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1 Department of Cell Biology, Institute for Virus Research, and
2 Department of Mammalian Regulatory Network, Graduate School of Biostudies, Kyoto University, Sakyo-ku, Kyoto 606-8507, Japan
3 Department of Bioscience, School of Science and Technology, Kwansei Gakuin University, Sanda, Hyogo 669-1337, Japan
4 Laboratory of Biochemistry II, School of Medicine, Shimane University, Izumo, Shimane 693-8501, Japan
5 Laboratory of Biochemistry and Molecular Biology, National Cancer Institute, National Institutes of Health, Bethesda, MA 20892, USA
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Abstract |
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 Top Abstract IntroductionResultsDiscussionExperimental proceduresNote added in proofReferences |
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Introduction |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresNote added in proofReferences |
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A major determinant of higher order chromatin structure is covalent modification of histone tails (Jenuwein & Allis 2001; Kouzarides 2007). In particular, histone tail methylation is considered to be an epigenetic marker of such structures (Lachner et al. 2003; Martin & Zhang 2005). Specific histone modifications are recognized and bound by various chromatin proteins, processes that lead to appropriate higher order chromatin structure (Kouzarides 2007). For example, methylation of histone H3 lysine 9 (H3K9) defines heterochromatin, and a conserved heterochromatin protein, HP1, recognizes and binds to methylated H3K9 (H3K9me) to form silent chromatin structures (Bannister et al. 2001; Lachner et al. 2001; Nakayama et al. 2001b).
Once established, higher order chromatin structures must be correctly duplicated during chromosome replication, and an important step in this process is nucleosome assembly onto newly replicated DNA. Parental nucleosomes are transiently disrupted during passage of the replication fork, and histone H3/H4 is transferred onto newly synthesized DNA. In addition, newly synthesized histone H3/H4 is assembled onto replicating DNA with the assistance of histone chaperones. Finally, H2A/B dimers are assembled to form intact nucleosomes (Gruss et al. 1993; Gasser et al. 1996). In addition to nucleosome assembly, histone tail modifications and associations with binding proteins must be re-established on replicated nucleosomes. This process appears to be coupled with chromosome replication. Indeed, some mutations affecting replication proteins, including components of the origin recognition complex (ORC), DNA polymerases and PCNA, also confer defects in heterochromatin formation in various organisms, suggesting that replication-associated mechanisms could be operative for heterochromatin maintenance (for review Wallace & Orr-Weaver 2005).
A highly conserved three-subunit complex, chromatin assembly factor I (CAF1), was initially isolated as a histone chaperone that performs replication-coupled nucleosome assembly in vitro (Smith & Stillman 1989). Further analysis indicated that interaction of CAF1 with PCNA, a "sliding-clamp" protein that confers high processivity to replicative DNA polymerases, is a mechanism by which CAF1 is targeted to newly synthesized DNA (Shibahara & Stillman 1999). An in vivo function for CAF1 in replication-coupled nucleosome assembly is indicated by the depletion of the one of its subunits in vertebrate cells. In human cells, knockdown of the largest subunit by the short interfering RNA technique results in an S phase block accompanied by checkpoint activation (Hoek & Stillman 2003), and knockdown of the second subunit causes rapid cell death (Nabatiyan & Krude 2004). Depletion of the largest subunit of CAF1 in chicken cells results in cell cycle arrest in S phase and decreased nucleosome assembly on newly replicated DNA (Takami et al. 2007). The co-localization of CAF1 with replication foci in S phase further supports a function of CAF1 in replication-coupled nucleosome assembly (Krude 1995; Shibahara & Stillman 1999; Taddei et al. 1999).
In addition to chromatin assembly, some evidence suggests a specific role for CAF1 in maintenance of silent chromatin. In several higher eukaryotes, inactivation of CAF1 was shown to cause misexpression of certain silent euchromatic genes or transposons, sometimes resulting in developmental defects (Kaya et al. 2001; Tchenio et al. 2001; Ono et al. 2006; Song et al. 2007). This observation suggests that CAF1 is involved in euchromatic gene regulation through the maintenance of silent states. Involvement of CAF1 in heterochromatin maintenance has also been suggested for mammalian cells, in which CAF1 is localized at replication foci at heterochromatin in late S phase (Krude 1995; Murzina et al. 1999; Quivy et al. 2004). In addition, the largest subunit of CAF1 has been shown to interact with HP1 (Murzina et al. 1999). However, these results are still circumstantial to indicate the direct role of CAF1 in heterochromatin maintenance, and molecular details have not been totally unknown. Specially, inhibition of S phase progression by depletion of CAF1 prevents an analysis of the role of CAF1 in heterochromatin replication, which occurs at the very-end of S phase.
