Current Biology 18, 1514–1519, October 14, 2008 ª2008 Elsevier Ltd All rights reserved
DOI 10.1016/j.cub.2008.08.044
Report
Spindle Positioning in Mouse
Oocytes Relies on a Dynamic
Meshwork of Actin Filaments
Jessica Azoury,1,4 Karen W. Lee,1,4 Virginie Georget,2
Pascale Rassinier,1 Benjamin Leader,3
and Marie-Hélène Verlhac1,*
1Unité Mixte de Recherche 7622
Centre National de la Recherche Scientifique
Université Pierre et Marie Curie
Bat. C, 5e, 9 Quai Saint Bernard
75005 Paris
France
2Institut de Biologie Intégrative
IFR83
Université Pierre et Marie Curie
Bat. B., 7e, 9 Quai Saint Bernard
75005 Paris
France
3Department of Genetics
Harvard Medical School
Howard Hughes Medical Institute
200 Longwood Avenue
Boston, Massachusetts 02115
Summary
Female meiosis in higher organisms consists of highly asymmetric divisions, which retain most maternal stores in the oocyte for embryo development. Asymmetric partitioning of the
cytoplasm results from the spindle’s ‘‘off-center’’ positioning, which, in mouse oocytes, depends mainly on actin filaments [1, 2]. This is a unique situation compared to most systems, in which spindle positioning requires interactions
between astral microtubules and cortical actin filaments
[3]. Formin 2, a straight-actin-filament nucleator, is required
for the first meiotic spindle migration to the cortex and cytokinesis in mouse oocytes [4, 5]. Although the requirement for
actin filaments in the control of spindle positioning is well established in this model, no one has been able to detect them
in the cytoplasm [6]. Through the expression of an F-actinspecific probe and live confocal microscopy, we show the
presence of a cytoplasmic actin meshwork, organized by
Formin 2, that controls spindle migration. In late meiosis I,
these filaments organize into a spindle-like F-actin structure,
which is connected to the cortex. At anaphase, global reorganization of this meshwork allows polar-body extrusion. In
addition, using actin-YFP, our FRAP analysis confirms the
presence of a highly dynamic cytoplasmic actin meshwork
that is tightly regulated in time and space.
Results and Discussion
Oocytes from all species are blocked in the ovary in prophase I
of meiosis. At this stage, they contain a big nucleus, the germinal vesicle (GV). After meiosis resumption, oocytes undergo
meiotic maturation, which consists of two consecutive
*Correspondence: [email protected]
4These authors contributed equally to this work
asymmetric divisions resulting in the formation of a large haploid oocyte and small polar bodies (Figure 1A). After the first
polar-body extrusion (PBE), the second meiotic spindle forms
rapidly and is anchored to the cortex via actin microfilaments
[1, 7, 8]. Vertebrate oocytes remain arrested in metaphase II
(MII) until fertilization.
Meiotic spindle migration is crucial for the asymmetry of the
first meiotic division. The meiosis I spindle forms slightly
off-center because of a slight GV off-centering, and the spindle
migrates along its long axis toward the nearest cortex (Figure 1A). This positioning mainly depends on actin filaments and
the straight-microfilament nucleator, Formin 2 (fmn2) [1, 2, 4, 5].
To visualize the actin filaments responsible for this migration, we used an F-actin-specific probe, Utr-GFP, which contains the calponin-homology domain of an actin-binding protein, utrophin [9, 10]. This probe has been demonstrated to
specifically label actin filaments in vitro and in vivo in many
systems [10, 11]. Using time-lapse video microscopy, we
imaged Utr-GFP-expressing mouse oocytes during meiotic
maturation. Utr-GFP efficiently labeled cortical actin filaments,
but low resolution prevented us from detecting any clear cytoplasmic labeling (Figure 1B). Thus, we imaged the Utr-GFP
labeling by using high-resolution live confocal microscopy.
In control immature oocytes, we observed actin microfilaments around the GV (Figure 1C, cont GV, indicated by arrowheads). These microfilaments were absent in fmn22/2 and
cytochalasin D (ccd)-treated oocytes; instead, large patches
of staining surrounding the GV were observed (Figure 1C,
fmn22/2 GV and cont + ccd GV, indicated by asterisks). The
absence of microfilament labeling after ccd treatment proves
that Utr-GFP specifically labels F-actin.
