Cotranslational assembly of the yeast SET1C histone methyltransferase complex
2009; Springer Nature; Volume: 28; Issue: 19 Linguagem: Inglês
10.1038/emboj.2009.240
ISSN1460-2075
AutoresAndré Halbach, Haidi Zhang, Agnieszka Wengi, Zofia Jabłońska, Isabel M. L. Gruber, Regula E Halbeisen, Pierre-Marie Dehé, Patrick Kemmeren, Frank C. P. Holstege, Vincent Géli, André P. Gerber, Bernhard Dichtl,
Tópico(s)Epigenetics and DNA Methylation
ResumoArticle27 August 2009free access Cotranslational assembly of the yeast SET1C histone methyltransferase complex André Halbach André Halbach Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Haidi Zhang Haidi Zhang Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Agnieszka Wengi Agnieszka Wengi Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Zofia Jablonska Zofia Jablonska Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Isabel M L Gruber Isabel M L Gruber Institute of Molecular Biology, University of Zürich, Zürich, Switzerland PhD Program in Molecular Life Sciences of the University of Zürich and the ETH Zürich, Zürich, Switzerland Search for more papers by this author Regula E Halbeisen Regula E Halbeisen Institute of Pharmaceutical Sciences, ETH Zürich, Zürich, Switzerland Search for more papers by this author Pierre-Marie Dehé Pierre-Marie Dehé Instabilité du Génome et Cancérogénèse (ICG), CNRS, Marseille, Cedex 20, France Search for more papers by this author Patrick Kemmeren Patrick Kemmeren Department for Physiological Chemistry, Genomics Laboratory, UMC Utrecht, Utrecht, The Netherlands Search for more papers by this author Frank Holstege Frank Holstege Department for Physiological Chemistry, Genomics Laboratory, UMC Utrecht, Utrecht, The Netherlands Search for more papers by this author Vincent Géli Vincent Géli Instabilité du Génome et Cancérogénèse (ICG), CNRS, Marseille, Cedex 20, France Search for more papers by this author André P Gerber André P Gerber Institute of Pharmaceutical Sciences, ETH Zürich, Zürich, Switzerland Search for more papers by this author Bernhard Dichtl Corresponding Author Bernhard Dichtl Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author André Halbach André Halbach Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Haidi Zhang Haidi Zhang Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Agnieszka Wengi Agnieszka Wengi Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Zofia Jablonska Zofia Jablonska Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Isabel M L Gruber Isabel M L Gruber Institute of Molecular Biology, University of Zürich, Zürich, Switzerland PhD Program in Molecular Life Sciences of the University of Zürich and the ETH Zürich, Zürich, Switzerland Search for more papers by this author Regula E Halbeisen Regula E Halbeisen Institute of Pharmaceutical Sciences, ETH Zürich, Zürich, Switzerland Search for more papers by this author Pierre-Marie Dehé Pierre-Marie Dehé Instabilité du Génome et Cancérogénèse (ICG), CNRS, Marseille, Cedex 20, France Search for more papers by this author Patrick Kemmeren Patrick Kemmeren Department for Physiological Chemistry, Genomics Laboratory, UMC Utrecht, Utrecht, The Netherlands Search for more papers by this author Frank Holstege Frank Holstege Department for Physiological Chemistry, Genomics Laboratory, UMC Utrecht, Utrecht, The Netherlands Search for more papers by this author Vincent Géli Vincent Géli Instabilité du Génome et Cancérogénèse (ICG), CNRS, Marseille, Cedex 20, France Search for more papers by this author André P Gerber André P Gerber Institute of Pharmaceutical Sciences, ETH Zürich, Zürich, Switzerland Search for more papers by this author Bernhard Dichtl Corresponding Author Bernhard Dichtl Institute of Molecular Biology, University of Zürich, Zürich, Switzerland Search for more papers by this author Author Information André Halbach1,‡, Haidi Zhang1,‡, Agnieszka Wengi1,‡, Zofia Jablonska1, Isabel M L Gruber1,2, Regula E Halbeisen3, Pierre-Marie Dehé4, Patrick Kemmeren5, Frank Holstege5, Vincent Géli4, André P Gerber3 and Bernhard Dichtl 1 1Institute of Molecular Biology, University of Zürich, Zürich, Switzerland 2PhD Program in Molecular Life Sciences of the University of Zürich