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.

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 (Bartek and Lukas, 2007; 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.

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 (M₁₅) (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 (S₁₅) (Günesdogan et al., 2010). In order to verify that the lack of histone transcription also results in a diminished pool of histone proteins in S₁₅, we compared the protein levels of histone H2B and H3 of wild type embryos that are in S₁₅ at 4–5 hr after egg laying (AEL) to sorted His^C mutant embryos that are still in S₁₅ 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 S₁₅ 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.

Figure 1 with 1 supplement see all

Download asset Open asset

Arrested His^C mutant cells express high levels of mitotic Cyclin B suggesting a cell cycle arrest in G2 phase of cycle 15 (G2₁₅) 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 S₁₅, we used BrdU incorporation assays to label newly synthesized DNA (Figure 2). The results show that most cells of His^C mutant embryos entered S₁₅, 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 S₁₅, whereas the lateral cells entered G2₁₅ and stopped BrdU incorporation. Dorsal epidermal cells had already passed through M₁₅ and incorporated BrdU in S₁₆. In contrast, the His^C mutant cells failed to reach G2₁₅ and the majority of the cells continued to incorporate BrdU at low levels in S₁₅ (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 S₁₅ (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 S₁₅ (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.

Figure 2 with 1 supplement see all

Download asset Open asset

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 M₁₅ (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 M₁₅ 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, embryos containing 12xHis-GUs were fully rescued and entered M₁₅ 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).

Figure 3 with 2 supplements see all

Download asset Open asset

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 S₁₆ (2N) and G2₁₅ (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 non-irradiated His^C mutant and wild type cells in early S₁₅ (Figure 4C,D). However, we noted a slight increase in γH2Av staining betwen early and late S₁₅ 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 S₁₅ at 5.5–6.5 hr AEL when compared to wild type sibling embryos (Figure 4E).

Figure 4 with 2 supplements see all

Download asset Open asset

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 example, UV irradiation or HU treatment (Zhao and Piwnica-Worms, 2001). This antibody is expected to cross-react with Drosophila GRP due to sequence similarity and detected a single band in Western blots (Figure 5—figure supplement 1) as well as a clear signal in immunofluorescence (Figure 5—figure supplement 2) upon HU treatment of Drosophila S2R+ tissue culture cells. 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 S₁₅ (Figure 5—figure supplement 5). Interestingly, UVC-treated embryos were able to enter M₁₅ 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.

Figure 5 with 6 supplements see all

Download asset Open asset

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 G2₁₅ cells of the dorsal epidermis, His^C mutant embryos failed to accumulate string in the corresponding cells although they showed normal upregulation of string expression in ventral epidermal cells prior to M₁₄ (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 S₁₅ (Figure 6C). However, when wild type embryos undergo M₁₆ 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 S₁₅ 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 M₁₅ in 2xHis-GU embryos closely resembled the pattern of M₁₆ 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.

Figure 6 with 1 supplement see all

Download asset Open asset

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 M₁₅, degraded Cyclin B and reaccumulated Cyclin B after mitosis, whereas cells lacking string expression remained arrested (Figure 6J–P). Mitotic progression in wild type 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.

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 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 (Bartek and Lukas, 2007) 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.

1. 1. Abdu U
   2. Brodsky M
   3. Schüpbach T

   (2002) Activation of a meiotic checkpoint during Drosophila oogenesis regulates the translation of Gurken through Chk2/Mnk

   Current Biology 12:1645–1651.

   https://doi.org/10.1016/S0960-9822(02)01165-X

   • Google Scholar
2. 1. Alabert C
   2. Groth A

   (2012) Chromatin replication and epigenome maintenance

   Nature Reviews Molecular Cell Biology 13:153–167.

   https://doi.org/10.1038/nrm3288

   • Google Scholar
3. 1. Annunziato AT

   (2012) Assembling chromatin: the long and winding road

   Biochimica et Biophysica Acta 1819:196–210.

   https://doi.org/10.1016/j.bbagrm.2011.07.005

   • Google Scholar
4. 1. Arama E
   2. Steller H

   (2006) Detection of apoptosis by terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling and acridine orange in Drosophila embryos and adult male gonads

   Nature Protocols 1:1725–1731.

