An essential DNA strand-exchange activity is conserved in the divergent N-termini of BLM orthologs
2010; Springer Nature; Volume: 29; Issue: 10 Linguagem: Inglês
10.1038/emboj.2010.61
ISSN1460-2075
Autores Tópico(s)Genomics and Chromatin Dynamics
ResumoArticle13 April 2010free access An essential DNA strand-exchange activity is conserved in the divergent N-termini of BLM orthologs Chi-Fu Chen Chi-Fu Chen Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Steven J Brill Corresponding Author Steven J Brill Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Chi-Fu Chen Chi-Fu Chen Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Steven J Brill Corresponding Author Steven J Brill Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA Search for more papers by this author Author Information Chi-Fu Chen1 and Steven J Brill 1 1Department of Molecular Biology and Biochemistry, Rutgers University, Piscataway, NJ, USA *Corresponding author. Department of Molecular Biology and Biochemistry, Rutgers University, CABM, Room 304, Piscataway, NJ 08854, USA. Tel.: +1 732 235 4197; Fax: +1 732 445 6186; E-mail: [email protected] The EMBO Journal (2010)29:1713-1725https://doi.org/10.1038/emboj.2010.61 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The gene mutated in Bloom's syndrome, BLM, encodes a member of the RecQ family of DNA helicases that is needed to suppress genome instability and cancer predisposition. BLM is highly conserved and all BLM orthologs, including budding yeast Sgs1, have a large N-terminus that binds Top3–Rmi1 but has no known catalytic activity. In this study, we describe a sub-domain of the Sgs1 N-terminus that shows in vitro single-strand DNA (ssDNA) binding, ssDNA annealing and strand-exchange (SE) activities. These activities are conserved in the human and Drosophila orthologs. SE between duplex DNA and homologous ssDNA requires no cofactors and is inhibited by a single mismatched base pair. The SE domain of Sgs1 is required in vivo for the suppression of hyper-recombination, suppression of synthetic lethality and heteroduplex rejection. The top3Δ slow-growth phenotype is also SE dependent. Surprisingly, the highly divergent human SE domain functions in yeast. This work identifies SE as a new molecular function of BLM/Sgs1, and we propose that at least one role of SE is to mediate the strand-passage events catalysed by Top3–Rmi1. Introduction The RecQ family of DNA helicases comprises five eukaryotic members that participate in homologous recombination to repair double-stranded DNA breaks (DSBs) and stalled replication forks. Defects in all five helicases are associated with genome instability, and defects in three (BLM, WRN and RecQ4) are known to cause cancer predisposition syndromes in humans (Wang et al, 2003; Hu et al, 2005; Chu and Hickson, 2009). Structurally, these enzymes are well characterized. All members contain a highly conserved DNA helicase domain (Supplementary Figure S1A), and most contain an RQC domain that participates in DNA binding and protein–protein interactions (Bernstein et al, 2003; Bennett and Keck, 2004) (Figure 1A). Some RecQ members also contain a C-terminal HRDC domain that assists in DNA binding (Bernstein and Keck, 2005) and is required for in vitro activity (Wu et al, 2005). These helicases efficiently unwind a variety of model recombination intermediates such as Holliday Junctions (HJs), D-loops, replication forks and G-quadruplex DNA (Sun et al, 1998; Karow et al, 2000; LeRoy et al, 2005; Bachrati et al, 2006; Ralf et al, 2006; Capp et al, 2009). However, a clear understanding of how these enzymes suppress inappropriate recombination in vivo is lacking. Figure 1.Identification of a ssDNA-binding activity in Sgs11−652 (A) Schematic representations of the full-length 1447 aa Sgs1 protein and Sgs11−652. Domains: TR, Top3–Rmi1 binding; B, C-terminal domain in Bloom's syndrome DEAD helicases (BDHCT) homology; RQC, RecQ C-terminal homology; HRDC, Human RecQ and RnaseD C-terminal homology. (B) EMSA assays contained the indicated concentrations of His6-tagged Sgs11−652 and 1 nM of either primed ssDNA (oligo #16 annealed to ∅X174 ssDNA) or a plasmid-based D-loop (oligo #17 annealed to