Myb-domain protein Teb1 controls histone levels and centromere assembly in fission yeast
2013; Springer Nature; Volume: 32; Issue: 3 Linguagem: Inglês
10.1038/emboj.2012.339
ISSN1460-2075
AutoresLuis P. Valente, Pierre-Marie Dehé, Michael Klutstein, Sofia Aligianni, Stephen Watt, Jürg Bähler, Julia Promisel Cooper,
Tópico(s)Plant Molecular Biology Research
ResumoArticle11 January 2013free access Source Data Myb-domain protein Teb1 controls histone levels and centromere assembly in fission yeast Luis P Valente Luis P Valente Cancer Research UK, London Research Institute, London, UK PDBEB, Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, PortugalPresent address: Epigenetics Mechanisms Laboratory, Instituto Gulbenkian de Ciência, 2780-156 Oeiras, Portugal Search for more papers by this author Pierre-Marie Dehé Pierre-Marie Dehé Cancer Research UK, London Research Institute, London, UKPresent address: Marseille Cancer Research Center (CRCM), U1068 Inserm, UMR7258 CNRS, Aix-Marseille University, Institut Paoli-Calmettes, Marseille 13009, France Search for more papers by this author Michael Klutstein Michael Klutstein Cancer Research UK, London Research Institute, London, UK Search for more papers by this author Sofia Aligianni Sofia Aligianni Department of Genetics, Evolution and Environment, UCL Cancer Institute, University College London, London, UKPresent address: Laboratory of Genetics, Salk Institute for Biological Studies, San Diego, CA 92186-5800, USA Search for more papers by this author Stephen Watt Stephen Watt Department of Genetics, Evolution and Environment, UCL Cancer Institute, University College London, London, UKPresent address: Regulatory Systems Biology Laboratory, Cancer Research UK Cambridge Research Institute, Li Ka Shing Centre, Robinson Way, Cambridge CB2 0RE, UK Search for more papers by this author Jürg Bähler Jürg Bähler Department of Genetics, Evolution and Environment, UCL Cancer Institute, University College London, London, UK Search for more papers by this author Julia Promisel Cooper Corresponding Author Julia Promisel Cooper Cancer Research UK, London Research Institute, London, UK Search for more papers by this author Luis P Valente Luis P Valente Cancer Research UK, London Research Institute, London, UK PDBEB, Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, PortugalPresent address: Epigenetics Mechanisms Laboratory, Instituto Gulbenkian de Ciência, 2780-156 Oeiras, Portugal Search for more papers by this author Pierre-Marie Dehé Pierre-Marie Dehé Cancer Research UK, London Research Institute, London, UKPresent address: Marseille Cancer Research Center (CRCM), U1068 Inserm, UMR7258 CNRS, Aix-Marseille University, Institut Paoli-Calmettes, Marseille 13009, France Search for more papers by this author Michael Klutstein Michael Klutstein Cancer Research UK, London Research Institute, London, UK Search for more papers by this author Sofia Aligianni Sofia Aligianni Department of Genetics, Evolution and Environment, UCL Cancer Institute, University College London, London, UKPresent address: Laboratory of Genetics, Salk Institute for Biological Studies, San Diego, CA 92186-5800, USA Search for more papers by this author Stephen Watt Stephen Watt Department of Genetics, Evolution and Environment, UCL Cancer Institute, University College London, London, UKPresent address: Regulatory Systems Biology Laboratory, Cancer Research UK Cambridge Research Institute, Li Ka Shing Centre, Robinson Way, Cambridge CB2 0RE, UK Search for more papers by this author Jürg Bähler Jürg Bähler Department of Genetics, Evolution and Environment, UCL Cancer Institute, University College London, London, UK Search for more papers by this author Julia Promisel Cooper Corresponding Author Julia Promisel Cooper Cancer Research UK, London Research Institute, London, UK Search for more papers by this author Author Information Luis P Valente1,2, Pierre-Marie Dehé1, Michael Klutstein1, Sofia Aligianni3, Stephen Watt3, Jürg Bähler3 and Julia Promisel Cooper 1 1Cancer Research UK, London Research Institute, London, UK 2PDBEB, Center for Neuroscience and Cell Biology, University of Coimbra, Coimbra, Portugal 3Department of Genetics, Evolution and Environment, UCL