Far upstream element-binding protein 1 and RNAsecondary structure both mediate second-stepsplicing repressionHuang Lia, Zhijia Wanga, Xuexia Zhoua, Yuanming Chenga, Zhiqin Xiea, James L. Manleyb,1, and Ying Fenga,c,1

aKey Laboratory of Food Safety Research, Institute for Nutritional Sciences, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences,University of Chinese Academy of Sciences, Shanghai 200031, China; bDepartment of Biological Sciences, Columbia University, New York, NY 10027;and cKey Laboratory of Food Safety Risk Assessment, Ministry of Health, Beijing 100021, China

Contributed by James L. Manley, June 4, 2013 (sent for review April 2, 2013)

Splicing of mRNA precursors consists of two steps that are almostinvariably tightly coupled to facilitate efficient generation ofspliced mRNA. However, we described previously a splicing sub-strate that is completely blocked after the first step. We have nowinvestigated the basis for this unusual second-step inhibition andunexpectedly elucidated two independent mechanisms. Oneinvolves a stem–loop structure located downstream of the 3′splicesite, and the other involves an exonic splicing silencer (ESS) situ-ated 3′ to the structure. Both elements contribute to the second-step block in vitro and also cause exon skipping in vivo. Impor-tantly, we identified far upstream element-binding protein 1(FUBP1), a single-stranded DNA- and RNA-binding protein not pre-viously implicated in splicing, as a strong ESS binding protein, andseveral assays implicate it in ESS function. We demonstrate usingdepletion/add-back experiments that FUBP1 acts as a second-steprepressor in vitro and show by siRNA-mediated knockdown andoverexpression assays that it modulates exon inclusion in vivo.Together, our results provide additional insights into splicing con-trol, and identify FUBP1 as a splicing regulator.

Removal of introns from pre-mRNAs by splicing is a preciseprocess required for the expression of nearly all genes in

human cells. Splicing takes place via two sequential transesteri-fication reactions in the spliceosome, a large complex consistingof several hundred proteins and five small nuclear RNAs (1, 2).During the first step, an adenosine residue designated the branchpoint, attacks the 5′ splice site (5′SS) to generate the splicingintermediates (free exon1 and lariat-exon2). In the second step,the first exon attacks the 3′ splice site (3′SS), yielding ligatedexons and a lariat intron.Splicing, like numerous other cellular processes, must be un-

der strict regulatory control. Indeed, aberrant splicing is involvedin a wide range of human diseases (3, 4). Besides the coresplicing signals, additional cis-regulatory elements play impor-tant roles in splicing and its control. These elements exist withinexons and introns and can function both positively and negatively(5–7). Two major classes of exonic elements have been identi-fied, known as exonic splicing enhancers (ESEs) and exonicsplicing silencers (ESSs), and both of these contribute to regu-lation of alternative splicing (AS). ESEs are usually bound bymembers of the serine/arginine-rich (SR) protein family to en-hance recognition of adjacent splice sites. By contrast, ESSsfunction to inhibit the use of adjacent splice sites, and are gen-erally dependent on the interaction with members of the het-erogeneous nuclear ribonucleoprotein(hnRNP) family, as wellas other splicing regulators such as NOVA1/2 and RBFOX1/2(8, 9).A good deal is now known about the structure and function

of ESSs. For example, several large-scale strategies have beendeveloped to identify and characterize ESSs (10–12). Thesequences of known and selected ESSs share little similarity witheach other, perhaps reflecting the diversity of their cognatebinding proteins. It has been thought that ESSs inhibit splicing

either by antagonizing the functions of ESEs or by recruitingfactors that interfere directly with the splicing machinery. Uponbinding of splicing repressors to ESSs, splicing is most oftenblocked at the stages of splice-site recognition and/or earlyspliceosome assembly (5, 6). However, inhibition can also occurat later stages of spliceosome assembly or activation. For ex-ample, in Drosophila, the sex-lethal protein (Sxl), an hnRNP-likesplicing repressor, autoregulates its own splicing by binding tothe pre-mRNA and causing exon exclusion by blocking splicingat the second step via interference with a splicing factor, SPF45,specifically required for the second step (13). However, examplesof splicing regulation resulting from second-step inhibition havenot been described in vertebrates.RNA secondary structures can also contribute to splicing

control. For example, RNA structure has been shown to affectsplicing efficiency by masking splice sites (14) or binding sites forsplicing factors (15). Indeed, an increasing amount of genomicdata in recent years indicate that RNA structures might bea common method to regulate AS by repressing exon inclusion(16, 17). Additionally, there is also evidence, although currentlyonly in yeast, that secondary structure can function to inhibit thesecond step of splicing. A stem–loop inserted directly after thebranch point of the ACT intron inhibits the second step in vitro(18). Likewise, an exonic stem–loop structure adjacent to the3′SS in an ACT-LacZ fusion gene primarily blocks the secondstep in vivo (19).We previously characterized the role of the serine/arginine-

rich splicing factor 10 (SRSF10) in regulation of AS in murineembryonic hearts (20). In that study, we showed that SRSF10

Significance

Splicing of mRNA precursors occurs in two sequential trans-esterification steps. We characterize a highly unusual inhibitionof splicing in which the reaction is blocked between the twosteps. We demonstrate that RNA secondary structure and anexonic splicing silencer element (ESS) can independently me-diate second-step splicing repression in vitro and cause exonexclusion in vivo. Importantly, we provide evidence that farupstream element-binding protein 1, a single-stranded DNA-and RNA-binding protein initially identified as a regulator ofMYC transcription and recently implicated in several cancers,binds the ESS and functions as a splicing regulator in vitro andin vivo.

Author contributions: H.L., J.L.M., and Y.F. designed research; Z.W., X.Z., Y.C., Z.X., andY.F. performed research; H.L., J.L.M., and Y.F. analyzed data; and H.L., J.L.M., and Y.F.wrote the paper.

The authors declare no conflict of interest.1To whom correspondence may be addressed. E-mail: jlm2@columbia.edu or fengying@sibs.ac.cn.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1310607110/-/DCSupplemental.

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controls AS of the pre-mRNA encoding the cardiac proteintriadin, by binding to and activating inclusion of exon 9. Duringthe course of these studies, we also examined splicing in vitro oftriadin exon 10 in the context of a chimeric β-globin substrate inwhich exon 10 was placed in the downstream exon (β-E10). Wehad used this assay previously with other exons to characterizethe role of SRSF10 as a sequence-specific splicing activator (21).Unexpectedly, we found that the β-E10 substrate underwent thefirst step of splicing efficiently but was completely blocked beforethe second step (20). This led us to suggest that exon 10 containssequences that can act as a potent ESS, but functioning to blocksplicing at the second step.In this study, we investigated the mechanism underlying the

second-step inhibition observed with the β-E10 substrate. Re-markably, we found that two independent mechanisms contrib-ute to the block. One is mediated by a serendipitously createdstem–loop structure downstream of the 3′SS that involvesβ-globin and triadin exon 10 sequences. This structure contributesto the second-step block in vitro and also induces exon skippingin vivo. The second mechanism involves an ESS in exon 10, whichalso induces a second-step block in vitro and exon skippingin vivo. Furthermore, we show that FUBP1, a single-strandedDNA- and RNA-binding protein that was implicated previouslyin regulation of MYC transcription (22) and aspects of RNAmetabolism other than splicing (23, 24) binds the exon 10 ESS.Depletion/add-back experiments show that FUBP1 can inducethe second-step block in vitro, whereas siRNA-mediated knock-down or overexpression of FUBP1 causes exon skipping and ASchanges in vivo. Our results thus indicate that both RNA sec-ondary structure and an ESS can repress splicing at the secondstep and that FUBP1 can function as an unusual second-stepsplicing repressor.

