To monitor histone dynamics in vivo, fusion genes encoding various histones and the green fluorescent protein (GFP) were constructed under the control of heat shock-inducible promoters. These constructs were transfected into exponentially growing Kc cells and induce. The deposition of histone H3-GFP in the nucleus parallels that of nucleotide analog incorporation into DNA. Localization of histone H3-GFP is completely blocked by pretreatment of cells with the DNA replication inhibitor aphidicolin, demonstrating that the deposition of histone H3 is strictly replication dependent. Detection of a component of the DNA replication machinery, PCNA, also confirms that deposition of histone H3-GFP is coupled to DNA replication: PCNA, BrdU, and H3-GFP give similar labeling patterns both in early S phase (when euchromatic DNA is replicating) and in late S phase (when heterochromatic DNA is replicating). BrdU and H3-GFP closely overlap because both are present for the entire 2 hr labeling period. PCNA labeling does not precisely overlap, since it provides a 'snapshot' of replication only at the time of fixation. In subsequent labeling experiments, PCNA as used to indicate the cell cycle stage (Ahmad, 2002).
Since histone H3 deposition is strictly replication dependent, it was reasoned that replication-independent deposition of histone H4 might be accompanied by the deposition of H3 variants to form variant nucleosomes. Centromeric histones are thought to be included in nucleosomes at centromeres, and it has been demonstrated that the Drosophila centromeric H3 variant Cid localizes to centromeres by a replication-independent pathway. Thus, it was expected that some sites showing H4 replication-independent deposition would be centromeres. Detection of centromeres in H4GFP-transfected cells demonstrates that four to six of the H4 replication-independent foci were indeed centromeres, consistent with the assembly of nucleosomes containing Cid and H4 at these sites. It was reasoned that the remaining H4 sites must be incorporating the final histone H3 variant, H3.3. Indeed, expression of H3.3-GFP in cells demonstrated that this variant does undergo both replication-coupled and replication-independent deposition. None of the H3.3-GFP foci coincided with centromeres, showing that centromeres use the Cid histone exclusively (Ahmad, 2002).
It was confirmed the H3.3-GFP is tightly bound to chromatin by extracting cells with 1.5 M salt before fixation. After this treatment, nuclei retain 48% of the H3.3-GFP but only 22% of the H2B-GFP. Such differential extraction is expected from the biochemical properties of these histones, and the proper behavior of GFP-tagged histones has been extensively documented (Ahmad, 2002).
To map the locations of the sites in the nucleus where replication-independent deposition of histone H3.3 and H4 occurs, mitotic figures were examined from cells transfected with histone-GFP constructs. The G2 phase in Kc cells is 4-6 hr long; thus, mitotic figures with H3-GFP labeling first appear 4-6 hr after heat-shock induction and show patterns consistent with histone-GFP production in late S phase, when heterochromatin is replicating. In contrast, labeled mitotic figures with H3.3-GFP and H4-GFP appear within 2 hr of induction. H4-GFP showed prominent labeling at a single extended site near the middle of an X chromosome. The pattern of H3.3-GFP is very similar to that of H4-GFP, showing the greatest labeling over an extended site on the X chromosome and at low levels specifically in euchromatin. These cells must have been in the G2 phase of the cell cycle when histone-GFP was produced. This was confirmed by the presence of H3.3 labeling on mitotic chromosomes that showed no incorporation of pulse-labeled nucleotides and by observing mitotic figures from aphidicolin-treated cultures that nevertheless displayed H3.3-GFP labeling. Thus, these mitotic labeling patterns with H3.3-GFP and H4-GFP must have resulted from replication-independent deposition (Ahmad, 2002).
The extended appearance and proximal location of the prominent H3.3 and H4 site on the labeled X chromosome suggested that it coincides with the large rDNA gene repeat array on this chromosome. In situ hybridization with probes to the 28S rDNA gene confirmed that this is so. Quantitative measurements of GFP signal over the rDNA array and over all of the chromosomes indicate that ~40% of all histone H3.3 in the cell is deposited at the rDNA locus. In Tetrahymena, a histone H3 replacement variant is enriched in the transcriptionally active macronucleus, suggesting that this Tetrahymena variant potentiates active chromatin (Allis, 1984). It is presumed that the high intensity of histone H3.3-GFP staining at the rDNA locus in Drosophila is due to the combination of its densely repeated genes with high transcriptional activity (Ahmad, 2002).
Notably, labeling with H3.3-GFP and H4-GFP was often observed of only one X chromosome. This is not due to absence of rDNA from other X chromosomes in these cells because the detection of 28S rDNA by in situ hybridization confirmed that rDNA arrays are present on each of the three X chromosomes. Other studies have pointed out that many Drosophila cell lines (including Kc) carry two distinguishable kinds of X chromosomes: a short one (XS) that resembles the normal X of flies, and a longer X (XL). The origin of XL has been attributed to an expansion of the rDNA locus on this chromosome, presumably as these cells adapted to culture conditions. It was observed that the rDNA array on XL is always labeled by H3.3-GFP, consistent with this locus being active in all cells. However, in some experiments, variable numbers of cells had additional labeling on XS chromosomes. To test whether some of this variability between experiments was due to differences in growth conditions, cells were transfected with the histone H3.3-GFP construct and then expression was induced in samples of this culture 16 or 24 hr later. It was found that many cells from exponentially growing cultures show replication-independent labeling on both XL and XS chromosomes, while metaphase spreads from the later time point, when culture growth had slowed, showed labeling on only the one XL. This change in frequency suggests that the smaller rDNA arrays on XS chromosomes are maintained in a transcriptionally silent state but can be activated (Ahmad, 2002).
The silencing of XS rDNA arrays might be due to heterochromatin-mediated silencing. Indeed, staining of metaphase spreads from cells expressing histone H3.3-GFP for the heterochromatin marker H3di-MethylK9 (H3Me) revealed that rDNA arrays labeled by replication-independent deposition of H3.3-GFP are depleted for H3Me, in spite of being flanked on both sides by heterochromatin. In every XS chromosome where the proximal region was labeled with H3.3-GFP, a corresponding gap in the H3Me pattern was found. That sites heavily labeled with H3.3-GFP are largely unlabeled with H3Me was confirmed in interphase nuclei. It is concluded that the chromatin state of rDNA arrays can be reversed in response to changes in growth conditions, and H3.3 accumulates de novo at activated genes (Ahmad, 2002).
Alternate interpretations of the phylogenetic history of the histone H3 family have been proposed. One analysis suggested that a replacement histone H3 variant was the common ancestor, but other interpretations have proposed that replacement histones have multiple independent origins. The presence of paralogous histone H3 genes in many organisms may preclude delineation of which sequence is ancestral. However, the findings of this study suggest that a replication-independent nucleosome assembly pathway is essential in all cells. This implies that, functionally, a replacement histone H3 has always been extant. In organisms that encode only one kind of canonical histone H3 protein that is used throughout chromatin, it is expected that this H3 variant must undergo both replication-coupled and replication-independent deposition. Fungal lineages are particularly intriguing in this regard because all ascomycetes, including laboratory yeasts and molds, carry only one canonical histone H3. Each of these is identical to animal H3.3 at positions 89 and 90, and often identical at position 31. Thus, by this criterion, it is proposed that the solitary histone H3 proteins in ascomycetes are equivalent to histone H3.3. Indeed, nucleosome assembly activity in the cell cycle gap phases has been detected in Saccharomyces. These fungi appear to have lost their ancestral H3, since genomes from the Basidiomycota sister clade have both H3 and H3.3. Histone H2A in Saccharomyces may have an analogous evolutionary history, since it now performs the functions of the H2A and the H2A.X variants in other organisms. Thus, both histone H3 and H2A in Saccharomyces appear to be evolutionary derivatives of replacement genes (Ahmad, 2002).
The lack of an H3 counterpart in yeasts and molds may provide insight into differences between simple fungi and complex multicellular eukaryotes in maintaining silent chromatin. Much of the Saccharomyces genome is continually in a transcriptionally competent state similar to H3.3-containing regions in complex genomes. Perhaps this relative lack of silent chromatin allowed the loss of the strictly replication-coupled histone substrate. Heterochromatic silencing in yeast may be needed only at special sites, such as silent mating type loci and telomeres, where SIR-based silencing has evolved. In multicellular eukaryotes, the need for maintaining most of the genome in a continuously silent state in differentiated cells may favor maintaining two distinct H3 histones (Ahmad, 2002).
Chromatin states can be distinguished by differential covalent modifications of histones or by utilization of histone variants. Chromatin associated with transcriptionally active loci becomes enriched for histones with particular lysine modifications and accumulates the H3.3 histone variant, the substrate for replication-independent nucleosome assembly. However, studies of modifications at particular loci have not distinguished between histone variants, so the relationship among modifications, histone variants, and nucleosome assembly pathways is unclear. To address this uncertainty, the relative abundance of H3 and H3.3 and their lysine modifications has been quantified. Using a Drosophila cell line system in which H3.3 has been shown to specifically package active loci, it was found that H3.3 accounts for 25% of total histone 3 in bulk chromatin, enough to package essentially all actively transcribed genes. MALDI-TOF mass spectrometry (MS) and antibody characterization of separated histone 3 fractions revealed that H3.3 is relatively enriched in modifications associated with transcriptional activity and deficient in dimethyl lysine-9, which is abundant in heterochromatin. To explain enrichment on alternative variants, it is proposed that histone modifications are tied to the alternative nucleosome assembly pathways that use H3 primarily at replication forks and H3.3 at actively transcribed genes in a replication-independent manner (McKittrick, 2004).
Until recently, H3.3 was thought to be interchangeable with H3, because there are only four amino acid differences between them. However, the demonstration that H3.3 is found at active loci raises the possibility that all active chromatin is packaged into H3.3-containing nucleosomes. To test whether H3.3 is sufficiently abundant, a separation method was used that would resolve the two variants. HPLC on a reverse-phase column has previously been shown to separate histone 3 variants in plants, despite the abundant tail modifications and the close similarity between the variant sequences. When acid-extracted histones from Drosophila Kc nuclei were resolved by reverse-phase HPLC, all H3 histones were found in two partially overlapping peaks well separated from other acid-extractable material. Peak fractions were pulled and subjected to complete proteolysis with Arg-C protease and MALDI-TOF mass spectrometry to identify the two HPLC peaks. It was possible to identify eight of the 10 expected peptides >500 Da and their known modified forms based on their predicted masses. Examination of peptide 27-40 revealed a 16-Da difference between corresponding mass peaks, identifying the earlier and larger HPLC peak as H3, which has an alanine at position 31, and the later and smaller peak as H3.3, which has a serine at position 31 (McKittrick, 2004).
