Assembling chromatin: The long and winding road

Biochimica et Biophysica Acta 1819 (2012) 196–210

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Biochimica et Biophysica Acta j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / b b a g r m

Assembling chromatin: The long and winding road☆ Anthony T. Annunziato ⁎ Biology Department, Boston College, 140 Commonwealth Avenue, Chestnut Hill, MA 02467, USA

a r t i c l e

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Article history: Received 17 May 2011 Received in revised form 29 June 2011 Accepted 4 July 2011 Available online 18 July 2011 Keywords: Chromatin Replication Assembly Histone Chaperone Factor

a b s t r a c t It has been over 35 years since the acceptance of the “chromatin subunit” hypothesis, and the recognition that nucleosomes are the fundamental repeating units of chromatin fibers. Major subjects of inquiry in the intervening years have included the steps involved in chromatin assembly, and the chaperones that escort histones to DNA. The following commentary offers an historical perspective on inquiries into the processes by which nucleosomes are assembled on replicating and nonreplicating chromatin. This article is part of a Special Issue entitled: Histone chaperones and Chromatin assembly. © 2011 Elsevier B.V. All rights reserved.

1. Prolog In January of 1974 I was a first-year graduate student at the University of Massachusetts, preparing to enter the laboratory of Christopher L. Woodcock. Just a month or so earlier, Chris [1], and Don and Ada Olins [2], had independently presented the first electron micrographs of “beads-on-a-string” chromatin fibers. We didn't call the beads nucleosomes in those days: “chromatin subunits” was our rather unadventurous term (“nu bodies” was also being used [2,3]). It seemed obvious that their study would be an exciting way to earn a degree. As Chris and I were considering possible dissertation topics, he presented me with a recently published PNAS article by Kriegstein and Hogness. It was titled “Mechanisms of DNA Replication in Drosophila Chromosomes…” [4]. In it was a series of electron micrographs, depicting Kleinschmidt spreads of DNA replication bubbles from the early cleavage stages of Drosophila embryos. Chris suggested that this would be an excellent system for preparing replicating chromatin fibers, to examine what was happening to nucleosomes during DNA synthesis. I thought it was a great idea. As it happened, so did Steve McKnight and Oscar Miller, Jr., and in the fall of 1977 they published their stunning pictures of chromatin caught in the act of replicating (Fig. 1) [5]. Although we were disappointed at having missed an opportunity to make a major contribution, the publication of this

☆ This article is part of a Special Issue entitled: Histone chaperones and Chromatin assembly. ⁎ Tel.: +1 617 552 3812; fax: +1 617 552 2011. E-mail address: [email protected]. 1874-9399/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.bbagrm.2011.07.005

article was not as personally unsettling as it might have been. By then I had already changed dissertation projects. It is not easy to convey the level of excitement felt in chromatin labs during the early days of the “subunit” hypothesis. And for someone like myself, a student in a laboratory that could be considered among the co-discoverers of the nucleosome, it was an especially heady time. Fundamental questions, such as whether or not nucleosomes were universally distributed, were still to be answered. (For a thorough and enlightening analysis of the multiple threads that led to the acceptance of the nucleosome theory, see K. E. van Holde's “Chromatin” [6]; Ada and Don Olins have also recounted their unique perspective of that period [7]). Eventually my zeal became tempered by the realization that electron microscopy was never going to be my strong suit. Nevertheless, I had become intrigued by the topic of chromatin assembly. Two research articles, one by Jackson et al. in 1975 [8], and another by Hildebrand and Walters in 1976 [9], helped me to formulate an approach to the question of histone deposition during DNA replication (both of those papers will be discussed below). Chris generously permitted me to shift the emphasis of my project to the biochemical investigation of nucleosome assembly, using the ciliate Tetrahymena pyriformis as a model system (this protozoan being the only dividing cell available to us at the time). The decision permanently influenced the path of my career. What follows is a survey of the questions and challenges surrounding investigations into chromatin assembly in the years following the discovery of nucleosomes. Given the scope of the topic, such a history cannot hope to be comprehensive. Instead, an attempt will be made to highlight the theories and controversies surrounding the analysis of chromatin replication and assembly, with particular attention paid to those earliest studies.

A.T. Annunziato / Biochimica et Biophysica Acta 1819 (2012) 196–210

Fig. 1. Electron micrograph of replicating chromatin (from Fig. 4 of McKnight, S.L. and Miller, O.L., Jr. (1977) Electron microscopic analysis of chromatin replication in the cellular blastoderm Drosophila melanogaster embryo Cell 12:795–804; used by permission).

2. Introduction The subjects of chromatin replication and nucleosome assembly encompass a broad range of interrelated topics. These include the structure of newly replicated chromatin, and the maturation of nascent chromatin following DNA synthesis; the segregation of parental histones at the replication fork; the sites of new histone deposition, and the de novo assembly of histone octamers; histone posttranslational modifications; and the identification of histone chaperones. In the following discussion these areas will be touched upon, focusing on the experiments that provided the foundation for subsequent investigations. 3. The structure of newly replicated chromatin It was obvious that the doubling of DNA during S phase would require the deposition of additional histones to rebuild functional chromosomes. A basic question concerning chromatin replication involved the timing of the assembly process with respect to DNA synthesis. Nucleases had become well established as probes of chromatin architecture, and it was natural that these should be used to dissect the structure of newly replicated chromatin. The radiolabeling of newly replicated DNA (typically with [ 3H]thymidine) provided a means of identifying the replicating regions, while standard pulse-chase protocols offered a way to examine chromatin assembly in vivo over time. Using this approach, Seale [10] showed that chromatin labeled for 1 min is preferentially susceptible to digestion by DNase-I, but after ~15 min recovers the nuclease resistance of mature bulk chromatin. Similar studies involving chromatin replicated in the presence of the protein synthesis inhibitor cycloheximide further demonstrated that protein (presumably histone) synthesis is required for chromatin maturation [11,12]. The finding that certain nucleases, including micrococcal nuclease (MNase), preferentially cut the linker DNA between nucleosomes [13–15], thereby releasing particles that could be separated on a sucrose gradient [15], provided another avenue for monitoring chromatin assembly in vivo. (As an aside, it also formed the basis of my own dissertation project [16]). Using these techniques, Hildebrand and Walters demonstrated that newly replicated chromatin is preferentially cleaved to mononucleosomes, relative to mature chromatin [9]. This observation was subsequently repeated in several laboratories [16–20]. Moreover, it was found that nearly normal nuclease resistance was regained after ~10 min of DNA synthesis [9], in good agreement with the earlier results of Seale, cited above [10]. In a series of elegant experiments using SV40 minichromosomes as a model system, DePamphilis, Wassarman and colleagues carefully mapped the positions of the first nucleosomes on either side of the advancing replication fork [21–23]. Their findings indicated that nucleosomes were present on both the forward and retrograde arms, within 125 to 300 bp of the site of DNA replication. Essentially identical results were later obtained by Sogo and colleagues, who used