In the cytoplasm of control metaphase I (MI) oocytes, actin
microfilaments formed a meshwork consisting of numerous
thin filaments and crossing points, from which several filaments emanated (Figure 1C, lower cont panel, indicated by arrowheads). This F-actin meshwork was extremely dynamic
(Movie S1 available online), with the crossing points moving
at approximately 0.14 mm/s (n = 17). In fmn22/2 and ccdtreated oocytes, this meshwork was absent (Figure 1C,
fmn22/2 and cont + ccd). Instead, patches of staining (of 2
to 6 mm in size) similar to the ones described around the GV
were observed (Figure 1C). These patches were absent in
wild-type oocytes, whereas they were present in fmn2+/2
(30%), in fmn22/2 (70%), and in all oocytes treated with ccd
(Figure 1D). They might correspond to aggregates of actin
in the absence of polymerization. Nevertheless, in fmn22/2
oocytes, the cortical staining was not altered (Figure 1E), a finding that is consistent with previous observations showing that
Formin 2 is not required for cortical F-actin [5].
During spindle migration, actin filaments progressively organized into a ‘‘spindle-like’’ structure (n = 80; Figure 2A and
Movie S2). Most of these filaments formed along the microtubules (Figures 2B and 2C, indicated by arrowheads). They assembled into an F-actin ‘‘cage’’ surrounding the microtubule
spindle, with some filaments also present inside it (Figures
2C and 2D and Movies S3 and S4). This organization was not
observed in the absence of the microtubule spindle, i.e., after
nocodazole treatment (n = 12; Figure 2E and Movie S5). Thus,
Formin 2-Organized Actin Meshwork in Mouse Oocytes
1515
Figure 1. In Vivo Observations of F-Actin in
Maturing Mouse Oocytes
(A) Scheme representing the process of meiotic
maturation in mouse oocytes. Chromosomes
appear in red, microtubules in blue, and cortical
actin filaments in green. GV, germinal vesicle;
BD, germinal vesicle breakdown; PBE, first
polar-body extrusion; MII, metaphase II arrest.
The timings of the different stages are given.
(B) Image showing the labeling from a wild-type
(cont) oocyte expressing Utr-GFP (green) and Histone-RFP (red) observed at BD + 6 hr by live video
microscopy. The scale bar represents 10 mm.
(C) Images showing the labeling from wild-type
(left, cont), fmn22/2 (middle, fmn22/2), and
wild-type treated with cytochalasin D (right,
cont + ccd) oocytes expressing Utr-GFP observed in GV (upper panel) and in the cytoplasm
at BD + 5 hr (lower panel) by live confocal microscopy. One Z plane is shown. The arrowheads
point toward actin filaments and crossing points.
The asterisks show patches of staining with the
Utr-GFP in fmn22/2 and control oocytes treated
with ccd. The scale bar represents 10 mm.
(D) Histogram showing the percentage of Utr-GFP
expressing oocytes from different strains (OF1,
fmn2+/2, and fmn22/2) showing patches of UtrGFP staining when raised under normal conditions
and treated with cytochalasin D (+ ccd). The
patches were counted by live video microscopy.
‘‘n’’ is the number of oocytes examined.
(E) Image showing the labeling from a fmn22/2
oocyte expressing Utr-GFP (green) and HistoneRFP (red) observed at BD + 6 hr by live video
microscopy. The scale bar represents 10 mm.
the F-actin ‘‘spindle’’ and the microtubule spindle are closely
associated. This is consistent with the fact that actin filaments
can bind microtubules in Xenopus egg extracts [12]. Indeed,
microtubule and F-actin interactions are required for a wide
variety of processes, including meiosis [11, 13–15]. For instance, in Xenopus oocytes, myosin X links F-actin and microtubules and is essential for spindle anchoring and asymmetric
division [16].
Late in meiosis I, we detected an accumulation of straight actin microfilaments connecting the closest spindle
pole to the cortex (Figures 2A and 2B, indicated by arrowheads). Chromosomes
are still able to migrate to the closest
cortex in nocodazole-treated oocytes
[2]. Under this condition, F-actin formed
in direct contact with the chromatin
mass, mostly in the region facing the
nearest cortex (fluorescence intensity
was approximately 42% higher, n = 12;
Figure 2E and Movie S5; see Supplemental Data for quantification). The
asymmetric formation of microfilaments
and their redistribution might correlate
with the asymmetric localization of
mPARD6a at the spindle pole closest
to the cortex and the redistribution of
mPARD6a toward the region of chromosomes facing the cortex in the absence
of the microtubule spindle [17]. Indeed,
a regulator of the PAR3/PAR6/aPKC complex, the RhoGTPase
Cdc42, has been reported to play a role in meiosis I spindle
migration [18].
