1 Department of Biology, University of Washington, Seattle, WA 98195-1800,
USA
2 Laboratory of Molecular Biology, University of Wisconsin, Madison, WI 53706,
USA
* Author for correspondence (e-mail: gerold{at}u.washington.edu)
Accepted 24 October 2003
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SUMMARY |
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Key words: Early embryonic mitosis, Cdk1-CycB, Drosophila, Metaphase, Interphase
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Introduction |
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Studies of early developmental events in Xenopus have identified
at least two important temporal phases. First, timing of the early synchronous
cycles is based on oscillation of Cdk1-CycB activity
(Murray and Kirschner, 1991).
Second, later cell cycles slow down and become asynchronous, depending on the
nucleo-cytoplasmic ratio (Newport and
Kirschner, 1982
; Kirschner et
al., 1985
). This transition point for both cell-cycle rate and
synchrony is known as mid-blastula transition
(Yasuda and Schubiger, 1992
).
As in Xenopus, the nucleo-cytoplasmic ratio also regulates the
cell-cycle rate in Drosophila embryos
(Edgar et al., 1986
). However,
loss of synchrony does not occur at a single time point during the syncytial
blastoderm cycles (cycle 10-13) in Drosophila embryos
(Foe and Alberts, 1983
).
Similarly, zygotic gene transcription begins gradually and as early as cycle 8
in a gene-specific manner (Pritchard and
Schubiger, 1996
). During this maternal-zygotic transition,
depletion of mitotic cyclins might cause the elongation of interphase, thus
permitting sufficient time for zygotic transcription to proceed
(Edgar and Datar, 1996
;
Shermoen and O'Farrell,
1991
).
Over the last 60 years, many studies have analyzed the timing of the first
13 cycles in Drosophila using two different approaches. First,
embryos were fixed at different times and stained to estimate the duration of
both the total cell cycle and cycle phases based on the percentages of embryos
in each specific cycle phase. Second, time-lapse recordings of living embryos
were made using either DIC or confocal microscopy. The former approach
generated variable, thus potentially unreliable, results. For example,
metaphase length before cycle 10 was estimated as between 0.3 and 0.7 minutes
by Rabinowitz (Rabinowitz,
1941) and between 2.0 and 2.4 minutes by Stiffler et al.
(Stiffler et al., 1999
).
Although the latter method is useful for estimating total cell-cycle length,
the duration of cycle phases cannot be defined because chromosomal morphology
is not discernable with DIC optics. Although time-lapse recordings of embryos
labeled with fluorescent markers using confocal microscopy have been used to
analyze interphase and total cell-cycle time of cycles 11-13
(Sibon et al., 1997
;
Yu et al., 2000
), this method
is ineffective for imaging nuclei prior to their migration to the cortex
(cycle 10). In addition, long-term imaging with standard confocal wavelengths
is detrimental to living embryos
(Squirrell et al., 1999
).
These limitations result in a significant gap in our understanding of the
specific timing of cell-cycle events during the early embryonic cycles,
particularly those before cycle 10.
Two-photon laser-scanning microscopy (TPLSM) circumvents the technical
problems mentioned above, as it permits high resolution imaging deep within
thick, light-scattering tissues (Centonze
and White, 1998), as well as long-term imaging of living specimens
(Squirrell et al., 1999
).
Here, by using TPLSM, we characterize cell-cycle progression of preblastoderm
cycles (before cycle 10) and compare it with syncytial blastoderm cycles
(after cycle 10) in live embryos. Furthermore, we show that crucial CycB
concentrations control both interphase duration and the timing of
metaphase-anaphase transition.
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Materials and methods |
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Two-photon laser scanning microscopy (TPLSM)
Embryos were manually dechorionated and lined up on a 22x30 mm gluey
cover glass (Schubiger and Edgar,
1994). This cover glass was then taped to a plastic holder that
resembled a 25x76 mm glass slide with a 20x26 mm hole cut in the
center. To prevent dehydration, we covered embryos with a thin layer of
halocarbon oil (HC-700, Halocarbon Products Corp.), which is oxygen
permeable.
The two-photon imaging was performed on a BioRad Radiance 2000 System
equipped with a Mai Tai laser (Spectra Physics) set at 900 nm and a Nikon
Eclipse E600FN microscope with a 40x/1.3 Nikon Plan Apo oil objective.
