|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online February 20, 2009
doi: 10.1242/10.1242/dev.031815
1 Program in Molecular Medicine and the Program in Cell Dynamics, University of
Massachusetts Medical School, 377 Plantation Street, Worcester, MA 01605,
USA.
2 Institut Jacques Monod, UMR7592, 2 place Jussieu, 75251 Paris, Cedex 05,
France.
3 Department of Molecular Genetics, University of Toronto, 1 King's College
Circle, Toronto, Ontario M5S 1A8, Canada.
4 Program in Developmental and Stem Cell Biology, Research Institute, Hospital
for Sick Children, TMDT Building, 101 College Street, Toronto, Ontario M5G
1L7, Canada.
5 Department of Cell and Systems Biology and Canadian Drosophila
Microarray Centre, University of Toronto, Mississauga, Ontario L5L 1C6,
Canada.
6 Department of Biochemistry, University of Toronto, 1 King's College Circle,
Toronto, Ontario M5S 1A8, Canada.
* Author for correspondence (e-mail: william.theurkauf{at}umassmed.edu)
Accepted 17 December 2008
 |
SUMMARY |
|---|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
Key words: Cell cycle, Midblastula transition, Transcription, Drosophila
|
INTRODUCTION |
|---|
|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
;
O'Farrell et al., 2004). The
cleavage stage is driven by mRNAs and proteins that are deposited into the
oocyte, and is therefore under maternal genetic control. Transfer of
developmental control to the genome of the embryo occurs during the
maternal-to-zygotic transition (MZT), when a subset of the maternal mRNAs is
eliminated and zygotic transcription is initiated
(Schier, 2007;
Tadros et al., 2007b;
Zurita et al., 2008). While
this transfer of genetic control is under way, the cleavage stage divisions
slow and cell cycle checkpoints are activated. The MZT ends with the
midblastula transition (MBT), which is defined as the first developmental
process that depends on zygotic rather than maternal mRNAs.
Typically, the MBT follows a fixed number of cleavage divisions. In
Drosophila, for example, the MBT occurs during interphase of nuclear
cycle 14. However, haploid Drosophila embryos undergo an additional
division before the MBT whereas Drosophila embryos injected with
exogenous DNA proceed through fewer divisions before the MBT
(Edgar et al., 1986).
Similarly, polyspermic Xenopus embryos undergo fewer cleavage
divisions before the MBT (Newport and
Kirschner, 1982). These studies suggested that titration of a
maternally deposited factor by nuclei or DNA – which increase
exponentially during the cleavage divisions – might determine the timing
of the MBT. However, the timing of maternal Cyclin E destruction in
Xenopus and degradation of a pool of maternal mRNAs in
Drosophila are independent of the cleavage divisions
(Bashirullah et al., 1999;
Howe and Newport, 1996;
Tadros et al., 2003). In
addition, RNAi-mediated knock down of cyclins leads to premature termination
of the Drosophila cleavage divisions, but does not alter the timing
of cellularization (McCleland and
O'Farrell, 2008). These findings suggest that the MBT is under the
control of a developmental clock, but the molecular mechanisms that drive this
clock remain to be defined.
In Drosophila, the embryonic cleavage divisions slow progressively
during the syncytial blastoderm stage, when the majority of nuclei are in a
monolayer at the cortex (Foe and Alberts,
1983). Following cleavage 13, the cortical nuclei are surrounded
by membranes in a process termed cellularization, which is the first
morphological transformation that requires zygotic gene expression and
therefore represents the MBT (Mazumdar and
Mazumdar, 2002; Merrill et
al., 1988; Schweisguth et al.,
1991; Wieschaus and Sweeton,
1988). DNA replication checkpoint mutants fail to slow the
syncytial blastoderm stage cell cycle, cellularize or initiate high-level
zygotic transcription, which suggested a direct role in the MZT
(Sibon et al., 1999;
Sibon et al., 1997). However,
the cellularization and transcriptional defects associated with checkpoint
mutants are dramatically suppressed by mnk (encoding checkpoint
kinase 2, Chk2), a mutation that disrupts DNA damage signaling
(Takada et al., 2003;
Takada et al., 2007).
Furthermore, DNA damaging agents can block cellularization and zygotic
transcription (Takada et al.,
2007). The replication checkpoint thus controls the syncytial
divisions and helps maintain DNA integrity, but does not appear to directly
control the MZT.
Maternal transcript destruction during the Drosophila MZT requires
the conserved RNA-binding protein Smaug
(Tadros et al., 2007a). Smaug
recruits the CCR4/POP2/NOT deadenylase complex to its target transcripts,
triggering poly-A tail removal and transcript destruction
(Semotok et al., 2005;
Semotok et al., 2008;
Zaessinger et al., 2006).
