|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online 23 June 2004
doi: 10.1242/dev.01244
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
ISREC (Swiss Institute for Experimental Cancer Research), 155, chemin des Boveresses, CH-1066 Epalinges/Lausanne, Switzerland
* Author for correspondence (e-mail: pierre.gonczy{at}isrec.unil.ch)
Accepted 30 April 2004
 |
SUMMARY |
|---|
 TOP SUMMARY IntroductionMaterials and methodsResultsDiscussionREFERENCES |
|---|
Key words: Meiotic cell cycle, AP polarity, E3 ligase, C. elegans, APC
|
Introduction |
|---|
|
TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
|---|
Progression through mitosis is governed by cyclin-dependent kinases (Cdks),
which drive cells into metaphase, and by the anaphase-promoting
complex/cyclosome (APC), a multi-subunit E3 ligase that poly-ubiquitinates
substrate proteins to target them for destruction by the proteasome (reviewed
by Nurse, 2002
;
Peters, 2002). APC substrates
include securins, whose destruction is essential for the metaphase to anaphase
transition, and B-type cyclins, whose destruction is essential for mitotic
exit. Although the APC is a universal regulator of mitosis, the situation
appears to be different during meiosis. In Xenopus laevis, the APC is
required for the metaphase to anaphase transition of meiosis II, but not for
that of meiosis I (Peter et al.,
2001; Taieb et al.,
2001). Conversely, in C. elegans, the APC is required at
meiosis I but seemingly not at meiosis II
(Golden et al., 2000;
Shakes et al., 2003;
Wallenfang and Seydoux, 2000).
It is likely that components other than the APC ensure progression through
meiosis I in X. laevis and meiosis II in C. elegans,
although their identity is not known.
Progression through the mitotic cell cycle also requires cullin-based E3
ligases, including Skp1-Cul1-F-box-protein complexes (SCF) and Elongin
C-Cul2-SOCS box complexes (ECS) (reviewed by
Deshaies, 1999). In
Saccharomyces cerevisiae, SCF activity is required at the G1 to S
transition to target Cdk inhibitors for degradation
(Schwob et al., 1994), and
plays a role at the G2 to M transition as well
(Goh and Surana, 1999). ECS
activity is thought to be required for the G1 to S transition, as germ cells
homozygous for a cul-2 deletion in C. elegans accumulate in
G1 (Feng et al., 1999).
Although embryos lacking cul-2 function often fail to extrude a
second polar body (Feng et al.,
1999), the exact requirement of cullin-based E3 ligases in meiotic
cell cycle progression remains to be elucidated.
Establishment of polarity along the anteroposterior (AP) embryonic axis in
C. elegans occurs shortly after exit from the meiotic cell cycle. The
two female meiotic divisions take place shortly after fertilization, which
occurs when the oocyte enters the spermatheca
(McCarter et al., 1999).
Because the nucleus is located distally in the mature oocyte, oocyte-derived
chromosomes and the meiosis I and II spindles are on the distal side of the
newly fertilized embryo, whereas sperm-derived chromosomes and centrosomes are
on the proximal side. AP polarity is initiated by a sperm component that
confers posterior character to the proximal side and becomes apparent during S
phase of the first mitotic cell cycle
(Cuenca, 2003;
Goldstein and Hird, 1996).
This results in a polarized distribution of maternally contributed PAR and MEX
proteins in the one-cell stage embryo (reviewed by
Lyczak et al., 2002;
Pellettieri and Seydoux, 2002;
Rose and Kemphues, 1998).
PAR-3 and PAR-6 localize to the anterior cortex, PAR-2 and PAR-1 localize to
the posterior cortex, whereas MEX-5 and MEX-6 are restricted to the anterior
cytoplasm. Polarity established by PAR and MEX proteins translates into
differential segregation of developmental factors. For instance, P granules
and PIE-1, which are both destined to the germ lineage, are segregated to the
posterior of the one-cell stage embryo. Polarity also translates into
asymmetric elongation of the mitotic spindle, resulting in an unequal first
division that generates a larger anterior blastomere and a smaller posterior
one.
The prevailing view is that the sperm component acting as polarity cue
corresponds to astral microtubules nucleated by paternally contributed
centrosomes. Relevant to this view are experiments conducted with conditional
mutants in APC components. At the restrictive temperature, such mutants arrest
at the metaphase to anaphase transition of meiosis I
(Golden et al., 2000;
Wallenfang and Seydoux, 2000).
Approximately 40% of these embryos establish inverted polarity, with the
posterior markers PAR-1, PAR-2 and PIE-1 enriched in the vicinity of the
persisting meiotic spindle, and the anterior marker PAR-3 enriched on the
opposite side (Wallenfang and Seydoux,
2000). The fraction of embryos with inverted polarity is halved
when microtubules are depolymerized with nocodazole, and PIE-1 is no longer
segregated when the meiotic spindle is lacking following inactivation of the
Cdc2 homologue ncc-1
(Wallenfang and Seydoux,
2000). These observations suggest that microtubules from the
persisting meiosis I spindle can act as a surrogate cue sufficient for
polarity establishment. It has also been proposed that astral microtubules may
be necessary, because polarity is not established in embryos with aberrant
sperm asters that nucleate few astral microtubules following the inactivation
of air-1, spd-2 or spd-5
(Hamill et al., 2002;
O'Connell et al., 2000;
Schumacher et al., 1998;
Wallenfang and Seydoux, 2000).
The APC itself has also been implicated in polarity establishment. Hypomorphic
APC mutants that allow meiotic cell cycle completion result in polarity
defects, which are also observed following mild inactivation of separase
(Rappleye et al., 2002).
