Prp40 pre-mRNA processing factor 40 homolog B (PRPF40B) associates with SF1 and U2AF⁶⁵ and modulates alternative pre-mRNA splicing in vivo

PRPF40B is a protein highly enriched in nuclear speckles

In this study, we initially identified PRPF40B as a member of a subset of factors containing tandem repeats of WW and FF domains. Multiple alignments of these related structural proteins using bioinformatic tools revealed that the primary sequence of PRPF40B contains two WW domains in the amino-terminal half and five FF repeat motifs in the carboxyl-terminal half. Human and murine PRPF40B protein sequences show a high degree of homology with 95% identity and 96.5% similarity. The human PRPF40B protein sequence has 10% homology and 30% identity or 22% and 45% similarity with yeast Prp40, using ungapped or gapped local alignment, respectively. An analysis of the Prp40 sequence revealed the presence of four FF repeat domains, whereas PRPF40B is predicted to encode five of these domains (data not shown). To investigate the subcellular localization of PRPF40B, we performed immunofluorescence experiments with confocal laser microscopy on various cell types using a rabbit IgG-purified polyclonal PRPF40B antibody (see Materials and Methods). We consistently observed a diffuse nucleoplasmic pattern with an increased signal in organized granule-like sites in HEK293T and HeLa cells (Fig. 1A, left panels). The staining pattern of PRPF40B was reminiscent of that of nuclear speckles, which are nuclear compartments enriched in pre-mRNA splicing factors located at the interchromatin region of the nucleoplasm in mammalian cells (Lamond and Spector 1993). To determine whether the nuclear staining pattern of the endogenous PRPF40B protein coincides with that of nuclear speckles, we performed immunofluorescence analyses using an antibody directed against the phosphorylated form of the essential splicing factor SRSF2 (formerly SC35), which is commonly used to define nuclear speckles (Fig. 1A, middle panels). Interestingly, the PRPF40B nuclear dots were found to overlap with speckles in both cell lines (Fig. 1A, right panels). To confirm these data, we performed semiquantitative analysis of the spatial relationship between the relative distribution of PRPF40B and SRSF2 as a way to determine the degree of colocalization (Fig. 1, right). The polyclonal antiserum used here recognizes PRPF40B and several other proteins of slower mobility (we failed to raise peptide-specific antibodies). The mobility of one of these proteins coincides with the predicted mobility of PRPF40A. Given the high homology between PRPF40B and PRPF40A (using EMBOSS global and local alignments, the identity and similarity percentages ranged between 51%–54% and 65–68% for PRPF40A and PRPF40B, respectively) and the fact that both proteins are expressed in the cell lines used in this study (data not shown), the antibodies may be recognizing both proteins in the nuclei. To confirm the spatial localization of PRPF40B, we performed indirect immunofluorescence experiments with green fluorescence protein (GFP)-tagged PRPF40B. A similar pattern of nuclear staining was also observed when enhanced green fluorescence protein (EGFP)-tagged PRPF40B cDNA was transfected into HEK293T and HeLa cells (Fig. 1B). These results confirm the presence of a population of PRPF40B that colocalizes with nuclear speckles and suggest that PRPF40B might be implicated in splicing regulation.

To identify the sequence element responsible for the nuclear localization of PRPF40B, the primary amino acid sequence of PRPF40B was analyzed for the presence of a potential nuclear localization signal (NLS) using the WoLF PSORT (an update of PSORT II) (Horton et al. 2007), NucPred (Brameier et al. 2007), and cNLS Mapper (Kosugi et al. 2009) computer programs. This computational analysis identified a consensus NLS, KRRRR, with the highest score at amino acid residues 732–736 in PRPF40B. To test the functionality of this predicted NLS, we generated a truncated derivative of PRPF40B fused to EGFP that lacked this sequence (Fig. 2, EGFP-PRPF40B [1–687]). When analyzed by immunofluorescence, HEK293T cells expressing this mutant construct displayed predominantly cytoplasmic localization (Fig. 2), suggesting that the deleted region contains the signal for PRPF40B nuclear localization. To further characterize PRPF40B, we performed immunofluorescence analysis with a series of EGFP-tagged PRPF40B deletion mutants. The constructs are diagrammed in Figure 2. All mutants retain the putative NLS signal found in the carboxyl region. The PRPF40B mutant with an amino-terminal truncation removing the two WW domains produced a speckled staining pattern within the nucleus that was similar to that observed for the full-length protein (EGFP-PRPF40B [263–871] and EGFP-PRPF40B [1–871], respectively, in Fig. 2). This result strongly implies that the WW domains play no significant role in the nuclear speckle localization of PRPF40B. To determine the importance of the FF domains in targeting PRPF40B to nuclear speckles, we further generated carboxyl-terminal deletions. Removal of the most upstream two FF domain sequences created a fusion protein that significantly accumulated in nuclear speckles (EGFP-PRPF40B [394–871]) (Fig. 2). Deletion of the next FF domain partially disrupted the localization to nuclear speckles (EGFP-PRPF40B [474–871]) (Fig. 2), suggesting that this domain might be required for targeting the protein to nuclear speckles. Further deletions within the carboxyl-terminal region of PRPF40B generated fusion proteins that aggregated in the nucleus (EGFP-PRPF40B [618–871]) (Fig. 2 and data not shown). To further analyze the speckle-targeting capacity of the FF domains, we generated an EGFP chimera with the FF4 domain fused to the carboxyl-terminal region containing the NLS signal. Cells expressing this chimera showed significant nuclear speckle fluorescence (EGFP-PRPF40B [474–553/672–871]) (Fig. 2). The same results were obtained when these EGFP-tagged constructs were transfected in HeLa cells (data not shown). Although PRPF40B is distributed throughout the nucleoplasm with nucleolar exclusion, deletion of the FF domains affected intranuclear distribution with increased accumulation in the nucleoli (Fig. 2). We further examined the distribution of the EGFP-tagged PRPF40B deletion mutants using an antibody against the nucleolar phosphoprotein NOLC1 (Meier and Blobel 1990). We observed that the fusion protein with a deletion of the most upstream two FF domains sequences [EGFP-PRPF40B (394–871)], which did not disrupt the localization to nuclear speckles (Fig. 2), localized to the nucleoli (Fig. 3). Larger deletions [EGFP-PRPF40B (474–871), [EGFP-PRPF40B (618–871)] that affected the speckle distribution of PRPF40B (Fig. 2), resulted in an increased overlapping signal at the nucleoli (Fig. 3). Interestingly, the expression and detection of the EGFP chimera with the FF4 domain showed significant attenuation of nucleolar accumulation and produced a speckled pattern of staining within the nucleus (Fig. 3), which overlaps with the SRSF2 signal (Fig. 2). These results suggest that the putative NLS is essential for PRPF40B nuclear localization and nuclear speckle localization, that the FF domains participate in nucleolar exclusion, and that the FF4 domain is sufficient for localization to speckles. However, other FF domains may also be important for the efficient targeting of PRPF40B to the speckle compartment.

