ARTICLE
Oocyte-derived microvilli control female fertility by
optimizing ovarian follicle selection in mice
Yan Zhang1,4, Ye Wang1,4, Xie’an Feng1, Shuo Zhang1, Xueqiang Xu1, Lingyu Li1, Shudong Niu1, Yingnan Bo1,
Chao Wang1, Zhen Li2, Guoliang Xia1,3 & Hua Zhang 1✉
Crosstalk between oocytes and surrounding somatic cells is crucial for mammalian oogenesis,
but the structural mechanisms on oocytes to control female reproduction remain
unknown. Here we combine endogenous-fluorescent tracing mouse models with a highresolution
live-cell imaging system to characterize oocyte-derived mushroom-like microvilli
(Oo-Mvi), which mediate germ-somatic communication in mice. We perform 3D live-cell
imaging to show that Oo-Mvi exhibit cellular characteristics that fit an exocrine function for
signaling communication. We find that deletion of the microvilli-forming gene Radixin in
oocytes leads to the loss of Oo-Mvi in ovaries, and causes a series of abnormalities in ovarian
development, resulting in shortened reproductive lifespan in females. Mechanistically, we find
that Oo-Mvi enrich oocyte-secreted factors and control their release, resulting in optimal
selection of ovarian follicles. Taken together, our data show that the Oo-Mvi system controls
the female reproductive lifespan by governing the fate of follicles.
https://doi.org/10.1038/s41467-021-22829-2 OPEN
1 State Key Laboratory of Agrobiotechnology, College of Biological Sciences, China Agricultural University, Beijing, China. 2 State Key Laboratory of Plant
Physiology and Biochemistry, College of Biological Sciences, China Agricultural University, Beijing, China. 3 Key Laboratory of Ministry of Education for
Conservation and Utilization of Special Biological Resources in Western China, College of Life Science, Ningxia University, Yinchuan, Ningxia, China. 4
These
authors contributed equally: Yan Zhang, Ye Wang. ✉email: huazhang@cau.edu.cn
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I
n animals with sexual reproduction, female oocytes are primarily
responsible for supporting the early development of
their offspring. Therefore, a robust and finely regulated
development of oocytes in adult life is the foundation of species
continuity1,2. In mammals, the oocyte is covered by surrounding
somatic cells (granulosa cells, GCs) to form the ovarian follicle as
the functional unit of female reproduction3. The synchronous
development of oocyte and GCs orchestrates programmed folliculogenesis
to guarantee the production of qualified egg2,4.
However, growing oocytes establish a glycoprotein egg coat, the
zona pellucida (ZP), blocking the direct cellular membrane
attachment between oocytes and GCs5,6.
To overcome the blockage of the ZP, microvilli-like structures
existing in the ZP are believed to construct communications
between oocyte and GCs, maintaining proper
folliculogenesis7. Previous results from electron microscopy
and cytoskeleton immunostaining studies showed that those
structures, named transzonal projections (TZPs), are derived
from the GCs of follicles in mammals8–10. Several reports
have also suggested that oocytes might form microvillus
structures to participate in follicle development11,12. The lack
of direct in vivo experimental evidence, however, made the
cellular origin of those communicating structures in the
mammalian ovary elusive13–15. Meanwhile, the inner mechanisms
regulating the establishment of those structures, and the
physiological significance of those microvilli in female reproduction
have not been well identified.
In the mammalian ovary, a complicated but orchestrated
regulating system determines the developmental fate of each
individual follicle, and the balance of follicle survival and death
determines the proper female reproductive lifespan3,16. Inside
the follicles, the pivotal role of oocytes governing follicle
development has been well accepted2,4. By releasing various
specific oocyte-secreted factors (OSFs), such as growth differentiation
factor 9 (GDF9) and bone morphogenic protein 15
(BMP15)17–19, oocytes control the proliferation and activity of
GCs to determine the fate of follicles2. However, as soluble
factors, OSFs need to pass through the ZP to activate downstream
signaling in GCs, but the existence of the ZP increases
the physical distance between oocytes and GCs, which might
reduce the efficiency of OSFs. The cellular structural basis of
the oocyte to efficiently release the OSFs is not clear, and the
mechanisms of how oocyte overcomes the ZP blockage to stimulate
GC activity is unknown.
In the current study, we report an oocyte-derived microvilli
system that mediates orderly and timely OSF release, which
contributes to the fine regulation of ovarian follicle development
and female reproductive lifespan maintenance. By combining
endogenous-fluorescent tracing mouse models with a highresolution
live-cell imaging system, we establish systems to
image the developmental details of mouse ovarian follicle cells at
the single-cell and subcellular levels. With this system, we trace
and describe the derivation and cellular behaviors of microvilli
structures in ovarian follicles in detail. Our imaging observations
show a cellular character of oocyte microvilli (Oo-Mvi), which
play regulating roles in the development of ovarian follicles. By
deleting Radixin (Rdx), which is a gene belonging to the
microvillus-forming related ERM (Ezrin/Radixin/Moesin) gene
family in oocytes20,21, we disrupt the formation of Oo-Mvi and
determine how these structures regulate the release of OSFs to
govern female reproduction in mice. With these experimental
data, we identify an elegant system by which Oo-Mvi in the ZP
determine the fate of individual follicles by regulating oocyte-GC
communication, which in turn adjusts the balance of follicle
survival and death to determine the reproductive lifespan in
female mammals.
Results
Imaging microvilli structures on oocyte by endogenousfluorescent
mouse models. As a conserved communicating
structure in the ZP, microvilli or TZPs have been well documented
to exist between oocytes and GCs22. To visualize these
structures and distinguish their cellular origin during follicle
development, an mTmG reporter mouse model in which
membrane-localized GFP (mG) is inducibly expressed in Crepositive
cells23, was introduced in this study. As illustrated in
Fig. 1a, by crossing the mTmG mice with oocyte-specific Gdf9-Cre
mice24 or granulosa cell-specific Foxl2-CreERT2 mice25, we
established a system to distinguish the derivation of microvilli by
drawing the outlines of the cellular membrane with mG.
We first examined the ovaries in tamoxifen-treated Foxl2CreERT2;mTmG
females (Supplementary Fig. 1a), intensive GCTZPs
were observed in the ZP of growing follicles (Fig. 1b,
arrows) under spinning-disc confocal microscopy. By modifying
tamoxifen dosage, which allowed a few of GCs to be labeled and
imaged, we reconstructed the 3D model of GC membrane and
revealed a messy tree root-like structure of GC-TZPs at the
single-cell level (Fig. 1b, right). These results are consistent with
previous observations of GC-TZPs10 and further demonstrated
that GC-TZPs play a supporting function in oocyte
development7.
Interestingly, in Gdf9-Cre;mTmG ovaries (Supplementary
Fig. 1b), a remarkably low density of microvilli structures derived
from oocytes were found in the ZP in growing follicles (Fig. 1c,
arrowheads). These Oo-Mvi exhibited a mushroom-like structure
that consisted of a slender handle and a swollen vesicle tip
(Fig. 1c, right). Histologically, we found that the establishment of
Oo-Mvi was consistent with the growth of follicles (Fig. 1d). With
the formation of the ZP, Oo-Mvi were first observed on the
membrane of oocytes in primary follicles (Fig. 1d, arrowheads)
and existed in the ZP at all growing stages (Fig. 1d, arrowheads),
showing that the function of Oo-Mvi should be related to the
growth of ovarian follicles. We next investigated the behaviors of
Oo-Mvi in living oocytes isolated from adult Gdf9-Cre;mTmG
ovaries. As shown in Fig. 1e, Oo-Mvi was clearly observed on the
surface of oocytes under a live-cell imaging system (Supplementary
Movie 1). Over the course of 50 min, no remarkable growth
or extension of Oo-Mvi was observed (Supplementary Movie 2),
showing that the formation of Oo-Mvi was not active in fully
grown oocytes. However, a clear breakdown of head vesicles in
some Oo-Mvi (Fig. 1e arrowheads and Supplementary Movie 3)
was detected in living oocytes, showing complex cellular
behaviors occurred in those Oo-Mvi.
Therefore, our imaging results showed that there are two types
of communicating structures in the ZP: highly dense TZPs
derived from GCs and sparse microvilli with vesicles formed by
oocytes. Unlike the well-observed GC-TZPs, Oo-Mvi exhibited a
series of characteristics and cellular behaviors. Since oocytes play
a central role in controlling follicular growth4, we therefore,
hypothesized that Oo-Mvi act as a regulator of ovarian follicle
development in female reproduction.
The microvilli-specific protein RADIXIN was specifically
expressed in oocytes. To clarify our hypothesis and investigate
the functional role of Oo-Mvi in folliculogenesis, we isolated the
ZP containing Oo-Mvi (Fig. 2a, arrowheads) from oocytes and
performed mass spectrometry (MS) analysis to identify potential
regulators of the formation of Oo-Mvi. In total, 66 proteins were
detected in ZP samples, including the well-studied ZP structure
proteins ZP1/2/326 and gap junction protein alpha 1 (GJA1)27,
fertilization-related protein CD912, and cytoskeletal proteins
β-actin and myosin heavy chain 9 (MYH9)27 (Fig. 2b).
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Furthermore, one ERM family member, RADIXIN (RDX), which
has been reported to control the formation of microvilli in many
organs28, was also detected in our MS results (Fig. 2b and Supplementary
Fig. 2). To confirm our findings, we performed
quantitative real-time PCR (QRT-PCR) to detect the mRNA
levels of different ERM family members in the ovary. The mRNA
level of Rdx in oocytes was approximately 700 times higher than
that in ovarian somatic cells and other tissues (Fig. 2c), whereas
the mRNA of the other two ERM family members, Ezrin and
Moesin, were not highly expressed in the oocytes (Supplementary
Fig. 3a, b). At the protein level, western blot results also showed
that RDX and its activated form phosphorylated (p-) ERM were
highly expressed in oocytes (Fig. 2d). These results showed that
RDX might be a specific microvillus-related protein in oocytes
that contributes to the formation of Oo-Mvi in oocyte
development.
