Journal of Cell Science 103, 389-396 (1992)
Printed in Great Britain © The Company of Biologists Limited 1992
389
Cell cycle progression of parthenogenetically activated mouse oocytes to
interphase is dependent on the level of internal calcium
C.VINCENT1, T.R. CHEEK2 and M.H. JOHNSON 1,*
1Department
2Department
*Author
of Anatomy, University of Cambridge, Downing Street,Cambridge CB2 3DY, England
of Zoology, University of Cambridge, Downing Street, Cambridge CB2 3EJ, England
for correspondence
Summary
Nuclear maturation of the mouse oocyte becomes
arrested in metaphase of the second meiotic division
(MII). Fertilization or parthenogenetic activation
induces meiotic completion, chromosomal decondensation and formation of a pronucleus. This completion of
meiosis is probably triggered by a transient increase in
cytosolic calcium ions.
When activated just after ovulation by a low concentration of the calcium ionophore A23187, the majority
of the mouse oocytes go through a metaphase to
anaphase transition and extrude their second polar body
but they do not proceed into interphase; instead their
chromatids remain condensed and a microtubular
metaphase spindle reforms (metaphase III). However, a
Introduction
Immature mouse oocytes are arrested at prophase of the
first meiotic division until just prior to ovulation. The
resumption of meiosis is characterized by germinal vesicle
breakdown (GVBD), the condensation of chromosomes into
distinct bivalents and subsequently the separation of homologous chromosomes and the emission of the first polar body
(Donahue, 1968). The oocytes then enter a second meiosis
but arrest their progression at metaphase II (referred to as
MII). During normal maturation, chromosome decondensation does not occur between metaphase I (MI) and MII.
Activation of an oocyte arrested in MII triggers the completion of meiosis, i.e. the extrusion of the second polar
body, the decondensation of the chromosomes in the cytoplasm and the formation of a pronucleus. This progression
of the cell cycle from MII to interphase can be induced by
the spermatozoon at fertilization or by parthenogenetic activation. Fertilization is accompanied by a series of transient
rises in the intracellular calcium concentration (Cuthbertson and Cobbold, 1985; Swann, 1990; Cheek et al. 1992),
and parthenogenetic activation can result from the release
of intracellular stores of calcium induced by calcium
ionophore (Steinhardt et al. 1974; Kline and Kline, 1992).
After exposure to parthenogenetic activating agents, the
high percentage of these oocytes will undergo a true
parthenogenetic activation assessed by the formation of
a pronucleus, when exposed to a higher concentration
of the calcium ionophore. The capacity of the mouse
oocyte to pass into metaphase III is lost with increasing
time post-ovulation. Direct measurement of intracellular calcium with Fura-2 reveals higher levels of cytosolic calcium in aged oocytes and/or using higher concentrations of calcium ionophore for activation. It is
concluded that the internal free calcium level determines the transition to interphase.
Key words: oocyte, parthenogenetic activation, cell cycle,
calcium.
proportion of activated oocytes, assessed by the formation
of a pronucleus, increases with the post-ovulatory age of
the oocytes (Kaufman, 1983). Indeed, the response of
oocytes activated immediately after ovulation can be defective: young mouse or rat oocytes extrude the second
polar body but do not progress into interphase, arresting
again in a new metaphase, called metaphase III (MIII;
Keefer and Schuetz, 1982; Kubiak, 1989; Zernicka-Goetz,
1991).
We have now used the calcium ionophore A23187 as a
parthenogenetic activator and show that MIII formation
depends not only on the age of the oocyte but also on the
concentration of calcium ionophore used for activation: the
majority of young oocytes which fail to undergo a full activation can nevertheless be forced to go into interphase by
increasing the concentration of ionophore.
By monitoring the intracellular calcium signal by the
video-imaging of Fura-2 (Moreton, 1991), it is possible to
obtain precise measurements of the calcium transient inside
the oocytes for the two different ionophore concentrations
used. Our results suggest that the cell cycle progression of
oocytes activated early after ovulation depends on the level
of internal calcium released during activation and that the
capacity of the oocyte to release cytosolic calcium in
response to ionophore increases with age.
