Histone supply regulates S phase timing and cell cycle progression

Eukaryotes package DNA into nucleosomes that contain a core of histone proteins. During DNA replication, nucleosomes are disrupted and re-assembled with newly synthesized histones and DNA. Despite much progress, it is still unclear why higher eukaryotes contain multiple core histone genes, how chromatin assembly is controlled, and how these processes are coordinated with cell cycle progression. We used a histone null mutation of Drosophila melanogaster to show that histone supply levels, provided by a defined number of transgenic histone genes, regulate the length of S phase during the cell cycle. Lack of de novo histone supply not only extends S phase, but also causes a cell cycle arrest during G2 phase, and thus prevents cells from entering mitosis. Our results suggest a novel cell cycle surveillance mechanism that monitors nucleosome assembly without involving the DNA repair pathways and exerts its effect via suppression of CDC25 phosphatase String expression. DOI: http://dx.doi.org/10.7554/eLife.02443.001


Introduction
Chromatin assembly during DNA replication is crucial for the repackaging of newly synthesized DNA and for maintaining or erasing histone modifications. During this process, pre-existing or so-called parental histones are recycled and assembled into nucleosomes together with de novo synthesized histones (Alabert and Groth, 2012;Annunziato, 2012). To compensate for the high demand of histone proteins during DNA replication, the canonical histones H1, H2A, H2B, H3, and H4, which are encoded by multiple gene copies in higher eukaryotes, are highly and exclusively expressed in S phase of the cell cycle (Marzluff et al., 2008).
The assembly of chromatin is mediated by an interplay of components of the DNA replication machinery and histone chaperones, which mediate the deposition of histones into nucleosomes (Alabert and Groth, 2012;Annunziato, 2012). Apparently, the pace of DNA synthesis is tightly coupled to the assembly of newly synthesized DNA into chromatin. Multiple studies showed that the depletion of the histone chaperones Asf1 and CAF-1 results in a slow down of DNA synthesis during S phase (Hoek and Stillman, 2003;Ye et al., 2003;Nabatiyan and Krude, 2004;Groth et al., 2007;Takami et al., 2007) preceding the accumulation of DNA damage in mammalian cells (Hoek and Stillman, 2003;Ye et al., 2003). Also, diminishing histone supply during S phase through knock down of SLBP, which is required for histone mRNA stability and translation, decreases the rate of DNA synthesis (Zhao et al., 2004). A recent study that targeted SLBP together with FLASH, a factor that is required for histone mRNA transcription and processing (Barcaroli et al., 2006;Yang et al., 2009), revealed that replication fork progression depends on nucleosome assembly potentially through a mechanism based on a feedback from the histone chaperone CAF-1 to the replicative helicase and/or the unloading of PCNA from newly synthesized DNA upon nucleosome assembly (Groth et al., 2007;Mejlvang et al., 2014).
The coupling of replication fork progression and nucleosome assembly might compensate for short-term fluctuations in histone availability (Mejlvang et al., 2014). However, it is still unclear whether chromatin integrity is monitored after or during DNA replication. Genome integrity during S phase is governed by the ATR/Chk1 and ATM/Chk2 checkpoint mechanisms that sense replication stress and DNA damage, respectively Cimprich and Cortez, 2008). Lack of CAF-1 or Asf1 function leads to accumulation of DNA damage and activation of the ATM/Chk2 pathway (Hoek and Stillman, 2003;Ye et al., 2003). These findings led to the hypothesis that chromatin assembly is monitored indirectly through accumulation of DNA lesions in response to stalled replication forks. However, since these chaperones have multiple functions such as unwinding of DNA during replication, in DNA repair (Gaillard et al., 1996;Green and Almouzni, 2003;Schöpf et al., 2012) as well as other nuclear processes (Quivy et al., 2004;Houlard et al., 2006). These multiple functions of these chaperones make it difficult to assess the direct effects of defective chromatin assembly.
