Splicing mainly takes place during transcription. Catalysis is achieved through the stepwise assembly of the spliceosome onto nascent RNA.
Recent in vivo studies have shown that the catalytic spliceosome is physically close to RNA polymerase II, underscoring that spliceosome assembly and splicing catalysis occur on similar timescales to transcription.
Transcription elongation rate and pausing influence splice site availability and consequent spliceosome assembly. DNA sequence, nucleosome positioning, chromatin structure and post-translational modifications of the RNA polymerase II carboxy-terminal domain influence transcription dynamics.
Splicing is coupled to other pre-mRNA processing events, such as 5′ end capping, 3′ end processing and RNA editing, thereby ensuring the efficient and precise production of mature mRNAs.
Efficient co-transcriptional processing of nascent RNA may be achieved by concentrating transcription and processing machineries in subnuclear membrane-less compartments.
Several macromolecular machines collaborate to produce eukaryotic messenger RNA. RNA polymerase II (Pol II) translocates along genes that are up to millions of base pairs in length and generates a flexible RNA copy of the DNA template. This nascent RNA harbours introns that are removed by the spliceosome, which is a megadalton ribonucleoprotein complex that positions the distant ends of the intron into its catalytic centre. Emerging evidence that the catalytic spliceosome is physically close to Pol II in vivo implies that transcription and splicing occur on similar timescales and that the transcription and splicing machineries may be spatially constrained. In this Review, we discuss aspects of spliceosome assembly, transcription elongation and other co-transcriptional events that allow the temporal coordination of co-transcriptional splicing.
Subscribe to Journal
Get full journal access for 1 year
only $4.92 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Sakharkar, M. K., Perumal, B., S., Sakharkar, K. R. & Kangueane, P. An analysis on gene architecture in human and mouse genomes. In Silico Biol. 5, 347–365 (2005).
Wahl, M. C., Will, C. L. & Lührmann, R. The spliceosome: design principles of a dynamic RNP machine. Cell 136, 701–718 (2009).
Brugiolo, M., Herzel, L. & Neugebauer, K. M. Counting on co-transcriptional splicing. F1000 Prime Reports 5, 9 (2013).
Carrillo Oesterreich, F. et al. Splicing of nascent RNA coincides with intron exit from RNA polymerase II. Cell 165, 372–381 (2016). Shows a tight match between progress in transcription and splicing completion in vivo.
Alpert, T., Herzel, L. & Neugebauer, K. M. Perfect timing: splicing and transcription rates in living cells. Wiley Interdiscip. Rev. RNA 8, e1401 (2016).
Carrillo Oesterreich, F., Bieberstein, N. & Neugebauer, K. M. Pause locally, splice globally. Trends Cell Biol. 21, 328–335 (2011).
Bieberstein, N. I., Carrillo Oesterreich, F., Straube, K. & Neugebauer, K. M. First exon length controls active chromatin signatures and transcription. Cell Rep. 2, 62–68 (2012).
Braunschweig, U., Gueroussov, S., Plocik, A. M., Graveley, B. R. & Blencowe, B. J. Dynamic integration of splicing within gene regulatory pathways. Cell 152, 1252–1269 (2013).
Moehle, E. A., Braberg, H., Krogan, N. J. & Guthrie, C. Adventures in time and space: splicing efficiency and RNA polymerase II elongation rate. RNA Biol. 11, 313–319 (2014).
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, 165–198 (2015).
Custódio, N. & Carmo-Fonseca, M. Co-transcriptional splicing and the CTD code. Crit. Rev. Biochem. Mol. Biol. 51, 395–411 (2016).
Saldi, T., Cortazar, M. A., Sheridan, R. M. & Bentley, D. L. Coupling of RNA polymerase II transcription elongation with pre-mRNA splicing. J. Mol. Biol. 428, 2623–2635 (2016).
Wallace, E. & Beggs, J. Extremely fast and incredibly close: co-transcriptional splicing in budding yeast. RNA 23, 601–610 (2017).
Lander, E. S. et al. International human genome sequencing consortium. Nature 409, 860–921 (2001).
Gould, G. M. et al. Identification of new branch points and unconventional introns in Saccharomyces cerevisiae. RNA 22, 1522–1534 (2016).
Qin, D., Huang, L., Wlodaver, A., Andrade, J. & Staley, J. P. Sequencing of lariat termini in S. cerevisiae reveals 5′ splice sites, branch points, and novel splicing events. RNA 22, 237–253 (2016).
Mercer, T. R. et al. Genome-wide discovery of human splicing branchpoints. Genome Res. 25, 290–303 (2015).
Taggart, A. J. et al. Large-scale analysis of branchpoint usage across species and cell lines. Genome Res. 27, 639–649 (2017).
Mount, S. M. A catalogue of splice junction sequences. Nucleic Acids Res. 10, 459–472 (1982).
Lim, L. & Burge, C. A computational analysis of sequence features involved in recognition of short introns. Proc. Natl Acad. Sci. USA 98, 11193–11198 (2001).
