Regulation of mitosis in response to damaged or incompletely replicated DNA require different levels of Grapes (Drosophila Chk1)

Eukaryotic cells employ a variety of surveillance mechanisms or checkpoints that detect damage to DNA and act to prevent the propagation of damage to the next generation by slowing cell cycle progress, and activating repair and apoptotic pathways (Zhou and Elledge, 2000). The canonical DNA damage checkpoint is made of four conserved checkpoint kinases that regulate cell cycle progress after DNA damage and incomplete replication. Two phosphatidylinositol 3-kinase (PI3K)-like protein kinases (PI3KKs), ATR and ATM, transduce signals to Chk1 and Chk2, which then act on a number of effectors (Motoyama and Naka, 2004; Rhind and Russell, 2000).

In response to different types of damage, these four checkpoint kinases are used in different combinations that vary among different model organisms (Melo and Toczyski, 2002; Rhind and Russell, 2000). For instance, Chk1 homologs in yeast seem to be mainly required for DNA damage checkpoints, but play redundant roles in DNA replication checkpoints. In S. cerevisiae, scChk1 is required for Securin-dependent regulation of metaphase-anaphase transition after DNA damage, but as of yet, has no established role in responding to incomplete DNA replication (Rhind and Russell, 2000; Sanchez et al., 1999). Similarly in S. pombe, spChk1 is required for cell cycle arrest after exposure to UV and ionizing radiation (IR), but not upon treatment with hydroxy urea, an inhibitor of DNA replication (Brondello et al., 1999; Walworth and Bernards, 1996). In both yeast systems, Chk1 appears to act downstream of the ATR homologs, Rad3 and Mec1 in checkpoint signaling pathways (Rhind and Russell, 2000). By contrast, the opposite trend is evident in Xenopus where Xchk1 is activated after exposure to UV or treatment with aphidicolin, an inhibitor of DNA replication, but not in response to double strand DNA ends (Guo et al., 2000). However, similar to yeast, Xchk1 acts exclusively downstream of Xatr (Guo et al., 2000). In mammals, whether Chk1 acts together with ATM or ATR may depend on the type of damage (Liu et al., 2000; Motoyama and Naka, 2004; Takai et al., 2000); Chk1 is activated in an ATR-dependent manner after UV and HU treatments, whereas ATM is hypothesized to be the upstream activator of Chk1 after IR treatment. In any case, it appears that in mammals, Chk1 is involved in replication and DNA damage checkpoints.

Adding another dimension to this complex picture is the fact that cell cycle regulation by checkpoints changes dramatically during metazoan development. Drosophila melanogaster has been an ideal model system to document such changes because of the well-characterized cell cycles that occur during development, the genetic tools that allow mutational analysis with both spatial and temporal control and because detailed cell biological characterizations are possible. We know, for example, that regulation of mitosis in response to damaged DNA can differ according to embryonic stage. During rapid cleavage cycles that occur in a common syncytium, so-called `syncytial cycles', DNA damage results not in a delay of entry into mitosis, but in a Chk2-dependent inactivation of centrosomes and disruption of mitotic chromosome segregation (Sibon et al., 2000). Thirteen syncytial cycles are followed by cellularization of the embryo, onset of general zygotic transcription and gastrulation, and the introduction of the first G₂ phase into cell cycles (Foe et al., 1993). During these cell cycles, DNA damage now results in a checkpoint-dependent delay of entry into mitosis (Su et al., 2000). Thus, regulation of the cell cycle by checkpoints can drastically change during development.

Drosophila Chk1, encoded by grp, along with the ATR homolog, mei-41, are required to delay the entry into mitosis when DNA replication is incomplete during syncytial cycles (Fogarty et al., 1997; Sibon et al., 1999) (Table 1). Neither grp nor mei-41, however, is needed for DNA damage responses in syncytial cycles (Sibon et al., 2000) (Table 1). After cellularization, cells of the embryonic ectoderm undergo three additional cell cycles (cycles 14-16) during gastrulation and arrest in G₁ until the embryo hatches into a larva. Both mei-41 and grp are required to delay mitosis in response to damaged DNA in the larva (Brodsky et al., 2000) (Table 1), and the role of mei-41 has been documented for the DNA damage and replication checkpoint in gastrulating embryos (Table 1). The role of grp in either DNA damage or replication checkpoints in the gastrula stage has not been investigated before and forms the focus of this work.