To ascertain the relative abundance of H3 and H3.3, peak areas were integrated, and it was found that H3.3 accounts for about one-fourth of the histone 3 in Kc cells. The abundance of H3.3 in Kc cells can be compared to the percentage of the genome that is estimated to be transcribed in the same cells. Microarray analysis has revealed that 68% of a large sample of genes are transcribed in Kc cells, which have a tetraploid female karyotype. The female Drosophila genome is estimated to encompass 176 Mb, of which 59 Mb is heterochromatin and 49 Mb is annotated as intergenic. This yields an overall estimate of 25% of the genome transcribed in Kc cells, remarkably close to the relative abundance of H3.3 (McKittrick, 2004).
H3.3 is known to increase in abundance in nondividing cells in vertebrates (Urban, 1983; Pina, 1987), presumably because H3 is not deposited outside of S phase, whereas H3.3 is deposited constitutively. It was asked whether replication-independent assembly of H3.3 in Kc cells also leads to accumulation of H3.3 when cells are induced to exit the cell cycle and differentiate. Kc cells are known to cease dividing and differentiate in response to the moulting hormone, ecdysone, and ecdysone was added and cells were harvested at various time intervals up to 16 days. Indeed, a gradual increase from 24% to >50% H3.3 was seen, whereas untreated cells remained close to 25% H3.3 as they reached stationary phase over a 6-day period. Thus, the close correspondence between H3.3 abundance and active transcription does not apply to differentiating cells, where accumulation of H3.3 likely represents replacement of nucleosomes that are lost. The increase of endogenous H3.3 levels in differentiating Kc cells confirms results (Ahmed, 2002) showing replication-independent deposition of tagged H3.3 introduced into this cell line (McKittrick, 2004).
The deposition of H3.3 at active loci in Kc cells (Ahmed, 2002), taken together with the demonstration that H3.3 is sufficiently abundant to account for all active transcription, is consistent with the notion that H3.3-containing nucleosomes comprise all active chromatin in dividing cells. If this were the case, covalent histone 3 modifications that have been correlated with active transcription to be enriched on H3.3 relative to H3 would be expected. The separation of these two forms allowed for identification and quantification of their modifications (McKittrick, 2004). Modifications on the variants were quantified using LC-ESI MS on ArgC-digested peptides. The area of each LC-ESI MS peak was measured and expressed as the ratio of that modified form to the total areas of all of the modified forms of that peptide. Peptide 73-83 in both H3 and H3.3 can be methylated at lysine 79, which has been associated with active chromatin regions. Consistent with the expectation that active modifications would be enriched on H3.3, it was found that dimethylated H3.3K79 is enriched 2-fold on H3.3 relative to H3. H3.3 was also enriched for monomethyl K79, whereas trimethylation of this residue, known to occur on active chromatin in budding yeast, was not observed in Kc cells on either histone (McKittrick, 2004).
Active chromatin is also correlated with the hyperacetylation of histone tails. Two acetylation sites are known to be present on peptide 18-26, at lysines 18 and 23. LC-ESI MS shows that H3.3 has more total acetylation, with more diacetyl (both 18 and 23) and less mono-acetyl (either 18 or 23, but not both) than H3. Therefore, modifications to lysine residues in both the core (K79) and tail (K18 and 23) that correlate with active chromatin are enriched on H3.3 (McKittrick, 2004).
Other peptides had too many possible modifications on lysines to readily ascertain relative abundances. Nevertheless, the spectrum of modifications was determined using tandem MS, in which individual ions are collected, fragmented, and reanalyzed, to assign modifications to specific lysines. In this way, it was found that peptide 27-40 displays complex methylation patterns for both H3 and H3.3, with different peptides containing 0, 1, 2, or 3 methyl groups on K27, combined with 0, 1, or 2 methyl groups on K36 and K37. These complex patterns precluded assignment of specific modifications to particular peptide masses, where LC-ESI MS spectra showed 0-6 methyl groups overall. Likewise, numerous and abundant modifications were identified on peptide 9-17, including 0-3 methyl groups and/or single acetyl groups on K9 and K14, with only low levels of unmodified peptide 9-17 (McKittrick, 2004).
Detection of abundant lysine modifications on H3 and H3.3 tails contrasts with an inability to detect any phosphorylated serines or threonines. This failure to detect phosphates does not appear to have resulted from their removal during acid extraction, because salt-extracted and SDS gel-purified histone 3 that is not exposed to acidic pH yields MALDI-TOF spectra that are indistinguishable from those obtained by acid extraction followed by HPLC and/or SDS gel purification. It was also found that phosphatase treatment before a second round of HPLC does not alter the mass spectra. Furthermore, it is unlikely that changes occur during nuclear isolation, because histones extracted from whole cells produced H3 and H3.3 spectra that are indistinguishable from those obtained from nuclei. Therefore, it appears that phosphates on H3 and H3.3 are not sufficiently abundant in Kc cells to be detected in bulk chromatin, and that the isolation procedures had no effect on modification patterns. The inability to detect phosphates by MS might simply be that phosphates on histones, such as H3S10 phosphate, are restricted to regulatory regions at interphase, and to mitotic chromosomes, so would be undetectable in bulk histone 3 by the method used. Phosphates on histone 3 have been seen in bulk chromatin only after in vivo inhibition of phosphatases by okadaic acid or after colchicine treatment, which arrests Kc cells after mitosis (McKittrick, 2004).
The limitations of peptide mass analysis that were encountered in trying to quantify certain modifications, including complexity and low abundance, led to use of modification-specific antibodies to further examine correlations between histone 3 subtypes and covalent modifications. Indeed, the links between particular modifications and active vs. inactive chromatin that motivated this study have been almost entirely based on such antibodies (McKittrick, 2004).
Slot and Western blot analyses were used to look for modification differences between H3 and H3.3. HPLC-purified H3 and H3.3 were applied to a slot blot and probed with antiacetyl-K9, antiacetyl-K14, and antidimethyl-K79. All three of these modifications are enriched in active chromatin, and it is clear that all three are enriched on H3.3. Dimethyl-K9 is known to be enriched in silent chromatin, and it is clearly enriched on H3. Di- and trimethyl-K4 are known to be enriched in active chromatin, and they are clearly enriched on H3.3. Such enrichments were also seen by LC-ESI MS (McKittrick, 2004).
ELISA was used to more effectively quantify the enrichment or depletion of specific modifications on H3.3 relative to H3. H3.3 was found to be enriched in both di- and trimethyl-K4 5-fold and in dimethyl-K79, acetyl-K9, and acetyl-K14 2-fold. H3.3 is depleted in dimethyl-K9 2-fold. These enrichments are unlikely to be caused by differential binding to the variants, because none of the sites assayed are close to sequence differences between H3 and H3.3. Therefore, these well established 'silent' and 'active' modifications correlate with H3 and H3.3, respectively, consistent with the hypothesis (Ahmed, 2002) that active chromatin consists of H3.3-containing nucleosomes (McKittrick, 2004).
Therefore active and silent chromatin can be distinguished both by differential histone modification and by the incorporation of alternative histone 3 variants. The lack of antibodies able to distinguish H3.3 from H3 has left open the question of whether modifications are specific for these alternative variants. This study shows that modifications are indeed enriched on H3.3 or H3 in accordance with previously demonstrated associations with active or silent chromatin. Specifically, it was found that 'active' modifications, including methylated K4 and K79 and acetylated K9, K14, and K18 + K23, are enriched on H3.3, and a 'silent' modification, methylated K9, is enriched on H3. Similarly, Waterborg (1990) showed enrichment of 'active' modifications in alfalfa replacement histone 3, even before such modifications were found to correlate with activity of specific loci. Therefore, these results using a Drosophila system appear to be general for plants and animals (McKittrick, 2004).
In no case were modifications found to be exclusive to H3.3 or H3. All sites of known modification are shared between the two variants. Furthermore, none of the modified residues for which differential enrichment was seen was close to any of the four sequence differences. Therefore, binding-site preferences of modification enzymes are unlikely to account for these differences. Rather, alternative nucleosome assembly pathways may be responsible, whereby modifying enzymes would be primarily associated with alternative assembly machines. These machines are separated in both space and time, with replication-coupled assembly depositing primarily H3 and replication-independent assembly depositing exclusively H3.3 (Ahmed, 2002). If modification enzymes are preferentially associated with alternative assembly machines, they could efficiently propagate the modification status of chromatin. "Silent" modifications would occur as the replication fork passes through, whereas "active" modifications would occur as H3.3-containing nucleosomes are assembled at transcriptionally active loci throughout the cell cycle (McKittrick, 2004).
The active modification of histone 3 and the replication-independent deposition of H3.3 may be independent events. For example, the association of both processes with transcription could explain their correlation. However, the possibility that modifications are primarily associated with alternative assembly pathways not only can account for this correlation but also can reconcile discrepancies in the burgeoning histone modification literature. A case in point is a finding (Labrador, 2003) that challenges the assumption that a modification classified as active by chromatin immunoprecipitation is relevant to gene activity. That study shows that H3K14 acetylation is enriched in polytene bands in Drosophila, rather than in interbands, which are sites of active transcription. It is noted that the polytene banding pattern reflects the chromatin density, so any marker that is uniformly distributed in chromatin will be enriched in bands. The enrichments seen for active modifications on H3.3 ranged from 2- to 5-fold, increases not likely to be sufficient to show an interband pattern even if H3.3 packages all active chromatin. Thus, the polytene banding pattern seen for H3K14 acetyl is consistent with its 2-fold enrichment on H3.3. Likewise, the widespread euchromatic localization of H3K4 dimethylation on polytene chromosomes, rather than a band-interband pattern, is consistent with its 5-fold enrichment on H3.3. Therefore, enrichment of active modifications on H3.3 can explain both the enrichment of these modifications on active chromatin detected by chromatin immunoprecipitation and the localization patterns seen on polytene chromosomes (McKittrick, 2004).
