The histone octamer is comprised of two copies of each of the four core histones H2A, H2B, H3, and H4, organized with two H2A/H2B dimers associated with a central core tetramer of histone H3 and H4 (Kornberg and Lorch, 1999; Kornberg and Thomas, 1974). The presence of two copies of each histone in the octamer (Luger et al., 1997) raises the potential for three distinct stoichiometries (0, 1 or 2) for any modification, and increasing evidence supports the existence of asymmetrically-modified nucleosomes (Li and Shogren-Knaak, 2008; 2009; Voigt et al., 2012). Using rational design, followed by in vivo optimization and selection, we recently developed a pair of H3 proteins – H3X and H3Y – that form obligate heterodimers (Ichikawa et al., 2017). Shortly thereafter, another group reported an asymmetric H3 interface (H3D and H3H) based on an electrostatic interaction between a single altered amino acid on each half of the heterodimeric pair (Zhou et al., 2017). These systems were used to probe the biological outcomes resulting from packaging of the genome with nucleosomes bearing single or double point mutations, and the findings in the two studies display many similarities. For example, both studies showed that a single H3 N-terminal tail carrying H3K36me3 is sufficient to suppress cryptic transcriptional initiation. Thus, asymmetric nucleosome systems represent a novel approach to mechanistically probe the role of histone stoichiometry in multiple aspects of chromatin biology.

Histone isoforms are an important source of chromatin diversity. Notably, there is a specialized H3 isoform termed CENP-A (or Cse4 in budding yeast) required for centromere function (Foltz et al., 2006; Meluh et al., 1998; Stoler et al., 1995; Sullivan, 2001). Here, we extended the utility of our asymmetric H3 interface to explore the stoichiometry of Cse4 domains required for centromeric nucleosome function. The question of Cse4 stoichiometry has received particular attention (reviewed in [Biggins, 2013]). Octomeric Cse4 nucleosomes, with two copies of Cse4 in place of H3 can be reconstituted from recombinant components in vitro (Camahort et al., 2009; Dechassa et al., 2011; Kingston et al., 2011) and display a canonical nucleosome-like structure when analyzed at high resolution via X-ray crystallography (Sekulic et al., 2010). Microscopy-based experiments (Aravamudhan et al., 2013; Wisniewski et al., 2014) have also been consistent with octomeric Cse4 nucleosomes. However, crosslinking and atomic force microscopy studies (Dalal et al., 2007), reconstitution experiments (Furuyama et al., 2013), and chromatin digestion experiments (Henikoff et al., 2014) had suggested that Cse4/CENP-A forms a tetrameric half-nucleosome, raising the question of which mode of Cse4 interaction is important for chromosome segregation. Using our tools, we confirm that an octomeric centromeric nucleosome exists and is essential for viability in budding yeast, and show that important residues on both the N- and C-terminal Cse4 tails function when present at one copy per pair.

Results and discussion

All eukaryotes use a specialized histone H3 variant (CENP-A, or Cse4 in budding yeast) to form centromeric nucleosomes. CENP-A proteins are essential because they provide the fundamental attachment between chromosomes and kinetochore proteins, thereby ensuring equal chromosome segregation at mitosis (Biggins, 2013; Buchwitz et al., 1999; Meluh et al., 1998; Palmer et al., 1987; Stoler et al., 1995). Cse4/CENP-A proteins have a histone-fold globular domain similar to that of H3, with more divergent N- and C-terminal extensions (Chen et al., 2000; Keith et al., 1999; Tachiwana et al., 2011). To explore the stoichiometry of domains required for Cse4 function, we have generated a heterodimeric ‘Cse4X’ and ‘Cse4Y’ pair (Figure 1A).

