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• W2009358412 abstract "Article26 April 2007free access Splicing factors stimulate polyadenylation via USEs at non-canonical 3′ end formation signals Sven Danckwardt Sven Danckwardt Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Isabelle Kaufmann Isabelle Kaufmann Biozentrum, University of Basel, Basel, SwitzerlandPresent address: Sir William Dunn School of Pathology, University of Oxford, UK Search for more papers by this author Marc Gentzel Marc Gentzel European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Konrad U Foerstner Konrad U Foerstner European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Anne-Susan Gantzert Anne-Susan Gantzert Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Niels H Gehring Niels H Gehring Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Gabriele Neu-Yilik Gabriele Neu-Yilik Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Peer Bork Peer Bork European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Walter Keller Walter Keller Biozentrum, University of Basel, Basel, Switzerland Search for more papers by this author Matthias Wilm Matthias Wilm European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Matthias W Hentze Corresponding Author Matthias W Hentze Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Andreas E Kulozik Corresponding Author Andreas E Kulozik Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Sven Danckwardt Sven Danckwardt Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Isabelle Kaufmann Isabelle Kaufmann Biozentrum, University of Basel, Basel, SwitzerlandPresent address: Sir William Dunn School of Pathology, University of Oxford, UK Search for more papers by this author Marc Gentzel Marc Gentzel European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Konrad U Foerstner Konrad U Foerstner European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Anne-Susan Gantzert Anne-Susan Gantzert Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Niels H Gehring Niels H Gehring Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Gabriele Neu-Yilik Gabriele Neu-Yilik Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Peer Bork Peer Bork European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Walter Keller Walter Keller Biozentrum, University of Basel, Basel, Switzerland Search for more papers by this author Matthias Wilm Matthias Wilm European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Matthias W Hentze Corresponding Author Matthias W Hentze Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany European Molecular Biology Laboratory, Heidelberg, Germany Search for more papers by this author Andreas E Kulozik Corresponding Author Andreas E Kulozik Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany Search for more papers by this author Author Information Sven Danckwardt1,2, Isabelle Kaufmann3, Marc Gentzel4, Konrad U Foerstner4, Anne-Susan Gantzert1,2, Niels H Gehring1,2, Gabriele Neu-Yilik1,2, Peer Bork4, Walter Keller3, Matthias Wilm4, Matthias W Hentze 1,4 and Andreas E Kulozik 1,2 1Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Germany 2Molecular Medicine Partnership Unit, EMBL and University of Heidelberg, Heidelberg, Germany 3Biozentrum, University of Basel, Basel, Switzerland 4European Molecular Biology Laboratory, Heidelberg, Germany *Corresponding authors: Department of Pediatric Oncology, Hematology and Immunology, University of Heidelberg, Im Neuenheimer Feld 156, 69120 Heidelberg, Germany. Tel.: +49 6221 564555; Fax: +49 6221 564559; E-mail: [email protected] European Molecular Biology Laboratory, Meyerhof str. 1, 69117 Heidelberg, Germany. Tel.: +49 6221 387501; Fax: +49 6221 387518; E-mail: [email protected] The EMBO Journal (2007)26:2658-2669https://doi.org/10.1038/sj.emboj.7601699 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The prothrombin (F2) 3′ end formation signal is highly susceptible to thrombophilia-associated gain-of-function mutations. In its unusual architecture, the F2 3′ UTR contains an upstream sequence element (USE) that compensates for weak activities of the non-canonical cleavage site and the downstream U-rich element. Here, we address the mechanism of USE function. We show that the F2 USE contains a highly conserved nonameric core sequence, which promotes 3′ end formation in a position- and sequence-dependent manner. We identify proteins that specifically interact with the USE, and demonstrate their function as trans-acting factors that promote 3′ end formation. Interestingly, these include the splicing factors