However, the view that the APC plays a direct role in polarity establishment
is controversial, because there is a strict correlation between defects in
meiotic cell cycle progression and defects in polarity, suggesting that the
latter may be a consequence of the former
(Shakes et al., 2003).
zyg-11 is another component that appears to play a role in meiotic
cell cycle progression and AP polarity. Analysis of live embryos from the
temperature-sensitive allele zyg-11(b2), and of fixed
specimens from the non-conditional allele zyg-11(mn40),
indicate that zyg-11 is required for progression through meiosis II
(Kemphues et al., 1986). The
division of zyg-11(mn40) one-cell stage embryos can be asymmetric as
in wild type, yielding a larger anterior blastomere and a smaller posterior
one, asymmetric but inverted, or symmetric
(Kemphues et al., 1986).
Moreover, P granules are mislocalized in zyg-11(mn40)
embryos, further indicating that zyg-11 is somehow required for
proper AP polarity (Kemphues et al.,
1986). Although defects are observed at later stages of
embryogenesis, they probably reflect an earlier requirement for
zyg-11 function, as the temperature-sensitive period of
zyg-11(b2) spans the time from fertilization until the end
of meiosis II (Kemphues et al.,
1986). zyg-11 encodes a 799 amino acid protein that bears
at least two leucine-rich repeats and one Armadillo repeat
(Carter et al., 1990). There
are ZYG-11 homologues in other metazoans, including Drosophila and
human, suggesting that this family of proteins performs an evolutionarily
conserved function.
Here, we establish that embryos with impaired zyg-11 function have a delayed metaphase to anaphase transition and M phase exit at meiosis II. We find an identical phenotype after inactivating cul-2 and suggest that ZYG-11 acts with a CUL-2-based E3 ligase during meiosis. We provide evidence that the B-type cyclin CYB-3 is a target of this E3 ligase at meiosis II. We uncover that PAR proteins and P granules become localized in an inverted manner during the meiosis II delay following zyg-11 or cul-2 inactivation, and establish that these two components can regulate polarity establishment independently of their role in cell cycle progression. Unexpectedly, we find that microtubules appear dispensable for ectopic polarity in embryos lacking zyg-11 function, as well as for polarity establishment in wild-type embryos. We propose a model in which the centrosome acts as a cue to polarize the embryo along the AP axis following exit from the meiotic cell cycle.
|
Materials and methods |
|---|
|
TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
|---|
): him-1(e879) I; mnC1
dpy-10(e128)
unc-52(e444)/zyg-11(mn40)
unc-4(e120) II (Kemphues
et al., 1986);
cul-2(ek1)/unc-64(e246) III
(Feng et al., 1999), as well
as unc-32(e189) cul-2(t1644) and
unc-32(e189) cul-2(t1664), both balanced
by qC1 dpy-19(e1259ts) glp-1(q339)
(Gönczy et al., 1999);
mat-1(ax161) I; him-8(e1489) IV and
mat-1(ax144) I; him-8(e1489) IV (raised at
16°C and shifted 8-10 hours to 25°C for restrictive conditions)
(Wallenfang and Seydoux,
2000); nDf29/unc-13(e1091)
spd-2(oj29) I (O'Connell
et al., 2000). Transgenic animals expressing GFP-histone2B
(GFP-HIS), GFP-ß-tubulin (GFP-TUB), GFP-PAR-2 and GFP-ZIF-1 have been
described (DeRenzo et al.,
2003; Strome et al.,
2001; Wallenfang and Seydoux,
2000).
For generation of GFP-ZYG-11 and GFP-CYB-3 transgenic animals, PCR-derived
genomic fragments (sequences of these and other primer pairs available upon
request) were subcloned into pSU25, a modified version of the
pie-1-gfp vector containing the unc-119 cDNA (a gift from
Michael Glotzer). Sequence-verified constructs were bombarded essentially as
described (Praitis et al.,
2001). For GFP-ZYG-11, two integrated and one non-integrated lines
were retained. For GFP-CYB-3, one non-integrated line was recovered; GFP-CYB-3
was present within nuclei of oocytes and embryonic blastomeres throughout the
cell cycle, and was released in the cytoplasm at NEBD (data not shown).
zyg-11 and cul-2 mutant alleles
Most of the zyg-11 coding sequence was sequenced following PCR
reactions; a C to T alteration at position 483 of the zyg-11 gene was
found in zyg-11(mn40) in two independent PCR reactions,
predicted to truncate ZYG-11 after amino acid 161.
Two alleles (t1664 and t1644) of a maternal-effect locus
mapping close to cul-2 give rise to a phenotype similar to that of
cul-2(RNAi) (Gönczy
et al., 1999). Sequencing of RT-PCR products and parts of the
cul-2 genomic sequence following PCR reactions revealed mutations in
the donor site of the third intron (G to A alteration at position 1200 of the
gene in t1664) or the fourth intron (G to A alteration at position
1472 of the gene in t1644). Both mutations result in the use of a new
donor site (G at position 1152 for t1664, G at position 1496 for
t1644) predicted to yield a protein truncation and an internal
deletion, respectively.
RNAi
RNAi feeding strains for zyg-11, cul-2 (a gift from Lionel
Pintard), cyb-3, elc-1, ncc-1 and tba-2 (a gift from Michael
Glotzer) were generated essentially as described
(Timmons et al., 2001). The
mat-1 feeding strain was from the chromosome I feeding library
(Fraser et al., 2000). RNAi
was performed by feeding L4 larvae for 24-36 hours essentially as described
(Kamath et al., 2001).
