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Human Molecular Genetics Advance Access originally published online on July 6, 2005
Human Molecular Genetics 2005 14(16):2289-2303; doi:10.1093/hmg/ddi233
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© The Author 2005. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oupjournals.org

Up-regulation of the ubiquitous alternative splicing factor Tra2ß causes inclusion of a germ cell-specific exon

Julian P. Venables1,*,{dagger}, Cyril F. Bourgeois2,{dagger}, Caroline Dalgliesh1, Liliane Kister2, James Stevenin2 and David J. Elliott1

1Institute of Human Genetics, University of Newcastle upon Tyne, International Centre for Life, Central Parkway, Newcastle upon Tyne NE1 3BZ, UK and 2Institut de Genetique et de Biologie Moleculaire et Cellulaire, CNRS/INSERM/ULP, 67404 Illkirch, C.U. Strasbourg, France

* To whom correspondence should be addressed. Tel: +44 1912418636; Fax: +44 1912418666; Email: j.venables{at}ncl.ac.uk

Received March 22, 2005; Accepted June 24, 2005

GenBank accession nos AY425955–AY425960


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
We have discovered a new exon of the homeodomain-interacting kinase HipK3 that incorporates a premature stop codon and is included only in the human testis. To investigate this, we tested the effects of transfecting cells with green fluorescent protein fusions of RNA-binding proteins implicated in spermatogenesis using a novel assay based on multi-fraction fluorescence-activated cell sorting (MF-FACS). This allows the effect of a controlled titration of any splicing factor on the splicing of endogenous genes to be studied in vivo. We found that Tra2ß recapitulates testis-specific splicing of endogenous HipK3 in a concentration-dependent manner and binds specifically to a long purine-rich sequence in the novel exon. This sequence was also specifically bound by hnRNP A1, hnRNP H, ASF/SF2 and SRp40, but not by 9G8. Consistent with these observations, in vitro studies showed that this sequence shifts splicing to a downstream 5' splice site within a heterologous pre-mRNA substrate in the presence of Tra2ß, ASF/SF2 and SRp40, whereas hnRNP A1 specifically inhibits this choice. By mutating the purine-rich sequence in the context of the HipK3 gene, we also show that it is the major determinant of Tra2ß- and hnRNP A1-mediated regulation. Tra2 is essential for sex determination and spermatogenesis in flies, and Tra2ß protein was most highly expressed in testis out of six mouse tissues, whereas hnRNP A1 is down-regulated during germ cell development. Therefore, our data imply an evolutionarily conserved role for Tra2 proteins in spermatogenesis and suggest that an elevated concentration of Tra2ß may convert it into a tissue-specific splicing factor.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Alternative splicing is an important means of regulating gene expression which controls the tissue specificity of gene expression of a distinct set of genes from transcriptional control (1Go). However it is only known in a handful of cases how tissue-specific isoforms arise as they are thought to mainly depend on complex subtle changes in the levels of multiple ubiquitous semi-redundant splicing factors, rather than on unique tissue-specific expression of individual splicing factors (2Go,3Go). However, a recent study in Drosophila cells has shown that targeted down-regulation of most individual splicing factors has specific effects on the splicing of unique subsets of genes (4Go). To understand the phenomenon of regulated tissue-specific alternative splicing, we have concentrated on human testis, as this is one of the tissues in which alternative splicing is most prevalent (5Go). However, although several testis-specific splicing factors and exons are known, so far no causal relationship between these has been demonstrated (6Go–8Go).

Traditionally, verification of alternative splicing activity is demonstrated by constructing a shortened ‘minigene’, which contains the regulated exons and splicing signals next to each other (9Go). The minigene is then transcribed and assayed with recombinant protein in vitro or co-transfected with candidate splicing factors. Minigenes can yield artefactual results due to removal or artificial creation of secondary structures (10Go–12Go) or by altering co-transcriptional splicing kinetics (13Go–15Go). Although it is possible to study coupled transcription/splicing in vitro (16Go), the production of recombinant RNA-binding proteins in correctly modified soluble form takes considerable optimization and is not always successful. The closest simulation of alternative splicing in cell lines would be to assay splicing changes of endogenous transcripts, rather than minigenes. With a minigene, the assumption is that it is successfully transfected into the same cells as the splicing factor being investigated. However, the reproducible study of endogenous splicing requires a high and reproducible transfection rate, but these are often considerably <50%, and there is a highly variable accumulation of splicing factor levels between individual transfected cells. To overcome these problems, we have developed a simple assay to test alternative splicing of endogenous genes in cell lines using automated multi-fraction fluorescence-activated cell sorting (MF-FACS) of cells transfected with green fluorescent protein (GFP) fusions to splicing factors. Taking advantage of automated separation of cell populations expressing distinct and largely non-overlapping levels of GFP-fused splicing factors has allowed controlled titrations of exogenous splicing factors in vivo for the first time.

We uncovered an alternative exon of the homeodomain-interacting protein kinase HipK3, a member of a gene family that has been implicated in malignant growth and apoptosis, and we showed that this exon is only expressed in human testis. Using our novel FACS-splicing assay, we showed that it is incorporated upon ectopic expression of the alternative splicing factor Tra2ß, which is one of a group of proteins containing several ‘RS’ di-peptides, including ‘SR’ proteins, that function in constitutive and regulated splicing pathways (17Go–20Go). Tra2ß mRNA has previously been shown to be ubiquitously expressed, although at a higher level in human testis (21Go). Here, we show in mouse tissues that Tra2ß is also most highly expressed at the protein level in testis and therefore that elevated concentrations of this ubiquitous splicing factor in human testis could be the cellular switch regulating the alternative splicing of HipK3.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Testis-specific alternative splicing of HipK3
Recently, considerable progress has been made in the detection of novel alternatively spliced transcripts by simply mass-screening candidate genes by RT–PCR, agarose gel analysis and sequencing. In one study, novel alternatively spliced transcripts were found in 70% of the genes assayed (22Go). We have used a similar method to discover alternative splices involved in spermatogenesis. We designed primers to amplify short internal candidate regions from 74 ubiquitous genes with important developmental functions from testis and brain cDNA. Products were electrophoresed on an agarose gel and six bands, other than the expected size found in testis but not in brain, were cloned and sequenced (GenBank accession nos AY425955–AY425960). Tissue specificity was then tested more rigorously on 10 human tissues, and although five of the six splices were variably expressed across the other eight tissues (data not shown), one of the alternative splices (AY425958) which was of the 1215 amino acid homeodomain- and Fas-interacting protein kinase HipK3 (unigene Hs.201918) was found only in testis (Fig. 1A, top panel). The organization of the HipK3 gene is shown in Figure 1C. The novel splice form introduces a 109 nucleotide extra exon (Fig. 1C) and the deduced amino acid sequence of the encoded protein is identical for the N-terminal 407 amino acids, after which four novel residues DRRY (followed by stop) are introduced. Database searching revealed that this was not the only alternatively spliced exon of HipK3. Another alternatively spliced exon peptide cassette (GenBank accession no. AF004849) encoding 21 extra amino acids in frame exists further downstream (Fig. 1C). The presence of this was tested by RT–PCR on the tissue panel and it appears to be incorporated in between one-third and two-thirds of the time in different tissues (Fig. 1A, bottom panel). However, although RT–PCR amplification of two alternative splices with a single primer pair can give a reasonable approximation of their actual levels (23Go), shorter products will be amplified faster and therefore, we have quantified our results as an indicative ratio of PCR products, but we cannot infer the actual proportions in the starting pool with any defined accuracy. The two alternative exons of HipK3 fit the typical profiles of two different general classes of alternative exons (24Go). The T exon is not conserved in the corresponding region of the HipK3 gene in mouse; it is close to the median size (104 bases) for ‘genome-specific’ exons, and like the majority of these it introduces a stop codon. The U exon is conserved in the mouse genome; its smaller size reflects the median size for conserved alternative exons (76 bases), and as for three-quarters of these it encodes an in-frame peptide cassette.