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psoralen cross-linking and electron microscopy to examine nucleosome positions during DNA replication [24]. These data fitted in quite well with the earlier electron microscopic observations of McKnight and Miller [5], which had revealed fully nucleosomal chromatin on both arms of the replication fork, with no stretches of free DNA in evidence. In addition, distinct nucleosomal subtypes (including H1and HMG protein-containing species) were detectable on new DNA within 30 seconds of DNA replication [20]. A model was emerging that called for the placement of histones on new DNA as soon as replication was completed. There was, however, an apparent inconsistency in the accumulating data. On the one hand it had been shown that newly replicated chromatin was preferentially susceptible to nuclease digestion for an extended period. Yet in seeming contrast, newly synthesized DNA very rapidly becomes fully nucleosomal as judged by electron microscopy [5,25], which implied that histone transfer and/or deposition occurred almost immediately after passage of the fork. It was of course possible that new nucleosomes contained structural transitions that resulted in increased nuclease sensitivity [20,26–28]. But it was also clear that fully formed nucleosomes containing new DNA could be isolated using the very same nucleases [9,20,29]. A potential resolution of this apparent problem was obtained through experiments involving chromatin replicated in the absence of concurrent histone synthesis, either in the presence of a translation inhibitor such as cycloheximide, or using isolated nuclei (or viral DNA) in vitro. In these cases it was still possible to detect nucleosomes on newly replicated DNA, presumably containing histones recycled from parental chromatin; moreover, approximately half of the new DNA remained unassembled [11,12,17,25,30–32]. It was concluded that at least some nascent nucleosomes (very likely ~ 50%) contained old histones transferred to new DNA at the fork. Evidently this process was very rapid. The remaining fraction of new DNA must therefore be assembled de novo, in a process requiring new protein (i.e., histone) synthesis [27]. Nevertheless, questions persisted about the timing of chromatin maturation. As discussed above, in no case had long stretches of naked or partially assembled DNA been observed at native replication forks. Yet the complete maturation of newly replicated chromatin could take 15 min (or longer) in vivo, and even de novo nucleosome assembly did not take that long, based on both biochemical and electron microscopic evidence. Promising clues to an explanation for the gradual maturation process lay in the discovery that newly synthesized H4 is posttranslationally modified by acetylation [33,34], and the later observation that histone acetylation can partially disrupt chromatin higher order structures [35–37]. Interestingly, deacetylation of new H4 to steady state levels was found to take approximately 20–30 min [33,34,38], not unlike the nascent chromatin maturation period. To test whether acetylation was partially responsible for the increased nuclease accessibility of newly assembled chromatin, the nuclease resistance of chromatin replicated in the presence of the deacetylase inhibitor sodium butyrate was compared to that replicated under control conditions. It was found that preventing deacetylation did not impair nucleosome assembly per se (as judged by the generation of normal MNase ladders and resistance), but that approximately 50% of the initial sensitivity to DNase-I persisted as long as cells remained exposed to the inhibitor [39]. Upon removal of butyrate, new chromatin matured within the normal time frame [39]. Subsequent experiments indicated that acetylated nascent chromatin had an altered association with histone H1 [40,41], similar to that seen with active genes [37,42]. After the many experiments performed during this period, a model was eventually developed in which the assembly of newly replicated chromatin occurred in distinct stages, involving 1) transfer of pre-fork histones to new DNA; 2) de novo assembly of new nucleosomes through a process that required histone synthesis; and 3) re-formation of chromatin higher order structures following histone deacetylation and the stable deposition of H1. The

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details of these steps were only vaguely understood, and turned out to be far more complicated than most investigators imagined. These topics will be discussed in the following sections. 4. Segregation of old histones to new DNA 4.1. Stability of the H32H42 tetramer As a class, histones are very long-lived proteins [43,44]. It therefore follows that in dividing cells pre-existing histones must somehow be transferred to daughter chromosomes. The experiments cited above, in which the properties of chromatin replicated in the absence of concurrent histone synthesis were examined, indicated that old histones are placed on new DNA relatively quickly, perhaps immediately after DNA synthesis. The process whereby old histones are distributed to new DNA came to be termed histone (or nucleosome) segregation. Initial questions concerning histone segregation fell into two broad categories: 1) do histone complexes (either octamers or H3/H4 tetramers) dissociate or remain intact? and 2) are parental histones shunted to one or both arms of the replication fork? The former question has recently received new interest, following the observation that some chromatin assembly complexes contain H3/H4 dimers, not tetramers as long assumed [45–47]. This topic has previously been reviewed by the author [48], and will not be discussed at length here. For the purposes of this article, it will suffice to note that typical nucleosomes can be found on DNA replicated in the absence of de novo nucleosome assembly [25,27,30,31,49], and that chemical cross-linking and other studies lend support to a model in which the H3/H4 tetramer (at least) remains intact over multiple generations [24,50–53]. Thus, if H3/H4 tetramers do dissociate during DNA replication, they evidently have the ability to quickly reform. Recent evidence also weighs against permanent tetramer splitting being a routine phenomenon, although exceptions may occur [54,55]. 4.2. Distribution of old histones at the replication fork It had become generally accepted that parental histones are distributed to approximately half of newly replicated DNA, where they associate to comprise typical histone octamers. It was thus logical to ask whether there were any rules governing the partitioning of old histones at the fork. The first forays into this problem involved micrococcal nuclease digests of chromatin replicated in the presence of cycloheximide, or in isolated nuclei in vitro [17,30,31]. These experiments revealed nascent nucleosomal ladders up to heptamers at least, which was taken as evidence of cooperative segregation of old histones to new DNA. This condition was usually referred to as conservative histone segregation. How the old histones were transferred to new DNA was unknown, and the partial disassembly of octamers (followed by reassembly) remained a possibility. Interestingly, SV40 minichromosomes that had been reconstituted with cross-linked octamers were later shown to be completely replicated in vitro (though with a reduced replication rate) [56]. Moreover, crosslinked octamers were efficiently segregated to new DNA, demonstrating that disassembly was not obligatory [56]. However, transient disruption of H3/H4 tetramers in vivo still cannot be excluded, especially given the potential involvement of the chaperone Asf1 as a histone acceptor during DNA replication [57] (as Asf1 binds an H3/H4 dimer [46,58]). The inherent asymmetry at the replication fork prompted investigations into whether parental histones are preferentially distributed to the leading or lagging arm. Seidman et al. addressed this problem by labeling replicating SV40 minichromosomes with [ 3H] thymidine in the presence of the protein synthesis inhibitor cycloheximide, to prevent de novo nucleosome assembly [59]. Nucleosomes were prepared by nuclease digestion (to eliminate any unassembled DNA), and the radiolabeled chromatin DNA was then

hybridized to strand-separated SV40 restriction fragments. A strong bias of histone segregation toward the leading arm was indicated, which in SV40 is the transcription template strand. In additional experiments using cellular chromatin, a preference for the transcribed strand was also seen [59]. It therefore appeared that the problem of histone segregation had been solved. However, Wassarman, DePamphilis and colleagues, using the same approach as Seidman et al., found evidence for random (or dispersive) segregation of old histones to new DNA during SV40 replication [60]. Dispersive segregation was also observed in other studies of replicating SV40 chromatin [32,61]. By labeling new DNA with BrdU in the absence of concurrent protein synthesis, Roufa and Marchionni [62] and Kirov et al. [63] tested the segregation of parental histones in cellular chromatin. As in previous experiments, unassembled DNA was digested with micrococcal nuclease. Density-labeled nucleosomal DNA was then purified in alkaline CsSO4 gradients, and hybridized to the sense and antisense strands of either integrated SV40 [62], or a mouse α-globin gene [63]. In one case [62], conservative segregation was observed, but with the non-coding strand of SV40, in partial conflict with the results of Seidman et al. Surprisingly, Kirov et al. observed that histones segregated dispersively to both the coding and non-coding parental DNA strands of the globin gene [63]. Experiments in which the distribution of density- or radiolabeled histones was followed for several generations with respect to density-labeled DNA also supported the dispersive segregation of parental histones (i.e., no selective association with leading or lagging strands) [8,64–67]. Still, the observations of nucleosomal oligomers containing old histones on new DNA (cited above), and the tendency of adjacent histone octamers to segregate together [68], remained to be accounted for. Electron microscopy provided further evidence that adjacent histone octamers tend to segregate cooperatively. Cremisi et al. [25] examined polyoma virus that had replicated in the absence of protein synthesis, and detected viral minichromosomes with approximately half the number of nucleosomes; moreover, the nucleosomes tended to be clustered. In a similar fashion, Sogo et al. [24] used psoralencrosslinking and electron microscopy to study SV40 minichromosomes that had replicated in cycloheximide. When chromatin is treated with psoralen, the paired DNA strands in linkers are crosslinked together, and thus cannot be denatured. Nucleosomal DNA, in contrast, is unaffected. When psoralen cross-linked DNA is deproteinized and spread on a water surface, denatured bubbles mark the positions of nucleosomes. Using this technique, Sogo et al. observed that, following replication in cycloheximide, parental histones formed nucleosome clusters on the replicated viral products, separated by stretches of non-nucleosomal DNA. As expected, the number of nucleosomes per minichromosome was reduced, the result of replication without concomitant de novo assembly. When considering the replication of cellular (as opposed to viral) chromatin, it can be seen that strictly conservative segregation in the absence of protein synthesis will result in new DNA that is either fully nucleosomal or histone-free. With the possible exception of the regions immediately surrounding replication origins, there should be little juxtaposition of nucleosomes and unassembled DNA. Restriction enzymes such as HaeIII selectively cut in the linker DNA between nucleosomes, generating nucleosome ladders whenever the recognition sequences are appropriately spaced. In contrast, it is expected that unassembled DNA will be cleaved at virtually all available sites, yielding a smear after gel electrophoresis of purified DNA. In accord with this prediction, chromatin labeled with [ 3H]thymidine under control conditions yielded typical nucleosomal ladders in gels after HaeIII digestion (although of course considerably less DNA was released than would be with MNase), while chromatin labeled in the presence of cycloheximide yielded randomly sized fragments [69]. Importantly, redigestion of heterogeneously sized “cycloheximidechromatin” with micrococcal nuclease produced typical nucleosomes, suggesting that contiguous regions of nucleosomal and unassembled