Slightly milder conditions were used for zyg-11(RNAi) embryos expressing GFP-HIS, GFP-TUB and GFP-PAR-2 to improve recovery. Although this reduced the frequency of chromosome segregation defects, inverted polarity was nevertheless always established during the meiosis II delay under these conditions (see Fig. 7). Of 25 embryos analyzed in this manner, 6 failed to develop until the two-cell stage and were not considered for analysis.
|
The following 26 genes were inactivated using RNAi by feeding to search for components required for meiosis II. Ten were genes whose reported phenotype was potentially similar to that of zyg-11(RNAi), C47B2.4 (pbs-2), F25H2.9 (pas-5), F33D11.10, F39H11.5 (pbs-7), F48E8.5 (paa-1), K07C11.2 (air-1), T09A5.9, Y45F10A.2 (puf-3), ZK1127.5, ZK520.4 (cul-2); 10 were putative cell cycle regulators, C09G4.3 (cks-1), H31G24.4 (cyb-2.2), K06A5.7a (cdc-25.1), T05A6.1 (cki-1), T05A6.2a (cki-2), T06E6.2 (cyb-3), W02A2.6 (rec-8), Y60A3A.12 (chk-2), ZC168.4 (cyb-1), ZK1307.6 (fzr-1); and six were genes with homologies to zyg-11, C33A12.12, Y9C9A.13, Y39G10AR.5, F44E5.2, F47D12.5, T24C4.6.
Microscopy
Pressure-free and isotonic conditions
(Shelton and Bowerman, 1996)
were used because embryos shortly after fertilization are fragile. To achieve
precise timing of events, embryos were imaged essentially as described
(Brauchle et al., 2003),
collecting 1 frame every 30 seconds either in utero, starting at
fertilization, or ex utero, starting before anaphase I. Duration of events
obtained with either method was comparable. For in utero recordings, 4-10
worms were soaked for 
45 minutes in M9 buffer, supplemented with 0.01%
tetramisole and 0.1% Tricaine (Kirby et
al., 1990), and mounted under a cover-slip on a 2% agarose pad
surrounded by vaseline. For visualizing chromosomes in live embryos expressing
GFP-PAR-2 and that are osmosensitive, samples were bathed in Hoechst 33342 for
four minutes and then washed for two minutes prior to imaging.
Fixation and indirect immunofluorescence were essentially as described
(Gönczy et al., 1999).
Primary antibodies were mouse anti-tubulin (DM1A; 1:400; Sigma) and the
following antibodies raised in rabbits: anti-PAR-1 (1:2000)
(Gönczy et al., 2001),
anti-PAR-2 (1:400) (Pichler et al.,
2000), anti-PAR-3 (1:60)
(Pichler et al., 2000),
anti-PGL-1 (1:2000) (Kawasaki et al.,
1998), anti-SAS-4 (1:1600)
(Leidel and Gönczy,
2003), anti-phosphorylated histone H3 (1:1200; Upstate) and
anti-GFP (1:600, a gift from Viesturs Simanis). Slides were counterstained
with Hoechst 33258 (Sigma) to reveal DNA. Indirect immunofluorescence was
imaged on an LSM510 Zeiss confocal microscope; quantification of GFP-CYB-3,
GFP-ZYG-11 and GFP-ZIF-1 signals was performed using a 12-bit CCD Camera and
Metamorph software (Universal Imaging). Images were processed with Adobe
Photoshop.
|
Results |
|---|
|
TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
|---|
|
|
|
30 minutes, as compared with 6 minutes in wild type.
During this delay, chromosomes lose their tight arrangement on the metaphase
plate and the spindle becomes less organized
(Fig. 2I). Thereafter,
anaphase-like figures are observed, although chromosome bridges are often
apparent and sister chromatids are not segregated properly
(Fig. 2J). Moreover, anaphase
II is lengthened. Chromosomes display the M phase-specific phosphorylated
histone H3 (Hsu et al., 2000)
throughout the meiosis II delay in zyg-11(RNAi) embryos,
including when anaphase-like configurations are apparent, indicating that it
corresponds to a prolonged M phase (Fig.
2L-N). Eventually, oocyte and sperm-derived chromosomes
decondense, pronuclei form and the first mitotic cell cycle ensues. We
conclude that zyg-11 is required for timely metaphase to anaphase
transition and M phase exit at meiosis II.
cyb-3 and cul-2 are also required for meiosis II progression
As the sequence of ZYG-11 does not offer obvious clues as to its
biochemical activity, we sought to identify other genes whose inactivation
results in delayed progression through meiosis II to gain insight into the
mechanism of action of ZYG-11. We examined the results of RNAi-based
functional genomic screens available as DIC movies through
www.wormbase.org.
Out of 5500 genes considered, we found 10 whose reported phenotype
resembles that of zyg-11(RNAi). We also selected 10 putative
cell cycle regulators and six ORFs that bear homology with zyg-11 for
further analysis (see list of genes in Materials and methods). We inactivated
these 26 genes using RNAi and analyzed meiotic cell cycle progression using
time-lapse DIC microscopy. We found two genes whose inactivation does not
affect meiosis I but results in delayed progression through metaphase of
meiosis II: the B type cyclin cyb-3 and the cullin
cul-2.
In cyb-3(RNAi) embryos
(Fig. 1,
Table 1; Table S1 and Movie 4
at
http://dev.biologists.org/supplemental/),
the delay is already apparent prior to metaphase, earlier than in the absence
of zyg-11 function. Another difference is that anaphase II is not
delayed in cyb-3(RNAi) embryos. By contrast, we found that
cul-2(RNAi) embryos (Fig.