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  		Figure 1. Alternative splicing of HipK3 in human testis. (A) Tissue distribution of the ‘T’ and ‘U’ exons. Alternative splicing of the T and U exons of HipK3 was studied by RT–PCR analysis of a panel of human tissue RNAs. Upper bands in each panel denote incorporation of the alternative exons. The T exon is specific to testis. Relative detection of the presence versus absence of the U exon is indicated below each lane as a molar ratio from the final products. (B) Tra2ß recapitulates the testis-specific alternative splicing of the T exon. HEK293 cells were transfected with GFP fusions to three splicing factors indicated and subjected to RT–PCR at the T exon as in (A). Brain and testis PCRs were also assayed for reference. (C) Structure of the HipK3 gene. Exons are shown as boxes, coding regions are shaded and introns are shown as line (localized scale bars are shown). The ATG starts the ORF at the third base of the long first exon shown; the upstream region remains to be definitively sequenced. The positions of the testis-specific (‘T’) and ubiquitous alternative (‘U’) exons are shown and their sequences are shown in boxes. Flanking intronic sequences are in lower case and in the T exon, an unused downstream consensus 5' splice site is indicated by a slash, the stop codon is underlined and the R-rich 30mer used in Figures 3 and 4 is indicated in bold.

 
A novel application of fluorescence-activated cell sorting demonstrates a Tra2ß-concentration dependence of the HipK3-‘T’ exon
Tissue-specific alternative splicing results from a tissue-specific complement of RNA-binding proteins (3Go), and several proteins have been strongly implicated in alternative splicing in testis (8Go,21Go,25Go–27Go). Therefore, we sought to establish whether there was any causal relationship between the germ cell expression of these factors and the alternative splicing. The putative splicing factors, T-STAR, RBM and Tra2ß, were cloned in frame downstream of enhanced green fluorescent protein and transfected into HEK293 cells, and splicing of endogenous HipK3 was analysed by RT–PCR. Untransfected cells showed no incorporation of the extra exon, nor did cells transfected with GFP-RBM or GFP-T-STAR. However, cells transfected with GFP-Tra2ß consistently showed a faint band (but of varying intensity) corresponding to incorporation of the testis-specific exon (Fig. 1B shows a good example).

To consolidate these results, we took advantage of FACS technology that allows sorting of cells into multiple fractions, according to precise boundaries of fluorescence intensity. Twenty-four hours after transfection, cells were sorted into eight fractions expressing discrete levels of GFP fusion protein successively differing by a constant ratio of ~2.37 and varying over three orders of magnitude (Fig. 2A). RNA was isolated from these eight fractions and analysed by RT–PCR for relative expression of the alternative isoforms of endogenous HipK3 (Fig. 2B). Consistent with the results from unsorted cells, RT–PCR analysis of FACS-sorted cells expressing GFP-T-STAR and GFP-RBM failed to show induction of the HipK3-T exon (data not shown). However, midway through the GFP-Tra2ß series, there was a concerted incorporation of HipK3-T. In contrast, there was a considerable reduction in the relative level of the HipK3-U exon commencing similarly midway through the GFP-Tra2ß concentration series (Fig. 2B).



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  		Figure 2. Tra2ß causes T exon inclusion above a concentration threshold. (A) Strategy for MF-FACS purification of transfected cells. Human HEK293 cells were transfected with GFP-Tra2ß and a typical cell population is shown in the cartoon. Note that many cells are untransfected and the GFP expression level is highly variable between transfected cells. These cells were automatically sorted into eight homogenous populations on a FACS DiVa cell sorter and eight fractions with a graded homogenous level of GFP expression increasing from left to right are shown. To demonstrate that distinct intensities of GFP-Tra2ß expression had been separated, a small aliquot of each sort was analysed by flow cytometry and the counts versus intensity were plotted on a graph and signals were superimposed and each represented by a different colour line (corresponding to the fractions above). As the eight fractions sit within the boundaries of three orders of magnitude, the relative expression level between each fraction can be estimated at 10001/8=2.37. (B) ‘T’ incorporation is up-regulated and ‘U’ incorporation is down-regulated by active concentrations of Tra2ß. Agarose gel analysis of RT–PCR across the T and U alternative exons from the numbered sorted cell fractions that express increasing amounts of GFP-Tra2ß (A). Relative detection of the alternative exons were calculated as ratios and plotted on the graph. Scales for the U and T exons are on the left and right, respectively. ‘Un’ denotes untransfected cells.

 
Tra2ß binds specifically to the HipK3-T exon
The Tra2ß RNA-binding domain was originally shown to bind to purine-rich sequences, especially those containing GAAGAA (28Go,29Go), and the HipK3-T exon contains a long central 50 nucleotide stretch containing 44 purine residues. Recently, 6% of the theoretically possible 4096 hexamer RNA sequences were predicted to be exonic splicing enhancers by the virtue of being over-represented both in exons and also in exons with poor splice sites (30Go). Web-based searching with the 109 nucleotide extra exon for these possible splicing enhancers (http://genes.mit.edu/burgelab/rescue-ese/) predicted 26 possible splicing enhancing hexamers (compared with the expected frequency of 7). Furthermore, 17 of these hexamers were located within a 30-nucleotide part of the purine-rich region (GGGAGGAAGAAATAGAAGATGCAGAAGAGG) with a GAAGAA motif (Fig. 1C). We therefore tested the binding of Tra2ß to this purine-rich 30-nucleotide stretch of the HipK3-T exon in a pulldown assay. First, the R-rich 30mer and a random sequence were transcribed, immobilized on agarose beads and incubated in whole cell extracts from cells transfected with GFP-Tra2ß or GFP alone. Proteins that bound to the RNAs were then separated by SDS–PAGE, western blotted and probed with an anti-GFP antibody. GFP-Tra2ß bound strongly to the R-rich sequence, but not to the negative control sequence ‘GST-89’ (Fig. 3A).