The association of PRPF40B with nuclear speckles is not affected by RNase treatment or transcription inhibition

Nuclear speckles are dynamic structures that are believed to act as storage, assembly, and modification compartments for splicing components, which can supply processing factors to nearby transcription sites. The structural architecture of nuclear speckles depends on interactions that are resistant to RNase treatment (e.g., SRSF2) and interactions that are RNase sensitive (e.g., Sm ribonucleoproteins). To evaluate the nature of the association of PRPF40B with nuclear speckles, we investigated whether RNA interaction was the molecular mechanism involved in concentrating PRPF40B in the nuclear speckle region. For these experiments, HEK293T cells were transiently transfected with EGFP-PRPF40B and treated with RNase A. Under these conditions, the staining pattern of PRPF40B and SRSF2 was unaffected, whereas Sm staining became completely diffuse throughout the nucleoplasm (Fig. 4A). Thus, like SRSF2, PRPF40B is not primarily localized to speckles through its binding to RNA. To determine the effect of transcriptional activity on the distribution of speckle-associated PRPF40B, we treated HEK293T cells with the RNA polymerase II inhibitor α-amanitin. Upon RNA polymerase II inhibition, speckles decrease in number, enlarge, and become rounded because of the accumulation of the splicing machinery (O'Keefe et al. 1994). Indeed, we observed a redistribution of PRPF40B into enlarged foci similar to that observed with SRSF2 upon transcriptional inhibition (Fig. 4B). We conclude that the localization of PRPF40B to nuclear speckles is not dependent on active transcription, and its accumulation in speckles shows a behavior similar to that of splicing factors.

PRPF40B associates with the splicing factors SF1 and U2AF⁶⁵

Prp40 directly interacts with Bbp, which is the ortholog of the splicing factor SF1, and associates with Mud2p, which is the ortholog of U2AF⁶⁵ (Abovich and Rosbash 1997). In addition, SF1 binds cooperatively with U2AF⁶⁵ to pre-mRNA introns at an early stage of spliceosome assembly (Berglund et al. 1998). Based on these data, we hypothesize that PRPF40B may be associated with SF1 and U2AF⁶⁵. To address this possibility, we first sought to investigate the spatial distribution of endogenous PRPF40B, SF1, and U2AF⁶⁵ by confocal microscopy. SF1 and U2AF⁶⁵ accumulate in speckles and show diffuse nucleoplasmic staining (Fig. 5). When we performed double-labeled immunofluorescence experiments using PRPF40B-specific antibodies and antibodies directed against the splicing factors SF1 (Fig. 5A) and U2AF⁶⁵ (Fig. 5B) in HEK293T or HeLa cells, we observed a high degree of colocalization of the proteins, thus demonstrating that these factors are in close proximity in the nucleus.