U2AF35, U2AF65 and hnRNPI. We show that these splicing factors not only modulate 3′ end formation via the USEs contained in the F2 and the complement C2 mRNAs, but also in the biocomputationally identified BCL2L2, IVNS and ACTR mRNAs, suggesting a broader functional role. These data uncover a novel mechanism that functionally links the splicing and 3′ end formation machineries of multiple cellular mRNAs in an USE-dependent manner. Introduction With the exception of some histone mRNAs, all eukaryotic mRNAs possess poly(A)-tails at their 3′ end, which are produced by a two-step reaction involving endonucleolytic cleavage and subsequent poly(A) tail addition (Colgan and Manley, 1997; Keller and Minvielle-Sebastia, 1997; Zhao et al, 1999; Gilmartin, 2005). The specificity and efficiency of 3′ end processing is determined by the binding of a multiprotein complex to the 3′ end processing signal. Most cellular pre-mRNAs contain two core elements. The canonical polyadenylation signal AAUAAA upstream of the cleavage site is recognized by the multimeric cleavage and polyadenylation specificity factor (CPSF). This RNA–protein interaction determines the site of cleavage 10–30 nt downstream, preferentially immediately 3′ of a CA dinucleotide. The second canonical sequence element is characterized by a high density of G/U or U residues and is located up to 30 nt downstream of the cleavage site. This downstream sequence element (DSE) is bound by the 64 kDa subunit of the heterotrimeric cleavage stimulating factor (CstF) that promotes the efficiency of 3′ end processing. Additional proteins, cleavage factors I and II (CF I and CF II), associate and the pre-mRNA is cleaved by CPSF 73 (Ryan et al, 2004; Dominski et al, 2005; Mandel et al, 2006). Subsequently, poly(A) polymerase (PAP) adds ∼250 A-nucleotides to the 3′ end in a template-independent manner. Finally, several molecules of the poly(A)-binding protein II (PABPN1) bind to the growing poly(A) tail and determine its length. These proteins remain attached to the poly(A) tail during nuclear export and enhance both, the stability and the translation of the mRNA (von der Haar et al, 2004). Therefore, defects of mRNA 3′ end formation can profoundly alter cell viability, growth and development by interfering with essential and well-coordinated cellular processes. Although almost all pre-mRNAs are constitutively polyadenylated, alternative and regulated poly(A) site selection represents an important regulatory mechanism for spatial and temporal control of gene expression (Zhao and Manley, 1996; Edwalds-Gilbert et al, 1997; Barabino and Keller, 1999; Zhao et al, 1999). Some 49% of human mRNAs contain more than one polyadenylation site (Yan and Marr, 2005). Alternative and regulated 3′ end processing serves to direct important cellular processes such as immunoglobulin class switch (Takagaki et al, 1996) or the regulated expression of the transcription factor NF-ATc during T-cell differentiation (Chuvpilo et al, 1999). The medical consequences of errors of 3′ end processing are exemplified by the molecular sequelae of a common prothrombotic mutation in the prothrombin (F2) mRNA (F2 20210G → A). This mutation affects the most 3′ nucleotide of the mature mRNA, where the pre-mRNA is endonucleolytically cleaved and polyadenylated; it reverts the physiologically inefficient F2 cleavage site into the most favorable CA dinucleotide context, increasing cleavage site recognition and resulting in the accumulation of correctly 3′ end processed F2 mRNA in the cytoplasm. From these studies, enhanced mRNA 3′ end formation efficiency emerged as a novel molecular principle underlying pathological gene expression and explaining the role of F2 20210G → A in the pathogenesis of thrombophilia (Gehring et al, 2001). Subsequent analyses of the F2 mRNA 3′ end revealed an unusual architecture of non-canonical 3′ end processing signals that explain the susceptibility of the F2 3′ UTR and 3′ flanking sequence to additional, clinically relevant gain-of-function mutations (Danckwardt et al, 2004, 2006a, 2006b). The presence of a sequence element that is located upstream (upstream sequence element (USE)) of the cleavage site within the 3′ UTR stimulates F2 3′ end processing. Moreover, this 15-nucleotide spanning element is both necessary and sufficient to enhance 3′ end processing when inserted into a heterologous β-globin mRNA 3′ UTR (Danckwardt et al, 2004). Unlike (retro-)viral RNAs (Gilmartin et al, 1995; Graveley et al, 1996), stimulatory USEs have been experimentally documented in only a few mammalian mRNAs such as