DNA replication times the cell cycle and contributes to the mid-blastula transition in Drosophila embryos

Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA 94143
The Journal of Cell Biology (Impact Factor: 9.83). 10/2009; 187(1):7-14. DOI: 10.1083/jcb.200906191
Source: PubMed


We examined the contribution of S phase in timing cell cycle progression during Drosophila embryogenesis using an approach that deletes S phase rather than arresting its progress. Injection of Drosophila Geminin, an inhibitor of replication licensing, prevented subsequent replication so that the following mitosis occurred with uninemic chromosomes, which failed to align. The effect of S phase deletion on interphase length changed with development. During the maternally regulated syncytial blastoderm cycles, deleting S phase shortened interphase, and deletion of the last of blastoderm S phase (cycle 14) induced an extra synchronous division and temporarily deferred mid-blastula transition (MBT) events. In contrast, deleting S phase after the MBT in cycle 15 did not dramatically affect mitotic timing, which appears to retain its dependence on developmentally programmed zygotic transcription. We conclude that normal S phase and replication checkpoint activities are important timers of the undisturbed cell cycle before, but not after, the MBT.


Available from: Patrick H O'Farrell
The Rockefeller University Press $30.00
J. Cell Biol. Vol. 187 No. 1 7–14
Correspondence to Patrick H. O’Farrell:
Abbreviation used in this paper: MBT, mid-blastula transition.
The rst 13 nuclear divisions in the Drosophila embryo have a
streamlined cell cycle in which nuclei synchronously oscillate
between S phase and mitosis independent of zygotic transcrip-
tion (Foe et al., 1993). Maternal RNAs and proteins deposited
during oogenesis guide cell cycle progression. Cycles 1–9 occur
deep in a common cytoplasm and are of equal duration. At the
close of cycle 9, nuclei reach the embryo surface forming a
blastoderm, and undergo 4 additional mitotic cycles where
interphase (and S phase) gets progressively longer. At cycle 14
bulk zygotic transcription begins, many maternal products are
degraded, nuclei are cellularized, and S phase significantly
lengthens (Edgar and Schubiger, 1986; Edgar et al., 1986). These
events in cycle 14 are collectively known as the mid-blastula
transition (MBT).
The MBT is accompanied by changes in cell cycle regula-
tion (O’Farrell, 2001). In cycle 14, maternal Cdc25 phosphatase
is degraded and cells arrest in the rst G2 due to inhibitory
phosphorylation of cyclin-dependent kinase 1 (Cdk1) (Edgar
and O’Farrell, 1989; Edgar et al., 1994b). Patterned zygotic
transcription of Cdc25
during G2 in cycle 14 relieves Cdk1
inhibition and directs mitosis in a matching pattern (Edgar and
O’Farrell, 1990). In contrast, Cdc25 is abundant during the ma-
ternally driven cycles and there is little inhibitory phosphoryla-
tion of Cdk1 (Edgar et al., 1994b; Stumpff et al., 2004). It has
been suggested that accumulation of cyclin to some critical
threshold times Cdk1 activation (Murray and Kirschner, 1989;
Edgar et al., 1994b; Stifer et al., 1999; Ji et al., 2004; Crest
et al., 2007). However, the bulk levels of mitotic cyclin exhibit
little oscillation during the Drosophila maternally driven cycles
(Edgar et al., 1994b), and recent experiments ruled out mitotic
cyclin accumulation as the direct timer of mitotic entry (McCleland
and O’Farrell, 2008; McCleland et al., 2009).
The fast maternally driven cycles have altered checkpoint
responsiveness. Arresting DNA replication during the maternal
cycles extends interphase, but does not block nuclei from enter-
ing mitoses (Raff and Glover, 1988; Sibon et al., 1997). Flies
mutant for genes in the DNA damage checkpoint, mei41, grapes
(grp), or mus304, are viable, but eggs laid from homozygous
mutant mothers fail to normally prolong interphase during cy-
cles 10–13 and ultimately enter a catastrophic mitosis in cycle
13 (Fogarty et al., 1997; Sibon et al., 1997, 1999). Models attri-
bute the normal interphase extension to prolongation of S phase
and consequent activation of the checkpoint genes. Although
the results indeed showed that Grapes is required to delay mito-
sis, the idea that the duration of DNA replication is the master
timer of the early cycles is inferred from the common involve-
ment of S phase in checkpoint activation. Thus, the contribution
e examined the contribution of S phase in tim-
ing cell cycle progression during Drosophila
embryogenesis using an approach that de-
letes S phase rather than arresting its progress. Injection
of Drosophila Geminin, an inhibitor of replication licens-
ing, prevented subsequent replication so that the follow-
ing mitosis occurred with uninemic chromosomes, which
failed to align. The effect of S phase deletion on inter-
phase length changed with development. During the ma-
ternally regulated syncytial blastoderm cycles, deleting
S phase shortened interphase, and deletion of the last of
blastoderm S phase (cycle 14) induced an extra synchro-
nous division and temporarily deferred mid-blastula tran-
sition (MBT) events. In contrast, deleting S phase after the
MBT in cycle 15 did not dramatically affect mitotic timing,
which appears to retain its dependence on developmen-
tally programmed zygotic transcription. We conclude that
normal S phase and replication checkpoint activities are
important timers of the undisturbed cell cycle before, but
not after, the MBT.