Taking advantage of a histone null mutation in a higher eukaryote that recently became available in Drosophila melanogaster (Günesdogan et al., 2010), we directly addressed the requirement of canonical histone supply for DNA replication and cell cycle progression in a developing organism. By reintroducing a defined number of transgenic histone genes into the histone null mutant background, we show that the rate of DNA replication is coupled to the number of histone genes present in the genome and that histone supply is critical to coordinate S phase length with the developmental program. Surprisingly, cells that completely lacked de novo histone synthesis replicate DNA at a reduced rate, but complete S phase and arrest in cell cycle without accumulating DNA damage. This cell cycle arrest is mediated by suppressing the accumulation of transcripts encoding the CDC25 phosphatase String and provides evidence for a chromatin assembly surveillance mechanism that is independent of the known S phase checkpoints. eLife digest As a cell prepares to divide, it goes through four distinct stages. First, it grows in size (G1 phase); next it copies its entire DNA content (S phase); then it grows some more (G2 phase); and, last, it splits into two new cells (M phase).
During S phase, groups of histone proteins that normally stick together to tightly package the DNA are pulled apart in order to make the DNA accessible for copying. After the DNA has been duplicated, both copies of the DNA strand need to be repackaged. Therefore, after copying the DNA the cell rapidly reassembles the DNA-histone complexes (called nucleosomes), using a combination of old and newly synthesized histones to do so. A cell can adjust how quickly it copies DNA according to the availability of these histone proteins, which is important because copying DNA without the resources to package it could expose the DNA to damage.
Here, Günesdogan et al. investigate how a cell controls these processes using a mutant of the fruit fly Drosophila melanogaster that completely lacks the genes required to make histones. Cells that lack histones copy their DNA very slowly but adding copies of histone genes back into these flies speeds up the rate at which DNA is copied. Günesdogan et al. ask whether the slower speed of DNA replication in cells without new histones is connected to preventing DNA damage. However, these cells can still copy all their DNA, despite being unable to package it, so the higher risk of making mistakes is not enough to stop S phase. In fact, indications suggest that DNA damage detection methods continue to work as normal in cells without histones: these cells can get all the way to the end of G2 phase without any problems.
To go one step further and start splitting in two, a cell needs to switch on another gene, called string in the fruit fly and CDC25 in vertebrates, which makes an enzyme required for the cell division process. Normal cells switch on string during G2 phase, but cells that lack histones do not-and therefore do not enter M phase. Günesdogan et al. show that turning on string by a genetic trick is sufficient to overcome this cell cycle arrest and drive the cells into M phase. String could therefore form part of a surveillance mechanism that blocks cell division if DNA-histone complexes are not assembled correctly.

Results
The histone null mutation in D. melanogaster, called Df(2L)His C , lacks all genes encoding the canonical histones (Günesdogan et al., 2010). Df(2L)His C homozygous mutant animals (hereafter referred to as His C mutants) that are derived from heterozygous parents contain only maternal histone mRNA and proteins, which are sufficient to complete the first 14 cell division cycles of the embryo (Günesdogan et al., 2010). His C mutant embryos arrest before the onset of mitosis in cycle 15 (M 15 ) (Günesdogan et al., 2010). This highly uniform phenotype is likely due to the degradation of maternal histone mRNAs during the first G2 phase of embryogenesis in cell cycle 14 (Marzluff et al., 2008;O'Farrell et al., 1989) combined with the complete lack of zygotic histone gene expression during S phase of cell cycle 15 (S 15 ) (Günesdogan et al., 2010). In order to verify that the lack of histone transcription also results in a diminished pool of histone proteins in S 15 , we compared the protein levels of histone H2B and H3 of wild type embryos that are in S 15 at 4-5 hr after egg laying (AEL) to sorted His C mutant embryos that are still in S 15 at 5.5-6.5 hr AEL (see below and Günesdogan et al., 2010) by quantitative Western blotting ( Figure 1A,B). The approximate twofold reduction in the histone levels of His C mutant embryos is consistent with the fact that these embryos lack synthesis of new histones in S 15 but still contain parental histones from chromatin that was assembled during cycle 14. To test whether the reduced supply of histones in His C mutant embryos leads to a decrease in nucleosome formation, we carried out Micrococcal Nuclease (MNase) digestion assays on chromatin from sorted His C mutant and wild type sibling embryos ( Figure 1C,D, Figure 1-figure supplement 1). The results show that chromatin from His C mutant embryos is more accessible to MNase than control chromatin, leading to a more rapid generation of mononucleosomal DNA fragments and reflecting a decrease in nucleosome occupancy in chromatin of the His C mutants.
