Dual chromatin recognition by the histone deacetylase complex HCHC is required for proper DNA methylation in Neurospora crassa

Shinji Honda; Vincent T. Bicocca; Jordan D. Gessaman; Michael R. Rountree; Ayumi Yokoyama; Eun Y. Yu; Jeanne M. L. Selker; Eric U. Selker

doi:10.1073/pnas.1614279113

Contributed by Eric U. Selker, August 26, 2016 (sent for review May 5, 2016; reviewed by Sarah Elgin and Mo Motamedi)

Significance

Modifications of chromatin proteins (e.g. histones) and DNA play vital roles in genome function. Both hypo- and hypermethylation of DNA are associated with human diseases, including cancers, but the underlying mechanisms are not well understood. Using the filamentous fungus Neurospora crassa, one of the simplest eukaryotes with DNA methylation, we report a DNA methylation pathway that depends partially on the histone deacetylase complex HCHC [heterochromatin protein 1 (HP1)–chromodomain protein 2 (CDP-2)–histone deacetylase 1 (HDA-1)– CDP-2/HDA-1–associated protein (CHAP)]. Genome-wide DNA methylation analyses revealed both hypo- and hyper-DNA methylation in strains with defective HCHC components. We show the interrelationship of HCHC components and genetically dissect the proteins to define domains critical for proper DNA methylation and centromeric silencing. This work provides insights into the crosstalk between DNA methylation and histone modifications.

Abstract

DNA methylation, heterochromatin protein 1 (HP1), histone H3 lysine 9 (H3K9) methylation, histone deacetylation, and highly repeated sequences are prototypical heterochromatic features, but their interrelationships are not fully understood. Prior work showed that H3K9 methylation directs DNA methylation and histone deacetylation via HP1 in Neurospora crassa and that the histone deacetylase complex HCHC is required for proper DNA methylation. The complex consists of the chromodomain proteins HP1 and chromodomain protein 2 (CDP-2), the histone deacetylase HDA-1, and the AT-hook motif protein CDP-2/HDA-1–associated protein (CHAP). We show that the complex is required for proper chromosome segregation, dissect its function, and characterize interactions among its components. Our analyses revealed the existence of an HP1-based DNA methylation pathway independent of its chromodomain. The pathway partially depends on CHAP but not on the CDP-2 chromodomain. CDP-2 serves as a bridge between the recognition of H3K9 trimethylation (H3K9me3) by HP1 and the histone deacetylase activity of HDA-1. CHAP is also critical for HDA-1 localization to heterochromatin. Specifically, the CHAP zinc finger interacts directly with the HDA-1 argonaute-binding protein 2 (Arb2) domain, and the CHAP AT-hook motifs recognize heterochromatic regions by binding to AT-rich DNA. Our data shed light on the interrelationships among the prototypical heterochromatic features and support a model in which dual recognition by the HP1 chromodomain and the CHAP AT-hooks are required for proper heterochromatin formation.

DNA methylation is an epigenetic mechanism involved in fundamental biological processes such as transcriptional regulation, genome defense, X chromosome inactivation, and genomic imprinting (1 ⇓⇓–4). In mammals, patterns of DNA methylation are established during embryonic development and are maintained during subsequent cell divisions (5). Abnormal DNA methylation is associated with human disease, including cancer (6, 7), but the events leading to abnormal DNA methylation are not well understood. A full understanding of aberrant methylation will first require a more complete understanding of normal methylation. Nevertheless, revelations during the last decade have provided clues to guide further research. Most importantly, studies in fungi, plants, and animals have revealed that histone modifications and RNA signals can influence, if not outright control, DNA methylation (8 ⇓–10).

Research using the filamentous fungus Neurospora crassa first revealed a link between DNA methylation and histone H3 lysine 9 (H3K9) methylation, which is a molecular hallmark of constitutive heterochromatin (11). Subsequent genetic and biochemical studies uncovered a direct pathway from H3K9 methylation to DNA methylation. The DIM-5 (defective in methylation-5) lysine methyltransferase (KMT) catalyzes trimethylation of H3K9 (H3K9me3) (12), which is recognized and bound by the chromodomain (CD) of heterochromatin protein 1 (HP1) (13). The DNA methyltransferase (DNMT) DIM-2 is directly recruited by HP1 through the chromoshadow domain of HP1 and two PxVxL-like motifs in DIM-2 (14).

In N. crassa, H3K9me3, HP1, and DNA methylation are colocalized and together define the regions of constitutive heterochromatin (15). The centromere regions, generally rich in transposon relics, account for the largest regions of constitutive heterochromatin, but telomeres and interstitial islands of transposon relics also have some features of heterochromatin. Most of these regions are A:T-rich as a result of the genome defense system RIP (repeat-induced point mutation). RIP detects duplicated sequences and induces G:C to A:T mutations in these regions during the sexual phase of the N. crassa life cycle (15 ⇓–17). The resulting A:T-rich sequences serve as potent signals for triggering H3K9me3 and DNA methylation de novo (15, 18, 19). We identified DIM-5, DIM-7, DIM-9, CUL4 (cullin 4), and DDB1 (DNA damage-binding protein 1) as components of a KMT complex, DCDC (DIM-5/-7/-9–CUL4–DDB1 complex), which is required for H3K9me3 and appears to operate by a two-step mechanism: DIM-7–dependent DIM-5 recruitment and CUL4/DDB1/DIM-9–dependent catalysis by the KMT DIM-5 (20, 21).

We previously identified a histone deacetylase (HDAC) complex, HCHC, which contains HP1, CD protein 2 (CDP-2), the HDAC HDA-1, and a CDP-2/HDA-1–associated protein, CHAP (22). The HCHC complex works in parallel with the DNMT complex DIM-2–HP1 to establish and maintain normal heterochromatin. In addition, the HCHC complex indirectly maintains proper DNA methylation at regions with moderate and heavy mutation by RIP, which respectively show hypo- and hypermethylation in cdp-2, hda-1, or chap mutants (22). Here we describe detailed functional interrelationships and domain functions of the components of the HCHC complex.

Results

HCHC Plays an Important Role in Centromere Function.

We previously demonstrated that mutants lacking HP1, but not DIM-2, exhibit sensitivity to the microtubule inhibitor thiabendazole (TBZ) and the topoisomerase I inhibitor camptothecin (CPT) and suffer from chromosome segregation defects (20). Because HP1 is present in both the HCHC and DIM-2/HP1 complexes (22) and because centromere regions are hypermethylated in mutants defective in components of HCHC (22), we wished to test if mutants lacking other components of HCHC show these hpo (HP1 gene) phenotypes. We found that the hda-1, cdp-2, and chap mutants did not display sensitivity to CPT comparable to that observed for the hpo mutant (Fig. 1A), suggesting that HP1 has functions other than its role in the HCHC and HP1–DIM-2 complexes. Like hpo strains, mutants lacking HDA-1 exhibited strong sensitivity to TBZ, whereas cdp-2 and chap mutants showed an intermediate level of TBZ sensitivity (Fig. 1A). Like hpo strains but unlike dim-2 strains, all the HCHC mutants showed numerous chromosome bridges (Fig. 1B) (20). These findings fit with our prior observation that HCHC mutants show centromeric silencing defects (22) and strengthen the conclusion that HCHC is important for centromere function.

Fig. 1.

HCHC is important for centromere function. (A) Serial dilutions of conidia from each of the indicated strains were spot-tested on medium with or without TBZ or CPT. Strains: N3753, N4922, 775, 903, and 949. (B) The distribution of the nuclear marker H2A-GFP in growing hyphae in wild-type, cdp-2, hda-1, and chap strains were examined microscopically. The frequency of observed chromatin bridges and the total number of nuclei are shown beside each representative micrograph. Strains: N5015, N5017, N5024, and N5026.

Whole-Genome Bisulfite Sequencing Analysis of the HCHC Mutants.

DNA methylation of HCHC mutants was previously assessed in selected genomic regions by Southern hybridization and methylated DNA immunoprecipitation analyses (15). To extend our understanding of the role of HCHC, we carried out whole-genome bisulfite sequencing (WGBS) of cdp-2, chap, and hda-1 mutants. Consistent with our prior analyses (22), a heat map display of the WGBS data revealed both hypomethylated and hypermethylated regions in the three HCHC mutants (Fig. S1). In addition, when methylated regions were sorted by increasing size, we found that shorter regions, which are generally heavily methylated in wild-type HCHC (average of 47.6% of C residues are methylated in the 50 shortest regions), tend to show significantly reduced methylation in the HCHC mutants (averages of 13.9, 14.8, and 22.7% in cdp-2, hda-1, and chap mutants, respectively) (Fig. 2A). Conversely, longer regions, most notably centromeres, are more lightly methylated in wild-type strains (average of 25.9% in the longest 50 regions) but tend to show moderately more methylation in the HCHC mutants (average of 30.7, 32.0, and 35.0% in hda-1, cdp-2, and chap mutants, respectively) (Fig. 2 A and B). Sequences near telomeres that are normally methylated were found to lose methylation in the mutants (Fig. 2B and Fig. S1). In addition, sequences with a low combined RIP index (CRI) (17) tend to show reduced methylation in the mutants, whereas sequences with higher CRI scores show increased methylation (Fig. 2C). The borders of normally methylated regions typically lose methylation and show a contraction of boundary methylation in the HCHC mutants (Fig. 2D).

Fig. 2.

WGBS analysis of HCHC mutants. (A) Heat map analyses showing the relative level of 5mC for all methylated regions, sorted from shortest to longest, for wild-type strains and the HCHC mutants. (B) Heat map analyses showing the relative level of 5mC at centromeres and methylated regions within 100 kb of telomeres for wild-type strains and HCHC mutants. (C) The CRI (x axis) and average methylation level (y axis) were calculated for 500-bp windows across the genome (Materials and Methods) and then were plotted for wild-type strains and each of the HCHC mutants. A mutant lacking HP1 (Δhpo) is used as a control for complete loss of DNA methylation. (D) Average methylation levels were calculated for 50-bp windows across the borders of methylated regions (Materials and Methods) and then were plotted for wild-type strains and HCHC mutants. Strains: N3752, N3615, N3612, N3435, and N4922.

Fig. S1.

WGBS profiles for WT and HCHC mutants. The average 5mC levels were calculated for 500-bp windows across the genome from the WGBS data derived for wild-type strains and the HCHC mutants (Materials and Methods) and displayed by the Integrative Genomics Viewer using the heatmap function. Genes (green vertical lines) are displayed on the x axis below the DNA methylation heatmap profiles. Methylated regions called by RSEG (Materials and Methods) are indicated by black bars above the wild-type row. N. crassa’s seven linkage groups are not displayed to scale. Strains: N3752, N3615, N3612, and N3435.

The CD of HP1 but Not That of CDP-2 Is Required for HCHC Complex Function.