DNA replication times the cell cycle and contributes
to the mid-blastula transition in Drosophila embryos
Mark L. McCleland, Antony W. Shermoen, and Patrick H. O’Farrell
Department of Biochemistry and Biophysics, University of California, San Francisco, San Francisco, CA 94143
© 2009 McCleland et al. This article is distributed under the terms of an Attribution–
Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publica-
tion date (see After six months it is available under a
Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license,
as described at
Page 1
JCB • VOLUME 187 • NUMBER 1 • 2009 8
interphase (Fig. 1 C and Video 2). Likewise, when Geminin was
injected at the beginning of cycle 14, defects appeared only 2.5
h later when cells began to collect in a prolonged mitosis 15
with unreplicated chromatids. The successful execution of cel-
lularization, gastrulation, mitosis 14, and germband extension
in these embryos implies that gene expression and general cell
biological functions are not disturbed by Geminin injection, at-
testing to the specicity of its action.
We used real-time records of YFP-PCNA to determine how
S phase deletion affects interphase duration. After Geminin in-
jection, interphase was shortened by 37% in cycle 12 and by 51%
in cycle 13 (Fig. 1 D). We note that a brief interphase remains in
the absence of S phase, suggesting that inhibitors such as phos-
phatases or low mitotic activators can limit progress to mitosis in
a brief post-mitotic period. However, the advancement of mitosis
shows that the levels of mitotic cyclin and other mitotic activators
are sufcient for mitosis midway through interphase. Thus, these
factors do not normally limit nal entry into mitosis during the
blastoderm cycles, consistent with previous analysis suggesting
that cyclin accumulation is not the direct timer of mitotic entry
(McCleland and O’Farrell, 2008; McCleland et al., 2009). Im-
portantly, the data suggest that DNA replication produces a nega-
tive signal that denes interphase length and directs its normal
extension during the blastoderm divisions.
Geminin injection promotes premature
chromatin condensation and mitosis with
unreplicated chromatids
Embryos coexpressing Cid-GFP and Histone-RFP were injected
with Geminin to examine whether deletion of S phase affected
chromatin dynamics. In the affected interphase, nuclei decon-
densed normally after telophase, but less than 3.5 min into inter-
phase chromatin began to recondense (Fig. 2 B and Video 4).
Within 1–2 min the chromatin was a compact mass in the center
of the nucleus. Shortly thereafter, nuclear envelope breakdown
and initial chromatid movements occurred. The early compac-
tion contrasts to wild-type embryos, wherein chromatin begins
to condense 8 min into cycle 12 or 13.5 min into cycle 13, just
before mitotic entry (Fig. 2 A, Video 3, and unpublished data).
Hence, deleting S phase in the early mitotic cycles promotes pre-
mature DNA condensation and advances mitotic entry.
Control metaphases exhibit eight stably aligned binemic
chromosomes whose paired centromeres are detected as two
dots of Cid-GFP (Fig. 2 and Video 3). In contrast, after S phase
deletion, mitotic chromosomes appeared uninemic, oscillated
unstably, and possessed a single Cid-GFP dot (Fig. 2 and Video
4). Thus, in a cycle after Geminin injection, nuclei enter mitosis
with single sister chromatids and fail to bi-orient. Real-time im-
aging of other mitotic proteins, GFP-Polo and Aurora B-GFP,
supports the conclusion that S phase deletion advances mitosis
with uninemic chromosomes (Fig. S2 and Videos 5 and 6).
Despite the lack of metaphase alignment, nuclei of
Geminin-injected embryos exited mitosis after a brief mitotic
delay. The total length of mitosis was nearly doubled in cycle 13
(Fig. 1 D), as chromosomes oscillated for 8 min before ana-
phase onset (Fig. 2 B and Video 4). We conclude that unrepli-
cated chromosomes delay mitotic progression.
of a normal S phase in timing the early embryonic cycles has not
been directly demonstrated.
In contrast to inhibition of DNA replication, which gener-
ally activates the S phase checkpoint, some modes of preventing
S phase do not arrest the cell cycle. Cells mutant in genes re-
quired for the initial licensing of origins, such as the yeast gene
CDC6 and the Drosophila gene double-parked (dup, a homo-
logue of yeast CDT1) fail to replicate their DNA, yet they enter
mitosis with unreplicated sister chromatids (Kelly et al., 1993;
Piatti et al., 1995; Whittaker et al., 2000). Addition of the Dup
inhibitor, Geminin, to Xenopus egg extracts or its overexpression
in ies prevents the loading of the MCM helicase complex, and
blocks the licensing step of DNA replication (McGarry and
Kirschner, 1998; Quinn et al., 2001). We reasoned that injection
of recombinant Geminin into syncytial Drosophila embryos
would similarly block DNA replication licensing, delete S phase
(Fig. 1 B), and thereby allow us to directly assess an S phase
contribution in timing the cell cycle.
Injection of Geminin prevented S phase in the ensuing
cycle and advanced mitotic entry during the syncytial blastoderm
divisions to give interphase durations similar to those in grp mu-
tant embryos. After S phase deletion, nuclei entered mitosis with
uninemic chromosomes, failed to achieve a metaphase align-
ment, and subsequently exited mitosis after a delay. We conclude
that normal S phase delays mitotic entry during the maternal
blastoderm divisions. Preventing S phase at the MBT in cycle 14
caused an extra syncytial division, uncovering a role for S phase
14 in the termination of the maternal mitotic program. Interest-
ingly, deleting S phase in cycle 15 did not signicantly alter the
timing of mitosis. We suggest that the shift in regulation of Cdk1
activation to dependency on transcription of Cdc25 at the MBT
supersedes the inputs from DNA replication.
Results and discussion
Geminin injection deletes S phase and
advances blastoderm mitoses
If Geminin injection blocks the licensing of origins, it should af-
fect the S phase occurring subsequent to licensing. Recruitment
of MCM proteins to anaphase chromosomes during the early
embryonic cycles marks the time of licensing (Su and O’Farrell,
1997). Geminin injection during interphase 13 did not affect the
progression of nuclei through mitosis 13, but did block the incor-
poration of dUTP in cycle 14, demonstrating the expected sec-
ond cycle effect (Fig. S1).