pSK+ DNA by RecA). Asterisks represent positions of 32P-labelling. The reaction mixtures were incubated for 20 min at room temperature under standard conditions as described in 'Materials and methods'. After incubation, the products were subjected to electrophoresis in composite 2.5% polyacrylamide/0.8% agarose gels followed by phosphorimager analysis. (C) Sgs11−652 (300 nM) was incubated together with 1 nM primed ssDNA and various concentrations of the four unlabelled versions of pSK+ DNA as indicated. Incubation and analysis was performed as in (B). M is a mock incubation in the absence of Sgs1 protein. (D) The indicated concentrations of Sgs11−652 were incubated with one of three 32P-labelled ssDNAs at a concentration of 1 nM: poly(dT)174, a 90 nt oligo of random sequence (oligo #18), or oligo(dT)60. Incubation and analysis was performed as in (B) except for the use of 10% PAGE. Download figure Download PowerPoint The only RecQ DNA helicase conserved in lower eukaryotes is the ortholog of the gene defective in Bloom's syndrome (BS) BLM. BS is a rare autosomal disease associated with elevated levels of sister chromatid exchange and susceptibility to a wide variety of cancers (Chaganti et al, 1974; German et al, 2007). Similar to human BLM, budding yeast Sgs1 forms a tight complex with two other subunits, Top3 and Rmi1 (Gangloff et al, 1994; Bennett et al, 2000; Wu et al, 2000; Fricke et al, 2001; Ui et al, 2001; Mullen et al, 2005). The hyper-recombinational phenotypes of BS cells and yeast sgs1Δ mutants (Gangloff et al, 1994) indicate that the BLM–TOP3–RMI1 complex functions as an anti-recombinase that may be related to its ability to dissolve double HJs and/or unwind D-loops in vitro (Wu and Hickson, 2003; Bachrati et al, 2006). However, the complex is known to be multi-functional. It interacts with the Rad51 strand-exchange (SE) protein (Wu et al, 2001; Bugreev et al, 2009), it is required for recombination in model systems such as fission yeast and Drosophila (Adams et al, 2003; Cromie et al, 2008), and it promotes 5′-end resection (Mimitou and Symington, 2008; Nimonkar et al, 2008; Zhu et al, 2008). Furthermore, BLM promotes single-strand DNA (ssDNA) annealing (SA) in vitro. Considerable evidence indicates that this SA activity is intrinsic to the RecQ helicase domain: annealase activity has been identified in all five RecQ members, including the two smallest (RecQ1 and RecQ5), and although it is ATP independent, it is typically inhibited by non-hydrolyzable ATP analogues (Garcia et al, 2004; Cheok et al, 2005; Machwe et al, 2005; Sharma et al, 2005; Macris et al, 2006; Muftuoglu et al, 2008; Xu and Liu, 2009). For RecQ5 and WRN, SA activity has been localized to RecQ C-terminal domains (Garcia et al, 2004; Muftuoglu et al, 2008). Similarly, the N-terminal extension of BLM is dispensible for an SA activity that maps to residues 642–1350 (Cheok et al, 2005). Finally, BLM shows DNA SE activities that are both ATP dependent (Machwe et al, 2005; Weinert and Rio, 2007) and ATP independent (Machwe et al, 2005; Bugreev et al, 2009). For reasons that are unclear, the ATP-independent SE activities are sensitive to non-hydrolyzable ATP analogues and certain DNA helicase mutations (Machwe et al, 2005; Bugreev et al, 2009). Apart from the DNA helicase domain, BLM/Sgs1orthologs contain a poorly characterized N-terminal domain of about 650 amino acids (aa) (Figure 1A). Functional analysis of the N-terminus has been hindered in part by the lack of sequence conservation between orthologs (Supplementary Figure S1B). In yeast, this domain (Sgs11−652) is known to be physiologically important (Mullen et al, 2000; Rockmill et al, 2003; Bernstein et al, 2009) and at least one essential role is to bind Top3 and Rmi1 through its N-terminal 100 aa. In searching for a biochemical function of Sgs11−652 we identified a sub-domain that shows ssDNA binding and SA activity. Despite the lack of sequence conservation, this domain is functionally conserved in multiple BLM orthologs. In addition, it shows an SE activity that is inactive on homologous templates containing a single mismatch. To determine the physiological role of this activity we characterized