Reed, R. & Maniatis, T. Intron sequences involved in lariat formation during pre-mRNA splicing. Cell 41, 95–105 (1985).
Moore, M., Query, C. & Sharp, P. in The RNA World (eds Gesteland, R. F. & Atkins, J. F.) 303–357 (Cold Spring Harbor Laboratory Press, 1993).
Will, C. L. & Lührmann, R. Spliceosome structure and function. Cold Spring Harb. Perspect. Biol. 3, a003707 (2011).
Cvitkovic, I. & Jurica, M. S. Spliceosome database: a tool for tracking components of the spliceosome. Nucleic Acids Res. 41, D132–D141 (2013).
Cordin, O. & Beggs, J. D. RNA helicases in splicing. RNA Biol. 10, 83–95 (2013).
Fabrizio, P. et al. The evolutionarily conserved core design of the catalytic activation step of the yeast spliceosome. Mol. Cell 36, 593–608 (2009).
Kotovic, K. M., Lockshon, D., Boric, L. & Neugebauer, K. M. Cotranscriptional recruitment of the U1 snRNP to intron-containing genes in yeast. Mol. Cell. Biol. 23, 5768–5779 (2003).
Görnemann, J., Kotovic, K. M., Hujer, K. & Neugebauer, K. M. Cotranscriptional spliceosome assembly occurs in a stepwise fashion and requires the cap binding complex. Mol. Cell 19, 53–63 (2005).
Lacadie, S. A. & Rosbash, M. Cotranscriptional spliceosome assembly dynamics and the role of U1 snRNA:5′ss base pairing in yeast. Mol. Cell 19, 65–75 (2005).
Tardiff, D. F., Lacadie, S. A. & Rosbash, M. A genome-wide analysis indicates that yeast pre-mRNA splicing is predominantly posttranscriptional. Mol. Cell 24, 917–929 (2006). Analyses co-transcriptional recruitment of U snRNPs onto nascent RNA.
Újvári, A. & Luse, D. S. Newly Initiated RNA encounters a factor involved in splicing immediately upon emerging from within RNA polymerase II. J. Biol. Chem. 279, 49773–49779 (2004).
Baejen, C. et al. Transcriptome maps of mRNP biogenesis factors define pre-mRNA recognition. Mol. Cell 55, 745–757 (2014). Systematic mapping of the binding sites of several factors involved in mRNP biogenesis on yeast transcripts.
Rigo, N., Sun, C., Fabrizio, P., Kastner, B. & Lührmann, R. Protein localisation by electron microscopy reveals the architecture of the yeast spliceosomal B complex. EMBO J. 34, 3059–3073 (2015).
Nguyen, T. H. et al. Cryo-EM structure of the yeast U4/U6. U5 tri-snRNP at 3.7 Å resolution. Nature 530, 298–302 (2016).
Nguyen, T. H. et al. The architecture of the spliceosomal U4/U6. U5 tri-snRNP. Nature 523, 47–52 (2015).
Wan, R. et al. The 3.8 Å structure of the U4/U6. U5 tri-snRNP: insights into spliceosome assembly and catalysis. Science 351, 466–475 (2016).
Liu, H. L. & Cheng, S. C. The interaction of Prp2 with a defined region of the intron is required for the first splicing reaction. Mol. Cell. Biol. 32, 5056–5066 (2012).
Rymond, B. C. & Rosbash, M. Cleavage of 5′ splice site and lariat formation are independent of 3′ splice site in yeast splicing. Nature 317, 735–737 (1985).
Yan, C., Wan, R., Bai, R., Huang, G. & Shi, Y. Structure of a yeast activated spliceosome at 3.5 Å resolution. Science 353, 905–911 (2016).
Rauhut, R. et al. Molecular architecture of the Saccharomyces cerevisiae activated spliceosome. Science 353, 1399–1405 (2016). Cryo-EM structure of the Bact complex, revealing details of spliceosome active site conformation before step I catalysis and suggesting that conformation rearrangement is needed for catalysis.
Schneider, C. et al. Dynamic contacts of U2, RES, Cwc25, Prp8 and Prp45 proteins with the pre-mRNA branch-site and 3′ splice site during catalytic activation and step 1 catalysis in yeast spliceosomes. PLoS Genet. 11, e1005539 (2015). Evaluates direct interactions between splicing proteins and pre-mRNA at different stages of spliceosome assembly.
Ruskin, B. & Green, M. Role of the 3′ splice site consensus sequence in mammalian pre-mRNA splicing. Nature 317, 732–734 (1985).
Wan, R., Yan, C., Bai, R., Huang, G. & Shi, Y. Structure of a yeast catalytic step I spliceosome at 3.4 Å resolution. Science 353, 895–904 (2016).
Galej, W. P. et al. Cryo-EM structure of the spliceosome immediately after branching. Nature 537, 197–201 (2016).