1, Table 1; Table
S1 and Movie 5 at
http://dev.biologists.org/supplemental/)
have a meiotic phenotype indistinguishable from that of
zyg-11(RNAi) or zyg-11(mn40) embryos. In
particular, the metaphase to anaphase transition of meiosis II is
significantly delayed, with embryos exhibiting metaphase-like figures for
33 minutes. Moreover, anaphase II is lengthened and often aberrant. We
found a similar phenotype in the deletion allele cul-2(ek1),
and in two maternal-effect embryonic lethal alleles of cul-2
(Table 1; see Materials and
methods). Importantly, inactivation of cul-2 by RNAi in
zyg-11(mn40) mutant embryos does not result in a more
prolonged meiosis II delay (Fig.
1, Table 1; see
Movie 6 at
http://dev.biologists.org/supplemental/),
compatible with the notion that zyg-11 and cul-2 act in a
common process. A phenotype similar to that observed by inactivating
zyg-11 or cul-2 has also been obtained by inactivating
Elongin C (elc-1) and Rbx (rbx-1), two other core components
of ECS E3 ligases (Liu et al.,
2004). Taken together, these results establish that a
CUL-2-containing E3 ligase is essential for timely metaphase to anaphase
transition and M phase exit at meiosis II, and suggest that ZYG-11 somehow
acts in concert with this ECS.
CYB-3 accumulates following zyg-11 inactivation, delaying M phase exit
Given that cyb-3 acts at meiosis II, and that degradation of B
type cyclins is generally required for M phase exit, we investigated whether
persistence of CYB-3 may be causing delayed M phase exit when the CUL-2-based
E3 ligase is inactivated. We generated a fusion protein between GFP and CYB-3;
in wild type, we found levels of GFP-CYB-3 to be high until metaphase I, and
significantly lower after the metaphase to anaphase transition of meiosis I
(data not shown). Similar low levels were observed during meiosis II
(Fig. 2O), although these low
levels precluded assessing whether levels drop further after the metaphase to
anaphase transition of meiosis II. Importantly, we found that GFP-CYB-3 levels
during the meoisis II delay in zyg-11(RNAi) and
cul-2(RNAi) embryos were high compared with wild-type
meiosis II embryos (Fig. 2P,Q).
To test whether CYB-3 accumulation was causing delayed M phase exit in the
absence of zyg-11 function, we examined
zyg-11(mn40) cyb-3(RNAi) embryos.
Strikingly, whereas the metaphase to anaphase transition of meiosis II is
still delayed in such embryos, the anaphase II delay is abolished
(Fig. 1,
Table 1; see Movie 7 at
http://dev.biologists.org/supplemental/).
Therefore, the persistence of CYB-3 appears to be responsible for delayed M
phase exit following zyg-11 inactivation.
ZYG-11 may be a substrate recruitment subunit of a meiosis-specific CUL-2-containing ECS
ECS E3 ligases contain a SOCS-box-containing protein that bridges Elongin C
and the substrate (Kile et al.,
2002). We considered whether ZYG-11 could be such a substrate
recruitment subunit, even though it does not contain an obvious SOCS-box.
Substrate recruitment subunits of the ECS and other cullin-based E3 ligases
are unstable and are targeted for degradation by the very same complex of
which they are a part (DeRenzo et al.,
2003; Galan and Peter,
1999; Geyer et al.,
2003; Pintard et al.,
2003b; Wirbelauer et al.,
2000; Zhou and Howley,
1998). To test whether ZYG-11 behaves in this manner, we generated
a transgenic line expressing GFP-ZYG-11 and compared GFP levels in wild-type
and cul-2(RNAi) animals. Although wild-type animals express
the fusion protein at very low levels in the germ-line and early embryos
(Fig. 3A,D), GFP-ZYG-11 levels
are dramatically increased when cul-2 is inactivated
(Fig. 3B,E,G). As anticipated,
analogous results were obtained after elc-1 inactivation
(Fig. 3G). Elevated levels of
GFP-ZYG-11 do not result merely from slower progression through meiosis II,
because GFP-ZYG-11 is not increased in cyb-3(RNAi) animals
(Fig. 3C,F,G). These
observations are compatible with ZYG-11 being a substrate recruitment subunit
of a CUL-2 containing ECS ligase.
|
). In
cul-2(RNAi) animals, levels of GFP-ZIF-1 are increased
compared with otherwise wild-type embryos
(Fig. 3H,J,K,M,N)
(DeRenzo et al., 2003). By
contrast, we found that levels of GFP-ZIF-1 are not altered in
zyg-11(RNAi) animals
(Fig. 3I,L,N). We conclude that
zyg-11 is not required for the function of all CUL-2-containing E3
ligases but plays a more restricted role during meiosis.
zyg-11 and cul-2 function redundantly with APC during meiosis I
In addition to an essential requirement at meiosis II, we uncovered a
non-essential function for zyg-11 and cul-2 at meiosis I.
The mat-1 locus encodes the APC component Cdc27/APC3 and embryos from
the conditional allele mat-1(ax161) raised at 25°C
arrest at the metaphase to anaphase transition of meiosis I
(Fig. 4A-C,
Table 2) (Golden et al., 2000;
Wallenfang and Seydoux, 2000).
Such arrested embryos fall into three categories on the basis of their
position with respect to the spermatheca in the animal (data not shown), and
the appearance of chromosomes and microtubules
(Fig. 4). First, embryos
closest to the spermatheca and thus the youngest, which have a canonical
metaphase I arrest configuration (Fig.
