Chromosoma DOI 10.1007/s00412-014-0504-7
REVIEW
Histone storage and deposition in the early Drosophila embryo Béatrice Horard & Benjamin Loppin
Received: 3 October 2014 / Revised: 17 December 2014 / Accepted: 18 December 2014 # Springer-Verlag Berlin Heidelberg 2015
Abstract Drosophila development initiates with the formation of a diploid zygote followed by the rapid division of embryonic nuclei. This syncytial phase of development occurs almost entirely under maternal control and ends when the blastoderm embryo cellularizes and activates its zygotic genome. The biosynthesis and storage of histones in quantity sufficient for chromatin assembly of several thousands of genome copies represent a unique challenge for the developing embryo. In this article, we have reviewed our current understanding of the mechanisms involved in the production, storage, and deposition of histones in the fertilized egg and during the exponential amplification of cleavage nuclei. Keywords Histone . Drosophila . Embryo . Chromatin . Storage . Nucleosome assembly
Introduction The organization of DNA into regular arrays of octameric nucleosomes is a distinctive and almost universal feature of eukaryotes. This type of chromatin organization was possibly selected to package large and segmented genomes into the confined nuclear environment. In addition, nucleosomes are much more than packaging structures. They combine key properties that altogether allow the genome to perform essential tasks such as DNA replication, chromosome condensation, and segregation. They also play a genome-indexing role for the repair of DNA lesions, the regulation of gene B. Horard : B. Loppin (*) Centre de Génétique et de Physiologie Moléculaire et Cellulaire—CNRS UMR5534, Université Claude Bernard Lyon 1, University of Lyon, 69100 Villeurbanne, France e-mail:
[email protected]
transcription, or the formation of heterochromatin. Histones are at the heart of this versatility. The existence of non-allelic histone variants combined with the expanding universe of histone post-translational modifications potentially provides an enormous source of nucleosome diversity whose functional relevance is far from being understood (Talbert and Henikoff 2010; Banaszynski et al. 2010). Another layer of complexity lies in the regulation of different histone biosynthesis and deposition pathways involved in various outcomes for chromatin metabolism. These different pathways are distinguished by the implication of specific histone chaperones, nucleosome assembly factors, and histone variants (De Koning et al. 2007). However, they also share common components at different levels of nucleosome assembly. Our understanding of how these different pathways are orchestrated for the timely assembly of different types of nucleosomes at the correct genomic position is still limited. Drosophila is one of the key model organisms for chromatin biology, and its contribution to the field is of primary importance. The relatively large size of Drosophila nuclei combined with a small number of chromosomes (2n=8) facilitates imaging approaches, and the existence of giant polytene chromosomes is a fly specialty very useful for chromatin biologists. Importantly, Drosophila melanogaster has one of the smallest sets of histones, with only five canonical histones and four variants known so far. The early fly embryo yet represents by itself a fascinating model for chromatin biology and histone dynamics in vivo. The syncytial phase of embryonic development is almost entirely under maternal control and is characterized by one of the most rapid nuclear amplification known in animals. In the virtual absence of zygotic transcription, these cleavage nuclei simply alternate S and M phases as they progressively invade the embryo periphery. This rapid amplification of cleavage nuclei thus represents a powerful in vivo model for the study of replication-coupled (RC) nucleosome assembly. Replication-independent (RI)
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repackaging of the paternal genome at fertilization is the other major chromatin assembly event at the onset of development. Although it occurs in a single nucleus, this unique wholegenome assembly is critical for the integration of paternal chromosomes in the zygote. In this article, we have reviewed the strategies employed by the early Drosophila embryo to produce, store, and deposit histones from fertilization to blastoderm formation, with a special focus on the more recent advances of this dynamic field.
Genomic organization and expression of Drosophila histone genes The minimal set of Drosophila histones Among multicellular model organisms, Drosophila has the smallest diversity of histones (Table 1). The five canonical or replicative histones (H1, H2A, H2B, H3, and H4) are
Table 1
Core
Linker
encoded by a cluster of histone genes located near the centromere of chromosome 2. Replicative histone genes are organized in gene units of about 5 kb containing one copy of each gene (Fig. 1). The D. melanogaster reference genome sequence indicates the presence of 23 units in the cluster (Flybase; Günesdogan et al. 2010). These intronless genes are transcribed during S phase for the production of canonical histones. Canonical histone messenger RNAs (mRNAs) are generally not polyadenylated. Instead, their 3′ end forms a stem loop secondary structure that is specifically recognized by stem loop binding protein (SLBP), which regulates their translation (reviewed in Marzluff et al. 2008). The fly replicative histone H3 is actually identical to human H3.2 and differs from human H3.1 (the main replicative histone) at a single residue (Fig. 1). Note that although these histones are commonly referred to as RC or replicative histones, at least some of them are also deposited in a RI manner. This is the case for H2B, which does not have a replacement variant and, presumably, for H4 (Talbert and Henikoff 2010). Interestingly,
The minimal set of Drosophila histones Drosophila
Human phylogenetic homolog
Phenotype in D. melanogaster
Deposition pathway
Identified (or putative) histone chaperones in D. melanogaster
H3/H3.2
H3.2
RC
CAF-1, ASF1 (Tyler et al. 2001)
H3.3
H3.3
Deletion of histone gene cluster causes embryonic lethality (Günesdogan et al. 2010). Double null mutants have reduced viability and are sterile (Sakai et al. 2009; Hödl and Basler 2009)
RI
HIRA/YEM (Loppin et al. 2005; Orsi et al. 2013) (ASF1) ATRX/XNP (Schneiderman et al. 2012) (Daxx-like-protein)
CID
CENP-A
Null mutants die during embryogenesis (Blower et al. 2006)
RI
CAL1 (Erhardt et al. 2008; Schittenhelm et al. 2010; Mellone et al. 2011; Chen et al. 2014)
H4
H4
Deletion of histone gene cluster causes embryonic lethality (Günesdogan et al. 2010)
RC/RI
CAF-1, ASF1 (Tyler et al. 2001) HIRA/YEM (Loppin et al. 2005; Orsi et al. 2013)
H4r H2B
H4 H2B
Unknown Deletion of histone gene cluster causes embryonic lethality (Günesdogan et al. 2010)
RI? RC/RI
Unknown NAP-1 (Ito et al. 1996a) (NLP) (Ito et al. 1996b; Crevel et al. 1997; Namboodiri et al. 2003)
H2A
H2A
Deletion of histone gene cluster causes embryonic lethality (Günesdogan et al. 2010)
RC/RI
NAP-1 (Ito et al. 1996a) (NLP) (Ito et al. 1996b; Crevel et al. 1997; Namboodiri et al. 2003)
H2A.Z (H2Av)
H2A.Z
RC/RI?
