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Previous Article | Next Article 
Molecular and Cellular Biology, April 2003, p. 2999-3007, Vol. 23, No. 8
0270-7306/03/$08.00+0 DOI: 10.1128/MCB.23.8.2999-3007.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.
Department of Biological Sciences, Lehigh University, Bethlehem, Pennsylvania 18015
Received 19 November 2002/ Returned for modification 3 January 2003/ Accepted 13 January 2003
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ABSTRACT |
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INTRODUCTION |
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Three classes of proteins are required for sister chromatid cohesion and proper chromosome segregation. Structural cohesion proteins (or cohesins) provide the glue that maintains sister chromatid pairing from G1/S to anaphase onset. In budding yeast, the cohesins include Smc1p, Smc3p, Mcd1p/Scc1p, Scc3p/Irr1p, and Pds5p (16, 19, 28, 35, 42, 52, 57). Recent data revealed that the structural cohesins form a ring. This ring structure is thought to hold sister chromatids together, but whether a single ring encircles both sisters or catanated rings encompass individual sisters is unknown (2, 17). Deposition factors load structural cohesin proteins onto chromatin. Deposition factors include Scc2p (Mis4p in Schizosaccharomyces pombe) and Scc4p, which combine to form a deposition complex separate from the cohesin complex. Scc2p and Scc4p are active throughout a large part of the cell cycle, but are required during S phase (4, 12, 57). Establishment factors appear to couple sister chromatid cohesion to DNA replication, but are not required for DNA replication per se (51, 57).
Analyses of CTF7/ECO1 mutant cells revealed that Ctf7p/Eco1p (herein called Eco1p) is essential for cohesion establishment and acts in a pathway unique from the structural and deposition cohesion factors. First, Eco1p is required during S phase when cohesion is established but not in mitosis when cohesion is maintained. Second, structural cohesins appear to form a complex and load normally in eco1 mutant cells. These findings indicate that Eco1p does not function in cohesin assembly, deposition, or cohesion maintenance (51, 57; R. V. Skibbens and D. Koshland, unpublished results). Eso1p, the fission yeast homolog of Eco1p, also functions in sister chromatid cohesion, revealing that cohesion establishment is conserved through evolution (56). Recent findings reveal that Eco1p provides acetyltransferase activity and that, at least in vitro, the structural cohesins Mcd1p/Scc1p, Scc3p/Irr1p, and Pds5p—as well as Eco1p itself—are acetylation targets. Currently, however, physiologically relevant substrates of Eco1p acetylation have yet to be documented (21). Thus, the molecular mechanism by which cohesion is established remains unknown.
Early studies temporally correlated cohesion establishment with the S-phase portion of the cell cycle (15, 46, 58). The first evidence that cohesion establishment and DNA replication may be intimately coupled was obtained by the findings that Eco1p is required only during S phase and that ECO1 genetically interacts with both POL30 (PCNA) and CHL12/CTF18 (51). PCNA is a homotrimeric sliding clamp that locks DNA polymerase onto double-stranded DNA (dsDNA) and promotes processive DNA replication. Chl12p/Ctf18p (herein termed Ctf18p) exhibits limited homology to Rfc1p. Rfc1p is a large subunit of the replication factor C (RFC) complex, which also contains Rfc2p to Rfc5p. This RFC complex loads PCNA onto dsDNA (22, 26). The interaction between an essential cohesion establishment factor, Eco1p, and two DNA replication factors of interdependent function, PCNA and an RFC homolog, suggested a model in which Eco1p acts to pair nascent sister chromatids as they emerge from the DNA replication fork (51). A link between DNA replication and sister chromatid cohesion was confirmed by the characterization of Trf4p. Trf4p (also called Pol 
and later renamed Pol 
) is a DNA polymerase that also functions in sister chromatid cohesion (3, 61).