To test whether PRPF40B could directly interact with SF1, we performed in vitro GST-binding assays. To this end, full-length, amino-terminal (amino acid residues 1–333), and carboxyl-terminal (amino acid residues 446–871) PRPF40B were purified from bacteria as GST fusions (Fig. 6A). As a positive control, we used the amino regions of TCERG1 (Goldstrohm et al. 2001) (TCERG1 Nt in Fig. 6B) and U2AF⁶⁵ (data not shown) (see also Fig. 6C), which are known to bind SF1, fused to GST. The GST protein and carboxyl region of TCERG1 (TCERG1 Ct in Fig. 6B) were used as negative controls. The proteins were bound to glutathione–agarose beads and incubated with purified T7-tagged SF1 recombinant protein. After extensive washing, the bead-bound proteins were eluted by boiling the samples with SDS-PAGE loading buffer and analyzing by SDS-PAGE and Western blotting to detect SF1. The results of the in vitro binding assay revealed an interaction between PRPF40B and SF1. In the GST-binding assay, the amino portion of PRPF40B displayed strong binding, whereas the carboxyl-terminal region showed weak binding (PRPF40B [1–333] and PRPF40B [446–871], respectively, in Fig. 6B). These results suggest that PRPF40B contains SF1-binding domains mainly in the amino-terminal segment. Further deletion analysis is required to precisely locate the PRPF40B residues involved in the SF1 interaction. To measure the strength of these interactions, we determined the effect of increasing salt concentrations on the binding between PRPF40B and SF1 (Fig. 6C). Recombinant SF1 was incubated with GST, GST-PRPF40B, and GST-U2AF⁶⁵ (as positive control) and bound to glutathione beads in the presence of 150, 200, 300, and 500 mM NaCl. Beads with bound proteins were extensively washed with a buffer containing the same salt concentration as that in the binding reaction. The bound proteins were eluted by boiling with SDS-PAGE loading buffer and then analyzed by SDS-PAGE and Western blotting. The interaction between PRPF40B and SF1 proved to be stable under high salt concentrations up to 500 mM, although binding was gradually reduced with increasing salt concentrations (Fig. 6C).

To investigate whether these proteins interact in living cells, we transfected HEK293T cells with T7-tagged PRPF40B or an empty vector as a negative control. After immunoprecipitation with anti-T7 antibodies, the samples were analyzed by immunoblotting with specific antibodies directed against endogenous SF1 and U2AF⁶⁵. The results showed that endogenous SF1 and U2AF⁶⁵ were detected in the immunoprecipitates, indicating an association between PRPF40B and SF1 and U2AF⁶⁵ (Fig. 7A). To confirm these results, we transiently transfected a T7-tagged SF1 expression vector and analyzed the immunoprecipitates with antibodies directed against endogenous PRPF40B and U2AF⁶⁵. We observed the presence of endogenous PRPF40B and U2AF⁶⁵ associated with SF1, confirming the observation that an association between SF1 and PRPF40B exists (Fig. 7B). In contrast, ERK2, an unrelated mitogen-activated protein kinase, could not be detected in the eluates of the immunoprecipitation experiments (Fig. 7A,B). We conclude that PRPF40B associates with the SF1 and U2AF⁶⁵ splicing factors. RNase A treatment of the extracts impaired the interaction between PRPF40B and SF1 and U2AF⁶⁵ (Fig. 7C,D), which may indicate that this interaction is stabilized by concomitant binding to specific mRNAs. To test whether the interaction between PRPF40B and the SF1 and U2AF⁶⁵ splicing factors was mediated by U1 snRNP, we used an antisense morpholino oligonucleotide (AMO) complementary to U1 snRNA to knock down U1 snRNP in HEK293T cells (Kaida et al. 2010; Berg et al. 2012). We confirmed that the U1 AMO inhibited the activity of U1 snRNP by testing its effect on in vivo Fas alternative splicing (data not shown). We then repeated the immunoprecipitation experiments in the presence of control or U1 AMOs and detected endogenous SF1 and U2AF⁶⁵ in the immunoprecipitates (Fig. 7E), which suggest that U1 snRNP does not mediate this interaction. Taken together, these data indicate that PRPF40B associates with the splicing factors SF1 and U2AF⁶⁵, two factors involved in 3′ splice site recognition.

PRPF40B influences alternative splice site selection

Based on the results obtained above, we conducted in vivo splicing assays to examine the ability of PRPF40B to influence alternative splicing. Reporter constructs containing alternative exons flanked by constitutive introns were transiently cotransfected with T7-tagged PRPF40B or an empty vector in HEK293T cells. We used minigenes that recapitulate alternative 5′ splice site choice (Bcl-x and Fas) (see Fig. 8 for a schematic representation of the minigenes). Human Bcl-x pre-mRNA undergoes alternative splicing at 5′ splice sites in exon 2 to produce the anti-apoptotic long (Bcl-x_L) and proapoptotic short (Bcl-x_S) isoforms (Boise et al. 1993). The death receptor Fas/CD95, which encodes a transmembrane death receptor, can be alternatively processed to exclude exon 6 from its mRNA to produce a soluble isoform whose function is to inhibit apoptosis (Cheng et al. 1994; Cascino et al. 1995).