and the ETH Zürich, Zürich, Switzerland 3Institute of Pharmaceutical Sciences, ETH Zürich, Zürich, Switzerland 4Instabilité du Génome et Cancérogénèse (ICG), CNRS, Marseille, Cedex 20, France 5Department for Physiological Chemistry, Genomics Laboratory, UMC Utrecht, Utrecht, The Netherlands ‡These authors contributed equally to this work. *Corresponding author. Institute of Molecular Biology, University of Zürich, Winterthurer Strasse 190, Zürich 8057, Switzerland. Tel.: +41 44 635 3160; Fax: +41 44 635 6811; E-mail: [email protected] The EMBO Journal (2009)28:2959-2970https://doi.org/10.1038/emboj.2009.240 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info While probing the role of RNA for the function of SET1C/COMPASS histone methyltransferase, we identified SET1RC (SET1 mRNA-associated complex), a complex that contains SET1 mRNA and Set1, Swd1, Spp1 and Shg1, four of the eight polypeptides that constitute SET1C. Characterization of SET1RC showed that SET1 mRNA binding did not require associated Swd1, Spp1 and Shg1 proteins or RNA recognition motifs present in Set1. RNA binding was not observed when Set1 protein and SET1 mRNA were derived from independent genes or when SET1 transcripts were restricted to the nucleus. Importantly, the protein–RNA interaction was sensitive to EDTA, to the translation elongation inhibitor puromycin and to the inhibition of translation initiation in prt1-1 mutants. Taken together, our results support the idea that SET1 mRNA binding was dependent on translation and that SET1RC assembled on nascent Set1 in a cotranslational manner. Moreover, we show that cellular accumulation of Set1 is limited by the availability of certain SET1C components, such as Swd1 and Swd3, and suggest that cotranslational protein interactions may exert an effect in the protection of nascent Set1 from degradation. Introduction Most proteins in a cell are integral components of heterogeneous multi-subunit complexes. Assembly pathways for such complexes have been studied, for example, for ribonucleoproteins that form on small nuclear or small nucleolar RNAs (Filipowicz and Pogacic, 2002; Paushkin et al, 2002; Staley and Woolford, 2009) and for membrane-bound complexes in organelles (Choquet and Vallon, 2000; Ackerman and Tzagoloff, 2005). The manner in which large structures such as ribosomal subunits are built poses another huge challenge for biologists (Fatica and Tollervey, 2002); moreover, very little is known about the formation of a large number of diverse multi-protein complexes in the cytosol. In particular, the mechanisms that ensure specific association of a certain set of interaction partners remain largely obscure. Chromatin modification enzymes such as histone methyltransferases (HMTases) are often contained in multi-protein assemblies (Kouzarides, 2007). In yeast, the Set1 protein is required for all methylation of histone 3 lysine 4 (H3K4) (Briggs et al, 2001; Roguev et al, 2001; for a recent review see Dehe and Geli, 2006). The protein stably associates with seven other subunits, which have been given multiple denominations: Swd1/Cps50p/Saf49p, Swd2/Cps35p/Saf35p, Swd3/Cps30p/Saf35p, Spp1/Cps40p/Saf41p, Bre2/Cps60p, Sdc1/Cps25p/Saf19p and Shg1 (Miller et al, 2001; Roguev et al, 2001; Nagy et al, 2002). The resulting SET1 complex is also known as COMPASS (Miller et al, 2001). For simplicity, we will use the standard nomenclature of the proteins as indicated by the Saccharomyces Genome Database (http://www.yeastgenome.org/). Only the Swd2 subunit of the complex is essential for cell viability, which is probably because of an independent role of the protein as a component of the cleavage and polyadenylation factor (CPF) complex (Cheng et al, 2004; Dichtl et al, 2004). H3K4 HMTases from yeast to humans carry, in addition to variable enzyme-specific subunits, a common set of associated proteins: the so-called ‘MLL core complex’. This complex consists of a catalytic SET domain containing subunit and WDR5, RpBP5 and Ash2L proteins, which are homologous to yeast Swd3, Swd1 and Bre2 (reviewed in Ruthenburg et al, 2007). Analysis of these core components in vivo and in vitro showed a central role for WDR5 in complex integrity