Further evidence has implicated a subset of RFC factors as important in sister chromatid cohesion. Currently, there are three known RFC complexes. Rfc1p associates with Rfc2p to Rfc5p to load PCNA onto DNA and thus promote processive DNA replication (22, 40). Rad24p associates with Rfc2p to Rfc5p to load the heterotrimeric Mec3p, Rad17p, and Ddc1p sliding clamp during activation of the DNA damage checkpoint mechanism (14, 22, 24, 30, 32, 37, 41). Ctf18p and two other cohesion factors, Ctf8p and Dcc1p, comprise an RFC complex that contains Rfc2p to Rfc5p, but the identity of an associated sliding clamp remains unknown. This Ctf18p-based RFC complex plays a role in cohesion but not DNA replication (5, 18, 33). Biochemical analyses revealed that Rfc1p, Ctf18p, and Rad24p all associate with Rfc2p to Rfc5p but not with each other, revealing the formation of three independent RFC complexes (33, 36). Characterization of both RFC complexes and DNA polymerases has led to a model in which DNA polymerases and RFC complexes switch in or out, depending on the DNA sites encountered at the replication fork (3, 5, 50, 60-62).
Sister chromatid cohesion is clearly fundamental to proper chromosome segregation. Thus, it is surprising that Rad24p, Ctf18p (and associated Ctf8p and Dcc1p), and Trf4p are nonessential for cell viability. Based on this observation, we inferred that these proteins perform an essential but redundant activity in cohesion establishment. For instance, while the three RFC complexes appear biochemically distinct, they are partially redundant for DNA repair and checkpoint functions (34, 36, 39, 45, 47, 54). In contrast, Eco1p is essential, raising the possibility that redundant cohesion activities may ultimately converge through a single Eco1p-dependent pathway. Here, we report that Eco1p interacts with all three alternate RFC complexes. In addition, we provide new evidence that an RFC subunit is required for sister chromatid cohesion.
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MATERIALS AND METHODS |
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lethality, produces a plasmid-dependent band of the appropriate molecular weight, and directs for elevated expression levels of Eco1p (51). YBS5 was transformed with pBS9, and pRS316-ECO1 was subsequently removed by counterselection with medium containing 5-fluoroorotic acid (5-FOA) (1). Eco1p sedimentation was determined for log-phase cells and cells synchronized in early S phase (hydroxyurea) or in mitosis (nocodazole). The experimentally treated cells were lysed by mechanical disruption with glass beads (100 mM KCl, 1 mM MgCl2, 0.1 mM CaCl2, 20 mM Tris [pH 7.6], 50 mM sucrose, 10% glycerol plus protease inhibitors), placed over a continuous sucrose gradient (10 to 40% sucrose in lysis buffer), and centrifuged with a Beckman SW28 rotor at 25,000 rpm for 24 h at 4°C. Approximately 1.2-ml fractions were harvested and trichloroacetic acid (TCA) precipitated, and the fractions containing Eco1-HAp were identified by Western blot analysis with 12CA5 monoclonal antibody directed against the HA epitope (Covance) and horseradish peroxidase (HRP)-conjugated goat anti-mouse antibody (Cappell). Molecular weight markers included Blue dextran 2000, thryoglobulin, ferritin, catalase, aldolase, and bovine serum albumin (BSA; Pharmacia). GST pull-down assay. To generate GST-Eco1p, the entire ECO1 open reading frame was digested from pBS6 (XhoI-SacI) and inserted in-frame behind glutathione S-transferase (GST) of pGEX4T-3 digested with XhoI-NotI. SacI and NotI were filled in to produce ligatable blunt ends. GST-ECO1 expression in Escherichia coli cells was induced with 2 µM isopropyl-ß-D-thiogalactopyranoside (IPTG) (Sigma) for 2 h at 37°C, and the cells were lysed by sonication. The whole-cell extract was centrifuged at 9,500 rpm for 5 min (Beckman JA-20), and the soluble and insoluble fractions were harvested. Western blot analysis revealed a plasmid-dependent band of the appropriate molecular weight in the soluble fraction visualized with a monocolonal antibody directed against the GST epitope (Santa Cruz) and HRP-conjugated goat anti-mouse antibody (Bio-Rad). E. coli cells harboring either pGEX4T-3 or pGEX4T-3-ECO1 were induced for 2 h with IPTG and lysed by sonication. GST versus GST-Eco1p proteins in bacterial extracts were then coupled to glutathione Sepharose 4B beads (Amersham-Pharmacia). Prior to incubation with yeast extracts, the bead matrices were washed several times in 1x phosphate-buffered saline (PBS).