(NAP-1) (NLP)
H1
H1
RC/RI?
Unknown
Big H1
–
Null mutants die at 3d instar larvae (van Daal and Elgin 1992) RNAi depletion causes late larval-pupal lethality associated with altered pericentric heterochromatin organization (Lu et al. 2009) Null mutants die at the blastoderm stage and show premature zygotic genome activation (Pérez-Montero et al. 2013)
RC/RI?
Unknown
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however, in addition to its 23 copies located in the histone gene cluster, H4 is also represented by an additional gene on chromosome 3 named H4r (H4 replacement) (Akhmanova et al. 1996). H4r has a standard gene organization, is abundantly expressed in all tissues, and encodes a histone H4 identical to canonical H4. Although H4r could potentially provide a source of H4 for RI assembly pathways, its function is currently unknown. Remarkably, Drosophila has only four histone variants (bigH1, H2A.Z, H3.3, and Cid) (Fig. 1). Among them, H3.3 is the most conserved histone variant and is identical to human
H3.3. As in mammals, Drosophila H3.3 is encoded by two single copy genes: His3.3A (on chromosome 2) and His3.3B (on chromosome X). The other H3 variant is the centromeric histone H3 encoded by the centromere identifier (cid) gene. The single fly histone H2A variant (van Daal and Elgin 1992; Leach et al. 2000) is phylogenetically related to H2A.Z (Talbert et al. 2012). Here, we chose to name it H2A.Z, but it is also commonly referred to as H2Av (for a recent review on H2A.Z/H2Av, see Baldi and Becker 2013). Although Drosophila H2A.Z shares extensive homology with other H2A.Z histones, it also presents an extended tail domain
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Fig. 1
Canonical histones and their variants. a Genomic organization of histone genes in Drosophila. A total of 23 histone gene repeat units, each containing a single His1, His2B, His2A, His4, and His3 gene, are clustered in a histone complex located near the centromere of chromosome 2L. Colored boxes represent coding regions and gray boxes represent UTRs. The five canonical intron-less histone genes are transcribed during S phase. Replicative histone mRNAs are usually not polyadenylated but, instead, end in a conserved stem loop. On the contrary, histone variant genes His3.3A, His3.3B, cid, BigH1, and His2A.Z are single-copy genes scattered throughout the genome. The histone variant genes possess introns and their polyadenylated mRNAs are expressed throughout the cell cycle. Their corresponding transcripts are shown; thin lines representing known or putative introns. The two H3.3 genes (His3.3A and His3.3B) encode the same protein, but the transcripts have distinct untranslated regions. The intron-less centromere identifier (cid) gene encodes the centromeric histone H3. The single-copy His4r gene encodes a histone H4 identical to canonical H4. His4r has a standard gene organization, its mRNA is polyadenylated, and it is expressed independently of DNA synthesis. b Schematic representation of histone H3, H2A, and H1 proteins and their variants. The major core histones and their variants contain a conserved histone-fold domain (HFD), a protein dimerization motif. In human, two versions of the canonical H3 exist: H3.1 and H3.2, which differ by only a single amino acid at position 96. The Drosophila replicative histone H3 is identical to mammalian H3.2. The histone variant H3.3 differs from H3.2 (H. s or D. m) by four amino residues: one resides in the N-terminus tail (31) and three others are located in the histone-fold domain (87, 89, and 90). Drosophila has only one H2A variant, which shares extensive homology with human H2A.Z histone. However, Drosophila H2A.Z (also known as H2Av) has an extended C-terminus domain containing a SQAY motif. This motif resembles the SQ[E/D]Φ motif found in the C-terminus of H2A.X. Drosophila linker histone H1 has a short amino-terminal domain, a globular winged-helix domain (GB) and a longer C-terminus domain. BigH1 has the same tripartite structure, with the GB domain showing a high degree of similarity (57 %) with respect to H1. Differences in amino acid sequences are essentially present in the extended N-terminal domain of BigH1 and, to a lesser extent, in its Cterminus and contribute to the relatively low overall homology to H1 (~30 % similarity) (see Pérez-Montero et al. 2013)
containing an SQAY motif resembling the SQ[E/D]Φ motif found in the C-terminus of H2A.X variants (Clarkson et al. 1999) (Fig. 1). In fact, Drosophila H2A.Z is phosphorylated at serine 137 (γH2A.Z) in response to DNA double-strand breaks, indicating that it independently acquired the same property as vertebrate H2A.X (Madigan et al. 2002). The Drosophila histone set is also unusual in lacking germline-specific histone variants. Indeed, animals and plants frequently posses testis-specific variants of H1, H2A, H2B, or H3 (Gaucher et al. 2010; Ingouff and Berger 2010; Rathke et al. 2014). In mammals, for instance, H1t, HILS1, H2AL1/2, TH2B, and H3t are all specifically incorporated in spermatid nuclei before or during the histone-to-protamine transition. However, despite the absence of testis-specific variants, Drosophila spermatid nuclei undergo a massive replacement of nucleosomes with sperm-specific nuclear proteins such as Mst77F, which is related to the mouse HILS1 gene, and Mst35Ba/b, two protamine-like proteins (Rathke et al. 2014). In addition, a new H1-related histone, BigH1, was recently discovered in Drosophila (Pérez-Montero et al.