and substrate interaction (Wysocka et al, 2005; Dou et al, 2006; Steward et al, 2006). It is noteworthy that the requirement for certain core components to maintain complex integrity seems to be conserved in yeast, as a mutation of the essential WD-repeat protein Swd2 (Dichtl et al, 2004) or a deletion of the genes encoding the non-essential Swd1 and Swd3 subunits resulted in a severe under-accumulation of Set1 (Dehe et al, 2006; Steward et al, 2006). Although a reconstitution of an MLL core enzyme was recently achieved with heterologously expressed proteins in vitro (Dou et al, 2006), little is known about the manner by which HMTase complexes are formed in vivo. Set1 harbours an evolutionarily conserved RNA-binding domain in its amino-terminal half, comprising two RNA recognition motifs (RRM1 and RRM2) (Tresaugues et al, 2006). We recently showed that, although this domain can bind RNA in vitro, mutations within RRM2 do not interfere with the HMTase activity of SET1C in vivo (Tresaugues et al, 2006). However, removal of the entire RRM domain or point mutations within RRM1 resulted in a specific loss of tri-methylation, consistent with the idea that RNA may have a role in regulating SET1C (Schlichter and Cairns, 2005; Tresaugues et al, 2006). Moreover, it was suggested that the catalytic SET domain and associated n-SET and post-SET domains can bind single-stranded DNA and RNA in vitro (Krajewski et al, 2005). Nevertheless, the physiological implications of nucleic acid binding by SET1C and related complexes remain unclear. This study was initiated by a search for RNA molecules that bind to SET1C. Our analyses showed that SET1 mRNA was associated with a SET1C sub-complex, which we refer to as SET1RC. Our experiments provide evidence that SET1RC assembles on nascent Set1 during translation. We propose that this cotranslational assembly couples the synthesis and accumulation of Set1, the catalytic subunit of the complex, with the formation of SET1C. Cotranslational processes, as described here, may be of wider relevance for the assembly of multi-protein complexes. Results Specific enrichment of SET1 mRNA in Shg1–TAP purifications To test for RNA associated with SET1C, we partially purified the complex from Shg1–TAP extracts; a non-tagged isogenic wild-type strain served as control. RNA isolated from extracts and from purified Shg1–TAP material was used to generate cDNA probes labelled with different fluorescent dyes, which were then mixed and hybridized to Saccharomyces cerevisiae oligonucleotide microarrays (Gerber et al, 2004; Supplementary data). The scatter plot shown in Figure 1B illustrates that SET1 RNA was highly enriched in Shg1–TAP affinity isolates (average enrichment ∼23-fold; P-value was 50% and hence were not likely to represent true targets. The microarray results were confirmed by real-time PCR (Figure 1C): SET1 mRNA was found to be approximately 50-fold enriched compared with ACT1 mRNA in Shg1–TAP isolates; transcripts encoding other SET1C subunits (SPP1, SDC1), ribosomal proteins (RPL29A and RPS7A) and histones (HHT1 and HTA2) were not enriched. Moreover, Figure 1D shows that a SET1 signal was obtained when cDNA synthesis was primed with oligo(dT) instead of random hexamers (which were used in Figure 1C), suggesting that the RNA contained poly(A). To exclude the fact that we detected short polyadenylated fragments, we used for cDNA synthesis a primer that anneals in the 3′ UTR (oligo UTR; see Figure 1A). We observed similar signal intensities for primer pairs PP1, PP2 and PP4, indicating that RNAs extended from the 3′ UTR to the 5′ end of the transcript (Figure 1E). Taken together, our analyses suggested that bona fide, full-length SET1 mRNA was associated with Shg1–TAP. Figure 1.Specific association of SET1 mRNA with Shg1–TAP. (A) Schematic presentation of the SET1 gene. Oligonucleotides (primer pairs PP1–PP6) used for qPCR analysis or reverse transcription (oligo UTR) and their location relative to the translational start codon (+1) are indicated. (B) Average RNA enrichment from four independent microarray hybridizations analysing Shg1–TAP affinity isolates (y-axis) and mock controls (x-axis). Data are shown for 6253 