an SGS1 allele lacking the SE domain. This sgs1-ΔSE allele showed a null phenotype in most in vivo assays including suppression of hyper-recombination. These data indicate that DNA SE is an essential conserved function of BLM/Sgs1 orthologs, and we speculate that its in vivo role is to promote DNA SE in conjunction with Top3–Rmi1 at double HJs and D-loops. Results Identification of a novel ssDNA-binding activity in BLM orthologs Structure–function analysis of Sgs1 showed earlier that deletion of the Top3–Rmi1-binding domain (TR; Figure 1A) creates a hypermorphic phenotype in yeast (Mullen et al, 2000; Bennett and Wang, 2001; Weinstein and Rothstein, 2008); that is, removal of the first 80–150 aa results in slow-growth and hyper-recombinational phenotypes that are more extreme than the sgs1Δ null. These hypermorphic phenotypes are suppressed either by a point mutation that eliminates Sgs1 DNA helicase activity or by deleting more of the N-terminus to aa 323 (Mullen et al, 2000; Weinstein and Rothstein, 2008). The simplest interpretation of this result is that the first 323 aa of Sgs1 contain a helicase-dependent activity that is toxic when untethered to Top3–Rmi1. As prior studies had failed to identify deoxyoligonucleotide (oligo)-binding activity in the N-terminus of Sgs1 or BLM (Cheok et al, 2005; Chen and Brill, 2007), we assayed it for structure-specific DNA-binding activity. To this end, we incubated Sgs11−652 protein with two large radiolablled probes consisting of either primed ∅X174 ssDNA or a plasmid-based D-loop. Analysis by electrophoretic mobility shift assay (EMSA) indicated that the migration of both the probes was retarded by Sgs11−652 (Figure 1B). Surprisingly, further characterization showed that the binding of Sgs11−652 to primed ∅X174 ssDNA was sensitive to unprimed plasmid-length ssDNA competitor (Figure 1C). This suggested that Sgs11−652 does bind ssDNA and that it might bind oligonucleotide substrates if they were unusually long. To test this hypothesis, we incubated Sgs11−652 with oligonucleotides of 60, 90 or 174 nt. As predicted, Sgs11−652 efficiently bound d(T)174, but bound the 90- and 60-mer oligos progressively less well (Figure 1D). We note that Sgs11−652 binds d(T)174 with high affinity because it could be detected at nM concentrations of both protein and substrate. As the D-loop is unlikely to contain significant ssDNA character, its binding by Sgs11−652 may involve different structural determinants. To further localize this activity, we expressed and purified sub-domains of Sgs11−652 as GST-fusion proteins and assayed them for ssDNA binding (Figure 2A). On the basis of these assays we determined that the C-terminal half of Sgs11−652 was dispensible for binding, and that Sgs1103−322 was the minimal region required for activity (Figure 2B). To confirm this result, and show that ssDNA binding could be detected by methods other than EMSA, we performed nitrocellulose filter-binding assays with His6-tagged proteins. These assays confirmed that ssDNA-binding activity could be detected in both Sgs11−322 and Sgs1103−322 (Figure 2C). Figure 2.The ssDNA-binding activity of Sgs11−652 maps to a sub-domain and is conserved in human and Drosophila BLM orthologs. (A) The following GST–Sgs1 fusion proteins were subjected to EMSA assay as in Figure 1D using 32P-labelled poly(dT)174 as probe: Sgs11−158, Sgs11−322, Sgs11−652 and Sgs1322−652 at 18, 37, 75, 150 and 300 nM; Sgs150−322, Sgs1103−322 and Sgs1158−484 at 100, 200, 400 and 800 nM; and Sgs1103−250 at 150, 300, 600 and 1200 nM. Dashes (−) indicate no-protein control lanes. (B) Summary of ssDNA binding and SA results. Symbols in the ssDNA-binding column represent the following results: +, strong; +/−, weak; and −, no ssDNA-binding activity. Symbols in the SA column represent the following results: +, strong; +/−, weak; and −, no SA activity. (C) The indicated His6-tagged proteins were assayed for ssDNA-binding using a nitrocellulose filter-binding assay. Reactions were performed as in Figure 1D, but were analysed by filtering through alkalai-treated nitrocellulose and quantifying the bound products by scintillation counting. The data are presented as a percentage