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. 2007 Apr;175(4):1549–1560. doi: 10.1534/genetics.106.068684

Interaction of Epe1 With the Heterochromatin Assembly Pathway in Schizosaccharomyces pombe

Sara Isaac *,1, Julian Walfridsson †,1, Tal Zohar *, David Lazar *, Tamar Kahan ‡, Karl Ekwall †, Amikam Cohen *,2
PMCID: PMC1855143  PMID: 17449867

Abstract

Epe1 is a JmjC domain protein that antagonizes heterochromatization in Schizosaccharomyces pombe. Related JmjC domain proteins catalyze a histone demethylation reaction that depends on Fe(II) and α-ketoglutarate. However, no detectable demethylase activity is associated with Epe1, and its JmjC domain lacks conservation of Fe(II)-binding residues. We report that Swi6 recruits Epe1 to heterochromatin and that overexpression of epe1+, like mutations in silencing genes or overexpression of swi6+, upregulates expression of certain genes. A significant overlap was observed between the lists of genes that are upregulated by overexpression of epe1+ and those that are upregulated by mutations in histone deacetylase genes. However, most of the common genes are not regulated by Clr4 histone methyltransferase. This suggests that Epe1 interacts with the heterochromatin assembly pathway at the stage of histone deacetylation. Mutational inactivation of Epe1 downregulates ∼12% of S. pombe genes, and the list of these genes overlaps significantly with the lists of genes that are upregulated by mutations in silencing genes and genes that are hyperacetylated at their promoter regions in clr6-1 mutants. We propose that an interplay between the repressive HDACs activity and Epe1 helps to regulate gene expression in S. pombe.

THE assembly of heterochromatin domains along eukaryotic chromosomes is essential for several chromosomal functions, including proper chromosome segregation, inhibition of deleterious recombination, and dosage compensation (reviewed in Allshire 1997; Karpen and Allshire 1997; Grewal 2000; Martin and Zhang 2005; Zardo et al. 2005). However, unrestricted spreading of heterochromatin is detrimental, as it may repress expression of essential genes and tumor-suppressor genes. The heterochromatin assembly mechanism is conserved from fission yeast (Schizosaccharomyces pombe) to mammals. This mechanism is linked to post-translational modification of histones and association of chromodomain proteins, such as HP1 in mammals and Swi6 in S. pombe, with methylated lysine 9 on histone H3 (H3-K9methyl) (Wang et al. 2000; Jenuwein and Allis 2001). In addition, RNA interference pathway enzymes and DNA-binding proteins participate in heterochromatin assembly (Hall et al. 2002; Volpe et al. 2002; Verdel and Moazed 2005; Yamada et al. 2005; Irvine et al. 2006).

Current models postulate that deacetylation of H3-K9acetyl and H3-K14acetyl is a prerequisite for methylation of H3-K9. At least three histone deacetylases (HDACs) are linked to heterochromatin assembly in S. pombe: Clr3 and Sir2 deacetylate H3-K14acetyl and H3-K9acetyl, respectively, and Clr6 is a broad range HDAC. Another HDAC, Hos2, promotes high expression of growth-related genes by deacetylating H4-K16acetyl in open reading frames (ORFs) (Wiren et al. 2005). Clr4 is the only known H3-K9-specific histone methyltransferase (HMTase). Methylation of H3-K9 by Clr4 generates a high-affinity binding site for Swi6, which promotes further propagation of methylated H3-K9 by recruiting Clr4 HMTase to the growing end of the heterochromatin structure (Nakayama et al. 2001).

Recently, it has been demonstrated that Swi6 is also involved in propagation of Clr3 along heterochromatin domains. Clr3 is recruited through an Atf1/Pcr1-dependent mechanism to a specific binding site at the mat locus and then spreads from its nucleation site by a mechanism that depends in its HDAC activity, Sir2, Swi6, and another chromodomain protein, named Chp2 (Yamada et al. 2005). In addition to their role in constitutive heterochromatin assembly, HDACs and Clr4 HMTase regulate transcription activity (Hansen et al. 2005; Wiren et al. 2005). However, whereas both HDACs and Clr4 are essential for constitutive heterochromatin assembly (Nakayama et al. 2001), HDAC-mediated repression of gene expression is often independent of H3-K9 methylation (Hansen et al. 2005).

Epe1 is a conserved Jumonji C (JmjC) domain nuclear protein that modulates heterochromatization (Ayoub et al. 2003). Inactivation of Epe1 enhances reporter gene silencing at the periphery of heterochromatin domains, promotes heterochromatin spreading across boundary elements, and partially suppresses clr1, clr3, and clr6 mutations (Ayoub et al. 2003). Conversely, overexpression of Epe1 abrogates heterochromatin structure and impairs centromere functions. Furthermore, Epe1 activity depends on the integrity of its JmjC domain (Ayoub et al. 2003).

The biochemical activity associated with Epe1 is not understood. However, Epe1 can be modeled onto the structure of a JmjC protein, named factor-inhibiting hypoxia-inducible factor (HIF), which catalyzes the hydroxylation of an asparagine residue on HIF through a Fe(II)- and α-ketoglutarate (α-KG)-dependent reaction (Hewitson et al. 2002). On the basis of this homology, it has been postulated that Epe1 is a histone demethylase that acts by hydroxylation of the methyl group on mono-, di-, or trimethyl lysine in a Fe(II)- and α-KG-dependent reaction (Trewick et al. 2005). Consistent with this hypothesis, several JmjC domain proteins display histone demethylase activity that depends on Fe(II) and α-KG (Fodor et al. 2006; Tsukada et al. 2006; Whetstine et al. 2006; Yamane et al. 2006). Yet, the relevance of these findings to the enzymatic activity of Epe1 is not clear, as no detectable demethylation activity was associated with purified Epe1, and Epe1 lacks conservation of two JmjC domain residues predicted to bind Fe(II) (Tsukada et al. 2006; Zofall and Grewal 2006).

To gain an insight into the mechanism underlying Epe1-mediated modulation of heterochromatization, we investigated Epe1 interaction with the heterochromatin assembly pathways. Our results indicate that Epe1 interacts with Swi6 and that this interaction is essential for localization of Epe1 to heterochromatin domains. Furthermore, analysis of global gene expression profiling suggests that Epe1 helps to regulate gene expression by counteracting the repressive effect of silencing proteins and that its interaction with the heterochromatin assembly pathway is at the stage of histone deactylation.

MATERIALS AND METHODS

Yeast strains and culture conditions:

All strains used in this study and their genotypes are listed in Table 1. Standard genetic crosses and transformation procedures (Moreno et al. 1991) were used in strain construction. The strain collection with random ura4+ insertions has been described before (Ayoub et al. 2003). Locations of the inserted ura4+ reporter gene in selected strains of this collection were determined by Genome Walker methodology, using BD GenomeWalker universal kit (BD Bioscience Clontech). The epe1∷ura4+ allele was constructed by inserting the ura4+ marker into chromosomal epe1+ at the XhoI site, upstream from the JmjC domain. The phenotype of epe1∷ura4+ mutants is indistinguishable from that of strains with the epe1-1 nonsense allele (data not shown). swi6+∷ECFP was crossed into the appropriate genetic background, using YA727 as a donor. YA727 was a gift from Hiroshi Iwasaki, Yokahama City University, Yokohama, Japan. Chromosomal epe1+∷YFP(venus) was constructed by transferring YFP(venus) from pSC2 (Nagai et al. 2002), using PCR amplification with long primers homologous to YFP, to flanking chromosomal sequences at the 5′-end of epe1+. nmt1 was inserted upstream of epe1+ by a similar methodology. Strains were grown on rich medium (YEA) (Moreno et al. 1991) and defined medium supplemented with the appropriate growth requirements (Moreno et al. 1991). AA−ura is a uracil-depleted defined medium. Cells were tested for the Ura− phenotype on 5-fluoroorotic acid (5-FOA) medium, since ura4+ expression in this medium leads to the synthesis of a toxic product from FOA (Boeke et al. 1987). FOA medium was a defined medium supplemented with 1 mg/ml FOA and 0.1 mg/ml uracil.

