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Department of Obstetrics and Gynecology, Washington University School of Medicine, St. Louis, Missouri 63110
Address all correspondence and requests for reprints to: Lisa M. Olson, Ph.D., Department of Obstetrics and Gynecology, Box 8064, Washington University School of Medicine, 4911 Barnes Hospital Plaza, St. Louis, Missouri 63110. E-mail: olson_l{at}kids.wustl.edu
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Introduction |
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NO has been shown to mediate its actions by binding to metal-containing centers of enzymes and regulating their activities in both positive and negative ways. For example, NO is a well recognized activator of guanylate cyclase, leading to an increase in cGMP levels in target cells (8, 9). In addition, NO has been shown to directly activate the mitogen-activated protein kinase family (12). In contrast, NO inhibits the actions of the P450 enzymes, including the P450 aromatase (13, 14, 15). NO is synthesized by rat (16, 17) and human (13, 18) ovarian cells and has been shown to influence ovulation, steroidogenesis, and apoptotic cell death of follicular cells (as a follicle survival factor) (2, 5, 6, 17, 19, 20). Recent studies have suggested that NO plays an important function during the periovulatory period. Ovarian nitrite levels increase during the periovulatory period in rodents, and inhibition of NOS with pharmacological inhibitors decreases the number of ovulated oocytes both in perfused ovaries and in vivo (1, 4, 5, 21).
Studies in our laboratory have demonstrated that the rat ovary expresses both eNOS and iNOS in a cell- and development stage-specific manner (6). Furthermore, we have shown specific eNOS staining on the surface of the oocytes in rat ovaries. We have also observed that rats treated with NOS inhibitors not only showed a lower ovulation rate, but also had abnormal oocyte meiotic maturation. Specifically, inhibition of NOS/NO resulted in a significantly lower percentage of mature oocytes in metaphase II and a greater percentage of atypical oocytes (21).
Numerous cell factors have been implicated in regulating meiosis in mammals. The resumption of meiosis, indicated by germinal vesicle (GV) breakdown (GVBD) and oocyte maturation, characterized by progression of the dictyate chromosomes through the first meiotic cycle, extrusion of the first polar body, and formation of metaphase II (22), are controlled by several categories of molecules. These include intracellular messengers, such as cAMP (23, 24, 25), calmodulin, calcium, and diacylglycerol (24) as well as peptides (22), steroids (26), purines (27), and a meiosis-arresting component in the oocyte membrane (24). Preceding ovulation, resumption of meiosis occurs in response to the preovulatory surge of gonadotropins (28). Entry and exit from metaphase are controlled by changes in the activity of maturation-promoting factor (29), which has been described in mitotic cells (30), as well as in oocytes from different species (29, 31). Results from our studies suggest that NO also plays an important role in oocyte meiosis. As the ovary expresses two isoforms of NOS, it is impossible to distinguish isoform-specific ovarian functions using pharmacological inhibitors, as these drugs are not isoform specific. The eNOS knock-out (eNOS-deficient) mice have provided an opportunity to further explore the role of NO in ovulation and oocyte meiotic maturation.
In the present study, we examined the specific role of eNOS-derived NO in ovarian function using eNOS knock-out mice. We found that eNOS knock-out breeding pairs had significantly fewer pups than heterozygote or wild-type (WT) breeding pairs. This lower fecundity supports the results of a superovulation study in which immature eNOS knock-out females had a reduced ovulatory efficiency relative to that of WT females. Furthermore, ovulated oocytes obtained from eNOS knock-out females had significant abnormalities in meiotic maturation and increased oocyte cell death relative to oocytes obtained from WT females. These studies indicate that follicular/oocyte NO is a key regulator of oocyte meiotic maturation.
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Materials and Methods |
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Pups were weaned at 21 days of age, at which time all pups were sexed. Female and male pups from heterozygote matings were ear notched for identification purposes, and a 1-cm segment of the tail (not including the keratinized tip) was removed and prepared for isolation of genomic DNA. The genotype of the offspring was analyzed by Southern blotting. For studies of the reproductive capability of the eNOS knock-out mice, we compared the litter size and the percent mortality of pups born to WT, eNOS knock-out, and heterozygote females.