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  		Figure 3. Analysis of proteins that bind to the HipK3-T R-rich 30mer. (A) Transcribed RNAs corresponding to the HipK3 R-rich 30mer (Fig. 1C), and the negative control (GST-89) were immobilized on beads and incubated in whole cell extracts either transfected with GFP alone (marked –) or with GFP-Tra2ß (marked +). A western blot of bound material is shown, probed with anti-GFP antibody. (B) Right panel: using a positive control RNA for Tra2ß binding (ASF-id), the same pulldown experiment was performed as in (A) except that RNAs were incubated in untransfected HeLa cell nuclear extracts, and bound endogenous proteins were probed with anti-Tra2ß antibody. The left panel shows the input nuclear with or without calf intestinal phosphatase treatment. (C) Pulldown samples were subjected to western blotting with splicing factor antibodies: upper panel, anti-phospho-SR protein antibody 10H3; asterisk represents the doublet ~40 kDa which probably corresponds to SRp40 (upper band) and phospho-Tra2ß (lower band); n.b. both bands were visible in the nuclear extract ~40 kDa upon longer exposure (data not shown). Lower panels show staining of the same samples with the antibodies indicated. (D) Silver-stained gel of proteins bound to the two RNA sequences. Three strong bands that differentially bound to the 30mer were excised and sequenced and these were found to be hnRNPs A1, A2 and H. (E) Competitive binding to the HipK3-T R-rich 30mer RNA. Highly labelled HipK3-T R-rich region RNA was incubated with whole cell extract with or without Tra2ß and recombinant hnRNP A1 or hnRNP H or with short RNA competitors specific for Tra2ß (containing GAA repeats) or hnRNP H (GGGAx4). The R-rich 30mer RNA was then cross-linked to any proteins bound to it with UV radiation, and partially digested to leave labelled RNA-binding proteins, which were then visualized by electrophoresis and autoradiography. Only three major bands are seen probably corresponding to GFP-Tra2ß, hnRNP H and hnRNP A1. N.b. His-tagged hnRNP H migrates slightly slower than endogenous protein and a presumed degradation product of recombinant hnRNP H is marked with an asterisk.

 
To confirm this interaction with endogenous protein, we immobilized the HipK3-T R-rich 30mer and control RNAs as before and we included another RNA as a positive control, the known Tra2ß-binding sequence ‘ASF-id’, which contains GAA repeat sequences (29Go,31Go). RNAs were then incubated in untransfected HeLa nuclear extract, and this time proteins bound to the RNA were probed with anti-Tra2ß antibody (Fig. 3B, right side). The input nuclear extract contained two protein species of ~35–40 kDa recognized by the anti-Tra2ß antibody and these were identified as the phosphorylated and unphosphorylated form of Tra2ß by treatment of the nuclear extracts with calf intestinal phosphatase, as this removed the upper phosphorylated form (Fig. 3B, left side). In the pulldown assay (Fig. 3B, right side), the endogenous Tra2ß was found bound to the HipK3-T R-rich 30mer and ASF-id, but not to the negative control sequence. It is worth noting that only phosphorylated Tra2ß is bound to the positive control RNA, whereas both forms are bound to the HipK3-T R-rich 30mer.

We next tested whether other splicing factors bound to the HipK3-T R-rich 30mer RNA using several antibodies to probe the bound material. The phospho-RS-domain antibody 10H3 recognized a doublet of proteins ~40 kDa and two 30 kDa SR protein species in the pulldown (Fig. 3C, top panel). As we have observed that the 10H3 antibody recognizes both SRp40 and Tra2ß (data not shown), it was difficult to know exactly which of these proteins bound to the HipK3 sequence. However, a careful examination of several independent experiments, using SRp40- or Tra2ß-enriched fractions, suggested that the doublet probably corresponds to both SRp40 (upper band) and phospho-Tra2ß (lower band). It is striking to see that Tra2ß was not significantly detected in the input nuclear extract with Mab 10H3, suggesting that it is highly enriched when it is bound to the purine-rich 30mer RNA. A specific antibody to the 30 kDa SR protein ASF/SF2 showed it also bound specifically to the HipK3-T R-rich 30mer (Fig. 3C), and western analysis also detected the specific binding of hnRNP A1 and hnRNP H. In contrast, the SR proteins 9G8 and SC35 (data not shown) did not bind preferentially to the R-rich 30mer.

A silver-stained gel was used as a loading control for the western blot analysis (Fig. 3D). Although the global pattern of proteins (mostly consisting of proteins sticking to the agarose beads) was similar in the two lanes, three abundant proteins differentially bound to the two RNA sequences. Mass spectrometry and database searching revealed that they were hnRNP A1, its close homologue hnRNP A2 and hnRNP H (Fig. 3D), confirming the western blotting results earlier. This is consistent with the presence of potential binding sites for these proteins within the HipK3-T R-rich 30mer sequence (32Go–34Go). Mass spectrometry did not detect Tra2ß or any of the SR proteins identified earlier by western blotting, probably because they are present at low concentrations in nuclear extracts and also because of the competition with more abundant hnRNP proteins (discussed subsequently).