4A, metaphase I). Second, embryos located further away from the
spermatheca, which have chromosomes with a looser arrangement and a less
organized spindle (Fig. 4B,
metaphase I late). Third, embryos located further still from the spermatheca
and thus the oldest, which have elongated chromosomes located in the embryo
center and a completely disassembled spindle
(Fig. 4C, metaphase I very
late).
|
|
;
Wallenfang and Seydoux, 2000),
we found that they exhibit meiosis I defects when, in addition, either
zyg-11 or cul-2 are inactivated, accumulating with
chromosomes and spindle configurations resembling those of 25°C
mat-1(ax161) embryos
(Fig. 4H-J, compare with
Fig. 4A-C; Table 2). Although some
15°C mat-1(ax161) zyg-11(RNAi) and
15°C mat-1(ax161) cul-2(RNAi) embryos
bypass the meiosis I block, the first polar body is small or absent (data not
shown) and metaphase II figures invariably exhibit supernumerary chromosomes,
which is indicative of defective anaphase I
(Fig. 4M,N). Similar results
were obtained using a distinct conditional mat-1 allele,
mat-1(ax144) (Table
2). These findings indicate that zyg-11 and
cul-2 also act at meiosis I, although this role is revealed only when
APC function is slightly compromised. Because zyg-11 behaves like
cul-2 in enhancing the mat-1 phenotype, these findings also
reinforce the notion that ZYG-11 and CUL-2 function in a common process. Interestingly, we noted that a sizeable fraction of 15°C mat-1(ax161) zyg-11(RNAi) and 15°C mat-1(ax161) cul-2(RNAi) metaphase II embryos have elongated chromosomes located in the center of the embryo and a completely disassembled spindle (Fig. 4O, Table 2), resembling the last category of 25°C mat-1(ax161) embryos (see Fig. 4C). Such configurations are never observed when zyg-11 and cul-2 are inactivated on their own (Fig. 2). Therefore, it appears that some embryos lacking zyg-11 or cul-2 function cannot exit meiosis II if APC function is compromised. Although this may be a consequence of the meiosis I defects, it could also reflect a requirement for APC at meiosis II.
Inverted polarity is established during the meiosis II delay in the absence of zyg-11 or cul-2 function
Next, we investigated the cause of the alterations in cleavage pattern and
P granule distribution that have been reported in
zyg-11(mn40) embryos
(Kemphues et al., 1986) by
analyzing the distribution of polarity markers. In wild-type meiosis II,
PAR-1, PAR-2 and PAR-3 proteins are not polarized, and P-granules are present
throughout the cytoplasm (Fig.
5A-D, Table 3). By
contrast, we found that 50% of zyg-11(RNAi) meiosis II embryos
exhibit polarized distribution of PAR proteins and P-granules
(Fig. 5E-H,
Table 3), although enrichment
of all four markers is typically less pronounced than for wild-type embryos
during the first mitotic cell cycle (Fig.
5Q-T). Strikingly, in all zyg-11(RNAi) meiosis II embryos
with polarized distribution, PAR-1, PAR-2 and P-granules are enriched in the
vicinity of the meiotic spindle, whereas PAR-3 is enriched on the opposite
side. These distributions are inverted compared with wild-type embryos during
the first mitotic cell cycle. We next examined cul-2(RNAi)
meiosis II embryos and found similar alterations in the distribution of PAR-1,
PAR-2, PAR-3 and P granules (Fig.
5I-L, Table 3). We
conclude that inverted polarity is established during meiosis II in the
absence of zyg-11 or cul-2 function.
|
|
Because the meiosis II delay in cyb-3(RNAi) embryos is
slightly less pronounced and of a different nature than that observed in
zyg-11(RNAi) or cul-2(RNAi) embryos, we
conducted two experiments to test whether zyg-11 and cul-2
can regulate polarity establishment independently of their requirement for
cell cycle progression during meiosis II. First, we made use of 25°C
mat-1(ax161) embryos, which are arrested in meiosis I with
evenly distributed P granules (Fig.
6A,B, Table 4)
(Wallenfang and Seydoux,
2000). Because P granules are polarized in an inverted manner in
50% of embryos lacking zyg-11 or cul-2 function
(Table 3), we performed a
molecular epistasis experiment using 25°C mat-1(ax161)
embryos subjected to zyg-11(RNAi) or
cul-2(RNAi). Interestingly, we found that 30% of
25°C mat-1(ax161) embryos lacking, in addition, either
zyg-11 or cul-2 function, exhibit an enrichment of P
granules in the vicinity of the meiotic spindle
(Fig. 6D,
Table 4). Because the
CUL-2-based E3 ligase plays a non-essential role at meiosis I, we considered
whether this may reflect a more robust cell cycle arrest in such embryos.
However, we found that the fraction of embryos in each of the three meiosis I
arrest categories (Fig. 4) is
the same in 25°C mat-1(ax161) embryos irrespective of
whether they have been subjected to zyg-11(RNAi) or
cul-2(RNAi) (Table
2), indicating that cell cycle progression is similarly
affected.
|
|
). Nevertheless, the time interval between fertilization and
the appearance of pronuclei is unchanged as the meiotic divisions are bypassed
(Boxem et al., 1999). We found
the average number of embryos in meiosis, as judged by examining chromosomes
with GFP-HIS, to be 1.5/gonad for ncc-1(RNAi) (n=17
gonads), 3.0/gonad for zyg-11(mn40) (n=23 gonads)
and 1.6/gonad for ncc-1(RNAi) zyg-11(mn40)
(n=25 gonads). This indicates that the time separating fertilization
from pronuclear appearance is not changed in zyg-11(mn40)
ncc-1(RNAi) embryos compared with
ncc-1(RNAi) embryos, enabling us to test the requirement of
zyg-11 in polarity establishment independently of that in meiotic
cell cycle progression. As shown in Fig.