Southern blot analysis
Genomic DNA was isolated from tails using the QIAamp Tissue Kit
(Qiagen, Santa Clarita, CA) and was digested with Spe1 (5
IU; Boehringer Mannheim, Indianapolis, IN). Equal amounts of DNA (15
µg) were electrophoresed on 1% agarose gels, denatured in 0.2
M HCl for 15 min, renatured by soaking in 0.4 N
NaOH, and subsequently transferred overnight onto GeneScreen membranes
(New England Nuclear, Boston, MA) using 0.4 N NaOH (33).
The membranes were prehybridized at 42 C in a hybridization buffer
containing 50% formamide (Boehringer Mannheim), 10% dextran sulfate
(Fisher Scientific, Fairlawn, NJ), 50 mM sodium phosphate
(Fisher), 5 x SSC (0.75 M sodium chloride and 0.075
M sodium citrate, pH 7.0; Fisher), 5 x Denhardt’s
solution [1 g/ml Ficoll 400 (Sigma Chemical Co., St. Louis, MO), 1
g/ml polyvinylpyrrolidone (Fisher), 1 g/ml BSA (fraction V; Sigma), and
100 µg/ml denatured salmon sperm DNA (Sigma)]. After 4 h, the
hybridization buffer was replaced with fresh hybridization buffer
containing 106 cpm/ml 32P-labeled eNOS genomic
DNA that had been previously labeled using [32P]deoxy
(d)-CTP (Amersham Life Science, Arlington Heights, IL) and the High
Prime DNA labeling kit (Boehringer Mannheim). After hybridization for
16 h at 42 C, membranes were washed sequentially twice at 42 C for
15 min each time with 2 x SSC and 0.1% SDS (Sigma) followed by
twice at 42 C with 0.2 x SSC and 0.1% SDS, and lastly twice at
65 C with 0.1 x SSC and 0.1% SDS. The membranes were allowed to
air-dry and then were wrapped with saran wrap, and the signals were
visualized by exposure to a phosphorimager screen. Visual inspection of
the phosphorimage determined which mice were wild-type, heterozygote,
and homozygous knock-out mice. Wild-type and homozygous eNOS knock-out
littermates were subsequently used for our experiments.
Superovulation and tissue collection
Immature (27 days old) wild-type (control) and eNOS knock-out
female mice were superovulated by injecting 5 IU PMSG (Calbiochem, La
Jolla, CA) ip followed 48 h later by an injection of 5 IU hCG
(Profasi) (6).
To determine whether mouse ovarian oocytes express eNOS, mice were weighed and killed before PMSG injection (unstimulated, control) and at 0 h (48 h after PMSG; preovulatory ovaries) and 8 h (ovulatory ovaries) after hCG administration (n = 3/genotype·time point). For each mouse, both ovaries were removed and frozen in Tissue-Tek OCT embedding compound (Miles, Elkhart, IN) using liquid nitrogen. The ovaries were stored at -70 C until immunohistochemistry was performed. To determine the expression of eNOS on ovulated oocytes, WT and eNOS-deficient mice were weighed and killed 16 h after hCG injection (n = 3/genotype). For each mouse, both ovaries were weighed, and oocytes collected from each ampulla were stained using immunofluorescent methods.
To examine whether eNOS deficiency affects ovarian weight, ovulation, oocyte meiotic maturation, and steroid concentration, mice were weighed and killed 16 h after hCG administration. Blood samples were collected by cardiac puncture, and plasma from these samples was stored at -20 C until assayed for progesterone and estradiol concentrations. For each mouse, both ovaries with oviducts were removed into prewarmed (37 C) Dulbecco’s PBS (DPBS; Sigma). Ovaries were weighed, and ovulated oocytes were collected from each ampulla.
To determine whether the effect of eNOS deficiency on ovarian weight was specific, we also weighed spleen, liver, kidney, and adrenal gland. All animals were killed by CO2 asphyxiation, followed by cervical dislocation.