Embryos expressing YFP-PCNA were used to follow
S phase in real time. PCNA, a sliding clamp for DNA poly-
merase, localizes to the nucleus during interphase and forms foci
at active sites of DNA replication (Hingorani and O’Donnell,
2000). Although the syncytial S phases of Drosophila are incred-
ibly rapid, distinct foci of YFP-PCNA were evident throughout
S phase and were especially prominent during the closing minutes
of cycle 13 in the apical region of nuclei (Fig. 1 C and Video 1).
Consistent with a second cycle block to S phase, Geminin
injection did not immediately disturb YFP-PCNA foci and a
successful mitosis followed (Video 2 and unpublished data).
However, YFP-PCNA foci were not evident in the subsequent
Page 2
checkpoint activation, delays mitosis to create the gradual
lengthening of blastoderm cycles (Sibon et al., 1997; Crest
et al., 2007). If the activity of a checkpoint gene, such as grp,
relies on DNA replication, the deletion of S phase by Geminin
S phase deletion does not advance mitosis
mutant embryos
It has been argued that the exponential increase in DNA ti-
trates replication components and slows S phase, which, via
Figure 1. Injection of Geminin during the Drosophila blastoderm divisions deletes S phase and advances mitosis. (A) Schematic illustrating progres-
sion through the blastoderm divisions and the MBT. The bar beneath the graph represents time. Cdc25
protein declines during the blastoderm cycles.
Zygotic transcription of Cdc25
during G2 of cycle 14 promotes mitosis. (B) Schematic illustrating DNA replication initiation and experimental expecta-
tion from Geminin injection. (C) Nuclei of YFP-PCNA–expressing embryos in the cycle after injection of Geminin or control buffer. YFP-PCNA exhibits
distinct nuclear foci in control injections. Arrows highlight foci in the apical region of nuclei at the end of S phase. Note that YFP-PCNA foci are absent
throughout S phase after Geminin injection. Frames 9:56 and 11:48 of the Geminin-injected embryo correspond to premature mitotic entry and dis-
persal of nuclear YFP-PCNA in the cytoplasm. (D) YFP-PCNA–expressing embryos were injected in cycle 11 or cycle 12 with Geminin and interphase
length and mitosis were measured in the subsequent cycle (cycle 12 or cycle 13, respectively). Interphase was defined by nuclear localization of PCNA
and mitosis was defined by its dispersal. Average cell cycle times are shown for cycle 12 (n = 7 embryos) and cycle 13 (n = 10 embryos). Error bars
represent SD.
Page 3
JCB • VOLUME 187 • NUMBER 1 • 2009 10
after S phase deletion, chromatin in grp mutant embryos
remain dispersed until the close of interphase (Fig. 3 A).
Furthermore, Geminin injection into grp mutant embryos re-
sults in chromatin condensation behavior indistinguishable from
Geminin injection into wild-type embryos. Thus, DNA replica-
tion promotes chromatin dispersal, and dispersal does not require
checkpoint function. In cycle 14, the dispersed interphase ap-
pearance of chromatin is not dependent on S phase (see below
and Fig. 4), suggesting a change in the dependence on S phase
for chromatin dispersal.
injection should prevent its activation. In this case, a grp mu-
tation will have no additional phenotype in a cycle lacking
S phase. Furthermore, if all of the effects of DNA replication
on mitosis are mediated by grp, the mitotic phenotype of grp
and deletion of S phase ought to be the same. To probe this
we compared grp mutant embryos to those in which S phase
was deleted.
Histone-GFP–expressing embryos, which were wild type,
mutant in grp, or injected with Geminin, were lmed at high
resolution. In contrast to the very early condensation of chromatin
Figure 2. Geminin injection promotes premature chromatin condensation and mitosis with unreplicated chromatids. Frames showing nuclei from embryos
expressing Histone-RFP and Cid-GFP in the cycle after injection with control buffer (A) or Geminin (B). In the Geminin-injected embryo, chromatin begins
to condense 3 min into interphase, kinetochores are unpaired in mitosis, and chromosomes fail to congress to the metaphase plate. Time is presented as
min:sec. (C) Higher magnification view of Cid-GFP during mitosis in control and Geminin injections. Bifurcated arrows in the control nucleus point to paired
kinetochores at metaphase. Single arrows in Geminin injection highlight unpaired kinetochores.
Page 4
11S PHASE AS A CELL CYCLE TIMER • McCleland et al.
DNA replication at the MBT is
required for prompt termination of the
syncytial divisions
The switch from maternal to zygotic control at the MBT is accom-
panied by changes in cell cycle regulation (O’Farrell, 2001). The
literature has emphasized that a G2 phase is added to the cell cycle,
and that zygotic transcription times subsequent mitoses. However,
the rst MBT change in the cell cycle is prolongation of S phase.
We directly tested whether this S phase has a role in the MBT.
Geminin was injected into embryos expressing Histone-GFP
during cycle 13 (Fig. 4 A). After progressing normally through mi-
tosis 13, nuclei at the injected pole synchronously entered mitosis
16.5 min after the onset of cycle 14 (Fig. 4, A and C; and
Video 7), whereas mitosis 14 normally occurs in a spatial program
beginning 70 min after mitosis 13. These data indicate that S phase
14 is required to terminate the syncytial mitotic cycles.
To assess the inuence of S phase 14 deletion on MBT
events, we monitored cellularization. Geminin was injected during
cycle 13 into embryos expressing Histone-RFP and Sqh-GFP.
