Cross-talk in transcription, splicing and chromatin

Signalling and Control from a Systems Perspective

Cross-talk in transcription, splicing and chromatin: who makes the first call? Ross Alexander and Jean D. Beggs1 Wellcome Trust Centre for Cell Biology, and Edinburgh Centre for Systems Biology, University of Edinburgh, King’s Buildings, Mayfield Road, Edinburgh EH9 3JU, U.K.

Abstract The complex processes of mRNA transcription and splicing were traditionally studied in isolation. In vitro studies showed that splicing could occur independently of transcription and the perceived wisdom was that, to a large extent, it probably did. However, there is now abundant evidence for functional interactions between transcription and splicing, with important consequences for splicing regulation. In the present paper, we summarize the evidence that transcription affects splicing and vice versa, and the more recent indications of epigenetic effects on splicing, through chromatin modifications. We end by discussing the potential for a systems biology approach to obtain better insight into how these processes affect each other.

Introduction When protein encoding transcripts are produced by RNAPII (RNA polymerase II), they are subject to processing at their 5 -ends (capping), 3 -ends (cleavage and polyadenylation) and internally (splicing to remove non-coding intron sequences). There has been a gradual move away from the dogma that these processes are independent events, and it is now widely accepted that some of these modifications occur cotranscriptionally, meaning while RNAPII is still elongating the nascent RNA. There are many excellent reviews on the subject [1–5]. In the present paper, the emphasis is on cross-talk between transcription, splicing and chromatin remodelling. Early electron microscopy studies showed that many, but not all, introns are removed in a co-transcriptional manner [6]. However, it was only relatively recently that the application of chromatin immunoprecipitation allowed the co-transcriptional recruitment of splicing factors to be monitored in real time [7–11]. Owing to their close proximity to the DNA template, it is possible to cross-link co-transcriptionally recruited splicing factors to the DNA by treating cells with formaldehyde. After cell lysis, chromatin is sheared and immunoprecipitated with antibodies raised against specific splicing factors. At this point, the crosslinks are reversed and the recovered DNA is amplified by PCR using specific primers. It is therefore possible to detect the sequential recruitment of splicing factors during co-transcriptional spliceosome assembly along the length of a gene as the nascent RNA is elongated by RNAPII. Nevertheless, it has been pointed out that, although early Key words: coupling, histone modification, nucleosome, RNA polymerase, small nuclear ribonucleoprotein particle (snRNP), spliceosome.. Abbreviations used: CTD, C-terminal domain; PTB, polypyrimidine-tract-binding protein; P-TEFb, positive transcription elongation factor b; RNAi, RNA interference; RNAPII, RNA polymerase II; SKIP, Ski-interacting protein; snRNP, small nuclear ribonucleoprotein particle; SR, serine/argininerich; Tat, transactivator of transcription; Tat-SF1, Tat-specific factor 1. 1 To whom correspondence should be addressed (email [email protected]).

Biochem. Soc. Trans. (2010) 38, 00–00; doi:10.1042/BST038xxxx

splicing factors may be recruited co-transcriptionally, splicing is not necessarily completed co-transcriptionally [2,12]. So, is it just by chance that spliceosome assembly sometimes occurs at the site of transcription or are the processes actively coupled and, if so, why? What are the benefits? In vitro studies to address these questions used HeLa nuclear extracts that have combined transcription and splicing activities. Whereas improved kinetics [13] and altered patterns of splicing [14] were observed in these physically coupled transcription/splicing systems, it was concluded that this may be largely attributed to RNAPII protecting nascent transcripts against nuclear degradation, rather than enhanced splicing activity [14,15]. In other words, there was no evidence for functional coupling of the two processes. In order to clarify discussion of this topic, it was proposed that the transcription and splicing reactions can be considered ‘coupled’ if properties of the splicing reaction are specifically altered in a transcription-dependent manner [15]. Two distinct mechanisms of coupling have been identified and are described in the present paper.

