site selection.
  95. Alsafadi, S. et al. Cancer-associated SF3B1 mutations affect alternative splicing by promoting alternative branchpoint usage. Nat. Commun. 7, 10615 (2016).
  96. Roca, X. & Krainer, A. R. Recognition of atypical 5′ splice sites by shifted base-pairing to U1 snRNA. Nat. Struct. Mol. Biol. 16, 176182 (2009).
  97. Roca, X. et al. Widespread recognition of 5′ splice sites by noncanonical base-pairing to U1 snRNA involving bulged nucleotides. Genes Dev. 26, 10981109 (2012).
  98. Rueter, S. M., Dawson, T. R. & Emeson, R. B. Regulation of alternative splicing by RNA editing. Nature 399, 7580 (1999).
  99. Shen, X. et al. Complementary signaling pathways regulate the unfolded protein response and are required for C. elegans development. Cell 107, 893903 (2001).
  100. Yoshida, H., Matsui, T., Yamamoto, A., Okada, T. & Mori, K. XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to produce a highly active transcription factor. Cell 107, 881891 (2001).
  101. Filipowicz, W. Making ends meet: a role of RNA ligase RTCB in unfolded protein response. EMBO J. 33, 28872889 (2014).
  102. Dergai, M. et al. Microexon-based regulation of ITSN1 and Src SH3 domains specificity relies on introduction of charged amino acids into the interaction interface. Biochem. Biophys. Res. Commun. 399, 307312 (2010).
  103. Quesnel-Vallieres, M., Irimia, M., Cordes, S. P. & Blencowe, B. J. Essential roles for the splicing regulator nSR100/SRRM4 during nervous system development. Genes Dev. 29, 746759 (2015).
  104. Wright, P. E. & Dyson, H. J. Intrinsically disordered proteins in cellular signalling and regulation. Nat. Rev. Mol. Cell Biol. 16, 1829 (2015).
  105. Rossbach, O. et al. Auto- and cross-regulation of the hnRNP L proteins by alternative splicing. Mol. Cell. Biol. 29, 14421451 (2009).
  106. Buckley, P. T., Khaladkar, M., Kim, J. & Eberwine, J. Cytoplasmic intron retention, function, splicing, and the sentinel RNA hypothesis. Wiley Interdiscip Rev. RNA 5, 223230 (2014).
  107. Sibley, C. R. Regulation of gene expression through production of unstable mRNA isoforms. Biochem. Soc. Trans. 42, 11961205 (2014).
  108. Jens, M. & Rajewsky, N. Competition between target sites of regulators shapes post-transcriptional gene regulation. Nat. Rev. Genet. 16, 113126 (2015).
  109. Dhir, A., Buratti, E., van Santen, M. A., Luhrmann, R. & Baralle, F. E. The intronic splicing code: multiple factors involved in ATM pseudoexon definition. EMBO J. 29, 749760 (2010).
  110. Pagani, F. et al. A new type of mutation causes a splicing defect in ATM. Nat. Genet. 30, 426429 (2002).
  111. Liu, N. et al. N6-methyladenosine-dependent RNA structural switches regulate RNA–protein interactions. Nature 518, 560564 (2015).
  112. Solomon, O. et al. Global regulation of alternative splicing by adenosine deaminase acting on RNA (ADAR). RNA 19, 591604 (2013).
  113. Lovci, M. T. et al. Rbfox proteins regulate alternative mRNA splicing through evolutionarily conserved RNA bridges. Nat. Struct. Mol. Biol. 20, 14341442 (2013).
  114. Bitton, D. A. et al. Widespread exon skipping triggers degradation by nuclear RNA surveillance in fission yeast. Genome Res. 25, 884896 (2015).
  115. de Koning, A. P., Gu, W., Castoe, T. A., Batzer, M. A. & Pollock, D. D. Repetitive elements may comprise over two-thirds of the human genome. PLoS Genet. 7, e1002384 (2011).
  116. Brouha, B. et al. Hot L1s account for the bulk of retrotransposition in the human population. Proc. Natl Acad. Sci. USA 100, 52805285 (2003).
  117. Jacob, F. Evolution and tinkering. Science 196, 11611166 (1977).
  118. Cowley, M. & Oakey, R. J. Transposable elements re-wire and fine-tune the transcriptome. PLoS Genet. 9, e1003234 (2013).
  119. Ule, J. Alu elements: at the crossroads between disease and evolution. Biochem. Soc. Trans. 41, 15321535 (2013).
  120. Brunet, T. D. & Doolittle, W. F. Multilevel selection theory and the evolutionary functions of transposable elements. Genome Biol. Evol. 7, 24452457 (2015).