Here, we show that smg mutant embryos fail to slow the syncytial
blastoderm cell cycle, terminate cleavage divisions, activate the DNA
replication checkpoint, cellularize or gastrulate. We also show that Smaug is
required for high-level zygotic transcription at the MZT. Significantly,
transgenic expression of Smaug in an anterior-to-posterior gradient induces an
anterior-to-posterior gradient in the onset of syncytial cell cycle delays,
maternal transcript destruction, zygotic gene activation, blastoderm
cellularization and even gastrulation. Smaug thus coordinates both the
cleavage-dependent and cleavage-independent aspects of the MZT, and may drive
a molecular clock that controls the timing of this key developmental
transition.
|
MATERIALS AND METHODS |
|---|
|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
) or
grpfs(A)4/grpfs(A)4
(Sibon et al., 1997) females
to wild-type Oregon R or w1118 males. The Smaug protein
gradient was generated using the UASp-smg-bcd3'UTR
(USB) transgenes (Tadros et al.,
2007a) and germline driver Nanos-Gal4:VP16 (NGV)
(Rorth, 1998). Females with
the genotype w1118; smg1 USB /smg1
NGV were crossed to wild-type Oregon R males. To assay for phenotypic
suppression by mnk, embryos were derived from w1118;
mnk6006/mnk6006;
smg1/smgDf(3L)Scf-R6 females crossed to
wild-type males (Abdu et al.,
2002). Unfertilized eggs were obtained from Oregon-R females
crossed to sterile males of the genotype T(Y;2)bw DRev#1, cn
bwDRev#11mr2/SM6a (Reed
and Orr-Weaver, 1997). The smg1, Df(3L)Scf-R6,
mnk6006 and Nanos-Gal4:VP16 stocks were obtained from
the Bloomington Drosophila Stock Center.
Time-lapse microscopy and checkpoint assays
Cell-cycle progression and replication checkpoint function were assayed by
time-lapse confocal microscopy using a Leica TCS-SP inverted laser-scanning
microscope, as described previously (Sibon
et al., 1997). Time-lapse DIC microscopy was performed using Zeiss
Axiovert–S100 inverted microscope and 20x Planapo lens. Images
were captured at 20-second intervals, and MetaMorph software was used for
image acquisition and processing. The length of cellularization was defined as
the time between the end of mitosis 13 and completion of membrane
invagination. Basic statistical analyses were performed in Excel
(Microsoft).
RNA in situ hybridization
Fluorescence in situ hybridization using RNA probes was performed as
described previously (Takada et al.,
2007). For slam antisense RNA probes, the DNA template
was amplified by PCR from wild-type genomic DNA, using the following sets of
primers: slam 5'-ctgttcagtccgattctcatcc-3' and
5'-T7-aatcttgtccatgtgctcgctg-3'. The T7 promoter sequence was:
5'-cgtaatacgactcactataggg-3'. For cyclin A and B
antisense RNA probes, the DNA templates were amplified by PCR from plasmids
containing the appropriate cDNA (Lehner
and O'Farrell, 1989; Lehner
and O'Farrell, 1990) using T3 and T7 primers. Antisense RNAs were
transcribed with the T7 polymerase. Images were acquired using a Leica TCS-SP
inverted laser-scanning microscope and processed with Photoshop.
Immunofluorescence labeling and quantification
For whole-mount immunostaining, embryos were fixed and immunolabeled as
described previously (Theurkauf,
1994). Rabbit anti-phospho-Histone H3 (P-H3) (Upstate Innovative
Cell Signaling Solution) was used at 1:500; mouse anti-phospho-Tyrosine
(P-Tyr) (Cell Signaling Technology) was used at 1:1000; guinea pig anti-Smaug
(Tadros et al., 2007a) was
used at 1:1000. The following fluorescent secondary antibodies were used: goat
anti-rabbit Alexa 488 (Molecular Probes, 1:500); donkey anti-mouse Alexa 488
(Molecular Probes, 1:500); donkey anti-guinea-pig FITC (Jackson
ImmunoResearch, 1:500). Embryos were labeled for DNA with TOTO3 (Molecular
Probes) at 0.1 µM and treated with RNase (Promega) at 10 U/ml during the
secondary antibody incubation. Images were acquired using a Leica TCS-SP
inverted laser-scanning microscope and processed with Photoshop.
To quantify the Smaug gradient, single plane images of cycle 13 embryos were captured using a Leica TCS-SP inverted laser-scanning microscope. Imaging was carried out under non-saturating conditions, and all genotypes were labeled and imaged under identical conditions. Pixel intensity (from 0 to 250) was extracted from raw images using MetaMorph software (Molecular Probes) and exported to Excel files. Immunolabeling and imaging of the embryo's interior is inefficient, owing to antibody penetration and light scattering. Analysis was therefore restricted to a four-pixel band at the cortex. Average pixel intensity as a function of position along the anterior-posterior axis was then calculated from an analysis of four independent embryos. To derive the profile of the gradient, the average cortical pixel intensity in wild-type embryos was subtracted from the average pixel intensity in wild-type embryos expressing the USB transgene.
Western blots
The following primary antibodies were used for western blotting:
anti-
-Tubulin (Sigma, 1:20,000, mouse), anti-β-Tubulin (E7 DSHB,
1:500, mouse), anti-large subunit of RNA polymerase II (ARNA-3a from Research
Diagnostics, 1:500, mouse), anti-phospho-ser-2 from the C-terminal domain of
RNA polymerase II (H5 from Research Diagnostic, 1:500, mouse) and anti-Smaug
(1:5000, guinea-pig). The following HRP-conjugated secondary antibodies were
used at a 1:1500 dilution: sheep anti-mouse (Amersham) and donkey
anti-guinea-pig (Jackson ImmunoResearch). Embryos were labeled and
hand-selected as described previously
(Edgar et al., 1994). The
equivalent of two embryos or one quarter of an ovary was loaded for each lane.
For western analysis of developmentally aged embryos, adult male and female
flies were placed in collection cages and fed yeast paste on grape juice agar
plates. Cages were kept at 25°C and plates were changed twice before
collection for 1 hour. The embryos were then used immediately (0-1 hour time
point), or aged 1 to 3 hours. The equivalent of 2.5 embryos of the desired age
was loaded for each lane. The ECL Plus detection system as used to detect
antibody binding (Amersham). Blots were imaged and quantified using a Kodak
Image Station and Excel.