TABLE 1.

S. pombe strains used in this study

Strains mat1 allele Insertions/deletions Auxotrophic markers Other mutations Source or reference
AP1102a mat1-M-smt0 L(HpaI)∷ade6+SPBC1683.07∷ura4+ leu1-32 his2 ade6-210 ura4-D18 epe1-1 This study
AP1107b mat1-M-smt0 L(HpaI)∷ade6+SPAC23H3.14∷ura4+ leu1-32 his2 ade6-210 ura4-D18 epe1-1 This study
AP1112c mat1-M-smt0 L(HpaI)∷ade6+SPAC23H3.14∷ura4+ leu1-32 his2 ade6-210 ura4-D18 epe1-1 This study
AP1066a mat1-P-Δ17∷LEU2 L(SacI)∷ade6+SPBC1683.07∷ura4+ leu1-32 his2 ade6-210 ura4-D18 This study
AP1170b mat1-P-Δ17∷LEU2 L(SacI)∷ade6+SPAC23H3.14∷ura4+ leu1-32 his2 ade6-210 ura4-D18 This study
AP1186a mat1-P-Δ17∷LEU2 L(SacI)∷ade6+SPBC1683.07∷ura4+ leu1-32 his2 ade6-210 ura4-D18 epe1-1 This study
AP1190b mat1-P-Δ17∷LEU2 L(SacI)∷ade6+SPAC23H3.14∷ura4+ leu1-32 his2 ade6-210 ura4-D18 epe1-1 This study
AP136 mat1-M-smt0 L(SacI)∷ade6+ leu1-32 his2 ade6-210 ura4-D18 Ayoub et al. (1999)
AP182 mat1-P-Δ17∷LEU2 L(SacI)∷ade6+ ade6-210 ura4-D18 leu1-32 his2 Ayoub et al. (2003)
AP2005 mat1-M-smt0 L(SacI)∷ade6+ leu1-32 his2 ade6-210 ura4-D18 epe1-1 Ayoub et al. (2003)
AP2048 h− epe1∷ura4+ ura4DS/E This study
AP938 mat1-M-smt0 L(SacI)∷ade6+swi6+∷ ECFP leu1-32 ade6-210 ura4-D18 This study
AP940 mat1-M-smt0 L(SacI)∷ade6+swi6+∷ ECFP epe1∷ura4+ leu1-32 ade6-210 ura4-D18 This study
AP944 mat1-M-smt0 L(SacI)∷ade6+swi6+∷ ECFP nmt1-epe1+ leu1-32 ade6-210 ura4-D18 This study
AP948 mat1-M-smt0 L(SacI)∷ade6+epe1-TAP(KanR) leu1-32 ade6-210 ura4-D18 his2 This study
AP952 mat1-M-smt0 L(SacI)∷ade6+epe1+∷YFP(venus)LEU2 leu1-32 ade6-210 ura4-D18 his2 This study
AP955 mat1-M-smt0 L(SacI)∷ade6+swi6Δ∷ura4+ epe1+∷YFP(venus)LEU2 leu1-32 ade6-210 ura4-D18 his2 This study
Hu652 h− ura4 DS/E This study
Hu969 h− nmt1-epe1+ ura4DS/E This study
YA727 h90 swi6+∷ECFPd leu1-32 arg3-D1 ura4-D18 his3-D1 H. Iwasaki
UM234 mat1-M-smt0 (HpaI)∷ade6+ C11E10.09c∷ura4+ ade6-210 ura4-D18 leu1-32 his2 epe1-1 Ayoub et al. (2003)
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a

The ura4+ insertion in this strain is ∼400 bp downstream from SPBC1683.07. L(Hpa1)∷ade6+, here and elsewhere, designates an ade6+ insertion at the HpaI site within the L region of the mat locus (see Figure 2).

b

The ura4+ insertion in this strain is ∼150 bp upstream of SPAC23H3.14.

c

The ura4+ insertion in this strain is ∼200 bp upstream of SPAC23H3.14.

d

Chromosomal swi6+∷ECFP encodes a functional Swi6 derivative with an ECFP tag fused to its C terminus.

Fluorescence microscopy:

Logarithmic-phase cells were applied to slide coverslips that had been pretreated with polylysine. For fixed-cell analysis (Figure 2C), samples were subjected to ethanol fixation and DNA was stained with 4′,6-diamidino-2-phenylindole (DAPI). Samples were analyzed by fluorescence microscopy, and images were processed with Metamorph (Universal Imaging, West Chester, PA). Filters used were CFP HE FS47, YFP FS46 HE, and DAPI FS02 HE (Zeiss).

Figure 2.—

Figure 2.—

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Localization of Epe1-YFP(venus) to heterochromatin domains depends on Swi6 activity. (A) At the mat locus of S. pombe, heterochromatin is located between the IRL and IRR elements (Noma et al. 2001; Thon et al. 2002). The locations of mat2, cenH, and the SacI site at the centromere-distal end of IRL are indicated. (B) Fusion of peptide tags to the C-terminal end of Epe1 enhances its activity and/or stability. Expression of ade6+ from the SacI site in cells of the indicated genotypes was monitored on low-adenine medium (YE), and association of H3-K9dimethyl with cenH was determined by ChIP analysis with anti-H3-K9dimethyl antibodies and cenH primers. (C) Localization of Epe1-YFP(venus) in swi6+ (AP952) and Δswi6 (AP955) cells. DAPI nuclear staining and fluorescence microscopy analysis of Epe1-YFP localization in fixed cells of isogenic swi6+ and Δswi6 strains are presented. (D) Epe1-YFP association with cenH, centromeric outer repeats, and a subtelomeric region (lth1). Association of Epe1-YFP with the respective regions was determined by ChIP analysis with anti-GFP antibodies and the corresponding primers (supplemental Table S1 at http://www.genetics.org/supplemental/).

Chromatin immunoprecipitation analysis:

Chromatin immunoprecipitation (ChIP) experiments with anti-GFP (Abcam) and anti-H3-K9dimethyl antibodies (Upstate Biotechnology, Lake Placid, NY) were conducted as described before (Ayoub et al. 2003). The polyclonal anti-GFP antibodies used in this study cross-react with enhanced cyan fluorescence protein (ECFP) and yellow fluorescence protein (YFP). The relative ratios of band intensities were corrected for the ratios in whole-cell extracts (WCE).