6F, we observed cortical enrichment of GFP-PAR-2 in the vicinity
of oocyte chromosomes in 60% of zyg-11(mn40)
ncc-1(RNAi) embryos prior to pronuclear appearance
(n=29), although to lower levels than in
zyg-11(mn40) embryos (data not shown). Taken together, our
findings indicate that zyg-11 and cul-2 can regulate
polarity establishment independently of their requirement for meiosis II cell
cycle progression.
Timing of inverted polarity establishment
We noted that only 25°C mat-1(ax161)
zyg-11(RNAi) or 25°C mat-1(ax161)
cul-2(RNAi) embryos of the `metaphase I late' and `metaphase
I very late' categories show polarized distribution of P granules
(Fig. 6D;
Table 4). Compatible with this
view, inverted distribution of GFP-PAR-2 in mat-1(RNAi) or
mat-1(RNAi) zyg-11(mn40) embryos is found
predominantly in embryos of these two categories
(Fig. 6H,J;
Table 4). Therefore, inverted
polarity is established some time after the initial metaphase I arrest in
embryos lacking both mat-1 and zyg-11 function.
By analogy, we considered whether inverted polarity might be established
some time after the initial metaphase II block in embryos lacking
zyg-11 or cul-2 function. We imaged GFPPAR-2 in
zyg-11(RNAi) and zyg-11(mn40) embryos, and
found that cortical enrichment in the vicinity of the meiotic spindle always
becomes apparent during the second half of the delay
(Fig. 1B, vertical white lines;
n=7). This is likely to explain why polarity inversion is observed in
50% of fixed zyg-11(RNAi) or
cul-2(RNAi) meiosis II embryos, and indicates that inverted
polarity is invariably established by the end of the meiotic cell cycle in the
absence of zyg-11 or cul-2 function.
Plasticity of polarity after the meiotic cell cycle in zyg-11(RNAi) embryos
We next investigated how polarity inversion by the end of meiosis II
generates the diverse cleavage patterns that occur in the absence of
zyg-11 function (Kemphues et al.,
1986). By live imaging of zyg-11(RNAi) embryos
expressing GFP-HIS and GFP-PAR-2, we found that a second GFP-PAR-2 domain,
which is variable in size, is always established on the opposite side of the
embryo at the onset of the first mitotic cell cycle, as in wild type (see
Movies 8-10 at
http://dev.biologists.org/supplemental/;
n=19). Therefore, zyg-11(RNAi) embryos experience
two polarizing signals, one during meiosis II and a second one at the onset of
the first mitotic cell cycle.
We found that these embryos can be placed into three classes according to the evolution of the two GFP-PAR-2 domains. In six embryos, the first GFP-PAR-2 domain diminishes in intensity, whereas the second one becomes more robust; embryos in this first class divide asymmetrically as in wild type, yielding a smaller blastomere on the side of the prevailing GFP-PAR-2 domain (Fig. 7A,D; see Movie 8 at http://dev.biologists.org/supplemental/). In 10 embryos, the first GFP-PAR-2 domain remains robust, whereas the second one diminishes in intensity; embryos in this second class divide asymmetrically, but in an inverted manner compared with wild type (Fig. 7B,D; Movie 9 at http://dev.biologists.org/supplemental/). In the remaining three embryos, both domains of GFP-PAR-2 become undetectable by the end of the first mitotic cell cycle; embryos in this third class divide symmetrically (Fig. 7C,D; Movie 10 at http://dev.biologists.org/supplemental/). These findings suggest that one PAR-2-containing domain at most can be present by the end of the first mitotic cell cycle and that this domain is predictive of spindle positioning during the first cleavage division.
We used GFP-HIS to follow sperm-derived chromosomes in the three classes of zyg-11(RNAi) embryos (n=14). In the first class, where the second GFP-PAR-2 domain prevails, sperm chromosomes remain in the position they occupy in wild type, opposite the first GFP-PAR-2 domain (Movie 8 at http://dev.biologists.org/supplemental/; 6/6 embryos). In most embryos of the second class, where the first GFP-PAR-2 domain prevails, sperm chromosomes are initially in their normal position, but then move towards the inverted GFP-PAR-2 domain (Movie 9 at http://dev.biologists.org/supplemental/; 5/6 embryos). In the third class, where both GFP-PAR-2 domains become undetectable, sperm chromosomes are also initially in their normal position, but then move towards the cell center (Movie 10 at http://dev.biologists.org/supplemental/; 2/2 embryos). These observations suggest that the cleavage pattern of zyg-11(RNAi) embryos is dictated primarily by the position of a sperm-derived component during the course of the first mitotic cell cycle.
To test this hypothesis further, we analyzed polarity in embryos lacking
both zyg-11 and spd-2, which is required for establishing
polarity induced by the sperm-derived component
(O'Connell et al., 2000). We
found that spd-2 is dispensable for establishing inverted polarity in
the absence of zyg-11 at meiosis II, as PAR-1 is still inverted in
50% of fixed spd-2(oj29) zyg-11(RNAi)
meiosis II embryos (Table 3).
In addition, the fraction of embryos with an inverted first division is
increased from 36% in zyg-11(RNAi) embryos to 60% in zyg-11(RNAi)
spd-2(oj29) (Table 3).
These findings indicate that a sperm-derived component that is in part
spd-2 dependent competes with the inverted polarity established
during the meiosis II delay to result in a single GFP-PAR-2 domain by the end
of the first mitotic cell cycle.