Gene expression profiling
Total RNA was extracted from staged fertilized or unfertilized eggs, as
well as from stage 14 oocytes as described previously
(Tadros et al., 2007a). To
assay mRNA quality, known stable (rpA1) and unstable (Hsp83)
transcripts were probed on northern blots. Total RNA was then reverse
transcribed with random primers (Tadros et
al., 2007a) and labeled as described in the Indirect Labeling of
Total RNA for Microarray Hybridization protocol at
http://www.flyarrays.com.
The fluorescently labeled cDNA probes were hybridized to 12Kv1 and 14Kv1
microarrays obtained from the Canadian Drosophila Microarray Centre
(http://www.flyarrays.com;
GEO platform accession numbers: GPL1467,
http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GPL1467;
and GPL3603:
http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GPL3603).
Hybridization and scanning were performed as previously described
(Neal et al., 2003). The 16
bit TIFF image files were quantified using QuantArray Version 3 (PerkinElmer),
using the adaptive quantification algorithm and analyzed using GeneTraffic
Duo3.2 (Iobion Informatics/Stratagene). The 12Kv1 arrays were normalized using
the rank invariant LOWESS extrapolation method
(Schadt et al., 2001) and the
14Kv1 arrays were normalized to a set of known stable transcripts: Rpl1,
RpL32, Rps5a, Rps3, RpL22, mRpS30, mRpS22, CG6764, bonsai, RpS29, RpL11,
mRpL1, CG6764, CG317, RpL37A and RpL40. Additional information on data
analysis can be provided on request. To profile expression of miRs, after
extraction with Trizol reagent, total RNA was further purified using the
RNeasy Mini Kit (50) (Qiagen) as recommended by Exiqon. RNA samples were then
sent to Exiqon for miRCURY LNA Array microRNA Profiling. mRNA and miRNA
expression profiling data are available from the Gene Expression Omnibus (GEO)
server (Accession Numbers GSE13287 and GSE13288, respectively).
miRNA northern blotting
miRCURY LNA Detection Probes were purchased from Exiqon. Northern blotting
was performed described at
http://www.exiqon.com/MicroRNA_northern_blot,
except that a 15% polyacrylamide gel was used. For all samples, 12 µg of
total RNA was loaded per lane. The hybridization temperature was 40°C for
all miRCURY LNA probe-to-membrane hybridizations.
|
|
RESULTS |
|---|
|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
). To
gain insight into the genesis of these defects, we directly assayed cycle
progression and spindle assembly in smg mutants. For these studies,
embryos were injected with rhodamine-Tubulin and the DNA dye OliGreen and
monitored by time-lapse confocal microscopy for cell cycle progression.
Interphase length normally increases from 6 to 14 minutes between division
cycles 11 and 13. In smg mutants, by contrast, cell cycle length did
not increase and interphase 13 averaged only 7 minutes
(Fig. 1A). The smg
mutant embryos also progressed through additional syncytial cycles after
mitosis 13 (Fig. 1A) and
cellularization appeared to fail. These defects were confirmed by time-lapse
differential interference contrast (DIC) microscopy (see Table S1, Movies 1
and 2 in the supplementary material), which reduces photo-damage and reveals
membrane invagination, as well as by labeling with an anti-phospho-tyrosine
antibody, which reveals cellular membranes in early Drosophila
embryos (see Fig. S1A-C in the supplementary material).
The DNA replication checkpoint is required to slow the cleavage divisions
(Sibon et al., 1999;
Sibon et al., 1997). To
determine whether smg mutations disrupt the replication checkpoint,
we injected syncytial blastoderm stage embryos with the DNA replication
inhibitor aphidicolin, OliGreen and rhodamine-Tubulin, and assayed progression
into mitosis by time-lapse confocal microscopy
(Sibon et al., 1997). In
wild-type embryos, aphidicolin induced progressively more significant
interphase delays during cycles 11 and 13
(Fig. 1B)
(Crest et al., 2007),
indicating that replication checkpoint activity increases progressively during
the syncytial blastoderm stage. In smg mutants, by contrast,
aphidicolin failed to trigger significant cell cycle delays during any of the
syncytial blastoderm divisions (Fig.
1B). Smaug is therefore required for replication checkpoint
activation during the Drosophila MZT.
Smaug is required for activation of the zygotic genome
We next assayed activation of zygotic gene expression by microarray-based
gene expression profiling. For these studies, we compared transcript abundance
in embryos 2-3 hours post-fertilization with transcript abundance in mature
oocytes. Zygotic gene expression is normally activated at high levels between
2 and 3 hours after fertilization, whereas mature oocytes contain the full
complement of maternally loaded mRNA. The zygotically transcribed genes fall
into three distinct classes (Fig.
2A; see Table S2 in the supplementary material). Class I genes are
strictly zygotic, as they are not present in oocytes but are expressed in 2-
to 3-hour-old embryos. The remaining two classes are present in mature
oocytes, and are therefore maternally loaded. Class II mRNAs are stable
through the MZT, but increase in abundance as transcription is initiated.
Class III transcripts are degraded during this transition, and then
re-expressed zygotically.