Arrested His C mutant cells express high levels of mitotic Cyclin B suggesting a cell cycle arrest in G2 phase of cycle 15 (G2 15 ) before mitosis (Günesdogan et al., 2010;Lehner and O'Farrell, 1990;Figure 2A). Consistent with this, we did not observe degradation of the mitotic Cyclin A or assembly of mitotic spindles in His C mutant cells (Figure 2-figure supplement 1). To assess DNA replication in S 15 , we used BrdU incorporation assays to label newly synthesized DNA ( Figure 2). The results show that most cells of His C mutant embryos entered S 15 , but the spatial pattern of replicating cells was different from the highly stereotyped wild type pattern ( Figure 2B-D). In wild type embryos, about 5 hr AEL ( Figure 2C,E), the ventral epidermal cells incorporated BrdU in S 15 , whereas the lateral cells entered G2 15 and stopped BrdU incorporation. Dorsal epidermal cells had already passed through M 15 and incorporated BrdU in S 16 . In contrast, the His C mutant cells failed to reach G2 15 and the majority of the cells continued to incorporate BrdU at low levels in S 15 ( Figure 2D,F). To test whether the extended S phase of His C mutant cells is caused by a reduced rate of DNA synthesis, we shortened the BrdU labelling pulses from 15 to 5 min. In wild type cells, BrdU incorporation was detectable during S 15 ( Figure 2I-K), whereas in His C mutant cells BrdU incorporation was detected only in a few cells with low Cyclin B levels indicating that they were still in early S 15 ( Figure 2L-N). Notably, in His C mutants ventral cells showed a uniform pattern and dorsal cells a punctuate pattern of BrdU incorporation ( Figure 2G,H), which is characteristic for the replication of euchromatin and heterochromatin in early and late S phase, respectively (Shermoen et al., 2010). Although we cannot exclude that the lack of histone synthesis in His C mutants interfered with firing of individual origins, the results suggest that both early and late replication origins are activated in His C mutants. In conclusion, the absence of de novo histone supply reduces the rate of DNA synthesis shortly after the initiation of DNA replication, resulting in an extended S phase in mutant cells.
It was previously shown that depletion of histones in cultured cells leads to a slow down of replication fork movement and it was proposed that this mechanism might avoid chromatin assembly defects due to short-term fluctuations in histone availability (Mejlvang et al., 2014). Thus, we asked whether S phase could be faithfully completed under diminished but constant histone supply and whether there is a direct dose dependent relation between histone synthesis and the length of S phase. We reintroduced defined numbers of histone gene units (His-GUs) into the genome of His C mutant embryos. Embryos carrying either two (2xHis-GU) or six (6xHis-GU) units completed cell cycle 15 as shown by the degradation of Cyclin B in M 15 ( Figure 3A,B). These embryos were not rescued and they died later towards the end of embryogenesis (Figure 3-figure supplements 1 and 2). Compared to wild type, the onset of M 15 was delayed in embryos with 2xHis-GUs and 6xHis-GUs by 2 hr and 1 hr, respectively, which was due to an extended S phase 15 ( Figure 3A-E, Figure 3-figure supplements 1 and 2). In contrast, Figure 1. Nucleosome density is affected in His C mutant cells. (A) Fluorescent Western blot analysis for histone H2B and H3 (green) using wild type embryos at 4-5 hr and sorted His C mutant embryos at 5.5-6.5 hr AEL, respectively. α-Tubulin (α-Tub; red) was used as loading control. A dilution series of each extract was loaded (1, 0.5, 0.25). Figure 1. Continued on next page embryos containing 12xHis-GUs were fully rescued and entered M 15 at the same time as the wild type cells, that is, 4.5-5 hr AEL as shown previously (Günesdogan et al., 2010). These results establish that the length of S phase directly correlates with the transgene-derived de novo histone supply. In addition, our data indicate that the mechanisms adjusting replication fork movement to the available histone supply allow the completion of S phase under conditions of permanently diminished new histone supply in vivo ( Figure 3F).