390
C. Vincent and others
Materials and methods
Collection and treatment of oocytes
Oocytes were collected from superovulated females 12.5 h after
hCG injection and denuded of cumulus cells as described previously (Vincent et al. 1990). All the incubations were performed
in medium H6 containing bovine serum albumin (BSA, 4 mg
ml-1; Nasr-Esfahani et al. 1990) in cavity blocks pre-warmed to
above 37°C. Removal of the zona pellucida was accomplished by
a 10 min exposure to 0.001% chymotrypsin (Sigma Type II)
(Boldt and Wolf, 1986), followed by two extensive washes.
After a 5 to 15 min incubation in H6 + BSA free of calcium,
oocytes were exposed to the calcium ionophore A23187 diluted
in calcium-free medium at different concentrations according to
schedules described in Results. As DMSO (dimethyl sulphoxide)
was used as a solvent in the ionophore stock solution, an equivalent dilution of the highest concentration (0.1 or 0.25% DMSO)
was used in control groups. After each treatment, oocytes were
washed 5 times over a 30 min period. The first three washes were
performed in calcium-free medium.
Staining of microtubules and chromosomes
Oocytes were carried through processing in specially designed
chambers as described by Maro et al. (1984). Cells were fixed at
37°C for 30 min in 3.7% formaldehyde in PBS in the presence
of 0.5% Triton X-100 (Sigma) for extraction and were washed in
phosphate-buffered saline (PBS). Tubulin was visualized with a
rat monoclonal anti-tubulin antibody (Kilmartin et al. 1982) followed by rhodamine-labelled anti-rat IgG. Chromosomes were
stained by incubation in Hoechst dye 33342 (10 µg ml -1 in PBS)
for 30 min.
Air-dried chromosome spreads were prepared by the procedure
of Tarkowski (1966) and stained with Giemsa for 20 min.
Cortical granule staining
The procedure used was derived from that used to stain hamster
oocyte cortical granules (Cherr et al. 1988). Oocytes were fixed
in 3% paraformaldehyde in PBS for 30 min and then washed
extensively in a blocking solution of 1 mg ml-1 BSA, 100 mM
glycine and 0.2% sodium azide in PBS. To visualize exclusively
the content of the cortical granules after extrusion, oocytes were
not permeabilized. Oocytes were incubated in 10 µg ml -1 Lens
culnaris agglutinin conjugated to fluorescein isothiocyanate
(FITC-LCA; United States Biochemical Corporation) in blocking
solution for 15 min and then washed extensively in the blocking
buffer.
Loading oocytes with Fura-2
Oocytes were collected from superovulated females and denuded
of cumulus cells and zona pellucida as described earlier. Oocytes
were washed and transferred to H6+ polyvinylpyrrolidone (PVP;
6 mg ml-1) on a coverslip which had been precoated with concanavalin A (ConA; 0.2 mg/ml in PBS) and which formed the
base of a metallic perfusion chamber (Moreton, 1991). Oocytes
were then loaded with Fura-2 (2 µM; Molecular Probes) for 20
to 30 min and washed extensively with H6 + PVP. The chamber
was then placed in a well on the stage of a Nikon Diaphot TMD
inverted epifluorescence microscope for imaging. Incubations
were all carried out via a system of continuous perfusion through
the perfusion chamber maintained at 37°C. After a 5 min incubation in H6 + PVP free of added calcium, oocytes were exposed
to the calcium ionophore A23187 diluted in the same buffer. The
concentration of the calcium ionophore used and the time of exposure are described in Results. After ionophore treatment, oocytes
were washed continuously in calcium-free medium.