features. SET1 data are depicted in red. (C) Quantitative RT–PCR analysis of the indicated target genes on RNA co-purifying with partially purified Shg1–TAP (solid black bars) or from a control extract (open white bars). Fold enrichment was determined as described in ‘Materials and methods’. (D) Quantitative RT–PCR analysis as described in (C) with the indicated primer pairs, with cDNA that was obtained after oligo(dT)-primed reverse transcription. (E) Quantitative RT–PCR analysis as described in (C) with the indicated primer pairs with cDNA that was obtained after reverse transcription that was primed with oligo UTR. Download figure Download PowerPoint SET1 mRNA is associated with a SET1C sub-complex Most of cellular Shg1 is associated with SET1C (Roguev et al, 2001). To test whether other SET1C subunits were also bound to SET1 mRNA, we affinity purified all eight SET1C subunits individually from extracts. An immunoblot analysis confirmed the correct expression of TAP-fusion proteins and enrichment of proteins after IgG-Sepharose adsorption and elution by TEV-protease cleavage (Supplementary Figure S1A). We observed an enrichment of SET1 mRNA with TAP–Set1, Spp1–TAP, Swd1–TAP and Shg1–TAP, but not with Bre2–TAP, Sdc1–TAP, Swd2–TAP and a non-tagged wild-type strain (Figure 2A). Swd3–TAP gave a signal that was slightly above background, indicating that it may either associate loosely with the complex or that this protein does not support efficient purification because of other limitations. No enrichment of RPS7A mRNA took place in any case. To evaluate whether the TAP-tag compromised the functionality of the fusion proteins and potentially influenced the efficiency of SET1 mRNA co-purification, we tested H3K4 methylation by western blot. Supplementary Figure S1B shows that all analysed strains had similar levels of H3K4 di- and tri-methylation, suggesting that TAP-fusion proteins were functional. Figure 2.SET1 mRNA is associated with a SET1C sub-complex. (A) Affinity purification was carried out with extracts obtained from the indicated strains. After RNA extraction and cDNA synthesis, qPCR analysis was performed to detect SET1 RNA (PP4; solid black bars) or a control RNA (RPS7A; open white bars). Values represent the mean of three to five comparable experiments for each indicated strain and error bars indicate s.d. (B) The diagram (top) depicts the domain structure of Set1 (RRM1, RRM2, n-SET, SET and post-SET; not in scale) and numbers delineate the interaction domains for Shg1, Spp1 and Bre2, as derived from yeast two-hybrid results. The region of interaction (aa 1–900; Dehe et al, 2006) of the Swd1–Swd3 heterodimer is indicated. Sdc1 forms a heterodimer with Bre2 and does not directly bind to Set1; no interaction domain is known for Swd2. The bottom panels show GST pull-down assays with GST–Set1 and in vitro translated 35S-methionine-labelled Swd1, Spp1 and Shg1 proteins. Input lanes show 10% of radioactive protein included in the binding reactions. The GST lane shows background binding. (C) Shg1–TAP and associated material were partially purified by affinity chromatography on IgG-Sepharose and the obtained material was then applied on a Superdex 200 column. The upper graph displays the obtained protein elution profile. The indicated fractions were then analysed by RT–PCR for the presence of SET1 mRNA (lower graph). The western blot at the bottom of the figure shows the distribution of Shg1 in the Superdex 200 fractions. The arrow indicates the void volume of the column. Download figure Download PowerPoint Next, we addressed whether Set1, Spp1, Swd1 and Shg1 associate with SET1 mRNA as constituents of the same complex. Results from yeast two-hybrid screening identified domains within Set1 that mediate interactions with Shg1 (amino acid (aa) 462–560) and Spp1 (aa 762–794) (BD, unpublished data; Figure 2B). Furthermore, Bre2 interacted with a Set1 domain encompassing aa 900–1081, consistent with previous results (Dehe et al, 2006). No interaction domains on Set1 are known for Swd1, Swd2 and Swd3, but the Swd1–Swd3 heterodimer does not require the C-terminal SET domain (aa 901–1081) for association with SET1C (Dehe et al, 2006). In