Several studies showed that mammalian tissue culture cells depleted for CAF-1 accumulate in S phase due to a decreased rate of DNA replication (Hoek and Stillman, 2003;Ye et al., 2003;Nabatiyan and Krude, 2004;Takami et al., 2007) followed by accumulation of DNA damage and activation of the conventional DNA damage checkpoints (Hoek and Stillman, 2003;Ye et al., 2003). However, it remained unclear whether this S phase arrest represents a direct consequence of a failure in chromatin assembly as yeast cells, for example, can complete one round of replication after the depletion of histone H4 (Kim et al., 1988). In order to address whether DNA replication can be completed in the absence of de novo histone synthesis, we quantified the amount of nuclear DNA in DAPI-stained His C mutant cells and compared it with wild type control cells during early S 16 (2N) and G2 15 (4N), respectively ( Figure 3G). The DNA content of His C mutant cells at 6.5-7.5 hr AEL corresponded to the 4N value of wild type nuclei in G2. This finding indicates that DNA replication in His C mutant cells has essentially been completed.
To further explore whether His C mutant cells accumulate DNA damage and activate the DNA damage checkpoints, we stained for the phosphorylated histone variant H2Av (γH2Av), which is like its vertebrate ortholog γH2AX a marker of DNA damage (Madigan et al., 2002). We found that γH2Av can be induced by ionizing irradiation that causes double strand breaks (DSBs) both in wild type and His C mutant embryos, showing that the ATM/Chk2 checkpoint mechanism is still functional in His C mutant cells ( Figure 4A,B). We did not observe a difference with respect to γH2Av staining between nonirradiated His C mutant and wild type cells in early S 15 ( Figure 4C,D). However, we noted a slight increase in γH2Av staining betwen early and late S 15 in His C mutant embryos (Figure 4-figure supplement 1). To further investigate this increase, we performed Western blot analysis for γH2Av. Drosophila S2R+ cells showed a dramatic increase in γH2Av upon treatment with hyrdoxyurea (HU) ( Figure 4E, Figure 4-figure supplement 1 and see below), which was not detectable in His C mutant embryos undergoing late S 15 at 5.5-6.5 hr AEL when compared to wild type sibling embryos ( Figure 4E).
To independently address the extend of DNA damage accumulation in His C mutant embryos, we used TUNEL assays which support that His C mutant cells do not accumulate significant levels of DNA damage until much later in development (≥10 hr AEL) when these embryos die (Figure 4-figure supplement 2). Finally, we performed genetic tests using mutants of loki (lok), the Drosophila DNA damage checkpoint kinase chk2 (Xu et al., 2001). Homozygous lok mutants are viable and fertile. In contrast, lok, His C double mutant embryos exhibited the His C phenotype ( Figure 4F-H). Hence, the cell cycle arrest in His C mutant embryos is not mediated by the chk2-dependent DNA damage checkpoint pathway.
Mutations in the Drosophila ortholog of Chk1 (GRP) show developmental defects prior to cell cycle 15, excluding genetic experiments as we performed for lok (Fogarty et al., 1994;Su et al., 1999). Thus, we tested ATR/Chk1 checkpoint activation by using a phosphospecific antibody that recognizes the ATR-dependent phosphorylation of S345 in human Chk1 in response to replicative stress, for (B) Quantification of Western blots as shown in (A). Fluorescence measurements of the histone signal was normalised to the α-Tubulin signal and the wild type (WT)/His C ratio is shown. The histone protein content is ∼twofold reduced in His C mutant embryos as compared to wild type. Mean values from three independent experiments are shown. Error bars indicate standard error. (C) Gel electrophoresis of time-course (0′-5′) microccocal nuclease (MNase) digestions using sorted His C mutant and wild type sibling embryos at 5.5-6.5 hr after egg laying, respectively. (D) Quantification of MNase digestion experiments as shown in (C). His C mutant chromatin is digested more rapidly into mononucleosomal DNA than control chromatin. Mean values from three independent experiments are shown. Error bars indicate standard error. DOI: 10.7554/eLife.02443.003 The following figure supplement is available for figure 1: To test whether the ATR/Chk1 checkpoint is functional in His C mutant embryos, we irradiated embryos with UV light (254 nm, UVC), which induces replication stress and replication fork uncoupling (Byun et al., 2005;Cimprich and Cortez, 2008). Wild type embryos and His C mutant embryos accumulated phosphorylated GRP protein (pGRP) in