We next detected the protein localization of RDX as well as
other ERM family members in the ovary. Consistent with the
QRT-PCR result, EZRIN and MOESIN were only detected in
non-follicular cells such as stromal cells or blood vessels in the
ovary (Supplementary Fig. 3c, arrows), and no expression was
found in oocytes (Supplementary Fig. 3c, arrowheads). In sharp
contrast, the immunofluorescent signals of RDX were detected in
both the membrane and cytoplasm of oocytes at all stages of
folliculogenesis (Fig. 2e, arrows and arrowheads), including
dormant primordial follicles (Fig. 2e, arrowheads). Meanwhile,
the localization of RDX in Oo-Mvi under high magnification
(Fig. 2f, arrowheads) indicated that RDX might participate in the
formation of Oo-Mvi. Compared to total RDX, activated p-ERM
was only localized on the cell membrane (Fig. 2g, arrows) and
Oo-Mvi in growing oocytes (Fig. 2h, arrowheads) but was not
observed in dormant oocytes of primordial follicles (Fig. 2g,
mT mG polyA
mG polyA
mTmG
Foxl2-CreER
T2
Gdf9-Cre
GCs
Oo
GCs
Oo
GC
Oo
Oo
Foxl2-CreER ;mTmG
Gdf9-Cre;mTmG
T2
Foxl2-CreER ;mTmG
T2
Gdf9-Cre;mTmG
a b
c
Oo-Mvi
0 min 10 min 20 min 30 min
e
d
AntralSecondaryPrimary
GCs
Oo
GCs
Oo
GCs
Oo
Follicle Single GC
mT/mG/Hoe/Inverted b/w
Fig. 1 Imaging cell-specific communicating structures in the ZP by endogenous-fluorescent labeling in follicles. a Illustration of the strategy to label the
derivation of the cellular communicating structures in the ZP by cellular specifically expressed CreERT2 or Cre recombinase. Membrane-localized redfluorescent
protein (mT) switches to green-fluorescent protein (mG) in GCs of Foxl2-CreERT2;mTmG follicles and oocytes of Gdf9-Cre;mTmG follicles.
b Images of Foxl2-CreERT2;mTmG ovarian sections showing the high density of GC-TZPs (arrows), and labeling of a single GC showing the messy tree rootlike
structure of GC-TZPs in detail (right). c Images of Gdf9-Cre;mTmG ovarian sections, the mushroom-like Oo-Mvi (arrowheads) distributed in a low
density (left) and high magnification showing the vesicle tips (arrowheads) of Oo-Mvi (right). d The developmental dynamics of Oo-Mvi (arrowheads),
showing the establishment of Oo-Mvi accompanied by follicle growth. e Time-lapse imaging of Oo-Mvi in living oocytes showing the breakdown of vesicles
(arrowheads) of Oo-Mvi (Supplementary Movie 3). All experiments were repeated at least three times, and representative images are shown. Oo oocyte.
GCs granulosa cells. ZP zona pellucida. b–e The colors were inverted to black/white (b/w) to highlight GC-TZPs (b) and Oo-Mvi (c–e). Scale bars: 5 μm
(b–d) and 2 μm (e).
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Protein name Score Significant
matches
β-actin
MYH9
ZP1
ZP2
ZP3
GJA1
CD9
RADIXIN
775
168
214
717
286
155
70
58
32
7
12
45
11
5
3
6
9
6
4
14
6
2
1
4
Significant
sequences
BF BF/mG
800
700
600
500
20
10
0
O
vary
O
o
G
C
Intestine
Lung
Oo GC Ovary
RADIXIN
p-ERM
α-TUBULIN
RelativemRNAlevelofRadixin
***
GCs
GCs
Oo
Oo
GCs
GCs
Oo
Oo
ZP
ZP
a b
c d
e
g
f
h
Isolated ZP
mG
RDX/Hoe
p-ERM/Hoe
RDX/Hoe
p-ERM/Hoe
72
95
kDa
72
95
43
55
Fig. 2 The microvilli-related protein RDX was specifically expressed in oocytes. a Isolated ZP from Gdf9-Cre;mTmG oocytes with Oo-Mvi (arrowheads)
kept in the samples for MS analysis. b MS analysis revealing the expression of RDX in the isolated ZP. c Relative mRNA levels of Rdx in different follicle
components and tissues showing high Rdx expression in oocytes. n = 3 and P-value (Ovary vs. Oo) = 0.0014. d Western blot showing that RDX and p-ERM
are highly expressed in oocytes. e Immunostaining showing RDX protein localized in the oocytes of follicles from the dormant primordial (arrowheads) to
growing (arrow) stages. g p-ERM highly expresses on the membrane of oocytes in growing follicles (arrows) but not on that of dormant oocytes in
primordial follicles (arrowheads). f, h High-magnification images showing the localization of RDX (f) or p-ERM (h) in the Oo-Mvi (arrowheads). All
experiments were repeated at least three times, and representative images are shown. Data are presented as the mean ± SD with experiments performed
in triplicate. Data were analyzed by two-tailed unpaired Student’s t-test and ***P < 0.001. Oo oocyte. GCs granulosa cells. ZP zona pellucida. Scale bars:
25 μm (a), 100 μm (e, g), 10 μm (f, h).
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arrowheads). This localization is consistent with the formation of
Oo-Mvi in follicle development, indicating that RDX in oocytes is
involved in the formation of Oo-Mvi, and the major function of
RDX and Oo-Mvi might be related to the growth of oocytes and
follicles in the ovary.
Oocytic Rdx deletion resulted in a failure of the formation of
Oo-Mvi and a shortened reproductive lifespan in females. To
study the function of Oo-Mvi, we generated a conditional Rdx
knockout mouse by flanking exons 4/5 with loxP sequences
(RdxloxP) (Fig. 3a). RdxloxP/loxP females were crossed with Gdf9Cre
males to delete Rdx in oocytes at the primordial follicle stage
(Fig. 3a). The resulting Gdf9-Cre;RdxloxP/loxP mice were referred
to as Oo-Rdx−/− mice, and the littermates of No-Cre;RdxloxP/loxP
females, referred to as Oo-Rdx+/+, were used as the control. To
validate the efficiency of deletion in Oo-Rdx−/− mice, we performed
immunofluorescent staining analysis to detect the
Oo-Rdx Oo-Rdx+/+ -/Oo-Rdx
;mTmG Oo-Rdx ;mTmG+/+ -/Living
oocyte (mG)
Oo-Rdx ;mTmG Oo-Rdx ;mTmG+/+ -/Ovarian
tissue section (mG/Hoe)
0
10
20
30
40
Oo-Rdx
+/+
Oo-Rdx
No.ofovulatedoocytes
**
a b
c d
e f
Oo-Rdx+/+
Oo-Rdx-/-
(n=9)
(n=8)
0
5
10
15
20
25
0 4 8 12 16 20 24 28 32 36 40
No.ofpupsperfemale
Weeks after mating
30
Exon loxP ~ 2060 bp
RdxloxP/loxP
Gdf9-Cre Gdf9 Cre
Oo-Rdx-/-
4 51 2 3 6 7
x
Oo-Rdx
+/+
Oo-Rdx
-/-RDX/Hoep-ERM/Hoe
1 2 3 6 7
UTR
50
-/Fig.
3 Deletion of Rdx in oocytes led to a failure of Oo-Mvi formation and shortened reproductive lifespan in females. a Schematic representation of the
deletion of Rdx exons 4 and 5 by Gdf9-Cre-mediated recombination in oocytes of Oo-Rdx−/−. b Immunofluorescence detection of RDX and p-ERM showing
successful deletion of RDX in oocytes (arrowheads) of Oo-Rdx−/−. n = 6 ovaries per group. c, d Imaging the living oocytes (c) and ovarian tissue sections
(d) showing a failure of the construction of Oo-Mvi on the oocyte surface in Oo-Rdx−/−;mTmG females. n = 40 Oo-Rdx+/+;mTmG and 16 Oo-Rdx−/−;mTmG
oocytes (c), 12 Oo-Rdx+/+;mTmG and 9 Oo-Rdx−/−;mTmG ovaries (d). e Fertility check showing a subfertility phenotype in the Oo-Rdx−/− females with
significantly decreased litter size and number and a shortened reproductive lifespan (n = 8) compared to that in Oo-Rdx+/+ mice (n = 9) during 40 weeks
of mating. f A significantly lower ovulated oocyte number in Oo-Rdx−/− females (n = 5) compared to that in Oo-Rdx+/+ mice (n = 4), indicating that the
abnormal development of oocytes caused subfertility in Oo-Rdx−/− females. P-value = 0.0036. Representative images are shown. Data are presented as
the mean ± SD. Data were analyzed by two-tailed unpaired Student’s t-test and **P < 0.01. Scale bars: 100 μm (b, f), 25 μm (c, d).
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expression of RDX in the ovaries. In the Oo-Rdx+/+ ovary, the
expression of both RDX and p-ERM was observed in oocytes of
follicles (Fig. 3b, arrowheads). In sharp contrast, neither RDX nor
p-ERM expression was detected in oocytes of Oo-Rdx−/− ovaries
(Fig. 3b, arrowheads), demonstrating a successful oocytic deletion
of Rdx.
Since RDX is highly expressed in the cortex of oocytes, we first
detected the construction of oocyte cortex in Oo-Rdx−/− females
followed a standard approach by labeling the actin with Alex488phalloidin
and detecting the cortical granules with LCA-FITC in
oocytes10,29,30. As a result, an identical cortex was found in the
mutant Oo-Rdx−/− oocytes and Oo-Rdx+/+controls (Supplementary
Fig. 4), showing that deletion of Rdx had no effect on the
cortical formation of oocytes. We next detected the cellular
architecture changes of oocytes in Oo-Rdx−/− mice by introducing
the mTmG reporter mice into the model to outline the cell
surface of the gene-modified oocytes (Supplementary Fig. 5). As
shown in Fig. 3c, normal Oo-Mvi with vesicles were found on the
surface of living oocytes in control Oo-Rdx+/+;mTmG females.