Figure 1 with 5 supplements see all

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We first attempted to place the X and Y asymmetric mutations we had developed in the canonical H3 protein onto wild-type Cse4. However, these constructs (termed Cse4X’ and Y’) would not support viability in the absence of a functional, homodimeric Cse4 protein (Chen et al., 2000) (Figure 1—figure supplement 1). We reasoned that the X and Y alterations may function only in the context of the canonical H3-H3 interface. Because the asymmetric Y alterations are present in both the α2 and α3 helices of the histone fold domain (Ichikawa et al., 2017; Figure 1A), we tested the X and Y alterations in a chimeric Cse4 in which key residues in these helices were substituted with canonical H3 residues. We used a chimeric Cse4 (‘Chimera’ in Figure 1A; originally termed the cse4-337 allele (Keith et al., 1999) which had previously been shown to support viability (Keith et al., 1999); also see Figure 1B). Inserting the X and Y asymmetric residues into this Chimera, we generated the Cse4X and Cse4Y proteins (Figure 1A). Notably, cells simultaneously expressing Cse4X and Cse4Y were viable, but those expressing a single one of these alone were not (Figure 1B). These data support the idea that at least some of Cse4 functions are performed as part of an octameric centromeric nucleosome, consistent with previous structural, biochemical and cell biological studies (Aravamudhan et al., 2013; Black and Cleveland, 2011; Camahort et al., 2009; Dechassa et al., 2011; Kingston et al., 2011; Wisniewski et al., 2014; Xiao et al., 2017; Zhang et al., 2012). Further, these data suggested that the centromeric H3 isoform is amenable to analysis via our asymmetric mutations.

We note that the Chimera protein itself conferred a partial temperature-sensitive growth phenotype, in which colony size was markedly reduced at 37°C, but much less so at 34°C (Figure 1C). Adding the X and Y alterations to the Chimera protein very modestly increased temperature sensitivity at 37°C, but did not increase the sensitivity of the cells to benomyl, a microtubule destabilizing drug (Davidse and Flach, 1977). To test whether an alternative protein design could provide a less perturbed setting for these studies, we compared the X-Y interface to a different pair of asymmetric H3 constructs (Zhou et al., 2017) that were described soon after our study was published. That study tested pairs of oppositely charged electrostatic residues at key contact points between nucleosomal H3 molecules, finding one pair of asymmetric H3s that would support viability. This pair is comprised of ‘H3^D’, which contains an A110D replacement, and ‘H3^H’, which contains a L130H replacement. We generated Cse4 derivatives with either the D or H single amino acid substitutions (Figure 1—figure supplement 2, Cse4D’ and Cse4H’), and we also made D or H substitutions in the context of the Chimera used above (Figure 1—figure supplement 2, Cse4D and Cse4H). Notably, we found that after removing homodimeric Cse4, cells expressing either the Cse4H’ or the Cse4H protein without any partner protein were viable and grew robustly, indicating significant homodimerization of this protein (Figure 1—figure supplement 2B). This led us to test the reported D-H interface in the context of canonical histone H3. We observed that H3^H would also support growth of cells lacking all other sources of histone H3, although growth was faster when both H and D were present (Figure 1—figure supplement 3). In contrast, as we had observed before (Ichikawa et al., 2017), no growth of cells expressing only H3X or H3Y was observed. Not only do our findings imply significant H3^H homodimerization in vivo, but we find no improvement in temperature sensitivity in the D-H system, as cells expressing H3^D + H3^H were also unable to grow at 37°C (Figure 1—figure supplement 4). We therefore performed all subsequent experiments on Cse4 heterodimerization using the Cse4X and Cse4Y pair.

We also tested for heterodimerization of Cse4X and Cse4Y via biochemical experiments. To do this, we expressed three distinct epitope-tagged X or Y proteins simultaneously in the same cell, and used co-immunoprecipitation to analyze dimerization. We had taken a similar approach in the analysis of H3 (Ichikawa et al., 2017). In the present case, the poor solubility of the Cse4 nucleosome (Skene and Henikoff, 2017) led us to use a different protocol for the preparation of cell extracts. Specifically, we modified existing protocols and generated high-concentration MNase-treated extracts (Furuyama and Biggins, 2007; Ichikawa et al., 2014; Nelson and Fangman, 1979) as the starting point for immunoprecipitation. Upon anti-FLAG epitope immunoprecipitation, we easily detected the expected heterodimeric interactions (i.e. HA-Cse4Y with FLAG-Cse4X in strain PKY5527 cells and V5-Cse4X with FLAG-Cse4Y in PKY5529 cells) but not homodimeric interactions (Figure 1—figure supplement 5). Together, our genetic experiments (Figure 1B) and the co-immunoprecipitation experiments (Figure 1—figure supplement 5) led us to conclude that we had designed a specific heterodimeric Cse4X-Cse4Y nucleosome.