Smaug is required for upregulation of 85% of the Class I genes and 90% of
the Class II genes, including the vast majority of the highly transcribed
zygotic genes (Fig. 2A). By
contrast, smg mutations blocked expression of only 15% of Class III
genes (Fig. 2A). Smaug is
required for destruction of the maternal pool of most Class III transcripts
(75%) (Tadros et al., 2007a),
so continued expression of these genes in smg mutant embryos could
reflect stabilization of the maternal pool, Smaug-independent transcription in
the embryo or a combination of these two factors. In summary, Smaug is
required for normal zygotic accumulation of transcripts from the vast majority
of protein-coding genes that are highly expressed during the MZT.
Transcript elongation is linked to phosphorylation of the large subunit of
RNA polymerase II (RNAPII) on serine 2 of the C-terminal domain (CTD)
(Majello and Napolitano,
2001). Accumulation of this phosphorylated form of RNAP II,
designated II0, also correlates with the increase in transcription during the
Drosophila MZT (Bellier et al.,
1997; Dantonel et al.,
2000; Leclerc et al.,
2000; Palancade et al.,
2001). However, II0 is not detectable in smg mutant
embryos (Fig. 2B). By contrast,
total RNAPII levels are similar in smg mutants and wild-type embryos
(Fig. 2B). The smg
mutation thus appears to block full activation of the transcription machinery
during the MZT.
|
), thus the transcription defects in smg mutants
could have been secondary to checkpoint failure and DNA damage. We therefore
assayed RNAPII CTD II0 phosphorylation in mnk; smg double-mutant
embryos (Fig. 2B). The defects
in II0 accumulation in smg mutants were only partially suppressed in
mnk; smg double mutants. In addition, these double-mutant embryos
consistently failed to cellularize or gastrulate (see Fig. S1F in the
supplementary material). By contrast, grp checkpoint mutant embryos
also showed reduced levels of CTD phosphorylation, but this was restored to
wild-type levels in mnk grp double mutants
(Fig. 2B). The transcriptional
block in smg mutants is therefore independent of checkpoint failure
and Chk2 activation.
To determine whether the defects in maternal transcript destruction in
smg mutants are secondary to checkpoint failure, we assayed maternal
mRNA destruction in grp and mnk grp checkpoint mutants.
Fluorescence in situ hybridization (FISH) studies showed that Hsp83
(Semotok et al., 2005;
Semotok et al., 2008) and
cyclin B mRNAs were stabilized in smg mutants
(Fig. 3D,F,J,L; and data not
shown), but degraded with normal kinetics in grp and mnk grp
double mutants (Fig. 3K; see
Fig. S2F in the supplementary material, data not shown). Cyclin A
mRNA was also degraded in grp and mnk grp double mutants,
but some transcripts appeared to persist
(Fig. 3E, data not shown).
These studies indicate that maternal transcript destruction is largely
independent of replication checkpoint control and DNA damage signaling.
miR-309-cluster microRNAs mediate Smaug-dependent destruction of maternal mRNAs
High-level zygotic expression of miR-309 cluster microRNAs (miRs)
directs destruction of a subset of maternal mRNAs at the MZT
(Bushati et al., 2008).
Hybridization of total RNA purified from staged eggs or embryos to a
microarray carrying probes for 68 known Drosophila miRs confirmed
that high-level miR-309-cluster transcription occurs only in embryos
and not in activated unfertilized eggs (see Fig. S3 in the supplementary
material). Northern blot analysis demonstrated that expression of three of the
miRs – miR-3, miR-6 and miR-286 – is disrupted
in smg mutants (Fig.
2C). Four-hundred and ten maternal mRNAs appear to require zygotic
expression of the miR-309 cluster for destabilization at the MZT
(Bushati et al., 2008).
Consistent with our observations, 
85% of these maternal transcripts are
stabilized in smg mutants (Fig.
2D). Smaug-dependent expression of the miR-309-cluster
thus leads to further destabilization of a subset of maternal mRNAs.
Smaug accumulation during the MZT
smg mRNA is translationally silent in mature oocytes, but is
translationally activated on egg activation
(Tadros et al., 2007a), and
Smaug protein is present at high levels in early embryos
(Dahanukar et al., 1999;
Smibert et al., 1999). To
define the precise kinetics of Smaug protein accumulation during the MZT, we
performed western blots on embryos that had been hand sorted according to cell
division cycle number (see Materials and methods). These studies showed that
Smaug levels progressively increase through the early cleavage divisions, peak
during syncytial blastoderm cycles 10 through 13, and then dramatically
decline during interphase of nuclear cycle 14
(Fig. 4). Thus, initiation of
Smaug expression correlates with initiation of maternal mRNA degradation,
whereas the peak in Smaug accumulation correlates with the onset of zygotic
gene expression and replication checkpoint activation.
|
A Smaug protein gradient triggers a temporal gradient in the MZT and MBT
To determine whether Smaug accumulation functions as a trigger for the MZT,
we expressed this protein in an anterior-to-posterior gradient and assayed
cell cycle progression, transcription, cellularization and gastrulation. If
Smaug functions as a trigger, changing protein levels will change the timing
of downstream processes rather than the extent to which these processes are
completed. By contrast, if Smaug has a direct role in multiple processes at
the MZT, the gradient should produce the equivalent of an allelic series,
which would change the extent of phenotypic rescue, but not the timing of
rescue. For example, different alleles of scraps produce
cellularization defects that differ in severity, but these alleles do not
appear to change the timing of this process
(Field et al., 2005).
To generate a Smaug gradient, we expressed a hybrid transgene in which the
smg open reading frame was fused to the bicoid
3'UTR, in the smg1 mutant background
(Tadros et al., 2007a)
(Fig. 5C). We quantified the
resulting gradient using immunolabeling for Smaug (see Materials and methods).
