InteractiveFly: GeneBrief

Suppressor of Under-Replication: Biological Overview | References

Eukaryotic DNA replicates asynchronously, with discrete genomic loci replicating during different stages of S phase. Drosophila larval tissues undergo endoreplication without cell division, and the latest replicating regions occasionally fail to complete endoreplication, resulting in underreplicated domains of polytene chromosomes. This study shows that linker histone H1 is required for the underreplication (UR) phenomenon in Drosophila salivary glands. H1 directly interacts with the Suppressor of UR (SUUR) protein and is required for SUUR binding to chromatin in vivo. These observations implicate H1 as a critical factor in the formation of underreplicated regions and an upstream effector of SUUR. It was also demonstrated that the localization of H1 in chromatin changes profoundly during the endocycle. At the onset of endocycle S (endo-S) phase, H1 is heavily and specifically loaded into late replicating genomic regions and is then redistributed during the course of endoreplication. The data suggest that cell cycle-dependent chromosome occupancy of H1 is governed by several independent processes. In addition to the ubiquitous replication-related disassembly and reassembly of chromatin, H1 is deposited into chromatin through a novel pathway that is replication-independent, rapid, and locus-specific. This cell cycle-directed dynamic localization of H1 in chromatin may play an important role in the regulation of DNA replication timing (Andreyeva, 2017).

This study demonstrated that virtually all major sites of UR throughout the Drosophila genome exhibit a substantial increase in salivary gland DNA copy number upon depletion of the linker histone H1, thus implicating H1 in the regulation of endoreplication. In control knockdown salivary glands, 46 underreplicated domains were identified. While these regions are in general agreement with previous efforts to map underreplicated domains by less sensitive microarray analyses, fewer underreplicated sites were identified than a recent report that used high-throughput sequencing of salivary gland DNA (Yarosh, 2014). Notably, the underreplicated domains that the current analyses failed to detect represent sites with the weakest degree of UR. One possible source of variation is the distinct technical approach that was used compared with Yarosh (2014), as simultaneous sequencing of a nonpolytenized (embryonic) genome as a means to normalize the reads from underrepresented sequences in polytenized tissues (Yarosh, 2014) likely provides additional sensitivity. Another potential explanation could lie in the relative sequencing depth of the respective assays (approximately fourfold lower in the current study), considered crucial for the analyses of next-generation sequencing data. However, this explanation is less likely, as subsampling of the current reads to much lower depths yielded no appreciable difference in the number and location of identified underreplicated sites or the change in copy number upon H1 knockdown (Andreyeva, 2017).

On average, a moderate knockdown of H1 led to an ~50% copy number gain at the center of underreplicated domains in intercalary heterochromatin (IH; large dense bands scattered in euchromatin comprising clusters of repressed genes. The copy number is not restored to the same degree as that in a SuUR genetic mutant. The difference is likely attributable to the incomplete depletion of H1. In fact, in an independent biological validation experiment that resulted in an ~95% depletion of H1, an almost complete restoration of copy number was observed. The observation of an almost complete reversal of UR in cells depleted of H1 (but still wild type for SuUR) strongly suggests an epistatic mechanism of action in which both H1 and SUUR act together in the same biochemical pathway (Andreyeva, 2017).

This study found that H1 and SUUR are also involved in UR of PH. For instance, both the mapped pericentric regions and TE sequences, which are highly abundant in pericentric regions, exhibit an increase of DNA copy number upon H1 knockdown. The SuURES mutation also results in a robust loss of UR at PH, as measured by changes in DNA copy number at TEs. The abrogation of H1 expression gives rise to a somewhat weaker effect on the UR of PH than that of IH, which is consistent with an almost complete elimination of SUUR protein from polytene chromosome arms in salivary glands depleted of H1 by RNAi but the persistence of residual SUUR at their PH. The role of H1 in maintaining the underreplicated state of PH may be relevant to its important regulatory functions in constitutive heterochromatin, where it recruits Su(var)3-9, facilitates H3K9 methylation, and maintains TEs in a transcriptionally repressed state. Recently, it was proposed that TE repression in ovarian somatic cells involves an H3K9 methylation-independent process through recruitment of H1 by Piwi-piRNA complexes, resulting in reduced chromatin accessibility. The current results also implicate UR of TE sequences in polytenized cells as yet another putative mechanism that contributes to regulation of their expression. Interestingly, it was shown previously that double mutants encompassing both the Su(var)3-9 and SuUR mutant alleles exhibit a synthetically increased predominance of novel band-interband structures at PH compared with the mutation of SuUR alone. While the evidence suggests a relationship between UR and transcriptionally repressive epigenetic states, such as H3K9 methylation, the nature of this relationship remains largely speculative (Andreyeva, 2017).

This study demonstrated that SUUR protein physically interacts with H1 in both a complex mixture of whole-cell extracts that contain endogenous native H1 and recombinant purified H1 polypeptides. Furthermore, the particular structural domains of the two proteins were delimited that are required for the interaction. SUUR protein contains several sequence features that have been implicated in regulation of UR and binding to specific proteins. Although SUUR possesses a putative bromodomain, it contains no identifiable DNA-binding domain, so the mechanism that allows SUUR to exhibit a preference for specific genomic underreplicated loci is unknown. The positively charged central region is both necessary and sufficient to interact with heterochromatin protein 1a (HP1a), which suggests a possible involvement of HP1a in tethering SUUR to H3K9me2/3-rich PH. However, the specific localization of SUUR to underreplicated IH, which is not enriched for H3K9me2/3, remains enigmatic. This study now demonstrates that the central region of SUUR is also sufficient for binding directly to H1 in vitro. Considering that the central region of SUUR is essential for the faithful localization of the protein to chromatin in vivo, including underreplicated IH, it seems likely that H1 directly mediates the tethering of SUUR to chromatin in underreplicated regions (Andreyeva, 2017).

The tripartite structure of H1 provides multiple binding interfaces for interacting proteins and thus allows H1 to mediate several biochemically separable functions in vivo. For instance, the globular domain and proximal 25% of the CTD are required for H1 loading into chromatin, while the proximal 75% of the CTD is needed for normal polytene morphology, H3K9 methylation, and physical interactions with Su(var)3-9. This study discovered a previously unknown function for the distal 25% of the H1 CTD, which is shown to be essential for binding to SUUR. Deletion of this region of H1 results in a near-complete loss of the interaction with SUUR. Thus, in addition to its critical functions in heterochromatin structure and activity, the CTD of H1 is likely also important in facilitating UR (Andreyeva, 2017).

One of the most striking findings in this study is the observation that the genomic occupancy of H1 undergoes profound changes during the endoreplication cycle. It also remains largely mutually exclusive with that of DNA polymerase clamp loader PCNA, which is consistent with the observed depletion of H1 in nascent chromatin compared with mature chromatin (Andreyeva, 2017).

H1 is heavily loaded into late replicating loci at the onset of replication (when these loci are silent for replication). Combined, the current observations indicate that the chromosome distribution of H1 during the endocycle is governed by at least three independent processes. Two of them [replication-dependent (RD) eviction of H1 and RD deposition of H1 after the passage of replication fork] are related to the well-recognized obligatory processes of chromatin disassembly and reassembly during replication. The third pathway, which directs early deposition of H1 into late replicating loci, has not been described previously. This process is (1) replication-independent (RI); (2) locus-specific, with a strong preference for late replicating sites; and (3) apparently more rapid than the RD deposition of H1, since very high levels of H1 occupancy are observed in all nuclei immediately after the initiation of endo-S. It is possible that the RI pathway of H1 loading into chromatin is mediated by a selective recruitment of H1 based on epigenetic core histone modification-dependent mechanisms. For instance, mammalian H1.2 was reported to recognize H3K27me3, and this modification is very abundant in IH (Andreyeva, 2017).