response to UVC, showing that the checkpoint response is functional in the mutant embryos ( Figure 5A-D). Without UVC treatment His C mutant embryos did not display elevated pGRP levels as compared to wild type ( Figure 5E-H), which was also verified by Western blotting of extracts from sorted His C mutant and wild type sibling embryos ( Figure 5I). In addition, treatment of His C mutant embryos with the ATR inhibitor VE-821 (Prevo et al., 2012) or the Chk1 inhibitor CHIR-124 (Tse et al., 2007) did not result in a release of the cell cycle arrest (Figure 5-figure supplements 3 and 4). In addition to DSBs, replicative stress can induce phosphorylation of H2Av (Figure 4-figure supplement 1K-P), either directly by ATR dependent phosphorylation (Ward and Chen, 2001;Joyce et al., 2011) or through interconversion of single-stranded DNA generated at stalled replication forks into DSBs (Cimprich and Cortez, 2008). Consistent with the notion that the DNA damage checkpoints are functional in His C mutant embryos we found accumulation of γH2Av in UVC-treated embryos to levels well above the background levels detected in untreated His C mutant embryos in late S 15 ( Figure 5-figure supplement 5). Interestingly, UVC-treated embryos were able to enter M 15 while displaying levels of γH2Av comparable or above to what we observed in untreated His C mutant embryos ( Figure 5-figure supplement 6). Taken together, these results strongly suggest that His C mutant cells complete DNA replication in S phase without inducing significant DNA damage or replication stress and that the cell cycle arrest at the G2/M transition in His C mutant cells is not mediated by the conventional S phase checkpoints. Progression from the G2 phase into mitosis critically depends on the dephosphorylation and activation of Cyclin/Cdk complexes, which is accomplished by a single gene in Drosophila embryos, encoding the CDC25 phosphatase String (Edgar and O'Farrell, 1990). In wild type, string mRNA accumulates during G2 phase and becomes rapidly degraded after cells exit mitosis (Edgar et al., 1994). string transcription is highly dynamic, dictates the pattern of cell divisions during embryogenesis and is controlled by the activity of developmentally regulated transcription factors binding to cis-regulatory sequences spread over >30 kb of the string locus (Edgar et al., 1994). In contrast to wild type embryos which accumulate string mRNA in G2 15 cells of the dorsal epidermis, His C mutant embryos failed to accumulate string in the corresponding cells although they showed normal more than one cell cycle behind as compared to wild type (see also    Figure 6A,B). Given the complex developmental regulation of string, we asked whether string transcription is disturbed in His C mutant embryos due to misregulation of key patterning genes. However, we observed normal temporal and spatial expression patterns of developmental genes such as the segmentation gene engrailed and the homeotic genes Ultrabithorax and Abdominal-B in His C mutant embryos ( Figure  6-figure supplement 1). Hence, the developmental programme progresses normally in His C mutant embryos up to the late embryonic stage when they eventually die (Figure 4-figure supplement 2). Similar to His C mutant embryos, string mRNA was not detectable in dorsal epidermal cells of 2xHis-GU embryos during their extended S 15 ( Figure 6C). However, when wild type embryos undergo M 16 at 6.5-7 hr AEL ( Figure 6D,E), His C mutant embryos still failed to express string ( Figure 6F,G) but 2xHis-GU embryos upregulated string expression ( Figure 6H,I). This result suggests that the failure of His C mutant embryos to upregulate string after they finished replication in S 15 is not simply a consequence of their extended S phase but rather due to a surveillance mechanism that blocks the G2/M transition because chromatin assembly is not completed. It is interesting to note that the pattern of M 15 in 2xHis-GU embryos closely resembled the pattern of M 16 in wild type embryos (Günesdogan et al., 2010;Figure 6D,H). This observation provides further support that the developmental programme of these embryos can progress normally, as string mRNA expression readjusts once DNA replication is completed. Thus, sufficient histone supply, as provided by multiple copies of His-GUs, is critical to the coordination of the developmental and cell division programmes during wild type embryogenesis of Drosophila. The results also explain why higher eukaryotes, which undergo rapid mitotic cell divisions during embryonic development, contain multiple His-GUs in their genomes.