However, in budding yeast, deletion of CAC1/RFL2, which encodes the largest subunit of CAF1, causes moderate defects in heterochromatic silencing at the mating locus and at telomeres, an effect which has been implicated in a defect in heterochromatin maintenance rather than establishment (Monson et al. 1997; Enomoto & Berman 1998). However, budding yeast heterochromatin is different from that of other eukaryotes as Sir proteins replace HP1 as heterochromatin components and because it lacks H3K9 methylation. Thus, the observation in budding yeast might not be directly applicable to HP1-based heterochoromatin in other eukaryotic cells.
Fission yeast provides a good model system for heterochromatin analysis, because like higher eukaryotes it is characterized by H3K9 methylation and by a heterochromatin binding protein, Swi6, a homologue of HP1 (Nakayama et al. 2001b). In this report, we identified fission yeast genes encoding proteins homologous to the three CAF1 subunits and showed that these proteins form a complex that associates with PCNA during S phase. Deletion analysis of each subunit showed moderate defects in heterochromatin at centromeres and the mating locus, accompanied with a decrease of Swi6 in heterochromatin. Importantly, CAF1 deletion enhanced the variegated phenotype of meta-stable heterochromatin, indicating that it functions in heterochromatin maintenance. We found that Swi6 that dissociated from heterochromatin during replication bound to CAF1. Moreover, CAF1 localizes to heterochromatic loci during replication. These data suggest that CAF1 actively participates in heterochromatin maintenance by delivering Swi6/HP1 to replicated heterochromatin.
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Results |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresNote added in proofReferences |
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In attempts to identify ORC-interacting proteins using the yeast two hybrid assay, we isolated a gene (SPBC29A10.03) that showed significant homology to the largest subunit of human and budding yeast CAF1 (CAF1p150 and Cac1, respectively; Fig. 1A) and designated it pcf1 (the first subunit of Schizosaccharomyces pombe CAF1). The biological significance of the interaction between Pcf1 and ORC was not clear at this stage. Pcf1 lacks the N-terminal region of CAFp150, which contains the PEST domain, but retains the highly charged K/E/R-rich and E/D-rich regions (Fig. 1A). The K/E/R-rich region of Pcf1 showed 35.4% and 42.1% identity to that of Cac1 and CAF1p150, respectively, whereas the E/D-rich region showed 32.6% and 30.6% identity to that of Cac1 and CAF1p150, respectively. In addition, the PCNA binding motif [Qxx(I/L/V)xx(F/Y)(F/Y)] (Warbrick 1998) is also conserved within amino acids residues 172–179 [QLKLNNFF] of Pcf1.
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pcf1, pcf2 and pcf3 cells) were all viable, indicating that spCAF1 is not essential for growth.
To determine whether Pcf2 and Pcf3 form complexes with Pcf1, we added the Flag-tag and HA-tag sequences to the C terminus of the endogenous pcf2 and pcf3 genes, respectively. The tagged strains did not show any defects in silencing at centromere and mating locus nor in sporulation observed in the pcf2 or pcf3 strains (see below), indicating that addition of a tag did not affect the function of either gene product. Using doubly-tagged strains, we performed co-immunoprecipitation experiments (Fig. 1B). Both Pcf1 and Pcf3-HA were co-immunoprecipitated with Pcf2-Flag (lanes 2 and 4). In addition, PCNA was detected in immunoprecipitates, indicating that PCNA interacts with the Pcf1-2-3 complex. When pcf1 was deleted, neither Pcf3-HA nor PCNA was co-immunoprecipitated with Pcf2-Flag (lanes 3 and 6). This result is consistent with previous findings that CAF1p150 directly binds to CAF1p60, CAF1p48 and PCNA (Kaufman et al. 1995; Shibahara & Stillman 1999). The level of Pcf3 is significantly reduced in pcf1 cells (Fig. 1B, lane 3), indicating that Pcf3 is de-stabilized in the absence of Pcf1.