HnRNP A1 and hnRNP H have been shown to bind competitively with SR proteins including ASF/SF2 (35Go–38Go). To verify the significance of the binding of hnRNP A1 and hnRNP H to the HipK3-T R-rich region and to determine whether they can interfere with the binding of Tra2ß, we performed ultraviolet (UV) cross-linking assays using the two whole cell extracts from GFP- or GFP-Tra2ß-transfected cells described earlier. No binding of any protein was detected to the GST80 control sequence (data not shown), but the R-rich sequence strongly bound two main proteins of 50–52 and 35–36 kDa in GFP-transfected extract (Fig. 3E, lane 11) consistent with the sizes of hnRNP H and hnRNP A1, respectively, and also with the assumption (mentioned earlier) that endogenous Tra2ß is insufficiently abundant in cell line extracts to be detected in this assay. In GFP-Tra2ß-transfected extracts, GFP-Tra2ß bound strongly to the R-rich region (Fig. 3E, lane 6), and this binding appeared to largely displace binding of hnRNP A1 and also to slightly reduce hnRNP H binding (Fig. 3E, compare lanes 11 with 6). Competitive binding of Tra2ß and hnRNPs to the R-rich region was then confirmed by the titration of purified recombinant hnRNP A1 and hnRNP H. As expected, recombinant hnRNP A1 bound to the R-rich region strongly in GFP-transfected extract (Fig. 3E, lanes 12 and 13), but less strongly in Tra2ß-transfected extract (Fig. 3E, lanes 7 and 8) and binding of GFP-Tra2ß was only slightly reduced with recombinant hnRNP A1 (Fig. 3E, lanes 6–8), implying that Tra2ß binds more strongly than hnRNP A1 to the R-rich 30mer. Binding of recombinant hnRNP H was largely independent of the presence of Tra2ß (Fig. 3E, lanes 9–10 and 14–15) and indeed it displaced some Tra2ß (Fig. 3E, lanes 6 and 9–10). To further confirm these results, we removed specific proteins from the complex with short RNA aptamers for hnRNP H or Tra2ß. The hnRNP H aptamer (containing four GGGAs) removed hnRNP H and increased the binding of GFP-Tra2ß (Fig. 3E, compare lanes 4, 5 and 1), whereas the Tra2ß aptamer (containing GAA repeats) removed Tra2ß and increased the binding of hnRNP A1 to the purine-rich 30mer (Fig. 3E, compare lanes 2, 3 and 1). We conclude that Tra2ß binds competitively with hnRNP H and hnRNP A1 to the HipK3-T purine-rich region (see Discussion).

Tra2ß and hnRNP proteins have opposite effects on splicing through the HipK3-T purine-rich sequence
To test the effects of Tra2ß and hnRNP A1 on the activity of the HipK3-T purine-rich 30mer, we constructed a minigene containing the HipK3-T exon and surrounding regions in between two constitutive ß-globin exons and transfected this into HEK293 cells (Fig. 4A and B). In the absence of exogenous RNA-binding proteins, the T exon was not incorporated; however, GFP-Tra2ß caused a significant inclusion of the exon. This incorporation appeared more efficient than in the endogenous gene, possibly because it is more easily recognized in an intron of just 1400 bases rather than 9 kb. Alternatively, it may be that higher levels of the minigene transcript, compared with the endogenous one, required less rounds of PCR which will select against the longer product, and that actually our assay underestimates the abundance of the HipK3-T exon. We then mutated every third base in the R-rich sequence (9Gs and an A) to cytosine. Consistent with the removal of repressor and enhancer sequences, the exon was weakly present without transfected hnRNP A1 (GFP only) and no longer greatly enhanced by Tra2ß. With hnRNP A1, this low level of expression was inhibited, suggesting that the binding of hnRNP A1 to the mutated region or flanking regions is partially preserved. We conclude that the HipK3-T exon is subject to both positive and negative regulations and the major determinant of HipK3-T exon up-regulation is the purine-rich 30mer.



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  		Figure 4. Mechanistic studies of splice activation at the HipK3T R-rich 30mer. (A) The genomic region encompassing the HipK3-T exon (shown in thick line) was cloned into a minigene containing exon 2, intron 2 and exon 3 of rabbit ß-globin. The sequence of the wild-type and a mutant construct are shown. (B) The two constructs were then co-transfected into HEK293 cells along with GFP, GFP-Traß or hnRNP A1. The cells were harvested and RNA was extracted for RT–PCR analysis using the primer pair whose positions are indicated in (A). ±T indicated T exon inclusion and exclusion, respectively. (C) Cartoon of the minigene constructs used which contains two identical 5' splice sites (39Go) with the R-rich 30mer inserted between the two splice sites (D and E) and also with the E1A BSE inserted (E). (D) In vitro splicing assays of radiolabelled RNA from the R-rich 30mer-containing minigene (C) in splicing limiting conditions with low levels of SR proteins (see Materials and Methods). Whole cell extract from cells transfected with GFP alone (left-hand side) or GFP-Tra2ß (right-hand side) was added. Various combinations of recombinant SR proteins indicated were also added and spliced products were analysed by urea–PAGE on a 6% polyacrylamide gel. Quantification of relative downstream/upstream 5' splice site utilization is calculated as a ratio. (E) Splicing assays in nuclear extract (non-limiting conditions). Pre-mRNA with wild-type, HipK3-T R-rich 30mer or E1A BSE linker sequences were used in this experiment. Splicing was competed by the addition of the amounts (in µg) of recombinant hnRNP indicated. Percent splicing efficiency is indicated as well as the ratio of downstream to upstream splice site utilization.

 
The upstream 3' splice site of the HipK3-T exon is a perfect match to the consensus (Fig. 1B), so presumably it is suboptimal use of the downstream 5' splice site (a weak match to the consensus) that is enhanced by Tra2ß. To investigate the mechanism of conflicting action of Tra2ß, SR proteins and hnRNPs on downstream 5' splice site selection, we placed the R-rich 30mer sequence into a minigene between two identical 5' splice sites (39Go). This minigene was transcribed into RNA, which was then incubated in splicing-limiting conditions in vitro (see Materials and Methods) with added whole cell extract that had either been transfected with GFP alone or with GFP-Tra2ß (Fig. 4D, left and right sides, respectively) and also with combinations of recombinant SR proteins, ASF/SF2, SRp40 and 9G8. In a corresponding experiment with the control parent construct (without the insertion of the R-rich 30mer), the overall splicing efficiency remained fairly weak, even with SR proteins, and the ratio between downstream and upstream 5' splice was not significantly changed by the various proteins added (data not shown). With the R-rich 30mer inserted between the splice sites, utilization of the downstream 5' splice site usage varied considerably (as shown by the ratio of the downstream site to the upstream site, which varies from 0.3 to 5.1), depending on the different exogenous proteins. If no SR protein was added, overall splicing was very inefficient and splicing occurred mainly from the upstream site (ds/us=0.6). Although 9G8 shifted splicing moderately to the upstream 5' splice site (the ds/us ratio changed from 0.6 to 0.3), the SR proteins, ASF/SF2 and SRp40, alone had no significant effect but they synergized and shifted splicing to the downstream site when added together (ds/us=1.2). This additive effect was further increased with Tra2ß, to reach a ds/us ratio of 5.1. Tra2ß alone, in contrast, had a very limited effect (ds/us=0.8). Importantly, in all cases where exogenous Tra2ß was added together with at least one SR protein (SF2/ASF or SRp40, but not 9G8), a considerable enhancement of the downstream 5' splice site occurred. Therefore, we conclude that Tra2ß plays a major role in 5' splice site activation downstream of the HipK3-T R-rich region, but efficient activation requires the presence of at least one SR protein.