Recruitment coupling There is evidence that certain splicing factors interact with the CTD (C-terminal domain) of the large subunit of RNAPII, thereby facilitating their co-transcriptional recruitment to the nascent transcript [3]. Although this may, in turn, promote co-transcriptional spliceosome assembly on nascent transcripts, it does not necessarily imply functional coupling of the two processes. The CTD consists of multiple repeats (26 in yeast, 52 in humans) of the heptad sequence YS2 PTS5 PS7 that is dynamically phosphorylated as RNAPII transcribes the gene (Figure 1). Typically, when RNAPII is at the promoter, the CTD is in a hypophosphorylated form. It then becomes phosphorylated on Ser5 (pSer5 ) to permit promoter clearance and initiation. The phosphorylation at Ser5 decreases towards the 3 -end of the gene as the CTD C The

C 2010 Biochemical Society Authors Journal compilation

1

2

Biochemical Society Transactions (2010) Volume 38, part 5

Figure 1 Co-transcriptional recruitment coupling RNA-processing factors, including capping enzymes (small ovals), splicing factors (parallelograms) and 3 -end-modifying factors (rectangles and hexagons) associate with the transcription machinery via the CTD of the largest subunit of RNAPII. The dynamic phosphorylation of serines 2, 5 and 7 (black circles) in the conserved heptad (YSPTSPS) repeats of the CTD are important for these and other interactions. mG represents the 7-methylguanosine 5 -cap.

becomes phosphorylated on Ser2 . Recent reports suggest that CTD Ser7 is also phosphorylated, in a manner resembling that of Ser5 . Currently, the only characterized function for pSer7 is related to 3 -end processing of snRNAs (small nuclear RNAs) [16,17]. Over 100 proteins have been shown to bind to the phosphorylated CTD [18], and it was proposed that the CTD acts as a ‘landing platform’ for RNA-processing factors, with the specificity of binding being determined by a ‘CTD code’ of post-translational modifications [19]. For example, the yeast U1 snRNP (small nuclear ribonucleoprotein particle) Prp40 was found to bind specifically to CTD that is doubly phosphorylated at Ser2 and Ser5 [20]. The central importance of the CTD was demonstrated in a study where truncation of the CTD led to defects in capping, polyadenylation and splicing [21]. This dynamic CTD phosphorylation suggests that RNAPII reorganizes its CTD interactions during transit along the DNA template to promote the sequential action of distinct RNA-processing events [4]. Recruitment coupling is also evident by the presence of dual-function proteins such as PGC-1 (peroxisome-proliferator-activated receptor γ co-activator 1), which is both a transcriptional coactivator and an alternative splicing regulator whose splicing activity is effective only when it is tethered to promoters through binding to a sequence-specific transcription factor [22].

‘The co-transcriptional race’ The second mode of interaction between transcription and splicing is referred to as kinetic coupling, which is particularly important in the context of alternative splicing. Alternative C The


splicing is a widespread means of producing polypeptide diversity from a single gene. The transcripts of 95% of human multi-exon genes undergo alternative splicing and there are ∼100 000 alternative splicing events in major human tissues [23]. Transcript elongation by RNAPII is not a uniform process that occurs at a steady rate. Rather, it fluctuates and is prone to pause, as dictated by the local sequence environment. The simplest way in which elongation rate can influence the process of alternative splicing is through the control of what has been described as the “window of opportunity” [5]. This is the time in which an upstream splice site can assemble in a functional spliceosome before it has to compete with a downstream splice site. A low elongation rate or paused polymerase would allow recognition and inclusion of a poor upstream 3 -splice site before a better downstream one is transcribed, thereby promoting inclusion of an alternative exon [3]. Essentially, it represents a race between RNAPII elongation and the time it takes for splicing factor recruitment and spliceosome assembly to occur. It is therefore not surprising to find other factors that are implicated in the control of RNAPII pausing and processivity, and which affect both constitutive and alternative splicing. These include promoter- and enhancer-associated transcription factors, elongation factors and co-activators (for a review, see [3]).