ResultsCharacterization of a Second-Step Splicing Block. We previouslyconstructed and analyzed chimeric β-globin substrates (β-E9 andβ-E10) in which triadin exon 9 and exon 10 were inserted 16 nt

downstream of the 3′SS, replacing the remaining sequences inthe second exon of the β-globin gene (20). Whereas the β-E9RNA was efficiently spliced in HeLa nuclear extract (NE) (Fig.1A, lanes 1–3), the β-E10 RNA only produced what appearedto be the 5′ exon cleavage product (Fig. 1A, lanes 4–6; see alsoref. 20), consistent with the splicing reaction stalling after com-pletion of the first step.We first characterized the apparent second-step inhibition in

greater detail. We initially asked whether inhibition indeed re-quired exon 10 sequences. To test this, we analyzed splicing of anRNA substrate that was prepared from β-E10 linearized usingAccI, as indicated in Fig. 1B. The resultant RNA lacks the E10sequences, containing only a 12-nt β-globin–derived second exon.This RNA was spliced efficiently, in contrast to the second-stepinhibition observed with the β-E10 RNA (Fig. 1B, lanes 1–3,compare with lanes 4–6), indicating that the E10 sequences arerequired for inhibition. Given that only one of the two splicingintermediates, the 5′ exon, was apparent, we next wished to con-firm that this was indeed a splicing-related reaction and that thisRNA was not simply a degradation product that was fortuitouslythe same size as 5′ exon. To this end, we performed in vitro-splicingassays as a function of time and in the presence or absence of ATPand resolved the products on a 12% (vol/vol) polyacrylamide de-naturing gel instead of the 6% (vol/vol) polyacrylamide gel usedabove. As shown in Fig. 1C, incubation of the β-E10 RNA undersplicing conditions resulted in time- and ATP-dependent accu-mulation of the lariat-3′ exon intermediate, which migrated moreslowly than the precursor RNA, as well as the free 5′ exon (lanes1–4, compare with lanes 5–8). As expected, first-step splicing ofthe β-E10 RNA was also Mg2+-dependent (Fig. S1). Additionally,using an S100-based splicing assay (21), first-step splicing of theβ-E10 RNA was enhanced by SR proteins, such as SRSF1 andSRSF2 (Fig. 1D, lanes 13–16). Together, these results demon-strate that the β-E10 RNA undergoes the first step of splicingefficiently in vitro but is unable to proceed to the second step,reflecting the presence of inhibitory sequences in E10 that spe-cifically block the second step.

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Fig. 1. Characterization of the second-step splicingblock. (A) In vitro splicing was performed in HeLaNE using β-E9 or β-E10 RNAs for the indicated time.Products of splicing were analyzed by denaturing6% (vol/vol) PAGE and autoradiography. Splicingproducts are indicated schematically. (B) The β-E10plasmid was linearized with the enzymes indicatedat the bottom, and the two RNA substrates wereanalyzed for splicing in NE as in A. (C) β-E9 or β-E10RNAs were incubated in NE in the presence (lanes1–3 and 5–7) or absence (lanes 4 and 8) of ATP forthe indicated time. Splicing products were resolvedon a 12% (vol/vol) denaturing PAGE. (D) His-SRSF1,His-SRSF2, or His-SRSF10 (100 ng) was incubatedwith β-E9 or β-E10 RNAs in HeLa S100 alone orsupplemented with NF40-60. Products were ana-lyzed as in A.

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E10 Sequences Induce Exon Skipping in Vivo. We next wished todetermine whether the second-step inhibition observed in vitrocould affect exon use in vivo. To address this, we used minigeneconstructs containing E9 or E10 sequences in a middle exonflanked by β-globin genomic sequences, driven by a CMV pro-moter (Fig. 2A; see Materials and Methods for details). Similarsubstrates have been used previously to analyze ESE-dependentexon inclusion (25). All four splice sites, as well as immediatelyadjacent sequences, were the same in all constructs. Splicing wasassayed following transient expression in HeLa cells. Total RNAwas extracted from transfected cells and exon inclusion/skippingwas analyzed by RT-PCR (Fig. 2B). Strikingly, whereas the E9-

containing exon was fully included, the corresponding E10 exonwas completely excluded. Thus, the strong second-step blockbrought about by E10 sequences in vitro correlates with com-plete exon exclusion in vivo.

A Secondary Structure Downstream the 3′SS Contributes to ExonSkipping in Vivo. We next set out to investigate the molecularbasis for the observed E10-induced splicing inhibition. Onepossibility we considered is that an inhibitory secondary structureforms in the β-E10 RNA. We initially used RNAfold (26) forRNA secondary structure prediction, which revealed a potentialstem–loop structure immediately downstream of the 3′SS, in-volving β-globin exon 2 sequences and the first 10 nt of E10 (Fig.3A). To test whether the putative secondary structure contributesto exon exclusion, we first made two 7-mer mutations in the E10-based construct, both of which disrupt the putative stem: onemutated UGCUGGU to CCGACGC (E10M1), and the other mu-tated the complementary sequence AUCAGUA to GCGUUGG(E10M2) (Fig. 3A, Upper). Following transfection and RT-PCRanalysis, we found that both mutations strongly activated E10inclusion (Fig. 3A, Lower, lanes 2–3, compare with lane 1).Furthermore, when the secondary structure was reconstituted bycombining the two mutations (E10M1/E10M2), E10 containingexon was again totally skipped (Fig. 3A, Lower, lane 4).To provide additional support for the importance of the stem

in inducing exon exclusion, we analyzed several mutants in whicha truncated version of E10 was substituted for the intact E10, asshown in Fig. 3B. The stem–loop structure is predicted to remainintact in the F19 and F30 constructs, whereas it is disrupted inL24 and M39. Consistent with the importance of the structurefor exon skipping, complete exon exclusion was detected withboth F19 and F30 constructs (Fig. 3B, Lower, lanes 2 and 4),whereas almost complete (M39) or partial (L24) inclusion wasobserved with the L24 andM39 (Fig. 3B, Lower, lanes 3 and 5). Inaddition, we analyzed another four sets of more subtle mutations,made in the F19-based construct and also disrupting/recon-stituting the stem. Although, in some cases, the effects observedwere partial, the results obtained with all four sets were entirelyconsistent with the existence/importance of the stem–loop struc-ture (Fig. S2). Together, these findings provide strong evidencethat exonic RNA secondary structure can block exon inclusion.

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(E10M2) Fig. 3. A secondary structure downstream the 3′SScontributes to exon skipping in vivo. (A, Upper)Putative stem–loop structure located downstreamof the 3′SS involving β-globin exon 2 (black) and E10(red) sequences. Two mutants (E10M1 and E10M2)disrupt the putative stem, which is reconstituted inthe double mutant (E10M1/M2). The 3′SS is in-dicated by an arrow. (A, Lower) HeLa cells weretransiently transfected with each of the E10 mutantconstructs, and RT-PCR was performed and productsanalyzed as described in Fig. 2B. (B, Upper) Diagramof several E10 truncations. (B, Lower) In vivo-splic-ing analysis of E10 truncations. (C, Upper) Diagramof a spacer inserted downstream the 3′SS of the F19construct. (C, Lower) F19 constructs with differentlength spacer were analyzed by RT-PCR as above.(D, Left) Diagram of a stem–loop structure in the E9/stem minigene. Mutations (red) were introduced inthe E9 construct as indicated. (D, Right) HeLa cellswere transfected with E9 or E9/stem constructs, andRT-PCR analysis was performed as above.