Yeast strains expressing candidate epitope-tagged proteins were first spheroplasted in 100T Zymolyase (Seikagaku), lysed by swelling and mechanical disruption (20 mM HEPES-HCl [pH 7.5], 5 mM MgCl2 plus protease inhibitors), and centrifuged at 9,500 rpm for 45 min (Beckman JA-20). The supernatant was removed, and the insoluble chromatin pellet was extracted with lysis buffer containing 1 M NaCl before recentrifugation. The salt-extracted supernatants were then harvested and divided into four equal aliquots, one of which was precipitated with TCA and then resuspended in Laemmli buffer. The other three aliquots were each diluted 10-fold in lysis buffer prior to incubation with one of the three bead matrices (glutathione Sepharose beads, or beads coupled to GST or GST-Eco1p). The treated beads were washed several times before eluting the specifically bound proteins by using reduced glutathione (Sigma). To test for DNA-based interactions, the pull-down buffer was supplemented to 10 mM MnCl2 and 500 ng of DNase I per ml, incubated for 1 h at 4°C, and then processed as described above. During Western blot analyses for bead-bound proteins, the presence of FLAG-tagged proteins was detected with a monoclonal anti-FLAG antibody, M2 (Sigma); HA-tagged proteins were detected with the 12CA5 monoclonal antibody (Babco); and MYC-tagged proteins were detected with the anti-cMYC 9E10 monoclonal antibody (Santa Cruz). For each type of detection, HRP-conjugated goat anti-mouse (Bio-Rad) antibody and ECL-Plus (Amersham-Pharmacia) were used for visualization.
Sister chromatid cohesion analyses. The strain YPH1477 (33), which contained Tet operator repeats (TetO) integrated proximal to the centromere and which expresses both green fluorescent protein (GFP)-labeled Tet repressor (TetR-GFP) and Pds1-13MYCp, was backcrossed four times into the S288C background to produce YBS1042. YBS1042 was then crossed with a mutant strain (TSY601) harboring the rfc5-1 allele (47) and sporulated. Spores YBS1058 and YBS1059 containing the rfc5-1 allele were identified by growth on selective media, visualization of loci via GFP, and by temperature sensitivity. Spore YBS1060 containing the wild-type RFC5 gene was identified by growth on selective media, visualization of loci via GFP, and growth at 37°C. To assay for a defect in cohesion, log-phase cells were placed into fresh medium containing 20 µg of nocodazole per ml and maintained at 37°C for 3 h. An optional step of synchronizing log-phase cells in early S phase by growth in 0.2 M hydroxyurea for 3 h at 23°C was also performed prior to incubating the cells at the permissive temperature in medium containing nocodazole. Digital images were captured on a Nikon Eclipse E800 microscope with a Coolsnapfx charge-coupled device camera (Photometrics) and IPLab software, 3.5.3 (Scanalytics). Flow cytometry (BD FACScan) and Western blot analysis were performed as described previously with minor modifications (13). For localization of Pds1-13MYC, cells arrested in mitosis were fixed with formaldehyde (3.7%) for 10 to 15 min, washed, and prepared for immunofluorescence as previously described (6). The MYC tag was visualized with the mouse monoclonal antibody 9E10 (Santa Cruz, Biotechnology) in combination with goat anti-mouse tetramethyl rhodamine isocyanate (TRITC)-labeled (Cappell) antibodies.