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DNA were present after replication in the absence of de novo chromatin assembly, as expected for dispersive segregation. This interpretation was in accord with the earlier observation that chromatin replicated in cycloheximide, and released using BspRI, possessed a buoyant density between that of control chromatin and naked DNA [70]. To resolve the apparently conflicting data regarding conservative and dispersive segregation, current models call for the segregation of parental histone octamers (or H3/H4 tetramers) to both arms of the fork, but in groups. This would explain the detection of nucleosomal oligomers following replication in the absence of protein synthesis, while avoiding an assignment of histones to either the leading or lagging arms. It would also account for the observation of contiguous regions of nucleosomal and unassembled DNA. How this is regulated is still unknown, but it seems likely that histone chaperones such as Asf1 play an important part [57]. It should also be kept in mind that protocols that interfere with the maturation of newly replicated DNA (for example the ligation of Okazaki fragments) may skew the segregation of pre-existing histones to the leading arm of the fork [71]. The question of histone segregation has received relatively little attention in the past 20 years. Although histone complexes (tetramers or octamers) tend to segregate cooperatively, the size of the clusters is highly variable, at least as measured by electron microscopy [24] or superhelical density [60] of replicating SV40 chromatin. As the sequences of the replicating plasmids used in these studies are all identical, this variability may reflect a process that is fundamentally stochastic. With respect to cellular chromatin, it is currently unknown whether specific genomic regions apply localized strategies to distribute old histones at the replication fork. Nevertheless, there is overall no multi-generational association of H3/H4 tetramers with either DNA strand [67]. As noted above, electron microscopy of SV40 minichromosomes in the process of DNA replication under physiological (or nearly physiological) conditions has revealed intact nucleosomes immediately preceding the replication fork, and the re-association of histone octamers within 200–300 bp of the fork on both growing arms [24]. This is in good agreement with results obtained using exonucleases as probes of replicating SV40 [21,22]. The absence of an extended lag between DNA synthesis and histone deposition has prompted inquiries into the restoration of nucleosome positioning following replication. In an experiment in which replicating rDNA was examined in S. cerevisiae, it was found that precise nucleosome positioning was rapidly reestablished immediately behind the fork [72]. Thus, translationally positioned histone alignments need not be abolished during chromatin replication, at least at some loci. In a recent global analysis of the retention and positioning of pre-existing histones through multiple rounds of replication in yeast, it was found that parental H3 tends to accumulate most readily at the 5′ end of relatively long, moderately transcribed genes [73]. To explain their observations, the authors introduced a computational model incorporating three parameters: 1) gradual 3′ → 5′ “pass-back” of histones during transcription; 2) short-range spreading of parental histones during DNA replication (by ~ 1-2 nucleosome lengths); and 3) replication-independent histone turnover. It is perhaps worth noting that in both budding and fission yeast, the sole H3 histone is equivalent to the replacement variant H3.3 [74], making it difficult to distinguish between replication-coupled and replication-independent assembly pathways. It is therefore unclear how well this model can be applied to multicellular systems. Nevertheless, as most budding yeast nucleosomes are well positioned [75], the overall impression one gets is that of a system that can readily rebound following polymeraseinduced disruptions. If histones serve as carriers of epigenetic information, the posttranslational modifications of old histones that are segregated to new DNA, and the manner of histone and nonhistone distribution,

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become topics of considerable interest [reviewed in 76,77,339]. In this regard, chromatin immunoprecipitation (ChIP) experiments have indicated that parental histones are not obligatorily acetylated to allow for DNA synthesis (potentially preserving local differences in histone marks) [78], and that at least some histone modifications, as well as polycomb proteins, may persist during replication [47,78,340]. Certain modifications have also been detected in histones associated with Asf1-MCM protein complexes [57]. Notably, in C. elegans histone demethylation has been shown to facilitate DNA replication, by reversing the repressive effect of HPL-2 (an HP1 homolog) [79]. The targeted loss and timely regeneration of specific histone marks during replication is clearly an area deserving further exploration [80,81]. Nevertheless, as only one nascent DNA duplex can receive parental histones over a specific sequence, and because H3/H4 tetramers tend to segregate in clusters, the inheritance of chromatin epigenetic states is likely to operate regionally, rather than at the mononucleosome level [73]. 5. Posttranslational modifications of newly synthesized histones Before discussing studies aimed at uncovering the mechanisms of chromatin assembly, it will be helpful to present information on newly synthesized histones, with particular emphasis on their posttranslational modifications. This is of course an immense topic, which stretches back to some of the earliest investigations of histone metabolism. What follows is an overview of some of the groundbreaking experiments in this now burgeoning area, to aid in the subsequent treatment of the assembly process. 5.1. Acetylation of newly synthesized H4 In 1974 Gordon Dixon and colleagues first described the reversible acetylation of newly synthesized H4 during trout spermatogenesis [82]. The acetylation of new H4 is very likely the first histone modification to be unequivocally linked to a specific cellular process, though today we are not much closer to assigning a function to this modification than were investigators 35 years ago. The earliest experiments were nevertheless successful in establishing the universality of the acetylation of new H4, and defining the specific sites that are modified. Two major studies of new H4 acetylation followed the initial discoveries of Louie et al. (cited above). In 1975 Allfrey and colleagues showed that new H4 in dividing duck erythroblasts was “dimodified” [33]. Moreover, it was observed that the modifications were not permanent, but were lost with a half-life of approximately 20 min. Within a year, Jackson et al., working with rat HTC cells, presented evidence that the dimodification was in fact acetylation [34]. Moreover, loss of the dimodified forms followed the essentially same kinetics as seen in the duck system. Subsequent experiments extended the diacetylated isoform of new H4 to Tetrahymena [83,84], Xenopus oocytes [85,86], developing sea urchins [87], Drosophila [88], and Physarum (though new H4 in Physarum was seen to be mostly monoacetylated) [89]. The acetylation of new H4 (and in the overwhelming number of cases its diacetylation) was therefore revealed to be extraordinarily conserved. A remarkable aspect of these experiments is that they were for the most part performed before the widespread use of histone deacetylase inhibitors. The next challenge was to determine the actual sites of diacetylation. At the time it was known that four lysine residues in the H4 Nterminal tail domain (K5, K8, K12 and K16) could be reversibly acetylated (these sites correspond to K4, K7, K11 and K15 in Tetrahymena, due to a deletion of the typical arginine at position 3). By microsequencing newly synthesized H4 that had been radiolabeled with [ 3H]lysine, it was discovered that new H4 in Tetrahymena is acetylated selectively at K4 and K11 [84]. In later experiments it was found that newly synthesized H4 in Drosophila and human (HeLa)