We previously demonstrated that the CD of CDP-2 efficiently binds to H3K9me3 in vitro (22). Considering that the HCHC complex harbors two proteins containing this domain, we wished to investigate the possibility that the chromodomains might be partially or fully redundant. To do so, we generated constructs to produce epitope-tagged proteins bearing mutations changing or deleting residues known to be critical for the CD function (CDP-2^W446G, CDP-2^ΔCD, and CDP-2^1-419) (Fig. 3A). All constructs were tagged with a C-terminal 3×FLAG epitope, and expression was driven by the native cdp-2 promoter. We inserted the constructs at the his-3 locus in a cdp-2 mutant strain and confirmed that their expression levels were all comparable to that in the wild-type strain (Fig. S2A). Curiously, all the cdp-2 constructs bearing CD mutations rescued the cdp-2 defects, namely region-specific hypomethylation (Fig. 3B), global hypermethylation (e.g., at centromere regions) (Fig. S2B), and derepression of centromere silencing (Fig. S2C), suggesting that the CD of CDP-2 is dispensable.

Fig. 3.

The HP1 CD but not the CDP-2 CD is required for normal DNA methylation. (A) Diagram of the CDP-2 mutations tested. Tryptophan 446 is predicted to be an aromatic cage residue essential for binding to H3K9me (49, 50). (B) Introduction of CDP-2 CD mutant constructs complements DNA hypomethylation defects at the 8:A6 methylated region in cdp-2–null mutants. Genomic DNA of the indicated strains was digested with 5mC-sensitive BfuCI (B) or its 5mC-insensitive isoschizomer DpnII (D) and was gel-fractionated, and DNA methylation was analyzed by Southern hybridizations with the 8:A6 probe, a region that is methylated in wild-type strains (17). The positions of size standards are shown at left. Strains: N150, N1877, N3615, N4006, N3992, N4088, and N4089. (C) Diagram of the HP1 CD mutation tested. (D) The introduction of the HP1 CD-deletion gene partially restores DNA methylation at the 8:A6 methylated region. Strains: N3753, N5580, N6166, and N6390. (E) The residual DNA methylation of the 8:A6 region in the HP1 CD-deletion mutation is not affected by the CDP-2 CD deletion. Strains: N3753, N5580, N3615, N6390, N6393, and N6394.

Fig. S2.

The CDP-2 CD is dispensable for normal HCHC function. (A) Expression of cdp-2 mutants was assessed by Western blotting with antibodies against FLAG. β-Tubulin was used as a loading control. Strains: N3343, N3992, N4088, and N4089. (B) Effects of cdp-2 mutations on global DNA methylation by ethidium bromide (EtBr) staining. Genomic DNA of indicated strains was digested with 5mC-sensitive BfuCI (B) or its 5mC-insensitive isoschizomer, DpnII (D) and were gel-fractionated, and the products were visualized by EtBr straining. The positions of size standards are shown at left. The enhanced, more slowly migrated DNA (∼10 kb) in cdp-2–null mutants indicates hypermethylation, but insertion of the CDP-2 CD mutants complements the methylation defects. Strains: N623, N1877, N3135, N3343, N3992, N4088, and N4089 (C) Effects of cdp-2 mutations on centromere silencing. Serial dilutions of conidia from each of the indicated strains harboring a centromeric bar construct were spot-tested on medium with or without basta. Strains: N4890, N4891, N4915, 888, 890, 892, and 894.

These findings did not eliminate the possibility that the CDP-2 CD has a redundant function, perhaps with HP1. We therefore generated C-terminal HAT-FLAG epitope-tagged HP1 constructs, similar to those created for CDP-2, that contain mutations changing or deleting residues critical for HP1 CD function (hpo^W98G and hpo^ΔCD) (Fig. S3A). These hpo promoter-driven constructs were inserted at the his-3 locus in a Δhpo mutant strain. DNA methylation was entirely lost in the Δhpo transformation host strain but was fully restored with the his-3 targeted hpo^WT construct (Fig. S3B). Surprisingly, both the HP1 CD mutant constructs, hpo^W98G and hpo^ΔCD, restored a low level of DNA methylation, indicating that the CD region is not absolutely required for DNA methylation. Similarly, these CD mutants partially alleviated sensitivity to TBZ (Fig. S3C). However, the mutants still showed sensitivity to CPT and defective centromeric silencing (Fig. S3 C and D).

Fig. S3.

The HP1 CD is required for centromeric silencing and for most but not all DNA methylation. (A) Diagram of HP1 and the CD mutation (tryptophan 98 to glycine) and deletion tested. Tryptophan 98 is predicted to be essential for binding to H3K9me (43, 44). (B) Southern analysis of the normally methylated 8:A6 region (17) demonstrating that insertion of either the hpo CD point mutation (lanes 4 and 6) or the hpo CD deletion (lanes 5 and 7) results in a partial restoration of the methylation lost at 8:A6 in a hpo-deletion strain (lane 2), whereas insertion of the wild-type hpo allele (lane 3) restores 8:A6 methylation to levels seen in a wild-type strain (N4909, lane 1). Genomic DNAs were digested with the 5mC-sensitive BfuCI (B) restriction enzyme or its 5mC-insensitive isoschizomer, DpnII (D). The positions of size standards are shown at left. Strains: N4909, N4922, N5898, N5869, N5870, N5871, and N5872. (C and D) Strains harboring the hpo CD mutation (hpo^W98G) or CD deletion (hpo^∆CD) slightly restore TBZ sensitivity but fail to restore CPT sensitivity or to silence the cenVIR::bar marker properly. Serial dilutions of conidia from each of the indicated strains were spot-tested on medium with the indicated drugs. Strains: N4890, N4922, N5898, N5869, and N5871.

The CDP-2 and HP1 CD mutations described above were expressed at the his-3 locus. To test the effect of simultaneously compromising the chromodomains of both CDP-2 and HP1, we inserted the CD-deletion constructs at their respective endogenous loci (Fig. 3). The new hpo constructs were tagged with the LexA DNA-binding domain (LexADBD) epitope at the C terminus. We confirmed that the strain carrying the tagged hpo^WT construct exhibited normal DNA methylation, indicating that the tagged protein was functional (Fig. 3D). As in our results described above (Fig. S3B), we found that deletion of the HP1 CD, unlike the loss of the whole protein (13), resulted in reduced DNA methylation rather than a complete loss of DNA methylation (Fig. 3D). Deletion of the CDP-2 CD did not accentuate the DNA methylation defect of the HP1 CD mutant (Fig. 3E), suggesting that, despite its ability to bind methylated H3K9 in vitro (22), the CDP-2 CD does not act redundantly with the HP1 CD.

CDP-2 Interacts Directly with the Chromoshadow Domain of HP1.

To gain insights into how HCHC operates without the CDP-2 CD, we investigated the organization of the HCHC subunits and tested the function of their prominent domains (22). We performed a yeast two-hybrid assay and found that HP1 and CDP-2 interact directly and that this interaction requires the chromoshadow domain of HP1 (Fig. 4A). CDP-2 did not interact with the HP1^Y244E mutant, which has a single amino acid substitution in the HP1 chromoshadow domain that prevents both dimerization of HP1 and interaction with the DIM-2 tandem PxVxL-like motifs (14, 23). To determine which CDP-2 sequences interact with the HP1 chromoshadow domain, we generated and tested a series of CDP-2 fragments. We found that amino acids 8–24 of CDP-2 are sufficient for binding HP1 (Fig. S4A). Inspection of the CDP-2 sequence revealed a PxVxL-like motif, (I/F/V)x(I/V)x(I/L/V), at amino acids 14–18 that is conserved among filamentous fungi (Fig. S4B). We generated and tested a mutant construct with alanines substituted for conserved residues at 14 and 15 (IE/AA) in the motif and found that the change abolished interaction with HP1 (Fig. 4B and Fig. S4A). To verify the specific interaction, we deleted a second PxVxL-like motif, ΔPPITL, found at amino acids 33–37, adjacent to the first PxVxL-like motif, and confirmed that it did not abolish the interaction (Fig. S4A). We next created the corresponding CDP-2 mutant strains and tested this interaction in vivo. Each protein included a 3×FLAG epitope tag, and expression was driven by the endogenous promoter. In line with our yeast two-hybrid results, coimmunoprecipitation (co-IP) experiments revealed that the IE/AA mutation abolished interaction with HP1 (Fig. 4C). Interestingly, the ΔPPITL mutation also eliminated interaction with HP1 in vivo even though it did not in vitro (Fig. 4C and Fig. S4A), suggesting CDP-2 might be similar to DIM-2 in requiring tandem PxVxL-like motifs to interact with HP1 (14). Together, these results support a model in which the N-terminal fragment of CDP-2 interacts directly with the chromoshadow domain of HP1.

Fig. 4.

HCHC component interactions. (A) Summary of yeast two-hybrid results for HCHC components expressed pairwise in yeast as galactose (Gal)-binding domain fusions (pGBDU) or Gal-activation domain fusions (pGAD). The indicated constructs were cotransformed into the PJ69-4A yeast cells. Transformants were tested on synthetic defined (SD) agar plates without adenine, histidine, leucine, or uracil; growth results are shown at left. (B) Diagram summarizing interactions between components of the HCHC complex. Detailed analyses are presented in Fig. S4. The chromoshadow domain of HP1 interacts with the most N-terminal PxVxL-like motif of CDP-2. An adjacent PxVxL-like motif on CDP-2 interacts with the HDAC domain of HDA-1. The Arb2 domain of HDA-1 interacts with the first zinc-finger motif of CHAP. (C and D) Verification of the interaction of CDP-2 with HP1 and HDA-1 via the N-terminal PxVxL-like motifs of CDP-2 in vivo. Co-IP experiments were performed with anti-FLAG antibodies in strains with the indicated tagged proteins. Input and immunoprecipitation samples were fractionated and analyzed by Western blotting with antibodies against the indicated epitopes. The asterisk indicates nonspecific bands. Strains: N3808, N3836, 3440, 3443, 3445, and 3447.

Fig. S4.

Yeast two-hybrid analyses among truncated or mutated components of HCHC. (A) Schematic diagram of tested truncated or mutated derivatives of CDP-2 fused to the Gal DNA-binding domain. All constructs were cotransformed with pGAD-HP1 prey-vector into the PJ69-4A yeast cells (51). (B) Sequence comparison of the HP1-interacting fragment of CDP-2 from N. crassa (Nc) and the corresponding region of its counterparts in Chaetomium globosum (Cg, accession number XP_001220390), Podospora anserina (Pa, XP_001912830), Gibberella zeae (Gz, XP_385206), and Magnaporthe grisea (Mg, XP_001414497). The residue identical in all CDP-2 proteins is indicated by an asterisk. Colons and periods indicate strong and weak conservation, respectively. Residues similar to the PxVxL consensus sequence implicated in HP1 binding are shown in white on a black background. Residues replaced by alanines (IE/AA) are indicated above. (C–F) Schematic diagrams of tested truncated or mutated derivatives of CDP-2, HDA-1, and CHAP fused to the Gal DNA-binding domain.

CDP-2 and CHAP Interact Directly with HDA-1.