If string activity is the only limiting factor that restricts cell cycle progression in His C mutant cells, its ectopic expression should drive the G2 to M transition. To test this hypothesis, His C mutant embryos were forced to express string in a striped pattern under the control of the prd-GAL4 driver using the GAL4/UAS system (Brand and Perrimon, 1993), which was visualised by coexpression of a UAS-EYFP transgene. The string expressing His C mutant cells entered M 15 , degraded Cyclin B and reaccumulated Cyclin B after mitosis, whereas cells lacking string expression remained arrested ( Figure 6J-P). Mitotic progression in wild type (E) Western blot for γH2Av using untreated (w/o HU) and HU-treated (HU) S2R+ cells as controls (two dilutions, 1 and 0.5) as well as His C mutant embryos and wild type embryos at 5.5-6.5 hr AEL. α-Tubulin (α-Tub) was used as a loading control. HU treatment results in a significant increase of γH2Av, which was not observed in His C mutant embryos. (F-H) BrdU pulse labelling for 15 min and staining with antibodies against BrdU and Cyclin B. lok, His C double mutant embryos showed a similar phenotype as His C mutant embryos (see Figure 1F) Research article embryos follows a stereotyped pattern characterized by (i) Cyclin B degradation and sister chromatid separation at the metaphase to anaphase transition, (ii) spindle elongation at the transition from anaphase-a into anaphase-b, (iii) the onset of chromatin decondensation in telophase, and eventually by (iv) cytokinesis ( Figure 6Q). The string-induced mitosis in His C mutant embryos was normal up to the metaphase to anaphase transition. During anaphase-b and telophase, however, we observed lagging chromosomes or chromatin bridges in almost all of the cells (95.8%, n = 24). These bridges were eventually resolved during cytokinesis, and the cells entered into interphase of cell cycle 16 ( Figure 6Q). Together, these data indicate that string transcription is indeed the limiting downstream factor that restricts cell cycle progression in the absence of de novo histone synthesis. In summary, our study shows that DNA replication and histone availability are tightly coupled. Lack of de novo histone synthesis causes a string-dependent cell cycle arrest in G2 phase, suggesting a novel chromatin assembly checkpoint monitoring chromatin integrity.

Discussion
We used a recently generated null mutation for canonical histones to address the consequences of histone deprivation during metazoan development. In addition to canonical histones, eukaryotes express histone variants that can replace canonical histones in a specific genomic context (Banaszynski et al., 2010). Our results show that these histone variants do not compensate for the lack of canonical histone synthesis with regard to chromatin assembly and cell cycle progression. This could be due to insufficient expression of variant histones from their endogenous promoters as it has been shown for the variant histone H3.3, which can fully replace its canonical counterpart, histone H3, but only if it is expressed from within a histone gene unit like the canonical histone (Hödl and Basler, 2012). Alternatively, it could reflect structural divergence of the histone variants as in the case of His2Av (van Daal et al., 1988) and dBigH1 (Perez-Montero et al., 2013). It will be interesting to test whether individual histone mutations, like a mutation in H2B which does not have a variant histone in Drosophila (Talbert et al., 2012), will cause a similar cell cycle arrest as the histone null mutation His C .
Our results provide evidence that canonical histone supply directly affects the rate of DNA synthesis ( Figure 2L-N). This observation is in line with studies that targeted either histone chaperones (Hoek and Stillman, 2003;Ye et al., 2003;Nabatiyan and Krude, 2004;Groth et al., 2007;Takami et al., 2007) or histone mRNA through SLBP or FLASH (Zhao et al., 2004;Barcaroli et al., 2006;Mejlvang et al., 2014) to interfere with chromatin assembly in tissue culture cells. However, previous work on SLBP in multicellular organisms revealed pleiotropic effects (Sullivan et al., 2001;Lanzotti et al., 2002;Pettitt et al., 2002). Our data illustrate that an extension of the S phase duration caused by diminished histone supply allows a faithful completion of S phase and transition from G2 into M phase of the cell cycle. This S phase extension is likely to be caused by a direct effect of lowered histone availability on replication fork progression (Groth et al., 2007;Mejlvang et al., 2014) and not by a lack of origin firing, although we cannot exclude this possibility completely. It was previously shown that postblastodermal development in Drosophila embryos proceeds largely uncoupled from progression through cell cycles 14-16 (Edgar et al., 1994;Meyer et al., 2002). Therefore, histone availability limits S phase duration and appears to be a critical link between cell division and development.