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RESULTS |
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null strains, indicating that the GST moiety did not adversely affect the essential function of Eco1p in vivo (data not shown).
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Eco1p associates with four small RFC subunits. Recent findings indicate that Ctf18p associates with Rfc2p, Rfc3p, Rfc4p, and Rfc5p (18, 33). To test whether Eco1p also associates with these RFC subunits, extracts of log-phase yeast cells harboring FLAG-tagged Rfc2p, Rfc3p, Rfc4p, or Rfc5p were incubated with either beads alone, beads linked to bacterially expressed GST, or beads linked to bacterially expressed GST-Eco1p. The three matrices were then washed, and the bound proteins were eluted. Western blot analyses revealed that Rfc2p, Rfc3p, Rfc4p, and Rfc5p all bound specifically to Eco1p but did not bind to GST-linked beads or beads alone (Fig. 2). Rfc3p consistently yielded the highest binding efficiency. To test the possibility that the FLAG tag was responsible for binding the RFC subunits to Eco1p, we also tested the ability of FLAG-tagged bacterial alkaline phosphatase to bind beads linked to either bacterially expressed GST or GST-Eco1p. Western blot analyses failed to reveal an interaction between Eco1p and bacterial alkaline phosphatase (BAP)-FLAGp, indicating that the FLAG tag did not participate in the protein interactions observed for Eco1p and Rfc2p-Rfc5p (Fig. 2). These findings reveal a new physical interaction between Eco1p and an RFC-based complex containing Ctf18p.
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DNA. The DNA was completely digested (Fig. 3D), attesting to the efficiency of the enzymatic treatment under these conditions. These results indicate that the binding of Eco1p to RFC complexes occurs independent of DNA.
Eco1p assembled with RFCs in vivo cosediment as a complex.
Recently, Ctf18p assembled in vivo with Rfc2p to Rfc5p was shown to sediment as a complex of approximately 12S. This sedimentation is completely coincident with Rfc1p-RFC sedimentation and overlaps that of Rad24p-RFC sedimentation (36). We thus decided to test whether Eco1p would cosediment with RFC complexes assembled in vivo by using Ctf18p as a fiduciary marker. To facilitate detection of Eco1p, we generated extracts from yeast cells in which Eco1-HAp expressed at elevated levels was the sole source of Eco1p function. This Eco1-HAp construct is fully functional and maintains viability of eco1 cells at wild-type growth rates (data not shown). Extracts from hydroxyurea-arrested cells expressing Eco1-HAp were placed over a sucrose gradient (10 to 40%) and subjected to centrifugation. Fractions were then harvested, and the sedimentation of Eco1p was determined by Western blot analyses. Consistent with an in vivo interaction, a significant fraction of Eco1p sedimented at approximately 12S (Fig. 4). To independently test for Ctf18p sedimentation, the membrane was stripped and reprobed with antibodies directed against endogenous Ctf18p. The results show that Ctf18p cosedimented exactly with Eco1p as a 12S complex when assembled in vivo (Fig. 4), recapitulating previously described Ctf18p sedimentation (36). A very slight decrease in sedimentation was observed for extracts derived from logarithmically growing cells (Fig. 4), which was still consistent with previously characterized sedimentations for all three RFC complexes (36). The finding that Eco1p assembled in vivo cosediments with RFC subunits provides strong support for the Eco1p-RFC associations detected in vitro.
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12S but not as a much larger complex (36). Thus, we decided to use this aberrant Eco1p sedimentation (presumably the result of overexpression) to ask whether Ctf18p would be pulled deeper into the gradient by virtue of its association with Eco1p. Indeed, Ctf18p persisted in cosedimenting with Eco1p deep in the sucrose gradient (Fig. 4). These observations suggest that Eco1p and Ctf18p not only physically associate in vivo but that this binding is of sufficient avidity to alter Ctf18p sedimentation.
Rfc5p is required for sister chromatid cohesion.