We further used the yeast two-hybrid assay to ask how other components of HCHC interact. We found that, in addition to interacting with HP1, CDP-2 interacts with HDA-1 but not with CHAP (Fig. 4A). Similarly, HDA-1 interacts with CHAP but not with HP1, and CHAP does not interact directly with HP1 (Fig. 4A). To identify the protein regions involved in the interactions, we generated and tested a series of CDP-2, HDA-1, and CHAP fragments. The experiments revealed that amino acids 24–54 of CDP-2 are sufficient for binding HDA-1 and that deletion of the PxVxL-related motif PPITL (amino acids 33–37) abolished its interaction with HDA-1 (Fig. 4B and Fig. S4C). In vivo co-IP analysis of this interaction, using the IE/AA and ΔPPITL constructs described above, revealed that only the PPITL motif is required for CDP-2’s interaction with HDA-1 and that the IE/AA mutation had no effect on their interaction (Fig. 4D). Further yeast two-hybrid analysis showed amino acids 87–474 of the HDA-1 HDAC domain were sufficient for binding CDP-2 (Fig. 4B and Fig. S4D), and amino acids 478–744 of the HDA-1 Arb2 (argonaute-binding protein 2) domain are sufficient for its direct interaction with CHAP (Fig. 4B and Fig. S4E). To test whether the AT-hook and zinc finger motifs of CHAP are involved in the interaction between HDA-1 and CHAP, we made CHAP point mutants [first AT-hook (ATh1); second AT-hook (ATh2); first zinc finger (Zf1); and second zinc finger (Zf2)] (Fig. S5A) and found that the Zf1 of CHAP is important for its interaction with HDA-1 (Fig. 4B and Fig. S4F). Unfortunately, we were unable to validate the roles of CHAP’s zinc fingers in vivo, because mutant constructs produced unstable CHAP protein (Fig. S5B). The yeast two-hybrid interactions of all the components of HCHC are summarized in Fig. 4B.

Fig. S5.

Sequence alignment of N. crassa CHAP with homologs from other filamentous fungi and de-stabilization of the CHAP zinc finger motif mutant proteins. (A) Proteins were aligned with the Clustal X algorithm. Highly and partially conserved residues are indicated by asterisks and colons, respectively. The predicted AT-hook and zinc finger motifs are indicated by thick lines. The residues that are conserved among species and are critical resides (the arginine residue for the AT-hook motif and the cysteine residue in the zinc finger motif) were changed to alanine (ATh1, R210A; ATh2, R250A; Zf1, C280A; Zf2, C317A). The accession numbers of the proteins are Chaetomium globosum, XP_001225541; Podospora anserina, XP_001909682; Magnaporthe grisea, XP_370514; Gibberella zeae, XP_384555; Sclerotinia sclerotiorum, XP_001585520; and Botryotinia fuckeliana, XP_001554742. (B) The CHAP zinc finger motif mutant proteins are unstable. Expression of chap mutants was assessed by Western blotting with antibodies against HA. β-Tubulin was used as a loading control. Strains: N3818, N3825, N3827, and N3846.

CDP-2 Recruits HDA-1 HDAC Activity to H3K9me3 Regions.

Based on the interaction map among the HCHC components (Fig. 4B), we hypothesized that CDP-2 might simply serve as a tether between HP1 and HDA-1, facilitating deacetylation of histones marked with H3K9me3. To test this idea, we first carried out co-IP experiments on extracts from strains with epitope-tagged HDA-1 and HP1 in cdp-2–null mutant strains and found that interactions between HDA-1 and HP1 indeed depend on CDP-2 (Fig. S6A). Interestingly, the interaction between HDA-1 and HP1 occurred in a dim-5 mutant, indicating that formation of the HCHC complex can occur before HP1 binding to H3K9me3 (Fig. S6A). To elucidate further the role of CDP-2 in tethering, we performed DamID (DNA adenine methyltransferase identification) by generating wild-type and cdp-2 mutant strains expressing HDA-1–Dam. DamID uses DpnI (which cuts specifically at adenine-methylated GATC sites) and DpnII (which cuts at GATC sites without adenine methylation) to assess adenine methylation catalyzed by Dam fusion proteins and can provide information on the genomic localization of proteins that are not detectable by ChIP (21, 24). At all heterochromatic regions tested, expression of HDA-1–Dam in a wild-type background produced low- and some intermediate-molecular-weight DpnI fragments, indicating that HDA-1–Dam localized to heterochromatin (Fig. 5A). Little DpnI digestion was detected at the euchromatic pan-1 gene. Compared with a wild-type strain, HDA-1–Dam localization to heterochromatin was reduced in cdp-2 strains, providing further evidence that CDP-2 is required for proper targeting of HDA-1 (Fig. 5A).

Fig. 5.

HCHC function depends on the HDA-1 HDAC. (A) Sequence-dependent localization of HDA-1–Dam depends on CDP-2 and CHAP. Genomic DNA from a wild-type strain with (+) or without (−) HDA-1–Dam, as well as wild-type, cdp-2, and chap strains expressing HDA-1–Dam, were incubated with (+) or without (−) DpnI, which cuts adenine-methylated GATC sites. As a control for completely digested DNA, genomic DNA from the wild-type strain was incubated with the 5mC-insensitive isoschizomer DpnII. Digested DNA was used for Southern hybridizations with probes for the methylated regions 8:A6, 8:G3, and cenVIIR as well as a euchromatic gene, pan-1. Strains: N3752, N3995, N4023, and N4082. (B) The position of the point mutation in the HDA-1 catalytic domain. Asparagine 263 is predicted to be essential for the HDAC activity (25). (C) The introduction of the hda-1 gene with the catalytic mutation does not complement DNA methylation defects in hda-1–null mutants. The experiment was carried out as described in Fig. 3B with the 8:A6 methylated region (17). Strains: N623, N1877, N3610, N3997, and N3998. (D and E) The HDA-1 catalytic mutation does not disrupt the HCHC complex. Co-IP experiments were performed with anti-FLAG antibodies in strains with (+) or without (−) the indicated tagged proteins. Input and immunoprecipitation samples were fractionated and analyzed by Western blotting with antibodies against the indicated epitopes. Strains: N3321, N4002, N4043, N3377, N4000, and N4699.

Fig. S6.

Co-IP analyses among each component of the HCHC complex in vivo. Extracts from the indicated tagged strains were immunoprecipitated with anti-HA or anti-FLAG antibodies. Input and immunoprecipitation samples were fractionated and analyzed by Western blotting with antibodies against the tagged proteins. The asterisk indicates nonspecific bands, and the arrow indicates a specific band. Strains: (A) N3704, N3735, N3733, and N3720; (B) N3836, N3835, N3838, and N3839; (C) N0808, N3852, N3853, and N3802; (D) N3803, N3842, N3843, and N3719; (E) N3805, N3800, N3717, and N3804; (F) N3730, N3806, N3841, and N3728.

In principle, the essential role of HDA-1 in HCHC may or may not depend on HDAC activity. To distinguish between these possibilities, we generated an hda-1 construct with a point mutation causing an amino acid substitution of a residue critical for HDAC activity, HDA-1^D263N (Fig. 5B) (25), and with a 3×FLAG epitope tag at the C terminus of the protein. The construct was driven by the native hda-1 promoter and was inserted at the his-3 locus of an hda-1 mutant. Insertion of a wild-type control hda-1–FLAG construct restored nearly normal patterns of DNA methylation, indicating that the HDA-1–FLAG fusion was functional (Fig. 5C). In contrast, HDA-1^D263N–FLAG failed to complement the methylation defects (Fig. 5C), even though it was expressed as well as wild-type HDA-1–FLAG (Fig. 5 D and E). Co-IP experiments verified that the mutation did not affect the stable formation of HCHC (Fig. 5 D and E), implying that HDAC activity is required for the HCHC function. Taking these results together with the previous observation that mutants lacking CDP-2 show striking hyperacetylation of histones H3 and H4 at heterochromatic regions (22), we conclude that CDP-2 serves as a bridge between HP1 and HDA-1 to recruit HDAC activity to methylated H3K9 regions.

HP1 and CDP-2 Localize to Heterochromatin Independently of HDA-1 and CHAP.

To investigate further the extent to which recruitment of HP1 and CDP-2 may depend on other members of the complex, we generated hda-1 and chap mutants expressing HP1–GFP or CDP-2–GFP and examined localization of the GFP-tagged proteins by microscopy. Because CDP-2 is destabilized in the absence of other components of the complex (Fig. S6 B–D), it was necessary to overexpress CDP-2 to test if its punctate localization depends on the other HCHC components. The nuclear foci that characterize normal HP1–GFP localization were lost when H3K9me3 was abrogated in a dim-5 mutant but were evident in cdp-2, hda-1, and chap mutants (Fig. S7A), consistent with the model that HP1 recruits CDP-2, HDA-1, and CHAP to chromatin marked with H3K9me3. Similarly, CDP-2 localization was unaltered in hda-1 and chap mutants but was dependent on HP1 and H3K9me3 (Fig. S7B). Furthermore, using the DamID assay, we verified that CDP-2 localization to heterochromatin depends on HP1 but not on HDA-1 and CHAP (Fig. S7C). These data suggest that CDP-2 is important for tethering HP1 to HDA-1/CHAP, as is consistent with our interaction map (Fig. 4B).

Fig. S7.

HP1 and CDP-2 localize to heterochromatin independently of HDA-1 and CHAP. (A) Punctate localization of HP1–GFP is independent of other HCHC components. Conidia of the indicated strains were examined microscopically using visible light (DIC) or UV fluorescence. Strains: N3321, N3431, N3649, N3757, and N3759. (B) CDP-2–GFP localization is independent of HDA-1 and CHAP. Strains: N3790, N3792, N3794, N3911, and N3913. (C) CDP-2 localization depends on HP1 but not on HDA-1 and CHAP. Genomic DNA from the wild-type strain, which does not express HDA-1–Dam, and wild-type, hpo, hda-1, and chap strains which express CDP-2–Dam were incubated with (+) or without (−) DpnI, which cuts adenine-methylated GATC sites. As a control for completely digested DNA, genomic DNA from the wild-type strain was incubated with its 5mC-insensitive isoschizomer DpnII. Digested DNA was used for Southern hybridizations with probes for the methylated regions 8:A6, 8:G3, and cenIVR and the euchromatic gene pan-1. Strains: N150, N4011, N4013, N4083, and N4085.

CHAP Is Required for the Residual DNA Methylation in the HP1 CD-Deletion Strain.

We next characterized the role of CHAP in vivo and found that mutants lacking CHAP had unstable interactions with the other HCHC components (Fig. S6 A–C) and reduced HDA-1–Dam localization to heterochromatin (Fig. 5A), as is consistent with importance of CHAP for histone deacetylation at heterochromatic regions (22). We also found that the stability of CHAP and its localization to DNA with repeat-induced point mutations were dependent on the other components of HCHC (Fig. S6 D–F and Fig. S8A). Given the similar residual DNA methylation in the HP1 CD mutant and in chap-null mutant strains (Fig. 6), we hypothesized that chromatin binding of the HCHC complex may rely partly on CHAP. We therefore tested a strain with both the HP1 CD deletion and a chap deletion and found that it showed complete loss of DNA methylation at regions that lose methylation in the hda-1 mutant (8:A6, 8:G3, and 2:B3) (22) but did not lose methylation at a region unaffected by hda-1 (8:F10) (Fig. 6). Taken together, these data provide evidence that CHAP, in conjunction with the CD of HP1, facilitates the recruitment of HCHC to some heterochromatic regions.