Previously, we have shown that cells of the Drosophila gastrula can regulate both the entry into and progress through mitosis in response to either incomplete DNA replication or DNA damage by IR (X-rays) or an alkylating agent (MMS) (Su and Jaklevic, 2001; Su et al., 2000). Here, we use live measurement of mitotic phase duration to establish that regulation of metaphase-anaphase transition (mitotic progress) occurs in response to IR during all three gastrula cell cycles. We find that grp¹ mutants are capable of regulating both the entry into mitosis and the metaphase-anaphase transition in response to IR, but are unable to delay either transition when DNA replication is inhibited (Table 1). These results indicate that DNA damage and DNA replication checkpoints during the same cell cycle at the same stage in development have distinct requirements for a Chk1 homolog.

All mutant alleles used here, mei-41^D12, grp¹, bubR1^KO3110, dup^a1 and dup^a3, have been described before (Fogarty et al., 1997; Laurencon et al., 2003; Whittaker et al., 2000). Fly stocks carrying Histone H2Av-GFP transgene has also been described previously (Clarkson and Saint, 1999). The chromosomal deficiency uncovering grp, Df(2L)H2O, was obtained from Bloomington stock center as was the deficiency uncovering dup, Df(2R)Jp1. Double mutants were generated by standard Drosophila techniques.

Polyclonal antibodies to Grp were generated at a commercial facility (Cocalico Biologicals Inc., Reamstown, PA). The C-terminus of Grp (amino acid 261 to the C-terminus) was expressed in bacteria and used as antigen. Antisera were purified against recombinant antigen immobilized onto nitrocellulose. Detailed purification protocol is available upon request.

To prepare extracts for western blotting, embryos were dechorinated with 50% bleach, washed extensively in double-distilled water, and homogenized in extract buffer (25 mM HEPES pH 7.6, 0.1 M EDTA, 12.5 mM MgCl2, 150 mM KCl, 10% glycerol, 150 mM KCl, 2 mM Na₃VO₄, 1 mM sodium fluoride, 10 mM β-glycerol-phosphate, 1 mM sodium metabisulfite, 1 mM benzamidine, 5 μg ml^-1 aprotinin, 5 μg ml^-1 leupeptin, 1.5 mM DTT). An equal volume of 2×SDS sample buffer was added to the sample and boiled for 10 minutes before separation by SDS-PAGE and western blotting. The blot was blocked with PBT (0.2%Tween-20 + PBS, 140 mM NaCl, 10 mM Na₂HPO₄, 2.6 mM KCl, 1.8 mM KH₂PO4) containing 5% milk, probed with affinity-purified anti-Grp antibody diluted 1:200 in PBT, washed in blocking solution, probed with HRP-conjugated secondary antibody (Amersham) diluted 1:2000 in PBT, and visualized by ECL (Supersignal; Pierce, Rockford IL). Homozygous grp¹ mutant embryos from heterozygous mothers were identified by the lack of GFP signal encoded by the balancer chromosome and processed for western blotting as above.

To examine Grapes localization, wild type (Sevelen) embryos were dechorinated in 50% bleach, washed extensively in double-distilled water and fixed for 20 minutes at room temperature in a two-phase mixture of heptane and PBS containing 10% formaldehyde. Embryos were devitallinized in methanol, washed with PBT, and blocked in PBT containing 10% normal goat serum (NGS). Embryos were stained with affinity-purified anti-Grp antibody diluted 1:50 in blocking solution. HRP-conjugated secondary antibody was used at 1:500 dilution in blocking solution and was visualized by tyramide amplification according to manufacturer's instructions (Perkin-Elmer Life Sciences, Boston MA). The embryos were also stained with Texas-Red-conjugated wheat germ agglutinin (WGA) to visualize the nuclear envelope and with Hoechst33258, at 10 μg ml^-1 in PBT, to visualize the DNA.

To compare immunofluoresence signals for Grp, embryos were collected from wild-type parents carrying a GFP transgene or grp¹ homozygyous parents and fixed as described above. The grp¹ mutant (GFP-negative) and wild-type (GFP-positive) embryos were mixed together prior to fixing and processed for Grp immunofluoresence in the same tube. This excludes sample-to-sample variations in antibody staining and allows a direct comparison of wild-type and mutant embryos on the same microscope slide.