RT–PCR analysis:

Cell cultures were grown in rich YES media to logarithmic phase and harvested by centrifugation at 4°. Cells were disrupted, using a Fastprep FP120 bead beater at a speed of 6.5, five times for 20 sec. Total RNA was extracted from crude lysates using QIAGEN (Chatsworth, CA) RNAeasy extraction and purification protocol (QIAGEN). Purified RNA was subjected to reversed transcription using Superscript first-strand synthesis to generate cDNA, according to the manufacturer's instructions (Invitrogen, San Diego). cDNA was amplified with primers specific for isp5, SPAC922.03, SPAC1039.06, SPAC1039.07, SPAC1039.08, and a moderately expressed gene, SPAPB1A10.11c, as a control. SPAPB1A10.11c encodes glutamyl tRNA synthetase (a role inferred from homology). Its expression is known not to be affected by meiosis, cell cycle, or environmental stress (Mata et al. 2002; Chen et al. 2003). PCR products were separated on agarose gels and stained with ethidium bromide, and band intensities were determined using Fujifilm Image Gauge software. The relative ratios for the epe1 mutant and the overexpressed Epe1 were determined by normalization to the wild-type ratio at the same locus.

Genomic sequence data:

All sequence and gene location data were retrieved from Welcome Trust Sanger Institute Schizosaccharomyces pombe genome center (Wood et al. 2002) (http://www.sanger.ac.uk/Projects/S_pombe).

Microarray analysis:

Genomewide expression profiling was performed as previously described (Xue et al. 2004). Briefly, cells were cultured in rich YES media to midlogarithmic growth phase, harvested, and disrupted. Total RNA was extracted from wild-type and respective mutants and labeled using RT–PCR with fluorescent cy3 and cy5 dyes (Amersham, Buckinghamshire, UK), respectively. Cy3- and cy5-labeled RNA preparations from mutant and wild-type cells, respectively, were pooled and together hybridized on the microarray slides. Each gene was represented by two spots on the microarray slide, the expression-profiling experiments were carried out in duplicates, and labeled RNA was dye swapped to correct for dye effects. After scanning and quantification of the microarrays, the data were analyzed using Gene Spring v. 7.2 (Silicon Genetics). The pEpe1 vs. wild-type data were normalized using Lowess “per spot per chip” intensity-dependent normalization according to our standard procedure (Xue et al. 2004). However, we found that Lowess was not suitable for epe1∷ura4+ vs. wild-type data sets since it corrects for a ratio bias in the low-intensity range of the expression profile and therefore underestimates downregulation of low-expression genes. Therefore, a “per chip” median normalization was applied to epe1∷ura4+ data sets (Table 4). The microarray data are deposited at http://natvet.sh.se/ekwall_lab/data1.htm.

TABLE 4.

Comparison between the list of genes that are downregulated by epe1:ura4+ and genes that are upregulated by overexpression of epe1+, mutations in silencing genes, meiosis, stress, and genes that are poorly expressed in growing cells

Mutation/condition No. of genes affected by the mutation/condition Intersection with downregulated genes in epe1∷ura4+ P-value
nmt-epe1+ (up) 270 55 1.31E-4
clr6-1 (up)a 442 86 1.03E-5
clr3-735 (up)a 186 35 0.0083
clr4-681 (up)a 109 23 0.0077
High IGR clr6-1 H4K5ac (up)ab 349 72 6.71E-5
High IGR clr6-1 H4K12ac (up)ab 238 59 2.4E-4
High IGR clr6-1 H3K9ac (up)ab 217 51 0.0099
Meiosis (up)c 1429 296 5.4E-22
Stress (up)d 862 158 1.7E-4
Low expression in rapidly growing cellsa 989 351 5.7E-103
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up, upregulated.

a

Data are from Wiren et al. (2005).

b

IGR indicates intergenic regions upstream from the affected genes; values are corrected for nucleosome density (Wiren et al. 2005).

c

Data are from Mata et al. (2002).

d

Data are from Chen et al. (2003).

Differentially expressed genes were defined as twofold upregulated (unless otherwise indicated) for at least three of four data points. Data acquisition and quality control are essentially as described before (Wiren et al. 2005). For comparison with expression profiles of strains in different genetic backgrounds, sporulation, stress conditions, and average wild-type expression, we used databases from the following articles: Mata et al. (2002); Chen et al. (2003); Gatti et al. (2004); Watson et al. (2004); Hansen et al. (2005); and Wiren et al. (2005).

The genetic interactions of Epe1 with heterochromatin assembly proteins were examined by grouping the interacting genes with partial overlaps and estimating the statistical significance of the overlaps among these groups. We used a common method for such analyses—the hypergeometric distribution (Lee et al. 2005), where P-value <0.05 is statistically significant (Tables 3 and 4). Clusters of Epe1 upregulated genes were defined as follows: the genomic sequence of each chromosome was scanned with overlapping windows, each containing 20 successive genes. For each such window, the number of upregulated genes was counted, and the statistical significance of this number was calculated using the hypergeometric distribution. Successive statistically significant (P-value <0.05) windows were united if and only if the union was statistically significant. Clustering P-values were calculated using hypergeometric distribution.

TABLE 3.

Comparison of upregulation of gene expression between overexpression of epe1+ and mutations in silencing genes, overexpression of swi6+, sporulation, and stress

Mutant/condition No. of genes upregulated by the mutation/condition Intersection with overexpression of epe1+f P-value
clr1-5a 151 59 1.48E-12
clr3-735b 186 75 1.33E-18
clr4-681a 109 47 3.45E-12
clr6-1b 442 70 6.50E-12
clr6-1Δclr3a 582 91 1.19E-12
Δsir2c 127 13 0.022
Δhos2c 451 42 0.0058
Δswi6c 10 5 3.71E-04
Δdcr1a 60 18 1.43E-09
Δago1a 65 13 4.00E-05
Δrdp1a 84 17 2.16E-06
swi6+ overexpressionc 244 77 2.11E-12
Meiosisd 945 144 3.81E-12
Stresse 474 132 5.09E-15
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a

Data are from Hansen et al. (2005).

c

Data are from Wiren et al. (2005).

d

Data are from Mata et al. (2002).

e

Data are from Chen et al. (2003) (environmental stress); Gatti et al. (2004) (cisplatin); Watson et al. 2004 (ionizing radiation).

f

Intersections are with the list of genes that were upregulated over twofold by overexpression of epe1+ (270 genes).

RESULTS

Overexpression of epe1+ impairs methylation of H3-K9 and Swi6 occupancy at heterochromatic domains:

Overexpression of epe1+ enhances methylation of H3-K4 and acetylation of H3-K9 and H3-K14 at the silent domain of the mat locus (Ayoub et al. 2003). To further elucidate the effect of overexpression of epe1+ on heterochromatin structure, we constructed strains that express chromosomal epe1+ from its endogenous promoter (AP936) or from an nmt1 promoter (AP944) and, in addition, express a functional Swi6-ECFP fusion protein. We then conducted a ChIP analysis with anti-GFP and anti-H3-K9dimethyl antibodies. Results (Figure 1A) indicate that overexpression of epe1+ from the nmt1 promoter lowers the level of H3-K9 methylation and Swi6 occupancy at constitutive heterochromatic domains in the mat locus and at a subtelomeric sequence. However, reduction at the centromeric outer repeats (otr) was less pronounced. We also employed fluorescence microscopy to determine the effect of epe1 genotype on nuclear localization of Swi6-ECFP (Figure 1B). Consistent with previous observations (Ekwall et al. 1995), Swi6-ECFP was localized in epe1+ cells (AP938) to a small number of distinct foci at heterochromatic domains. Fluorescent foci were also visible in cells of an epe1∷ura4+ mutant (AP940) and in cells that overexpress epe1+ (AP944). However, while mutational inactivation of Epe1 had no detectable effect on the number of fluorescence foci, overexpression of epe1+ decreased this number significantly (Figure 1C). These data suggest that overexpression of Epe1 partially reduced H3-K9 methylation and Swi6 association with constitutive heterochromatin domains.