   https://doi.org/10.1038/nprot.2006.235

   • Google Scholar
5. 1. Banaszynski LA
   2. Allis CD
   3. Lewis PW

   (2010) Histone variants in metazoan development

   Developmental Cell 19:662–674.

   https://doi.org/10.1016/j.devcel.2010.10.014

   • Google Scholar
6. 1. Barcaroli D
   2. Bongiorno-Borbone L
   3. Terrinoni A
   4. Hofmann TG
   5. Rossi M
   6. Knight RA
   7. Matera AG
   8. Melino G
   9. De Laurenzi V

   (2006) FLASH is required for histone transcription and S-phase progression

   Proceedings of the National Academy of Sciences of USA 103:14808–14812.

   https://doi.org/10.1073/pnas.0604227103

   • Google Scholar
7. 1. Bartek J
   2. Lukas J

   (2007) DNA damage checkpoints: from initiation to recovery or adaptation

   Current Opinion in Cell Biology 19:238–245.

   https://doi.org/10.1016/j.ceb.2007.02.009

   • Google Scholar
8. 1. Bernard P
   2. Maure JF
   3. Partridge JF
   4. Genier S
   5. Javerzat JP
   6. Allshire RC

   (2001) Requirement of heterochromatin for cohesion at centromeres

   Science 294:2539–2542.

   https://doi.org/10.1126/science.1064027

   • Google Scholar
9. 1. Brand A
   2. Perrimon N

   (1993)

   Targeted gene expression as a means of altering cell fates and generating dominant phenotypes

   Development 118:401–415.

   • Google Scholar
10. 1. Byun TS
    2. Pacek M
    3. Yee MC
    4. Walter JC
    5. Cimprich KA

    (2005) Functional uncoupling of MCM helicase and DNA polymerase activities activates the ATR-dependent checkpoint

    Genes & Development 19:1040–1052.

    https://doi.org/10.1101/gad.1301205

    • Google Scholar
11. 1. Cimprich KA
    2. Cortez D

    (2008) ATR: an essential regulator of genome integrity

    Nature Reviews Molecular Cell Biology 9:616–627.

    https://doi.org/10.1038/nrm2450

    • Google Scholar
12. 1. Edgar BA
    2. Lehman DA
    3. O'Farrell PH

    (1994)

    Transcriptional regulation of string (cdc25): a link between developmental programming and the cell cycle

    Development 120:3131–3143.

    • Google Scholar
13. 1. Edgar BA
    2. O'Farrell PH

    (1990) The three postblastoderm cell cycles of Drosophila embryogenesis are regulated in G2 by string

    Cell 62:469–480.

    https://doi.org/10.1016/0092-8674(90)90012-4

    • Google Scholar
14. 1. Fogarty P
    2. Kalpin RF
    3. Sullivan W

    (1994)

    The Drosophila maternal-effect mutation grapes causes a metaphase arrest at nuclear cycle 13

    Development 120:2131–2142.

    • Google Scholar
15. 1. Gaillard PH
    2. Martini EM
    3. Kaufman PD
    4. Stillman B
    5. Moustacchi E
    6. Almouzni G

    (1996) Chromatin assembly coupled to DNA repair: a new role for chromatin assembly factor I

    Cell 86:887–896.

    https://doi.org/10.1016/S0092-8674(00)80164-6

    • Google Scholar
16. 1. Green CM
    2. Almouzni G

    (2003) Local action of the chromatin assembly factor CAF-1 at sites of nucleotide excision repair in vivo

    The EMBO Journal 22:5163–5174.

    https://doi.org/10.1093/emboj/cdg478

    • Google Scholar
17. 1. Groth A
    2. Corpet A
    3. Cook AJ
    4. Roche D
    5. Bartek J
    6. Lukas J
    7. Almouzni G

    (2007) Regulation of replication fork progression through histone supply and demand

    Science 318:1928–1931.

    https://doi.org/10.1126/science.1148992

    • Google Scholar
18. 1. Günesdogan U
    2. Jäckle H
    3. Herzig A

    (2010) A genetic system to assess in vivo the functions of histones and histone modifications in higher eukaryotes

    EMBO Reports 11:772–776.