At anaphase, the F-actin spindle became more prominent
(Figures 3A and 3B, lower right corner). Moreover, the density
of the cytoplasmic meshwork increased remarkably (w50%
increase in intensity) while the cortical staining decreased
(w20%; Figure 3B and Supplemental Data for quantification).
Current Biology Vol 18 No 19
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Figure 2. In Vivo Observations of F-Actin by Live
Confocal Microscopy of Control Oocytes during
Spindle Migration
(A) Representative sequence of images from control wild-type oocytes expressing Utr-GFP (gray
levels) and Histone-RFP (red) observed during
spindle migration at the end of meiosis I. The
lower panel shows a zoom of selected time
frames. One Z plane is shown. The arrowhead
points toward the anchoring filaments. Times
after BD (time 0) are indicated in hours. Scale
bars represent 10 mm.
(B) Representative image from a control wildtype oocyte expressing Utr-GFP (green) and
Map7-RFP (red) showing the anchoring of spindle poles via actin filaments to the cortex (indicated by the arrowhead) at the end of metaphase
I. The maximal projection of three Z planes
(Z step, 1 mm) is shown. The scale bar represents
10 mm.
(C and D) Representative images from a control
wild-type oocyte expressing Utr-GFP (green)
and Map7-RFP (red) showing the organization
of F-actin and microtubules in the metaphase I
spindle. In (C), the same spindle is shown from
a top section (upper panel) and a middle section
4 mm apart (lower panel). Arrowheads point toward F-actin. In (D), the maximal projection of
three Z planes (Z step, 1 mm) from a metaphase
I spindle orientated perpendicular to the plane
of observation shows cross-sections of actin
microfilaments, both surrounding (indicated by
arrowheads) and inside (indicated by the asterisk) the microtubule spindle. Scale bars represent
5 mm.
(E) Representative sequence of images from control oocytes expressing Utr-GFP (green) and Histone-RFP (red) that were treated with nocodazole
(cont + noco) and observed during chromosome
migration at the end of meiosis I. One Z plane is
shown. The nearest cortex appears on the upper
left corner. The dotted white ellipse indicates the
position of chromosomes at the beginning of the
time lapse. Times after BD (time 0) are indicated
in hours. The scale bar represents 10 mm.
In fact, the cortex thickness dropped from w2 mm to 1 mm, and
a subcortical F-actin layer of 2 mm thick formed (n = 7; Figure 3B, lower panel cortex). These global changes of cortical
and cytoplasmic F-actin during anaphase might ensure successful PBE. In MII, the F-actin spindle reappeared below
the cortical differentiated zone (CDZ; Figure 3C, indicated by
the arrowhead).
We did not observe any F-actin spindle in fmn22/2 (n = 37;
Figure 3D) nor in ccd-treated oocytes (n = 22; Figure 3E). Under
these conditions, the microtubule spindle organizes normally,
and anaphase takes place without cytokinesis [5]. Hence, the
absence of an F-actin spindle in these oocytes is an additional
proof that the Utr-GFP binds to F-actin specifically. The meshwork-density rise of F-actin at anaphase also did not occur in
fmn22/2 oocytes (Figure 3D). Instead, in both fmn22/2 and
ccd-treated oocytes, we observed patches of staining close
to chromosomes, some of which remained between the
two sets of chromatin (Figures 3D and 3E and Movies S6 and
S7). Interestingly, active phospho-myosin II also mislocalizes
between the two sets of chromosomes at anaphase in
fmn22/2 oocytes [5].
Using an F-actin-specific probe, we showed the presence of
a cytoplasmic actin meshwork that had never been visualized
[6]. In fact, previous studies with oocytes expressing actinGFP (or its spectral variants) had failed in visualizing this meshwork because of the high background from diffusible fluorescent actin. However, these probes are well established for
studying actin dynamics in living cells with fluorescence recovery after photobleaching (FRAP) [19–21]. Hence, we used
actin-YFP to study actin dynamics during meiosis I spindle
migration.
The cortex of MII oocytes is known to be rich in polymerized
actin filaments, particularly in the CDZ overhanging the chromosomes (Figure 4A) [1, 22]. Because it was the first time
that quantitative FRAP was performed in mouse oocytes, we
used cortical regions of MII oocytes as a control. In these regions, the fluorescence intensity did not reach its initial value
after photobleaching, showing the presence of a polymerized
immobile fraction (IF; Figure 4B). In addition, fitting of the FRAP
data showed that the mobile-fraction recovery corresponds to
a biexponential function (Supplemental Data). Therefore,
FRAP analysis detected three fractions of actin, an IF and
two mobile fractions: a fast mobile fraction (FMF) corresponding to the diffusing G-actin and a slower mobile fraction (SMF)
representing F-actin integrated into dynamic microfilaments.