of input CPM. (D) The N-termini of Sgs1, hsBLM and dmBlm are presented schematically with putative domain boundaries indicated by dotted lines. SE, strand-exchange domain. (E) EMSA assays were performed as above using GST–hsBLM1−294 at 0, 100, 200, 400 and 800 nM, and dmBlm1−380 at 0, 140, 280, 560 and 840 nM. Download figure Download PowerPoint To determine whether these results generalized to other BLM orthologs, we assayed comparable regions of human and Drosophila BLM for ssDNA binding. Pairwise amino acid sequence alignments were of limited usefulness in identifying homologous regions in these orthologs. However, vertebrate BLM orthologs contain a conserved 40 aa region of unknown function, BDHCT (InterPro: IPR012532; hsBLM372−411) that showed weak similarity to the fly and yeast orthologs in multiple sequence alignments (Supplementary Figure S1B). Using this alignment we chose to express hsBLM1−294 and dmBLM1−380 as approximations of Sgs11−322 and Sgs11−386, respectively (Figure 2D). These domains were then purified as GST-fusion proteins for use in an EMSA assay. As shown in Figure 2E, titrations of both metazoan proteins resulted in a mobility shift of the d(T)174 probe. When the reaction products were incubated with an antibody to GST before electrophoresis, the resulting signals were further retarded indicating that the GST–Sgs1 and GST–hsBLM fusion proteins are responsible for this activity (Supplementary Figure S2). The GST portions of the proteins did not contribute to this activity as hexahistidine-tagged versions of all three proteins bound ssDNA (Supplementary Figure S3). Characterization of a strand annealing activity We assayed GST-tagged Sgs1 proteins for SA activity by incubating them with two partially homologous oligos one of which was 32P-labelled. Compared with mock treatment, Sgs1 proteins that included residues 103–322 accelerated the rate of strand pairing (Figure 3A, upper). By assaying a variety of sub-domains we observed a correlation between ssDNA binding and SA activity (Figure 2B). Sgs1103−322 is the minimal domain required for this activity and, for reasons described below, we hereafter refer to it as the SE domain. SA activity was conserved in the human and fly domains as His6-tagged versions of all three proteins accelerated strand pairing (Figure 3A, lower). The His6-tagged proteins, which showed SA activity at concentrations as low as 5 nM, were judged to be superior to the GST-tagged versions presumably because of the smaller size of the epitope tag. Figure 3.The SE domains from BLM/Sgs1 orthologs show strand annealing activity. (A) SA assays contained the indicated concentrations of GST- (upper) or His6-tagged (lower) proteins plus 1 nM each of a 32P-labelled 50 nt oligo (#1) and an unlabelled 50 nt oligo (#2) that share 25 nt of perfect complementarity. The reactions were incubated at 37°C for 5 min under standard conditions as described in 'Materials and methods'. Reactions were stopped and the products were resolved by 10% PAGE followed by phosphorimager analysis. M is a mock reaction lacking protein. (B) GST–Sgs11−322 (50 nM) or Sgs1103−322 (20 nM) were assayed as in (A) except that the reactions were stopped at the indicated times before analysis. (C) The indicated SE domain proteins (50 nM) were assayed as in (A) except that the reactions contained either no competitor (−) or a 10-, 100- or 1000-fold excess of oligo #16 before the addition of proteins. (D) The indicated SE domain proteins (50 nM) were assayed as in (A) except that the reaction contained 1 mM Mg2+ and either no additions (−) or 1 mM of the indicated cofactor. Throughout, all proteins are His6-tagged unless indicated as GST-tagged. Asterisks represent positions of 32P-labelling. Download figure Download PowerPoint Further characterization of the SA activity indicated that it is rapid. In contrast to spontaneous annealing, which was just detectable at 20 min, the enzyme-catalysed reaction was complete within 1 min (Figure 3B). The effect of non-homologous competitor ssDNA was tested by including high concentrations of an unrelated oligo in the reaction. As shown in Figure 3C, SA was resistant to a 100-fold excess of