Immunofluorescent localization of eNOS in ovarian and ovulated
oocytes
Staining of ovarian oocytes and ovulated oocytes was performed
using a polyclonal antiserum and immunofluorescent detection methods,
as previously described (6). Briefly, ovarian cryosections and ovulated
oocytes (denuded from cumulus cells) from WT and eNOS knock-out mice
were mounted on poly-L-lysine-coated slides and fixed in
Carnoy’s solution (ethanol-chloroform-glacial acetic acid, 6:3:1). All
slides were incubated with eNOS polyclonal antiserum (a gift from
Monsanto/Searle, St. Louis, MO) at a 1:100 dilution. Negative control
slides were incubated in normal rabbit serum (Vector Laboratories,
Burlingame, CA) in place of the primary antibody. Ovarian sections and
ovulated oocytes were counterstained with propidium iodide (PI;
Boehringer Mannheim) to visualize all cellular nuclei or oocyte
chromosomes, respectively.
For ovaries, the staining procedure was conducted on several sections for the primary antibody, and three sections were used for a negative control staining (with normal rabbit serum) from each ovary for each mouse. Ovarian sections from eNOS knock-out mice served as an additional negative control. For ovulated oocytes, 80 oocytes collected from WT mice and 30 oocytes from eNOS knock-out mice were stained with eNOS antiserum. Thirty oocytes collected from WT mice were incubated with normal rabbit serum.
For ovarian and ovulated oocytes, incident light fluorescence was monitored with a Nikon light microscope (Nikon Corp., Melville, NY) equipped with a mercury lamp and a dual wavelength filter (Boyce Scientific, St. Louis, MO) to permit the simultaneous detection of green and red fluorescence. In addition, cytoplasm of ovulated oocytes was analyzed with a laser scanning confocal fluorescent microscope (Carl Zeiss, Thornwood, NY). Oocytes were sampled by collecting serial images, spaced 2 µm apart, through the full thickness of the oocyte.
Ovulation efficiency and oocyte classification
The enlarged and translucent ampulla of each oviduct was excised
and trimmed, and the ovulated oocyte-cumulus complexes were recovered
into a petri dish with DPBS. Cumulus cells were removed with
hyaluronidase (0.25 mg/ml; type III; Sigma) diluted in DPBS and by
gently mouth-pipetting the oocytes through a glass micropipette. The
denuded oocytes were pipetted through three wash drops (200 µl each)
of DPBS supplemented with 5 mg/ml BSA (fraction V) covered with mineral
oil (Sigma). After washes, oocytes from each mouse were pooled and
placed in a final droplet of 200 µl DPBS with BSA covered with
mineral oil. Oocytes were counted, and data are expressed as the mean
number of ovulated oocytes per mouse/genotype.
To determine whether lack of NO synthesis influences oocyte meiotic maturation, all ovulated oocytes were carefully examined with an inverted microscope and classified as being at the following stages: 1) metaphase I, healthy oocytes showing GVBD or at metaphase I (the outline of the GV had disappeared, but no polar body had been released); 2) metaphase II, healthy oocytes with a polar body; and 3) atypical oocyte showing degenerative changes or atypical morphology: cytoplasmic fragmentation, loss of spherical shape, etc. The percentage of oocytes in each stage of meiosis was calculated by dividing the number of oocytes at each stage by the total number of ovulated oocytes for each animal.
To further confirm the nuclear maturation of oocytes, all oocytes were first examined for stage of meiosis and then incubated with Giemsa solution to stain chromosomes (34). Briefly, recovered oocytes were placed in 1% sodium citrate for 5–10 min at 25 C. With the help of a mouth-controlled pipette, a microdrop of sodium citrate together with an oocyte(s) was placed on the glass slide, spread by the dropwise addition of freshly prepared acetic alcohol (3 parts of absolute ethyl alcohol to 1 part of glacial acetic acid), and dried by gentle blowing. The dried spreads were fixed in acetic alcohol for 5 min and stained with 10% Giemsa stain (Sigma) diluted in PBS (pH 7.4) for 30 min. After staining, oocytes were gently rinsed with distilled water, allowed to air-dry, and mounted with Permount. Each oocyte was analyzed for the chromosome arrangement characteristic of each stage of meiosis.