pull-down assays, Swd1, Spp1 and Shg1 efficiently bound to GST–Set1 but not to GST alone (Figure 2B), consistent with the view that the three proteins directly associate with Set1. Furthermore, a Superdex 200 gel-filtration analysis of material associated with Shg1–TAP after IgG purification suggested that SET1 mRNA was associated with a large molecular weight complex that eluted with the void volume of the column (Figure 2C). Taken together, our results showed the existence of a SET1 mRNA containing complex that we will henceforth refer to as SET1RC (SET1 mRNA-associated complex). Swd1, Spp1 and Shg1 are dispensable for SET1 mRNA binding To determine the requirements for SET1RC formation and stability, we initially tested SET1 mRNA association to TAP-tagged SET1RC subunits in the absence of other SET1RC proteins. As shown in Figure 2A, SET1 mRNA association was observed with TAP–Set1 in Δswd1, Δspp1 and Δshg1 backgrounds, with Spp1–TAP in Δswd1 and Δshg1 backgrounds, with Shg1–TAP in Δswd1 and Δspp1 backgrounds and with Swd1–TAP in the Δshg1 background. The signal obtained with the TAP–Set1 Δspp1 strain was elevated in this analysis, but remained within the limits of variation that we observed in numerous such experiments. The only combination that gave no signal was Swd1–TAP in Δspp1, indicating that the absence of Spp1 weakens the binding of Swd1–TAP to the complex. The absence of Bre2 and Sdc1, which we found not to associate with SET1RC, was tested in Spp1–TAP and Shg1–TAP backgrounds, respectively, and did not interfere with the binding of SET1 mRNA. We, along with others, previously found that inactivation of Swd1, Swd2 and Swd3 resulted in a severe under-accumulation of Set1 (Dichtl et al, 2004; Dehe et al, 2006; Nedea et al, 2008), and western blot analysis confirmed strongly reduced levels of TAP–Set1 in Δswd1 and Δswd3 cells (Figure 6A). Remarkably, co-purification of SET1 mRNA with TAP–Set1 occurred with the same efficiency, irrespective of the presence or absence of either Swd1 or Swd3 (Figure 2A). This indicated that the amount of TAP–Set1 associated with SET1RC was not significantly reduced, despite strongly reduced cellular levels of the protein. In summary, we found that SET1RC formation can occur in the absence of Swd1, Spp1 and Shg1, suggesting that only Set1 is required for SET1 mRNA binding. mRNA binding is not associated with a specific Set1 domain Set1 carries RRM1- and RRM2 RNA-binding motifs in the N-terminal half and the catalytic SET domain at its C-terminus (Figure 3A). To test which domains are required for SET1 mRNA binding we fused Set1 fragments to ProteinA and assayed whether encoding mRNAs co-purified. The data in Figure 3B illustrate that both the N- and the C-terminal half of the protein (residues 1–571 and 572–1081) bound their encoding mRNA with an efficiency similar to full-length protein (residues 1–1081). Thus, SET1 mRNA binding occurred both with a Set1 protein, which carried the RRM domains, and the C-terminal part of the protein, which lacked RRMs. Moreover, proteins carrying mutations in RRM1 (1–571YF/AA) or RRM2 (1–571H/A), which strongly reduced RNA binding in vitro (Tresaugues et al, 2006), retained the ability to interact with cognate mRNA (Figure 3B). Thus, RNA binding of the N-terminal domain did not depend on intact RRMs, consistent with the observation that the C-terminal fragment, which completely lacked these motifs, also bound RNA. Figure 3.Characteristics of SET1 mRNA association. (A) Schematic presentation of Set1 (not in scale); numbers indicate approximate amino-acid positions of RRM1, RRM2 and SET domains. Also indicated are fragments that were expressed as plasmid-encoded ProteinA fusions in Δset1 strains. A strain carrying the empty ProteinA-fusion vector was analysed for control. The 1-571 YT/AA fragment carries two point mutations in RRM1, and 1–571 H/A carries a single point mutation in RRM2 (asterisks indicate the presence of point mutations; Tresaugues et al, 2006). (B) RT–PCR analysis of RNA isolated from IgG purifications of the indicated ProteinA-fusion proteins. PP2 and PP4 were used to detect SET1 mRNA or fragments thereof (primer locations are indicated in panel C). RPS7A