Also, the RI mechanism for deposition of H1 probably does not involve de novo nucleosome assembly, as H1 is known to exhibit a mutually exclusive distribution with RI core histone variants, and there is no known nuclear process during early S phase that requires core histone turnover. In the future, it will be interesting to further confirm that RI nucleosome assembly does not take place during early replication in salivary gland polytene chromosomes. Finally, the locus-specific RI deposition of H1 in early endo-S chromatin may be conserved in the normal S phase of diploid tissues, and it will require independent experimentation with sorted mitotically dividing cells to confirm this possibility (Andreyeva, 2017).

This study also provides cytological evidence that the functions of H1 and SUUR are biochemically linked. Specifically, it was demonstrated that SUUR localizes to a subset of H1-positive bands and requires H1 for its precise distribution in polytene chromosomes, nuclear localization, and stability in salivary gland cells. These observations implicate H1 as an upstream effector of SUUR functions in vivo and an essential component of the biological pathway that maintains loci of reduced ploidy in polytenized cells. Importantly, this finding adds to a growing list of biochemical partners of H1 that mediate their chromatin-directed functions in an H1-dependent fashion (Andreyeva, 2017).

Interestingly, even a moderate depletion of H1 (to ~30% of normal) results in a complete removal of SUUR from chromosome arms. Thus, H1-dependent localization of SUUR requires high concentrations of the linker histone in chromatin. This conclusion is also consistent with SUUR colocalization with polytene loci that are the most strongly stained for H1. In contrast, elimination of the H3K9me2 mark from polytene spreads requires very extensive depletion of H1, whereas the moderate depletion of H1 does not strongly affect H3K9 dimethylation in the chromocenter or polytene arms. Therefore, the robust effect of even moderate H1 depletion on SUUR localization in chromatin is unlikely to be mediated indirectly through disorganization of heterochromatin structure (Andreyeva, 2017).

Unexpectedly, the cell cycle-dependent temporal pattern of H1 localization is not identical to that of SUUR. In contrast to H1, SUUR protein (1) is only weakly present in IH during early endo-S phase, (2) achieves the maximal occupancy at IH loci only in the late endo-S, and (3) colocalizes with PCNA at certain sites. The observations made in this study and in previous works can be summarized in the following model for H1-mediated regulation of SUUR association with chromatin. The initiation of the deposition of SUUR in chromosomes is strongly dependent on H1. More specifically, SUUR is preferentially localized to chromatin domains that are highly enriched for H1. For instance, the tremendously elevated concentration of H1 in IH of early endo-S cells promotes and nucleates the initiation of deposition of SUUR into these regions. However, the pattern of SUUR occupancy at these sites does not occur temporally in parallel with that of H1. Initially, the exceptionally high abundance of H1 in late replicating loci during early endo-S is not paralleled by a simultaneous comparable increase of SUUR occupancy. Rather, loading of SUUR into these sites lags significantly behind H1 occupancy. Thus, the rate of SUUR localization to H1-rich IH appears to be much slower than that of the RI deposition of H1 into these loci. After the initial recruitment, further loading of SUUR does not require H1, and SUUR continues (in a slower fashion) to accumulate at IH throughout the endo-S phase even when H1-enriched domains dissipate in the course of DNA endoreplication. The additional loading of SUUR in chromatin is likely facilitated by its self-association through dimerization of the N terminus and physical interactions with the replication fork, as proposed previously. In this fashion, SUUR achieves its maximal concentration in IH loci by the late endo-S (Andreyeva, 2017).

This study has demonstrated that H1 has a pivotal function in the establishment of UR of specific IH loci in polytenized salivary gland cells. The findings that H1 interacts directly with SUUR in vitro and is required for SUUR localization to late replicating IH in polytene chromosomes in vivo strongly suggest that the H1-mediated recruitment of SUUR promotes UR by obstructing replication fork progression in its cognate underreplicated loci but does not affect replication origin firing. However, the remarkable temporal pattern of H1 distribution in endoreplicating polytene chromosomes suggests that it may also play a direct SUUR-independent role in regulation of endoreplication. This is especially plausible considering that the temporal distribution patterns of SUUR and H1 are dissimilar (Andreyeva, 2017).

In contrast to the role of SUUR in slowing down the replication fork progression during late endo-S phase, H1 (acting in the absence of SUUR during early endo-S) may function to repress the initiation of endoreplication, as proposed in several studies. DNA-seq analyses also suggest this mechanism. Compared with the relatively smooth, flat profiles of DNA copy numbers in SuURES mutant salivary glands, the profiles in H1-depleted cells exhibit a jagged, uneven appearance, indicative of aberrant local initiation of replication. Unfortunately, the experimental system (cytological analyzes of salivary glands) cannot be used to further confirm this idea. First, an extensive depletion of H1 results in the loss of polytene morphology; second, since the staging of endo-S progression is based on PCNA staining, a spurious activation of ectopic replication origins would result in an incorrect calling of the stage. To further complicate these analyses, polytenized cells are not amenable to other methods of cell cycle staging, such as fluorescence-activated cell sorting (FACS). In the future, it will be important to examine the role of H1 in regulation of DNA replication timing in sorted Drosophila diploid cells (Andreyeva, 2017).

Similarity in replication timing between polytene and diploid cells is associated with the organization of the Drosophila genome

Morphologically, polytene chromosomes of Drosophila melanogaster consist of compact 'black' bands (ruby bands, rb-bands) alternating with less compact 'grey' bands and interbands. This study developed a comprehensive approach that combines cytological mapping data of FlyBase-annotated genes and novel tools for predicting cytogenetic features of chromosomes on the basis of their protein composition and determined the genomic coordinates for all black bands of polytene chromosome 2R. By a PCNA immunostaining assay, the replication timetable was obtained for all the bands mapped. The results allowed comparison of replication timing between polytene chromosomes in salivary glands and chromosomes from cultured diploid cell lines and to observe a substantial similarity in the global replication patterns at the band resolution level. In both kinds of chromosomes, the intervals between black bands correspond to early replication initiation zones. Black bands are depleted of replication initiation events and are characterized by a gradient of replication timing; therefore, the time of replication completion correlates with the band length. The bands are characterized by low gene density, contain predominantly tissue-specific genes, and are represented by silent chromatin types in various tissues. The borders of black bands correspond well to the borders of topological domains as well as to the borders of the zones showing H3K27me3, SUUR, and LAMIN enrichment. In conclusion, the characteristic pattern of polytene chromosomes reflects partitioning of the Drosophila genome into two global types of domains with contrasting properties. This partitioning is conserved in different tissues and determines replication timing in Drosophila (Kolesnikova, 2018).

This work used the four-state chromatin model, previously published data on the chromatin localization of proteins, and in situ hybridization of annotated genes and identified the locations of all black bands of polytene chromosome 2R on a genome map (Kolesnikova, 2018).

The special feature of the four-state chromatin model is the generalization of data obtained from four cell lines. This generalization resulted in identification of two chromatin types -- aquamarine and ruby -- which show stable properties in all these cell lines. Starting the current mapping effort with identification of domains that have ruby chromatin in them and that are flanked by aquamarine chromatin, the genome is roughly divided into constitutively active and constitutively inactive zones (Kolesnikova, 2018).

To take into account the tissue-specific features of chromatin organization in salivary gland polytene chromosomes, the morphology of polytene chromosomes, an extensive pool of 'experimental cytology' data, and data on gene expression in salivary glands were examined and, whenever deemed necessary, corrections were introduced into initial predictions. The results of this study revealed that this approach works well (Kolesnikova, 2018).

Yet another unique feature of the four-state chromatin model is that it reveals the chromatin type (specifically, aquamarine) that is enriched with interband-specific proteins; no other models of clustering chromatin proteins can do this. According to the most recent high-resolution Hi-C data from embryos, TAD boundaries correspond with high resolution to polytene chromosome interbands, whereas black and gray bands are the visualization of topological domains with different types of DNA folding. Thus, the four-state chromatin model allows the boundaries of physical (not epigenetic) domains to be found. The work by Hou (2012) clearly indicates that these boundaries are not always the same (Kolesnikova, 2018).