However, a smooth surface with few microvilli was observed on
oocytes of Oo-Rdx−/−;mTmG ovaries, showing that the deletion
of RDX successfully disrupted the formation of microvilli on
oocytes (Fig. 3c). Meanwhile, histological analysis showed that
Oo-Mvi were hardly detected in the ZP of Oo-Rdx−/−;mTmG
follicles, confirming that Rdx deletion led to a failure of the
establishment of Oo-Mvi during follicle development (Fig. 3d).
We next focused our study on the effect of the loss of Oo-Mvi on
female fertility. Surprisingly, the absence of Oo-Mvi did not lead to
complete infertility in Oo-Rdx−/− females. Most Oo-Rdx−/−
females gave birth from 4 weeks after mating, but all of them
became infertile before 24 weeks after mating, showing a phenotype
of a remarkably shortened reproductive lifespan compared to their
Oo-Rdx+/+ littermates (Fig. 3e). Moreover, the time of first labor
was delayed, and the litter frequency and litter size were
significantly decreased in Oo-Rdx−/− females compared to those
in the control mice (Fig. 3e), demonstrating a subfertile phenotype
in female Oo-Rdx−/− mice. Meanwhile, Oo-Rdx−/− females only
ovulated 18.40 ± 2.66 oocytes after superovulation, which was in
sharp contrast to the 33.75 ± 2.39 ovulated oocytes in Oo-Rdx+/+
females (Fig. 3f), indicating that abnormal folliculogenesis is the
major cause of female subfertility in Oo-Rdx−/− females.
Loss of Oo-Mvi led to a retardation of oocyte and follicle
development in Oo-Rdx−/− females. To investigate the inner
mechanisms of female subfertility in Oo-Rdx−/− mice, we first
detected the development of Oo-Rdx−/− ovaries at different ages
(Fig. 4a and Supplementary Fig. 6). At Postnatal Day (PD) 5, a
similar morphology of ovaries was found in Oo-Rdx−/− mice and
control mice (Supplementary Fig. 6a), showing that oocyte deletion of
Rdx does not affect the formation or survival of dormant primordial
follicles. At PD35, although the size of ovaries did not differ between
Oo-Rdx−/− and control females, a remarkably lower density of follicles
was observed in Oo-Rdx−/− ovaries than in the Oo-Rdx+/+
female littermates (Fig. 4a, PD35). At 2 months, the ovaries in
mutant females were significantly smaller than those in the controls
(Figs. 4a, 2 months), and more atresia follicles were observed in the
Oo-Rdx−/− ovaries (Fig. 4a, arrows and Supplementary Fig. 6b,
arrows). Consistent with the phenotype of premature ovarian
insufficiency, the Oo-Rdx−/− ovaries no longer displayed normal
ovarian morphology at 8 months (Fig. 4a, 8 months), and no corpus
luteum but many atresia follicles (Fig. 4a, arrows) were observed in
the mutant ovaries. Quantification of follicles in Oo-Rdx−/− and
control females confirmed the accelerating follicle loss in Oo-Rdx−/−
ovaries (Fig. 4b), and the peak of follicle loss occurred in Oo-Rdx−/−
ovaries before adulthood (Fig. 4b), which is a period of fast activation
and growth of ovarian follicles in the female reproductive lifespan25.
Therefore, we hypothesized that loss of Oo-Mvi might decrease
the developmental capability of growing follicles in ovaries, leading
to increased follicle loss and premature ovarian insufficiency in
females.
To test our hypothesis, we isolated and cultured ovarian
cortical pieces or follicles from Oo-Rdx−/− and Oo-Rdx+/+
females and traced the developmental dynamics of growing
oocytes and follicles in vitro. By culturing the ovarian cortical
pieces from Oo-Rdx−/−;mTmG and Oo-Rdx+/+;mTmG females,
we found significant retardation of the oocyte diameter increase
in Oo-Rdx−/−;mTmG females compared to that in controls
during 7 days of in vitro growth (Fig. 4c, d, 1.28 ± 0.09 times vs.
1.79 ± 0.07 times), showing that loss of Oo-Mvi by Rdx deletion
disrupted the growth of oocytes in the follicles. Next, we isolated
and cultured follicles from Oo-Rdx−/− ovaries and Oo-Rdx+/+
ovaries. Consistent with the retardation of oocyte growth, the
increase in follicle diameters in Oo-Rdx−/− females (117.60 ± 1.50
to 248.50 ± 15.25 μm) was also significantly slower than that in
Oo-Rdx+/+ females (121.90 ± 1.39 to 322.50 ± 21.79 μm) during
8 days of in vitro culture (Fig. 4e, f). These results clearly showed
that loss of Oo-Mvi might decrease the developmental capability
of growing follicles in ovaries.
Loss of Oo-Mvi affected the development of follicle GCs in OoRdx−/−
females. Based on the behaviors of Oo-Mvi in living oocytes
and the general phenotypes of Oo-Rdx−/− females, we proposed that
the function of Oo-Mvi might be to regulate the development of GCs
that support the growth of follicles. To test our hypothesis, we
detected the developmental status of GCs in Oo-Rdx−/− females.
Although GC proliferation rate varies greatly among follicles, we
found that the rate of GC proliferation in primary and secondary
follicles was significantly lower in Oo-Rdx−/− females than in controls
(Fig. 5a, arrows and b, 14.92% ± 2.53% vs. 34.74% ± 2.52% in the
primary stage, 34.47% ± 2.17% vs. 43.83% ± 2.65% in the secondary
stage). Meanwhile, TUNEL detection also showed a remarkable
increase in GC apoptosis in primary and secondary follicles of OoRdx−/−
ovaries (Fig. 5c, arrows and d, 16.44% ± 3.52% vs. 0 in the
primary stage, 29.98% ± 4.87% vs. 13.51% ± 0.89% in secondary
stage). These results showed that loss of Oo-Mvi decreased the
developmental capability of GCs in early growing follicles in ovaries.
To further investigate the effect of the loss of Oo-Mvi on the
development of GCs, we next tested the formation of GC-TZPs in
oocytes lacking Oo-Mvi in Oo-Rdx−/− females. Oocytes were
collected from Oo-Rdx−/− or Oo-Rdx+/+ ovaries at PD35 and Factin
was stained to show the TZPs within the ZP10. As expected,
a high density of TZPs covering the oocyte surface was observed
in the ZP of control oocytes (Fig. 5e, Oo-Rdx+/+). However, only
a small portion of GCs containing TZPs existed as groups in the
ZP (Fig. 5e, Oo-Rdx−/−, arrows), and significantly fewer GCTZPs
on the surface of oocytes were detected in Oo-Rdx−/−
females than in controls (Figs. 5f, 28.14 ± 3.94 vs. 121.70 ± 9.03).
This result showed a failure of GC-TZP network construction in
the follicles without Oo-Mvi, indicating a controlling role of OoMvi
in the formation of GC-TZPs.
Previous studies reported that OSFs, such as GDF9, control the
formation of GC-TZPs10. To further investigate whether the
effect of the loss of Oo-Mvi on follicle development is related to
OSF release, we cultured ovarian cortical pieces from Oo-Rdx−/−
females with or without GDF9 and examined the formation of
GC-TZPs. After 24 h of culture with GDF9 (100 ng/mL), the
failure of TZP network construction was well rescued by
GDF9 supplementation (Fig. 5g, Oo-Rdx−/−+GDF9). TZPcounting
results confirmed our observations that supplying
GDF9 significantly increased the density of GC-TZPs in OoARTICLE
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Rdx−/− follicles (Fig. 5h, 60.05 ± 5.72 in Oo-Rdx−/−+GDF9,
compared to 26.32 ± 3.00 in Oo-Rdx−/−, and 69.25 ± 4.42 in OoRdx+/+).
Moreover, we found a significantly decreased expression
of Ptgs2 and Tnfnip6, which were reported as the GDF9downstream
genes18 in Oo-Rdx−/− females (Supplementary
Fig. 7), indicating deletion of Rdx led to a GDF9 related
deficiency of ovaries. These results supported our hypothesis that
Oo-Mvi regulate GC function and follicle development, and this
regulation relates to the release of OSFs in the ovary.
Oo-Mvi were involved in the release of OSFs to manipulate the
development of follicles. Our in vivo and in vitro experimental
evidence strongly indicated that Oo-Mvi might regulate OSF
PD352months8months
Day 0 Day 7
**
*
100
200
300
400
Folliclediameters(μm)
Oo-Rdx+/+
Oo-Rdx-/-
Oo-Rdx+/+
Oo-Rdx-/Ooctyes
(mG)
Oo-Rdx+/+
Oo-Rdx-/Day
0
Ovarian follicles (BF)
d
1
*
***
1.5
2
Therateofoocytegrowh Oo-Rdx+/+
Oo-Rdx -/-
e
0
0 3 7 Day
0 4 8 Day
a
b c
f
ControlMutant
Day 4 Day 8
7,000
5,000
3,000
1,000
0
PD
5PD
352M
8M
No.offollicles
Oo-Rdx+/+
Oo-Rdx-/-
**
***
*
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release to control the crosstalk between oocytes and GCs. To
confirm our hypothesis, the expression of GDF9 was detected as a
representative molecule in the oocytes of PD35 ovaries. By costaining
of RDX and GDF9 in ovaries, we found that the signal of
GDF9 was clearly observed in the vesicles of Oo-Mvi on oocytes
(Fig. 6a, arrowheads, Supplementary Fig. 8). This result showed
that GDF9 distributed in the tip of Oo-Mvi, indicating OSFs were
transported to the vesicles of Oo-Mvi to be released.