For analysis of dup and grp dup double mutants, embryos were collected for 30 minutes and aged for various amounts of time from 4.5 to 10.5 hours, dechorinated, and fixed as above with 3% formaldehyde in PBS. Fixed embryos were blocked in PBT containing 10% NGS and stained with a rabbit antibody to phosphorylated histone H3 (Upstate Biologicals), PH3, to visualize mitotic cells and a mouse anti-βgal antibody (Roche) to identify homozygous mutants that lack β-gal encoded by the balancer chromosome. PH3 was visualized by staining with a Rhodamine-conjugated secondary antibody diluted from 1:50 to 1:150 (Jackson). β-Gal was identified by staining with an HRP-conjugated secondary antibody diluted 1:2000, followed by Tyramide-FITC amplification. Embryos were also counterstained with the DNA dye, Hoechst33258.

For embryos irradiated in early interphase: for live analysis of metaphase length, embryos were collected for 30 minutes and aged for 330 minutes to reach interphase of cycle 16 (Foe et al., 1993). Embryos were irradiated with 8.3 Gy of X-rays in a TORREX X-ray generator (Astrophysics Research) set at 5 mA and 115 kV, and then allowed to rest for 30 minutes. Embryos were dechorionated as described above and transferred to a glass coverslip with a soft brush. A thin layer of halocarbon 700-oil (Sigma) was placed over embryos and the coverslip was inverted and placed over a hole in the center of a microscope slide such that oil-covered embryos were otherwise exposed to air. Live analysis began as close to 40 minutes after IR as possible. Time-lapsed images were taken with 100× objective lens on a Leica microscope using spinning-disc confocal illumination and Ultraview software for image analysis (PerkinElmer). Images were taken every 30 seconds up to 10 minutes at 150 milliseconds exposure.

For embryos irradiated in late interphase/early mitosis, embryos were collected and aged as above. However, embryos were transferred to a coverslip and prepared for visualization prior to irradiation to minimize the time between irradiation and analysis (approximately ∼2-8 minutes).

Homozygous embryos from heterozygous mothers were identified by lack of immuno-staining for β-gal encoded by the balancer chromosome. After live analysis, the halocarbon oil was removed by rolling the embryo on a glass coverslip. Each embryo was fixed separately and blocked in PBT containing 10% normal goat serum, stained for 60 minutes with the primary antibody against β-gal diluted 1:300 in blocking solution, washed in blocking solution and stained with FITC-conjugated secondary antibody diluted 1:50 in blocking solution. Embryos were also counterstained with the DNA dye, Hoechst33258.

For mitotic index, embryos were collected for 10 minutes, aged for 325 minutes at 25°C to reach interphase of embryonic cycle 16 (Foe et al., 1993), irradiated, and allowed to rest for 20 minutes before fixing in 37% formaldehyde for 5 minutes. For metaphase index, embryos were collected for 60 minutes and aged for 330 minutes at 25°C before fixing (for non-irradiated controls) or aged for 310 minutes, irradiated, and rested for 40 minutes to allow cells to enter mitosis before fixing. Embryos were irradiated with 8.3 Gy of X-rays. Fixed embryos were blocked in PBT containing 3%NGS and stained with a rabbit anti-PH3 to visualize mitotic cells and a mouse anti-βgal antibody to identify homozygous mutants. Embryos were also counterstained with Hoechst33258 as above.

P-values were generated using the standard Student's t-test available with Microsoft Excel.

Live analyses show a delay in the metaphase-anaphase transition in response to DNA damage in all three cellular cycles. Drosophila embryogeneis commences with 13 nuclear division cycles that consists of S and M phases only. These are followed by cellularization of the embryo and three additional cell cycles (cycles 14-16) that consist of S, G₂ and M phases. Most cells of the embryonic ectoderm then arrest in G₁ until the embryo hatches into a larva. We reported previously that DNA damage by X-rays or MMS affects two cell cycle transitions in cellularized Drosophila embryos. The first affects the entry into mitosis, whereas the second affects the metaphase-anaphase transition and requires Cyclin A, a known inhibitor of the metaphase-anaphase transition (Su and Jaklevic, 2001; Su et al., 2000). The delay in metaphase-anaphase transition was expressed as an increase in metaphase index of fixed embryos, which is defined as the ratio of metaphase cells to those in anaphase and telophase (M/A+T). This response was documented in the first two cell cycles of the cellularized embryo, cycles 14 and 15, and has been corroborated in direct measurement of metaphase length during cycle 14 in live embryos expressing a GFP-tagged Histone H2Av transgene (Su and Jaklevic, 2001). Here, we first confirmed the regulation of metaphase-anaphase transition in cycle 15 using live analysis and, second, extended it to cycle 16 (Fig. 1A,B). All three cellular cycles of the embryonic ectoderm now appear to show this response, allowing us to analyze mutants in the latest cellular cycle of embryogenesis. This is especially useful for studies of homozygous embryos from heterozygous mothers that deposit wild-type gene products into the embryo; we can, thereby, maximize the depletion of deposited maternal product.