In human cells, the third subunit of CAF1 was also identified as a protein that interacts with the Rb protein, named Rbp48/46 (Qian et al. 1993; Qian & Lee 1995); later it was found as a component of other protein assemblies related to chromatin metabolism such as HDAC complexes (Taunton et al. 1996) and the human Mis18 complex (Hayashi et al. 2004; Fujita et al. 2007). Therefore, we asked if Prw1, Mis16 or Pcf3 could form complexes with Pcf1 and Pcf2. We added the HA-tag to the C terminus of the three paralogues and performed co-immunoprecipitation experiments (Fig. 1C). When each of the three HA-tagged proteins was precipitated with the anti-HA antibody, only Pcf3-HA co-precipitated with Pcf1 and PCNA (upper panel), indicating that only Pcf3 forms a complex with these proteins. Consistent with this result, when Pcf2-Myc was precipitated, only Pcf3-HA co-precipitated (lower panel). We could not rule out the possibility that the HA-tagging of Mis16 or Prw1 disturbs the complex formation with Pcf1 and Pcf2. However, the Pcf1-2-3 complex seems to be a major counterpart of CAF1 in other organisms because the strains harboring HA-tagged Mis16 or Prw1 did not show the similar defects observed in pcf1 or pcf2 cells. Thus, hereafter we refer to the Pcf1-2-3 complex as spCAF1.
Phenotypes of spCAF1 deletion
Strains singly disrupted for spCAF1 subunits showed the same morphology and growth rate as wild-type cells. Double disruptants (pcf1pcf2, pcf2pcf3, pcf1pcf3) and the triple disruptant (pcf1pcf2pcf3) also did not show growth defects (data not shown). In chicken and mammalian cells, depletion of CAF1 delays S phase progression and ultimately causes cell death (Hoek & Stillman 2003; Nabatiyan & Krude 2004; Takami et al. 2007). In fission yeast, however, loss of Pcf1 did not affect the progression of S phase, as judged by flow cytometry (data not shown). Furthermore, in contrast to what was seen for budding yeast (Kaufman et al. 1997; Game & Kaufman 1999), pcf1 cells did not show increased sensitivity to genotoxins including UV irradiation, methanesulfonate and hydroxyurea (data not shown).
We noticed that deletion of pcf1 or pcf2 resulted in defects in sporulation. In homothallic (h90) cultures of fission yeast, mating type switches every generation, resulting in a mixture of different mating type cells (Klar et al. 1998). Thus, in sporulation medium, h90 cells mate and undergo meiosis to form spores. In h90
pcf1 and h90
pcf2 cells, sporulation was severely inhibited (Fig. S1A in the Supporting Information). The observed defects were not because of defects in meiosis, because wild type and pcf1 cells harboring the pat1-114 mutation, which induces meiosis at non-permissive temperature even in haploid cells (Iino & Yamamoto 1985), underwent meiosis at the same level (Supporting Information Fig. S1B). We also found by flow cytometry that deletion of spCAF1 subunits did not affect entry into the G0 state induced by nitrogen starvation, which is a prerequisite to mating (Supporting Information Fig. S1C). These data suggested that the cells lacking a subunit of spCAF1 have defects in mating or in mating type switching.
spCaf1 is required for heterochromatin maintenance
A current model suggests that CAF1 is involved in the maintenance of heterochromatin (Maison & Almouzni 2004). Therefore, we analyzed the effects of deletion of spCAF1 genes on heterochromatin (Fig. 2). Fission yeast has heterochromatin at the peri-centromeric and subtelomeric regions, and at the mating locus. We used strains in which ura4 gene was inserted at one of these three heterochromatic regions. The expression of the ura4 is silenced by heterochromatin structure spreading into the inserted locus, resulting in poor growth on uracil-depleted medium and resistance to 5-fluoro-orotic acid (FOA), which is toxic to cells expressing the ura4 gene product. Deletion of one of the spCAF1 subunits resulted in a significant derepression of ura4 at the peri-centromeric region and at the mating locus, although the extent of derepression was lower than that seen for clr4 cells (Fig. 2A,B), in which heterochromatin is completely erased by the absence of the heterochromatin-specific histone methyltransferase Clr4. Each single disruptant showed a similar extent of derepression at both peri-centromeric and mating locus heterochromatin. Furthermore, the triple disruptant showed the same extent of derepression as the single disruptants. These results are consistent with the assumption that Pcf1, Pcf2 and Pcf3 function as a complex, spCAF1. In contrast, deletion of pcf1 did not affect telomeric silencing (Fig. 2C). In addition, pcf1pcf2 cells were derepressed to a similar extent as single disruptants (data not shown). This result suggests that depletion of spCAF1 does not affect telomeric heterochromatin formation.