Cancer Institute, University College London, London, UK *Corresponding author. Cancer Research UK, London Research Institute, 44 Lincoln's Inn Fields, London WC2A 3LY, UK. Tel.:+44 20 7269 3415; Fax:+44 20 7269 3258; E-mail: [email protected] The EMBO Journal (2013)32:450-460https://doi.org/10.1038/emboj.2012.339 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 TTAGGG motif is common to two seemingly unrelated dimensions of chromatin function—the vertebrate telomere repeat and the promoter regions of many Schizosaccharomyces pombe genes, including all of those encoding canonical histones. The essential S. pombe protein Teb1 contains two Myb-like DNA binding domains related to those found in telomere proteins and binds the human telomere repeat sequence TTAGGG. Here, we analyse Teb1 binding throughout the genome and the consequences of reduced Teb1 function. Chromatin immunoprecipitation (ChIP)-on-chip analysis reveals robust Teb1 binding at many promoters, notably including all of those controlling canonical histone gene expression. A hypomorphic allele, teb1-1, confers reduced binding and reduced levels of histone transcripts. Prompted by previously suggested connections between histone expression and centromere identity, we examined localization of the centromeric histone H3 variant Cnp1 and found reduced centromeric binding along with reduced centromeric silencing. These data identify Teb1 as a master regulator of histone levels and centromere identity. Introduction The regulation of histone levels is a multi-faceted phenomenon crucial to genomic integrity. While DNA is generally packaged into nucleosomes comprising two copies each of histones H2A, H2B, H3, and H4, histones stand as a double-edged sword for DNA metabolism. Although their extreme basicity greatly favours strong interaction with DNA, any non-specific binding can lead to dramatic outcomes that threaten cell survival (Gunjan et al, 2005). Therefore, a major challenge is reached when cells undergo chromosomal replication, when histone production and nucleosome assembly must be coordinated with replication fork progression (Schumperli, 1986). A delay between DNA synthesis and histone deposition triggers DNA damage, chromosomal rearrangements and loss of viability (Han et al, 1987; Kim et al, 1988). Conversely, an excess of histones is also ill tolerated, as even a slight or transient excess of histone synthesis threatens the integrity of chromosome structure and gene expression (Meeks-Wagner and Hartwell, 1986; Singh et al, 2010). Regulation of histone levels also appears to play a key role in controlling the centromeric deposition of the histone H3 variant Centromere Protein A (CENP-A) (Castillo et al, 2007), which in turn nucleates the assembly of the kinetochores that connect chromosomes with spindle microtubules. While CENP-A and the kinetochore proteins are highly conserved, centromeric DNA sequences are poorly conserved and in many species, centromeres are assembled in a largely sequence-independent manner. Indeed, in mammals, flies, worms, and yeasts (Candida albicans and Schizosaccharomyces pombe), 'neocentromeres' can arise and fulfill proper kinetochore assembly in chromosomal regions devoid of normal centromere sequences (Williams et al, 1998; Ishii et al, 2008; Marshall et al, 2008; Ketel et al, 2009; Yuen et al, 2011). These observations suggest that epigenetic mechanisms control centromere identity. In all cases, perpetuation of the epigenetic mark depends on CENP-A binding, which is both necessary and sufficient for centromere inheritance (Mendiburo et al, 2011). The lack of sequence specificity also allows the spreading of CENP-A past its usual boundaries in cells overproducing CENP-A (Heun et al, 2006; Castillo et al, 2007). Conversely, overproduction of canonical histone H3 in fission yeast compromises centromere function by displacing CENP-A from the centromere central core (Castillo et al, 2007). The importance of the balance of core histone levels is further underscored by experiments in which the level of H3 relative to H4 is increased, leading to impaired CENP-A loading. In contrast, CENP-A loads normally if the H3/H4 ratio is decreased, suggesting that H3 and CENP-A