Schwer, B. A conformational rearrangement in the spliceosome sets the stage for Prp22-dependent mRNA release. Mol. Cell 30, 743–754 (2008).
Ohrt, T. et al. Molecular dissection of step 2 catalysis of yeast pre-mRNA splicing investigated in purified system. RNA 19, 902–915 (2013).
Yan, C., Wan, R., Bai, R., Huang, G. & Shi, Y. Structure of a yeast step II catalytically activated spliceosome. Science 355, 149–155 (2017). Cryo-EM structure of C* complex, revealing details of spliceosome active site conformation before step II catalysis and suggesting that conformation rearrangement is needed for catalysis.
Fica, S. M. et al. Structure of a spliceosome remodelled for exon ligation. Nature 542, 377–380 (2017).
Bertram, K. et al. Cryo-EM structure of a human spliceosome activated for step 2 of splicing. Nature 542, 318–323 (2017).
Smith, C., Chu, T. & Nadal-Ginard, B. Scanning and competition between AGs are involved in 3′ splice site selection in mammalian introns. Mol. Cell. Biol. 13, 4939–4952 (1993).
Gahura, O., Hammann, C., Valentová, A., Puta, F. & Folk, P. Secondary structure is required for 3′ splice site recognition in yeast. Nucleic Acids Res. 39, 9759–9767 (2011).
Meyer, M., Plass, M., Peréz-Valle, J., Eyras, E. & Vilardell, J. Deciphering 3′ss selection in the yeast genome reveals an RNA thermosensor that mediates alternative splicing. Mol. Cell 43, 1033–1039 (2011). Analyses the role of RNA secondary structure in 3′SS selection.
Martinez-Rucobo, F. W. et al. Molecular basis of transcription-coupled pre-mRNA capping. Mol. Cell 58, 1079–1089 (2015).
Fourmann, J. B. et al. Dissection of the factor requirements for spliceosome disassembly and the elucidation of its dissociation products using a purified splicing system. Genes Dev. 27, 413–428 (2013).
Kwak, H. & Lis, J. T. Control of transcriptional elongation. Annu. Rev. Genet. 47, 483–508 (2013).
Jonkers, I. & Lis, J. T. Getting up to speed with transcription elongation by RNA polymerase II. Nat. Rev. Mol. Cell Biol. 16, 167–177 (2015).
Neves, L. T. et al. The histone variant H2A.Z promotes efficient cotranscriptional splicing in S. cerevisiae. Genes Dev. 31, 702–717 (2017).
Nissen, K. E. et al. The histone variant H2A.Z promotes splicing of weak introns. Genes Dev. 31, 688–701 (2017).
Zaborowska, J., Egloff, S. & Murphy, S. The pol II CTD: new twists in the tail. Nat. Struct. Mol. Biol. 23, 771–777 (2016).
Harlen, K. M. & Churchman, L. S. The code and beyond: transcription regulation by the RNA polymerase II carboxy-terminal domain. Nat. Rev. Mol. Cell Biol. 18, 263–273 (2017).
Allison, L., Moyle, M., Shales, M. & Ingles, C. Extensive homology among the largest subunits of eukaryotic and prokaryotic RNA polymerases. Cell 42, 599–610 (1985).
Corden, J., Cadena, D., Ahearn, J. & Dahmus, M. A unique structure at the carboxyl terminus of the largest subunit of eukaryotic RNA polymerase II. Proc. Natl Acad. Sci. USA 82, 7934–7938 (1985).
Kim, H. et al. Gene-specific RNA polymerase II phosphorylation and the CTD code. Nat. Struct. Mol. Biol. 17, 1279–1286 (2010).
Mayer, A. et al. Uniform transitions of the general RNA polymerase II transcription complex. Nat. Struct. Mol. Biol. 17, 1272–1278 (2010).
Mayer, A. et al. CTD tyrosine phosphorylation impairs termination factor recruitment to RNA polymerase II. Science 336, 1723–1725 (2012).
Harlen, K. M. et al. Comprehensive RNA polymerase II interactomes reveal distinct and varied roles for each phospho-CTD residue. Cell Rep. 15, 2147–2158 (2016). Analyses phospho-specific Pol II C-terminal domain interactomes by mass spectrometry.
Milligan, L. et al. Strand-specific, high-resolution mapping of modified RNA polymerase II. Mol. Syst. Biol. 12, 874 (2016). Identifies distinct Pol II C-terminal domain PTMs associated with transcription initiation, and early and late elongation.
Singh, G., Pratt, G., Yeo, G. W. & Moore, M. J. The clothes make the mRNA: past and present trends in mRNP fashion. Annu. Rev. Biochem. 84, 325–354 (2015).
Müller-McNicoll, M. & Neugebauer, K. M. How cells get the message: dynamic assembly and function of mRNA-protein complexes. Nat. Rev. Genet. 14, 275–287 (2013).
König, J. et al. iCLIP reveals the function of hnRNP particles in splicing at individual nucleotide resolution. Nat. Struct. Mol. Biol. 17, 909–915 (2010).