    https://doi.org/10.1038/embor.2010.124

    • Google Scholar
19. 1. Hödl M
    2. Basler K

    (2012) Transcription in the absence of histone H3.2 and H3K4 methylation

    Current Biology 22:2253–2257.

    https://doi.org/10.1016/j.cub.2012.10.008

    • Google Scholar
20. 1. Hoek M
    2. Stillman B

    (2003) Chromatin assembly factor 1 is essential and couples chromatin assembly to DNA replication in vivo

    Proceedings of the National Academy of Sciences of USA 100:12183–12188.

    https://doi.org/10.1073/pnas.1635158100

    • Google Scholar
21. 1. Houlard M
    2. Berlivet S
    3. Probst A
    4. Quivy JP
    5. Héry P
    6. Almouzni G
    7. Gérard M

    (2006) CAF-1 is essential for heterochromatin organization in pluripotent embryonic cells

    PLOS Genetics 2:e181.

    https://doi.org/10.1371/journal.pgen.0020181

    • Google Scholar
22. 1. Jacobs HW
    2. Knoblich JA
    3. Lehner CF

    (1998) Drosophila Cyclin B3 is required for female fertility and is dispensable for mitosis like Cyclin B

    Genes & Development 12:3741–3751.

    https://doi.org/10.1101/gad.12.23.3741

    • Google Scholar
23. 1. Jin Y
    2. Wang Y
    3. Walker DL
    4. Dong H
    5. Conley C
    6. Johansen J
    7. Johansen KM

    (1999) JIL-1: a novel chromosomal tandem kinase implicated in transcriptional regulation in Drosophila

    Molecular Cell 4:129–135.

    https://doi.org/10.1016/S1097-2765(00)80195-1

    • Google Scholar
24. 1. Joyce EF
    2. Pedersen M
    3. Tiong S
    4. White-Brown SK
    5. Paul A
    6. Campbell SD
    7. McKim KS

    (2011) Drosophila ATM and ATR have distinct activities in the regulation of meiotic DNA damage and repair

    The Journal of Cell Biology 195:359–367.

    https://doi.org/10.1083/jcb.201104121

    • Google Scholar
25. 1. Karr TL
    2. Alberts BM

    (1986) Organization of the cytoskeleton in early Drosophila embryos

    The Journal of Cell Biology 102:1494–1509.

    https://doi.org/10.1083/jcb.102.4.1494

    • Google Scholar
26. 1. Kim UJ
    2. Han M
    3. Kayne P
    4. Grunstein M

    (1988)

    Effects of histone H4 depletion on the cell cycle and transcription of Saccharomyces cerevisiae

    The EMBO Journal 7:2211–2219.

    • Google Scholar
27. 1. Knoblich JA
    2. Sauer K
    3. Jones L
    4. Richardson H
    5. Saint R
    6. Lehner CF

    (1994) Cyclin E controls S phase progression and its down-regulation during Drosophila embryogenesis is required for the arrest of cell proliferation

    Cell 77:107–120.

    https://doi.org/10.1016/0092-8674(94)90239-9

    • Google Scholar
28. 1. Kusch T
    2. Florens L
    3. Macdonald WH
    4. Swanson SK
    5. Glaser RL
    6. Yates JR III
    7. Abmayr SM
    8. Washburn MP
    9. Workman JL

    (2004) Acetylation by Tip60 is required for selective histone variant exchange at DNA lesions

    Science 306:2084–2087.

    https://doi.org/10.1126/science.1103455

    • Google Scholar
29. 1. Lanzotti DJ
    2. Kaygun H
    3. Yang X
    4. Duronio RJ
    5. Marzluff WF

    (2002) Developmental control of histone mRNA and dSLBP synthesis during Drosophila embryogenesis and the role of dSLBP in histone mRNA 3' end processing in vivo

    Molecular and Cellular Biology 22:2267–2282.

    https://doi.org/10.1128/MCB.22.7.2267-2282.2002

    • Google Scholar
30. 1. Lehner CF
    2. O'Farrell PH

    (1990) The roles of Drosophila cyclins A and B in mitotic control

    Cell 61:535–547.