Microtubules and polarity establishment during meiosis II and the first mitotic cell cycle
We next investigated whether establishment of inverted polarity during the
meiotic II delay is microtubule dependent. We eliminated microtubules from
zyg-11(mn40) mutant embryos using RNAi against the alpha
tubulin gene tba-2, thus likely inactivating all alpha tubulin genes
owing to cross-RNAi between highly related sequences
(Wright and Hunter, 2003). In
such embryos, all microtubule processes, including the two meiotic divisions,
are defective, and all oocyte chromosomes are found approximately two-thirds
of the way down the length of the egg
(Yang et al., 2003)
(Fig. 8B and data not shown).
Importantly, we found that 65% of zyg-11(mn40)
tba-2(RNAi) meiosis II embryos have a patch of cortical
GFP-PAR-2 (Fig. 8B; n=18), which is comparable to the fraction of
zyg-11(RNAi) meiosis II embryos exhibiting inverted PAR-2
cortical localization (Table
3). Similar findings were made examining PAR-1 distribution (data
not shown). Moreover, P granules tend to be enriched in the vicinity of
cortical GFPPAR-2 (Fig. 8B).
Interestingly, we noted also that the patch of GFP-PAR-2 is located invariably
on the cortex closest to oocyte chromosomes
(Fig. 8C), raising the
possibility that they may somehow influence positioning of cortical GFP-PAR-2.
Together, these observations indicate that microtubules are not essential for
imparting ectopic polarity in the absence of zyg-11 function.
|
|
Discussion |
|---|
|
TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
|---|
),
reveals that zyg-11 is likely to act with a CUL-2-containing E3
ligase at meiosis. Could ZYG-11 be a substrate of this ECS, given that
GFP-ZYG-11 levels are dramatically increased in the absence of cul-2?
This possibility seems unlikely, given that inactivation of the ECS and of its
substrate are expected to yield opposite, not identical, phenotypes. Although
other scenarios can be envisaged, a plausible hypothesis is that ZYG-11 is the
substrate recruitment component of this ECS. Although this seems at odds with
the fact that ZYG-11 does not contain a recognizable SOCS box, this motif can
be very divergent, as is the case for ZIF-1
(DeRenzo et al., 2003), and
Drosophila Zyg11 contains a potential SOCS box (R.S. and P.G.,
unpublished). Whether C. elegans ZYG-11 truly acts as a substrate
recruitment subunit remains to be ascertained biochemically.
Regulated protein degradation by way of E3 ligases plays a crucial role in
other aspects of early C. elegans development. Another ECS that uses
ZIF-1 as a substrate recruitment subunit is essential for removing several
CCCH finger proteins from somatic lineages
(DeRenzo et al., 2003).
Moreover, a CUL-3-based complex is essential for degradation of the
meiosis-specific microtubule-severing protein MEI-1 during the first mitotic
cell cycle (Kurz et al., 2002;
Pintard et al., 2003a). The
activity of these two E3 ligases is dependent on the DYRK kinase MBK-2
(Pellettieri et al., 2003).
This does not seem to be the case for the ECS described in this work, as
embryos lacking mbk-2 function complete the meiotic divisions
normally and have no apparent polarity defects
(Pellettieri et al.,
2003).
The APC and an ECS together ensure progression through the two meiotic divisions
Previous work established that the APC is essential for the metaphase to
anaphase transition of meiosis I in C. elegans
(Golden et al., 2000;
Wallenfang and Seydoux, 2000).
Hypomorphic APC mutants that affect meiosis I without arresting cell cycle
progression exhibit defective sister chromatid segregation at meiosis II
(Shakes et al., 2003).
However, this may result from the aberrant meiosis I, as no semi-permissive
conditions were found that yield a meiosis II arrest, raising the possibility
that APC is not required at meiosis II
(Shakes et al., 2003).
Compatible with this view, the APC component FZY-1 localizes to chromosomes
during meiosis I but not meiosis II
(Kitagawa et al., 2002). Here,
we establish that a CUL-2-based E3 ligase is required for progression through
meiosis II. Though unlikely given the distinct phenotypes of APC hypomorphic
mutants and of zyg-11 or cul-2 inactivation, the possibility
that this ECS is an essential positive regulator of the APC cannot be excluded
for meiosis II. By contrast, this cannot be the case at meiosis I, because
inactivation of zyg-11 or cul-2 does not affect meiosis I,
whereas that of the APC results in metaphase I arrest. Although embryos
lacking zyg-11 or cul-2 do not arrest at meiosis I,
compromising APC activity slightly in these embryos results in meiosis I
defects, indicating that the ECS plays a non-essential role at meiosis I.
Interestingly, progression through the meiotic cell cycle in other
organisms also rests on distinct E3 ligases at meiosis I and meiosis II. In
Saccharomyces pombe, the Fizzy/Cdc20-related APC activator mfr1 is
required specifically at meiosis II for degradation of the B-type cyclin cdc13
(Blanco et al., 2001).
Similarly, in Drosophila, the Cdc20/Fizzy protein Cortex is required
for timely metaphase to anaphase transition at meiosis II
(Chu et al., 2001;
Page and Orr-Weaver, 1996).
Thus, whereas meiosis II in S. pombe and Drosophila relies
on specific APC variants, it requires the activity of an ECS in C.
elegans.
What substrates of the CUL-2-based E3 ligase must be targeted for
degradation to ensure progression through meiosis II in C. elegans?