served as control. Note that PP2 will only detect full-length and N-terminal fragments and PP4 will detect only full-length and C-terminal fragments. (C) Schematic representation of N- and C-terminal halves of Set1, which were expressed as ProteinA fusions from plasmids in strains carrying a chromosomal SET1-MYC gene (1–1081-Myc). The location of primer pairs used for qPCR is also shown. Wavy lines represent mRNAs that correspond to Set1 proteins as indicated. Double-headed arrows indicate interactions between proteins and RNA that were tested. The crossed-out double-headed arrows indicate interactions that were not observed. (D) RT–PCR analysis of RNA associated with the indicated ProteinA-fusion proteins. The 1–571 mRNA was detected with PP1 and 572–1081 mRNA was detected with PP4. 1–1081-Myc mRNA was detected with PP6 in the 1–571 strain and with PP1 in the 572–1081-expressing strain, as indicated. (E) RNA analysis of SHG1-TAP strains carrying plasmids encoding the 5′ and 3′ halves of the SET1 mRNA, under control of a T7 RNA polymerase promoter. RT–PCR was carried out with primers specific for T7 RNAs (T7-5′ and T7-3′ primer pairs, respectively) to determine the fold-increase of T7 RNA compared with that of chromosomally encoded SET1 mRNA. (F) RT–PCR analysis of RNA co-purifying with Shg1–TAP strains as described in (E). Endogenous SET1 mRNA was detected with PP1 and PP4 (solid black bars; note that PP1 does not detect the T7-3′ fragment and that PP4 does not detect the T7-5′ fragment). The T7-derived RNAs were detected with T7-5′ and T7-3′ primer pairs (open white bars; note that T7 primer pairs specifically detect T7 transcripts and not endogenous SET1 mRNA). (G) Sub-cellular fractionation of Shg1–TAP cells. Total extract (XT), post-nuclear supernatant (PNS) and nuclear (Nc) fractions were analysed by western blot with the indicated antibodies. H3K4me3 is a nuclear-specific marker, whereas Pgk1 is cytoplasmic. Download figure Download PowerPoint Next, we asked whether the interaction may be restricted to mRNAs and to their derived proteins, or whether Set1 proteins were able to bind SET1 RNA derived from a second SET1 locus. For this purpose, we expressed the ProteinA fusions of the N- and C-terminal Set1 halves in a strain that also carries a chromosomal full-length SET1-MYC gene (Figure 3C). We found that both Set1 fragments efficiently co-purified their encoding mRNA, but neither of the fragments bound to SET1-MYC mRNA (Figure 3D). This suggested that binding was restricted to Set1 proteins and their encoding mRNAs. To address the requirements of sub-cellular RNA localization for association within SET1RC, we asked whether nuclear-restricted SET1 RNAs were included in the complex. RNAs corresponding to the N- and C-terminal halves of SET1 were expressed in SHG1-TAP strains with nuclear localized T7 RNA polymerase (Dower and Rosbash, 2002). As termination of these transcripts was mediated by a T7 terminator, the RNAs will not be exported to the cytoplasm (Dower and Rosbash, 2002). Figure 3E shows that we achieved efficient transcription of T7-derived RNAs (approximately 180-fold relative to endogenous SET1 mRNA levels). Analysis of RNA co-purifying with Shg1–TAP showed, however, that neither of the T7 RNAs was enriched with this procedure (Figure 3F). In contrast, chromosomally encoded SET1 mRNA was bound, as expected (Figure 3F). These results suggested that nuclear-restricted RNAs do not qualify for association with SET1RC. The above data are consistent with the possibility that SET1RC formation may rely on a cytoplasmic localization of its components. To test this hypothesis, we performed fluorescence microscopy on strains carrying GFP-tagged alleles of SHG1 and SET1. We observed nuclear localization of these SET1C components as expected (Supplementary Figure S4; Huh et al, 2003). However, we could not conclude about cytoplasmic protein levels, as the signal strength was very weak compared with auto-fluorescence obtained with a GFP-only control strain (Supplementary Figure S4). To circumvent this problem, we used Shg1–TAP-expressing cells and performed sub-cellular fractionation. Figure 3G shows that Shg1–TAP distributed