The smg1 allele produces a truncated 60 kDa protein that
is detected by the anti-Smaug antibody
(Fig. 5E). We therefore
estimated the shape of the gradient by quantifying immunofluorescence labeling
along the anterior-posterior axis in wild-type embryos expressing the
USB transgene, and then subtracting the uniform fluorescence observed
in control wild-type embryos (Fig.
5D). Immunofluorescence labeling intensity may not be linear with
respect to protein concentration, thus these data provide only a rough
estimate of the shape of the gradient, not precise information on protein
concentration. Nonetheless, these studies indicate that Smaug is expressed
above wild-type levels at the anterior pole, drops below average wild-type
levels between 75 and 80% egg length (EL), and progressively declines between
75% and 0% egg length (0% and 100% EL represent the posterior and anterior
pole, respectively).
|
|
) (see Movie 1 in the supplementary material). In
smg-USB embryos, by contrast, mitosis was delayed at the anterior,
leading to posterior-to-anterior waves of nuclear division (see Movies 3 and 4
in the supplementary material). To confirm these observations, we analyzed the
distribution of phosphorylated histone H3, a marker for mitosis, along the
axis of the embryos. Wild-type embryos showed uniform H3 phosphorylation,
labeling at both poles or labeling through the center
(Fig. 6C, data not shown). In
smg-USB embryos, however, phosphorylated-histone-H3-labeled
mitotic nuclei were often observed at the posterior and unlabeled interphase
nuclei were present near the anterior pole, indicating a delay in mitosis at
the anterior (Fig. 6D).
Expressing Smaug in an anterior-to-posterior gradient thus triggers
anterior-to-posterior delays in cell cycle progression. The Smaug gradient also consistently led to a gradient in the timing of cellularization (see Movies 3 and 4 in the supplementary material; Fig. 6F). We found that 100% of wild-type embryos initiated cellularization synchronously along the anteroposterior axis, with a mean mid-cellularization time of 93 minutes after syncytial interphase 10 (see Movie 1 in the supplementary material; Fig. 6E; Fig. S4 in the supplementary material). By contrast, 100% of embryos expressing Smaug in a gradient showed a striking anterior-to-posterior cellularization gradient (n=12, Fig. 6F; see Movies 3 and 4 in the supplementary material). In these embryos, cellularization was initiated over the anterior 20 to 25% of egg length with essentially wild-type timing (mean of 94 minutes after syncytial mitosis 10, see Fig. S4 in the supplementary material). Over the remaining 75% of egg length, where Smaug drops below wild-type levels, cellularization was progressively delayed and mid-cellularization was not observed until 117 minutes after interphase 10 at the posterior pole (see Fig. S4 in the supplementary material). In addition, embryos derived from hemizygous females, which express Smaug at 50% of wild-type levels, proceeded through the normal number of syncytial divisions, but cellularized 10 minutes later than embryos derived from wild-type diploid mothers (data not shown). Reducing Smaug expression thus leads to progressive delays in the MBT.
In 8 of 12 live recordings, the entire embryo completed 13 syncytial divisions before cellularizing in a graded manner (see Movie 3 in the supplementary material). In 4 out of 12 embryos, however, division at the anterior pole terminated after mitosis 12, whereas the rest of the embryo progressed through the normal 13 syncytial divisions (see Movie 4 in the supplementary material; Fig. 6B). However, the timing of cellularization at the anterior pole was the same in both classes of embryos. Smaug overexpression thus triggers premature arrest of the cleavage divisions, but does not advance the MBT.
In wild-type embryos, the gastrulation movements of germband extension and dorsal migration of the pole cells were observed before the cephalic furrow could be clearly identified by DIC (see Movie 1 in the supplementary material). In embryos expressing Smaug in a gradient, by contrast, cephalic furrow formation initiated before cellularization was completed at the posterior pole (see Movies 3 and 4 in the supplementary material). Higher resolution time-lapse DIC imaging confirmed that cellularization and gastrulation were significantly delayed, but these processes appeared to be completed (see Fig. S5, Movies 5 and 6 in the supplementary material), and 19% of embryos expressing the Smaug gradient hatched (n=300). Reducing Smaug thus delays the early developmental program, but does not appear to block execution of this program.
To determine the influence of the Smaug gradient on zygotic gene
activation, we used FISH to assay zygotic expression of slam and
runt (Fig. 7), which
are required for cellularization and segmentation, respectively
(Kania et al., 1990;
Lecuit et al., 2002). In
wild-type interphase 14 embryos proceeding through cellularization,
slam transcripts were uniformly expressed and runt was
expressed in seven stripes (Fig.
7B,F). slam expression dropped rapidly as cellularization
was completed and gastrulation was initiated, while the seven-stripe pattern
of runt expression persisted (data not shown). In
smg-USB embryos undergoing graded cellularization,
slam was first expressed in an anterior cap and runt was
expressed in a single anterior stripe (Fig.
7C,G). In later embryos, slam was expressed at peak
levels at the posterior and at lower levels at the anterior pole, and the full
seven-stripe runt pattern was established
(Fig. 7D,H). The Smaug gradient
thus triggers a wave of slam and runt transcription that
starts at the anterior pole and progresses to the posterior pole. To assay for
maternal transcript destruction, we used FISH to monitor maternal cyclin
B mRNA levels. This transcript is normally degraded over most of the
embryo, but is retained in the pole cells
(Fig. 7I-J). In
smg-USB embryos, by contrast, cyclin B mRNA was degraded in
an anterior-to-posterior gradient (Fig.