Testing the same two trancripts in non-limiting conditions in full nuclear extract (Fig. 5E) showed that the transcript containing an insertion of the R-rich sequence between the two 5' splice sites not only stimulated the global splicing efficiency (from 23 to 45%), but most importantly specifically enhanced the utilization of the downstream site. In that sense, the HipK3-T R-rich element behaves like the previously described E1A bidirectional splicing enhancer (BSE), which is also shown in Figure 4E. However, the R-rich region is not as potent as the BSE in downstream 5' splice site activation, possibly due to the binding of competitive repressor proteins. We therefore looked at the effects of hnRNP A1 and hnRNP H to see whether they would inhibit the use of the downstream site. Addition of hnRNP A1 had two significant effects. First, we observed a general decrease of splicing efficiency, independent of the nature of the linker sequence between the two competing 5' splice sites. This could result from the non-specific binding of hnRNP A1 to the different splicing substrates. However, with the HipK3 R-rich pre-mRNA, only the downstream site was inhibited by hnRNP A1, whereas splicing at the upstream site remained unaffected. This dramatically changed the ratio of downstream to upstream site usage from 2.1 to 1.0. In contrast, for the control construct, the ratio did not change. However, hnRNP A1 could not shift the splice site choice when the linker was the BSE, indicating that the inhibitory mechanism is likely to involve the binding of hnRNP A1 to the HipK3-T R-rich sequence. Finally, hnRNP H showed similar effects to hnRNP A1, but was far weaker in its inhibition such that the addition of both hnRNP A1 and hnRNP H did not show any difference from the effect of hnRNP A1 alone. We conclude that Tra2ß, with a subset of SR proteins, can stimulate splicing through the R-rich sequence from the HipK3-T exon and that hnRNP A1 has the opposite effect through its binding to the same sequence. Therefore, splicing of the HipK3-T exon is likely to be regulated by a balance of these factors.



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  		Figure 5. Function of Tra2ß-dependent alternative splicing in testis. (A) Tra2ß protein is highly expressed in testis. Western blot with anti-Tra2ß N-terminus is shown above a ß-actin control for six mouse tissues. The ratio of combined Tra2ß band intensities to ß-actin is shown as a bar chart. (B) Inhibiting NMD moderately stabilizes the T exon. Two cell lines were treated with Upf1 siRNA or a control siRNA or water only before transfection with GFP-Tra2ß and single-fraction FACS purification. Samples were subjected to nested RT–PCR across the T exon. Lower panel: bar chart showing relative detection of the T exon in six fractions. (C) HipK3-T protein is mis-localized in the nucleus. GFP fusions to HipK3 and HipK3-T were expressed in HeLa cells. Both proteins were nuclear as they co-localized with DAPI staining but the truncated alternatively spliced protein is localized diffusely unlike the full-length protein which has a punctate distribution. Scale bars, 10 µm.

 
Tra2ß is up-regulated in testis
Tra2ß mRNA is ubiquitous, but most highly expressed in testis and in developing brain (21Go,40Go–42Go). The expression levels of Tra2ß in six mouse tissues were determined by western blotting (Fig. 5A). This showed that Tra2ß protein is ubiquitously expressed but, by a factor of at least 2, the highest relative level of total Tra2ß protein compared with ß-actin was seen in testis. The lower band, possibly corresponding to unphosphorylated Tra2ß, was also relatively overexpressed in testis.

HipK3-T could be a substrate for nonsense-mediated decay
Incorporation of the T exon was consistently lower than that of the U exon and this could be explained by the fact that the T exon contains a stop codon, which makes it a potential target for down-regulation by nonsense-mediated decay (NMD) (43Go). To test whether transcripts containing the T exon can be subjected to NMD, we chose to partially inhibit this pathway by siRNA down-regulation of the essential NMD component Upf1 (44Go). If the T exon is down-regulated by this pathway, we should observe increased steady-state levels of HipK3-T. A moderate up-regulation of endogenous HipK3-T by Upf1 compared with control siRNA or water alone was indeed observed in both HeLa cells and HEK293 cells transfected with GFP-Tra2ß (Fig. 5B).

Protein encoded by HipK3-T fails to localize properly
Although HipK3-T can be degraded by NMD, stabilization was minimal when compared with that of a known substrate for NMD, U2AF35 (44Go), and clearly some HipK3-T is not degraded (Fig. 1A), so we sought to investigate the function of the truncated protein it potentially encodes. HipK3 and HipK3-T were cloned into pGFP3 and expressed in HEK293 cells (Fig. 5C). The full-length protein localized in a nuclear punctate or speckled pattern as observed previously (45Go). The truncated protein was also nuclear, but it failed to localize properly and was found diffused throughout the nucleus with holes in the staining representing nucleoli.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Tra2ß is a mammalian homologue of the prototypical alternative splicing factor Tra2 that is responsible for sexual differentiation and spermatogenesis in flies (3Go). It is likely to be an important developmental regulator of alternative splicing that acts by binding to purine-rich elements in or near exons (29Go), and evidence is accumulating from many different tissues that mutation of these purine-rich elements or disruption of Tra2ß expression patterns can lead to disease (46Go) (Table 1).


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  		Table 1. RNA targets of human Tra2 homologues and their overexpression in disease
 
We show here that Tra2ß binds to and enhances the incorporation of a novel testis-specific alternative exon of the homeodomain-interacting protein kinase HipK3 (HipK3-T) and that Tra2ß promotes exclusion of another exon in the HipK3 gene, the ubiquitous alternative (U) exon. A similar situation was recently reported, whereby Tra2ß promotes inclusion of exon 10 and exclusion of exon 2 of the important neural protein Tau (47Go). The mechanism of enhancement of exon 10 is thought to involve direct binding of Tra2ß, whereas inhibition of exon 2 involves interaction and interference with the directly bound enhancer proteins, SRp30c and SRp55. Consistent with a direct and predominant role in T exon inclusion, Tra2ß is overexpressed in mouse testis relative to other tissues, whereas other RNA-binding proteins bound to the HipK3-U may be primarily responsible for its alternative splicing, as Tra2ß protein levels do not strictly mirror the level of ‘U’ exon exclusion (Figs 1A and 5A).

The T exon contains a putative 30 nucleotide purine-rich splicing enhancer (Fig. 1C), and we showed that this is specifically bound by Tra2ß and the SR proteins, SRp40 and ASF/SF2, but not by 9G8 (Fig. 3). In agreement with a role of Tra2ß as a splicing enhancer acting at this R-rich 30mer, we found that, with at least one SR protein, Tra2ß caused a strong enhancement of a downstream 5' splice site in vitro (Fig. 4D). This is consistent with the situation in exon 10 of tau where mutations that increase Tra2ß binding cause exon incorporation by stabilizing U1 snRNP binding to the downstream 5' splice site of the exon (48Go). As the best activation was observed when ASF/SF2 and SRp40 were added together with Tra2ß, these results imply that Tra2ß cooperates with the very subset of SR proteins that are also bound to the R-rich 30mer (Fig. 3C). 9G8 failed to enhance the use of the downstream 5' splice site consistent with it not binding specifically to the exon (Fig. 3C).