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We also investigated how proximity of the secondary structureelement to the 3′ SS affects its inhibitory activity. For this, weused the F19 construct, which allowed straightforward insertionof increasing length spacers downstream of the 3′SS (Fig. 3C,Upper). If the secondary structure directly blocks recognition ofthe 3′SS, insertion of longer spacers would perhaps disrupt itsinhibitory effect and increase inclusion of the internal exon. Inagreement with this, whereas insertion of 2 nt had, at most,a minimal effect, 5- and 8-nt spacers significantly increased exoninclusion, to nearly 40% and 60%, respectively (Fig. 3C, Lower,lanes 3–4, compare with lane 1).We next asked whether introduction of a comparable structure

in a naturally included exon would be sufficient to induce skip-ping. To this end, mutations were made in the E9 construct toallow for formation of a stem just downstream of the 3′SS in themiddle exon (Fig. 3D, Left). When RNA from transfected cellswas analyzed by RT-PCR, ∼50% of the mutated exon was ex-cluded, compared with the total inclusion observed with the wild-type E9 construct (Fig. 3D, Right). These results suggest that a3′SS-proximal secondary structure is sufficient to induce exonskipping. However, although the secondary structure was almostidentical to that in E10, exclusion was only partial, suggestingthat other elements contribute to the complete exclusion ob-served with the E10 construct (see below).

Secondary Structure Contributes to but Is Insufficient for the Second-Step Block in Vitro. We next wished to investigate whether any ofthe mutations that led to exon inclusion in vivo could also pre-vent the second-step block and allowed splicing in vitro. To thisend, we analyzed several of these mutations in the context of theβ-globin–based splicing substrates used in our initial in vitroassays (Fig. 1). For the β-F30 and β-E10M1/M2 RNAs, second-step splicing inhibition was observed, in both NE and S100 plusNF40-60 [Fig. 4A, lanes 1–2 and 9–10; NF40-60 is a nuclearfraction that contains sufficient SR proteins to allow splicing,albeit weak, of β-globin–based substrates such as β-E9 RNA (Fig.1D, lanes 1 and 2, and ref. 21)]. This is consistent with the factthat these two mutants displayed exon exclusion in vivo. Incontrast, when incubated in NE, efficient splicing was detectedwith β-M39, β-E10M1 and β-E10M2 substrates (Fig. 4A, lanes 4,6, and 8), consistent with their ability to promote exon inclusionin vivo. Unexpectedly, however, when these three mutants wereanalyzed in NF40-60 plus S100, splicing remained blocked afterthe first step (Fig. 4A, lanes 3, 5, and 7). This inhibition suggeststhe presence of a second inhibitory element in exon 10, whichcan function independently of the secondary structure. It ispossible, for example, that when secondary structure was dis-rupted, binding of an unknown factor to an ESS was sufficient toresult in a second-step block in S100 plus NF40-60, perhaps

reflecting limiting amounts of SR proteins (21) and/or a relativeexcess of the putative second-step repressor.

FUBP1 Binds to a Potential ESS in E10. To identify a possible ESS inE10, we first examined E10 and active derivatives for sequencesthat match previously described ESSs. We found a sequence,AUAUAUGAU, located downstream of the stem-forming re-gion (Fig. 5A) that is similar to several ESS motifs identified bya SELEX approach, which likely constitute binding sites for in-hibitory protein(s) (11). Next, we sought to identify nuclearproteins that might bind to the AU-rich sequence. To this end,we prepared an [α-32P]UTP-labeled 30-nt RNA spanning thestem-forming region and AU-rich consensus of E10 (F30) andperformed UV-crosslinking assays with HeLa NE. After RNasetreatment and separation by SDS/PAGE, a prominent product of∼70 kDa was detected (Fig. 4B). To identify the bound protein,we performed RNA-affinity chromatography using a 5′ biotin-labeled version of the RNA used in crosslinking. After SDS/PAGE and Coomassie staining, one of two specifically boundproteins closely matched the protein observed after UV cross-linking (Fig. 4C). The indicated protein was excised and identi-fied by mass spectrometry (MS) as the FUBP1 (also known asFBP). FUBP1 is a largely nuclear protein that appears to possessboth RNA- and DNA-binding activity. It has been shown, on theone hand, to regulate transcription of the MYC protooncogene(22, 27) and, on the other, to bind to AU-rich sequences in the3′ UTRs of several transcripts (23, 28, 29). However, it has notbeen implicated previously in splicing. The other bound protein,of ∼35 kDa, was identified by MS as hnRNPD (Fig. 4C). Giventhat hnRNPD was not detected by UV crosslinking (Fig. 4B), andalso that the other mammalian FBP family member (KH-typesplicing regulatory protein, KSRP) (30) and the Drosophila ho-molog of the FBPs (P-element somatic inhibitor, PSI) have beenpreviously linked to splicing regulation (31), we decided to in-vestigate further the possible role of FUBP1 in splicing repression.

Binding of FUBP1 Correlates with ESS Activity. We next set out toexamine in more detail the role of the AU-rich element inbinding of FUBP1 and in splicing inhibition. To this end, we firstanalyzed the effects of a number of mutations in the AU se-quence on FUBP1 binding to F30 RNA. As shown in Fig. 5A,one mutant RNA (F30M1) contained continuous replacement ofAU with GC, the second mutant (F30M2) had four U-to-Ctransitions, and the third (F30M3) had four A-to-C substitutions.Fig. 5B shows the results of a gel-shift experiment in which in-creasing amounts of purified baculovirus-expressed His-taggedFUBP1 (Fig. S3) were incubated with F30 RNA and with each ofthree mutated derivatives. Whereas FUBP1 binding to F30 wasreadily detected, F30M1 and F30M2 RNAs were bound much

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less efficiently (Fig. 5B, lanes 4–6 and 7–9, compare with lanes 1–3).In contrast, the F30M3 mutant displayed only slightly weakenedbinding (Fig. 5B, lanes 10–12). We also estimated the Kd ofFUBP1 for F30 RNA by gels shift, which suggested a value of∼50 nM (Fig. S4). Although this is typical of other sequence-specific RNA-binding proteins (21), it is significantly higher thanvalues reported for single-stranded DNA (32, 33). Whether thisreflects differences in affinity of FUBP1 for DNA versus RNA,or that the F30 RNA site is not the optimal sequence, remains tobe determined. To extend the above analysis, we examined theinteraction of purified FUBP1 and RNA using UV-crosslinkingassays. Results shown in Fig. 5C revealed a sequence preferencefor FUBP1 binding essentially identical to that observed in thegel-shift assays. Taken together, these data indicate that FUBP1preferably binds to sequences highly enriched for U residues andthat the AU-rich sequence is critical for efficient binding ofFUBP1 to the F30 RNA.We reasoned that if binding of FUBP1 to the AU sequence is

responsible for the observed inhibitory effects on splicing, thenthe mutations that disrupt FUBP1 binding should activatesplicing. To test this, we first introduced each of the three abovemutations into the minigene construct containing F30 and testedthe effects on exon skipping in vivo. Fully in line with the aboveRNA-binding results, the F30M1 and F30M2 mutations thatdisrupted FUBP1 binding strongly impaired exon skipping,leading to more than 90% F30 inclusion (Fig. 5D, compare lanes2–3 with lane 1). In contrast, the F30M3 mutant displayed onlya slight effect on exon skipping (lane 4), consistent with its weakeffect on FUBP1 binding. We next introduced the same muta-tions into the β-F30–splicing substrate and tested their effects onin vitro splicing (Fig. 5E). Strikingly, and consistent with both theRNA-binding and in vivo-splicing results, efficient splicing wasobserved with the β-F30M1 and β-F30M2 RNAs when incubatedin NE (Fig. 5E, lanes 3–4 and 5–6), whereas β-F30M3 splicing wasstill blocked at the second step (lanes 7–8). Taken together, theresults provide strong evidence that the AUAUAUGAU consensusbinds FUBP1 and functions as an ESS capable of blocking splicingat the second step in vitro and inducing exon skipping in vivo.