  121. Feschotte, C. Transposable elements and the evolution of regulatory networks. Nat. Rev. Genet. 9, 397405 (2008).
  122. Roy, M., Kim, N., Xing, Y. & Lee, C. The effect of intron length on exon creation ratios during the evolution of mammalian genomes. RNA 14, 22612273 (2008).
  123. Pickrell, J. K., Pai, A. A., Gilad, Y. & Pritchard, J. K. Noisy splicing drives mRNA isoform diversity in human cells. PLoS Genet. 6, e1001236 (2010).
  124. Lopez-Bigas, N., Audit, B., Ouzounis, C., Parra, G. & Guigo, R. Are splicing mutations the most frequent cause of hereditary disease? FEBS Lett. 579, 19001903 (2005).
  125. Daguenet, E., Dujardin, G. & Valcarcel, J. The pathogenicity of splicing defects: mechanistic insights into pre-mRNA processing inform novel therapeutic approaches. EMBO Rep. 16, 16401655 (2015).
  126. Singh, R. K. & Cooper, T. A. Pre-mRNA splicing in disease and therapeutics. Trends Mol. Med. 18, 472482 (2012).
  127. Supek, F., Minana, B., Valcarcel, J., Gabaldon, T. & Lehner, B. Synonymous mutations frequently act as driver mutations in human cancers. Cell 156, 13241335 (2014).
  128. Xiong, H. Y. et al. RNA splicing. The human splicing code reveals new insights into the genetic determinants of disease. Science 347, 1254806 (2015).
    In this study, machine learning reveals the effect of distal sequence variants on splicing outcome, the predictive value of which will be important when considering genomic variation at non-canonical splicing elements.
  129. Meili, D. et al. Disease-causing mutations improving the branch site and polypyrimidine tract: pseudoexon activation of LINE-2 and antisense Alu lacking the poly(T)-tail. Hum. Mutat. 30, 823831 (2009).
  130. Ferlini, A. et al. A novel Alu-like element rearranged in the dystrophin gene causes a splicing mutation in a family with X-linked dilated cardiomyopathy. Am. J. Hum. Genet. 63, 436446 (1998).
  131. Sowalsky, A. G. et al. Whole transcriptome sequencing reveals extensive unspliced mRNA in metastatic castration-resistant prostate cancer. Mol. Cancer Res. 13, 98106 (2015).
  132. Yuan, H. et al. A chimeric RNA characteristic of rhabdomyosarcoma in normal myogenesis process. Cancer Discov. 3, 13941403 (2013).
  133. Greer, K. et al. Pseudoexon activation increases phenotype severity in a Becker muscular dystrophy patient. Mol. Genet. Genom. Med. 3, 320326 (2015).
  134. Buratti, E., Dhir, A., Lewandowska, M. A. & Baralle, F. E. RNA structure is a key regulatory element in pathological ATM and CFTR pseudoexon inclusion events. Nucleic Acids Res. 35, 43694383 (2007).
  135. Highsmith, W. E. et al. A novel mutation in the cystic fibrosis gene in patients with pulmonary disease but normal sweat chloride concentrations. N. Engl. J. Med. 331, 974980 (1994).
  136. Chen, X. et al. Intronic alterations in BRCA1 and BRCA2: effect on mRNA splicing fidelity and expression. Hum. Mutat. 27, 427435 (2006).
  137. Lualdi, S. et al. Multiple cryptic splice sites can be activated by IDS point mutations generating misspliced transcripts. J. Mol. Med. (Berl.) 84, 692700 (2006).
  138. Sathasivam, K. et al. Aberrant splicing of HTT generates the pathogenic exon 1 protein in Huntington disease. Proc. Natl Acad. Sci. USA 110, 23662370 (2013).
  139. Ghosal, S., Das, S., Sen, R., Basak, P. & Chakrabarti, J. Circ2Traits: a comprehensive database for circular RNA potentially associated with disease and traits. Frontiers Genet. 4, 283 (2013).
  140. Akker, S. A. et al. Pre-spliceosomal binding of U1 small nuclear ribonucleoprotein (RNP) and heterogenous nuclear RNP E1 is associated with suppression of a growth hormone receptor pseudoexon. Mol. Endocrinol. 21, 25292540 (2007).
  141. Vorechovsky, I. Transposable elements in disease-associated cryptic exons. Hum. Genet. 127, 135154 (2010).
  142. Madan, V. et al. Aberrant splicing of U12-type introns is the hallmark of ZRSR2 mutant myelodysplastic syndrome. Nat. Commun. 6, 6042 (2015).
  143. Edery, P. et al. Association of TALS developmental disorder with defect in minor splicing component U4atac snRNA. Science 332, 240243 (2011).