the human complement C2 (Moreira et al, 1998), lamin B2 (Brackenridge and Proudfoot, 2000), cyclooxygenase-2 (Hall-Pogar et al, 2005) and the collagen genes (Natalizio et al, 2002). Biocomputational analyses now predict that USEs may represent a common and evolutionarily conserved feature of mammalian 3′ end formation signals (Legendre and Gautheret, 2003; Hu et al, 2005), suggesting a broad role of USEs in cellular 3′ end mRNA processing. We systematically analyzed the F2 USE and determined its mechanism of function. We show that several splicing factors, CPSF and CstF components specifically bind to the highly conserved USE. The functional characterization of these RNA-binding proteins by RNAi reveals a specific stimulatory effect of known splicing factors on the 3′ end processing of the F2 and C2 USE-containing pre-mRNAs as well as the biocomputationally predicted targets BCL2L2, IVNS and ACTR mRNAs. We propose a model of USE-directed 3′ end processing that involves a novel mRNP that integrates different nuclear pre-mRNA processing steps. Our data also implicate USE-dependent RNP complex formation in the physiology of important cellular processes such as hemostasis (and other thrombin-dependent processes) and the regulation of C2 gene expression as a component of innate immunity. Results The F2 USE increases mRNA 3′ end processing efficiency in a position- and sequence-dependent manner To systematically define the F2 USE and study its mechanism of function, we established an internally controlled in vivo 3′ end processing assay (Danckwardt et al, 2004) and generated constructs that contain a tandem array of 3′ end formation signals, with modifications of the F2 USE within the 5′ site (Figure 1A). In contrast, the unmodified downstream site consists of sequences originating from the wild-type F2 3′ UTR and its 3′ flanking sequences. Thus, the smaller mRNA isoform detected in the poly(A) test (PAT) analysis has been cleaved and polyadenylated at the 5′ site, whereas the longer isoform has been processed at the 3′ site. This experimental setting enabled us to directly compare the processing efficiency of the (manipulated) 5′ site in relation to the control 3′ site, providing an internal control for other potential variables such as transcription or splicing efficiency, which could influence the abundance of the mRNA encoded by the transfected constructs. Figure 1.The F2 USE stimulates mRNA expression and 3′ end formation in a position- and sequence-dependent manner. (A) Schematic representation of the HBB (human β-globin)—F2 hybrid gene construct with a tandem array of two F2 3′ end formation signals used in transient transfection experiments (Ter: stop codon). The F2 USE was either completely or partially replaced by an unrelated nucleotide sequence, or displaced downstream and upstream of its original position as depicted. (B) In vivo assay carried out by transient transfection of a HBB-F2 hybrid gene construct with modifications of the USE as depicted in (A). The bar diagram shows the fold difference of the mRNA ratio processed at the 5′ or the 3′ site (5′/3′) relative to the F2 WT construct (highlighted)±s.e. (at least four independent experiments). (C) In vivo assay carried out by transient transfection of a HBB-F2 hybrid gene construct with modifications of the USE as depicted. The bar diagram shows the fold difference of the mRNA ratio processed at the 5′ or the 3′ site (5′/3′) relative to the F2 WT construct (highlighted)±s.e. (at least four independent experiments). Download figure Download PowerPoint The in vivo assay carried out in transiently transfected HeLa cells (Figure 1B) indicates that the replacement of the entire USE (Unrel., lane 2) almost completely abolishes 3′ end formation at the affected 5′site, when compared to F2 WT (USE, lane 1). In contrast, partial replacement of the first, second or third nucleotide quintett of the USE motif by an unrelated sequence reduces the 3′ end formation capacity at the respective site by ∼2-fold (Figure 1B, lanes 3–5), although significant 3′ end formation was still observed. Because of the critical spatial relationship of canonical 3′ end formation signals to each other, we next analyzed the positional requirements of the USE on mRNA expression and 3′ end formation. For this purpose, it is important to note that the 15-nucleotide spanning USE per se is sufficient to enhance 3′ end processing even in a heterologous 3′ UTR in a context-independent manner (Danckwardt et al, 2004). Displacing the USE, therefore, assays the positional requirements of USE function and