Regulation of alternative splicing and its connections to cancer
Mo Chen
Submitted for the partial fulfillment of the requirements
for the degree of Doctor of Philosophy in the
Graduate School of Arts and Sciences
Columbia University
2011
© 2011 Mo Chen All rights reserved
ABSTRACT
Regulation of alternative splicing and its connections to cancer
Mo Chen
This thesis presents two separate pieces of work pertaining to pre-mRNA splicing in
mammalian cells. The first piece, as the main research project of the thesis, consists of two
related parts. The first part identified the regulators of the alternative splicing of the PKM gene
in cancer cells while the second part elucidates the molecular mechanism of how this mutually
exclusive alternative splicing is regulated. The second piece investigates the molecular
mechanism of how SRp38 functions as a splicing activator when phosphorylated.
Cancer cells uniformly alter key aspects of their metabolism, including their glucose
usage. In contrast to quiescent cells, which use most of their glucose for oxidative
phosphorylation when oxygen is present, under the same conditions, most of the glucose
consumed by cancer cells is converted to lactate. This phenomenon is known as aerobic glycolysis, and is critical for cancer cell growth. The pyruvate kinase isoform expressed by the cell is a key determinant of glucose usage. Pyruvate kinase in most tissues is produced from the
PKM gene, which is alternatively spliced to produce the PKM1 or PKM2 isoforms, which contain exons 9 or 10 respectively. Adult tissues, such as skeletal muscle and brain, express predominantly the PKM1 isoform, which is universally reverted to the embryonic PKM2 isoform in cancer cells. PKM2 expression promotes aerobic glycolysis.
In Chapter 3, I describe a mechanism by which cancer cells promote switching to PKM2.
We show that PKM exon 9 is flanked by binding sites for the RNA-binding proteins hnRNP
A1/A2 and PTB. These proteins bind to exon 9 and repress its inclusion in the mRNA, resulting in PKM2 production. Additionally, we show that hnRNP A1/A2 and PTB are all overexpressed in cancers in a way that precisely correlates with the expression of PKM2. Finally, we show that the oncogenic transcription factor c-Myc promotes PKM2 expression by transcriptionally
upregulating the genes encoding hnRNP A1/A2 and PTB.
In Chapter 4, I provide additional insights into how PKM AS is regulated and a novel discovery
that general splicing repressors can repress either one of the two mutually exclusive exons at different
expression levels, through protein-protein interactions of these proteins bound on different sets of binding
sites on and flanking each. First, using a splicing minigene construct that recapitulates PKM splicing in HeLa cells, we identified additional PTB and hnRNP A1/ A2 ISSs in intron 9 necessary for full exclusion of exon 9. More importantly, we found two ESSs in exon 9, absent from exon 10, that match the hnRNP A1 consensus, and which are critical for exon 9 exclusion.
We show that these ESSs function cooperatively to facilitate hnRNP A1 binding to an intronic splicing silencer in intron 9 described in Chapter 3. I also elucidated the mechanism of how exon 10
is excluded when exon 9 is derepressed and show that hnRNP A1 and PTB, when their protein levels are
reduced, release the inhibition of exon 9 but repress exon 10 inclusion, through binding sites present in
introns 9 and 10. This mechanism, coupled with nonsense mediated decay, function to prevent the
appearance of PKM mRNA containing both exon 9 and exon 10.
In the second piece of work, presented in Chapter 5, I, based on the findings from a
previous post doctor that SRp38 functions as a sequence-specific splicing activator, showed that
SRp38 promotes spliceosomal complex A formation. I examined the mechanism of spliceosomal
A complex formation and found that SRp38 promotes the recruitment of U1 and U2 snRNPs to
splicing substrates that contain high-affinity SRp38 binding sites.
TABLE OF CONTENTS
Table of Contents ………………………………………………………………………………. i
Acknowledgements ……………………………………………………………………………..iii
Dedication ……………………………………………………………………………………… iv
Preface ...... v
Chapter 1: Mechnisms of alternative splicing regulation: insights from molecular and genomics approaches …………………………………………………………………... 1
Preface …………………………………………………….………………….…………….…… 3
Introduction………………………….………………….…………….…………………………...3
Splice site recognition and selection .……………………………………………………………. 5
Regulation by U1 and U2 snRNP pairing .…………………………………………………...….11
Transcription–coupled alternative splicing ……………………………………………………...15
Alternative splicing and tissue specificity.………..……………………………………………..17
Conclusions and perspectives ………………………………………………………………….. 22
References ……………………………………………………………………………………… 25
Chapter 2: Tumor metabolism: hnRNP proteins get in on the act .……………………..… 49
References ……………………………………………………………………………………… 52
Chapter 3: HnRNP proteins controlled by c-Myc deregulate pyruvate kinase mRNA splicing in cancer ……………………………………………………………………… 54
Abstract ………………………………………………………………………………………... 56
Methods summary …………………………………………………………………………….. . 63
References …………………………………………………………………………………….... 64
Full Methods …………………………………………………………………………………… 70
Supplementary information ……………………………………………………………………..79
i
Chapter 4: HnRNP proteins control mutually exclusive splicing of pyruvate kinase M pre- mRNA ………………………………………………………………………………….. 93
Abstract ………………………………………………………………………………………… 95
Introduction ……………………………………………………………………………………. .96
Results ………………………………………………………………………………………..... 98
Discussion …………………………………………………………………………………...…111
Methods ……………...…………………………………………………………………………117
References ……………………………………………………………………………………...122
Figure legends ………………………………………………………………………………….129
Figures ………………………………………………………………………………………….137
Chapter 5: Phosphorylation switches the general splicing repressor SRp38 to a sequence- specific activator.………………………………….………………………………………….. 152
Abstract……………………………….………………………………………………………...154
Introduction……………………………………………………………………………………..155
Results…………………………………………………………………………………………..157
Discussion………………………………………………………………………………………166
Methods…………………………………………………………………………………………171
References………………………………………………………………………………………172
ii
Acknowledgements
I would like to first thank my advisor Jim Manley for years of patient support. I have
learned a great deal from him over the past several years. I also have to thank Jim for introducing
me to a couple of very interesting projects. My committee members Larry Chasin and Carol
Prives have been the source of helpful suggestions, for which I think them. I would also like to thank my outside committee members, Robert Darnell and Livio Pellizzoni for taking the time to be on my committee.
Members of the Manley lab have been indispensable during my PhD. I learned in vitro splicing from Ying Feng, who also initiated the SRp38 project described in Chapter 5. I learned
FPLC chromatography from Chuck David and we also collaborated on the first part of the PKM project described in chapter 3. Discussions with Patricia Richard, Emanuel Rosonina, Jing-ping
Hsin were also helpful. I am very indebted to Emanuel Rosonina and Kensai Nishida, Dafne
Campigli Di Giammartino and Tristan Coady who spent time listening to me talk. I thank Jenn
Aoki for moral support.
Finally, I wouldn’t be completing a PhD if it weren’t for members of my family. My grandmother has supported me all these years without any reservation. My dad passed his nature of curiosity to me and has taught me to be a modest person. Finally, I owe the biggest debt of
gratitude scientifically and personally to my fiancé, Charles David. I would certainly not be
completing this without him!
iii
To Chuck
iv
Preface
This thesis is divided into five chapters. The Chapters 1 is a review article on the regulation of alternative splicing written for Nature Reviews Molecular Cell Biology, provided in their original print format. Chapter 2 is a review article written for Cell Cycle, summarizing the discovery described in chapter 3, provided in its original print format. Chapter 3, about identifying the regulators of pyruvate kinase M (PKM) alternative splicing, is provided in its original form as published in Nature. Chapter 4 is a manuscript, further elucidating the molecular mechanism of the mutually exclusive alternative splicing of PKM. Chapter 4 is about the mechanism of how SRp38 functions as a sequence-specific splicing activator, provided in its original print format published in Nature Structural Molecular Biology.
v
1
Chapter 1: Mechanism of Alternative splicing regulation: insights from molecular and genomic approaches.
2
Mechanisms of Alternative Splicing Regulation: Insights from Molecular
and Genomics Approaches
Mo Chen and James L. Manley
Department of Biological Sciences, Columbia University, New York, NY 10027
3 Preface
Alternative splicing of mRNA precursors provides an important avenue of gene control,
and constitutes a critical step in the expression of a majority of genes. Alternative
splicing profoundly impacts human development, and its mis-regulation underlies many
human diseases. The mechanisms of alternative splicing have been studied extensively,
but not until the past few years have we begun to realize fully the diversity and
complexity of alternative splicing regulation by an intricate protein-RNA network.
Significant progress has been made by studying individual transcripts and through
genome-wide approaches, which together provide a better picture of the mechanistic
regulation of alternative splicing.
Introduction
Alternative splicing (AS) is a critical mechanism for gene regulation and generation of
proteomic diversity. Recent estimates indicate that expression of nearly 95% of human
multi-exon genes involves alternative splicing1, 2. In metazoans, alternative splicing plays an important role in generating different protein products that function in diverse cellular processes, including cell growth, differentiation and death.
Splicing is carried out by the spliceosome, a massive structure in which five small nuclear ribonucleoprotein particles (snRNPs) and a large number of auxiliary proteins cooperate to accurately recognize the splice sites and catalyze the two steps of the splicing reaction1, 2 (BOX 1). Spliceosome assembly (BOX 1) begins with the
recognition of the 5’-splice site (ss) by U1 snRNP, and binding of splicing factor 1 (SF1)
to the branch point3 and factor U2AF to the polypyrimidine tract (py tract) and 3’
2 4 terminal AG4, 5, in an ATP-independent manner to form the E complex. The E complex is converted into the ATP-dependent pre-spliceosomal A complex by the replacement of
SF1 by U2 snRNP at the branch point. Further recruitment of the U4·U6/U5 tri-snRNP
leads to the formation of B complex, which is converted to the catalytically active C
complex after extensive conformational change and remodeling.
The decision of which exon is removed and which exon is included involves
RNA sequence elements and protein regulators. Depending on the position and function
of the cis-regulatory elements, they are divided into four categories: exonic splicing
enhancers (ESEs), exonic splicing silencers (ESSs), intronic splicing enhancers (ISEs)
and intronic splicing silencers (ISSs). ESEs are usually bound by members of the SR
protein family (BOX 2)6-8. ISSs and ESSs are commonly bound by hnRNP proteins
(TABLE 1), which have one or more RNA-binding domains and protein-protein
interaction domains9, 10. ISEs are not as well characterized as the other three types of
element, although recently several proteins, such as hnRNP F/H, Nova, Fox1/2, have
been shown to bind to ISEs and stimulate splicing11-14.
Choices of alternative splicing were traditionally thought to be made at the
stages of splice site recognition and early spliceosome assembly, and indeed this is
frequently the case1. However, several recent studies have shown that the decision can be made at different stages of spliceosome assembly, and even during conformational changes between the two transesterification steps15-17. In addition, mounting evidence of
coupling of RNA transcription and splicing regulation has emerged18-22.
Alternative splicing contributes to genomic diversity and tissue specificity23.
Understanding tissue-specific AS requires understanding the regulatory network of
3 5 protein-protein, protein-RNA and RNA-RNA interactions involved in tissue-specific AS
events1, 24. Tissue-specific AS is thought to be regulated by differentially expressed splicing regulators1, 25, and/or different concentrations and/or activity of ubiquitously expressed splicing factors26, 27. Additionally, tremendous progress made using high- throughput methods has not only revealed many more novel AS events28-30, but also
accelerated the process of understanding AS regulation in different tissues, by examining
the expression levels of protein regulators and helping to define cis-regulatory elements31,
32.In this review, we will discuss mechanisms of AS control from different perspectives.
At which stage of spliceosome assembly is the decision of whether to include an
alternative exon made? What protein factors are involved? How does RNA polymerase II
function in AS regulation? Are the kinetics of spliceosome assembly important for AS
regulation? Which mechanisms control tissue-specific AS? We will conclude by
summarizing the diversity of the mechanisms of AS regulation and speculate on the
future directions of AS research.
Splice site recognition and selection
Most human genes contain multiple exons, and the average length of exons (50-250 bp)
is much shorter than that of intervening sequences (frequently thousands of base pairs).
Early stages of spliceosome assembly are built around the exons due to the large size of
introns33. This kind of exon-centered splice site (ss) recognition is referred as “exon
definition”34 (BOX 1). Exon definition must eventually be converted to intron definition,
which occurs via cross-intron interactions between U1 and U2 snRNP35, 36. The best
4 6 studied mechanisms of AS regulation involve controlling ss recognition by facilitating or interfering with the binding of the U1 or U2 snRNP to the splice sites.
Facilitating splice site recognition
SR proteins play important roles in facilitating splice site recognition. For example, they
recruit U1 snRNP to the 5’ ss, and U2AF and U2 snRNP to the 3’ss, by binding to an
ESE and directly interacting with protein targets37-40 (FIG. 1a). These interactions are
mediated by RS domains41, 42 and require their proper phosphorylation and
dephosphorylation1, 43, 44. SR proteins also cooperate with other positive factors to form
larger enhancing complexes by interacting with other RS domain-containing proteins, such as Tra2 and SRm160/30045-47 (FIG. 1a). The mechanism of binding and recruiting can also be achieved by intronic binding proteins. For example, TIA-1 binds to a U-rich sequence downstream of weak 5’splice sites to recruit U1 snRNP48, 49. Sam68 binds and recruits U2AF to the 3’ss of exon V5 of the transmembrane glycoprotein CD44 pre- mRNA50.
SRp38, also known as TASR51, NSSR52 and SRrp4053, has been characterized as
a general splicing repressor that is activated by dephosphorylation54, 55 (see below).
However, recent studies indicate that it also functions as a sequence-dependent splicing
activator when phosphorylated39. It was found that in vitro SRp38 activates A complex
formation and splicing, by facilitating recruitment of U1 and U2 snRNPs to the pre-
mRNA and stabilizing 5’ss and branch site recognition (FIG. 3a). Notably, SRp38, unlike
other SR proteins, is unable to complement cytoplasmic S100 extracts (which contain all
factors required for splicing except SR proteins) to activate splicing56,. While an A-like
5 7 complex is formed in S100 plus SRp38, it is stalled or inactive, and to proceed requires a
specific coactivator (FIG. 3a), the identity of which is currently unknown39. In vivo,
SRp38 was shown to favor inclusion of the Flip exon of the GluR-B pre-mRNA52, while the mutually exclusive Flop exon is included when SRp38 is absent39. Interestingly, both exons contain SRp38 binding sites, and it was proposed that differential binding by
SRp38 (stronger to the Flip exon) and intracellular concentrations of SRp38 influence the decision to include either the Flip or Flop exon39.
Inhibiting splice site recognition
Inhibition of ss recognition can be achieved in multiple ways. First, when splicing
silencers are located close to splice sites or to splicing enhancers, inhibition can occur by
sterically blocking the access of snRNPs or positive-acting factors. For example,
polypyrimidine -tract binding protein (PTB), also named hnRNP I, binds to the py tract
and blocks the binding of U2AF to regulated exons57-59; hnRNP A1 binds to ISSs located
upstream of HIV Tat pre-mRNA exon 3 and prevents binding of U2 snRNP60; tissue- specific splicing factors Fox1/2 inhibits E’ complex formation by binding to an intronic sequence to prevent SF1 (splicing factor 1) from binding to the branch site of the calcitonin/CGRP pre-mRNA61 (FIG. 1b).
Splicing inhibitors also sterically block the binding of activators to enhancers: Hu
proteins inhibit U1 snRNP binding by competing with TIA-1 binding to an AU-rich sequence downstream of the 5’ss of exon 23a of neurofibromatosiss type 1 pre-mRNA62;
Fox1/2 also inhibits E complex formation by binding to an exonic sequence in the
calcitonin/CGRP pre-mRNA in close proximity to the ESE that Tra2 and the SR protein
6 8 SRp55 bind to, preventing recruitment of U2AF by the activators61 (FIG. 1b); hnRNP A1
binds to an ESS upstream of the Tra2-dependent ESE in exon 7 in the SMN2 pre-mRNA,
possibly inhibiting formation or stabilization of the U2 snRNP complex63, 64 (FIG.1c).
Some silencers can be over 100-200 bp away from enhancers, and a simple “bind
and block” model thus cannot explain their inhibitory effect. One explanation for the
activity of such splicing inhibitors is that they function by masking ss recognition through
multimerization along the RNA58. Another model proposes that the alternative exon (AE)
might be “looped out” in a process involving protein-protein interactions between RNA
binding proteins bound at sites spanning the AE58, 65-67, and loop formation may sterically
interfere with further spliceosome assembly, even though splice site recognition may not
be inhibited67. For example, hnRNP A1 binds to elements upstream and downstream of
exon 7B in its own pre-mRNA to promote skipping of exon 7B68. HnRNP A1 has also
been shown to bind to exonic and intronic silencers in SMN2 exon 7 and intron 7, and it
is proposed that an interaction between hnRNP A1 molecules is required to fully suppress
inclusion of exon 764, 69, 70. PTB has also been shown to bind to sites flanking the c-src N1
exon, and mutating one of the PTB sites affects the binding of PTB to the other site58, 71.
Combinatorial effects of activators and inhibitors
Splicing of individual pre-mRNAs is frequently controlled by combinatorial or competitive effects of both activators and inhibitors. The final decision of whether an AE is included or not is determined by the concentration or activity of each kind of regulator, often SR proteins and hnRNP proteins72-74. For example, the SR protein 9G8 and hnRNP
F/H regulate splicing of -tropomyosin exon 2 by competing for binding to the same
7 9 element75; hnRNP A1 and the SR proteins ASF/SF2 and SC35 have antagonistic
functions in splicing of -tropomyosin exon 6B76; CELF (CUGBP- and ETR3-like
factor)-family proteins ETR3 and CUG-BP activate the SM exon of -actinin splicing by
displacing PTB77. A recent study showed that at least some Drosophila SR proteins and
hnRNP A family proteins do not have as many common targets as had been thought78.
Using siRNAs to deplete individual splicing factors followed by splicing sensitive microarray analysis, the authors compared genes regulated in opposite directions by hnRNP proteins and two SR proteins (dASF and BC52). Surprisingly, less than 5% of the genes overlap. However, a systematic analysis of more SR and hnRNP proteins will be necessary to make a conclusion as to whether antagonizing effects between these proteins is a major mode of AS regulation.
Position-dependent splicing regulation
The nature of the activity of cis-acting elements and their cognate binding proteins in some cases depends on their position relative to regulated exons. Several proteins, such as Nova 1/2, Fox1/2, hnRNP L, hnRNPL-like protein and hnRNP F/H, have been shown to act as either repressors or activators depending on the location of their binding site11-14,
79-83 . For example, Nova-1 binds to an ISE in GABAAR2 pre-mRNA and promotes
inclusion of exon 984, but it binds to the ESS in the alternative exon 4 of its own pre-
mRNA and prevents exon 4 from being included81. Similar to Nova, hnRNP L can
activate or repress upstream AEs, likely dependent on the location of its binding site
relative to the regulated 5’ss12. hnRNP H promotes formation of ATP-dependent
spliceosomal complexes when it binds to G-runs that are located downstream of the
8 10 5’ss85, but it inhibits splicing when the G-runs are located in exons86. By using
information from validated Nova1/2-targeted transcripts and searching YCAY clusters
(Nova1/2 binding sequences) around regulated exons, Darnell and colleagues drew an
‘RNA map’ that includes the location of Nova1/2-binding sties and consequence of each
binding event11. This map provides insight into the mechanisms underlying Nova1/2’s
effects on splicing. Binding of Nova1/2 to an ESS inhibits formation of prespliceosomal
E complex by altering the composition of the pre-spliceosome complex before the binding of hnRNP proteins and inhibits the binding of U1 snRNP11. By contrast,
Nova1/2 binding to an ISE downstream of the AE promotes formation of spliceosomal
complexes A, B and C11. A novel technique, combining CLIP and high-throughput
sequencing, called HITS-CLIP80 (CLIP-seq)13, 87, 88, not only verified the reproducibility
of the RNA map, but also provided genome-wide information on Nova1/2 and Fox2
target genes and possibly mechanisms of regulation80.
Why does the position of splicing regulatory elements determine the action of
cognate splicing factors? It is possible that enhancers are in positions such that when the
splicing factors bind to them, the splice sites of the AE are better presented to the splicing
machinery by changing local RNA structure. In contrast, silencing elements function in
the opposite manner, by competing with components of the splicing machinery or by
changing RNA structure to impede ss recognition.
Roles for RNA in AS regulation
Splice site selection can also be influenced by RNA secondary structure within the pre-
mRNA, and this can occur in multiple different ways. Perhaps the most striking example
9 11 is the complex AS of Drosophila Dscam pre-mRNA. In the case of the exon 6 cluster,
which consists of 48 mutually exclusive exons, it appears that pairing between a
conserved sequence located downstream of constitutive exon 5 (the docking site) and
another conserved sequence, a variant of which is located upstream of each exon 6
variant (the selector sequence), allows inclusion of only one exon 6 variant89, with exclusion of the remainder insured by binding of hrp36, a Drosophila hnRNP A homologue90. Secondary structures can affect AS by masking splice sites91 or binding
sites for splicing factors92, 93. For example, secondary structure was shown to sequester
alternative exon 6B of the chicken -tropomyosin pre-mRNA, leading to its exclusion94.
The IDX exon of H-ras can form a secondary structure with an ISS (rasISS1), preventing the binding of hnRNP H to rasISS1. Unwinding this secondary structure by an RNA helicase, P68, exposes the binding site, partially explaining the exclusion of IDX93.
Riboswitches, well known to control gene expression in prokarotes95, also have the
potential to modulate AS. Splicing of the NMT1 pre-mRNA in Neurospora crassa was
found to respond to the coenzyme thiamine pyrophosphate via a riboswitch-like
structure96. It remains to be seen however whether this intriguing mechanism operates in
higher eukaryotes. In mammals, small nucleolar RNAs (snoRNAs) have also been
97 implicated in AS regulation . For example, snoRNA HBII-52, regulates AS of 5-HT2cR
pre-mRNA by binding to a silencing element in exon Vb to promote inclusion of Vb98.
Regulation by U1 and U2 snRNP pairing
After the 5’ss and 3’ss are recognized and exons are defined, exon definition must be converted to intron definition, which involves cross-intron interaction between U1 and
10 12 U2 snRNPs, to form a functional spliceosome. Precisely when this occurs and when the
commitment to ss pairing happens have been intensively investigated over the past few
years. Several studies have shown that the commitment to splicing of at least some AEs
occurs during ss pairing in the A complex35, 36. For example, ATP hydrolysis is required
for ss pairing, which locks splice sites into a splicing pattern after U2 snRNP binding to
the branch site36. Additional studies, discussed below, also provided evidence that
binding of U1 and U2 snRNPs to splice sites to define an exon does not necessarily
commit the exon to splicing15, 16, 99.
Regulation by protein factors
A novel splicing inhibition mechanism was demonstrated recently by Lynch and colleagues, who found that binding of hnRNP L to an ESS can inhibit the pairing of U1 and U2 snRNPs15. An ATP-dependent spliceosomal-like complex, A-like exon-definition
complex (AEC), was found to form across alternative exon 4 of the CD45 pre-mRNA,
even when its inclusion is inhibited. The AEC contains U1 and U2 snRNPs and displays
the same gel mobility as A complex, but progression into B complex is inhibited. They
proposed a model such that when hnRNP L is not present, an A-like complex forms
across exons, then the U4·U6/U5 tri-snRNP is recruited to the intron-defined A complex
to form B complex. However, when hnRNP L is present, an hnRNP L-containing AEC
prevents the U1 or U2 snRNPs bound to the splice sites of exon 4 from cross-intron
pairing with the adjacent U2 or U1, resulting in exon 4 skipping. There are two possible
ways in which the binding of hnRNP L might interfere with the U snRNP pairing. One is
that binding of hnRNP L physically shields the interaction between the snRNPs, while
11 13 another is that hnRNP L induces a change in the conformation of the pre-mRNA that
prevents cross-intron pairing of the snRNP-bound AE. It is intriguing that the AEC, at
least superficially, resembles the “stalled” A complex formed by SRp38 in S100 (see
above). While the significance of this is unknown, the following example suggests that
such complexes may be more widespread than realized.
PTB is another inhibitory splicing factor that has been shown to function, in some
cases, by blocking the transition from exon definition to intron definition16, 65 (FIG. 2a).
Sharma et al. studied the mechanism of PTB inhibition by comparing the active and inactive spliceosomal complexes from neuronal WERI-1 nuclear extract (NE), where c- src N1 exon is included, and HeLa NE, where N1 is excluded16. PTB is highly expressed
in HeLa cells, whereas a less repressive brain paralogue, nPTB, is expressed in WERI-1
cells100-103 (see below). Similar to the example provided by hnRNP L, ATP-dependent
exon definition complexes form in both NEs. The protein compositions of the exon
definition E and A complexes (EDE and EDA, respectively) formed on constitutive exon
4 from both NEs were very similar, and PTB was only found in complexes formed in
HeLa NE. By contrast, the EDE and EDA complexes formed on the substrate containing
both exon N1 and exon 4 in WERI-1 NE and in HeLa NE differed in their properties and protein compositions. The WERI-1 EDE can transit to functional A, B and C complexes upon ATP addition, whereas the EDE formed in HeLa NE can only transit to a “dead-end
A” complex. Several proteins only exist in the functional A complex formed in WERI-1
NE, such as the Prp19 complex104, 105 and SRm160/30046, 106, which are likely to be
important for 3’ and 5’ss bridging and exon N1 inclusion, and are excluded from the
spliceosome by PTB. This study provides a novel mechanism for how different protein
12 14 compositions in different tissues can help determine the AS pattern, through a silencing
factor and its interactions other splicing factors to prevent intron definition. This study
also for the first time revealed that the protein composition of different exon and intron
definition complexes can vary, and thus begins to decipher a novel mechanism for AS
regulation and tissue specificity.
RBM5 (also known as Luca-15 and H37) is a putative tumor suppressor protein
107, 108 that promotes exclusion of exon 6 of the Fas receptor pre-mRNA109. Valcarcel and colleagues found that RBM5 interacts with sequences in exon 6, but does not affect the association of U1 and U2 snRNPs to the adjacent splice sites, but instead inhibits incorporation of the U4·U6/U5 tri-snRNP on the introns flanking exon 6, thereby
blocking maturation of prespliceosomes. In addition, RBM5 also promotes pairing of U1
and U2 at the distal splice sites, contributing further to exclusion of exon 6 (FIG. 2b).
In addition to AS regulation by inhibition of intron definition, it is also possible
that AS is stimulated by activation of intron definition. In vitro experiments with
substrates containing expanded introns have shown that the presence of binding sites for
hnRNP proteins near intron boundaries can facilitate splicing82. This presumably involves
cross-intron interactions between hnRNP proteins that help bring together the ends of the
introns, and suggests a possible positive role for hnRNP proteins in splicing.
Cis-acting elements have recently been identified that affect U1 and U2 snRNP
pairing by modulating 5’ss competition. Nilsen and colleagues performed an in vitro
screen for splicing silencers (ESSs and ISSs) that alter 5’ss selection by choosing a distal,
weak 5’ss over a proximal, strong 5’ss99. The isolated silencers did not affect whether U1
snRNP binds to the 5’ss, but rather appear to alter in some way the conformation of the
13 15 proximal U1 snRNP-5’ss complex so that it loses its advantage to compete with the distal
U1 snRNP-5’ss complex to pair with the U2 snRNP-3’ss complex. This study suggests a
new mechanism for how silencers can subtly affect ss selection, not by sequestering splice sites, but by changing the conformation of the snRNP/pre-mRNA complex.
Moreover, the silencers do not affect the rate-limiting step of splicing per se but rather affect how well U1 and U2 snRNPs pair, which can in turn influence ss choice.
Transcription-coupled regulation of AS
Two models have been proposed to explain the role of RNA polymerase II in AS regulation110. In one, the recruitment model, RNA polymerase II (RNAP II) and
transcription factors interact, directly or indirectly, with splicing factors22, 111, 112, thereby increasing or decreasing the efficiency of splicing. Kornblihtt and colleagues have shown that promoter structure can affect the splicing pattern of the fibronectin pre-mRNA by in some way facilitating differential recruitment of ASF/SF2113. It is conceivable that this
reflects the ability of different transcription factors to influence recruitment of different
splicing factors to the nascent pre-mRNA, resulting in inclusion or exclusion of the AE.
PGC-1, a transcription coactivator recruited to target genes by specific transcription
factors, can modulate AS of nascent transcripts by interacting, via its RS domain, with
other splicing factors114.
Another study showed that differential recruitment of transcription coactivators
to progesterone- and estrogen-responsive elements upstream of reporter genes, such as
CD44, alters AS of the resultant transcripts112. Specifically, recruitment of activating
signal cointegrator (ASC)-1 and -2 and associated proteins to these elements was found
14 16 both to activate transcription of the reporter genes and to affect splicing, but in opposite
ways. The ASC-1 associated protein CAPER contains an RS domain and two RRM
domains, similar to SR proteins, while an ASC-2 associated protein, CoAA, is
structurally related to hnRNP A1. It was proposed that the antagonistic effects by ASC-1
and 2 complexes are mediated by these factors.
A second, kinetic, model proposes that the rate of transcription elongation
influences the inclusion of AEs by affecting whether the splicing machinery is recruited
sufficiently quickly for spliceosome assembly/splicing to occur. In support of this model,
an RNAP II with a reduced elongation rate caused by point mutation was found to greatly
stimulate the inclusion of an AE that has weak splice sites115. Specifically, it was found that slow transcription favors inclusion of the fibronectin EDI exon, which was excluded when transcription was more rapid. Consistent with this, Kornblihtt and colleagues recently showed that changes in RNAP II elongation rate upon UV irradiation could lead to AS changes that occur in response to DNA damage116.
One way of changing transcription rate is through changing the phosphorylation
status of RNAP II. The C-terminal domain of the RNAPl II largest subunit (CTD)
consists of up to 52 tandem repeats of the heptapeptide consensus sequence YSPTSPS117.
Excess Ser5 phosphorylation on the CTD is associated with RNAP II stalling downstream of the promoter region, whereas Ser2 phosphorylation is associated with elongation through the gene117, 118. Batsche et al. showed that a subunit of the human
SWI/SNF complex, Brm, regulates changes in AS of CD44 pre-mRNA stimulated by T- cell activation18. Upon T-cell stimulation, Ser5 phosphorylated RNAP II pauses at the
variant exon region of the CD44 gene, by a mechanism requiring Brm association.
15 17 Interestingly, this also results in increased association of Brm with components of the
splicing machinery and splicing factor Sam68, leading to inclusion of the V5 exon. This
study not only provides direct support of the kinetic model, but also demonstrates that the
mechanism for AS regulation by transcription can result from a combination of
transcription elongation-related effects and differential recruitment of splicing factors.
Alternative splicing and tissue specificity
AS regulation by tissue-specific splicing factors
AS plays an important role in defining tissue specificity. Recent high-throughput studies have shown that, of the human tissues examined, 50% or more of AS isoforms are differently expressed among tissues29, indicating that most AS is subject to tissue-specific
regulation.