Fig. 6.

CHAP is essential for the residual DNA methylation in the HP1 CD mutant. Southern blot analysis was carried out as in Fig. 3B. The three upper panels are the hypomethylated 8:A6, 8:G3, and 2:B3 regions, and the bottom panel is the intact methylated 8:F10 region in hda-1–null mutants. Mutants lacking the HP1 CD or CHAP show the residual DNA methylation at the 8:A6, 8:G3, and 2:B3, whereas the double mutants show complete loss of DNA methylation. DNA methylation at the 8:F10 region is unchanged in mutants lacking the HP1 CD and/or CHAP. Strains: N3753, N5580, N6166, N6390, N6392, and N6391.

Fig. S8.

CHAP localization to heterochromatin depends on the other components of HCHC. (A) Sequence-dependent localization of CHAP–Dam depends on HDA-1 and CDP-2. DNA isolated from the indicated strains was analyzed by Southern hybridizations as in Fig. 3B. Strains: N4045, N4687, and N4690. (B) Expression of chap AT-hook motif mutants was assessed by Western blotting with antibodies against HA. β-Tubulin was used as a loading control. Strains: N3752, N3818, N3820, N3822, and N3845. (C) Normal HCHC complex formation in the CHAP AT-hook motif mutants. Co-IP experiments were performed with anti-HA antibodies in strains with (+) or without (−) the indicated tagged proteins. Input and immunoprecipitation samples were fractionated and analyzed by Western blotting with antibodies against the indicated epitopes. Strains: N3319, N3730, and N3986.

The CHAP AT-Hook Motifs Are Required for Normal DNA Methylation.

To evaluate the possible role of the AT-hooks of CHAP, we generated a series of constructs with point mutations to change critical residues in these motifs and with a 3×HA epitope tag at the C terminus of the protein (CHAP–HA) (Fig. 7A). All constructs were driven by the native chap promoter and were inserted at the pan-2 locus of a chap-deletion strain. We confirmed that mutations in the AT-hook motifs (CHAP^ATh1, CHAP^ATh2, and CHAP^ATh1&2) did not affect the level of CHAP protein (Fig. S8B) and that insertion of a wild-type chap–HA construct into a chap-deletion strain restored nearly normal patterns of DNA methylation, indicating that the CHAP–HA fusion is functional (Fig. 7B). Strains expressing CHAP^ATh1 showed moderate restoration of DNA methylation, and strains expressing CHAP^ATh2 exhibited almost full restoration. However, strains bearing mutations in both AT-hook motifs (CHAP^ATh1&2) showed marked defects in DNA methylation (Fig. 7B). Co-IP experiments verified that the AT-hook mutations do not affect the stable formation of HCHC (Fig. S8C). We conclude that the CHAP AT-hook motifs are required for normal DNA methylation, perhaps through the AT-rich DNA-binding activity expected of such motifs (26).

Fig. 7.

The CHAP AT-hook motifs are required for DNA methylation. (A) Diagram of the CHAP AT-hook motif point mutations tested. (B) Effects of chap AT-hook motif mutations on DNA methylation at 8:A6 region and peak 33 (22). DNA isolated from indicated strains was analyzed by Southern hybridizations as in Fig. 3B. Strains: N3752, N1877, N3643, N3818, N3820, N3822, and N3845.

The CHAP AT-Hook Motifs Specifically Bind AT-Rich DNA with High Numbers of Repeat-Induced Point Mutations.

To test whether the CHAP AT-hook motifs bind AT-rich DNA that has repeat-induced point mutations, we performed in vitro DNA-affinity purification with the recombinant CHAP N terminus (residues 1–274) containing the two AT-hook motifs and analyzed the purified DNA with high-throughput sequencing. To complement this approach, we also assessed the binding of CHAP in vivo with DamID sequencing using the CHAP–Dam strain. Together, these techniques gave us a detailed genomic view of the specific localization and binding of CHAP to AT-rich DNA that has repeat-induced point mutations, which is nearly coincident with methylated DNA regions (Fig. 8 A and B). We carried out band-shift assays to test further the binding of CHAP to AT-rich DNA and the role of its AT-hooks, using two representative probe sequences containing distinct AT contents: the middle segment of the heterochromatic region peak 33 (probe 1, 75.1% A+T) and a segment adjacent to the region (probe 2, 43.5% A+T) (Fig. 8B). The CHAP AT-hook motifs bound strongly to the AT-rich DNA that has repeat-induced point mutations (probe 1) but not to the control region (probe 2), and the binding was essentially abolished by mutations of the AT-hook motifs (CHAP-N^ATh1&2) (Fig. 8 C and D). We conclude that CHAP, through its AT-hook motifs, binds AT-rich DNA that has repeat-induced point mutations (15).

Fig. 8.

The CHAP AT-hooks specifically bind to DNA that has repeat-induced point mutations. (A) Genome-wide distribution of DNA methylation (5mC) in the wild-type stain (Top Row), in vitro CHAP-binding distribution mapped with DNA affinity-purified by the recombinant wild-type N-terminal CHAP protein (Middle Row), and in vivo CHAP distribution determined by DamID sequencing with CHAP–Dam (Bottom Row). The bottom row indicates GC content. Strains: N3752 and N4045. (B) Distribution of DNA methylation and CHAP localization at the peak 33 region. GATC sites are indicated at the bottom to explain the gaps in CHAP localization as determined by DamID sequencing and to illustrate the limitations of the technique. The horizontal black bars and numbers identify the regions used for gel mobility shift assays. (C) The CHAP AT-hook motifs bind AT-rich DNA that has repeat-induced point mutations. Gel mobility shift assays were performed using recombinant wild-type N-terminal CHAP protein (CHAP^WT-N) or CHAP AT-hook mutant protein (CHAP^ATh1&2-N). Probe regions 1 and 2 shown in B were amplified with radiolabeled dCTP by PCR and tested for binding as indicated by the gel shift. (D) Dissociation constant for N-terminal CHAP protein (CHAP^WT-N) and the probe for AT-rich DNA that has repeat-induced point mutations (probe 1) (n = 3).

Discussion

DNA methylation, a prototypical epigenetic mark, is widely thought to be stably maintained by a simple copying system at symmetric methylated sites, as proposed by Riggs (27) and Holliday and Pugh (28) more than 30 years ago. However, it has become apparent that maintenance of methylation patterns reflects the product of a variety of processes involving a multitude of proteins. In addition to DNA methyltransferases and other enzymes that interact with DNA to convert or excise 5-methylcytosine (5mC) residues, chromatin remodelers and histone modification enzymes impact the distribution and intensity of DNA methylation (29). Indeed, in some organisms, such as N. crassa, DNA methylation is dependent on the methylation of a particular residue of histone H3, H3K9 (11). Other histone modifications, such as methylation of H3K4, phosphorylation of H3S10 (30, 31), and histone acetylation, also influence DNA methylation (32, 33). We previously demonstrated that HCHC mutants of N. crassa show increased acetylation of histone H3 and H4 at larger heterochromatin domains, such as centromeres, and speculated that the increased acetylation might provide an enhanced environment for the HP1–DIM-2 complex, leading to the increased DNA methylation observed in the large domains of constitutive heterochromatin in centromere regions. Our WGBS analyses on a wild-type strain confirmed that shorter regions tend to be more methylated than longer regions (Fig. 2A), whereas HCHC mutants show hypomethylation of shorter regions and hypermethylation of longer regions (Fig. 2 A and B). The current study also demonstrated that the AT-hook motifs of CHAP are important for proper DNA methylation and bind specifically to AT-rich DNA that has repeat-induced point mutations (Fig. 8), which is particularly prevalent at centromere regions. This finding raises the possibility that CHAP binding contributes to stronger recruitment of HCHC at centromeres, at the expense of the HP1–DIM-2 complex, leading to the characteristic low levels of DNA methylation in these regions. It is interesting that, in contrast to the importance of DNA methylation in silencing short heterochromatic regions, DNA methylation is unnecessary for silencing at centromere regions (22).

The HCHC complex possesses two CD proteins, HP1 and CDP-2, which one might imagine could operate semiredundantly. Consistent with this possibility, we found that although the CD of CDP-2 binds efficiently to methylated H3K9 in vitro (22), this domain is not required for normal DNA methylation and centromere silencing in vivo (Fig. 3B and Fig. S2 B and C). We therefore considered the possibility that the CDP-2 CD in the HCHC complex might mediate the association of this complex with methylated H3K9 in the absence of HP1 binding. However, the CDP-2 CD mutants did not show additional DNA methylation defects in the HP1 CD-mutant background (Fig. 3E), suggesting that the CDP-2 CD does not work redundantly with the HP1 CD in HCHC. Instead, our findings suggest that CDP-2 serves as a bridge between HP1 and HDA-1, ensuring the proper recruitment of the HDA-1 HDAC domain to chromatin (Fig. 9B). We show that CDP-2, like DIM-2, interacts directly with the HP1 chromoshadow domain through the PxVxL-like motifs near the N terminus (Fig. 4 B and C). This observation is consistent with our previous observation that HP1 forms physically and functionally distinct complexes with DIM-2 and CDP-2 (22).

Fig. 9.

Model for the interrelationship of the components of HCHC. (A) The HP1 chromoshadow domain (CSD) is predicted to dimerize, creating a binding pocket for the PxVxL-like motif of CDP-2. The adjacent region of CDP-2 interacts with the HDA-1 HDAC domain. The HDA-1 Arb2 domain interacts with the CHAP zinc finger motif (Zf). The CD of HP1 and CDP-2 and the CHAP AT-hooks bind to trimethylated H3 (red spheres) and DNA that has repeat-induced point mutations (black line), respectively. Proper formation and chromatin recognition by HCHC is required for the full HDA-1 HDAC activity, which removes acetyl groups (AC) from chromatin. (B–D) The CDP-2 CD is not required for the HCHC function (B), but the CHAP AT-hooks and the HP1 CD are important (C and D). mC, methyl-cytosine.

We demonstrated that in the absence of the HP1 CD, N. crassa still has residual DNA methylation in the regions with repeat-induced point mutations that depend on HDA-1. This surprising residual DNA methylation is dependent on CHAP (Fig. 6), which apparently serves as an additional means to recruit HCHC that is independent of the chromodomains (Fig. 9D). Therefore, we propose that dual chromatin recognition of heterochromatin by the HP1 CD and by the CHAP AT-hook motifs is responsible for HCHC function (Fig. 9A). Curiously, we still observed DNA methylation in double mutants lacking the HP1 CD and CHAP at the region 8:F10, which has repeat-induced point mutations (Fig. 6), raising the possibility that another element of HP1 recognizes a heterochromatic signal. In mammals and fission yeasts, HP1 has been shown to bind to RNA through the hinge region in addition to binding methylated H3K9 through the CD (34, 35). Although the RNAi pathway is not involved in heterochromatin formation in N. crassa (36), bivalent recognition via the CD and hinge region of HP1 seems possible.