Four-dimensional data were collected using Direct Detection System and
LaserSharp software (BioRad) with the zoom set at 1.7. Single slow scans of
two optical sections (5 µm apart) were taken every 10 seconds. The images
were compiled into time-lapse recordings using 4D Turnaround-Java software
(Thomas et al., 1996) and then
analyzed with 4D Viewer software (Thomas
et al., 1996
).
Embryos were imaged for up to 2 hours. The room temperature was 21.6±0.4°C (n=21 days). We were able to detect the histone-GFP signal as early as cycle 4. Two hours of laser scanning did not reduce the hatching rates: 86% of the imaged embryos hatched (n=21 randomly chosen embryos) compared with 89% in non-imaged histone-GFP control embryos (n=210). Time-lapse recordings from unhatched embryos were not used.
A two-photon optical workstation was used to generate bright field
transmission images (Fig.
6A,B). This transmitted light imaging is achieved by simply
allowing the scanning infrared laser beam (900 nm) to pass through the live
embryo and detecting it by an infrared photodiode
(Wokosin et al., 2003).
|
With fixed embryos, the duration of a cell-cycle phase was estimated as the percentage of embryos in the cell-cycle phase within a specific cycle. This fraction, also known as the mitotic index, represents the percentage of the total cell-cycle time spent in that phase. The total cell cycle duration for a given cycle was determined from the TPLSM recordings, thus the time of the cell-cycle phase could be calculated. To obtain meaningful estimations, several hundred embryos of each cell cycle were analyzed. The results were analyzed with the exact distribution of Wilcoxon-Mann-Whitney statistic test (StatXact 4.0 by Cytel Software).
CycB quantification
We collected images of embryos immunostained with the anti-CycB antiserum
Rb271 (Whitfield et al., 1990)
using a BioRad MRC-600 confocal microscope system as described in Stiffler et
al. (Stiffler et al., 1999
).
Images of mid-sections through each nucleus were obtained with an Olympus
microscope using a 60x oil objective and the confocal settings of Zoom
3.0 and Kalman 6. To outline energids (Fig.
3A), we used StackViewer, software written by Eli Meir
(http://www.beakerware.org/stackviewer).
With the same software we calculated the average pixel intensity of each
energid.
|
The ranking analysis is based on measurements from two-dimensional (2D)
images of mid-nuclear sections instead of three-dimensional (3D) data sets.
With confocal microscopy, fluorophore excitation occurs above and below the
plane of focus, resulting in fluorophore bleaching even outside the focal
plane (White et al., 2001).
Indeed, we observed substantial photobleaching when collecting 3D images for
each energid, making quantification of such data sets potentially unreliable.
Furthermore, when we performed a sample analysis of measurements from 3D data
sets we found greater differences in intensity among different areas compared
to the 2D measurements, but the results of the ranking among the three regions
remained the same, indicating that our method of measuring the 2D data sets is
justified.
Measurement of velocity of sister-chromatid separation in anaphase
High-resolution (1024x1024 pixels) time-lapse recordings were
collected with TPLSM settings described above. The recordings were analyzed by
using ImageJ
(http://rsb.info.nih.gov/ij/)
to measure both distance and time elapsed.
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Results |
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We analyzed 39 histone-GFP embryos and found that total cell-cycle time
steadily increases with every cycle after cycle 7
(Fig. 1I), which is three
cycles earlier than previously reported
(Foe and Alberts, 1983;
Warn and Magrath, 1982
;
Zalokar and Erk, 1976
).
Comparing the time of different cell-cycle phases, we found that only
interphase increases (Fig. 1I).
Other cell-cycle phases were unchanged: prophase-metaphase durations remained
about 230 seconds (s.d.=20 seconds) and anaphase-telophase duration about 160
seconds (s.d.=20 seconds). These observations were confirmed by analyses of
fixed embryos between cycles 6 and 9 (data not shown).
The interphase extension between cycles 7-10 is not dependent on a DNA replication checkpoint
In wild-type embryos, a depletion of factor(s) involved in DNA replication,
may lead to slower DNA replication and longer interphase (S-phase). In
DNA-replication checkpoint mutant embryos, such as grapes
(grp) or Mei-41, the rapid cycles continued after cycle 11
(Sibon et al., 1997;
Sibon et al., 1999
). Thus, it
has been proposed that interphase extension after cycle 10 depends on proper
function of the DNA-replication checkpoint
(Nyberg et al., 2002
;
Sibon et al., 1999
;
Sibon et al., 1997
).