Tissue-specific AS events can be explained in part by tissue-specific expression
of splicing factors, and the corresponding regulation of their target transcripts31, 119, 120. In
keeping with this, a number of tissue-specific AS regulators have now been identified
(TABLE 2). Among all human tissues, brain is the most functionally diverse tissue, with
the highest occurrence of tissue-specific AS isoforms. Accordingly, a number of brain-
specific factors have been identified, such as nPTB101, 121, Nova1/281, 84, 122 and Hu/Elav
family proteins123-125. In addition, region- and cell type-specific expression of most of the >300 RNA-binding proteins examined was observed in proliferating versus post- mitotic mouse brain126.
PTB is expressed in neural progenitor cells, but its expression level is greatly
downregulated in differentiated neurons, where nPTB, also called brPTB or PTBP2, is
16 18 upregulated100, 101. Recent experiments provided evidence that the PTB-to-nPTB switch
provides a post-transcriptional mechanism that is important for programming neuronal
differentiation101. Using microarray analysis, it was shown that siRNA-mediated PTB
depletion in N2A neuroblastoma cells caused upregulation of nPTB and an altered AS
pattern. Most of the observed changes were also detected when P19 cells (derived from
an embryonal carcinoma) were differentiated into neuronal cells. In neuronal cells, expression of nPTB and downregulation of PTB explains ~25% of neural system (NS)-
specific AS101. However, the molecular mechanism that allows NS-specific AEs to be included in neuronal cells even when bound by nPTB is still unclear. It may reflect differences in the ability of nPTB and PTB to interact with other splicing factors and/or in the presence or absence of other splicing factors, for example Nova proteins, in neuronal and non-neuronal cells.
Other brain-specific factors, including the Nova proteins, may be involved in fine tuning the programming of different types of neuronal cells. The expression of Nova1 and Nova2 in postnatal mouse brain appears reciprocal, especially in the neocortex and hippocampus where Nova2 is highly expressed while Nova-1 is expressed primarily in
hindbrain and spinal cord127. Nova-1 null mice die postnatally from a motor deficit
associated with apoptotic death of spinal and brainstem neurons whereas Nova2
conditional knockout mice showed that Nova2 regulates ~7% of brain-specific splicing in
the neocortex and Nova2-dependent AS regulates mRNAs that encode synaptic
functions122, indicating that these proteins regulate AS events that have specific functions
in the brain. Similar to the roles of PTB and nPTB in helping to define non-neuronal and
neuronal tissues, expression of Nova1 and Nova2 may contribute to different functions of
17 19 different brain regions. However, due to the limited number of exons analyzed and the
overlap between PTB/nPTB and Nova targets, more detailed studies are needed to obtain
a more complete understanding of tissue-specific AS regulation by these factors.
Tissue-specific AS factors have recently been shown to be important in
controlling expression of epithelial-specific exons. Carstens and colleagues identified two
paralogues, RBM35a (ESRP 1) and RBM35b (ESRP 2), to be important for inclusion of
epithelial cell-specific exons in several transcripts128. In addition, downregulation of
RBM35a was found to coincide with loss of epithelial splicing during the epithelial to
mesenchymal transition, and ectopic expression of RBM35a in mesenchymal cells
restored epithelial splicing. These data show that RBM35a/b contributes to defining the
distinguishing characteristics of epithelial cells.
AS regulation by constitutive splicing factors
SR proteins were originally discovered by biochemical methods as general, or non-
sequence-specific, splicing activators7. However, more recent findings indicate that
individual SR proteins can act as specific AS regulators in different cells and tissues.
Heart-specific disruptions of ASF/SF2 and SC35 genes have shown that they play
important but distinct roles in tissue development129, 130. In addition, a recent study showed that mice with complete ablation of SRp38 survived through early embryogenesis, and strikingly, displayed only cardiac defects, and only limited differences in AS were observed.131.
Core spliceosomal proteins (CSPs) are also involved in AS regulation. Analysis of
microarray-based expression profiles from mouse, chimpanzee and human tissues
18 20 revealed that snRNP proteins are differentially expressed in particular tissues132. This is
consistent with results from an RNAi screen in Drosophila, which showed that changing
levels of CSPs leads to changes in AS26. These CSPs include components of U1, U2 and
U4/U6 snRNPs, and also U2AF proteins. Further evidence was provided by RNAi
knockdown of isoforms of U2AF35, U2AF35a and/or U2AF35b, and a subunit of
splicing factor SF3B, SAP155, in human cells44, 133, 134. Knockdown of these CSPs was
found to affect only AS of a subset of transcripts: for example, mRNAs encoding cell
cycle phosphatases in the case of U2AF35 and 5’ss selection of Bcl-x pre-mRNA in
response to ceramide in the case of SAP155. In addition, evidence from budding yeast,
showing differences in splicing patterns in response to different kinds of stress, also suggests that CSPs are involved in AS regulation135.
The survival of motor neurons (SMN) protein, which is part of the SMN complex,
has recently been shown to regulate AS in multiple mouse tissues27. The SMN complex is
important for efficient assembly of snRNPs, and depletion of SMN in HeLa cells leads to
a decrease in snRNP levels. SMN-deficient mice showed tissue-specific alterations in
snRNAs, and different snRNPs were affected in different tissues, leading to an altered
stoichiometry of snRNPs27. In addition, microarray analysis of RNA samples from
different tissues of SMN-deficient mice revealed changes in a number of AS events in
various tissues27. The mechanism of AS alteration by SMN deficiency is unknown, but it
is likely that the resulting changes in snRNP levels directly affected AS of specific
transcripts. It will also be important to understand how, and if, these changes in AS
contribute to spinal muscular atrophy, which is caused by SMN deficiency136, 137.
19 21 AS regulation by post-translational modifications of splicing factors
Differences in protein expression levels of either tissue-specific splicing regulators or
CSPs may not fully explain how cells are able to rapidly change AS patterns, for example, in response to cellular stress. Mounting evidence has shown that post-translational modification of splicing factors can affect AS. The best studied modification is phosphorylation, and consistent with this, several well-studied cell signaling pathways have been shown to be involve AS regulation (reviewed in refs, 138, 139). Phosphorylation has been shown to affect the local concentration of splicing factors adjacent to RNA substrates by altering their intracellular localization140-144, protein-protein43 and protein-
RNA interaction s50, 145 and even intrinsic splicing activity39, 54.
Phosphorylation can change the ability of splicing factors to interact with other proteins or RNA substrates, leading to changes in ss selection. Phosphorylation of RS domains in SR proteins affects interactions with CSPs43 and is necessary for sequence- specific RNA binding in vitro146. Phosphorylation of TIA-1/TIAR by Fas-activated serine-threonine kinase (FAST K) enhances TIA-1 mediated recruitment of U1 snRNP to a suboptimal 5’ss, leading to the inclusion of Fas exon 6147. Tyrosine phosphorylation of
Sam68 by Fyn kinase favors formation of the anti-apoptotic Bcl-xL mRNA by interfering with the interaction between Sam68 and hnRNP A1, and with their interaction with the pre-mRNA148. Phosphorylation of RS motifs of PTB-associated splicing factor PSF by
SR kinases inhibits PSF RNA binding145.
Phosphorylation can also change the intracellular localization of splicing factors.
Osmotic shock stresses cells and activates the MEK3/6-p38 signaling pathway, leading to relocalization of hnRNP A1 to the cytosol as a result of hyperphosphorylation. This
20 22 change in hnRNP A1 localization can alter the AS pattern of an adenovirus E1A reporter transcript141, 144. Ischemia triggers changes in Ca2+ concentration, leading to
hyperphosphorylation of a Tra2 isoform, Tra2-1, and to its localization to the
cytosoplasm143. Protein kinase A phosphorylates PTB on ser16, which leads to its
translocation to the cytoplasm149, 150.
Phosphorylation status can also, in one case, determine whether a splicing factor
functions as a splicing repressor or activator. SRp38 functions as a global splicing repressor when dephosphorylated in M phase of the cell cycle and following heat shock54,
55. However, it becomes a sequence-specific activator when phosphorylated39. Shi and
Manley elucidated the detailed regulatory mechanism of SRp38 phosphorylation in
response to heat shock151. At normal temperatures, two mechanisms ensure that SRp38 remains phosphorylated (FIG. 3c). One is that SRp38 is bound and protected by 14-3-3 proteins, while the other is that the phosphatase PP1, shown to target SRp38, is masked by associated proteins, including NIPP1, which had previously been implicated in splicing control152, 153. At elevated temperatures, 14-3-3 proteins dissociate from SRp38
and PP1 is released from NIPP1, thereby freeing PP1 to dephosphorylate SRp38. Unlike other SR proteins, SRp38 is a very poor substrate for the SR protein kinases Clk/Sty and
SRPK1. Therefore, after dephosphorylation, SRp38 remains dephosphorylated and thus able to repress splicing events. Other SR proteins, if dephosphorylated, are rapidly re- phosphorylated.
Conclusions and perspectives
21 23 The studies described here illustrate the complexity of AS regulation. As we have
seen, AS can be regulated at different steps of spliceosome assembly by different splicing
factors, both general and specific, and by multiple mechanisms AS relies on cis-acting
elements. Although AEs are shorter than CEs and are flanked by longer introns, AEs are
more conserved than CEs, especially the exon-intron junctions, and these conserved
regions often extend into flanking introns for 80-100 nucleotides154,where cis-regulatory elements are embedded. Correct AS also depends on the stoichiometry and interactions of
positive and negative regulatory proteins, including CSPs. Each cell type has a unique
repertoire of SR proteins and hnRNP proteins, and moderate changes in their relative
stoichiometry can have significant effects on AS pattern1, 63. It is possible that changes in
the stoichiometry of snRNPs perturb the complex network of splicing factors and the
interactions between splicing factors and CSPs. Therefore, AS regulatory networks
constitute such an exquisite architecture that perturbation of any single step can lead to
AS misregulation.
Diverse mechanisms are utilized to ensure tissue and cell type-specific splicing
regulation. Increasing evidence has shown that AS plays an important role in defining
tissue specificity. The action of sequence-specific transcription factors has been
considered the most robust ways of defining tissue specificity155. Significantly, >2,500
transcription factors have been identified in humans156, while the number of sequence-
specific AS factors reported is <50. Given that it now appears that AS is as prevalent and
perhaps as important a mechanism as transcriptional control, what might be the
explanation for this? One possibility is that many more splicing regulators remain to be
discovered. However, the total number of putative RNA-binding proteins (RBPs) (in
22 24 mouse) has been estimated to be less than 400126, and some fraction of these will not be
involved in splicing. Another possibility is that there are fundamental differences in how
splicing and transcription are regulated. For example, individual splicing regulators
control much larger groups of genes than do specific transcription factors. This is
consistent with the large numbers of neuronal transcripts thought to be controlled by PTB
and Nova80, 101, 103, 122. It is also likely that significant regulation is achieved by
combinations of abundant regulators with limited sequence-specificity (that is, the SR
and hnRNP proteins), which act in concert to regulate different transcripts in different tissues dependent on their relative concentrations.
Important goals of future studies of AS regulation include understanding how regulators switch important splicing events during development and in response to environmental stimuli, and how misregulation of AS leads to disease. Complete characterization of tissue-specific patterns of expression is of great importance to defining mechanisms of AS regulation in different cell types. De novo identification of regulatory motifs31 and HITS-CLIP80 are two complementary approaches to achieve this
goal. Until now though, only a limited number of AS regulators and tissues have been
studied by these methods. In addition to Nova and Fox2, CLIP-seq also provided a
landscape of potential ASF/SF2 target transcripts, and characterized a purine-rich
consensus motif87, 88 that is nearly identical to a consensus obtained previously by an in
vitro SELEX approach157.A database that includes more comprehensive information on
AS regulatory protein expression patterns, definition of potential target transcripts and
positions of binding motifs on these transcripts, will facilitate searches for regulatory
23 25 proteins that control specific splicing events, such as with genes that are involved in
disease, and possibly provide insights into underlying mechanisms.
Due to the dynamic nature of spliceosome assembly and the potential for
regulation at multiple points, detailed proteomic analysis will be important in completely
elucidating the molecular mechanisms of AS regulation. For example, how do RNA
binding proteins interact with other factors and core splicing factors? When and where do
splicing factors function to regulate the spliceosome? How do posttranslational
modifications influence these events? Obtaining a full understanding of the mechanisms
underlying alternative splicing and its role in defining tissue specificity will require
multiple approaches and methods.
Acknowledgements
Work from the authors' laboratory was supported in part by grants from the National Institutes of Health. We thank Charles David for comments on the manuscript.
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30 32 112. Auboeuf, D. et al. Differential recruitment of nuclear receptor coactivators may determine alternative RNA splice site choice in target genes. Proc Natl Acad Sci U S A 101, 2270-4 (2004). 113. Cramer, P. et al. Coupling of transcription with alternative splicing: RNA pol II promoters modulate SF2/ASF and 9G8 effects on an exonic splicing enhancer. Mol Cell 4, 251-8 (1999). 114. Monsalve, M. et al. Direct coupling of transcription and mRNA processing through the thermogenic coactivator PGC-1. Mol Cell 6, 307-16 (2000). 115. de la Mata, M. et al. A slow RNA polymerase II affects alternative splicing in vivo. Mol Cell 12, 525-32 (2003). 116. Munoz, M.J. et al. DNA damage regulates alternative splicing through inhibition of RNA polymerase II elongation. Cell 137, 708-20 (2009). 117. Phatnani, H.P. & Greenleaf, A.L. Phosphorylation and functions of the RNA polymerase II CTD. Genes Dev 20, 2922-36 (2006). 118. Lin, P.S., Marshall, N.F. & Dahmus, M.E. CTD phosphatase: role in RNA polymerase II cycling and the regulation of transcript elongation. Prog Nucleic Acid Res Mol Biol 72, 333-65 (2002). 119. David, C.J. & Manley, J.L. The search for alternative splicing regulators: new approaches offer a path to a splicing code. Genes Dev 22, 279-85 (2008). 120. Ohno, G., Hagiwara, M. & Kuroyanagi, H. STAR family RNA-binding protein ASD-2 regulates developmental switching of mutually exclusive alternative splicing in vivo. Genes Dev 22, 360-74 (2008). 121. Li, Q., Lee, J.A. & Black, D.L. Neuronal regulation of alternative pre-mRNA splicing. Nat Rev Neurosci 8, 819-31 (2007). 122. Ule, J. et al. Nova regulates brain-specific splicing to shape the synapse. Nat Genet 37, 844-52 (2005). 123. Perrone-Bizzozero, N. & Bolognani, F. Role of HuD and other RNA-binding proteins in neural development and plasticity. J Neurosci Res 68, 121-6 (2002). 124. Zhu, H., Hasman, R.A., Barron, V.A., Luo, G. & Lou, H. A nuclear function of Hu proteins as neuron-specific alternative RNA processing regulators. Mol Biol Cell 17, 5105-14 (2006). 125. Soller, M., Li, M. & Haussmann, I.U. Regulation of the ELAV target ewg: insights from an evolutionary perspective. Biochem Soc Trans 36, 502-4 (2008). 126. McKee, A.E. et al. A genome-wide in situ hybridization map of RNA-binding proteins reveals anatomically restricted expression in the developing mouse brain. BMC Dev Biol 5, 14 (2005). 127. Yang, Y.Y., Yin, G.L. & Darnell, R.B. The neuronal RNA-binding protein Nova- 2 is implicated as the autoantigen targeted in POMA patients with dementia. Proc Natl Acad Sci U S A 95, 13254-9 (1998). 128. Warzecha, C.C., Sato, T.K., Nabet, B., Hogenesch, J.B. & Carstens, R.P. ESRP1 and ESRP2 are epithelial cell-type-specific regulators of FGFR2 splicing. Mol Cell 33, 591-601 (2009). 129. Ding, J.H. et al. Dilated cardiomyopathy caused by tissue-specific ablation of SC35 in the heart. EMBO J 23, 885-96 (2004).
31 33 130. Xu, X. et al. ASF/SF2-regulated CaMKIIdelta alternative splicing temporally reprograms excitation-contraction coupling in cardiac muscle. Cell 120, 59-72 (2005). 131. Feng, Y. et al. SRp38 Regulates Alternative Splicing and Is Required for Ca(2+) Handling in the Embryonic Heart. Dev Cell 16, 528-538 (2009). 132. Grosso, A.R. et al. Tissue-specific splicing factor gene expression signatures. Nucleic Acids Res 36, 4823-32 (2008). 133. Massiello, A., Roesser, J.R. & Chalfant, C.E. SAP155 Binds to ceramide- responsive RNA cis-element 1 and regulates the alternative 5' splice site selection of Bcl-x pre-mRNA. FASEB J 20, 1680-2 (2006). 134. Pacheco, T.R., Moita, L.F., Gomes, A.Q., Hacohen, N. & Carmo-Fonseca, M. RNA interference knockdown of hU2AF35 impairs cell cycle progression and modulates alternative splicing of Cdc25 transcripts. Mol Biol Cell 17, 4187-99 (2006). 135. Pleiss, J.A., Whitworth, G.B., Bergkessel, M. & Guthrie, C. Rapid, transcript- specific changes in splicing in response to environmental stress. Mol Cell 27, 928- 37 (2007). 136. Monani, U.R. Spinal muscular atrophy: a deficiency in a ubiquitous protein; a motor neuron-specific disease. Neuron 48, 885-96 (2005). 137. Gabanella, F. et al. Ribonucleoprotein assembly defects correlate with spinal muscular atrophy severity and preferentially affect a subset of spliceosomal snRNPs. PLoS ONE 2, e921 (2007). 138. Shin, C. & Manley, J.L. Cell signalling and the control of pre-mRNA splicing. Nat Rev Mol Cell Biol 5, 727-38 (2004). 139. Tarn, W.Y. Cellular signals modulate alternative splicing. J Biomed Sci 14, 517- 22 (2007). 140. Huang, Y., Yario, T.A. & Steitz, J.A. A molecular link between SR protein dephosphorylation and mRNA export. Proc Natl Acad Sci U S A 101, 9666-70 (2004). 141. van der Houven van Oordt, W. et al. The MKK(3/6)-p38-signaling cascade alters the subcellular distribution of hnRNP A1 and modulates alternative splicing regulation. J Cell Biol 149, 307-16 (2000). 142. Habelhah, H. et al. ERK phosphorylation drives cytoplasmic accumulation of hnRNP-K and inhibition of mRNA translation. Nat Cell Biol 3, 325-30 (2001). 143. Daoud, R. et al. Ischemia induces a translocation of the splicing factor tra2-beta 1 and changes alternative splicing patterns in the brain. J Neurosci 22, 5889-99 (2002). 144. Guil, S., Long, J.C. & Caceres, J.F. hnRNP A1 relocalization to the stress granules reflects a role in the stress response. Mol Cell Biol 26, 5744-58 (2006). 145. Huang, C.J., Tang, Z., Lin, R.J. & Tucker, P.W. Phosphorylation by SR kinases regulates the binding of PTB-associated splicing factor (PSF) to the pre-mRNA polypyrimidine tract. FEBS Lett 581, 223-32 (2007). 146. Tacke, R., Chen, Y. & Manley, J.L. Sequence-specific RNA binding by an SR protein requires RS domain phosphorylation: creation of an SRp40-specific splicing enhancer. Proc Natl Acad Sci U S A 94, 1148-53 (1997).
32 34 147. Izquierdo, J.M. & Valcarcel, J. Fas-activated serine/threonine kinase (FAST K) synergizes with TIA-1/TIAR proteins to regulate Fas alternative splicing. J Biol Chem 282, 1539-43 (2007). 148. Paronetto, M.P., Achsel, T., Massiello, A., Chalfant, C.E. & Sette, C. The RNA- binding protein Sam68 modulates the alternative splicing of Bcl-x. J Cell Biol 176, 929-39 (2007). 149. Ma, S., Liu, G., Sun, Y. & Xie, J. Relocalization of the polypyrimidine tract- binding protein during PKA-induced neurite growth. Biochim Biophys Acta 1773, 912-23 (2007). 150. Xie, J., Lee, J.A., Kress, T.L., Mowry, K.L. & Black, D.L. Protein kinase A phosphorylation modulates transport of the polypyrimidine tract-binding protein. Proc Natl Acad Sci U S A 100, 8776-81 (2003). 151. Shi, Y. & Manley, J.L. A complex signaling pathway regulates SRp38 phosphorylation and pre-mRNA splicing in response to heat shock. Mol Cell 28, 79-90 (2007). 152. Van Eynde, A. et al. Molecular cloning of NIPP-1, a nuclear inhibitor of protein phosphatase-1, reveals homology with polypeptides involved in RNA processing. J Biol Chem 270, 28068-74 (1995). 153. Cohen, P.T. Protein phosphatase 1--targeted in many directions. J Cell Sci 115, 241-56 (2002). 154. Kim, E., Goren, A. & Ast, G. Alternative splicing: current perspectives. Bioessays 30, 38-47 (2008). 155. Hallikas, O. et al. Genome-wide prediction of mammalian enhancers based on analysis of transcription-factor binding affinity. Cell 124, 47-59 (2006). 156. Babu, M.M., Luscombe, N.M., Aravind, L., Gerstein, M. & Teichmann, S.A. Structure and evolution of transcriptional regulatory networks. Curr Opin Struct Biol 14, 283-91 (2004). 157. Tacke, R. & Manley, J.L. The human splicing factors ASF/SF2 and SC35 possess distinct, functionally significant RNA binding specificities. EMBO J 14, 3540-51 (1995). 158. Kent, O.A., Ritchie, D.B. & Macmillan, A.M. Characterization of a U2AF- independent commitment complex (E') in the mammalian spliceosome assembly pathway. Mol Cell Biol 25, 233-40 (2005). 159. Shen, H. & Green, M.R. RS domains contact splicing signals and promote splicing by a common mechanism in yeast through humans. Genes Dev 20, 1755- 65 (2006). 160. Huang, Y., Gattoni, R., Stevenin, J. & Steitz, J.A. SR splicing factors serve as adapter proteins for TAP-dependent mRNA export. Mol Cell 11, 837-43 (2003). 161. Zhang, Z. & Krainer, A.R. Involvement of SR proteins in mRNA surveillance. Mol Cell 16, 597-607 (2004). 162. Sanford, J.R., Gray, N.K., Beckmann, K. & Caceres, J.F. A novel role for shuttling SR proteins in mRNA translation. Genes Dev 18, 755-68 (2004). 163. Cartegni, L. & Krainer, A.R. Disruption of an SF2/ASF-dependent exonic splicing enhancer in SMN2 causes spinal muscular atrophy in the absence of SMN1. Nat Genet 30, 377-84 (2002).
33 35 164. Cartegni, L., Hastings, M.L., Calarco, J.A., de Stanchina, E. & Krainer, A.R. Determinants of exon 7 splicing in the spinal muscular atrophy genes, SMN1 and SMN2. Am J Hum Genet 78, 63-77 (2006).
Box 1 Splicing and spliceosome assembly
Pre-mRNA splicing is a process in which intervening sequences (introns) are removed from an mRNA precursor. Splicing consists of two transesterification steps, each involving a nucleophilic attack on terminal phosphodiester bonds of the intron, by the 2’ hydroxyl of the branch-point (usually adenosine) in the first step and by the 3’ hydroxyl of the upstream (5’) exon in the second step1, 2. This process is carried out in the
spliceosome, a dynamic molecular machine the assembly of which involves sequential
binding and release of snRNPs and numerous protein factors as well as the formation and
disruption of RNA-RNA, protein-RNA and protein-protein interactions.
The basic mechanics of spliceosome assembly are well known. Briefly, the
process begins with the base pairing of U1 snRNA to the 5’-splice site (ss) and the
binding of splicing factor 1 (SF1) to the branch point3 in an ATP-independent manner to
form the E’ complex (see the figure). The E’ complex can be chased into the E complex
by the recruitment of U2 auxiliary factors (U2AF) heterodimer (comprising U2AF65 and
U2AF35) to the polypyrimidine tract (py tract) and 3’ terminal AG158. The ATP-
independent E complex is converted into the ATP-dependent pre-spliceosome A complex
by the replacement of SF1 by U2 snRNP at the branch point. Further recruitment of the
U4·U6/U5 tri-snRNP leads to the formation of B complex, which contains all
spliceosomal subunits to carry out splicing. This is followed by an extensive
34 36 conformational change and remodeling, including the loss of U1 and U4 snRNPs,
resulting in the formation of C complex, the catalytically active splicesome.
Box 2 SR proteins
The SR (Ser-Arg) proteins are a family of nuclear factors that play multiple important
roles in splicing of mRNA precursors in metazoan organisms, functioning in both
constitutive and alternative splicing7. They are involved in multiple steps of splicing regulation, by binding ESEs through their RRM domains, and mediating protein-protein41,
42, and perhaps protein-RNA159 interactions through their RS domains. All canonical SR
proteins share common characteristics (see table). They display similar structures with
one or two RNP-type RNA-binding domains at their N-termini and a variable-length
domain enriched in Ser-Arg dipeptides at their C-termini (the RS domain). RS domains
are extensively phosphorylated and they function in splicing, usually as activators. Most
SR proteins function as pivotal regulators in multiple aspects of mRNA metabolism, such
as mRNA nuclear export160, nonsense-mediated mRNA decay161 and translation162. A
number of additional RS domain-containing proteins have been identified and those
known to be involved in AS are listed in the table.
35 37
Figure legends
Figure 1│ Mechanisms of alternative splicing by splice site selection. a│ Mechanisms
of splicing activation. SR proteins bind to ESEs to stimulate U2AF binding to the
upstream 3’ss, or U1 snRNP binding to the downstream 5’ss. SR proteins function with
other splicing coactivators, such as Tra2 and SRm160/300. TIA-1 binds to U-rich
sequences (ISEs) immediately downstream of 5’ss to facilitate U1 binding. CELF
proteins, such as ETR-3, bind to similar sequences as does PTB, thereby activating
splicing by competing with PTB. b│ Fox1/2 inhibits inclusion of calcitonin/CGRP exon
4 by blocking binding of SF1 to the branch site (top panel) and Tra2 and SRp55 to ESEs
(bottom panel) thereby inhibiting spliceosome assembly at two stages, the E’ and E
complexes. Arrow indicates that SRp55 and Tra2 promote binding of U2AF.c│ Single nucleotides differences in the SMN2 gene compared to SMN1 create hnRNP A1 (or A2) binding sites in exon 7 and in the downstream intron in SMN2 pre-mRNA. hnRNP
A1/A2 may then inhibit formation or stabilization of the U2 snRNP complex, either directly or by blocking the activity of the downstream Tra2-dependent ESE. Note that it has also been suggested that the base change in exon 7 destroys an ASF/SF2-dependent
ESE163, 164 (although see ref. 69).
Figure 2│ Mechanisms of AS regulation at the transition from exon-definition to
intron-definition. a│ PTB inhibits c-src exon N1 inclusion by inhibiting U1 and U2
snRNP interactions and intron definition. In both WERI-1 and HeLa cells, the N1 exon is
36 38 defined by U1 snRNP binding to the 5’ssand U2 snRNP to the branch site (red dots). In
WERI-1 cells, in the absence of PTB, U1 and U2 snRNPs bound to the N1 exon interact
with the U2 and U1 snRNP on adjacent constitutive exons, respectively, thereby allowing efficient spliceosome assembly on introns flanking N1. In HeLa cells, PTB binds to sequences flanking N1 and prevents the cross-intron interactions that occur in WERI- cells, thereby excluding exon N1. b│ RBM5 regulates Fas AS by inhibiting inclusion of
exon 6. It does so not at the stage of U1 and U2 snRNP binding, but instead by promoting
tri-snRNP assembly on the intron-defined spliceosomal complex between exon 5 and
exon 7 while blocking tri-snRNP recruitment to complexes that would result in inclusion
of exon 6. Double-headed arrows indicate intron definition.
Figure 3│ Phosphorylation switches the general splicing repressor SRp38 into a
sequence-specific activator. a│ Phosphorylated SRp38 activates splicing by recruiting
U1 and U2 snRNPs to splice sites. SRp38 binds SRp38-dependent ESEs in target
transcripts and facilitates association of U1 and U2 snRNPs with the pre-mRNA to
stabilize 5’ss and branch site recognition by interacting with U1 and U2 snRNPs,
respectively. However, the spliceosomal A complex formed is stalled in S100, where an
SRp38-specific cofactor from NF40-60 is absent, which is required to proceed through
the splicing pathway. Reproduced with permission from REF.39.b│ SRp38 enhances
inclusion of the Flip exon of GluR-B pre-mRNA relative to the mutually exclusive Flop
exon. Both exons contain SRp38-binding sites (indicated by black bars under exon 14
(Flop) and exon 15 (Flip)), but the site in Flip is stronger (indicated by the thicker bar)
and Flip inclusion is therefore favored in the presence of SRp38. c│ Phosphatase PP1
37 39 dephosphorylates SRp38 upon heat shock. Under normal conditions, phosphorylated
SRp38 is associated with 14-3-3 proteins, which help to protect SRp38 from dephosphorylation, while PP1 activity toward SRp38 is inhibited by associated proteins, including NIPP1. During heat shock, PP1 dissociates from NIPP1 and directly binds to and dephosphorylates SRp38, which has dissociated from 14-3-3 proteins. Reproduced
with permission from REF.151.
38 40
Table 1. Ribonucleoproteins proteins that are involved in pre-mRNA splicing
Name Other domains Binding sequences Target genes names hnRNP A1 NA RRM, RGG UAGGGA/U SMN2, c-H-ras and G hnRNP NA RRM, RGG (UUAGGG)n HIV Tat, IKBKAP A2/B1 and G hnRNP AUF1 RRM U-rich β-amyloid receptor C1/C2 hnRNP F NA RRM, RGG GGGA, G-rich, PLP, c-SRC, Bcl-x and GY hnRNP G NA RRM and AAGU, SMN2, α-tropomyosin SRGY hnRNP DSEF-1 RRM, RGG, GGGA, G-rich PLP, HIV tat, Bcl-x H/H’ GYR and GY hnRNP I PTB RRM UCUU, CUCUCU PTB, nPTB, c-SRC, Fas, cTNT, CGRP, NMDA, hnRNP L NA RRM C and A rich eNOS, CD45 hnRNP LL SRRF RRM C and A rich CD45 hnRNP M NA RRM, GY ND FGFR2 hnRNP Q NA RRM, RGG CC(A/C) SMN2
Domains: RRM, RNA recognition motif; GY, glycineand tyrosine-rich domain; GYR, glycine-, tyrosine-, and arginine-rich domain; RGG, RGG box: arginineglycine-glycine repeats; SRGY, motif enriched in serine, arginine, glycine, and tyrosine.
Binding sequences: n, motif repetitions; ND, not determined.
39 41
TABLE 2. Tissue specific alternative splicing factors.