To clarify the direct relationship between Oo-Mvi and OSF
release, we next performed oocyte microinjection to introduce
GDF9 coupled with red-fluorescent rhodamine (R-GDF9) using
the NHS-rhodamine amine-specific labeling method. In this
system, the localization of injected R-GDF9 in oocytes could be
directly visualized in culture. As the negative control, rhodaminelabeled
bovine serum albumin (R-BSA) (50 μg/mL) was first
injected into oocytes from wild-type females, and a dispersed redfluorescent
signal (Fig. 6b, R-BSA) showed a uniform distribution
of non-specific protein in the oocytes. In sharp contrast, the RGDF9
(50 μg/mL) signal aggregated as cloudy structures in the
cytoplasm of oocytes (Fig. 6b, R-GDF9), and various R-GDF9
spots were observed in the ZP of injected oocytes (Fig. 6b, RGDF9,
arrowheads, Supplementary Movies 4 and 5), which were
similar to the distribution of the vesicles of Oo-Mvi in Gdf9-Cre;
mTmG oocytes (Supplementary Fig. 9, arrows). Interestingly, we
found a clear reduction of Rho-fluorescent intensity in the
oocytes with the increase of time after R-GDF9 injection from 15
to 30 min (Fig. 6b, c), showing a fast release of the R-GDF9 from
oocytes. With the decrease of the R-GDF9 signal in oocytes, a
time-dependent increase of the R-GDF9 spot number (Fig. 6b,
arrowheads, Supplementary Movies 4 and 5) was observed on
oocytes after R-GDF9 injection. As we showed in Fig. 6d, the
average number of R-GDF9 spots was increased 8 ± 4 at ~15 min
to 25 ± 9 at ~30 min after injection. Therefore, these results
supplied direct experimental evidence showing that the
injected exogenous OSF was enriched and released through the
Oo-Mvi.
Since the secretory proteins including the OSFs are transported
through the endoplasmic reticulum (ER) to the plasma
membrane31, we next detected the localization of ER in GV
oocytes by the ER-tracker with a high-resolution approach. We
found that the ER exhibited an aggregative and cloudy
distribution in the oocyte cytoplasm (Fig. 6e, top panel), which
was consistent with the previously reports32,33. Interestingly, we
also found lots of ER-tracker labeled bubbles (ER bubbles) in the
ZP (Fig. 6e, top panel, arrowheads), which were similar as the
distribution of Oo-Mvi and the R-GDF9 bubbles (Fig. 6b, e,
arrowheads and Supplementary Movie 6). The counting results
showed that a comparable number of ER bubbles (107 ± 6) and
mG-labeled vesicles (110 ± 7) on Oo-Mvi of Gdf9-Cre;mTmG
ovaries (Fig. 6f). Furthermore, the quantitation of the fluorescent
intensity showed that these ER bubbles exhibited a significantly
higher signal intensity than that in the oocyte cytoplasm (Fig. 6g
and Supplementary Fig. 10). Therefore, these results supplied
evidence suggesting the enrichment of ER-transported protein in
the vesicles of Oo-Mvi, which was in consistent with our
conclusion about the OSF release function of Oo-Mvi.
Finally, we detected the mRNA levels of several well-studied
OSFs, including Gdf9, Bmp15, and Fgf834, in fully grown oocytes
from Oo-Rdx−/− and Oo-Rdx+/+ ovaries. We found that the
mRNA levels of these OSFs in Oo-Rdx−/− oocytes were
significantly higher (Gdf9, 2.33 ± 0.28 times, Bmp15, 2.20 ± 0.38
times, and Fgf8, 2.91 ± 1.54 times) than those in the Oo-Rdx+/+
control (Fig. 6h). Since the majority of oocyte mRNA transcripts
occur at the early stage of follicle development35, the increased
levels of mRNAs in ovulated oocytes showed that the consumption
of storage OSF mRNAs was dramatically reduced in oocytes
without Oo-Mvi. Therefore, the data confirmed our finding that
Oo-Mvi contributed to the release of OSFs.
Taken together, our data provide direct evidence that oocytes
establish a microvillus network to control the development of
GCs in follicles. These Oo-Mvi shorten the physical distance
between oocytes and surrounding GCs, and enrich and release
OSFs in their vesicle tips to maintain normal follicle development
and survival in the ovary. The loss of Oo-Mvi in Oo-Rdx−/−
females decrease the efficiency of OSF stimulation, thus leading to
abnormal GC development and finally affecting the survival of
follicles and shortening the reproductive lifespan in females
(Fig. 6i).
Discussion
The developing regulation of oocytes is a complex and multi-cell
participating process in all vertebrate species. In mammals, each
oocyte is covered by surrounding somatic GCs to form a relatively
independent structure called ovarian follicles as the fundamental
unit of female reproduction3. Active bidirectional communications
between oocytes and GCs are essential for oogenesis in adult
ovaries, and oocytes have been well accepted to dominate intercellular
conversation during folliculogenesis7. By timely and
orderly secretion of the oocyte-secreted factors, oocytes govern
the proliferation and differentiation of GCs to intermediate the
germ-somatic crosstalk4,36. Controversially, oocyte also formed a
glycoprotein egg coat, called the zona pellucida (ZP) to block the
direct communication with the follicle somatic cells5. Till now,
the accurate cellular constructs on oocytes supporting these fineregulated
dialogs are unclear. Here, by combining endogenousfluorescent
mouse models with a high-resolution imaging system,
we identified and imaged two types of microvillus structures in
detail in the mouse ZP, i.e., GC-TZPs and Oo-Mvi. Moreover, we
supply direct evidence showing that oocytes forming Oo-Mvi act
as a cellular antenna to regulate OSF release, therefore maintaining
the proper reproductive lifespan in females.
At the single-cell level, our data showed that each GC in the
inner layer of follicles establishes a high-density GC-TZP network
to connect to oocytes directly. This observation is consistent with
previous views that GC-TZPs support the growth of oocytes2,36.
Meanwhile, our results confirm previous findings that the formation
of GC-TZPs is controlled by the secretion of OSFs (such
Fig. 4 The failure of Oo-Mvi formation in Oo-Rdx−/− females led to abnormal folliculogenesis. a Histological analysis of the ovarian morphological
changes in Oo-Rdx−/− females, showing a decrease in ovarian size and an increase in follicle death (arrows) in the mutant ovaries. b Follicle counting
confirmed a fast follicle loss in Oo-Rdx−/− ovaries. P-value: 0.98 (PD5, n = 3), 0.0015 (PD35, n = 5), 0.0008 (2 M, n = 3), 0.03 (8 M, n = 3). c, d Tracing
the growth of oocytes in cultured ovarian pieces of Oo-Rdx+/+;mTmG and Oo-Rdx−/−;mTmG (c). A significant retardation of oocyte growth in Oo-Rdx−/−;
mTmG (n = 11) ovaries compared to that in Oo-Rdx+/+;mTmG (n = 11) ovaries (d). P-value: 0.015 (day 3), 0.0002 (day 7). e, f Tracing the growth of
ovarian follicles from Oo-Rdx−/− and Oo-Rdx+/+ ovaries in vitro (e), showing significant retardation of the increase in follicle diameter in Oo-Rdx−/− mice
(n = 9, red) compared to that in controls (n = 6, black). P-value: 0.0057 (day 4), 0.014 (day 8) (f). Representative images are shown. Data are presented
as the mean ± SD. Data were analyzed by two-tailed unpaired Student’s t-test and ***P < 0.001, **P < 0.01, *P < 0.05. Scale bars: 500 μm (black) and 100
μm (red) (a), 25 μm (c), and 200 μm (e).
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as GDF9) from oocytes10,36. Therefore, these findings further
support the classical theory that oocytes govern the growth of
follicles in mammals.
Consistent with previous studies that demonstrated that oocytes
also form the communicating structures in different species such as
surf clams37, frogs38, rodent39–41, bovine42–44, and humans45,46,
our results clearly described the oocyte-derived microvilli during
the process of mammalian oogenesis. Compared to GC-TZPs, our
high-resolution living cell observation of Oo-Mvi with endogenousfluorescent
labeling showed the oocyte-derived communicating
structures exhibit several interesting and previously undefined
cellular characteristics, supplying important indications for the
functional role of such Oo-Mvi. First, the Oo-Mvi, at a relatively
low density, is not directly connected to the GC membrane, indicating
that these structures do not act as channels for nutrient
transport. Moreover, the fact that the majority of Oo-Mvi
a
Oo-Rdx+/+
Oo-Rdx-/-
PFSF
PCNA/Hoe
%ofPCNApositiveGCs
0
20
PF SF
40
60
80
100
Oo-Rdx
+/+
Oo-Rdx
-/-
***
**
b
Oo-Rdx-/-
+/+
Oo-Rdx
%ofTUNELpositivefollicles
PF SF
0
20
40
10
30
d
0
50
100
150
200
No.ofTZPs
***
f
Oo-Rdx-/-
+/+
Oo-Rdx
Oo-Rdx+/+
Oo-Rdx-/-
F-actin
e
Oo-Rdx+/+
Oo-Rdx-/-
PFSF
c TUNEL/Hoe
Oo-Rdx+/+
Oo-Rdx-/-
Oo-Rdx-/+
GDF9
Oocytes in cultured ovarian pieces
g
F-actin
0
100
No.ofTZPs
20
40
60
80
***
n.s.
h
Oo-Rdx-/-
+/+
Oo-Rdx
GDF9 +
***
*
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contained vesicle heads and their breakdown indicates an exocrine
function of Oo-Mvi in the regulation of folliculogenesis. Indeed,
our functional study strongly supports the idea that Oo-Mvi act as
a regulating structure since the failure of Oo-Mvi formation leads
to the retardation of oocyte and follicle growth, and accelerated
follicle loss in the ovary. Interestingly, although our results showed
that Oo-Mvi play an important role in folliculogenesis, we found
that the formation of Oo-Mvi is not indispensable for the survival
of oocytes and follicles in vivo. Even though the construction of
Oo-Mvi is destroyed by Rdx deletion, some of the oocytes can still
develop to ovulation in young adulthood. Therefore, the mutant
females are fertile but have a shortened reproductive lifespan, just
like the clinical symptom in patients with premature ovarian
insufficiency47. With our experimental data, we proposed that the
role of Oo-Mvi is an optimized system to select the qualified follicles,
especially the early growing follicles. Meanwhile, although
our results showed that OSFs are released through Oo-Mvi, there is
no clear evidence to show a lack of secreted GDF9 in Oo-Rdx−/−
females. Therefore, we also hypothesized that the release of OSFs
does not completely rely on Oo-Mvi since the folliculogenesis never
passed the secondary stage in Gdf9−/− females17. Also, inconsistency
of the phenotypes between Gdf9−/− and Oo-Rdx−/−
females implied complex underlying mechanisms of either Oo-Mvi
forming regulations or the OSF release controlling on oocytes for
future study. Interestingly, a previous study reported that oocytes
also form membrane fusion with GCs to mediate the secretion of
GDF948, and these observations further support our hypothesis
that Oo-Mvi play a regulating role in controlling folliculogenesis.