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clr4 cells (Fig. 2D) (Nakayama et al. 2001b). Deletion of pcf1 resulted in decrease in H3K9 methylation and Swi6: 76 ± 12% and 62 ± 6% of wild type, respectively. In contrast, the hypoacetylated state was maintained in pcf1 disruptants. Similarly, the level of H3K9 methylation and Swi6 at mating locus decreased in pcf1 cells, whereas those at subtelomere was not affected (Fig. 2D). These results indicated that the decrease of Swi6 and H3K9 methylation caused the silencing defects at centromere and mating locus observed in Fig. 2A,B. Consistent with results of the ChIP assay, nuclear Swi6 dots, which correspond to heterochromatic loci (Ekwall et al. 1995), were decreased in number and slightly diffuse in pcf1 cells, as observed by immunofluorescence (Fig. 2D). Notably, the overall levels of Swi6 in cells were not affected by the loss of spCAF1; the relative amounts of Swi6 in pcf1 cells to that in wild-type cells were 1.05 ± 0.14 at G2 cells, 0.96 ± 0.08 at G1/S phase and 0.98 ± 0.12 at early S phase (Fig. 5). These results suggest that the structure of heterochromatin, in particular Swi6 association, is disturbed by the loss of spCAF1, that correlates with a partial disruption of silencing of a marker gene inserted at the centromere and at the mating-type loci.
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::ura4 construct (Grewal & Klar 1996). At the mating locus, there are two heterochromatin nucleation mechanisms, which involves either RNA interference (RNAi) machinery acting through centromere-homologus repeat cenH or DNA-bindng factors bound to silencer DNA elements (Grewal & Jia 2007). In K::ura4 cells, the cenH sequence is replaced with the ura4 gene, which causes variegation of ura4 gene expression (Grewal & Klar 1996) (Fig. 3A). In this strain, two types of epigenetic chromatin states, a ura4-On and a ura4-Off, can be observed; the former represents the transcriptionally active state of ura4 and the latter the heterochromatic state, in terms of Swi6 localization and H3K9 methylation (Nakayama et al. 2000; Hall et al. 2002). Each epigenetic state is meta-stable and converts to the other state at a low frequency (Grewal & Klar 1996) (Fig. 3A). Using this system, we could measure the stability of the heterochromatic state as well as that of the euchromatic state (Fig. 3A, Experimental procedures). We deleted either pcf1 or pcf2 in K::ura4 cells and measured the conversion rates of the ura4-On and ura4-Off states. Deletion of pcf1 and pcf2 resulted in a more than 10-fold increase of the conversion rate from ura4-Off to ura4-On (Fig. 3B, left), indicating that heterochromatin became unstable by the loss of spCAF1 components. Interestingly, the conversion rate from ura4-On to ura4-Off also increased more than threefold in spCAF1 mutants (Fig. 3B, right). This result shows that spCAF1 is also involved in the maintenance of the euchromatic state, although the extent of involvement is lower than for heterochromatin maintenance. The pcf1pcf2 double mutant showed the same phenotype as either single disruptant, supporting the idea that Pcf1 and Pcf2 function in the spCAF1 complex to maintain higher order chromatin structure.
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We then analyzed the cellular localization of each spCAF1 subunit by indirect immunofluorescence (Fig. 4). The tagged proteins Pcf2-Flag and Pcf3-HA showed a nuclear localization that was diminished in the absence of Pcf1 (Fig. 4A). As deletion of pcf1 did not affect the cellular level of Pcf2 (Fig. 1B), this result indicates that the nuclear localization of Pcf2 depends on Pcf1. The loss of the Pcf3-HA signal can be explained, at least in part, by the decrease in the level of Pcf3-HA in pcf1 cells (Fig. 1B). We next analyzed the localization of Flag-tagged Pcf1 expressed from the endogenous pcf1 locus, which was also functional (data not shown), because our polyclonal antibody against Pcf1 was not suitable for immunofluorescence. Approximately 10% of the cells showed a weak but clear nuclear localization whereas others showed no significant signals (Fig. 4B). Cells exhibiting Pcf1 nuclear signals did not correspond to a particular cell cycle stage, as judged by cell morphology (data not shown). As continuous localization of Pcf2 totally depended on Pcf1, we speculated that Pcf1 also localized to nuclei throughout the cell cycle but that association with other proteins partially blocked access of the antibody to the C-terminal Flag epitope.