2013). BigH1 is abundant in early embryos where it plays an essential role in repressing zygotic genome activation (ZGA) until its replacement by H1 at cellularization. Although bigH1 is not specifically expressed in gonads, it is nevertheless restricted to primordial germ cells throughout embryo development where it delays ZGA.
Histone production and storage in the early embryo The Drosophila egg is a gigantic cell that contains all the stockpiles of proteins, RNAs, ribosomes, lipids, and metabolites required for the syncytial phase of early embryo development. Indeed, the bulk of zygotic transcription essentially begins by the time of blastoderm cellularization, about 150 min after fertilization (Foe et al. 1993). The blastoderm is formed by the progressive outward migration of rapidly dividing cleavage nuclei. After 13 syncytial nuclear divisions, about 4000 nuclei are formed exclusively from maternally provided components, including histones. ZGA and cellularization then occur at the onset of cycle 14. The early nuclear cycles in Drosophila embryos are among the fastest known in animals. In the absence of distinctive G phases, S and M phases alternate synchronously, each cycle lasting about 9 min at 25 °C. This phase of nuclear amplification thus represents a unique and abundant source of nuclei undergoing “pure” replication-coupled chromatin assembly. The exponential accumulation of replication forks at each cycle supposes a constant supply of histones and chromatin assembly factors to nuclei in S phase. In contrast to the general context of tight temporal coupling of histone production and DNA replication (reviewed in Jasencakova and Groth 2010), the syncytial embryo must use its maternal supplies without any additional transcription of histone genes. Despite the extremely large volume of the egg cell, the accumulation of excess histones proteins is potentially toxic. In fact, overexpression of histones during Drosophila oogenesis, like in the abnormal oocyte (abo) maternal effect mutant, is associated with embryonic lethality (Berloco et al. 2001). On the contrary, a full deletion of the histone gene cluster is zygotic lethal immediately following the exhaustion of maternal histone mRNAs at nuclear cycle 14 of embryogenesis (Günesdogan et al. 2010). Thus, the supply of maternal histones must be tightly regulated to provide enough material for the assembly of thousands of nuclei and, at the same time, to avoid detrimental effects of histone overproduction. Translational regulation of histone biosynthesis The storage of histone mRNAs in the mature oocyte represents the first level of regulation for the maternal supply of histones to the developing embryo. In Drosophila, canonical
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histone gene expression is coupled to DNA synthesis except during late oogenesis. Indeed, histone mRNAs are massively synthesized after the last cycle of DNA replication in nurse cell nuclei (stage 10 of oogenesis) and then transported to the growing oocyte where they accumulate (Ambrosio and Schedl 1985; Walker and Bownes 1998). A large proportion of these accumulating mRNAs are associated to polysomes without delay after their synthesis (Ruddell and Jacobs-Lorena 1985). Like the histone genes in other metazoans, the canonical histone mRNAs are not primarily polyadenylated but instead end with a stem-loop sequence which is specifically bound by the conserved SLBP protein (Sullivan et al. 2001; Lanzotti et al. 2002). The binding of SLBP to the 3′ end of canonical histone mRNAs not only participates in histone mRNA processing but also in their translation (Sanchez and Marzluff 2002). SLBP is expressed at all stages of Drosophila life cycle, consistently with a function in all cycling cells, but is particularly abundant in developing egg chambers where it is required for the maternal production of mature histone mRNAs and histone proteins. Interestingly, SLBP is not detected in syncytial preblastoderm embryos, suggesting that the histone supply during early nuclear cycles does not rely on the translation of histone mRNAs but on a pool of histone proteins (Lanzotti et al. 2002, 2004; Iampietro et al. 2014). However, females bearing a hypomorphic Slbp allele (Slbp10) in trans to a deficiency are viable but sterile (Sullivan et al. 2001). Eggs laid by Slbp10/Df females have a dramatically reduced pool of maternal histone mRNA and protein, and mutant embryos exhibit mitotic defects and loss of cortical nuclei (nuclear fallout) (Sullivan et al. 2001; Lanzotti et al. 2002). Recently, it was demonstrated that the nuclear fallout defect is at least partially caused by impaired synthesis of zygotic histone proteins. Indeed, disruption of SLBP function was associated with nuclear retention of early zygotic histone mRNAs at cellularization (Iampietro et al. 2014). This mRNA nuclear retention leads to depletion of zygotic histone proteins when maternal histone proteins may become limiting, with devastating consequences for nuclear integrity (Iampietro et al. 2014). Histone storage in lipid droplets In addition to the translational regulation of stored histone mRNAs, Drosophila embryos have an original way to regulate the maternal histone supply at the protein level. Indeed, in early embryos, 50 % or more of the total embryonic pool of histones H2A, H2A.Z, and H2B are associated with lipid droplets and ubiquitous fat storage organelles (Cermelli et al. 2006; Welte 2007). These maternal histones are anchored to lipid droplets via a droplet-associated protein named Jabba (Li et al. 2012). The Jabba-dependent association of histones with droplets begins in the germline nurse cells, where these structures are initially produced. It is thus likely that histones are