Only one extra synchronous division was observed, and this
Despite differences in chromatin condensation, Geminin-
injected embryos and grp mutant embryos exhibited similar mi-
totic entry times (Fig. 3 B). Furthermore, grp mutants injected
with Geminin exhibited similar interphase duration as wild-type
embryos injected with Geminin (Fig. 3). This result suggests
that Grapes has no activity after Geminin injection, and that the
shortening of interphase in the cycle after Geminin injection
might be attributable to the absence of the checkpoint pathway
when S phase is deleted. These ndings support the previous
inference that S phase delays entry into mitosis until its comple-
tion by activating Grapes-dependent checkpoint function.
Although Geminin injection into grp mutants barely al-
tered interphase length, it still prolonged mitosis, presumably
because it induced spindle checkpoint function in response to
uninemic chromatids. However, the mitotic delay caused by
Geminin injection into grp embryos was almost twice that
caused by Geminin injection into wild-type embryos (Fig. 3 B).
Perhaps the additional mitotic extension in grp mutants re-
flects a role of Grapes in mitosis or the spindle checkpoint
(Su et al., 1999).
Figure 3. Direct cell cycle comparison between embryos injected with Geminin and grp mutant embryos. (A) Either wild-type or grp mutant embryos
expressing Histone-GFP were injected with control buffer or Geminin. Time is presented in min:sec. (B) Quantification of interphase chromatin dynamics
and mitotic duration from embryos in A. Nuclear envelope formation (NEF) to the beginning of chromatin condensation was measured and the time from
chromatin condensation to the onset of mitosis was measured (onset of mitosis was defined by disappearance of the Histone-GFP halo around the chromatin
and the first chromatid movements). Average cell cycle times are shown for wild-type (n = 8 embryos), grp (n = 18 embryos), Geminin injected (n = 10
embryos), and Geminin injected into grp (n = 11 embryos). Error bars represent SD.
Page 5
JCB • VOLUME 187 • NUMBER 1 • 2009 12
Other observations also support an interpretation that zy-
gotic programming of post-MBT mitoses is independent of
S phase. First, although deletion of S phase 14 caused rapid entry
into an extra mitosis, this was followed by MBT events and a mi-
tosis that closely followed the normal temporal program of mitosis
14. Second, previous analyses of dup mutant embryos, which lack
S phase 16, show a normally patterned mitosis 16 (Garner et al.,
2001). Post-MBT divisions are normally promoted by Cdc25
expression (Edgar and O’Farrell, 1990), which is guided by the
patterning regulators (Edgar et al., 1994a). This zygotic program-
ming of divisions appears relatively resistant to S phase deletion.
As in the earlier cycles, deletion of S phase resulted in a
prolonged mitosis; however, the duration of the prolonged mito-
ses increased from 8.3 min in mitosis 14 to 20.5 min in mi-
tosis 15. We do not know the basis of this difference; however,
the mitotic delay in both cycles depended on the spindle check-
point protein Mad2 (Fig. 5), as has similarly been observed in
dup mutants in cycle 16 (Garner et al., 2001).
S phase as a mitotic timer
We developed a means to examine the contribution of a normal
S phase in timing mitotic entry during early Drosophila em-
bryogenesis. Previous work suggests that an S phase checkpoint
is activated during a normal S phase (Grallert and Boye, 2008),
and that S phase extension is key to the gradual prolongation of
was followed by cellularization, only slightly delayed compared
with the uninjected end of the embryo (Video 8). Surprisingly, a
subsequent division exhibited a normal mitosis 14 domain pat-
tern (unpublished data). Apparently, S phase 14 forestalls mito-
sis, but other MBT events, which introduce zygotic programming
of gastrulation and the cell cycle, continue in its absence (Edgar
and Datar, 1996).
S phase is not required for the zygotic
programming of a post-MBT mitosis
Geminin injection in cycle 14 had no effect until the next cycle,
when cells entered a normally patterned mitosis 15 with unrep-
licated chromatids (Fig. 4, B and C; Fig. S3 and Video 9). To
probe for subtle timing defects, we tracked individual cells dur-
ing cycle 15. The fth group of cells to initiate mitosis 14, do-
main 5, does so relatively synchronously (Foe et al., 1993), but
the program is more complex at entry into mitosis 15 (Fig. S3
and Video 10). Cells proximal to the cephalic furrow divide ear-
liest at 60.5 min after mitosis 14. Geminin-injected embryos
showed a slight advancement of mitosis in these cells to 52.75
min. Notably, deletion of S phase 15 did not normalize inter-
phase duration in domain 5, as cells anterior to the rst-dividing
cells still showed a later mitosis (Fig. S3 and Video 9). Thus,
deletion of S phase may have slightly shortened cycle 15, but it
did not override inputs that spatially pattern mitosis.
Figure 4. Geminin injection induces an extra syncytial division, but does not affect interphase length after the MBT. (A) Histone-GFP–expressing embryos
were injected at one pole (left) during cycle 13 with Geminin. Note that nuclei synchronously and prematurely enter mitosis 14 in the injected region.
(B) Histone-GFP–expressing embryos were injected with Geminin during interphase 14 before cellularization. Bifurcated arrows highlight normal ana-
phases during mitosis 14 and arrows show unreplicated chromosomes in mitosis 15 that fail to reach the metaphase plate. (C) Quantification of interphase
and mitotic duration from embryos as in A and B. Average interphase and mitotic times are shown for cycle 14 (n = 13 embryos) and cycle 15 in domain 5
(n = 33 embryos). Error bars represent SD.
Page 6
13S PHASE AS A CELL CYCLE TIMER • McCleland et al.