Previously we found that mei-41 (Drosophila ATR) is required to delay metaphase-anaphase transition after DNA damage in cellularized embryos (Laurencon et al., 2003) and Table 1). In those studies, mei-41^D12 mutants, in contrast to wild type, failed to show a robust increase in metaphase index in embryos fixed after DNA damage. The mei-41^D12 allele is a hypomorphic loss of function allele that can be maintained as a homozygous stock. The mei-41 null mutants are maternal-effect lethal; embryos from homozygous null mutant mothers die during syncytial nuclear divisions and do not reach cellular stages. By contrast, embryos from mei-41^D12 homozygous mothers progress to cellular stages where metaphase-anaphase delay has been documented. Here we use live analysis to corroborate our conclusions from analysis of fixed embryos. Cells of the dorso-lateral ectoderm (e.g. enclosed in brackets in Fig. 3A) are analyzed in these studies because of their tendency to lie flat on a coverslip. The increase in length of metaphase in these cells after DNA damage is significantly shorter in mei-41^D12 mutants than in wild type, confirming that mei-41 is required to delay the metaphase-anaphase transition after DNA damage (Fig. 1C and Table 1).

Most mammalian cells do not show a delay in metaphase-anaphase transition following DNA damage, and this has led to the suggestion that the delay observed in Drosophila embryos is specific to Drosophila (Mikhailov et al., 2002; Smits et al., 2000). We address here the alternate explanation that these differences are instead due to differences in the radiation dose and the cell cycle phase at the time of irradiation. In our experiments, we irradiated 310-370-minute-old embryos in which lateral ectodermal cells are in cell cycle 16, with 8.3 Gy, which represents a repairable dose since around 20% of the treated embryos survive (our unpublished data) and will be called `LD₈₀'. We observed the most reproducibly robust increase in metaphase index at 40 minutes after irradiation (Fig. 2A). These data suggested that embryos must be irradiated during a specific phase of the cell cycle that occurs approximately 40 minutes prior to mitosis, to produce a subsequent delay in mitotic progress. Cells of the lateral ectoderm divide asynchronously, but the length of each cell cycle appears to be similar among cells and from embryo to embryo (Foe et al., 1993). The 15th and 16th cell cycles are composed of M phase (10 minutes), directly followed by S phase (approximately 35-45 minutes) and a well-defined G₂ phase (approximately 10 minutes) (Foe et al., 1993) (Fig. 2B). Following irradiation, interphase (G₂+S) increases by about 10 minutes; whether this is due to a slowing of S phase or increase in G₂ length is unknown (Su et al., 2000). This information and our data from analysis of fixed embryos (Fig. 2A) led us to hypothesize that cells must be 40 minutes prior to mitosis, i.e. in S phase, at the time of irradiation to produce a subsequent delay in metaphase-anaphase transition. To address this more directly in live analysis, embryos were irradiated and allowed to rest for 40 minutes prior to analysis of metaphase length (to examine those in S phase at the time of irradiation) or analyzed without a rest and after only the time needed for processing, which averaged about 5 minutes (to examine those in late G₂/early M at the time of irradiation). We find that the first set of embryos, which were irradiated in S phase, showed a delay in metaphase-anaphase transition in mitosis whereas the second set that were irradiated in late G₂/early M, did not (Fig. 2C). These data suggest that damage incurred in late G₂/early M cannot cause a delay in metaphase-anaphase transition, whereas the same level of radiation applied earlier in the cell cycle can. These observations are in agreement with results from mammalian cells in which doses of radiation that readily produced a G₂/M delay if applied in interphase had no effect on mitotic progress if applied in early M (Rieder and Cole, 1998). It is possible that damage caused by radiation at G₂/M is not recognized by the checkpoint or that the damage is sensed but that cells are unable to activate the checkpoint at such a late point in the cell cycle.