7K,L). Graded expression of Smaug thus appears to trigger an
anterior-to-posterior gradient in the transition from maternal to zygotic
control of development.
|
|
DISCUSSION |
|---|
|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
|
), and an additional factor (`Y') has been proposed to act
with Smaug to trigger decay (Tadros et
al., 2007a). Factor Y or a component of the CCR4/POP2/NOT complex
could become limiting for transcript destruction and the MBT when Smaug is
overexpressed. Alternatively, Smaug expression at wild-type levels could
saturate the binding sites on target transcripts and the deadenylation
machinery could be present in excess. Under these conditions, wild-type levels
of Smaug would lead to transcript destruction at maximal rates, which in turn
would determine the minimum time for the MZT. Smaug also represses translation
of specific targets (e.g. nanos), and this function could also have a
role in coordinating the MZT.
Based on the present studies and earlier work, we favor the simple
hypothesis that Smaug-dependent maternal transcript destruction triggers the
MZT by coordinately downregulating a suite of maternal proteins that suppress
zygotic transcription and the replication checkpoint. Consistent with this
speculation, Smaug is required for maternal cyclin B transcript
destruction, and overexpression of Cyclin B suppresses checkpoint activation
and leads to additional rapid syncytial divisions (Crest et al., 2006).
Similarly, Smaug triggers destruction of maternal mRNA encoding Tramtrack,
which represses transcription of a subset of genes until near the end of the
MZT (Pritchard and Schubiger,
1996; Tadros et al.,
2007a).
An initial wave of Smaug-dependent zygotic gene expression appears to
trigger a series of positive and negative feedback loops that drive completion
of the MZT (Fig. 8). For
example, Smaug is required for zygotic transcription of fruhstart
(frs), which promotes destruction of maternal string mRNA
(Grosshans et al., 2003).
String activates Cyclin-B-Cdk1, and string mRNA destruction in
response to Frs expression could cooperate with Smaug-dependent
cyclinB mRNA destruction to terminate the rapid cleavage stage
divisions (Donzelli and Draetta,
2003). Smaug is also required for zygotic expression of
transcriptional activators, including cyclin T (see Table S2 in the
supplementary material), cdk7 and cdk9 (C.H.H., J.T.W. and
H.D.L., unpublished), and CyclinT-Cdk9 complex catalyzes phosphorylation of
serine 2 on the RNAPII CTD repeat, which is linked to Smaug-dependent
transcription at the MZT (Fig.
3) (Bellier et al.,
1997; Palancade et al.,
2001; Shim et al.,
2002). Transcription of cyclin T and cdk9
concomitant with destruction of maternal RNAs encoding transcriptional
repressors, could therefore accelerate activation of the zygotic gene
expression. Finally, Smaug is required for zygotic expression of the
miR-309 cluster, which promotes a second, more rapid phase of
maternal transcript destruction (Bushati et
al., 2008) that terminates maternal genetic control of
embryogenesis (Fig. 8). A
recent study has shown that a maternally deposited transcription factor,
Zelda, is also required for expression of the miR-309 cluster and
number of other early zygotic genes (Liang
et al., 2008). There is no evidence that Zelda regulates the
timing of the MZT, but Zelda and Smaug could function cooperatively to promote
transcription and maternal transcript destruction during this transition.
|
;
Sibon et al., 1997). These
observations, along with the present studies, suggest a simple model in which
Smaug accumulation drives a maternal clock that determines timing of the MZT
and cellularization, while the replication checkpoint monitors the N/C ratio
and thus controls the number of cleavage divisions that can be completed
before the clock runs down. In this model, haploid embryos contain less DNA,
and therefore activate the checkpoint later and proceed through an additional
division before cellularization. Embryos containing additional DNA, by
contrast, trigger the checkpoint earlier and complete fewer divisions within
the window provided by the clock. This model thus explains the link between
the MBT and the N/C ratio, and control of the timing of this transition by a
cleavage-independent clock. |
Footnotes |
|---|
Supplementary material for this article is available at http://dev.biologists.org/cgi/content/full/136/6/923/DC1
|
REFERENCES |
|---|
|
TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
|---|
Abdu, U., Brodsky, M. and Schupbach, T. (2002).
Activation of a meiotic checkpoint during Drosophila oogenesis regulates the
translation of Gurken through Chk2/Mnk. Curr. Biol.
12,1645
-1651.[CrossRef][Medline]
Bashirullah, A., Halsell, S. R., Cooperstock, R. L., Kloc, M.,
Karaiskakis, A., Fisher, W. W., Fu, W., Hamilton, J. K., Etkin, L. D. and
Lipshitz, H. D. (1999). Joint action of two RNA degradation
pathways controls the timing of maternal transcript elimination at the
midblastula transition in Drosophila melanogaster. EMBO
J. 18,2610
-2620.[CrossRef][Medline]
Bellier, S., Chastant, S., Adenot, P., Vincent, M., Renard, J.
P. and Bensaude, O. (1997). Nuclear translocation and
carboxyl-terminal domain phosphorylation of RNA polymerase II delineate the
two phases of zygotic gene activation in mammalian embryos. EMBO
J. 16,6250
-6262.[CrossRef][Medline]
Bushati, N., Stark, A., Brennecke, J. and Cohen, S. M.
(2008). Temporal reciprocity of miRNAs and their targets during
the maternal-to-zygotic transition in Drosophila. Curr.
Biol. 18,501
-506.[CrossRef][Medline]
Crest, J., Oxnard, N., Ji, J. Y. and Schubiger, G.