Descostes, N. et al. Tyrosine phosphorylation of RNA polymerase II CTD is associated with antisense promoter transcription and active enhancers in mammalian cells. eLife 3, e02105 (2014).
Schüller, R. et al. Heptad-specific phosphorylation of RNA polymerase II CTD. Mol. Cell 61, 305–314 (2016).
Suh, H. et al. Direct analysis of phosphorylation sites on the Rpb1 C-terminal domain of RNA polymerase II. Mol. Cell 61, 297–304 (2016).
Nojima, T. et al. Mammalian NET-seq reveals genome-wide nascent transcription coupled to RNA processing. Cell 161, 526–540 (2015).
Mayer, A. et al. Native elongating transcript sequencing reveals human transcriptional activity at nucleotide resolution. Cell 161, 541–554 (2015).
Veloso, A. et al. Rate of elongation by RNA polymerase II is associated with specific gene features and epigenetic modifications. Genome Res. 24, 896–905 (2014).
Nedelcheva-Veleva, M. N. et al. The thermodynamic patterns of eukaryotic genes suggest a mechanism for intron–exon recognition. Nat. Commun. 4, 2101 (2013).
Schor, I. E., Fiszbein, A., Petrillo, E. & Kornblihtt, A. R. Intragenic epigenetic changes modulate NCAM alternative splicing in neuronal differentiation. EMBO J. 32, 2264–2274 (2013).
Tilgner, H. et al. Nucleosome positioning as a determinant of exon recognition. Nat. Struct. Mol. Biol. 16, 996–1001 (2009).
Schwartz, S., Meshorer, E. & Ast, G. Chromatin organization marks exon–intron structure. Nat. Struct. Mol. Biol. 16, 990–995 (2009).
Huff, J. T., Zilberman, D. & Roy, S. W. Mechanism for DNA transposons to generate introns on genomic scales. Nature 538, 533–536 (2016). Explains the correlation between internal exon length and nucleosome DNA length.
Beckmann, J. & Trifonov, E. Splice junctions follow a 205-base ladder. Proc. Natl Acad. Sci. USA 88, 2380–2383 (1991).
Venkatesh, S. & Workman, J. L. Histone exchange, chromatin structure and the regulation of transcription. Nat. Rev. Mol. Cell Biol. 16, 178–189 (2015).
Weber, C. M., Ramachandran, S. & Henikoff, S. Nucleosomes are context-specific, H2A.Z-modulated barriers to RNA polymerase. Mol. Cell 53, 819–830 (2014). Describes the effects of nucleosomes on transcription elongation dynamics in vivo.
Churchman, L. S. & Weissman, J. S. Nascent transcript sequencing visualizes transcription at nucleotide resolution. Nature 469, 368–373 (2011). Describes the NET-seq method, which enables the analysis of transcription elongation dynamics at high resolution.
Kwak, H., Fuda, N., Core, L. J. & Lis, J. T. Precise maps of RNA polymerase reveal how promoters direct initiation and pausing. Science 339, 950–953 (2013). Describes the PRO-seq assay, which enables the analysis of transcription elongation dynamics at high resolution.
Voong, L. N. et al. Insights into nucleosome organization in mouse embryonic stem cells through chemical mapping. Cell 167, 1555–1570.e15 (2016).
Fuchs, G., Hollander, D., Voichek, Y., Ast, G. & Oren, M. Cotranscriptional histone H2B monoubiquitylation is tightly coupled with RNA polymerase II elongation rate. Genome Res. 24, 1572–1583 (2014).
Luco, R. F. & Misteli, T. More than a splicing code: integrating the role of RNA, chromatin and non-coding RNA in alternative splicing regulation. Curr. Opin. Genet. Dev. 21, 366–372 (2011).
Kfir, N. et al. SF3B1 association with chromatin determines splicing outcomes. Cell Rep. 11, 618–629 (2015).
Hnilicová, J. et al. Histone deacetylase activity modulates alternative splicing. PLoS ONE 6, e16727 (2011).
Herissant, L. et al. H2B ubiquitylation modulates spliceosome assembly and function in budding yeast. Biol. Cell 106, 126–138 (2014).
Carrillo Oesterreich, F., Preibisch, S. & Neugebauer, K. M. Global analysis of nascent RNA reveals transcriptional pausing in terminal exons. Mol. Cell 40, 571–581 (2010).
Chathoth, K. T., Barrass, J. D., Webb, S. & Beggs, J. D. A splicing-dependent transcriptional checkpoint associated with prespliceosome formation. Mol. Cell 53, 779–790 (2014).
Alexander, R. D., Innocente, S. A., Barrass, J. D. & Beggs, J. D. Splicing-dependent RNA polymerase pausing in yeast. Mol. Cell 40, 582–593 (2010).
Újvári, A. & Luse, D. S. RNA emerging from the active site of RNA polymerase II interacts with the Rpb7 subunit. Nat. Struct. Mol. Biol. 13, 49–54 (2006).