    https://doi.org/10.1016/0092-8674(90)90535-M

    • Google Scholar
31. 1. Madigan JP
    2. Chotkowski HL
    3. Glaser RL

    (2002) DNA double-strand break-induced phosphorylation of Drosophila histone variant H2Av helps prevent radiation-induced apoptosis

    Nucleic Acids Research 30:3698–3705.

    https://doi.org/10.1093/nar/gkf496

    • Google Scholar
32. 1. Marzluff WF
    2. Wagner EJ
    3. Duronio RJ

    (2008) Metabolism and regulation of canonical histone mRNAs: life without a poly(A) tail

    Nature Reviews Genetics 9:843–854.

    https://doi.org/10.1038/nrg2438

    • Google Scholar
33. 1. Mejlvang J
    2. Feng Y
    3. Alabert C
    4. Neelsen KJ
    5. Jasencakova Z
    6. Zhao X
    7. Lees M
    8. Sandelin A
    9. Pasero P
    10. Lopes M
    11. Groth A

    (2014) New histone supply regulates replication fork speed and PCNA unloading

    The Journal of Cell Biology 204:29–43.

    https://doi.org/10.1083/jcb.201305017

    • Google Scholar
34. 1. Meyer CA
    2. Kramer I
    3. Dittrich R
    4. Marzodko S
    5. Emmerich J
    6. Lehner CF

    (2002)

    Drosophila p27Dacapo expression during embryogenesis is controlled by a complex regulatory region independent of cell cycle progression

    Development 129:319–328.

    • Google Scholar
35. 1. Nabatiyan A
    2. Krude T

    (2004) Silencing of chromatin assembly factor 1 in human cells leads to cell death and loss of chromatin assembly during DNA synthesis

    Molecular and Cellular Biology 24:2853–2862.

    https://doi.org/10.1128/MCB.24.7.2853-2862.2004

    • Google Scholar
36. 1. Nonaka N
    2. Kitajima T
    3. Yokobayashi S
    4. Xiao G
    5. Yamamoto M
    6. Grewal SI
    7. Watanabe Y

    (2002) Recruitment of cohesin to heterochromatic regions by Swi6/HP1 in fission yeast

    Nature Cell Biology 4:89–93.

    https://doi.org/10.1038/ncb739

    • Google Scholar
37. 1. O'Farrell PH
    2. Edgar BA
    3. Lakich D
    4. Lehner CF

    (1989) Directing cell division during development

    Science 246:635–640.

    https://doi.org/10.1126/science.2683080

    • Google Scholar
38. 1. Perez-Montero S
    2. Carbonell A
    3. Moran T
    4. Vaquero A
    5. Azorin F

    (2013) The embryonic linker histone H1 variant of Drosophila, dBigH1, regulates zygotic genome activation

    Developmental Cell 26:578–590.

    https://doi.org/10.1016/j.devcel.2013.08.011

    • Google Scholar
39. 1. Pettitt J
    2. Crombie C
    3. Schümperli D
    4. Müller B

    (2002)

    The Caenorhabditis elegans histone hairpin-binding protein is required for core histone gene expression and is essential for embryonic and postembryonic cell division

    Journal of Cell Science 115:857–866.

    • Google Scholar
40. 1. Prevo R
    2. Fokas E
    3. Reaper PM
    4. Charlton PA
    5. Pollard JR
    6. McKenna WG
    7. Muschel RJ
    8. Brunner TB

    (2012) The novel ATR inhibitor VE-821 increases sensitivity of pancreatic cancer cells to radiation and chemotherapy

    Cancer Biology & Therapy 13:1072–1081.

    https://doi.org/10.4161/cbt.21093

    • Google Scholar
41. 1. Quivy JP
    2. Roche D
    3. Kirschner D
    4. Tagami H
    5. Nakatani Y
    6. Almouzni G

    (2004) A CAF-1 dependent pool of HP1 during heterochromatin duplication

    The EMBO Journal 23:3516–3526.

    https://doi.org/10.1038/sj.emboj.7600362

    • Google Scholar
42. 1. Schöpf B
    2. Bregenhorn S
    3. Quivy JP
    4. Kadyrov FA
    5. Almouzni G
    6. Jiricny J

    (2012) Interplay between mismatch repair and chromatin assembly

    Proceedings of the National Academy of Sciences of USA 109:1895–1900.