To address whether the DNA-replication checkpoint function accounts for
interphase extension before cycle 10, we analyzed grp1
embryos from grp1/grp1; histone-GFP mothers
with TPLSM. We found that interphase increased from cycle 7 to cycle 10
(Fig. 1J), as in control
embryos, indicating that interphase extension between cycle 7 and cycle 10 is
DNA-replication checkpoint independent. This observation was confirmed by
estimating interphase time in fixed grp1 embryos (data not
shown). It should be noted that interphase duration after cycle 10 in
grp1 embryos extends, but less dramatically than in
wild-type embryos (Fig. 1J),
confirming previous observations (Sibon et
al., 1997; Sibon et al.,
1999
; Yu et al.,
2000
).
The preblastoderm cycles are metasynchronous in wild-type embryos
When analyzing fixed wild-type embryos, we observed embryos with nuclei at
different cell-cycle phases, indicating metasynchronous mitoses before cycle
10 (e.g. Fig. 2H). Thus, we
asked whether nuclei in different regions extend interphase coordinately. To
exclude fixation artifacts and to measure cell-cycle phases, we examined live
preblastoderm embryos with TPLSM. We determined the cell-cycle phases of
nuclei in different regions of single embryos at cycle 7. At this stage,
nuclei are spread along the anteroposterior axis.
Fig. 2A-F show an embryo with
regional differences in cell-cycle phases. We followed two nuclei, one located
posteriorly and the other in the middle of the embryo
(Fig. 2B). Both entered mitosis
(prophase) at the same time (Fig.
2B). Surprisingly, 220 seconds later the posterior nucleus entered
anaphase, while the medial one remained in metaphase
(Fig. 2C), entering anaphase 40
seconds later (Fig. 2D). Both
nuclei were in telophase by 520 seconds
(Fig. 2E) and entered the next
interphase at the same time (Fig.
2F). Thus, in this particular case, prophase-metaphase of the
medial nucleus was 40 seconds longer than that of the more posterior nucleus
although interphase showed little regional difference. Similar metasynchrony
was observed in all of the nine embryos analyzed at cycle 7. The average
prophase-metaphase time of medial nuclei was 20 seconds longer compared with
nuclei in the posterior region (P=0.0289; n=9,
Fig. 2G). This temporal
difference was compensated by a 20 second shorter anaphase-telophase in the
middle region so that nuclei entered the next interphase at similar times
(Fig. 2F,G). Therefore, total
cell-cycle duration of medial and posterior nuclei were not different (total
cell-cycle duration: medial=620±40 seconds, posterior=600±50
seconds, n=9).
|
To confirm these observations, the cell-cycle phases in four different regions (anterior, anterior medial, posterior medial and posterior) of embryos fixed between cycles 5 and 10 were determined. For each cycle, 700 to 1000 fixed embryos were immunostained and analyzed. This method has two advantages over live analysis: the entire embryo is accessible for analyses and all cycle phases can be identified. Between cycles 5 and 8, more metaphase and fewer anaphase nuclei were observed in the two middle regions compared with nuclei at the two polar regions (Fig. 2H), indicating longer metaphase and shorter anaphase in the middle than at the two polar regions. By contrast, there was little or no regional difference in the number of interphase, prophase and telophase nuclei (data not shown), supporting the data from live embryos that difference in metaphase duration between the polar regions and the middle is largely compensated by a concomitant change in anaphase duration. Interestingly, at cycles 9 and 10, cell-cycle phases were more synchronous than earlier cycles (data not shown).
As distances between two daughter nuclei at telophase showed no regional difference, we can assume that sister chromatids migrate the same distance in anaphase. Therefore, we tested whether differences in velocity of sister-chromatid migration in anaphase could account for the regional difference in anaphase-telophase timing (Fig. 2G). As shown in Table 1, between cycles 6 and 8, sister chromatids migrate significantly faster in the middle region of the embryo compared with the posterior region, accounting for the differences in anaphase duration between the regions.