Lai, D., Proctor, J. R. & Meyer, I. M. On the importance of cotranscriptional RNA structure formation. RNA 19, 1461–1473 (2013).
Liu, S. R., Hu, C. G. & Zhang, J. Z. Regulatory effects of cotranscriptional RNA structure formation and transitions. Wiley Interdiscip. Rev. RNA 7, 562–574 (2016).
Buratti, E. & Baralle, F. E. Influence of RNA secondary structure on the pre-mRNA splicing process. Mol. Cell. Biol. 24, 10505–10514 (2004).
Warf, M. B. & Berglund, J. A. Role of RNA structure in regulating pre-mRNA splicing. Trends Biochem. Sci. 35, 169–178 (2010).
Eperon, L., Graham, I., Griffiths, A. & Eperon, I. Effects of RNA secondary structure on alternative splicing of pre-mRNA: is folding limited to a region behind the transcribing RNA polymerase? Cell 54, 393–401 (1988).
Goguel, V., Wang, Y. & Rosbash, M. Short artificial hairpins sequester splicing signals and inhibit yeast pre-mRNA splicing. Mol. Cell. Biol. 13, 6841–6848 (1993).
Deshler, J. & Rossi, J. Unexpected point mutations activate cryptic 3′ splice sites by perturbing a natural secondary structure within a yeast intron. Genes Dev. 5, 1252–1263 (1991).
Charpentier, B. & Rosbash, M. Intramolecular structure in yeast introns aids the early steps of in vitro spliceosome assembly. RNA 2, 509–522 (1996).
Nilsen, T. Internal mRNA methylation finally finds functions. Science 343, 1207–1208 (2014).
Reenan, R. Molecular determinants and guided evolution of species-specific RNA editing. Nature 434, 409–413 (2005).
Peng, Z. et al. Comprehensive analysis of RNA-seq data reveals extensive RNA editing in a human transcriptome. Nat. Biotechnol. 30, 253–260 (2012).
Rodriguez, J., Menet, J. S. & Rosbash, M. Nascent-seq indicates widespread cotranscriptional RNA editing in Drosophila. Mol. Cell 47, 27–37 (2012).
Wang, I. X. et al. RNA–DNA differences are generated in human cells within seconds after RNA exits polymerase II. Cell Rep. 6, 906–915 (2014). Shows that RNA nucleotides can be edited as soon as they emerge from Pol II.
Rieder, L. E. & Reenan, R. A. The intricate relationship between RNA structure, editing, and splicing. Semin. Cell Dev. Biol. 23, 281–288 (2012).
Laurencikiene, J., Källman, A. M., Fong, N., Bentley, D. L. & Öhman, M. RNA editing and alternative splicing: the importance of co-transcriptional coordination. EMBO Rep. 7, 303–307 (2006).
Bratt, E. & Öhman, M. Coordination of editing and splicing of glutamate receptor pre-mRNA. RNA 9, 309–318 (2003).
Solomon, O. et al. Global regulation of alternative splicing by adenosine deaminase acting on RNA (ADAR). RNA 19, 591–604 (2013).
Hawkins, J. A survey on intron and exon lengths. Nucleic Acids Res. 16, 9893–9908 (1988).
Deutsch, M. & Long, M. Intron–exon structures of eukaryotic model organisms. Nucleic Acids Res. 27, 3219–3228 (1999).
Wang, Z. & Burge, C. B. Splicing regulation: from a parts list of regulatory elements to an integrated splicing code. RNA 14, 802–813 (2008).
Zhang, X. H.-F., Leslie, C. & Chasin, L. Dichotomous splicing signals in exon flanks. Genome Res. 15, 768–779 (2005).
Barash, Y. et al. Deciphering the splicing code. Nature 465, 53–59 (2010).
Berget, S. Exon recognition in vertebrate splicing. J. Biol. Chem. 270, 2411–2414 (1995).
Robberson, B., Cote, G. & Berget, S. Exon definition may facilitate splice site selection in RNAs with multiple exons. Mol. Cell. Biol. 10, 84–94 (1990).
Talerico, M. & Berget, S. Intron definition in splicing of small Drosophila introns. Mol. Cell. Biol. 14, 3434–3445 (1994).
Sterner, D., Carlo, T. & Berget, S. Architectural limits on plit genes. Proc. Natl Acad. Sci. USA 93, 15081–15085 (1996).
Izaurralde, E. et al. A nuclear cap binding protein complex involved in pre-mRNA splicing. Cell 78, 657–668 (1994).
Izaurralde, E. et al. A cap-binding protein complex mediating U snRNA export. Nature 376, 709–712 (1995).
Rottman, F., Shatkin, A. & Perry, R. Sequences containing methylated nucleotides at the 5′ termini of messenger RNAs: possible implications in processing. Cell 3, 197–199 (1974).
Ramanathan, A., Robb, G. B. & Chan, S. H. mRNA capping: biological functions and applications. Nucleic Acids Res. 44, 7511–7526 (2016).