Terminal Deoxynucleotidyl Transferase (TdT)-Mediated dUTP Nick End
Labeling (TUNEL) staining of ovulated oocytes
To determine whether the abnormal ovulated oocytes were dying,
TUNEL was performed using the In Situ Cell Death Detection
Kit, POD (Boehringer Mannheim). Ovulated oocytes collected from WT and
eNOS-deficient mice (n = 5 mice/genotype) were denuded from
cumulus cells as described above, mounted on
poly-L-lysine-coated slides, and fixed with 4%
paraformaldehyde (in PBS, pH 7.4) for 30 min at 25 C. Slides were
rinsed with PBS and treated for 30 min with 0.3% hydrogen peroxide in
methanol (Fisher). After rinsing, the slides were incubated in
permeabilization solution [0.1% Triton X-100 (Sigma) in 0.1% sodium
citrate] for 2 min on ice. After washes in PBS, slides were incubated
(60 min, 37 C) with TdT enzyme in the presence of fluorescein
isothiocyanate-conjugated dUTP (TUNEL reaction mixture). Oocytes were
counterstained with PI (1:400 in PBS) to visualize chromosomes.
Negative controls, which were included in each staining replicate,
consisted of incubating slides in TUNEL reaction mixture without TdT
enzyme. For positive controls, oocytes were fixed and first
permeabilized with deoxyribonuclease I (DNase I; 1 mg/ml; Boehringer
Mannheim) for 10 min to induce DNA strand breaks followed by the TUNEL
method.
Progesterone and estradiol RIAs
Progesterone and estradiol concentrations were determined in
plasma using commercial RIA kits [Diagnostic Products Corp. (Los
Angeles, CA) for progesterone and Abbott Laboratories (Abbott Park, IL)
for estradiol]. Steroids were determined in plasma from WT and eNOS
knock-out mice killed 16 h after hCG administration. Samples were
analyzed singly or in duplicate when sufficient plasma was available.
The inter- and intraassay coefficients of variation for the
progesterone assay were 5.1% and 2.6%, and those for the estradiol
assay were 4.5% and 4.1%, respectively.
Statistical analysis
All data are presented as the mean ± SEM.
Significant differences between wild-type and eNOS knock-out mice were
determined for various parameters using independent t test.
For comparison of three groups (wild-type, heterozygous, and eNOS
knock-out mice in the breeding studies), data were evaluated by ANOVA,
and multiple comparisons were made with Fisher’s least significant
procedures (35).
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Results |
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). The intact eNOS gene identified in WT
mice shows a 7-kb hybridizing fragment, whereas the disrupted eNOS gene
shows a 4-kb hybridizing fragment (Fig. 1). Analysis of the genotype
(females and males) of offspring (n = 280 pups) from heterozygote
breeding pairs revealed that 23.4% were WT, 59.3% were heterozygote,
and 17.3% were eNOS knock-out mice. The overall distribution deviates
somewhat (P < 0.05) from a 1:2:1 Mendelian
distribution, with the major deviation due to the lack of eNOS
knock-out pups.
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). Furthermore, eNOS knock-out females
delivered not only fewer pups relative to WT females but these pups
also had higher (P < 0.05) initial mortality rates
(number of pups at birth vs. number of weaned pups) than
pups born to either heterozygote or WT females (42%, 15%, and 5%
mortality, respectively; Table 1). In addition, the sex distribution
(ratio of females to males) within each litter of eNOS knock-out
females was lower than that in heterozygote or WT females (Table 1).
These results show that although eNOS-deficient females can reproduce,
homozygous eNOS knock-out breeding pairs show a significant reduction
(by 60%) in reproductive performance relative to either heterozygote
or WT breeding pairs.
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, for 48 h post-PMSG treatment). We also observed punctuate staining
for eNOS in granulosa cells however, the intensity of staining was
weaker than that in other follicular cells. PI counterstaining
identified all cellular nuclei, as indicated by red fluorescence. This
pattern of staining was observed in ovaries at all three time points
tested: before PMSG injection (unstimulated; data not presented),
48 h after PMSG, and 8 h after hCG injection (data not
presented). All negative control slides stained with normal rabbit
serum instead of primary antiserum were negative (data not presented).