to both nuclear and post-nuclear fractions. In contrast, histone H3 tri-methylated at lysine 4 (H3K4me3) was exclusively nuclear and the glycolysis enzyme 3-phosphoglycerate kinase (Pgk1) localized to the post-nuclear supernatant. We conclude that some cellular Shg1–TAP was associated with the cytoplasmic compartment. SET1 mRNA binding is sensitive to EDTA and puromycin One of the simplest explanations to account for our observation in Figures 1, 2 and 3 is that the SET1 mRNA interactions depended on nascent Set1 peptides that were associated with their mRNAs during translation. Therefore, we wanted to test whether SET1RC formation was indeed translation dependent. For this purpose, we initially performed RNA co-purification experiments with Shg1–TAP extracts in the presence of EDTA. This treatment dissociates ribosomes and we found that SET1 mRNA binding was indeed abolished under these conditions (Figure 4A). However, EDTA could also disrupt binding independently of ribosome dissociation. Therefore, we performed affinity purification in the presence and absence of the translation inhibitor puromycin. Figure 4B shows that the drug reduced co-purification of SET1 mRNA by more than six-fold, suggesting that SET1 mRNA binding required translation. Figure 4.SET1 mRNA binding is translation dependent. (A) RT–PCR analysis of RNA associated with Shg1–TAP, which was purified in the presence of 10 mM EDTA. PCR was carried out with PP4 to detect SET1 mRNA; RPS7A served as control. Shown are results from two independent extracts (EDTA XT 1 and EDTA XT 2). (B) Shg1–TAP was purified from identical extracts on IgG-Sepharose in the absence (XT+H2O) or presence of 200 μg/ml puromycin (XT+puromycin). Shown is a representative RT–PCR using PP4 to detect SET1 mRNA; RPS7A served as control. (C) A254 recordings of sucrose-gradient fractionations of extracts from a prt1-1 strain expressing plasmid-born ProtA-SET1, after growth at 27°C or after a shift to 37°C for 35 min, as indicated. (D) RT–PCR analysis of RNA associated with ProtA–Set1 in extracts that were obtained as described in (C). PCR was carried out with PP4 to detect SET1 mRNA; RPS7A served as control. Values represent the mean of at least three independent extracts produced for each strain at the indicated conditions and error bars indicate s.d. (E) Co-immunoprecipitation experiment with extracts (XT) prepared from the indicated TAP-tagged strains that also carried a plasmid-expressing HA–Set1. After adsorption of material on IgG-Sepharose, bound material was eluted by TEV-protease cleavage (eluate). The obtained fractions were analysed by western blot using anti-HA (top) and anti-TAP (bottom) antibodies, respectively. Migration of molecular weight marker (M) and HA- and TAP-tagged proteins is indicated on the right. Download figure Download PowerPoint To further scrutinize the relationship of SET1RC and translation, we initially tested the efficiency of SET1 mRNA binding when glucose was withdrawn from the medium, a condition that rapidly inhibits the initiation of protein synthesis (Ashe et al, 2000). Sucrose-gradient fractionation showed that most polysomes disappeared on glucose withdrawal, but a minor polysomal signal remained (we estimate less than 5%; Supplementary Figure S2A), in agreement with previous reports (Ashe et al, 2000). The bulk of RPL29 mRNA exited polysomal fractions (8–12) on glucose withdrawal and relocated to early fractions (1–5), whereas some mRNA remained associated with fractions 7–9, overlapping the 80S monosome peak (Supplementary Figure S2B). In contrast, SET1 mRNA distribution was very similar in the presence and absence of glucose (Supplementary Figure S2B). Consistent with this, we also found that binding of Shg1–TAP to SET1 mRNA was not sensitive to glucose withdrawal (Supplementary Figure S2C). We conclude that control of SET1 mRNA translation does not respond to glucose withdrawal and therefore occurs uncoupled from most cellular mRNAs. To interfere with translation initiation using an alternative approach, we analysed SET1 mRNA binding in prt1-1 temperature-sensitive strains expressing plasmid encoded ProtA–
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