Name Other Binding Binding Tissue expression Target genes names domain motif nPTB brPTB RRM CUCUCU Neurons, c- Bin1, GlyR 2, PTBP2 myoblasts and PMCA1, Mef2, Nasp, testis Spag9, c-src Nova N/A KH YCAY Neurons of the GABA A2, GlyR2 1 hindbrain and and NOVA1 spinal cord Nova N/A KH YCAY Neurons of the GIRK, APLP2, 2 cortex, gephyrin, hippocampus and JNK2, neogenin, dorsal spinal cord NMDAR1, PLC4 Fox1 A2bp1 RRM (U)GCAUG Muscle, heart and -actinin, EWS, neurons FGFR2, fibronectin, SRC Fox2 RBM9 RRM (U)GCAUG Nuscle, heart, EWS, FGFR2, neurons and fibronectin, nonmuscle embryo myosin LC, SRC ESRP Rbm35a RRM GU-rich Epithelia cells FGFR2, CD44, p120- 1 Catenin, hMena ESRP Rbm35b RRM GU-rich Epithelia cells FGFR2, CD44, p120- 2 Catenin, hMena TIA-1 mTia1 RRM U-rich Brain, spleen, MYPT1, Fas, CGRP, testis FGFR2, TIAR, IL-8, VEGF; NF1; type II procollagen gene (COL2A1) TIAR Tial1, RRM U-rich Brain, spleen, TIA1, CGRP, TIAR; mTIAR lung, liver, testis NF1; Fas; calcitonin/CGRP Slm-2 Khdrbs3 KH UAAA Slm-2: brain, CD44, VEGF-A; , testis, heart T-STAR Quaki QK, KH ACUAAY[ Brain MAG, PLP; ng QKL …]UAAY HuB HuC, RRM AU-rich Neurons Calcitonin/CGRP HuD, elements Fas; NF1 ELAV family MBN N/A C3H1 YGCU(U/G Muscles, uterus, cTnT, Insulin receptor, L Znf )Y ovary Clcn1, Tnnt3
40 42 famil y CUG Bruno2 RRM U/G-rich Brain cTNT, Insulin BP1 Receptor ETR- CELF2, RRM U/G-rich Heart, skeletal cTnT, Tau, Cox-2 3 Brunol3 muscle, brain CELF Brunol4 RRM U/G-rich muscle Mtmr1, cTnT; 4 CELF Brunol5 RRM U/G-rich Heart, skeletal -actinin, cTNT, 5 NAPOR muscle and brain NMDAR1 CELF Brunol6 RRM U/G-rich Kidney, brain and cTnT 6 testis
Other names: N/A, not applicable; Domains: KH, RNA-binding domain; RRM, RNA recognition motif; RRMH; Z, Zinc finger; C3H1 Znf, CCCH zinc finger domain; Binding motif: […], spacer sequence of 1 to 20 nucleotides; n, motif repetitions; ND, not determined; R, purine; Y, pyrimidine
41 43 Glossary small nuclear ribonucleoprotein particle (snRNPs): U1, U2,U4, U5 and U6, which contain U-rich snRNAs and both snRNP specific and common proteins, are core components of the spliceosome.
SR protein family: The SR (Ser-Arg) proteins are a family of nuclear factors that play multiple important roles in splicing of mRNA precursors in metazoan organisms, functioning in both constitutive and alternative splicing. arginine/serine-rich (RS) domain: A protein domain that is variable in length and enriched in Ser-Arg dipeptides and appear to be involved in protein-protein and protein- RNA interaction.
RRM domain: A protein domain that is frequently involved in sequence-specific single- strand RNA binding. Also known as an RNP-type RNA binding domain. hnRNP protein: pre-mRNA/mRNA binding proteins that associate with transcripts during or after transcription and influence their function and fate. Some shuttle in and out of nuclei, while some are constitutively nuclear.
Hu proteins: ELAV/Hu proteins constitute a family of nervous system-specific RNA- binding proteins that specifically bind to AU-rich sequences. alternative exon: exons that are included in mature mRNA in certain cellular contexts but excluded in others.
CLIP: A method that combines crosslinking and immunoprecipitation to identify in vivo targets for RNA-binding proteins.
SELEX: Systematic Evolution of Ligands by Exponential Enrichment, is a technique to determine the DNA or RNA sequence that is specifically recognized by a protein ligand. The method involves multiple rounds of binding to an initially random sequenceuntil a high affinity, consensus sequence emerges.
14-3-3 proteins: A family of conserved phospho-serine/phospho-threonine binding proteins that are encoded by seven genes in most mammals. They bind diverse regulatory proteins, including kinases, phosphatases and transmembrane receptors. branch point: is a nucleotide, usually an adenosine, within a variably conserved ‘branch point sequence’ upstream of the 3’ splice site, the 2’ hydroxyl of which attacks the 5’ splice site in the first step of splicing.
42 BOX 1. Splicing and spliceosome assembly 44
Pre-mRNA splicing is a process in which intervening sequences (introns) are removed from an mRNA precursor. Splicing consists of two transesteri cation steps, each involving a nuceophilic attack on terminal phosphodiester bonds of the intron, by the 2’ hydroxyl of the branch-point (usually adenosine) in the rst step and by the 3’ hydroxyl of the upstream (5’) exon in the second step3, 4. This process is carried out in the spliceosome, a dynamic molecular machine the assembly of which involves sequential binding and release of snRNPs and numerous protein factors as well as the formation and disruption of RNA-RNA, protein-RNA and protein-protein interactions. The basic mechanics of spliceosome assembly are well known. Brie y, the process begins with the base pairing of U1 snRNA to the 5’-splice site (ss) and the binding of splicing factor 1 (SF1) to the branch point2 in an ATP- independent manner to form the E’ complex. The E’ complex can be chased into the E complex by the recruitment of U2 auxiliary factors heterdimer (U2AF, containing U2AF65 and U2AF35) to the polypyrimidine tract (py tract) and 3’ terminal AG130. The ATP-independent E complex is converted into the ATP-dependent pre-spliceosome A complex by the replacement of SF1 by U2 snRNP at the branch point. Further recruitment of the U4•U6/U5 tri-snRNP leads to the formation of B complex, which contains all spliceosomal subunits to carry out splicing. This is followed by an extensive conformational change and remodeling, including the loss of U1 and U4 snRNPs, resulting in the formation of C complex, the catalytically active splicesome.
5’ss 5’ss 3’ss 3’ss
branch site py tract
U1 U1 E’ complex SF1
exon de nition
U2AF U2AF E complex U1 SF1 U1 SF1 U1
? exon de nition intron de nition
U1 U2U2AF ? U2U2AF U1 A complex
intron de nition ?
tri-snRNP
U2U2AF B complex U1
Rearrangement to C complex and catalysis 45
BOX 2 SR proteins
The SR (Ser-Arg) proteins are a family of nuclear factors that play multiple important roles in splicing of mRNA precursors in metazoan organisms, functioning in both constitutive and alternative splicing7. They are involved in multiple steps of splicing regulation, by binding ESEs through their RRM domains, and mediating protein-protein41, 42, and perhaps protein-RNA159 interactions through their RS domains. All canonical SR proteins share common characteristics (see table). They display similar structures with one or two RNP-type RNA-binding domains at their N-termini and a variable-length domain enriched in Ser-Arg dipeptides at their C-termini (the RS domain). RS domains are extensively phosphorylated and they function in splicing, usually as activators. Most SR proteins function as pivotal regulators in multiple aspects of mRNA metabolism, such as mRNA nuclear export160, nonsense-mediated mRNA decay161 and translation162. A number of additional RS domain-containing proteins have been identi ed and those known to be involved in AS are listed in the table.
Name Domains Binding sequences Target genes Canonical SR proteins SRp20 RRM, RS GCUCCUCUUC SRp20, CT/CGRP, IR SC35 RRM, RS UGCUGUU AChE GluR1-4 ASF/SF2 RRM, RGAAGAAC HipK3, RRMH, RS CaMKIIδ, HIV RNAs; GluR1-4 SRp40 RRM, AGGAGAAGGGA HipK3, RRMH, RS PKCβ-II, Fibronectin SRp55 RRM, GGCAGCACCUG cTnT, CD44 RRMH, RS SRp75 RRM, GAAGGA FN1, E1A, RRMH, RS CD45 9G8 RRM, Z, RS (GAC)n Tau, GnRH, 9G8 SRp30c RRM, CUGGAUU Bcl-x, Tau, RRMH, RS hnRNP A1 SRp38 RRM, RS AAAGACAAA GluR-B, Tradin Additional SR proteins SRp54 RRM, RS ND Tau Domains: SRp46 RRM, RS ND N/A RNPS1 RRM, serine- ND Tra2-beta RRM, RNA recognition motif; rich RRMH; RRM homology; SRrp35 RRM, RS ND N/A RS, arginine-serine repeats-containing domain; SRrp86, RRM, RS ND N/A Z, Zinc finger; SRrp508 hTra2α RRM, RS×2 GAAARGARR DSX Binding sequences: n, motif repetitions; hTra2β RRM, RS×2 (GAA)n SMN1, ND, not determined; CD44, Tau N/A, not applicable RBM5 RRM, RS ND Fas CAPER RRM, RS ND VEGF R, purine a 46 tra2 SRm160/300 PTB
tra2 SR U2 proteins U1 TIA-1 U2AF ETR-3
ESEs ISEs PTB and ETR-3 5’ss 3’ss 5’ss 3’ss constitutive exon alternative exon binding site constitutive exon
SF1 b + Fox Fox U1 SF1 U1 calcitonin/CGRP ESEs ESEs exon 1 5’ss 3’ss 5’ss 3’ss exon 4 exon 1 exon 4 E’ complex
U2AF tra2
SRp55 Fox Fox U1 SF1 + U1 ESEs 5’ss ESEs exon 1 3’ss 5’ss exon 4 exon 1 3’ss exon 4
E complex U2AF SF1 tra2 c SRp55
SR SR SR SR SR proteins proteins proteins proteins proteins tra2 tra2 tra2 tra2
U2 U2 A1 hnRNP U2AF U2AF U1 U1 ESE ESS ESE ISSA1 hnRNP
inclusion of exon7 in SMN1 exclusion of exon7 in SMN2
Figure 1 47 a exon-de nition exon-de nition
u1 u2 u1 u2 u1 c-src 3 N1 4 5’ss branch site 5’ss WERI-1 and Hela cells
intron de nition intron de nition intron de nition intron de nition
u1 u1 u2 u1 u2 u1 PTB u2 u1 PTB u2 u1 nPTB 3 N1 nPTB 4 3 N1 4 5’ss 5’ss 5’ss 5’ss intron de nition
3 N1 4 3 4
WERI-1 cells Hela cells
b exon-de nition u1 u2 u1 u2 Fas 5 6 7
intron-de nition intron-de nition intron-de nition tri-snRNP tri-snRNP tri-snRNP
u1 u2 5 7 u1 u2 u2 u1 RBM5 5 6 tri-snRNP 7 tri-snRNP u2 u1 6
Figure 2 48
1 a U1 5’ 3’ 5’ 1 ESE 2 +S100 3’ 2 U2 ESE
U1 SRp38 5’ 1 1 U2 U1 5’ SRp38 A complex +SRp38 U2 ESE 2 ESE 2 3’ 3’
1 1 ? U1 Catalytically active +NF40-60 SRp38 U2 coactivator spliceosome ESE 2 3’
b
GluR-B 13 14 15 16
weak strong
WT DT40 cells coactivator SRp38-/- DT40 cells
SRp38 U1 U1 U2 U2 U2 U1 13 15 16 13 14 15 16 U2 U1 14
Flop 13 14 16 Flip 13 15 16 c
p 14-3-3 14-3-3 14-3-3 14-3-3 p p p SRp38 p Heat shock p M phase of cell cycle SRp38 p p SRp38 Kinases p
PP1 PP1 NIPP1 PP1
NIPP1
Figure 3 49
Chapter 2: Tumor metabolism: hnRNP proteins get in on the act
50 Tumor metabolism: hnRNP proteins get in on the act
Mo Chen1, Charles J. David1, and James L. Manley1
1Department of Biological Sciences, Columbia University, New York, NY 10027.
Correspondence to: James L. Manley1. Correspondence and requests for materials should
be addressed to J.L.M. ([email protected])
A long observed attribute of tumor cells, which sets them apart from most normal cells, is a high uptake of glucose. The glucose subsequently enters the glycolytic pathway and is converted to lactate, even in the presence of oxygen. This phenomenon, known as aerobic glycolysis1, is thought to allow growing cells to use glycolytic intermediates for biosynthetic processes critical for proliferation. The glycolytic phenotype of cells hinges on the fate of pyruvate, the intermediate converted to lactate in cancer cells, but enters the mitochondria to fuel oxidative phosphorylation in resting cells.
The enzyme that produces pyruvate, pyruvate kinase, has been implicated as a critical determinant of metabolic phenotype. In mammals, most tissues express the pyruvate kinase M (PKM) gene. PKM can produce two different isoforms by mutually exclusive inclusion of exon 9 or 10 during splicing of the mRNA precursor. The form expressed in most adult tissues, PKM1, contains exon 9, while the form expressed in growing (e.g., embryonic) cells, PKM2, contains exon 10. In tumor cells, or other cells induced to proliferate, reversion to PKM2 occurs, and is near universal and complete. Cantley and colleagues showed that when tumors are forced to express the PKM1 isoform rather than PKM2, aerobic glycolysis is reduced, oxidative phosphorylation is increased, and tumor growth is impaired, indicating that PKM isoform expression is a key determinant of how cells use glucose, and for proliferation of tumor cells2.
Given their critical and opposing roles in determining the cell’s glycolytic phenotype, the regulation of switching between PKM1 and PKM2 splicing is of great interest. To investigate this, David et al.3 set out to identify potential regulatory proteins. Alternative splicing is regulated by proteins that bind to specific RNA sequences, including members of the heterogeneous nuclear ribonucleoprotein (hnRNP) protein family, such as hnRNP A1, hnRNP A2 and PTB (also known as hnRNP I)4. Using UV crosslinking and RNA affinity chromatography, it was observed that these three proteins bind to intronic sequences flanking PKM exon 9 but not exon 10. Furthermore, the binding sequences are similar to their known consensus binding sites5, 6 and are conserved between mouse and human. To examine whether hnRNP A1, hnRNP A2 and PTB are functionally involved in PKM alternative splicing regulation, the three proteins were depleted by siRNAs in HeLa cells and other cancer cell lines. PKM1 mRNA levels were significantly increased and PKM2 mRNA levels were concomitantly decreased, indicating that hnRNP A1, hnRNP A2 and PTB expression levels are the critical 51 determinant of the expression of PKM2 isoform in transformed cells, likely by repressing exon 9 inclusion.
The mouse myoblast cell line C2C12 can be induced to differentiate into myotubes, a process that brings about switching from PKM2 to PKM1 expression, and David et al. observed a decrease in PTB and hnRNP A1 protein levels3. This result further supports the model that high expression levels of PTB and hnRNP A1 repress the inclusion of exon 9 to maintain the expression of PKM2 in proliferating cells. Because of the importance of PKM2 in cancer cell growth2, different classes of human glioma tumor samples were examined for a correlation between PTB, hnRNP A1 and hnRNP A2 expression levels and PKM splicing. Consistently, all tumor samples with high PKM2 expression, most notably the aggressive glioblastoma multiforme, were found to have elevated levels of PTB, hnRNP A1 and hnRNP A2.
Given the key role of PTB, hnRNP A1 and hnRNP A2 in promoting PKM2, and the tight coupling of PKM2 splicing and proliferation, David et al. next asked whether the expression of the three hnRNP proteins is under control of a proliferation-associated transcription factor. c- Myc has been shown to bind to PTB, hnRNP A1 and hnRNP A2 promoters and to upregulate their expression levels7-9. Significantly, c-Myc expression was found to correlate with PTB, hnRNP A1 and hnRNP A2 expression almost perfectly in human glioma tumor samples, and c- Myc directly stimulates transcription of hnRNP A13. Most importantly, siRNA-mediated reductions in c-Myc levels in NIH3T3 cells not only led to decreased mRNA and protein levels of PTB, hnRNP A1 and hnRNP A2, but also switched PKM splicing from PKM2 to PKM13. These results demonstrated a role of c-Myc in directly activating transcription of PTB, hnRNP A1 and hnRNP A2 genes and ensuring high levels of PKM2 in proliferating cells (Figure 1). As c-Myc is upregulated in numerous cancers10 , this provides a satisfying mechanism for how this important aspect of cell growth control is deregulated in cancer.
PTB, hnRNP A1 and hnRNP A2 have been observed to be upregulated in a wide variety of cancers11-13. Repression of PKM exon 9 inclusion and therefore the dominance of the PKM2 isoform in proliferating cells is at least one important functional consequence of this phenomenon. The results of David et al. indicate that overexpression of some combination of these three proteins in cancer is, like PKM2 expression, likely to be a general phenomenon. The fact that c-Myc depletion did not bring about PKM1 to PKM2 switching in all cell lines tested, including Hela cells3 suggests that additional transcription factors are capable of promoting PTB, hnRNP A1 and hnRNP A2 upregulation in some tumor cells. ChIP- seq data indicate that E2F family transcription factors as well as AP-1 proteins such as c-Fos bind to the promoters of the hnRNP A1, A2 and PTB genes, suggesting that multiple proliferation-associated pathways exist that bring about high expression of these proteins in a variety of cancers14. The intimate link between PKM alternative splicing, cell proliferation, and the RNA-binding proteins identified by David et al., casts the familiar hnRNP A1, A2 and PTB proteins in a new role: proliferation- associated splicing factors. 52
1. Warburg O. On the origin of cancer cells. Science 1956; 123:309‐14. 2. Christofk HR, Vander Heiden MG, Harris MH, Ramanathan A, Gerszten RE, Wei R, Fleming MD, Schreiber SL, Cantley LC. The M2 splice isoform of pyruvate kinase is important for cancer metabolism and tumour growth. Nature 2008; 452:230‐3. 3. David CJ, Chen M, Assanah M, Canoll P, Manley JL. HnRNP proteins controlled by c‐Myc deregulate pyruvate kinase mRNA splicing in cancer. Nature; 463:364‐8. 4. Chen M, Manley JL. Mechanisms of alternative splicing regulation: insights from molecular and genomics approaches. Nat Rev Mol Cell Biol 2009; 10:741‐54. 5. Burd CG, Dreyfuss G. RNA binding specificity of hnRNP A1: significance of hnRNP A1 high‐affinity binding sites in pre‐mRNA splicing. EMBO J 1994; 13:1197‐204. 6. Spellman R, Rideau A, Matlin A, Gooding C, Robinson F, McGlincy N, Grellscheid SN, Southby J, Wollerton M, Smith CW. Regulation of alternative splicing by PTB and associated factors. Biochem Soc Trans 2005; 33:457‐60. 7. Chen X, Xu H, Yuan P, Fang F, Huss M, Vega VB, Wong E, Orlov YL, Zhang W, Jiang J, Loh YH, Yeo HC, Yeo ZX, Narang V, Govindarajan KR, Leong B, Shahab A, Ruan Y, Bourque G, Sung WK, Clarke ND, Wei CL, Ng HH. Integration of external signaling pathways with the core transcriptional network in embryonic stem cells. Cell 2008; 133:1106‐17. 8. Shiio Y, Donohoe S, Yi EC, Goodlett DR, Aebersold R, Eisenman RN. Quantitative proteomic analysis of Myc oncoprotein function. EMBO J 2002; 21:5088‐96. 9. Schlosser I, Holzel M, Hoffmann R, Burtscher H, Kohlhuber F, Schuhmacher M, Chapman R, Weidle UH, Eick D. Dissection of transcriptional programmes in response to serum and c‐Myc in a human B‐cell line. Oncogene 2005; 24:520‐4. 10. Eilers M, Eisenman RN. Myc's broad reach. Genes Dev 2008; 22:2755‐66. 11. He X, Pool M, Darcy KM, Lim SB, Auersperg N, Coon JS, Beck WT. Knockdown of polypyrimidine tract‐binding protein suppresses ovarian tumor cell growth and invasiveness in vitro. Oncogene 2007; 26:4961‐8. 12. Zerbe LK, Pino I, Pio R, Cosper PF, Dwyer‐Nield LD, Meyer AM, Port JD, Montuenga LM, Malkinson AM. Relative amounts of antagonistic splicing factors, hnRNP A1 and ASF/SF2, change during neoplastic lung growth: implications for pre‐mRNA processing. Mol Carcinog 2004; 41:187‐96. 13. Hanamura A, Caceres JF, Mayeda A, Franza BR, Jr., Krainer AR. Regulated tissue‐specific expression of antagonistic pre‐mRNA splicing factors. RNA 1998; 4:430‐44. 14. Birney E, et al. Identification and analysis of functional elements in 1% of the human genome by the ENCODE pilot project. Nature 2007; 447:799‐816.
53
8 9 10 11
Proliferating cells Most adult tissues c-Myc
PTB,hnRNP A1 and hnRNP A2
8 9 11 8 10 11 PKM1 PKM2
pyruvate is tuned pyruvate goes into into lactic acid citric acid cycle ( also known as tricarboxylic acid cycle, TCA cycle) 54
Chapter 3: HnRNP proteins controlled by c-Myc deregulate pyruvate kinase mRNA splicing in cancer.
Mo Chen and Charles David contributed equally to this work
55
HnRNP proteins controlled by c-Myc deregulate pyruvate kinase mRNA splicing in cancer
Charles J. David1,*, Mo Chen1,*, Marcela Assanah2, Peter Canoll2, and James L. Manley1
1Department of Biological Sciences, Columbia University, New York, NY 10027. 2Department of
Pathology and Cell Biology, Columbia University, New York, NY,10032. *These authors contributed equally to this work.
Correspondence to: James L. Manley1. Correspondence and requests for materials should be addressed to J.L.M. ([email protected])
56 When oxygen is abundant, quiescent cells efficiently extract energy from glucose
primarily by oxidative phosphorylation, while under the same conditions tumor cells consume glucose more avidly, converting it to lactate. This long-observed phenomenon is
known as aerobic glycolysis1, and is now understood to be important for cell growth2, 3.
Because aerobic glycolysis is only useful to growing cells, it is tightly regulated in a proliferation-linked manner4, in part through control of pyruvate kinase (PK) isoform
expression. The embryonic isoform, PKM2, is almost universally re-expressed in cancer2, and promotes aerobic glycolysis, while the adult isoform, PKM1, promotes oxidative phosphorylation2. These two isoforms result from mutually exclusive alternative splicing of the PKM pre-mRNA, reflecting inclusion of either exon 9 (PKM1) or exon 10 (PKM2).
Here we show that three hnRNP proteins, PTB, hnRNPA1 and hnRNPA2, bind repressively to sequences flanking exon 9, resulting in exon 10 inclusion. We also demonstrate that the oncogenic transcription factor c-Myc upregulates transcription of
PTB/A1/A2, ensuring a high PKM2:PKM1 ratio. Establishing a relevance to cancer, we show that gliomas overexpress c-Myc and PTB/A1/A2 in a manner that correlates with
PKM2 expression. Our results thus define a pathway that regulates an alternative splicing event required for tumor cell proliferation.
Alternative splicing of PKM plays an important role in determining the metabolic phenotype of mammalian cells. The single exon difference imparts the enzymes produced with important functional distinctions. For example, PKM2, but not PKM1 is regulated by the binding of tyrosine phosphorylated peptides, which results in release of the allosteric activator fructose 1-6 bisphosphate and inhibition of PK activity5, a property that might allow growth-
factor initiated signaling cascades to channel glycolytic intermediates into biosynthetic processes.
1 57 The importance of tumor reversion to PKM2 was underscored by experiments in which
replacement of PKM2 with PKM1 in tumor cells resulted in markedly reduced growth2.
Consistent with a critical role in proliferation, re-expression of PKM2 in tumors is robust2, although little is known about the regulation of this process.
We set out to identify RNA binding proteins that might regulate PKM alternative splicing.
To this end, we prepared an [α-32P]-UTP labeled 250 nucleotide (nt) RNA spanning the E9 5’
splice site (EI9), previously identified as inhibitory to E9 inclusion6, as well as a labeled RNA
from a corresponding region of E10 (EI10) (Fig. 1b), and performed ultraviolet (UV)
crosslinking assays with HeLa nuclear extracts (NE)7. After separation by SDS-PAGE, multiple
proteins from 35-40 kDa appeared using the EI9 substrate, while little binding was observed
using the EI10 substrate (Fig. 1b). Strong binding was mapped to a 19 nt region we named
EI9(50-68) that spans the E9 5’ splice site (Supplementary Fig. 1). To identify the bound
proteins, we performed RNA affinity chromatography using a 5’ biotin-labeled RNA
corresponding to EI9(50-68). After SDS-PAGE and Coomassie staining, the pattern of
specifically bound proteins closely matched that observed after UV crosslinking (Fig. 1c). The
four indicated proteins between 35-40 kDa were excised, and identified by mass spectrometry as
isoforms of hnRNPA1 and hnRNPA2, RNA binding proteins with well established roles as
sequence-specific repressors of splicing (e.g., refs. 7, 8). This result was confirmed by
immunoblotting with antibodies against hnRNPA1 (Supplementary Fig. 2).
The sequence immediately downstream of the E9 5’ splice site contains a UAGGGC
sequence that is highly related to the consensus hnRNPA1 high affinity binding site identified by
9 SELEX, UAGGG(A/U) (Fig. 1d). Consistent with previous mutational studies of an identical
2 58 A1 binding site8, mutation of the G3 nucleotide of this motif to C led to a large decrease in hnRNPA1/A2 binding (Fig. 1d; Supplementary Fig. 3). The G3C mutation resulted in increased splicing in vitro when introduced into a splicing substrate containing E9 (Supplementary Fig. 4), and led to increased E9 inclusion in a minigene construct in vivo (Supplementary Fig. 5). These data confirm the presence of an inhibitory hnRNPA1/A2 binding site immediately downstream of the E9 5’ splice site.
To explore the possibility that other splicing regulators bind upstream of E9 or E10, we constructed crosslinking substrates (48 nt) that span the region upstream of each exon. Using these RNAs for UV crosslinking showed strong binding of a 55 kDa protein to the I8 RNA probe, but not to the I9 probe (Fig. 1e). Inspection of the polypyrimidine tract upstream of E9 revealed two potential PTB (polypyrimidine tract binding protein, or hnRNPI) binding sequences
10 (UCUUC) within 35 nucleotides of the intron/exon boundary, while no such sequence exists in the E10 polypyrimidine tract. PTB frequently functions as a splicing repressor10, often by binding repressively to the polypyrimidine tract11. Immunoprecipitation confirmed that the 55 kDa crosslink observed using I8 RNA is PTB (Fig. 1e), and we observed strong binding of PTB to a biotinylated version of I8 (Supplementary Fig. 6). In addition, mutation of the two putative
PTB binding sites from UCUUC to UGUUC significantly diminished binding (Fig. 1f;
Supplementary Fig. 7). Our data indicate that the splicing repressor PTB binds specifically to the polypyrimidine tract of E9.
Because the locations of hnRNPA1/A2 and PTB binding sites flanking E9 overlap elements critical to exon inclusion (the polypyrimidine tract for PTB11, the site of U1snRNA-pre- mRNA base-pairing for A1/A212), we speculated that these proteins are inhibitors of E9
3 59 inclusion. To examine this possibility, we used siRNA to deplete hnRNPA1, hnRNPA2 and/or
PTB from HeLa cells. We assayed the PKM mRNA isoform ratio using RT-PCR followed by
exon-specific restriction digestion (Fig. 2a). Knockdown of hnRNPA1 or hnRNPA2 in HeLa
cells resulted in little change in splicing pattern (Supplementary Fig. 8). Because we have previously observed functional redundancy of hnRNPA1/A27, we next simultaneously depleted both proteins (Fig. 2b). This resulted in an increase in PKM1 mRNA, from 2% to 29%, and a concomitant decrease in PKM2 mRNA (Fig. 2c; Supplementary Fig. 8). PTB knockdown also increased the PKM1 isoform, to 16% (Fig. 2c; Supplementary Fig. 8), consistent with earlier
observations13. Next, we simultaneously depleted all three factors, which further increased
PKM1 levels, to about 48% (Fig. 2c). Similar results were obtained using 293 cells, with the
triple knockdown resulting in an increase from 5% to 67% PKM1 (Fig. 2d). Increases in PKM1
mRNA upon A1/A2/PTB knockdown were observed in all cell lines tested, including the breast
cancer cell line MCF-7 and the glioblastoma cell line U87 (Supplementary Fig. 9). Knockdown of two other cancer-associated splicing factors in HeLa cells, the SR proteins ASF/SF2 and
SRp20, while also resulting in slowed growth, failed to significantly affect PKM1/2 ratios
(Supplementary Fig. 8 and 10), indicating that the effects seen in PTB/A1/A2 depleted cells on
PKM splicing are specific and not the result of pleiotropic effects due to changes in cell growth.
Together, our results indicate that PTB/A1/A2 expression is the critical determinant of PKM isoform in transformed cells.
We next wished to determine whether PTB/A1/A2 expression levels and PKM1/2 alternative splicing are correlated. We first examined whether changes in PTB/A1/A2 levels correlate with changes in PKM splicing during switching from growth to quiescence. To this end, we used the mouse myoblast cell line C2C12, which, when grown to confluence and then
4 60 switched to low-serum medium, undergoes myogenic differentiation, a process that includes
PKM2 to PKM1 switching14. We differentiated C2C12 cells for 6 days, and used RT-PCR
followed by restriction digestion to assess the PKM1/2 ratio each day. We observed a large
increase in PKM1 and a corresponding decrease in PKM2 mRNA during differentiation (Fig. 3a).
We then prepared lysates of C2C12 cells at time points throughout differentiation and examined
protein levels by immunoblotting (Fig. 3b). PTB expression dropped over 70% by day 3 of
differentiation, after which it remained stable, consistent with previous studies15. We also
observed an approximately 50% decrease in hnRNPA1 levels by day 3 of differentiation, though no significant changes were observed in the level of hnRNPA2. This result is consistent with a
role for PTB/A1 in maintaining high PKM2 levels in proliferating C2C12 cells.
Because of the importance of the PKM2 isoform to the growth of cancer cells, we next
examined human glioma tumor samples for a correlation between PTB/A1/A2 expression and
PKM splicing. We first assayed PKM1/2 mRNA levels as described earlier. Normal brain tissue
ranged from 4-13% PKM2, pilocytic astrocytomas (PA) samples expressed approximately 66-
77% PKM2, low grade astrocytomas (LGA) ranged from 7-73%, and glioblastoma multiforme
(GBM) samples expressed 72-86% PKM2 (Fig. 3c). To explore a potential correlation between
elevated PKM2 mRNA levels and expression of the regulatory proteins we identified, we
performed immunoblots for PTB, hnRNPA1 and hnRNPA2. Significantly, all high-PKM2
tumors expressed elevated levels of PTB/A1/A2, with the most striking overexpression in GBMs
(Fig. 3d). Consistent with their uniformly high PKM2 expression, all four PA samples also showed overexpression of the PTB/A1/A2. In LGAs the two high PKM2 tumors showed elevated expression of the three proteins, while the two low PKM2 tumors showed expression levels similar to normal brain. Immunoblotting for four other splicing factors (ASF/SF2, Tra2β,
5 61 TLS/FUS, and hnRNPK) revealed no correlation with PKM2 expression (Supplementary Fig.
11), indicating that the correlation between an elevated PKM2/1 mRNA ratio and overexpression of PTB/ A1/ A2 is specific and not reflective of a general property of splicing factors.