targeted to lipid droplets shortly after their synthesis and stored into the oocyte already bound to Jabba. In the absence of Jabba, maternal histone mRNA synthesis is unaffected, but H2A and H2B proteins do not accumulate in the egg, likely as the result of degradation of unbound cytoplasmic histones (Li et al. 2012). Despite this initial lack of maternal histone deposits, the overall amount of histone protein is similar in wild type and in embryos produced by Jabba mutant females (called Jabba embryos for simplicity) as they reach the blastoderm stage when zygotic transcription resumes (Li et al. 2012). The viability of Jabba mutant embryos suggests that the maternal pool of histones H2A, H2A.Z, and H2B is not essential for early development. Analysis of double Jabba slbp mutants however revealed that when the amount of stored histone mRNAs is mildly reduced, histones stored in lipid droplets become essential to ensure early embryogenesis (Li et al. 2012). Jabba embryos are thus able to compensate for defective histone protein storage by increasing histone synthesis from the stored maternal histone mRNAs. A regulatory crosstalk between storage and synthesis of histones likely participates in the fine-tuning of maternal histone supply during cleavage divisions. Interestingly, in their most recent work, Li and coworkers (2014) reported an additional role of lipid droplets in regulating the balance between H2A and its related variant H2A.Z in embryos. In the absence of Jabba, cycles 10 to 13 embryos display an increased level of H2A.Z in their nuclei and this is associated with elevated mitotic defects and reduced viability and hypersensitivity to H2A.Z overexpression (Li et al. 2014). It thus appears that lipid droplets, by transiently sequestering H2A.Z at the end of syncytial development, control the H2A.Z/H2A ratio in the nuclei for appropriate chromatin assembly. It now remains to understand how these storage organelles regulate their differential association with H2A and H2A.Z and how they release histones for their timely deposition at DNA replication forks. Since charge interactions seem to play an essential role in localizing histones to the droplets, one can wonder whether post-translational modifications, e.g., acetylation or phosphorylation that bring negative charges, could control histone-droplets dissociation (Cermelli et al. 2006), yet proteomic analyses have not detected any histone chaperone associated with droplet-localized histones (Li et al. 2012). Nevertheless, Jabba-bound histones must be handed over to downstream cytoplasmic histone chaperones on the route to replicating nuclei. Nucleosome assembly protein 1 (Nap1) is a conserved histone chaperone that could potentially play this role. It preferentially binds H2A and H2B in vivo and promotes import of H2A-H2B dimers from the cytoplasm to the chromatin assembly machinery in the nucleus (Ito et al. 1996a; Keck and Pemberton 2012). Although the role of NAP1 during Drosophila embryogenesis has not been studied in details, its maternal expression is important for viability (Lankenau et al. 2003). However, NAP1 has been recently shown to have
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additional role in spermiogenesis that could be distinct from its histone chaperoning activity (Kimura 2013). Drosophila nucleoplasmin-like protein (NLP) is another H2A-H2B histone chaperone abundant in early embryos, which is related to Xenopus nucleoplasmin (NP), the founding member of the family (Philpott et al. 1991; Ito et al. 1996b; Crevel et al. 1997). NP is present in large amounts in frog oocytes where it serves as a storage protein for H2A/H2B (Finn et al. 2012). Structural studies have proposed that both NP and NLP assemble into pentameric or decameric structures to stabilize all four core histones (Dutta et al. 2001; Namboodiri et al. 2003), and Drosophila NLP facilitates nucleosome assembly in vitro (Ito et al. 1996b; Crevel et al. 1997). More recently, and quite unexpectedly, NLP was identified as a key mediator of centromere clustering at the periphery of the nucleolus in somatic cells (Padeken et al. 2013). It will thus be difficult to understand the actual role of this histone chaperone in embryos without a proper functional analysis. Finally, the chromatin remodeling Tip60 complex and associated factor remodeling and spacing factor (RSF) can mediate H2A.Z replacement on chromatin (Kusch et al. 2004; Hanai et al. 2008). However, these factors interact with H2A.Z on chromatin and likely do not play any role in chaperoning H2A.Z in the cytosol.