FUBP1 Inhibits Pre-mRNA Splicing at the Second Step. We nextwished to analyze directly the function of FUBP1 in splicing, todetermine if it indeed contributes to the observed second-step

inhibition. To this end, we first added increasing amounts ofpurified recombinant FUBP1 to splicing reactions in NE con-taining each of the E10-derived substrates. As shown in Fig.6A, with two mutant substrates that retained the AU-rich ele-ment, β-E10M1 and β-M39, addition of increasing amounts ofFUBP1 caused 50–80% inhibition of the second step, estimatedfrom three independent experiments (lanes 1–3 and 4–6). How-ever, only a very modest effect on splicing of the β-F30M2 RNA,which lacks an intact ESS, was observed (Fig. 6A, lanes 7–9).Furthermore, FUBP1 had no effect on the stalled first-stepsplicing reaction observed with the β-E10 RNA itself (Fig. 6A,lanes 10–12). Together, these results provide evidence thatFUBP1 functions as a sequence-dependent second-step repressor.To provide additional evidence that FUBP1 functions in

splicing repression of β-E10 RNA, we wished to deplete theprotein from NE to determine whether such a depleted extractbecomes competent for the second-step of splicing. Our initialattempts to do this by using anti-FUBP1 antibodies were un-successful, because we were unable to deplete the protein effi-ciently. We, therefore, decided to attempt to deplete FUBP1 byanother approach, RNA affinity. The AU-rich sequence shownto bind to FUBP1 efficiently was used to produce a 30-nt bio-tinylated RNA containing three copies of the consensus motif(Bio-AU), and this was used to deplete FUBP1 from NE withstreptavidin beads. RNA affinity reduced the levels of FUBP1 toless than 10% of the amount in mock-depleted extracts (Fig. 6B,compare lane 2 with lane 1), with no effect on KSRP proteinlevels, as demonstrated by Western blotting using KSRP-specificantibodies (Fig. S5A). We then compared the splicing activity ofthe mock- and Bio-AU–depleted extracts with untreated NEusing the β-F30 substrate. Whereas the mock and untreatedextracts were inactive for the second step, the Bio-AU–depletedextract indeed displayed activity, albeit weak, in facilitating theexon ligation step (Fig. 6C, compare lane 3 with lanes 1–2).Furthermore, addition of purified FUBP1 specifically inhibitedappearance of fully spliced RNA (Fig. 6C, lanes 4–6). We rea-soned that the weak effect observed could reflect the presence ofthe inhibitory secondary structure within the β-F30 RNA. To testthis possibility, we constructed a substrate, β-M18, which con-tained 18 nt spanning the AUAUAUGAU consensus but lackedthe stem-forming region (Fig. 6D, Lower). Indeed, this sequencewas sufficient to induce the second-step block (Fig. 6D, lane 1).

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Fig. 5. FUBP1 binding correlates withESS activity. (A) Sequence of F30 RNA andmutant derivatives. The stem-forming re-gion (5′-aucaguaugc-3′) and putativeFUBP1-binding sites (5′-auauaugau-3′) areunderlined. Mutations were made withinthe FUBP1-binding sites as indicated. Notethat the stem-forming nucleotides wereunchanged. (B) The indicated 32P-labeledRNAs were incubated with increasingamounts of recombinant His-FUBP1 (100and 300 ng), and complexes were re-solved by nondenaturing PAGE. (C) Eachof the indicated RNA probes was incu-batedwith 150 ng of recombinant FUBP1,subjected to UV crosslinking, and ana-lyzed as in Fig. 4B. (D) HeLa cells weretransfected with constructs containingF30 or each of the indicated mutantderivatives. RNA was isolated and ana-lyzed by RT-PCR as in Fig. 2B. (E) Splicingassays in NE with RNA substrates con-taining F30 or the indicated mutantderivatives were performed and analyzedas in Fig. 1.

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However, most importantly, the Bio-AU–depleted extract nowdisplayed significant second-step activity (Fig. 6D, comparelane 3 with lanes 1–2), and addition of increasing amounts ofFUBP1 again specifically restored second-step inhibition (Fig.6D, lanes 4–6).

FUBP1 Causes Exon Skipping in Vivo. We next wished to determinewhether FUBP1 plays a role in exclusion of the E10 exon in vivo.To address this, siRNAs were used to reduce the levels ofFUBP1 in HeLa cells. As shown in Fig. 6E, both #1 and #3siRNA resulted in 70–80% decrease in FUBP1 accumulationcompared with the control (lanes 2 and 4, compare with lane 1),whereas no effects on FUBP1 expression were observed withsiRNA #2 (compare lane 3 with lane 1). KSRP protein levelswere not affected by any of the siRNAs (Fig. S5B). To examinethe effect of FUBP1 depletion independent of the E10 secondarystructure, we prepared a minigene construct analogous to thoseused above, except containing the 18-nt M18 sequence in themiddle exon. Cells transiently cotransfected with this constructand the control siRNA gave rise to predominantly exon-skippedRNA, as determined by RT-PCR, although exclusion was notcomplete, perhaps reflecting the absence of the secondarystructure (Fig. 6F, lane1). Strikingly, knockdown of FUBP1 byeither siRNA #1 or #3 significantly increased exon inclusioncompared with the control or siRNA #2 (Fig. 6F, compare lanes2 and 4 with lanes 1 and 3), indicating that FUBP1 indeedinduces exon exclusion.

FUBP1 Regulates Alternative Splicing of Endogenous Pre-mRNAs. Wenext investigated whether FUBP1 can regulate splicing of en-dogenous transcripts. To this end, we designed primer pairs thatdetect alternative exons in 10 transcripts regulated by other AU-rich binding proteins, such as HuR and TIA-1/TIAR (34–37), aswell as a number of other potential target transcripts, chosenbased on their cancer relevance, reflecting the emerging role ofFUBP1 in cancer (see below), and used these to analyze RNAisolated from control and each of the FUBP1 siRNA-treatedcells. Although there were no obvious changes in the splicingpatterns of 51 transcripts examined, 4 revealed significant dif-ferences. Specifically, inclusion of exon 14 of the pre-mRNAencoding ATP citrate lyase (ACLY) and exons 4–7 of the cas-pase 9 pre-mRNA were increased upon FUBP1 knockdown (Fig.6F, compare lanes 2 and 4 with lanes 1 and 3). On the otherhand, increased exclusion of PTBP2 exon 10 and ENAH/MENAexon 11 were observed in the FUBP1-depleted cells (Fig. 6F,compare lanes 2 and 4 with lanes 1 and 3). Significantly, over-expression of Flag-tagged FUBP1 switched splicing in the oppositedirection (Fig. S6). The effects of overexpression were relativelymodest, likely reflecting the fact that FUBP1 is highly abundant inHeLa cells (∼3 × 106 molecules per cell; Fig. S7). These dataprovide evidence that FUBP1 functions in control of AS of en-dogenous transcripts, and, as discussed below, can function bothpositively and negatively in regulating exon inclusion.