Meanwhile, similar structures of Oo-Mvi were reported on the
membrane of cleaved bovine eggs at 2 or 3 cells stage49, implying
that this kind of Mv might be a conserved structure during oocyte
and early embryo development. Therefore, we believed that the
major function of Oo-Mvi are to make the release of OSFs timely
and orderly, guaranteeing a precise and efficient regulation to
determine the developmental fate of ovarian follicles.
Taken together, the observations in the current study identify
an elegant microvillus system in the ZP of growing follicles that
maintains proper folliculogenesis in mice. After the awakening of
dormant follicles, oocytes produce ZP proteins to establish the
egg coat, which is essential for oocyte development and
maturation26. At the same time, RDX-related Oo-Mvi form and
elongate on the surface of oocytes to shorten the physical distance
between oocytes and GCs. With the formation of Oo-Mvi, OSFs
are transported to the tips of Oo-Mvi to construct OSF-rich
vesicles. Although the mechanism by which OSFs are transported
to the vesicles in Oo-Mvi is still unknown, as these vesicles
breakdown, the soluble OSFs are efficiently released to the adjacent
space of the ZP. In response to OSF stimulation, a series of
cellular events take place in GCs, including the increase of proliferation,
the decrease of apoptosis, and the construction of GCTZPs,
which connect with the oocyte membrane to finally support
the robust development of oocytes.
Methods
Mice. C57BL/6 mice were from the Laboratory Animal Center of the Institute of
Genetics (Beijing, China). Gdf9-Cre, Foxl2-CreERT2, and mTmG mice were generated
as previously reported23–25. Foxl2-CreERT2 mice were a gift from Dr. Liu
Kui. The RdxloxP/loxP and Gdf9−/− mice were generated using CRISPR/Cas9mediated
genome engineering by the Model Animal Research Center of Nanjing
University (MARC), Nanjing BioMedical Research Institute of Nanjing University,
Nanjing, China. Generally, exons 4 and 5 of the Rdx gene were selected as the
targeted region to delete in the RdxloxP/loxP mice, and the exon 2 of Gdf9 gene was
deleted in the Gdf9−/− mice. All mutant mouse strains were on the C57BL/6
background.
In the Foxl2-CreERT2;mTmG females, upon a single injection of tamoxifen
(75648, Sigma-Aldrich) at a dosage of 5 μg/kg body weight (BW) at PD8, CreERT2
recombinase-mediated recombination to label a single GC after 1 week50. With
tamoxifen at a high dosage of 20 μg/kg BW for three injections at PD8/10/12,
almost all GCs were labeled after 1 week of treatment25,50.
All mice were housed in mouse facilities under 16/8-h light/dark cycles at 26 °C
and humidity 40–70% with access to chow and water at libitum. The animal
experiments conformed to the guidelines and regulatory standards of the
Institutional Animal Care and Use Committee of China Agricultural University,
No. AW8012020-2-3.
Histological section staining and follicle counting. For morphological analysis
and fluorescent detection at the tissue level, ovarian samples were fixed in 4%
paraformaldehyde (PFA, Santa Cruz, 30525-89-4) for 8 h, embedded in paraffin,
and sectioned serially at 5 μm. To count the follicle number, tissue sections were
stained with hematoxylin (Santa Cruz, sc-24973A). Primordial follicles were
counted in every fifth section and multiplied by five to calculate the number of all
primordial follicles in each ovary. The growing follicles were counted by scanning
all sections and only the follicles with clear oocyte nuclei were counted to exclude
the effect of residual structures after oocyte ablation in the ovaries. The number of
primordial follicles and growing follicles was summed to the total number of
follicles. For fluorescent detection, after deparaffinization, sections were stained
with different antibodies and the Hoechst 33342 (Sigma-Aldrich, 14533) was used
as a counter-stain to check the cell nucleus. The sections were sealed with an antifade
fluorescence mounting medium (Applygen, C1210) by coverslips.
High-resolution imaging of the isolated oocytes to detect the subcellular
structure or protein/factor localization. All high-resolution images of isolated
oocytes were acquired by an Andor Dragonfly spinning-disc confocal microscope
equipped with a ×63 or ×100, 1.44 N.A. oil objective, a scientific complementary
metal-oxide semiconductor (sCMOS) camera (Andor Zyla 4.2), and the 405-nm
(Hoe), 488-nm (mG, Alexa Fluor 488-phalloidin, and LCA-FITC) and 568-nm
(mT, ER-tracker, and rhodamine) lines of the Andor Integrated Laser Engine (ILE)
system with a spinning-disc confocal scan head (Andor Dragonfly 500). Images
were acquired by Fusion 2.1 software (https://andor.oxinst.com/products/
dragonfly#fusion).
To detect the Oo-Mvi in oocytes, denuded oocytes at GV stages (average
diameter around 65 μm) of late secondary or pre-antral follicles were collected by
tearing the ovaries with syringe needle from wild type females or Oo-Rdx−/−;
mTmG females and control. The oocytes were moved with mouth pipette to ~20 μL
minimum MEMα-FBS-ITS medium: essential medium α (MEMα; Gibco, 32-571-
036) with 10% FBS (Gibco, 10-099-141) and 1% insulin-transferrin-selenium (ITS;
Sigma-Aldrich, 13146), covered with mineral oil (Sigma-Aldrich, M8410) and
photographed in a living cell workstation (Okolab) at 37 °C, 5% CO2. Images were
typically acquired at a proper temporal interval (10 min) with an optical slice
thickness of 0.5 μm and covered ~40 μm of oocytes.
To observe the cortex actin of oocytes or the GC-TZPs, denuded oocytes at the
GV stage (average diameter around 65 μm) were collected from fresh ovaries or
GDF9-cultured ovarian pieces of Oo-Rdx+/+ and Oo-Rdx−/− females. Then the
oocytes were fixed with 4% PFA in PBS for 15 min. After washing with PBS, the
fixed oocytes were incubated with Alexa Fluor 488-phalloidin (1:100 dissolved in
MEMα-FBS-ITS medium, ThermoFisher Scientific, A12379) with (to detect oocyte
Fig. 5 Loss of Oo-Mvi disrupted OSF-regulated GC development. a, b PCNA staining (arrows) of GC proliferation (a), showing a significant decrease in GC
proliferation in the early growing follicles of Oo-Rdx−/− ovaries (b). Each point represents the GC-PCNA rate in a single follicle (PF: control n = 105, Oo-Rdx−/−
n = 77, P-value = 0.0000011; SF: control n = 46, Oo-Rdx−/− n = 49, P-value = 0.0072). c, d TUNEL detection (arrows) of apoptosis (c), showing a significantly
increased ratio of follicles with TUNEL-positive GCs in Oo-Rdx−/− ovaries (n = 6). P-value = 0.0007 (PF), 0.02 (SF) (d). e F-actin staining GC-TZPs (arrows)
showing a failure of GC-TZP construction in Oo-Rdx−/− oocytes. f Quantification of GC-TZPs confirmed a dramatically decreased density of GC-TZPs in OoRdx−/−
oocytes (n = 18) compared to that in Oo-Rdx+/+ controls (n = 7). P-value = 0.0000008. g Supplying GDF9 rescued the defect of GC-TZP formation in
Oo-Rdx−/− ovaries. h Quantification of GC-TZPs in Oo-Rdx+/+ (n = 20), Oo-Rdx−/− (n = 19), and Oo-Rdx−/−+GDF9 (n = 21) oocytes, showing the successful
rescue of GC-TZP defect by GDF9 supplementation. P-value: 0.000000027 (Oo-Rdx+/+ vs. Oo-Rdx−/−); 0.21 (Oo-Rdx+/+ vs. Oo-Rdx−/−+GDF9). Data are
presented as the mean ± SD. PF primary follicle, SF secondary follicle. Representative images are shown. Data were analyzed by two-tailed unpaired Student’s ttest
and ***P < 0.001, **P < 0.01, *P < 0.05, n.s. P ≥ 0.05. Scale bars: 25 μm.
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RDX/GDF9 RDX GDF9 Merge
a
R-BSA
15 min 30 min
R-GDF9
0
10
20
30
40
0
20
40
60
80
15 min 30 min 15 min 30 min
Theintensityofoocyte
Thenumberofspots
*** ***
Oo-Rdx+/+
Oo-Rdx-/GCs
GCs
Oo Oo
ZP ZP
GC
Oocyte
OSFs
Gdf9 Bmp15 Fgf8
0
1
2
3
4
5
RelativemRNAlevel
*** ***
**Oo-Rdx
Oo-Rdx
+/+
-/b
c d
e f g
h
ER-trackerOo-Mvi
i
Single Z 3D view Magnification
TheERfluorescenceintensity
0
200
400
600
800
C
ytoplasm
Bubble
***
0
50
100
150
Thenumberofbubbles
ER
O
o-M
vi
n.s. 1000
Fig. 6 OSFs were enriched in Oo-Mvi to manipulate the development of follicles. a GDF9 was localized in the vesicles of Oo-Mvi. Immunostaining
showing GDF9 protein (Red, arrowheads) localized in the oocytes and the vesicles of Oo-Mvi (Green, RDX). n = 29 ovarian follicles. b Rhodamine-labeled
GDF9 (R-GDF9) or BSA (R-BSA) was microinjected into living oocytes at GV stage (n = 15 per group), and the R-GDF9 intensity and distribution were
detected at 15 or 30 min after injections (Supplementary Movies 4 and 5). The R-GDF9 spots surrounding the oocytes were pointed by arrowheads. c, d A
time-dependent decrease of R-GDF9 intensity in oocytes (c) and increase of the number of R-GDF9 spots (d) were observed after injections. P-value:
0.0000014 (c), 0.00064 (d). e ER-tracker staining showed a cloudy ER distribution (top) in the oocyte cytoplasm, and the many ER bubbles (top,
arrowheads) surrounding oocytes (Supplementary Movie 6), which were similar to the R-GDF9 distribution (b, arrowheads) and Oo-Mvi vesicles (bottom,
arrowheads). f Quantification of the number of ER bubbles (n = 17) and Oo-Mvi vesicles (n = 14). P-value = 0.81. g Quantification of the fluorescence
intensity of ER in oocyte cytoplasm and bubbles (n = 20). P-value = 0.00000009. h Relative mRNA levels of Gdf9, Bmp15, and Fgf8 in Oo-Rdx+/+ and OoRdx−/−
oocytes showing a decreased consumption of OSFs in the oocytes of Oo-Rdx−/− females. P-value: 0.000006 (Gdf9), 0.0001 (Bmp15), 0.0019
(Fgf8). i The proposed model of the function of Oo-Mvi in follicle development and female reproduction, showing that Oo-Mvi in the ZP determine the fate
of individual follicles by regulating oocyte-GC communication. Data are presented as the mean ± SD with experiments performed in triplicate, and
representative images are shown. Data were analyzed by two-tailed unpaired Student’s t-test and ***P < 0.001, **P < 0.01, and n.s.P ≥ 0.05. Scale bars: 5
μm (a); 20 μm (b, e).