Analysis of ovaries from eNOS knock-out mice, which should not be
expressing any eNOS, showed a complete lack of immunofluorescent
staining. Figure 2B shows representative staining for eNOS in an
ovarian section obtained from an eNOS knock-out mouse 48 h after
PMSG administration.
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. This oocyte exhibits positive green
immunofluorescent staining for eNOS in the cytoplasm (Fig. 2D). No
positive staining was observed when the primary antibody was omitted
(Fig. 2E) or with ovulated oocytes collected from eNOS knock-out mice
(Fig. 2F). These data demonstrate that eNOS is expressed on the surface of oocytes in ovaries from WT mice, but not on oocytes from eNOS knock-out mice. Furthermore, oocytes from WT mice retain their expression of eNOS following ovulation.
Effects of eNOS deficiency on ovarian weight, ovulation efficiency,
and steroidogenesis
Ovarian function of eNOS knock-out female mice was assayed by
determining the ovary’s response to superovulation with exogenous
gonadotropins. Ovaries obtained from eNOS knock-out female mice were
significantly smaller (P < 0.001) than those from WT
females even though body weights were not different in the two groups
(Table 2). Furthermore, the ratio of
ovarian weight to body weight shows that ovaries from eNOS knock-out
females were approximately half the size (P < 0.0001)
of ovaries from WT females after superovulation (Table 2). There was no
influence of eNOS deficiency on spleen, liver, kidney, and adrenal
gland weights, indicating the specificity of the effect of eNOS
deficiency on the ovary (Table 2).
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). The mean number of ovulated
oocytes per mouse was 16.70 ± 2.00 for eNOS knock-out females and
44.50 ± 2.10 for WT females (Table 3). Overall, the ovulation
rate was reduced by 63% in eNOS-deficient mice relative to that in WT
mice. These results indicate that NO is necessary for optimum ovulation
in rodents.
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). In contrast, the estradiol level was significantly
higher (P < 0.01) in plasma collected from
eNOS-deficient mice than in that from WT mice (47.41 ± 9.29
vs. 24.49 ± 1.79 pg/ml; Table 3). These data show that
estradiol levels are elevated in eNOS knock-out mice during the
ovulatory period and suggest that NO functions to negatively regulate
estradiol synthesis and/or secretion.
Abnormal oocyte meiotic maturation in eNOS-deficient mice
After ovulation, the majority of oocytes in the ampulla have
released their first polar body and are arrested at metaphase II (36).
During the course of the studies examining the effect of eNOS
deficiency on ovulation, we observed that the lack of eNOS affected not
only the ovulation rate but also oocyte meiotic maturation. For this
reason, freshly isolated unstained ovulated oocytes obtained from eNOS
knock-out and WT mice were carefully examined and classified as being
at the following stages of meiosis: GVBD/metaphase I, metaphase II, or
atypical, showing evidence of degeneration (Fig. 3). We did not observe any ovulated
oocytes at the GV stage of meiosis regardless of genotype, indicating
that the resumption of meiosis involving GVBD was unaffected in
eNOS-deficient mice. Although, oocytes collected from eNOS knock-out
mice resumed meiosis normally, they showed a pattern of abnormal
meiotic maturation at the later stages of meiosis. Namely, fewer
oocytes (P < 0.001) from eNOS knock-out mice achieved
metaphase II of meiosis (14.4 ± 1.1%) compared with those from
WT mice (68.1 ± 0.6%; Fig. 3). Furthermore, a greater percentage
(P < 0.002) of oocytes from eNOS knock-out mice
remained in metaphase I (46.3 ± 0.3%) or were atypical
(39.3 ± 0.8%) relative to those in WT mice (27.7 ± 0.7%
and 4.2 ± 1.2%, respectively; Fig. 3).
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). The
classification of ovulated oocytes presented above was confirmed by
staining of oocytes obtained from eNOS knock-out mice with Giemsa
solution to visualize chromosomes. Figure 5 shows an oocyte at metaphase II with
the first polar body (Fig. 5A), oocytes still remaining at metaphase I
of meiosis (Fig. 5, B–D), and an oocyte at metaphase I but with
disorganized chromosomes (Fig. 5C).