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cells is diacetylated at K5 and K12, in direct correspondence to the sites used in Tetrahymena [90]. The K5/K12 diacetylation of new H4 is now seen as a hallmark of chromatin assembly. It may be one of the most highly conserved histone modification patterns (as opposed to an individual modified site) among eukaryotes. One approach to uncovering the role of the K5/K12 diacetylation of new H4 involves mutating lysines 5 and 12 of H4 to unacetylatable amino acids (usually arginine to maintain positive charge, or glutamine, glycine, etc. to provide uncharged residues) [reviewed in 91,92]. These experiments have typically been performed in yeast. In some cases the changes are accompanied by additional amino acid substitutions, or deletions of the H3 tail. While this extensive literature cannot be reviewed in full here, some generalizations can be made. For example, it has been shown that the K5,K12 → R substitution increases the length of S phase by ~50% [93], and that the triple K5,K8,K12 → G mutation is lethal when combined with deletion of the H3 amino-terminal domain [94]. In addition, yeast with combined mutations of acetylation sites (including the deposition sites) of H3 and H4 have sluggish growth rates that place them at a clear selective disadvantage [95,96]. Acetylation may also facilitate the import of H3/H4 dimers into the nucleus [96–98]. Recent studies using Physarum as a model system have provided strong evidence that the K5/K12 diacetylation of H4 is required for efficient nucleosome assembly in vivo [98]. It therefore seems reasonable to propose that the rigorous conservation of the diacetylation of nascent H4 reflects a universal role at some stage in the import/assembly process. 6. Modifications of newly synthesized H3 6.1. Histone H3 variants Human cells contain four different histone H3 variants, termed H3.1, H3.2, H3.3, H3.4, which differ only slightly in amino acid sequence (a fifth variant, CenH3 (CENP-A), is centromere-specific [99–103]). H3.4 appears to be expressed solely in testis [104]. Synthesis of the major H3 variant (H3.1) is replication-coupled (RC), as it rises sharply in S phase, and is linked to DNA replication [105,106]. Synthesis of H3.2 is also RC, but it is distinguished from H3.1 by a single amino acid change at position 96 (cysteine in H3.1, serine in H3.2) [103,107]. Synthesis of the replication-independent (RI) variant H3.3 is not coupled to DNA replication, but continues at a basal level throughout the cell cycle [105,106]. In mammals, H3.3 differs by only 4–5 amino acids from H3.1 [107,108]. H3.3 can be deposited in the absence of DNA synthesis, often in association with transcription [109–116]. As will be detailed below, replication-coupled (RC) deposition of H3.1/H4 dimers onto new DNA is mediated by the assembly factor CAF-1; the H3/H4-escort protein Asf1 appears to assist CAF-1 in this process [91,117–121]. The RI deposition of H3.3/H4 into chromatin is carried out by HIRA [45], in agreement with evidence that HIRA [109] and the yeast HIR complex [122] can effect RI nucleosome assembly. It has recently been shown that DAXX (a death-domain-associated protein) binds to histone H3.3, and can serve as a HIRA-independent chaperone during RI chromatin assembly in vivo, in conjunction with the remodeling factor ATRX [123–125]. Thus there is evidence that RC and RI histone deposition pathways utilize several different assembly factors, including CAF-1, Asf1, HIRA and DAXX [119,126,127]. The centromere-specific H3 variant CenH3 (CENP-A/CID/Cse4/Cnp1) is deposited by the orthologous factors Scm3 (in yeast) and HJURP (in human cells) [reviewed in 128–130]. 6.2. Posttranslational modifications of newly synthesized H3 The modification state of newly synthesized H3 has turned out to be considerably more variable than that of H4. In Tetrahymena, K9 and K14 are the predominant acetylated sites, while in Drosophila K14 and

K23 are preferred [90]. In the budding yeast S. cerevisiae, K9 is the most preferred residue, but significant acetylation is also detected at K14 and K27 [131]. Thus far the reason for this variability remains unexplained. Early reports of the modification state of nascent mammalian H3 indicated that it was unmodified, even when synthesized in the presence of the HDAC inhibitor sodium butyrate [38,132]. When the acetylation state of newly synthesized human H3 was examined by microsequencing, it showed relatively little acetylation at any lysine in the N-terminal tail domain [90]. In fact, ≥75% of each acetylatable amino-tail lysine of all nascent human H3 appeared unacetylated. This was true for both H3.1 and H3.3, although new H3.3 was the more modified of the two [47,90]. Strikingly similar results were obtained when the modifications of predeposition (cytosolic) H3.1 and H3.3 were later examined (~ 80% of H3.1, and ~ 65% of H3.3 were unmodified) [133]. In the same report it was also found that 35% of predeposition H3.3 was monomethylated at K9 [133]. A low level of acetylation of the H3 tail has also been observed in human Asf1histone complexes [134]. As a consequence, there appears to be no specific “deposition” acetylation pattern for new H3 in human cells, at least as far as the N-terminal domain is concerned. Another possibility for the deposition-related modification of H3 involves the lysine residue at position 56. Work from several laboratories has established that in S. cerevisiae the acetylation of H3 at K56 facilitates nucleosome assembly and DNA repair, and promotes the association of (new) H3 with the assembly factor CAF-1 [135–139]. H3K56 → R mutants exhibit slow growth and heightened sensitivity to DNA damaging agents [135–137,140], while the K56 → E mutation permits CAF-1 binding [141]. Notably, acetylation at K56 does not act redundantly with the acetylation of the tail domains of H3 or H4 [136]. H3K56 acetylation is a conserved modification, found abundantly in Drosophila and Tetrahymena, and to a lesser degree in cultured human cells [142–145]. The function of H3K56 acetylation in metazoan systems has been controversial [reviewed in 146]. For example, it has been reported that acetylation of H3 at K56 in human cells takes place in S phase, requires Asf1, and is necessary for genomic stability [147]. In this study, S-phase delay and increased phosphorylation of H2A.X was observed when H3.1K56 → R and H3.1K56 → Q were overexpressed, while WT-H3.1 was repressed using shRNA (thereby flooding the cells with mutant H3.1) [147]. Whether the delay was due to a defect in chromatin assembly or to DNA damage was unclear. It has also been shown that H3K56 acetylation increases in human cells in response to gamma irradiation [143]. In contrast, it has been observed that the level of H3K56 acetylation changes minimally during the cell cycle, and decreases in response to DNA damage [145]. Whether these differences are due to the antibodies used for analysis, the methods applied to induce DNA damage, or to other factors, remains to be determined. Knockdown of CAF-1 blocks the incorporation of acetylated H3K56 into chromatin in Drosophila S2 cells, and it has been proposed that acetylation of H3K56 facilitates chromatin assembly in metazoan systems [143]. However, there is as yet no evidence that new H3 is generally acetylated at K56 in mammalian cells. In HeLa cells only 1– 2% of Asf1-associated H3 is acetylated at K56 [134]. This may be due in part to the association of Asf1 with (unacetylated) parental histones during chromatin replication [57]. 7. Chromatin assembly in vivo The finding (discussed above) that chromatin replicated in the absence of protein synthesis remains partially histone-free led to the logical conclusion that ~ 50% of newly replicated DNA is assembled with newly synthesized histones. Based on early models of conservative histone segregation, it seemed that events at the replication fork were relatively straightforward: one side of the fork (e.g., the