In the absence of de novo histone synthesis, we find that cells arrest in G2 phase of the cell cycle without activating the known ATM/Chk2 and ATR/Chk1 checkpoints. This observation is in contrast to previous studies on CAF-1, which found that cells arrest in S phase and accumulate DNA damage (Hoek and Stillman, 2003;Ye et al., 2003). This discrepancy might in part be explained by the fact that histone chaperones also have a direct function in DNA repair (Schöpf et al., 2012); and thus, in the presence of an intact DNA repair/chromatin assembly machinery in His C mutants, DNA is replicated without the accumulation of damage, even when histone supply is restricted to the parental load of histones. Alternatively, the accumulation of DNA damage in histone chaperone-depleted cells might be the consequence of a prolonged replication slow down, since it was shown that neither ATM/Chk2 nor ATR/Chk1 are activated as an immediate consequence of histone deprivation but only after prolonged incubation times (>48 hr) (Mejlvang et al., 2014). Based on our DNA quantification experiments, we found that the bulk of DNA replication in His C mutants is completed by about 2 hr  (E-H) Without irradiation, His C mutant cells did not show elevated staining for pChk1 compared to wild type, indicating that mutant cells did not activate the ATR/Chk1 checkpoint. (I) Western blot detecting pGRP by an antibody to phosphorylated Chk1 (pChk1) and α−Tubulin (α-Tub). Extracts were prepared from SR2+ tissue culture cells that were either untreated (w/o HU) or treated with HU (HU). Embryos were either sorted wild type controls or after entry into S phase, which might differ from the timeframe required to develop significant DNA damage. Interestingly, we find that His C mutant cells become TUNEL positive by about 6 hr after they enter S phase 15, which might reflect secondary DNA damage and/or cell death. Nevertheless, we found a moderate increase of γH2Av staining during late S phase in His C mutant embryos. Our data indicate, however, that cells that resolved UVC-induced DNA damage, and therefore entered mitosis can do so with levels of γH2Av comparable to those we observe in His C mutants. Thus, it is plausible that the slight increase in γH2Av in His C mutants could result from incomplete turnover of γH2Av rather than directly reflect DNA damage that could activate the S phase checkpoints. Turnover of γH2Av was shown to require the Tip60 chromatin-remodelling complex (Kusch et al., 2004), which may be affected by the altered chromatin structure in His C mutants. Alternatively, H2Av was recently shown to be phosphorylated independent of ATM/ATR by the chromosomal tandem kinase JIL-1 (Jin et al., 1999;Thomas et al., 2014), which may also be influenced by the changed chromatin topology in His C mutants.
Both, the ATM/Chk2 and ATR/Chk1 checkpoints are known to act on CDC25 phosphatases by phosphorylation and protein destabilization  and it was shown in Drosophila that string transcripts accumulate normally in embryos that suffered from DNA damage (Su et al., 2000). In contrast, we find that His C mutant cells fail to accumulate string transcripts when arrested in G2. This finding was surprising since it was shown that the temporal and spatial expression pattern of string is essentially unchanged in embryos that are arrested in G2 by mutations in string or in mitotic Cyclins (Edgar et al., 1994). Thus, this difference is likely due to the failure of His C mutant embryos to assemble chromatin, resulting in a diminished nucleosome density as shown by the presence of excess MNase hypersensitive DNA. Although we cannot rule out that the lower abundance of histone proteins itself directly contributes to the G2 arrest, this possibility seems unlikely since histone levels rapidly decrease in G2 cells where the chromatin assembly surveillance should act (Marzluff et al., 2008). It remains unclear how the presence of unassembled chromatin is linked to the regulation of string, but the effect is specific, since string transcript accumulation is the only limiting factor to overcome the G2 arrest in His C -mutant embryos. The subsequent mitosis in His C mutants is completed and cells enter into the next cell cycle. Given that His C mutant cells enter mitosis with presumably about half of the nucleosomes present in wild type chromatin, the mitotic defects like lagging anaphase chromosomes appear surprisingly mild. These defects could reflect problems in loading of structural components that are required for chromosome condensation and sister chromatid cohesion, like Cohesins and Condensins, which are proposed to require contact to chromatin rather than naked DNA (Bernard et al., 2001;Nonaka et al., 2002;Tada et al., 2011).