compete for the available H4 histones. Histone concentrations are regulated at multiple levels including that of transcription. In budding and fission yeast, histone genes are arranged in a tail-to-tail organization with a common intergenic sequence. Schizosaccharomyces pombe contains a single gene encoding H2Aβ (hta2+), the H2Aα-H2B-encoding gene pair (hta1+-htb1+), and three copies of the gene pair encoding H3 and H4 (hht1-hhf1+, hht2+-hhf2+, hht3+-hhf3+). The levels of all histone transcripts except hht2+ increase during S phase (Takayama and Takahashi, 2007). Repression of histone gene transcription outside S phase or in response to hydroxyurea treatment is accomplished by members of the HIRA histone chaperone complex (Blackwell et al, 2004; Takayama and Takahashi, 2007). During S phase, the GATA-like transcription factor Ams2 is required for histone upregulation (Takayama and Takahashi, 2007). Ams2 is thought to bind a 17-bp consensus sequence known as the AACCCT box, located in the common promoter of each divergent pair of histone genes. Ams2 protein levels oscillate through the cell cycle, being maximal in S phase and decreasing dramatically in G2 via ubiquitin-mediated degradation. This degradation is linked to passage through S phase by phosphorylation of Ams2, which is accomplished by the S-phase kinase DDK and is requisite for the interaction between Ams2 and the Pof3 subunit of the SCF ubiquitin ligase. These observations suggest a regulatory loop in which Ams2 is synthesized in G1 to favour histone transcription during S phase and degraded in a DDK-dependent manner (Takayama et al, 2010). Ams2 is thought to play roles in chromatin assembly not only via modulation of histone levels, but also via assembly of CENP-A chromatin. Fission yeast centromeres comprise the pericentric heterochromatin region, which assembles at the so-called outer repeats (otr), and the central (cnt) and inner-most (imr) regions characterized by the presence of Cnp1CENP-A. Proper loading of Cnp1CENP-A at the central core requires both the establishment of heterochromatin at the otr and Ams2 function. Indeed, Ams2 was originally identified as a multicopy suppressor of the temperature-sensitive CENP-A mutant cnp1-1. A biphasic model for incorporation of Cnp1CENP-A into centromeric chromatin has been proposed in which Ams2 is required to favour Cnp1CENP-A loading during S phase while Cnp1CENP-A loading during G2 occurs through an Ams2-independent backup mechanism (Takayama et al, 2008). Teb1, also known as SpX or Mug152, was initially reported as a potential telomeric factor as it harbours two helix-loop-helix dsDNA binding domains of the homeodomain subset of Myb domains (Figure 1A), which are recurrently found in telomeric proteins; later it was identified as the product of a meiotically upregulated gene (Vassetzky et al, 1999; Spink et al, 2000; Martin-Castellanos et al, 2005). Curiously, however, Teb1 was shown to have higher affinity for the vertebrate telomere repeat, TTAGGG, than to fission yeast telomere repeats in vitro (Vassetzky et al, 1999; Spink et al, 2000). Here, we investigate the function of this essential protein and show that Teb1 binds in vivo and regulates the activities of many promoters, including those controlling the expression of all four types of canonical histones. We also find that Teb1 is involved in the centromeric loading of Cnp1CENP-A and maintenance of centromere identity. Moreover, Teb1 regulates the expression of a protease capable of histone clipping. Hence, Teb1 is a newly recognized general transcription factor with prominent roles in controlling histone levels and stability. Figure 1.Teb1 is essential and localizes to the nucleus. (A) Teb1 contains two Myb domains near its N-terminus. (B) Tetrad dissection of sporulated homozygous (teb1 +/+) and heterozygous (teb1+/−) diploids. While teb1+ spores are fully viable, teb1Δ spores germinate but die, forming microcolonies (arrow). (C) Endogenously GFP-tagged Teb1 localizes to the nucleus (counterstained with Hoechst 33342). (D) Serial dilution assay on complete media. teb1-1 and teb1-2 behave as hypomorphic alleles at 25°C and are inviable at 36°C. Rows A, B and D contain other teb1 TS mutants