RESEARCH ARTICLE
923
Development 136, 923-932 (2009) doi:10.1242/dev.031815
An essential role for the RNA-binding protein Smaug during
the Drosophila maternal-to-zygotic transition
Beatrice Benoit1,2, Chun Hua He3,4, Fan Zhang1, Sarah M. Votruba3, Wael Tadros3,4, J. Timothy Westwood5,
Craig A. Smibert3,6, Howard D. Lipshitz3,4 and William E. Theurkauf1,*
Genetic control of embryogenesis switches from the maternal to the zygotic genome during the maternal-to-zygotic transition
(MZT), when maternal mRNAs are destroyed, high-level zygotic transcription is initiated, the replication checkpoint is activated and
the cell cycle slows. The midblastula transition (MBT) is the first morphological event that requires zygotic gene expression. The
Drosophila MBT is marked by blastoderm cellularization and follows 13 cleavage-stage divisions. The RNA-binding protein Smaug is
required for cleavage-independent maternal transcript destruction during the Drosophila MZT. Here, we show that smaug mutants
also disrupt syncytial blastoderm stage cell-cycle delays, DNA replication checkpoint activation, cellularization, and high-level
zygotic expression of protein coding and micro RNA genes. We also show that Smaug protein levels increase through the cleavage
divisions and peak when the checkpoint is activated and zygotic transcription initiates, and that transgenic expression of Smaug in
an anterior-to-posterior gradient produces a concomitant gradient in the timing of maternal transcript destruction, cleavage cell
cycle delays, zygotic gene transcription, cellularization and gastrulation. Smaug accumulation thus coordinates progression through
the MZT.
INTRODUCTION
In oviparous organisms that deposit their eggs into the environment,
embryogenesis is initiated by a cleavage stage characterized by a
rapid, simplified S-phase/M-phase cell cycle and little or no zygotic
transcription (Etkin, 1988; 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
1
Program in Molecular Medicine and the Program in Cell Dynamics, University of
Massachusetts Medical School, 377 Plantation Street, Worcester, MA 01605, USA.
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. 4Program in Developmental and Stem Cell
Biology, Research Institute, Hospital for Sick Children, TMDT Building, 101 College
Street, Toronto, Ontario M5G 1L7, Canada. 5Department of Cell and Systems
Biology and Canadian Drosophila Microarray Centre, University of Toronto,
Mississauga, Ontario L5L 1C6, Canada. 6Department of Biochemistry, University of
Toronto, 1 King’s College Circle, Toronto, Ontario M5S 1A8, Canada.
2
*Author for correspondence (e-mail: [email protected])
Accepted 17 December 2008
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 highlevel 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
DEVELOPMENT
KEY WORDS: Cell cycle, Midblastula transition, Transcription, Drosophila
RESEARCH ARTICLE
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-toposterior 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
Drosophila stocks and genetics
smg and grp mutant embryos were generated by crossing homozygous
smg1/smgDf(3L)Scf-R6 (Dahanukar et al., 1999) 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 laserscanning microscope, as described previously (Sibon et al., 1997). Timelapse DIC microscopy was performed using Zeiss Axiovert –S100 inverted
microscope and 20⫻ 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 laserscanning 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.
Development 136 (6)
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-αTubulin (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
DEVELOPMENT
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Maternal control of the MZT
RESEARCH ARTICLE
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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
Smaug is required for replication checkpoint
activation during the MZT
Mutations in smaug (smg) disrupt nuclear organization during the
syncytial blastoderm stage (Dahanukar et al., 1999). 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 microarraybased 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 Cterminal 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.
DEVELOPMENT
Fig. 1. Replication checkpoint activation during the MZT. The DNA replication checkpoint is required for increases in interphase length during
syncytial blastoderm divisions 11-13. (A) Interphase length was assayed by injection of rhodamine-conjugated Tubulin and time-lapse confocal
imaging. Embryos mutant for smg, like grp replication checkpoint mutants, do not show increases in interphase length during the syncytial
blastoderm divisions. (B) Replication checkpoint function was assayed by co-injection of rhodamine-Tubulin and aphidicolin or carrier control. In
wild-type embryos, aphidicolin induced progressively longer interphase delays during cycles 11-13. By contrast, smg mutants showed only minimal
interphase delays in response to aphidicolin, and the delays did not increase with division cycle number. Each bar represents the mean interphase
length (in minutes) with standard deviations. The number of individual embryos scored is given in brackets.