Although N. crassa has a relatively simple DNA methylation pathway centered on H3K9 methylation serving as a signal for the direct recruitment of the HP1–DIM-2 complex (13, 14), reinforcing loops involving H3K9me3, HP1, and DNA methylation occur. Recent studies using N. crassa and Arabidopsis uncovered mutants that fail to modulate these reinforcing loops properly (37, 38). The mutants exhibit abnormal silencing of essential genes by aberrant DNA methylation and H3K9 methylation, resulting in growth defects. In N. crassa, aberrant H3K9me3 depends on DNA methylation, revealing the existence of feedback pathway from DNA methylation to H3K9me3 (37). In addition, our WGBS analyses revealed that HDA-1 and CHAP are required for the spreading of DNA methylation (Fig. 2D), presumably through their binding of HDAC and AT-rich DNA that has repeat-induced point mutations. In summary, we describe multifaceted interrelationships among AT-rich DNA that has repeat-induced point mutations, H3K9me3, HP1, histone deacetylation, and DNA methylation that together result in the observed establishment and maintenance of heterochromatic domains.

The N. crassa HCHC complex shares features with the Schizosaccharomyces pombe HDAC complex SHREC (25, 39), which also functions in centromeric silencing. Although there are obvious differences between the HDAC complexes in fission yeast and N. crassa (e.g., N. crassa HCHC does not contain a homolog of the chromatin remodeler Mit1), it would be interesting to know if mammals use a similar mechanism to control proper heterochromatin domains, especially at AT-rich peri-centromeric heterochromatin.

Materials and Methods

N. crassa Strains and Molecular Analyses.

All N. crassa strains and primers used in this study are listed in Tables S1 and S2, respectively. Strains were grown, crossed, and maintained according to standard procedures (40). N. crassa strains were built according to methods described previously (41). Detailed methods for strain building, including plasmid and primers used are included in SI Materials and Methods. DNA isolation, Southern blotting, Western blotting, co-IP, and fluorescence microscopy were performed as previously described (14). The following antibodies were used: anti-FLAG (Sigma, F3165; MBL, M185-3), anti-HA (University of Oregon monoclonal facility; Roche, 3F10; MBL, M180-3), anti-GFP (Abcam, ab290; MBL, 598), and anti-tubulin (Sigma, T6199). Specific HP1, CDP-2, HDA-1, and CHAP mutations were made with a QuikChange site-directed mutagenesis kit (Stratagene) and a PCR-based mutagenesis with the In-Fusion HD cloning system (Takara).

View this table:

Table S1.

Neurospora crassa strains used in this study

View this table:

Table S2.

Primers used in this study

Assessment of Chromosome Bridges.

The frequency of chromosome bridges was quantified using with GFP-tagged histone H2A (20). Conidia were plated on a thin layer of Vogel's medium solidified with 2% agar and supplemented with required nutrients and were grown at 32 °C overnight. A square of the culture was placed on a slide and covered with a drop of Vogel's medium and a coverslip. Hyphae observed using a 100× oil-immersion objective in a Zeiss Axioplan 2 fluorescence microscope with differential interference contrast (DIC) showed cytoplasmic streaming. Two methods were used to count chromosome bridges that were visualized with a GFP filter. (i) For tips with at least four nuclei, those having a chromosome bridge were scored as positive. (ii) In other hyphae (in which the tips were not obvious), the number of bridges was recorded relative to the total number of nuclei observed. To combine the results from the two counting schemes, the number of tips multiplied by 4 was used as the number of nuclei for the first counting method. Results were expressed as the percentage of nuclei showing bridges relative to the total number of nuclei. For each nucleus there are two possible outcomes: having a bridge or not having a bridge. Thus, we used the statistical test for a binomial distribution. The 95% confidence intervals were calculated for the binomial parameter p, the probability of a bridge in a given strain, using the formula (p(1−p)/n)(1.96) where n is the total number of nuclei observed.

WGBS.

WGBS was performed and reads were mapped as previously described (42). Sequencing reads can be downloaded from the National Center for Biotechnology (NCBI) database (accession no. GSE81129). Normally methylated regions with a minimum size of 200 bp were determined using the RSEG software package (smithlabresearch.org/software/rseg/). To display the bisulfite sequencing data, the average 5mC level was determined for specified step-wise window sizes across the genome using the MethPipe program (smithlabresearch.org/software/methpipe/) (43). The resulting file was renamed with an .igv file extension to allow display on the Integrated Genome Viewer (software.broadinstitute.org/software/igv/) (44). Similarly, the MethPipe (ROI function) was used to calculate the average 5mC level over the normally methylated regions found in the wild-type strain (N3752) as determined using RSEG software and sequences immediately flanking these regions. The CRI was calculated for 500-bp windows across the N. crassa genome using a custom Perl script (15).

CHAP–DamID Sequencing.

Whole-genome DamID sequencing was performed using a procedure adapted from ref. 46. Briefly, genomic DNA from the Dam-tagged CHAP strain was digested with DpnI. Digested DNA was ligated to adapters and amplified using a biotin-tagged primer. The amplified DNA was fragmented by sonication to 100- to 500-bp products and purified using streptavidin-conjugated beads (Sigma). Bound DNA was eluted using a DpnII digestion. Purified DNA was prepared for sequencing using the Illumina TruSeq ChIP Sample Preparation Kit. Sequence alignments were performed as previously described (47), except that the reads were mapped to the N. crassa OR74A (NC12) genome (N. crassa Sequencing Project, Broad Institute of Harvard and MIT; www.broadinstitute.org/), and read densities then were averaged over 25-bp windows to generate all tiled data files. Sequencing reads can be downloaded from the NCBI database (accession no. GSE81129).

Construction of HA-Tagged CHAP Fusion Constructs Expressed at the pan-2 Locus.

We amplified a fragment of HA-tagged chap gene with its native promoter by PCR with primers 2090 and 2497 from the genomic DNA of a strain expressing CHAP–HA from its native locus (created using the knock-in system described above). The PCR products were digested with NotI and XhoI, inserted into the pan-2 targeting vector pRATT42b (the gift of R. Aramayo, Texas A&M University), linearized, and inserted at the pan-2⁻::hph⁺::tk⁺ locus of the chap-null mutant (N3642).

Generation of Recombinant CHAP Proteins and Gel Mobility Shift Assays.

The chap ORF (amino acids 1–274) was amplified with primers 3011 and 3069 and inserted between the EcoRI and BamHI sites of pMALc2 (New England Biolabs). The plasmids were transformed into E. coli. strain BL21, and recombinant proteins were purified as described by the manufacturer of pMALc2. Recombinant maltose binding protein (MBP)-CHAP^1–274 protein was incubated for 30 min at room temperature in a 20-µL volume of EMSA binding buffer [20 mM Hepes (pH 7.9), 50 mM KCl, 4 mM MgCl₂, 25 μM ZnCl₂, and 1 mM DTT], 1 μg of BSA, and a radiolabeled DNA probe. A 100-pM DNA probe was used for K_d determination. Double-stranded DNA probes were produced using PCR primers (probe 1: primers 3019 and 3020; probe 2: primers 2483 and 2484) in reactions supplemented with [α-³²P]dCTP. Following incubation, EMSA reactions were analyzed on 5–20% Mini-Protean TGX gels (Bio-Rad); after electrophoresis, gels were dried and autoradiographed.

DNA–Protein Affinity Purification.

DNA affinity purification using recombinant MBP-CHAP was performed using a protocol adapted from ref. 48. Amylose resin (New England Biolabs) containing immobilized MBP-CHAP was incubated with sonicated wild-type genomic DNA (∼250-bp fragments) in binding buffer [20 mM Hepes (pH 7.5), 80 mM NaCl, 37.5 mM Imidazole, 0.7 mM MgCl₂, 0.35 mM EDTA, 0.7 mM DTT, and 17.8% glycerol] for 2 h at 4 °C. Beads were washed six times with 1 mL of wash buffer [10 mM Tris (pH 7.5), 50 mM NaCl, 1 mM MgCl₂, 0.5 mM EDTA, and 4% glycerol]. Following washes, DNA was eluted with TES [20 mM Tris (pH 8.0), 10 mM EDTA, and 1% SDS] by heating for 10 min at 65 °C. Eluted samples were treated with proteinase K, and DNA was purified with MinElute columns (Qiagen). Purified DNA was prepared for high-throughput sequencing using the TruSeq ChIP Sample Preparation Kit (Illumina).

SI Materials and Methods

Construction of a C-Terminal Dam Fusion Vector for his-3 Targeting and Fusion Constructs for CDP-2, HDA-1, and CHAP.

Escherichia coli dam was amplified from pCmyc-Dam (24) with primer 3058, which contains a PacI site and a 10×Gly tail, and primer 3059, which contains an EcoRI site. The PCR products were digested with PacI and EcoRI and inserted into PacI- and EcoRI-digested pCCG::C-3×FLAG (41) to replace the 3×FLAG region with the 10×Gly–Dam segment, yielding pCCG::C-Gly::Dam.

To express the CDP-2–Dam fusion construct from its endogenous promoter, the cdp-2 coding region was transferred from pTTK26 with NotI and PacI into pCCG::C-Gly::Dam. We assembled FLAG- and Dam-tagged hda-1 constructs for targeting to the his-3 locus by PCR with primers 3062 and 2093 and cloned the product into pCR2.1 using a TA cloning kit (Invitrogen). A fragment of the hda-1 promoter region was amplified by PCR with primers 2091 and 3086 and was transferred with NotI and SpeI into pCCG::C-Gly::3×FLAG (41) to replace the ccg-1 promoter with the hda-1 promoter, yielding phda-1::C-Gly::3×FLAG. We amplified the his-3 3′-flanking sequence along with the hda-1 promoter from phda-1::C-Gly::3×FLAG using primers 3131 and 3126. Similarly, the 5′ his-3 region with a 3×FLAG or a Dam tag was amplified by PCR with primers 3125 and 3128 and pCCG::C-Gly::3×FLAG or pCCG::C-Gly::Dam, respectively, as templates. The three products, containing the 3′ his-3– flanking sequence with the hda-1 promoter, the 5′ partial his-3 sequence with the epitope tag, and the cloned hda-1 ORF sequence, were mixed and assembled by PCR with primers 3128 and 3131, were gel-purified, and were introduced at the his-3 locus of the hda-1–null mutant by electroporation (45). To express Dam-tagged CHAP from its endogenous promoter, a fragment of the chap coding region with its promoter was amplified by PCR, digested with NotI and PacI, and inserted into pCCG::C-Gly::Dam using the same enzymes, yielding pCHAP::C-Gly::Dam. The pCHAP::C-Gly::Dam then was linearized and inserted at his-3 in the chap-null mutant (45).

Construction of the C-Terminal HAT/FLAG–Tagged HP1 CD Constructs Expressed at his-3.