Is coupling reciprocal? Most of the available evidence demonstrates influences of the transcription machinery on factors involved in pre-mRNA processing. However, there is a growing body of evidence to suggest a reciprocal (bi-directional coupling) relationship between the two processes. For example, a functional 5 -splice site in close proximity to a promoter was found to enhance transcriptional output [24], and it was shown that the presence of the U1 snRNP at a 5 -splice site can actively recruit general transcription factors and RNAPII to the promoter in a splicing-dependent manner [25]. Similarly, Fong and Zhou [26] demonstrated that the U1 snRNP can regulate transcription via stimulation of RNAPII elongation. The effect is mediated through interaction with the elongation factor Tat-SF1 [Tat (transactivator of transcription)-specific factor 1], which itself interacts with P-TEFb (positive transcription elongation factor b), the kinase responsible for phosphorylation of Ser2 on the CTD of RNAPII. Additionally, in human cell extracts, U1 snRNPs, but not other snRNPs, were shown to co-purify with RNAPII [27]. More recently, the mammalian transcriptional regulator SKIP (Ski-interacting protein) was demonstrated to have a role in Tat-dependent transcription [28]. Interestingly, SKIP and its yeast counterpart, Prp45p, are components of spliceosomes [29,30], as is Cus2p, the yeast counterpart of Tat-SF1 [31]. These findings point towards a conserved structural and functional intertwining of the transcription and splicing machineries. The SR (serine/arginine-rich) family of proteins is a class of RNA-binding proteins that are capable of binding nascent transcripts, committing pre-mRNA to the splicing pathway


through contact with the U1 and U2 snRNPs (reviewed in [32]). SR proteins have been demonstrated to co-purify with RNAPII [27], and the SC35 family member has been implicated in stimulation of transcriptional elongation through interaction with P-TEFb [33]. SC35 depletion induced RNAPII accumulation within the gene body and attenuated elongation. This work suggests that SC35, and possibly other SR proteins, may act at an earlier stage than was first thought, functioning as single-strand-RNA-binding proteins to facilitate transcriptional elongation, even on intronless genes (reviewed in [34]). Whereas the evidence discussed so far is strongly indicative of functional links between transcription and splicing, recent studies suggest a more complex scenario. As reviewed by Kornblihtt et al. [35], the recognition of exons by the splicing machinery might need “a little help from a chromatin friend”.

Figure 2 How nucleosomes may affect transcription and splicing Nucleosomes (circles and ovals) are preferentially associated with exons (1.5-fold greater enrichment on exons compared with introns). Exon marking by nucleosome positioning may act as ‘speed bumps’ or ‘road blocks’ to RNAPII elongation. Bioinformatic analyses have shown H3K36me3 and other histone modifications to increase 5 →3 in genes as illustrated. Such histone modifications may act as signals to affect nucleosome positioning and/or affect splicing factor recruitment to the gene. Parallelograms represent splicing factors attached to RNAPII CTD. Exons are thick black lines. The thin black line in the middle represents an intron. mG represents 7-methylguanosine 5 -cap.