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The level of Swi6 and H3K9 methylation at centromeric repeats (dg223) decreased in asynchronous pcf1 cells (Fig. 4C), indicating that loss of spCAF1 caused reduction of Swi6 and H3K9 methylation at not only ura4 gene inserted at centromere (otr::ura4, Fig. 2D) but also centromeric repeats. Interestingly, the level of Swi6 at centromeric heterochromatin significantly decreased at the G1/S transition in both wild-type and pcf1 cells (Fig. 4C). We did not see such a decrease in wild-type cells incubated at high temperature or in cells with a cdc25 mutation induced to undergo G2 arrest at high temperature (Fig. S2 in the Supporting Information), indicating that the decrease of Swi6 was not because of high temperature. As the level of Swi6 did not change at the G1/S transition (Fig. 5), the decrease of Swi6 observed in the ChIP assay indicates that Swi6 does not bind to heterochromatin at this stage. We also found by immunofluorescence analysis that Swi6 foci were dispersed in G1/S-arrested cells (Fig. 4F), further supporting the conclusion that Swi6 dissociates at this time. In contrast, the level of dimethylation of H3K9 did not change at any time in either strain (Fig. 4C), indicating de-methylation of H3K9 does not cause dissociation of Swi6 at G1/S phase. In early S phase, Swi6 bound to heterochromatin at the same level as in G2 phase in both wild-type and pcf1 cells, although the amount of Swi6 in pcf1 cells was less than that in wild-type cells as observed in asynchronous cells. This suggests that Swi6 can be recruited by spCAF1-independent pathway(s) in pcf1 cells inefficiently. Furthermore, our analyses suggest that the dynamics of Swi6 localization at subtelomertic regions is different from that at centromeres, as significant levels of Swi6 could be detected at a subtelomeric site during G1/S phase (Fig. 4E). This difference might reflect multiple mechanisms operating to recruit heterochromatin factors at subtelomeric regions (Kanoh et al. 2005). Notably, the level of histone H3 on heterochromatin and euchromatin did not change during the cell cycle and was unaffected by deletion of pcf1 (Fig. 4C–E).
spCAF1 interacts with Swi6 and PCNA in S phase
Mammalian CAF1 was shown to interact with HP1, and this interaction has been implicated in HP1 assembly at heterochromatin during replication (Murzina et al. 1999). Thus, we analyzed the spCAF1–Swi6 interaction by a co-immunoprecipitation assay at various phases of the cell cycle (Fig. 5). Swi6 was co-precipitated with Pcf2-Flag at G1/S and S-phase in pcf1-dependent manner (Fig. 5A). Thus, spCAF1 interacts with Swi6 both at the G1/S transition and in early S-phase but not in G2 phase. In addition, PCNA co-precipitated with Pcf2-Flag in S phase but not at G1/S phase (Fig. 5A). The co-precipitation of PCNA and Swi6 in S-phase cells was much efficient than that in asynchronous cells (Fig 5A. long exposure), suggesting that spCAF1 specifically interacts with PCNA in S phase. When Swi6 was precipitated from extracts prepared from asynchronous cells (mainly G2 phase) (Fig. 5B), Pcf2-Flag was not co-precipitated. But at the G1/S transition and in early S phase, we detected interaction between Pcf2 and Swi6. These interactions depended on Pcf1, as Pcf2 did not co-precipitate with Swi6 in pcf1 cells.