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Fig. 6. FUBP1 inhibits pre-mRNA splicing at the second step in vitro and modulates exon skipping in vivo. (A) In vitro splicing was performed in NE in thepresence of increasing amounts of recombinant FUBP1 (100 and 300 ng) with indicated RNAs. The presence or absence of the intact ESS in the RNAs is in-dicated above. (B) A biotinylated RNA (Bio-AU) 20-mer containing the E10 AU-rich element was used to deplete FUBP1 from NE. Mock depletion was donewith beads alone. The resultant extracts were analyzed by Western blotting with anti-FUBP1 and anti-actin antibodies. (C) Extracts from B were used forsplicing with β-F30 RNA (lanes 2 and 3), and standard NE was used for comparison (lane 1). FUBP1-depleted extracts were supplemented with increasingamount of FUBP1 (0, 100, and 300 ng) and analyzed for splicing (lanes 4–6). (D) In vitro splicing with the β-M18 RNA was performed essentially as above. The18-nt sequence is shown at the bottom, with the FUBP1-binding site underlined. (E) Whole-cell lysates were prepared from FUBP1 siRNA- and control siRNA-transfected cells and analyzed by Western blotting using an anti-FUBP1 antibody. (F) The M18-containing minigene plasmid and each siRNA werecotransfected into HeLa cells, and total cellular RNA was analyzed by RT-PCR as above (top gel). Alternatively spliced isoforms of endogenous ACLY, Caspase9,ENAH, and PTBP2 RNAs from each of the FUBP1 siRNA- and control siRNA-transfected cells were analyzed by RT-PCR (bottom gels). Quantification of theirRNA products was measured as inclusion/exclusion (In/Ex) ratio with SD and indicated below each gel. RNA products are indicated schematically on the right.

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DiscussionIn this paper, using a chimeric β-globin–triadin exon 10 RNAsubstrate (β-E10), we have shown that a previously describedsecond-step inhibition results from two separate inhibitory ele-ments. The first is a serendipitously created stem–loop structuredownstream of the 3′SS, and the other is an ESS situated 3′ tothe structure that depends on the RNA/DNA-binding proteinFUBP1 (Fig. 7). Our data suggest that these two independentmechanisms operate simultaneously to block splicing at thesecond step, because disrupting either of them rescues the sec-ond-step defect (Fig. 7). We also demonstrated that these twoelements, and FUBP1, contribute to exon skipping in vivo. Be-low, we discuss the features of FUBP1-regulated splicing com-pared with previously reported examples of splicing regulation,how this function of FUBP1 might be relevant to its role in diseaseand the function of RNA secondary structure in splicing regula-tion. It is striking that both of these mechanisms result in an un-usual second-step inhibition, and we discuss the basis for this andhow this type of inhibition may relate to exon exclusion in vivo.Our data have shown that an AU-rich sequence in triadin exon

10 acts as an FUBP1-dependent ESS. Although FUBP1 wasoriginally identified as a ssDNA-binding protein that modulatesMYC expression (22, 38, 39), the protein was later demonstrated tobe a member of the AU-rich element (ARE)-binding proteinfamily (40) and has subsequently been found to be involved inseveral aspects of RNA metabolism other than splicing. For ex-ample, FUBP1 has been reported to interact with the poly(U) tractof the hepatitis C virus 3′ UTR and is required for its efficientreplication (28). FUBP1 may also function in stabilization of cer-tain ARE-containing transcripts (29, 40) and was recently shown tobind the 3′ UTR of nucleophosmin mRNA to repress translation(23) and to the 5′UTR of CDKN1BmRNA to enhance translation(24). These findings suggest that FUBP1 can function in biologicalprocesses occurring in both the cytoplasm and the nucleus.The involvement of FUBP1 in splicing regulation was un-

anticipated. However, several other ARE-binding proteins havepreviously been implicated in regulation of AS, in addition totheir roles in RNA metabolism events occurring in the cytoplasm(34, 36, 41). HuR, well characterized for its role in post-translational regulation of AU- and U-rich mRNAs (42), wasrecently found to enhance skipping of alternative exons inZNF207 and PTBP2 pre-mRNAs, likely via interactions withintronic AU-rich elements (34). Likewise, TIA-1/TIAR has beenshown to regulate translation of various mRNAs by binding to

AU-rich elements located in the 3′ UTR. In the nucleus, theseproteins act as splicing regulators of several alternatively splicedpre-mRNAs (36, 43, 44). Our data have demonstrated that FUBP1,like these ARE-binding proteins, functions as an AS regulator.Significantly, FUBP1 can either increase or decrease exon in-clusion. This property likely depends on the location of its bindingsite within the pre-mRNA; such position-dependent activity hasbeen shown previously for a number of splicing regulators (5, 7).FUBP1, like HuR, regulates skipping of exon 10 of the pre-

mRNA encoding the splicing regulator PTBP2 (also known asnPTB). It is intriguing although that HuR contributes to exon 10exclusion, whereas FUBP1 facilitates inclusion. Skipping of thisexon causes nonsense-mediated decay, and is thought to be usedin the cross-regulation of expression of PTB itself (45). Ourresults suggest a possible interplay involving FUBP1 and thesesplicing regulators in AS control. Reduced levels of FUBP1 in-crease skipping of nPTB exon 10, resulting in decreased nPTBexpression, which, in turn, up-regulates PTB expression. A switchin expression of nPTB/PTB has been proposed to be critical forcell-fate determination (46), and it will, thus, be of interest todetermine how the cross-regulatory network involving thesefactors, including FUBP1, functions in development. For exam-ple, inclusion of triadin exon 10 varies during cardiac develop-ment in the mouse (20), and it will be important to determinewhether FUBP1 functions in this process.Deregulation of FUBP1 expression has recently been impli-

cated in several cancers. For example, two groups observed el-evated expression of FUBP1 in more than 70% of human hepa-tocellular carcinoma (HCC) samples and also provided evidencethat FUBP1 plays an important role in tumor growth (47, 48).Notably, they also revealed no significant correlations betweenFUBP1 and MYC expression in HCC cells, suggesting thatFUBP1-dependent regulation of MYC plays a minor role inhepatocarcinogenesis, although evidence for a role in renal car-cinomas has been presented (49). In addition, FUBP1 targetgenes other than MYC such as those encoding the microtubule-destabilizing protein stathmin (48, 50) and the cell-cycle regu-lator CDKN1A (47), were found to be involved in tumor-rele-vant functions. More recently, mutations in FUBP1 were shownto contribute to another type of cancer, oligodendroglioma (51).All these findings indicate that multiple functions of FUBP1 maybe critical for its role in tumorigenesis, and our data suggest thatderegulation of cancer-related AS events, such as Caspase9 (52)and ENAH/MENA (53), by FUBP1 might be involved in thisprocess. Aberrant AS events have been implicated in many types

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Fig. 7. Model for splicing regulation by FUBP1binding to the ESS and by RNA secondary structure.(A) Schematic representation of the β-E10 pre-mRNAin which the intron can be processed only throughthe first catalytic step of splicing. The positions of 5′SS, branch point (BP), and 3′SS, the stem–loopdownstream of the 3′SS, and the FUBP1-dependent(blue oval) ESS are shown. The green arrow repre-sents first-step splicing. (B) RNA secondary structureand binding of FUBP1 to the ESS both block thesecond step, leading to accumulation of free exon 1,and exon 2-intron in a lariat configuration. Thegreen arrow represents the second-step, and Xindicates blockage of this step. These two elementscontribute to exon skipping in vivo. (C) Disruption ofthe secondary structure can relieve the inhibitoryeffect caused by binding of FUBP1 to the ESS andthus activate the second step, leading to exon in-clusion in vivo. (D) Disruption of the ESS or depletionof FUBP1 binding enhances the second step even inthe presence of RNA secondary structure, resultingin partial exon inclusion in vivo.