Figure 1.—

Figure 1.—

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The effect of the epe1 genotype on methylation of H3-K9 and association of Swi6 with heterochromatin. swi6+-ECFP derivatives, expressing an epe1+ allele (AP938), an epe1∷ura4+ allele (AP940), or chromosomal epe1+ from an nmt1 promoter (AP944), were subjected to ChIP analysis and fluorescence microscopy. (A) ChIP analysis of H3-K9dimethyl and Swi6-ECFP localization at mat2, the cenH region of the mat locus, the otr, and a subtelomeric sequence (sub-tel). DNA isolated from immunoprecipitated fractions, using anti-H3-K9dimethyl or anti-GFP antibodies, or from WCEs was analyzed by PCR with the respective primers (supplemental Table S1 at http://www.genetics.org/supplemental/). “Ratio” is defined as the ratio of signal intensities between the electrophoretically separated bands of the immunoprecipitated fractions, normalized to that in WCE. (B) Localization of Swi6-ECFP fluorescent foci in strains of the indicated epe1 genotypes. The same cells subjected to fluorescence and phase microscopy are shown with superimposed images. (C) Analysis of the number of Swi6-ECFP foci/cells in the different epe1 genotypes (B). The percentage of cells with the indicated number of foci is presented. Foci were counted in 125 epe1+ cells, 191 epe1-1 cells, and 295 nmt1-epe1+ cells. Focus-number distributions were compared by χ2 test (d.f. = 4). The difference in focus numbers between nmt1-epe1+ and epe1+ cells is statistically significant (P ≪ 0.0001). However, the distribution of foci numbers in epe1+ and epe1-1 cells is statistically indistinguishable (P > 0.2).

Swi6 recruits Epe1 to constitutive heterochromatic domains:

To determine the nuclear localization of Epe1, we constructed strains that express an Epe1-YFP (Venus) fusion protein from the native chromosomal epe1 promoter. We then assessed the activity of the fusion protein by monitoring ade6+ expression from the SacI site at the centromere-distal end of the IRL boundary element at the mat locus (Figure 2A). Expression of ade6+ from this locus is affected by the epe1 genotype: An epe1-1 mutant (AP2005) exhibits an Ade− phenotype [red colonies on low-adenine (YE) medium], a strain that overexpresses epe1+ (AP944) is Ade+ (white colonies), and an epe1+ strain (AP136) exhibits a variegated Ade phenotype (Figure 2B) (Ayoub et al. 2003). The Ade+ phenotype [white colonies on low-adenine (YE) medium] of the epe1+-YFP strain (AP952) (Figure 2B) indicated that the fusion protein is functional. Furthermore, it suggests that the YFP tag at the C-terminal end of Epe1 enhances Epe1 activity and/or stability. A similar phenotype was observed with an isogenic strain expressing an epe1+-TAP fusion protein (AP948) (Figure 2B). Consistent with this indication, the degree of H3-K9 methylation at a cenH sequence within the mat locus was lower in cells that express epe1-YFP than in cells of an isogenic epe1+ strain (Figure 2B). Next, we subjected cells of a strain that expresses epe1-YFP to fluorescence microscopy. The observed pattern of fluorescent foci indicated that Epe1, like Swi6, is localized at a small number of nuclear foci (Figure 2C). To ascertain whether the observed foci reflect enrichment of Epe1-YFP at constitutive heterochromatin domains, we conducted ChIP analysis with anti-GFP antibodies and primers of cenH, centromeric outer repeats, and subtelomeric sequences. The observed enrichment of Epe1-YFP at these loci (Figure 2D) confirms that Epe1 is preferentially localized to constitutive heterochromatin domains.

Results of a yeast two-hybrid assay indicated physical interaction of Swi6 with Epe1 (Trewick et al. 2005). Since both Swi6 and Epe1 are associated with heterochromatin domains, we asked whether targeting of Epe1 to heterochromatic domains depends on its interaction with Swi6. To this end, we determined the effect of a swi6Δ mutation on Epe1-YFP localization. Remarkably, deletion of swi6+ abolished Epe1-YFP nuclear foci (Figure 2C), thus indicating dependence of Epe1 localization on Swi6. To further substantiate this indication, we compared ChIP analysis data with anti-GFP antibodies on swi6+ and Δswi6 derivatives of an epe1-YFP strain. Consistent with the fluorescence microscopy data, deletion of swi6+ markedly reduced Epe1 association with all three heterochromatin domains (Figure 2D). These data indicate that recruitment of Epe1 to constitutive heterochromatic domains depends on Swi6. Dependence of Epe1 localization on Swi6 has been recently demonstrated independently by Zofall and Grewal (2006).

Genetic interactions of Epe1 with silencing proteins:

Given that overexpression of epe1+, as with mutations in silencing genes, derepresses reporter genes within constitutive heterochromatin domains (Ayoub et al. 2003) and that mutations in silencing genes upregulate gene expression (Hansen et al. 2005; Wiren et al. 2005), we asked whether overexpression of epe1+ would upregulate gene expression at endogenous euchromatic locations. To this end, we subjected total RNA preparations of cells that express epe1+ from its endogenous promoter (Hu652) or from an nmt1 promoter (nmt1-epe1+) (Hu969) to expression profiling by spotted DNA microarray. This analysis revealed that ∼5% of S. pombe genes (270 genes) are upregulated by overexpression of epe1+ (supplemental data at http://www.genetics.org/supplemental/). Statistical analysis indicated significant clustering of upregulated genes at eight chromosomal locations (Table 2 and Figure 3A). However, none of these clusters was located within 100 kb of the telomeres. RT–PCR analysis of gene expression within one of the clusters (Figure 3B) confirmed that overexpression of epe1+ enhances transcription within the cluster.

TABLE 2.

Clusters of genes that are upregulated by overexpression of epe1+

Chromosome no./cluster Chromosomal locationa Length (bp) Total no. of genes No. of genes upregulated by nmt1-epe1+ P-value
I/1 309706 95,673 65 19 1.21E-08
I/2 2824851 78,236 30 10 4.13E-06
I/3 5587187 44,635 21 7 1.26E-04
II/4 343760 43,144 20 5 0.010
II/5 4352245 108,112 46 12 2.61E-06
II/6 4469502 40,569 14 5 2.26E-04
III/7 1455033 42,810 17 6 2.3E-04
III/8 1801848 39,354 21 6 8.4E-04
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a

Distance parameters according to http://www.sanger.ac.uk/Projects/S_pombe/. Numbers indicate the location of the cluster start point in base pairs.