Materials and methods
Fly stocks
Drosophila melanogaster strains were grown on standard cornmeal-
yeast medium. Flies expressing Histone H2AvD-GFP, GFP-Polo (Fly Trap
#CC01326), Sqh-GFP, Cid-GFP, or Histone H2AvD-RFP were used for live-
embryo analysis (Clarkson and Saint, 1999; Royou et al., 2004; Buszczak
et al., 2007; Schuh et al., 2007). For checkpoint deficient experiments, the
following stocks were generated: w; grp
; H2AvD-GFP and w; H2AvD-
GFP/Cyo; mad2
(Buffin et al., 2007).
The PCNA ORF was amplified using primers 5-CACCATGTTC-
and cloned into the gateway system pENTR-D-TOPO vector (Invitrogen) ac-
cording to the manufacturer’s instructions. The resulting clone was used to
recombine the PCNA ORF into pPVW (UASp promoter with N-terminal
venus fusion from the Drosophila Gateway collection; T. Murphy Laboratory,
Carnegie Institute of Washington, Baltimore, MD). P{UASp-YFP-PCNA}
lines were obtained by standard germ-line transformation (BestGene, Inc.).
Resulting lines were crossed to Da-Gal4 flies (Daughterless promoter) for
YFP-PCNA expression.
Similarly, P{UASp-Aurora B-GFP} lines were constructed. The Aurora
B ORF was amplified using primers 5-CACCATGACGCTTTCCCGCGC-
GAAG-3 and 5-ATTTCTGGCCGTGTTCTCC-3 and ultimately recom-
bined into pPWG (UASp promoter with C-terminal GFP fusion). Resulting
lines were crossed to Tub-Gal4 ies (Lee and Luo, 1999) for Aurora
B-GFP expression.
Embryo manipulation
Drosophila embryos were collected on agar plates containing grape juice,
aged for the appropriate developmental time, dechorionated for 2 min in
50% bleach, and then extensively washed in water. For injections, em-
bryos were aligned on agar plates and transferred to coverslips. Embryos
were desiccated for 8–10 min and then overlaid with Halocarbon oil
(Sigma-Aldrich). Alexa 546–conjugated dUTP (Invitrogen) was injected at
a needle concentration of 100 µM.
Geminin protein purification
The Drosophila Geminin ORF was amplified using primers 5-AAACATAT-
GACCTTGTCCTC-3 and cloned into Pet28a as an NdeI–Xho fragment.
6XHis-Geminin was expressed in BL-21 DE3 pLysS bacteria (Agilent Tech-
nologies) and purified on nickel agarose beads according to the manu-
facturers instructions (QIAGEN). Geminin protein was dialyzed into
40 mM Hepes, pH 7.4, and 150 mM KCl and concentrated to 20 mg/ml
before injection.
Image acquisition and processing
Embryos were imaged on an inverted microscope (DM 1RB; Leica)
equipped with a spinning-disk confocal unit (CSU10; Yokagowa), 100x
Plan Fluotar 1.3 NA and 40x Plan Fluotar 0.7 NA objectives (Lecia), a
camera (Orca AG; Hamamatsu Photonics), and Volocity 4 acquisition soft-
ware (PerkinElmer). Image stacks were collected using 1.5-µm steps over
an 8–12-µm range using a controlled stage (MS-2000; Applied Scientific
Instrumentation). All images and videos were processed in Volocity 4 soft-
ware and presented as extended focused images. Image capture rates are
indicated per video. Time is displayed in hours:minutes:seconds.
At least three independent experiments were performed for each ex-
periment shown. In each experiment an X-Y stage facilitated the filming of
multiple embryos (usually greater than five embryos).
Fixed analysis and immunofluorescence
Embryos injected with Geminin were incubated for the appropriate time,
fixed in a mixture of 37% formaldehyde and heptane for 10 min, and
immediately hand devitellinized. Embryos were washed into phosphate-
buffered saline containing 2 µg/ml Hoescht 33258 (Invitrogen) and 2 µg/ml
Oregon green–conjugated wheat germ agglutinin (Invitrogen).
Online supplemental material
Fig. S1 illustrates the absence of DNA replication after Geminin injection.
Fig. S2 highlights the dynamics of mitotic proteins during the blastoderm
division after the deletion of S phase. Fig. S3 provides a wide-field
perspective of the post-MBT divisions after deletion of S phase 15. Vid-
eos 1–6 show real-time videos of blastoderm embryos progressing
through interphase and mitosis after deletion of S phase. Videos 7–8
highlight the effects of S phase deletion on MBT events. Videos 9–10
the maternally driven blastoderm cycles (Fogarty et al., 1997;
Sibon et al., 1997, 1999); however, before the results shown
here, the role of an undisturbed S phase in timing mitotic entry
had not previously been demonstrated. Our results show that a
normal S phase is an important timer of interphase length dur-
ing the syncytial blastoderm cycles.
The role of S phase as a mitotic timer during the maternal
divisions does not persist in the zygotically controlled divisions
after the MBT. We suggest that this change reects complete tran-
sition to a different mitotic timer. Before the MBT, Cdc25 and
other mitotic activators are abundant throughout the cell cycle,
there is no evident gap between S phase and mitosis, and nuclei
rely on S phase to forestall mitotic entry. After the MBT, many
mitotic activators are degraded upon mitotic exit, and restoration
of one of these, Cdc25
, requires new zygotic transcription
(Lehner and O’Farrell, 1989; Edgar and O’Farrell, 1990). Mitosis
awaits Cdc25
expression, which usually occurs after comple-
tion of S phase, thus introducing a G2 and absolving DNA repli-
cation of responsibility for timing mitosis. Notably, this proposal
offers an explanation for why the checkpoint genes, which couple
mitosis to the completion of S phase, are essential during the
early embryonic divisions but subsequently dispensable.