Phosphorylation of RS domains can be another important mechanism of controlling alternative splicing (49Go,50Go) and this is another level at which the control of HipK3 splicing by Tra2ß could be modified. Phosphorylation of Tra2ß increases its interaction with the putative spermatogenic splicing factor RBM and also affects its localization and function (21Go,51Go,52Go). Although both forms of Tra2ß bound to the R-rich 30mer, the dephosphorylated form was relatively enriched, suggesting that dephosphorylation may be a mechanism of stimulating incorporation of the HipK3-T exon. This is consistent with previous data showing that phosphorylation of Tra2ß inhibits alternative splicing and binding of its own message (53Go). Consistent with the high affinity of hypo-phosphorylated Tra2ß for the HipK3-T exon relative to an artificial substrate (Fig. 3B), Tra2ß also appears to be relatively hypo-phosphorylated in testis (Fig. 5A).

Although Tra2ß up-regulation in testis appears to be solely responsible for HipK3-T exon inclusion, it is not clear why that exon is not selected in somatic cells as it has a perfect consensus 3' splice site and although it has a suboptimal 5' splice site, there is another perfect 5' splice site just 30 nucleotides downstream which is not used (Fig. 1C). The abundant heterogeneous nuclear ribonucleoproteins (hnRNPs) that coat nascent transcripts often inhibit splicing, and the effect of Tra2ß on the splicing of tau and SMN is antagonized by hnRNP A1 and hnRNP G (21Go,33Go,54Go–57Go). Therefore, we sought to establish what proteins were bound to the putative HipK3-T splicing element. Silver staining and mass spectrometry of proteins bound to the HipK3-T R-rich 30mer revealed abundant proteins specifically associated with this RNA element, especially hnRNP H, hnRNP A1 and its close homologue hnRNP A2. HnRNP H binds GGGG and GGGA motifs, which are found in the R-rich 30mer as well as upstream in the T exon (Fig. 1B) (34Go,58Go,59Go). HnRNP H can either enhance or antagonize the effects of hnRNP A1 on splicing that are usually negative (35Go,60Go,61Go). Indeed hnRNP A1 down-regulation alone is thought to be responsible for protein 4.1R exon inclusion during red blood cell development (62Go). In fact, hnRNP A1 is expressed at a very low level in rat testis which has the lowest hnRNP A1 to ASF/SF2 ratio of any tissue (63Go). It has also been shown that hnRNP A1 is completely switched off at the onset of meiosis during spermatogenesis in mice (64Go); however, hnRNP A2 is expressed in meiotic germ cells (65Go), suggesting that it could be the combined up-regulation of Tra2ß and down-regulation of hnRNP A1 that causes the testis-specificity of HipK3-T. It has been shown that hnRNP A1 can inhibit Tra2ß by binding at a separate site nearby (33Go); however, we found that Tra2ß and hnRNP A1 compete for binding to the purine-rich 30mer sequence (Fig. 3E) and act there to have opposite effects on splice site selection (Fig. 4 and see model Fig. 6).



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  		Figure 6. Model of HipK3 regulation. Different cell types contain a different complement of RNA-binding proteins. In somatic cells, hnRNP A1 inhibits the downstream 5' splice sites of the HipK3-T exon. Tra2ß concentration is low and it is mostly phosphorylated. In male germ cells, hnRNP A1 is down-regulated and Tra2ß is up-regulated and relatively hypo-phosphorylated allowing Tra2ß occupancy of the HipK3-T exon (shown as box) and exon definition by stimulation of the proximal downstream 5' splice site.

 
A full understanding of the function of alternative splicing of HipK3 in testis awaits further investigation; however, HipK3-T contains a premature stop codon that can be subjected to degradation by NMD (Fig. 5B), so that incorporation of the T exon could be used in a stage-specific manner to down-regulate the HipK3 gene. However, stabilization of the HipK3-T without Upf1 was considerably less pronounced than that of another gene U2AF35 (44Go) and we have found by template mixing that the abundance of HipK3 RNA is not reduced by overexpression of Tra2ß (data not shown). It therefore remains unproven whether the NMD pathway is the key molecular change in HipK3 regulation or indeed whether NMD functions effectively in testis at all, as several testis-specific transcripts with pre-mature stop codons are detectable at significant levels, including those of AKAP1 and Cyclin L2 (7Go,8Go). If HipK3-T mRNA is stable enough to encode a protein in testis, and its presence there suggest it is, then this protein is likely to have dramatically altered and possibly antagonistic properties, as it lacks the corresponding region of HipK1 and HipK2 that bind androgen receptor, homeodomains, Fas and p53 (66Go,67Go). The C-terminus of HipK2 contains a 95 amino acid ‘speckle retention sequence’ (68Go) and consistent with this, we have found that HipK3-T protein lacks the ability to localize into its normal punctate pattern in the nucleus (45Go,69Go) (Fig. 5C).

HipK3 was originally isolated as an overexpressed gene in multi-drug-resistant cells (70Go) and it phosphorylates FADD (71Go), which reduces its interaction with Caspase 8 and thereby protects prostate carcinoma cells from Fas-mediated apoptosis (72Go). Fas activation is also known to cause SR protein dephosphorylation (73Go), so there may be complex regulatory networks controlling apoptosis involving alternative splicing of HipK3 (74Go). A homologue of HipK3, the dual specificity kinase DYRK1 (75Go), strongly phosphorylates Tra2ß as well as a recently identified cyclin, L2, which was identified as a splicing factor with an RS domain which is specifically removed from the protein by alternative splicing in testis (7Go). Along with our results presented here, these findings suggest that, like HipK3, Tra2ß may be part of a complex regulatory network involving alternative splicing in testis.

During this work, we have developed a technique of multi-fraction (MF-FACS) purification of splice factor transformed cells. MF-FACS RT–PCR should be widely used for in the study of alternative splicing of endogenous transcripts as it allows a concentration gradient of any splicing factor to be studied in an in vivo experimental setting. As well as being a tool for studying the effect of tissue-specific splicing factors, it should be possible to extend this technology to study combinatorial models (2Go,3Go,76Go) by sorting cells transfected with two different coloured splicing factors to create two-dimensional alternative splicing matrices. This method has allowed us to demonstrate a Tra2ß-concentration dependence of HipK3 alternative splicing that may have important consequences for the control of spermatogenesis.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Tissue expression studies
Human tissue RNAs were obtained from Clontech, Ambion and Origene, and DNase treated and extracted with phenol/chloroform and then with chloroform, then precipitated with ammonium acetate and isopropanol, and resuspended in RNase-free water with one part per hundred ‘Superasin’ RNase inhibitor (Ambion). RNAs were reverse-transcribed using first-strand cDNA synthesis kit and random primers as instructed (Amersham). Bands were manually quantified densitometrically with automatic background correction and manual nearby rectangle background subtraction using GeneTools software (Syngene, Cambridge, UK). Ratios of products were calculated from band intensities assuming inverse proportionality of molarity with product length. Extraction and western blotting of mouse tissue samples using anti-Tra2ß N-terminal antibody (77Go) were as previously described (25Go). Protein bands were quantified using GeneTools as for PCRs mentioned earlier.