Site-directed mutagenesis using overlapping forward and reverse primers (3081 and 3082) was used to create a point mutation (T to G) in the CD region of HP1 resulting in a codon change of W to G at the 98th residue. The his-3–targeting plasmid 2899 (phpo::hpo-HAT-FLAG) was used as the template, and the creation of the point mutation was confirmed by sequencing.

To generate a deletion of the entire hpo CD, we amplified two fragments of the hpo gene by PCR with one fragment immediately upstream of the CD (primers 5246 and 5247) and the other immediately downstream of the CD (primers 5248 and 5249) using plasmid 2899 (phpo::hpo-HAT-FLAG) as the template. These fragments were combined through stitching PCR, generating a deletion of the HP1 CD. This stitched PCR fragment then was digested with BsiWI and PacI and was cloned into the BsiWI/PacI-digested plasmid 2899. The deletion of the CD was confirmed by sequencing.

These HP1 CD mutant constructs, along with the HP1 wild-type construct, were linearized and transformed into N5430 (his-3; Δhpo strain). His⁺ transformants were selected and checked by Southern analysis for the proper integration of the HP1 constructs at the his-3 locus. Positive transformants then were crossed to the N4909 (cenVIR::bar; his-3; trp-2) strain to generate homokaryotic strains containing cenVIR::bar; his-3::hpo^WT ^or ^CDmutant; Δhpo::hph; trp-2.

Construction of CDP-2, HDA-1, and CHAP Mutant Constructs.

We created the other mutant constructs by site-direct mutagenesis similarly with the following primer pairs: primers 3187 and 3188 for cdp-2^I14A, ^E15A; primers 3056 and 3057 for cdp-2^W466G; primers 3138 and 3139 for cdp-2^ΔPPITL; primers 3171 and 3172 for cdp-2^{ΔCD(444–459aa)}; primers 3054 and 3055 for hda-1^D263N; primers 2433 and 2434 for CHAP^ATh1(R210A); primers 2435 and 2436 for CHAP^ATh2(R250A); primers 2437 and 2438 for CHAP^Zf1(C280A); and primers 2439 and 2440 for CHAP^Zf2(C327A).

Construction of LexADBD-Tagged HP1 CD-Deletion Fusion Constructs Expressed at the Native hpo Locus.

We amplified the hpo gene with the CD deleted by PCR with primers 4525 and JGP123 from N5869. Additionally the LexADBD with an 8× glycine linker was amplified by PCR from plasmid 3015 using primers JGP62 and JGP63. The hygromycin resistance cassette P_trpC-hph was amplified from PCR from plasmid 2409 using primers JGP60 and JGP61. Also, the flanking region downstream of the hpo gene was amplified by PCR using primers JGP124 and JGP125. These PCR products were combined by PCR stitching and integrated at the native hpo locus of N5643.

Construction of CDP-2 CD-Deletion Constructs Expressed at the Native cdp-2 Locus.

We amplified the CD-deleted cdp-2 gene by PCR with primers N3064 and JGP281 from plasmid 2973 (created for ectopic CDP-2^ΔCD expression from the his-3 locus). Additionally, the flanking region downstream of the cdp-2 gene was amplified by PCR using primers JGP282 and 3145. These PCR products were combined through PCR stitching with a central trpC–nat-1 antibiotic resistance cassette from plasmid 3130. This PCR product then was integrated at the native cdp-2 locus of N2930.

Acknowledgments

We thank Tamir Khalafallah and Paula Grisafi for technical support. This work was funded by NIH Grants GM025690 (to E.U.S) and CA180468 (to V.T.B). J.D.G. was supported by NIH Genetics Training Grant GM007413. S.H. received support from the Competitive Funds in the Program to Disseminate Tenure Tracking System of the Ministry of Education, Culture, Sports, Science, and Technology, Japan.

Footnotes

↵¹S.H., V.T.B., J.D.G., and M.R.R. contributed equally to this work.
↵²Present address: Nzumbe, Inc., Portland, OR 97201.
↵³To whom correspondence should be addressed. Email: selker{at}uoregon.edu.

Author contributions: S.H., V.T.B., J.D.G., M.R.R., and E.U.S. designed research; S.H., V.T.B., J.D.G., M.R.R., A.Y., E.Y.Y., and J.M.L.S. performed research; S.H., V.T.B., J.D.G., M.R.R., J.M.L.S., and E.U.S. analyzed data; and S.H., V.T.B., J.D.G., M.R.R., J.M.L.S., and E.U.S. wrote the paper.
Reviewers: S.E., Washington University; and M.M., Harvard Medical School.
The authors declare no conflict of interest.
Data deposition: The sequence reported in this work has been deposited in the National Center for Biotechnology Information database (accession no. GSE81129).
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1614279113/-/DCSupplemental.

References

↵
1. Heard E,
2. Disteche CM
(2006) Dosage compensation in mammals: Fine-tuning the expression of the X chromosome. Genes Dev 20(14):1848–1867.
.
OpenUrl Abstract/FREE Full Text
↵
1. Reik W,
2. Walter J
(2001) Genomic imprinting: Parental influence on the genome. Nat Rev Genet 2(1):21–32.
.
OpenUrl CrossRef PubMed
↵
1. Slotkin RK,
2. Martienssen R
(2007) Transposable elements and the epigenetic regulation of the genome. Nat Rev Genet 8(4):272–285.
.
OpenUrl CrossRef PubMed
↵
1. Weber M,
2. Schübeler D
(2007) Genomic patterns of DNA methylation: Targets and function of an epigenetic mark. Curr Opin Cell Biol 19(3):273–280.
.
OpenUrl CrossRef PubMed
↵
1. Reik W,
2. Dean W,
3. Walter J
(2001) Epigenetic reprogramming in mammalian development. Science 293(5532):1089–1093.
.
OpenUrl Abstract/FREE Full Text
↵
1. Feinberg AP
(2007) Phenotypic plasticity and the epigenetics of human disease. Nature 447(7143):433–440.
.
OpenUrl CrossRef PubMed
↵
1. Robertson KD
(2005) DNA methylation and human disease. Nat Rev Genet 6(8):597–610.
.
OpenUrl CrossRef PubMed
↵
1. Cedar H,
2. Bergman Y
(2009) Linking DNA methylation and histone modification: Patterns and paradigms. Nat Rev Genet 10(5):295–304.
.
OpenUrl CrossRef PubMed
↵
1. Rountree MR,
2. Selker EU
(2010) DNA methylation and the formation of heterochromatin in Neurospora crassa. Heredity (Edinb) 105(1):38–44.
.
OpenUrl CrossRef PubMed
↵
1. Chan SW,
2. Henderson IR,
3. Jacobsen SE
(2005) Gardening the genome: DNA methylation in Arabidopsis thaliana. Nat Rev Genet 6(5):351–360.
.
OpenUrl CrossRef PubMed
↵
1. Tamaru H,
2. Selker EU
(2001) A histone H3 methyltransferase controls DNA methylation in Neurospora crassa. Nature 414(6861):277–283.
.
OpenUrl CrossRef PubMed
↵
1. Tamaru H, et al.
(2003) Trimethylated lysine 9 of histone H3 is a mark for DNA methylation in Neurospora crassa. Nat Genet 34(1):75–79.
.
OpenUrl CrossRef PubMed
↵
1. Freitag M,
2. Hickey PC,
3. Khlafallah TK,
4. Read ND,
5. Selker EU
(2004) HP1 is essential for DNA methylation in Neurospora. Mol Cell 13(3):427–434.
.
OpenUrl CrossRef PubMed
↵
1. Honda S,
2. Selker EU
(2008) Direct interaction between DNA methyltransferase DIM-2 and HP1 is required for DNA methylation in Neurospora crassa. Mol Cell Biol 28(19):6044–6055.
.
OpenUrl Abstract/FREE Full Text
↵
1. Lewis ZA, et al.
(2009) Relics of repeat-induced point mutation direct heterochromatin formation in Neurospora crassa. Genome Res 19(3):427–437.
.
OpenUrl Abstract/FREE Full Text
↵
1. Selker EU
(1990) Premeiotic instability of repeated sequences in Neurospora crassa. Annu Rev Genet 24:579–613.
.
OpenUrl CrossRef PubMed
↵
1. Selker EU, et al.
(2003) The methylated component of the Neurospora crassa genome. Nature 422(6934):893–897.
.
OpenUrl CrossRef PubMed
↵
1. Miao VP,
2. Freitag M,
3. Selker EU
(2000) Short TpA-rich segments of the zeta-eta region induce DNA methylation in Neurospora crassa. J Mol Biol 300(2):249–273.
.
OpenUrl CrossRef PubMed
↵
1. Tamaru H,
2. Selker EU
(2003) Synthesis of signals for de novo DNA methylation in Neurospora crassa. Mol Cell Biol 23(7):2379–2394.
.
OpenUrl Abstract/FREE Full Text
↵
1. Lewis ZA, et al.
(2010) DNA methylation and normal chromosome behavior in Neurospora depend on five components of a histone methyltransferase complex, DCDC. PLoS Genet 6(11):e1001196.
.
OpenUrl CrossRef PubMed
↵
1. Lewis ZA,
2. Adhvaryu KK,
3. Honda S,
4. Shiver AL,
5. Selker EU
(2010) Identification of DIM-7, a protein required to target the DIM-5 H3 methyltransferase to chromatin. Proc Natl Acad Sci USA 107(18):8310–8315.
.
OpenUrl Abstract/FREE Full Text
↵
1. Honda S, et al.
(2012) Heterochromatin protein 1 forms distinct complexes to direct histone deacetylation and DNA methylation. Nat Struct Mol Biol 19(5):471–477, S1.
.
OpenUrl CrossRef PubMed
↵
1. Thiru A, et al.
(2004) Structural basis of HP1/PXVXL motif peptide interactions and HP1 localisation to heterochromatin. EMBO J 23(3):489–499.
.
OpenUrl Abstract/FREE Full Text
↵
1. Vogel MJ,
2. Peric-Hupkes D,
3. van Steensel B
(2007) Detection of in vivo protein-DNA interactions using DamID in mammalian cells. Nat Protoc 2(6):1467–1478.
.
OpenUrl CrossRef PubMed
↵
1. Sugiyama T, et al.
(2007) SHREC, an effector complex for heterochromatic transcriptional silencing. Cell 128(3):491–504.
.
OpenUrl CrossRef PubMed
↵
1. Aravind L,
2. Landsman D
(1998) AT-hook motifs identified in a wide variety of DNA-binding proteins. Nucleic Acids Res 26(19):4413–4421.
.
OpenUrl Abstract/FREE Full Text
↵
1. Riggs AD
(1975) X inactivation, differentiation, and DNA methylation. Cytogenet Cell Genet 14(1):9–25.
.
OpenUrl CrossRef PubMed
↵
1. Holliday R,
2. Pugh JE
(1975) DNA modification mechanisms and gene activity during development. Science 187(4173):226–232.
.
OpenUrl FREE Full Text
↵
1. Du J,
2. Johnson LM,
3. Jacobsen SE,
4. Patel DJ
(2015) DNA methylation pathways and their crosstalk with histone methylation. Nat Rev Mol Cell Biol 16(9):519–532.
.
OpenUrl CrossRef PubMed
↵
1. Adhvaryu KK,
2. Selker EU
(2008) Protein phosphatase PP1 is required for normal DNA methylation in Neurospora. Genes Dev 22(24):3391–3396.
.
OpenUrl Abstract/FREE Full Text
↵
1. Adhvaryu KK,
2. Berge E,
3. Tamaru H,
4. Freitag M,
5. Selker EU
(2011) Substitutions in the amino-terminal tail of Neurospora histone H3 have varied effects on DNA methylation. PLoS Genet 7(12):e1002423.
.
OpenUrl CrossRef PubMed
↵
1. Selker EU
(1998) Trichostatin A causes selective loss of DNA methylation in Neurospora. Proc Natl Acad Sci USA 95(16):9430–9435.
.
OpenUrl Abstract/FREE Full Text
↵
1. Smith KM, et al.
(2010) H2B- and H3-specific histone deacetylases are required for DNA methylation in Neurospora crassa. Genetics 186(4):1207–1216.
.
OpenUrl Abstract/FREE Full Text
↵
1. Keller C, et al.
(2012) HP1(Swi6) mediates the recognition and destruction of heterochromatic RNA transcripts. Mol Cell 47(2):215–227.
.
OpenUrl CrossRef PubMed
↵
1. Muchardt C, et al.
(2002) Coordinated methyl and RNA binding is required for heterochromatin localization of mammalian HP1α. EMBO Rep 3(10):975–981.
.
OpenUrl Abstract/FREE Full Text
↵
1. Freitag M, et al.
(2004) DNA methylation is independent of RNA interference in Neurospora. Science 304(5679):1939.
.
OpenUrl FREE Full Text
↵
1. Honda S, et al.
(2010) The DMM complex prevents spreading of DNA methylation from transposons to nearby genes in Neurospora crassa. Genes Dev 24(5):443–454.
.
OpenUrl Abstract/FREE Full Text
↵
1. Saze H,
2. Shiraishi A,
3. Miura A,
4. Kakutani T
(2008) Control of genic DNA methylation by a jmjC domain-containing protein in Arabidopsis thaliana. Science 319(5862):462–465.
.
OpenUrl Abstract/FREE Full Text
↵
1. Motamedi MR, et al.
(2008) HP1 proteins form distinct complexes and mediate heterochromatic gene silencing by nonoverlapping mechanisms. Mol Cell 32(6):778–790.
.
OpenUrl CrossRef PubMed
↵
1. Davis RH
(2000) Neurospora: Contributions of a Model Organism (Oxford Univ Press, Oxford, UK), p 333.
.
↵
1. Honda S,
2. Selker EU
(2009) Tools for fungal proteomics: Multifunctional neurospora vectors for gene replacement, protein expression and protein purification. Genetics 182(1):11–23.
.
OpenUrl Abstract/FREE Full Text
↵
1. Klocko AD, et al.
(2015) Neurospora importin α is required for normal heterochromatic formation and DNA methylation. PLoS Genet 11(3):e1005083.
.
OpenUrl CrossRef PubMed
↵
1. Song Q, et al.
(2013) A reference methylome database and analysis pipeline to facilitate integrative and comparative epigenomics. PLoS One 8(12):e81148.
.
OpenUrl CrossRef PubMed
↵
1. Robinson JT, et al.
(2011) Integrative genomics viewer. Nat Biotechnol 29(1):24–26.
.
OpenUrl CrossRef PubMed
↵
1. Margolin BS,
2. Freitag M,
3. Selker EU
(1997) Improved plasmids for gene targeting at the his-3 locus of Neurospora crassa by electroporation. Fungal Genet Newsl 44:34–36.
.
OpenUrl CrossRef
↵
1. Zhou VW
(2012) Methods for Global Characterization of Chromatin Regulators in Human Cells (Harvard Univ Press, Cambridge, MA).
.
↵
1. Jamieson K,
2. Rountree MR,
3. Lewis ZA,
4. Stajich JE,
5. Selker EU
(2013) Regional control of histone H3 lysine 27 methylation in Neurospora. Proc Natl Acad Sci USA 110(15):6027–6032.
.
OpenUrl Abstract/FREE Full Text
↵
1. Jolma A, et al.
(2010) Multiplexed massively parallel SELEX for characterization of human transcription factor binding specificities. Genome Res 20(6):861–873.
.
OpenUrl Abstract/FREE Full Text
↵
1. Fischle W, et al.
(2005) Regulation of HP1-chromatin binding by histone H3 methylation and phosphorylation. Nature 438(7071):1116–1122.
.
OpenUrl CrossRef PubMed
↵
1. Nielsen PR, et al.
(2002) Structure of the HP1 chromodomain bound to histone H3 methylated at lysine 9. Nature 416(6876):103–107.
.
OpenUrl CrossRef PubMed
↵
1. James P,
2. Halladay J,
3. Craig EA
(1996) Genomic libraries and a host strain designed for highly efficient two-hybrid selection in yeast. Genetics 144(4):1425–1436.
.
OpenUrl Abstract/FREE Full Text
1. Kouzminova EA,
2. Selker EU
(2001) Dim-2 encodes a DNA-methyltransferase responsible for all known cytosine methylation in Neurospora. EMBO J 20:4309–4323.
.
OpenUrl Abstract/FREE Full Text