The tight coupling of PKM2 expression to proliferation suggests that the expression of the PKM splicing regulatory proteins we identified might be under the control of a proliferation- associated regulatory mechanism. A strong candidate to control this is the oncogenic transcription factor c-Myc, which, like PTB/A1/A2, is upregulated in GBMs16, and has been shown to bind the PTB/A1/A2 promoters17, 18 and upregulate the expression of all three19, 20.
Consistent with a role for c-Myc in PTB/A1/A2 regulation, we observed a near perfect correlation between the levels of c-Myc and PTB/A1/A2 in gliomas and differentiating C2C12 cells (Fig. 3b and 3d). In addition, the transcription factor N-myc, which is closely related to c-
Myc21, was upregulated in PAs and to a lesser extent in GBMs (Supplementary Fig. 11),
indicating that this protein may in some cases contribute to PTB/A1/A2 upregulation.
We next examined directly c-Myc’s involvement in PTB/A1/A2 expression and PKM
splicing regulation. We first asked whether decreasing c-Myc levels can affect PTB/A1/A2
levels and the PKM1/PKM2 mRNA ratio. To this end, we transfected NIH-3T3 cells with
vectors bearing a puromycin resistance marker that express either a c-Myc-targeting shRNA or a
control shRNA. Immunoblotting showed a reduction in c-Myc levels in cells stably transfected
with c-Myc shRNA, compared to control cells (Fig. 4a). PTB/A1/A2 protein levels were also
significantly reduced after depletion of c-Myc, in contrast with two other RNA processing
factors not implicated in PKM splicing regulation, ASF/SF2 and CPSF73 (Fig. 4a). PTB/A1/A2
mRNA levels were also significantly reduced in the knockdown cells (Fig. 4b), supporting the
6 62 idea that c-Myc regulates transcription of these genes. Importantly, the cells stably expressing the c-Myc shRNA showed a pronounced increase in the PKM1/2 ratio, expressing 33% PKM1 mRNA compared to 7% in the control (Fig. 4c). A separate line stably expressing a second c-
Myc shRNA revealed a similarly elevated PKM1/2 ratio, as well as reduced levels of
PTB/A1/A2, showing that the observed effects were not due to off-target effects of the c-Myc shRNA (Supplementary Fig. 12). Additionally, we co-transfected an hnRNPA1 promoter- luciferase construct with a c-Myc expression vector22, which resulted in a dose- and c-Myc binding site- dependent increase in promoter activity (Supplementary Fig. 13).
The above results demonstrate a direct role for c-Myc in maintaining high PTB/A1/A2 levels in NIH-3T3 cells. In contrast, c-Myc knockdown in HeLa cells revealed only a small decrease in PTB/A1/A2 levels, and no change in the PKM1/2 ratio (Supplementary Fig. 14), suggesting that factors other than c-Myc might promote PTB/A1/A2 expression in these cells.
One possibility is the transcription factor E2F1, which like c-Myc binds upstream of all three genes18. However, knockdown of E2F1, or of Rb, a negative regulator of E2F family transcription factors23, resulted in little change in PTB/A1/A2 levels (unpublished data).
However, since the E2F and Rb families exhibit redundancy, this result does not rule out involvement of the E2F/Rb pathway in PTB/A1/A2 regulation. Indeed, because of their importance to proliferating cells, it is likely that PTB/A1/A2 can be upregulated by proliferation- associated factors in addition to c-Myc.
The fact that PTB/A1/A2 depletion results in switching to the PKM1 isoform suggests that RNA binding proteins can control the outcome of a mutually exclusive splicing event by simultaneously acting as repressors of one exon (E9) and activators of the other (E10)
7 63 (Fig. 4d). While it is easy to envision how these proteins exclude E9, how might PTB/A1/A2
promote E10 inclusion? A variety of RNA binding proteins, including hnRNPA1/A2, have been
shown to stimulate splicing of an adjacent exon through intronic binding sites24. One proposed
mechanism for this is intron definition, in which intron-binding proteins induce intronic
structures conducive to inclusion of the neighboring exon24. We propose that, like many
alternatively spliced exons, PKM E10 is poorly recognized by the splicing machinery in the
absence of adjacent intron definition, and such a structure is promoted by PTB/A1/A2 binding
(Fig. 4d).
We have demonstrated a critical functional consequence for observations connecting
PTB/A1/A2 upregulation with cell proliferation25, 26, transformation27, 28, and a wide variety of
cancers (e.g., refs. 26, 27, 29, 30). Given the critical role of these proteins in promoting PKM2
production in tumors, overexpression of some combination of them is, like PKM2 expression,
likely to be a general phenomenon in cancer. The fact that the proteins show some redundancy in
promoting PKM2 splicing may ensure robust re-expression of PKM2 in tumors.
Methods summary
UV crosslinking substrates were cloned into pcDNA3 vector (Invitrogen) and UV
crosslinking was performed as previously described7. Mutations were introduced in EI9 by PCR-
based site-directed mutagenesis. Biotinylated RNAs for affinity purification were purchased
from Dharmacon, and RNA affinity chromatography was carried out as described7.
Immunoprecipitations were carried out using protein A-agarose beads (Roche). RNAi was performed as described7. We transfected 50 pmol of hnRNPA1 siRNA and 25 pmol of other
siRNA duplex in a 24-well plate. After 72 hours, we collected cells for RNA isolation and
8 64 immunoblotting. C2C12 cells were grown in DMEM (Invitrogen) supplemented with 20% fetal bovine serum (FBS) (Hyclone) at 37°C in 5% CO2. For differentiation treatment, C2C12 were plated on gelatin coated plates, allowed to reach confluence, and then switched to DMEM 2% donor equine serum (Hyclone). Human brain and glioma samples were obtained from the Bartoli
Brain Tumor Bank at the Columbia University Medical Center. Samples were homogenized and used for Trizol RNA extraction and western blotting as described30. In all cases, immunoblots were scanned and quantified using the LI-COR Odyssey system. c-Myc shRNA DNA sequences were purchased from Invitrogen and cloned into the pRS vector (Origene). shRNA constructs were transfected into NIH3T3 cells and stable cell lines were selected with puromycin for RNA isolation and immunoblotting. PKM1/PKM2 ratio was analyzed by extracting total RNA from cells and tissue samples and preforming by RT-PCR followed by PstI, Tth111I, or EcoNI digestion. qPCR for PTB/A1/A2 in control and c-Myc knockdown cells was performed with
SYBR green from Fermentas using the Applied Biosystems 7300 real-time PCR system. hnRNPA1 promoter sequence for dual luciferease reporter (DLR) assay was cloned into PGL3- enhancer vector (Promega) and DLR assays were performed using Dual Luciferase Reporter
Assay System (Promega)
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Acknowledgements We thank D. Black for BB7 antibody, T. Kashima for hnRNP A1/A2
siRNA, R. Prywes and E. Henckles for C2C12 cells, M. Sheetz and X. Zhang for NIH-3T3 cells,
12 68 C. Prives, T. Barsotti, and L. Biderman for MCF-7 cells, Rb, and E2F-1 siRNA, R. Dalla-Favera and Q. Shen for anti-c-Myc antibodies and c-Myc expression vector, R. Eisenmann for N-myc antibody, and members of the Manley laboratory for discussions. This work was supported by grants from the Avon Foundation and the NIH.
Author contributions C.J.D., M.C., and J.L.M. conceived the project and designed experiments,
C.J.D. and M.C. carried out experiments, P.C. and M.A. provided tumor samples, C.J.D., M.C. and J.L.M. interpreted data and wrote the paper.
Figure legends
Figure 1: hnRNP proteins bind specifically to sequences flanking E9. a, Schematic diagram of PKM splicing. b, Position of probes spanning the E9 or E10 5’ splice sites (top). After UV crosslinking, proteins were detected by autoradiography (bottom). c, Affinity chromatography using EI9(50-68). Bound proteins were separated by SDS-PAGE and Coomassie stained. Bands excised for mass spectrometry are indicated. d, Sequence of EI9(50-68), putative hnRNPA1/A2 binding site indicated in bold italics (top). UV crosslinking with wild-type RNA, or RNA with a mutation in the putative hnRNPA1/A2 binding site (bottom). e, Position of I8 and I9 (top). UV crosslinking using I8 or I9 substrates (bottom left). UV crosslinking reactions were IPed with either α-PTB (BB7) or α-HA antibodies (bottom right). f, UV crosslinking with I8 and mutant derivative I8mu, sequences indicated above. Putative PTB binding sites in I8 are underlined.
Figure 2: PTB, hnRNPA1, and hnRNPA2 are required for high PKM2:PKM1 mRNA ratios. a, Scheme for assaying PKM1/PKM2 ratios in human cells. b, Immunoblots showing protein levels after the indicated siRNA treatment. Protein bands were quantified after LI-COR
13 69 Odyssey scanning and normalized to GAPDH. c, The indicated splicing factors were depleted by siRNA, followed by PKM splicing assay outlined in (a). Products corresponding to M1 and M2 are indicated with arrows. The PKM1 percentage is indicated below. d, PKM1/2 levels assayed after the indicated siRNA treatment in 293 cells.
Figure 3: Expression of PTB/A1/A2 and c-Myc correlates with PKM2 expression in C2C12 cells and tumors. a, PKM splicing assay after the indicated number of days of C2C12
differentiation. b, Immunoblots for the indicated proteins were performed throughout
differentiation, and normalized to GAPDH (Day 0 = 1). c, RNA was extracted from brain tissue or tumor samples and assayed for PKM mRNA isoforms. d, Lysates were immunoblotted for
PTB, hnRNPA1, hnRNPA2 or c-Myc and normalized to Actin. Sample order is the same for
RT-PCR and immunoblotting.
Figure 4: c-Myc upregulates PTB/A1/A2 alters PKM splicing. a, Immunoblotting using
NIH-3T3 cells stably expressing control or c-Myc-targeting shRNAs. Signals were quantified and normalized to actin. b, RT-PCR using the same cell lines as in (a). Realtime RT-PCR was performed separately to quantify the relative levels of PTB/A1/A2 mRNAs in control and c-Myc
knockdown cells, using RPL13A as a reference gene. Relative levels of each are shown below
each panel, with s.d. indicated (n=3). c, PKM1/2 ratios in control and c-Myc knockdown cells
determined as in Fig. 2a., d, A model for PKM splicing regulation. Top, in adult tissues, low
expression of PTB/A1/A2 allows for recognition of E9 by the splicing machinery and disrupts intronic structures favorable for E10 inclusion. Bottom, in embryonic and cancer cells,
PTB/A1/A2 are upregulated, bind to splicing signals flanking E9 and repress its inclusion.
14 70 Binding of these proteins around E9 and possibly to other sites creates an intronic structure
favorable to E10 inclusion.
Methods
Plasmid constructs. Long UV crosslinking substrates (EI9, EI10) were prepared by amplifying
fragments from HeLa genomic DNA using Pfu turbo (Stratagene), and cloning the products into
pcDNA3 (Invitrogen). EI9(1-20), EI9(21-49), and EI9(50-68, I8, I8mu, and I9 DNA sequences
were ordered from Invitrogen and cloned into pcDNA3. Primers used to amplify genomic DNA fragments were: EI9 forward, CGC GGA TCC TTC TTA TAA GTG TTT AGC AGC AGC T , reverse, CGG AAT TCA CTG AGC CAC AGG ACC CTT TG ; EI10 forward, CGC GGA TCC
CTC CTT CAA GTG CTG CAG TG , reverse, CGG AAT CCT GGG CCC AGG GAA GGG G;
I8E9 forward, CCC AAG CTT AAA TTC CCC ATT CTG TCT TCC CAT G , reverse, CGG
GAT CCC TGC CAG ACT CCG TCA GAA CT; I9E10 forward, CCC AAG CTT CTG TCC
GGT GAC TCT TCC CC , reverse, CGG GAT CCC TGC CAG ACT TGG TGA GGA CG.
Mutations were introduced in EI9 by PCR-based site-directed mutagenesis. Mouse c-Myc and control shRNA DNA sequences were ordered from Invitrogen and cloned into pRS vector
(Origene) with BamH I and Hind III. The hnRNPA1 promoter region, either wild-type or the E box mutant, was cloned into PGL3-enhancer vector (Promega).
Antibodies. The following antibodies were used in this study: BB7 for human PTB IP (gift from
Dr. Douglas Black, UCLA), 3H8 for mouse/human PTB immunoblots (Sigma), MC3 for
15 71 U2AF65 (Sigma), α-HA (Covance) DP3D3 for hnRNPA2 (Abcam), N-262 for c-Myc (Santa
Cruz), α-Actin (Sigma), α-GAPDH (Sigma), 9H10 for hnRNPA1 (Sigma), mAb104 for SRp20.
UV crosslinking, RNA affinity purification, and immunoprecipitation assays. We carried out
ultraviolet crosslinking as previously described7. Briefly, we linearized the UV crosslinking
plasmids with an appropriate restriction enzyme and synthesized the RNAs with [32p]-UTP or
[32p]-CTP. We incubated 1X105 c.p.m. RNAs with 10 g HeLa or C2C12 NE in buffer D in a 20
l reaction at 30°C for 15 minutes, then irradiated the samples with ultraviolet light in a
Stratalinker 1800 (Stratagene), digested them with RNase A(10µg/ml) and resolved them by
SDS-PAGE. The RNA affinity pull-down experiment and immunoprecipitation was preformed as described7. The 5’biotinylated EI9(50-68) and I8 RNA oligonucleotides were purchased from
Dharmacon. Antibodies were bound to protein A-agarose beads prior to IP. We used the following antibodies for IP: BB7 for PTB, and MC3 for U2AF65.
In vitro and in vivo splicing assays. Minigene containing PKM gene exon 8, exon 9, exon 10 , exon 11 and flanking regions was cloned into pcDNA3 vector (Invitrogen). G to C mutation was introduced in Minigene by PCR-based site-directed mutagenesis11. Wildtype and mutated
minigene vectors were transfected into HeLa cells. 24 hours after transfection, cells were
collected and PKM1/M2 ratio was analyzed using RT-PCR followed by Pst I digestion. In vitro
splicing substrates were constructed by replacing the first exon and downstream intronic
sequence of AdML pre-mRNA with PKM exon 9 and downstream intron 9 sequences or
sequence with mutated hnRNP A1 binding site. pre-mRNA substrates were synthesized by in
vitro transcription using T7 RNA polymerase (Promega) following product protocol. In vitro
16 72 splicing of the wildtype and mutated pre-mRNA was carried out using HeLa nuclear extract as
described31.
RNA interference. We carried out RNA interference of PTB and hnRNPA1/A2 as described7.
Briefly, we plated HeLa, 293, MCF-7, or U87 cells at 2.5-3x104 cells per well in 24-well plates.
The next day, we mixed 50 pmole of hnRNPA1 duplex RNA and 25 pmole of the other duplex
RNAs with 1.5 l lipofectamine 2000 transfection reagent (Invitrogen) plus 100 l of Opt-MEM
medium and added this to cells after RNA duplex-lipid complex formation. For double and triple
knockdowns in HeLa and 293 cells, RNA duplexes were transfected simultaneously. The control
RNA duplex was used to ensure that parallel experiments had equal amounts of RNA. In MCF-7
and U87 cells, the second and third RNA duplexes were transfected 6 hours after the previous
transfection. 72 hours after transfection, we collected cells for RNA isolation and
immunoblotting. We used the following siRNAs (Dharmacon; the sense strand sequences are
given): human hnRNPA1, CAGCUGAGGAAGCUCUUCA; human hnRNPA2,
GGAACAGUUCCGUAAGCUC; human PTB, GCCUCAACGUCAAGUACAA. ASF/SF2
depletion was performed as previously described7.
c-Myc shRNA stable cell lines. Stable cell lines expressing c-Myc shRNAs or control shRNA
were obatined by transfecting pRS-shRNA vectors into NIH3T3 cells followed by drug selection.
Cells were plated in 10cm plates. The next day, transfected cells were diluted and medium was
replaced with medium containing a final concentration 3 g/ml puromycin. After 7-10 days, a
mixture of fast- and slow-growing colonies appeared in cells transfected with c-Myc shRNA,
while only fast-growing colonies appeared in cells transfected with control shRNA. Single slow-
growing colonies were isolated and cultured for c-Myc expressing cells. c-Myc expression was
17 73 examined by immunoblotting. Positive colonies were collected for RT-PCR and western blotting.
The following sense shRNA sequences were used:
control, gat ccG AGG CTT CTT ATA AGT GTT TAC TCG AGT AAA CAC TTA TAA GAA
GCC TCT TTT Ta ; Mouse c-Myc shRNA1, gat ccC ATC CTA TGT TGC GGT CGC TAC
TCG AGT AGC GAC CGC AAC ATA GGA TGT TTT Ta; Mouse c-Myc shRNA2, gat ccC
GGA CAC ACA ACG TCT TGG AAC TCG AGT TCC AAG ACG TTG TGT GTC CGT TTT
Ta; human c-Myc shRNA, gat ccC CAT AAT GTA AAC TGC CTC AAC TCG AGT TGA
GGC AGT TTA CAT TAT GGT TTT Ta.
RT-PCR. Total RNA was extracted from tissue culture and human brain tumor samples using
Trizol (Invitrogen) according to the manufacturer’s instructions. Total RNA (2.5–5 μg) was used
for each sample in a 20 μl reaction with 0.5μL of SuperScript III RT (Invitrogen). 1μl of the
cDNA library was used in a 50 μl PCR reaction containing 3 μCi [32p]-dCTP. 10μl of the PCR
products were digested by Pst I and Tth111 I (human PKM) or EcoN I (mouse PKM) and the
products were resolved by 6% non-denaturing PAGE. Primers used in the PCR reactions were:
human PKM exon8 forward, CTG AAG GCA GTG ATG TGG CC; human PKM exon11 reverse, ACC CGG AGG TCC ACG TCC TC; mouse PKM exon 8 forward, CAA GGG GAC
TAC CCT CTG G; mouse PKM exon11 reverse, ACA CGA AGG TCG ACA TCC TC, human
µglobulin: forward, GGC TAT CCA GCG TAC TCC AAA, reverse, CGG CAG GCA TAC
TCA TCT TTT T; mouse µglobulin: forward, TTC TGG TGC TTG TCT CAC TGA, reverse,
CAG TAT GTT CGG CTT CCC ATT C. qRT-PCR was performed using the following primers:
mouse hnRNPA1: forward, TGG AAG CAA TTT TGG AGG TGG, reverse, GGT TCC GTG
GTT TAG CAA AGT; mouse hnRNPA2: forward, AAG AAA TGC AGG AAG TCC AAA GT,
18 74 reverse, CTC CTC CAT AAC CAG GGC TAC; mouse PTB: forward, AGC AGA GAC TAC
ACT CGA CCT, reverse, GCT CCT GCA TAC GGA GAG G; mouse RPL13A forward, GGG
CAG GTT CTG GTA TTG GAT, reverse, GGC TCG GAA ATG GTA GGG G. Relative
amounts of mRNA were calculated using the comparative Ct method.
Cell culture and differentiation. C2C12 cells were grown in DMEM (Invitrogen) supplemented
with 20% fetal bovine serum (FBS) (Hyclone) at 37°C in 5% CO2. For differentiation treatment,
C2C12 were plated on gelatin coated plates, allowed to reach confluence, and then switched to
DMEM 2% donor equine serum (Hyclone). HeLa and 293 cells were grown in DMEM, 10%
FBS. NIH3T3 cells were grown in DMEM, 10% bovine calf serum (BCS) (Hyclone).
Human brain tumor samples. De-identified brain and glioma samples were obtained from the
Bartoli Brain Tumor Bank at the Columbia University Medical Center. Non-cancerous samples removed from epileptic patients were used for normal brain. Approximately 25-200 mg of each sample was obtained. Half of the homogenate was used for Trizol RNA extraction, the other half of each sample was processed for immunoblotting as described30.
Dual Luciferase Reporter (DLR) Assay. c-Myc expression vector and hnRNP A1 promoter vector were co-transfected into HeLa cells. 24 hours after transfection, cells were collected and
DLR assays were preformed using Dual Luciferase Reporter Assay System (Promega) following product protocol.
31 Krainer, A.R., Maniatis, T., Ruskin, B., & Green, M.R., Normal and mutant human beta- globin pre-mRNAs are faithfully and efficiently spliced in vitro. Cell 36, 993-1005 (1984)
19 75 a b 9 1100 c
EI9 EI10 EI9(50-68)No RNA
8 9 10 11
EI9 EI10
75
8 8 11 9 11 10 50 50 PKM1 PKM2 37 Adult Embryo/Tumor 37 25 hnRNP A1/A2 isoforms
25 UV crosslinking
d e f
I9: aaagctctgtccccctctcgtccctctggacggatgttgctccccctag
EI9(50-68) sequence: 9 1100 I8: tcctcctccttccctcttccttgccccctcttcccctaaaccttacag Exon Intron 1 2 3 4 I8 I9 I8mu: tcctcctccttccctGttccttgccccctGttcccctaaaccttacag aggtagggccctaagggca IP
I8mu Supernatant I8 I9 I8 WT
G3C
BB7
BB7 Mock I8 I9 Mock
PTB
PTB hnRNPA1/A2
Figure 1 76 a b
PTB
A1/A2
PTB/A1/A2 siRNA mock Human PTB Tth111 I PTB/GAPDH 100 4.3 58.1 8.5 PKM1 8 9 11
hnRNP A1 PKM 2 8 10 11
hnRNP A1/GAPDH 100 72.0 17.9 26.2 Pst I
hnRNP A2
hnRNP A2/GAPDH 100 89.0 10.6 16.3
GAPDH
Immunoblotting c d
siRNA
PTB mock PTB/A1/A2
PTB/A1/A2
mock A1/A2
M1 M1
M2 M2
M1% 1.8 16.4 29.2 47.8 M1% 5 67
cut by [ Pst I ] cut by [ Pst I ]
RT-PCR RT-PCR
Figure 2 77 a b Day Day 6 Day Day 0 Day 1 Day 3 Day 5 Day Day 4
PTB Days of 0 1 2 3 4 5 6 di erentiation PTB/GAPDH 1.00 0.86 0.27 0.29 0.21 0.14 M1 hnRNP A1
A1/GAPDH 1.00 0.87 0.51 0.56 0.53 0.40
hnRNP A2
M2 1 2 3 4 5 6 7 A2/GAPDH 1.00 1.00 0.73 1.13 0.97 0.84
c-Myc %M1 9.5 14.4 24.7 41.1 54.9 55.0 79.7 cut by [ Pst I ] c-Myc/GAPDH 1.00 0.56 0. 43 0.49 0.59 0.58
GAPDH RT-PCR
Immunoblotting c Pilocytic Normal Brain Low grade Glioblastoma astrocytomas astrocytomas mutiforme
M1
M2
M1% 95.2 96.1 89.0 87.5 32.8 33.9 22.6 34.4 92.9 44.3 83.8 27.5 18.9 19.3 13.7 28.0 cut by [ Pst I ] RT-PCR
Pilocytic Low grade Normal Brain Glioblastoma d astrocytomas astrocytomas mutiforme
PTB
PTB/Actin 1.0 0.5 0.3 1.0 18.7 8.7 12.3 18.2 0.9 10.0 0.9 6.5 58.7 37.9 21.4 12.2
hnRNP A1 1 0.83 0.15 0.46 1.82 5.40 4.70 2.69 0.69 1.29 0.34 1.59 2.10 4.39 2.16 2.02 3.41 A1/Actin 1.0 1.1 0.8 1.9 12.5 6.3 5.9 4.1 2.4 15.2 0.8 8.9 21.6 14.1 13.5 10.2
hnRNP A2 1 0.80 0.02 0.95 2.95 4.61 2.86 1.30 0.43 1.99 0.79 2.15 9.00 21.95 17.0 8.88 13.24 A2/Actin 1.0 1.2 0.4 1.5 3.0 1.1 2.0 2.5 0.5 4.0 0.2 1.9 4.8 5.1 3.3 2.1
c-Myc
c-Myc/Actin 1 1.2 1.2 0.5 4.1 4.8 3.0 4.1 0.2 1.6 2.2 1.7 4.1 6.8 3.6 2.0
Actin
Immunoblotting Figure 3 78
c a b shRNA shRNA
c-Myc1
control shRNA c-Myc1 control
c-Myc1 control c-Myc M1 c-Myc c-Myc/Actin 100 6.3
PTB PTB PTB/Actin 100 42.7 100 29.3+/-3.1 M2 hnRNP A1 hnRNP A1 hnRNP A1/Actin 100 20.3 M1 % 7.0 32.7 cut by [ Pst I ] hnRNP A2 100 48.7+/-3.1 RT-PCR hnRNP A2/Actin 100 2.4 hnRNP A2 ASF/SF2 d 100 6.7+/-1.2 ASF/SF2/Actin 100 76.8 intron de nition intron de nition CPSF73 mglobulin U1 U2 U1 U2 Adult 11 8 9
CPSF73/Actin 100 90.5 10 RT-PCR Actin
Proliferating cells intron de nition intron de nition
A1 PTB A2 U1 U2 U1 U2 Immunoblotting 8 10 11
A1/A2 c-MycMyc
9 PTB
Figure 4 79
9
1-20
21-49
EcoN I 50-68 1 2
37 Bsu36 I 1a
1 2 3
1- 20 21- 49 50-68
Supplementary Figure 1 80
-68)
EI9(50 No RNA hnRNP A1
Supplementary Figure 2 81
EI9 (1a) WT EI9(1a) G3C cold competitor 0 0
hnRNP A1/A2
1 2 3 4 5 6 7 8
Supplementary Figure 3 82
AdML 1 2
taggg
WT 9 2
taCgg
G3C 9 2
WT G3C
9 2
9 2
Supplementary Figure 4 83
WT G3C WT G3C
8 9 10 11
8 9/109 11 8 9 11
Uncut Cut by Pst I
Supplementary Figure 5 84
a b
I8
No RNA supernatant beads
I8
No RNA
I8 No RNA 75 PTB/hnRNP K PTB PTB 50
PCBP1/2 37 1 2 3 4
Immunoblotting 25
1 2
SDS-PAGE
Supplementary Figure 6 85
cold Competitor 8b WT 8bmu
0 0
PTB
1 2 3 4 5 6 7 8
Supplementary Figure 7 mglobulin PKM
Mock ASF A1 A2 PTB A1/A2 M2 M1 cut by [ Tth111I ] [ Tth111I Mock ASF A1 A2 PTB A1/A2 [ Pst I] Supplementary FigureSupplementary 8 Mock ASF A1 A2 PTB A1/A2 M2 M1 86 87
MCF-7 U87
control ASF
ASF
control
PTB/A1/A2 siRNA PTB/A1/A2
M1
M2
M1% 5 .0 2.2 10.2 9.2 6.5 18.8
[ cut by Pst I ]
Supplementary Figure 9 88
a
b shRNA
SRp20 mock
M1
shRNA mock SRp20
SRp20
M2 Actin
1 2 1 2 M1% 0.9 3.5
Immunoblotting Cut by [ Pst I ]
RT-PCR
Supplementary Figure 10 89
Normal Brain Pilocytic Low grade Glioblastoma astrocytomas astrocytomas mutiforme
ASF/SF2
Tra2b
Tra2b/Actin 1.0 1.3 1.3 1.2 0.5 0.5 0.6 0.6 0.5 0.2 0.4 0.2 0.4 0.3 0.4 0.3
TLS
TLS/Actin 1.0 1.2 0.9 1.1 1.8 1.0 2.0 2.1 0.4 3.8 0.5 1.4 5.5 10.5 0.6 0.9
hnRNP K hnRNP K/Actin 1.0 1.2 0.8 1.5 1.2 1.0 2.0 1.5 0.4 6.2 0.2 1.6 5.3 5.8 0.6 1.4
n-Myc
Actin
Supplementary Figure 11 90 a b c
control c-Myc2 shRNA shRNA control c-Myc2 shRNA control c-Myc2
c-Myc c-Myc M1
c-Myc/Actin 100 33.6
PTB PTB
PTB/Actin 100 45.4 hnRNP A1 M2 hnRNP A1 1 2 hnRNP A1/Actin 100 48.6 hnRNP A2 M1% 7 30
cut by [ Pst I ] hnRNP A2 mglobulin hnRNP A2/Actin 100 48.7 RT-PCR
ASF/SF2 ASF/SF2/Actin 100 96.5 RT-PCR
CPSF73
CPSF/Actin 100 122.7
Actin
Immunoblotting
Supplementary Figure 12 91
a E Box E Box
pGL3-A1p CACGTG CACGTG
hnRNP A1 promoter Luciferase Poly A SV40 Enahncer
E Box E Box
pGL3-A1pMu CAgcTG CAgcTG
hnRNP A1 promoter Luciferase Poly A SV40 Enahncer b
2.5
2
1.5
1
0.5 Relative luciferase activity luciferase Relative
0
PGL3-A1p + + + - - - PGL3-A1pMu - - - + + +
c-Myc - -
c-Myc
Actin
1 2 3 4 5 6
Supplementary Figure 13 92
a b
shRNA control c-Myc shRNA control c-Myc
c-Myc M1
PTB
hnRNP A1 M2
hnRNP A2
1 2 Actin M1% 4.8 5.0
Cut by [ Pst I ] 1 2
Immunmoblotting RT-PCR
Supplementary Figure 14 93
Chapter 4: HnRNP proteins control mutually exclusive splicing of pyruvate kinase M pre-mRNA
94
HnRNP proteins control mutually exclusive splicing of pyruvate kinase M pre-mRNA
Mo Chen1, Charles J. David2 and James L. Manley1.*
1 Department of Biological Sciences, Columbia University, New York, NY 10027 2 Current address: Cancer Biology and Genetics Program, Howard Hughes Medical Institute, Memorial Sloan-Kettering Cancer Center, P.O. Box 116, 1275 York Avenue, New York, NY 10021
*Correspondence and requests for materials should be addressed to J.L.M. ([email protected])
95 Abstract
Expression of the pyruvate kinase M (PKM) gene, to generate the PKM1 and PKM2 isoforms, provides an important example of mutually exclusive splicing. We showed previously that hnRNP A1/ A2 and PTB, are critical for expression of PKM2. In this study, we further investigate the molecular mechanism of PKM mutually exclusive alternative splicing. We show that additional binding sites for these proteins in intron 9 are required for full repression of exon
9 inclusion. Moreover, we identify two critical repressive hnRNP A1/A2 binding sites in exon 9, which facilitate cooperative binding to a site overlapping the exon 9 5’ splice site. Strikingly, these two exonic sites, which are lacking in the homologous exon 10, are sufficient, when recreated in exon 10 by mutation, to induce exon 10 exclusion. Paradoxically, we found that hnRNP A1/A2 and PTB binding sites in introns 9 and 10 are required to prevent exon 10 inclusion when exon 9 is de-repressed by reduction in the levels of the three proteins. This represents a novel, concentration-dependent modulation of hnRNP function, reflecting redistribution of occupancy along sites in the PKM pre-mRNA, and, together with nonsense mediated decay, functions to prevent appearance of PKM mRNA including both exons. 96 Introduction
Alternative splicing (AS) is a common mechanism for regulating gene expression and
generating proteomic diversity. Recent estimates indicate that as many as 95% of human multi-
exon genes produce transcripts that are alternatively spliced 1,2. Precursor mRNAs are spliced
and exons are re-joined in different ways to produce proteins that are tailored for specific
functions to meet environmental and tissue-specific requirements. Whether an alternative exon is
included or excluded during splicing is often determined by cis-regulatory elements which
harbor information that is recognized by trans-acting regulatory proteins 3. Two groups of
classical AS regulators are the serine/arginine-rich (SR) proteins, which usually function as
splicing activators, and hnRNP proteins, which often act as splicing repressors. SR proteins
frequently function by recognizing exonic splicing enhancers (ESEs), while hnRNP proteins bind
to exonic splicing silencers (ESSs) and intronic splicing silencers (ISSs) to inhibit splicing.