The choice of aquamarine chromatin as potential band boundaries is supported well by comparing coordinates with the distribution of SUUR and H3K27me3 on polytene chromosomes: these two are markers of black bands and display sharp changes on their boundaries. Thus, the approach used in the current work can be conveniently used to identify the coordinates of specific polytene-chromosome black bands with high accuracy. For most black bands, accuracy of 2-10 kb is attainable (Kolesnikova, 2018).

With the band coordinates inferred, a detailed comparison was performed of replication profiles from diploid cells and replication patterns observed in polytene chromosomes and analyzed the properties of black bands (Kolesnikova, 2018).

A considerable degree of similarity in replication timing has been demonstrated between salivary gland polytene chromosomes and diploid cells. In both object types, the zones between black bands correspond to early replication initiation zones. This result is consistent with the observation that most ORC2-binding sites are in aquamarine chromatin corresponding to interbands. Ruby-containing polytene chromosome bands (Rb-bands) in different cell types have a U-shaped replication profile, which implies that replication in them proceeds from the boundaries to the center, leading to a local delay in replication completion, this delay being proportional to the band length (Kolesnikova, 2018).

The averaged boundary replication profiles in INTs that were built from previously published cell culture data are consistent with the prediction that very few sites within the intervals between rb-bands initiate replication in each replication cycle. The typical size of replicons, 80 kb, originating from the early replication initiation zone (data from cell cultures) fits this model well too. Analysis of stretched DNA fibers in D. nasuta polytene chromosomes has revealed that the replicons initiated in the early S phase are each 64 μm in size on average, which should amount to more than 120 kb. The fact that the replicons are that long provides further support to the hypothesis that the replication origins fired during one cycle in a particular DNA molecule should be well spaced. While analyzing replication in partially denatured polytene chromosomes, researchers observed temporal and spatial asynchrony in replication initiation in parallel fibers and proposed that this asynchrony is one of the main reasons for continuous labeling in polytene chromosomes. Although these data come from a Drosophila species irrelevant to this study and the typical sizes and genomic distances may be different to some extent, it is proposed that the organization of replication in polytene chromosomes is conserved across Drosophila species (Kolesnikova, 2018).

Replication patterns change in polytene chromosomes, and these patterns are linked to events in DNA sequences. At the beginning of the S phase, replication is initiated in INTs, which may contain a large number of potential replication initiation sites. In each DNA strand, an initiation event occurs only once per INT in a random interband. Initiation events in different interbands can occur in asynchrony, either in different INTs or in the same INT on the parallel DNA strands of a polytene chromosome. Replication forks move through INTs in the opposite directions from the site of replication initiation and eventually enter the nearest rb-bands. After all INTs have completed replication, the replication fork should be detectable only in rb-bands. This situation is consistent with the observed inverted PCNA pattern, when all black bands produce the signal that the intervals do not (Kolesnikova, 2018).

By analysis of stretched DNA fibers in D. nasuta polytene chromosomes, it has been demonstrated that the rates of replication fork movement in polytene chromosomes during the late S phase are on average one-tenth of those in the early S phase. The authors believe that upon entering polytene chromosome rb-bands, replication forks slow down. That is why, although some rb-bands are shorter than the flanking intervals, all 'black' bands undergoing replication at once. Replication in these bands goes on until the forks moving toward each other meet. In the longest rb-bands, forks fail to meet before the end of the S phase, leading to under-replication. In D. melanogaster, replication rates depend on SUUR. This study demonstrated that all rb-bands are enriched with SUUR both in salivary gland polytene chromosomes and in diploid cells. According to another study, local artificial tethering of SUUR to an early replicating region of a salivary gland polytene chromosome causes delayed replication there. It can be proposed that this protein plays an important role in delayed replication associated with all rb-bands genome-wide (Kolesnikova, 2018).

Evidence exists that the S phase of the endocycle is quite different from that in diploid cells. The former is distinguished by under-replication of a large part of the genome and low expression of genes involved in replication. The presence of the intra-S checkpoint in salivary gland cells is questionable, and so is activation of any late-firing origins. Nevertheless, the results reveal a substantial similarity in replication timing for the euchromatic arm of the whole chromosome. What underlies this similarity is thought to be the organization of the Drosophila genome. The genome consists of alternations of domains capable of initiating early replication (INTs; the interval between black/ruby bands) and domains with the potential to initiate replication late in the S phase. These late domains vary in size, but seldom are they longer than a few hundred kilobases. In diploid cells, these relatively short domains are replicated by replication forks coming from border origins of replication and complete replication before the classic late S phase, which is when late-firing origins activate. Thus, replication of a large portion of a euchromatic arm is, in the classic sense, early replication. Only the most extended bands and regions of pericentric heterochromatin initiate replication in the late S phase. The question of whether replication initiation events occur in the bands is not easy to answer. Schwaiger (2009) analyzed replication profiles and concluded that extended late-replication zones in cell cultures contain origins initiating replication shortly before the end of the S phase. It can be assumed that the replication origins located in rb-bands do not bind all proteins required for independent initiation of replication, and replication on these origins cannot be initiated before a fork comes from outside; these properties are typical of regions showing a U-shaped replication profile in mammals. The same is suggested by recent studies of the genome-wide distribution of the Mcm2-7 helicase complex in D. melanogaster (Kolesnikova, 2018).

Multiple published comparisons of replication profiles for different tissues of the same organism suggest that each cell type has its own schedule of origin activation. One study on individual IH regions indicates that all the 60 analyzed regions are late replicating in cultured cells, but inside those regions, there are local zones of early replication. After artificial induction of transgene expression in IH, there are also local changes in replication timing (Kolesnikova, 2018).

In cell cultures, early-replication zones within rb-bands can be identified by analysis of the outliers in the boxplots of the averaged boundary replication profile. Among all the rb-bands, only two had early-replication zones spanning them from end to end. It can be theorized that in different tissues, most bands similar in size undergo replication within a similar time interval in the S phase, and gene activation in these bands makes the corresponding fragment of the band earlier replicating (Kolesnikova, 2018).

It can be concluded that the alternation of rb-bands and INTs forms the basis of the pattern of replication timing in D. melanogaster. This organization is conserved in eukaryotes. It has been demonstrated that a substantial portion of the mammalian genome represents the alternation of replication initiation zones, in which early master origins lie, and U-shaped replication zones, in which initiation occurs at virtually random positions and in a cascadelike manner, shaping the profile accordingly. The initiation zones are notable for active transcription and high gene density. The boundaries of these zones correspond to those of topological domains (Kolesnikova, 2018).

IH regions represent a separate fraction of black bands, grossly corresponding to the most extended and late-replicating bands (group LR5). This study demonstrated that all rb-bands, including small ones, share a large number of properties with IH regions. This is direct evidence that among all genomic regions, IH regions do not stand out as some special type of sequences. Genes in any rb-band tend to be expressed in a limited number of tissues and, according to GO analysis, these regions are enriched with tissue-specific genes. By contrast, the intervals between black bands are enriched with genes that are highly expressed in most tissues chosen for analysis here. Each rb-band appears as a combination of repressed chromatin types; however, open chromatin can be found in its boundary regions, pointing to a similarity between bands and TADs. It is confirmed that the boundaries of black bands correspond to those of topologically associating domain or sub-domains. TADs represent a stable level of genome organization during development both in mammals and in Drosophila. It has been demonstrated that the partitioning of genomes into physical domains correlates with gene density and transcription distribution. These features are closely associated with replication timing. That late replication correlates with LADs has been demonstrated in both Drosophila and mammals (Kolesnikova, 2018).