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of cancer, including HCC and gliomas (54, 55). Thus, it will beimportant to determine whether alterations in these AS eventscaused by deregulated expression of FUBP1 contribute to tumorcell proliferation.There are now a number of examples of RNA secondary struc-

tures that modulate AS. Such structures often function by blockingor interfering with recognition of the core splicing signals (16). Inaddition, several genome-wide studies investigating the potential ofRNA secondary structure formation near such core elements sup-ports the idea that these structures can indeed be involved insplicing regulation (56, 57). Our findings that an exonic stem–loopclose to the 3′SS induced a splicing block in vitro and exon skippingin vivo provide further support for this theory. A unique feature ofthis regulation in our case is that the second step is specificallyaffected, as opposed to a more typical first-step inhibition (16).Traditional models of splicing repression target early stages of

splice-site recognition and/or early spliceosome assembly, andindeed many examples of this type of regulation, have been de-scribed (5, 6). However, several studies have shown that in-hibition can also occur at later stages of spliceosome assemblyand even during conformational changes between the twotransesterification steps (13, 58, 59). This is further exemplifiedby the unique functions of FUBP1 as a second-step repressordescribed in our study. Similar to the role of Drosophila Sxl insplicing autoregulation (13), we speculate that binding of FUBP1interferes with functions of protein factors required for the secondstep, thus preventing exon ligation by the splicing machinery. It isintriguing that second-step inhibition was also caused by sec-ondary structure close to the 3′SS with the β-E10 RNA usedhere. It is possible that the structure might block a helicase ac-tivity required for the exon–exon ligation step, as was shownpreviously with a yeast ACT-LacZ fusion transcript with a struc-ture similar to that of β-E10 RNA at the 3′SS (19). How bothRNA secondary structure and the FUBP1-dependent ESS resultin an unusual second-step block with the β-E10 RNA is currentlyunclear and will be an important focus of future studies.In summary, the results presented here have provided additional

insights into mechanisms of splicing control. FUBP1 representsa distinct type of splicing regulatory protein in that it binds towhat appears to be a typical ESS but then inhibits splicing at thesecond step, which presumably underlies its ability to induce ESS-dependent exon skipping in vivo. A nearby exonic RNA structurewas also shown to induce exon skipping in vivo and to blocksplicing at the second step in vitro. It will be of interest in thefuture to learn how widely these two mechanisms are used toregulate AS in human cells, as well as to investigate further therole of FUBP1 as a splicing regulator in development and disease.

Materials and MethodsPlasmids Constructions. β-E9 and β-E10 plasmids were described previously(20). All of the β-E10 truncations were constructed by placing indicatedsequences (F30, L24, M39, and M18) between AccI and BamHI sites to sub-stitute for E10 sequences. Mutations on the β-E10 and β-F30 plasmids werecreated by site-directed mutagenesis, with specific sequences shown in thefigures. Plasmids were linearized by BamHI or by AccI as indicated and usedfor making RNAs for in vitro-splicing assays. For transcribing RNAs for gel-shift and UV-crosslinking assays, corresponding sequences (F30, F30M1,F30M2, and F30M3) were inserted between HindIII and BamHI sites in theβ-globin construct. In vivo minigene plasmids were constructed by subclon-ing modified β-globin genomic sequences into pcDNA3.1(+). Specifically,4 bp of the first exon of β-globin adjacent to the 5′ splice site, the first intronand the second exon were inserted downstream of the β-E9 or β-E10 plas-mid, generating a construct containing three exons with E9 or E10 in themiddle exon flanked by two identical introns. Indicated sequences wereinserted in the internal exon for splicing assays in vivo. The length of E10 was54 nt; F30 represents the first 30 nt within E10, and L24 represents the last 24nt of E10. M39 and M18, containing 39 and 18 nt, respectively, located in themiddle region of E10. The underlined ATATATGAT consensus is the putativeFUBP1-binding site: M39, 5′-GCATTCTGTCGATATATGATTGACATGTTTGTC-CATGGG; M18, 5′- TCTGTCGATATATGATTG.

In Vitro Transcription and Splicing. In vitro transcription and splicing assays inHeLa NE was performed essentially as described previously (20, 21).

RNA Gel-Shift and UV-Crosslinking Assay. 32P-labeled F30 RNA and each of itsderivatives were incubated with recombinant His-tagged human FUBP1 (100and 300 ng) as described previously (4, 60).

RNA Affinity and FUBP1 Depletion. For RNA-affinity assays, 1 nmol of 5′ biotin-labeled F30 RNA was mixed with 100 μL of HeLa NE in 500 μL of bindingbuffer [10 mM Hepes (pH7.9), 100 mM KCl, 10% glycerol, 2 mM MgCl2, 0.75mM ATP, 25 mM creatine phosphate, 30 μg/mL tRNA]. After incubation at30 °C for 40 min, reaction mixtures were briefly centrifuged and 50 μL ofstreptavidin agarose (Sigma) was added to the supernatant. Reaction mix-tures were then incubated at 4 °C on a rotating wheel for 1 h. Proteins wereeluted from the beads with SDS loading buffer, resolved by SDS/PAGE, andstained with Coomassie blue. Protein bands were excised and analyzed byMS as described previously (61).

FUBP1 depletion was conducted using an RNA-affinity assay. A 5′ biotin-labeled RNA (Bio-AU) was synthesized containing three copies of the puta-tive FUBP1-binding sequence (5′-AUAUAUGAU); 5 nmol of Bio-AU RNAbound to 60 μL of streptavidin agarose beads was pelleted, and 200 μL of NEwas added and rotated at 4 °C overnight. Mock depletion was done withbeads alone. Depletion efficiency was determined by Western blot.

Splicing Extracts and Recombinant Proteins. NE, S100, NF40-60, His-SRp38,SRSF1, and SRSF2 were prepared essentially as described (20, 21). HumanFUBP1 cDNA was first cloned into pQE80L vector (Qiagen) containing a His6tag sequence and then subcloned into pFastBacTM HT B (Invitrogen) for His-FUBP1-baculovirus production. Sf9 and High-Five cells were used for bacu-lovirus production and His-FUBP1 expression, respectively. His-FUBP1 waspurified by Ni2+ agarose under native conditions and dialyzed againstbuffer D.