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leading arm) would continuously receive old octamers, while the other side would be assembled de novo with entirely new histones. However, as with the earlier studies on histone segregation, experiments on the mode and sites of nucleosome assembly yielded an array of conflicting results and interpretations, some of which still remain unresolved. Although the allied questions of where and how histones are deposited are inevitably linked, it will be convenient to discuss these topics separately. 7.1. Assembly of newly replicated DNA Initially, the most fundamental question asked was “Are new histones deposited onto new DNA?” To address this it would be critical to somehow distinguish between newly replicated and nonreplicating chromatin regions. This was accomplished in several ways, including: 1) density labeling of new DNA using “heavy” thymidine analogs; 2) exploiting inherent density differences between old and new chromatin; 3) using the characteristic properties of newly replicated chromatin, such as increased sensitivity to nucleases and altered solubility properties; and 4) separation of replicating and nonreplicating SV40 minichromosomes, based on their distinctive sedimentation rates. In each of these approaches, new histones were radiolabeled with [ 3H]lysine, [ 3H]arginine, or both. In experiments in which new DNA was density-labeled with heavy analogs such as BrdU, new and old chromatin was separated in CsCl gradients [8,64,148–150], in a manner reminiscent of the experiments of Meselson and Stahl [151]. In the case of chromatin, however, this required chemical cross-linking (typically with formaldehyde) to prevent dissociation of histones from DNA in the high ionic environment of the gradients. In some cases chromatin was first isolated (e.g., by MNase digestion or shearing), then fixed. In other cases whole cells were fixed, and chromatin isolated by sonication, much as the case in current ChIP experiments [64,152]. Rather surprisingly, the earliest experiments of this type indicated that new histones were deposited mostly, if not exclusively, on non-replicating DNA [8,148,149]. This would obviously require a major rearrangement of histones throughout S phase. Subsequent studies, in which the histones of fractionated chromatin were electrophoretically resolved, presented a more nuanced scenario. New H3 and H4 were selectively targeted to new DNA; new H2A and H2B were less likely to be assembled on new DNA; new H1 was randomly deposited [64]. Similar results were obtained when new and old chromatin were separated by a naturally occurring density difference after fixation [152]. It was possible that the manner of chromatin isolation, including formaldehyde fixation, was somehow randomizing histone distribution. To overcome this objection, several researchers used metrizamide (an iodinated aminoglycoside) or D2O/sucrose gradients to resolve density-substituted nascent chromatin [65,153–156]. In all of these cases it was found that radiolabeled new proteins separated with both new and old nucleosomes, although in one case [65] up to 74% of the [ 3H]arginine label fractionated with newly replicated DNA (note that the use of [ 3H]arginine would weight the labeling toward H3/H4). When individual histones were examined in gels, it was again found that only new H3 and H4 were selectively deposited onto new DNA, in agreement with previous studies using fixed chromatin [153,156]. The replicating forms of SV40 minichromosomes sediment in sucrose gradients more rapidly than the mature nonreplicating forms. When this difference in S value was used to monitor the deposition of new histones during viral replication, the selective deposition of new H3/H4 onto replicating DNA was observed; new H2A/H2B was found on new and mature minichromosomes, and in some experiments was skewed toward the nonreplicating form [64,157,158]. In one case this was interpreted as representing the delayed deposition of new H2A/H2B following viral maturation [158]. Yet this was difficult to

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reconcile with the observed rapidity of nucleosome assembly [e.g., 5,24,25,159]. As discussed in Section 3, newly replicated chromatin is more sensitive to digestion with MNase than is bulk chromatin [9,19,20,160]. It also exhibits a shorter repeat length, and altered solubility properties [17,20,27,154,161]. When these distinguishing characteristics were applied to the analysis of histone deposition, it was again found that new H3/H4 is preferentially assembled onto new DNA, but that new H2A/H2B, and especially H1, become associated with bulk chromatin [18,161,162]. The preferential digestibility of newly replicated chromatin by micrococcal nuclease permits the partial separation of nascent mononucleosomes from mature chromatin in a sucrose gradient [9,16,18,160]. In one case it was found that new H3/H4 co-sedimented with the rapidly digested nascent mononucleosomes, while new H2A, H2B and H1 sedimented with more nuclease-resistant bulk chromatin [18]. In this case the results were interpreted to indicate sequential assembly of all new core histones onto newly replicated DNA, with the deposition of H2A and H2B coinciding with the loss of heightened nuclease susceptibility. Yet an equally valid interpretation could be that the histone pairs are deposited independently, onto both replicating (H3/H4) and nonreplicating (H2A/H2B) chromatin regions. From this it can be seen that a pressing question becomes whether the increased nuclease sensitivity of newly replicated chromatin persists after the deposition of H2A and H2B. Notably, nascent chromatin retains this immature characteristic for 10 min or longer, while de novo nucleosome assembly proceeds considerably more rapidly (as judged by the absence of extended regions of free DNA at the site of native replication forks [5,24,25]). To this point must be added the results from density-labeling studies (cited above), which found evidence for considerable deposition of new H2A and H2B on nonreplicating DNA. The potential pitfalls in interpreting experiments on histone deposition are highlighted in these studies. However, there is another facet to the problem of histone deposition, and that involves the analysis of octamers alone, without considering the sites of deposition. This work will be discussed in the following section. 7.2. Histone octamer assembly The assembly of histone octamers entirely from newly synthesized histones has been called conservative assembly (it can be seen that the term conservative has become quite a workhorse). It has usually been studied by labeling new histones with dense and radioactive amino acids, and separating isolated octamers in density gradients. To analyze octamers in the absence of their associated DNA, it is necessary to cross-link the histones together. This is typically done using either formaldehyde (often at pH 9.0) or Lomant's reagent (Dithiobis[succinimidyl propionate], or DSP). Much of the work on octamer assembly has been performed by Leffak and colleagues, using the density-labeling techniques described above (and DSP as the crosslinking reagent) [52,68,163,164]. In all of these experiments the results have been consistent with the conservative assembly of new octamers entirely from new histones. The one exception to this involved octamers isolated from a magnesium-soluble fraction of chromatin that was enriched in transcriptionally active sequences [165]. In this case hybrid density octamers were detected, suggesting the mixing of new and old histones (although the proteins involved were not examined). Moreover, pulse-chase experiments provided evidence for the longterm stability of conservatively assembled octamers [52]. It is now clear that if octamers are conservatively assembled, new and old histones must be deposited together, and not independently on both new and old DNA. In light of this, another look at the assembly question was taken by Jackson and colleagues. In these experiments new histones were labeled with dense and radioactive amino acids, and whole nuclei were cross-linked with formaldehyde (typically at pH 9.0, to help release histone tails from DNA) [53,166–168]. Cross-