competitor, whereas a 1000-fold excess resulted in inhibition. Thus, high levels of non-homologous ssDNA are required to inhibit SA activity. We next examined the role that cofactors may have in SA. As shown in Figure 3D, the SA activities of all three orthologous SE domains were unaffected by ATP, ADP or the non-hydrolyzable analogue AMPPNP. Thus, the BLM/Sgs1 SA activity identified here behaves differently than activities identified in the full-length protein, which are presumably dependent on the RecQ helicase domain. The SE domain shows DNA SE activity The SE domain was tested for the ability to catalyse SE between a duplex DNA substrate containing one labelled strand and a complementary ssDNA oligo. A similar ATP-dependent or ATP-stimulated reaction has been observed with multiple RecQ homologs (Machwe et al, 2005; Weinert and Rio, 2007; Xu and Liu, 2009). One version of this reaction uses an excess of recipient ssDNA that has the same sense as the duplex's labelled strand. Denaturation of the fork is expected to result in annealing of the unlabelled complementary strands and release of the free 32P-labelled ssDNA oligo. This reaction is essentially unidirectional as there is little chance of the duplex's unlabelled strand exchanging back to the less abundant labelled strand. Therefore, our substrates consisted of a synthetic forked donor DNA with a radiolabelled top strand plus a five-fold molar excess of unlabelled top strand as recipient. As shown in Figure 4A, SE domains from all three species promoted the exchange reaction. SE was catalysed by SE protein, and did not result from spontaneous denaturation of the duplex, because neither incubation with non-specific protein (GST), nor excess complementary DNA alone, resulted in SE (Figure 4A). To eliminate the possibility that the donor DNA was simply melted by SE after which it passively annealed to the recipient oligo during the protease step before electrophoresis, we included a high concentration of a second recipient oligo during the protease incubation. The failure to detect annealing to this larger 94 nt oligo during the protease reaction indicates that ssDNA was not present after the assay or during protease treatment (Supplementary Figure S4A). Moreover, the SE protein lacked detectable nuclease activity that might result in an artefactual DNA SE activity (Supplementary Figure S4B). Thus, the simplest explanation for SE is that the SE domain melts double-stranded DNA while it simultaneously anneals complementary DNA strands. Such a coordinated reaction might explain why higher protein concentrations were required for SE (200–400 nM) than for SA (5–20 nM). Figure 4.The SE domains from BLM/Sgs1 orthologs show DNA SE activity. (A) The SE assay is illustrated at the top of the panel. Reactions contained the indicated SE domain proteins at 0 (−), 50, 200 or 400 nM plus 2 nM forked DNA (where oligo #1 is 32P-labelled) plus 10 nM oligo #1. Substrate DNAs were incubated with GST at 50, 200 or 800 nM as negative control. The reactions were incubated at 37°C for 30 min under standard conditions and the products were analysed by 8% PAGE and phosphorimaging. The first lane (Δ) contains 32P-labelled oligo #1 as marker. (B) Sgs1103−322 (0, 100, 200, 400, 800 and 1600 nM) was used in an SE assay using blunt-ended substrate as indicated in the reaction at the top of the panel. Reactions contained 1 nM duplex DNA plus the indicated amounts of oligo 3. Assays were performed as in (A) except for use of 10% PAGE. (C) Sgs1103−322 was assayed using 5′-tail duplex DNA as substrate as in (B). (D) Sgs1103−322 was assayed using 3′-tail duplex DNA as substrate as in (B). (E) The SE reactions shown in (B–D) (20 nM unlabelled oligo) were quantified and are presented as a function of protein concentration. Sequences of the indicated oligos are presented in Supplementary Table 1. Throughout, all proteins are His6-tagged. Asterisks represent positions of 32P-labelling. Download figure Download PowerPoint The substrate requirements for SE were examined by preparing radiolabelled duplex substrates whose ends were either flushed or contained a free 5′-tail or 3′-tail. Sgs1103−322 was then titrated into