The failure of stable modification patterns to correlate with transcription
and silencing genome-wide makes them unlikely candidates for the propagation of
epigenetic information. Rather, the nucleosome assembly process provides a
straightforward mechanism for inheritance of an active chromatin state through
multiple rounds of cell division. H3.3 deposits at active regions (Ahmed,
2002), and it has been shown that H3.3 abundance corresponds closely to the
extent of transcription; therefore, modifications that generally follow H3.3
deposition patterns would be classified as active. This active chromatin would
be diluted 2-fold by new H3-containing nucleosomes during DNA replication, but
the remaining H3.3 might be enough to maintain the active state. Reinitiation of
transcription would lead to replication-independent replacement, restoring
H3.3-containing nucleosomes with their active modifications over entire
transcription units (McKittrick, 2004).
The enrichment of H3.3 in gene-rich interbands suggests that RI deposition occurs during transcription. Previous experiments with rDNA or artificial gene arrays could not distinguish whether RI assembly occurs during or after transcription of individual genes (Ahmad, 2002; Janicki 2004). To determine when RI assembly occurs, changes were sought in histone composition during controlled induction of HSP loci in polytene chromosomes. The HSP70 loci at cytological bands 87A and 87C are compacted before induction, but expand as large puffs during induction and gene expression. The summed intensities of histone-GFP signals were measured to compare puffs of different sizes. Strikingly, it was found that the fully expanded active puffs show little H3-GFP above background. Instead, the puffs are dramatically enriched for the H3.3-GFP histone. Other heat-shock-inducible loci also show enrichment of H3.3 during induction, indicating that H3.3 enrichment is a common feature of these highly induced genes. It is concluded that induced loci are predominantly packaged with H3.3 while they are transcribed. Because the H3 subtype is largely absent from puffs, 'H3' modifications that are enriched in polytene puffs must actually be modifications of the H3.3 subtype. Indeed, immunodetection of H3S10-phospho and H3K14-acetyl epitopes show that these modifications coat puffs that contain no detectable H3 signal (Schwartz, 2005).
Quantitative comparison of puffs during induction of the HSP70 genes suggests that changes in histone composition begin in the first few minutes of induction. The amounts of histone-GFP at the HSP70 loci could not be quantitated before induction because they are too small to resolve from flanking bands, but intermediate puffs (25 min into the induction) show moderate amounts of H3.3, while maximal puffs (20 min) display more H3.3. The rapid gain in H3.3 signals implies that RI deposition is occurring during transcription. Enrichment at the HSP70 genes occurs substantially faster than was observed at multicopy gene arrays in interphase nuclei (Janicki 2004), and may be due to the greater sensitivity of polytene chromosomes. Alternatively, fast gain may be due to the higher rates of transcription at induced HSP70 genes. Such a relationship could explain why the intensities of H3.3-GFP signal vary between different HSP loci (Schwartz, 2005).
It is expected that since induced loci acquire H3.3 as a puff expands, H3-GFP signals should reciprocally decrease. However, the small size of the HSP70 genes (~2 kb) hampered measuring how much tagged histone was present before induction. Instead, the much larger ecdysone-responsive (EcR) genes E74 and E75, which each encompasses 80100 kb across characteristic sets of DAPI-stained bands, were examined. Measurable amounts of both H3 and H3.3 are present in these intervals before induction. The E74 and E75 genes also generate chromosomal puffs when they are induced. Just as was observed for heat-shock loci, the transcribing EcR puffs are heavily labeled with H3.3. At these larger genes, the H3-subtype histones show reciprocal changes after induction: H3 amounts decrease in the puff, while H3.3 amounts increase. These experiments support observations with the heat-shock-inducible genes, and confirm that induced genes become depleted for H3 and enriched for H3.3 during transcription (Schwartz, 2005).
The observations of loss of H3 and gain of H3.3 at expanding puffs are not due to epitope differences or accessibility as chromatin decompacts, because the same epitope (GFP) was used and opposite results in puffs were seen with each histone. However, to demonstrate that quantitation of chromosomal puffs correctly reported changes in histone composition, a heat-shock-inducible construct was used to produce a pulse of new tagged H3.3 at the same time as the endogenous HSP70 loci was induced. It was predicted that producing new H3.3 and inducing the HSP70 genes would result in deposition only at those loci. Indeed, it was found that the newly produced H3.3-GFP localizes exclusively to the HSP loci, demonstrating that new nucleosomes are being assembled from new histones at transcribed sites. A similar pulse of H3 protein does not incorporate into chromatin at this time, confirming that the new nucleosomes only use H3.3. It is concluded that RI histone deposition is initiated by gene induction and occurs as the gene is being transcribed. The consequence of replacement is that activated loci lose H3 and become enriched for H3.3 (Schwartz, 2005).
The overall behavior of H3.3 differs from that of many markers of active
transcription. While RNA polymerase II components are enriched at HSP
loci and depleted from most other loci during the heat-shock response, H3.3-GFP,
which was deposited earlier in development remains
distributed throughout chromosomes. As a
heat-shock response terminates, active RNA polymerase II rapidly leaves these
loci and transcription of the HSP70 genes shuts off. In contrast, tagged
H3.3 remains stably incorporated at the HSP70 genes hours after a
transient induction. Signals at HSP70 genes
are undiminished even 48 h after the induction. It is concluded
that H3.3 remains incorporated even after transcription has ceased. In
differentiated cells where the cessation of DNA replication precludes expression
of the H3 histone and replication-coupled deposition, H3.3 enrichment will be
permanent (Schwartz, 2005).
In budding yeast, specific nucleosomes within the promoters of some genes are dissembled during transcriptional induction, and then reassembled when repressed (Adkins 2004). In contrast, increases in H3.3 signals at induced genes are observed in Drosophila during active transcription, suggesting that at least some H3.3 deposition is distinct from promoter nucleosome changes. Indeed, H3.3-GFP signals are distributed throughout induced gene puffs, supporting the argument that new nucleosome assembly is not limited to promoters. This is especially clear at the E74 and E75 puffs, which encompass 80 and 100 kb of transcribed DNA, respectively. At the E75 gene, probes to the 5'- and 3'-ends of the transcription unit label the ends of the puff, and the bulk of the H3.3 signal lies between these probes, confirming that H3.3 has been enriched in transcribed chromatin (Schwartz, 2005).
Artificial promoter arrays were used to test if H3.3 enrichment at the
HSP70 genes could be accounted for by replacement of promoter
nucleosomes. These arrays contain ~40
tandem copies of HSP70 promoter elements, but do not support RNA
polymerase firing. Accordingly, the arrays bind
extremely large amounts of transcriptional activator, but show little active RNA
polymerase II (compared to that at the endogenous HSP70 genes in the same
spreads). While the
endogenous genes are heavily labeled after a pulse of H3.3-GFP,
promoter arrays show little or no labeling above background. It is concluded
that transcription factor binding and
promoter activation are insufficient to stimulate H3.3 deposition. These results
suggest that transcriptional elongation is required, and observation of H3.3
throughout the very long transcription units of EcR genes is consistent with
this idea (Schwartz, 2005).
If transcription triggers replication independent nucleosome assembly by first displacing histones from the template, it was of interest to see what happens to these released histones. Recent experiments have characterized an activity that performs a histone H2A/H2B dimer exchange reaction between nucleosomes and histone chaperones. In vivo, H2A/H2B dimers release from chromatin and rebind to other sites, indicating that they can exchange and be reused. However, these experiments detected little or no exchange of the H3 or H4 histones. The experiments here show that replication independent assembly can use newly produced H3.3, suggesting that old, displaced H3 subtype histones might be destroyed. Indeed, when histone-GFP signals in whole-mount polytene nuclei were compared from larvae induced 0.5 to 3 d earlier, it was observed that H3.3 signals decrease approximately four times faster than H3 signals. However, in this time period polytene cells continue to duplicate their chromatin by replication-coupled nucleosome assembly, which complicates measures of histone stability. The possibility that H3.3 is unstable was directly tested by measuring chromatin-bound histones after a pulse of production. To measure only RI-deposited histones, the pulse was produced in adult flies. The vast majority of cells in the adult are post-mitotic, and these cells should not synthesize H3 or perform replication-coupled nucleosome assembly. The main exception is the germline lineages, and therefore the abdomens of adult flies were discarded before analysis (Schwartz, 2005).
Chromatin-bound histones were distinguished from soluble pools by their resistance
to extraction with 400 mM salt. Induction of
histone-GFP constructs produces soluble pools of protein that are available for
deposition for several hours. As expected for
differentiated cells, a pulse of H3-GFP in adults was not incorporated.
In contrast, a pulse of H3.3-GFP rapidly entered
chromatin: this must have been due to RI assembly. While
the amounts of chromatin-bound H3.3 greatly increased within 8 h after
production, this enrichment was transient, and after 4 d the amounts of
chromatin-bound H3.3 have returned to a low level. It is estimated that the bulk
chromatin-bound H3.3 had a half-life of ~24 h. This is in contrast to the stability of chromatin-bound H3,
where a half-life of hundreds of days has been estimated.
The decay of H3.3-GFP is not due to a defect in the fusion protein,
because an identical pulse of H2B-GFP becomes chromatin-bound in adults but does
not significantly decrease over the chase period,
and H3-GFP incorporated during development is also stable.
Instead, a short half-life appears to be a general feature of replacement
histones, because the replacement H3 subtype in plants also disappears rapidly.
The progressive degradation of H3.3 suggests that some
step in RI replacement consumes the histone, and distinguishes RI replacement
from a simple exchange process (Schwartz, 2005).
The estimate of H3.3 turnover is based on bulk measurements of all the tagged histone in a cell. However, since transcriptional rates vary between sites in the nucleus, histone half-lives at each site might differ. Indeed, a pulse of H3.3-GFP shows labeling with a wide range of intensities in the nucleus (Ahmad, 2002). These intensity differences might indicate densities of H3.3 in chromatin, but an alternative explanation is that intense sites are where the pulse of tagged H3.3 rapidly enters chromatin (Schwartz, 2005).