The results of this work suggest that Drosophila polytene chromosomes can serve as vivid visualization of the organization of the eukaryotic genome, which is conserved between Drosophila and mammals. The characteristic pattern of polytene chromosomes -- the compacted black bands alternating with less compact grey bands and interbands -- reflects the partitioning of the Drosophila genome into domains with contrasting properties (Kolesnikova, 2018).

Rif1 inhibits replication fork progression and controls DNA copy number in Drosophila

Control of DNA copy number is essential to maintain genome stability and ensure proper cell and tissue function. In Drosophila polyploid cells, the SNF2-domain-containing SUUR protein inhibits replication fork progression within specific regions of the genome to promote DNA underreplication. While dissecting the function of SUUR's SNF2 domain, an interaction between SUUR and Rif1 was identified. Rif1 has many roles in DNA metabolism and regulates the replication timing program. Repression of DNA replication is dependent on Rif1. Rif1 localizes to active replication forks in a partially SUUR-dependent manner and directly regulates replication fork progression. Importantly, SUUR associates with replication forks in the absence of Rif1, indicating that Rif1 acts downstream of SUUR to inhibit fork progression. These findings uncover an unrecognized function of the Rif1 protein as a regulator of replication fork progression (Munden, 2018).

The SUUR protein is responsible for promoting underreplication of heterochromatin and many euchromatin regions of the genome. Although SUUR was recently shown to promote underreplication through inhibition of replication fork progression, the underlying molecular mechanism has remained unclear. Through biochemical, genetic, genomic and cytological approaches, this study has found that SUUR recruits Rif1 to replication forks and that Rif1 is responsible for underreplication. This model is supported by several independent lines of evidence. First, SUUR associates with Rif1, and SUUR and Rif1 co-localize at sites of replication. Second, underreplication is dependent on Rif1, although Rif1 mutants have a clear pattern of late replication in endo cycling cells. Third, SUUR localizes to replication forks and heterochromatin in a Rif1 mutant, however, it is unable to inhibit replication fork progression in the absence of Rif1. Fourth, Rif1 controls replication fork progression and phenocopies the effect loss of SUUR function has on replication fork progression. Fifth, SUUR is required for Rif1 localization to replication forks. Critically, using the gene amplification model to separate initiation and elongation of replication, it was shown that Rif1 can affect fork progression without altering the extent of initiation. Based on these observations, this study defines a new function of Rif1 as a regulator of replication fork progression (Munden, 2018).

This work suggests that the SNF2 domain of SUUR is critical for its ability to localize to replication forks. This is based on the observation that deletion of this domain results in a protein that is unable to localize to replication forks, but still localizes to heterochromatin. SUUR has previously been shown to dynamically localize to replication forks during S phase, but constitutively binds to heterochromatin (Kolesnikova, 2013; Nordman, 2014). SUUR associates with HP1 and this interaction occurs between the central region of SUUR and HP1 (Pindyurin, 2008). Therefore, it is speculated that the interaction between SUUR and HP1 is responsible for constitutive SUUR localization to heterochromatin, while a different interaction between the SNF2 domain and a yet to be defined component of the replisome, or replication fork structure itself, recruits SUUR to active replication forks during S phase (Munden, 2018).

Uncoupling of SUUR's ability to associate with replication forks and heterochromatin also provides a new level of mechanistic understanding of underreplication. Overexpression of the C-terminal two-thirds of SUUR is capable of inducing ectopic sites of underreplication. In contrast, overexpression of the SUUR's SNF2 domain, in the presence of endogenous SUUR, suppresses SUUR-mediated underreplication (Kolesnikova, 2005). Together with the data presented in this study, it is suggested that overexpression of the SNF2 domain interferes with recruitment of full-length SUUR to replication forks, by saturating potential SUUR binding sites at the replication fork. Although the C-terminal region of SUUR is necessary to induce underreplication (Kolesnikova, 2005), the C-terminal portion of SUUR remains associated with heterochromatin in the SUURΔSNF construct, but this protein is not sufficient to induce underreplication. It is suggested that at physiological levels, the affinity of SUUR for replication forks is substantially diminished in the absence of the SNF2 domain. This work raises questions about the biological significance of SUUR binding to heterochromatin, since without the SNF2 domain SUUR is still constitutively bound to heterochromatin, yet unable to induce underreplication. Additionally, SUUR dynamically associates with heterochromatin in mitotic cells although heterochromatin is fully replicated (Munden, 2018).

While trying to uncover the molecular mechanism through which SUUR is able to inhibit replication fork progression, this study has uncovered an interaction between SUUR and Rif1. Through subsequent analysis, it was demonstrated that Rif1 has a direct role in copy number control and that Rif1 acts downstream of SUUR in the underreplication process. Although underreplication is largely dependent on SUUR, there are several sites that display a modest degree of underreplication in the absence of SUUR. In a Rif1 mutant, however, these sites are fully replicated and there is no longer any detectable levels of underreplication within any regions of the genome. It is possible that Rif1 is capable of promoting underreplication through a mechanism independent of SUUR. Therefore, it is concluded that Rif1 is a critical factor in driving underreplication (Munden, 2018).

Further emphasizing the critical role Rif1 plays in copy number control, this study has shown that Rif1 acts downstream of SUUR in promoting underreplication. SUUR is still able to associate with chromatin in the absence of Rif1 but is unable to promote underreplication. Underreplicated regions of the genome, including heterochromatin, tend to be late replicating, raising the possibility that changes in replication timing in a Rif1 mutant suppresses underreplication. Rif1 mutant endocycling cells of Drosophila display a cytological pattern of late replication, where heterochromatin is discretely replicated. While Rif1 controls replication timing in Drosophila and is necessary for the onset of late replication at the mid-blastula transition (Seller, 2018), it is argued that the changes in copy number associated with loss of Rif1 function are not solely due to a loss of late replication. This is supported by the clear pattern of late replication of heterochromatin in Rif1 mutant endocycling cells, although heterochromatin appears to be fully replicated in these cells. Previous work in mammalian polyploid cells has shown that underreplication is dependent on Rif1, which was attributed to changes in replication timing (Hannibal, 2016). It is important to note that Rif1-dependent changes in replication timing were not measured in this system and that many genomic regions transition from early to late replication in a Rif1 mutant (Foti, 2016). This work raises the possibility that Rif1 has a direct role in mammalian underreplication through a mechanism similar to that of Drosophila and may not simply be due to indirect changes in replication timing. Future work will be necessary to define the role of mammalian Rif1 in underreplication (Munden, 2018).

This analysis of amplification loci demonstrates that Rif1 controls replication fork progression independently of initiation control, thus demonstrating that Rif1 has a specific effect on replication fork progression. Therefore, this study has uncovered a new role for Rif1 in DNA metabolism as a regulator of replication fork progression and copy number control. Rif1 has been identified as part of the replisome in human cells by nascent chromatin capture, a technique that identifies proteins associated with newly synthesized chromatin. Multiple studies have assessed whether loss of Rif1 function affects replication fork progression in yeast, mouse and human cells, but have come to different conclusions. DNA fiber assays have been used to measure fork progression in these studies and nearly all have shown that Rif1 mutants have a slight increase in replication fork progression, although not always statistically significant. There could be several reasons for these differing results; Rif1 may control replication fork progression in specific genomic regions that may be underrepresented in some assays, Rif1 function could vary among different cell types, or sample sizes may have been too small to reach significance. These observations, taken together with these previous studies, leave open the possibility that Rif1-mediated control of replication fork progression could be an evolutionarily conserved function of Rif1. It is not suggested that Rif1 is constitutively associated with replication forks in all cell types. Rather, Rif1 could be recruited to replication forks at a specific time in S phase, or in specific developmental contexts, to modulate the progression of replication forks and provide an additional layer of regulation of the DNA replication program (Munden, 2018).