is not expected to be compounded by a potential disruption of the surrounding mRNA architecture. The successive shift of the USE downstream towards the polyadenylation signal enhances 3′ end processing (Figure 1B, compare lane 1 with lanes 6 and 7). In contrast, shifting the USE further upstream (by 10, 20 and 30 nucleotides, respectively) resulted in a successive down-modulation of mRNA expression through loss of function of 3′ end processing (Figure 1B, compare lane 1 with lanes 8, 9 and 10). Furthermore, the (relative) changes of the efficiency of the 5′ poly(A) site upon modification (in the context of the tandem construct) were also reflected on the level of absolute mRNA abundance (Supplementary Figure S1), which indicates that the results of the PAT analysis as shown here are not compounded by the fact that the normal F2 3′ end processing site is <100% efficient. Thus, the position of the USE with respect to the canonical polyadenylation signals seems to be a quantitative determinant of its function in 3′ end processing. Previously published data suggest that USEs might stimulate 3′ end processing, at least in part, by recruiting components of the canonical CstF complex (Moreira et al, 1998), which, under normal circumstances, critically depends on the density of U residues. We therefore analyzed whether the F2 USE activity depends on its uridine (U) content or on a more specific sequence context. To this end, we tested constructs with increasing number of U residues within the USE core region (Figure 1C). While decreasing the number of U residues caused a gradual reduction of the 3′ end processing efficiency (Figure 1C, lanes 1–7 and lane 9), increasing the number of U residues also reduced the 3′ end formation efficiency (Figure 1C, lanes 10 and 11), eventually even ablating 3′ end maturation completely (lane 12). Furthermore, the USE of the L3 mRNA that is bound by hFip1 (Kaufmann et al, 2004) was less efficient as the wild-type F2 USE (Figure 1C, lanes 8 and 9). However, duplicating the wild-type USE motif had a stimulatory impact on 3′ end formation by ∼2-fold (Figure 1C, lane 13). These effects seem to be independent of a specific cell type, as similar results were obtained both in transfected HUH-7 and HeLa cells (not shown). These results show that USE function is sequence and position sensitive, and that its potency is not simply determined by its U content. Because CstF binding at U-rich DSEs depends on the density of U-residues, these findings suggest that the F2 USE plays a specific role and does not simply compensate for the absent DSE in the F2 pre-mRNA. In this respect, the F2 mRNA appears to differ from the otherwise similar C2 mRNA (Moreira et al, 1998). Finally, a sequence alignment revealed that the F2 USE is highly conserved among higher eukaryotes and is located at similar positions 17–22 nucleotides upstream of the poly(A) signal (Figure 2A). It comprises two highly conserved overlapping 3′ UTR motifs (UAUUUUU and UUUUGU) belonging to the top 10 out of 106 highly conserved 3′ UTR motifs, with a cross-species conservation rate of 30 and 24%, respectively (Xie et al, 2005). Interestingly, the disruption of either motif individually and/or the presence of only one motif highly correlated with loss of function (Figure 1B and C; Supplementary Figure S2, and data not shown), which emphasizes the importance of both sequence elements. It seems likely, therefore, that the F2 USE has evolved as an optimal sequence context that includes a nonameric core sequence (Figure 2A) in a functionally important region up to 40 nucleotides upstream of the poly(A) site (previously designated as core upstream element (CUE); Hu et al, 2005) to promote 3′ end processing. It is noteworthy that the USE as identified here does not include the tetramers UGUA and UAUA that have recently been shown to account for 3′ end formation at another non-canonical poly(A) site by recruiting the human 3′ processing factor CFIm (Venkataraman et al, 2005). Figure 2.The F2 USE is highly conserved among higher eukaryotes. (A) Sequence comparison of the 3′ ends of vertebrate F2 genes (encompassing the entire 3′ UTR until the poly(A) signal). Shaded sequences denote identity. The graphical representation of the nucleic acid multiple sequence alignment (shown below) highlights the sequence conservation of the F2 USE according to the WebLogo 3 algorithm (Materials and methods), which contains a composite of two highly conserved sequence motifs (TATTTTTGT, highlighted; Xie et al, 2005). (B) Applying a sequence search algorithm that takes