Disruption of alternative splicing is a frequent cause of disease. This can occur in several
ways, including mutation of RNA sequence elements or deregulation of RNA binding proteins 4,5.
Mutations can destroy or weaken essential splicing signals or regulatory elements 6. In some cases, mutations create cis-elements that can be bound by splicing factors to result in changes in
AS. For example, the SMN2 gene is nearly identical to the nearby SMN1 gene, but single nucleotide changes in SMN2 exon 7 and intron 7 result in creation of binding sites for hnRNP
A1/A2 proteins, which repress inclusion of exon 7, creating a non-functional protein 7,8, leading to the neurodegenerative disease Spinal Muscular Atrophy. Likewise, changes in expression of hnRNP or SR proteins, or other splicing factors, can have significant effects on AS patterns.
Indeed, a variety of diseases have been shown to involve changes in the expression levels of these and other splicing factors 4,5,9.
Regulation of PKM mRNA splicing provides an important example of an AS event
critical for disease, and which reflects changes in the levels of hnRNP splicing regulators 10-12.
The PKM gene encodes a primary transcript that contains two mutually exclusive exons, exon 9 97 and exon 10, and inclusion of one or the other leads to two different isoforms, PKM1 and PKM2,
respectively 13. PKM2 is expressed in embryonic cells while PKM1 is expressed in most adult
tissues 14,15. Reversion from PKM1 to PKM2 is observed in most cancers, partially explaining
the Warburg effect observed in tumor cells 16 and ensuring maximal turmorigenicity 15,17.
Recently, David et al. elucidated a pathway that regulates PKM AS 10. Expression of three
hnRNP proteins, hnRNP A1, hnRNP A2 and PTB, was found to be upregulated in cancer cells
by the oncogene c-Myc, and this was shown to alter splicing of PKM pre-mRNA to promote
formation of PKM2. These three hnRNP proteins exclude exon 9 by binding to sequences
flanking exon 9. A subsequent study by Clower et al. also showed that hnRNP A1/A2 and PTB
are important for the PKM splicing switch in cancer cells 12.
A large number of previous studies have suggested important roles for hnRNPA1/A2 and
PTB in cell growth control and disease, especially cancer. All three proteins are known to be
expressed in a proliferation-associated manner and in a wide range of cancers 5,10,18-21. In addition, they are all involved in AS of transcripts produced from cancer-related genes 10,22-25.
More recently, the studies of PKM AS mentioned above demonstrated the critical functional consequences for the numerous observations of PTB and hnRNP A1/A2 upregulation in proliferating cells, transformed cells and various cancers. Therefore, further analysis of the detailed molecular mechanism of PKM AS is important and should shed light on how these hnRNP proteins regulate AS, both in general and in cancer cells.
A number of studies have elucidated aspects of the mechanisms of splicing regulation by hnRNP A1/A2 and PTB. Although some studies have shown activation effects from these proteins 23,26, the splicing-repressive activity is predominant 3,27 (ref 2 reviews, including yours).
In some cases, binding to ESSs and ISSs sterically blocks splicing activators from binding to
ESEs and this in turn prevents recruitment of the general splicing machinery 28. hnRNP A1 can also function by binding multiple ISSs flanking an alternative exon, which is proposed to forme a loop structure and thereby repressing the inclusion of the internal alternative exon 29-31. Multiple 98 studies have also provided evidence for cooperative binding of hnRNP A1, and that this can be important for splicing repression 28,32-35. The mechanism of AS regulation by PTB has also been
well studied 36 and its repressive functions can occur at different stages of spliceosome assembly
3. For example, it has been shown to inhibit splicing by binding to the polypyrimidine tract to
block binding of U2AF 37,38 and also by preventing the pairing of U1 and U2 snRNPs 39.
In this study, we provide additional insights into how PKM AS is regulated. First, using a splicing minigene construct that accurately recapitulates PKM splicing in HeLa cells, we identified additional PTB and hnRNP A1/ A2 ISSs in intron 9 necessary for full exclusion of exon 9. More importantly, we found two ESSs in exon 9, absent from exon 10, that match the hnRNP A1 consensus, and which are critical for exon 9 exclusion. We show that these ESSs function cooperatively to facilitate hnRNP A1 binding to a previously described ISS in intron 9
10. Unexpectedly, precisely swapping exon 9 and exon 10 did not affect splicing outcome; exon
10 was included in both cases. However, mutating the ESSs in exon 9 and creating them in exon
10 was sufficient to completely switch splicing from exon 10 inclusion to exon 9 inclusion. We
also elucidated the mechanism of how exon 10 is excluded when exon 9 is derepressed and show
that hnRNP A1 and PTB, when their protein levels are reduced, release the inhibition of exon 9
but repress exon 10 inclusion, through binding sites present in introns 9 and 10. This mechanism, coupled with nonsense mediated decay, function to prevent the appearance of PKM mRNA containing both exon 9 and exon 10. Together, our results provide a detailed picture of how an important mutually exclusive splicing event is regulated.
Results
PTB and hnRNP A1/A1 binding sites in PKM intron 9 contribute to exon 9 exclusion.
We previously showed that hnRNP A1/A2 (A1/A2) and PTB inhibit PKM exon 9
inclusion by binding to intronic sequences flanking exon 9. PTB recognizes two UCUU elements
upstream of the 3’ splice site of exon 9 and A1/A2 bind to UAGGGC (ISS1) 10, which is 99 immediately downstream of the exon 9 5’ splice site 10 (Fig. 1a). We first wished to investigate
whether additional intronic cis-elements are involved in regulating PKM AS. To this end, we
constructed a minigene splicing construct containing PKM sequences from exon 8 to exon 11 with 200-400 nucleotide (nt) intronic sequences flanking each exon and with an intact 401 nt intron 9 (Fig. 1a). This construct accurately recapitulates PKM AS in HeLa cells (see below).
Apart from the elements identified previously 10, sequence examination revealed a
number of additional sequences in intron 9 that resemble A1/A2 and PTB binding sites (Fig. 1b).
To examine whether these potential A1/A2 and PTB sites are important for PKM splicing, we
mutagenized several of them and examined effects on PKM splicing. We first mutated the C
residues in the putative PTB sites into G’s as indicated in Fig. 1b, as C to G mutation has been
shown to significantly reduce PTB binding to the ISS1 site in intron 9 10. To analyze PKM
splicing, we transfected wild-type (WT) and mutated (PTBMu) splicing constructs (Fig. 1c, right
panel) into HeLa cells, isolated total cellular RNA and performed RT-PCR assays with [α-
32P]dCTP (primers and diagrams for predicted products are shown in Fig. 1c, left panel). Two
major bands were detected (indicated in Fig. 1c, right panel). One, which we dubbed SI (single
inclusion), contained either exon 9 or 10 (see below), while the other, DI (double inclusion),
contained both of the mutually exclusive exons.
We next determined the efficiency of exon inclusion with WT and mutant constructs.
Results from three separate experiments were quantified, by dividing the amount of each product
by the sum of both products, and are graphed in Fig. 1d. With the WT construct, the majority
(~97%) of the pre-mRNA produced was spliced into an SI RNA product, while only 3.0% was
DI (Fig. 1c, lane 1). Additionally, similar to endogenous PKM splicing in HeLa cells, all SI RNA
contained exon 10 (see Fig. 1e below). Significantly, C to G mutations in putative PTB binding
sites increased the amount of DI product, from 3.0% to 12%, indicating that these sequences
contribute to exon 9 exclusion (Fig. 1c and d, lanes 1 and 3). Why this leads to more DI rather
than SI with exon 9 is addressed below. 100 Next, putative A1/A2 binding sites, which contain a core ‘TAG’ element 40, were mutated
to ‘TAC’, which has been shown to abrogate A1/A2 binding 8,10,25. The resulting mutants were
named A1Mu1-4 (contains mutations in bindings sites 1 to 4; site 1 is the exon 9 proximal site,
ISS1, described previously 10); A1Mu1-6, A1Mu2-6 and A1+PTBMu (contains all mutations in
A1Mu1-4 and PTBMu). Mutating either all six putative A1/A2 binding sites (A1Mu1-6) or only
the first four (A1Mu1-4) drastically increased the DI product, from 3.0% to 66 (Fig. 1c and 1d,
lanes 2 and 5). In addition, mutation of ISS1 by itself also had a dramatic effect, increasing DI to
48% (see ref. 10 and Supplementary Fig. 1), similar to the effect observed with A1Mu2-6 (Fig.
1c and 1d, lane 6). Finally, the A1+PTBMu construct displayed the greatest effect, increasing DI
from 3.0% in WT to 75% (Fig. 1, lane 4). These findings suggest that efficient exon 9 exclusion
involves multiple A1/A2 and PTB binding sites in intron 9.
We also examined the amounts of exon 9 inclusion in the SI products produced from the
intron 9-mutated splicing constructs. While the total amount of SI went down in all cases, did
any of the remaining SI contain exon 9; i.e., correspond to PKM1 mRNA? For this, we designed
exon 9-specific and E10-specific primers to amplify exon 9 SI and DI or exon 10 SI and DI,
respectively (Fig. 1e, left panel). Significantly, in all cases all the exon 9-included product was
DI (Fig. 1e, middle panel), suggesting that the release of repression of exon 9 inclusion by mutation of intron 9 ISSs led to increased DI but not exon 9-containing SI. While the mechanisms resulting in PKM1 mRNA instead of DI are investigated below, these results provide further evidence that multiple A1/A2 and PTB binding sites in intron 9 contribute to exon 9 exclusion.
Exonic sequences are involved in PKM alternative splicing regulation
Exonic sequences are of course also often involved in AS regulation. Although our previous experiments had failed to identify potential regulatory proteins binding to exon 9 or 10
(11), we nonetheless set out to investigate whether cis-elements in exon 9 or 10 function in PKM
AS regulation. To this end, we first precisely swapped exon 9 and exon 10 in the WT splicing 101 construct (named Eswap) to determine if changing the intronic sequences flanking the exons would affect PKM AS (Fig. 2a upper panel). If exon sequences were not important and the ISS elements are the sole determinant of exon 9 exclusion, then exon 10 would be excluded and exon
9 included in transcripts produced from Eswap. We transfected WT and Eswap constructs into
HeLa cells and performed PKM splicing assays as described above. We determined the SI identity by digesting the PCR products with Pst I (specifically digests exon 10) (Fig. 2a) and the results were confirmed by PCR using exon-specific primers (data not show). Surprisingly, however, exon swapping did not alter the outcome of PKM AS; exon 10-containing RNA remained the only SI product (Fig. 2a, lanes 2 and 4). Furthermore, the small amount of DI product detected with WT was eliminated in Eswap (Fig. 2a, lane 2), indicating exon 9 was excluded even more efficiently in this downstream position. We conclude that despite the importance of intronic sequences in intron 9 (and 8), these ISSs are not sufficient to exclude any adjacent exon. Therefore, we hypothesize that either ESEs in exon 10 function to overcome the repressive hnRNP proteins, and/or ESSs in exon 9 function to repress exon 9 inclusion regardless of its position.
To investigate the above possibilities further, we next examined exon 9 and exon 10 splicing independent of each other. First, we precisely deleted exon 9 from WT and Eswap constructs to generate E9del1 and E9del2, respectively (Fig. 2b, top panel). Following transfection and RNA analysis as above, we found that exon 10 was included efficiently in both positions (Fig. 2b). We also constructed E10del1 and E10del2 derivatives in which exon 10 was precisely deleted from WT and Eswap, respectively, and found that exon 9 was largely excluded in both cases (Fig. 2c). These results confirm our hypothesis that exonic elements exist in exon 9 and possibly exon 10. Inclusion or exclusion of either exon was independent of its position, but rather dependent on the exonic sequences. In addition, exon 9 was partly included in E10del1
(24%) whereas it was completely excluded in E10del2 (Fig. 2c, lanes 2 and 3), suggesting that the putative ESSs in exon 9 cooperate with ISSs more or less effectively when exon 9 is 102 positioned differently in PKM transcripts, and that possible ISSs may exist in intron 10 that could function with ESSs in exon 9 (see below).
Exonic splicing silencers are important for intronic binding of hnRNP A1/A2
We next set out to examine the possible role of exonic sequences in protein binding to
PKM RNA regulatory sequences, such as ISS1. Our previous studies showed that an RNA consisting of the 3’ part of exon 9 and 5’ part of intron 9, called EI9, was bound by A1/A2, while the corresponding EI10 RNA was not 10. However, the apparent A1/A2 binding site, ISS1, is
very similar to the corresponding sequence in intron 10, ISS1-10 (Fig. 3a, top panel). Two
possible reasons can explain why ISS1 was bound by A1/A2 whereas ISS1-10 was not. One is
that the intrinsic sequence of ISS1 determines that it is a better A1/A2 binding site, while the
other is that both ISS1 and ISS1-10 are potential A1/A2 binding sites, but the context facilitates
binding to ISS1. Considering the results from the exon swapping splicing assays, we hypothesize
that exonic sequences contribute to the binding profiles of EI9 and EI10.
To test the above hypothesis, we carried out ultraviolet (UV) crosslinking assays, initially
using uniformly labeled exon 9- or exon 10-containing RNAs. We first prepared two RNAs for
crosslinking, EI9s and EI10s, which contain the exonic region of EI9 or EI10 and a very short
intronic extension that contains only ISS1 or ISS1-10 and several additional nucleotides to
ensure protein binding to the ISSs (Fig. 3a, top panel). ISS1 (TAGGGC) and ISS1-10
(TAGGAG) differ only by the two most 3’ nucleotides, so we mutated the last two nts of ISS1
into AC or AG so that they either resemble or are the same as ISS1-10, and named them EI9sM1
and EI9sM2. Similarly, we mutated the last two nts of ISS1-10 into GG or GC, generating
EI10sM1 and EI10sM2 (Fig. 3a, top panel). We synthesized [α-32P]-UTP labeled RNAs from
these constructs, and performed UV crosslinking assays with HeLa nuclear extract (NE) 41. EI9s showed the same pattern observed previously with the EI9 RNA, one major band that was identified by IP and mass spectrometry as A1/A2 10 (Fig. 3a, lane 1). Interestingly, both EI9sM1
and EI9sM2 were also bound strongly by A1/A2 (Fig. 3a, lanes 2 and 3). Moreover, mutations 103 “improving” ISS1-10 did not promote binding of A1/A2 to the EI10s substrates (lanes 4-6).
Because the intronic sequences of EI10sM2 and EI9s were the same, the difference in binding
(lanes 1 and 6) must therefore have been due to exonic sequences. In addition, EI9sM2 was bound by A1/A2 more strongly than EI9s (lanes 1 and 3), which is consistent with our finding that in splicing assays more exon 9 was included in E10del1 than in E10del2 (Fig. 2c). It is possible that sequences in exon 9 cooperate with ISS1-10 more effectively than ISS1, so that repression of exon 9 was more complete. Our results suggest that ISS1-10 is intrinsically as good an A1/A2 binding site as ISS1, but that exonic sequences cooperate with ISS1 to enable A1/A2 binding to EI9s but not EI10s.
We next examined exonic sequences and found two conserved TAG sequences (which we refer to as ESS1 and ESS2) in exon 9 that are absent in homologous exon 10 (66% sequence identity). hnRNP A1 has been shown to bind cooperatively to RNAs with multiple binding sites
28,32-34, so we propose that the binding of A1/A2 to EI9s requires all three elements, ESS1, ESS2 and ISS1 (Fig. 3b , upper panel). To investigate this hypothesis, we mutated the TAG to TAC in
ESS1 (EI9sE9G3C1) or in both ESSs (EI9sE9G3C2) (Fig. 3bc, top left), and first analyzed uniformly labeled RNAs by UV crosslinking as above. Consistent with the effects of a similar
G3C mutation in ISS1 10, the G3C mutation in ESS1 decreased A1/A2 binding to EI9s RNA by half (Fig. 3b, lane 3), while the double mutant EI9sE9G3C2 reduced the amount of A1 crosslinked to approximately 25% of the EI9S RNA (Fig. 3b, lane 2). To rule out the possibility that this reduction was a nucleotide-specific effect, the ESSs were mutated to TAA instead of
TAC (Fig. 3b, top right); the G3A mutations resulted in similar reductions in A1/A2 binding (Fig.
3b, lanes 4-6). Thus, although intronic sequences immediately downstream of both exon 9 and exon 10 contain putative A1/A2 binding sites, exon 9 harbors two elements that enable A1/A2 to interact with EI9s RNA, and this suggests the importance of cooperation between ISS1 and ESSs in A1/A2 binding and perhaps a role in exon 9 exclusion. hnRNP A1/A2 bind cooperatively to exonic and intronic splicing silencers 104 In order to test the hypothesis that the putative ESSs facilitate binding of A1/A2 to ISS1,
we next performed UV-crosslinking experiments with site-specifically labeled EI9s and the ESS
mutants, first using HeLa NE. In these experiments, the G in ISS1 at position 3 was 5’ labeled
with 32P to determine whether the G3C mutations in the two ESSs affect binding of A1/A2 to
ISS1. Importantly, the results (Fig. 3c) show that while EI9s was bound strongly by A1, binding
was reduced to ~25% of that to EI9s with EI9sE9G3C1 and ~19% with EI9sE9G3C2. These
results indicate that the ESSs cooperate with ISS1 to promote binding of A1/A2 to ISS1 (Fig. 3c).
We next set out to examine the apparent cooperative binding in more detail. Given that
the above experiments were performed with NE, it was possible that proteins in addition to
A1/A2 might be involved. We therefore wished to determine whether purified hnRNP A1 binds
cooperatively to the EI9s RNA, and if so what domains of the protein are necessary. To this end,
we purified GST-hnRNP A1 and several truncations (diagrams in Fig. 3d, left) from E. coli (Fig.
3d, right), cleaved off the GST motif, and performed UV-crosslinking assays with site-
specifically labeled wild-type and mutant EI9s RNAs (Fig. 4e). Full-length hnRNP A1 (Fig. 3e,
lanes 1-3) bound to the RNAs more strongly than derivatives with only one RRM (RRM2-A1)
(lanes 4-6) or without the C-terminal domain (Up1) (lanes 7-9), while the RRM2 domain alone
did not bind to the RNAs (lanes 10-12). This indicates that both the C-terminal glycine-rich domain and RRMs are important for maximal RNA binding, as previously shown 42-44.
Importantly, not only full-length A1 but also Up1 and RRM2-A1 binding to ISS1 was reduced by the ESS mutations, to 38%, 6.1% and 18%, respectively, compared to the binding to WT
RNA (Fig. 3e, lanes 1 to 9, quantification indicated below). These results indicate first that hnRNP A1 alone is sufficient for the cooperative binding to ISS1 detected in NE, and second that the C-terminal domain is not essential for cooperative binding, and that RRM1 can be sufficient. While the C-terminal domain has been previously shown to be involved in protein- protein interactions 34,35 and it contributes to optimal binding to EI9s, our results indicate that
cooperativity can be achieved in its absence. 105 The exon 9 hnRNP A1/A2 binding sites are critical for exon exclusion
We next wanted to test whether the two exon 9 A1 binding sites defined above are important in regulating PKM splicing. To this end, we mutated both TAGs to TACs in the WT and A1Mu1-4 constructs (Fig. 4a), and performed PKM splicing assays as shown in Fig. 1. Since the most dramatic effect of the intron 9 binding site mutations was to increase DI, we quantified the DI percentages as in Fig. 1c, and a bar graph derived from three independent experiments is shown in Fig. 4b. Strikingly, the ESS mutations in the WT construct sharply increased DI product, from 6.1% to 90% (Fig. 4a and 4b, lanes 1 and 2), while the same mutations in A1Mu1-
4 also increased DI, from 69% in A1Mu1-4 to 100% (Fig. 4a and 4b, lanes 3 and 4). To determine whether the ESS mutations enhanced SI (PKM1) production (i.e., exon 10 exclusion), we used exon 9-specific primers to examine if the ESS mutations changed the composition of the remaining SI in the WT construct. We found that 8.3% of the exon 9-containing products were in the form of PKM1 with the ESS mutant, compared to 0% in WT (Fig. 4c, lanes 1 and 2).
Together, these results indicate that the ESSs in exon 9 play a significant role in exon 9 exclusion.
We next wanted to investigate whether other exonic sequences besides the ESSs in exon
9 are important for PKM AS regulation. One way to address this is to create TAG elements in exon 10 while simultaneously mutating the TAGs in exon 9 into the corresponding sequences in exon 10, and determining whether this is sufficient to switch PKM splicing from M2 to M1. If simply switching the position of the A1/A2 binding sites is sufficient to switch PKM splicing, this would indicate that no other exonic elements are required for exclusion of exon 9 and inclusion of exon 10. Therefore, we introduced the two ESSs into exon 10 in the same position as in exon 9, which required only 4 nt changes, and mutated 4 nts in exon 9 to create the original exon 10 sequence (ESSMu, Fig. 4b, left panel, sequences indicated on top). Strikingly, using the same splicing assay as above, we found that the mutations in ESSMu completely switched splicing from PKM2 (exon 10 inclusion) to PKM1 (exon 9 inclusion) (Fig. 4d, lanes 3 and 6). In addition, no DI inclusion was detected with either Eswap or ESSMu constructs. This is 106 consistent with our results (see Fig. 3a) suggesting that ISS1-10 provides a stronger potential
A1/A2 binding site than ISS1, so that the two ESSs in exon 9 (in the case of Eswap) or in exon
10 (in the case of ESSMu) cooperated better with ISS1-10. Most importantly, these results
indicate that the two TAG elements are the only exonic sequences critical for determining
whether exon 9 or exon 10 is included in proliferating cells.
Intron 10 plays a role in preventing double inclusion PKM mRNA splicing
The above experiments have provided a detailed picture of how exon 9 is excluded from
PKM mRNA to generate PKM2 mRNA. However, when exon 9 inclusion was increased by
mutating ISSs or ESSs, most of the product detected was in the form of DI RNA, rather than SI
(PKM1) RNA. One explanation for this is that sequences in intron 8 and/or 10 missing in the
minigene construct contribute to exon 10 exclusion. To investigate this, we first made a construct
containing full-length intron 8 (WT-In8) and several derivatives containing mutations that
increase exon 9 inclusion. However, none of these displayed an increase in the amount of PKM1
mRNA, although the amount of DI was somewhat decreased (results not shown; see also below),
perhaps due to the presence of additional repressive PTB sites in the added sequences 25.
We next asked whether splicing to generate PKM1 would be increased by the presence of full-length intron 10. We therefore inserted additional intron 10 sequences into WT-In8, to obtain a construct containing both full-length intron 8 and intron 10 (WT-In8-In10), and into a derivative containing the two ESS mutations that increase exon 9 inclusion (E9G3C-In8-In10).
Following transfection and RT-PCR as above, we found that addition of In10 did not by itself significantly change PKM AS relative to that obtained with WT-In8 in HeLa cells (Fig. 5a, lanes
5 and 6). To ask whether PKM1 splicing was increased when exon 9 inclusion was enhanced, we transfected the exon 9 mutated derivatives into HeLa cells and analyzed PKM splicing by RT-
PCR. Significantly, with E9G3C-In8-In10, compared with E9G3C-In8, we observed an increase in PKM1 levels, from 2.4% to 13% (Fig. 5b, lanes 2 and 3). This suggests that full-length intron
10 plays a role in producing PKM1 mRNA when exon 9 repression is released. 107 To examine this hypothesis further, we next wished to investigate how intronic sequences
contribute to exon 10 exclusion when A1/A2 and PTB levels were reduced. To this end, we transfected WT-In8-In10 and WT constructs into PTB/A1/A2 siRNA- or control siRNA-treated
HeLa cells (knockdown efficiency was estimated by western blotting; Supplementary Fig. 2), and analyzed PKM splicing by RT-PCR as above. Consistent with the results described with the
PTBMu and A1Mu1-4 constructs, PTB and A1/A2 knockdown greatly increased the amount of
DI in WT, from 9% to 90% (Fig. 5c, lanes 1 and 2), with the de-repressed exon 9 mainly found in DI (96%) rather than PKM1 (4.1%) (Fig. 5c, lane 6). WT-In8-In10 showed less DI in both control and knockdown experiments than WT, which was due to the repression function of full- length intron 8 (Fig. 5c, compare DI in lanes 3 and 4 to lanes 1 and 2). Interestingly, however, following PTB and A1/A2 depletion, WT-In8-In10 showed significantly increased M1 levels and concurrently reduced amount of DI relative to WT (lanes 6 and 8). Addition of full-length intron
10 increased the amount of PKM1 in the PTB/A1/A2 depleted cells from 4.1% to 40%. The enhanced PKM1 splicing (i.e., exon 10 exclusion) appeared to be due solely to the expanded exon 10 sequences, as splicing of WT-In8 in PTB/A1/A2-depleted cells did not result in increased M1 production (Supplementary Fig. 3). To confirm this, we constructed and analyzed
WT-In10 and E9G3C-In10, which contain intact intron 10 but not intron 8. WT-In10 showed a similar effect in reducing DI and increasing SI (PKM1) (38% of PKM1) when PTB/A1/A2 were depleted (Fig. 5d, lanes 6 to 8) as did WT-In8-In10 (lane 7) (35% of PKM1). WT-In10 did not reduce DI in the control siRNA-treated cells whereas WT-In8 did, consistent with our results showing that intact In8 further repressed exon 9 inclusion (Supplementary Fig. 3, lanes 1, 3 and lanes 4, 6). Likewise, full-length intron 10 significantly increased SI (PKM1) with E9G3C-In10 compared to E9G3C, from 8.4% to 23% (Fig. 5b, lanes 1 and 4). Together, these results indicate
that full-length intron 10 enhances PKM1 production by reducing exon 10 inclusion when
A1/A2/PTB levels are reduced. 108 We next wished to identify specific sequences, if any, responsible for the In10 effect. To this end, we first made minigene constructs with In10 truncations containing the 5’ or 3’ half of
In10 in WT (WT-In10 1/2 and WT-In10 2/2, respectively; both constructs contained intact 5’ and 3’ splice sites) and performed splicing assays as above. The results showed that both of these truncations resulted in reduced PKM1 levels when compared to constructs containing the full length In10 (Figure 5d, lanes 8 to 10). To exclude the possibility that an important sequence is located in the middle of In10 and destroyed in the above truncations, we constructed WT-In10
1/3, WT-In10 2/3 and WT-In10 2/3, each containing one third of In10. Using these constructs,
PKM1 levels were further reduced with a shorter intron 10 (Figure 5e lanes 7 to 10). These results suggest that PKM1 production in cells with reduced PTB/A1/A2 depends on a full-length intron 10.
Intron 9 cooperates with intron 10 to produce PKM1 when PTB/A1/A2 levels are reduced
We next asked whether intron 9 sequences play a similar role in preventing double inclusion. We first deleted three ~100 nt long fragments in intron 9 (leaving splice sites intact) separately from WT-In10, generating WT-In10-In9Del1, WT-In10-In9Del2 and WT-In10-
In9Del3 (Fig. 6a, left). Interestingly, each deletion resulted in a large reduction in PKM1 production, with a concomitant increase in DI when PTB/A1/A2 levels were reduced (Fig. 6a, lanes 3 to 5). This result indicates that PKM1 production enhanced by full-length intron 10 also depends on full-length intron 9.
Due to the importance of intron 9 in PKM1 production, we next cloned full-length intron
10 into the various intron 9 mutants analyzed in Figure 1 and examined if the A1/A2 and PTB binding sites in intron 9 play a role in exon 10 exclusion when PTB/A1/A2 levels are reduced.
First, when PTB/A1/A2 levels were high, A1Mu1-4-In10 and PTBMu-In10 produced similar amounts of DI as the versions with truncated exon 10 (Supplementary Fig. 4). Interestingly, consistent with the intron deletion result, A1/A2 and PTB binding site mutations in intron 9 greatly reduced PKM1 level in minigenes containing full-length intron 10 when PTB/A1/A2 109 levels were reduced (Figure 6b, compare lane 8 and lanes 10 and 12). When both A1/A2 and
PTB binding sites were mutated in intron 9 in A1+PTBMu-In10, PKM1 levels were reduced to the same level as observed using constructs lacking the additional intron 10 sequences
(Supplementary Fig. 5). These results indicate that A1/A2 and PTB binding sites in intron 9 are important for exon 9 exclusion when PTB/A1/A2 levels are high, but when PTB/A1/A2 levels are reduced by siRNAs to lower than 10% for A2 nd PTB and to ~20% for A1 (Supplementary
Fig. 2), these sites cooperate with intron 10 to inhibit exon 10 inclusion, to produce PKM1 mRNA.
Because A1Mu1-4 and A1Mu1-6 had a similar effect in exon 9 inclusion (Figure 1e, lanes 2 and 5), it appears that the fifth and sixth putative A1/A2 binding sites in intron 9 are not important for exon 9 exclusion, possibly because they are farther away from exon 9. To examine their potential role in exon 10 exclusion when PTB/A1/A2 levels are low, we constructed
A1Mu5+6-In10 construct that only contains the fifth and sixth A1/A2 binding sites but not the first four. As expected, A1Mu5+6-In10 did not result in increased inclusion of exon 9 as
A1Mu1-4-In10 did (Fig. 6c, lane 3). However, similar to A1Mu1-4-In10, this mutation reduced
PKM1 production when PTB/A1/A2 levels were reduced (Fig. 6c, lanes 10 to 12). In addition, we also preformed UV crosslinking-immunoprecipitation experiments with antibodies against hnRNP A1 and PTB, and showed that hnRNP A1 and PTB bind to intron 9 sequences
(Supplementary Fig. 6). Together,these results further confirm the importance of putative A1/A2 and PTB binding sites in intron 9 in preventing the production of DI when A1/A2/PTB levels are low.
Reduced levels of hnRNP A1 and PTB exclude exon 10
The results above suggest the intriguing possibility that A1/A2 and PTB have dual roles in PKM AS regulation. When PTB/A1/A2 levels are high, the proteins bind strongly to sites in exon 9 and intronic regions flanking exon 9 to repress its inclusion. However, when PTB/A1/A2 110 levels are reduced, their binding to these sequences is reduced, allowing exon 9 to be included.
At the same time, binding to intronic sites flanking exon 10 functions to prevent double inclusion.