How could SUUR and Rif1 function in concert to inhibit replication fork progression? This study has shown that Rif1 retention at replication forks is dependent on SUUR. Additionally, underreplication depends on Rif1's PP1-binding motif, raising the possibility that a Rif1/PP1 complex is necessary to inhibit replication fork progression. Rif1/PP1 dephosphorylates DDK-activated helicases to control replication initiation. Recently, however, DDK-phosphorylated MCM subunits have been shown to be necessary to maintain DNA-unwinding enzyme Cdc45~MCM~GINS (CMG) association and stability of the helicase (Alver, 2017). This result suggests that continued phosphorylation of the helicase is necessary for replication fork progression (Alver, 2017). It is proposed that SUUR recruits Rif1/PP1 to replication forks where it is able to dephosphorylate MCM subunits, ultimately inhibiting replication fork progression. Although this mechanism needs to be tested biochemically, it provides a framework to address the underlying molecular mechanism responsible for controlling DNA copy number and could provide new insight into the mechanism(s) Rif1 employs to regulate replication timing. (Munden, 2018).

Functional dissection of Drosophila melanogaster SUUR protein influence on H3K27me3 profile

In eukaryotes, heterochromatin replicates late in S phase of the cell cycle and contains specific covalent modifications of histones. SuUR mutation found in Drosophila makes heterochromatin replicate earlier than in wild type and reduces the level of repressive histone modifications. SUUR protein was shown to be associated with moving replication forks, apparently through the interaction with PCNA. The biological process underlying the effects of SUUR on replication and composition of heterochromatin remains unknown. This study performed a functional dissection of SUUR protein effects on H3K27me3 level. Using hidden Markow model-based algorithm SuUR-sensitive chromosomal regions were revealed that demonstrated unusual characteristics: They do not contain Polycomb and require SUUR function to sustain H3K27me3 level. This study tested the role of SUUR protein in the mechanisms that could affect H3K27me3 histone levels in these regions. SUUR did not affect the initial H3K27me3 pattern formation in embryogenesis or Polycomb distribution in the chromosomes. The possible effect of SUUR on histone genes expression and its involvement in DSB repair were also ruled out. These results support the idea that SUUR protein contributes to the heterochromatin maintenance during the chromosome replication (Posukh, 2017).

SuUR mutation affects two processes in the repressed regions of polytene chromosomes-their polytenization and repressed histone modifications maintenance. In SuUR mutants, the replication in these regions becomes more efficient; however, the levels of H3K27me3 and H3K9me3 decrease significantly. A 'differential diagnosis' was performed for the effects of SuUR mutation on H3K27me3 level in polytene chromosomes. Successive conclusions allowed exclusion of SUUR involvement in certain mechanisms that, to this point, obscured the assessment of its function. Fur potential explanations were tested for the remarkable, but insufficiently studied, effects that SUUR has on chromosome replication and chromatin. The study revealed that H3K27 methylation in SuUR-sensitive regions (SSRs) of wild type chromosomes does not happen in response to DSB formation during under-replication as was shown in other model systems. Indeed, many regions that are 100% polytenized in wild type contain H3K27me3 that is sensitive to SuUR mutation. It was also shown that SuUR mutation does not affect Polycomb DamID profile in salivary gland and is not involved in the initial placement of H3K27me3 mark early in embryogenesis. These results indicate that the effect of SuUR mutation on H3K27me3 level develops during the ontogenesis. Finally, the possibility of SUUR protein regulating the expression of histone genes was excluded (Posukh, 2017).

In a previous work, a hypothesis was proposed that SUUR protein is involved in the maintenance of repressive histone modifications during replication in Drosophila. It was suggested that SUUR protein could function in the replication-coupled re-establishment of repressed histone modifications in polytene chromosomes. According to this model, SUUR impedes the progression of the replication complex through heterochromatin regions until the pattern of repressed histone marks is properly re-established on the newly synthesized DNA strands (or until the context for future chromatin maturation is properly formed). In the absence of this regulation, replication forks progress through heterochromatin regions more efficiently, but at the expense of the significant depletion of H3K27me3 and H3K9me3. This model combines all major effects of SUUR protein and provides a causal link between them. In this study, necessary experiments were performed to test this model in context of SUUR effect on H3K27me3 (Posukh, 2017).

New data obtained in this study add fascinating details to the well-known effects of SUUR protein. Analysis of the published H3K27me3 profile in salivary gland chromosomes revealed two distinct types of H3K27me3-containing regions-SSRs and SNRs-that differ in H3K27me3 levels, sensitivity to SuUR mutation and the presence of Pc. Intriguingly, the reduction in H3K27me3 levels upon SuUR mutation is observed only in regions that are moderately enriched with H3K27me3 and lack Pc, whereas highly enriched regions remain unaffected. Although, SUUR DamID signal in SNRs is even higher than in SSRs, which is consistent with early cytological studies. Thus, SUUR function is required to preserve H3K27me3 levels at the regions, which are devoid of Pc (Posukh, 2017).

The majority of SSRs detected in this study overlap with the BLACK chromatin type described in Kc167 cells. Genomic regions corresponding to BLACK chromatin were recently shown to contain H3K27me2 in Sg4 cells. Given the known cross-reactivity of the antibodies, which were used in ChIP experiments that detected SuUR effect on H3K27 methylation level, it could be suggested that SSRs mainly contain di-methylated H3K27 and SuUR mutation affects the level of this modification. This suggestion contradicts with the previous immunostaining results that showed no effect of SuUR mutation on H3K27me2 level; however, the effect may be too subtle to be detected with the cytological methods. The present study proved that the previously observed effect of SuUR mutation is specifically directed on tri-methylated H3K27; however, to further address the mechanism of SUUR action in chromatin it would be useful to study the effects of this protein considering a wider spectrum of histone modifications (Posukh, 2017).

It is highly plausible that Pc-G proteins maintain H3K27 methylation at their target regions by temporarily over-producing H3K27me3-marked histones prior to replication, as reported earlier. Hence, the regions of high H3K27me3 enrichment resist the effect of SuUR mutation (Posukh, 2017).

Although the presence of Polycomb protein in SNRs apparently compensates the effect of SuUR mutation on H3K27me3 level, it fails to neutralize the effect of SUUR protein on the replication in these regions. Indeed, the very first characterized under-replicated region (89DE) contains Bithorax complex, which is densely covered with Pc, but is still under-replicated in wild type and fully polytenized in SuUR mutants. Similar situation is observed in the Antennapedia complex. Both Bithorax complex and Antennapedia complex are as large as 200–300 kb, so it seems that under-replication normally occurs at H3K27me3-enriched regions that exceed a certain length and lack internal replication origins. This suggestion is in line with recently discovered negative correlation between the length of the under-replicated regions and their polytenization levels. Hence, the selectivity of SuUR mutation effect on H3K27me3 level turns out to be associated with the presence of Pc protein. However, the effect of SuUR mutation on under-replication apparently is largely dependent on the size of the repressed domain, rather than its overall level of H3K27me3. These conclusions are consistent with earlier cytological observations based on immunostaining. Discovered SUUR protein effects on replication and chromatin in polytene chromosomes are schematically summarized in a Scheme explaining the effects of SUUR on H3K27me3 maintenance and under-replication in polytene chromosomes. (Posukh, 2017).

The fact that SSRs do not bind Pc suggests two plausible mechanisms of how SUUR could maintain the level of H3K27me3 in these regions. On the one hand, SUUR could mediate the interaction between the replication complex and H3K27-specific methylase PRC2 and possibly other histone-modifying enzymes. On the other hand, SUUR could regulate the incorporation of parental modified histones (or those over-produced by PRCs at their binding sites) into nascent chromatin of SSRs. Notably, a recent study suggests that linker histone H1 may be involved in this process. Future studies will elucidate the exact mechanism of this process (Posukh, 2017).