Fura-2 imaging
Intracellular free calcium activity [Ca2+]i was imaged through a
Nikon CF-Fluor 20× objective and intensified CCD camera
(Extended ISIS, Photonic Science, Robertsbridge, UK), by calculating the ratio of Fura-2 fluorescence at 510 nm, excited by UV
light alternately at 340 and 380 nm from twin xenon arc lamps
and grating monochromators. Excitation wavelengths were alternated by a rotating chopper mirror attached to a stepper-motor,
which was driven in synchrony with the video signal from the
camera, to switch wavelengths at the end of each video frame.
The resulting video signals were combined by an “Imagine” digital image processor (Synoptics Ltd., Cambridge, UK) using a
lookup table to implement the formula of Grynkiewicz et al.
(1985). The calculation was done in real time, to give a “live”
image of [Ca2+]i, which was updated every 80 ms, and smoothed
by recursive filtering with a 200 ms time-constant to reduce the
noise (for further details, see Moreton, 1991; O’Sullivan et al.
1989). The live image was recorded continuously on video tape,
and subsequently played back and re-digitized into a frame-store,
using software written in the semper language (Synoptics Ltd.) to
sample selected oocytes and to record and plot mean [Ca2+]i readings at regulat time intervals. In all cases data were sampled at 4
s intervals.
Results
Parthenogenetic activation of mouse oocytes after
treatment with different concentrations of calcium
ionophore
Freshly ovulated zona-free oocytes (recovered 12.5 h after
the hCG injection) were pipetted several times in order to
remove the first polar body and then exposed to calcium
ionophore at two concentrations (2 µM for 2 min and 5 µM
for 5 min) in calcium-free medium. After washing, each
oocyte was cultured in an individual microdrop and examined periodically for evidence of the second polar body
extrusion, which generally took place within an hour. Apart
from experiment (4), the percentage of oocytes showing
polar body extrusion after exposure to 5 µM ionophore
appeared high (from 90 to 98%), but only slightly superior
to that observed after 2 µM treatment (from 70 to 87%)
(Table 1).
The state of the chromatin organization was analysed at
least 4 hours after the ionophore treatment. Those oocytes
which had not shown any second polar body extrusion
(whether control or ionophore-treated) were in MII and
their condensed chromosomes were organized in a
metaphase plate (Fig. 1A, A′, A′′). The oocytes, from which
polar body extrusion had been recorded, showed different
behaviour not only according to the experiment but also
according to the concentration of ionophore used (Table 1).
Regardless of the concentration of ionophore, the incidence
of pronucleus formation varied from one experiment to
another. However, apart from experiment (4), the percentage of pronucleus formation in each experiment was always
significantly higher (P < 0.005; χ2 test) after exposure to 5
µM ionophore (from 27 to 76%) than after exposure to 2
µM ionophore (0 to 19%) (Table 1). Thus after exposure
to a low concentration of ionophore (2 µM), most of the
oocytes that had extruded their second polar body did not
form pronuclei. As these oocytes did not undergo full acti-
Cell cycle of activated oocytes
391
Table 1. Response of mouse oocytes to different concentrations of calcium ionophore
No. of oocytes with polar body
No. of oocytes
Incubation conditions
(1) Control (0.25% DMSO)
2 µM ionophore
5 µM ionophore
(2) Control (0.25% DMSO)
2 µM ionophore
5 µM ionophore
(3) Control (0.25% DMSO)
2 µM ionophore
5 µM ionophore
(4) Control (0.25% DMSO)
2 µM ionophore
5 µM ionophore
(5) Control (0.25% DMSO)
2 µM ionophore
5 µM ionophore†
Observed
Showing polar
body II (%)
Observed
With pronucleus
(%)
16
113
41
−
133
60
71
121
89
32
43
48
48
67
61
0 (0)
97 (86)
38 (93)
−
106 (80)
55 (92)
0 (0)
85 (71)
80 (90)
3 (9)
23 (54)
40 (87)
0 (0)
58 (87)
60 (98)
−
41
22
−
32
45
−
49
58
−
22
42
−
53
42
−
6 (15)
15 (68)*
−
6 (19)
24 (53)*
−
0 (0)
16 (27)*
−
10 (45)
20 (48)
−
8 (15)
32 (76)*
With condensed chromatin
(% of these that show a
metaphase spindle)
−
35 (80)
7 (43)
−
26 (58)
21 (67)
−
49 (100)
42 (98)
−
12 (100)
22 (95)
−
−
−
*Significant difference between 2 and 5 µM ionophore; P<0.005 (χ 2 test).