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linked octamers were then extracted with 0.2 M sulfuric acid, and subjected to density gradient centrifugation. The positions of newly synthesized histones were determined by gel electrophoresis and fluorography, after reversal of the cross-links. In contrast to the results obtained by Leffak and colleagues, it was observed that new H2A/H2B were assembled into octamers distinct from those receiving new H3/H4. To account for this discrepancy, Jackson proposed that Lomant's reagent could yield spurious results due to (dense) nonhistone proteins cosedimenting with histone complexes [169]. In counterargument, it was claimed that the procedures used by Jackson and colleagues caused histone rearrangement [164]. Whether the differences seen by these investigators are due to differing techniques, different chromatin fractions analyzed, or other factors, remains unresolved. However, when unfixed nucleosomes were studied, the formation of hybrid octamers containing both new and old histones was observed [170]. It must be stressed that there is much that investigators into histone deposition and octamer assembly agree upon. First, it is now generally accepted that new H3.1/H4 dimers are selectively deposited onto newly replicated DNA. It is also now accepted that linker histones are deposited more randomly with respect to new core histones [171]. In addition, as noted above, even proponents of strictly conservative assembly have reported hybrid octamers on a subset of nucleosomes [165]. Thus in the end what separates one camp from another is the behavior of a fraction of H2A/H2B dimers during replication-coupled chromatin assembly (and to some degree whether the replication-independent deposition of H3.3/H4 dimers always requires new H2A/H2B). While not all aspects of this topic can be addressed here, a few points may bear analysis. First, there is relatively little known about the normal deposition of H2A and H2B histones during DNA replication, although it appears that the chaperone Nap1 may be involved [172,173]. Also, there is a considerable literature on the greater lability of H2A/H2B relative to H3/H4 in vivo [e.g., 53,174,175], and on H2A/H2B exchange being facilitated by transcription and the acetylation of H4 [47,176–178] [reviewed in 179]. Although the deposition of H3/H4 onto new DNA is mediated by the association of CAF-1 with the replication processivity factor PCNA [180], to the author's knowledge there is as yet no known mechanism that targets new H2A/H2B dimers to nascent chromatin. It therefore follows that the association of new H2A/H2B with new H3/H4 (which are imported as a heterodimer [45,47,181]), will be governed by the pool size of exchanging H2A/H2B dimers. This, in turn, will depend on the functional state of the nucleus, including the level of transcription, and the participation of other factors that drive H2A/H2B exchange [182,183]. 8. Chromatin assembly in vitro; chromatin assembly factors The processes of chromatin assembly were turning out to be more complicated than initially envisaged. While investigations continued into the sites and mode of histone deposition and octamer assembly in vivo, parallel lines of research were being explored that relied on more simplified systems. The most straightforward method of assembling chromatin in vitro would be to simply add histones to DNA. However, the great affinity of histones for DNA results in the production of nonspecific aggregates, when the purified components are mixed under physiological ionic conditions. Initial studies of chromatin reconstitution therefore involved introducing histones to DNA in the presence of very high concentrations of NaCl and urea, followed by dialysis to allow nucleosomes to form gradually as the salt and urea are removed [e.g., 184–187]. Living cells must of course employ different assembly methods, and so it was reasonable to search for chaperones that could assemble chromatin in vitro under conditions approaching those within the nucleus. These experiments typically involved extracts prepared from oocytes, eggs or somatic cells, and had as aims the identification of histone escort factors, and the steps in

nucleosome assembly. The in vitro systems themselves could be subdivided into those that support DNA replication and those that do not. In the following sections it will be convenient to treat the various assembly factors independently, though there is some inevitable overlap. The subject of histone chaperones has been widely reviewed [e.g., 119,129,188,189]. The following discussion will therefore primarily touch upon early experiments. 8.1. Nucleoplasmin and N1/N2 Virtually all studies of chromatin assembly in vitro employ two complementary assays to demonstrate the presence of nucleosomes: micrococcal nuclease digestion (with the generation of either mononucleosomes or a nucleosomal ladder), and the introduction of supercoils into closed circular DNA. Electron microscopy is also sometimes used. The supercoiling assay was developed in 1975 by Germond et al. [190], who had observed the retention of one negative supercoil for every nucleosome present on SV40 minichromosomes after topoisomerase treatment. (Why two left-handed turns should yield only one negative supercoil has been called the linking number paradox [reviewed in 191]). To assemble nucleosomes in the absence of DNA replication (and hence radiolabeled nascent DNA) requires an abundant histone pool. For this researchers initially turned to the oocytes or eggs of several vertebrate and invertebrate species, including urodele amphibians [192], and the fruit fly Drosophila [193,194]. Possessing a sufficient histone pool to assemble ~20,000 nuclei [195], the oocytes and eggs of the African clawed frog Xenopus laevis became highly utilized systems for studying chromatin assembly in vitro. Notably, the stored histones are posttranslationally modified, and in particular H4 in the germinal vesicle is diacetylated [85]. Although extracts prepared from eggs that are fertilized or “activated” (e.g., by pricking) support DNA replication, many of the early studies relied on unstimulated oocytes. In a series of groundbreaking experiments, Laskey and coworkers used extracts prepared from Xenopus oocytes and eggs to assemble of nucleosomes on purified DNA [196–198]. It had thus been demonstrated that chromatin could be assembled in a test tube under physiological conditions. There were two interrelated questions that stemmed from these experiments: 1) what was protecting Xenopus chromatin from the potentially deleterious effects of the very large histone pool? and 2) what factor(s) could assemble chromatin in vitro? The acidic protein nucleoplasmin, the most abundant protein in Xenopus oocytes and eggs, appeared to provide an answer to both questions [198]. Nucleoplasmin was shown to effect chromatin assembly in vitro [196,199,200], and given its abundance, could possibly be acting as a storage protein for the oocyte histone pool. Still, it was not certain that nucleoplasmin was naturally found associated with all four core histones, and there was some uncertainty concerning the degree to which nucleoplasmin was a histone escort protein in somatic cells [201–203]. It was later discovered that the proteins N1 and N2, originally described by Bonner [204] and Gurdon and colleagues [205], were associated with H3 and H4 in oocytes, and that nucleoplasmin was the storage protein for H2A and H2B [206,207]. Both complexes were also shown to participate in nucleosome assembly [208,209]. It is now known that nucleoplasmin is a member of the nucleophosmin/nucleoplasmin (NPM) protein family. Nucleoplasmin itself is exclusive to eggs and oocytes, where (in addition to histone storage) it is involved in the disassembly and remodeling of sperm chromatin after fertilization [reviewed in 210]. Other NPM family members include nucleophosmin/NPM1 (B23) and NPM3 (NO29), which are involved to different degrees in ribosome assembly, histone chaperoning and chromatin decondensation [210]. N1/N2, on the other hand, is related to the NASP (nuclear autoantigenic sperm protein) family of proteins, with homologues identified in yeast

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[211,212], C. elegans [213], and mammalian cells [214–217]. In mammalian systems, NASP is often considered an H1-specific chaperone [215–217]. However, there is evidence that mammalian NASP may also bind H3/H4, in line with the specificity of N1/N2 in Xenopus oocytes [218,219]. The ability of the Xenopus oocytes and eggs to assemble chromatin came to be widely exploited [197,207,209,220,221], which fostered a succession of experiments into the assembly of transcriptionally active chromatin structures [reviewed in 222]. As noted above, activated Xenopus eggs or extracts will support DNA replication [223–228]. Moreover, in this system exogenous single-stranded DNA is both replicated and assembled into nucleosomes [208]. Taking advantage of this, Almouzni and Méchali showed that DNA synthesis greatly facilitated the chromatin assembly process, yielding regularly spaced nucleosomes on the replicated template [229]. In subsequent work it was further demonstrated that nucleosome assembly proceeded in a stepwise fashion, with H2A/H2B deposited onto an intermediate particle that likely contained histones H3 and H4 [230]. This supported the findings of Dilworth et al. [208], who had observed that two separate histone-containing complexes were required for efficient chromatin assembly in Xenopus oocytes, and further helped to establish a fundamental understanding of how nucleosomes are assembled in vivo. 8.2. Nap1 In contrast to the manner in which nucleoplasmin and N1/N2 were discovered, the chromatin assembly factor Nap1 was first observed in somatic (cultured mammalian) cells. Somewhat ironically, the trail to Nap1 started with the study of histone synthesis in the absence of DNA replication. Although histone synthesis decreases when DNA replication is blocked, even replication-dependent variants continue to be synthesized at reduced rates for two or more hours [231–234]. Under these conditions, the nascent H2A/H2B and H3/H4 pairs have different behaviors. A significant fraction of newly synthesized H2A and H2B continues to be stably deposited into chromatin, through a process that involves histone exchange [47,155,177,232–234]. In contrast, new H3 and H4 preferentially accumulate in a soluble pool, which can be detected in cytosolic extracts [47,177,233,235,236]. (However, not all investigators find evidence for the nonconservative deposition of new histones during hydroxyurea treatment [164].) In 1980 Senshu and Yamada observed that a cytoplasmic extract from cells pre-treated with the replication inhibitor hydroxyurea could assemble nucleosome-like particles, and that extracts from control cells had a similar capability when exogenous histones were provided [237]. In subsequent studies, proteins in cytosolic extracts from untreated cells were fractionated chromatographically, and tested for their ability to supercoil DNA when supplemented with free histones. Although the purified assembly preparation contained three different polypeptides, it was found that the active factor had a molecular weight of 59-kDa, and that it could form a 10S complex with all four core histones [238]. Notably, an additional ~ 6S complex was also observed, containing the 59-kDa protein and H2A and H2B alone [237]. The factor was characterized as a histone-binding chaperone with nucleosome assembly capability, and named AP-1 [239,240]. To avoid confusion with the AP-1 transcription factor, it was subsequently renamed Nap1 (nucleosome assembly protein 1), in a study in which the yeast homolog was described [241]. It was also determined that Nap1 had a greater affinity for H2A/H2B than for H3/H4 [240]. Many negatively charged macromolecules can assemble chromatin in vitro, including RNA [242] and polyglutamic acid [243]. The key question regarding Nap1 was whether it was a universal chromatin assembly factor (or histone binding protein) in living cells. The biology of Nap1 has turned out to be surprisingly complex, and explorations of its distribution and properties have yielded an almost