Figure 3.—

Figure 3.—

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Overexpression of epe1+ upregulates chromosomal gene expression. (A) The distribution of genes that are upregulated twofold or more by overexpression of Epe1 along S. pombe chromosomes. The locations of gene clusters are indicated by yellow frames. (B) Enhancement of transcription activity within a cluster by overexpression of epe1+. Transcription activity of the indicated genes within cluster 1 (A), in epe1+ (Hu652), epe1∷ura4+ (Hu968), and nmt1-epe1 (Hu969) strains was determined by RT–PCR with the SPAPB1A10.11c gene, encoding glutamyl tRNA synthetase (a role inferred from homology) as a control.

Comparison of the list of genes that are upregulated by overexpression of epe1+ with the list of genes that are upregulated by mutations in silencing genes (clr1, clr3, clr6, hos2, sir2, or clr4) revealed extensive overlap (Figure 4A). This result suggests that Epe1 shares gene targets with silencing enzymes. Furthermore, overexpression of epe1+, as with mutations in silencing genes (Hansen et al. 2005), upregulates meiotic genes and stress-responsive genes (Table 3). Hypergeometric distribution analysis indicated different degrees of overlapping between the list of genes that are upregulated by overexpression of epe1+ and the lists of genes that are upregulated by mutations in different HDAC genes (Table 3). About 40% of the genes that are upregulated by a clr3 mutation are also upregulated by overexpression of epe1+. Yet, only 4% of the genes that are upregulated by a hos2 mutation, 9% of the genes that are upregulated by a clr6 mutation, and 10% of the genes that are upregulated by a sir2 mutation are also upregulated by overexpression of epe1+.

Figure 4.—

Figure 4.—

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Genetic interactions of Epe1 with heterochromatin assembly proteins. (A) Most genes that are upregulated by overexpression of epe1+ are also upregulated by mutations in silencing genes. The list of genes that are upregulated by mutations in one or more silencing genes was compared to the list of genes that are upregulated by overexpression of epe1+. (B and C) The list of genes that are upregulated by overexpression of epe1+ was compared to the list of genes that upregulated by mutations in HDAC genes, clr3, and clr4. The majority of the genes that are upregulated by overexpression of epe1+ and by mutations in HDAC genes (B) or clr3 (C) are not affected by clr4 genotype. (D) A significant overlap is observed between the lists of genes that are upregulated by overexpression of epe1+, overexpression of swi6+, and mutations in HDAC genes. P-value indicates the statistical significance of the overlap between all intersecting gene lists.

About 85% of the genes that are upregulated by mutations in HDAC genes are not affected by clr4 mutations (Hansen et al. 2005). Therefore, to examine the possibility that Epe1 affects HDAC-mediated gene repression, independently of a possible effect on H3-K9 methylation, we compared gene expression profiles of strains that overexpress epe1+ to those of clr4 and HDAC mutants. About half of the genes that are upregulated by both clr4 mutations and mutations in one or more HDAC genes (clr3, clr6, sir2, and/or hos2) are also upregulated by overexpression of epe1+. However, most genes that are upregulated by mutations in HDAC genes and overexpression of epe1+ are not affected by a clr4 mutation (Figure 4B). Furthermore, close to 80% of the genes that are upregulated by a clr3 mutation and overexpression of epe1+ are not affected by a clr4 mutation (Figure 4C). These data suggest that Epe1 can antagonize gene silencing by interaction with the heterochromatin assembly pathway at the stage of histone deacetylation, independently of a possible interaction at the stage of H3-K9 methylation.

Only 10 genes are upregulated by a swi6 mutation (Wiren et al. 2005). Yet, 5 of these genes are also upregulated by overexpression of epe1+ (Table 3). Unexpectedly, overexpression of swi6+ upregulates a larger number of genes than a swi6 mutation (Wiren et al. 2005), and a significant fraction of these genes is upregulated by overexpression of epe1+ (Table 3). Furthermore, 85% of the genes that are upregulated by both overexpression of epe1+ and overexpression of swi6+ are also upregulated by mutations in HDAC genes (Figure 4D). Possible implications of this observation are discussed below.

Epe1 regulates gene expression:

The observation that a high dosage of Epe1 antagonizes the repressive effect of silencing enzymes does not necessarily imply that Epe1 can regulate gene expression under physiological conditions. To address this possibility, we screened 500 clones of a strain collection with random chromosomal insertions of ura4+ in an epe1-1 genetic background for growth on a 5-FOA medium, which counterselects against ura4+ expression (Boeke et al. 1987). We then examined epe1+ and epe1-1 derivatives of the FOA-resistant clones for ura4+ repression by plating serial dilutions of the respective cultures on FOA-supplemented and uracil-depleted media. Three epe1-1 clones were partially resistant to FOA, but their isogenic epe1+ derivatives were FOA sensitive (Figure 5A). The respective ura4+ insertion sites were determined by sequence analysis of flanking regions and compared to the genomewide map of genes that are upregulated by overexpression of epe1+ (Figure 5B).

Figure 5.—

Figure 5.—

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An epe1-1 mutation downregulates the expression of a ura4+ reporter gene at two chromosomal loci. (A) Expression of ura4+ in epe1+ and epe1-1 derivatives of two isolated clones is presented. Expression of ura4+ was determined by serial dilutions of cultures on nonselective medium (N/S), medium lacking uracil (AA-URA), and medium supplemented with FOA (FOA) that counterselects against ura4+ expression. Serial dilutions of ura4+ (UM234) and ura4-D18 (AP182) strains are presented as controls. The locations of ura4+ in the respective clones are indicated in B.

Remarkably, all three insertions were located in close proximity to endogenous genes that are upregulated by overexpression of epe1+. Two were 150 and 200 bp upstream from SPAC23H3.14 (in strains AP1112 and AP1107, respectively), but in opposite orientation to each other. Expression profiling data (supplemental data at http://www.genetics.org/supplemental/) indicate that expression of SPAC23H3.14 is not affected by the epe1 genotype. However, SPAC23H3.15c, which is located within 2 kb from the ura4+ insertion site, is upregulated by overexpression of epe1+ (supplemental data at http://www.genetics.org/supplemental/), as well as by mutations in clr3, clr4, clr6, and dcr1 (Hansen et al. 2005; Wiren et al. 2005). The third insertion (in AP1102) was located 400 bp downstream from SPBC1683.07, within one of the identified clusters (cluster 6, Figure 3A). SPBC1683.07 is upregulated by clr6, clr1, and clr4 mutations (Hansen et al. 2005; Wiren et al. 2005), but not by overexpression of epe1+. However, SPBC1683.06c, which is located 3 kb from the insertion site, is upregulated by overexpression of epe1+ (supplemental data at http://www.genetics.org/supplemental/), as well as by mutations in clr1, ago1, dcr1, and rpd1 (Hansen et al. 2005; Wiren et al. 2005). Notably, all three genes within a distance of 3 kb from this ura4+ insertion (SPBC1683.06c, SPBC1683.07, and SPBC1683.08) are upregulated during meiosis (Mata et al. 2002). These data suggest regional regulation of gene expression by the interplay between silencing proteins and Epe1.