How then does S phase contribute during the beginning of
cycle 14 at the MBT? The prolongation of interphase is an im-
portant part of the MBT, allowing cellularization and providing
enough time for complete transcription of genes with long tran-
scription units (Shermoen and O’Farrell, 1991; Foe et al., 1993).
The prolongation of interphase and production of G2 phase
requires down-regulation of mitotic activators, particularly the
activating phosphatases encoded by string and twine. For exam-
ple, the inhibitors of String action, Tribbles and Früshstart,
which promote Cdc25
destruction and inhibit cyclin:Cdk1,
respectively, are transcribed during the long S phase of cycle 14
(Grosshans and Wieschaus, 2000; Seher and Leptin, 2000).
Although it has been attractive to view the MBT as a sharp
switch from maternal to zygotic control, we suggest that the re-
tooling of cell cycle regulation requires considerable time within
interphase 14 and that the prolonged S phase 14 enforces Cdk1
inactivation while MBT transition events progress.
Figure 5. The mitotic delay in response to unreplicated sister chromatids
is dependent on the spindle checkpoint. Embryos that were wild type or
mad2 mutant and expressing Histone-GFP were injected with Geminin pro-
tein. Mitotic duration was recorded during cycle 14 and cycle 15 in which
mitoses occurred with unreplicated sister chromatids. Mitosis 15 duration
was measured in multiple mitotic domains and identical results were found.
Average mitotic duration is shown for cycle 14 (n = 6 embryos) and cycle
15 (n = 12 embryos). Error bars represent SD.
Page 7
JCB • VOLUME 187 • NUMBER 1 • 2009 14
Lee, T., and L. Luo. 1999. Mosaic analysis with a repressible cell marker for
studies of gene function in neuronal morphogenesis. Neuron. 22:451–
461. doi:10.1016/S0896-6273(00)80701-1
Lehner, C.F., and P.H. O’Farrell. 1989. Expression and function of Drosophila
cyclin A during embryonic cell cycle progression. Cell. 56:957–968.
McCleland, M.L., and P.H. O’Farrell. 2008. RNAi of mitotic cyclins in
Drosophila uncouples the nuclear and centrosome cycle. Curr. Biol.
18:245–254. doi:10.1016/j.cub.2008.01.041
McCleland, M.L., J.A. Farrell, and P.H. O’Farrell. 2009. Inuence of cyclin type
and dose on mitotic entry and progression in the early Drosophila em-
bryo. J. Cell Biol. 184:639–646. doi:10.1083/jcb.200810012
McGarry, T.J., and M.W. Kirschner. 1998. Geminin, an inhibitor of DNA replica-
tion, is degraded during mitosis. Cell. 93:1043–1053. doi:10.1016/
Murray, A.W., and M.W. Kirschner. 1989. Cyclin synthesis drives the early em-
bryonic cell cycle. Nature. 339:275–280. doi:10.1038/339275a0
O’Farrell, P.H. 2001. Triggering the all-or-nothing switch into mitosis. Trends
Cell Biol. 11:512–519. doi:10.1016/S0962-8924(01)02142-0
Piatti, S., C. Lengauer, and K. Nasmyth. 1995. Cdc6 is an unstable protein whose
de novo synthesis in G1 is important for the onset of S phase and for pre-
venting a ‘reductional’ anaphase in the budding yeast Saccharomyces
cerevisiae. EMBO J. 14:3788–3799.
Quinn, L.M., A. Herr, T.J. McGarry, and H. Richardson. 2001. The Drosophila
Geminin homolog: roles for Geminin in limiting DNA replication, in ana-
phase and in neurogenesis. Genes Dev. 15:2741–2754. doi:10.1101/
Raff, J.W., and D.M. Glover. 1988. Nuclear and cytoplasmic mitotic cycles con-
tinue in Drosophila embryos in which DNA synthesis is inhibited with
aphidicolin. J. Cell Biol. 107:2009–2019. doi:10.1083/jcb.107.6.2009
Royou, A., C. Field, J.C. Sisson, W. Sullivan, and R. Karess. 2004. Reassessing
the role and dynamics of nonmuscle myosin II during furrow formation in
early Drosophila embryos. Mol. Biol. Cell. 15:838–850. doi:10.1091/
Schuh, M., C.F. Lehner, and S. Heidmann. 2007. Incorporation of Drosophila
CID/CENP-A and CENP-C into centromeres during early embryonic
anaphase. Curr. Biol. 17:237–243. doi:10.1016/j.cub.2006.11.051
Seher, T.C., and M. Leptin. 2000. Tribbles, a cell-cycle brake that coordinates
proliferation and morphogenesis during Drosophila gastrulation. Curr.
Biol. 10:623–629. doi:10.1016/S0960-9822(00)00502-9
Shermoen, A.W., and P.H. O’Farrell. 1991. Progression of the cell cycle through
mitosis leads to abortion of nascent transcripts. Cell. 67:303–310.
Sibon, O.C., V.A. Stevenson, and W.E. Theurkauf. 1997. DNA-replication check-
point control at the Drosophila midblastula transition. Nature. 388:93–97.
Sibon, O.C., A. Laurençon, R. Hawley, and W.E. Theurkauf. 1999. The
Drosophila ATM homologue Mei-41 has an essential checkpoint function
at the midblastula transition. Curr. Biol. 9:302–312. doi:10.1016/
Stifer, L.A., J.Y. Ji, S. Trautmann, C. Trusty, and G. Schubiger. 1999. Cyclin A
and B functions in the early Drosophila embryo. Development.
Stumpff, J., T. Duncan, E. Homola, S.D. Campbell, and T.T. Su. 2004. Drosophila
Wee1 kinase regulates Cdk1 and mitotic entry during embryogenesis.