We next used our asymmetric proteins to analyze Cse4 domain structure. The long N-terminal tail of Cse4 contains an Essential N-terminal Domain (END) between residues 28 and 60 (Chen et al., 2000; Keith et al., 1999, Figure 2A), which is a binding site for the Ctf19-Mcm21-Okp1 ‘COMA’ kinetochore protein complex (Chen et al., 2000). Complete removal of the END domain via deletion of the N-terminal 70 residues of homodimeric, wild-type Cse4 generated a protein (Cse4 Δ70) that does not support viability (Figure 2B), consistent with previous observations (Chen et al., 2000; Keith et al., 1999). The END domain was also essential in the context of the Cse4 X-Y system, as cells expressing paired Cse4 proteins containing the Δ70 deletion on both X and Y were also inviable. In contrast, cells with one intact END domain on either Cse4X or Cse4Y were viable. To further assess the key biological function of the centromeric nucleosome, we measured centromeric plasmid maintenance, which depends on a cloned centromeric sequence (CEN; [Murray and Szostak, 1983]). CEN is the binding site for the Cse4 nucleosome, and is thereby essential for kinetochore-mediated chromosome segregation (Camahort et al., 2009; Meluh et al., 1998); reviewed in [Biggins, 2013]). We observed that cells with a single END-Cse4 construct displayed plasmid loss rates that were statistically indistinguishable from those observed in the presence of the ‘pseudo-wildtype’ Cse4 X + Y pair (Figure 3). These data indicate that a single END domain per Cse4 nucleosome is sufficient for building functional kinetochores. However, these cells displayed partial temperature sensitivity at 34°C and sensitivity to benomyl, suggesting that this architecture is suboptimal under stress conditions (Figure 2C). These data confirm that asymmetric Cse4 proteins, like asymmetric H3 proteins, can be used to probe domain stoichiometry in living cells.

Figure 2

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Figure 3

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Next, we analyzed motifs at the Cse4 C-terminus. The terminal three residues of Cse4, QFI, are distinct from the ERS residues at the end of H3 (Figure 4A), and comprise part of the interaction surface for the centromeric protein CENP-C (Mif2 in budding yeast; (Kato et al., 2013), which is critical for kinetochore function (Meeks-Wagner et al., 1986; Przewloka et al., 2011). We first exchanged ERS residues onto the C-termini of the homodimeric Chimera Cse4 protein and onto the heterodimeric Cse4X and Cse4Y constructs (Figure 4A). We observed that cells remained viable in the absence of QFI residues, consistent with previous studies (Keith et al., 1999). However, cells expressing only Cse4 proteins with ERS C-termini (e.g. the Chimera-ERS and X-ERS +Y ERS strains) displayed slow growth at 30°C and extreme sensitivity to elevated temperatures or benomyl (Figure 4B). X-ERS +Y ERS cells also displayed significantly elevated plasmid loss rates compared to the Cse4 X + Y pseudo-wildtype cells (Figure 3). In contrast, X-ERS +Y and X + Y ERS strains with a single ERS domain per Cse4 nucleosome did not display temperature or benomyl sensitivity (Figure 4B) or elevated plasmid loss rates (Figure 3). Thus, our asymmetric Cse4 tools allowed us to conclude that a single QFI tail per Cse4 nucleosome was sufficient to maintain normal centromere function and prevent stress sensitivity phenotypes.