(Nabeel-Shah et al. 2014), thus paving the way for the investigation of its role during syncytial development. The implication of very different mechanisms to store and handle maternal stockpiles of H2A-H2B and H3.2-H4, respectively, is intriguing. Indeed, these different core histones will ultimately associate at equal stoichiometry to form nucleosome particles. Nucleosome assembly is initiated with the formation of a (H3-H4)2 tetramer on DNA followed by the deposition of two H2A-H2B (or H2A.Z-H2B) dimers to complete the octameric particle. The storage of H3.2-H4 and H3.3-H4 dimers with chaperones, rather than in lipid droplets, could possibly exert a more direct control on their delivery to their respective nucleosome assembly pathways, as detailed in the next section. In contrast, H2A-H2B or H2A.Z-H2B dimers stored in lipid droplets could be indistinguishably mobilized for any mode of nucleosome assembly. Finally, it should be mentioned that histones can be released from lipid droplets in the presence of intracellular bacteria to mount an efficient antimicrobial defense (Anand et al. 2012). Jabba mutant embryos indeed poorly survive infections with at least four different pathogenic microbes. This most surprising role of histones nicely illustrates the remarkable adaptability of the Drosophila embryo to unexpected fluctuations in histone supply.
Chaperone-dependent storage of histone H3 and H4? A surprising aspect of histone storage in lipid droplets is the absence of H3 and H4 in these structures (Cermelli et al. 2006). Maternal H3 and H4 are obviously also abundant in embryos, and their presence is independent of Jabba (Li et al. 2012). This implies that alternative mechanisms are operating to store H3 and H4 before their deposition into chromatin. Numerous studies carried out either in yeast or in human cells have identified several H3-H4 cytosolic histone chaperones (Keck and Pemberton 2012). Among the identified subcomplexes, the nuclear autoantigenic sperm protein (NASP) appears critical to maintain a reservoir of soluble H3-H4 (Campos and Reinberg 2010; Finn et al. 2012). NASP is a homodimeric chaperone that binds H3.1-H4 dimers in human cells and controls the availability of newly synthesized histones that are handed over to another histone chaperone, antisilencing factor 1 (ASF1). The fine-tuning of the soluble H3-H4 reservoir by NASP involves a dynamic balance between histone protection and degradation by chaperonemediated autophagy (Cook et al. 2011). Interestingly, NASP shares extensive sequence homology with the Xenopus N1/N2 histone chaperone implicated in histone H3/H4 storage in oocytes (Kleinschmidt et al. 1985). Furthermore, the knockout of the mouse NASP gene causes embryonic lethality (Richardson et al. 2006). NASP could be a good candidate chaperone for the storage of maternally expressed H3.2 and H4 histones in the early fly embryo. Recently, an extensive phylogenetic analysis identified a NASP-related protein in Drosophila
Nucleosome assembly pathways operating in the early Drosophila embryo Replication-coupled nucleosome assembly The early embryo is obviously an interesting model for the study of in vivo chromatin assembly. In the absence of transcription, histone deposition during cleavage divisions essentially occurs through a RC pathway. The conserved heterotrimeric complex chromatin assembly factor-1 (CAF1) is responsible for H3.1/2 and H4 deposition at DNA replication forks and during DNA repair (Ridgway and Almouzni 2000; Loyola and Almouzni 2004) and is represented in Drosophila by the 3 subunits p180, p105, and p55; a portion of p105 is proteolytically processed to p75 (Tyler et al. 2001). Null alleles of all three Caf1 genes are zygotic lethal at the larval stage, and all show severe growth defects (Song et al. 2007; Klapholz et al. 2008; Anderson et al. 2011; Wen et al. 2012; Yu et al. 2013). Mutant larvae die after the exhaustion of the maternally contributed CAF-1 subunits, which are most likely required for RC nucleosome assembly in cleavage nuclei. In addition, analysis of ovarian germline Caf1-180 mutant clones revealed an essential requirement of CAF-1 for oogenesis (Song et al. 2007; Klapholz et al. 2008). Similarly, female germline knockdown of the Caf1-180 or Caf1-55 genes prevent oogenesis and egg deposition (our
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own unpublished observations). Functional analysis of CAF1-dependent nucleosome assembly in early embryogenesis thus represents a technical challenge for the reasons mentioned above. In fact, recent studies on the complex have actually focused on additional functions of CAF-1 which can be studied later during development, such as heterochromatin-mediated gene silencing, Polycombdependent cell determination, DNA repair, and Notch signaling (Song et al. 2007; Klapholz et al. 2008; Anderson et al. 2011; Wen et al. 2012; Yu et al. 2013). As expected, however, nucleosome assembly and mitotic proliferation are affected, although not completely abolished, in p180 mutant larvae (Song et al. 2007; Klapholz et al. 2008). Although the distribution of CAF-1 subunits in the early embryos has not been reported