|
Dosage effects of Cdk1-CycB activity on cell-cycle phases
Although immunocytochemistry can detect local difference of CycB levels, it
is not possible to measure regional differences of Cdk1-CycB activity within a
living embryo. Therefore, we varied Cdk1-CycB levels in the entire embryo by
changing maternal cycB gene copy number and then analyzed these
embryos for alterations in cell-cycle-phase time using TPLSM. We found that
higher CycB levels correlated with longer prophase-metaphase between cycles 6
and 10 (Fig. 4A).
|
When we analyzed anaphase-telophase duration in live embryos with more CycB, we did not observe shorter anaphase-telophase as expected (data not shown). It is possible that the time differences are too small to detect among embryos with different amounts of CycB. For this reason, we analyzed fixed embryos. Indeed, we estimated that anaphase in four cycB embryos was 5 seconds shorter than in two cycB embryos and 13 seconds shorter than in one cycB embryos (Fig. 4C). Thus, the longer metaphase in four cycB embryos was compensated up to 50% by a shorter anaphase. Varying CycB levels produced incomplete compensation, indicating that anaphase duration is more sensitive to local differences in CycB concentration than global differences. Nevertheless, these results suggest that anaphase duration is affected by differing CycB levels, whether these differences occur within an embryo or among embryos of different maternal genotypes. Perhaps, there are other unknown factors that can also affect anaphase duration.
Interestingly, we found that increased CycB levels correlated with shorter interphase after cycle 7 (Fig. 4B) in live embryos. These observations were confirmed from data calculated from fixed embryos (Fig. 4C). Thus, longer metaphase is almost completely compensated by shorter anaphase and interphase.
Furthermore, interphase extension begins earlier in embryos with less
maternal CycB than in those embryos with more CycB
(Fig. 4B), supporting the idea
that interphase extension occurs when CycB becomes limited
(Edgar et al., 1994). A
crucial level of CycB would be reached earlier in embryos receiving less
maternal CycB.
Velocity of sister chromatid separation in anaphase correlates with CycB levels
We used one cycB and four cycB embryos to test whether
increased amounts of CycB resulted in shorter anaphase time because sister
chromatids moved faster. As shown in Table
1, whenever more CycB was present, sister chromatids moved
significantly faster, whether the difference of CycB occurred within an embryo
or among embryos of different maternal genotypes.
Cell cycle progression after cycle 10
We observed that metasynchrony before cycle 10 resulted from regional
differences in metaphase and anaphase time. As metasynchronous mitoses were
also observed after cycle 10 (Foe and
Alberts, 1983), we asked whether it was also a result of longer
metaphases in the middle region of the embryo. We analyzed live embryos with
TPLSM and found that nuclei at the two polar regions enter the succeeding
blastoderm cycle earlier than nuclei in the middle, confirming observations
made by Foe and Alberts (Foe and Alberts,
1983
). Surprisingly, we further found that both interphase and
total cycle of cycles 11 and 12 were longer in the middle regions than at the
posterior poles (Fig. 5A-D),
although no regional difference was observed in any other cell-cycle phases.
At cycle 11, interphase was 370 seconds in the middle (s.d.=30 seconds) and
350 seconds in the posterior region (s.d.=30 seconds, n=36). At cycle
12, this regional difference increased: interphase was 510 seconds in the
middle (s.d.=50 seconds) and 460 seconds in the posterior region (s.d.=50
seconds, n=17, Fig.
5E). Similar observations were made for cycle 13 (data not shown).
Thus nuclei in the posterior region entered a blastoderm cycle earlier than
nuclei in the middle. With each blastoderm cycle, this regional difference
increased, up to a difference of 120 seconds at the beginning of cycle 13.
These observations clearly demonstrate that the reason for earlier entry of
succeeding blastoderm cycle at the two poles is a lengthening of interphase in
the middle region of the embryo.