the reactions that contained different concentrations of unlabelled recipient DNA. In all cases, SE required a five-fold excess of recipient DNA (Figure 4B–D). SE also took place using the blunt donor; however, lower levels of protein were required when the donor duplex contained a 3′-tail (Figure 4E). The stimulation by a 3′-tail suggests that unwinding and annealing has a specific polarity. To further characterize the Sgs1103−322 protein, we used an SE reaction in which the recipient DNA was complementary to the labelled strand of the duplex but larger in size. Thus, the appearance of a retarded signal in native gel electrophoresis is diagnostic for SE. Using these substrates, the SE domains from three BLM orthologs efficiently converted the donor DNA signal into a slower-migrating form (Figure 5A). Time-course experiments confirmed that the three orthologs had similar kinetics and that the reactions were essentially complete within 5 min (Figure 5B). Figure 5.Characterization of SA and SE reactions. (A) The indicated SE reaction was performed by titrating Sgs1103−322, hsBLM1−294 or dmBLM1−380 into the standard SE assay. After incubation at 37°C for 30 min, the products were analysed as in Figure 4. Δ, boiled substrate; M, mock reaction without protein. (B) Time courses of the SE reaction illustrated in (A) were carried out with Sgs1103−322 (2.4 μM), hsBLM1−294 (1.2 μM) or dmBLM1−380 (1.2 μM). A, annealed oligos were obtained by slow cooling and used as marker. (C) The indicated SA reaction was performed by incubating 2 nM each of oligo #6 (32 nt) and oligo 11 (94 nt) together with various concentrations of either E. coli SSB or yeast RPA. Reactions were assembled on ice before incubation at 37°C for 5 min. (D) An SE reaction was performed using the indicated duplex DNA (0.5 nM) as donor and a 94-nt ssDNA (2.5 nM) as recipient. Reactions were assembled on ice before incubation at 37°C for 30 min. Throughout, all proteins are His6-tagged. Asterisks represent positions of 32P-labelling. Download figure Download PowerPoint The impact of ssDNA-binding proteins on SA and SE activities was then assessed. The annealing of complementary 32- and 94-nt oligos (2 nM each) was inhibited by 8 nM Escherichia coli SSB and 16 nM yeast RPA (Figure 5C). Under these conditions, we estimate that both ssDNA-binding proteins occlude 30 nt of ssDNA, so that there are 8 nM of binding sites in the substrate. Thus, access of Sgs1103−322 to ssDNA appears to be blocked by excess concentrations of RPA and SSB. The corresponding substrates were then assayed in an analogous SE reaction: a 32 bp duplex (0.5 nM) and 94-nt recipient oligo (2.5 nM). E. coli SSB again inhibited the reaction at 8 nM, which is expected to saturate the 7.5 nM binding sites. Higher levels of RPA partially inhibited the SE reaction (Figure 5D). Thus, although there are quantitative differences, Sgs1103−322-promoted SE is inhibited by high levels of both ssDNA-binding proteins. Two additional experiments were performed to characterize the SE reaction. First, because the above experiments were performed in the presence of EDTA, we tested whether it was influenced by divalent cations. The results (Supplementary Figure S5A and B) indicated that the SE reaction was unaltered by physiological levels of Mg2+. Second, to examine its stoichiometry we titrated Sgs1103−322 into an SE reaction and quantified the products. The results indicated that 40–54% of the flush-end DNA duplex was exchanged onto the recipient ssDNA over a range of 0.4–1.6 μM Sgs1103−322 (Supplementary Figure S5C). On the basis of these values, SE requires a minimum of one molecule of protein for each 7 nts of ssDNA. The SE domain is required for Sgs1 function in vivo To determine the in vivo function of the SE domain, we constructed the sgs1-ΔSE allele that lacks the SE coding region. This allele, which expresses Sgs1Δ103−322 from its own promoter, was tested for its ability to complement sgs1Δ phenotypes. To eliminate the possibility that sgs1-ΔSE phenotypes were due to a defect in Top3–Rmi1 binding, we first assayed the interaction by immunoprecipitating (IP'ing) Sgs1Δ103−322 and immunoblotting for Top3 and Rmi1. As