Given the physical association of Eco1p with RFC complexes (this study), a likely model was that all RFC subunits would play a key role in cohesion establishment. Both ctf7 and rfc5 temperature-sensitive mutant strains are rescued by elevated levels of POL30 (51, 54). Thus, we decided to first test whether Rfc5p—a component of each RFC complex—functioned in cohesion. The rfc5-1 allele was crossed into a strain that contains Tet operator repeats (TetO) integrated proximal to the centromere. This strain also expresses GFP-tagged Tet repressor protein (TetR-GFP), allowing for visualization of the centromere-proximal locus. Visualization of the GFP signal was then used to determine the position of one sister chromatid relative to the other (33, 35). Log-phase wild-type and rfc5-1 cells were shifted to 37°C for 3 h (to inactivate rfc5p function in the mutant strain) in medium supplemented with nocodazole to inhibit the onset of anaphase. An optional synchronization step of arresting cells in early S phase with hydroxyurea was also used. Parallel cell samples were then assayed for DNA content, cell morphology, and sister chromatid cohesion. Wild-type cells treated with nocodazole were predominantly large budded and contained a 2C DNA content, indicative of a mitotic arrest. When GFP-tagged loci were viewed by epifluorescent microscopy, wild-type cells were found to contain tightly paired sister chromatids, such that few (9%) sisters were dissociated. rfc5-1 mutant cells treated with nocodazole also were predominantly large budded and contained a 2C DNA content. In contrast to wild-type cells, however, rfc5-1 mutant cells contained a significant increase in the number of separated sisters (20%) (Fig. 5). Both wild-type and rfc5-1 strains exhibited similarly low levels (5%) of separated sisters in early S phase. These results reveal that the incidence of two GFP spots in mitotic rfc5 mutant cells was not due to aneuploidy present early in the cell cycle but instead was due to a loss of sister chromatid cohesion.
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We next tested whether Rad24p played a role in cohesion establishment. A rad24 deletion strain (ResGen) was crossed into the cohesion assay strain (TetO integration and expressing TetR-GFP and Pds1-MYCp) modified from reference 33. The resulting diploid was sporulated, and haploid cells containing the appropriate markers were identified. To independently assess for loss of Rad24p function, we confirmed that the resulting strains exhibited sensitivity to UV light (data not shown). Wild-type and rad24 mutant cells were grown for 3 h in the presence of nocodazole. As before, the number of GFP spots was then used to determine the position of one sister chromatid, relative to the other, in large-budded cells that retained Pds1p. While a reproducible increase in sister chromatid separation was observed in rad24-null cells relative to wild-type cells, this increase was only minimally above background (data not shown). Independent analyses of rad24 mutant cells revealed a similar variability, such that a cohesion defect was considered not to be significant (M. Mayer and P. Hieter, personal communication). While the extent to which Rad24p functions in cohesion remains unknown, a likely possibility is that Rad24p may perform a key role in cohesion, but that this activity occurs only along very short tracts of DNA (possibly during nucleotide excision repair) and thus was undetected under the conditions tested.
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In this report, we provide new evidence that Eco1p physically associates in complexes comprised of seven other proteins—Rfc1p to Rfc5p, Ctf18p, and Rad24p—all of which are components of the DNA replication/repair fork machinery. First, GST chromatographic methodologies revealed that Eco1p associates with each of these RFC subunits in a DNA-independent manner. In contrast, no interaction was detected with Mcd1p, a chromatin-associated cohesin, suggesting that Eco1p-RFC associations are indeed specific. Second, Eco1p-containing complexes assembled in vivo were found to cosediment exactly with RFC complexes, with Ctf18p used as a fiduciary mark. Previous findings revealed that the Ctf18p- and Rad24p-RFC complex sedimentations are completely coincident with that of Rfc1p-RFC (36)—a likely outcome due to the fact that each complex is in part comprised of common components. In addition, Rad24p sedimentation is completely coincident with Ctf18p (although the peak sedimentation is very slightly decreased in Rad24p) (36). Thus, Ctf18p is an appropriate marker for Ctf18p-Rad24p- and Rfc1p-RFC complexes. We noted a very slight mobility shift of Eco1p when obtained from logarithmically growing cells versus hydroxyurea-treated cells (Fig. 4), but both are within range of all three RFC complex sedimentations. Thus, both in vitro and in vivo results are consistent with a model in which Eco1p associates with each of the three RFC complexes.