In the majority of mammalian cells, irradiation in late G₂/early M with extremely high doses (approximately 8-16-fold over what is needed to elicit a G₂/M delay if applied earlier) does induce a subsequent delay in mitotic progress (Mikhailov et al., 2002). This delay is independent of the DNA checkpoint functions ATM and ATR, and appears to be mediated by damaged centromeres/kinetochores because it requires a functional spindle checkpoint. To determine if we could induce a similar delay in mitotic progress in Drosophila simply by increasing the dose of radiation, we irradiated wild-type embryos in late G₂/early M with a 10-fold higher dose of X-rays (10×LD₈₀). In contrast to cells irradiated in late G₂/early M with a 10×LD₈₀ dose, cells irradiated with 10×LD₈₀ lingered in mitosis (Fig. 2C). Furthermore, the delay in mitotic progress caused by 10×LD₈₀ still occurred in mei-41^D12 embryos that are unable to delay metaphase-anaphase transition after irradiation in S phase with LD₈₀ (Fig. 2D). We conclude that the regulation of mitotic progress after irradiation in late G₂/early M is not dependent on a mei-41-mediated checkpoint as the regulation in response to irradiation earlier in the cell cycle is. We used mutants in bubR1 to address the to role of the spindle checkpoint in the mitotic delay caused by 10×LD₈₀. Unfortunately, bubR1 mutants have significant pre-existing defects in mitotic progress without irradiation treatment, thus precluding unambiguous interpretation of the data.

To further analyze the mei-41-dependent checkpoint that regulates mitotic progress after DNA damage, we examined mutants in the Chk1 homolog, grp. Grp is required during early embryogenesis and larval cycles to delay cell cycle progress: grp acts in the same pathway as mei-41 in responding to incomplete replication in syncytial embryos (Sibon et al., 1999) (Table 1). Like mei-41, grp is also needed to regulate the entry into mitosis in response to ionizing radiation in the larvae (Brodsky et al., 2000; Jaklevic and Su, 2004) (Table 1). To address the role of Grp in the DNA damage checkpoint in cellular stage embryos, we quantified mitotic index (for regulation of mitotic entry) and metaphase length (for regulation of mitotic progress) in grp¹ mutants after irradiation.

The grp¹ allele is the strongest extant allele of grp. Homozygous or hemizygous grp¹ animals from heterozygous parents survive to adulthood, presumably by using maternally supplied gene products. These females, however, are sterile; they produce embryos that fail to complete syncytial cycles and do not progress into the cellular stage (Sibon et al., 1997). The grp¹ is reported to be a genetic null allele because it lacks detectable levels of mRNA and because hemizygotes behave similarly to homozygotes; both mutants exhibit the same degree of nuclear abnormalities in the early syncytial cycles (Fogarty et al., 1994).

We performed our studies of checkpoints during the cellular cycles in grp¹ homozygous mutant embryos from heterozygous mothers. To analyze mitotic entry we quantified the mitotic index in embryos fixed 20 minutes after irradiation – at which time cells of irradiated wild-type embryos are still delayed before M16. A decrease in mitotic index reveals a delay in mitotic entry and activation of the mitotic entry checkpoint. The assay for metaphase-anaphase transition is performed by fixed and live measurement at 40 minutes after irradiation, at which time cells have recovered from the pre-mitotic delay and have progressed into M16 (see Fig. 1). We find that grp¹ mutants can regulate the entry into M16; mitotic index decreased in mutant embryos at 20 minutes after irradiation similar to wild type (Fig. 3A,B). Similarly, metaphase index in fixed embryos and metaphase length in live embryos indicate that grp¹ mutants are as capable of regulating metaphase-anaphase transition as wild type after DNA damage (Fig. 3C-E). Similar results were obtained from hemizygous mutant embryos (Df/grp¹) generated from a cross of grp¹ heterozygotes with heterozygotes for a chromosomal deficiency that includes the grp gene (not shown). Furthermore, embryos homozygous for this deficiency (thus completely devoid of the grp gene and is a true genetic null) were also capable of delaying the metaphase-anaphase transition (Fig. 3D). Thus, these data suggest that mutations in grp do not affect the ability to regulate the metaphase-anaphase transition in cellular stage embryos when DNA is damaged.