Overall, the summed intensity of H3.3-GFP within nuclei decreases over a time course. This is similar to what was observed in polytene nuclei, and is consistent with turnover of the histone variant in interphase cells. However the deposition rate hypothesis predicts that the dimming of individual sites will differ, with the brightest foci of H3.3-GFP being the first sites to lose the tagged histone. To detect turnover rates at particular loci, the subnuclear distribution of H3.3-GFP immediately was tracked after a pulse of production and subsequent time points. Labeled nuclei were classed into 'nucleolar' (Nu), 'euchromatic' (Eu), and 'heterochromatic' (Hete) patterns. Previous experiments have shown that nucleolar patterns only result from RI deposition, and heterochromatic ones from replication-coupled deposition. Both early S-phase replication and RI deposition produce euchromatic labeling. Two hours after producing tagged H3.3 in an asynchronous cell culture, 40% of labeled cells show focal staining within the nucleolus. These rDNA foci are approximately five times brighter than the next brightest sites in euchromatin. The rest of the labeled nuclei show replication-coupled H3.3 deposition patterns. However, over the next few hours the nucleolar pattern becomes rare, and as well, the intensity of GFP within nucleoli diminishes. By 20 h after the pulse there are no obvious labeled rDNA foci, and the brightest site within nucleoli averages ~0.8 the intensity of euchromatic sites in the same nucleus. This change in nucleolar intensities is statistically significant. Thus, the amounts of H3.3-GFP at active rDNA genes drop at least sixfold over the 20-h chase. It is concluded that the rDNA foci do gain and then lose the tagged histone more rapidly than other sites in the nucleus. The estimate of the nucleolar turnover rate must be an underestimate, because soluble H3.3 is available for at least the first 8 h of the time course. In spite of this, the turnover rate at the highly transcribed rDNA genes appears to be much faster than the bulk rate, suggesting that this rate corresponds to transcriptional activity (Schwartz, 2005).
If the early intense signals at nucleolar foci are due to the kinetics of RI assembly in the nucleus, continual production of H3.3-RFP should be deposited throughout the nucleus. Indeed, it was found that euchromatin and nucleolar foci are labeled with roughly uniform intensities, while a pulse of H3.3-GFP in the same cells intensely labels nucleolar foci. Thus, with continual production of the labeled histone, it appears that all sites within the nucleus reach a steady state. Notably, at later times the H3.3-GFP signals within the nucleolus are always dimmer than the constitutive H3.3-RFP signals, as if the pulse of protein does not freely exchange within the nucleus. It is concluded that the early intense labeling after pulsed expression results from the more rapid deposition of the histone at these sites. This suggests that the bulk measurements of histone turnover are the average of many sites undergoing replacement, with some sites rapidly replacing histones while many sites do so more slowly (Schwartz, 2005).
In sexually reproducing animals, a crucial step in zygote formation is the decondensation of the fertilizing sperm nucleus into a DNA replication-competent male pronucleus. Genome-wide nucleosome assembly on paternal DNA implies the replacement of sperm chromosomal proteins, such as protamines, by maternally provided histones. This fundamental process is specifically impaired in sésame (ssm; Loppin, 2000), a unique Drosophila maternal effect mutant that prevents male pronucleus formation (Loppin, 2005).
ssm is a point mutation in the Hira gene; the histone chaperone protein HIRA is required for nucleosome assembly during sperm nucleus decondensation. In vertebrates, HIRA has been shown to be critical for a nucleosome assembly pathway independent of DNA synthesis that specifically involves the H3.3 histone variant. This study shows that nucleosomes containing H3.3, and not H3, are specifically assembled in paternal Drosophila chromatin before the first round of DNA replication. The exclusive marking of paternal chromosomes with H3.3 represents a primary epigenetic distinction between parental genomes in the zygote, and underlines an important consequence of the critical and highly specialized function of HIRA at fertilization (Loppin, 2005).
With the exception of its four centromeric regions, the needle-like, extremely condensed Drosophila sperm nucleus is devoid of the core histones (H2A, H2B, H3 and H4) that form the nucleosome particle. Indeed, in elongating spermatids, histones are replaced by sperm-specific chromosomal proteins including protamines. At fertilization, de novo nucleosome assembly in the male pronucleus is a crucial process that determines the ability of the paternal genome to participate in the formation of the diploid zygote. However, the molecular basis of this process in Drosophila is largely unknown. sésame (ssm), a unique maternal effect, embryonic-lethal mutation specifically affects the formation of the male pronucleus (Loppin 2000). In eggs laid by homozygous ssm females, maternal histones are not deposited in the male pronucleus, which remains abnormally condensed. Consequently, haploid embryos develop with only maternal chromosomes and invariably die before hatching (Loppin, 2000). This dramatic phenotype suggests that ssm is involved in the replacement of protamines with histones in the male pronucleus (Loppin, 2001). To identify the ssm gene, the previously defined ssm genetic region (7B6-7C1, X chromosome) was aligned relative to the annotated genome. Of the ten candidate genes predicted in this region, the Hira gene was first tested in rescue experiments. A single copy of a Hira transgene completely rescued the fertility of homozygous ssm females, thereby demonstrating that Hira is the gene affected by the ssm mutation (Loppin, 2005).
Analysis of Hira transcripts by polymerase chain reaction with reverse transcription (RT-PCR) revealed no significant difference in expression between wild-type and ssm flies. HIRA proteins are characterized by seven conserved WD repeats in their amino-terminus, which are predicted to assemble into a ß-propeller structure known to mediate protein-protein interactions. Sequencing of the Hira gene amplified from ssm flies revealed a single point mutation that results in substitution of the highly conserved arginine 225 with a lysine residue (R225K). This residue is located between the fourth and fifth WD repeats of HIRA. The 100% penetrant phenotype of ssm arises from this subtle mutation, indicating that this residue sits in a crucial functional domain of the protein (Loppin, 2005).
HIRA has been shown to be critical for a replication-independent nucleosome assembly pathway, distinct from the major replication-coupled chromatin assembly that occurs during genome duplication (Ray-Gallet, 2002). In the decondensing sperm nucleus, chromatin assembly takes place before the onset of the first zygotic S phase. It was thus reasoned that this process could represent a peculiar case of replication-independent nucleosome assembly at the scale of a whole genome. To test this hypothesis, the distribution of maternal HIRA was studied in fertilized eggs. Transgenic flies were establised expressing HIRA fused to the 3 x Flag tag (HIRA-Flag). Anti-Flag Western blot analysis of early embryos from transgenic females revealed a single band corresponding to the expected size of HIRA-Flag. This fusion protein is functional; the Hira-Flag transgene fully rescues the fertility of ssm females (Loppin, 2005).
Immunofluorescence of eggs from transgenic females crossed with wild-type y w67c males revealed bright HIRA-Flag staining in the male nucleus from the initiation of decondensation, when the sperm nucleus is still elongated, until the end of pronuclear formation. In contrast, HIRA-Flag was never detected in maternal chromosomes at any stage of meiosis or pronuclear formation, and was also absent from embryonic nuclei during early development. It was also verified that sperm from transgenic males did not contribute any detectable level of HIRA-Flag when crossed with wild-type females. Thus, the highly specific distribution of maternal HIRA in the male nucleus is consistent with the observed mutant phenotype (Loppin, 2005).
This result was confirmed using antibodies directed against two peptides from the Drosophila HIRA protein. In wild-type w1118 flies, anti-HIRA antibodies stained the male nucleus exactly like HIRA-Flag did. In addition, anti-HIRA antibodies stained the sperm flagellum non-specifically, as is often the case for rabbit antibodies. In conclusion, the data show that maternal HIRA very efficiently targets the male nucleus immediately after sperm entry. This highly specific distribution of HIRA on paternal chromatin well before the onset of the first S phase makes a strong case for its potential role in replication-independent nucleosome assembly during male pronucleus formation (Loppin, 2005).
In eggs from mutant females stained with anti-HIRA antibodies, it was found that the mutant HIRA is still detected in the male nucleus at all stages of zygote formation. A version of the Hira-Flag transgene was constructed containing the R225K substitution (Hirassm-Flag). As expected, this construct was unable to rescue the ssm phenotype, despite the fact that HIRAssm-Flag brightly stained the male nucleus in mutant eggs. In conclusion, the ssm mutation does not affect the ability of HIRA to localize to the male nucleus (Loppin, 2005).
Increasing evidence supports an important role for histone H3 variants in specifying modes of nucleosome assembly. Whereas the major H3 histones (H3.1 and H3.2) are synthesized during S phase and are deposited on DNA strictly during DNA replication, the histone H3 replacement variant H3.3 is synthesized and deposited onto DNA throughout the cell cycle. Additionally, H3.3 deposition has been shown to mark transcriptionally active chromatin in vivo, suggesting that H3.3 nucleosomes may confer epigenetic inheritance of active chromatin states. Recently, the purification from human cells of complexes for replication-coupled deposition of H3.1 and replication-independent deposition of H3.3 has demonstrated that these alternative nucleosome assembly pathways depend on different chromatin assembly factors: the CAF-1 complex during DNA replication or DNA repair, and the HIRA complex, which is independent of DNA synthesis (Loppin, 2005).
The evidence that vertebrate HIRA is specialized for the deposition of H3.3 nucleosomes on DNA prompted the possible implication of this histone variant in male pronucleus formation in Drosophila. H3, the unique Drosophila S phase variant, and H3.3 differ by only four amino acids, and no specific H3.3 antibody is currently available. Transgenic lines were thus established expressing 3 x Flag-tagged versions of both histone variants. Both constructs use the regulatory sequences of the Drosophila His3.3A gene, resulting in similar levels of both recombinant histones in eggs (Loppin, 2005).
The distribution of H3.3-Flag during fertilization and zygote formation was examined by crossing transgenic females with y w67c males. In all cases, maternal H3.3-Flag specifically accumulated in the male nucleus at all stages of pronuclear formation. In contrast, the polar bodies and female pronucleus did not stain for H3.3-Flag, except for a weak staining that appeared by the time the pronuclei apposed. During the first zygotic mitosis, H3.3-Flag still strongly labelled paternal chromosomes, but the staining faded away after a few nuclear divisions. As expected, H3.3-Flag was absent or barely detected in the male nucleus in eggs from ssm mutant females (Loppin, 2005).
In clear contrast to H3.3-Flag, maternal H3-Flag was never detected in the decondensing male nucleus. However, shortly before pronuclear apposition, both pronuclei and the polar bodies were intensely labelled with H3-Flag. This stage corresponds to the onset of the first S phase in all nuclei, as confirmed by immunostaining of the replication factor PCNA. Thus, as expected, H3-Flag is strictly deposited during S phase. H3-Flag was not detected in maternal meiotic chromosomes, presumably because the transgene is not sufficiently expressed during oocyte differentiation to compete with the S-phase expression of the numerous endogenous H3 genes. After the first nuclear cycle, H3-Flag labelled all nuclei throughout embryonic development. Together, these results indicate that although both variants are present in the egg cytoplasm at fertilization, only H3.3 is deposited in the decondensing male nucleus, and this process requires HIRA (Loppin, 2005).