demonstrated earlier for WT Sgs1 (Mullen et al, 2005), epitope-tagged versions of Top3 and Rmi1 were present in precipitates of FLAG-tagged Sgs1Δ103−322, and Sgs1Δ103−322 was found in Rmi1 precipitates (Figure 6A). In a side-by-side comparison, approximately equal amounts of the Top3 and Rmi1 subunits were co-IP'd with Sgs1-WT and Sgs1Δ103−322, respectively (Figure 6B). On the basis of these results and the data presented below, we conclude that Sgs1Δ103−322 interacts properly with Top3 and Rmi1. Figure 6.Sgs1Δ103−322 physically interacts with Top3–Rmi1. (A) Yeast strains were constructed to express integrated versions of Top3-V5, Rmi1-HA, and either Sgs1Δ103−322-FLAG (ΔSE) or no Sgs1 (−) as the sole copies of these subunits. Cell extracts were prepared and either immunoblotted directly (extract) or subjected to IP with α-FLAG beads (IP-FLAG) before immunoblotting with α-FLAG, α-V5 and α-HA antibodies. Similar to WT Sgs1-FLAG, Sgs1Δ103−322-FLAG is insufficiently abundant to be detected in crude cell extracts (Mullen et al, 2005). In the lower panel, extracts were IP'd with α-HA and immunoblotted with α-FLAG to detect Sgs1Δ103−322-FLAG. (B) Cell extracts were prepared from strains expressing Top3-V5, Rmi1-HA and either Sgs1-FLAG (WT) or Sgs1Δ103−322-FLAG (ΔSE) as above. Extracts containing 2 mg of total protein were subjected to IP with α-FLAG beads (IP-FLAG) and immunoblotted with α-FLAG, α-V5 and α-HA antibodies. (C) Cells of the indicated genotype were grown in liquid YPD at 30°C and doubling times were determined. Shown are the average values±s.d. (D) Cells of the indicated genotype were resuspended at OD=3, serially diluted in three-fold increments and approximately 5 μl were spotted onto YPD plates with or without 0.03% MMS. Plates were photographed after 2 (YPD) or 3 days (MMS) growth at 30°C. Download figure Download PowerPoint Different alleles of SGS1 have been shown to confer distinct slow-growth phenotypes (Weinstein and Rothstein, 2008). For example, the sgs1Δ null strain grows more slowly than WT, but a strain lacking the Top3–Rmi1-binding domain of Sgs1 (e.g. sgs1-ΔN158 encoding Sgs1159−1447) grows even more slowly than sgs1Δ (Figure 6C). As previously observed (Mullen et al, 2000), this hypermorphic phenotype was eliminated by a larger N-terminal truncation (e.g. sgs1-ΔN322 encoding Sgs1323−1447). In contrast to these mutants, the doubling time of the sgs1-ΔSE strain was identical to WT (Figure 6C). Furthermore, when we examined its ability to complement the MMS sensitivity of sgs1Δ, the sgs1-ΔSE allele conferred a WT level of resistance (Figure 6D). This result is consistent with other internal deletions made within the Sgs1 N-terminus (Ui et al, 2001) and indicates that the sgs1-ΔSE allele functions similar to WT in these assays. The sgs1-ΔSE allele was then used to test whether it would complement the sgs1Δ hyper-recombination phenotype. Intrachromosomal recombination was measured using a marker-excision assay in which CAN1 and URA3 are inserted between direct-repeat (DR) sequences at LYS2 and the rDNA, respectively (Mullen et al, 2000). Compared with WT, the sgsΔ null strain showed a 2.9-fold increase in recombination frequency at LYS2 and a 4.2-fold increase at the rDNA (Figure 7A), and consistent with its hypermorphic phenotype, sgs1-ΔN158 showed 7- and 32-fold increases at these loci. The sgs1-ΔSE allele generated recombination rates that were elevated by 3.8- and 4.2-fold, respectively. These levels closely match those obtained with sgs1Δ and sgs1-ΔN322. Thus, the SE domain is required to suppress hyper-recombination. Figure 7.The SE domain is required for multiple SGS1 functions. (A) Yeast strains of the indicated genotype were assayed for excision recombination at the LYS2 and rDNA loci as described earlier (Mullen et al, 2000). Recombination frequencies were determined and are presented as fold increase over WT. (B) Strain NJY728 (sgs1Δ top3Δ plus pJM555 (TOP3/URA3/ADE3/CEN)) was transformed with the indicated SGS1 alleles in pRS415 (LEU2/CEN). Transformants were streak purified on SD-leu plates, resuspended to OD600=3.0 and serially diluted in five-fold increments. Approximately 5 μl were spott
Referência(s)