yet, the complex likely follows the same dynamic nuclear localization as DNA replication factors. For instance, DNA polymerases and proliferating cell nuclear antigen (PCNA), enter S phase nuclei at each cycle and return in the cytoplasm during mitosis (Yamaguchi et al. 1991; Kuroda and Ueda 2005). In vertebrate cells, the CAF-1 complex is targeted to replication forks through the interaction of the large subunit with PCNA (Takami et al. 2007). Drosophila CAF-1 also interacts biochemically and genetically with the histone chaperone ASF1 (Tyler et al. 2001; Klapholz et al. 2008). Like PCNA, ASF1 enters replicating nuclei at each embryo nuclear division (Moshkin et al. 2002). It is not known whether PCNA precedes CAF-1 and ASF1 at DNA replication forks in these extremely rapid embryonic S phases. Structural and biochemical evidences support a role for ASF1 in providing H3.1/2-H4 dimers to the CAF-1 complex (Tagami et al. 2004; Tang et al. 2006). In addition, in human cells, ASF1 is involved in regulating the flux of old and newly synthesized histones at replication forks (Groth et al. 2007). However, the precise implication of ASF1 in RC nucleosome assembly in early Drosophila embryos still awaits a proper investigation. Replication-independent nucleosome assembly in the male pronucleus In contrast to RC nucleosome assembly, which takes place at replication forks and at damaged sites undergoing DNA synthesis, RI histone deposition occurs in various chromatin and developmental contexts. A unifying theme for these different processes is the use of the histone variant H3.3 as the unique source of RI histone H3, with the exception of CID at centromeres (see “Assembly of centromeric nucleosomes”). Although H3.3 in Drosophila was first characterized in the male germline using a specific antibody (Akhmanova et al. 1997), its implication in RI processes was established in cultured embryonic Drosophila Kc cells when H3.3 was shown to be incorporated in transcriptionally active chromatin independently of DNA synthesis (Ahmad and Henikoff 2002).
This enrichment of H3.3 in active chromatin occurs through a transcription-coupled (TC) mechanism where H3.3-H4 replaces histones evicted by the passage of the RNA polymerase complexes (Schwartz and Ahmad 2005). Although this aspect of H3.3 biology initially elicited considerable interest, a more global picture of the versatility of this histone variant has since emerged (Orsi et al. 2009; Elsaesser et al. 2010; Szenker et al. 2011; Filipescu et al. 2013). For instance, TC assembly does not contribute to any significant H3.3 deposition in the transcriptionally silent early Drosophila embryo. H3.3 deposition nevertheless massively occurs during RI assembly of paternal chromatin to replace sperm nuclear basic proteins (SNBPs) following fertilization. This unique RI assembly process was originally discovered after the functional characterization of the histone H3.3 chaperone HIRA (Loppin et al. 2005). Mutations affecting the Hira gene are maternal effect embryonic lethal and induce the development of non-viable gynogenetic haploid embryos (Loppin et al. 2000). In eggs laid by mutant females, the fertilizing sperm nucleus partially decondenses after the elimination of its SNBPs but fails to assemble its chromatin (Loppin et al. 2001; Bonnefoy et al. 2007). Surprisingly, paternal chromatin assembly is apparently the only essential function of the HIRA complex in Drosophila (Bonnefoy et al. 2007). As expected, this highly specialized role of HIRA is conserved in mammals (Inoue and Zhang 2014; Lin et al. 2014) and other vertebrates (Zhao et al. 2011), but the chaperone is also critically required at later stages of mouse embryo development (Roberts et al. 2002). The mammalian HIRA complex also comprises two other subunits, namely Ubinuclein1 and Cabin1 (Tagami et al. 2004). Cabin1 does not seem to have a clear ortholog in Drosophila whereas Ubinuclein1 is related to the single yemanuclein (yem) gene (Balaji et al. 2009). It was recently shown that YEM is, like HIRA, required for paternal chromatin assembly, and the phenotype of yem and Hira mutant embryos is indistinguishable (Orsi et al. 2013). The implication of YEM in paternal chromatin assembly provides an interesting clue about the possible targeting mechanism of the HIRA complex to the fertilizing sperm nucleus. YEM was indeed originally characterized as an oocyte-specific DNA binding protein (Aït-Ahmed et al. 1992), and more recent work indicates that it is necessary for the accumulation of HIRA in the male nucleus (Orsi et al. 2013). The death-associated protein DAXX is another histone chaperone involved in H3.3 RI deposition in human cells (Drané et al. 2010). DAXX cooperates with the alpha-thalassemia X-linked mental retardation protein (ATRX) for the deposition of H3.3-H4 at telomeres and pericentric regions (Wong et al. 2010; Goldberg et al. 2010; Drané et al. 2010; Lewis et al. 2010). Interestingly, in Drosophila somatic cells, HIRA and XNP, the Drosophila ATRX homolog, are redundant for H3.3 deposition at nucleosome-depleted chromatin gaps, suggesting that these factors can bind exposed DNA (Schneiderman et al. 2012).