The appearance of mitotic chromosomes in irradiated grp mutants is similar to those in irradiated mei-41 mutants (not shown), even though the delay in metaphase-anaphase transition is present in the former but not the latter. In other words, gross chromosome anomalies, that might otherwise explain the delay in metaphase-anaphase transition, are not observed in grp mutants.

We have documented above that grp¹ mutant embryos are able to regulate both the entry into mitosis and mitotic progress after irradiation during cellular stages. Chk1 homologs in Xenopus and mammalian systems are involved in replication checkpoints and as mentioned above, Grp is also implicated in a replication checkpoint in syncytial cycles that occur earlier in embryogenesis. Therefore, we asked if the DNA replication checkpoint is functional in grp¹ mutants.

We blocked DNA replication using two alleles of double park, a homolog of an essential replication factor, Cdt1. Mutants carrying either allele complete the first 15 embryonic cell cycles, presumably using the maternally supplied Dup. The dup^a3 mutants initiate but fail to complete S16 and show a mei-41-dependent delay in the entry into M16 (Garner et al., 2001; Whittaker et al., 2000) (Table 1). Conversely, dup^a1 mutants show no detectable DNA synthesis in cycle16 and do not significantly delay the entry into M16, consistent with the findings from other systems that initiation of DNA synthesis is necessary to activate a checkpoint that blocks mitosis (D'Urso et al., 1995; Piatti et al., 1995). Additionally, both dup^a1 and dup^a3 mutants subsequently arrest during M16 (Whittaker et al., 2000).

The dup alleles allow us to determine the requirement for grp in blocking the entry into mitosis in response to incomplete replication (in dup^a3) and in regulating mitotic progress in response to the same signal (in both alleles). To determine the role of grp in regulating mitotic entry we compared M16 in dup^a3 and grp¹ dup^a3 mutants. At 7-7.5 hours after egg deposition (AED), cells of the lateral ectoderm of dup^a3 are delayed in entering M16 as shown by the near absence of PH3 staining, in agreement with previous reports (Garner et al., 2001; Whittaker et al., 2000) (Fig. 4A). By contrast, the corresponding cells of age-matched grp¹ dup^a3 mutants have entered M16 and show robust PH3 staining (Fig. 4A). We conclude that grp is needed to delay the entry into mitosis when DNA replication is incomplete as in dup^a3 mutants, and that the failure to execute this delay is apparent in grp¹ mutants (Table 1). Similarly, previous work with mei-41^D3 dup^a3 mutants showed that mei-41 is required to delay mitotic entry after incomplete replication (Garner et al., 2001) (Table 1).

Because grp affected the time of entry into M16 in dup^a3 mutants, we chose not to examine the role of grp in regulating mitotic progress in this allele of dup. We reasoned that the effects we see on mitotic progress might be secondary consequences of altered timing in mitotic entry. Instead, we used dup^a1 to determine the role of grp in mitotic progress. Previous work has shown that dup^a1 mutant embryos enter M16 on time but become delayed in M16; mitotic cells are readily visible at least until embryonic stages 13 and 14, about 3-4 hours later when mitotic activity has ceased in wild type or heterozygous siblings (Whittaker et al., 2000) (Fig. 4B). In grp¹ dup^a1 double mutant embryos, we find that PH3 staining is reduced at stages when mutants hemizygous for dup^a1 and a deficiency that removes dup (dup^a1/Df(2R)Jp1) are still arrested in M16 (8.5-9-hour-old embryos are shown in Fig. 4B). That is, grp¹ mutation rescued the elevation of mitotic index seen in dup^a1 mutants.

The above finding could be because either grp¹ dup^a1 double mutants are progressing through M16 and exiting mitosis prematurely (indicating a role for grp in regulating mitotic progress) or because they never entered M16. To distinguish between the above possibilities, we compared nuclear densities in each of the three thorasic segments in grp¹ dup^a1 double mutants and dup^a1 single mutants. We find that nuclear density in dup^a1 single mutants is about 60% of that in heterozygous controls (consistent with a delay in M16) while the nuclear density in grp¹ dup^a1 double mutants increased to levels seen in controls (Fig. 4C). In other words, cells of grp¹ dup^a1 embryos completed M16 and show nuclear density that is normal for the stage in embryogenesis. These results support the first possibility, that double mutant cells have exited M16 while the dup^a1 cells are still arrested in M16. We therefore infer that grp is needed to delay the progress through mitosis when DNA replication is incomplete (Table 1).