Chromatin joins the pre-mRNA processing party In eukaryotes, genomic DNA is packed into nucleosomes, octamers of histone proteins around which the DNA is wrapped and from which histone ‘tails’ protrude. This chromatin environment is in a constant state of flux, being altered by specific modifications to the histone tails, including acetylation, methylation, phosphorylation, ubiquitination and proline isomerization [36,37]. These modifications convert chromatin between ‘closed’, transcriptionally repressed, heterochromatin and more ‘open’, transcriptionally accessible, euchromatin states [38]. A general link between nucleosome density and gene exon/intron architecture was first proposed by Beckmann and Trifonov [39], who made the striking observation that the mean distance between consecutive 5 or 3 -splice sites showed a periodicity reminiscent of the 147 nt length of DNA required to wrap around a nucleosome. They suggested that nucleosomes are somehow positioned in conjunction with the elements that promote intron removal. More recently, a number of reports suggest a link between chromatin structure and exon/intron architecture. Kolasinska-Zwierz et al. [40] showed that trimethylation of histone 3 Lys36 (H3K36me3), a mark previously associated with transcription elongation, is found preferentially on exons relative to introns of actively transcribed genes in nematodes and mammals. It was suggested that H3K36me3 on expressed exons may represent a marking mechanism, providing a dynamic link between transcription and splicing. Further bioinformatic analyses of published experimental data derived from deep sequencing of human and Caernorhabditis elegans DNA fragments generated through micrococcal nuclease digestion have provided further insights. It was demonstrated that exons are differentially marked from introns, both in terms of nucleosome occupancy (approx. 1.5-fold higher in exons than in introns) and in specific histone modifications [41–44]. This suggests that the H3K36me3 pattern reported by Kolasinska-Zwierz et al. [40] can be explained at least in part by the more general phenomenon of increased nucleosome density over exons.

Other epigenetic marks such as monomethylation of H3K79, H4K20 and H2BK5 and mono-, di- and tri-methylation of H3K27 are also enriched on exons, with an increase in amplitude as gene expression levels increase (Figure 2). Conceivably, nucleosomes carrying specific histone modifications may interact with splicing factors to enhance exon recognition [42,45]. It was also found that the ends of introns contain sequences disfavoured by nucleosomes, that may shift nucleosome occupancy to exons, and nucleosome enrichment was most pronounced for exons with weak splice sites [42–44]. It was suggested that nucleosomes may function as ‘speed bumps’ to slow RNAPII, thereby improving the selection of exons by increasing the time (or window of opportunity) for newly synthesized splice signals in the nascent pre-mRNA to be recognized by splicing factors [42]. Indeed, recent biophysical evidence has been obtained for the nucleosome behaving as a fluctuating barrier that affects RNAPII movement [46]. In this way, nucleosome positioning may aid exon definition, especially for long genes. Another intriguing connection between nucleosome positioning and splicing is through chromatin remodelling. This was suggested after the discovery that the SWI/SNF chromatin-remodelling ATPases influence splicing [47]. This work demonstrated that alternative splicing was influenced by differences in transcription elongation rates in a manner regulated by SWI/SNF and CTD phosphorylation, and involving the creation of ‘road blocks’ to transcription (reviewed in [48]). C The


3

4


RNAPII is thought to mediate the cross-talk between chromatin and the exon/intron architecture of RNA. The nucleosome-specific histone methyltransferase Set2 is responsible for methylation of H3 at Lys36 and is recruited through the selective action of pSer2 on RNAPII [49]. Intriguingly, this suggests that perturbations of the phosphorylation status of RNAPII would have an effect on epigenetic marking specifically of the H3K36me3 mark, with an altered chromatin landscape possibly controlling alternative splicing events. Indeed, Luco et al. [50] have demonstrated a direct role for histone modifications in splice site selection in human cells. Using the human FGFR2 (fibroblast growth factor receptor 2) gene, which has been used extensively to study alternative splicing, they observed that modulating the level of H3K36me3 through overexpression of Set2 resulted in a significant increase in H3K36me3 globally and reduced inclusion of PTB (polypyrimidinetract-binding protein)-dependent exons. Conversely, downregulation of Set2 by RNAi (RNA interference) promoted the inclusion of normally repressed PTB-dependent exons. These results suggest a role for histone modifications in alternative splicing control through the existence of an adaptor system, containing histone modifications, a chromatin-binding protein that reads the histone marks and an interacting splicing regulator. It was proposed that such a system could transmit epigenetic information to the premRNA-processing machinery by promoting the recruitment of specific splicing factors (reviewed in [51]).