†Oocytes were exposed to the 5 µM ionophore for 30 min.
vation, their position in the cell cycle was examined by both
microtubule and chromatin staining at different times after
ionophore activation.
Metaphase III formation in oocytes that did not undergo
full activation
Before ionophore exposure (Fig. 1 A, A′, A′′), the chromosomes consisting of two chromatids (Fig. 1A′′) were
organized on a metaphase plate (MII, Fig. 1A′), and the
microtubules were organized in a short anastral barrelshaped spindle (Fig. 1A) (Maro et al. 1985). After
ionophore treatment, the oocytes underwent cell cycle progression through anaphase, separation of the sister chromatids and emission of the second polar body (Fig. 1B, B′,
B′′). Thereafter, in the majority of the oocytes (Table 1) a
new spindle appeared (MIII spindle; Fig. 1C) around the
unichromatid chromosomes (Fig. 1C′′). In the majority of
the cases, the chromosomes were scattered along the spindle (Fig. 1C′) which appeared longer and thinner (Fig. 1C)
than that observed in the MII arrested oocytes (Fig. 1A),
and sometimes the MIII spindle was abnormally shaped
(pointed-shape spindle, detached microtubules, multipolar
spindle). Observations 24 hours later (data not shown)
showed only a disorganization of the spindle microtubules
but no further progression through the cell cycle, emphasizing that the arrest of oocytes at this MIII stage is durable,
as it is in MII before activation. Metaphase III spindles were
observed in oocytes that had not undergone pronuclear formation after activation in either 2 or 5 µM ionophore
(Table 1).
Metaphase III formation does not occur in aged oocytes
Both the proportion of activated oocytes (assessed by the
extrusion of the second polar body) and the proportion of
oocytes showing a pronucleus increased with the post-ovulatory age of the oocytes: 16 h post-hCG, treatment with 2
µM ionophore induced 94% (49/52) polar body extrusion
and 98% (38/39) pronucleus formation. It is important to
note that in the case of such aged oocytes, decreasing the
concentration of ionophore used (1 µM and 0.5 µM) less ened the percentage of oocytes showing the extrusion of
the second polar body (respectively, 32% (11/34) and 3%
(1/39)) but failed to induce the formation of MIII; all the
oocytes which were activated formed a pronucleus.
Cortical granule extrusion
Cortical granule exocytosis is an event that takes place
within the few minutes following fertilization (Cran, 1988;
Fukuda and Chang, 1978) or parthenogenetic activation
(Gulyas and Yuan, 1985). However, recent studies reported
that the exocytosis of cortical granules might occur in
oocytes without parthenogenetic activation, either during in
vitro maturation (Ducibella et al. 1990) or during exposure
to 1.5 M DMSO (Vincent et al. 1991). We have investigated the occurrence of cortical granule extrusion when full
parthenogenetic activation was not achieved. The study was
performed on freshly ovulated oocytes after 2 µM
ionophore treatment when the incidence of MIII formation
was high (Table 1) and after 0.5 µM ionophore treatment
when activation assessed by polar body extrusion did not
occur (data not shown). The oocytes were not permeabilized before the lectin staining in order to assess
directly the amount of cortical granule content extruded
onto the oocyte surface (Table 2). After exposure to 2 µM
ionophore (Fig. 2C,C′), an extensive extrusion of granules
(++) had occurred in the majority of the oocytes (Table 2),
while no evidence of extrusion (Fig. 2A,A′; (−) Table 2) or
a low-density cortical granule extrusion (Fig. 2B,B′; (+)
Table 2) was found in the majority of the control oocytes
or in oocytes treated with 0.5 µM ionophore.