The absence of HIRA from maternal nuclei suggests that replication-coupled chromatin machinery, presumably involving the CAF-1 complex, is responsible for the limited deposition of H3.3 observed in the female pronucleus and polar bodies. In contrast, H3 is strictly deposited during DNA replication in all nuclei, independent of their origin. Considering that HIRA is expressed in all tissues, it is speculated that the ssm mutation only affects the function of HIRA in the male pronucleus. Indeed, H3.3-Flag deposition in other tissues is not affected by the mutation. A recent study has confirmed that in ssm eggs, the paternal chromatin remains associated with protamines (Jayaramaiah Raja, 2005). A model is whereby favored that the HIRA complex fulfils both protamine removal and replication-independent deposition of H3.3-H4 tetramers in the male nucleus. The R225K mutation would only affect the removal of protamines, thus preventing histone deposition. Stronger Hira alleles are expected to affect viability, as is the case in mouse (Roberts, 2002), for which the Hira-null mutation is embryonic-lethal (Loppin, 2005).
The HIRA-dependent deposition of H3.3 during sperm nucleus decondensation establishes an unsuspected level of epigenetic distinction between parental genomes within the diploid zygote. Considering the fact that most animal sperm nuclei must assemble nucleosomes independent of replication in the egg, it is expected that the HIRA-H3.3 chromatin assembly pathway is widely conserved for this process. This is supported by a recent study in mouse that pointed out the absence of H3.1 in the male pronucleus (van der Heijden, 2005). Moreover, it is well established that in mouse, histone H3 post-translational methylations are differentially distributed between the male and female pronuclei. In particular, di- or tri-methylation of lysine 4 and 9 of H3 are only found in the female pronucleus, at least during the earliest stages of pronuclear formation. Remarkably, these four H3 modifications (di- and tri-methyl K4, and di- and tri-methyl K9) are also restricted to maternal chromosomes in Drosophila eggs and are thus absent on paternal H3.3. Thus, the epigenetic distinction between parental genomes is remarkably conserved, both at the level of H3 variants and H3 lysine methylation. Future investigations should establish the functional significance of this H3.3 epigenetic mark in mammals with respect to the specific loss of paternal DNA methylation at fertilization or the inactivation of the paternal X chromosome in cleavage embryos. Finally, this work underlines the value in using the male pronucleus as a unique model for studying genome-wide replication-independent chromatin assembly in vivo (Loppin, 2005).
Histones of multicellular organisms are assembled into chromatin primarily during DNA replication. When chromatin assembly occurs at other times, the histone H3.3 variant replaces canonical H3. A new strategy has been introduced for profiling epigenetic patterns on the basis of H3.3 replacement, using microarrays covering roughly one-third of the Drosophila melanogaster genome at 100-bp resolution. Patterns of H3.3 replacement were identified over active genes and transposons. H3.3 replacement occurred prominently at sites of abundant RNA polymerase II and methylated H3 Lys4 throughout the genome and is enhanced on the dosage-compensated male X chromosome. Active genes are depleted of histones at promoters and are enriched in H3.3 from upstream to downstream of transcription units. It is proposed that deposition and inheritance of actively modified H3.3 in regulatory regions maintains transcriptionally active chromatin (Mito, 2005).
Cellular memory is maintained at homeotic genes by cis-regulatory elements whose mechanism of action is unknown. Drosophila homeotic gene clusters have been examined by measuring, at high resolution, levels of histone replacement and nucleosome occupancy. Homeotic gene clusters display conspicuous peaks of histone replacement at boundaries of cis-regulatory domains superimposed over broad regions of low replacement. Peaks of histone replacement closely correspond to nuclease-hypersensitive sites, binding sites for Polycomb and trithorax group proteins, and sites of nucleosome depletion. These results suggest the existence of a continuous process that disrupts nucleosomes and maintains accessibility of cis-regulatory elements (Mito, 2007).
Chromatin can be differentiated by the replication-independent replacement of one histone variant with another. For example, histone H3.3 is deposited throughout the cell cycle, replacing H3 that is deposited during replication. Unlike replication-coupled assembly of H3, which occurs in gaps between old nucleosomes on daughter helices, the insertion of H3.3 is preceded by disruption of preexisting histones during transcription and other active processes. H3.3 replacement profiles resemble those for RNA polymerase II, which suggests that gradual replacement of H3.3 occurs in the wake of transiting polymerase to repair disrupted chromatin. This study asked whether histone replacement and nucleosome occupancy are also distinctive at cis-regulatory elements (Mito, 2007).
Log-phase Drosophila S2 cells were induced to produce biotin-tagged H3.3 for two or three cell cycles. DNA was extracted from streptavidin pull-down assay and input material, labeled with Cy3 and Cy5 dyes, and cohybridized to microarrays. To provide a standard, biotin-tagged H3-containing chromatin was profiled in parallel. Analysis of H3.3/H3 levels over the entire 3R chromosome arm revealed that the 350-kb bithorax complex (BX-C) region displays the lowest H3.3/H3 ratio of any region of comparable size on 3R, and the Antennapedia homeotic gene complex (ANTP-C) also displays an unusually low H3.3/H3 ratio. Low H3.3/H3 ratios at the homeotic gene clusters are attributable to infrequent histone replacement, and not to low nucleosome occupancy, because H3.3 levels at the BX-C are far below the median for all of 3R, whereas H3 levels are slightly above the median overall. Even the heterochromatic chromosome 4 includes only shorter (100-kb) stretches that are as depleted in H3.3 as the BX-C (Mito, 2007).
A close-up view of the BX-C iab region reveals the presence of several prominent H3.3 peaks. Notably, the seven highest peaks correspond to the functional boundaries of the seven proximal-to-distal cis-regulatory domains that regulate the abd-A (iab2 to iab4) and Abd-B (iab5 to iab8) homeotic genes successively from anterior to posterior in the abdomen. Conspicuous peaks of H3.3 also correspond to the bxd Polycomb response element (PRE) and to promoters within the Abd-B gene, which is known to be active in S2 cells. Therefore, each of the most prominent H3.3 peaks in the region corresponds to a previously defined cis-regulatory element. These findings are likely to be general, because in budding yeast, promoters and boundaries are also sites of intense histone replacement (Mito, 2007).
A characteristic feature of both boundaries and PREs in the BX-C is that they span deoxyribonuclease I (DNaseI)-hypersensitive sites in a variety of cell types, including S2 cells. To better delineate histone replacement patterns in the vicinity of hypersensitive sites, the entire BX-C was tiled at 20-bp resolution. The bxd, Mcp, Fab-7, and Fab-8 PRE-boundaries each encompass conspicuous peaks of H3.3 abundance that closely correspond to all the known nuclease-hypersensitive sites within the region. Nuclease hypersensitivity identifies sites of relatively accessible DNA, so that their correspondences to peaks of histone replacement suggest that continuous disruption of nucleosomes exposes cis-regulatory DNA relative to surrounding regions (Mito, 2007).
PRE-boundary elements in the BX-C and other regions are binding sites for multiple Polycomb group (PcG) proteins, which have been mapped in an S2 cell line at high resolution. If the process that disrupts nucleosomes also facilitates PcG binding, then a correspondence would be expected between peaks of PcG binding and peaks of H3.3. Indeed, when H3.3 profiles were compared with those for Enhancer-of-zeste (EZ) and Posterior-sex-combs (PSC) PcG proteins, all 10 peaks of PcG binding in the abdominal region were found to be local peaks of H3.3. Likewise at the ANTP-C, all 13 peaks of PcG binding in the Scr-Antp region correspond to high levels of H3.3. H3.3 enrichment at PcG-binding sites is not attributable to higher nucleosome occupancy, because essentially identical results were obtained for H3.3/H3 profiles (Mito, 2007).
Not all PREs in the BX-C are found to be sites of PcG binding; for example, neither Fab-7 nor Fab-8 is detectably bound by EZ or PSC. The fact that all PcG sites are peaks of histone replacement, but not vice versa, suggests that histone replacement at PREs and boundaries is constitutive and independent of the expression of the homeotic genes that they regulate. For example, Abd-B is expressed at high levels in S2 cells and displays the typical H3.3 5' peak for an active gene, whereas Ubx and abd-A are nearly inactive, yet the PREs and boundaries regulating all three genes are sites of conspicuous histone replacement over a low background (Mito, 2007).
Histone replacement averaged over the 175 genomewide EZ+PSC peaks outside of the BX-C and ANTP-C was examined and an H3.3 peak was observed centered over the PcG maximum. Therefore, the strong association between PcG protein binding and histone replacement is not limited to homeotic gene clusters. The genomewide H3.3 peak is higher than that for the BX-C and ANTP-C, presumably because other PcG-binding sites are not superimposed over such deep H3.3 valleys (Mito, 2007).
The colocalization of PcG-binding sites and local peaks of H3.3 suggests that the process that disrupts nucleosomes locally maintains the accessibility of cis-regulatory DNA to PcG proteins. If so, then there should be a lower average occupancy of nucleosomes over sites of PcG protein binding than over their surrounding regions. To test this possibility, nucleosomal DNA and fragmented genomic DNA were hybridized on the same microarrays, and nucleosomal/genomic DNA log ratios were measured. Around peak regions of EZ+PSC binding, nucleosomal DNA was clearly depleted on average, similar to the depletion seen for active gene promoters, and essentially the same results were obtained with different methods for genomic DNA fragmentation. It is concluded that the correspondence between histone replacement and nucleosome depletion is a genomewide feature of PcG-binding sites (Mito, 2007).
In Drosophila, many cis-regulatory elements, including PREs and boundaries, are bound by the trxG proteins, Zeste and GAGA factor (GAF). To test the possibility that histone replacement is enhanced and nucleosome occupancy is reduced where Zeste protein preferentially binds, 390 Zeste-binding sites identified by high-resolution chromatin immunoprecipitation (ChIP) combined with tiling microarrays (ChIP-chip profiling) were aligned, and log ratios of H3.3/H3 and nucleosome occupancy were averaged. A prominent maximum of histone replacement and a sharp minimum of nucleosome occupancy was observed centered over the point of alignment. Similar results were obtained for predicted GAF sites, which suggests that nucleosome disruption is a general feature of trxG protein DNA-binding sites. H3.3 enrichment at PcG- and trxG protein-binding sites results from a replication-independent replacement process, because essentially identical profiles were obtained for H3.3core, which lacks the N-terminal tail and does not assemble during replication (Mito, 2007).