Total RNA was isolated from cells transfected with the minigenes under the conditions of PRPF40B overexpression or gene knockdown, and splicing was analyzed by RT-PCR using specific primers to detect the different transcript isoforms. Different post-transfection times were used between the overexpressed/knockdown samples to visualize the splicing effects under each condition, which affected the alternative splicing in the mock condition (see Fig. 8A,B, siEGFP versus Mock, and Materials and Methods). To note and because of the high homology between PRPF40B and PRPF40, we determined that the siRNAs against PRPF40B did not affect the PRPF40A level using RT-qPCR assays (data not shown). In the case of the Bcl-x minigene, increased levels of PRPF40B produced more efficient utilization of the Bcl-x_S 5′ splice site, resulting in a decreased ratio of Bcl-x_L to Bcl-x_S (Fig. 8A), while the amount of the Bcl-x_L isoform was increased upon PRPF40B knockdown (Fig. 8A). For the Fas minigene, PRPF40B enhanced the amount of the short anti-apoptotic Fas isoform (Fig. 8B), while the absence of PRPF40B resulted in a slight increase in the long proapoptotic Fas isoform (Fig. 8B, see below). We obtained the same results using a second siRNA against PRPF40B (data not shown; see Materials and Methods for siRNA sequences). Taken together, these results show that PRPF40B can modulate alternative splicing events.

To further study the function of PRPF40B in regulating Fas alternative splicing, we first analyzed the splicing pattern of the endogenous Fas gene by RT-PCR. Consistent with previous results, we observed that PRPF40B overexpression reduced the level of exon 6 inclusion, giving rise to increased amounts of the short anti-apoptotic isoform. The absence of PRPF40B promoted the inclusion of exon 6, provoking a robust increase in the long proapoptotic Fas isoform (Fig. 8C). We repeated the analysis by qPCR using primers that exclusively amplify the transmembrane receptor isoform and obtained the same results (Fig. 8D).

PRPF40B activity in Fas alternative splicing requires exon 6 sequences and associated 5′ and 3′ splice sites

As a first step to understand the regulation of Fas alternative splicing by PRPF40B, we sought to determine the exonic and intronic regulatory sequences important for PRPF40B function. Although it is assumed that yeast Prp40 bridges the 5′ and 3′ ends of the intron by interacting with Bbp and U1 snRNP without interacting directly with the pre-mRNA, a systematic study of the sequence elements that affect PRPF40B function would provide useful information regarding the mechanism by which PRPF40B modulates alternative splicing. For this goal, we took advantage of minigene constructs in which the exon 6 sequences were divided into three segments of control sequences of approximately the same length (e1, e2, and e3 in Fig. 9A), which were previously used to delineate the responsive elements of the putative tumor suppressor RBM5 within Fas exon 6 (Bonnal et al. 2008). Replacement of the e3 sequence did not compromise PRPF40B function (Fig. 9B, lanes 5,6). Replacement of the e1 and e2 sequences rendered reduced levels of exon skipping by PRPF40B (Fig. 9B, lanes 3,4 and 7–10), which reveals that Fas exon 6 sequences are important for PRPF40B function. Segment e2 corresponds to a uridine-rich exonic silencer (URE6), which is the binding region for the splicing repressor PTB (Izquierdo et al. 2005). Previous data have shown that the URE6 element contains an exonic splicing silencer that mediates repression by the splicing factor PTB and that the silencing effect relies on its pyrimidine-rich content (Izquierdo et al. 2005). To further evaluate the function of PRPF40B through the URE 6 element, we performed RT-PCR experiments with constructs containing three different mutations in URE6 (i.e., a cytidine and adenosine-rich sequence, another uridine-rich sequence that maintains silencer activity, and a scrambled sequence of balanced composition [m0, m1, and m2 in Fig. 9A, respectively]) (Izquierdo et al. 2005; Bonnal et al. 2008). We observed that only the m0 mutation significantly reduced the effects of PRPF40B overexpression (Fig. 9C, lanes 3–4), which supports the previous results with the mutation in the e2 segment and further demonstrate that uridine-rich sequences within the exonic splicing silencer of URE6 are important for PRPF40B function.

Next, the influence of splice site strength on PRPF40B function was analyzed. Strengthening either the 3′ splice site by inserting a strong polypyrimidine tract (mutant Py) or the 5′ splice site by improving its base-pairing with the U1 snRNA (mutant U1C) splice site associated with exon 6 compromised PRPF40B regulation of Fas alternative splicing (Fig. 9D), suggesting that a strong 5′ or 3′ splice site in exon 6 limits the response to PRPF40B. Taken together, these data indicate that in the case of the regulation of Fas alternative splicing, the presence of exonic sequences and weak 5′ and 3′ splice sites are required for PRPF40B function in alternative exon selection.

To evaluate whether the recruitment of PRPF40B to URE6 is sufficient to mediate the observed effects, we used an MS2 tethering assay. Previous data showed that the substitution of the URE6 sequence with binding sites for the MS2 bacteriophage protein resulted in efficient exon skipping upon MS2-PTB overexpression (Fig. 9E; Izquierdo et al. 2005). PRPF40B overexpression did not induce exon skipping in this minigene construct (Fig. 9E), which further supports the importance of exon 6 sequences in the PRPF40B-mediated regulation of Fas alternative splicing. However, this minigene was insensitive to the expression of an MS2-PRPF40B fusion protein (Fig. 9E). We conclude that PRPF40B recruitment to the URE6 element is insufficient to induce Fas exon 6 skipping.