into consideration both the strand specificity and the typical length distribution for 3′ UTR motifs (peak >8-mers after exclusion of miRNAs target sites; Xie et al, 2005), more than 1700 human transcripts were identified to contain the nonameric USE core sequence motif. Number of hits are shown according to the localization of the sequence element within the mRNAs (bar diagram, x-axis, 5′ to 3′, left to right). Positive hits were filtered according to their relative location with respect to the poly(A) signal (AATAAA and ATTAAA, respectively). The identity of transcripts that contained the USE core sequence in close proximity to the poly(A) signal (<30 nucleotides upstream of the poly(A) signal (90 and 61 transcripts upstream of the AATAAA or ATTAAA, respectively)) is depicted in Supplementary Tables I and II. Download figure Download PowerPoint We next analyzed if other mRNAs that contain the nonameric USE core sequence can be identified. By using a sequence search algorithm that takes into consideration both the strand specificity and the typical length distribution for 3′ UTR motifs (peak >8-mers after exclusion of miRNAs target sites; Xie et al, 2005), we identified more than 1500 human transcripts that contain the nonameric USE core sequence (Figure 2B). Remarkably, a considerable amount of positive hits were identified in human transcripts with unusually long 3′ UTRs (>1000 nucleotides, not shown). Filtering hits according to the localization of the sequence element within transcripts showed a polar distribution toward their 3′ end (Figure 2B), with more than 500 transcripts that contained the USE motif in the ultimate (tenth) part (0.9–1.0) in a 5′ to 3′ direction. Considering the critical spatial relationship of this sequence element for its function, we identified more than 150 human transcripts that contained the USE core sequence in close proximity to the poly(A) signal (less than 30 nucleotides upstream of the AATAAA and ATTAAA, respectively; see Supplementary Tables I and II). Interestingly, with the exception of four transcripts, all of them contained the USE core sequence motif in their 3′ UTRs. These finding suggest, therefore, that USE-dependent 3′ end processing plays a more general role in many transcripts. Identification of specific nuclear USE-binding proteins To identify trans-acting factors that specifically interact with the F2 USE to promote 3′ end processing, we next performed electromobility shift assays (EMSA) and UV crosslinking experiments. We used a 32P-5′ end-labeled 21-mer RNA oligonucleotide probe including the 15 nucleotide USE core sequence that is both necessary and sufficient to promote 3′ end processing when inserted into a heterologous β-globin gene context (Danckwardt et al, 2004). Incubation of the USE probe with nuclear extract elicits a specific shift (Figure 3A, lanes 2–5 and 6–9). A 21-mer RNA oligonucleotide in which the USE core was replaced by a non-functional unrelated sequence fails to revert the observed shift (Unrel. comp cold, lane 10), whereas an RNA oligonucleotide containing the hFip1-binding site of the L3 mRNA (see above) competes for the formation of the shifted complex (Fip comp cold, lane 11), indicating that the hFip1-binding site interacts with at least one protein that is essential for the F2 USE gel shift. However, recombinant hFip1 failed to result in a shift of the USE oligonucleotide under physiological conditions, nor could it be identified as an interacting protein on the (entire) F2 3′ UTR by RNAse H protection analysis (not shown). No significant shift was observed when the USE probe was incubated with equal amounts of cytoplasmic extract (S100, lanes 12 and 13), indicating that at least one essential protein bound by the USE is nuclear. Figure 3.The F2 USE specifically interacts with nuclear proteins. (A) EMSA carried out with a F2 USE containing 21-mer RNA oligonucleotide (lane 1, free probe) after incubation in HeLa nuclear extract (NE, lanes 2–11) or cytoplasmatic extract (S100, lanes 12 and 13), respectively. Specificity of the RNA–protein interaction is shown by coincubation of an unlabeled F2 USE-containing 21-mer RNA oligonucleotide as cold competitor (lanes 6–9), of an unrelated competitor (lane 10) and a competitor, including the hFip1 binding site of the L3 RNA (lane 11). (B) UV crosslinking study carried out with the same USE-containing or competitor RNA oligonucleotides (lane 0, free probe) after incubation in HeLa nuclear extract (NE, lanes 1–11 (lane 1 without UV light exposure)) or cytoplasmatic extract (S100, lanes 12 and 