In other words, PTB/A1/A2 binding to sites in and around exon 9 may be more dramatically
reduced than is their binding to the intronic binding sites that prevent DI. To test this hypothesis, we performed crosslinking-immunoprecipitation (CLIP) using an hnRNP A1 antibody to detect hnRNP A1 (A1) binding in cells at different positions on PKM mRNA in both control and
PTB/A1/A2 siRNA-treated HeLa cells. hnRNP A1 binding at different positions on PKM pre- mRNA was quantified by dividing RT-PCR from IP signal by input RNA signal and we arbitrarily set binding at the exon 9 5’ ss (primer set 9f5r1) in control siRNA-treated HeLa cells as 1.0. The amplicons amplified by different primer sets are indicated in Figure 6d. First, consistent with our in vitro UV crosslinking results, A1 binding to the exon 9 5’ss was higher than to the exon 10 5’ss, consistent with its role in repressing exon 9 inclusion. In addition, binding of A1 to the 5’ss of exon 9 was reduced by about 50% after A1/A2 levels were reduced
(Fig. 6e and Supplementary Fig. 7). Interestingly, however, A1 binding to the region amplified by In9f3r3, which includes A1 binding sites 5 and 6, did not decrease after A1/A2 depletion.
Moreover, A1 binding to three regions in intron 10 containing putative A1 binding sites actually increased, with ~2 fold increase for primer set In10f1r1 and 1.5 fold increase for In10f2r2, in line with a potential role for these proteins in preventing exon 10 inclusion at lower levels (Fig. 6e and Supplementary Fig. 7). A likely cause of this dramatically different behavior at the distinct sites is discussed below (see Discussion).
We also examined binding of PTB in introns 8 and 9. Primer set In8f1r1 amplifies the region containing the PTB binding sites described earlier 10 and primer sets In9PTBf1r1 and
In9PTBf2r2 amplify two regions containing putative PTB binding sites described in Figure 1. As
expected, binding of PTB to the site immediately upstream of exon 9 was reduce by about 50%
after PTB/A1/A2 knockdown, whereas binding to sites in intron 9 was not reduced (Fig. 6f and
Supplementary Fig. 8). These results suggest that, as with A1/A2, exon 9-proximal PTB binding 111 was significantly more sensitive to a reduction in PTB levels than PTB binding deep within
introns. The fact that the hnRNP proteins remain bound to the intron 9/10 sites after their partial
depletion, along with the functional importance of the PTB/A1/A2 binding sites in preventing
double inclusion (see above), strongly suggests a role for these proteins in the prevention of DI
when they are expressed at reduced levels by inhibiting exon 10 inclusion (see Discussion).
Nonsense-mediated decay helps prevent appearance of PKM double-included mRNA
While the above data shows that sequences in the introns flanking exon 10 function to reduce double inclusion, significant amount of DI mRNA was nonetheless still detected, even using the full length wild-type minigene. In contrast, double included product is never detected from the endogenous gene after PTB/A1/A2 knockdown. Exon 10 is 167 nt long, so its inclusion in a DI mRNA results in the disruption of the reading frame, resulting in a premature stop codon in exon 11. This suggests that DI mRNA produced from the endogenous PKM gene is likely subject to nonsense mediated decay (NMD). One possibility is that some level of DI PKM
mRNA is naturally produced when exon 9 inclusion is activated, but this is removed NMD.
NMD has in fact been shown to influence AS outcome in several studies 45-48. To investigate
whether NMD plays a role in preventing accumulation of endogenous DI PKM mRNA, we treated HeLa cells with cycloheximide, a translation inhibitor known to block NMD, when PTB and A1/A2 were depleted using siRNAs and analyzed PKM mRNA levels by RT-PCR. A DI product indeed appeared in the presence but not the absence of cycloheximide (Fig. 6g, lanes 4).
No DI mRNA was detected when cells were treated with cycloheximide but not siRNAs (lane 2).
This result was confirmed by depleting Upf1, a factor required for NMD 49, together with PTB
and hnRNP A1/A2 depletion (data not shown). We conclude that NMD prevents accumulation of
DI PKM transcript in cells in which exon 9 exclusion is de-repressed.
Discussion 112 In this study, we have provided a detailed analysis of the mechanisms involved in
regulation of an important mutually exclusive alternative splicing event, the switch that
determines whether PKM1 or PKM2 mRNA is made. Our data suggests that this mechanism is
on the one hand simple, requiring only the activity of several well-characterized RNA binding
splicing repressors, while on the other hand complex, involving multiple cis elements in and
around the two alternative exons that respond differently to varying levels of these proteins (see
Fig. 7). Unexpectedly, two exonic hnRNPA1/A2 binding sites are critical for exon 9 exclusion,
functioning to facilitate cooperative binding to a key intronic site overlapping the adjacent 5’
splice site. Importantly, these and other experiments indicate that the splicing repressors hnRNP
A1/A2 and PTB are sufficient to control exon 9 exclusion and exon 10 inclusion in proliferating
cells. When the levels of these proteins are reduced, or their binding sites mutated, exon 9 is
included and exon 10 excluded. Perhaps most interestingly, we show that PTB/A1/A2 play a
dual role in PKM splicing: when their levels are high, they promote skipping of exon 9 by repressively binding to sites in and around exon 9. When their levels are reduced, they are
displaced from binding sites flanking exon 9, but remain bound to intronic regions flanking exon
10, where they act to prevent exon 10 inclusion. Exon 10 exclusion however is not complete, and
NMD functions to remove DI PKM mRNA (Fig. 7). Below we discuss these mechanisms in
more detail, as well as the implications of our results with respect to AS control more generally.
One key finding of our work is that multiple exonic and intronic sequences, located in
exon 9 and intron 9 and bound by hnRNP A1/A2, play a critical role in suppressing PKM exon 9
inclusion. While the details of the mechanism differ from other well-studied examples of hnRNP
A1 repression, overall it is consistent with a number of previous observations, and appears to
hinge on the ability of hnRNP A1 to dimerize or multimerize 43,50. Several examples of how
hnRNP A1 molecules cooperate with each other to repress exon inclusion have been described.
For example, Chabot and colleagues provided evidence that when hnRNP A1 binds to two
intronic binding sites flanking HNRNPA1 exon 7b, it “loops out” the intervening RNA to repress 113 exon inclusion 29,50. In the case of SMN2 splicing, an exonic and an intronic hnRNP A1 binding
site have been proposed to cooperate to repress SMN2 exon 7 inclusion, likely also by RNA
looping 8. PKM exon 9 repression bears some similarity to both of these, more so to SMN2.
However, exclusion of exon 9 is unlikely to involve an RNA loop as the hnRNP A1 binding sites
are in close juxtaposition. Indeed, while PKM intron 9 contains several hnRNP A1 binding sites,
the site most upstream overlaps the 5’ ss and the ESSs in exon 9 are also close to this 5’ ss. It is
likely that hnRNP A1, through cooperative interactions, forms a multimer that spans the 5’ ss
and sterically blocks U1 snRNP binding. Although it has been shown that ISSs situated near a 5’
ss can interfere with U1 snRNP function but not binding 51, in our case the overlapping nature of the sites strongly suggests that binding is inhibited 52.
Another repression model, known as the propagation model, is also based on hnRNP A1
cooperative binding, and suggests that A1 molecules spread along RNAs following binding to a
high affinity site 28,35. While this is unlikely to apply to PKM splicing, as the principal
mechanism of exon 9 exclusion involves cooperative binding to closely spaced sites, our finding
that additional A1 sites situated downstream in intron 9 are important for full repression suggests
that a more extensive array of bound A1 molecules contributes to exclusion. Indeed, efficient
exon 9 exclusion involves an extensive network of hnRNP proteins and cis-regulatory elements.
Although the cooperative binding of A1 just described plays a critical role, crosstalk between
A1/A2 and PTB proteins also contributes. For example, our data, here and previously 10, indicates that interactions involving a number of PTB sites in intron 8 contribute to repression by interfering with U2AF binding to the exon 9 3’ ss 10.
Our exon swapping results provide additional novel insight into hnRNP -mediated
splicing repression. We were initially surprised to find that when the positions of exon 9 and
exon 10 were switched, splicing outcome was not affected, but simply transplanting the two
ESSs from exon 9 to exon 10 was sufficient to reverse this. These findings indicate that the
repressive context formed by the network of hnRNP binding sites is more extensive than we 114 previously envisioned. During evolution, the acquisition of negative elements in exon 9 was sufficient to bring this exon under the negative control of the intronic hnRNP network. This is
reminiscent of the recently duplicated SMN1/2 genes, where SMN2 exon 7 and intron 7 have
acquired hnRNP A1 binding sites that cooperate to prevent exon 7 inclusion 7,8. Our data thus
show that the precise location of the exon within the inhibitory network is secondary to the
presence or absence of ESS sequences within the exon, indicating a high degree of flexibility in
the evolution of AS regulation.
Our studies have provided one of the most complete pictures to date of how mutually
exclusive splicing can be controlled. Another well-studied example of this mechanism is
provided by the FGFR2 gene. FGFR2 transcripts contain two mutually exclusive (ME) exons,
exon iiib and exon iiic, which are differentially included in two different cell types. As with
PKM AS, FGFR2 AS regulation involves hnRNP A1 and PTB. HnRNP A1 binds to an ESS in
exon iiib 25,53 and PTB binds to ISSs flanking exon iiib 54,55. Artificial recruitment of A1 to exon
iiib leads to its exclusion 25, and siRNA-mediated PTB depletion resulted in partial inclusion of
exon iiib 56. However A1 and PTB levels do not appear to change in cells differentially
expressing the two isoforms, and the small increase of exon iiib inclusion by PTB depletion does
not explain the almost complete switch during EMT. FGFR2 AS is thus unlike PKM AS in that
A1 and PTB do not play critical regulatory roles. The most important regulators of FGFR2 AS
appear to be two cell-specific proteins, ESRP1 and ESRP2. Similar to A1/A2 and PTB in PKM
AS regulation, levels of ESRP1/2 correlated with FGFR iiib expression, and knockdown of these
proteins completely switched AS from iiib to iiic inclusion 57. ESRP1/2 thus function as
activators of exon iiib in epithelial cells, and in fact both exons are repressed by not only A1 and
PTB, but also hnRNP F/H 58 and hnRNP M 59, and activation of exon iiic in mesenchymal cells
may require another currently unidentified activator. In contrast, our data show that A1/A2 and
PTB and changes in their expression levels are sufficient for PKM AS regulation. Our study has thus elucidated a remarkably simple mechanism of regulating ME AS. An interesting possibility 115 is that this type of mechanism may be important in control of AS for widely expressed genes
such as PKM.
An additional interesting aspect of our work is our demonstration that A1/A2 and PTB
play dual roles in PKM splicing, depending on their expression levels. Our data showed that
exon 10 exclusion was enhanced by the presence of full-length intron 10 and intron 9. This is similar to an effect observed in studies of SMN2 pre-mRNA splicing, in which exclusion of
SMN2 exon 7 was more efficient in the presence of a full-length downstream intron 8, which was
suggested to reflect the presence of additional repressive A1/A2 sites. Similarly, mutations in
A1/A2 or PTB binding sites present in intron 9 resulted in increased DI and reduced PKM1
production. Additionally, PTB and A1 binding to sequences that immediately flank exon 9 was reduced, whereas binding to sites deep in introns 9 or 10 remained the same or was even increased after depletion of PTB/A1/A2. This supports a dual-function model for the actions of
A1/A2 and PTB in PKM splicing: at high levels, they function to repress exon 9 inclusion, while
at low levels,exon 10 inclusion is inhibited.
How might changes in the levels of hnRNP proteins result in redistribution of their binding sites
within a pre-mRNA? We suggest the following model (Figure 7). When A1/A2 levels are high,
these proteins bind to multiple sites in intron 9 and intron 10, and also bind strongly to exon 9
and the exon 9 5’ss, resulting in exon 9 exclusion. However, when A1/A2 levels are reduced,
they less effectively compete with the splicing machinery for binding to the exon 9 5’ss and is
thus displaced. In contrast, A1/A2 bound to sites deep in intron 9 are not subject to competition
with the splicing machinery, and do not significantly change, even when A1/A2 levels are
reduced to ~20% of the level in proliferating HeLa cells. Strikingly, PTB behaves in an identical
manner, with exon 9-proximal binding more sensitive to low PTB levels, while intronic binding
is maintained, even when PTB levels are reduced by more than ~90%. Interestingly, A1 binding
to intron 10 sites is actually increased when A1/A2 levels are reduced. Given the high degree of
cooperativity in hnRNP A1 RNA binding, shown here and elsewhere. we hypothesize that this 116 increased binding results from cooperative interactions between A1 molecules bound to intron 9.
When A1/A2 levels are high, A1/A2 molecules bound in and around exon 9 interact with A1/A2
bound to intron 9. After displacement of A1/A2 from the exon 9 5’ ss, intron 9-bound A1/A2
molecules interact with and promote A1/A2 binding to the intron 10 sites, which explains the
increased binding of A1/A2 to intron 10 observed after partial depletion of PTB/A1/A2. Such
cross-exon interactions likely form an RNA loop, resulting in exclusion of exon 10. An
additional important implication of the new work presented here is the likelihood that exonic
hnRNP binding is likely more sensitive to changes in cellular hnRNP concentration than intronic sites. This concept is likely to have broad implications for the regulation of alternative splicing in general.
Accumulation of double inclusion PKM mRNA is also limited by NMD. NMD is a surveillance mechanism that can be activated when a premature termination codon (PTC) is created 60. Up to one third of AS events create a PTC 60. While our data has established a role for
this process in PKM AS regulation, an interesting question is whether NMD is commonplace in
ME AS regulation. To address this, we searched the ME splicing events identified by Wang et al.
1, and analyzed the 34 genes in which two alternative spliced exons were annotated in the UCSC
Genome browser. Among the 34 ME AS events, 25 transcripts would contain a PTC which
triggers NMD when both exons are included. Of the nine transcripts that would not create a PTC, six of them contain a very short intron between the two ME exons (from 1 nt to 64 nts). Given that a minimum size of ~66 nt is required for splicing in mammalian cells 61, this small size is by
itself sufficient to prevent DI. Among the 25 transcripts theoretically targeted by NMD, 23 have introns between the alternative exons longer than 66 nts. Only three transcripts neither contain a
short intron nor create a PTC when both exons are included. This analysis suggests that NMD
and short intron size provide two mechanisms generally used to prevent DI during ME
alternative splicing. As we found with PKM, this implies that regulation of inclusion/exclusion
of only one of the two ME exons must be stringently regulated during ME AS. 117 In conclusion, we have provided a detailed picture of how mutually exclusive alternative
splicing can be regulated. We have shown that hnRNP A1/A2 and PTB are sufficient to regulate
PKM AS, and that this occurs through a network of binding sites that respond differently to
varying levels of the proteins, with two exonic sites in one of the exons orchestrating the process.
Our data also highlight the role that NMD can play in ME AS, by preventing the appearance of
double included mRNAs. These results together illustrate how an important AS event that occurs
in a variety of different cell types and tissues can be precisely controlled by a small number of
general factors. It will be of interest in the future to learn how widely this simple mechanism, or
related ones, is used to regulate mutually exclusive alternative splicing.
Methods
Plasmid constructs
Ultraviolet crosslinking substrates EI9s and EI10s were prepared by inserting an HpaI site downstream of ISS1 and ISS1-10 in EI9 and EI10 constructs 10 by PCR-based site-directed
mutagenesis with Pfu Turbo (Stratagene). The primers were:
EI9HpaIF: ggagtctggcaggtagggcccgttaactaagggcaggtaacactg;
EI9HpaIR: cagtgttacctgcccttagttaacgggccctacctgccagactcc;
EI10HpaIF: caagtctggcagtaggaggcgttaacggcagcggctccctggaatg;
EI10HpaIR: cattccagggagccgctgccgttaacgcctcctacctgccagacttg.
EI9s and EI10s mutants were prepared with site-directed mutagenesis. The constructs for
synthesizing the 5’ RNA oligos for PCR-based site-specific labeling were prepared by inserting a
BstZ17I site at the 4th position intron 9 in EI9 construct. The primers used were:
EI9BSTZF: gacggagtctggcaggtatacgggccctaagggcaggt;
EI9BSTZR: acctgcccttagggcccgtatacctgccagactccgtc. 118 The WT splicing construct was constructed as described 10. Intronic and exonic mutants were
prepared by PCR-based site-directed mutagenesis with primers containing desired mutations in
the center and complementary sequences on both sides of the point of mutations. E9del1 and
E10del1 were prepared by site-directed mutagenesis using primers whose 3’ half is complementary to the 5’ side of exon 9 or 10 and 5’ half is complementry to the 3’ side of exon
9 or 10. Eswap was constructed by four steps of site-directed mutagenesis: 1. deleting exon 9 from WT to generate E9del1; 2. exon 10 was introduced in the position of exon 9 with two steps.
First, exon 10 was amplified with a forward primer annealing to the 5’ end exon 10 with an overhang that annealed to the intron sequence upstream of exon 9, and a reverse primer annealing to the 3’ end of exon 10 with an overhang that annealed to the intron sequence downstream of exon 9. Second, using the PCR product from step one as primers to introduce exon 10 into the position of exon 9 by site-directed mutagenesis; 3. Remove the original exon 10 as described in 1; 4. Introduce exon 9 into the position of original exon 10 as described in 3.
E9del2 and E10del2 were prepared by removing exon 9 or exon 10 from Eswap by site-directed mutagenesis.
Transfection and splicing assays
We transiently transfected PKM-derived plasmids into HeLa cells with Lipofectamine
2000 (Invitrogen) as described 7. After 24 hours, we isolated total RNA and reverse-transcribed 5
μg RNA, using 0.5 μg of oligo (dT)15 primer and Maxima reverse transcriptase (Fermentas).
Resultant cDNAs were PCR amplified with PKM-specific forward primer 8F and a plasmid- specific reverse primer BGHR. Primer sequences are as follows:
8F, ctgaaggcagtgatgtggcc;
BGHR, tagaaggcacagtcgaggct;
E9F, cgcggatccttcttcttataagtgtttagcagcagct;
T7F, gactcactatagggagaccc;
E10R, cgggatccctgccagacttggtgaggacg; 119 11R, acccggaggtccacgtcctc.
For quantitative PCR assays, we amplified 1 μl of cDNAs in 50 μl of standard reaction
mixtures containing 0.6 μCi [α-32P]dCTP (3,000 Ci/mmol; Amersham, Pittsburgh, PA) for 26
cycles. PCR products were resolved on gels buffered with 4% polyacrylamide and 90 mM
Tris/89 mM boric acid/2 mM EDTA, pH 8.3, visualized by autoradiography and quantitated by using a PhosphorImager (Molecular Dynamics, Piscataway, NJ).
RNA interference
We carried out RNA interference as described 10 with minor modifications. Briefly, we reverse transfected with 25 pmol of hnRNPA1 duplex RNA and 12.5 pmol of the other duplex
RNAs into HeLa cells, cultured in DMEM supplemented with 10% FBs (Hyclone), at 1-2 × 104 cells per well in 24-well plates with lipofectamine RNAi Max reagent (Invitrogen). The next day, siRNA transfection was repeated and cells were split into 12-well plates. siGENOME non- targeting siRNA (Dharmacon) was used as a control to ensure that parallel experiments had equal amounts of RNA. To express splicing constructs in PTB and hnRNP A1/A2 depleted cells, we transfected plasmids after 48 hours of the initial siRNA knockdown. Seventy-two hours after the initial siRNA transfection, we collected cells for total RNA isolation and immunoblotting.
We used the following siRNAs (Dharmacon; the sense strand sequences are given): hnRNPA1,
5′-cagcugaggaagcucuuca-3′; hnRNPA2, 5′-ggaacaguuccguaagcuc-3′; PTB, 5′- gccucaacgucaaguacaa-3′.
UV crosslinking assays and immunoprecipitation
We carried out ultraviolet (UV) crosslinking assays as described in ref. 7, with some
modifications. EI9s and EI10s were synthesized in vitro by T7 RNA polymerase (Promega) with
[32P]-UTP from HpaI-linearized plasmids. We incubated ~10 fmol (1× 105 c.p.m.) of RNA with
2 μl of HeLa NE in 10 μl reaction mixtures. Reaction mixtures were incubated at 30°C for 15 min, and samples were then irradiated with UV light using a Stratalinker (Stratagene), treated
with RNaseA (USB), and proteins resolved by 10% SDS/PAGE. We carried out site-specific 120 labeling as described in ref. 62. The 3’ RNA oligo was purchased from Dharmacon and the
5’RNA oligos was transcribed in vitro with T7 RNA polymerase from BstZ17I-linearized plamids. The 3' oligo fragment was 5'-end-labeled with [ -32P]-ATP and T4 polynucleotide kinase (NEB). The two RNAs were annealed to a DNA bridging oligonucleotide complementary to the 3' end of the 5' RNA fragment and the 5' end of the 3' RNA fragment and ligated using T4
DNA ligase for 4 hours at room temperature. The ligation product was purified following electrophoresis on a 10% denaturing polyacrylamide gel. The sequences of the 3’ oligo and bridging DNA are as follows: 3’ oligo, gggcccuaag; bridging DNA, cccttagggccctacctgccagactccgtcagaactatcaaagc. UV crosslinking-immunoprecipitation was described in ref.10.
Recombinant proteins
hnRNP A1 cDNA and truncations were cloned intothe pGEX6p-1 vector. GST-tagged hnRNP A1, UP1, RRM2-A1 and RRM2 were purified from E. coli using glutathione–Sepharose
4B (GE healthcare), and the GST tag was cleaved off with Prescission protease (GE healthcare).
Purity and concentration of proteins were determined by Coomassie blue staining of SDS gels.
Primers used for cloning hnRNP A1 and truncations were as follows:
hnRNP A1 forward, cggaattcatgtctaagtcagagtctcctaaagagc;
hnRNP A1 reverse, ccgctcgagttaaaatcttctgccactgccatagctac;
Up1 forward, cggaattcatgtctaagtcagagtctcctaaagagc;
Up1 reverse, ccgctcgagtcgacctctttggctggatgaagc ;
RRM2-A1 forward, cggaattcatgtctcaaagaccaggtgcccacttaac;
RRM2-A1 reverse, ccgctcgagttaaaatcttctgccactgccatagtac;
RRM2 forward, cggaattcatgtctcaaagaccaggtgcccacttaac;
RRM2 reverse, ccgctcgagtcgacctctttggctggatgaagc.
Crosslinking-immunoprecipitation. 121 HeLa cells were cross-linked in vivo, and the RNA-binding targets for hnRNP A1 and
PTB were obtained as described (refs, Ule and Darnell, two papers) , with the following
modifications. In brief, 15 cm dishes were cross-linked at 400 mJ cm-2, cells were collected and total cell extracts were prepared in lysis buffer (1xPBS, 0.5% (v/v) Triton-X100). After DNase treatment and a mild digestion with RNase, the extracts were immunoprecipitated for 4 h at 4 °C with the monoclonal antibody 4B10 (Immuquest) coupled to protein A–Sepharose beads (GE) and BB7 (ATCC) bound to protein G-Sepharose beads (GE). After extensive washing with wash buffer (1xPBS, 0.5% (v/v) Triton-X100, 0.1% (w/v) SDS, 0.5% (w/v) sodium deoxycholate for
4B10 and ) and PK buffer (50 mM Tris-HCl (pH 7.5), 50 mM NaCl, 10mM EDTA), the cross- linked RNAs were eluted with 4mg ml-1 Proteinase K (USB) in PK buffer and then PK buffer
containing 7M Urea. RNAs were then precipitated and treated with DNase RQ1 (Promega) and
purified again before subjected to RT-PCR analysis as described before. The primers used in
PCR are as follows:
9f5, gcagcagctttgatagttctgacgg;
9r1; ggctggttatcctaaca;
10f5, tggggccataatcgtcctcacca;
10r1, ccactgagcagggcatt;
In8f1, tgttgtgtctcgtttttttcctcctcc
In8r1, ctcacgagctatctgtaaggtttagg;
I9PTBf1, cacttggtgaaggactggtttctgtgg;
I9PTBr1, gaggagcccaatcactggagattctg,
I9PTBf2, gttcctcaccagctgtccggtga;
I9PTBr2, ggacagagctttgtcagagctttgtc;
In9f1, ggccctaagggcaggtaacac;
In9r1, ggttgggcctggctgtcttc; 122 In9f2, ccagcctcttgctccacctg;
In9r2, gacctccctggcggtgttc;
In9f3, gaccaaagggtcctgtggctc;
In9r3, cagtccttcaccaagtgcctgtg;
In10f1, ggcccaggacattgagttaatg;
In10r1, ccctgtggacagggacaagg;
In10f 2, cccactcttggctgaacagc;
In10r2, gcaagaactggaccttttaagaaaggt;
In10f4, ccagttcttgctgtttggctattttaag;
In10r4, cctgcacttcacgcttggcc;
In10f5, gggcctttatagcccagttgct;
In10r5,ctgggaatcaacatcaggcttctct.
Acknowledgements
We thank T. Kashima for hnRNP A1/A2 siRNA, E. Rosonina for the help with quantitative PCR assays, D. Campigli Di Giammartino for discussion with RNA immunoprecipitation experiment, P. Richard for help with 5’ labeling, and members of the
Manley laboratory for discussions. This work was supported by a grant from the NIH R01
GM048259.
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Figure legends
Figure 1. Mutations of intron 9 sequences derepress exon 9 inclusion. (a) Schematic diagram of PKM splicing construct containing exon 8 to exon 11. “//” indicates deletions of intron 130 sequences. Mutually exclusive AS of exon 9 and exon 10 is indicated. Solid black boxes flanking
exon 9 indicate binding sites for hnRNP A1/A2 and PTB, described previously 10. (b) Schematic
diagram of PKM intron 9. Vertical lines indicate putative hnRNP A1/ A2 (above the line
indicating intron 9) and PTB (below the line) binding sites (BSs). Mutations of BSs are indicated
above or below wild-type (WT) BSs in red. (c) Schematic diagram of splicing construct and
possible products are indicated on the left panel. Black arrows indicate primers used to amplify
PKM AS products. RT−PCR assays of RNA isolated from transient transfections of WT and mutated splicing constructs. The positions of splicing products are indicated on the left. The percentages of DI in total products are indicated under the lane numbers. (d) Bar graphs show percentages of DI/total (left) and SI/total (right) using WT and mutated splicing constructs with standard deviation, n=3. Lane numbers correspond to lane numbers in Figure 1c, and the same lane numbers represent the same constructs. (e) Left panel, scheme indicates the positions of exon 9- and exon 10-specific primers. E9F, which anneals to exon 9, and vector-specific primer
BGHR were used to amplify exon 9-containing products. Vector-specific primer T7F and E10R were used to amplify exon 10-containing products. Right, RT-PCR assays with primers that amplify only exon 9-containing products to analyze splicing products from intron 9-mutated splicing constructs. Splicing constructs are indicated above, and splicing products are indicated on the left. Lane numbers correspond to those in Figure 1c, and the same lane numbers represent the same constructs.
Figure 2. Exonic sequences are involved in PKM alternative splicing regulation. (a) Top panel, diagram of WT construct and Eswap construct, in which the positions of exon 9 and exon
10 are switched. Lower panel, RT-PCR assays of RNAs extracted from transfected HeLa cells, using primers 8F and BGHR (lower left). Bands representing PKM2 are indicated with arrows.
PCR products were digested with Pst I (lower right). (b) Top panel, diagrams indicate splicing constructs with exon 9 deleted from WT (E9del1) and from Eswap (E9del2). RT-PCR assays of 131 RNA isolated from HeLa cells tranfected with indicated plasmids, with primers 8F and BGHR
(lower left). PCR products were digested with Pst I (lower right). PKM splicing products are
indicated on the left. (c) Top panel, diagrams indicate constructs with exon 10 deleted from WT
(E10del1) and from Eswap (E10del2). RT-PCR assays of RNA isolated from HeLa cells
tranfected with indicated splicing constructs with primers 8F and BGHR (lower left). PCR
products were digested with Pst I (lower right). PKM splicing products are indicated on the left.
DS, products that contains only exon 8 and exon 11.
Figure 3. hnRNP A1/A2 bind cooperatively to exonic and intronic elements. (a) UV
crosslinking of EI9s, EI10s and variants in HeLa nuclear extract (NE). Upper panel, diagrams of
UV-crosslinking substrates EI9s, EI10s and variants. ISS1 and the corresponding sequence in
intron 10 are indicated above each RNA. Mutations are indicated with capital letters. Lower
panel, RNA constructs shown in the upper panel were incubated with NE, crosslinked, and
analyzed by SDS-PAGE. The position of hnRNP A1 is indicated on the left. (b) Upper panel,
Schematic diagrams of EI9s and variants. Putative exonic hnRNP A1 binding sites are indicated
as black box and sequence indicated above. TAG was mutated into TAC or TAA as indicated.
Lower panel, UV crosslinking assays with RNAs shown in the upper panel. hnRNP A1/A2
positions are indicated on the left. Quantification of hnRNP A1/A2 binding is indicated below. (c)
Site-specific label (SSL) of EI9s and variants with exonic TAG mutations. Upper panel,
diagrams of EI9s and its variants. Asterisks indicate the nucleotide labeled with 32P. Site- specifically labeled EI9s and mutants were incubated in NE, crosslinked, and analyzed by SDS-
PAGE. hnRNP A1/A2 is indicated on the left. Asterisk, background, non-specific band.
Quantification of the binding of hnRNP A1/A2 is indicated below. (d) Left panel, schematic diagrams indicate hnRNP A1 and truncated derivatives. hnRNP A1, full-length hnRNP A1. UP1, contains the N-terminal RRM1 and RRM2. RRM2-A1, hnRNP A1 with RRM1 deleted. RRM2 contains only the second RRM domain. Right panel, recombinant hnRNP A1 and truncations are 132 purified from E. coli, analyzed on SDS-PAGE, and commassie stained. (e) UV-crosslinking assays with SSL EI9s and variants. RNAs were incubated with 0.3 μM hnRNP A1 (lanes 1-3),
RRM2-A1 (lanes 4-6), Up1 (lanes 7-9) and RRM2 (lanes 10-12), UV crosslinked and analyzed by SDS-PAGE. Quantification of binding is indicated at the bottom.
Figure 4. Exonic hnRNP A1/A2 binding sites are critical for exon exclusion. (a) Left top, schematic indicates E9G3C mutations in exon 9. Right, RT-PCR assays of RNAs isolated from plasmids transfected HeLa cells. PCR products are indicated on the left. Splicing constructs used for transfection are indicated on the top. The percentages of DI in total products are indicated under lane numbers. (b) Bar graphs show percentages of DI and SI with standard deviation calculated from three independent experiments. Lane numbers correspond to lane numbers in
Figure 3a, and the same lane numbers represent the same constructs. (c) RT-PCR with splicing constructs with ESS mutations using exon 9-specific primers. (d) Left panel, diagrams depict splicing constructs. ESSs of exon 9 and corresponding sequences of exon 10 are indicated on top of the exons. Right panel, RT-PCR assays of RNAs extracted from HeLa cells transfected with
WT (lanes 1, 4, 7), Eswap (lanes 2, 5, 8) and ESSMu (lanes 3, 6, 9). PCR products were digested with Pst I (lanes 4-6) or Tth111 I (lanes 7-9). Splicing products are indicated with arrows.