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Discussion |
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A model to explain the axial expansion process (between cycle 4 and cycle
8) is based on solation-contraction of the microfilament network within the
embryo (for details, see von Dassow and
Schubiger, 1994). Briefly, according to this model, local solation
(disassembly) of the contractile microfilament network in the center of embryo
will cause the microfilament network attached to the cortex to contract away
from the solated site in the center of the embryo. Nuclei and cytoplasm in the
center of the embryo move towards the two polar regions because the tension
generated by the contraction of the microfilament network is greatest along
the anteroposterior axis (von Dassow and
Schubiger, 1994
). This poleward cytoplasmic movement in the
interior would force cortical material to flow from poles towards the middle
region, where the cytoplasm then moves inwards. This then generates two
circular cytoplasmic movement during prophase and metaphase
(Fig. 6B,C). However, the
nature of the `solating agent' remains unknown in this model
(von Dassow and Schubiger,
1994
). We observed that Cdk1-CycB affects both microtubule and
microfilament dynamics (Ji et al.,
2002
) and that CycB is higher in the middle region from prophase
to anaphase (Fig. 3E),
suggesting that Cdk1-CycB is a likely candidate for the solating agent that
initiates axial expansion. Therefore, we see a positive feedback loop between
CycB distribution and cytoplasmic flow
(Fig. 6). This feedback loop is
disrupted by CycB degradation at anaphase, when we observe a slight backward
cytoplasmic flow.
According to this scenario, the global cytoplasmic movement and local
oscillation of mitotic cyclin concentration are the two key factors generating
metasynchronous mitoses during preblastoderm cycles. Interestingly, axial
expansion only occurs between cycle 4 and 8
(Baker et al., 1993), the same
period during which we observe regional differences in metaphase and anaphase
duration. This global cytoplasmic movement is not observed after cycle 8,
correlating with the observation of little regional difference in metaphase
and anaphase duration after cycle 8. Direct observation of CycB movement in
embryos with CycB-GFP fusion proteins
(Huang and Raff, 1999
) would
be ideal. However, we were unable to detect CycB-GFP signal prior to cycle
10.
Different control mechanisms of interphase extension before and after cycle 10
Interphase extension after cycle 10 has been explained in two ways. Edgar
et al. (Edgar et al., 1994)
observed that decreasing CycB correlates with longer interphase after cycle
10, thus proposed that interphase extension after cycle 10 was due to CycB
limitation. However, based on the observation that fast cycles continue after
cycle 10 in grp mutant embryos, Sibon et al.
(Sibon et al., 1997
) proposed
that in wild-type embryos depletion of factors involved in DNA replication
causes longer interphase after cycle 10 and the interphase extensions are
regulated by the DNA-replication checkpoint pathway.
Several of our observations might resolve this controversy. We report here
that interphase extension occurs in grp mutant embryos before cycle
10 (Fig. 1J), thus we propose
that interphase extension before cycle 10 is solely due to CycB limitation.
This is further supported by the following observations. First, interphase
extension occurs at an earlier cycle when maternal CycB is reduced and later
when CycB is increased (Fig.
4B). Second, looking within a specific cycle, interphase is longer
when CycB is lower and shorter when CycB is higher
(Fig. 4B,C). Third, global CycB
levels start to oscillate at the beginning of cycle 6 or 7 in wild-type
embryos (Edgar et al., 1994),
exactly the same time when interphase duration starts to increase
(Fig. 1I).
We also propose that after cycle 10, interphase extension is under control
of both CycB limitation and the DNA-replication checkpoint. It was reported
that grp1 is a null allele
(Fogarty et al., 1997). We
observed that interphase continuously extends in grp1
embryos after cycle 10, although this extension is not as extensive as in
wild-type embryos (Fig. 1J).
This observation supports the idea that limitation of CycB is responsible for
this increase.
With these two proposals in mind, we re-examined the interphase extension.
Before cycle 10, interphase extension occurs coordinately in all nuclei and
the nuclei doubling time shows no regional difference. After cycle 10, nuclei
divide slower in the middle because interphase in this region is longer, which
correlate with an increase of nuclear density in this region of the embryo
after cycle 10 (Blankenship and Wieschaus,
2001) (G. K. Yasuda, PhD thesis, University of Washington, 1992).
In many organisms, a higher nucleocytoplasmic ratio correlates with a slower
cell cycle (Sveiczer et al.,
2001
). A venerable hypothesis is that higher nuclear density could
result in an earlier depletion of factors necessary for DNA replication, such
as deoxynucleotide triphosphates. This may result in slower DNA replication,
thereby specifically prolonging interphase and ultimately total cell-cycle
length.
How do Cdk1-CycB levels affect the velocity of sister chromatid separation?