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Discussion |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresNote added in proofReferences |
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::ura4 locus in cells lacking a subunit of spCAF1 (hereafter, we use "caf1 cells" for cells lacking a subunit of spCAF1). Significantly, we observed a small but significant increase in the conversion rate from the ura4-On to ura4-Off state in caf1 cells. This result indicates that silent state can be established in the absence of spCAF1. If spCAF1 had defects in heterochromatin establishment, the On-to-Off rate would had decreased as observed in other mutants involved in heterochromatin establishment (Grewal & Klar 1996). Thus, spCAF1 is not involved in the establishment of silent-state in caf1 cells and the observed instability of ura4-Off state in caf1 cells is caused by the defects in maintenance. Furthermore, the stronger effect of caf1 on the silent state than the active state suggests that spCAF1 has specific role(s) in the maintenance of heterochromatin. spCAF1 co-localized with RP-A at centromeric heterochromatin and ars2004 in early S phase when those loci replicates but not with non-replicating act1 locus and subtelomeric heterochromatin. This suggested that spCAF1 localizes to replication forks. A Pcf1-dependent and S phase-specific interaction between spCAF1 and PCNA (Fig. 5), could be a molecular basis for localization at replication forks, as suggested for mammalian cells (Murzina et al. 1999; Shibahara & Stillman 1999).
We assume that spCAF1 at replication forks contributes to the quick re-formation of heterochromatin on newly replicated chromatin through multiple mechanisms, as shown in Fig. 6. We found that Swi6 dissociates from heterochromatin in G1/S phase (Figs 4, 6A). In mammals, HP1 dissociates from heterochromatin in M phase (Murzina et al. 1999). The decrease of Swi6 levels at heterochromatin during mitosis was also observed in fission yeast, which seems to be induced by phosphorylation of histone H3 serine 10 (Nakayama et al. 2000; Chen et al. 2008). Thus, Swi6 dissociated during mitosis seems not to re-bind to centromeric heterochromatin until G1/S transition. As H3K9me was maintained (Fig. 4C) and phosphorylation of H3S10 was diminished (K.D. and Y.M., unpublished data) in cells arrested at G1/S boundary, some other mechanisms might prevent the binding of Swi6 to heterochromatin. The release of Swi6 from heterochromatin before its replication would make it easier to replicate heterochromatin in early S phase. In contrast, Swi6 does not dissociate from subtelomeric heterochromatin in G1/S and early S phase (Fig. 4E), suggesting that regulation of subtelomeric heterochromatin is different from that of centromeric heterochromatin. At the G1/S boundary, spCAF1 interacts with Swi6 but does not associate with PCNA or heterochromatin. Thus spCAF1 seems to associate with released Swi6 at this time (Fig. 6B). spCAF1 is not necessary for the dissociation of Swi6 from chromatin because deletion of pcf1 does not prevent this (Fig. 4C). It is possible that there is a mechanism that releases HP1 from heterochromatin just before its replication to facilitate synthesis, even in higher eukaryotes. Further analysis is required to reveal the mechanism for Swi6 release.
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As Pcf1 lacks the HP1/Swi6-binding motif (MIR; Murzina et al. 1999), which is conserved among CAF1 homologues in higher eukaryotes, we assume that this motif is not required for transferring HP1/Swi6 to newly replicated chromatin. Consistent with this assumption, chicken CAF1p150 lacking HP1 binding activity because of a MIR point mutation can support normal cell growth without detectable defects in heterochromatin structure (Takami et al. 2007). In addition, MIR is not required for the targeting of CAF1 to heterochromatic replication foci, but it is required for the association of CAF1p150 with heterochromatin outside S phase (Murzina et al. 1999). From these data, we propose that the weak and probably G1/S-specific transient interaction with HP1/Swi6 is important for CAF1 function at heterochromatic replication forks, whereas a strong interaction mediated by MIR might be required to retain CAF1 at heterochromatic regions and/or for other functions.
Chromatin bound Swi6 might recruit, directly or indirectly, H3K9 specific histone methyl transferase, Clr4 and HDACs to modify the newly assembled chromatin (Fig. 6D). Indeed, Swi6 interacts with the HDAC complex SHREC (Yamada et al. 2005; Sugiyama et al. 2007), and mammalian HP1 was shown to interact with SUV39H, the human homologue of Clr4 (Aagaard et al. 1999), although interaction between Swi6 and Clr4 has not been reported. H3K9me was decreased in pcf1 cells (Figs 2 and 4). We suspect that this is because of the decrease of Swi6, which might result in insufficient recruitment of Clr4. Alternatively, spCAF1 might recruit Clr4 to replication forks, as suggested for human cells; human CAF1 forms an S phase-specific complex containing the H3K9-specific histone methyl transferase SETDB1 (Sarraf & Stancheva 2004), although we have not so far detected association of CAF1 with Clr4 (K.D. and Y.M. unpublished results).