Lateral thinking: a systems biology approach Considering the growing body of evidence suggesting a threeway cross-talk between transcription, splicing and chromatin, it would be beneficial to adopt a systems biology approach to integrate the large amount of complex and diverse biological information that is available. Through mathematical modelling, it should be possible to generate biological predictions that can be tested experimentally. There has already been a move towards a systems level approach to understand coupling and coordination in gene expression systems [52], focused primarily on protein interactions. Maciag et al. [53] developed computational methods to analyse protein coupling in the gene-expression machineries of yeast, with extrapolation of their findings to humans. Using this approach, they were able to confirm known coupling such as that of transcription and RNA processing with export, and predict further coupling with translation and nonsense-mediated decay. Efforts have been made by Darzacq et al. [54] to measure elongation rates of RNAPII in situ. By following the synthesis of RNA in real time and through the use of deterministic computational models constrained by extensive data analysis, they were able to make predictions that were tested experimentally through the use of transcriptional inhibitors. Multistep models of transcript synthesis have also been developed [55] and studied theoretically [56,57]. Modelling techniques were applied to the analysis of cytoplasmic mRNA turnover C The


in Saccharomyces cerevisiae, yielding insights into mRNA metabolism that would not readily have been obtained from conventional RNA analyses alone [58]. RNA turnover is a simpler pathway than transcription and splicing, but this illustrates the potential for generating novel insights. Although both transcription and mRNA turnover have been extensively modelled, these models have yet to incorporate the splicing reaction or take into account the effects of chromatin. Many intriguing questions remain, such as the following. Is the order of RNA processing events important? How is information transferred from the DNA to the RNA level and vice versa? With regard to the chromatin link, what happens first? Does the splicing machinery first signal to the chromatin where a functional intron is present, causing chromatin remodelling and/or histone modification to establish the intron mark? Or conversely, does epigenetic marking first determine splicing by signalling to the polymerase and splicing machinery? It should be possible to address some of these questions by using genetic or RNAi knockdown of relevant factors in combination with high-resolution kinetic analyses of transcript elongation, splicing, the recruitment of transcription and splicing factors, CTD kinases and phosphatases, and chromatin-modifying and -remodelling factors during the very early stages of de novo transcription of a gene. As a beginning, a recent quantitative analysis of the expression of long genes in human cells following removal of a chemical inhibitor of transcription has permitted more accurate measurements of the kinetics of transcription and splicing and provided direct evidence for co-transcriptional splicing, even of long introns [59]. This demonstrates the feasibility of obtaining the kinds of quantitative real-time in vivo data required for mathematical modelling. Undoubtedly, new modelling approaches will also be required to describe this very complex multi-component and multi-dimensional system, but the insights gained should be worth the effort.

Acknowledgements We are grateful to Steve Innocente for helpful discussion and comments on the manuscript.

Funding R.A. was supported by the European Union-funded Framework Programme 6 RiboSys project [grant number 518280] and by the Biotechnology and Biological Sciences Research Council and the Engineering and Physical Sciences Research Council through funding [grant number BB/D019621/1] to the Edinburgh Centre for Systems Biology. J.D.B. is the Royal Society Darwin Trust Research Professor.

References 1 Manley, J.L. (2002) Nuclear coupling: RNA processing reaches back to transcription. Nat. Struct. Biol. 9, 790–791