Cell culture
The cloning of Tra2ß into pGFP3 and also cell culture and transfection were as described previously (25Go). For the inhibition of NMD, cells were grown in media containing Upf1 siRNA as previously described (44Go) except GFP-Tra2ß was transfected into cells on day 5. On day 6, cells were FACS purified before harvesting into Tri-Reagent (Sigma) and subjecting to RT–PCR as in MF-FACS discussed subsequently. For subcellular imaging, cells were grown on slides and then transfected and fixed using 4% paraformaldehyde at 4°C for 1 h. Slides were then mounted with Vectashield mounting medium with DAPI (Vector Laboratories H-1200) and viewed using a Zeiss ‘Axioplan 2’ microscope. DAPI was visualized using Zeiss filter set 02 (excitation 365 nm and emission 420 nm) and GFP was detected with Zeiss filter set 13 (excitation 470/20 nm and emission 505–530 nm).

Minigene splicing experiment
The HipK3-T extra exon and approximately 270 nucleotides of intronic flanking region at each end were amplified from human DNA with the primers AAAAAAAAGAATTCGCATGGCGTCGTGGAAATTGAGCGAT and AAAAAAAAGAATTCCCTCTCCATGAGACTCTAAGTTCCAT. The PCR product was cleaved with EcoRI and cloned into the Mfe1 site in pXJ41 (39Go), which is exactly midway through the 757 nucleotide rabbit ß-globin intron 2. The R to C mutation was made by overlap PCR with the additional primers CAACTACAACATCCACAACAGCATGGACTAATTGATGGAGCAGAGTCT and GCTGTTGTGGATGTTGTAGTTGTTGCTGCCTCTGAGCTCTCTCCCCACCA. About 600 ng pGFP3 or 200 ng splicing factor plasmid and 100 ng minigene were transfected into HEK293 cells. After 24 h, RNA was harvested in Tri-Reagent (Sigma). An aliquot of 1 µl was used for ‘Superscript’ one-step RT–PCR (Invitrogen) with the program at 50°C for 30 min, 94°C for 2 min, then 35 cycles of 94°C for 30 s, 55°C for 30 s, 72°C for 45 s followed by 10 min at 72°C, with the pXJ forward and reverse primers GCTCCGGATCGATCCTGAGAACT and GCTGCAATAAACAAGTTCTGCT.

Multi-fraction fluorescence-activated cell sorting
Twenty-four hours after transfection with GFP-splicing factors, cells were sorted into eight discrete sample intensities in high-purity sort mode on an FACS DiVa Cell Sorter with a 488 nm Argon ion laser (Beckton Dickinson). The laser was run at 80 mW and a 100 µm nozzle was used. The machine was set to produce ~30 000 drops/s with cells being detected at ~9000 cells/s and with a stream pressure of 13 p.s.i. GFP fluorescence was detected using a 530/30 band-pass filter. First, all cells expressing detectable GFP were sorted and then these were sorted again within eight contiguous sets of limits covering three orders of magnitude. About 50 000 cells were collected in each fraction and lysed in 50 µl of ‘Cells to cDNA’ (Ambion) reagent and frozen as 7 µl aliquots. These were thawed and heated to 70°C with random decamers and cooled on ice before reverse transcription, as per protocol but omitting the heat inactivation step. An aliquot of 1 µl was used in a hot start PCR with 35 cycles of 94°C for 30 s, 59°C for 30 s, 72°C for 45 s followed by 10 min at 72°C, then run on a well-set 1.5–2% agarose gel. Alternatively, FACS fractions were suspended in 100 µl Tri-Reagent (Sigma) extracted as per protocol and resuspended in 10 µl water with one part per 100 ‘Superasin’ RNase inhibitor (Ambion). A total of 1 µl was used for ‘Superscript’ one-step RT–PCR (Invitrogen) with the program at 50°C for 30 min, 94°C for 2 min, then 35 cycles of 94°C for 30 s, 59°C for 30 s, 72°C for 45 s followed by 10 min at 72°C. Primers used for T exon analysis were GGGTCGGCCAGTCATGTATC and GCGCTACATCATCCAGACTGTTGA for long products (490/609 bp) or GCTCTACCCAGGAGCCTTGGAGT and CCTGGCAAACCTTGAGTCTGAGAA for short products (66/175 bp). Primers for the U exon were GCCTCAGCCTGCCACTACC and GGACTCGGGGAGTCGGCAA (268/333 bp).

RNA affinity assays
The HipK3-T R-rich 30mer was cloned by insertion of two hybridized oligos 5'-CGGGAGGAAGAAATAGAAGATGCAGAAGAGG-3' and 5'-GATCCCTCTTCTGCATCTTCTATTTCTTCCTCCCGGTAC-3' between the KpnI and the BamHI sites of the pBluescript-SK II(+) plasmid (Stratagene). The non-specific sequences (GST-80 and -89) were described previously (31Go). Plasmids were linearized with XbaI (HipK3-T R-rich) or XmaI (controls) and transcribed using T7 RNA polymerase. The protocol for immobilization of RNA on beads was derived from (78Go). First, 0.5 nmol of each RNA was incubated for 1 h in the dark in a 300 µl mixture containing 100 mM sodium acetate (pH 5.0) and 6.7 mM sodium m-periodate (Sigma). After precipitation by the addition of 200 mM sodium acetate (pH 5.0) and three volumes of ethanol, RNAs were centrifuged and resuspended in 0.1 M sodium acetate (pH 5.0). For each sample, an aliquot of 150 µl of adipic acid dihydrazide agarose beads (50% slurry) (Sigma) was washed four times with large volumes of 0.1 M sodium acetate (pH 5.0), mixed with the periodate-treated RNA and rotated overnight at 4°C. Beads were washed twice with 2 M NaCl and twice with buffer D [20 mM HEPES, pH 7.9, 20% glycerol, 100 mM KCl, 0.2 mM EDTA, 0.5 mM dithiothreitol (DTT)].