To further explore the possibility that Epe1 plays a role in regulation of gene expression, we asked, how does an epe1∷ura4+ mutation affect the global pattern of gene expression? To this end, we compared RNA expression profiles from cells of isogenic epe1+ and epe1∷ura4+ strains by hybridization to microarrays displaying >99% of all known and predicted ORFs. Analysis of the microarray data indicated that expression of 659 S. pombe genes (12.5%) was downregulated at least twofold by the epe1∷ura4+ mutation (supplemental data at http://www.genetics.org/supplemental/). Next, we compared the list of genes that were downregulated by the epe1 mutation to the lists of genes that were upregulated by overexpression of epe1+ or by mutations in silencing genes (Hansen et al. 2005; Wiren et al. 2005). This analysis revealed a significant overlap between the list of genes that were downregulated by epe1∷ura4+ and the lists of genes that were upregulated by mutations in silencing genes or by overexpression of epe1+ (Table 4). This suggests that, under physiological conditions, some targets are shared by silencing proteins and Epe1. Together with the data on overexpression of epe1+ (Table 3), it suggests that Epe1 can potentially regulate the level of gene expression in a dosage-dependent manner.

The effect of a clr6-1 mutation on the global patterns of gene expression and genomewide acetylation indicated that Clr6 is involved in promoter localized repression by deacetylating intergenic regions (IGRs), upstream from the affected genes (Wiren et al. 2005). To explore the possibility that Epe1 is a potential gene expression regulator at Clr6 targets, we compared the list of genes that were downregulated by epe1∷ura4+ to earlier data on IGR acetylation in a clr6-1 mutant (Wiren et al. 2005). Remarkably, the list of genes that were downregulated by the epe1∷ura4+ mutation overlapped significantly the list of IGRs that were hyperacetylated at histones H3-K9, H4-K5, or H4-K12 in a clr6-1 mutant. These results further substantiate the indications that Epe1 shares gene targets with Clr6 in epe1+ cells. It also suggests that expression from the shared targets may be regulated by the interplay between Clr6 deacetylase and Epe1.

Next we asked whether there is a significant relationship between potential gene regulation by Epe1 and the level of gene expression in logarithmically growing cells. Comparison of the list of genes that are downregulated by an epe1∷ura4+ mutation to earlier expression profiling experiments (Wiren et al. 2005) revealed a highly significant overlap between the lists of low-expression genes and genes that are downregulated by an epe1∷ura4+ mutation. Significant overlaps were also observed between the list of genes that were downregulated by epe1∷ura4+ and the lists of genes that were upregulated in meiosis or in response to environmental stress (Table 4).

DISCUSSION

Epe1 was first identified as a JmjC domain protein that modulates heterochromatization (Ayoub et al. 2003). However, the mechanism underlying Epe1 activity remained elusive. Here we report that Swi6 recruits Epe1 to constitutive heterochromatin domains and that overexpression of epe1+, like mutational inactivation of silencing genes and overexpression of swi6+, upregulates expression of S. pombe genes. We also report that mutational inactivation of Epe1 downregulates low-expression genes and genes that are upregulated in meiosis and stress conditions. The significant overlap between the lists of genes that are upregulated by overexpression of epe1+ and by mutations in HDAC genes, but not by a clr4 mutation, suggests that Epe1 interacts with the heterochromatin assembly pathway at the stage of histone deacetylation. Furthermore, the observation that a significant proportion of the genes that are upregulated by mutations in HDAC genes are downregulated by an epe1 mutation suggests that Epe1 may modulate the repressive effect of HDACs in logarithmically growing cells.

Evidence that Epe1 interacts with the heterochromatin assembly pathway at the stage of histone deacetylation:

Results of earlier genetic analyses were taken as evidence that Epe1 interacts with the heterochromatin assembly pathway upstream of methylation of H3-K9 (Ayoub et al. 2003). However, because of the interdependency of histone modifications, interaction of Epe1 at the stage of histone deacetylation could not be distinguished from interactions at other stages of the pathway. The indication that histone deacetylation is a common way of gene regulation and is mostly independent of H3-K9 methylation (Hansen et al. 2005) provided a handle for examining the effect of Epe1 activity on histone deacetylation, independently of a possible effect at a later stage of the heterochromatin assembly pathway. Indeed, the observation that most genes that are upregulated by both HDAC mutations and overexpression of epe1 are not affected by clr4 mutations suggested that Epe1 antagonizes histone deacetylation independently of the methylated state of H3-K9. The significant overlap between the list of genes that are downregulated by epe1∷ura4+ and IGRs that are highly acetylated in clr6-1 mutants, relative to clr6+ cells, is also consistent with this conclusion.

The mechanism through which Epe1 antagonizes the repressive activity of HDACs is not clear. It may involve direct interaction of Epe1 with the deacetylation machinery. For example, high dosage of the enzymatically inactive JmjC protein Epe1 may interfere with the activity of other histone lysine demethylases that normally act at HDAC targets, leading to reduced histone deacetylation. An effect of enzymatically inactive histone demethylase on deacetylation has been recently reported. The mammalian Lsd1 histone demethylase BHC110 is functionally and physically linked to HDACs, and ectopic expression of an enzymatically dead mutant form of BHC110 displayed decreased deacetylation activity (Lee et al. 2006). Alternatively, Epe1 may affect chromatin structure and organization by promoting transcription activity at or near repressed regions. Evidence for competition between transcriptionally active and inactive chromatin states has been reported in studies of position-effect variegation at subtelomeric regions of Saccharomyces cerevisiae (Aparicio and Gottschling 1994) and at the mat locus of S. pombe (Ayoub et al. 1999). Moreover, Epe1 derepresses transcription activity within constitutive heterochromatin domains (Ayoub et al. 2003; Zofall and Grewal 2006).

A role for Epe1 in gene regulation:

By employing a genetic screen, we identified two chromosomal regions that confer partial silencing on inserted reporter genes in an epe1-1 genetic background, but not in epe1+ cells. Furthermore, endogenous genes in the identified regions are upregulated by overexpression of epe1+ and by mutations in genes that encode heterochromatin assembly proteins. This suggests that, potentially, physiological dosage of Epe1 can regulate gene expression by antagonizing the repressive effect of silencing proteins. Genomewide analyses of gene expression in epe1+ and epe1∷ura4+ strains further substantiate this conclusion: ∼12% of S. pombe genes are downregulated by the epe1∷ura4+ mutation and the list of these genes overlaps significantly the lists of genes that are upregulated by mutations in heterochromatin assembly genes. Furthermore, a significant proportion of genes that are hyperacetylated in a clr6-1 mutant are downregulated by epe1∷ura4+.

The significant correlation between the lists of low-expression genes and genes that are downregulated by epe1∷ura4+ imply that, normally, Epe1 counteracts silencing of repressed genes. A significant correlation was also observed between meiotic or stress-induced genes that are downregulated in rapidly growing cells and genes that are upregulated by Epe1. These observations, together with earlier indications that HDACs play a role in repressing meiotic and stress-induced genes in rapidly growing cells (Hansen et al. 2005; Wiren et al. 2005), suggest that Epe1 provides a fine-tuning mechanism to the repressive effect of HDACs in rapidly growing cells. It is tempting to speculate that Epe1 also plays a role in the escape from the repressed state under stress conditions or nitrogen starvation. However, such a role is not likely to be related to a change in the level of epe1+ expression, as no significant change is observed in response to environmental stress (Chen et al. 2003) and only a minor change is observed late in sporulation (Mata et al. 2002). Thus, if Epe1 does play a role in gene regulation during meiosis or in response to environmental stress, its activity must be controlled by a mechanism that is independent of de novo synthesis, such as post-translational modifications or interaction with other proteins.