926
RESEARCH ARTICLE
Development 136 (6)
Fig. 2. Smaug is required for zygotic transcription during the MZT. (A) Microarray analysis of zygotic gene expression in 2- to 3-hour-old wild-type
(wt) and smg mutant embryos. Expression is relative to mature, stage 14 oocytes, which contain the full maternal pool of mRNA. Maternal genes that
are not transcribed at the MZT represent ~80% of transcripts in early embryos, and are not shown. Class I zygotic genes are not present in oocytes (not
colored) and show high levels of expression in 2- to 3-hour-old embryos. One-hundred and forty-two of the 166 Class I genes required Smaug for
zygotic expression. Class II genes produce maternal transcripts that are stable in unfertilized eggs and show significantly increased expression in 2- to 3hour-old post-fertilization embryos. Three-hundred and fifty-eight of 395 Class-II genes require Smaug for zygotic expression. Class III genes produce
maternal transcripts that are degraded and then re-expressed in 2-3 hour post-fertilization embryos. Sixty-five of 408 Class III genes require Smaug for
expression. (B) Serine 2 phosphorylation of the RNA polymerase II CTD is linked to active transcription. Western blots reveal a dramatic increase in ser 2
phosphorylation between 2 and 4 hours of embryogenesis in wild type (wt) and mnk grp, but not in grp, smg or mnk; smg mutants. For each
genotype, lane 1 shows 0- to 1-hour-old embryos, lane 2 shows 1- to 2-hour-old embryos, lane 3 shows 2- to 3-hour-old embryos and lane 4 shows 3to 4-hour-old embryos. For each genotype, the top panel shows the hyperphosphorylated (II0, Ser2-P) and the hypophosphorylated (IIa) forms, detected
with an anti-RNA polII (ARNA3). The middle panel shows phosphorylation on Ser2 (II0) detected using the phospho-epitope specific H5 antibody. The
bottom panel shows α-Tubulin, which was used as a loading control. (C) Northern blots for miR-6, miR-286 and miR-3 revealed a large increase in wildtype 2- to 4-hour-old embryos, whereas expression was not detected smg mutants. (D) Heat map showing the behavior of 406 of the 410 miR-309dependent maternal mRNAs (from Bushati et al., 2008) in embryos from wild-type and smg-mutant females relative to wild-type stage 14 oocyte
reference RNA. These transcripts are unstable (green) in wild type, whereas almost 85% are stabilized in smg mutants.
miR-309-cluster microRNAs mediate Smaugdependent 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). Fourhundred 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
DEVELOPMENT
DNA damage and Chk2 activation can inhibit zygotic
transcription and cellularization (Takada et al., 2007), 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.
Maternal control of the MZT
RESEARCH ARTICLE
927
Fig. 3. Maternal mRNA degradation is
independent of the replication
checkpoint. Embryos mutant for smg are
defective in replication checkpoint activation
and maternal transcript destruction. (A-L) To
determine whether checkpoint defects lead
to a block in maternal transcript destruction,
grp checkpoint mutant embryos were
assayed for maternal cyclin A (A-F) and
cyclin B (G-L) mRNA expression by wholemount in situ hybridization. In wild-type
controls, both transcripts are expressed in
syncytial blastoderm embryos (S; A,G) and
are degraded during interphase of cycle 14
(14 D,J). The smg mutation blocks
destruction of these transcripts (C,F,I,L), but
the grp mutations does not (B,E,H,K).
Embryos are orientated with their anterior
pole facing leftwards. S, syncytial
blastoderm; 14, cycle 14. Scale bar: 100 μm.
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
Fig. 4. Smaug protein expression during early embryogenesis.
(A) Smaug protein expression in wild-type ovaries and embryos. Lane 1,
ovary (o); lanes 2-9, embryos were fluorescently labeled, hand selected
for specific division cycles (cycles indicated above lane) and subjected to
western blotting (E and L indicate early and late interphase 1); lanes 1017, western blots of pooled embryos aged 0 to 1 hour (1), 1 to 2 hours
(2), 2 to 3 hours (3) and 3 to 4 hours (4); lanes 10-13 are fertilized
embryos; lanes 14-17 are activated unfertilized eggs (unf). α-Tubulin is
used as a loading control. (B) Quantification of Smaug expression
relative to α-Tubulin for each of the lanes shown in A. In fertilized
embryos, Smaug protein levels increase progressively through the early
cleavage divisions, peak during cycles 11-13, and decline rapidly during
interphase 14. In unfertilized eggs, the protein accumulates rapidly
through the first 3 hours post egg deposition, and persists for 4 hours
(lanes 14-17).
DEVELOPMENT
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.