Like Fab-7 and Fab-8, heat shock gene promoters are prominent sites of GAF binding, nuclease hypersensitivity, and reduced nucleosome occupancy. Heat shock protein Hsp70 genes are constitutively 'poised' for rapid induction, but do not produce detectable mRNAs in the uninduced state. Hsp70 genes were aligned at their 5' ends and H3.3 and H3 profiles were averaged. For comparison, similarly aligned H3.3 and H3 profiles were averaged for all 2165 genes on 3R with known 5' and 3' ends, divided into quintiles based on expression levels. H3.3 patterns were similar to those of highly active genes, with histone replacement levels peaking on either side of heat shock promoters. As do transcriptionally active gene promoters, heat shock genes display prominent H3.3 and H3 dips in abundance that are attributable to partial nucleosome depletion. Constitutive histone replacement also appears to be a feature of poised promoters in vertebrates, because H3.3 is strongly enriched in the upstream region of the chicken folate receptor gene, regardless of whether the gene is active or inactive (Mito, 2007).
What process maintains the chromatin of cis-regulatory elements in a state of flux? Many DNA-binding and chromatin-binding proteins involved in gene regulation display short residence times on DNA, and some mouse transcription factors show dynamic behavior at their functional binding sites. A model for this process has been proposed, involving alternating cycles of nucleosome disruption by a Brahma-related SWI/SNF chromatin-remodeler and transcription factor binding. The binding of PcG and trxG proteins is also dynamic, and it is proposed that a similar cycle of nucleosome disruption and factor binding takes place at boundaries and PREs. Nucleosome disruption by SWI/SNF remodeling complexes would occasionally evict nucleosomes and transiently expose DNA, which would become available to other diffusible factors, including PcG proteins. The continued local presence of nucleosome remodelers would result in another cycle of remodeling, nucleosome depletion, nuclease hypersensitivity, and histone replacement at the site. This model could account for the diversity of trxG proteins, which include DNA-binding proteins (Zeste and GAF), nucleosome remodelers (Brahma and Kismet), and histone methyltransferases (Trithorax and Ash1) that are specific for H3K4, a modification that is highly enriched on H3.3. The resulting dynamic process would allow for proteins that promote opposite epigenetic outcomes to act at common cis-regulatory sites (Mito, 2007).
Epigenetic maintenance of the expression state of the genome is critical for development. Drosophila GAGA factor interacts with FACT and modulates chromatin structure for the maintenance of gene expression. This study shows that the GAGA factor-FACT complex (Fact is a heterodimer of dSPT16 and dSSRP1; Shimojima, 2003) and its binding site just downstream from the white gene are crucial for position effect variegation. Interestingly there is a dip of histone H3 Lys 9 methylation and a peak of H3 Lys 4 methylation at this site. The GAGA factor and FACT direct replacement of histone H3 by H3.3 through association of HIRA at this site, and maintain white expression under the heterochromatin environment. Based on these findings it is proposed that the GAGA factor and FACT-dependent replacement of Lys 9-methylated histone H3 by H3.3 counteracts the spreading of silent chromatin (Nakayama, 2007).
This study shows that the GAGA factor-FACT complex is present on the GAGA factor-binding DNase-hypersensitive site d1, a site just downstream from w, and participates in PEV. d1 appears to be a peculiar site where histone H3 K4 methylation peaks and H3 K9 methylation dips, and necessary and sufficient to counteract the heterochromatin spreading. The GAGA factor and FACT contribute to replacement of histone H3 by H3.3 through association of histone H3.3 chaperone HIRA to d1, and maintains w expression under the heterochromatin environment. Based on these data, the following model is proposed for the maintenance of the active state against the spreading of silent chromatin. Heterochromatin is marked by K9-methylated histone H3 and its binding protein HP1, and has a tendency to spread into neighboring regions. Histone H3.3 replacement is thought to be achieved through either eviction of a nucleosome and deposition of a H3.3-containing nucleosome or stepwise disassembly-reassembly without eviction of a nucleosome. Since the GAGA factor-FACT complex facilitates chromatin remodeling and the GAGA factor is known to generate a nucleosome-free region around its binding site, it is most likely that eviction or disassembly of a nucleosome occurs at the DNase-hypersensitive site of d1. The GAGA factor and FACT participate association of HIRA to d1 and the histone replacement would be accomplished by subsequent deposition or reassembly of a H3.3-containing nucleosome. This process would be repeated constantly to eliminate K9-methylated histone H3 at d1 and counteract the spreading of silent chromatin (Nakayama, 2007).
It has been reported that histone H3.3 replacement is triggered by transcription elongation. However, genome-wide profiling has shown histone H3.3 replacement from upstream of to downstream from transcription units. Although some of the replacement may be explained by elongation during intergenic transcription, the histone H3.3 replacement at d1 appears to occur independent of transcription elongation. Thus, the present study indicates a distinct pathway for histone H3.3 replacement (Nakayama, 2007).
Transcription of the w adjacent gene CG32795 has been reported to start immediately after the GAGA factor-binding sequence of d1, suggesting that d1 is a part of the promoter region of CG32795. Therefore, the effect was examined of Trl and spt16 mutations on expression of CG32795. The reduction of a single dose of Trl or spt16 affect the CG32795 expression in the wm4 context but not in the w+ context. These data are consistent with the idea that d1 is a functional promoter element of CG32795 in the w+ context, although Trl and spt16 become haplo-insufficient only when the accessibility of the GAGA factor-FACT complex to d1 decreased under the heterochromatin environment. This raises the possibility that the protection from heterochromatin spreading by the GAGA factor and FACT at d1 is a consequence of their function within the CG32795 promoter. However, conventional promoters do not have a barrier function against heterochromatin silencing. For example, the presence of GAL4 (or E2F) on a promoter carrying GAL4 (or E2F)-binding site did not modify PEV of the attached reporter gene. Genome-wide profiling of H3.3 replacement in Drosophila has revealed the clear dip of H3.3-containing nucleosomes at immediately upstream of the transcription start sites of active genes. This is in sharp contrast with the case of d1, where peaks were observed of both the H3.3/H3 ratio and the actual H3.3 level, and illuminates the difference between d1 and ordinary promoters. Furthermore, the GAGA factor-dependent histone H3.3 replacement was detected also at the DNase HS1 in the Fab-7 boundary of Abd-B, where no promoter activity has been demonstrated. These findings indicate that the GAGA factor and FACT-dependent histone H3.3 replacement can occur without promoter functions. Nevertheless, the barrier function could be assisted by the putative promoter activity of d1 such as formation of a transcription initiation complex (Nakayama, 2007).
The GAGA factor-binding sequence at d1 consists of (GA)8. Since GAGA factor forms an oligomer through its BTB domain, the factor can make a cooperative and stable binding to closely spaced GAGAG elements. This is presumably the reason why d1 gave a prominent signal among the GAGAG sequences around w in the ChIP assay. Because the GAGA factor occupies many closely spaced GAGAG sequences within the Drosophila genome including the Polycomb/trithorax response elements of Hox genes, the proposed mechanism may operate not only in loci juxtaposed with heterochromatin but also in other loci such as the regulatory regions of Hox genes. Indeed GAGA factor and FACT-dependent histone H3.3 replacement were observed in the Fab-7 boundary of Abd-B. High levels of histone H3.3 have been also reported at the locus control region of the chicken folate receptor gene, suggesting that the barrier function against the chromatin silencing via histone H3.3 replacement may be evolutionarily conserved up to vertebrates (Nakayama, 2007).
The organization of chromatin affects all aspects of nuclear DNA metabolism in eukaryotes. H3.3 is an evolutionarily conserved histone variant and a key substrate for replication-independent chromatin assembly. Elimination of chromatin remodeling factor CHD1 in Drosophila embryos abolishes incorporation of H3.3 into the male pronucleus, renders the paternal genome unable to participate in zygotic mitoses, and leads to the development of haploid embryos. Furthermore, CHD1, but not ISWI, interacts with HIRA in cytoplasmic extracts. These findings establish CHD1 as a major factor in replacement histone metabolism in the nucleus and reveal a critical role for CHD1 in the earliest developmental instances of genome-scale, replication-independent nucleosome assembly. Furthermore, these results point to the general requirement of adenosine triphosphate (ATP)-utilizing motor proteins for histone deposition in vivo (Konev, 2007).
Histone-DNA interactions constantly change during various processes of DNA metabolism. Recent studies have highlighted the importance of histone variants, such as H3.3, CENP-A (centromere protein A), or H2A.Z, in chromatin dynamics. Incorporation of replacement histones into chromatin occurs throughout the cell cycle, whereas nucleosomes containing canonical histones are assembled exclusively during DNA replication. A thorough understanding of the replication-independent mechanisms of chromatin assembly, however, is lacking (Konev, 2007).
In vitro, chromatin assembly requires the action of histone chaperones and adenosine triphosphate (ATP)-utilizing factors. Histone chaperones may specialize for certain histone variants. For example, H3.3 associates with a complex containing HIRA, whereas canonical H3 is in a complex with CAF-1 (chromatin assembly factor 1) (Tagami, 2004). The molecular motors known to assemble nucleosomes are ACF (ATP-utilizing chromatin assembly and remodeling factor), CHRAC (chromatin accessibility complex), and RSF (nucleosome-remodeling and spacing factor), which contain the Snf2 family member ISWI as the catalytic subunit, and CHD1, which belongs to the CHD subfamily of Snf2-like adenosine triphosphatases (ATPases). These factors have not been shown to mediate deposition of histones in vivo. It has been demonstrated that CHD1, together with the chaperone NAP-1, assembles nucleosome arrays from DNA and histones in vitro (Lusser, 2005). This study investigated the role of CHD1 in chromatin assembly in vivo in Drosophila (Konev, 2007).