Calcium profile in oocytes activated at different ages and
with different concentrations of ionophore
After collection, the oocytes were either loaded with Fura2 in the chamber for analysis (freshly ovulated oocytes) or
left in culture in H6 + BSA under oil before loading (aged
392
C. Vincent and others
Metaphase II
Extrusion of the second
polar body
Metaphase III
Fig. 1. Analysis of the MII to MIII transition of mouse oocytes after exposure to 2 µM ionophore. Each oocyte was fixed and double
stained with both anti-tubulin antibody (A,B,C) and DNA-specific dye Hoechst 33342 (A′,B′,C′) or air-dried prepared and stained for
chromatin with Giemsa (A′′,B′′,C′′). (A,A′,A′′) Control oocytes before treatment. The oocyte presents a barrel-shaped spindle of
microtubules (A) around the chromosomes organized on a metaphase plate (A′). Each chromosome consists of two chromatids (A′′).
(B,B′,B′′) Oocytes observed 1.5 h after ionophore treament. The oocyte has gone through an anaphase, separation of the sister chromatids
and extrusion of the second polar body. (C,C′,C′′) Oocytes observed 4 h after ionophore treatment. The oocytes are arrested in a new
metaphase (MIII). The spindle is rather long and thin (C). The chromosomes are dispersed along the spindle (C′) and composed of single
chromatids (C′′).
oocytes). After 1 min incubation in H6 + PVP containing
1.2 mM calcium, oocytes were washed extensively with calcium-free medium for 5 min before the calcium ionophore
A23187 was added. The same concentrations of calcium
ionophore (2 µM and 5 µM) and the same times of expo sure (respectively, 2 min and 5 min) as the ones used for
the activation study (Table 1) were used for the calcium
analysis (Table 3; Fig. 3). After the ionophore treatment,
oocytes were washed continuously in calcium-free medium.
The results presented in this section came from a single
experiment. However, similar results have been obtained in
other replicates with similar numbers of oocytes in each
group. Exposure to control levels (0.25%) of DMSO did
not induce a calcium response.
As a result of the ionophore exposure in calcium-free
medium, only the internal stores of calcium are mobilized
for release (Stauderman and Pruss, 1989). Consequently,
one calcium transient is seen (Fig. 3) and the length of time
during which the oocytes are incubated with the ionophore
Table 2. Comparison of the cortical granule extrusion
occurring in mouse oocytes after treatment with different
concentrations of calcium ionophore
Number of oocytes according to the
density of cortical granule extrusion
Control (0.1% DMSO)
0.5 µM ionophore
2 µM ionophore
Total
−
+
++
58
24
78
38
23
8
15
1
13
5
0
57
Oocytes were classified according to the density of the cortical granule
contents observed on their surface.
−, Almost no cortical granules (Fig. 2A,A′); +, very low density of cortical
granules (Fig. 2B,B′); ++, high cortical granule density (Fig. 2C,C′).
Cell cycle of activated oocytes
−
+
393
++
Fig. 2. Fluorescence micrographs of cortical granule contents on the surface of mouse oocytes. Each oocyte was fixed and double stained
with Lens culnaris agglutinin (A, A′, B, B′, C, C′) and DNA-specific dye Hoechst 33342 (A′′,B′′,C′′). (A,B,C) oocytes viewed en face;
(A′,B′,C′) cross-section of the same oocytes. (A,A′,A′′) Almost no cortical granules are visible at the surface of the MII mouse oocyte (−).
(B,B′,B′′) Few cortical granules are detected at the surface of the MII mouse oocyte (+). (C, C′,C′′) High cortical granule concentration is
visible at the surface of the mouse oocyte after activation with 2 µM ionophore (++).
does not seem to influence the response; free cytosolic calcium declines even though the ionophore is still present
(Fig. 3B and D).