PRPF40B depletion increases Fas/CD95 receptor number

The absence of PRPF40B promoted the inclusion of the Fas exon 6, causing an increase in the long proapoptotic Fas isoform (Fig. 7B–D), which encodes the membrane-bound form of the Fas receptor. To determine whether the observed change in the alternative splicing of Fas was related to the expression of the transmembrane receptor protein, we examined the proportion of Fas receptors in control and PRPF40B-depleted cells by flow cytometry using a specific anti-Fas/CD95 antibody. The flow cytometric data shown in Figure 10A are displayed as frequency histograms of Fas/CD95 immunofluorescence, detected using an Alexa 647-conjugated secondary antibody. The data are presented on a logarithmic scale and a shift to the right in the histogram indicates an increase in Fas/CD95-associated immunofluorescence. We observed a higher amount of membrane-bound Fas receptor upon PRPF40B-depletion (Fig. 10A), which correlates with previous data regarding the expression of the proapoptotic isoform in these cells. Statistical analyses using the nonparametric Kolmogorov–Smirnov test for comparing two samples revealed a small P value (≤0.001), which indicates that the distribution did not occur due to chance (Fig. 10B).

PRPF40B affects cell apoptosis

The previous result indicates that the absence of PRPF40B stimulates cell apoptosis. Several experiments were carried out to determine whether PRPF40B influences apoptosis. We chose to perform the experiments in the same HEK293T cells, although they have high resistance to cell death and therefore the effects obtained are small but significant. First, we measured the percentage of dead cells in control and PRPF40B-depleted cells using propidium iodide (PI). We found that PRPF40B knockdown increased the number of dead cells (Fig. 11A). However, this technique does not distinguish between apoptotic and necrotic cells. To better quantify the apoptotic cells, we performed cell cycle experiments to determine the number of cells in the sub-G1 phase, which corresponds to apoptotic cells. We observed a slight but significant increase in the percentage of cells in the sub-G1 population in PRPF40B-depleted cells compared with control cells (Fig. 11B). These results suggest that PRPF40B depletion increases the proportion of apoptotic cells. To lend additional support to these data, we also performed annexin-V binding assays. Annexin-V specifically binds phosphatidylserine when this molecule is located at the cell surface under apoptotic conditions. We first determined cell viability in control and PRPF40B-depleted cells and observed that the knockdown of PRPF40B resulted in a decrease in the number of viable cells (Fig. 11C). Next, we measured the number of annexin-V-positive cells by flow cytometry in the cells. In agreement with our previous data, we observed significantly augmented annexin-positive cells upon PRPF40B knockdown (Fig. 11C). We also analyzed caspase-3 activity, which is currently used as a marker of apoptosis. To measure active caspase-3 in the cells, we used an assay based on the addition of a luciferase substrate that is cleaved in the presence of active caspase-3, leading to a luminescent signal. The number of luciferase units directly correlates with the amount of active caspase-3 present in the cell. We observed a low (less than twofold) but significant increase in the quantity of active caspase-3 in PRPF40B knockdown cells (Fig. 11D). Caspase-3 is expressed as a 35 kDa inactive precursor from which the 17 kDa (p17) and 11 kDa (p11) subunits are proteolytically generated during apoptosis. The caspase-3 precursor is first cleaved to generate a p11 subunit and a p20 peptide, which is subsequently cleaved to produce the mature p17 subunit. The active caspase-3 enzyme is a heterodimer composed of two p17 peptides and two p11 subunits. Using specific antibodies against caspase-3, we observed significantly more p20 and p17 upon PRPF40B knockdown (Fig. 11E). The amount of the caspase-3 precursor was consequently diminished in PRPF40B-depleted cells (Fig. 11E). These findings confirm that the inhibition of PRPF40B causes an increase in caspase-3 activation. The expression of the anti-apoptotic regulator Bcl-2 was also analyzed. Interestingly, this protein showed reduced expression upon PRPF40B depletion (Fig. 11E). Together, these data show that PRPF40B modulates apoptosis and suggest a functional link between the control of alternative splicing and molecular events leading to cell death by apoptosis.