Figure 5. . Full-length intron 10 is required for exon 10 exclusion. (a) RT-PCR assays with
T7 and 11R primers. Splicing products are indicated on the left. In8, the intron 8 sequences that were not included in WT. In10, the rest of intron 10 sequence that is not included in WT. (b) RT-
PCR assays with exon-specific primers. Splicing products are indicated on the left.
M1/(M1+DI)% represents the percentage of M1 in all exon 9 included products. (c) Lanes 1-4,
PCR with 8F and BGHR primers to amplify M1, M2 and DI products and DI/total% is indicated below. Lanes 5-8, exon 9-specific primers, the percentages of M1 in all exon 9 included products are indicated below. (d) RT-PCR assays with exon 9-specific primers and total RNA extracted 133 from HeLa cells transfected with WT, WT-In8-In10, WT-In10, WT-In10 1/2 or WT-In10 2/2 in
either control or hnRNP A1/A2 and PTB depleted conditions. Splicing products are indicated on
the left. M1/(M1+DI)% represents the percentage of M1 in all exon 9 included products. (e) RT-
PCR assays with exon 9-specific primers and total RNA extracted from HeLa cells transfected
with WT, WT-In10, WT-In10 1/3, WT-In10 2/3 or WT-In10 3/3 in either control or hnRNP
A1/A2 and PTB depleted conditions. Splicing products are indicated on the left. M1/(M1+DI)%
represents the percentage of M1 in all exon 9 included products.
Figure 6. Two mechanisms prevent double inclusion PKM mRNA when hnRNP A1/A2 and
PTB levels are low. (a)-(f) hnRNP A1 and PTB bound in intron 9 and intron 10 loop out exon 10 when
their levels are reduced. (a) ) Left panel, scheme indicates the positions deletions in intron 9. Right
panel, RT-PCR assays with exon 9-specific primers. Splicing products are indicated on the left.
(b) RT-PCR assays with exon 9-specific primers and total RNA extracted from HeLa cells
transfected with WT, WT-In10, A1Mu1-4, A1Mu1-4-In10, PTBMu and PTBMu-In10 in either control or hnRNP A1/A2 and PTB depleted conditions. Splicing products are indicated on the
left. (c) RT-PCR assays with primers 8F and BGHR (top panel) and exon 9-specific primers
(lower panel) and total RNA extracted from HeLa cells transfected with WT-In10, A1Mu1-4-
In10 and A1Mu5+6-In10 in either control or hnRNP A1/A2 and PTB depleted conditions.
Splicing products are indicated on the left. (d) Scheme indicates the positions of amplicons by primers used in CLIP assays. (e) CLIP of hnRNP A1 in HeLa cells treated with control or hnNRP
A1/A2/PTB siRNAs. Primers used are indicated under each column. Immunoprecipitations were performed using antibody against hnRNP A1 (4B10). The precipitated RNAs were subjected to RT-PCR.
Data show the mean +/- S.D. of triplicates from three independent experiments. (f) CLIP of PTB in HeLa cells treated with control or hnNRP A1/A2/PTB siRNAs. Primers used are indicated under each column.
Immunoprecipitations were performed using antibody against PTB (BB7). The precipitated RNAs were subjected to RT-PCR. Data show the mean +/- S.D. of triplicates from three independent experiments (g)
PKM double inclusion mRNA is subject to NMD. RT-PCR with RNA extracted from HeLa cells
treated with cycloheximide (as indicated) after PTB and hnRNP A1/A2 were depleted by 134 siRNAs (as indicated), using primers 8F and 11R to amplify endogenous PKM. PCR products were resolved in 1.5% agarose gel and stained with ethidium bromide. Products are indicated on the left.
Figure 7. A model for PKM mutually exclusive splicing. (a) PKM2 mRNA production. In the presence of elevated levels of hnRNPA1/A2 and PTB, hnRNP A1/A2 bind cooperatively to two
ESS elements in exon 9 and ISS1 in intron 9, excluding exon 9 (WT). PTB and additional hnRNPA1/A2 molecules bind to both intron 8 (PTB only) and intron 9. This further inhibits exon
9 inclusion, through formation of an inhibitory network of hnRNPs. The ESSs play a critical role, as when the positions of exon 9 and 10 are swapped, hnRNP A/A2 still binds to exon 9, enabling hnRNP A1/A2 to bind to intron 10, and exon 9 is still excluded (Eswap). Indeed the ESSs are sufficient, to induce exclusion, as when they are artificially created in exon 10, they repress exon
10 inclusion (ESSMu). (b) PKM1 mRNA production. In the absence of high levels of hnRNPA1/A2 and PTB, exon 9 is efficiently included because the reduced binding of hnRNP proteins flanking exon 9. Exon 10 is partially excluded by sequences bound by hnRNP A1/A2 and PTB in intron 9 and full-length intron 10. Unlike with exon 9 in the presence of the hnRNPs, exon 10 exclusion is not complete. Therefore, double inclusion mRNA containing both exon 9 and exon 10 is produced, but it is degraded by NMD.
Supplementary Figures
Supplementary Figure 1. RT-PCR assays with primers 8F and BGHR and total RNA extracted from HeLa cells transfected with splicing constructs indicated above. DI/total% is indicated below.
Supplementary Figure 2. Western blotting with antibodies against PTB, hnRNP A1 and hnRNP A2 to estimate siRNA depletion efficiency. Actin was used as loading control. Lane 1, lysate from control siRNA transfected HeLa cells. Lane2, lysate from triple siRNA transfected 135 HeLa cells. Depletion efficiencies of each hnRNP protein is indicated below each panel, with the control siRNA-treated sample set as 1.
Supplementary Figure 3. Intron 8 does not play a role in reducing double inclusion product.
RT-PCR assay was performed with E9-specific primers and total RNA extracted from HeLa
cells transfected with WT, WT-In8-In10 or WT-In10. Lanes 1-3, HeLa cell treated with control
siRNA. Lanes 4-6, HeLa cells treated with PTB, hnRNP A1 and hnRNP A2 siRNAs. Splicing
products are indicated on the left.
Supplementary Figure 4. RT-PCR assays with primers 8F and BGHR and total RNA extracted
from HeLa cells transfected with WT, WT-In10, A1Mu1-4, A1Mu1-4-In10, PTBMu and
PTBMu-In10 in either control or hnRNP A1/A2 and PTB depleted conditions. Splicing products
are indicated on the left.
Supplementary Figure 5. RT-PCR assays with exon 9-specific primers and total RNA extracted
from HeLa cells transfected with A1Mu1-4-In10, PTBMu-In10 and A1+PTBMu-In10 in either
control or hnRNP A1/A2 and PTB depleted conditions. Splicing products are indicated on the
left.
Supplementary Figure 6. UV-crosslinking-Immunoprecipitation of intron9 truncations Del1,
Del2, Del3 in HeLa NE. RNA oligos used are indicated on top. (a) Scheme indicates the
positions of oligos used in UV-crosslinking experiment. (b) UV-crosslinking experiment using
RNA oligos indicated in (a) and PTB antibody (BB7). PTB is indicated on the left. (c) UV-
crosslinking experiment using RNA oligos indicated in (a) and hnRNP A1 antibody (4B10).
hnRNP A1 is indicated on the left
136 Supplementary Figure 7. RT-PCR assays with [α-32P]dCTP and RNAs from CLIP experiments using hnRNP A1 antibody (4B10) as describe in Figure 6e. Primers used are indicated on the left.
Supplementary Figure 7. RT-PCR assays with [α-32P]dCTP and RNAs from CLIP experiments using PTB antibody (BB7) as describe in Figure 6f. Primers used are indicated on the left.
137
a PTB hnRNP A1/A2 TCTT...TCTT TAGGG
8 9 10 11 b
hnRNP A1BSs ISS1 2 3 4 5 6 Mutants TACGG TACGA TACGA TACGA TACGC TACAT WT TAGGG TAGGA TAGGA TAGGA TAGGC TAGAT 9 10 TCTT PTB BSs WT TCTT TCTT TCTT TCTT TCTT TCCCTCTCCTTTT TGTT
PTBMu TGTT TGTTTGTT TGTT TGTTTGGGTGTGGTTTT
uMBTP+1A
6-2uM1A 4-1uM1A
c 6-1uM1A uMBTP
8F TW
8 9 10 11
T7 promoter BGHR DI SI
double inclusion product (DI) 8 9 10 11
PKM1 8 9 11 single inclusion products (SI) or PKM2 8 10 11 1 2 3 4 5 6 DI/total % 3.0 66 12 75 66 28 SI/total % 97 34 88 25 34 72
[ 8F+BGHR ] d e E9F
8 9 10 11 BGHR DI/total% T7F 90 8 9 10 11 80 70 E10R
60 uMBTP+1A
50 6-2uM1A
4-1uM1A 6-1uM1A
40 uMBTP 30
TW 20 10 0 1 2 3 4 5 6 DI
SI/total% 120 1 2 3 4 5 6 100 [ E9F+BGHR ] 80
60 40 DI 20
0 1 2 3 4 5 6 M2
1 2 3 4 5 6 [ T7F+E10R ] Figure-1 (Manley) 138
a WT 8 9 10 11
Eswap 8 10 9 11
WT
WT
Eswap Eswap
DI SI
M2
1 2 3 4 uncut Pst I b c
WT 8 9 10 11 WT 8 9 10 11
E10del1 8 9 11 E9del1 8 10 11
E10del2 8 9 11
E9del2 8 10 11
E9del1
WT
WT
E9del1 E9del2 WT
WT
E10del1
E10del1
E10del2
E10del2 E9del2
DI M2 M1 M1 DS
M2 M2
1 2 3 4 5 6 1 2 3 4 5 6 Pst I uncut uncut Pst I Figure-2 (Manley) 139
a ESS1 ESS2 b 12 3 TAG TAG ISS1 ISS1-10 gtagggc gtaggag EI9s EI9s 9 EI10s 10 TAC TAG TAA TAG EI9sE9G3C1 EI9sE9G3A1 TAC TAC TAA TAA gtaggAc gtaggGg EI9sM1 9 EI10sM1 10 EI9sE9G3C2 EI9sE9G3A2
gtaggAG gtaggGC EI9sM2 9 EI10sM2 10
EI9sE9G3C1
EI9sE9G3C2 EI9s
EI9sE9G3A1
EI9sE9G3A2 EI9s
EI10s
EI9sM2 EI9s
EI10sM1 EI9sM1
EI10sM2
hnRNP A1/A2 hnRNP A1/A2 hnRNP A1/A2
1 2 3 4 5 6 Relative 1 2 3 Relative 4 5 6 binding 100 25 50 binding 100 25 50 c d
Up1
hnRNP A1 RRM2-A1
RRM2 ESS1 ESS2 12 3 hnRNP A1 RRM1 RRM2 C-Ter TAG TAG * gtagggc 50 EI9s Up1 RRM1 RRM2 TAC TAG * 50 37 gtagggc EI9sE9G3C1 RRM2-A1 RRM2 C-Ter 37 25 TAC TAC * gtagggc RRM2 EI9sE9G3C2 RRM2 25
EI9sE9G3C2 EI9s
EI9sE9G3C1
e hnRNP A1 RRM2-A1 Up1 RRM2 EI9s
EI9sE9G3C2
EI9s EI9s EI9s
EI9sE9G3C1
EI9sE9G3C2 EI9sE9G3C2 EI9sE9G3C2
EI9sE9G3C1
EI9sE9G3C1 EI9sE9G3C1 hnRNP A1/A2
1 2 3 Relative binding 100 25 19
1 2 3 4 5 6 7 8 9 10 11 12 Relative binding 100 63 38 100 19 6.1 100 23 18 NA site-speci c label
Figure-3 (Manley) 140
a b TAG TAG DI/total% 120 WT 8 9 10 11 100
TAC TAC 80
E9G3C 8 9 10 11 60 40
20
0
1 2 3 4
A1MuE9G3C A1Mu1-4
WT E9G3C c DI
SI
WT
A1Mu1-4
A1Mu+E9G3C E9G3C
1 2 3 4 DI DI/total 6.1 90 69 100 M1
1 2 3 4 [ E9F+BGHR ]
d
WT
WT
ESSMu
Eswap
Eswap ESSMu
TAG TAG GCA TCG
WT 8 9 10 11 SI M1
GCA TCG TAG TAG
Eswap 8 10 9 11
GCA TCG TAG TAG M2
ESSMu 8 9 10 11
1 2 3 4 5 6 [ 8F +BGHR ]
uncut Pst I
Figure-4 (Manley) 141
a c
WT
WT-In8-In10
WT WT-In8-in10 Triple KD - + - + - + - + WT A1Mu1-4 WT-In8 WT-In8-In10 A1Mu1-4-In8-In10 A1Mu1-4-In8 DI DI DI DI SI M1 M1 SI
1 2 3 4 5 6 7 8 1 2 3 4 5 6 DI/total% 9.0 90 0 28 M1/(M1 +DI)% 1.7 4.1 NA 40 + + + 1.5 +- 6.4 - 0 - 8.5 + [ 8F+BGHR ] - +- 1..1 - 1.4 +- 4.3 [ 8F+BGHR ] [ E9F+BGHR ]
b d
WT-In10 2/2
WT-In10 1/2
WT-In10
WT
WT-In10 2/2
WT-In10 1/2
WT-In10
WT
WT-In8-In10 WT-In8-In10 E9G3C E9G3C-In8 E9G3C-In8-In10 E9G3C-In10 Triple KD - - - - - + + + + +
DI M1
1 2 3 4 M1/(M1 +DI)% 8.4 2.4 13 23 1 2 3 4 5 6 7 8 9 10 M1/(M1 +DI)% 3.1 35 38 18 5.1 [ E9F + BGHR ]
e
WT
WT
WT-In10 2/3
WT-In10 3/3
WT-In10 1/3
WT-In10 3/3
WT-In10 2/3
WT-In10 1/3
WT-In10 WT-In10 Triple KD - - - - - + + + + +
DI M1
1 2 3 4 5 6 7 8 9 10 M1/(M1 +DI)% 1.4 26 9.7 3.0 4.4 [ E9F + BGHR ]
Figure-5 (Manley) 142
a
WT-In10
WT-In10-In9Del3
WT-In10-In9Del1 WT-In10-In9Del2
WT 9 10 WT c
WT-In10
A1Mu5+6 -In10 A1Mu5+6
WT-In10
A1Mu5+6 -In10 A1Mu5+6 A1Mu1-4 -In10 A1Mu1-4 Triple KD + + + + + -In10 A1Mu1-4 Triple KD - - - + + + In9Del1 9 10 DI DI In9Del2 9 10 M1 SI In9Del3 9 10 1 2 3 4 5 [ E9F+BGHR ] 1 2 3 4 5 6 DI/total% 5.3 76 12 28 76 68 [ 8F+BGHR ]
b
WT-In10
WT-In10
WT
A1Mu1-4-In10
A1Mu1-4
PTBMu -In10 PTBMu PTBMu
WT
A1Mu1-4 -In10 A1Mu1-4 A1Mu1-4
PTBMu -In10 PTBMu PTBMu
Triple KD ------+ + + + + +
WT-In10
A1Mu5+6 -In10 A1Mu5+6
WT-In10
A1Mu5+6 -In10 A1Mu5+6
A1Mu1-4 -In10 A1Mu1-4 A1Mu1-4 -In10 A1Mu1-4 Triple KD - - - + + + DI
M1 DI M1 1 2 3 4 5 6 7 8 9 10 11 12 7 8 9 10 11 12 [ M1/(M1 E9F+BGHR ] +DI)% 67 27 29 [ E9F+BGHR ]
d In8f1r1 9f5r1 In9f2r2 In9PTB f1r1 10f5r1 In10f1r1 In10f4r4 In10f5r5
PKM 9 10
In9f1r1 In9f3r3 In9PTBf2r2 In10f2r2
e f hnRNP A1 binding PTB binding 1.4 4 1.2 3.5 1 3 0.8 2.5 0.6 control 2 control KD 0.4 KD 1.5
0.2 1
0 9f5r1 10f5r1 In9f1r1 In9f2r2 In9f3r3 In10F1r1 In10f2r2 In10f5r5 0.5 0 In8f1r1 In9PTBf1r1a In9ptbf2r2 IN10f1r1 In10f2r2 In10f4r4
cycloheximide - + - + Triple KD - - + + g * DI
SI
1 2 3 4 [ 8F+11R ] Figure-6 (Manley) 143
a
PKM2 production A1/A2 U1 PTB PTB U2 PTB A1/A2 A1/A2A1/A2 PTB A1/A2PTB A1/A2PTB PKM WT 8 9 10 11
PTB PTB At high expression levels A1/A2A1/A2 8 10 11
b
A1/A2 PTB PTB PKM1 production 8 9 A1/A2 PTB 10 A1/A2PTBA1/A2 11
PTBPTB At low expression levels
A1/A2A1/A2
U1 PTC U2 8 9 11 A1/A2 A1/A2 8 9 10 11 PTB PTB DI product 10
8 9 11 Degraded by NMD PKM1
Figure-7 (Manley) 144
WT
A1Mu1-4 A1Mu1
DI
SI
1 2 3 DI/total % 7.4 46 75 SI/total % 93 54 25 [ 8F+BGHR ]
Supplementary Figure-1 (Manley) 145
siRNA control Triple
PTB
1 0.05 hnRNP A1
1 0.25 hnRNP A2
1 0.07
Actin
Western blotting
Supplementary Figure-2 (Manley)
146
01nI-8nI- TW 01nI-8nI-
01nI-8nI- TW 01nI-8nI-
8nI- TW 8nI-
8nI- TW 8nI-
TW TW
Triple KD - - - + + +
DI
M1
1 2 3 4 5 6 M1/(M1 +DI)% 2.6 27 1.4 [ E9F + BGHR ]
Supplementary Figure-3 (Manley)
147
WT
WT
WT-In10
A1Mu1-4
PTBMu A1Mu1-4 -In10 A1Mu1-4
PTBMu -In10 PTBMu
WT-In10
A1Mu1-4
PTBMu A1Mu1-4 -In10 A1Mu1-4 PTBMu -In10 PTBMu Triple KD ------+ + + + + +
DI SI
1 2 3 4 5 6 7 8 9 10 11 12
[ 8F+BGHR ]
Supplementary Figure-4 (Manley)
148
A1Mu -In10 A1Mu
PTBMu -In10 PTBMu A1+PTBMu -In10 A1+PTBMu
PTBMu -In10 PTBMu
A1Mu -In10 A1Mu A1+PTBMu -In10 A1+PTBMu
Triple KD - + - + - +
DI M1
1 2 3 4 5 6
[ E9F+BGHR ]
Supplementary Figure-5 (Manley) 149
a
WT 9 10
In9Del1 In9Del2 In9Del3 b In9Del1 In9Del2 In9Del3 IP sup IP sup IP sup Mock PTB Mock PTB Mock PTB Mock PTB Mock PTB Mock PTB
PTB PTB
1 2 3 4 5 6 7 8 9 10 11 12
c In9Del1 In9Del2 In9Del3 sup IP sup IP sup IP A1 A1 A1 A1 A1 Mock Mock Mock Mock A1 Mock Mock
A1 A1
1 2 3 4 5 6 7 8 9 10 11 12
Supplementary Figure-6 (Manley) 150
Input 4b10
Rnase A 1:5k 1:5k Triple KD - + - +
9f5r1
10f5r1
In9f1r1
In9f2r2
In9f3r3
In10f1r1
In10f2r2
In10f5r5
Supplementary Figure-7 (Manley) 151
Input BB7
Rnase A 1:5k 1:5k Triple KD - + - +
In8f1r1
In9PTBf1r1
In9PTBf2r2
In10f1r1
In10f2r2
In10f4r4
1 2 3 4
Supplementary Figure-8 (Manley) 152
Chapter 5: Phosphorylation switches the general splicing repressor SRp38 to a sequence-specific activator.
Mo Chen designed and performed the experiments in Figure 3 and Figure 4
153
Phosphorylation switches the general splicing repressor SRp38 to a sequence-
specific activator
Ying Feng, Mo Chen and James L. Manley
Department of Biological Sciences
Columbia University
New York, NY, 10027
154 Abstract
SRp38 is an atypical SR protein that functions as a general splicing repressor when
dephosphorylated. We now show that phosphorylated SRp38 can function as a splicing activator with RNA substrates that contain high-affinity SRp38 binding sites. Unlike characterized splicing activators, SRp38 functions in the absence of other SR proteins, but requires a specific cofactor for activity. Unexpectedly, SRp38 was sufficient to induce formation of splicing complex A in the absence of the cofactor, which was necessary for progression to complexes B and C. Mechanistically, we provide evidence that SRp38 function involves strengthening the ability of the U1 snRNP to stably recognize the 5′ splice site and U2 snRNP to recognize the branch site. Given these properties of SRp38, we hypothesized that the protein might regulate specific alternative splicing events in vivo. Indeed, analysis of alternative splicing of pre-mRNA encoding the glutamate receptor B in vitro and in vivo revealed that SRp38 alters its splicing pattern in a sequence-specific manner. Together, our data demonstrate that SRp38, in addition to its role as a global splicing repressor, can function as an unusual sequence-specific splicing activator.
1
155 Introduction
Alternative splicing is a common mechanism for regulating gene expression and
increasing protein diversity in metazoan organisms1. In humans, greater than 70% of primary
transcripts are estimated to undergo alternative splicing2. Splicing is carried out in the
spliceosome, in which five small nuclear ribonucleoprotein particles (snRNPs; U1, U2, U4/U6,
and U5) and a large number of auxiliary proteins cooperate to accurately recognize the splice
sites and catalyze the two steps of the splicing reaction3.
The inclusion of a specific exon in the mature mRNA is largely dependent upon the
recognition and usage of the flanking splice sites by the splicing machinery4. This appears to be
governed by the dynamic formation of protein complexes on the pre-mRNA. Specific sets of
splicing regulatory proteins assemble on different pre-mRNAs, generating a "splicing-" or
"mRNP”-code that determines exon recognition5,6. Cis-elements known as exonic splicing
enhancers (ESEs) modulate the assembly of regulatory proteins on pre-mRNA and therefore
contribute to splice site choice. The majority of characterized ESEs have been found adjacent to
introns containing (a) weak splice site(s), despite the finding that constitutive exons are also
frequently enriched in potential binding sites for splicing regulatory proteins7. ESEs were
initially identified as purine-rich sequences8-10, but different motifs have been identified by different strategies11,12.
ESEs are often recognized and bound by SR proteins, a family of highly conserved
splicing factors that play key roles in spliceosome activation and in regulation of splice site
selection13-15. SR proteins contain one or two N-terminal RNP-type RNA binding domains (RBD)
and a C-terminal arginine- and serine-rich (RS) domain. The RBDs of SR proteins are necessary
and sufficient for RNA binding, while the RS domains are involved in protein-protein16,17 and
2
156 perhaps protein-RNA18 interactions. SR proteins function as essential general splicing factors,
necessary for spliceosome assembly in in vitro assays13,15,19.
Many studies have identified ESEs that activate 3’ or 5’ splice sites by binding SR proteins20. The ESE-bound SR proteins can stimulate splicing by recruiting the general splicing
factor U2AF to weak polypyrimidine tracts via a direct interaction between the RS domains of
the SR proteins and U2AF21,22. They are also involved in bridging interactions between ESEs
and spliceosomal components, which is likely mediated by the SRm160 and SRm300 splicing
coactivators23,24. These co-activators contain RS domains but without RBD domains, and can
form multiple interactions with snRNPs and enhancer-bound SR proteins. In addition, a series of
sequential RS domain-RNA contacts at the branch point and the 5’splice site during splicing
complex assembly have been documented, suggesting that RS domain-RNA interactions might
also contribute to splicing activation by ESE-bound SR proteins18. ESEs also activate 5’ splice
sites, as exemplified by fruitless pre-mRNA splicing in Drosophila25.
We have previously described an unusual member of the SR protein family, SRp38, which functions differently from standard SR proteins. Although the domain organization of
SRp38 is typical of SR proteins, SRp38 is unable to activate splicing in standard in vitro assays.
In contrast to other SR proteins, SRp38 functions as a general splicing repressor26,27. Its repression activity is turned on by tightly regulated dephosphorylation28 and it is required for
global inhibition of splicing both in M phase of the cell cycle and following heat shock29.
Although present at high levels in a number of cell types and tissues, no function has as yet been assigned to phosphorylated SRp38.
In this report, we provide a function for phosphorylated SRp38 by showing that it functions as a sequence-specific splicing activator in in vitro assays. We show that, unlike other
3
157 characterized sequence-specific regulators, such as Tra-230, SRp38 does not require other SR proteins to function, but unlike typical SR proteins, requires a nuclear cofactor for activity.
Remarkably, SRp38 is sufficient to induce formation of spliceosomal complex A in a cell extract lacking SR proteins (S100), but the complexes are stalled and require the cofactor to progress to
active splicing complexes. By analysis of alternative splicing of a specific pre-mRNA in vitro
and in vivo, we demonstrate that SRp38 can affect the selection of mutually exclusive exons in a
sequence-specific manner, reflecting affinity for SRp38. We therefore conclude that SRp38 is a
novel type of splicing factor capable of switching from a general repressor to a sequence-specific
activator and regulator of alternative splicing.
Results
SRp38 functions as a sequence-specific activator of splicing.
Previous work from our lab has shown that SRp38 functions as a general splicing repressor when dephosphorylated. Although that work showed that phosphorylated SRp38 cannot function like other SR proteins, as a general splicing activator27,29, we decided to test whether it might be able to function as a sequence-specific activator. To this end, we took advantage of the high-affinity binding of SRp38 to its consensus recognition sequence
(AAAGACAAA), previously determined by SELEX27, and constructed a modified β-globin pre- mRNA substrate in which three copies of this sequence were used to replace sequences in the downstream exon (designated β-SRp38, Fig. 1a). In parallel, we also constructed a control substrate in which random sequences of the same size were inserted into the same position as the
SRp38 consensus sequence (Fig. 1a). We then tested whether phosphorylated his-tagged SRp38, purified from recombinant baculovirus-infected insect cells27, could activate splicing of the β-
4
158 SRp38 RNA in Hela S100 extract. However, no splicing was detected (Fig. 1b, lanes 3-4), even
when high concentrations of SRp38 were used (see below).
One explanation for the inactivity of SRp38 in the above assay is that a cofactor not
present in S100 is required for activity. To test this possibility, we prepared ammonium sulfate
fractions of Hela nuclear extract (NE), and tested these together with SRp38 and S100. Strikingly,
in the presence of a 40-60% saturation cut of NE (NF40-60), strong activation by SRp38 was
observed with the β-SRp38 but not the β-control RNA (Fig. 1b, lanes 5-6, compare with lanes
11-12). Notably, as shown in Fig. 1b, activation of splicing was dose-dependent, with significant
splicing observed with as little as 20 ng of purified his-tagged SRp38. These results indicate that
three copies of the SRp38 binding sequence can function as an SRp38-specific ESE, and that
SRp38 can indeed function as a splicing activator.
Characteristics of SRp38-dependent splicing activation.
We next tested what properties of the SRp38 RS region affect the protein’s activity in
splicing. Purified GST derivatives of SRp38, a splice variant, SRp38-2, containing a short
version of the RS domain27, SRp38 RBD, lacking the whole RS domain, and dephosphorylated
SRp38 (dSRp38) were each added to splicing reactions with S100 or S100 plus NF40-60 and the
β-SRp38 RNA (Fig. 2a). While SRp38 RBD was completely inactive (lane 7), splicing activation
was observed with SRp38-2, again in a manner dependent on the presence of NF40-60, but
splicing efficiency was significantly reduced compared to full-length SRp38 (lane 5, compare to
lane 3). Significantly, dSRp38 was unable to activate splicing of the β-SRp38 RNA (lane 9). In
fact, splicing activated by 20 ng phosphorylated SRp38 was completely repressed by very low
amounts of dSRp38 (2-4 ng; Fig. 2b, lanes 1-4), consistent with its previously described
5
159 properties as a general splicing repressor27. These results indicate that the phosphorylated RS
domain of SRp38 is required for splicing activation. (The phosphorylation status of both SRp38
and dSRp38 did not change during the splicing reaction; Supplementary Fig. 1 online). Thus, while phosphorylation/dephosphorylation might serve as an on/off switch for standard SR proteins31, its function in the case of SRp38 is to switch from splicing activation to repression.
We next wished to investigate the specificity that the SRp38 RBD provides in
recognizing the β-SRp38 RNA. To this end, we added purified GST-SRp38 RBD, His-SC35
RBD and GST-hnRNP G RBD to β-SRp38 RNA-containing splicing reactions in S100 plus
SRp38 and NF 40-60. The SC35 RBD shares the highest identity with the SRp38 RBD (46%),
while the hnRNP G RBD displays only limited identity. The results (Fig. 2c) show that the
SRp38 RBD, but not the SC35 or hnRNP G RBDs, inhibited splicing brought about by SRp38
plus NF40-60. Notably, the amount of SRp38 RBD sufficient to block splicing was roughly
equivalent to the amount of SRp38 added. The inhibition of splicing thus reflects a specific,
competitive interaction between full-length SRp38 and the SRp38 RBD, providing evidence that
the SRp38 RBD-RNA interaction is indeed highly specific.
Next we examined the possible role of standard SR proteins in SRp38 activity. This was
tested by determining whether splicing of the β-SRp38 RNA in S100 activated by increasing
amounts of purified SR proteins could be enhanced by SRp38 (Fig. 2d). The results show that
SRp38 was not able to function with the SR proteins to increase splicing beyond the levels
provided by the SR proteins alone (compare lanes 2-4 with 5-7), indicating that SR proteins
cannot provide the coactivator activity present in NF40-60.
SRp38 is distinct from other SR proteins in requiring a nuclear fraction to activate
splicing in S100. To investigate this requirement further, we examined how NF40-60 influences
6
160 the activity of standard SR proteins. When ASF/SF2 and SC35 (50 ng) were added to splicing
reactions containing the β-SRp38 RNA, splicing was also enhanced by NF40-60, but relatively
weakly (Fig. 2e, lanes 1-7). Highlighting the difference between SRp38 and other SR proteins,
ASF/SF2 and SC35 plus NF40-60 gave rise to comparable levels of splicing with the β-control
RNA, while SRp38 was completely inactive (Fig. 2e, lanes 8-14). Given that classical SR
proteins typically function in S100 in the absence of a nuclear fraction, we next asked whether
higher levels of all three proteins (300 ng) could activate splicing of β-SRp38 (Fig. 2f) or β- control (data not shown) RNA in S100 without NF40-60. Significantly, while ASF/SF2 and
SC35 activated splicing of both substrates, SRp38 was entirely inactive. These results indicate that activation by SRp38, unlike that by other SR proteins, is entirely dependent on a coactivator.
SRp38 facilitates the formation of spliceosomal complex A.
We next wished to gain insight into the mechanism by which SRp38 stimulates splicing.
To this end, we first performed spliceosome assembly assays utilizing the β-SRp38 RNA and
S100 extract alone or S100 containing SRp38 in the presence or absence of NF40-60 (Fig. 3a).
Unexpectedly, the results of a time course showed that SRp38 in the absence of NF40-60 was able to bring about very efficient assembly of what appears to be spliceosomal complex A (Fig.