2 Neugebauer, K.M. (2002) On the importance of being co-transcriptional. J. Cell Sci. 115, 3865–3871 3 Kornblihtt, A.R., de la Mata, M., Fededa, J.P., Munoz, M.J. and Nogues, G. (2004) Multiple links between transcription and splicing. RNA 10, 1489–1498 4 Pandit, S., Wang, D. and Fu, X.D. (2008) Functional integration of transcriptional and RNA processing machineries. Curr. Opin. Cell Biol. 20, 260–265 5 Perales, R. and Bentley, D. (2009) “Cotranscriptionality”: the transcription elongation complex as a nexus for nuclear transactions. Mol. Cell 36, 178–191 6 Beyer, A.L. and Osheim, Y.N. (1988) Splice site selection, rate of splicing, and alternative splicing on nascent transcripts. Genes Dev. 2, 754–765 7 Kotovic, K.M., Lockshon, D., Boric, L. and Neugebauer, K.M. (2003) Cotranscriptional recruitment of the U1 snRNP to intron-containing genes in yeast. Mol. Cell. Biol. 23, 5768–5779 8 Gornemann, J., Kotovic, K.M., Hujer, K. and Neugebauer, K.M. (2005) Cotranscriptional spliceosome assembly occurs in a stepwise fashion and requires the cap binding complex. Mol. Cell 19, 53–63 9 Listerman, I., Sapra, A.K. and Neugebauer, K.M. (2006) Cotranscriptional coupling of splicing factor recruitment and precursor messenger RNA splicing in mammalian cells. Nat. Struct. Mol. Biol. 13, 815–822 10 Lacadie, S.A. and Rosbash, M. (2005) Cotranscriptional spliceosome assembly dynamics and the role of U1 snRNA:5 ss base pairing in yeast. Mol. Cell 19, 65–75 11 Lacadie, S.A., Tardiff, D.F., Kadener, S. and Rosbash, M. (2006) In vivo commitment to yeast cotranscriptional splicing is sensitive to transcription elongation mutants. Genes Dev. 20, 2055–2066 12 Moore, M.J., Schwartzfarb, E.M., Silver, P.A. and Yu, M.C. (2006) Differential recruitment of the splicing machinery during transcription predicts genome-wide patterns of mRNA splicing. Mol. Cell 24, 903–915 13 Das, R., Dufu, K., Romney, B., Feldt, M., Elenko, M. and Reed, R. (2006) Functional coupling of RNAP II transcription to spliceosome assembly. Genes Dev. 20, 1100–1109 14 Hicks, M.J., Yang, C.R., Kotlajich, M.V. and Hertel, K.J. (2006) Linking splicing to Pol II transcription stabilizes pre-mRNAs and influences splicing patterns. PLoS Biol. 4, e147 15 Lazarev, D. and Manley, J.L. (2007) Concurrent splicing and transcription are not sufficient to enhance splicing efficiency. RNA 13, 1546–1557 16 Egloff, S., O’Reilly, D., Chapman, R.D., Taylor, A., Tanzhaus, K., Pitts, L., Eick, D. and Murphy, S. (2007) Serine-7 of the RNA polymerase II CTD is specifically required for snRNA gene expression. Science 318, 1777–1779 17 Kim, M., Suh, H., Cho, E.J. and Buratowski, S. (2009) Phosphorylation of the yeast Rpb1 C-terminal domain at serines 2, 5, and 7. J. Biol. Chem. 284, 26421–26426 18 Phatnani, H.P., Jones, J.C. and Greenleaf, A.L. (2004) Expanding the functional repertoire of CTD kinase I and RNA polymerase II: novel phosphoCTD-associating proteins in the yeast proteome. Biochemistry 43, 15702–15719 19 Egloff, S. and Murphy, S. (2008) Cracking the RNA polymerase II CTD code. Trends Genet. 24, 280–288 20 Phatnani, H.P. and Greenleaf, A.L. (2006) Phosphorylation and functions of the RNA polymerase II CTD. Genes Dev. 20, 2922–2936 21 McCracken, S., Fong, N., Yankulov, K., Ballantyne, S., Pan, G., Greenblatt, J., Patterson, S.D., Wickens, M. and Bentley, D.L. (1997) The C-terminal domain of RNA polymerase II couples mRNA processing to transcription. Nature 385, 357–361 22 Fededa, J.P. and Kornblihtt, A.R. (2008) A splicing regulator promotes transcriptional elongation. Nat. Struct. Mol. Biol. 15, 779–781 23 Pan, Q., Shai, O., Lee, L.J., Frey, B.J. and Blencowe, B.J. (2009) Addendum: deep surveying of alternative splicing complexity in the human transcriptome by high-throughput sequencing. Nat. Genet. 41, 762 24 Furger, A., Binnie, J.M.O., Lee, B.A. and Proudfoot, N.J. (2002) Promoter proximal splice sites enhance transcription. Genes Dev. 16, 2792–2799 25 Damgaard, C.K., Kahns, S., Lykke-Andersen, S., Nielsen, A.L., Jensen, T.H. and Kjems, J. (2008) A 5 splice site enhances the recruitment of basal transcription initiation factors in vivo. Mol. Cell 29, 271–278 26 Fong, Y.W. and Zhou, Q. (2001) Stimulatory effect of splicing factors on transcriptional elongation. Nature 414, 929–933 27 Das, R., Yu, J., Zhang, Z., Gygi, M.P., Krainer, A.R., Gygi, S.P. and Reed, R. (2007) SR proteins function in coupling RNAP II transcription to pre-mRNA splicing. Mol. Cell 26, 867–881