Rasmussen, E. & Lis, J. T. In vivo transcriptional pausing and cap formation on three Drosophila heat shock genes. Proc. Natl Acad. Sci. USA 90, 7923–7927 (1993).
Gonatopoulos-Pournatzis, T. & Cowling, V. H. Cap-binding complex (CBC). Biochem. J. 457, 231–242 (2014).
Konarska, M. M., Padgett, R. & Sharp, P. A. Recognition of cap structure in splicing in vitro of mRNA precursors. Cell 38, 731–736 (1984).
Pabis, M. et al. The nuclear cap-binding complex interacts with the U4/U6.U5 tri-snRNP and promotes spliceosome assembly in mammalian cells. RNA 19, 1054–1063 (2013). Shows an association between the cap-binding complex and snRNPs.
Lewis, J., Izaurralde, E., Jarmolozski, A., McGuigan, C. & Mattaj, I. A nuclear cap-binding complex facilitates association of U1 snRNP with the cap-proximal 5′ splice site. Genes Dev. 10, 1683–1698 (1996).
Qiu, Z. R., Chico, L., Chang, J., Shuman, S. & Schwer, B. Genetic interactions of hypomorphic mutations in the m7G cap-binding pocket of yeast nuclear cap binding complex: an essential role for Cbc2 in meiosis via splicing of MER3 pre-mRNA. RNA 18, 1996–2011 (2012).
Proudfoot, N. J. Transcriptional termination in mammals: stopping the RNA polymerase II juggernaut. Science 352, 1291–1300 (2016).
Connelly, S. & Manley, J. L. A functional mRNA polyadenylation signal is required for transcription termination by RNA polymerase II. Genes Dev. 2, 440–452 (1988).
Logan, J., Falck-Pedersen, E., Darnell, J. & Shenk, T. A poly(A) addition site and a downstream termination region are required for efficient cessation of transcription by RNA polymerase II in the mouse βmaj-globin gene. Proc. Natl Acad. Sci. USA 84, 8306–8310 (1987).
Whitelaw, E. & Proudfoot, N. α-Thalassemia caused by a poly(A) site mutation reveals that transcriptional termination is linked to 3′ end processing in the human α2 globin gene. EMBO J. 5, 2915–2922 (1986).
Cooke, C., Hans, H. & Alwine, J. Utilization of splicing elements and polyadenylation signal elements in the coupling of polyadenylation and last-intron removal. Mol. Cell. Biol. 19, 4971–4979 (1999).
Rigo, F. & Martinson, H. G. Functional coupling of last-intron splicing and 3′-end processing to transcription in vitro: the poly(A) signal couples to splicing before committing to cleavage. Mol. Cell. Biol. 28, 849–862 (2008).
Kyburz, A., Friedlein, A., Langen, H. & Keller, W. Direct interactions between subunits of CPSF and the U2 snRNP contribute to the coupling of pre-mRNA 3′ end processing and splicing. Mol. Cell 23, 195–205 (2006).
Millevoi, S. et al. A novel function for the U2AF 65 splicing factor in promoting pre-mRNA 3′-end processing. EMBO Rep. 3, 869–874 (2002).
Vagner, S., Vagner, C. & Mattaj, I. The carboxyl terminus of vertebrate poly(A) polymerase interacts with U2AF 65 to couple 3′-end processing and splicing. Genes Dev. 14, 403–413 (2000).
Kaida, D. The reciprocal regulation between splicing and 3′-end processing. Wiley Interdiscip. Rev. RNA 7, 499–511 (2016).
Dye, M. & Proudfoot, N. J. Terminal exon definition occurs cotranscriptionally and promotes termination of RNA polymerase II. Mol. Cell 3, 371–378 (1999).
Davidson, L. & West, S. Splicing-coupled 3′ end formation requires a terminal splice acceptor site, but not intron excision. Nucleic Acids Res. 41, 7101–7114 (2013). Shows that early stages of spliceosome assembly are sufficient to couple splicing to 3′ end processing.
Bento Martins, S. et al. Spliceosome assembly is coupled to RNA polymerase II dynamics at the 3′ end of human genes. Nat. Struct. Mol. Biol. 18, 1115–1123 (2011).
Bird, G., Zorio, D. A. & Bentley, D. L. RNA polymerase II carboxy-terminal domain phosphorylation is required for cotranscriptional pre-mRNA splicing and 3′-end formation. Mol. Cell. Biol. 24, 8963–8969 (2004).
Gunderson, S., Polycarpou-Schwarz, M. & Mattaj, I. U1 snRNP inhibits pre-mRNA polyadenylation through a direct interaction between U1 70K and poly(A) polymerase. Mol. Cell 1, 255–264 (1998).
Kaida, D. et al. U1 snRNP protects pre-mRNAs from premature cleavage and polyadenylation. Nature 468, 664–668 (2010). Shows that U1 snRNP protects the transcriptome by suppressing premature cleavage and polyadenylation from cryptic poly(A) sites.