Article Alerts

Email Article

Citation Tools

Request Permissions

Article Classifications

Biological Sciences
Genetics

Proceedings of the National Academy of Sciences: 113 (41)

Table of Contents

Submit

[1] ↵
Heard E,
Disteche CM
(2006) Dosage compensation in mammals: Fine-tuning the expression of the X chromosome. Genes Dev 20(14):1848–1867.
.
OpenUrl Abstract/FREE Full Text

[2] Heard E,

[3] Disteche CM

[4] ↵
Reik W,
Walter J
(2001) Genomic imprinting: Parental influence on the genome. Nat Rev Genet 2(1):21–32.
.
OpenUrl CrossRef PubMed

[5] Reik W,

[6] Walter J

[7] ↵
Slotkin RK,
Martienssen R
(2007) Transposable elements and the epigenetic regulation of the genome. Nat Rev Genet 8(4):272–285.
.
OpenUrl CrossRef PubMed

[8] Slotkin RK,

[9] Martienssen R

[10] ↵
Weber M,
Schübeler D
(2007) Genomic patterns of DNA methylation: Targets and function of an epigenetic mark. Curr Opin Cell Biol 19(3):273–280.
.
OpenUrl CrossRef PubMed

[11] Weber M,

[12] Schübeler D

[13] ↵
Reik W,
Dean W,
Walter J
(2001) Epigenetic reprogramming in mammalian development. Science 293(5532):1089–1093.
.
OpenUrl Abstract/FREE Full Text

[14] Reik W,

[15] Dean W,

[16] Walter J

[17] ↵
Feinberg AP
(2007) Phenotypic plasticity and the epigenetics of human disease. Nature 447(7143):433–440.
.
OpenUrl CrossRef PubMed

[18] Feinberg AP

[19] ↵
Robertson KD
(2005) DNA methylation and human disease. Nat Rev Genet 6(8):597–610.
.
OpenUrl CrossRef PubMed

[20] Robertson KD

[21] ↵
Cedar H,
Bergman Y
(2009) Linking DNA methylation and histone modification: Patterns and paradigms. Nat Rev Genet 10(5):295–304.
.
OpenUrl CrossRef PubMed

[22] Cedar H,

[23] Bergman Y

[24] ↵
Rountree MR,
Selker EU
(2010) DNA methylation and the formation of heterochromatin in Neurospora crassa. Heredity (Edinb) 105(1):38–44.
.
OpenUrl CrossRef PubMed

[25] Rountree MR,

[26] Selker EU

[27] ↵
Chan SW,
Henderson IR,
Jacobsen SE
(2005) Gardening the genome: DNA methylation in Arabidopsis thaliana. Nat Rev Genet 6(5):351–360.
.
OpenUrl CrossRef PubMed

[28] Chan SW,

[29] Henderson IR,

[30] Jacobsen SE

[31] ↵
Tamaru H,
Selker EU
(2001) A histone H3 methyltransferase controls DNA methylation in Neurospora crassa. Nature 414(6861):277–283.
.
OpenUrl CrossRef PubMed

[32] Tamaru H,

[33] Selker EU

[34] ↵
Tamaru H, et al.
(2003) Trimethylated lysine 9 of histone H3 is a mark for DNA methylation in Neurospora crassa. Nat Genet 34(1):75–79.
.
OpenUrl CrossRef PubMed

[35] Tamaru H, et al.

[36] ↵
Freitag M,
Hickey PC,
Khlafallah TK,
Read ND,
Selker EU
(2004) HP1 is essential for DNA methylation in Neurospora. Mol Cell 13(3):427–434.
.
OpenUrl CrossRef PubMed

[37] Freitag M,

[38] Hickey PC,

[39] Khlafallah TK,

[40] Read ND,

[41] Selker EU

[42] ↵
Honda S,
Selker EU
(2008) Direct interaction between DNA methyltransferase DIM-2 and HP1 is required for DNA methylation in Neurospora crassa. Mol Cell Biol 28(19):6044–6055.
.
OpenUrl Abstract/FREE Full Text

[43] Honda S,

[44] Selker EU

[45] ↵
Lewis ZA, et al.
(2009) Relics of repeat-induced point mutation direct heterochromatin formation in Neurospora crassa. Genome Res 19(3):427–437.
.
OpenUrl Abstract/FREE Full Text

[46] Lewis ZA, et al.

[47] ↵
Selker EU
(1990) Premeiotic instability of repeated sequences in Neurospora crassa. Annu Rev Genet 24:579–613.
.
OpenUrl CrossRef PubMed

[48] Selker EU

[49] ↵
Selker EU, et al.
(2003) The methylated component of the Neurospora crassa genome. Nature 422(6934):893–897.
.
OpenUrl CrossRef PubMed

[50] Selker EU, et al.

[51] ↵
Miao VP,
Freitag M,
Selker EU
(2000) Short TpA-rich segments of the zeta-eta region induce DNA methylation in Neurospora crassa. J Mol Biol 300(2):249–273.
.
OpenUrl CrossRef PubMed

[52] Miao VP,

[53] Freitag M,

[54] Selker EU

[55] ↵
Tamaru H,
Selker EU
(2003) Synthesis of signals for de novo DNA methylation in Neurospora crassa. Mol Cell Biol 23(7):2379–2394.
.
OpenUrl Abstract/FREE Full Text

[56] Tamaru H,

[57] Selker EU

[58] ↵
Lewis ZA, et al.
(2010) DNA methylation and normal chromosome behavior in Neurospora depend on five components of a histone methyltransferase complex, DCDC. PLoS Genet 6(11):e1001196.
.
OpenUrl CrossRef PubMed

[59] Lewis ZA, et al.