Currently, there are two major mechanisms proposed to regulate sister
chromatid separation in anaphase. First, disassembly of microtubules in
kinetochore regions shortens kinetochore microtubules and generates the force
that pulls the sister chromatids apart once cohesin is cleaved by separase
(Compton, 2002). Second, the
disassembly of spindle microtubules at the centrosomal region induces the
poleward microtubule movement, which then generates the force that separates
the sister chromatids (Compton,
2002
). In syncytial blastoderm Drosophila embryos (cycles
10 to 13), it has been documented that the poleward microtubule movement is
the key component that separate the sister chromatids in anaphase A, whereas
the disassembly of microtubules at the kinetochore is a minor factor
(Maddox et al., 2002
).
How does Cdk1-CycB affect velocity of sister chromatid separation? We have
shown that CycB levels negatively affect microtubule stability: higher
Cdk1-CycB levels lead to less stable microtubules and, correspondingly, lower
Cdk1-CycB levels lead to more stable microtubules
(Stiffler et al., 1999;
Ji et al., 2002
). It also
takes a longer time to form a stable metaphase configuration when CycB levels
are elevated. Furthermore, when more CycB is present the microtubules of the
metaphase spindle are weaker than when less CycB is available. We speculate
that the disassembly of spindle microtubule is faster or more efficient at the
centrosomal regions because there are fewer microtubules at the centrosomal
regions in embryo with higher Cdk1-CycB activity
(Stiffler et al., 1999
),
contributing to the faster poleward microtubule movement. Alternatively,
Cdk1-CycB might affect the sister chromatid movement via its target proteins
in the centrosomal regions and/or the midzone where the interpolar
microtubules overlap. For example, Cdk1-CycB phosphorylates p93dis1, which is
enriched in the centrosomal regions
(Nabeshima et al., 1995
) and
kinesin Eg5 (Drosophila homolog KLP61F), which accumulates on the
midzone after phosphorylation (Blangy et
al., 1995
; Sawin and
Mitchison, 1995
; Sharp et al.,
1999
). Mutation of p93dis1 in fission yeast results in failure in
sister chromatid separation (Nabeshima et
al., 1995
), while the phosphorylated bipolar kinesin KLP61F is
thought to be involved in sister chromatid separation by regulating sliding of
the interpolar microtubules in anaphase
(Sharp et al., 1999
).
Our observation that Cdk1-CycB affects the velocity of sister chromatid movement during anaphase supports the idea that microtubule dynamics contribute to the mechanical force for sister chromatid separation. This novel observation provides an entry point to further investigate the molecular mechanism that leads to disassembly of spindle microtubules by analyzing, for example, the possible functions of the target proteins of Cdk1-CycB in the centrosomal region.
Limitations and opportunities using the TPLSM
The TPLSM provides new opportunities for precisely analyzing the timing of
biological events (Squirrell et al.,
1999). Its major advantages over conventional confocal imaging are
that the two-photon excitation generates less photo damage to thick living
objects, and is particularly successful at imaging fluorescent signals deep in
the specimen (Centonze and White,
1998
; Squirrell et al.,
1999
; White et al.,
2001
). Indeed, we found that with traditional confocal microscopy,
GFP signals prior to cycle 10 were difficult to detect. In addition, we
frequently observed cell-cycle arrest and chromosome bridges in the region of
focus, indicating phototoxic effects, confirming the observations made by
others (Clarkson and Saint,
1999
).
Using TPLSM, we found that in the earliest embryonic stages, an abundance of many maternal proteins obscure the observation of the product at the target. However, after the first few cycles, depending on the protein, the maternal storage declines and the GFP signal from the fusion protein becomes target specific. The time point of detection depends on the localization, the amount and function of the protein. For example, histone-GFP on chromosome is recognized after cycle 4, while tubulin-GFP can only be recognized on microtubules after cycle 8 (J.-Y.J., J.M.S. and G.S., unpublished).
The ability to detect fluorescent label within the preblastoderm embryo and
follow changes in this signal over a relatively long period of time without
affecting viability provides opportunity to study this early developmental
stage that has been previously inaccessible. For example, there are an
abundance of maternal effect mutations that develop apparently normal up to
cycle 5 or 6, after which development arrests, a phenomenon referred to as
`epigenetic crisis' (Counce,
1973). This phenomenon is observed in many vertebrates and
invertebrates, indicating that mid-cleavage is a critical developmental stage
(Counce, 1973
). Although some
of the Drosophila mutations have been molecularly identified, a
phenotypic analysis using the TPLSM to assess the function of these genes will
increase our understanding of this developmentally critical period.
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ACKNOWLEDGMENTS |
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