Baserga, S. & Steitz, J. in The RNA World (eds Gesteland, R. F. & Atkins, J. F.) 359–381 (Cold Spring Harbor Laboratory Press, 1993).
Almada, A. E., Wu, X., Kriz, A. J., Burge, C. B. & Sharp, P. A. Promoter directionality is controlled by U1 snRNP and polyadenylation signals. Nature 499, 360–363 (2013).
Berg, M. G. et al. U1 snRNP determines mRNA length and regulates isoform expression. Cell 150, 53–64 (2012).
Li, W., Notani, D. & Rosenfeld, M. G. Enhancers as non-coding RNA transcription units: recent insights and future perspectives. Nat. Rev. Genet. 17, 207–223 (2016).
El Kaderi, B., Medler, S., Raghunayakula, S. & Ansari, A. Gene looping is conferred by activator-dependent interaction of transcription initiation and termination machineries. J. Biol. Chem. 284, 25015–25025 (2009).
Mukundan, B. & Ansari, A. Srb5/Med18-mediated termination of transcription is dependent on gene looping. J. Biol. Chem. 288, 11384–11394 (2013).
Agarwal, N. & Ansari, A. Enhancement of transcription by a splicing-competent intron is dependent on promoter directionality. PLoS Genet. 12, e1006047 (2016).
Moabbi, A. M., Agarwal, N., El Kaderi, B. & Ansari, A. Role for gene looping in intron-mediated enhancement of transcription. Proc. Natl Acad. Sci. USA 109, 8505–8510 (2012). Shows that introns help gene looping and hence transcription in yeast.
Huranová, M. et al. The differential interaction of snRNPs with pre-mRNA reveals splicing kinetics in living cells. J. Cell Biol. 191, 75–86 (2010).
Phair, R. & Misteli, T. High mobility of proteins in the mammalian nucleus. Nature 404, 604–609 (2000).
Courchaine, E. M., Lu, A. & Neugebauer, K. M. Droplet organelles? EMBO J. 35, 1603–1612 (2016).
Kwon, I. et al. Poly-dipeptides encoded by the C9orf72 repeats bind nucleoli, impede RNA biogenesis, and kill cells. Science 345, 1139–1145 (2014).
He, C. et al. High-resolution mapping of RNA-binding regions in the nuclear proteome of embryonic stem cells. Mol. Cell 64, 416–430 (2016). Identifies and characterizes new protein domains, and unannotated and/or disordered regions that interact with RNA.
Castello, A. et al. Comprehensive identification of RNA-binding domains in human cells. Mol. Cell 63, 696–710 (2016).
Hnisz, D., Shrinivas, K., Young, R. A., Chakraborty, A. K. & Sharp, P. A. A. Phase separation model for transcriptional control. Cell 169, 13–23 (2017).
Neugebauer, K. M., Stolk, J. A. & Roth, M. B. A conserved epitope on a subset of SR proteins defines a larger family of pre-mRNA splicing factors. J. Cell Biol. 129, 899–908 (1995).
Gueroussov, S. et al. Regulatory expansion in mammals of multivalent hnRNP assemplies that globally control alternative splicing. Cell 170, 324–339 (2017).
Ying, Y. et al. Splicing activation by RBfox requires self-aggregation through its tyrosine-rich domain. Cell 170, 312–323 (2017).
Machyna, M., Neugebauer, K. M. & Stane˘k, D. Coilin: the first 25 years. RNA Biol. 12, 590–596 (2015).
Kwon, I. et al. Phosphorylation-regulated binding of RNA polymerase II to fibrous polymers of low-complexity domains. Cell 155, 1049–1060 (2013).
Yu, Y. & Reed, R. FUS functions in coupling transcription to splicing by mediating an interaction between RNAP II and U1 snRNP. Proc. Natl Acad. Sci. USA 112, 8608–8613 (2015).
Sun, S. et al. ALS-causative mutations in FUS/TLS confer gain and loss of function by altered association with SMN and U1-snRNP. Nat. Commun. 6, 6171 (2015).
Radó-Trilla, N. & Albà, M. Dissecting the role of low-complexity regions in the evolution of vertebrate proteins. BMC Evol. Biol. 12, 155 (2012).
Callan, H. Lampbrush Chromosomes Vol. 36 (Springer-Verlag Berlin Heidelberg, 1986).
Custódio, N., Vivo, M., Antoniou, M. & Carmo-Fonseca, M. Splicing- and cleavage-independent requirement of RNA polymerase II CTD for mRNA release from the transcription site. J. Cell Biol. 179, 199–207 (2007).
Custódio, N. et al. Inefficient processing impairs release of RNA from the site of transcription. EMBO J. 18, 2855–2866 (1999).
Bhatt, D. M. et al. Transcript dynamics of proinflammatory genes revealed by sequence analysis of subcellular RNA fractions. Cell 150, 279–290 (2012).