Chd1 alleles were generated by P element-mediated mutagenesis. Two excisions, Df(2L)Chd1 and Df(2L)Chd1, deleted fragments of the Chd1 gene and fragments of unrelated adjacent genes. Heterozygous combinations, however, of Chd1 or Chd1 with Df(2L)Exel7014 affect both copies of the Chd1 gene only. Also a single point mutation was identified that results in premature translation termination of Chd1 (Q1394*) in a previously described lethal allele, l(2)23Cd[A7-4]. Hence, l(2)23Cd[A7-4] was renamed Chd1 (Konev, 2007).
Analysis of Western blots of embryos from heterozygous Chd1 fruit flies revealed the presence of a truncated polypeptide besides full-length CHD1. No truncated polypeptides were detected in heterozygous Chd1 or Chd1 embryos. Therefore, the corresponding deficiencies result in null mutations of Chd1. Crosses of heterozygous Chd1 mutant alleles with Df(2L)JS17/CyO or Df(2L)Exel7014/CyO produced subviable adult homozygous mutant progeny. Both males and females were sterile. Homozygous null females mated to wildtype males laid fertilized eggs that died before hatching. Therefore, maternal CHD1 is essential for embryonic development (Konev, 2007).
When the chromosome structure of 0- to 4-hour-old embryos laid by Chd1-null females were examined, it was observed that, during syncytial mitoses (cycles 3 to 13), the nuclei appeared to be abnormally small. The observed numbers of anaphase chromosomes suggested that they were haploid. To confirm this observation, wild-type or Chd1-null females were mated with males that carried a green fluorescent protein (GFP) transgene. Embryonic DNA was amplified with primers detecting male-specific GFP and a reference gene, Asf1. In wild-type embryos, both primer pairs produced polymerase chain reaction (PCR)-amplified products, whereas only the Asf1 fragment was amplified in the mutants. Thus, Chd1 embryos develop with haploid, maternally derived chromosome content (Konev, 2007).
To investigate the causes of haploidy in mutant animals, distributions of various developmental stages were compared in samples of wildtype and Chd1-null embryos. The lack of maternal CHD1 dramatically changed this distribution. Most notably, at 0 to 4 hours after egg deposition, the majority of Chd1 embryos (56%) remained at a very early stage of development in contrast to the wild type (24%) (Konev, 2007).
In Drosophila eggs, meiosis gives rise to four haploid nuclei. When the egg is fertilized, one of them is selected as a female pronucleus; the remaining three form the polar body. After breakdown of the sperm nuclear envelope, the compacted sperm chromatin is decondensed, and sperm-specific protamines are replaced with maternal histones. The male and female pronuclei juxtapose in the middle of the embryo and undergo one round of separate haploid mitoses. The resulting products fuse with their counterparts to give two diploid nuclei. In the majority of Chd1 embryos, partial decondensation of the sperm chromatin and normal apposition of parental pronuclei were observed. Then, however, one pronucleus underwent mitosis; the other one did not. Considering the subsequent loss of paternal DNA, it is concluded that mitotic progression of the male pronucleus is hindered in Chd1 embryos (Konev, 2007).
Because CHD1 can assemble nucleosomes in vitro, it was asked whether the absence of CHD1 affects histone incorporation into the male pronucleus. Embryos from wild-type or Chd1-null females were stained with an antibody against histone H3. In wild-type embryos, uniform staining was observed in both parental pronuclei. In contrast, in Chd1-null embryos only the female chromatin was brightly stained. The male pronucleus contained considerably less histone H3. These observations indicate that CHD1 is necessary for nucleosome assembly during sperm decondensation (Konev, 2007).
Sperm DNA does not replicate during decondensation, and histones are deposited by replication-independent assembly mechanisms, which involve the variant histone H3.3 but not canonical H3. It has been shown in Drosophila and mice that H3.3 is specifically present in the male pronucleus (Loppin, 2005; Torres-Padilla, 2006). The distribution of H3.3 was analyzed in embryos derived from Chd1-null females that carry a FLAG-tagged H3.3 transgene. In wild-type embryos, colocalization of the H3.3-FLAG signal was observed with male pronuclear DNA during migration and apposition. No H3.3-FLAG was detectable in the maternal pronucleus. In Chd1-null embryos, the male pronucleus showed altered H3.3-FLAG staining. The signal did not co-localize with the DNA but remained constrained to the nuclear periphery in a sacshaped pattern (Konev, 2007).
These findings suggest that in the earliest phases of Drosophila development CHD1 is essential for the incorporation of H3.3 and normal assembly of paternal chromatin. In contrast, CHD1 does not appear to affect the organization of maternal chromatin. It is concluded that CHD1 is required for replication-independent nucleosome assembly in the decondensing male pronucleus, but is dispensable for replication-coupled incorporation of H3 (Konev, 2007).
It has recently been shown that the sèsame (ssm) mutation of Drosophila histone chaperone HIRA causes the development of haploid embryos and abolished H3.3 deposition into the male pronucleus. Chd1 and ssm mutants, however, differ profoundly in the manifestation of this phenotype. In ssm embryos, H3.3 is absent from the male pronucleus. In contrast, in Chd1-null embryos, H3.3 delivery to the male pronucleus appears to be unaffected. Thus, these observations allow the roles of CHD1 and HIRA to be mechanistically discerned. Whereas HIRA is essential for histone delivery to the sites of nucleosome assembly, CHD1 directly facilitates histone deposition. These findings are consistent with observations in vitro that histone chaperones either do not assemble nucleosomes or assemble them at a greatly reduced rate in the absence of ATP-utilizing factors. These data provide evidence that histone deposition in vivo also transpires through an ATP-dependent mechanism (Konev, 2007).
CHD1 has been implicated in transcription elongation-related chromatin remodeling (Sims, 2004). This study demonstrates that CHD1 functions in nucleosome assembly in the early Drosophila embryo, which is transcriptionally silent. The biological role of CHD1, therefore, is not confined to transcription-related processes. The Schizosaccharomyces pombe homolog of CHD1, Hrp1, has been shown to function in loading of the centromere-specific H3 variant CENP-A (Walfridsson, 2005). Similarly to H3.3, incorporation of CENP-A into chromatin is not restricted to S phase. Therefore, CHD1 may have a general role in replication-independent nucleosome assembly (Konev, 2007).
Sperm decondensation involves not only histone incorporation, but also eviction of protamines. To discern whether CHD1 has a role in this process, the fate of protamine B (Mst35Bb) was analyzed in Chd1-null embryos. Although GFP-tagged Mst35Bb was detected in the sperm head immediately upon fertilization, no Mst35Bb-GFP signal was detected in the male pronucleus. Thus, like HIRA, CHD1 is dispensable for protamine removal. This study has shown that the male pronucleus in Chd1-null embryos contains very low amounts of histones, yet the DNA is not packaged with protamines. It remains an open question whether other DNA-protein complexes exist in the male pronucleus (Konev, 2007).
Drosophila eggs contain stores of both known chromatin assembly factors CHD1 and ISWI. Nevertheless, ISWI is unable to substitute for CHD1 in the deposition of H3.3. To examine whether CHD1 and ISWI differ in their ability to interact with the H3.3 chaperone HIRA, coimmunoprecipitation experiments were performed with extracts from embryos expressing FLAG-HIRA. CHD1 signal was readily detected in FLAG-specific immunoprecipitates, whereas ISWI did not coimmunoprecipitate with HIRA. Thus, a fraction of CHD1, but not ISWI, physically associates with HIRA. This property of CHD1 may account for its unique function in the H3.3 deposition process (Konev, 2007).
A subpopulation of Chd1 mutant haploid embryos survives beyond apposition stage. Therefore, it was asked whether H3.3 deposition is altered in Chd1 mutant embryos during later developmental stages. In wild-type nuclei, the H3.3-FLAG signal originating from the male pronucleus becomes undetectable after 2 to 3 divisions. Most maternal H3.3 remains distributed diffusely throughout the syncytium. After cycle 11 (roughly correlating with the onset of zygotic transcription) H3.3-FLAG is redistributed into the nuclei, where it colocalizes with the DNA. In contrast, incorporation of H3.3 into Chd1 mutant nuclei was impaired. H3.3-FLAG produced a speckled staining with numerous bright dots that poorly overlapped with the maxima of DNA staining. It is important to note that, in the ssm (HIRA) mutant, H3.3 incorporation defects in tissues or developmental stages other than the apposition stage were not observed. This result is consistent with the idea that misincorporation of H3.3 in Chd1 embryos is a direct effect of CHD1 deletion rather than a consequence of haploid development. It is also concludes that CHD1 functions in H3.3 deposition during later stages of embryonic development, possibly in a HIRA-independent fashion (Konev, 2007).
This study provides evidence that ATP-dependent mechanisms are used for histone deposition during chromatin assembly in vivo. Thus, molecular motor proteins, such as CHD1, function not only in remodeling of existing nucleosomes but also in de novo nucleosome assembly from DNA and histones. Finally, this work identifies CHD1 as a specific factor in the assembly of nucleosomes that contain variant histone H3.3 (Konev, 2007).
The differentiation of post-meiotic spermatids in animals is characterized by a unique reorganization of their nuclear architecture and chromatin composition. In many species, the formation of sperm nuclei involves the massive replacement of nucleosomes with protamines, followed by a phase of extreme nuclear compaction. At fertilization, the reconstitution of a nucleosome-based paternal chromatin after the removal of protamines requires the deposition of maternally provided histones before the first round of DNA replication. This process exclusively uses the histone H3 variant H3.3 and constitutes a unique case of genome-wide replication-independent (RI) de novo chromatin assembly. Previous studies have shown that the histone H3.3 chaperone HIRA plays a central role for paternal chromatin assembly in Drosophila. Although several conserved HIRA-interacting proteins have been identified from yeast to human, their conservation in Drosophila, as well as their actual implication in this highly peculiar RI nucleosome assembly process, is an open question. This study shows that Yemanuclein (YEM), the Drosophila member of the Hpc2/Ubinuclein family, is essential for histone deposition in the male pronucleus. yem loss of function alleles affect male pronucleus formation in a way remarkably similar to Hira mutants and abolish RI paternal chromatin assembly. In addition, it was demonstrated that HIRA and YEM proteins interact and are mutually dependent for their targeting to the decondensing male pronucleus. Finally, this study shows that the alternative ATRX/XNP-dependent H3.3 deposition pathway is not involved in paternal chromatin assembly, thus underlining the specific implication of the HIRA/YEM complex for this essential step of zygote formation (Orsi, 2013).