DNA replication in nurse cell polytene chromosomes of Drosophila melanogaster otu mutants

Drosophila cell lines are used extensively to study replication timing, yet data about DNA replication in larval and adult tissues are extremely limited. To address this gap, DNA replication in polytene chromosomes from nurse cells of Drosophila melanogaster otu mutants was traced using bromodeoxyuridine incorporation. Importantly, nurse cells are of female germline origin, unlike the classical larval salivary glands, that are somatic. In contrast to salivary gland polytene chromosomes, where replication begins simultaneously across all puffs and interbands, replication in nurse cells is first observed at several specific chromosomal regions. For instance, in the chromosome 2L, these include the regions 31B-E and 37E and proximal parts of 34B and 35B, with the rest of the decondensed chromosomal regions joining replication process a little later. It was observed that replication timing of pericentric heterochromatin in nurse cells was shifted from late S phase to early and mid stages. Curiously, chromosome 4 may represent a special domain of the genome, as it replicates on its own schedule which is uncoupled from the rest of the chromosomes. Finally, it is report that SUUR protein, an established marker of late replication in salivary gland polytene chromosomes, does not always colocalize with late-replicating regions in nurse cells (Koryakov, 2014).

DNA copy-number control through inhibition of replication fork progression

Proper control of DNA replication is essential to ensure faithful transmission of genetic material and prevent chromosomal aberrations that can drive cancer progression and developmental disorders. DNA replication is regulated primarily at the level of initiation and is under strict cell-cycle regulation. Importantly, DNA replication is highly influenced by developmental cues. In Drosophila, specific regions of the genome are repressed for DNA replication during differentiation by the SNF2 domain-containing protein Suppressor of Under-Replication (SuUR) through an unknown mechanism. This study demonstrates that SuUR is recruited to active replication forks and mediates the repression of DNA replication by directly inhibiting replication fork progression instead of functioning as a replication fork barrier. Mass spectrometry identification of SUUR-associated proteins identified the replicative helicase member CDC45 as a SUUR-associated protein, supporting a role for SUUR directly at replication forks. These results reveal that control of eukaryotic DNA copy number can occur through the inhibition of replication fork progression (Nordman, 2014).

Drosophila SUUR protein associates with PCNA and binds chromatin in a cell cycle-dependent manner

Drosophila SUUR protein was shown to regulate the DNA replication elongation process in endocycling cells. This protein is also known to be the component of silent chromatin in polyploid and diploid cells. To mark the different cell cycle stages, immunostaining patterns of PCNA, the main structural component of replication fork, were used. It was demonstrated that SUUR chromatin binding is dynamic throughout the endocyle in Drosophila salivary glands. SUUR chromosomal localization changed along with PCNA pattern and these proteins largely co-localized during the late S-phase in salivary glands. The hypothesized interaction between SUUR and PCNA was confirmed by co-immunoprecipitation from embryonic nuclear extracts. These findings support the idea that the effect of SUUR on replication elongation depends on the cell cycle stage and can be mediated through its physical interaction with replication fork (Kolesnikova, 2013).

Conservation of domain structure in a fast-evolving heterochromatic SUUR protein in drosophilids

Different genomic regions replicate at a distinct time during S-phase. The SuUR mutation alters replication timing and the polytenization level of intercalary and pericentric heterochromatin in Drosophila melanogaster salivary gland polytene chromosomes. This study analyzed SuUR in different insects, identified conserved regions in the protein, substituted conserved amino acid residues, and studied effects of the mutations on SUUR function. SuUR orthologs were identified in all sequenced drosophilids, and a highly divergent ortholog was found in the mosquito genome. SUUR was shown to evolve at a very high rate comparable with that of Transformer. Remarkably, upstream ORF within 5' UTR of the gene is more conserved than SUUR in drosophilids, but it is absent in the mosquito. The domain structure and charge of SUUR are maintained in drosophilids despite the high divergence of the proteins. The N-terminal part of SUUR with similarity to the SNF2/SWI2 proteins displays the highest level of conservation. Mutation of two conserved amino acid residues in this region impairs binding of SUUR to polytene chromosomes and reduces the ability of the protein to cause DNA underreplication. The least conserved middle part of SUUR interacting with HP1 retains positively and negatively charged clusters and nuclear localization signals. The C terminus contains interlacing conserved and variable motifs. These results suggest that SUUR domains evolve with different rates and patterns but maintain their features (Yurlova, 2009).

Interaction between the Drosophila heterochromatin proteins SUUR and HP1

SUUR (Suppressor of Under-Replication) protein is responsible for late replication and, as a consequence, for DNA underreplication of intercalary and pericentric heterochromatin in Drosophila melanogaster polytene chromosomes. However, the mechanism by which SUUR slows down the replication process is not clear. To identify possible partners for SUUR a yeast two-hybrid screen was performed using full-length SUUR as bait. This identified HP1, the well-studied heterochromatin protein, as a strong SUUR interactor. Furthermore, this study has determined that the central region of SUUR is necessary and sufficient for interaction with the C-terminal part of HP1, which contains the hinge and chromoshadow domains. In addition, recruitment of SUUR to ectopic HP1 sites on chromosomes provides evidence for their association in vivo. Indeed, the distributions of SUUR and HP1 on polytene chromosomes are interdependent: both absence and overexpression of HP1 prevent SUUR from chromosomal binding, whereas SUUR overexpression causes redistribution of HP1 to numerous sites occupied by SUUR. Finally, HP1 binds to intercalary heterochromatin when histone methyltransferase activity of SU(VAR)3-9 is increased. It is proposed that interaction with HP1 is crucial for the association of SUUR with chromatin (Pindyurin, 2008).

The SU(VAR)3-9/HP1 complex differentially regulates the compaction state and degree of underreplication of X chromosome pericentric heterochromatin in Drosophila melanogaster

In polytene chromosomes of Drosophila melanogaster, regions of pericentric heterochromatin coalesce to form a compact chromocenter and are highly underreplicated. Focusing on study of X chromosome heterochromatin, it was demonstrated that loss of either SU(VAR)3-9 histone methyltransferase activity or HP1 protein differentially affects the compaction of different pericentric regions. Using a set of inversions breaking X chromosome heterochromatin in the background of the Su(var)3-9 mutations, it was shown that distal heterochromatin (blocks h26-h29) is the only one within the chromocenter to form a big 'puff'-like structure. The 'puffed' heterochromatin has not only unique morphology but also very special protein composition as well: (1) it does not bind proteins specific for active chromatin and should therefore be referred to as a pseudopuff and (2) it strongly associates with heterochromatin-specific proteins SU(VAR)3-7 and SUUR, despite the fact that HP1 and HP2 are depleted particularly from this polytene structure. The pseudopuff completes replication earlier than when it is compacted as heterochromatin, and underreplication of some DNA sequences within the pseudopuff is strongly suppressed. So, it was shown that pericentric heterochromatin is heterogeneous in its requirement for SU(VAR)3-9 with respect to the establishment of the condensed state, time of replication, and DNA polytenization (Demakova, 2007).

In Su(var)2-5 and Su(var)3-7 mutants the chromocenter in salivary gland nuclei looks relatively loose and diffuse (Spierer, 2005). Loss or drastic reduction in the HMTase activity of SU(VAR)3-9 also results in strong decompaction of pericentric heterochromatin (PH) in polytene chromosomes. Most likely, only the functional complex of all these proteins can form compact pericentric heterochromatin. It was of interest to find out to what degree the compact state of heterochromatin is dependent on specific heterochromatin proteins, particularly SU(VAR)3-9. Using a set of chromosome rearrangements placing different parts of X chromosome heterochromatin adjacent to euchromatin, it was discovered that different portions of the polytene Xh are very differently affected by loss of SU(VAR)3-9: only the distal part of heterochromatin (heterochromatic blocks h26-h29 of the metaphase chromosome map becomes decondensed to a varying extent, while morphology of the heterochromatic blocks h30-h34 appears unaffected (Demakova, 2007).