(2007). Onset of the DNA replication checkpoint in the early
Drosophila embryo. Genetics
175,567
-584.
Dahanukar, A., Walker, J. A. and Wharton, R. P.
(1999). Smaug, a novel RNA-binding protein that operates a
translational switch in Drosophila. Mol. Cell
4, 209-218.[CrossRef][Medline]
Dantonel, J. C., Quintin, S., Lakatos, L., Labouesse, M. and
Tora, L. (2000). TBP-like factor is required for embryonic
RNA polymerase II transcription in C. elegans. Mol.
Cell 6,715
-722.[CrossRef][Medline]
Donzelli, M. and Draetta, G. F. (2003).
Regulating mammalian checkpoints through Cdc25 inactivation. EMBO
Rep. 4,671
-677.[CrossRef][Medline]
Edgar, B. A., Kiehle, C. P. and Schubiger, G.
(1986). Cell cycle control by the nucleo-cytoplasmic ratio in
early Drosophila development. Cell
44,365
-372.[CrossRef][Medline]
Edgar, B. A., Sprenger, F., Duronio, R. J., Leopold, P. and
O'Farrell, P. H. (1994). Distinct molecular mechanisms
regulate cell cycle timing at successive stages of Drosophila embryogenesis.
Genes Dev. 8,440
-452.
Etkin, L. D. (1988). Regulation of the
mid-blastula transition in amphibians. Dev. Biol.
5, 209-225.[Medline]
Field, C. M., Coughlin, M., Doberstein, S., Marty, T. and
Sullivan, W. (2005). Characterization of anillin mutants
reveals essential roles in septin localization and plasma membrane integrity.
Development 132,2849
-2860.
Foe, V. E. and Alberts, B. M. (1983). Studies
of nuclear and cytoplasmic behaviour during the five mitotic cycles that
precede gastrulation in Drosophila embryogenesis. J. Cell
Sci. 61,31
-70.[Abstract]
Grosshans, J., Muller, H. A. and Wieschaus, E.
(2003). Control of cleavage cycles in Drosophila embryos by
fruhstart. Dev. Cell 5,285
-294.[CrossRef][Medline]
Howe, J. A. and Newport, J. W. (1996). A
developmental timer regulates degradation of cyclin E1 at the midblastula
transition during Xenopus embryogenesis. Proc. Natl. Acad. Sci.
USA 93,2060
-2064.
Kania, M. A., Bonner, A. S., Duffy, J. B. and Gergen, J. P.
(1990). The Drosophila segmentation gene runt encodes a novel
nuclear regulatory protein that is also expressed in the developing nervous
system. Genes Dev. 4,1701
-1713.
Leclerc, V., Raisin, S. and Leopold, P. (2000).
Dominant-negative mutants reveal a role for the Cdk7 kinase at the
mid-blastula transition in Drosophila embryos. EMBO J.
19,1567
-1575.[CrossRef][Medline]
Lecuit, T., Samanta, R. and Wieschaus, E.
(2002). slam encodes a developmental regulator of polarized
membrane growth during cleavage of the Drosophila embryo. Dev.
Cell 2,425
-436.[CrossRef][Medline]
Lehner, C. F. and O'Farrell, P. H. (1989).
Expression and function of Drosophila cyclin A during embryonic cell cycle
progression. Cell 56,957
-968.[CrossRef][Medline]
Lehner, C. F. and O'Farrell, P. H. (1990). The
roles of Drosophila cyclins A and B in mitotic control.
Cell 61,535
-547.[CrossRef][Medline]
Liang, H. L., Nien, C. Y., Liu, H. Y., Metzstein, M. M., Kirov,
N. and Rushlow, C. (2008). The zinc-finger protein Zelda is a
key activator of the early zygotic genome in Drosophila.
Nature 456,400
-403.[CrossRef][Medline]
Majello, B. and Napolitano, G. (2001). Control
of RNA polymerase II activity by dedicated CTD kinases and phosphatases.
Front. Biosci. 6,D1358
-D1368.[Medline]
Mazumdar, A. and Mazumdar, M. (2002). How one
becomes many: blastoderm cellularization in Drosophila melanogaster.
BioEssays 24,1012
-1022.[CrossRef][Medline]
McCleland, M. L. and O'Farrell, P. H. (2008).
RNAi of mitotic cyclins in Drosophila uncouples the nuclear and centrosome
cycle. Curr. Biol. 18,245
-254.[CrossRef][Medline]
Merrill, P. T., Sweeton, D. and Wieschaus, E.
(1988). Requirements for autosomal gene activity during
precellular stages of Drosophila melanogaster.
Development 104,495
-509.
Neal, S. J., Gibson, M. L., So, A. K. and Westwood, J. T.
(2003). Construction of a cDNA-based microarray for Drosophila
melanogaster: a comparison of gene transcription profiles from SL2 and Kc167
cells. Genome 46,879
-892.[Medline]
Newport, J. and Kirschner, M. (1982). A major
developmental transition in early Xenopus embryos: I. characterization and
timing of cellular changes at the midblastula stage.
Cell 30,675
-686.[CrossRef][Medline]
O'Farrell, P. H., Stumpff, J. and Su, T. T.
(2004). Embryonic cleavage cycles: how is a mouse like a fly?
Curr. Biol. 14,R35
-R45.[Medline]
Palancade, B., Bellier, S., Almouzni, G. and Bensaude, O.
(2001). Incomplete RNA polymerase II phosphorylation in Xenopus
laevis early embryos. J. Cell Sci.