Smaug protein levels drop dramatically during early interphase
14, when transcription increases dramatically. In unfertilized eggs,
which do no initiate zygotic transcription, Smaug protein
accumulates with normal kinetics but then persists at high levels
(Fig. 4). Our gene expression profiling data show that maternal smg
mRNA is stable through the MZT. Smaug-dependent transcription
of early zygotic genes may therefore block smg mRNA translation
or decrease Smaug protein stability. This may prevent destruction of
zygotic mRNAs carrying Smaug-binding sites, and thus promote the
burst of transcription at the end of the MZT.
RESEARCH ARTICLE
Fig. 5. Generating a Smaug protein gradient. (A) Schematic
representation of the UAS-smg-bcd transgene (USB). The yeast
upstream activator sequence (UAS), Smaug coding sequence (Smaug
ORF, bold line) and bcd 3⬘ UTR (3⬘-UTR-bcd) are indicated.
(B,C) Immunolocalization of Smaug protein in embryos derived from
wild-type (wt) and smg females expressing the USB transgene. Some of
the immunolabeling in the latter is due to antibody recognition of a
truncated from of Smaug expressed in the smg1 mutants (see E).
Embryos are oriented with their anterior pole facing leftwards.
(D) Single confocal mid-section showing Smaug immunolabeling in an
embryo from a wild-type female expressing the USB transgene. Average
cortical pixel intensity in wild-type embryos (red line, n=4) was
subtracted from average cortical pixel intensity in embryos from wildtype females expressing USB (blue line, n=4). 100% designates the
anterior pole and 0% represents the posterior pole. (E) Western-blot
showing Smaug protein expression in embryos from wild type (wt),
smg-mutant females and smg-mutant females expressing the USB
transgene. Embryos were 0-3 hours old. Full-length Smaug (Smg) and a
truncated form of the protein (Smg*) expressed in smg1 mutants are
indicated. β-Tubulin (β-Tub) is a loading control.
anterior pole, respectively).
To assess the dynamics of the syncytial divisions and
cellularization, we analyzed early development by time-lapse
differential interference contrast (DIC) microscopy. In wild-type
syncytial blastoderm stage embryos, mitosis is synchronous or is
initiated at the poles and progresses in converging waves toward the
center (Foe and Alberts, 1983) (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
Development 136 (6)
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 midcellularization 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
DEVELOPMENT
928
Maternal control of the MZT
RESEARCH ARTICLE
929
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.
Smaug overexpression at the anterior pole did not advance
cellularization, suggesting that an additional process or component
defines the timing of cellularization under these conditions. To test
this hypothesis, we assayed cellularization and maternal transcript
destruction in embryos derived from females carrying extra copies
of a wild-type smg transgene, which uniformly overexpress Smaug
(see Fig. S4 in the supplementary material). These embryos
cellularized at the same time as wild type, and FISH analyses
indicated that destruction of maternal cyclin B mRNA was
completed synchronously over most of the embryo during
interphase 14 (data not shown). The timing and pattern of
cellularization and maternal transcript destruction was also normal
in embryos derived from wild-type females carrying the USB
transgene (see Fig. S4 in the supplementary material). These
embryos significantly overexpress Smaug at the anterior pole, and
levels drop to near wild type at the posterior pole. At or below wildtype levels, Smaug thus determines the kinetics of maternal
transcript destruction, zygotic transcription and cellularization.
Above wild-type levels, by contrast, an additional process or factor
appears to become limiting and determines the timing of the MBT.
DISCUSSION
Here, we have shown that Smaug is required to slow the cleavage
divisions, activate the DNA replication checkpoint and initiate high
level expression of the vast majority of genes that are transcribed
during the final stages of the MZT. Furthermore, Smaug is essential
for blastoderm cellularization, which marks the MBT. Significantly,
Smaug protein begins to accumulate during the early cleavage
DEVELOPMENT
Fig. 6. Smaug protein gradient triggers graded cell cycle delays and cellularization. (A,B) Embryos co-stained with a phospho-Tyrosine
antibody (grayscale and green) and TOTO3 (red), to visualize membranes and nuclei at cellularization. (A) Nuclear density is uniform along the
anterior-posterior axis of wild-type interphase 14 embryos undergoing cellularization. (B) An embryo expressing a Smaug gradient. The anterior pole
cellularizes at cell cycle 13 nuclear density (ANT inset), the middle cellularizes at cycle-14 density (MID inset) and the posterior pole is disorganized
(POS inset). (C,D) Embryos labeled for mitotic nuclei with anti-phospho-Histone-3 antibody (grayscale and green) and for DNA with TOTO3 (red).