Curr. Biol. 14:2143–2148. doi:10.1016/j.cub.2004.11.050
Su, T.T., and P.H. O’Farrell. 1997. Chromosome association of minichromosome
maintenance proteins in Drosophila mitotic cycles. J. Cell Biol. 139:13–
21. doi:10.1083/jcb.139.1.13
Su, T.T., S.D. Campbell, and P.H. O’Farrell. 1999. Drosophila grapes/CHK1 mu-
tants are defective in cyclin proteolysis and coordination of mitotic
events. Curr. Biol. 9:919–922. doi:10.1016/S0960-9822(99)80399-6
Whittaker, A.J., I. Royzman, and T.L. Orr-Weaver. 2000. Drosophila double
parked: a conserved, essential replication protein that colocalizes with the
origin recognition complex and links DNA replication with mitosis and
the down-regulation of S phase transcripts. Genes Dev. 14:1765–1776.
highlight developmental and cell cycle behavior upon deletion of S
phase 15. Online supplemental material is available at
We thank members of the O’Farrell laboratory for helpful comments.
This research was supported by National Institutes of Health training
grant GM078710 to M.L. McCleland and National Institutes of Health grant
GM037193 to P.H. O’Farrell.
Submitted: 30 June 2009
Accepted: 8 September 2009
Bufn, E., D. Emre, and R.E. Karess. 2007. Flies without a spindle checkpoint.
Nat. Cell Biol. 9:565–572. doi:10.1038/ncb1570
Buszczak, M., S. Paterno, D. Lighthouse, J. Bachman, J. Planck, S. Owen, A.D.
Skora, T.G. Nystul, B. Ohlstein, A. Allen, et al. 2007. The carnegie pro-
tein trap library: a versatile tool for Drosophila developmental studies.
Genetics. 175:1505–1531. doi:10.1534/genetics.106.065961
Clarkson, M., and R. Saint. 1999. A His2AvDGFP fusion gene complements
a lethal His2AvD mutant allele and provides an in vivo marker for
Drosophila chromosome behavior. DNA Cell Biol. 18:457–462.
Crest, J., N. Oxnard, J.Y. Ji, and G. Schubiger. 2007. Onset of the DNA replica-
tion checkpoint in the early Drosophila embryo. Genetics. 175:567–584.
Edgar, B.A., and S.A. Datar. 1996. Zygotic degradation of two maternal Cdc25
mRNAs terminates Drosophila’s early cell cycle program. Genes Dev.
10:1966–1977. doi:10.1101/gad.10.15.1966
Edgar, B.A., and P.H. O’Farrell. 1989. Genetic control of cell division patterns
in the Drosophila embryo. Cell. 57:177–187. doi:10.1016/0092-
Edgar, B.A., and P.H. O’Farrell. 1990. The three postblastoderm cell cycles of
Drosophila embryogenesis are regulated in G2 by string. Cell. 62:469–
480. doi:10.1016/0092-8674(90)90012-4
Edgar, B.A., and G. Schubiger. 1986. Parameters controlling transcriptional
activation during early Drosophila development. Cell. 44:871–877.
Edgar, B.A., C.P. Kiehle, and G. Schubiger. 1986. Cell cycle control by the
nucleo-cytoplasmic ratio in early Drosophila development. Cell. 44:365–
372. doi:10.1016/0092-8674(86)90771-3
Edgar, B.A., D.A. Lehman, and P.H. O’Farrell. 1994a. Transcriptional regulation
of string (cdc25): a link between developmental programming and the
cell cycle. Development. 120:3131–3143.
Edgar, B.A., F. Sprenger, R.J. Duronio, P. Leopold, and P.H. O’Farrell. 1994b.
Distinct molecular mechanism regulate cell cycle timing at successive
stages of Drosophila embryogenesis. Genes Dev. 8:440–452. doi:10.1101/
Foe, V.E., G.M. Odell, and B.A. Edgar. 1993. Mitosis and morphogenesis in the
Drosophila embryo: point and counterpoint. In The Development of
Drosophila melanogaster. M. Bate and A. Martinez-Arias, editors. Cold
Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 149–300.
Fogarty, P., S.D. Campbell, R. Abu-Shumays, B.S. Phalle, K.R. Yu, G.L. Uy,
M.L. Goldberg, and W. Sullivan. 1997. The Drosophila grapes gene is
related to checkpoint gene chk1/rad27 and is required for late syncytial
division fidelity. Curr. Biol. 7:418426. doi:10.1016/S0960-
Garner, M., S. van Kreeveld, and T.T. Su. 2001. mei-41 and bub1 block mitosis
at two distinct steps in response to incomplete DNA replication in
Drosophila embryos. Curr. Biol. 11:1595–1599. doi:10.1016/S0960-
Grallert, B., and E. Boye. 2008. The multiple facets of the intra-S checkpoint.
Cell Cycle. 7:2315–2320.
Grosshans, J., and E. Wieschaus. 2000. A genetic link between morphogenesis
and cell division during formation of the ventral furrow in Drosophila.
Cell. 101:523–531. doi:10.1016/S0092-8674(00)80862-4
Hingorani, M.M., and M. O’Donnell. 2000. Sliding clamps: a (tail)ored t. Curr.
Biol. 10:R25–R29. doi:10.1016/S0960-9822(99)00252-3
Ji, J.Y., J.M. Squirrell, and G. Schubiger. 2004. Both cyclin B levels and DNA-
replication checkpoint control the early embryonic mitoses in Drosophila.
Development. 131:401–411. doi:10.1242/dev.00944
Kelly, T.J., G.S. Martin, S.L. Forsburg, R.J. Stephen, A. Russo, and P. Nurse.