Plasmid constructs

A commercial intermediate vector encoding the human cDNA for PRPF40B (IMAGE clone 664467, Source Bioscience) was used to generate the PRPF40B constructs described below. The mammalian expression vector pEFBOST7-PRPF40B (1–871) and the corresponding deletion expression constructs pEFBOST7-PRPF40B (1–687), pEFBOST7-PRPF40B (263–871), pEFBOST7-PRPF40B (394–871), pEFBOST7-PRPF40B (474–871), and pEFBOST7-PRPF40B (618–871) were obtained by inserting the appropriate PCR fragments using XbaI/ClaI or XbaI/BstBI sites at the ends into the XbaI/BstBI sites of the parental vector pEFBOST7, which encodes the 11-amino acid T7 epitope tag at its amino terminus, using standard cloning procedures (Suñé and Garcia-Blanco 1999). pEFBOS/EGFP/TCERG1 (Sánchez-Álvarez et al. 2010) was used to generate pEFBOS/EGFP/T7PRPF40B (1–871), pEFBOS/EGFP/T7PRPF40B (1–687), and an amino-terminal deletion series by subcloning appropriate EcoRI fragments obtained from pEFBOST7 parental vectors. pEFBOS/EGFP/T7PRPF40B (474–553/672–871) was generated by digesting the expression vector pEFBOS/EGFP/T7PRPF40B (474–871) with XbaI/BamHI and inserting the PCR-amplified fragment PRPF40B (474–553) into the XbaI/BamHI restriction sites. The cDNA encoding human SF1 was amplified from the pGEM-SF1-Bo vector, which was kindly provided by Angela Kramer (University of Geneva, Geneva, Switzerland). PCR fragments containing XbaI and BstBI restriction sites were cloned into corresponding sites in the pEFBOST7 vector. The bacterial expression vectors pGEX2TK-PRPF40B (1–871), pGEX2TK-PRPF40B (1–333), and pGEX2TK-PRPF40B (446–871) were generated by inserting PCR-amplified products digested by BglII/EcoRI into a BamHI/EcoRI-digested pGEX2TK vector (Amersham Pharmacia Biotech). The bacterial expression vectors pGEX2TK-TCERG1 (1–662) and pGEX2TK-TCERG1 (631–1098) have been previously described (Suñé et al. 1997; Carty et al. 2000). The Bcl-x and Fas splicing reporter minigenes were kindly provided by Benoit Chabot (University of Sherbrooke, Québec, Canada) and Juan Valcárcel (Centre for Genomic Regulation, Barcelona, Spain), respectively. The Fas minigene derivatives (e1, e2, e3, m0, m1, m2, Py, and U1C) have been previously described (Izquierdo et al. 2005; Bonnal et al. 2008) and were kindly provided by Juan Valcárcel.

Antibodies

The polyclonal PRPF40B antibody used in this study was generated in rabbits using the GST-PRPF40B (1–871) construct. The GST-PRPF40B fusion protein was expressed as previously reported (Faber et al. 1998). The antiserum was directly used for immunoblotting analysis at a dilution of 1:10,000. For immunofluorescence analysis, IgGs were purified by protein A sepharose columns (GE Healthcare Life Science) under the conditions suggested by the manufacturer, and the purified IgGs were used at 1:1000. An antibody directed against the T7 tag (Bethyl Laboratories) was used at 1:20,000 for immunoblotting. Antibodies directed against ERK2 (C-14, Santa Cruz Biotechnology) were used at 1:2000 for immunoblotting. Antibodies directed against NOLC1 (Abcam) were used at 1:500 for immunofluorescence analysis. Antibodies directed against U2AF⁶⁵ were used at a dilution of 1:100 for immunofluorescence analysis and were kindly provided by Juan Valcárcel. For immunoblotting, we used a commercially available anti-U2AF⁶⁵ antibody (sc-48804, Santa Cruz Biotechnology) at a dilution of 1:2000. The mouse monoclonal antibody directed against the splicing factor SF1 (Abnova) was used at 1:100,000 and 1:40,000 for immunoblotting and immunofluorescence analysis, respectively. An antibody directed against the splicing factor SC35 (S4045, Sigma-Aldrich) was used at 1:4000. Antibodies directed against caspase-3 (sc-7148) and Bcl-2 (sc-7382) (Santa Cruz Biotechnology) were used at a dilution of 1:500 for immunoblotting. Antibodies against the Fas/CD-95 receptor (BioLegend) were used at 1:100 for flow cytometry and were kindly provided by Mari Carmen Ruiz (CIBM, Granada). For immunoblotting, primary antibodies were detected using HRP-conjugated secondary antibodies (PerkinElmer Life Science) at 1:5000. Alexa 488-conjugated goat anti-mouse (Molecular Probes), Alexa 647-conjugated goat anti-mouse (Molecular Probes), and Alexa 647-conjugated goat anti-rabbit (Life Technologies) secondary antibodies were used to detect primary antibodies at 1:500 and 1:5000 for immunofluorescence and flow cytometry, respectively.

Cell culture and transfections

HEK293T and HeLa cells were grown and maintained as previously described (Sánchez-Álvarez et al. 2006). For immunoblotting, reverse transcriptase-PCR (RT-PCR) analysis, and apoptosis experiments, transfections were performed in 35-mm plates (Falcon, Fisher Scientific). Each plate was seeded with ∼1 × 10⁶ cells 20 h prior to transfection. Cells were grown to ∼60%–70% confluence and transfected with appropriate amounts of the indicated constructs using calcium phosphate. Approximately 40 h after transfection, the cells were harvested and processed. For RNA interference (RNAi) knockdown experiments, ∼0.5 × 10⁶ HEK293T cells were seeded in 35 mm plates. The cells were transfected using the Lipofectamine 2000 reagent (Invitrogen) according to the manufacturer's protocol with 60 nM (final concentration) of either of the following small interfering RNA (siRNA) duplexes: siEGFP: 5′-CUACAACAGCCACAACGE-3′, siPRPF40B: 5′-GCAGUUCUGGACAGCAUCA-3′, and siPRPF40B(2): 5′-AAUCAGAGACCACCAGCUAUCUU-3′. The cells were harvested and processed 48 or 72 h after transfection for RT-PCR or apoptosis experiments, respectively. For double transfections, the cells were transfected with the reporter minigene 48 h after siRNA transfection using the LipoD 293T reagent (SignaGen Laboratories) according to the manufacturer's protocols. Cells were harvested at 24 h after transfection for the Fas experiments and at 48 h for the Bcl-x experiments. After total RNA extraction, the splice variants were amplified by RT-PCR. For immunoprecipitation analysis, transfections were performed in 100 mm plates (Falcon). Each plate was seeded with ∼1.5 × 10⁶ cells 20 h prior to transfection. The cells were grown to ∼80%–90% confluence and transfected with 6 mg of DNA using calcium phosphate. The cells were harvested and processed ∼40 h later. For U1 snRNA blocking, we transfected the AMOs at a final concentration of 3 μM at ∼24 h after DNA transfection using the EndoPorter reagent (Gene Tools), according to the manufacturer's protocol. The sequence of U1 AMO and control AMO (Gene Tools) are as follows: 5′-GGTATCTCCCCTGCCAGGTAAGTAT-3′ and 5′-CCTCTTACCTCAGTTACAATTTATA-3′, respectively. For immunofluorescence, cells were grown on coverslips to 50%–60% confluence, transfected 20 h later using calcium phosphate, and processed ∼24 h after transfection.