  144. He, H. et al. Mutations in U4atac snRNA, a component of the minor spliceosome, in the developmental disorder MOPD I. Science 332, 238240 (2011).
  145. Merico, D. et al. Compound heterozygous mutations in the noncoding RNU4ATAC cause Roifman Syndrome by disrupting minor intron splicing. Nat. Commun. 6, 8718 (2015).
  146. Yoshida, K. et al. Frequent pathway mutations of splicing machinery in myelodysplasia. Nature 478, 6469 (2011).
  147. Menzies, F. M., Fleming, A. & Rubinsztein, D. C. Compromised autophagy and neurodegenerative diseases. Nat. Rev. Neurosci. 16, 345357 (2015).
  148. Argente, J. et al. Defective minor spliceosome mRNA processing results in isolated familial growth hormone deficiency. EMBO Mol. Med. 6, 299306 (2014).
  149. Bachmayr-Heyda, A. et al. Correlation of circular RNA abundance with proliferation — exemplified with colorectal and ovarian cancer, idiopathic lung fibrosis, and normal human tissues. Scientif. Rep. 5, 8057 (2015).
  150. Wang, Y. H., Yu, X. H., Luo, S. S. & Han, H. Comprehensive circular RNA profiling reveals that circular RNA100783 is involved in chronic CD28-associated CD8+ T cell ageing. Immun. Ageing 12, 17 (2015).
  151. Li, J. et al. Circular RNAs in cancer: novel insights into origins, properties, functions and implications. Am. J. Cancer Res. 5, 472480 (2015).
  152. Memczak, S., Papavasileiou, P., Peters, O. & Rajewsky, N. Identification and characterization of circular RNAs as a new class of putative biomarkers in human blood. PLoS ONE 10, e0141214 (2015).
  153. Dvinge, H. & Bradley, R. K. Widespread intron retention diversifies most cancer transcriptomes. Genome Med. 7, 45 (2015).
  154. Jung, H. et al. Intron retention is a widespread mechanism of tumor-suppressor inactivation. Nat. Genet. 47, 12421248 (2015).
  155. Romano, M., Buratti, E. & Baralle, D. Role of pseudoexons and pseudointrons in human cancer. Int. J. Cell Biol. 2013, 810572 (2013).
  156. Hsu, T. Y. et al. The spliceosome is a therapeutic vulnerability in MYC-driven cancer. Nature 525, 384388 (2015).
    In this study, the spliceosome is found to be a target of oncogenic stress in MYC-dependent breast cancers, providing an opportunity for genetic or pharmacological spliceosome inhibition.
  157. Ilagan, J. O. et al. U2AF1 mutations alter splice site recognition in hematological malignancies. Genome Res. 25, 1426 (2015).
  158. Milde-Langosch, K., Kappes, H., Riethdorf, S., Loning, T. & Bamberger, A. M. FosB is highly expressed in normal mammary epithelia, but down-regulated in poorly differentiated breast carcinomas. Breast Cancer Res. Treat. 77, 265275 (2003).
  159. Rickman, D. S. et al. SLC45A3ELK4 is a novel and frequent erythroblast transformation-specific fusion transcript in prostate cancer. Cancer Res. 69, 27342738 (2009).
  160. Zhang, Y. et al. Chimeric transcript generated by cis-splicing of adjacent genes regulates prostate cancer cell proliferation. Cancer Discov. 2, 598607 (2012).
  161. Bonnal, S., Vigevani, L. & Valcarcel, J. The spliceosome as a target of novel antitumour drugs. Nat. Rev. Drug Discov. 11, 847859 (2012).
  162. Koh, C. M. et al. MYC regulates the core pre-mRNA splicing machinery as an essential step in lymphomagenesis. Nature 523, 96100 (2015).
  163. Dominski, Z. & Kole, R. Restoration of correct splicing in thalassemic pre-mRNA by antisense oligonucleotides. Proc. Natl Acad. Sci. USA 90, 86738677 (1993).
  164. Hua, Y. et al. Peripheral SMN restoration is essential for long-term rescue of a severe spinal muscular atrophy mouse model. Nature 478, 123126 (2011).
  165. McClorey, G. & Wood, M. J. An overview of the clinical application of antisense oligonucleotides for RNA-targeting therapies. Curr. Opin. Pharmacol. 24, 5258 (2015).
  166. Goyenvalle, A. et al. Rescue of dystrophic muscle through U7 snRNA-mediated exon skipping. Science 306, 17961799 (2004).
  167. Gorman, L., Suter, D., Emerick, V., Schumperli, D. & Kole, R. Stable alteration of pre-mRNA splicing patterns by modified U7 small nuclear RNAs. Proc. Natl Acad. Sci. USA 95, 49294934 (1998).