[60] ↵
Lewis ZA,
Adhvaryu KK,
Honda S,
Shiver AL,
Selker EU
(2010) Identification of DIM-7, a protein required to target the DIM-5 H3 methyltransferase to chromatin. Proc Natl Acad Sci USA 107(18):8310–8315.
.
OpenUrl Abstract/FREE Full Text

[61] Lewis ZA,

[62] Adhvaryu KK,

[63] Honda S,

[64] Shiver AL,

[65] Selker EU

[66] ↵
Honda S, et al.
(2012) Heterochromatin protein 1 forms distinct complexes to direct histone deacetylation and DNA methylation. Nat Struct Mol Biol 19(5):471–477, S1.
.
OpenUrl CrossRef PubMed

[67] Honda S, et al.

[68] ↵
Thiru A, et al.
(2004) Structural basis of HP1/PXVXL motif peptide interactions and HP1 localisation to heterochromatin. EMBO J 23(3):489–499.
.
OpenUrl Abstract/FREE Full Text

[69] Thiru A, et al.

[70] ↵
Vogel MJ,
Peric-Hupkes D,
van Steensel B
(2007) Detection of in vivo protein-DNA interactions using DamID in mammalian cells. Nat Protoc 2(6):1467–1478.
.
OpenUrl CrossRef PubMed

[71] Vogel MJ,

[72] Peric-Hupkes D,

[73] van Steensel B

[74] ↵
Sugiyama T, et al.
(2007) SHREC, an effector complex for heterochromatic transcriptional silencing. Cell 128(3):491–504.
.
OpenUrl CrossRef PubMed

[75] Sugiyama T, et al.

[76] ↵
Aravind L,
Landsman D
(1998) AT-hook motifs identified in a wide variety of DNA-binding proteins. Nucleic Acids Res 26(19):4413–4421.
.
OpenUrl Abstract/FREE Full Text

[77] Aravind L,

[78] Landsman D

[79] ↵
Riggs AD
(1975) X inactivation, differentiation, and DNA methylation. Cytogenet Cell Genet 14(1):9–25.
.
OpenUrl CrossRef PubMed

[80] Riggs AD

[81] ↵
Holliday R,
Pugh JE
(1975) DNA modification mechanisms and gene activity during development. Science 187(4173):226–232.
.
OpenUrl FREE Full Text

[82] Holliday R,

[83] Pugh JE

[84] ↵
Du J,
Johnson LM,
Jacobsen SE,
Patel DJ
(2015) DNA methylation pathways and their crosstalk with histone methylation. Nat Rev Mol Cell Biol 16(9):519–532.
.
OpenUrl CrossRef PubMed

[85] Du J,

[86] Johnson LM,

[87] Jacobsen SE,

[88] Patel DJ

[89] ↵
Adhvaryu KK,
Selker EU
(2008) Protein phosphatase PP1 is required for normal DNA methylation in Neurospora. Genes Dev 22(24):3391–3396.
.
OpenUrl Abstract/FREE Full Text

[90] Adhvaryu KK,

[91] Selker EU

[92] ↵
Adhvaryu KK,
Berge E,
Tamaru H,
Freitag M,
Selker EU
(2011) Substitutions in the amino-terminal tail of Neurospora histone H3 have varied effects on DNA methylation. PLoS Genet 7(12):e1002423.
.
OpenUrl CrossRef PubMed

[93] Adhvaryu KK,

[94] Berge E,

[95] Tamaru H,

[96] Freitag M,

[97] Selker EU

[98] ↵
Selker EU
(1998) Trichostatin A causes selective loss of DNA methylation in Neurospora. Proc Natl Acad Sci USA 95(16):9430–9435.
.
OpenUrl Abstract/FREE Full Text

[99] Selker EU

[100] ↵
Smith KM, et al.
(2010) H2B- and H3-specific histone deacetylases are required for DNA methylation in Neurospora crassa. Genetics 186(4):1207–1216.
.
OpenUrl Abstract/FREE Full Text

[101] Smith KM, et al.

[102] ↵
Keller C, et al.
(2012) HP1(Swi6) mediates the recognition and destruction of heterochromatic RNA transcripts. Mol Cell 47(2):215–227.
.
OpenUrl CrossRef PubMed

[103] Keller C, et al.

[104] ↵
Muchardt C, et al.
(2002) Coordinated methyl and RNA binding is required for heterochromatin localization of mammalian HP1α. EMBO Rep 3(10):975–981.
.
OpenUrl Abstract/FREE Full Text

[105] Muchardt C, et al.

[106] ↵
Freitag M, et al.
(2004) DNA methylation is independent of RNA interference in Neurospora. Science 304(5679):1939.
.
OpenUrl FREE Full Text

[107] Freitag M, et al.

[108] ↵
Honda S, et al.
(2010) The DMM complex prevents spreading of DNA methylation from transposons to nearby genes in Neurospora crassa. Genes Dev 24(5):443–454.
.
OpenUrl Abstract/FREE Full Text

[109] Honda S, et al.

[110] ↵
Saze H,
Shiraishi A,
Miura A,
Kakutani T
(2008) Control of genic DNA methylation by a jmjC domain-containing protein in Arabidopsis thaliana. Science 319(5862):462–465.
.
OpenUrl Abstract/FREE Full Text

[111] Saze H,

[112] Shiraishi A,

[113] Miura A,

[114] Kakutani T

[115] ↵
Motamedi MR, et al.
(2008) HP1 proteins form distinct complexes and mediate heterochromatic gene silencing by nonoverlapping mechanisms. Mol Cell 32(6):778–790.
.
OpenUrl CrossRef PubMed

[116] Motamedi MR, et al.

[117] ↵
Davis RH
(2000) Neurospora: Contributions of a Model Organism (Oxford Univ Press, Oxford, UK), p 333.
.

[118] Davis RH

[119] ↵
Honda S,
Selker EU
(2009) Tools for fungal proteomics: Multifunctional neurospora vectors for gene replacement, protein expression and protein purification. Genetics 182(1):11–23.
.
OpenUrl Abstract/FREE Full Text

[120] Honda S,

[121] Selker EU

[122] ↵
Klocko AD, et al.
(2015) Neurospora importin α is required for normal heterochromatic formation and DNA methylation. PLoS Genet 11(3):e1005083.
.
OpenUrl CrossRef PubMed

[123] Klocko AD, et al.

[124] ↵
Song Q, et al.
(2013) A reference methylome database and analysis pipeline to facilitate integrative and comparative epigenomics. PLoS One 8(12):e81148.
.
OpenUrl CrossRef PubMed

[125] Song Q, et al.

[126] ↵
Robinson JT, et al.
(2011) Integrative genomics viewer. Nat Biotechnol 29(1):24–26.
.
OpenUrl CrossRef PubMed

[127] Robinson JT, et al.

[128] ↵
Margolin BS,
Freitag M,
Selker EU
(1997) Improved plasmids for gene targeting at the his-3 locus of Neurospora crassa by electroporation. Fungal Genet Newsl 44:34–36.
.
OpenUrl CrossRef

[129] Margolin BS,

[130] Freitag M,

[131] Selker EU

[132] ↵
Zhou VW
(2012) Methods for Global Characterization of Chromatin Regulators in Human Cells (Harvard Univ Press, Cambridge, MA).
.

[133] Zhou VW

[134] ↵
Jamieson K,
Rountree MR,
Lewis ZA,
Stajich JE,
Selker EU
(2013) Regional control of histone H3 lysine 27 methylation in Neurospora. Proc Natl Acad Sci USA 110(15):6027–6032.
.
OpenUrl Abstract/FREE Full Text

[135] Jamieson K,

[136] Rountree MR,

[137] Lewis ZA,

[138] Stajich JE,

[139] Selker EU

[140] ↵
Jolma A, et al.
(2010) Multiplexed massively parallel SELEX for characterization of human transcription factor binding specificities. Genome Res 20(6):861–873.
.
OpenUrl Abstract/FREE Full Text

[141] Jolma A, et al.

[142] ↵
Fischle W, et al.
(2005) Regulation of HP1-chromatin binding by histone H3 methylation and phosphorylation. Nature 438(7071):1116–1122.
.
OpenUrl CrossRef PubMed

[143] Fischle W, et al.

[144] ↵
Nielsen PR, et al.
(2002) Structure of the HP1 chromodomain bound to histone H3 methylated at lysine 9. Nature 416(6876):103–107.
.
OpenUrl CrossRef PubMed

[145] Nielsen PR, et al.

[146] ↵
James P,
Halladay J,
Craig EA
(1996) Genomic libraries and a host strain designed for highly efficient two-hybrid selection in yeast. Genetics 144(4):1425–1436.
.
OpenUrl Abstract/FREE Full Text

[147] James P,

[148] Halladay J,

[149] Craig EA

[150] Kouzminova EA,
Selker EU
(2001) Dim-2 encodes a DNA-methyltransferase responsible for all known cytosine methylation in Neurospora. EMBO J 20:4309–4323.
.
OpenUrl Abstract/FREE Full Text

[151] Kouzminova EA,

[152] Selker EU

Main menu

User menu

Search

Significance

Abstract

Results

HCHC Plays an Important Role in Centromere Function.

Whole-Genome Bisulfite Sequencing Analysis of the HCHC Mutants.

The CD of HP1 but Not That of CDP-2 Is Required for HCHC Complex Function.

CDP-2 Interacts Directly with the Chromoshadow Domain of HP1.

CDP-2 and CHAP Interact Directly with HDA-1.

CDP-2 Recruits HDA-1 HDAC Activity to H3K9me3 Regions.

HP1 and CDP-2 Localize to Heterochromatin Independently of HDA-1 and CHAP.

CHAP Is Required for the Residual DNA Methylation in the HP1 CD-Deletion Strain.

The CHAP AT-Hook Motifs Are Required for Normal DNA Methylation.

The CHAP AT-Hook Motifs Specifically Bind AT-Rich DNA with High Numbers of Repeat-Induced Point Mutations.

Discussion

Materials and Methods

N. crassa Strains and Molecular Analyses.

Assessment of Chromosome Bridges.

WGBS.

CHAP–DamID Sequencing.

Construction of HA-Tagged CHAP Fusion Constructs Expressed at the pan-2 Locus.

Generation of Recombinant CHAP Proteins and Gel Mobility Shift Assays.

DNA–Protein Affinity Purification.

SI Materials and Methods

Construction of a C-Terminal Dam Fusion Vector for his-3 Targeting and Fusion Constructs for CDP-2, HDA-1, and CHAP.

Construction of the C-Terminal HAT/FLAG–Tagged HP1 CD Constructs Expressed at his-3.

Construction of CDP-2, HDA-1, and CHAP Mutant Constructs.

Construction of LexADBD-Tagged HP1 CD-Deletion Fusion Constructs Expressed at the Native hpo Locus.

Construction of CDP-2 CD-Deletion Constructs Expressed at the Native cdp-2 Locus.

Acknowledgments

Footnotes

References

Citation Manager Formats

Article Classifications

Sign up for Article Alerts

Jump to section

You May Also be Interested in

Similar Articles

Articles

PNAS Portals

Information