The role of Eco1p-RFC associations is further supported by physiological evidence. First, our analyses of rfc5 mutants revealed that Rfc5p, a component of each RFC complex, is required for sister chromatid cohesion. This result greatly extends the previously documented roles for Rfc5p in DNA replication and replication checkpoint activity (8, 11, 37, 47, 53, 54). Do cells harboring mutations in other RFC subunits exhibit cohesion defects? Recent studies showed that budding yeast rfc4 mutant strains and fruit fly larva rfc2 mutants both exhibit sister chromatid cohesion defects (27, 33). Our quantification of cohesion defects in rfc5 mutant cells is nearly identical to that reported for rfc4 mutants (33). Importantly, RFC subunits perform distinct mechanochemical functions even within an RFC complex. For instance, based on similarities to the E. coli clamp loader, it has been postulated that the eukaryotic RFC complex required for the bulk of DNA replication is composed of three subcomplexes: Rfc1p, Rfc2p to Rfc4p, and Rfc5p. All are AAA+ family members but likely serve very different molecular functions corresponding to a motor (Rfc2p to Rfc4p), stator (Rfc5p), and wrench (Rfc1p) (10, 40). Thus, the assignment of a new function for any RFC subunit is nontrivial. Our characterization of Rfc5p as a cohesion factor completes the list in that at least one member of each subcomplex (Ctf18p as a wrench, Rfc2p and Rfc4p as motor components, and Rfc5p as a stator) has now been characterized as playing an important role in cohesion (18, 33; this study). Finally, independent analyses support our data that RFC complexes beyond those including Ctf18p function in cohesion. For instance, cells harboring defects in both Ctf18p and Rad24p are viable (36), indicating that Rfc1p-based RFC complexes, in association with Eco1p, most probably are competent to establish cohesion. Thus far, we have been unable to unambiguously observe cohesion defects in Rad24p-deficient cells, but this may be due to limitations in detecting cohesion loss along short chromosome segments and during DNA repair when Rad24p is active. In combination, these findings provide the basis for a new understanding regarding the mechanical linkage between cohesion establishment and DNA replication or repair.
Previous models of DNA replication fork dynamics proposed DNA polymerase handoff or switching mechanisms (3, 33, 59, 61). In terms of cohesion, a DNA polymerase switch is thought to occur when the DNA polymerase that performs the bulk of DNA replication (Pol 
or 
) encounters a site destined for cohesion establishment. At this juncture, the replication polymerase switches out for a polymerase that functions in cohesion establishment, Pol (previously termed Pol ). In conjunction with the DNA polymerase handoff, a similar switching mechanism has been postulated for RFC complexes such that the Rfc1p-RFC complex switches out for the Ctf18p-RFC complex upon encountering a cohesion site or the Rad24p-RFC complex upon encountering DNA damage (5). Our data greatly extend the current model. We posit that each of the RFC complexes is competent to establish cohesion such that Eco1p rides, in piggyback fashion, the RFC DNA replication machinery. In this way, Eco1p is free to establish cohesion during DNA replication, but not participate in DNA replication (Fig. 6). This new model sheds light on the functional redundancy of RFC complexes in cohesion establishment and helps explain how nonessential factors (Trf4p, Rad24p, Ctf18p, and associated cohesion factors Ctf8p and Dcc1p) can participate in cohesion, a process fundamental to cell survival.
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ACKNOWLEDGMENTS |
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M.A.K. was supported in part by funding from the Vice Provost of Research and Department of Biology, Lehigh University; R.V.S. is supported by the National Science Foundation (MCB-0212323).
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FOOTNOTES |
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