Previous work has shown that homozygous grp¹ mutants from heterozygous mothers (the mutants used in our experiments) lack detectable grp mRNA. However, since females deposit both mRNA and protein into early embryos, it is possible that wild-type maternal Grp protein persists in grp¹ mutants. To address this possibility, we raised polyclonal antibodies to Grp and compared protein levels in wild type and grp¹ mutants.

Affinity purified Grp antibodies recognize a band at the predicted MW of ∼55 kDa. Grp protein, similar to transcript expression, is present in newly deposited embryos and is at the highest levels during 2-8 hours AED, which loosely corresponds to cellular cycles 14-16 (Fogarty et al., 1997) (Fig. 5A). Grp levels decrease as most embryonic cells exit the cell cycle into G₁ of 17th cellular cycle and are almost absent at 12 hours AED through the rest of embryogenesis. The Grp signal is severely reduced in homozygous grp¹ mutants from heterozygous mothers, attesting to the specificity of the antibody (Fig. 5B). We find a similar level of persistent Grp protein in embryos homozygous for a deficiency that removes the entire grp coding region and grp¹ embryos from heterozygous mutant females of approximately 3% of Grp found in wild-type embryos of similar age (Fig. 5C and data not shown). We conclude that either Grp is not required to delay mitotic progress after irradiation or only very low levels of Grp are sufficient to do so. We cannot, however, rule out the possibility that the remaining signal in these mutants comes from a cross-reacting protein that is of exactly the same molecular mass as Grp. We found that 0-2-hour-old embryos (before the onset of zygotic gene expression) from homozygous grp¹ mothers have significantly reduced (∼5%) but not abolished levels of Grp (Fig. 5D). This could be either because grp¹, caused by a transposon insertion in the non-coding region, is a severe loss-of-function mutation and not a null, or because the cross-reacting protein is also present in pre-cellular stage embryos. We cannot distinguish between these possibilities at this point.

Immuno-staining of fixed embryos demonstrates that Grp localizes to the nucleus in interphase (Fig. 6A). Grp protein is present in syncytial embryos and is enriched in the interphase nucleus during cortical syncytial cycles (embryonic divisions 10-13). Upon nuclear envelope breakdown, Grp signal disperses into the cytoplasm for the duration of mitosis. Only after the reformation of the nuclear envelope in late telophase, does Grp re-localize to the nucleus. This is in contrast to MCMs, a family of chromatin-binding proteins, which localizes to the nucleus in interphase, disperse during mitosis, but re-accumulate onto chromosomes in anaphase prior to the reformation of the nuclear membrane (Su and O'Farrell, 1997). The kinetics of Grp nuclear localization are different and more consistent with nuclear localization via import rather than by association with chromosomes. Similar localization of Grp is observed in cycles 14-16 (not shown).

We have used live measurements of metaphase length to document the regulation of metaphase-anaphase transition in all three cell cycles of the cellularized Drosophila embryos. Many features of cell cycle responses to radiation in Drosophila appear to be conserved in mammalian cells. In both systems, irradiation in late G₂/early M does not cause a delay in mitotic progress, even though similar doses applied earlier in the cell cycle cause a checkpoint-dependent delay in entry into mitosis. In addition, in Drosophila embryos, irradiation during S results in a delay in the subsequent metaphase-anaphase transition. In both systems, 10-fold or higher doses of radiation applied in late G₂/early M did cause a delay in mitotic progress but this delay occurs independently of a DNA structure checkpoint. We speculate that additional cellular structures besides DNA are damaged to produce this delay. These results can account for previously reported apparent discrepancies between the behavior of Drosophila and mammalian cells, and indicate instead that the cells of the two systems behave similarly. Furthermore, it leaves open the possibility that irradiation, if applied earlier in the cell cycle such as during S phase, may induce a metaphase-anaphase delay in mammalian cells also.