that were not further characterized. (E) Sequence alignment of myb domains from several proteins. Hypomorphic teb1-1 and teb1-2 display single point mutations in conserved regions of the first (teb1-1, Arg184Gly, black square) or second (teb1-2, Arg92Cys, grey square) Myb domains. (F) Southern blot analysis of telomere length. No size changes are conferred by the teb1-1 mutation. The same results were obtained for the other ts alleles (unpublished observations). Download figure Download PowerPoint Results Teb1 is an essential nuclear protein To investigate the roles of Teb1, a loss-of-function approach was taken in which one copy of the teb1+ gene in a diploid strain was replaced by a G418 resistance marker (Table I). Sporulation of the resulting heterozygous teb1+/teb1Δ diploid revealed that teb1Δ haploids are inviable, forming microcolonies of elongated cells that divide only a few times before ceasing division (Figure 1B). Fusion of a GFP tag with the Teb1 C-terminus at its endogenous locus yields fully viable strains harbouring Teb1-GFP, which yields a diffuse nuclear localization pattern (Figure 1C). Table 1. Strains used in this work JCF number Genotype Mating type 1 Wt h+ 2 Wt h− 24 ade6-M210 leu1-32 ura4-D18 h− 1942 ade6-M210 leu1-32 ura4-D18 teb1-1-3HA-KanMX6 h− 1944 ade6-M210 leu1-32 ura4-D18 teb1-2-3HA-KanMX6 h− 1969 ade6-M? leu1-32 ura4-D18 hht1-CFP-KanMX6 sid4-YFP-KanMx6 teb1-1-3HA-KanMX6 mis6-mcherry-natMX6 h+ 7401 cnt:arg3 cnt3:ade6 otr2: ura4 tel1L:his3 ade6-210 arg3-D4 his3-D1 leu1-32 ura4-D18 teb1-3HA-KanMX6 h− 7404 cnt:arg3 cnt3:ade6 otr2: ura4 tel1L:his3 ade6-210 arg3-D4 his3-D1 leu1-32 ura4-D18 teb1-1-3HA-KanMX6 h− 7419 teb1-1-3HA-KanMX6 h+ 7429 teb1-3HA-KanMX6 h+ 7433 cnt:arg3 cnt3:ade6 otr2: ura4 tel1L:his3 ade6-210 arg3-D4 his3-D1 leu1-32 ura4-D18 teb1-3HA-Kan (gift from R Allshire) h+ 7450 ade6-M210 leu1-32 ura4-D18 teb1-3HA-KanMX6 h− 7452 cnt1:arg3 cnt3:ade6 otr2: ura4 tel1L:his3 ade6-210 arg3-D3 his3-D1 leu1-32 ura4-D18 rik1::LEU2+ (gift from R Allshire) h? 7453 cnt1:arg3 cnt3:ade6 otr2: ura4 tel1L:his3 ade6-210 arg3-D3 his3-D1 leu1-32 ura4-D18 mis6-302 (gift from R Allshire) h? 7872 teb1-1-3HA::KanMX Ams2-13MYC::Hyg VKS 201 teb1-GFP-KanMX6 h? To generate a tool for studying Teb1 function, we screened for conditional alleles. Random mutations were generated by PCR amplification of a cassette harbouring the teb1+ open reading frame and a kanR marker that confers resistance to G418. A wild-type (wt) strain was transformed with the PCR products and G418-resistant transformants were screened for temperature-sensitive growth. Several hypomorphic mutant alleles that display sickness at the permissive temperature (25°C) and lethality at the restrictive temperature (36°C) were identified (Figure 1D). Two of these conditional alleles, teb1-1 and teb1-2, harbour mutations of conserved arginine residues within one or the other Myb domain (Figure 1E). Backcrossing teb1-1 with a wt strain recapitulates sickness at 25°C and inviability at 36°C. When teb1-1 cells were transformed with a plasmid-borne library of overexpressed fission yeast genes and grown at 36°C, the only plasmid conferring viability at 36°C encoded the wt teb1+ sequence, confirming that the reduced viability of teb1-1 strains is due to loss of Teb1 function (unpublished observations). To investigate whether Teb1 plays a role at telomeres, we examined the telomeres of teb1-1 cells grown at 25°C or following shift to the restrictive temperature of 36°C for 16 h. Southern blot analysis of terminal restriction fragments revealed that both teb1-1 and wt cells harbour telomeres of 300±50 bp at both temperatures (Figure 1F). Therefore, the teb1-1 mutation does not affect telomere length. Teb1 binds to the promoters of many genes The in vitro binding specificity of Teb1 for the vertebrate telomeric repeat sequence (TTAGGG) (Vassetzky et al, 1999) along with the presence of tandem copies of this repeat in the promoters of several fission yeast genes suggested that Teb1 might bind the corresponding promoters. To investigate this, we immunoprecipitated (IP) endogenously haemaglutinin (HA)-tagged Teb1 and hybridized the IP with the oligonucleotide 4 × 44K Chromatin immunoprecipitation (ChIP)-on-chip