In order to analyse the response of the oocytes incubated
in different conditions, two parameters were chosen to characterize the shape of the calcium spike: in addition to the
amplitude (peak calcium reached; ∆ [Ca2+]i), the response
duration was quantified by measuring the width of the transient at 75% of the peak [Ca2+]i reached (full-width, 3/4
maximum, FWTM) (Thager and Miller, 1990). These parameters indicate differences in the total amount of [Ca2+]i
release during the incubation with the calcium ionophore.
Table 3. Characteristics of the calcium transient in freshly ovulated (12.5-13 h post-hCG) and aged (15-15.5 h post-hCG)
mouse oocytes activated by low (2 M) or high (5 M) concentration of ionophore
Measurements from individual oocyte
Ionophore
concentration
Freshly ovulated oocytes
2 µM
5 µM
Aged oocytes
2 µM
5 µM
(1)
(2)
(3)
(4)
(5)
Mean
∆Ca2+ (nM)
FWTM (s)
∆Ca2+ (nM)
FWTM (s)
145
98
226
106
151
98
223
120
96
85
192
113
177
85
303
70
127
77
237
120
139(a,b)
89(c)
236(a)
106(d)
∆Ca2+ (nM)
FWTM (s)
∆Ca2+ (nM)
FWTM (s)
223
141
367
134
295
141
433
197
272
127
257
127
186
169
124
197
178
183
254
296
231(b)
152(c)
287
190(d)
Means with similar letters are significant different (Student’s t-test): (a) P=0.001; (b) P=0.04; (c) P=0.009; (d) P=0.05.
394
C. Vincent and others
500
A
500
200
200
100
100
50
50
20
20
10
0
500
B
10
0
1000
500
Time (s)
C
500
200
100
100
50
50
20
20
500
1000
Time (s)
D
500
200
10
0
1000
Time (s)
10
0
500
1000
Time (s)
Fig. 3. [Ca 2+] responses induced in freshly ovulated (12.5-13 h post-hCG) and aged (15-15.5 h post-hCG) mouse oocytes by low (2 µM)
or high (5 µM) concentrations of ionophore. The graph plotted for each cell is representative of a typical response in each group (Table
3). (A) Freshly ovulated oocyte activated by 2 µM ionophore. (B) Freshly ovulated oocyte activated by 5 µM ionophore. (C) Aged oocyte
activated by 2 µM ionophore. (D) Aged oocyte activated by 5 µM ionophore. The arrowheads indicate the transition from medium
containing calcium (1.2 mM) to calcium-free medium. The bar represents the length of time during which the calcium ionophore was
applied (2 min for 2 µM ionophore, 5 min for 5 µM ionophore).
The amplitude of the spike was significantly higher in the
presence of 5 µM ionophore (Fig. 3B and D; Table 3) than
in the presence of 2 µM (Fig. 3A and C; Table 3), but only
for fresh, not aged, oocytes. The same concentration of
ionophore, whether 2 µM or 5 µM, stimulates a signifi cantly wider calcium transient in oocytes analysed after in
vitro ageing (Fig. 3C and D; Table 3) than in oocytes
analysed shortly after ovulation (Fig. 3A and B; Table 3).