What might be the purpose of delaying metaphase-anaphase transition? The delay is modest, lengthening metaphase by about 50%. Cell cycle delay by checkpoints are thought to allow time for DNA repair, but it is unclear if efficient repair can occur in such a short time on condensed chromosomes. It is possible that the delay acts as a signal to target damaged cells to death. It is in the interest of an organism to cull damaged cells and the presence of broken DNA during mitosis and this checkpoint may act to fulfill that purpose. If so, further work will be needed to address the mechanism that links a mei-41-dependent mitotic delay to cell death.

We present evidence that grp¹ mutants can delay the entry into mitosis and the metaphase-anaphase transition following IR, but are unable to delay these cell cycle transitions when replication is incomplete. We conclude that grp is essential for the DNA replication checkpoints in cellular stage embryos, whereas the DNA damage checkpoint requires little to no Grp. This conclusion agrees well with roles for Grp earlier in embryogenesis; Grp is required for a replication checkpoint in syncytial stage embryos, but not when DNA is damaged by irradiation (Fogarty et al., 1997; Sibon et al., 1997; Takada et al., 2003). Grp plays a redundant role with Chk2 to regulate the entry into mitosis after DNA damage checkpoint in the larvae (Brodsky et al., 2004). Thus it is possible that Grp plays a redundant role with Chk2 in regulation of metaphase-anaphase transition in the embryo. Analysis of grp chk2 double mutants would be informative in this regard.

A recently published report documents the regulation of metaphase-anaphase transition by checkpoints in response to DNA double-strand breaks in Drosophila larvae (Royou et al., 2005). This checkpoint, first described in embryos, is not limited to the embryo. This checkpoint in larvae, however, is abolished in grp¹ mutants, the same allele used in this study, indicating that the larval checkpoint requires a higher level of Grp than what is present in grp¹ mutants and is able to support a similar checkpoint in the embryo.

A bubR1-dependent spindle checkpoint has been shown to contribute to the mitotic arrest in dup mutants (Garner et al, 2001). Incomplete replication of centromeres is proposed to interfere with kinetochore function in these mutants, thereby activating a spindle checkpoint. Our current findings in grp¹ dup^a1 mutants suggest that a grp-dependent DNA structure checkpoint also contributes to this arrest. The involvement of both the spindle and DNA checkpoints in mitotic arrest upon incomplete DNA replication has precedence. In budding yeast, both MAD2, a spindle checkpoint protein, and Mec1, an ATR homolog, contribute to mitotic arrest in response to incomplete DNA replication (Garber and Rine, 2002; Krishnan et al., 2004).

ATR and Chk1 homologs act in the same checkpoint pathways in yeast and frogs, but can act together or in separate pathways in mammals depending on the type of damage. In Drosophila, Mei-41 and Grp appear to function together in most responses to DNA defects (Table 1). Both are required for a DNA damage checkpoint in larvae and to delay entry into mitosis in response to incomplete replication in syncytial embryos (Fogarty et al., 1997; Sibon et al., 1997). Delaying entry into mitosis in gastrula cycles in response to incomplete DNA replication in dup mutants requires mei-41 (Garner et al., 2001) and grp (this report). Grp is also needed for cells of dup^a1 mutants to remain in mitosis (this report). We have not been able to conclusively determine the role of mei-41 in the latter response because nuclei of dup^a1 mei-41 double mutant embryos are severely fragmented, making determination of nuclear density as in Fig. 4 unreliable.

Chk1 homologs exist in all eukaryotes examined and contribute to the regulation of mitosis in response both to DNA damage by ionizing radiation and to incomplete DNA replication. The differential importance of a Chk1 homolog to these two different responses has been compared only in fission yeast previously. In fission yeast, Chk1 is essential for delaying the entry into mitosis in response to IR damage, but has a redundant, non-essential role in delaying the entry into mitosis in response to incomplete DNA synthesis (Boddy et al., 1998; Brondello et al., 1999; Walworth and Bernards, 1996). Our results suggest a reverse situation in fly embryos; the regulation of mitotic progress after irradiation still occurs when levels of Grp are too low to support the regulation of mitosis in the presence of incomplete DNA replication. It would be of interest to see which of these intricate Chk1 requirements, that of yeast or that of Drosophila, are conserved in mammals.

We thank the Orr-Weaver lab for fly stocks and Mark Winey and members of the Su lab for critical comments on the manuscript. This work was supported by a grant from the NIH (GM66441) to T.T.S. A.P. and M.G.C. were partially supported by pre-doctoral training grants from the NIH.