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cortex) or without (to detect the GC-TZPs) 0.1% Triton X-100 for 30 min to
visualize the GC-TZPs by F-actin staining as previously reported10,51.
To observe the cortical granules of oocytes, Oo-Rdx+/+ and Oo-Rdx−/− oocytes
at the GV stage (average diameter around 65 μm) were fixed in 4% PFA in PBS for
15 min and permeabilized in 0.5% Triton X-100 for 20 min at room temperature.
Then, oocytes were blocked with 1% BSA (Sigma-Aldrich, V900933) in PBS for 1 h
and incubated with LCA-FITC 2 h (1:100 dissolved in PBS, ThermoFisher
Scientific, L32475, a gift from Dr. Bo Xiong in Nanjing Agricultural University,
China) at room temperature30. After washing with PBS, the stained oocytes were
imaged by an Andor Dragonfly spinning-disc confocal microscope as previously
described indexes.
After the acquisition, Movies or single time-point images were processed by
ImageJ (http://rsbweb.nih.gov/ij/) for projection of all z-stacks and merged color
channels. To clearly highlight the structures of the Oo-Mvi, the mG (488 nm)
channel was inverted to black and white by ImageJ software in Fig. 1. To show OoMvi,
R-GDF9, or ER of the oocyte, the rotary 3D Supplementary Movies were
processed by Imaris (https://imaris.oxinst.com/) software (Supplementary
Movies 1, 4, 5, and 6). To show the cellular behaviors of Oo-Mvi on oocytes, the
time-lapse Supplementary Movies were processed by ImageJ software
(Supplementary Movies 2 and 3).
The detailed protocol is also available in Protocol Exchange52.
High-resolution imaging of the ER in oocytes. To observe the ER distribution of
oocytes, denuded living oocytes at GV stage (average diameter around 65 μm) from
fresh wild-type ovaries were incubated with ER-tracker Red (1:1000, Beyotime,
C1041) for 30 min at 37 °C, 5% CO2
32,33. After washing with PBS, the stained
oocytes were imaged by an Andor Dragonfly spinning-disc confocal microscope as
previously described indexes. To the detection of ER fluorescent intensity of oocyte
cytoplasm and bubbles, the single Z of mean ER fluorescent intensity of oocyte
cytoplasm and bubbles were measured by ImageJ software.
Immunofluorescence staining and observation. The ovarian sections were
deparaffinized, rehydrated, and subjected to high-temperature (95–98 °C) antigen
retrieval for 16 min with 0.01% sodium citrate buffer (pH 6.0). The sections were
then blocked with 10% normal donkey serum (Jackson ImmunoResearch, 017-000-
121) for 60 min at room temperature and incubated with primary antibodies
overnight at 4 °C. Subsequently, the sections were incubated with fluorophoreconjugated
donkey secondary antibodies (1:200, Life Technologies) for 2 h at 37 °C.
The homologous IgG (Goat, normal control, A7007, Beyotime) with the primary
antibodies were used to be the negative control to guarantee the specificity of
staining.
To detect ERM family or GDF9 protein expression in the ovary, we incubated
sections with the following primary antibodies: the anti-RDX antibody (rabbit,
1:100, ab52495, Abcam), anti-p-ERM antibody (rabbit, 1:100, mAb#3726, Cell
Signaling Technologies), anti-EZRIN antibody (rabbit, 1:100, ab4069, Abcam),
anti-MOESIN antibody (rabbit, 1:100, ab52490, Abcam), and anti-GDF9 antibody
(goat, 1:50, AF739, R&D). Cell proliferation of GCs was assessed by staining with
the anti-proliferating cell nuclear antigen (PCNA) antibody (mouse, 1:100, Santa
Cruz, sc-25280). The percentage of PCNA-positive GCs was quantified as the
number of GCs with PCNA-positive signals divided by the total number of GCs in
primary follicles and secondary follicles. Cell apoptosis was detected by TUNEL
staining (In Situ Cell Death Detection Kit, Fluorescein, Roche, 11684795910). The
percentage of TUNEL-positive follicles was quantified as the number of follicles
with TUNEL-positive signal divided by the total number of follicles in the primary
and secondary stages.
All general morphologies were examined and photographed by a Nikon Eclipse
Ti digital fluorescence microscope.
For the high-resolution detections of GDF9, the co-localization of GDF9 with
vesicles of Oo-Mvi was acquired as a z-stack model with an optical slice thickness
of 0.3 μm after co-staining of GDF9 and RDX (to label Oo-Mvi), and confocal
sections covered ~8 μm tissue thickness. Both the IgG and the ovarian sections
from Gdf9−/− females were used as the negative control to guarantee the specificity
of staining. The images were acquired by an Andor Dragonfly spinning-disc
confocal microscope equipped with a ×100, 1.44 N.A. oil objective (Leica HC PL
APO), a scientific complementary metal-oxide semiconductor (sCMOS) camera
(Andor Zyla 4.2), and the 488-nm (anti-RDX, Oo-Mvi) and 568-nm (anti-GDF9
and IgG) lines of the Andor Integrated Laser Engine (ILE) system with a spinningdisc
confocal scan head (Andor Dragonfly 500). In detail, images were acquired
with laser 488-nm and laser 568-nm around 10–15%, exposure time 100–200 ms.
Images were acquired by Fusion 2.1 software (https://andor.oxinst.com/products/
dragonfly#fusion). After the acquisition, images were processed by ImageJ (http://
rsbweb.nih.gov/ij/) for projection of all z-stacks and merged color channels.
ZP collection and MS analysis. To collect empty ZP from oocytes, C57BL/6 mice
at PD23 with 5 IU of pregnant mare serum gonadotropin (PMSG; Sansheng
Biological Technology) for 46 h and the ovaries were punctured to obtain cumulusoocyte
complexes. The cumulus cells were digested with 0.3% hyaluronidase
medium (Merck, MR-051-F) to obtain denuded oocytes, which were then incubated
with 10% high osmotic glucose solution to separate the ZP and the oocyte.
Under the microinjection system, the ZP was torn to release the oocyte, and then
the empty ZP was collected. The protein (~5 μg) from ~3000 pieces of ZP was
extracted by RIPA extraction buffer for a single protein MS assay, which was
repeated three times.
Protein digestion was performed using the filter-aided sample preparation
(FASP) method with modifications as previously described53,54. Nanospray
electrospray (ESI)-MS was performed on a Thermo Q-Exactive high-resolution
mass spectrometer (Thermo Scientific) with a 70,000 MS scan resolution, 17,500
MS/MS scan resolution, and top-10 MS/MS selection. Raw data from the mass
spectrometer were preprocessed with Mascot Distiller 2.4 for peak picking. The
resulting peak lists were searched against the Uniport Mouse database using the
Mascot 2.5 search engine. Scaffold PTM was used to evaluate phosphorylation sites
of the Mascot search results using the Ascore algorithm.
Mouse fertility and ovulation analysis. To detect mouse fertility, 5-week-old
female Oo-Rdx−/− mice and control mice were continuously mated with 8-weekold
C57BL/6 fertile males for 40 weeks. The numbers of pups and litters were
recorded, from which fertility rates were determined.
For superovulation, Oo-Rdx−/− and control mice (PD23) were intraperitoneally
injected with 5 IU of PMSG, (Sansheng Biological Technology, Ningbo, China)
followed by 5 IU human chronic gonadotropin (hCG, Sansheng Biological
Technology, Ningbo, China) 46 h later. After an additional 16 h, oocytes were
collected from oviducts, and the numbers of oocytes for each animal were counted
after digestion with 0.3% hyaluronidase (Merck, MR-051-F).
In vitro culture of ovarian cortical pieces. The in vitro attached culture of
ovarian pieces was modified from the method in the previous study55. In details,
ovarian cortical pieces from Oo-Rdx+/+;mTmG and Oo-Rdx−/−;mTmG ovaries at
PD4 (oocyte diameter tracing experiment) were cut into approximately 2 × 1 ×
0.5 mm3 pieces and cultured in MEMα (Gibco, 32-571-036) supplemented with
10% FBS (Gibco, 10-099-141). After 4 days of adherent culture without changing
the medium, the cortical pieces were attached firmly to the bottom of dishes for the
experiments, and then the culture media was half-changed every day to maintain
the development of tissues.