Taken together, our results suggest that incomplete chromatin assembly is monitored by a novel surveillance mechanism that can block cell cycle progression at the G2/M transition in Drosophila. Our findings now pave the way to address key questions regarding the orchestration of DNA synthesis and chromatin formation as well as the control of chromatin integrity during cell cycle progression.

Irradiation of embryos, hydroxyurea treatment, treatment with inhibitors
Embryos were collected on apple-juice agar plates and aged to 4-5.5 hr AEL (w 1118 , wild type) or 6-7 hr AEL (His C mutant embryos). This procedure yielded wild type embryos with cells in G2 15 and His C mutant embryos that were arrested in cell cycle progression. After irradiation at 60 Gray in a Torrex 150D (Astrophysics Research Corp., City of Industry, CA) embryos from both collections were aged for 20 min on the apple-juice agar plates and mixed before fixation. UV irradiation (254 nm) was done at 200 mJ/cm 2 as described (Zhou and Steller, 2003), and embryos were aged 45 min before fixation. His C mutant embryos were identified in the stained samples based on their cell division arrest phenotype.
For inhibitor treatment, we used the same procedure as for BrdU incorporation (Günesdogan et al., 2010), replacing the BrdU with 10 µM VE-821 (Selleckchem, Houston, TX) or 10 µM CHIR-124 (Selleckchem) and incubation of 45 min at RT before fixation. S2R+ tissue culture cell were cultured in Schneiders Medium (Life Technologies), with 10% Fetal Calf Serum. Hydroxyurea (Sigma-Aldrich) was added to a final concentration 10mM and incubated for 12 hr. Extracts were prepared in SDS sample buffer after addition of phosphatase inhibitors (PhosSTOP, Roche) and protease inhibitors (cOmplete EDTA-free, Roche) at approximately 5 × 10 8 cells per ml.

DNA quantification
Embryos were fixed by heat/methanol treatment and stained for Cyclin B. DNA was stained with DAPI (1:1000; Life Technologies). Cyclin B staining was used to distinguish homozygous mutant Df(2L)His C embryos and control embryos and to define the cell cycle stage of each cell. Stacks of nuclei were acquired with a 63× objective and a 10× optical zoom with a Leica TCS-SP5 AOBS confocal laserscanning microscope (z-axis increment: 0.1 µm, 8 bit images, 512 × 512 pixel, 400 Hz scan speed). The gain and offset were adjusted once and then used for one complete experiment, avoiding saturation. Fluorescent measurements were carried out using ImageJ software. To define nuclear circumferences, we used the 'isodata thresholding' algorithm followed by manual inspection. This threshold we used in a customized macro (Source code 1) that utilizes the 'connected threshold grower' plugin of the ImageJ 3D toolkit to determine nuclear staining intensities in all slices of the z-stack. The absolute nuclear fluorescence intensities were calculated by integration of individual nuclear fluorescence distributed over the image stack. For background detection five regions in between nuclei were analysed and their average was used for background subtraction.

TUNEL assay
TUNEL assays were done as described (Arama and Steller, 2006) with some deviations. DIG-labelled nucleotides were detected with a sheep anti-DIG antibody (Roche) and a donkey biotinylated antisheep secondary antibody (Jackson ImmunoResearch). For signal amplification, embryos were incubated for 45′ with ABC reagents (Vector Laboratories), followed by a 5′ incubation with TSA Flourescein reagents (Perkin Elmer) diluted 1:50. Embryos were mounted in ProlongGold (Life Technologies).