  168. Uchikawa, H. et al. U7 snRNA-mediated correction of aberrant splicing caused by activation of cryptic splice sites. J. Hum. Genet. 52, 891897 (2007).
  169. Blazquez, L. et al. In vitro correction of a pseudoexon-generating deep intronic mutation in LGMD2A by antisense oligonucleotides and modified small nuclear RNAs. Hum. Mutat. 34, 13871395 (2013).
  170. Goyenvalle, A., Babbs, A., van Ommen, G. J., Garcia, L. & Davies, K. E. Enhanced exon-skipping induced by U7 snRNA carrying a splicing silencer sequence: promising tool for DMD therapy. Mol. Ther. 17, 12341240 (2009).
  171. Garcia-Blanco, M. A., Baraniak, A. P. & Lasda, E. L. Alternative splicing in disease and therapy. Nat. Biotechnol. 22, 535546 (2004).
  172. Xu, L. et al. CRISPR-mediated genome editing restores dystrophin expression and function in mdx mice. Mol. Ther. 24, 564569 (2015).
  173. Nelson, C. E. et al. In vivo genome editing improves muscle function in a mouse model of Duchenne muscular dystrophy. Science 351, 403407 (2015).
  174. Tabebordbar, M. et al. In vivo gene editing in dystrophic mouse muscle and muscle stem cells. Science 351, 407411 (2015).
  175. Puttaraju, M., DiPasquale, J., Baker, C. C., Mitchell, L. G. & Garcia-Blanco, M. A. Messenger RNA repair and restoration of protein function by spliceosome-mediated RNA trans-splicing. Mol. Ther. 4, 105114 (2001).
  176. Koller, U., Wally, V., Bauer, J. W. & Murauer, E. M. Considerations for a successful RNA trans-splicing repair of genetic disorders. Mol. Ther. Nucleic Acids 3, e157 (2014).
  177. Chao, H. et al. Phenotype correction of hemophilia A mice by spliceosome-mediated RNA trans-splicing. Nat. Med. 9, 10151019 (2003).
  178. Petkovic, S. & Muller, S. RNA circularization strategies in vivo and in vitro. Nucleic Acids Res. 43, 24542465 (2015).
  179. Davidson, L., Kerr, A. & West, S. Co-transcriptional degradation of aberrant pre-mRNA by Xrn2. EMBO J. 31, 25662578 (2012).
  180. Shen, S. et al. Widespread establishment and regulatory impact of Alu exons in human genes. Proc. Natl Acad. Sci. USA 108, 28372842 (2011).
  181. Tajnik, M. et al. Intergenic Alu exonisation facilitates the evolution of tissue-specific transcript ends. Nucleic Acids Res. 43, 1049210505 (2015).
  182. Rybak-Wolf, A. et al. Circular RNAs in the mammalian brain are highly abundant, conserved, and dynamically expressed. Mol. Cell 58, 870885 (2015).
  183. Kellis, M. et al. Defining functional DNA elements in the human genome. Proc. Natl Acad. Sci. USA 111, 61316138 (2014).
  184. Naftelberg, S., Schor, I. E., Ast, G. & Kornblihtt, A. R. Regulation of alternative splicing through coupling with transcription and chromatin structure. Annu. Rev. Biochem. 84, 165198 (2015).
  185. Pulyakhina, I. et al. SplicePie: a novel analytical approach for the detection of alternative, non-sequential and recursive splicing. Nucleic Acids Res. 43, e80 (2015).
  186. Chuang, T. J. et al. NCLscan: accurate identification of non-co-linear transcripts (fusion, trans-splicing and circular RNA) with a good balance between sensitivity and precision. Nucleic Acids Res. 44, e29 (2015).
  187. Szabo, L. et al. Statistically based splicing detection reveals neural enrichment and tissue-specific induction of circular RNA during human fetal development. Genome Biol. 16, 126 (2015).
  188. McPherson, A. et al. deFuse: an algorithm for gene fusion discovery in tumor RNA-seq data. PLoS Comput. Biol. 7, e1001138 (2011).
  189. Maher, C. A. et al. Transcriptome sequencing to detect gene fusions in cancer. Nature 458, 97101 (2009).
  190. Konig, J. et al. iCLIP reveals the function of hnRNP particles in splicing at individual nucleotide resolution. Nature Struct. Mol. Biol. 17, 909915 (2010).
  191. Alioto, T. S. U12DB: a database of orthologous U12-type spliceosomal introns. Nucleic Acids Res. 35, D110D115 (2007).
  192. Crooks, G. E., Hon, G., Chandonia, J. M. & Brenner, S. E. WebLogo: a sequence logo generator. Genome Res. 14, 11881190 (2004).