28 Bres, V., Gomes, N., Pickle, L. and Jones, K.A. (2005) A human splicing factor, SKIP, associates with P-TEFb and enhances transcription elongation by HIV-1 Tat. Genes Dev. 19, 1211–1226 29 Bessonov, S., Anokhina, M., Will, C.L., Urlaub, H. and Luhrmann, R. (2008) Isolation of an active step I spliceosome and composition of its RNP core. Nature 452, 846–850 30 Albers, M., Diment, A., Muraru, M., Russell, C.S. and Beggs, J.D. (2003) Identification and characterization of Prp45p and Prp46p, essential pre-mRNA splicing factors. RNA 9, 138–150 31 Yan, D., Perriman, R., Igel, H., Howe, K.J., Neville, M. and Ares, Jr, M. (1998) CUS2, a yeast homolog of human Tat-SF1 rescues function of misfolded U2 through an unusual RNA recognition motif. Mol. Cell. Biol. 18, 5000–5009 32 Long, J.C. and Caceres, J.F. (2009) The SR protein family of splicing factors: master regulators of gene expression. Biochem. J. 417, 15–27 33 Lin, S., Coutinho-Mansfield, G., Wang, D., Pandit, S. and Fu, X.D. (2008) The splicing factor SC35 has an active role in transcriptional elongation. Nat. Struct. Mol. Biol. 15, 819–826 34 Zhong, X.Y., Wang, P., Han, J., Rosenfeld, M.G. and Fu, X.D. (2009) SR proteins in vertical integration of gene expression from transcription to RNA processing to translation. Mol. Cell 35, 1–10 35 Kornblihtt, A.R., Schor, I.E., Allo, M. and Blencowe, B.J. (2009) When chromatin meets splicing. Nat. Struct. Mol. Biol. 16, 902–903 36 Mellor, J. (2006) It takes a PHD to read the histone code. Cell 126, 22–24 37 Kouzarides, T. (2007) Chromatin modifications and their function. Cell 128, 693–705 38 Li, B., Carey, M. and Workman, J.L. (2007) The role of chromatin during transcription. Cell 128, 707–719 39 Beckmann, J.S. and Trifonov, E.N. (1991) Splice junctions follow a 205-base ladder. Proc. Natl. Acad. Sci. U.S.A. 88, 2380–2383 40 Kolasinska-Zwierz, P., Down, T., Latorre, I., Liu, T., Liu, X.S. and Ahringer, J. (2009) Differential chromatin marking of introns and expressed exons by H3K36me3. Nat. Genet. 41, 376–381 41 Andersson, R., Enroth, S., Rada-Iglesias, A., Wadelius, C. and Komorowski, J. (2009) Nucleosomes are well positioned in exons and carry characteristic histone modifications. Genome Res. 19, 1732–1741 42 Schwartz, S., Meshorer, E. and Ast, G. (2009) Chromatin organization marks exon-intron structure. Nat. Struct. Mol. Biol. 16, 990–995 43 Spies, N., Nielsen, C.B., Padgett, R.A. and Burge, C.B. (2009) Biased chromatin signatures around polyadenylation sites and exons. Mol. Cell 36, 245–254 44 Tilgner, H., Nikolaou, C., Althammer, S., Sammeth, M., Beato, M., Valcarcel, J. and