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Recent experiments indicate that in addition to the C-terminus, unique residues within Cse4 Loop1, between the α1 and α2 alpha helices of the histone fold domain, are required for optimal Mif2 binding to Cse4 (Xiao et al., 2017). Therefore, we investigated how Loop1 mutations synergize with ERS substitutions in our asymmetric system. Notably, Loop1 in Cse4 is three residues longer than in H3, with an insertion of residues KDQ, and there is also a substitution of an evolutionarily conserved H3 phenylalanine for a tryptophan residue (Figure 4—figure supplement 1A). Previous studies have shown that simultaneous deletion of the KDQ residues and conversion of the W to an F residue is tolerated, but results in a ~ five fold increase in chromosome loss rates (Keith et al., 1999). In our experiments, combination of the Loop1 and ERS mutations (termed ERS*) in the context of Cse4-Chimera resulted in lethality, as did the presence of the ERS* combination on both halves of the X + Y asymmetric pair (Figure 4—figure supplement 1B). In contrast, the combined ERS* mutations could support viability if present only on a single half of the asymmetric Cse4 pair, although there was an increase in plasmid instability, temperature sensitivity and benomyl sensitivity in these strains (Figure 3, Figure 4—figure supplement 1C). These data are consistent with our earlier conclusion that a single intact Mif2 binding site per centromeric nucleosome is sufficient for viability.

We also tested the effects of combining a single N-terminal deletion (Δ70) and a single set of ERS* substitutions in cis or trans configurations within the asymmetric Cse4 nucleosome (Figure 4—figure supplement 2). Viable cells in both the cis and trans mutants could be obtained in the initial selection against the plasmid encoding homodimeric Cse4 (Figure 4—figure supplement 2A). We note that the two ‘trans’ mutant combinations may produce FOA-resistant colonies at different frequencies, perhaps because of combining the X mutations in alpha helix 3 with the C-terminal ERS mutations in the same molecule. In contrast to any small differences between the trans strains, the major observation was that none of the cis or trans strains could be propagated for further analysis (Figure 4—figure supplement 2B). These results suggest that simultaneous removal of one interaction site for the COMA complex at the Cse4 N-terminus and one interaction site for CENP-C at the Cse4 C-terminus resulted in a highly compromised Cse4 nucleosome. In other words, a single wild-type N-terminus and C-terminus per Cse4 nucleosome appears insufficient to prevent lethal levels of genome instability.

Additionally, because some of our altered Cse4 proteins were affecting viability and plasmid stability, we wondered if the asymmetric interface was undergoing sufficient negative selection during the course of these experiments to select for revertants different from the input constructs. To test for this, we amplified the CSE4 genes from the indicated strains listed in Supplementary file 1 (Figure 4—figure supplement 3). In no case did we observe alterations from the input plasmid sequence. We conclude that the phenotypes observed throughout this work result from the designed constructs and not due to intragenic suppressor mutations in CSE4.

In summary, we show here that our asymmetric nucleosome technology is well-suited to explore histone H3 variants. In analogy to our H3 studies, these reagents can be used to study Cse4 modifications, for example S33 phosphorylation (Hoffmann et al., 2018), K37 methylation (Samel et al., 2012) or phosphorylation by Ipl1 (aurora kinase) (Boeckmann et al., 2013). Of course, CENP-A stoichiometry and modification is also important in mammalian cells, but involves much more complex centromeres (Müller and Almouzni, 2017), and it will be of great interest to explore nucleosomal symmetry in those systems.

All data generated or analysed during this study are included in the manuscript and supporting files.

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Author details

1. Yuichi Ichikawa

   1. Department of Molecular, Cell, and Cancer Biology, University of Massachusetts Medical School, Worcester, United States
   2. Division of Cancer Biology, The Cancer Institute of JFCR, Tokyo, Japan

   Present address

   Division of Cancer Biology, The Cancer Institute of Ariake, Tokyo, Japan

   Contribution

   Conceptualization, Investigation, Methodology, Writing—original draft, Writing—review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-6773-2455

2. Noriko Saitoh

   Division of Cancer Biology, The Cancer Institute of JFCR, Tokyo, Japan

   Contribution

   Supervision, Funding acquisition, Writing—review and editing

   Competing interests

   No competing interests declared

3. Paul D Kaufman

   Department of Molecular, Cell, and Cancer Biology, University of Massachusetts Medical School, Worcester, United States

   Contribution

   Conceptualization, Supervision, Funding acquisition, Investigation, Writing—original draft, Writing—review and editing

   For correspondence

   paul.kaufman1@umassmed.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-3089-313X

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