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Cell death in ovulated oocytes
As oocytes obtained from eNOS knock-out mice showed either delayed
or abnormal meiotic maturation, we questioned the viability of these
oocytes. To identify dying oocytes, we looked for the presence of
fragmented DNA using the TUNEL method and fluorescent detection. All
oocytes were counterstained with PI (red fluorescence) to visualize
oocyte chromosomes. Initially we stained oocytes of both genotypes that
were arrested at metaphase II and observed that although the maternal
chromosomes were negative for TUNEL staining, the chromosomes of the
first polar body showed strong positive staining. Figure 6A shows the positive (bright
green/yellow fluorescence) staining of the polar body chromosomes and
the negative staining (red fluorescence) of the maternal chromosomes of
mature oocytes from an eNOS knock-out mouse (data not shown for
WT).
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, B–F). Figure 6, B and C, represents
oocytes with two polar bodies, both of which contain chromosomes
showing strong positive staining for TUNEL. Furthermore, maternal
chromosomes in these oocytes also show positive staining. In addition,
we observed that many oocytes from eNOS knock-out mice that were
arrested in metaphase I were also dying, and thus would never have
progressed through meiosis (Fig. 6D). Positive controls included
oocytes incubated with DNase I that showed strong positive staining in
both maternal and polar body chromosomes (Fig. 6E). All negative
control slides incubated without TdT showed no staining (Fig. 6F). Although TUNEL staining does not distinguish between apoptotic and necrotic cell death, these data indicate that a lack of NO during meiotic maturation results in abnormal oocyte development and premature oocyte cell death, possibly due to apoptosis.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Supporting a role for eNOS-derived NO in oocyte and thus follicular development is the oocyte-specific expression of eNOS both within the follicle as well as after ovulation. The positive staining for eNOS was observed not only in oocytes, but also in somatic cells in the follicles. Furthermore, a recent study in patients undergoing in vitro fertilization demonstrated that circulating nitrite/nitrate levels increase with follicular development (18, 39) and are correlated with follicular size (18). We have also reported previously that rats treated with NOS inhibitors during the periovulatory period show distinct abnormalities in oocyte development (21). Studies on oocyte growth have shown that the interactions between granulosa cells associated with the oocyte (cumulus cells) and the oocyte are essential for normal development of GV stage oocytes (40). For example, stage-specific patterns of protein phosphorylation in oocytes depend on interactions with granulosa cells, and the state of differentiation of granulosa cells affects patterns of kinase activity in oocytes (41). Conversely, oocytes have a paracrine influence on granulosa cell development by producing a factor that enables cumulus granulosa cells to undergo mucification, or expansion, in response to FSH (42). Thus, the relationship between the NO pathway and follicular growth could be necessary for both oocyte and follicle development.
Several lines of evidence also support the role of NO in the ovulatory process. Recent studies demonstrated that inhibition of NOS with nonselective competitive inhibitors led to fewer ovulations in vivo and in vitro in rats and rabbits (1, 4, 7, 21, 37). In the present investigation, we demonstrated that deleting the gene for eNOS lowers ovarian weight and the number of ovulated oocytes compared with those in WT mice. The abundance of eNOS in thecal cell layers and stroma reported for rat ovaries (6, 20) and in the present study for mouse ovaries suggests that NO may function as a vasodilator promoting the increased ovarian blood flow that is important for ovulation (4, 43). In addition, ovarian NO is important for optimum PG synthesis during the periovulatory period (7, 44). It is also likely that oocyte NO may function as a signal for somatic cells to properly modulate the theca and tunica albuginea of the follicle wall necessary for the oocyte to exit the ovary at ovulation.
The reduction in ovulation rate in eNOS-deficient mice compared with that in WT mice was also associated with a higher concentration of estradiol without significant changes in the plasma progesterone level. NO has been shown to inhibit aromatase activity directly in both human granulosa/lutein cells (14) and mature porcine granulosa cells (15), and has been suggested to function as a local modulator of follicular development. Similarly, a recent study demonstrated that inhibition of NOS in perfused rabbit ovaries resulted in an elevation of estradiol with no change in the progesterone level (7). It has also been demonstrated that NO negatively regulates steroidogenesis in human granulosa/lutein cells (13, 14), rat ovaries in vitro (4, 5), rodent testes (45), and cultured Leydig cells (46). Although the specific importance of estradiol to the rupture process is not currently understood, it is possible that estradiol functions to control blood pressure during the dramatic changes that occur in the ovulating ovary (47).