Cell Culture, Transfection, RT-PCR, and Western Blot. HeLa cells were culturedas adherent cells in Dulbecco’s modified Eagle medium with 10% FBS.Minigene constructs were transfected by using lipofectamine (Invitrogen)following the manufacturer’s instructions. RNA extraction and reversetranscription were all carried out as described previously (20). For PCRanalysis of minigene splicing, the forward (F) primer on exon 1 was 5′-ACTTAAGCTTGCTTACATTTGC, and the reverse (R) primer on exon 3 was 5′-ACTCAAAGAACCTCTGGGTC. Sequences for primer sets used in the studywere as follows: ACLY-F, 5′-CAAACTTCCTCCTCAACGC; ACLY-R, 5′-GAGGG-TGGTGCTCTTTCC; Caspase9-F, 5′-GACCAGTGGACATTGGTTC; Caspase9-R, 5′-GGTCCCTCCAGGAAACAA; ENAH-F, 5′-TGCTGGCCAGGAGGAGAAG; ENAH-R,5′-ACTGGGCTGTGATAAGGGTG; PTBP2-F, 5′-GGCAATACAGTCCTGTTGGT; andPTBP2-R, 5′-ATGGCAAGTTGTGATTGGTT.

Sequences for other primer pairs will be provided upon request. Westernblot was carried out as described previously (20). Primary antibodies used,mouse anti-FUBP1 (catalog no. sc-136137) and mouse anti-actin, werepurchased from Santa Cruz Biotechnology. The monoclonal anti-KSRPantibody (62) was a kind gift from D. Black (University of California, LosAngeles, CA).

RNA Interference. siRNA sequences were designed using Clontech RNAi TargetSequence Selector. All siRNAs were synthesized at GenePharma, and thefollowing sequences were sense strands of chosen siRNA duplexes: siRNA-FUBP1 (#1), GGAGGAGUUAACGACGCUUTT; siRNA-FUBP1 (#2), GCAGCAA-AGCAGAUCUGUATT; siRNA-FUBP1 (#3), CUGGAACACCUGAAUCUGUTT; andsiRNA control, UUCUCCGAACGUGUCACGUTT. siRNA duplexes were trans-fected at 120 pmol per well (12-well plate) by using Lipofectamine 2000(Invitrogen) following manufacturer’s instructions. Transfected cells wereharvested for RNA isolation or protein extraction 48 h after siRNA trans-fection. siRNA and minigene coupled transfection assay was carried out asdescribed previously (63). In brief, siRNA transfection (120 pmol per well;12-well plate) was performed when HeLa cells reached 30–40% conflu-ence. Minigene plasmids (0.6 μg per well; 12-well plate) were transfected24 h after siRNA transfection. After another 24 h, cells were collected forfurther analysis.

RNA Secondary Structure Prediction. RNA secondary structure prediction wasconducted using the Vienna RNAfold Web server (26).

ACKNOWLEDGMENTS. We thank Dr. D. Black for his generous gifts ofmonoclonal anti-KSRP antibodies. This work was supported by NationalInstitutes of Health Grant GM48259 (to J.L.M.) and by grants from the

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Ministry of Science and Technology of China (973 Program 2012CB524900),National Natural Science Foundation Grants 31170753 and 31070704, the

One Hundred Talents Program of the Chinese Academy of Sciences (Y.F.),and Shanghai Pujiang Program Grant 10PJ1411100.

1. Zhou Z, Licklider LJ, Gygi SP, Reed R (2002) Comprehensive proteomic analysis of thehuman spliceosome. Nature 419(6903):182–185.

2. Jurica MS, Moore MJ (2003) Pre-mRNA splicing: Awash in a sea of proteins. Mol Cell12(1):5–14.

3. Singh RK, Cooper TA (2012) Pre-mRNA splicing in disease and therapeutics. TrendsMol Med 18(8):472–482.

4. David CJ, Chen M, Assanah M, Canoll P, Manley JL (2010) HnRNP proteins controlledby c-Myc deregulate pyruvate kinase mRNA splicing in cancer. Nature 463(7279):364–368.

5. ChenM, Manley JL (2009) Mechanisms of alternative splicing regulation: Insights frommolecular and genomics approaches. Nat Rev Mol Cell Biol 10(11):741–754.

6. Nilsen TW, Graveley BR (2010) Expansion of the eukaryotic proteome by alternativesplicing. Nature 463(7280):457–463.

7. Licatalosi DD, Darnell RB (2010) RNA processing and its regulation: Global insightsinto biological networks. Nat Rev Genet 11(1):75–87.

8. Ule J, et al. (2006) An RNA map predicting Nova-dependent splicing regulation.Nature 444(7119):580–586.

9. Zhou HL, Lou H (2008) Repression of prespliceosome complex formation at twodistinct steps by Fox-1/Fox-2 proteins. Mol Cell Biol 28(17):5507–5516.

10. Wang Z, et al. (2004) Systematic identification and analysis of exonic splicing silencers.Cell 119(6):831–845.

11. Yu Y, et al. (2008) Dynamic regulation of alternative splicing by silencers thatmodulate 5′ splice site competition. Cell 135(7):1224–1236.

12. Zhang XH, Chasin LA (2004) Computational definition of sequence motifs governingconstitutive exon splicing. Genes Dev 18(11):1241–1250.

13. Lallena MJ, Chalmers KJ, Llamazares S, Lamond AI, Valcárcel J (2002) Splicingregulation at the second catalytic step by Sex-lethal involves 3′ splice site recognitionby SPF45. Cell 109(3):285–296.

14. Grover A, et al. (1999) 5′ splice site mutations in tau associated with the inheriteddementia FTDP-17 affect a stem-loop structure that regulates alternative splicing ofexon 10. J Biol Chem 274(21):15134–15143.

15. Hiller M, Zhang Z, Backofen R, Stamm S (2007) Pre-mRNA secondary structuresinfluence exon recognition. PLoS Genet 3(11):e204.

16. Warf MB, Berglund JA (2010) Role of RNA structure in regulating pre-mRNA splicing.Trends Biochem Sci 35(3):169–178.

17. Jin Y, Yang Y, Zhang P (2011) New insights into RNA secondary structure in thealternative splicing of pre-mRNAs. RNA Biol 8(3):450–457.

18. Halfter H, Gallwitz D (1988) Impairment of yeast pre-mRNA splicing by potentialsecondary structure-forming sequences near the conserved branchpoint sequence.Nucleic Acids Res 16(22):10413–10423.

19. Lin J, Rossi JJ (1996) Identification and characterization of yeast mutants thatovercome an experimentally introduced block to splicing at the 3′ splice site. RNA2(8):835–848.

20. Feng Y, et al. (2009) SRp38 regulates alternative splicing and is required for Ca(2+)handling in the embryonic heart. Dev Cell 16(4):528–538.

21. Feng Y, Chen M, Manley JL (2008) Phosphorylation switches the general splicingrepressor SRp38 to a sequence-specific activator.Nat StructMol Biol 15(10):1040–1048.

22. Duncan R, et al. (1994) A sequence-specific, single-strand binding protein activatesthe far upstream element of c-myc and defines a new DNA-binding motif. Genes Dev8(4):465–480.

23. Olanich ME, Moss BL, Piwnica-Worms D, Townsend RR, Weber JD (2011) Identificationof FUSE-binding protein 1 as a regulatory mRNA-binding protein that repressesnucleophosmin translation. Oncogene 30(1):77–86.

24. Zheng Y, Miskimins WK (2011) Far upstream element binding protein 1 activatestranslation of p27Kip1 mRNA through its internal ribosomal entry site. Int J BiochemCell Biol 43(11):1641–1648.

25. Labourier E, et al. (1999) Recognition of exonic splicing enhancer sequences by theDrosophila splicing repressor RSF1. Nucleic Acids Res 27(11):2377–2386.