Guigo, R. (2009) Nucleosome positioning as a determinant of exon recognition. Nat. Struct. Mol. Biol. 16, 996–1001 45 Allemand, E., Batsche, E. and Muchardt, C. (2008) Splicing, transcription, and chromatin: a menage ´ a` trois. Curr. Opin. Genet. Dev. 18, 145–151 46 Hodges, C., Bintu, L., Lubkowska, L., Kashlev, M. and Bustamante, C. (2009) Nucleosomal fluctuations govern the transcription dynamics of RNA polymerase II. Science 325, 626–628 47 Batsche, E., Yaniv, M. and Muchardt, C. (2006) The human SWI/SNF subunit Brm is a regulator of alternative splicing. Nat. Struct. Mol. Biol. 13, 22–29 48 Kornblihtt, A.R. (2006) Chromatin, transcript elongation and alternative splicing. Nat. Struct. Mol. Biol. 13, 5–7 49 Xiao, T., Hall, H., Kizer, K.O., Shibata, Y., Hall, M.C., Borchers, C.H. and Strahl, B.D. (2003) Phosphorylation of RNA polymerase II CTD regulates H3 methylation in yeast. Genes Dev. 17, 654–663 50 Luco, F.R., Pan, Q., Tominaga, K., Blencowe, B.J., Pereira-Smith, O. and Misteli, T. (2010) Regulation of alternative splicing by histone modification. Science 327, 996–1000 51 Fox-Walsh, K. and Fu, X.D. (2010) Chromatin: the final frontier in splicing regulation? Dev. Cell 18, 336–338 52 Komili, S. and Silver, P.A. (2008) Coupling and coordination in gene expression processes: a systems biology view. Nat. Rev. Genet. 9, 38–48 53 Maciag, K., Altschuler, S.J., Slack, M.D., Krogan, N.J., Emili, A., Greenblatt, J.F., Maniatis, T. and Wu, L.F. (2006) Systems-level analyses identify extensive coupling among gene expression machines. Mol. Syst. Biol. 2, 2006.0003 54 Darzacq, X., Shav-Tal, Y., de, T., V, Brody, Y., Shenoy, S.M., Phair, R.D. and Singer, R.H. (2007) In vivo dynamics of RNA polymerase II transcription. Nat. Struct. Mol. Biol. 14, 796–806 C The


5

6


55 Weber, A., Liu, J., Collins, I. and Levens, D. (2005) TFIIH operates through an expanded proximal promoter to fine-tune c-myc expression. Mol. Cell. Biol. 25, 147–161 56 Voliotis, M., Cohen, N., Molina-Paris, C. and Liverpool, T.B. (2008) Fluctuations, pauses, and backtracking in DNA transcription. Biophys. J. 94, 334–348 57 Voliotis, M., Cohen, N., Molina-Paris, C. and Liverpool, T.B. (2009) Backtracking and proofreading in DNA transcription. Phys. Rev. Lett. 102, 258101

C The


58 Cao, D. and Parker, R. (2003) Computational modeling and experimental analysis of nonsense-mediated decay in yeast. Cell 113, 533–545 59 Singh, J. and Padgett, R.A. (2009) Rates of in situ transcription and splicing in large human genes. Nat. Struct. Mol. Biol. 16, 1128–1133

Received 16 June 2010 doi:10.1042/BST038xxxx