In addition to reduced ovulations, our study on meiotic maturation of oocytes from eNOS knock-out mice has shown that eNOS deficiency results in at least three obvious defects: fewer oocytes entered metaphase II, a greater percentage of oocytes remained in metaphase I, and significantly more oocytes showed dramatic atypical morphological changes. Furthermore, many oocytes that remained in metaphase I were found to die, possibly through apoptosis, which indicates that these oocytes would never have progressed through meiosis. The present report extends our previous study in rats in which inhibition of NOS/NO with pharmacological inhibitors impaired oocyte meiotic maturation (21). There are many mechanisms through which NO may affect oocyte meiotic maturation. Several studies have indicated that NO binds to and activates soluble guanylyl cyclase, and increases cGMP levels in target cells (48). cGMP has been localized in granulosa cells of the rat ovary and is involved in the resumption of meiosis in rat and hamster oocytes (49, 50, 51). It has been suggested that cGMP lowers the cAMP level by activating oocyte cAMP-phosphodiesterase (cAMP-PDE) and thus permitting oocyte maturation to continue (52). The evidence implicating cAMP, cGMP, and cAMP-PDE in meiotic maturation combined with the effect of NO on oocyte maturation raise the possibility that NO regulates oocyte cAMP level either by increasing cGMP synthesis and/or by stimulating cAMP-PDE activity during the transition from metaphase I to metaphase II of meiosis.
Among the important structural alterations that occur during maturation are the microtubule reorganizations, including organelle aggregation during GVBD (53), meiotic spindle assembly (54, 55), and chromosome segregation (56). A variety of morphological atypical characteristics were noted with oocytes from eNOS-deficient mice. We observed oocytes with an unusually large first polar body, suggesting abnormal cell division, which may be due to a failure of the metaphase spindle to properly translocate to the cell surface. Another difference we observed was the presence of two polar bodies, which could be derived from cleavage of the first polar body or from premature release of the second polar body. Mature mouse oocytes remain in metaphase II until fertilization due to an elevated level of maturation-promoting factor, which is now identified as the protooncogene c-mos protein (Mos) (29, 31, 57, 58). Some of the abnormalities in meiotic maturation in eNOS knock-out mice are very similar to those observed in mice deficient of Mos (58, 59). For example, oocytes from Mos-deficient females produce an abnormally large polar body, which persists instead of degrading and sometimes undergoes an additional cleavage. Furthermore, these oocytes often fail to arrest at metaphase II and undergo parthenogenetic activation (59). Therefore, the evidence suggests that NO may be a key regulator of the signal transduction cascade that controls progression between metaphase I and metaphase II as well as the secondary arrest at metaphase II.
The major findings of the present study were the absence of oocyte-specific eNOS expression, the reduced number of ovulated oocytes, and the abnormal oocyte meiotic maturation observed in eNOS-deficient mice in vivo. These data suggest that ovarian/oocyte NO production plays an important role in ovulation and oocyte meiotic maturation. When oocyte eNOS is lacking as in eNOS knock-out mice, ovulation and oocyte maturation are impaired. Further studies using an in vitro system of oocyte maturation will be necessary to understand more fully the mechanism underlying defects in ovulation and meiotic maturation in mice lacking the eNOS gene.
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Acknowledgments |
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Footnotes |
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Received December 29, 1997.