13), respectively. Download figure Download PowerPoint We next investigated the USE-binding proteins by UV crosslinking (Figure 3B). The USE-specific 21-mer RNA was specifically UV crosslinked to at least five proteins of ∼30–100 kDa. These crosslinks can be competed by cold USE and hFip1-binding site-specific 21-mers (lanes 6–9 and lane 11). Crosslinking studies with cytoplasmic extracts (lanes 12 and 13) showed that some of the USE-binding proteins also appear to be present in the cytoplasm, but the overall pattern of crosslinks is distinct from that generated with nuclear extracts. The affinity of the interaction between the USE and the crosslinking cytoplasmic proteins, however, does not appear to be sufficient to cause a shift in the EMSA. These results demonstrate that the F2 USE directly interacts with at least five different proteins that are predominantly located in the nuclear compartment. Furthermore, the functional significance of this interaction is highlighted by RNA–protein interaction studies using a template with a triple point mutation within the 15-nt USE core affecting the highly conserved nonamer (USEmut). This manipulation results in loss of protein binding (Supplementary Figure S2D and E), which highly correlates with loss of function of the USE (Supplementary Figure S2B, USE and USEmut; lanes 3 and 5). Splicing factors and 3′ end processing proteins bind to the USE We next aimed to identify the F2 USE-binding proteins by affinity purification followed by mass spectrometry. For this purpose, we first ascertained that the 3′biotin-TEG (triethylenglycol)-linker-modification used for immobilization of the RNA bait does not interfere with protein binding to the short 21-mer RNA oligonucleotides (Supplementary Figure S2A). As a specificity control, we used a template with a triple point mutation within the 15-nt USE core (USEmut), which results in similar loss of function as the replacement of the entire USE does (Supplementary Figure S2B, USE, USEmut and Unrel.; lanes 2, 3 and 5). RNA–protein interaction studies based on EMSA and UV crosslinking revealed that this loss of function correlates highly with loss of protein binding (Supplementary Figure S2D and E). The point mutated 21-mer USE sequence (USEmut) thus qualifies as an excellent specificity control for non-functional RNA–protein interactions during the affinity purification procedure. As an additional control for nonspecific RNA–protein interactions, an immobilized 21-mer RNA oligonucleotide with an unrelated sequence was used (Supplementary Figure S2C). Affinity purification yielded several bands that were specific for the USE bait compared with the controls (Figure 4A, lanes 1–3). Comparison of the USE affinity purification-specific band pattern with the patterns of UV crosslinking experiments revealed that the size of some of the affinity-purified proteins corresponds to the size of the proteins identified by UV crosslinking; these proteins thus likely interact with the USE motif directly (Figure 4A, lanes 4 and 5, that is, PSF, p54nrb, U2AF65, hnRNPI, UFAF35). This analysis revealed that the F2 USE-binding proteins include factors known to be involved in 3′ end processing and, surprisingly, in splicing (Table I). Figure 4.Affinity purification followed by mass spectroscopy identifies proteins that specifically interact with the F2 USE. (A) Silver-stained SDS–PAGE polyacrylamide gel of protein samples derived from affinity purification with immobilized 3′ biotinylated 21-mer RNA oligonucleotides with the F2 USE motif (USE, lanes 1 and 5), with a mutated F2 USE motif (USEmut, lane 2) or with an unrelated sequence context (Unrel., lane 3). Lanes 1–3 show protein samples eluted with up to 2000 mM NaCl (starting concentration 150 mM NaCl). Lane 4 shows the autoradiograph of a UV crosslink that highlights the size of direct interaction partners of the F2 USE (dotted arrows indicate putative direct USE-binding proteins that could not be unequivocally assigned by the mass spectrometry data) in comparison to the band pattern of directly and indirectly interacting proteins derived from affinity purification (lane 1). Silver-stained bands in lane 1 were cut out and subjected to mass spectrometry (protein names are only indicated at the respective size where the respective peptide score was maximal; canonical 3′ end processing factors are highlighted in yellow; p54nrb peptides of unexpected small size are highlighted in light gray; for mass spectrometry data, see also Table I). The analysis a" @default.
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