114,2483
-2489.
Pritchard, D. K. and Schubiger, G. (1996).
Activation of transcription in Drosophila embryos is a gradual process
mediated by the nucleocytoplasmic ratio. Genes Dev.
10,1131
-1142.
Reed, B. H. and Orr-Weaver, T. L. (1997). The
Drosophila gene morula inhibits mitotic functions in the endo cell cycle and
the mitotic cell cycle. Development
124,3543
-3553.[Abstract]
Rorth, P. (1998). Gal4 in the Drosophila female
germline. Mech. Dev. 78,113
-118.[CrossRef][Medline]
Schadt, E. E., Li, C., Ellis, B. and Wong, W. H.
(2001). Feature extraction and normalization algorithms for
high-density oligonucleotide gene expression array data. J. Cell
Biochem. 37 Suppl.,120
-125.
Schier, A. F. (2007). The maternal-zygotic
transition: death and birth of RNAs. Science
316,406
-407.
Schweisguth, F., Vincent, A. and Lepesant, J. A.
(1991). Genetic analysis of the cellularization of the Drosophila
embryo. Biol. Cell 72,15
-23.[CrossRef][Medline]
Semotok, J. L., Cooperstock, R. L., Pinder, B. D., Vari, H. K.,
Lipshitz, H. D. and Smibert, C. A. (2005). Smaug recruits the
CCR4/POP2/NOT deadenylase complex to trigger maternal transcript localization
in the early Drosophila embryo. Curr. Biol.
15,284
-294.[CrossRef][Medline]
Semotok, J. L., Luo, H., Cooperstock, R. L., Karaiskakis, A.,
Vari, H. K., Smibert, C. A. and Lipshitz, H. D. (2008).
Drosophila maternal Hsp83 mRNA destabilization is directed by multiple SMAUG
recognition elements in the open reading frame. Mol. Cell
Biol. 28,6757
-6772.
Shim, E. Y., Walker, A. K., Shi, Y. and Blackwell, T. K.
(2002). CDK-9/cyclin T (P-TEFb) is required in two postinitiation
pathways for transcription in the C. elegans embryo. Genes
Dev. 16,2135
-2146.
Sibon, O. C., Stevenson, V. A. and Theurkauf, W. E.
(1997). DNA-replication checkpoint control at the Drosophila
midblastula transition. Nature
388, 93-97.[CrossRef][Medline]
Sibon, O. C., Laurencon, A., Hawley, R. and Theurkauf, W. E.
(1999). The Drosophila ATM homologue Mei-41 has an essential
checkpoint function at the midblastula transition. Curr.
Biol. 9,302
-312.[CrossRef][Medline]
Smibert, C. A., Lie, Y. S., Shillinglaw, W., Henzel, W. J. and
Macdonald, P. M. (1999). Smaug, a novel and conserved
protein, contributes to repression of nanos mRNA translation in vitro.
RNA 5,1535
-1547.[Abstract]
Tadros, W., Houston, S. A., Bashirullah, A., Cooperstock, R. L.,
Semotok, J. L., Reed, B. H. and Lipshitz, H. D. (2003).
Regulation of maternal transcript destabilization during egg activation in
Drosophila. Genetics
164,989
-1001.
Tadros, W., Goldman, A. L., Babak, T., Menzies, F., Vardy, L.,
Orr-Weaver, T., Hughes, T. R., Westwood, J. T., Smibert, C. A. and Lipshitz,
H. D. (2007a). SMAUG is a major regulator of maternal mRNA
destabilization in Drosophila and its translation is activated by the PAN GU
kinase. Dev. Cell 12,143
-155.[CrossRef][Medline]
Tadros, W., Westwood, J. T. and Lipshitz, H. D.
(2007b). The mother-to-child transition. Dev.
Cell 12,847
-849.[CrossRef][Medline]
Takada, S., Kelkar, A. and Theurkauf, W. E.
(2003). Drosophila checkpoint kinase 2 couples centrosome
function and spindle assembly to genomic integrity.
Cell 113,87
-99.[CrossRef][Medline]
Takada, S., Kwak, S., Koppetsch, B. S. and Theurkauf, W. E.
(2007). grp (chk1) replication-checkpoint mutations and DNA
damage trigger a Chk2-dependent block at the Drosophila midblastula
transition. Development
134,1737
-1744.
Theurkauf, W. E. (1994). Immunofluorescence
analysis of the cytoskeleton during oogenesis and early embryogenesis.
Methods Cell Biol. 44,489
-505.[Medline]
Wieschaus, E. and Sweeton, D. (1988).
Requirements for X-linked zygotic gene activity during cellularization of
early Drosophila embryos. Development
104,483
-493.
Zaessinger, S., Busseau, I. and Simonelig, M.
(2006). Oskar allows nanos mRNA translation in Drosophila embryos
by preventing its deadenylation by Smaug/CCR4.
Development 133,4573
-4583.
Zurita, M., Reynaud, E. and Aguilar-Fuentes, J.
(2008). From the beginning: the basal transcription machinery and
onset of transcription in the early animal embryo. Cell Mol. Life
Sci. 65,212
-227.[CrossRef][Medline]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
Related articles in Development:
This article has been cited by other articles:
|
|
 |
|
  W. Tadros and H. D. Lipshitz The maternal-to-zygotic transition: a play in two acts Development, September 15, 2009; 136(18): 3033 - 3042. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Budirahardja and P. Gonczy Coupling the cell cycle to development Development, September 1, 2009; 136(17): 2861 - 2872. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