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Author information

  1. These authors contributed equally to this work.

    • Christopher R. Sibley &
    • Lorea Blazquez


  1. Department of Molecular Neuroscience, University College London Institute of Neurology, Russell Square House, Russell Square, London WC1B 5EH, UK.

    • Christopher R. Sibley,
    • Lorea Blazquez &
    • Jernej Ule
  2. Department of Medicine, Division of Brain Sciences, Imperial College London, Burlington Danes, DuCane Road, London W12 0NN, UK.

    • Christopher R. Sibley

Competing interests statement

The authors declare no competing interests.

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Author details

  • Christopher R. Sibley

    Christopher R. Sibley is a junior group leader in the Division of Brain Sciences, Imperial College London, UK. During his postdoctoral work with Jernej Ule he studied post-transcriptional processing of RNA in the nervous system using the functional genomics approaches of individual-nucleotide-resolution crosslinking and immunoprecipitation (iCLIP) and RNA sequencing. This led to the discovery of the first reported mammalian recursive splice sites. He continues to research the numerous unannotated and non-canonical features of the transcriptome discussed in this Review. In addition, he uses functional genomics and systems biology approaches to study master regulators in neurological diseases.

  • Lorea Blazquez

    Lorea Blazquez completed her Ph.D. at the Biodonostia Institute, San Sebastian, Spain, where she worked on identification, functional characterization and therapeutics of splicing mutations. She then studied strategies to inhibit gene expression with small RNA molecules at Center for Applied Medical Research (CIMA), Pamplona, Spain. She joined the Ule laboratory to study the function and regulation of recursive splicing.

  • Jernej Ule

    Jernej Ule is Professor of Molecular Neuroscience at the University College London Institute of Neurology, London, UK. He has a long-standing interest in mechanisms of ribonucleoprotein (RNP) complexes and RNA structure in the regulation of pre-mRNA processing and mRNA translation. His group developed iCLIP (individual-nucleotide-resolution crosslinking and immunoprecipitation) and hiCLIP (RNA hybrid and iCLIP), which are methods to study in vivo protein–RNA and RNA–RNA contacts in a transcriptome-wide manner. Recently, the group has begun to focus on the regulation and functions of cryptic splicing events, such as those generated by transposable elements or recursive splice sites. Jernej Ule's homepage.

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