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bewildering array of homologues, family members, functions and binding partners [reviewed in 244,245]. Nevertheless, the role of Nap1 as a histone chaperone has been verified. Early indications (in addition to those cited above) that Nap1 functioned as a histone escort came from investigations of chromatin assembly using factors purified from Drosophila embryo extracts [194]. A 56-kDa protein was isolated that assisted in the assembly of regularly spaced nucleosomes on plasmid DNA. Initially termed d-CAF4, the protein was later identified as the Drosophila Nap1 homolog [172]. Drosophila Nap1 was demonstrated to bind histones H2A/H2B in vivo, and to shuttle from the cytoplasm to the nucleus during S phase [172]. A human homolog of Nap1, Nap2, has a similar cell-cycle dependent behavior [246]. Further evidence that Nap1 associates specifically with H2A/H2B came from the immunoprecipitation of cytosolic extracts from human (HeLa) cells [173], lending further support to a model in which H2A/H2B were deposited onto DNA independently from H3/H4. The association of Nap1 with H2A in vivo was also confirmed through twohybrid screens in the yeast Saccharomyces cerevisiae [247]. Nap1 can facilitate in vitro transcription by binding and transferring H2A/H2B dimers from nucleosomal templates [248–250]; [reviewed in 245,251]. Also, in accord with its action as a histone escort during chromatin assembly, Nap1 has been found in a complex together with H2A/H2B and the karyopherin (importin) Kap114, the factor responsible for transporting H2A/H2B into the nucleus [252,253]. In Drosophila, Nap1 can interact with heterochromatin protein HP2 and the chromatin remodeling complex NURF, suggesting that Nap1 may be involved in the establishment of transcriptional silencing [254]. Additional evidence for this is that the heterozygous knockout of Nap1 causes loss of position effect variegation in Drosophila melanogaster [255]; notably, the homozygous knockout causes either embryonic lethality or poorly viable adult flies [255]. NAP-1 also appears to serve as a chaperone for linker histones in vivo and in vitro, at least in some circumstances [256–259]. 8.3. CAF-1 The discovery of CAF-1 was a direct consequence of the search for a factor that could selectively assemble nucleosomes on newly replicated DNA. The approach was based on in vitro DNA replication protocols developed from human cells, that could replicate SV40 origin-containing plasmids in the presence of T antigen [260–262]. These replicating systems relied on nuclear and cytosolic extracts, and in the presence of both introduced negative supercoils into the replicated products [260–262]. In 1986 Bruce Stillman demonstrated that in such systems nucleosomes were assembled preferentially onto newly replicated DNA, but not on nonreplicated plasmids [263]. A strategy based on the supercoiling assay was developed to identify the assembly factor activity, which was first described by Smith and Stillman [264,265]. The replication-coupled assembly reaction relies on the small pool of H3/H4 histones in the cytoplasmic extract [264,266], which includes newly synthesized histones in transit to the nucleus [234,266,267]. The isolated assembly factor was named CAF-1 (chromatin assembly factor 1) [264]. CAF-1 assembles new DNA in a step-wise fashion, specifically depositing H3/H4. H2A/H2B are deposited subsequently in a separate step, and need not be derived from the cytoplasmic extract: purified nuclear H2A/H2B work equally well [266]. CAF-1 is a complex of three proteins: p150, p60 and p48 [268]. The p48 subunit is identical to the Rb-associated protein RbAP48, and can bind H3/H4 in the absence of the other two subunits [269]. The in vivo association of H3 and H4 in a somatic predeposition complex had first been demonstrated through the use of anti-acetylated H4 antibodies, to immunoprecipitate new H4 from cytosolic extracts [177]. It was later established that H3/H4, but not H2A/H2B, interact directly with the CAF-1 complex [268,269]. However, the specific acetylation state of newly synthesized H4 does not appear to be a factor in CAF-1

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binding, as H3 and H4 lacking their N-terminal tail domains can still be assembled into nucleosomes by CAF-1 [270]. CAF-1 activities have been detected in a wide range of organisms, including Drosophila [194,271], Xenopus [271,272], and budding yeast [273]. Deletion of the three yeast genes encoding the CAF-1 subunits (CAC1, CAC2 and CAC3) does not affect cell viability or prevent chromatin assembly [273], but does cause increased sensitivity to UV light, a reduction of telomeric silencing, and aberrant activation of the silent mating loci [273–277]. CAF-1 is also required for chromatin assembly during nucleotide excision repair in multicellular organisms [272,278,279]. Additional links between CAF-1 and transcriptional silencing lie in the findings that the largest subunit of CAF-1 (p150) can interact directly with the heterochromatin protein HP1α [280], as well as with the methyl-CpG-binding protein MBD1 [281]. These associations are consistent with the interaction of HP2 and the assembly factor Nap1 in Drosophila (discussed above [282]), and suggest a general theme for the escort of silencing factors to epigenetically modified histones and DNA. In line with this, the staging of DNA synthesis, chromatin assembly, histone acetylation/ deacetylation, and HP1 deposition are temporally coordinated [283]. CAF-1 is capable of selectively targeting nascent H3/H4 histones to newly replicated (or newly repaired) DNA. The basis for this specificity lies in the ability of the p150 subunit to recognize and bind PCNA, the DNA polymerase processivity clamp [180,278,284]. CAF-1 is also specific for the H3.1/H4 heterodimer [45], which is found in both nuclear and cytosolic extracts [45,47]. The CAF-1-H3.1/H4 complex, termed CAC (i.e., Chromatin Assembly Complex), is able to promote chromatin assembly in H3/H4-depleted cytosolic extracts [269]. Notably, the histones in CAC are posttranslationally modified, but in patterns that are somewhat unexpected. Most H3 (~60%) is unmodified; the remaining fraction is “monomodified” (likely by acetylation), as judged by its electrophoretic mobility in acetic acidurea gels [269]. Approximately 65% of CAC-associated H4 is acetylated at lysines 5, 8, and 12, in a mixture of mono-, di-, and trace triacetylated isoforms; up to one third of CAC-H4 shows no modifications in gels [269]. This PTM pattern is clearly different from the K5/K12 diacetylation of newly synthesized H4, and may be a consequence of nuclear post-deposition events (CAC is isolated from nuclear extracts). If this is so, heterogeneity in the modifications of CAC-associated histones may reflect the assembly of functionally distinct chromatin regions. In contrast to the case in budding yeast, CAF-1 is essential for proper replication-coupled chromatin assembly and/or cell cycle progression in human and Drosophila cells [285– 289]. 8.4. Asf1 The histone chaperone Asf1 (anti-silencing function 1) was initially described as a derepressor of transcriptionally silent chromatin when overexpressed in yeast [290,291]. Tyler et al. [292] later identified Asf1 as member the RCAF (replication-coupling assembly factor) complex, through its ability to complement CAF-1dependent chromatin assembly in depleted Drosophila embryo extracts. The RCAF complex comprises Asf1 and histones H3 and H4. Notably, H3 in RCAF is acetylated at K14 (but not K4 and K9), and H4 is diacetylated at K5 and K12 [292]. These sites correspond to those acetylated in newly synthesized H3 and H4 in Drosophila cultured cells [90]. This suggests that Drosophila Asf1 (dAsf1) can assist CAF-1 in vivo by providing an additional source of nascent H3/H4 histones. Asf1 is also termed CIA [reviewed in 189,293]. Loss of Asf1 function is lethal in the fission yeast S. pombe [294], and causes growth defects and sensitivity to DNA damaging agents in S. cerevisiae [290,292,295,296]. Newly synthesized H3 and H4 are associated with each other prior to chromatin assembly [177,268,269]. Based largely on the behavior of purified histones in solution [e.g., 297–299], it had been assumed that