Whereas the IF was greater in the CDZ than in the cortex
Formin 2-Organized Actin Meshwork in Mouse Oocytes
1517
Figure 3. In Vivo Observations of F-Actin by Live
Confocal Microscopy of Control Oocytes,
Fmn22/2 Oocytes, and Control Oocytes Treated
with Cytochalasin D during Anaphase
(A) Image from control wild-type oocytes expressing Utr-GFP (green) and Histone-RFP (red)
that were observed at anaphase. One Z plane is
shown. The scale bar represents 10 mm.
(B) Representative images from control wild-type
oocytes expressing the Utr-GFP that were observed 30 min before (left) and at anaphase (right)
to show the changes in F-actin organization, both
in the cytoplasm (cyto, upper panel) and in the
cortex (cortex, lower panel). One Z plane is
shown. Scale bars, 5 mm. An image of a spindlelike structure observed at anaphase is shown in
the lower right corner. Utr-GFP appears in green,
and Histone-RFP appears in red. One Z plane is
shown. The scale bar represents 10 mm.
(C) Zoom from a control wild-type oocyte expressing Utr-GFP (green) and Histone-RFP (red)
that was observed in MII. One Z plane is shown.
The arrowhead points toward the MII spindle,
which is located below the cortical differentiated
zone, perpendicular to the plane of observation.
The asterisk marks the polar body. The scale
bar represents 10 mm.
(D) Representative sequence of images from
fmn22/2 oocytes expressing Utr-GFP (green)
and Histone-RFP (red) that were observed at
the end of meiosis I. One Z plane is shown. Times
after BD (time 0) are indicated in the upper right
corner in hours. The scale bar represents 10 mm.
(E) Representative sequence of images from control oocytes treated with cytochalasin D expressing Utr-GFP (green) and Histone-RFP (red) that
were observed at the end of meiosis I. One Z
plane is shown. Times after BD (time 0) are indicated in the upper right corner in hours. The scale
bar represents 10 mm.
away from chromosomes (‘‘cortex away’’), the FMF was
greater in the cortex away than in the CDZ (Figure 4F). In addition, the polymerized actin fraction (SMF) was more dynamic
in the cortex away than in the CDZ (ts of 8.44 s compared to
16.68 s, Figure 4E). Taken together, our data show that microfilaments in the CDZ are more abundant and less dynamic than
those present in other cortical regions. These results demonstrate for the first time differences in microfilament stability
in these cortical regions, differences that are consistent with
what was previously described [1, 22].
The technique was also sensitive enough to detect microfilaments in the cytoplasm of MII oocytes. In fact, like in the
cortex, the three fractions of actin were detected, confirming
the presence of a meshwork of cytoplasmic F-actin. The distribution of these fractions and their dynamics showed that the
cytoplasm located near chromosomes is richer in stable microfilaments than the cytoplasm away from chromosomes
(Figures 4C, 4E, and 4F). This enrichment might play a role in
MII spindle anchoring, as previously suggested [1, 7, 8].
When we analyzed the recovery profiles of actin-YFP in the cytoplasm of MI
oocytes during migration (GV breakdown + 6 hr [BD + 6 hr]), no significant
difference was observed in either the
distribution or the dynamics of the actin
fractions within the cytoplasm, both
near and away from chromosomes (compare orange and green
curves in Figure 4D; also see Figures 4E and 4F). Hence, a different level of actin-cytoskeleton regulation, such as actomyosin
contractions, might govern the spindle migration at this
stage [23].
Our live confocal data showed an intensive reorganization of
the F-actin meshwork at anaphase. Similarly, our FRAP analysis also indicated that significant variations in the amount and
dynamics of polymerized microfilaments took place at this
stage (BD + 8 hr). Whereas more stable actin microfilaments
accumulate in the cytoplasm near chromosomes, polymerized
microfilaments become more dynamic away from chromosomes (Figures 4D–4F). Hence, the increase in stability close
to chromatin might ensure chromosome anchoring to the cortex. This reorganization observed by FRAP reinforces our live
confocal data.