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It is thus tempting to propose that the DNA binding property of YEM could be involved in the recognition of these histonefree regions by the HIRA complex. Interestingly, a study in human cells reported that all three subunits of the HIRA complex (HIRA, Cabin1, and UBN1) bind DNA in vitro, but not other tested chaperones or assembly factors (Ray-Gallet et al. 2011). This work suggests that the general affinity of the complex for DNA could be at play in the recognition of nucleosome gaps. Similarly, in the fertilized egg, the YEM-HIRA complex could bind exposed paternal DNA immediately after the rapid and genome-wide eviction of SNBPs. Unfortunately, this event likely occurs before egg deposition in Drosophila and is not practically accessible to direct observation. De novo assembly of nucleosomes is obviously critical but certainly not sufficient per se for the proper decondensation and higher order organization of paternal chromatin. Chromodomain helicase DNA binding protein 1 (CHD1) is a conserved ATP-dependent motor protein of the SNF2 protein family, which has both nucleosome assembly and remodeling properties in vitro (Lusser et al. 2005). Interestingly, chd1 mutant females are sterile and, like Hira or yem mutants, produce gynohaploid embryos (Konev et al. 2007). However, the clear presence of histones, including H3.3, in the male nucleus in chd1 mutant eggs (Orsi et al. 2009) indicates that this factor is not essential for paternal chromatin assembly. Instead, the distinctive, aberrant morphology of this nucleus suggests a different role for CHD1, such as, for instance, the regular spacing of newly assembled H3.3-containing nucleosomes at the genome-wide scale. Although the critical requirement of the HIRA complex in paternal chromatin assembly is well established, the actual role of H3.3 at fertilization has not been directly investigated. Deletion of both His3.3A and His3.3B genes led to the unexpected conclusion that this highly conserved histone variant is not absolutely essential for viability in Drosophila (Hödl and Basler 2009; Sakai et al. 2009). Double mutant animals are nevertheless male and female sterile. In the male germline, the absence of H3.3 compromises meiotic divisions and gamete production (Sakai et al. 2009). Similarly, oogenesis is defective in double mutant females, thus precluding the observation of male pronuclear formation (B.H and B.L. unpublished observations). Thus, the endogenous expression of His3.2 gene copies is not sufficient to cope with the absence of H3.3 in germ cells. More recently, however, Hödl and Basler reported the surprising observation that the sterility phenotype of double His3.3 mutants could be efficiently rescued by a transgene expressing H3.2 under the control of the His3.3B promoter (Hödl and Basler 2012). Thus, the H3.2 histone can functionally replace H3.3 in germ cells and gametes as long as appropriate transcriptional regulation drives the expression of the canonical H3. RI paternal chromatin assembly (which is required for embryo viability) can thus surprisingly occur with H3.2 as the unique source of core H3. Our preliminary
observations indicate that the dynamics of male pronuclear decondensation is nevertheless affected in this rescued context (B.H and B.L, unpublished observations). Although it remains to be properly investigated, the functional replacement of H3.3 with H3.2 in the zygote suggests that HIRA can adapt to a different H3 subtype to perform its most crucial developmental task. In addition, H3.3 can fully compensate for the loss of H3.2 when expressed in S phase, suggesting that the CAF-1 complex can also accommodate a different H3 for RC assembly (Hödl and Basler 2012). In wild-type eggs, however, the HIRA complex strictly uses H3.3 to assemble paternal nucleosomes (Loppin et al. 2005). Interestingly, this stringent use of the H3 replacement variant in this context does not apply to H2A and H2A.Z. Although the presence of γH2A.Z foci in the decondensing male nucleus indicates that H2A.Z is deposited along with H3.3 during de novo assembly of paternal nucleosomes (Delabaere et al. 2014), we actually observed the incorporation of both types of H2A during this RI assembly process (Fig. 2). In the mouse zygote, limited deposition of canonical H2A is observed in both pronuclei although H2A.X is the most abundant H2A used at this stage (Nashun et al. 2010). These observations confirm the general idea that the canonical and replacement H2A histones do not strictly follow RC and RI deposition pathways, at least in the developing embryo. In contrast, H3.2 does not seem to be used for RI assembly as long as H3.3 is normally available. RI nucleosome assembly factors can nevertheless adapt to the lack of H3.3 by utilizing H3.2, as also reported in somatic cells (Sakai et al. 2009). Assembly of centromeric nucleosomes The centromeric histone CID (Henikoff et al. 2000) is the epigenetic determinant of centromere identity in fly (Heun et al. 2006) and is required in mitosis and meiosis for the assembly of kinetochores (Blower and Karpen 2001; Blower et al. 2006; Pauleau and Erhardt 2011). Although it is well established that CID is both necessary and sufficient for centromere formation (Mendiburo et al. 2011), the transmission of paternal chromosomes at fertilization yet provides an additional and natural illustration of the absolute requirement of CID for the maintenance of centromere identity through generations. Indeed, CID is so far the only histone known to survive the massive replacement of nucleosomes with SNBPs at the canoe stage of spermiogenesis (Loppin et al. 2001; Dunleavy et al. 2012; Raychaudhuri et al. 2012). In mature sperm, CID is present at four discrete nuclear foci that correspond to the centromeres of these haploid gametes and paternal CID is indeed transmitted at fertilization. Remarkably, elimination of CID protein in post-meiotic male germ cells induces, through a paternal effect, the loss of paternal chromosomes at the first zygotic mitosis (Raychaudhuri et al. 2012). Interestingly, the inner kinetochore component