Wild-type cycle 11 embryos divide synchronously along the anterior-posterior axis (C). Embryos expressing Smaug in a gradient show cell cycle
delays at the anterior pole (D). In this example, the posterior is in mitosis while the anterior is in late interphase or prophase (D). (E,F). Time-lapse
DIC microscopy of cellularization in wild-type and USB embryos. (E) Wild-type embryos consistently cellularize synchronously along the anteriorposterior axis (see Movie 1 in the supplementary material). (F) USB embryos, by contrast, consistently initiate cellularization at the anterior pole, and
membrane invagination progresses in a wave towards the posterior pole (see Movie 3 in the supplementary material). Region of nuclear dropout is
indicated by an asterisk. Embryos are orientated with their anterior pole leftwards. High-magnification views at anterior (ANT), middle (MID) and
posterior (POS) regions are shown below each whole embryo image. Arrows indicate the position of the celluarization front.
930
RESEARCH ARTICLE
Development 136 (6)
Fig. 7. A Smaug gradient triggers graded transcription and maternal mRNA destruction. (A-H) Zygotic expression of runt and slam in wildtype and USB embryos. In wild-type controls, both genes are expressed at low levels during cycle 13 (A,E) but are significantly upregulated during
interphase 14 (B,F). As cellularization is initiated, USB embryos express slam (C) and runt (G) only at the anterior pole, where Smaug expression is
highest. (I-L). In later embryos, slam expression is highest at the posterior pole and has begun to decline at the anterior, whereas the striped pattern
of runt expression has extended to the posterior pole (D,H). In wild-type embryos, maternal cyclin B mRNA is uniformly expressed during interphase
13 (I) and degraded throughout the embryo during interphase 14 (J). Only the pole cells retain maternal cyclin B transcript during interphase 14 (J).
(I) In USB embryos, cyclin B mRNA is degraded in an anterior-to-posterior gradient during interphase 13 (K,L). All transcripts were detected by
fluorescent whole-mount in situ hybridization, and embryos where co-stained with TOTO3 (red) to visualize nuclear density. Embryos are orientated
with anterior towards the left. Insets show higher magnification images of RNA (green) and DNA (red) in the anterior region of each embryo. Scale
bar: 100 μm.
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
miR-309-cluster
Zygotic
Transcription
Smaug
Transcription
activators
Maternal
mRNA
Replication
Checkpoint
Cell cycle
Fig. 8. Model for Smaug-dependent control of the MZT. We
propose that Smaug-dependent destruction of maternal mRNAs
encoding transcriptional repressors and cell cycle activators leads to
coordinated activation of the basal transcription machinery and the
replication checkpoint. An initial wave of transcription then produces
proteins and miRNAs that feed back to enhance maternal transcript
destruction (e.g. mir-309) and activate additional genes, thus
completing the transition to zygotic control of embryogenesis. The
replication checkpoint coordinately couples the cell cycle to the N/C
ratio and thus determines the number of divisions that are completed
before cellularization.
DEVELOPMENT
divisions, when maternal transcript destruction is initiated, and
Smaug levels peak during the syncytial blastoderm stage, when the
replication checkpoint is activated and high-level zygotic
transcription begins. In addition, expression of Smaug in an anteriorto-posterior gradient triggers an anterior-to-posterior gradient in the
timing of transcript destruction, checkpoint activation,
cellularization and gastrulation. Smaug is therefore the first maternal
or zygotic factor shown to control the timing of the MZT.
Over-expression of Smaug does not accelerate maternal transcript
destruction or cellularization, indicating that an additional factor or
process becomes limiting under these conditions. Smaug triggers
mRNA destruction by recruiting the CCR4/POP2/NOT complex,
which catalyzes poly-A tail removal (Semotok et al., 2005), 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
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 Smaugdependent 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.
S-phase length increases during the later cleavage stage divisions,
suggesting that maternally deposited replication factors are titrated as
the N/C ratio increases. The DNA replication checkpoint is activated
as S-phase slows, and thus senses the increasing N/C ratio (Sibon et
al., 1999; 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.
We thank Carla Klattenhoff, Saeko Takada, Seong-ae Kwak, Byeong Jik Cha,
Hanne Varmark and Birgit Koppetsch for critical comments on the manuscript
and technical advice on injection, time-lapse imaging, and FISH. Zak Razak,
Dahlia Kasimer, Linan Chen and Aaron Goldman provided guidance and
assistance with the analysis of the microarray data. H.D.L., C.A.S. and J.T.W.
are members of the Canadian Institutes for Health Research (CIHR) Team in
mRNP Systems Biology (CTP-79838). This work was supported by an operating
grant to H.D.L. from the CIHR (MOP 14409), the CIHR Team Grant, and by
grants to W.E.T. from the National Institute of General Medical Sciences,
National Institutes of Health (R01 GM50898) and the National Institute for
Child Health and Human Development, National Institutes of Health (R01
HD049116). Deposited in PMC for release after 12 months.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/136/6/923/DC1
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