In human cells, the HIRA core complex is composed of at least three subunits, including HIRA, UBN1 and CABIN1 (Amin, 2011). This complex is functionally involved in a large diversity of cellular and developmental processes that require dynamic histone turnover or de novo assembly of nucleosomes, independently of DNA synthesis. Although the HIRA complex mediates the deposition of the highly conserved H3.3 histone variant, its subunits display a comparatively weak overall conservation in animals. For instance, Drosophila does not seem to have any CABIN1 homolog and the highest conservation between UBN1 and YEM is mainly restricted to the small HRD domain. Despite this poor conservation, this work establishes Yemanuclein as a bona fide ortholog of Ubinuclein, by demonstrating its physical interaction with the HIRA histone chaperone and its critical requirement for H3.3 deposition during male pronucleus decondensation (Orsi, 2013).
In contrast to the knock-out of the Hira gene in mouse, which is zygotic lethal in early embryos, null mutants of Drosophila Hira are viable but homozygous females are completely sterile. This indicates that only the maternal contribution of Hira is essential, at least to form the male pronucleus. Characterization of a null yem2 allele allowed led to the same conclusion for YEM. Remarkably, the phenotype of the male pronucleus in eggs laid by yem mutant females appeared indistinguishable to what was previously reported for Hira mutants. In both cases, RI deposition of H3.3-containing nucleosomes is practically abolished, typically preventing the full decondensation of the male nucleus and its integration into the zygotic nucleus. Thus, YEM and HIRA are equally required to assemble paternal nucleosomes at fertilization. This unique and major function of the HIRA complex is most likely conserved in animal groups where histones, and notably H3 and H4, are replaced with sperm-specific nuclear basic proteins (SNBPs) in sperm. This is for instance the case of mammals, where protamines package about 95% and 85% of mouse and human sperm DNA, respectively. In fact, HIRA has been previously detected in the decondensing male nucleus at fertilization in mouse, which incorporates H3.3 before the first round of DNA replication. It is thus expected that Ubinuclein1/2 is also involved in paternal chromatin assembly in mammals. In apparent contradiction with this prediction, a transgene expressing human UBN1 in the female germline could not rescue the sterility of yem mutant females. However, this absence of complementation of YEM and UBN1 can be explained by the strong divergence of these orthologous proteins at the primary sequence level and it suggests that UBN1 can only function within its native, human HIRA complex. The apparent lack of a CABIN1 homolog in Drosophila also underlines the central role played by the HIRA-UBN1/YEM pair in the complex. Interestingly, while the implication of HIRA and UBN1 for RI deposition of H3.3 in vivo was recently demonstrated in human cells, CABIN1 seemed to play only an auxiliary role in this context. Possibly, CABIN1 could be important for human-specific functions of the HIRA complex, such as the formation of senescence-associated heterochromatin foci (Orsi, 2013).
Previous studies have shown that HIRA specifically accumulates in the sperm nucleus shortly after its delivery in the egg cytoplasm. This study has established that maternally expressed YEM similarly accumulates in the male nucleus at fertilization and until pronuclear apposition. Strikingly, it was also shown that HIRA and YEM are mutually dependent for their targeting to the male nucleus, strongly suggesting that these proteins physically interact during the assembly of paternal nucleosomes. However, nothing is known about the mechanism responsible for their rapid and specific localization in the fertilizing sperm nucleus, which is delivered in the cytoplasm of the gigantic egg cell. Previous studies have established that the HIRA-dependent assembly of paternal nucleosomes occurs after the removal of sperm protamines. This opens the simple possibility that the HIRA complex could recognize exposed sperm DNA immediately after the removal of SNBPs. Interestingly, pioneer work on YEM by Ait-Ahmed (1992) had established that this maternal protein was able to bind DNA in vitro. This property could be important to efficiently target the HIRA complex to sites of de novo nucleosome assembly in the decondensing male nucleus. This hypothesis has recently received indirect experimental support in human cultured cells (RayGallet, 2011). That study established that HIRA, UBN1 and CABIN1 were all individually able to bind DNA in vitro, and it was proposed that this remarkable property could allow the HIRA complex to target naked DNA for H3.3 deposition. Accordingly, this HIRA-dependent nucleosome gap-filling mechanism has been shown to participate in the maintenance of genome integrity, but could also be employed, at the genome-wide scale, for de novo assembly of paternal chromatin at fertilization (Orsi, 2013).
Finally, the observation that YEM accumulates in discrete nuclear regions in both the male nucleus (this study) and the oocyte karyosome (Meyer, 2010) opens the possibility that YEM could perform additional roles not related to nucleosome assembly (Orsi, 2013).
Despite its expression in the female germline, this study found that Drosophila ATRX/XNP is not targeted to the male nucleus and does not seem to play any role in male pronucleus formation. Among the 17 SNF2 type chromatin remodelers present in Drosophila, the Chromodomain-helicase-DNA-binding protein 1 (CHD1) is the only one that has been implicated in the remodeling of paternal chromatin at fertilization. In contrast to Hira and yem, mutations in chd1 do not drastically affect H3.3 incorporation in paternal chromatin but still severely compromise the decondensation of the male nucleus, which appears aberrant in shape. In contrast to a previously reported HIRA/CHD1 interaction, this study could not detect any interaction between these proteins in ovaries, using experimental conditions that permitted co-immunoprecipitation of HIRA and YEM. These results thus suggest that the role of CHD1 in the male nucleus is distinct from the nucleosome assembly process mediated by the HIRA complex (Orsi, 2013).
Although the implication of the HIRA histone chaperone in paternal chromatin assembly was firmly established a few years ago, it has remained unclear until now if this highly specialized RI assembly process also involved other subunits of the HIRA complex or other histone deposition pathways. In fact,previous work has shown that the histone chaperone ASF1, which is known to interact with both the CAF1 and HIRA complexes, is actually absent from the decondensing male nucleus (Bonnefoy, 2007). Although the role, if any, of ASF1 in paternal chromatin assembly awaits a proper functional characterization, it is not expected that this histone chaperone is directly involved in the assembly of nucleosomes on paternal DNA. Accordingly, ASF1 has been previously shown to be dispensable for direct de novo RC or RI histone deposition in Xenopus egg extracts (Orsi, 2013).
The complete failure of the male nucleus to assemble its chromatin in Hira or yem mutant eggs demonstrates that no other nucleosome assembly machinery can substitute for the HIRA-YEM complex in this peculiar context. However, the functional requirement of H3.3 itself in this process is not known. In Drosophila, H3.3 is not absolutely required for survival but it is essential for both male and female fertility. Viability of His3.3A; His3.3B double null mutants could be explained by the fact that, in the absence of H3.3, canonical H3 can be assembled in a RI manner. Although the mode of RI deposition of replicative H3 in these mutants is not known, it opens the possibility that HIRA could use canonical H3 in certain critical circumstances, such as a limiting availability of H3.3. This compensatory mechanism, however, is apparently not possible in Drosophila spermatocytes, where H3.3 is required for the correct segregation of chromosomes during meiotic divisions, underlining the importance of this variant for sexual reproduction. Similarly, future work should aim at determining whether H3.3 is specifically required for the assembly of paternal nucleosomes at fertilization (Orsi, 2013).
Both HIRA and YEM proteins, which are presumably expressed from germinal nurse cells, display a remarkable accumulation in the oocyte nucleus during oogenesis. Most of the volume of the large germinal vesicle is devoid of DNA as the maternal genome is tightly packaged within the karyosome. The presence of HIRA and YEM in the nucleoplasm of the GV is thus not related to nucleosome assembly. However, the fact that HIRA and YEM are mutually dependent for their accumulation in the GV suggests that they are stored in this compartment as a complex. In contrast to the null alleles, point mutations do not affect HIRA/YEM localization in the GV, suggesting that the mechanisms controlling their recruitment to the GV or to the male pronucleus are distinct. This could reflect the fact that the HIRA complex is active in the male pronucleus where these proteins are in a chromatin environment in contrast to their nucleoplasm distribution in the germinal vesicle (GV). Whether or not this transient accumulation of HIRA/YEM in the GV plays any role in the maturation of the complex before paternal chromatin assembly at fertilization remains to be tested. Interestingly, it has been proposed that in human cells, formation of senescence-associated heterochromatin foci by HIRA requires its prior localization to promyelocytic leukemia nuclear bodies, suggesting that these structures could participate in the formation of the HIRA complex before its translocation to chromatin. It should be mentioned, however, that dATRX/XNP also accumulates in the GV despite its dispensability for paternal chromatin assembly. A recent study (Singer, 2011) reported the presence of several nuclear proteins in the GV with no known function in the oocyte, suggesting that this structure could serve as a storage compartment for a large number of nuclear proteins (Orsi, 2013).
In conclusion, this characterization of Drosophila Yemanuclein demonstrates that this protein is a functional partner of HIRA in vivo. It also establishes that HIRA and YEM directly cooperate in the male nucleus for the genome-wide replacement of sperm protamines with H3.3-containing nucleosomes. The specific requirement of the HIRA complex in this unique developmental chromatin assembly process implies the existence of specific properties not shared with other H3.3-deposition pathways. In this regard, future work should explore the potentially conserved DNA binding property of the HIRA complex and its potential role in targeting the fertilizing sperm nucleus in animals (Orsi, 2013).
Meiosis is characterized by two chromosome segregation rounds (Meiosis I and II), which follow a single round of DNA replication, resulting in haploid genome formation. Chromosome reduction occurs at meiosis I. It relies on key structures, such as chiasma, which is formed by repair between homologous chromatids of a double-strand break (DSB) in one of them; to function for segregation of homologues chiasma in turn relies on maintenance of sister chromatid cohesion. In most species, chiasma formation requires the prior synapsis of homologous chromosome axes, which is signaled by the Synaptonemal Complex (SC), a tripartite proteinaceous structure specific to prophase I of meiosis. Yemanuclein (YEM) is a maternal factor that is crucial for sexual reproduction. It is required in the zygote for chromatin assembly of the male pronucleus as a histone H3.3 chaperone in complex with HIRA. This study reports YEM association to the SC and the cohesin complex. A genetic interaction between yem1 (V478E) and the Spo11 homologue mei-W68, added to a yem1 dominant effect on crossover distribution suggest an early role in meiotic recombination. This is further supported by the impact of yem mutations on DSB kinetics. Hira mutant showed a similar effect presumably through disruption of HIRA-YEM complex (Meyer, 2014).
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