1993. The ssion yeast cdc18+ gene product couples S phase to START
and mitosis. Cell. 74:371–382. doi:10.1016/0092-8674(93)90427-R
Page 8
  • Source
    • "Embryonic cells replicate their entire genome in a short time period from a large number of randomly placed origins (Blumenthal et al. 1974). This pattern changes at the mid-blastula transition when S-phase length increases and the replication program is established (Newport and Kirschner 1982; Hyrien et al. 1995; McCleland et al. 2009). The analysis of replication timing in pluripotent stem cells undergoing differentiation has revealed that different cell-type-specific lineages change their gene expression pattern and chromatin architecture as well as their pattern of replication timing (Hiratani et al. 2010; Ryba et al. 2010; Chandra et al. 2012). "
    [Show abstract] [Hide abstract] ABSTRACT: DNA replication is a dynamic process that occurs in a temporal order along each of the chromosomes. A consequence of the temporally coordinated activation of replication origins is the establishment of broad domains (>100 kb) that replicate either early or late in S phase. This partitioning of the genome into early and late replication domains is important for maintaining genome stability, gene dosage, and epigenetic inheritance; however, the molecular mechanisms that define and establish these domains are poorly understood. The modENCODE Project provided an opportunity to investigate the chromatin features that define the Drosophila replication timing program in multiple cell lines. The majority of early and late replicating domains in the Drosophila genome were static across all cell lines; however, a small subset of domains was dynamic and exhibited differences in replication timing between the cell lines. Both origin selection and activation contribute to defining the DNA replication program. Our results suggest that static early and late replicating domains were defined at the level of origin selection (ORC binding) and likely mediated by chromatin accessibility. In contrast, dynamic domains exhibited low ORC densities in both cell types, suggesting that origin activation and not origin selection governs the plasticity of the DNA replication program. Finally, we show that the male-specific early replication of the X chromosome is dependent on the dosage compensation complex (DCC), suggesting that the transcription and replication programs respond to the same chromatin cues. Specifically, MOF-mediated hyperacetylation of H4K16 on the X chromosome promotes both the up-regulation of male-specific transcription and origin activation.
    Full-text · Article · Jul 2014 · Genome Research
  • Source
    • "We note that, sex-lethal, which plays a critical role in the fundamental biological process of dosage compensation in Drosophila, also exhibits a quick shutdown, at a time similar to that of hb shutdown [65]. Metazoan embryos share a common feature in that they start with a series of rapid, synchronous cell divisions, followed by a sudden slowing in division and onset of asynchronous divisions and morphogenic movements [66]–[68]. This period is referred to as the mid-blastula transition (MBT), which corresponds to the cycle 14A interphase in Drosophila [66]–[68]. "
    [Show abstract] [Hide abstract] ABSTRACT: Anterior-posterior (AP) patterning in the Drosophila embryo is dependent on the Bicoid (Bcd) morphogen gradient. However, most target genes of Bcd also require additional inputs to establish their expression domains, reflective of the operation of a cross-regulatory network and contributions of other maternal signals. This is in contrast to hunchback (hb), which has an anterior expression domain driven by an enhancer that appears to respond primarily to the Bcd input. To gain a better understanding of the regulatory logic of the AP patterning network, we perform quantitative studies that specifically investigate the dynamics of hb transcription during development. We show that Bcd-dependent hb transcription, monitored by the intron-containing nascent transcripts near the P2 promoter, is turned off quickly-on the order of a few minutes-upon entering the interphase of nuclear cycle 14A. This shutdown contrasts with earlier cycles during which active hb transcription can persist until the moment when the nucleus enters mitosis. The shutdown takes place at a time when the nuclear Bcd gradient profile in the embryo remains largely intact, suggesting that this is a process likely subject to control of a currently unknown regulatory mechanism. We suggest that this dynamic feature offers a window of opportunity for hb to faithfully interpret, and directly benefit from, Bcd gradient properties, including its scaling properties, to help craft a robust AP patterning outcome.
    Full-text · Article · Apr 2013 · PLoS ONE
  • Source
    • "PCNA associates with DNA polymerase at replication forks, and GFP-tagged PCNA accumulates into visible foci during replication. The formation of these foci requires replication, as none are observed in embryos that are prevented from licensing their origins and initiating replication (McCleland et al. 2009). Thus, we used GFP-PCNA foci as an approximation of the progress of replication. "
    [Show abstract] [Hide abstract] ABSTRACT: The Drosophila midblastula transition (MBT), a major event in embryogenesis, remodels and slows the cell cycle. In the pre-MBT cycles, all genomic regions replicate simultaneously in rapid S phases that alternate with mitosis, skipping gap phases. At the MBT, down-regulation of Cdc25 phosphatase and the resulting inhibitory phosphorylation of the mitotic kinase Cdk1 create a G2 pause in interphase 14. However, an earlier change in interphase 14 is the prolongation of S phase. While the signals modifying S phase are unknown, the onset of late replication-where replication of constitutively heterochromatic satellite sequences is delayed-extends S-phase 14. We injected Cdc25 mRNA to bypass the developmentally programmed down-regulation of Cdc25 at the MBT. Introduction of either Cdc25 isoform (String or Twine) or enhanced Cdk1 activity triggered premature replication of late-replicating sequences, even after their specification, and thereby shortened S phase. Reciprocally, reduction of Cdk1 activity by knockdown of mitotic cyclins extended pre-MBT S phase. These findings suggest that high Cdc25 and Cdk1 contribute to the speed of the rapid, pre-MBT S phases and that down-regulation of these activities plays a broader role in MBT-associated changes than was previously suspected.
    Full-text · Article · Mar 2012 · Genes & development
Show more