In the experiment of oocyte diameter tracing, the developmental dynamics of
oocytes were measured by the increase of the diameter, which was labeled by mG in
Oo-Rdx+/+;mTmG and Oo-Rdx−/−;mTmG ovaries. The diameter changes were
recorded with a Nikon Eclipse Ti digital fluorescence microscope every other day
for 7 continuous days. To ensure the detected oocytes were in a fast growth period,
the oocytes with an initial diameter of around 16–35 μm were traced in this
experiment. To compare the developmental dynamics of oocytes with different
initial diameter, we normalized the actual diameters to the growth ratio to show the
dynamics change of oocyte growth in different groups. Totally 151 oocytes from 27
cortical pieces in Oo-Rdx+/+;mTmG and 250 oocytes from 21 cortical pieces in
Oo-Rdx−/−;mTmG ovaries were traced. In the GDF9-supplying experiment, OoRdx+/+
and Oo-Rdx−/− ovaries at PD21 (GDF9-supplying experiment) were cut to
approximately 2 × 1 × 0.5 mm3 pieces and cultured in MEMα supplemented with
10% FBS, with or without GDF9 (500 ng/mL, R&D, 739-G9-010/CF) for 24 h, and
the oocytes (average diameter around 65 μm at GV stages) were collected by
tearing the ovaries with a syringe needle and digested with 0.3% hyaluronidase
medium to be used for the detection of GC-TZP density.
In vitro culture of ovarian follicles. Follicles were separated by tearing PD21
ovaries and mechanical isolation with a syringe needle. The follicles with the
diameter of around 130 μm (average diameter: 134.72 ± 13.73 μm in Oo-Rdx+/+,
n = 64 and 129.73 ± 20.21 μm in Oo-Rdx−/−, n = 77) were used in the experiments.
To monitor the growth of follicles, the isolated follicles were cultured in a
Matrigel (BD, 354234) culture system to support a 3D development of follicles,
which was modified from previously described56–58. Generally, the Matrigel was 3:1
diluted with pre-cooling MEMα-FBS-ITS medium-plus follicle-stimulating hormone
(ovine FSH, 10 ng/mL, NHPP) on ice. Add 20 μL Matrigel/microdroplet in
the 6 well culture plate and 6 microdroplets every well on ice keeping the gel in
liquid. Put the 6 well culture plate at 37 °C for 20 min to trans Matrigel from the
liquid phase to solid. Then, the isolated follicles were gently seeded into the gel with
a mouse pipette, respectively. Adding the 2 μL liquid Matrigel to seal the hole and
put the plate into the incubator at 37 °C for 10 min to make the gel sealed. After
this step, the cultured follicles were completely surrounded by the gel. The density
of seeding was around 1.6 per cm2 to guarantee sufficient space for growth. Supplemented
1 mL MEMα-FBS-ITS medium-plus follicle-stimulating hormone
(ovine FSH, 10 ng/mL, NHPP) into the culture plate. During culture, the media was
half-changed every day, and the diameter of follicles was recorded every 24 h for 8
continuous days by a Nikon Eclipse Ti digital fluorescence microscope in a bright
field channel.
The detailed protocol is also available in Protocol Exchange59.
Rhodamine-conjugated GDF9 and living oocyte injection. Recombinant GDF9
protein (50 μg/mL, R&D, 739-G9-010/CF) or BSA (50 μg/mL, Sigma-Aldrich,
V900933) as a control were labeled with rhodamine using the Pierce NHSrhodamine
antibody labeling kit (53031, ThermoFisher) according to the
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manufacturer’s protocol60. To collect the oocytes, the COCs were isolated by
puncturing the antral follicles from C57BL/6 mice at PD23 after 46 h of PMSG
treatment (5 IU, i.p. injection). After treated with 0.3% hyaluronidase (Merck, MR-
051-F) in medium, denuded oocytes were performed microinjection, and
approximately 10 pL of labeled protein or rhodamine solution were injected into
one oocyte with a FemtoJet 4× electric microinjector (Eppendorf). After 15 or 30
min of injections, the oocyte was imaged under an Andor Dragonfly spinning-disc
confocal microscope with the previously described indexes.
Gene expression analysis. To detect gene expression in mouse materials, the
mRNA from different tissues or ovary compartments was extracted by TRIZOL
Reagent (Thermo-Ambion, 15596018) according to the manufacturer’s protocol.
The quantity and quality of the total RNA were determined using a Nanodrop
(Thermo Scientific). Reverse transcription (TAKARA, RR047Q) was performed
using 1 μg total RNA per sample. QRT-PCR reactions were performed in 96-well
plates (Applied Biosystems, 4316813) in 15 μL reaction volumes and analyzed by
an Applied Biosystems 7500 Real-Time PCR System (Applied Biosystems,
4472908) using the following parameters: 10 min at 95 °C, followed by 40 cycles of
15 s at 95 °C and 1 min at 60 °C. Data were normalized to β-actin. The primer list is
provided in Supplementary Table 1.
Western blot. To detect protein expression in mouse materials, total proteins were
extracted using WIP Tissue and Cell lysis solution (BioChip, 110000) according to
the manufacturer’s protocol. Electrophoresis was performed with 50 μg total proteins
separated by 10% SDS-PAGE and transferred to polyvinylidene fluoride
membranes (Millipore, ISEQ00005). The membranes were incubated overnight at
4 °C with the appropriate primary antibodies listed below: anti-RDX (78 kDa,
1:500, ab52495, Abcam), anti-p-ERM (78 kDa, 1:500, mAb#3726, Cell Signaling
Technologies). The appropriate secondary antibody (ZB-2301, ZB-2305 from
ZSGB-BIO) was diluted 1:5000 in TBST (TBS plus 0.5% Tween 20). Tubulin
(1:5000, Beyotime, AF5012) was used as an internal control. The membranes were
visualized using the SuperSignal chemiluminescent detection system (Thermo
Scientific, 32109). Original blots can be found in the Source data file.
Statistical analysis. All experiments were repeated at least three times using
different mice. Data are presented as the mean ± standard deviation (SD) of each
experiment. Data were analyzed by Student’s t-test and were considered statistically
significant at P < 0.05. P is indicated as follows: *(P < 0.05), **(P < 0.01), ***(P <
0.001), and n.s.(not significant, P ≥ 0.05). Statistics and graphs were obtained using
Prism 5 (GraphPad Software, La Jolla).
Data availability
The mass spectrometry proteomics data have been deposited in the ProteomeXchange
Consortium via the partner repository iProx61 under the data set identifier
IPX0002846000 for Fig. 2b and Supplementary Fig. 2 associated raw data (https://www.
iprox.org/page/project.html?id=IPX0002846000). This is also found at the Proteome
Xchange: see http://proteomecentral.proteomexchange.org/cgi/GetDataset?ID=
PXD024931. All data are available from the corresponding author upon reasonable
request. Source data are provided with this paper.
Received: 15 July 2020; Accepted: 31 March 2021;
References
1. Li, L., Zheng, P. & Dean, J. Maternal control of early mouse development.
Development 137, 859–870 (2010).
2. Matzuk, M. M., Burns, K. H., Viveiros, M. M. & Eppig, J. J. Intercellular
communication in the mammalian ovary: oocytes carry the conversation.
Science 296, 2178–2180 (2002).
3. Hsueh, A., Kawamura, K., Cheng, Y. & Fauser, B. Intraovarian control of early
folliculogenesis. Endocr. Rev. 36, er20141020 (2014).
4. Eppig, J. J. Oocyte control of ovarian follicular development and function in
mammals. Reproduction 122, 829–838 (2001).
5. Dietl, J. The Mammalian Egg Coat: Structure and Function 1st edn (1989).
6. Wassarman, P. M. & Litscher, E. S. Biogenesis of the mouse egg’s extracellular
coat, the zona pellucida. Curr. Top. Dev. Biol. 102, 243–266 (2013).
7. Li, R. & Albertini, D. F. The road to maturation: somatic cell interaction and
self-organization of the mammalian oocyte. Nat. Rev. Mol. Cell Biol. 14,
141–152 (2013).
8. Albertini, D. F. & Rider, V. Patterns of intercellular connectivity in the
mammalian cumulus-oocyte complex. Microsc. Res. Tech. 27, 125–133 (1994).
9. Schroeder, T. E. Microfilament-mediated surface change in starfish oocytes in
response to 1-methyladenine: implications for identifying the pathway and
receptor sites for maturation-inducing hormones. J. Cell Biol. 90, 362–371
(1981).
10. El-Hayek, S., Yang, Q., Abbassi, L., FitzHarris, G. & Clarke, H. J. Mammalian
oocytes locally remodel follicular architecture to provide the foundation for
germline-soma communication. Curr. Biol. 28, 1124–1131 (2018).
11. Chastant-Maillard, S. et al. The canine oocyte: uncommon features of in vivo
and in vitro maturation. Reprod. Fert. Dev. 23, 391–402 (2011).
12. Runge, K. E. et al. Oocyte CD9 is enriched on the microvillar membrane and
required for normal microvillar shape and distribution. Dev. Biol. 304,
317–325 (2007).
13. Bernareggi, A. et al. On the mechanism of the electrophysiological changes
and membrane lesions induced by asbestos fiber exposure in Xenopus laevis
oocytes. Sci. Rep. 9, 2014 (2019).
14. Burgess, D. & Schroeder, T. E. Polarized bundles of actin filaments within
microvilli of fertilized sea urchin eggs. J. Cell Biol. 74, 1032–1037 (1977).
15. Makabe, S., Naguro, T. & Stallone, T. Oocyte–follicle cell interactions during
ovarian follicle development, as seen by high resolution scanning and
transmission electron microscopy in humans. Microsc. Res. Tech. 69, 436–449
(2006).
16. Zhang, H. & Liu, K. Cellular and molecular regulation of the activation of
mammalian primordial follicles: Somatic cells initiate follicle activation in
adulthood. Hum. Reprod. 10, 887–895 (2015).
17. Dong, J. et al. Growth differentiation factor-9 is required during early ovarian
folliculogenesis. Nature 383, 531–535 (1996).
18. Peng, J. et al. Growth differentiation factor 9:bone morphogenetic protein 15
heterodimers are potent regulators of ovarian functions. Proc. Natl Acad. Sci.
USA 110, 776–785 (2013).