Tress, M. L., Abascal, F. & Valencia, A. Alternative splicing may not be the key to proteome complexity. Trends Biochem. Sci. 42, 98–110 (2017).
Tilgner, H. et al. Comprehensive transcriptome analysis using synthetic long-read sequencing reveals molecular co-association of distant splicing events. Nat. Biotechnol. 33, 736–742 (2015).
Taliaferro, J. M. et al. Distal alternative last exons localize mRNAs to neural projections. Mol. Cell 61, 821–833 (2016).
Floor, S. N. & Doudna, J. A. Tunable protein synthesis by transcript isoforms in human cells. eLife 5, e10921 (2016).
Berkovits, B. D. & Mayr, C. Alternative 3′ UTRs act as scaffolds to regulate membrane protein localization. Nature 522, 363–367 (2015).
Gowda, N. et al. Cytosolic splice isoform of Hsp70 nucleotide exchange factor Fes1 is required for the degradation of misfolded proteins in yeast. Mol. Biol. Cell 27, 1210–1219 (2016).
Pu, S., Wong, J., Turner, B., Cho, E. & Wodak, S. J. Up-to-date catalogues of yeast protein complexes. Nucleic Acids Res. 37, 825–831 (2009).
Huh, W. et al. Global analysis of protein localization in budding yeast. Nature 425, 686–691 (2003).
Dosztányi, Z., Csizmók, V., Tompa, P. & Simon, I. IUPred: web server for the prediction of intrinsically unstructured regions of proteins based on estimated energy content. Bioinformatics 21, 3433–3434 (2005).
Dosztányi, Z., Csizmók, V., Tompa, P. & Simon, I. The pairwise energy content estimated from amino acid composition discriminates between folded and intrinsically unstructured proteins. J. Mol. Biol. 347, 827–839 (2005).
Birse, C., Minvielle-Sebastia, L., Lee, B., Keller, W. & Proudfoot, N. J. Coupling termination of transcription to messenger RNA maturation in yeast. Science 280, 298–301 (1998).
Baejen, C. et al. Genome-wide analysis of RNA polymerase II termination at protein-coding genes. Mol. Cell 66, 38–49 (2017).
The authors are grateful to H. Herzel and K. Reimer for comments on the manuscript. This work was supported in part by NIH R01GM112766 from the National Institute of General Medical Sciences (to K.M.N.) and by T32GM007223 (to T.A.). Its contents are solely the responsibility of the authors and do not necessarily represent the official views of the US National Institutes of Health.
The authors declare no competing financial interests.
- 5′ end capping
The addition of an untemplated guanosine to the 5′ end of an RNA polymerase II transcript followed by its methylation at the N7 position. Capping protects the mRNA 5′ end from endonucleases.
- 3′ end cleavage and polyadenylation
Endonucleolytic cleavage that defines the 3′ ends of RNA polymerase II transcripts by cleavage and polyadenylation specificity factor (CPSF) and other factors, followed by the addition of non-templated poly(A) tails by poly(A) polymerase.
- Nascent RNA
RNA that is tethered to DNA by any elongating RNA polymerase.
- Gene architecture
The ensemble of cis-regulatory, coding and non-coding elements of a gene, including length, position and sequence.
- Ultraviolet (UV) crosslinking
UV irradiation-induced covalent bonds that link amino acids with nucleic acids.
- SR proteins
RNA-binding proteins with long repeats of arginine (Arg) and serine (Ser) residues that are involved in the regulation of alternative splicing and other steps of gene expression.
- Intrinsically disordered regions
Protein regions that contain little amino acid diversity and appear to lack well-defined secondary and tertiary structures.
Membrane-less subnuclear granules that are enriched in splicing factors, particularly the SR proteins.
- Cajal bodies
Membrane-less subnuclear compartments (2–4 per cell) that are the sites of small nuclear RNA modification and small nuclear ribonucleoprotein assembly. Cajal bodies are not the sites of splicing.
Membrane-less cytoplasmic compartments that are involved in mRNA turnover.
- Lampbrush chromosomes
Giant meiotic chromosomes that are formed in oocyte nuclei owing to the looping of chromosomal regions that are highly transcribed and coated with nascent RNA.
About this article
Cite this article
Herzel, L., Ottoz, D., Alpert, T. et al. Splicing and transcription touch base: co-transcriptional spliceosome assembly and function. Nat Rev Mol Cell Biol 18, 637–650 (2017). https://doi.org/10.1038/nrm.2017.63
Two sides of the same medal: Noncoding mutations reveal new pathological mechanisms and insights into the regulation of gene expression
WIREs RNA (2021)
Integrated requirement of non‐specific and sequence‐specific DNA binding in Myc‐driven transcription
The EMBO Journal (2021)
Biochimica et Biophysica Acta (BBA) - Gene Regulatory Mechanisms (2021)
Identification of a Putative DNA-Binding Protein in Arabidopsis That Acts as a Susceptibility Hub and Interacts With Multiple Pseudomonas syringae Effectors
Molecular Plant-Microbe Interactions® (2021)