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Fig. 3 CID and the transgenerational maintenance of centromere identity. Schematic representation of spermiogenesis (blue) and zygote formation (green). In the wild-type situation (left), CID (green spots) is retained at centromeres in mature sperm after the histone-to-protamine replacement. After fertilization, paternal CID is required for the formation of functional kinetochores and for paternal chromosome segregation in anaphase of the first zygotic cycle. When CID is experimentally degraded in post-meiotic male germ cells (right), the epigenetic identity of paternal centromeres is lost and paternal chromosomes fail to segregate at the first mitosis (see Raychaudhuri et al. 2012)
Fig. 2 Histone deposition in the male pronucleus and in the zygote. Confocal images showing fertilized eggs and the first zygotic metaphase stained with an anti-FLAG antibody to detect the indicated tagged histone (green) and for DNA (red). The transgenes used were promHis3.3A-H3.2::Flag (H3.2::Flag), promHis3.3A-H3.3::Flag (H3.3::Flag), promHis2A.Z-H2A::Flag (H2A::Flag), and promHis2A.ZH2A.Z::Flag (H2A.Z::Flag). A zoom of the anti-Flag staining in the decondensing male pronucleus is shown (insets). While RI assembly of the male pronucleus strictly uses H3.3 and not the canonical histone H3.2, H2A and H2A.Z are both incorporated in the male pronucleus. After the first S phase, canonical H2A and H3.2 decorate both parental sets of chromosomes, whereas the H2A.Z and H3.3 variants are enriched in paternal chromosomes. Following the first zygotic division, both H2A.Z and H3.3 variants disappear after a few cycles
CENP-C, another key protein for centromere function, is not present at sperm centromeres. This study thus demonstrates that the maintenance of CID at sperm centromeres is critical for the epigenetic propagation of paternal centromere identity in the zygote, as it is also likely the case in vertebrates, where the centromeric histone CENP-A is retained in mature sperm in various species (Palmer et al. 1990; Milks et al. 2009) (Fig. 3). Of note, it is however currently unknown if CID is retained in sperm as nucleosome particles or in another configuration. After fertilization, paternally transmitted CID is retained on paternal centromeres throughout the first zygotic cycle and then rapidly disappears after the first mitosis. Surprisingly, maternal CID is detected on paternal chromosomes during the decondensation of the sperm nucleus before the first zygotic S phase (Raychaudhuri et al. 2012). This contrasts with the fact that, during the subsequent nuclear cycles, CID is loaded on chromosomes in anaphase (Schuh et al. 2007). Studies in cultured cells have however revealed that the loading of CID is dependent on cell type and is more complex and dynamic than previously anticipated (Mellone et al. 2011;
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Lidsky et al. 2013). What actually determines the deposition of CID at centromeres during mitosis remains an open question. The identification of the factor responsible for CID loading on centromeres is nevertheless a major achievement of the recent years. Chromosome alignment defect 1 (CAL1), which is found only in Drosophilidae, is functionally related to yeast SMC3 and human HJURP (Erhardt et al. 2008; Dunleavy et al. 2009; Müller and Almouzni 2014). CAL1 plays an essential and limiting role for the assembly of CID-containing nucleosomes and the deposition of CENP-C at centromeres (Schittenhelm et al. 2010; Mellone et al. 2011; Chen et al. 2014). Interestingly, the observation that CAL1 associates with centromeres in prophase before CID deposition suggests that CID is targeted to centromeres through CAL1 binding. In support of this model, it has been recently demonstrated that artificial tethering of CAL1 to chromatin is sufficient to trigger the formation of an ectopic functional centromere (Chen et al. 2014). Finally, proper centromere targeting of CID is also facilitated by CAL1-mediated mono-ubiquitylation of centromeric CID and proteasome-mediated degradation of noncentromeric CID (Moreno-Moreno et al. 2006; Bade et al. 2014), the latter process being specifically regulated by the F-box protein partner of paired (Moreno-Moreno et al. 2011). In addition, the conserved histone-fold protein CHRA C14, a subunit of the ATP-dependent chromatin remodeling complex CHRAC, prevents ectopic incorporation of Cid after DNA damage (Mathew et al. 2014). Chrac14 mutant flies are indeed hypersensitive to DNA damage, and mutant embryos displayed increased levels of CID in chromatin. The elevated incidence of mitotic defects in these embryos suggests that these ectopic centromeres perturb mitotic divisions and cause aneuploidy. The authors propose that, in the absence of CHRAC14 regulation, the ability of CID to be deposited in a RI manner could favor its natural incorporation at DNA lesions for the transient repair of damaged chromatin.
Conclusion This overview illustrates the interest of the early fly embryo for studying histone dynamics in a highly integrative manner. The building up of thousands of nuclei from the gigantic egg cell includes many unique developmental constraints, such as large-scale molecular trafficking, complex histone biosynthesis and storage regulation, ultrafast chromatin assembly and absence of cellular partition. Still, the formation of the male nucleus and the streamlined cleavage nuclear cycles represent archetypal RI and RC chromatin assembly processes, respectively, from which much can be learned. The availability of efficient techniques to specifically knockdown genes in the female germline and the mass production of gene knockouts
using the Crispr/Cas9 targeting system open promising horizons for functional genetics in the early embryo (Ni et al. 2011; Kondo 2014). Our understanding of histone biology and nucleosome assembly in a developing animal will certainly benefit from these methodological breakthroughs. Acknowledgments We thank Raphaëlle Dubruille for her critical reading of the manuscript. This work was supported by a grant from the Agence Nationale de la Recherche (ZygoPat - ANR-12-BSV6-0014).
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