whole genome DNA microarray platform (Agilent), which covers the majority of the fission yeast genome. The resulting ChIP-chip results show a distinct and reproducible pattern of Teb1 binding (Figure 2A; Supplementary Figure S1). As expected based on the in vitro binding data, the Teb1 binding sites often contain runs of TTAGGG repeats; when found in histone gene promoters, a slightly permuted 17-bp version of these repeats has been referred to as the AACCCT box (Matsumoto and Yanagida, 1985; Figure 2B). Accordingly, ChIP-chip analysis detected Teb1 binding to histone promoters. We found that using an enrichment value of two-fold as the criterion for Teb1 binding, Teb1 can be seen to bind all five promoters of the nine canonical histone genes (Figure 3; note that four pairs of histone genes share divergent promoters; see also Supplementary Table I). Hence, Teb1 binds in vivo to AACCCT-like boxes in all histone gene promoters. Figure 2.Teb1 binds to specific genomic loci. (A) Teb1 binding map of Chromosome 1, represented along the X axis. Teb1 enrichment values, based on ChIP-chip analysis of endogenously tagged functional Teb1, are represented on the Y axis. (B) The AACCCT box found in all canonical histone gene promoters is bound by Teb1. Download figure Download PowerPoint Figure 3.Teb1 binds all histone promoters, and this binding is lost in teb1-1 cells. (A–E) Teb1 is specifically enriched at histone gene promoters. ChIP-chip data showing Teb1 binding at the indicated promoters. (F) ChIP of the indicated Teb1-HA variants at 25°C analysed by qPCR with primers for the histone hht2+/hhf2+ promoter region and the ade6+ control region. The values of 'relative enrichment' correspond to Teb1 binding at the hht2+/hhf2+ promoter normalized to the ade6+ promoter and to wt (non-tagged). The experiment was performed in triplicate and error bars correspond to the standard deviation. Unlike teb1+cells, teb1-1 cells do not harbour detectable levels of Teb1 binding to the hht2+/hhf2+ promoter. Download figure Download PowerPoint Expression profiling reveals a role for Teb1 in transcriptional regulation Given that Teb1 binds a number of gene promoters, we sought to determine whether the phenotypes of teb1-1 cells stem from altered transcription of these genes. We used microarray technology to assess the transcriptional profile of the teb1-1 strain and compare it to that of wt. RNA from wt and teb1-1 cells grown at 25°C was extracted and differentially labelled by reverse transcription with Cy3- and Cy5-labelled dCTP. The resulting labelled cDNA was hybridized to in-house-constructed glass slide microarrays containing probes for 99.3% of all known and predicted S. pombe genes (Lyne et al, 2003). This procedure was repeated for cells that had been subjected to growth for 1 h at the restrictive temperature of 36°C. The data were analysed with GenePix 6.0 software, with a two-fold difference in gene expression as the minimal change scored as upregulation or downregulation. A comprehensive list of the expression profile results can be found in Supplementary Table II. Upon inhibition of Teb1 function, many genes become upregulated whereas only a few are downregulated (Supplementary Table III). Intriguingly, the downregulated genes can be classified into gene ontology groups. For example, one such group comprises four genes involved in iron homeostasis, all of which are downregulated by teb1-1 (Supplementary Table IV). In a random distribution of genes, one would expect only ∼0.2% of genes from this group in the list, as only around 0.2% of genes in the genome are involved in iron homeostasis. However, this group represents ∼5 and ∼10% of the genes downregulated by the teb1-1 mutation at 25 and 36°C, respectively. Nonetheless, it is unclear why these four genes (str1, str3, fip1 and frp1) are downregulated, as the only member of this group that shows detectable Teb1 promoter binding is str3 (Supplementary Table I). Hence, Teb1 may act indirectly on these genes, perhaps by regulating the expression of a transcription factor that controls the iron homeostasis genes. Alternatively, the levels of Teb1 binding to the str1, fip1 and frp1 promoters may be significant but below the