Another function of spCAF1 would be to recruit newly synthesized histones H3/H4 onto replicated DNA (Fig. 6E) at replication fork in both heterochromatin and euchromatin. Indeed we found that spCAF1 localized at replicating euchromatin (ars2004) at early S phase. This observation is consistent with the results that spCAF1 contributed to the maintenance of an active chromatin structure (the ura4-On state of the K::ura4 construct). In vertebrate cells, depletion of CAF1 results in a delay of chromatin assembly on newly replicated DNA and activates S phase checkpoint, resulting in S phase arrest and ultimately cell death (Hoek & Stillman 2003; Nabatiyan & Krude 2004; Takami et al. 2007). In contrast, fission yeast pcf1 cells did not show a detectable delay of S phase (data not shown) and no decrease of histone H3 on replicated DNA by the ChIP assay (Fig. 4C). However, it is possible that there might be a marginal "delay" in chromatin assembly on replicated DNA in cells lacking spCAF1. This delay would permit the action of inappropriate chromatin modifier proteins or misassembly of chromatin proteins, inducing instability of not only silent but also active chromatin structures.
The relatively small effect of depletion of spCAF1 could be because of the complementation of CAF1 defects by other chromatin assembly systems, including histone chaperones such as HIRA and Asf1. Our preliminary data indicate that a double mutation of one of CAF1 subunits and hip1, a fission yeast HIRA homologue, confers synergistic growth defects. Similarly, mutation of spCAF1 in conjunction with the asf1 temperature sensitive mutation confers synergistic heterochromatin and growth defects (K.D. and Y.M., unpublished results). A similar participation of spCAF1 and other histone chaperones in silent chromatin maintenance has been observed in budding yeast (Sharp et al. 2001; Huang et al. 2005). Interestingly, the replicative DNA polymerase Pol
was shown to interact with Swi6, and a Pol mutation that disturbs this interaction but not polymerase activity disrupts heterochromatin, indicating that Pol plays a specific role in heterochromatin assembly (Nakayama et al. 2001a). This Pol-dependent pathway might represent an alternative pathway to recruit Swi6 onto replicated heterochromatin. Thus, there are several pathways to ensure the maintenance of heterochromatin (and probably euchromatin). Another possible reason for the relatively weak silencing defect is that the de novo heterochromatin "establishing" activity compensates for defects in the spCAF1-dependent maintenance system by quick re-establishment of heterochromatin (Fig. 2C,D). This would also explain the fact that loss of spCAF1 did not affect telomeric heterochromatin, which has two independent heterochromatin-establishing systems, an RNAi-dependent system and a system dependent on the telomeric DNA binding protein, Taz1 (Kanoh et al. 2005).
In this study, taking advantage of fission yeast simple and conserved heterochromatin, we present the strong evidence that spCAF1 is involved in replication-coupled maintenance of HP1/Swi6-based heterochromatin. Mutational analyses in several organisms suggest that CAF1 plays a role in a wide range of nuclear functions, such as epigenetic regulation of gene expression (Kaya et al. 2001; Tchenio et al. 2001; Ono et al. 2006; Song et al. 2007), homologous recombination (Endo et al. 2006; Kirik et al. 2006) and genomic instability (Kolodner et al. 2002). Our preliminary results suggest spCAF1 is also involved in homologous recombination (K.D. and Y.M., unpublished results). Therefore, further analysis of spCAF1 will shed light on molecular mechanisms connecting maintenance of higher order chromatin structure and these nuclear functions.
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Experimental procedures |
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TopAbstractIntroductionResultsDiscussionExperimental proceduresNote added in proofReferences |
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All strains used in this study are listed in Table 1. Media and genetic methods for S. pombe experiments were essentially as described (Moreno et al. 1991). Yeast cells were cultured in YES, EMS at 30 °C, unless otherwise indicated. Comparative plating and serial dilution experiments for measuring silencing at heterochromatin were performed as previously described (Allshire et al. 1994). The deletion mutant of spCAF1 subunits were constructed by two-step PCR methods (Krawchuk & Wahls 1999) and entire ORF region of each subunit was replaced with selective marker gene. Disruption of pcf1 and pcf2 were carried out in both diploid and haploid cells with similar efficiency and the diploid disruptants generated viable spores harboring disruption of spCAF1 subunits with expected ratio, indicating that pcf1 and pcf2 were non-essential genes for growth. Deletion of pcf3 gene was carried out in haploid cells with similar efficiency to that of pcf1 or pcf2 gene, suggesting pcf3 is a non-essential gene. In addition, during all genetic crosses to make double or triple disruptants for spCAF1 genes, we did not see any evidence for the existence of suppressor mutations.