    https://doi.org/10.1073/pnas.1106696109

    • Google Scholar
43. 1. Shermoen AW
    2. McCleland ML
    3. O'Farrell PH

    (2010) Developmental control of late replication and S phase length

    Current Biology 20:2067–2077.

    https://doi.org/10.1016/j.cub.2010.10.021

    • Google Scholar
44. 1. Su TT
    2. Campbell SD
    3. O'Farrell PH

    (1999) Drosophila grapes/CHK1 mutants are defective in cyclin proteolysis and coordination of mitotic events

    Current Biology 9:919–922.

    https://doi.org/10.1016/S0960-9822(99)80399-6

    • Google Scholar
45. 1. Su TT
    2. Walker J
    3. Stumpff J

    (2000) Activating the DNA damage checkpoint in a developmental context

    Current Biology 10:119–126.

    https://doi.org/10.1016/S0960-9822(00)00300-6

    • Google Scholar
46. 1. Sullivan E
    2. Santiago C
    3. Parker ED
    4. Dominski Z
    5. Yang X
    6. Lanzotti DJ
    7. Ingledue TC
    8. Marzluff WF
    9. Duronio RJ

    (2001) Drosophila stem loop binding protein coordinates accumulation of mature histone mRNA with cell cycle progression

    Genes & Development 15:173–187.

    https://doi.org/10.1101/gad.862801

    • Google Scholar
47. 1. Tada K
    2. Susumu H
    3. Sakuno T
    4. Watanabe Y

    (2011) Condensin association with histone H2A shapes mitotic chromosomes

    Nature 474:477–483.

    https://doi.org/10.1038/nature10179

    • Google Scholar
48. 1. Takami Y
    2. Ono T
    3. Fukagawa T
    4. Shibahara K
    5. Nakayama T

    (2007) Essential role of chromatin assembly factor-1-mediated rapid nucleosome assembly for DNA replication and cell division in vertebrate cells

    Molecular Biology of the Cell 18:129–141.

    https://doi.org/10.1091/mbc.E06-05-0426

    • Google Scholar
49. 1. Talbert PB
    2. Ahmad K
    3. Almouzni G
    4. Ausio J
    5. Berger F
    6. Bhalla PL
    7. Bonner WM
    8. Cande WZ
    9. Chadwick BP
    10. Chan SW
    11. Cross GA
    12. Cui L
    13. Dimitrov SI
    14. Doenecke D
    15. Eirin-López JM
    16. Gorovsky MA
    17. Hake SB
    18. Hamkalo BA
    19. Holec S
    20. Jacobsen SE
    21. Kamieniarz K
    22. Khochbin S
    23. Ladurner AG
    24. Landsman D
    25. Latham JA
    26. Loppin B
    27. Malik HS
    28. Marzluff WF
    29. Pehrson JR
    30. Postberg J
    31. Schneider R
    32. Singh MB
    33. Smith MM
    34. Thompson E
    35. Torres-Padilla ME
    36. Tremethick DJ
    37. Turner BM
    38. Waterborg JH
    39. Wollmann H
    40. Yelagandula R
    41. Zhu B
    42. Henikoff S

    (2012) A unified phylogeny-based nomenclature for histone variants

    Epigenetics & Chromatin 5:7.

    https://doi.org/10.1186/1756-8935-5-7

    • Google Scholar
50. 1. Thomas CJ
    2. Kotova E
    3. Andrake M
    4. Adolf-Bryfogle J
    5. Glaser R
    6. Regnard C
    7. Tulin AV

    (2014) Kinase-mediated changes in nucleosome conformation trigger chromatin decondensation via poly(ADP-ribosyl)ation

    Molecular Cell 53:831–842.

    https://doi.org/10.1016/j.molcel.2014.01.005

    • Google Scholar
51. 1. Tse AN
    2. Rendahl KG
    3. Sheikh T
    4. Cheema H
    5. Aardalen K
    6. Embry M
    7. Ma S
    8. Moler EJ
    9. Ni ZJ
    10. Lopes de Menezes DE
    11. Hibner B
    12. Gesner TG
    13. Schwartz GK