Discussion
The formation of the MIII stage has been reported in freshly
ovulated mouse oocytes activated by ethanol (Kubiak,
1989). In that report, it was inferred that such oocytes were
not able to undergo the transition to interphase when activated by ethanol. Our results show that fresh ovulated
oocytes may also fail to complete a normal cell cycle when
activated by a short exposure to a low concentration of
ionophore. However, the majority of such freshly ovulated
oocytes could be forced into interphase by elevating the
concentration of ionophore used. Measurement of the calcium in the cytosol supports the idea that use of higher
ionophore concentrations causes release of more calcium
inside the cell. This result suggests that the MIII formation
seen in the majority of freshly ovulated oocytes which fail
to undergo a full activation can be related to a failure to
release enough cytosolic calcium. However, the limited calcium transient did successfully induce the release of cortical granules as is observed after fertilization or complete
parthenogenetic activation. The conclusion that entry to
interphase requires a larger calcium transient is reinforced
by the results on older oocytes. It has been known for a
long time that the proportion of activated oocytes increases
with post-ovulatory age (Kaufman, 1983; Whittingham and
Siracusa, 1978). We confirm, as previously reported by
Kubiak (1989), that the appearance of MIII is a phenomenon observed only in freshly ovulated oocytes and that the
Cell cycle of activated oocytes
proportion of oocytes showing a pronucleus increases with
the post-ovulatory age of the oocytes. Older oocytes also
give a wider calcium transient and show a higher calcium
amplitude in response to a lower ionophore concentration,
but the reason for these results is unknown. It is possible
that the ionophore partitions more effectively into the membrane lipid bilayers as the oocytes get older, increasing the
quantity of calcium released from the internal store to the
cytosol. Implicit in this conclusion is a change in the lipid
composition of the oocyte membranes occuring with aging
(Pratt, 1978). Indeed, when we perfused the oocytes at the
end of each run with a buffer containing calcium, it was
possible to see a small transient of [Ca2+]i in the group of
old oocytes (but not in fresh oocytes) that had been activated with 5 µM ionophore (results not shown), indicating
that ionophore is retained in old oocytes, but not fresh ones,
after washing. Direct analysis of the first calcium transient
induced in mouse oocytes by a fertilizing spermatozoon
shows a duration of between 2 and 3 min and an average
amplitude of 300 nM (Cheek et al. 1992). Thus this single
transient provides a sufficient rise in intracellular calcium
to activate entry into interphase, and the subsequent calcium transients observed after fertilization in mammals
might have some other role (Ozil, 1990).
We can conclude from our experiments that the cell cycle
progression of mouse oocytes appears to be dependent on
the profile of internal calcium release. In Xenopus, the calcium transient that follows oocyte activation is thought to
be involved in the inactivation of MPF (maturation-promoting factor) activity due to the degradation of the cyclin
B (Murray et al. 1989), and of CSF (cytostatic factor)
activity, linked to the destruction of the c-mos proto-oncogene product (Watanabe et al. 1989). There is evidence
from studies on Xenopus oocyte extracts in vitro that cyclin
B destruction can occur at a lower calcium concentration
than that which is required to destroy c-mos (Lorca et al.
1991). Moreover, whereas a Ca2+-calmodulin-dependent
process has been implicated in cyclin B destruction, the proteolysis of c-mos is probably caused by the calcium-dependent cysteine protease calpain (Lorca et al. 1991; Watanabe et al. 1989). In addition, work on Xenopus (Lorca et
al. 1991; Watanabe et al. 1991) and mouse (Weber et al.
1991) oocytes released from meiosis has shown that the
CSF inactivation is not the primary cause of MPF inactivation. A mouse oocyte which passes into metaphase III
must have exited from the metaphase II arrest, and to
achieve this should have decreased its MPF activity. However, the fact that its chromatin remains condensed argues
that sufficient CSF activity is still present to stabilize residual or newly formed MPF activity. Thus, the differential
sensitivity of MPF and CSF to destruction by calcium
observed in in vitro extracts of Xenopus (Lorca et al. 1991)
may have an in vivo counterpart in the mouse metaphase
III model.
We thank M. George and B. Doe for technical assistance, J.
Bashford and colleagues for photographic work, J. Kilmartin for
the anti-tubulin antibody, and S.J. Pickering, M.J. Berridge, R.B.
Moreton and J. McConnell for helpful discussion. This work was
supported by a Medical Research Council programme grant to
395
M.H. Johnson and P.R. Braude, and by a grant to M.H. Johnson
from the Wellcome Trust.
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(Received 19 November 1991 - Accepted, in revised form, 23 July 1992)