The metaphase to anaphase transition is generally triggered when securin is
targeted for degradation. IFY-1 is a C. elegans destruction-box
protein that has properties expected from a securin
(Kitagawa et al., 2002), and
it will be interesting to test whether the CUL-2-based E3 ligase mediates
IFY-1 degradation at meiosis II. Furthermore, M phase exit is generally
triggered when B type cyclins are targeted for degradation. Our findings that
cyb-3 inactivation prevents the anaphase delay of embryos lacking
zyg-11 function strongly suggests that CYB-3 is a target of the
CUL-2-based E3 ligase at meiosis II.
zyg-11 and cul-2 prevent polarity establishment
In wild type, the first signs of polarized GFP-PAR-2 distribution occur
shortly after meiosis II. Whether polarity is established when a set time is
reached (e.g. after fertilization), or when a given cell cycle stage is
encountered, has not been addressed prior to this work. We found that
asymmetric distribution of polarity markers in cyb-3(RNAi)
embryos is not established during the meiosis II delay, but instead during the
first mitotic cell cycle (R.S. and P.G., unpublished). Therefore, polarity
establishment does not occur at a fixed time after fertilization or meiosis I,
but is coupled instead to exit from the meiotic cell cycle and the onset of
the first mitotic cell cycle.
In contrast to the situation in cyb-3(RNAi) embryos,
inverted polarity is established during the meiosis II delay in embryos
lacking zyg-11 or cul-2 function. We show that
zyg-11 and cul-2 can regulate polarity establishment
independently of promoting progression through meiosis II, suggesting that
there are distinct polarity substrates that must be degraded by the
CUL-2-based E3 ligase to prevent polarity establishment during meiosis II. As
inverted polarity is also observed in metaphase I-arrested embryos lacking APC
function (Wallenfang and Seydoux,
2000), it is tempting to speculate that at least some of these
polarity substrates are shared between the two E3 ligases.
Interestingly, cortex mutant embryos in Drosophila have
impaired polyadenylation of bicoid and Toll mRNAs, resulting
in inefficient translation and ensuing defective embryonic polarity
(Lieberfarb et al., 1996).
Although the mechanisms by which APC promotes polyadenylation of these mRNAs
remains to be clarified, it is remarkable that E3 ligases that ensure
progression through meiosis II also serve to promote correct establishment of
embryonic axes in diverse metazoan organisms.
Mechanisms of polarity establishment in C. elegans
It has been proposed that astral microtubules nucleated by the sperm aster
are essential for establishing AP polarity in C. elegans embryos
(O'Connell et al., 2000;
Wallenfang and Seydoux, 2000).
Our findings challenge this view. We establish that PAR-1, GFP-PAR-2,
GFP-PAR-6, P granules and GFPPIE-1 (R.S. and P.G., unpublished) all localize
correctly in the first mitotic cell cycle in tba-2(RNAi)
embryos. These results raise the possibility that microtubules are dispensable
for AP polarity.
We recognize that we cannot exclude that minute microtubules undetectable
by immunofluorescence or damaged during fixation may play a role.
Nevertheless, we note that embryos lacking spd-2, air-1 or
spd-5 function, while exhibiting delayed microtubule nucleation
compared with wild type, have a more extensive microtubule network than
tba-2(RNAi) embryos do
(Hamill et al., 2002;
O'Connell et al., 2000;
Schumacher et al., 1998;
Wallenfang and Seydoux, 2000).
How can these apparently discrepant observations be reconciled? In wild type,
the onset of GFP-PAR-2 accumulation at the cortex coincides with that of
GFP-TUB on asters, early in the cell cycle
(Cuenca et al., 2003). It may
be that, at that early stage, minute microtubules are present in
tba-2(RNAi) embryos, but not in embryos lacking spd-2,
air-1 or spd-5 function. If this were the case, our findings
would merely demonstrate that microtubules are dispensable for the expansion
phase of the posterior PAR-2 cortical domain and not for the preceding
initiation phase (Cuenca et al.,
2003). However, we favour an alternative explanation in which
microtubules are dispensable throughout the process, and in which spd-2,
air-1 and spd-5 have a requirement for polarity establishment
that is independent from their role in microtubule nucleation.
As sperm chromosomes are not essential for AP polarity
(Sadler and Shakes, 2000), our
findings lead us to suggest that, rather than astral microtubules, the sperm
component acting as the polarity cue is the centrosome. Compatible with
centrosomes being key, spd-2, spd-5 and air-1 are all
required for centrosome maturation (Hamill
et al., 2002; Hannak et al.,
2001; O'Connell et al.,
2000), and the corresponding proteins localize to the centrosome
(Hamill et al., 2002;
Schumacher et al., 1998;
Hannak et al., 2001;
Kemp et al., 2004). It will be
interesting to investigate whether such a hypothetical polarity cue resides in
centrioles or the surrounding pericentriolar material. Equally interesting
will be to elucidate how a component somehow correlated with the position of
oocyte chromosomes, which do not have associated centrosomes, can act as a
surrogate polarity cue during meiosis II.
Our findings taken together suggest a working model (Fig. 9) in which a CUL-2-based E3 ligase targets polarity substrates for degradation during meiosis II; as a result, polarity cannot be established. After exit from the meiotic cell cycle, polarity substrates accumulate, perhaps due to diminished E3 ligase activity, and thus the centrosome can induce polarity. We postulate that in the absence of zyg-11 or cul-2, substrate accumulation occurs during the meiotic cell cycle, resulting in polarity establishment in response to a surrogate polarity cue.
|
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
|
REFERENCES |
|---|
|
TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
|---|
Blanco, M. A., Pelloquin, L. and Moreno, S. (2001).
-tubulin (A-C,G-J), or 