Interaction of Swi6 with histone modification proteins and Epe1:

Whereas overexpression of swi6+ enhances gene silencing in constitutive heterochromatin domains (Ayoub et al. 1999; Nakayama et al. 2000), it mainly upregulates gene expression at endogenous euchromatic locations (Wiren et al. 2005). Swi6 binds transiently to heterochromatin and euchromatin, and its binding dynamics are dependent on several parameters, including growth status, Clr4, and Rik1 (Cheutin et al. 2004). Enhancing swi6+ expression is likely to enhance the association of Swi6 with potential chromosomal targets and, subsequently, recruitment of Swi6-associated proteins to these targets. Swi6 recruits at least three proteins that affect chromatin structure: Clr3, Clr4, and Epe1. Accordingly, preferential recruitment of Clr3 and Clr4 would enhance heterochromatization and gene silencing by catalyzing deacetylation of H3-K14acetyl and methylation of H3-K9, respectively. Conversely, preferential recruitment of Epe1 would antagonize heterochromatization and enhance gene expression. While Swi6 recruits all three proteins to constitutive heterochromatin domains, the equilibrium between the recruited proteins favors heterochromatization. However, increased Epe1 dosage upsets this equilibrium and leads to disruption of heterochromatin structure (Ayoub et al. 2003).

The relatively minor effect of swi6 mutations on gene expression profiling (Wiren et al. 2005) suggests that chromatin-mediated repression at euchromatic locations is largely independent of Swi6 activity and that, at normal Swi6 dosage, HDACs do not share many euchromatic targets with Swi6. Furthermore, at high dosage, Swi6 acts as an enhancer of gene expression, rather than as a silencer. The different effects of enhanced swi6+ expression in euchromatic and heterochromatic environments may reflect preferential interactions of Swi6 with different Swi6-associated proteins at different chromosomal locations. This may be linked to a difference in the availability of the respective Swi6-binding proteins or to regulation of Swi6 activity through alternative modifications (Lomberk et al. 2006). Thus, if Swi6 preferentially interacts with Epe1, rather than with Clr3 or Clr4, and this interaction helps direct Epe1 to euchromatic loci, overexpression of Swi6 may have an effect on the pattern of gene expression similar to that of overexpression of Epe1. Other possible mechanisms may involve sequestering of HDACs from their natural acting sites by free Swi6 or causing a stress condition that leads to induction of stress-responsive genes. We note that the observed effect of chromosomal environment on Swi6 activity has interesting precedents. Depending on chromosomal location, HP1, the metazoan homolog of Swi6, forms different complexes with Su(var)3–9 that are associated with specific sets of developmental genes (Greil et al. 2003). Furthermore, HP1 enhances gene silencing at the periphery of heterochromatin domains in a dosage-dependent manner, but at different euchromatic locations it acts either as a gene silencer or as an activator of gene expression (Cryderman et al. 2005).

Acknowledgments

We are grateful to Hiroshi Iwasaki for YA727; Kathleen L. Gould, Nabieh Ayoub, and Jürg Bähler for plasmids; Sigal Ben-Yehuda for advice and support with fluorescence microscopy; and Valerie Wood, Martin Aslett, Juan Mata, Jürg Bähler, Paul Bevan, and Katja Kivinen of the Sanger Center for help in bioinformatics analysis. K.E. is a Royal Swedish Academy of Sciences Research Fellow supported by grants from the Knut and Alice Wallenberg Foundation, Swedish Cancer Society, Swedish Research Council (Vetenskapsrådet), and the EU “Epigenome” network of excellence. This research was supported by a grant from the Israel Science Foundation (438/04).