Immunofluorescence analysis and treatments

Immunofluorescence studies were performed as previously described (Sánchez-Hernández et al. 2012). To analyze the endogenous SF1 and U2AF⁶⁵ proteins, a specific protocol was performed: The cells were washed twice in phosphate-buffered saline (PBS) (pH 7.4) and permeabilized with 0.5% Triton X-100 in CSK buffer (100 mM NaCl, 300 mM sucrose, 10 mM PIPES, 3 mM MgCl₂, and 1 mM EGTA [pH 6.8]) for 1 min on ice. Subsequently, the cells were fixed with 3.5% paraformaldehyde in PBS for 5 min at room temperature and rinsed with PBS. The cells were incubated with blocking buffer (1% fetal bovine serum in PBS) for 10 min at room temperature and subsequently with primary antibodies in blocking buffer for 1 h at room temperature. After three 15-min washes in PBS, the cells were incubated with the appropriate secondary antibody in blocking buffer for 1 h at room temperature. After three washes with PBS, the coverslips were mounted in ProLong Gold Antifade Reagent (Molecular Probes) onto glass slides. Treatment with 25 mg/mL α-amanitin was performed for 6 h at 37°C prior to immunofluorescence processing. RNase was used at a concentration of 0.1 mg/mL for 2 h at 37°C after the fixation and permeabilization steps. Images were acquired using a Leica SP5 spectral laser confocal microscope and processed using the LAS AF software v2.3.6. The images were digitally processed for presentation using the Adobe Photoshop CS3 extended v10.0 software.

Preparation of cell extracts and immunoprecipitation

In vivo interaction experiments were performed with whole-cell extract (WCE). Cells were washed in cold PBS, pelleted, and lysed in 500 mL of T7 buffer (20 mM HEPES [pH 7.9], 150 mM NaCl, 5 mM EDTA, 1% NP-40, 1 mM dithiothreitol, 1 mM phenylmethylsulfonyl fluoride (PMSF), and protease inhibitor mixture [Complete Roche]) for 30 min at 4°C. The cell extracts were centrifuged at maximum speed for 5 min. For RNase A treatment, the cell lysates were incubated with 100 mg/mL of RNase A for 1 h at 4°C. Ten percent fractions of WCE were boiled in SDS-PAGE loading buffer and saved for immunoblotting analysis. The remaining cell lysates were diluted to a 1-mL final volume with T7 buffer. Diluted extracts were added to 50 µL of anti-T7 tag monoclonal antibody covalently coupled to cross-linked agarose beads (Novagen) and incubated with end-over-end rotation for 4–6 h at 4°C. After five washes with T7 buffer, the proteins bound to the antibody resin were eluted by boiling the samples with SDS-PAGE loading buffer, separated on 10% SDS-PAGE gels, and analyzed by Western blotting.

Protein purification and in vitro interaction assay

GST fusion proteins were expressed in BL21 (D3) cells and purified on glutathione–agarose beads (Sigma) following standard procedures. The elution of bound proteins was performed by affinity competition using saturating levels of reduced L-glutathione (Sigma) in a final volume of 1 mL. Proteins were dialyzed against PBS containing 1 mM PMSF and concentrated with an Ultra 0.5 Centrifugal Filter (Millipore). The SF1 protein was expressed in HEK293T cells and purified as previously described (Cazalla et al. 2005). In vitro binding of SF1 to GST fusion proteins was performed as previously described (Xiao and Manley 1997; Rain et al. 1998) with minor modifications. Briefly, 1 mg of GST or GST fusion proteins was bound to 50 mL of glutathione–agarose beads in 300 mL of NETN buffer (20 mM Tris–HCl at pH 8.0, 150 mM NaCl, 0.5% NP-40, and 0.5 mM EDTA) for 30 min with end-over-end rotation at 4°C. The beads were washed twice with 500 mL of NETN and incubated with 1 mg of SF1 in 200 mL of NETN for 30 min with end-over-end rotation at 4°C. After six washes with 200 µL NETN, the bound proteins were eluted by boiling in SDS-PAGE loading buffer for 5 min. Two independent fractions of each sample were separated by 10% SDS-PAGE and analyzed by silver staining and Western blotting.