19. Juengel, J. L. & McNatty, K. P. The role of proteins of the transforming growth
factor-β superfamily in the intraovarian regulation of follicular development.
Hum. Reprod. Update 11, 144–161 (2005).
20. Yonemura, S. & Tsukita, S. Direct involvement of Ezrin/Radixin/Moesin
(ERM)-binding membrane proteins in the organization of microvilli in
collaboration with activated ERM proteins. J. Cell Biol. 145, 1497–1509 (1999).
21. Hoeflich, K. P. & Ikura, M. Radixin: cytoskeletal adopter and signaling
protein. Int. J. Biochem. Cell 36, 2131–2136 (2004).
22. Clarke, H. J. Regulation of germ cell development by intercellular signaling in
the mammalian ovarian follicle. Wires Dev. Biol. 7, e294 (2017).
23. Muzumdar, M. D., Tasic, B., Miyamichi, K., Li, L. & Luo, L. A global doublefluorescent
Cre reporter mouse. Genesis 45, 593–605 (2010).
24. Lan, Z.-J., Xu, X. & Cooney, A. J. Differential oocyte-specific expression of Cre
recombinase activity in GDF-9-iCre, Zp3cre, and Msx2Cre transgenic mice.
Biol. Reprod. 71, 1469–1474 (2004).
25. Zheng, W. et al. Two classes of ovarian primordial follicles exhibit distinct
developmental dynamics and physiological functions. Hum. Mol. Genet. 23,
920–928 (2014).
26. Bleil, J. D. & Wassarman, P. M. Structure and function of the zona pellucida:
Identification and characterization of the proteins of the mouse oocyte’s zona
pellucida. Dev. Biol. 76, 185–202 (1980).
27. Sommersberg, B. Gap junction communication and connexin 43 gene
expression in a rat granulosa cell line: regulation by follicle-stimulating
hormone. Biol. Reprod. 63, 1661–1668 (2000).
28. Sauvanet, C., Wayt, J., Pelaseyed, T. & Bretscher, A. Structure, regulation, and
functional diversity of microvilli on the apical domain of epithelial cells. Annu.
Rev. Cell Dev. Biol. 31, 593–621 (2015).
29. Chaigne, A. et al. A soft cortex is essential for asymmetric spindle positioning
in mouse oocytes. Nat. Cell Biol. 15, 958–966 (2013).
30. Xiong, B., Zhao, Y. G., Beall, S., Sadusky, A. B. & Dean, J. A unique egg
cortical granule localization motif is required for ovastacin sequestration to
prevent premature ZP2 cleavage and ensure female fertility in mice. PLoS
Genet. 13, e1006580 (2017).
31. Rapoport, T. Transport of proteins across the endoplasmic reticulum
membrane. Science 258, 931–936 (1992).
32. FitzHarris, G., Marangos, P. & Carroll, J. Changes in endoplasmic reticulum
structure during mouse oocyte maturation are controlled by the cytoskeleton
and cytoplasmic dynein. Dev. Biol. 305, 133–144 (2007).
33. Ma, R. et al. Involvement of Rab6a in organelle rearrangement and
cytoskeletal organization during mouse oocyte maturation. Sci. Rep. 6, 23560
(2016).
34. Valve, E., Penttil, T.-L., Paranko, J. & Harkonen, P. FGF-8 is expressed during
specific phases of rodent oocyte and spermatogonium. Development 232,
173–177 (1997).
35. Pan, H., O’Brien, M. J., Wigglesworth, K., Eppig, J. J. & Schultz, R. M.
Transcript profiling during mouse oocyte development and the effect of
gonadotropin priming and development in vitro. Dev. Biol. 286, 493–506
(2005).
36. Combelles, C. M., Carabatsos, M. J., Kumar, T. R., Matzuk, M. M. & Albertini,
D. F. Hormonal control of somatic cell oocyte interactions during ovarian
follicle development. Mol. Reprod. Dev. 69, 347–355 (2004).
NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-021-22829-2 ARTICLE
NATURE COMMUNICATIONS | (2021)12:2523 | https://doi.org/10.1038/s41467-021-22829-2 | www.nature.com/naturecommunications 13
37. Rebhun, L. L. Electron microscope studies on the vitelline membrane of the
surf clam Spisula solidissima. J. Ultra Res. 6, 107–122 (1962).
38. Courjaret, R. et al. The Ca2+-activated Cl- channel Ano1 controls microvilli
length and membrane surface area in the oocyte. J. Cell Sci. 129, 2548–2558
(2016).
39. Trujillo-Cenóz, O. & Sotelo, J. Relationships of the ovular surface with follicle
cells and origin of the zona pellucida in rabbit oocytes. J. Biophys. Biochem.
Cytol. 5, 347–350 (1959).
40. Benammar, A., Ziyyat, A., Lefèvre, B. & Wolf, J.-P. Tetraspanins and mouse
oocyte microvilli related to fertilizing ability. Reprod. Sci. 24, 1062–1069 (2016).
41. Sezen, S. C. & Cincik, M. Oocyte membrane maturation and oocyte-sperm
relationship: transmission electron microscopy study. Arch. Androl. 49,
297–305 (2003).
42. Makita, M. & Miyano, T. Steroid hormones promote bovine oocyte growth
and connection with granulosa cells. Theriogenology 82, 605–612 (2014).
43. Hyttel, P., Fair, T., Callesen, H. & Greve, T. Oocyte growth, capacitation and
final maturation in cattle. Theriogenology 47, 23–32 (1997).
44. Fair, T., Hulshof, S. C. J., Hyttel, P., Greve, T. & Boland, M. Oocyte
ultrastructure in bovine primordial to early tertiary follicles. Anat. Embryol.
195, 327–336 (1997).
45. Dale, B., Tosti, E. & Iaccarino, M. Is the plasma membrane of the human oocyte
reorganised following fertilisation and early cleavage? Zygote 3, 31–36 (1995).
46. Luigia, S., Mina, A., Beth, E. T., Jacques, C. & Brian, D. Is the human oocyte
plasma membrane polarized? Hum. Reprod. 7, 999–1003 (1992).
47. Qin, Y., Xue, J., Leigh, S. J. & Chen, Z.-J. Genetics of primary ovarian
insufficiency: new developments and opportunities. Hum. Reprod. Update 6,
787–808 (2015).
48. Komatsu, K. & Masubuchi, S. Mouse oocytes connect with granulosa cells by
fusing with cell membranes and form a large complex during follicle
development. Biol. Reprod. 99, 527–535 (2018).
49. Suzuki, H., Ju, J.-C., Parks, J. E. & Yang, X. Surface ultrastructural
characteristics of bovine oocytes following heat shock. J. Reprod. Dev. 44,
345–351 (1998).
50. Hayashi, S. & McMahon, A. P. Efficient recombination in diverse tissues by a
tamoxifen-inducible form of Cre: a tool for temporally regulated gene
activation/inactivation in the mouse. Dev. Biol. 244, 305–318 (2002).
51. Schuh, M. & Ellenberg, J. A new model for asymmetric spindle positioning in
mouse oocytes. Curr. Biol. 18, 1986–1992 (2008).
52. Zhang, Y., Wang, Y., & Zhang, H. Imaging the communicating structures on
oocyte through high-resolution spinning-disc microscope. Protoc. Exch.
https://doi.org/10.21203/rs.3.pex-1381/v1 (2021).
53. Wiśniewski, J. R., Zougman, A., Nagaraj, N. & Mann, M. Universal sample
preparation method for proteome analysis. Nat. Methods 6, 359–362 (2009).
54. Zhao, X., Bai, X., Jiang, C. & Li, Z. Phosphoproteomic analysis of two
contrasting maize inbred lines provides insights into the mechanism of saltstress
tolerance. Int. J. Mol. Sci. 20, 1886 (2019).
55. Kouji, K., Akira, I., Tomohiko, M. & Satoru, M. Ovarian tissue culture to
visualize phenomena in mouse ovary. J. Vis. Exp. 19, 57794 (2018).
56. Adriaens, I., Cortvrindt, R. & Smitz, J. Differential FSH exposure in preantral
follicle culture has marked effects on folliculogenesis and oocyte
developmental competence. Hum. Reprod. 19, 398–408 (2004).
57. Higuchi, C. M., Maeda, Y., Horiuchi, T. & Yamazaki, Y. A simplified method
for three-dimensional (3-D) ovarian tissue culture yielding oocytes competent
to produce full-term offspring in mice. PLoS ONE 10, e0143114 (2015).
58. Green, L. J. & Shikanov, A. In vitro culture methods of preantral follicles.
Theriogenology 86, 229–238 (2016).
59. Zhang, Y., Feng, X.-A., Zhang, S., & Zhang, H. Tracing ovarian follicles
growth in vitro by three-dimensional culture system. Protoc. Exch. (2021).
60. Goding, J. W. Conjugation of antibodies with fluorochromes: modifications to
the standard methods. J. Immunol. Methods 13, 215–226 (1976).
61. Ma, J. et al. iProX: an integrated proteome resource. Nucleic Acids Res. 47,
D1211–D1217 (2019).
Acknowledgements
We are grateful to Xin Li for drawing the illustrations in the manuscript and Dr. Lei Li,
Dr. Bo Xiong, and Dr. Youqiang Su for kindly suggestions. This study was supported by
the National Key Research and Development Program of China to Y.Z. and H.Z.
(2018YFC1003800; 2018YFC1003700; 2017YFC1001100) and the National Natural Science
Foundation of China (81873815 and 31571542).
Author contributions
Y.Z., Y.W., G.X., and H.Z. designed the research; Y.Z., Y.W., X.F., S.Z., X.X., S.N., L.L.,
and Y.B. performed the experiments; Y.Z., Y.W., C.W., Z.L., and H.Z. analyzed the data;
Y.Z., Y.W., and H.Z. wrote the paper. All authors have seen and approved the final
version.
Competing interests
The authors declare no competing interests.
Additional information
Supplementary information The online version contains supplementary material
available at https://doi.org/10.1038/s41467-021-22829-2.
Correspondence and requests for materials should be addressed to H.Z.
Peer review information Nature Communications thanks Juliano Da Silveira and the
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