H3 and H4 formed a tetramer before being deposited onto DNA. The observation that nuclear and cytosolic H3/H4 assembly complexes contain heterotypic dimers was therefore rather surprising [45,47], and engendered new questions concerning the manner in which nucleosomes are assembled [reviewed in 48,300]. As with the assembly factors CAF-1 and HIRA [45], Asf1 also binds an H3/H4 heterodimer [46,58,301]. Asf1 binds directly to the p60 subunit of CAF-1 [121,302,303], and also interacts with the C-terminal region of histone H3 [58,301,304,305]. Moreover, Asf1 stimulates the acetylation of H3K56 by the Rtt109 histone acetyltransferase (as does the histone chaperone Vps75) [135,140,306–310]. In Drosophila and human cells, Asf1 facilitates H3K56 acetylation by p300/CBP [143]. An important link between Asf1 and DNA replication lies in its interaction with Replication Factor C, which loads the DNA polymerase clamp PCNA onto DNA [311]. Significantly, Asf1 can interact with MCM helicases [57], and localizes to replication forks [312], where it may help to displace pre-existing H3/H4 dimers during replication. In addition, siRNA-mediated knockdown of Asf1 delays S phase progression [313] and causes inefficient chromatin assembly in human and chicken cells [303,314]. It is now clear that Asf1 (CIA) is a critical histone chaperone during nucleosome assembly and DNA repair, in a wide range of systems and cell types. Its deletion is lethal in cultured vertebrate (chicken) cells [303].

9. Replication-independent chromatin assembly and histone exchange 9.1. H3.3 and HIRA Hir1 was initially identified as an 88-kDa protein involved in transcriptional repression of the H2A and H2B genes in S. cerevisiae [315–317]. The human gene had originally been called TUPLE1, and was first described as a potential transcriptional regulator within the DiGeorge syndrome critical region (DGCR) of chromosome 22 [318]. The protein was later renamed HIRA, based on sequence homology with yeast Hir1p and Hir2p [319]. In 1998 Kaufman et al. demonstrated that deletion of HIR1 had a synergistic and negative effect on telomeric silencing, when combined with a deletion of CAC2 (which encodes the p60 subunit of CAF-1) [320]. HIRA was also shown to interact with core histones [321,322]. The Xenopus homolog is able to assemble nucleosomes on nonreplicating DNA [109], as can the HIR complex of S. cerevisiae in conjunction with Asf1 [122]. In Drosophila, HIRA is required for chromatin assembly during decondensation of the male pronucleus following fertilization [323,324]. It has also been reported that HIRA is essential for normal murine embryogenesis [325]. It is now known that CAF-1 and HIRA mediate different chromatin assembly pathways, on replicating and nonreplicating DNA, respectively: H3.1 is deposited onto replicating DNA by CAF-1, and H3.3 is deposited in a replication-independent fashion by HIRA [45]. The H3.1 and H3.3 complexes have been separately isolated, and Asf1 is found in both complexes. The association of HIRA (or Hir family proteins) with Asf1/CIA has been repeatedly confirmed both biochemically as well as genetically [109,326–330]. However, the presence of Asf1 proteins in CAF-1 and HIRA complexes poses somewhat of a conundrum, as these complexes (purified using epitope-tagged H3.1 or H3.3) have been shown to chaperone H3/H4 dimers, not tetramers [45]. Given that the epitope-tagged species represented only ~ 5–10% of total H3, it is highly unlikely that the only complexes purified were those in which both Asf1 and HIRA (or CAF-1) were simultaneously associated with different tagged-H3 dimers. It therefore appears that only one of the chaperones actually binds H3/H4 in the isolated complexes. An alternative possibility is that Asf1 and HIRA physically interact with the same H3/H4 dimer in a four-protein complex, as HIRA and Asf1 interact with different surfaces of H3 [189,328,329].

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There is also evidence that DAXX (a death-domain-associated protein) can act as an H3.3 chaperone during replication-independent chromatin assembly, in conjunction with the chromatin remodeling factor ATRX [123–125]. Interestingly, the DAXX/ATRX pathway is involved in the assembly of specific chromosomal and nuclear regions, including PML bodies and telomeres [123–125]. In Drosophila embryos, eliminating CHD1 (an ATP-dependent chromatin remodeling factor) prevents the deposition of H3.3 into the male pronucleus [331]. This is consistent with models of nucleosome assembly that involve ATP-dependent processes [332]. 9.2. H2A/H2A.Z/H2B exchange As presented above in the discussion of Nap1, there is a large body of literature describing the exchange of H2A, H2B and H2A variants in living cells, with much of the replacement driven by transcription [47,53,105,174–178,231]; [reviewed in 129,179,251,333]. Although an in-depth analysis of H2A/H2B exchange is beyond the scope of this review, a few observations with respect to chromatin assembly can be made. First, there is ample evidence that the histone variant H2A.Z (Htz1 in yeast) can replace canonical H2A through an ATP-driven process that involves specialized factors, such as the SWR1 complex and Chz1 [182,334–336]. Second, it is uncertain how (or if) deposition of H2A and H2B during chromatin replication differs mechanistically from transcription-driven H2A/H2B exchange. The finding that a significant fraction of free H2A.Z/H2B dimers is associated with Nap1 [182,183,335], and that Nap1 may be a member of the SWR1 complex [182], somewhat blurs the distinction between exchange and de novo assembly. Finally, although the rate and extent of H2A/H2B replacement may vary among systems (including extensive genome-wide exchange in yeast [337]), it is clear that nucleosome disassembly/assembly at replication forks and replication-independent histone replacement must be temporally coincident. Discovering how this macromolecular traffic is controlled and targeted should be a fascinating topic for future investigation. 10. Afterword The model and demonstration of semi-conservative DNA replication provided an immediately discernable and esthetically pleasing answer to the riddle of genetic inheritance [151,338]. In contrast, the details of epigenetic inheritance, including the transfer and deposition of histone proteins at the replication fork, have often been less easily predicted, and less readily revealed. Of course, the actual enzymatic events surrounding the precise copying of DNA proved to be far more complicated than one might have initially predicted. Likewise, the surprising (and seemingly ever proliferating) number of chromatin assembly factors and histone escorts surely underlies the need to properly reconstruct the chromatin fiber following DNA synthesis, from one generation to the next. A consideration of this impressive feat once again calls to mind the striking electron micrographs of replicating chromatin, published 35 years ago (Fig. 1) [5]. The image of a cell copying and rebuilding a functional chromosome offers a tantalizing glimpse into the mechanisms of eukaryotic genetic transmission, mechanisms that have so fruitfully provided matters for study and conjecture. It is certain that these topics will continue to offer opportunities and challenges to future investigators of chromatin biosynthesis. Acknowledgements This work is dedicated in thanks to Dr. Christopher L. F. Woodcock, and to the past and present members of the author's research team. Research in the author's laboratory is supported by a grant from the National Science Foundation (award number 0744590) to ATA.

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