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 3JR, 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, 1251–1256; doi:10.1042/BST0381251
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
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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
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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
Signalling and Control from a Systems Perspective
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]).
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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 co-ordination 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
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Authors Journal compilation 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.
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Received 16 June 2010
doi:10.1042/BST0381251