26. Gruber AR, Lorenz R, Bernhart SH, Neubock R, Hofacker IL (2008) The Vienna RNAwebsuite. Nucleic Acids Res 36(Web Server issue):W70–W74.

27. He L, et al. (2000) Loss of FBP function arrests cellular proliferation and extinguishes c-myc expression. EMBO J 19(5):1034–1044.

28. Zhang Z, Harris D, Pandey VN (2008) The FUSE binding protein is a cellular factorrequired for efficient replication of hepatitis C virus. J Virol 82(12):5761–5773.

29. Irwin N, Baekelandt V, Goritchenko L, Benowitz LI (1997) Identification of twoproteins that bind to a pyrimidine-rich sequence in the 3′-untranslated region of GAP-43 mRNA. Nucleic Acids Res 25(6):1281–1288.

30. Min H, Turck CW, Nikolic JM, Black DL (1997) A new regulatory protein, KSRP,mediates exon inclusion through an intronic splicing enhancer. Genes Dev 11(8):1023–1036.

31. Labourier E, Adams MD, Rio DC (2001) Modulation of P-element pre-mRNA splicingby a direct interaction between PSI and U1 snRNP 70K protein. Mol Cell 8(2):363–373.

32. Cukier CD, et al. (2010) Molecular basis of FIR-mediated c-myc transcriptional control.Nat Struct Mol Biol 17(9):1058–1064.

33. Benjamin LR, et al. (2008) Hierarchical mechanisms build the DNA-binding specificityof FUSE binding protein. Proc Natl Acad Sci USA 105(47):18296–18301.

34. Lebedeva S, et al. (2011) Transcriptome-wide analysis of regulatory interactions of theRNA-binding protein HuR. Mol Cell 43(3):340–352.

35. Förch P, et al. (2000) The apoptosis-promoting factor TIA-1 is a regulator of alter-native pre-mRNA splicing. Mol Cell 6(5):1089–1098.

36. Förch P, Puig O, Martínez C, Séraphin B, Valcárcel J (2002) The splicing regulator TIA-1interacts with U1-C to promote U1 snRNP recruitment to 5′ splice sites. EMBO J 21(24):6882–6892.

37. Zhu H, Hasman RA, Young KM, Kedersha NL, Lou H (2003) U1 snRNP-dependentfunction of TIAR in the regulation of alternative RNA processing of the humancalcitonin/CGRP pre-mRNA. Mol Cell Biol 23(17):5959–5971.

38. Liu J, et al. (2000) The FBP interacting repressor targets TFIIH to inhibit activatedtranscription. Mol Cell 5(2):331–341.

39. Braddock DT, Louis JM, Baber JL, Levens D, Clore GM (2002) Structure and dynamics ofKH domains from FBP bound to single-stranded DNA. Nature 415(6875):1051–1056.

40. Dean JL, Sully G, Clark AR, Saklatvala J (2004) The involvement of AU-rich element-binding proteins in p38 mitogen-activated protein kinase pathway-mediated mRNAstabilisation. Cell Signal 16(10):1113–1121.

41. Ince-Dunn G, et al. (2012) Neuronal Elav-like (Hu) proteins regulate RNA splicing andabundance to control glutamate levels and neuronal excitability. Neuron 75(6):1067–1080.

42. Hinman MN, Lou H (2008) Diverse molecular functions of Hu proteins. Cell Mol Life Sci65(20):3168–3181.

43. Gal-Mark N, Schwartz S, Ram O, Eyras E, Ast G (2009) The pivotal roles of TIA proteinsin 5′ splice-site selection of alu exons and across evolution. PLoS Genet 5(11):e1000717.

44. Izquierdo JM, et al. (2005) Regulation of Fas alternative splicing by antagonisticeffects of TIA-1 and PTB on exon definition. Mol Cell 19(4):475–484.

45. Spellman R, Llorian M, Smith CW (2007) Crossregulation and functional redundancybetween the splicing regulator PTB and its paralogs nPTB and ROD1. Mol Cell 27(3):420–434.

46. Boutz PL, et al. (2007) A post-transcriptional regulatory switch in polypyrimidine tract-binding proteins reprograms alternative splicing in developing neurons. Genes Dev21(13):1636–1652.

47. Rabenhorst U, et al. (2009) Overexpression of the far upstream element bindingprotein 1 in hepatocellular carcinoma is required for tumor growth. Hepatology50(4):1121–1129.

48. Malz M, et al. (2009) Overexpression of far upstream element binding proteins: Amechanism regulating proliferation and migration in liver cancer cells. Hepatology50(4):1130–1139.

49. Weber A, et al. (2008) The FUSE binding proteins FBP1 and FBP3 are potential c-mycregulators in renal, but not in prostate and bladder cancer. BMC Cancer 8:369.

50. Singer S, et al. (2009) Coordinated expression of stathmin family members by farupstream sequence element-binding protein-1 increases motility in non-small celllung cancer. Cancer Res 69(6):2234–2243.

51. Bettegowda C, et al. (2011) Mutations in CIC and FUBP1 contribute to humanoligodendroglioma. Science 333(6048):1453–1455.

52. Shultz JC, et al. (2011) SRSF1 regulates the alternative splicing of caspase 9 via a novelintronic splicing enhancer affecting the chemotherapeutic sensitivity of non-small celllung cancer cells. Mol Cancer Res 9(7):889–900.

53. Di Modugno F, et al. (2012) Splicing program of human MENA produces a previouslyundescribed isoform associated with invasive, mesenchymal-like breast tumors. ProcNatl Acad Sci USA 109(47):19280–19285.

54. Berasain C, et al. (2010) Impairment of pre-mRNA splicing in liver disease: Mechanismsand consequences. World J Gastroenterol 16(25):3091–3102.

55. David CJ, Manley JL (2010) Alternative pre-mRNA splicing regulation in cancer:Pathways and programs unhinged. Genes Dev 24(21):2343–2364.

56. Shepard PJ, Hertel KJ (2008) Conserved RNA secondary structures promote alternativesplicing. RNA 14(8):1463–1469.

57. Zhang J, Kuo CC, Chen L (2011) GC content around splice sites affects splicing throughpre-mRNA secondary structures. BMC Genomics 12:90.

58. House AE, Lynch KW (2006) An exonic splicing silencer represses spliceosome assemblyafter ATP-dependent exon recognition. Nat Struct Mol Biol 13(10):937–944.

59. Sharma S, Kohlstaedt LA, Damianov A, Rio DC, Black DL (2008) Polypyrimidine tractbinding protein controls the transition from exon definition to an intron definedspliceosome. Nat Struct Mol Biol 15(2):183–191.

60. Zuo P, Manley JL (1994) The human splicing factor ASF/SF2 can specifically recognizepre-mRNA 5′ splice sites. Proc Natl Acad Sci USA 91(8):3363–3367.

61. Shi Y, Manley JL (2007) A complex signaling pathway regulates SRp38 phos-phorylation and pre-mRNA splicing in response to heat shock. Mol Cell 28(1):79–90.

62. Hall MP, Huang S, Black DL (2004) Differentiation-induced colocalization of the KH-type splicing regulatory protein with polypyrimidine tract binding protein and thec-src pre-mRNA. Mol Biol Cell 15(2):774–786.

63. Kashima T, Manley JL (2003) A negative element in SMN2 exon 7 inhibits splicing inspinal muscular atrophy. Nat Genet 34(4):460–463.

Li et al. PNAS | Published online July 1, 2013 | E2695

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