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Anti-Pcf1 antisera were prepared from rabbits injecting with GST-tagged Pcf1 protein produced in Escherichia coli. The anti-Swi6 rabbit polyclonal antibody and anti-dimethyl-histone H3-K9-dimethyl monoclonal antibody were previously described (Nakayama et al. 2000; Kato et al. 2005). Anti-H3K9-acetyl and anti-H3K14-acetyl antibodies were purchased from Upstate Biotechnology, Waltham, MA (cat #06-942 and 06-599). The anti-H3 antibody was purchased from Abcam, Cambridge, MA (cat # ab1791). Anti-PCNA and anti-RPA antibodies were gift from T. Tsurimoto and H. Masukata, respectively.
ChIP analysis was performed essentially as previously described (Nakagawa et al. 2002). The primer sets used were: ura4-DS/E-FW (5'-GAGGGGATGAAAAATCCCAT-3'), ura4-DS/E-RV (5'-TTCGACAACAGGATTACGACC-3'); act1-FW (5'-GAAG TACCCCATTGAGCACGG-3'), act1-RV (5'-CAATTTDADG TTCGGCGGTAG-3'), dg223-FW (5'-TGGTAATACGTACTAG CTCTCG-3'), dg223-RV (5'-AACTAATTCATGGTGATTGATG-3'), B15E12-FW (5'-CGATGCTCTCGACAAAGCCGTTCT-3') B15E12-RV (5'-CCATCTCAAACTTCTGTTCAACATT-3'), ars2004F (5'-ATGGTAGATGGAGAAACGGG-3') and ars2004R (5'-CACGGCATCTTTCTTCACGA-3'). Precipitated DNA was assayed by PCR in the presence of [32P]dCTP and separated on 5% native polyacrylamide gels, which were then dried. Bands were quantified using a PhosphorImager (Fuji BAS2000, Fuji Medical System, Valencia, CA). Each ChIP experiments were repeated at least three times to confirm their reproducibility.
Immunoprecipitation and immunoblotting
For immunoprecipitation, cells (3 x 108) were washed with cold water and suspended in EB buffer (50 mM HEPES–NaOH [pH 7.5], 50 mM KOAc [pH 7.5], 5 mM EGTA, 1% Triton X-100, 1 mM PMSF, protease inhibitor cocktail [Nakarai Tesque]). After disruption with glass beads, cells were centrifuged at 20 000 g for 15 min to obtain total extracts. A portion of the extracts was incubated at 4 °C for 3 h with 25 mL magnetic beads conjugated with an anti-mouse IgG antibody or an anti-rabbit IgG antibody (Dynal Biotech, Carlsbad, CA) that were preincubated with the antibody against a target protein. The magnetic beads were washed several times with cold EB buffer and suspended in SDS sample buffer. Immunoblotting was performed with appropriate antibodies conjugated with horseradish peroxidase and detected with the ECL system (GE Health Care).
Indirect immunofluorescence
Indirect immunofluorescence was performed essentially as previously described (Nakayama et al. 2001b) except that cells were incubated with the primary antibody for 30–60 min and with secondary antibody for 15–30 min.
Fluctuation assay
The ura4-Off and ura4-On states of K::ura4 cells were selected by growing cells in minimal medium containing FOA and in minimal medium lacking uracil (-Ura), respectively. The cells were transferred to non-selective media at 30 °C and grown for 10 generations and plated onto non-selective plates, -Ura and FOA plates. The ura4-On and ura4-Off phenotypes were scored after 6 days incubation at 30 °C and conversion rates were calculated.
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aPresent address: Graduate School of Natural Sciences, Nagoya City University, Mizuho-ku, Nagoya 467-8601, Japan.
bPresent address: GL Sciences Inc., 237-2 Sayamagahara, Iruma, Saitama 358-0032, Japan.
* Correspondence: yota{at}virus.kyoto-u.ac.jp |
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Received: 1 May 2008
Accepted: 10 July 2008
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