The extent of heterochromatin decompaction depends on several different factors. The pseudopuff is more pronounced in males than in females. Although MSL2 protein, the core component of the dosage compensation complex (DCC), was not found in the pseudopuff, it is suggested that dosage compensation, a process equalizing X-linked gene expression in both sexes, might be responsible for this effect. It is known that DCC is distributed rather discretely, particularly, skipping over intercalary heterochromatin (IH) regions along the male X. Nevertheless, the whole X has a decondensed appearance and underreplication in IH regions is highly suppressed due to DCC function. So, it can be proposed that in males the 'puffed' portion of pericentric Xh relocated into the vicinity of euchromatin becomes dependent on dosage-compensating mechanisms, similarly to IH regions. Another possibility is that the Y chromosome, which represents an additional factor competing for the compaction proteins, might also contribute to the decompaction of Xh and to pseudopuff formation. To note, the fully heterochromatic Y chromosome comprises 40.9 Mb of DNA, while Xh contains ∼20 Mb (Demakova, 2007).

Among the inversions analyzed, w^m4 produces the largest pseudopuff. A good explanation for this effect is currently missing; possibly, the chromatin environment in this eu-heterochromatin junction might contribute to Xh puffing, or some of the DNA sequences might be differentially represented in Xh in different inversions. And finally, decondensation is most strongly expressed on the background of two mutations, Su(var)3-9 and SuUR, despite the fact that SuUR mutation itself does not induce puffing of Xh. The SuUR mutation results in additional polytenization of some of the Xh regions and thus it might increase the amount of Xh material capable of forming the pseudopuff. So, it is believed that loss of both proteins, SUUR and SU(VAR)3-9, has an additive effect on the sizes of the decompacted region (Demakova, 2007).

The region of decondensed heterochromatin that can be called the pseudopuff does not demonstrate signs of true transcriptionally active puffs: the proteins characteristic for active decondensed chromatin (Z4, MSL2, JIL-1, and H3Me3K4) were not detected in the pseudopuff, with the exception of a very weak signal of PolIIo. At the same time, in the Su(var)3-9 mutants, HP1 and HP2 are weakly associated with the region 20F1-4, whereas SUUR and SU(VAR)3-7, in contrast, intensively bind the entire body of the pseudopuff. Recruitment of SU(VAR)3-7 into the pseudopuff in the absence of HP1 and SU(VAR)3-9 appears to be a very specific characteristic of pseudopuff heterochromatin since HP1 and SU(VAR)3-7 proteins cooperate closely in chromosome organization and development (Spierer, 2005). Presence of the SUUR and SU(VAR)3-7 in decompacted chromatin of the pseudopuff might indicate that they do not participate in the process of compaction of this heterochromatic material or that they act in this direction only in the complex with functional SU(VAR)3-9 (Demakova, 2007).

It is interesting to note that different parts of Xh respond differentially to the removal of this complex. The question then, is which features of organization permit proximal heterochromatin to maintain its dense packing in the absence of HP1 and SU(VAR)3-9 activities? It might be proposed that these regions are under control of protein complexes of another composition. For example, these complexes might not utilize HMTase SU(VAR)3-9. However, data on position-effect variegation contradict this idea, since gene inactivation induced by chromosome rearrangements in proximal heterochromatin also depends on the SU(VAR)3-9. It could be suggested that these complexes include some additional compacting proteins. It is known that, in contrast to the distal part of PH, its proximal part is enriched with satellite sequences that are associated with some specific proteins. For, example, the AT-hook protein D1 specifically binds to AT-rich satellites in deep Xh (Demakova, 2007).

In the course of investigating pseudopuff replication it was found that underreplication of the heterochromatic sequences was suppressed in the region of the eu-heterochromatic junction of the w^m4 inversion in Su(var)3-9 mutant. Thus, full polytenization of at least a 45-kb fragment adjacent to euchromatin occurs. The pseudopuff region, in general, completes replication before the end of S-phase; in other words, it does so earlier than the bulk of PH and even some IH regions. Still, a notable fraction of X chromosome heterochromatin sequences remains underreplicated in the polytene chromocenter. It can be assumed that some Xh regions not only do not complete replication but also do not start it. Probably, these regions are separated from replicating chromatin by some barriers that prevent progression of replication forks from adjacent replicons. Therefore, this study demonstrates that Su(var)3-9⁰⁶ may act as a suppressor of underreplication. However, it is not known how this mutation can affect underreplication in other heterochromatic regions (Demakova, 2007).

Replication timing of the pseudopuff is notably changed in the absence of essential changes in transcriptional activity (the PolIIo painting of the pseudopuff looks no more intensive than that of the chromocenter). Moreover, the H3K4 methylation mark characteristic of active regions was not found in the pseudopuff at all. At the same time there exists a correlation between the shift to earlier replication and chromatin decompaction. This observation is interesting in relation to cause-effect relationships among replication timing, transcriptional activity, and decompaction of chromatin (Demakova, 2007).

It is interesting to note that the effect of the SuUR mutation, known as suppression of underreplication, was found not only in PH but also in all IH regions and that these regions complete replication earlier in SuUR mutants than in wild-type ones. The pseudopuff material in SuUR mutants is virtually at the same level of polytenization as in the Su(var)3-9 mutant. However, the SuUR mutation does not involve decompaction of the distal Xh. The same was noted for IH regions. Even if SuUR does induce decompaction of high-level chromatin structures, this is not detected by cytological means. Therefore, SuUR mutation affects replication timing differently compared to Su(var)3-9 (Demakova, 2007).

Summing up, it can be concluded that the reaction of pericentric heterochromatin to loss of SU(VAR)3-9 and HP1 varies in different regions of X heterochromatin. Only the distal part of it undergoes decondensation and forms a new structure called a pseudopuff, which has a specific organization, demonstrating some characteristics of active chromatin: decompaction and, concomitantly, earlier completion of replication. At the same time, the pseudopuff does not contain proteins of active chromatin but does contain several heterochromatic proteins (Demakova, 2007).

High-resolution analysis of Drosophila heterochromatin organization using SuUR Su(var)3-9 double mutants

The structural and functional analyses of heterochromatin are essential to understanding how heterochromatic genes are regulated and how centromeric chromatin is formed. Because the repetitive nature of heterochromatin hampers its genome analysis, new approaches need to be developed. This study describes how, in double mutants for Su(var)3-9 and SuUR genes encoding two structural proteins of heterochromatin, new banded heterochromatic segments appear in all polytene chromosomes due to the strong suppression of under-replication in pericentric regions. FISH on salivary gland polytene chromosomes from these double mutant larvae allows high resolution of heterochromatin mapping. In addition, immunostaining experiments with a set of antibodies against euchromatic and heterochromatic proteins reveal their unusual combinations in the newly appeared segments: binding patterns for HP1 and HP2 are coincident, but both are distinct from H3diMetK9 and H4triMetK20. In several regions, partial overlapping staining is observed for the proteins characteristic of active chromatin RNA Pol II, H3triMetK4, Z4, and JIL1, the boundary protein BEAF, and the heterochromatin-enriched proteins HP1, HP2, and SU(VAR)3-7. The exact cytological position of the centromere of chromosome 3 was visualized on salivary gland polytene chromosomes by using the centromeric dodeca satellite and the centromeric protein CID. This region is enriched in H3diMetK9 and H4triMetK20 but is devoid of other proteins analyzed (Andreyeva, 2007).

Functional dissection of the Suppressor of UnderReplication protein of Drosophila melanogaster: identification of domains influencing chromosome binding and DNA replication

The Suppressor of UnderReplication (SuUR) gene controls the DNA underreplication in intercalary and pericentric heterochromatin of Drosophila melanogaster salivary gland polytene chromosomes. This work investigated the functional importance of different regions of the SUUR protein by expressing truncations of the protein in an UAS-GAL4 system. SUUR was found to have at least two separate chromosome-binding regions that are able to recognize intercalary and pericentric heterochromatin specifically. The C-terminal part controls DNA underreplication in intercalary heterochromatin and partially in pericentric heterochromatin regions. The C-terminal half of SUUR suppresses endoreplication when ectopically expressed in the salivary gland. Ectopic expression of the N-terminal fragments of SUUR depletes endogenous SUUR from polytene chromosomes, causes the SuUR- phenotype and induces specific swellings in heterochromatin (Kolesnikova, 2005).