Using two different approaches, we show the presence of
polymerized actin microfilaments in the cytoplasm of mouse
oocytes throughout maturation. Perhaps these differences in
Current Biology Vol 18 No 19
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Figure 4. Spatiotemporal Study of Actin-YFP Dynamics in Mouse Oocytes
(A) MII oocyte expressing actin-YFP (green) and Histone-RFP (red). The schematic on the right shows the corresponding regions: the cortex above (CDZ)
and away from chromosomes (cortex away) and the cytoplasm near (cytoplasm near) and away from chromosomes (cytoplasm away). A point bleach was
performed in each defined region (colored points). The scale bar represents 10 mm.
(B–D) Profiles of actin-YFP fluorescence recovery after photobleaching over time in the (B) cortex (the CDZ and the cortex away from chromosomes) and (C)
cytoplasm (near and away from chromosomes) of MII oocytes and in the (D) cytoplasm (near and away from chromosomes) of MI oocytes at 6 hr and 8 hr
after BD (BD+6 and BD+8). In (B), both the mean curves before fitting (light gray and light orange) and the fitted curves (black and red) are shown. The number
in parenthesis corresponds to the number of oocytes analyzed. See the Supplemental Experimental Procedures for FRAP analysis.
(E) Table showing the measures of ts and tf in different regions at different stages of meiotic maturation. See Supplemental Experimental Procedures for the
signification of the ts and tf. The statistical significance of differences was assessed with a t test with a confidence interval of 95% (**p < 0.001 and *p < 0.05).
(F) Histogram showing the distribution of the three actin fractions in different regions at different stages of meiotic maturation. See the Supplemental
Experimental Procedures for the signification of IF, SMF, and FMF. The statistical significance of differences was assessed with a t test with a confidence
interval of 95% (**p < 0.001 and *p < 0.05).
(G) Model for the F-actin meshwork required for spindle migration of mouse oocytes. The F-actin is in green, and the chromosomes are in red. The light green
represents the subcortical layer observed at anaphase.
the amount of polymerized actin in different areas and/or at
different times during meiotic maturation play a role in the
asymmetry of meiosis I division.
We propose that, in late meiosis I, spindle migration depends on the organization of a dense cytoplasmic meshwork
of dynamic actin filaments (Figure 4G). Antibody injection in
Formin 2-Organized Actin Meshwork in Mouse Oocytes
1519
mouse oocytes has previously suggested that myosin IIA is
required for this migration [23]. Myosin II is known to interact
with straight actin filaments during cytokinesis, and active
phospho-myosin II is mislocalized in fmn22/2 oocytes [5].
Therefore, it is reasonable to assume that myosin IIA could
be involved in the contractility required for the F-actin meshwork to position the spindle [2]. Later during migration, straight
actin filaments anchoring the closest spindle pole to the cortex
progressively accumulate and stabilize. This stabilization
could be correlated to the accumulation of RacGTP in the
cortex overhanging the chromosomes at the end of meiosis
I, especially because this accumulation has been shown to
be involved in meiotic spindle anchoring to the cortex [8]. At
anaphase, the oocyte undergoes a global reorganization of
its actin cytoskeleton, both in the cortex and in the cytoplasm
(Figure 4G).
The presence of an F-actin spindle is consistent with previous work suggesting that F-actin is present in metaphase
spindles of many models [24–26]. Recently, Woolner et al.
(2008) also showed that dynamic F-actin cables surrounding
the mitotic spindle and extending between the spindle and
the cortex are required for spindle anchoring and length
maintenance [11].
Formin 2 organizes both the cytoplasmic meshwork and
F-actin spindle. In its absence, neither of them forms, and
hence the spindle remains stationary. For future studies, it
will be interesting to determine whether Formin 2 activity
and/or localization are regulated at anaphase, allowing the observed global changes in F-actin organization and dynamics
occurring at this stage (Figure 4G).
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
Supplemental Data
17.
Supplemental Data include Supplemental Experimental Procedures and
seven movies and can be found with this article online at http://www.
current-biology.com/cgi/content/full/18/19/1514/DC1/.
Acknowledgments
We thank William M. Bement for the kind gift of the Utr-GFP probe. We thank
Christophe Klein for his help with the analysis of the FRAP data. This work
was supported by grants from the Association pour la Recherche sur le Cancer (ARC3877 and ARC4968 to M.H.V.) and from the Agence Nationale pour
la Recherche (ANRNT05-1-43120 to M.H.V.). J.A. is a recipient of a fellowship
from the Ministère délégué à la Recherche et aux Nouvelles Technologies.
K.W.L. is supported by the Agence Nationale pour la Recherche
(ANRNT05-1-43120).
18.
19.
20.
21.
22.
Received: May 15, 2008
Revised: July 23, 2008
Accepted: August 14, 2008
Published online: October 9, 2008
23.
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