detection limit used in our ChIP-chip analysis. Another group that is overrepresented in the list of genes downregulated in a teb1-1 background at 25°C is the ribosomal protein (RP) genes (Supplementary Table II). Moreover, many ribosomal genes that fail to emerge as downregulated using a two-fold threshold show a reduction in expression of just under two-fold. As for the iron homeostasis group, no evidence of Teb1 binding to their promoters was found. Interestingly, the Candida albicans Myb-domain protein Tbf1 plays a role in controlling this same subset of ribosomal genes (Hogues et al, 2008). Even more striking was the downregulation of expression of the canonical histone genes induced by Teb1 mutation (Supplementary Table V), considered below. Teb1 is involved in histone transcriptional regulation The correspondence between Teb1 promoter binding and significant downregulation by teb1-1 mutation for all nine canonical histone genes prompted us to examine this group in more detail. We confirmed the ChIP-chip data by quantitative real-time PCR using primers for the promoter driving hht2+ and hhf2+ (encoding histones H3.2 and histone H4.2), which showed significant enrichment (Figure 3F). To further probe the relationship between Teb1 promoter binding and transcriptional regulation, we investigated the ability of the mutant Teb1-1 protein to bind histone promoters in vivo. ChIP analysis shows a drastic reduction in Teb1-1 binding to the hht2 and hhf2 promoter at both permissive (Figure 3F) and restrictive (unpublished observations) temperatures, highlighting the critical importance of the single mutated arginine residue within the Myb domain for the DNA-binding ability of this protein. Moreover, the vastly reduced binding of Teb1-1 at 25°C provides a plausible explanation for why teb1-1 cells grow slowly, even at permissive temperature. While the genes encoding all four types of canonical histones were among those downregulated by the teb1-1 mutation (Supplementary Table V), the level of downregulation was more modest for hta1+ and htb1+, which are regulated by the same divergent promoter, than for the other histones, being well under the two-fold cutoff used. Intriguingly, the slight downregulation of these two copies of histones is alleviated when cells are grown for 1 h at the restrictive temperature, 36°C. This suggests that Teb1 exerts only a minor effect on the regulation of hta1+/htb1+, and that additional factors promote transcription of these histones at high temperatures. Unlike the canonical histone genes, Teb1 does not appear to bind or regulate the expression of the H4 variant SPBC800.13, the H2 variant pht1 or the H3 variant cnp1 (Supplementary Figure S2; Supplementary Table II). Transcriptional upregulation of the canonical histone genes in S phase is under the control of the GATA-like transcription factor Ams2. While it has been suggested that Ams2 binds histone promoters via the AACCCT motif, it remains unclear whether Ams2 binds directly or is a part of a complex, as there are no GATA sequences within the conserved motif. Our observation that Teb1 binds these motifs at sites known to be regulated by Ams2 suggests Teb1 as a mediator of Ams2 binding. To test this possibility, we fused a 13Myc tag with the C-terminus of Ams2 and HA tags at the C-termini of wt Teb1 and Teb1-1; all fusions were at the endogenous loci and yielded functional protein. As previously published, we observe higher levels of Ams2 in S phase; we also observe moderately elevated levels of Teb1 in S-phase cells, suggesting that this protein is also upregulated during the time window when canonical histones are transcribed (Figure 4A). Importantly, Ams2 levels are not altered by the teb1-1 mutation; our microarray data also failed to detect Teb1 binding to the ams2+ promoter and revealed no changes in ams2+ expression in the teb1-1 background. However, ChIP assays reveal that while Ams2 strongly binds the hht2+/hhf2+ and hta1/htb1+ promoters in a wt background, binding is significantly reduced by the teb1-1 mutation (Figure 4B). Hence, Ams2 binding to histone promoters requires the presence of wt Teb1. Figure 4.Binding of Ams2 to histone promoters is partially lost in teb1-1 