References

  1. Allshire, R. C., 1997. Centromeres, checkpoints and chromatid cohesion. Curr. Opin. Genet. Dev. 7: 264–275. [DOI] [PubMed] [Google Scholar]
  2. Aparicio, O. M., and D. E. Gottschling, 1994. Overcoming telomeric silencing: a trans-activator competes to establish gene expression in a cell-cycle-dependent way. Genes Dev. 8: 1133–1146. [DOI] [PubMed] [Google Scholar]
  3. Ayoub, N., I. Goldshmidt and A. Cohen, 1999. Position effect variegation at the mating-type locus of fission yeast: a cis-acting element inhibits covariegated expression of genes at the silent and expressed domains. Genetics 152: 495–508. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Ayoub, N., K. Noma, S. Isaac, T. Kahan, S. I. Grewal et al., 2003. A novel jmjC domain protein modulates heterochromatization in fission yeast. Mol. Cell. Biol. 23: 4356–4370. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Boeke, J. D., J. Trueheart, G. Natsoulis and G. R. Fink, 1987. 5-Fluoroorotic acid as a selective agent in yeast molecular genetics. Methods Enzymol. 154: 164–175. [DOI] [PubMed] [Google Scholar]
  6. Chen, D., W. M. Toone, J. Mata, R. Lyne, G. Burns et al., 2003. Global transcriptional responses of fission yeast to environmental stress. Mol. Biol. Cell 14: 214–229. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Cheutin, T., S. A. Gorski, K. M. May, P. B. Singh and T. Misteli, 2004. In vivo dynamics of Swi6 in yeast: evidence for a stochastic model of heterochromatin. Mol. Cell. Biol. 24: 3157–3167. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Cryderman, D. E., S. K. Grade, Y. Li, L. Fanti, S. Pimpinelli et al., 2005. Role of Drosophila HP1 in euchromatic gene expression. Dev. Dyn. 232: 767–774. [DOI] [PubMed] [Google Scholar]
  9. Ekwall, K., J.-P. Javerzat, K. Lorentz, H. Schmidt, G. Cranston et al., 1995. The chromodomain protein Swi6: a key component at fission yeast centromeres. Science 269: 1429–1431. [DOI] [PubMed] [Google Scholar]
  10. Fodor, B. D., S. Kubicek, M. Yonezawa, R. J. O'Sullivan, R. Sengupta et al., 2006. Jmjd2b antagonizes H3K9 trimethylation at pericentric heterochromatin in mammalian cells. Genes Dev. 20: 1557–1562. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Gatti, L., D. Chen, G. L. Beretta, G. Rustici, N. Carenini et al., 2004. Global gene expression of fission yeast in response to cisplatin. Cell. Mol. Life Sci. 61: 2253–2263. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Greil, F., I. van der Kraan, J. Delrow, J. F. Smothers, E. de Wit et al., 2003. Distinct HP1 and Su(var)3-9 complexes bind to sets of developmentally coexpressed genes depending on chromosomal location. Genes Dev. 17: 2825–2838. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Grewal, S. I. S., 2000. Transcriptional silencing in fission yeast. J. Cell. Physiol. 184: 311–318. [DOI] [PubMed] [Google Scholar]
  14. Hall, I. M., G. D. Shankaranarayana, K. Noma, N. Ayoub, A. Cohen et al., 2002. Establishment and maintenance of a heterochromatin domain. Science 297: 2232–2237. [DOI] [PubMed] [Google Scholar]
  15. Hansen, K. R., G. Burns, J. Mata, T. A. Volpe, R. A. Martienssen et al., 2005. Global effects on gene expression in fission yeast by silencing and RNA interference machineries. Mol. Cell. Biol. 25: 590–601. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Hewitson, K. S., L. A. McNeill, M. V. Riordan, Y. M. Tian, A. N. Bullock et al., 2002. Hypoxia-inducible factor (HIF) asparagine hydroxylase is identical to factor inhibiting HIF (FIH) and is related to the cupin structural family. J. Biol. Chem. 277: 26351–26355. [DOI] [PubMed] [Google Scholar]
  17. Irvine, D. V., M. Zaratiegui, N. H. Tolia, D. B. Goto, D. H. Chitwood et al., 2006. Argonaute slicing is required for heterochromatic silencing and spreading. Science 313: 1134–1137. [DOI] [PubMed] [Google Scholar]
  18. Jenuwein, T., and C. D. Allis, 2001. Translating the histone code. Science 293: 1074–1080. [DOI] [PubMed] [Google Scholar]
  19. Karpen, G. H., and R. C. Allshire, 1997. The case for epigenetic effect on centromere identity and function. Trends Genet. 13: 489–496. [DOI] [PubMed] [Google Scholar]
  20. Lee, H. K., W. Braynen, K. Keshav and P. Pavlidis, 2005. ErmineJ: tool for functional analysis of gene expression data sets. BMC Bioinformatics 6: 269. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Lee, M. G., C. Wynder, D. A. Bochar, M. A. Hakimi, N. Cooch et al., 2006. Functional interplay between histone demethylase and deacetylase enzymes. Mol. Cell. Biol. 26: 6395–6402. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Lomberk, G., D. Bensi, M. E. Fernandez-Zapico and R. Urrutia, 2006. Evidence for the existence of an HP1-mediated subcode within the histone code. Nat. Cell Biol. 8: 407–415. [DOI] [PubMed] [Google Scholar]
  23. Martin, C., and Y. Zhang, 2005. The diverse functions of histone lysine methylation. Nat. Rev. Mol. Cell Biol. 6: 838–849. [DOI] [PubMed] [Google Scholar]
  24. Mata, J., R. Lyne, G. Burns and J. Bahler, 2002. The transcriptional program of meiosis and sporulation in fission yeast. Nat. Genet. 32: 143–147. [DOI] [PubMed] [Google Scholar]
  25. Moreno, S., A. J. S. Klar and P. Nurse, 1991. Molecular genetic analysis of fission yeast Schizosaccharomyces pombe. Methods Enzymol. 194: 795–823. [DOI] [PubMed] [Google Scholar]
  26. Nagai, T., K. Ibata, E. S. Park, M. Kubota, K. Mikoshiba et al., 2002. A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications. Nat. Biotechnol. 20: 87–90.11753368 [Google Scholar]
  27. Nakayama, J., A. J. Klar and S. I. Grewal, 2000. A chromodomain protein, Swi6, performs imprinting functions in fission yeast during mitosis and meiosis. Cell 101: 307–317. [DOI] [PubMed] [Google Scholar]
  28. Nakayama, J., J. C. Rice, B. D. Strahl, C. D. Allis and S. I. Grewal, 2001. Role of histone H3 lysine 9 methylation in epigenetic control of heterochromatin assembly. Science 292: 110–113. [DOI] [PubMed] [Google Scholar]
  29. Noma, K., C. D. Allis and S. I. Grewal, 2001. Transitions in distinct histone H3 methylation patterns at the heterochromatin domain boundaries. Science 293: 1150–1155. [DOI] [PubMed] [Google Scholar]
  30. Thon, G., P. Bjerling, C. M. Bunner and J. Verhein-Hansen, 2002. Expression-state boundaries in the mating-type region of fission yeast. Genetics 161: 611–622. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Trewick, S. C., P. J. McLaughlin and R. C. Allshire, 2005. Methylation: Lost in hydroxylation? EMBO Rep. 6: 315–320. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Tsukada, Y., J. Fang, H. Erdjument-Bromage, M. E. Warren, C. H. Borchers et al., 2006. Histone demethylation by a family of JmjC domain-containing proteins. Nature 439: 811–816. [DOI] [PubMed] [Google Scholar]
  33. Verdel, A., and D. Moazed, 2005. RNAi-directed assembly of heterochromatin in fission yeast. FEBS Lett. 579: 5872–5878. [DOI] [PubMed] [Google Scholar]
  34. Volpe, T. A., C. Kidner, I. M. Hall, G. Teng, S. I. Grewal et al., 2002. Regulation of heterochromatic silencing and histone H3 lysine-9 methylation by RNAi. Science 297: 1833–1837. [DOI] [PubMed] [Google Scholar]
  35. Wang, G., A. Ma, C. M. Chow, D. Horsley, N. R. Brown et al., 2000. Conservation of heterochromatin protein 1 function. Mol. Cell. Biol. 20: 6970–6983. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Watson, A., J. Mata, J. Bahler, A. Carr and T. Humphrey, 2004. Global gene expression responses of fission yeast to ionizing radiation. Mol. Biol. Cell 15: 851–860. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Whetstine, J. R., A. Nottke, F. Lan, M. Huarte, S. Smolikov et al., 2006. Reversal of histone lysine trimethylation by the JMJD2 family of histone demethylases. Cell 125: 467–481. [DOI] [PubMed] [Google Scholar]
  38. Wiren, M., R. A. Silverstein, I. Sinha, J. Walfridsson, H. M. Lee et al., 2005. Genomewide analysis of nucleosome density histone acetylation and HDAC function in fission yeast. EMBO J. 24: 2906–2918. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Wood, V., R. Gwilliam, M. A. Rajandream, M. Lyne, R. Lyne et al., 2002. The genome sequence of Schizosaccharomyces pombe. Nature 415: 871–880. [DOI] [PubMed] [Google Scholar]
  40. Xue, Y., S. A. Haas, L. Brino, A. Gusnanto, M. Reimers et al., 2004. A DNA microarray for fission yeast: minimal changes in global gene expression after temperature shift. Yeast 21: 25–39. [DOI] [PubMed] [Google Scholar]
  41. Yamada, T., W. Fischle, T. Sugiyama, C. D. Allis and S. I. Grewal, 2005. The nucleation and maintenance of heterochromatin by a histone deacetylase in fission yeast. Mol. Cell 20: 173–185. [DOI] [PubMed] [Google Scholar]
  42. Yamane, K., C. Toumazou, Y. I. Tsukada, H. Erdjument-Bromage, P. Tempst et al., 2006. JHDM2A, a JmjC-containing H3K9 demethylase, facilitates transcription activation by androgen receptor. Cell 125: 483–495. [DOI] [PubMed] [Google Scholar]
  43. Zardo, G., F. Fazi, L. Travaglini and C. Nervi, 2005. Dynamic and reversibility of heterochromatic gene silencing in human disease. Cell Res. 15: 679–690. [DOI] [PubMed] [Google Scholar]
  44. Zofall, M., and S. I. Grewal, 2006. Swi6/HP1 recruits a JmjC domain protein to facilitate transcription of heterochromatic repeats. Mol. Cell 22: 681–692. [DOI] [PubMed] [Google Scholar]

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