    (2007) CHIR-124, a novel potent inhibitor of Chk1, potentiates the cytotoxicity of topoisomerase I poisons in vitro and in vivo

    Clinical Cancer Research 13:591–602.

    https://doi.org/10.1158/1078-0432.CCR-06-1424

    • Google Scholar
52. 1. van Daal A
    2. White EM
    3. Gorovsky MA
    4. Elgin SC

    (1988) Drosophila has a single copy of the gene encoding a highly conserved histone H2A variant of the H2A.F/Z type

    Nucleic Acids Research 16:7487–7497.

    https://doi.org/10.1093/nar/16.15.7487

    • Google Scholar
53. 1. Ward IM
    2. Chen J

    (2001) Histone H2AX is phosphorylated in an ATR-dependent manner in response to replicational stress

    The Journal of Biological Chemistry 276:47759–47762.

    https://doi.org/10.1074/jbc.M009785200

    • Google Scholar
54. 1. Xu J
    2. Xin S
    3. Du W

    (2001) Drosophila Chk2 is required for DNA damage-mediated cell cycle arrest and apoptosis

    FEBS Letters 508:394–398.

    https://doi.org/10.1016/S0014-5793(01)03103-9

    • Google Scholar
55. 1. Yang XC
    2. Burch BD
    3. Yan Y
    4. Marzluff WF
    5. Dominski Z

    (2009) FLASH, a proapoptotic protein involved in activation of caspase-8, is essential for 3' end processing of histone pre-mRNAs

    Molecular Cell 36:267–278.

    https://doi.org/10.1016/j.molcel.2009.08.016

    • Google Scholar
56. 1. Ye X
    2. Franco AA
    3. Santos H
    4. Nelson DM
    5. Kaufman PD
    6. Adams PD

    (2003) Defective S phase chromatin assembly causes DNA damage, activation of the S phase checkpoint, and S phase arrest

    Molecular Cell 11:341–351.

    https://doi.org/10.1016/S1097-2765(03)00037-6

    • Google Scholar
57. 1. Zhao H
    2. Piwnica-Worms H

    (2001) ATR-mediated checkpoint pathways regulate phosphorylation and activation of human Chk1

    Molecular and Cellular Biology 21:4129–4139.

    https://doi.org/10.1128/MCB.21.13.4129-4139.2001

    • Google Scholar
58. 1. Zhao X
    2. McKillop-Smith S
    3. Müller B

    (2004) The human histone gene expression regulator HBP/SLBP is required for histone and DNA synthesis, cell cycle progression and cell proliferation in mitotic cells

    Journal of Cell Science 117:6043–6051.

    https://doi.org/10.1242/jcs.01523

    • Google Scholar
59. 1. Zhou L
    2. Steller H

    (2003) Distinct pathways mediate UV-induced apoptosis in Drosophila embryos

    Developmental Cell 4:599–605.

    https://doi.org/10.1016/S1534-5807(03)00085-6

    • Google Scholar

Author details

1. Ufuk Günesdogan

   1. Abteilung Molekulare Entwicklungsbiologie, Max-Planck-Institut für biophysikalische Chemie, Göttingen, Germany
   2. Wellcome Trust/Cancer Research UK Gurdon Institute, University of Cambridge, Cambridge, United Kingdom

   Contribution

   UG, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   No competing interests declared.

2. Herbert Jäckle

   Abteilung Molekulare Entwicklungsbiologie, Max-Planck-Institut für biophysikalische Chemie, Göttingen, Germany

   Contribution

   HJ, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   HJ: Herbert Jäckle has been Vice President of the Max Planck Society–one of eLife's three founding funders–and has attended until recently several meetings of eLife's board of directors.

3. Alf Herzig

   1. Abteilung Molekulare Entwicklungsbiologie, Max-Planck-Institut für biophysikalische Chemie, Göttingen, Germany
   2. Abteilung Zelluläre Mikrobiologie, Max-Planck-Institut für Infektionsbiologie, Berlin, Germany

   Contribution

   AH, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   herzig@mpiib-berlin.mpg.de

   Competing interests

   No competing interests declared.

• 6,123

  views

• 553

  downloads

• 63

  citations

Views, downloads and citations are aggregated across all versions of this paper published by eLife.