cells. (A) Western blot showing the levels of Teb1-HA and Ams2-Myc in asynchronously growing and cultures synchronized in S phase by treatment with 15 mM HU for 4 h. Ponceau staining confirmed equal loading in each lane (data not shown). (B) ChIP analysis with an anti-Myc antibody and qPCR using primers for the histone hht2/hhf2 and hta1/htb1 promoter regions or the adh1 locus (which harbours no Teb1 binding; Supplementary Table I) in cells synchronized in S phase with 15 mM HU. Ams2 binding is lost in the teb1-1 background. Download figure Download PowerPoint Teb1 is essential for maintenance of centromere identity The regulation of histone levels has been implicated in the maintenance of centromere identity. For instance, the identification of Ams2 as a multicopy suppressor of the Cnp1CENP-A ts allele cnp1-1 (Chen et al, 2003) could be attributed either to direct binding of Ams2 to the centromere central core or to the altered histone levels imparted by Ams2 overexpression. Hence, we wished to explore the possibility that Teb1 plays a role in Cnp1CENP-A recruitment, a role that could comprise at least part of its essential function. Centromeric Cnp1CENP-A confers local silencing of transcription; hence, expression of a marker gene inserted in the centromeric central core provides a readout for centromeric Cnp1CENP-A assembly (Pidoux et al, 2003). To assess the involvement of Teb1 in the Cnp1CENP-A deposition, we introduced the teb1-1 mutation into strains with an arg3+ marker integrated into the centromeric central core. To separate general effects on gene silencing from specific effects on centromere structure, we used a strain that also harbours markers in other silent regions of the genome, the pericentromeric outer repeats (otr-ura4+) and the telomere (telo-his3+). Two controls were also utilized, the rik1Δ mutation, which abolishes silencing at the centromeric outer repeats and the telomere, and the mis6-302 mutation, which abolishes silencing at the centromeric central core (Ekwall et al, 1996; Partridge et al, 2000). Efficient silencing of reporter genes in the outer repeats and telomere was observed in both wt and teb1-1 backgrounds. However, the reporter gene at the centromeric central core displayed a clear loss of silencing in teb1-1 cells at 25°C (Figure 5A). Figure 5.Teb1 is required for normal Cnp1 loading. (A) Silencing at the centromeric central core is specifically reduced in teb1-1 cells. Strains of the indicated genotypes harbouring a ura4+ marker in the otr region, an arg3+ marker in the central core and a his3+ marker in the subtelomere were analysed by serial dilution assay. (B) Cnp1 binding to the central core decreases in teb1-1 cells compared to wt. ChIP was performed with an anti-Cnp1 antibody. The experiment was performed in triplicate and error bars correspond to the standard deviation. (C) Indirect immunofluorescence shows that while Mis6 and Cnp1 colocalize at wt centromeres, Cnp1 foci fail to appear in many teb1-1 cells; Mis6 localization is not affected (white arrowheads). Download figure Download PowerPoint The central core-specific silencing defect of teb1-1 cells suggests a problem with Cnp1CENP-A deposition and kinetochore formation. To address these latter parameters, we performed ChIP of Cnp1CENP-A in wt and teb1-1 cells and qPCR for central core sequences. Remarkably, levels of central core enrichment in Cnp1CENP-A ChIP were significantly reduced by the teb1-1 mutation (Figure 5B). Interestingly, Cnp1CENP-A levels are slightly higher at the control act1+ or ade6+ loci in teb1-1 cells (Figure 5B). Elevated levels of Cnp1CENP-A at non-centromeric sites may result from reduced loading at the centromere. Two independent pathways of Cnp1CENP-A loading have been reported in fission yeast (Takayama et al, 2008), an S-phase pathway that depends on Ams2 and a G2 pathway dependent upon Mis6. To assess whether these pathways are differentially affected by the teb1-1 mutation, we carried out indirect immunofluorescence microscopy using antibodies against Cnp1CENP-A or Mis6 tagged with mCherry at its endogenous locus. Log-phase cultures, which comprise mainly G2 cells, were grown
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