Western blot analysis

A fraction of the transfected cells was lysed in cold T7 buffer. The proteins were separated by 10% or 12.5% SDS-PAGE, transferred to a nitrocellulose membrane (Amersham Biosciences), and then incubated with specific antibodies. After washing, the membrane was incubated with peroxidase-conjugated secondary antibodies, and the bound antibodies were detected by enhanced chemiluminescence (PerkinElmer Life Science). Densitometric analysis was performed using the QuantityOne software v4.6.5 (Bio-Rad). The intensity of the bands was calculated relative to the internal control (ERK-2) for each lane and each sample.

RNA extraction and RT-PCR analysis

Total RNA was extracted from transfected cells with peqGOLD TriFast (peQlab) according to the manufacturer's protocol. For plasmid-derived RNA, 800 ng of RNA was reverse transcribed by the Moloney murine leukemia virus RT (Invitrogen) for 1 h at 37°C using the primers csnPT2 (5′-AAGCTTGCATCGAATCAGTAG-3′) and RT-SVeda (5′-GGGAAGCTAGAGTAAGTAG-3′) for the Fas and Bcl-x reporter minigenes, respectively. One quarter of the resulting cDNA was amplified by PCR using the oligonucleotides csnPT1 (5′-GTCGACACTTGCTCAAC-3′) and csnPT2 for Fas and X34 (5′-AGGGAGGCAGGCGACGGCGACGAGTTT-3′) and XAgeIR (5′-GTGGATCCCCCGGGCTGCAGGAATTCGAT-3′) for Bcl-x. The intensity of the bands was quantified using the Quantity One software. For endogenous Fas RNA, 1 µg of RNA was reverse transcribed using the qScript cDNA Supermix (Quanta Biosciences) following the manufacturer's protocol. Fas endogenous transcripts were analyzed by conventional PCR, using the oligonucleotides FasE5Fw (5′-GTGAACATGGAATCATCAAGG-3′) and FE7 (5′-TCCTTTCTGTGCTTTCTGCAT-3′). The intensity of the bands was quantified using the Quantity One software. Quantification of Fas endogenous transcripts by real-time PCR was performed by using the iQ SYBR Green Supermix (Bio-Rad) and the iCycler thermal cycler station (Bio-Rad) with the oligonucleotides FE6 (5′-TAACTTGGGGTGGCTTTGTC-3′) and FE7. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as an internal control gene and was amplified with the oligonucleotides GAPDH-F (5′-ATGGGGAAGGTGAAGGTCG-3′) and GAPDH-R (5′-GGGTCATTGATGGCAACAATATC-3′).

The statistical analysis was performed using the Prism 5.0 software (GraphPad). A two-tailed Student's t-test was used to compare the means between samples and their respective controls. The P values are represented in the figures by asterisks (* P < 0.05; ** P < 0.01; *** P < 0.005). The absence of an asterisk indicates that the change was not statistically significant.

Cell death and cell cycle assays

HEK293T cells were trypsinized, collected in polystyrene tubes and washed twice with PBS. For the cell death assays, one half of the cells were incubated in PBS containing 40 μg/mL propidium iodide (Sigma) at 37°C for at least 20 min. For the cell cycle analysis, the remainder of the cells was fixed with 70% ethanol at 4°C for at least 15 min. After washing with PBS, the cells were incubated for 20 min at 37°C in a solution of PBS containing 100 μg/mL RNase A (Roche) and 40 μg/mL propidium iodide. The samples were analyzed by flow cytometry. The data were processed using the FlowJo v7.6.5 software.

Annexin-V binding assay

HEK293T cells were trypsinized, collected in polystyrene tubes and washed with PBS. The cells were incubated in 100 mL of a solution containing 5 mL of annexin-V conjugated to fluorescein isothiocyanate (FITC) (Immunostep S.L.) for 15 min at room temperature. The fluorescence was measured by flow cytometry. The data were analyzed using the FlowJo v7.6.5 software. The statistical analysis was performed using the Prism 5.0 software.

Active caspase-3 measurement

Approximately 1 × 10⁴ HEK293T cells were collected to measure caspase-3 activation. Quantifications were performed using the Caspase-Glo 3/7 assay (Promega) according to the manufacturer's protocol. The luminescent signal (relative light units, RLUs), which was directly proportional to caspase activation, was measured using an Infinite F200 TECAN Luminometer.

Analysis of the expression of membrane-bound Fas/CD95 receptor

HEK293T cells were trypsinized, collected in polystyrene tubes and washed with PBS. The cells were incubated with the primary antibody anti-Fas/CD95 in PBS for 30 min at 4°C in the dark. After washing twice with PBS, the cells were incubated with the corresponding secondary antibody for 30 min at 4°C in the dark. The cells were washed with PBS, and the fluorescence was measured on a FACSCalibur flow cytometer (BD Biosciences Clontech). The data were analyzed using the Cell Quest Pro v1.b.1f4b software.