For the pulldown experiment, nuclear extract as prepared (39Go) (125 µl, 1 mg of proteins for each RNA sample) was pre-incubated for 15 min at 30°C in a 600 µl mixture containing (final concentrations): 10 mM HEPES, pH 7.9, 10% glycerol, 50 mM KCl, 2 mM MgCl2, 0.75 mM ATP, 25 mM creatine phosphate, 0.1 mM EDTA, 0.25 mM DTT, with 30 µg of Escherichia coli tRNA (Roche) and 18 µg of bovine serum albumin (BSA) (N.E.B.). This mixture was then added to ~300 pmol of immobilized RNA, and the binding reaction was carried out for 30 min at 30°C with constant mixing. After centrifugation at low speed (microfugation for 2 min at 3000 r.p.m. 800g), beads were washed four times with buffer D and eluted by addition of 40 µl of protein sample buffer and heated for 10 min at 95°C. Aliquots of 8 µl of the eluted bound proteins were resolved by SDS–PAGE (10% acrylamide) and analysed by western blotting with anti-Tra2ß (77Go), anti-hnRNP A1 antibody (4B10), anti-phospho-SR monoclonal (10H3), anti-9G8 monoclonal antibody, anti-hnRNP H polyclonal antibody and an affinity-purified polyclonal antibody directed against the N-terminal part of ASF/SF2. Dephosphorylation of 10 µl nuclear extracts was performed by incubation with 1 U/µl calf intestinal phosphatase (Promega) with 0.5 mM MgCl2 at 37°C for 30 min. The binding assay with transfected whole cell extracts was performed essentially as with HeLa nuclear extract; HEK293 EBNA cells were transfected with 2 µg of GFP-Tra2ß or control GFP alone plasmid, together with 10 µg of pBluescript-SK II(+) (Stratagene) and harvested 48 h later and anti-GFP antibody was used to probe the proteins bound to 80 pmol of RNA from 200 µl (between 2 and 2.5 mg of total proteins) whole cell extracts, prepared as previously described (79Go).

Mass spectrometric analysis of T exon R-rich 30mer-bound protein
Eluted complexes were separated on a 11% SDS–PAGE gel and silver stained (80Go). Digestion of excised bands and peptide extraction were performed as previously described (81Go). Peptide extracts were mixed with an equal volume of saturated 2,5-dihydroxybenzoic acid (Sigma), dissolved in 20% acetonitrile and applied to the target. MALDI mass measurements were carried out on a Bruker Reflex IV MALDI-TOF spectrometer. The acquisition mass range was 800–3000 kDa with low mass gate set at 700 kDa. Internal calibration was performed using autolytic trypsin peptides (MH+: m/z=842.51 and 2211.11). Monoisotopic peptide masses were assigned using the Bruker Flex Analysis software. The Profound program was used for database searching (http://prowl.rockefeller.edu/), with a mass tolerance of 75 p.p.m.

In vitro splicing assays
A BamHI fragment containing the HipK3-T R-rich sequence was obtained by hybridization of two complementary oligonucleotides 5'-GATCTCGAGGGTGAGGAGATCTGTGGGAGGAAGAAATAGAAGATGCAGAAGAG-3' and 5'-GATCCTCTTCTGCATCTTCTATTTCTTCCTCCCACAGATCTCCTCACCCTCGA-3' (the R-rich sequence is underlined) and used to replace the BamHI–BamHI fragment of the E1A-derived pD/D plasmid (39Go). Plasmids were linearized with HindIII and transcribed using SP6 RNA polymerase and [32P]CTP. In vitro splicing assays were performed in standard conditions as previously described, for 90 min at 31°C (31Go,82Go). In limiting conditions, a mixture of 7 µl of HeLa S100 cytoplasmic extract and 2 µl of HeLa nuclear extract, with 3 µl of GFP- or GFP-Tra2ß-transfected HEK293-EBNA whole cell extract, which showed similar intensity with anti-GFP (data not shown), was supplemented by 400–600 ng of baculovirus-purified ASF/SF2, 9G8 or SRp40. Quantifications of splicing assays were made using a Typhoon 8600 imager and the ImageQuant software (Amersham Pharmacia Biotech). Counts corresponding to the mRNA and the exon 1 intermediate for each alternative reaction were corrected for the background and also with respect to their respective number of labellable cytosine residues.

UV cross-linking
UV cross-linking assays were derived from Cavaloc et al. (31Go). An aliquot of 3 µl of whole cell extracts from cells transfected with GFP-Tra2ß or GFP alone were pre-incubated with 3 or 6 pmol RNA competitors for 5 min at room temperature or complemented with 250 or 500 ng of purified recombinant protein. RNAs were in vitro transcribed with T7 RNA polymerase. ASFid [Tra2ß-binding, containing (GAAGAAGAA)X2] has been described previously (31Go) and UGGGACUGGGACUGGGACUG (hnRNP H-binding site) was cloned into KpnI and BamHI sites of pBSKS (Stratagene). Extracts and competitors were then further incubated with 600 000 c.p.m. of short highly 32P-labelled RNA. The reactions were performed with 13 mM HEPES (pH 7.9), 65 mM KCl, 0.5 mM MgCl2, 0.38 mM ATP, 12.5 mM creatine phosphate, 0.77 mM DTT, 0.13 mM EDTA, 0.05% NP40, 15.7% glycerol, 0.25 µg BSA, 0.75 µg E. coli tRNA and 20 U RNasine (Promega), for 20 min at 31°C. The reactions were then transferred into a 96-well plate and exposed to UV light (254 nm) for 15 min, at a distance of ~1–2 cm, on ice. RNA was digested by incubation with 500 ng RNase A and 200 U RNase T1 for 45 min at 37°C. Samples were finally prepared by adding 5x concentrated Laemmli loading buffer and analysed by SDS–PAGE (11% acrylamide) and autoradiography of the dried gel.


   ACKNOWLEDGEMENTS
 
The authors thank Ian Harvey and Brian Shenton for assistance with FACS and Manuela Argentini for mass spectrometry; Christiane Branlant and Doug Black for recombinant purified hnRNP A1 and His-tagged hnRNP H, respectively; Stefan Stamm for anti-Tra2ß N-terminal antibody, Gideon Dreyfuss for hnRNP A1 antibody, Doug Black for hnRNP H antibody and M. Oulad for monoclonal anti-GFP antibody; Nick McGlincy and Chris Smith for siRNA reagents and advice. This work was supported by the Wellcome Trust (J.P.V., C.D. and D.J.E.) and by the Institut National de la Santé et de la Recherche Médicale, the Centre National de la Recherche Scientifique and the Association pour la Recherche contre le Cancer (C.F.B., L.K. and J.S).

Conflict of Interest statement. None declared.


   FOOTNOTES
 
{dagger} The authors wish it to be known that, in their opinion, the first two authors should be regarded as joint First Authors. Back


   REFERENCES
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 ABSTRACT
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 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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