Influence of the SuUR gene on intercalary heterochromatin in Drosophila melanogaster polytene chromosomes

Salivary gland polytene chromosomes of Drosophila melanogaster have a reproducible set of intercalary heterochromatin (IH) sites, characterized by late DNA replication, underreplicated DNA, breaks and frequent ectopic contacts. The SuUR mutation has been shown to suppress underreplication, and wild-type SuUR protein is found at late-replicating IH sites and in pericentric heterochromatin. This study shows that the SuUR gene influences all four IH features. The SuUR mutation leads to earlier completion of DNA replication. Using transgenic strains with two, four or six additional SuUR(+) doses (4-8xSuUR(+)) this study showed that wild-type SuUR is an enhancer of DNA underreplication, causing many late-replicating sites to become underreplicated. The underreplication sites were mapped, and it was shown that their number increases from 58 in normal strains (2xSuUR(+)) to 161 in 4-8xSuUR(+) strains. In one of these new sites (1AB) DNA polytenization decreases from 100% in the wild type to 51%-85% in the 4xSuUR (+) strain. In the 4xSuUR(+) strain, 60% of the weak points coincide with the localization of Polycomb group (PcG) proteins. At the IH region 89E1-4 (the Bithorax complex), a typical underreplication site, the degree of underreplication increases with four doses of SuUR(+) but the extent of the underreplicated region is the same as in wild type and corresponds to the region containing PcG binding sites. It is conclude dthat the polytene chromosome regions known as IH are binding sites for SuUR protein and in many cases PcG silencing proteins. It is proposed that these stable silenced regions are late replicated and, in the presence of SuUR protein, become underreplicated (Zhimulev, 2003).

Search PubMed for articles about Drosophila SuUR

Alver, R. C., Chadha, G. S., Gillespie, P. J. and Blow, J. J. (2017). Reversal of DDK-Mediated MCM Phosphorylation by Rif1-PP1 Regulates Replication Initiation and Replisome Stability Independently of ATR/Chk1. Cell Rep 18(10): 2508-2520. PubMed ID: 28273463

Andreyeva, E. N., Kolesnikova, T. D., Demakova, O. V., Mendez-Lago, M., Pokholkova, G. V., Belyaeva, E. S., Rossi, F., Dimitri, P., Villasante, A. and Zhimulev, I. F. (2007). High-resolution analysis of Drosophila heterochromatin organization using SuUR Su(var)3-9 double mutants. Proc Natl Acad Sci U S A 104(31): 12819-12824. PubMed ID: 17640911

Andreyeva, E. N., Bernardo, T. J., Kolesnikova, T. D., Lu, X., Yarinich, L. A., Bartholdy, B. A., Guo, X., Posukh, O. V., Healton, S., Willcockson, M. A., Pindyurin, A. V., Zhimulev, I. F., Skoultchi, A. I. and Fyodorov, D. V. (2017). Regulatory functions and chromatin loading dynamics of linker histone H1 during endoreplication in Drosophila. Genes Dev 31(6): 603-616. PubMed ID: 28404631

Demakova, O. V., et al. (2007). The SU(VAR)3-9/HP1 complex differentially regulates the compaction state and degree of underreplication of X chromosome pericentric heterochromatin in Drosophila melanogaster. Genetics 175(2): 609-20. PubMed ID: 17151257

Foti, R., Gnan, S., Cornacchia, D., Dileep, V., Bulut-Karslioglu, A., Diehl, S., Buness, A., Klein, F. A., Huber, W., Johnstone, E., Loos, R., Bertone, P., Gilbert, D. M., Manke, T., Jenuwein, T. and Buonomo, S. C. (2016). Nuclear Architecture Organized by Rif1 Underpins the Replication-Timing Program. Mol Cell 61(2): 260-273. PubMed ID: 26725008

Hannibal, R. L. and Baker, J. C. (2016). Selective Amplification of the Genome Surrounding Key Placental Genes in Trophoblast Giant Cells. Curr Biol 26(2): 230-236. PubMed ID: 26774788

Kolesnikova, T. D., Makunin, I. V., Volkova, E. I., Pirrotta, V., Belyaeva, E. S. and Zhimulev, I. F. (2005). Functional dissection of the Suppressor of UnderReplication protein of Drosophila melanogaster: identification of domains influencing chromosome binding and DNA replication. Genetica 124(2-3): 187-200. PubMed ID: 16134332

Kolesnikova, T. D., Posukh, O. V., Andreyeva, E. N., Bebyakina, D. S., Ivankin, A. V. and Zhimulev, I. F. (2013). Drosophila SUUR protein associates with PCNA and binds chromatin in a cell cycle-dependent manner. Chromosoma 122(1-2): 55-66. PubMed ID: 23149855

Kolesnikova, T. D., Goncharov, F. P. and Zhimulev, I. F. (2018). Similarity in replication timing between polytene and diploid cells is associated with the organization of the Drosophila genome. PLoS One 13(4): e0195207. PubMed ID: 29659604

Koryakov, D. E. and Zhimulev, I. F. (2014). DNA replication in nurse cell polytene chromosomes of Drosophila melanogaster otu mutants. Chromosoma [Epub ahead of print]. PubMed ID: 25256561

Munden, A., Rong, Z., Sun, A., Gangula, R., Mallal, S. and Nordman, J. T. (2018). Rif1 inhibits replication fork progression and controls DNA copy number in Drosophila. Elife 7. PubMed ID: 30277458

Nordman, J. T., Kozhevnikova, E. N., Verrijzer, C. P., Pindyurin, A. V., Andreyeva, E. N., Shloma, V. V., Zhimulev, I. F. and Orr-Weaver, T. L. (2014). DNA copy-number control through inhibition of replication fork progression. Cell Rep 9: 841-849. PubMed ID: 25437540

Pindyurin, A. V., Boldyreva, L. V., Shloma, V. V., Kolesnikova, T. D., Pokholkova, G. V., Andreyeva, E. N., Kozhevnikova, E. N., Ivanoschuk, I. G., Zarutskaya, E. A., Demakov, S. A., Gorchakov, A. A., Belyaeva, E. S. and Zhimulev, I. F. (2008). Interaction between the Drosophila heterochromatin proteins SUUR and HP1. J Cell Sci 121(Pt 10): 1693-1703. PubMed ID: 18445687

Posukh, O. V., Maksimov, D. A., Laktionov, P. P., Koryakov, D. E. and Belyakin, S. N. (2017). Functional dissection of Drosophila melanogaster SUUR protein influence on H3K27me3 profile. Epigenetics Chromatin 10(1): 56. PubMed ID: 29191233

Schwaiger, M., Stadler, M. B., Bell, O., Kohler, H., Oakeley, E. J. and Schubeler, D. (2009). Chromatin state marks cell-type- and gender-specific replication of the Drosophila genome. Genes Dev 23(5): 589-601. PubMed ID: 19270159

Seller, C. A. and O'Farrell, P. H. (2018). Rif1 prolongs the embryonic S phase at the Drosophila mid-blastula transition. PLoS Biol 16(5): e2005687. PubMed ID: 29746464

Spierer, A., Seum, C., Delattre, M. and Spierer, P. (2005). Loss of the modifiers of variegation Su(var)3-7 or HP1 impacts male X polytene chromosome morphology and dosage compensation. J Cell Sci 118(Pt 21): 5047-5057. PubMed ID: 16234327

Yarosh, W. and Spradling, A. C. (2014). Incomplete replication generates somatic DNA alterations within Drosophila polytene salivary gland cells. Genes Dev 28(16): 1840-1855. PubMed ID: 25128500

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date revised: 9 January 2019

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