3a, lanes 10-12). It was especially striking that nearly the entire non-specific H complex was
converted to the A-like complex by SRp38 alone, which is atypical and considerably more
efficient than when splicing was in fact activated by SRp38 plus NF40-60 (lanes 14-16). No
complex was detected in reactions containing β-SRp38 RNA with S100 (lanes 2-4), with NF40-
60 (lanes 6-8) alone, or with dSRp38 (Supplementary Fig. 2a online). These results suggest that
7
161 phosphorylated SRp38 promotes an early step in the splicing pathway by forming early
spliceosomal complexes.
We next wished to confirm the identities of the complexes described above, as well as
extend our findings to a second RNA substrate. To this end, we constructed a modified
adenovirus major late substrate RNA (AdML-SRp38) containing three copies of the SRp38
binding sites in its second exon. SRp38 gave rise to the same sequence-dependent splicing
activation with this RNA as observed with β-SRp38 RNA, and again required NF40-60 for
activity (Fig. 3b). We then examined the effect of SRp38 on spliceosome assembly with the
AdML-SRp38 RNA. As shown in Fig. 3c, efficient formation of a spliceosomal complex was
observed in S100 supplemented with SRp38 alone (lanes 2-3) but not in S100 (lanes 8-9) or with
SRp38 alone (lanes 11-12). Importantly, the mobility of the complex was identical to the A
complex observed in NE (compare lanes 2-3 with 5-6). Significantly, B and C complexes were
again not detected (lanes 1-3; longer exposure in lanes 13-15). To provide evidence that the
apparent stalled A complex could give rise to B and C complexes in the presence of NF 40-60,
we performed spliceosome assays in which NF 40-60 was added to reaction mixtures containing
SRp38 plus S100. Significantly, B and C complexes were formed in addition to A complex
(lanes 16-18). These results suggest that SRp38 promotes formation of spliceosomal A complexes, but these are stalled in the absence of NF40-60.
The above experiments provide evidence that SRp38 can facilitate A complex formation
in S100. To provide additional evidence that these complexes indeed correspond to A complex,
we first analyzed whether snRNPs were involved in their formation. For this, we utilized
antisense RNA oligos complementary to U1, U2, U5 and U6 snRNAs to test which if any of
these could inhibit complex formation in S100 plus SRp38. As shown in Fig. 3d, complex
8
162 formation with the AdML-SRp38 RNA was efficiently blocked by antisense oligos against U1
(lane 2) and U2 (lane 3) snRNAs but not against U5 (lane 4) or U6 (lane 5) snRNAs, indicating
that U1 and U2 snRNPs were involved in complex formation, consistent with the properties of A
complex. In addition, depletion of ATP (lane 6), lack of Mg2+ (lane 7) or incubation of splicing
reactions on ice (lane 8) completely inhibited complex formation, ruling out the possibility that the observed complex might be related to the ATP-independent E complex. We employed the same strategy to characterize the complex formed on the β-SRp38 substrate (Fig. 3a).
Importantly, the anti-U1 and anti-U2 oligos blocked complex formation, while the anti-U5 oligo did not, indicating that A complex but not B complex was formed on this substrate
(Supplementary Fig. 2b online).Together, these results demonstrate that SRp38 facilitates formation of A complex in splicing competent S100 extracts, but the complex is stalled in the absence of a cofactor.
SRp38 strengthens 5’ splice-site and branch site recognition by U1 and U2 snRNPs in a cooperative manner.
We next investigated the mechanism by which SRp38 facilitates complex A formation.
One possibility is that it enhances interactions of U1 and/or U2 snRNPs with the pre-mRNA. To investigate this, we carried out gel-shift assays with purified U1 and U2 snRNPs and SRp38 with the β-SRp38 RNA used in splicing (Fig. 4). As expected, SRp38 bound to the RNA tightly (Fig.
4a, lane 3), while under the conditions used17, U1 snRNP alone did not interact with β-SRp38
mRNA (lane 2). However, the presence of SRp38 stimulated formation of a stable (heparin-
resistant) complex of U1 snRNP, SRp38 and RNA (lanes 4-5). The SRp38 RBD, while able to
form an RNA-protein complex by itself, was unable to facilitate formation of a U1 snRNP
9
163 ternary complex (lanes 6-9), indicating that, as with splicing activation, the RS-domain is
necessary for complex formation. Importantly, the SRp38-U1 snRNP-RNA complex only
formed with β-SRp38 mRNA, and not the β-control mRNA (lanes 10-14). A similar cooperative
interaction was observed between SRp38 and U2 snRNP (Fig. 4b). These data suggest that
SRp38 facilitates U1 and U2 snRNPs association with the pre-mRNA to stabilize 5’ splice-site
and branch site recognition, respectively. We also performed gel shift assays using the β-SRp38
RNA with ASF/SF2. No complex was observed when 100 ng of ASF/SF2 was added to reaction
mixtures, in the presence or absence of snRNPs (Fig. 4c). These data indicate that SRp38, in a
binding-site dependent manner, can facilitate recruitment of U1 and U2 snRNPs with a pre-
mRNA.
We next set out to investigate whether the SRp38 interactions with U1 and U2 snRNPs
are functionally significant. To this end, we utilized a splicing inhibition assay designed to
measure, albeit indirectly, recruitment of U1 and U2 snRNPs to the 5’ splice site and branchsite,
respectively, during splicing. The assay measures the sensitivity of splicing to inhibition by
antisense RNA oligos. For example, an RNA oligo containing a polypyrimidine stretch was
shown to block the U2 snRNP-branchsite interaction by competing with the 3’ splice site for
binding to U2AF32. We predicted that if the interactions between SRp38 and U1/U2 snRNPs
were functionally relevant, then SRp38-activated splicing might show greater resistance to RNA
oligos that interfered with 5’ splice site and/or branch site recognition than would splicing
activated by other pathways. We measured the sensitivity of β globin-SRp38 splicing to several
different RNA oligos in nuclear extract (NE), and in S100 plus NF40/60 and either 50 ng SRp38 or 300 ng ASF/SF2. Strikingly, an RNA oligo containing both the 5’ splice site consensus and a polypyrimidine stretch strongly inhibited splicing in NE and in S100 in the presence of ASF/SF2,
10
164 but was much less effective in S100 plus SRp38 (Fig. 4d, compare lanes 1-5 and lanes 6-15).
Other RNA oligos tested, such as anti-U2 (Supplementary Fig. 3a online) and anti-U6
(Supplementary Fig. 3b online) oligos, led to equivalent splicing repression in NE and SRp38- activated splicing. Taken together, these results provide evidence that SRp38 stimulates splicing by facilitating recruitment of U1 snRNP and U2 snRNP to the 5’ splice-site and branchsite.
SRp38 activates in vitro splicing of a natural RNA containing SRp38 binding sites.
Given that SRp38 functions as a sequence-specific splicing activator, we next wished to identify possible natural targets. One potential substrate is that the alternatively spliced transcript is encoded by the murine AMPA receptor subunit GluR-B gene33. Exon 14 and 15 (referred to as
Flip and Flop) are mutually exclusive and impart different properties on currents evoked by L-
glutamate or AMPA34. Transient transfection experiments showed previously that SRp38
isoforms can differentially influence Flip/Flop splicing33.
In light of the above, we examined the sequences of both Flip and Flop for the presence
of potential SRp38 binding sites. Strikingly, we found the motifs GACAAA in the Flop exon and
AGACAA in the Flip exon that constitutes good matches to the core of the SRp38 consensus
sequence, AAAGACAAA (Fig. 5a). To determine whether these sequences might be functional,
we performed gel shift assays with both Flop and Flip RNAs and purified GST-tagged SRp38, and compared binding with an RNA containing three copies of the SRp38 consensus (Fig. 5b).
The results show that both Flop and Flip RNAs bind SRp38, but with different affinities. The
Flip RNA showed an affinity for SRp38 (~20 nM Kd) comparable to the SRp38 consensus RNA
(compare lanes 5-8 with lanes 9-12), while binding to the Flop RNA was significantly weaker
11
165 (Kd ~60 nM; lanes 1-4). These RNAs displayed no affinity for SC35 (Fig. 5c), indicating that the
binding was specific for SRp38.
We next asked whether the sequences in Flop and Flip could bring about SRp38-
dependent splicing and whether the differential binding affinity for SRp38 might be functionally
significant in splicing activated by SRp38. Following the same strategy used to measure SRp38- dependent splicing described above, we constructed modified β-globin substrates containing
Flop or Flip sequences in the second exon, which we named β-Flop and β-Flip (Fig. 5d). We tested these RNAs in S100 activated by SRp38 alone or by SRp38 plus NF40-60. Significantly, the Flip containing RNA was efficiently spliced, and again in a manner dependent on NF40-60
(lanes 13-16). On the other hand, SRp38-activated splicing of the Flop-containing substrate was detectable but much weaker, consistent with its lower affinity for SRp38 (lanes 5-8).
SRp38 promotes inclusion of Flip in GluR-B pre-mRNA splicing in DT40 cells.
We next wished to determine whether SRp38 favors inclusion of Flip in GluR-B pre- mRNA splicing in vivo. To address this, we constructed a modified GluR-B mini-gene plasmid in which GluR-B sequences were preceded by the chicken β-actin promoter and followed by an
SV40 poly(A) site as illustrated in Fig. 6a. This plasmid was stably transfected into chicken
DT40 cells with the genetic background SRp38(+/+) or SRp38(-/-)26. Several stably transfected
colonies were isolated and total RNAs were extracted and analyzed first by RT-PCR. Three
transcript variants are expected to be produced by alternative splicing from GluR-B minigene
transcripts35: Flop (612 bp), Flip (612 bp) and Truncated (lacking both the Flop or Flip exons;
about 500 bp), as shown in Fig. 6a. The pattern and intensity of RT-PCR products were identical
with RNA samples from SRp38-containing and SRp38-lacking DT40 cells (Fig. 6b, lanes 1-2;
12
166 data not shown). We then took advantage of the fact that there is a unique Stu I site in the Flop
exon, and treated PCR products with Stu I followed by agarose gel analysis. Strikingly, we
observed significantly more of the Flop variant in the SRp38(-/-) cells (Fig. 6b, lanes 3-4),
suggesting that loss of SRp38 promotes inclusion of Flop in GluR-B pre-mRNA splicing. To
confirm this, we performed an RNase protection assay using an antisense RNA targeted against full-length exon 13-14 sequences as shown in the Fig. 6a. Consistent with the above RT-PCR
results, a significant difference was observed between SRp38(+/+) and SRp38(-/-) cells, such
that there was an increase in the amount of Flop exon in the absence of SRp38, and a decrease in
Flop in the presence of SRp38 (Fig. 6c, lanes 1-2).
Finally, we wished to confirm that the increased Flop mRNA levels were caused directly
by loss of SRp38 in the SRp38(-/-) cells. To this end, we transfected the GluR-B reporter plasmid
into SRp38(-/-) cells that expressed HA-tagged SRp3826. RNA was isolated and analyzed by
RNase protection assay. Significantly, expression of SRp38 reduced Flop mRNA to levels
observed in the SRp38(+/+) cells (lane 3). For confirmation, we analyzed several additional HA-
SRp38 expressing cell lines that contain the stably transfected GluR-B mini-gene, and found that
all have decreased Flop mRNA levels (data not shown). These results demonstrate that SRp38 influences alternative splicing of the GluR-B Flop and Flip exons in vivo.
Discussion
In this paper, we have shown that phosphorylated SRp38, unlike its dephosphorylated counterpart, activates splicing. Our results have revealed a novel mechanism for splicing activation brought about by SRp38. Our data suggests a model in which U1 and U2 snRNP are unable to associate stably with certain pre-mRNA substrates in the absence of SRp38. Binding of
13
167 these snRNPs to the pre-mRNA is stabilized by SRp38, but the A complex formed is stalled and
unable to proceed in the splicing pathway in the absence of a specific coactivator (see Fig. 7).
This activator function of SRp38 is not only opposite from the function of dephosphorylated
SRp38, which acts as a general splicing repressor29, but also distinct from the mechanism of
standard SR or SR-related proteins in activating splicing10,30,36. Extending these findings, we
observed that SRp38 affects the selection of mutually exclusive exons in the GluR-B pre-mRNA in a sequence- and affinity-dependent manner. Below we discuss the features of SRp38-activated splicing compared to previously reported examples of ESE-dependent splicing activation, as well as the role of SRp38 in the regulation of alternative splicing.
We previously reported that SELEX-determined sequences optimal for ASF/SF2, SRp40 or Tra2 binding can function as splicing enhancers when placed downstream of an enhancer- dependent intron10,30,36. Artificial tethering experiments have also shown that RS domains are
sufficient to activate splicing in nuclear extracts or S100 plus SR proteins, when recruited to
ESE37,38. Therefore, it seems to be a general feature of SR proteins that they can interact with
downstream ESEs and activate splicing of weak upstream introns. However, SRp38-mediated
activation appears to have different requirements. In early experiments, we failed to detect any
stimulation by SRp38 of substrates in which three copies of the SRp38 binding site were placed
downstream of weak introns (unpublished data). We initially thought that perhaps this sequence
cannot function as an SRp38-dependent ESE, which would be consistent with the unusual
properties of SRp38. Additionally, this would be analogous to the behavior of SELEX selected sequences for the related protein SC35, which cannot function as an ESE in vitro10. Alternatively, given that SRp38 is an atypical SR protein, the selected sequence might in fact be able to function as an ESE but in a different way from sequences for standard SR proteins. This lead us
14
168 to test whether the SRp38 sequences might function with pre-mRNA containing consensus splice
sites, such as the -globin and AdML substrates, which allowed us to demonstrate that the
SRp38 consensus sequence can indeed function as an ESE. However, unlike standard SR
proteins, it may be that SRp38 functions preferentially on substrates containing introns with
strong splice sites.
Our data has demonstrated that SRp38 requires a specific cofactor (or cofactors) for
activity. It was previously shown that a 100 kD non-SR protein, in addition to SR proteins, is
required for activation of the weak 3’ splice site of -tropomyosin exon 239. Together with our
results, this suggests that the activity of certain ESEs can be modulated by the assembly of
additional proteins on the enhancer element, in addition to specific SR proteins. This is similar to
the role played by the sex determination genes, tra and tra-2, in Drosophila; the proteins
encoded by the two genes form a stable enhancer dependent complex with SR proteins,
stimulating the splicing of the female-specific exon of the doublesex transcript40. Likewise,
human Tra2 requires the activity of SR proteins for sequence-specific activation30,41. Notably,
this is distinct from the mechanism of SRp38, which does not require additional SR proteins.
We do not yet know the identity of the coactivator. However, SRp38 was not able to cooperate with general coactivators such as SRm160/30023, because no splicing inhibition was observed when SRm160/300 antibody was added in the SRp38-activated splicing reaction
(unpublished data). Another SR protein, 9G8, which has been shown to synergize with ASF concentrations at lower concentrations42, was also ruled out as the coactivator because it was not
present in partially purified fractions that retain cofactor activity. Consistent with the antibody
data, SRm160/300 were also absent from this fraction (unpublished data).
15
169 The mechanism of U1 snRNP and U2 snRNP recruitment by SRp38 appears distinct
from the mechanism of canonical SR proteins in recruiting U1 snRNP to the 5’splice site17 or stabilizing U2 snRNA interaction with the branchsite32. First, the stabilization by SRp38 is
sequence-dependent in that SRp38 is only capable of recruiting U1 and U2 snRNPs to a pre-
mRNA containing its high-affinity binding site. This is distinct from ASF/SF2, which recruits
U1 snRNP in an apparently sequence-independent manner17. Second, canonical SR proteins are
sufficient to facilitate formation of fully active spliceosomes in S100 extracts. In contrast, SRp38
facilitates A complex formation in the absence of other SR proteins, but the A complex is stalled and unable to proceed to later spliceosomal complexes in the absence of a cofactor.
How might the SRp38 cofactor function? One possibility is that it recruits the U4/U6·U5
tri-snRNP to the pre-formed A complex. Standard SR proteins have in fact been shown to play a
role in the recruitment of the tri-snRNP into the spliceosome43,44. Also, there are at least two RS domain-containing proteins in the tri-snRNP (27K and U5-100K) that have the potential to interact with SR proteins45,46. It is thus possible that SRp38 is distinct from other SR proteins in
that it is not able by itself to recruit the tri-snRNP to the A complex. It is also possible that tri-
snRNPs are able to bind transiently to the early spliceosome complex and form an unstable “B-
like” complex, but a conformational rearrangement brought about by the cofactor, is necessary
for stability. Another possibility is that the spliceosomal A complex formed by SRp38 is not
functionally active. In this situation, the cofactor may affect the conformation of the A complex
formed with SRp38, so that the tri-snRNPs will be able to bind to it productively and allow splicing to proceed. Identification of the cofactor will likely provide considerable insight into why SRp38-activated spliceosome are stalled at A complex, and is of course an important goal of future experiment.
16
170 Consistent with its role as a sequence-specific splicing activator, SRp38 is capable of acting as a regulator of alternative splicing, influencing selection of mutually exclusive exons of the GluR-B pre-mRNA. The mutually exclusive Flip and Flop exons are both flanked by what appear to be strong splice sites. Although SRp38 binds to both exons, the affinities are different.
Therefore, it is reasonable to propose that the differential binding by SRp38 influences the decision to include Flip or Flop, and may reflect in part intracellular concentrations of SRp38.
SC35 did not bind to either exon, but consistent with the general activation function of SR proteins, could activate splicing of substrates containing either the Flop or Flip exon but without preference (unpublished data). We note that this contradicts previous data in which transiently overexpressed SC35 (as well as ASF/SF2) was found to increase the Flop to Flip ratio, and
SC35-responsive elements in the Flop exon were identified47. Another group reported that the two splice variants of SRp38 (NSSR1 and NSSR2, or SRp38 and SRp38-2) had opposite effects on Flip inclusion33. However, our in vitro data indicates that both variants of SRp38 have similar effects on Flip vs Flop splicing activation, although SRp38-2 displayed reduced activity
(unpublished data). The basis for these differences is unclear, but they may reflect the use of transient overexpression assays in the previous experiments.
In summary, SRp38 represents a distinct type of splicing regulatory protein.
Phosphorylation/dephosphorylation switches the protein from a general splicing repressor to a sequence-specific activator. SRp38 activates splicing by facilitating formation of early splicing complexes that cannot proceed to active spliceosomes without the aid of a specific cofactor.
Furthermore, SRp38 can regulate alternative splicing, and has the potential to play important roles in a variety of physiologically significant processes. Future studies on both its mechanism and target transcripts should be informative.
17
171 Methods
Plasmid constructions, cell culture and transfection: All plasmids (β-SRp38, β-control,
β-Flop, β-Flip, AdML-SRp38) used to produce substrates for in vitro splicing were constructed
by replacement of sequences between Acc I site and BamH I site in the second exon with indicated sequences. For in vitro gel-shift assays and RNase protection assays, plasmids were constructed by insertion of indicated sequences into pBluescript SK(+) plasmid. The GluR-B reporter minigene was constructed as described previously33 except that pEXpress plasmid48 was
used as the backbone. DT40 cells of different background including wild-type, SRp38(-/-) and
SRp38(-/-) containing exogenously expressed HA-tagged SRp38 were maintained essentially as
described previously26. Transfection of the GluR-B reporter minigene into DT40 cells was also
performed as described previously26.
Recombinant proteins: His-SRp38, his-SC35 and his-ASF/SF2 and GST-SRp38 were prepared from recombinant baculovirus-infected High Five cells (Invitrogen). His-tagged recombinant proteins were purified under denaturing conditions by Ni2+agarose chromatography
and renatured by dialysis10,27. Dephosphorylated SRp38 was prepared by incubating recombinant
SRp38 with CIP and repurified by agarose chromatography27. GST-tagged SRp38 was purified by using glutathione–Sepharose 4B26. (His- and GST-tagged SRp38 behaved indistinguishably in all assays tested.) GST-SRp38 RBD and hnRNP G RBD proteins were prepared from E. coli
JM101 using glutathione-Sepharose 4B and his-SC35 RBD was prepared from E. coli BL2110.
Purity and concentration of proteins were determined by Coomassie blue staining of SDS gels.
In vitro splicing and spliceosome assembly assays: In vitro splicing was performed essentially as described10. Native SR proteins purified from Hela cells were obtained from Dr. T.
Kashima. Spliceosome assembly assays were also performed as described49. In the splicing
18
172 inhibition and spliceosome assembly assays, the anti-snRNAs sequences are: U1(1-14),
UGCCAGGUAAGUAU; U2(2-15), GGCCGAGAAGCGAU; U5(68-88),
UUGGGUUAAGACUCAGAGUUG; U6(78-95), CGCUUCACGAAUUUGCGU. 5’ss-Py,
UCACAGGUAAGUACUUAUUUUCCCAGGCC. Antisense RNAs were pre-incubated with
S100 at 30°C for 15 min before spliceosome assays.
Gel-shift assays: Gel-shift assays were preformed essentially as described17. Briefly,
radiolabelled pre-mRNA, U1 snRNP (300 ng and 600 ng) and SRp38 (150 ng/l) were incubated under splicing conditions: reaction mixtures were incubated at 30°C for 5 min. Heparin was then added to a concentration of 0.8 mg/ml and reaction mixtures were incubated at 30°C for an additional 5 min. Products were analyzed by 5% nondenaturing PAGE and autoradiography.
RT-PCR analysis and RNase protection assay: Total RNA was extracted by using
TRIzol reagent (Invitrogen). RT-PCR analysis was performed using Transcriptor Reverse
Transcriptase following the instructions provided by the supplier (Roche). RNase protection assays were performed as described50. Briefly, labeled RNA probes were synthesized by in vitro
transcription with SP6 RNAP. After hybridization and RNase digestion, protected RNAs were
resolved by 6% denaturing PAGE.
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175 42. Li, X., Shambaugh, M.E., Rottman, F.M. & Bokar, J.A. SR proteins Asf/SF2 and 9G8 interact to activate enhancer-dependent intron D splicing of bovine growth hormone pre-mRNA in vitro. Rna 6, 1847-58 (2000). 43. Roscigno, R.F. & Garcia-Blanco, M.A. SR proteins escort the U4/U6.U5 tri- snRNP to the spliceosome. Rna 1, 692-706 (1995). 44. Tarn, W.Y. & Steitz, J.A. Modulation of 5' splice site choice in pre-messenger RNA by two distinct steps. Proc Natl Acad Sci U S A 92, 2504-8 (1995). 45. Fetzer, S., Lauber, J., Will, C.L. & Luhrmann, R. The [U4/U6.U5] tri-snRNP- specific 27K protein is a novel SR protein that can be phosphorylated by the snRNP-associated protein kinase. Rna 3, 344-55 (1997). 46. Teigelkamp, S., Mundt, C., Achsel, T., Will, C.L. & Luhrmann, R. The human U5 snRNP-specific 100-kD protein is an RS domain-containing, putative RNA helicase with significant homology to the yeast splicing factor Prp28p. Rna 3, 1313-26 (1997). 47. Crovato, T.E. & Egebjerg, J. ASF/SF2 and SC35 regulate the glutamate receptor subunit 2 alternative flip/flop splicing. FEBS Lett 579, 4138-44 (2005). 48. Arakawa, H., Lodygin, D. & Buerstedde, J.M. Mutant loxP vectors for selectable marker recycle and conditional knock-outs. BMC Biotechnol 1, 7 (2001). 49. Das, R. & Reed, R. Resolution of the mammalian E complex and the ATP- dependent spliceosomal complexes on native agarose mini-gels. Rna 5, 1504-8 (1999). 50. Wang, J. & Manley, J.L. Overexpression of the SR proteins ASF/SF2 and SC35 influences alternative splicing in vivo in diverse ways. Rna 1, 335-46 (1995).
Acknowledgments
We thank R. Lührmann for providing purified U1 and U2 snRNP, T. Kashima for SR proteins and C. Shin for his-dSRp38. We thank Allison Yang for help with the manuscript. We also thank members of the Manley laboratory for helpful discussions and comments. This work
was supported by NIH grant NIH GM48259.
Figure Legends
Figure 1. SRp38 functions as a sequence-specific activator of splicing. (a) Schematic
representation of β-globin derivatives containing either three copies of SRp38 consensus
sequence or a control sequence. (b) Activation of β-SRp38 pre-mRNA splicing by SRp38.
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176 Splicing was performed in S100 supplemented with 50 ng (lanes 3, 5, 9, 11) or 100 ng (lanes 4, 6,
10, 12) of baculovirus-produced his-SRp38 together with or without a nuclear fraction (NF40-
60), as indicated at the top. Products of splicing were analyzed by denaturing PAGE and autoradiography. Splicing products are indicated schematically. (c) Dose-dependent splicing activation of SRp38. The indicated amounts of his-SRp38 were added to splicing reactions performed in S100 supplemented with NF40-60.
Figure 2. Characteristics of SRp38-dependent splicing activation. (a) RS domain requirements for activation. β-SRp38 pre-mRNA was incubated in S100 activated by 50 ng of
GST-SRp38, GST-SRp38-2, GST-SRp38 RBD or 8 ng of dephosphorylated his-SRp38
(dSRp38), respectively, in the absence (lanes 2, 4, 6, 8) or presence (lanes 3, 5, 7, 9) of NF40-60.
(b) Repression of splicing by dSRp38. His-dSRp38 (0, 2, 4 and 20ng) was incubated with S100 plus phosphorylated his-SRp38 (20 ng) and NF 40-60 (lanes 10-12). (c) Role of the RBD in
SRp38-dependent activation. The indicated amounts of purified GST-SRp38 RBD, His-SC35
RBD or GST-hnRNP G RBD were added to SRp38-dependent splicing assays as indicated. (d)
SR proteins do not cooperate with SRp38 for splicing activation. Increasing amounts of purified
SR proteins were added to reactions performed in S100 alone (lanes 2- 4) or in the presence of
50 ng of GST-SRp38 (lanes 5-7). (e) NF 40-60 coactivator activity is specific for SRp38. 50 ng of His-SRp38, His-ASF or His-SC35 was incubated with -SRp38 or -SRp38 RNAs in S100 alone or supplemented with NF40-60. (f) Splicing stimulation by ASF/SF2 or SC35 at high concentrations. 300 ng His-tagged SRp38, ASF/SF2 and SC35 were incubated with -SRp38
RNA in S100.
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177 Figure 3. SRp38 promotes formation of spliceosomal complex A. (a) Spliceosome
assembly assays were carried out in S100 complemented with the indicated components and the
β-SRp38 pre-mRNA. Splicing complexes were resolved on a 1.5% low-melting agarose gel. (b)
SRp38 activates AdML-SRp38 in in vitro splicing assays. Splicing was performed in S100
supplemented with 50 ng of GST-tagged SRp38 together with (lane 1) or without (lane 3) NF40-
60. Products of splicing were analyzed by denaturing PAGE and autoradiography. (c)
Spliceosome assembly was performed the same as in (a) except with the AdML-SRp38 pre- mRNA. (d) Spliceosome assembly assays were carried out as in (c) except with the additions indicated at the top. Anti-U1, U2, U5 or U6 snRNA oligos (final concentration 5 M) were added to reaction mixtures. Endogenous ATP was depleted by pre-incubating S100 at 30ºC for
40 min.
Figure 4. SRp38 interacts with both U1 and U2 snRNP complexes on the SRp38 substrate. Purified U1 snRNP (a) or U2 snRNP (b), GST-tagged SRp38 (150 ng) or GST-tagged
SRp38 RBD (250 ng) were incubated with β-SRp38 pre-mRNA (a and b, lanes 1-9) or β-control pre-mRNA (a and b, lanes 10-14), and following addition of heparin (0.8 mg/ml), complexes were resolved by 5% nondenaturing PAGE. Increasing amounts of U1 snRNP (a) or U2 snRNP
(b) (300, 600) were incubated with SRp38 and β-SRp38 RNA (a and b, lanes 4, 5 and 6). The complexes formed are indicated with brackets. 250 ng GST-tagged SRp38 RBD was incubated with β-SRp38 RNA (a and b, lanes 6), and increasing amounts of U1 or U2 snRNPs (150, 300 or
600 ng) were added (a and b, lanes 7-9). 150 ng SRp38 was incubated with β-control pre-mRNA alone (a and b, lanes 13) or β-control pre-mRNA plus U1 or U2 snRNPs (300 ng) (a and b, lanes
14). (c) His-tagged ASF/SF2 was incubated with β-SRp38 in the absence (lane 2) or in the
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178 presence of U1 snRNP (lane 3) or U2 snRNP (lane 4). (d) Splicing reactions were performed with the β-SRp38 RNA. The U1-Py oligo was pre-incubated with NE or with S100 plus NF40-
60 and SRp38 before splicing reactions were carried out. Final concentrations of the oligo are 0,
0.1, 0.5, 1, 2 M. Note that 300 ng ASF/SF2 was used in the splicing reactions in order to achieve similar splicing levels as in NE.
Figure 5. SRp38 binds to and activates splicing of RNA substrates containing the
GluR-B Flip or Flop exon. (a) Comparison of a putative SRp38 binding motifs in Flip and Flop exons from the murine GluR-B transcript with the SRp38 consensus recognition sequence. (b)
Flop and Flip exons of the GluR-B pre-mRNA bind SRp38 specifically. Gel shift assays with the indicated radiolabeled RNAs were performed with increasing amounts of GST-SRp38 (25, 75,
225 ng). Complexes were resolved by nondenaturing PAGE. (c) Flop and Flip exons of the
GluR-B pre-mRNA do not bind His-SC35. Increasing amounts of purified His-SC35 (25, 75, 225 ng) were added to gel-shift assays containing the identical RNAs. (d) SRp38 activates splicing of
RNA substrates containing Flip or Flop exons. The downstream β-globin exon was replaced with
Flip or Flop exons, and splicing reactions were performed in S100 with increasing amounts of
GST-SRp38 (20, 40, 100 ng) in the presence or absence of NF40-60 as indicated.
Figure 6. Loss of SRp38 promotes inclusion of the Flop exon in vivo. (a) Diagram of reporter plasmids containing murine GluR-B truncated genomic sequences and three alternative spliced products. The pair of primers used in RT-PCR (b) are shown as two reverse arrows. The
RNA probe used in RNase protection assay (c) is indicated at the top of the three products, and the length of protected probe is on the side. (b) Comparison of alternatively spliced products
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179 from SRp38(+/+) and SRp38(-/-) DT40 cells. RT-PCR was performed with RNAs extracted from stably transfected DT40 cells, and analyzed directly on a 1.5% agarose gel (lanes 1-2), or digested with Stu I followed by agarose gel analysis (lanes 3-4). (c) RNase protection assay.
RNA was extracted from stably transfected DT40 cells in the background of SRp38(+/+),
SRp38(-/-) and SRp38(-/-) expressing exogenous HA-SRp38. RNase protection assay was performed using the radiolabeled RNA probe indicated in (a). Products were resolved on 6% denaturing PAGE.
Figure 7. Model for SRp38-dependent splicing activation. SRp38 binds the SRp38- dependent ESE in target transcripts and facilitates U1 and U2 snRNPs association with the pre-
mRNA to stabilize 5’ splice-site and branch site recognition by interacting with U1 and U2
snRNPs, respectively. However, the spliceosomal A complex formed is stalled and requires an
SRp38-specific cofactor to proceed through the splicing pathway.
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