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B. Lu, C. Poirier, T. Gaspar, C. Gratzke, W. Harrison, D. Busija, M. M. Matzuk, K.-E. Andersson, P. A. Overbeek, and C. E. Bishop A Mutation in the Inner Mitochondrial Membrane Peptidase 2-Like Gene (Immp2l) Affects Mitochondrial Function and Impairs Fertility in Mice Biol Reprod, April 1, 2008; 78(4): 601 - 610. [Abstract] [Full Text] [PDF]
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A. T Grazul-Bilska, C. Navanukraw, M. L. Johnson, D. A Arnold, L. P Reynolds, and D. A Redmer Expression of endothelial nitric oxide synthase in the ovine ovary throughout the estrous cycle. Reproduction, October 1, 2006; 132(4): 579 - 587. [Abstract] [Full Text] [PDF]
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K.R. Barnett, C. Schilling, C.R. Greenfeld, D. Tomic, and J.A. Flaws Ovarian follicle development and transgenic mouse models Hum. Reprod. Update, September 1, 2006; 12(5): 537 - 555. [Abstract] [Full Text] [PDF]
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L.-J. Huo, C.-G. Liang, L.-Z. Yu, Z.-S. Zhong, Z.-M. Yang, H.-Y. Fan, D.-Y. Chen, and Q.-Y. Sun Inducible nitric oxide synthase-derived nitric oxide regulates germinal vesicle breakdown and first polar body emission in the mouse oocyte Reproduction, April 1, 2005; 129(4): 403 - 409. [Abstract] [Full Text] [PDF]
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 L. M. Mitchell, C.R. Kennedy, and G. M. Hartshorne Pharmacological manipulation of nitric oxide levels in mouse follicle cultures demonstrates key role of extrafollicular control of ovulation Hum. Reprod., August 1, 2004; 19(8): 1705 - 1712. [Abstract] [Full Text] [PDF]
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R.C. Manser, H.J. Leese, and F.D. Houghton Effect of Inhibiting Nitric Oxide Production on Mouse Preimplantation Embryo Development and Metabolism Biol Reprod, August 1, 2004; 71(2): 528 - 533. [Abstract] [Full Text] [PDF]
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 A. K. Nath, J. Enciso, M. Kuniyasu, X.-Y. Hao, J. A. Madri, and E. Pinter Nitric oxide modulates murine yolk sac vasculogenesis and rescues glucose induced vasculopathy Development, May 15, 2004; 131(10): 2485 - 2496. [Abstract] [Full Text] [PDF]
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L. M. Mitchell, C.R. Kennedy, and G. M. Hartshorne Expression of nitric oxide synthase and effect of substrate manipulation of the nitric oxide pathway in mouse ovarian follicles Hum. Reprod., January 1, 2004; 19(1): 30 - 40. [Abstract] [Full Text] [PDF]
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S. K. Sharan, A. Pyle, V. Coppola, J. Babus, S. Swaminathan, J. Benedict, D. Swing, B. K. Martin, L. Tessarollo, J. P. Evans, et al. BRCA2 deficiency in mice leads to meiotic impairment and infertility Development, January 1, 2004; 131(1): 131 - 142. [Abstract] [Full Text] [PDF]
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H. Yang, G. K. Bhat, R. Wadley, K. L. Wright, B. M. Chung, J. A. Whittaker, A. M. Dharmarajan, and R. Sridaran Gonadotropin-Releasing Hormone-Agonist Inhibits Synthesis of Nitric Oxide and Steroidogenesis by Luteal Cells in the Pregnant Rat Biol Reprod, June 1, 2003; 68(6): 2222 - 2231. [Abstract] [Full Text] [PDF]
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 C. Tamanini, G. Basini, F. Grasselli, and M. Tirelli Nitric oxide and the ovary J Anim Sci, February 1, 2003; 81(14_suppl_2): E1 - 7. [Abstract] [Full Text] [PDF]
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Y. Nakamura, Y. Yamagata, N. Sugino, H. Takayama, and H. Kato Nitric Oxide Inhibits Oocyte Meiotic Maturation Biol Reprod, November 1, 2002; 67(5): 1588 - 1592. [Abstract] [Full Text] [PDF]
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K. Mitsube, U. Zackrisson, and M. Brannstrom Nitric oxide regulates ovarian blood flow in the rat during the periovulatory period Hum. Reprod., October 1, 2002; 17(10): 2509 - 2516. [Abstract] [Full Text] [PDF]
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R. Gyurko, S. Leupen, and P. L. Huang Deletion of Exon 6 of the Neuronal Nitric Oxide Synthase Gene in Mice Results in Hypogonadism and Infertility Endocrinology, July 1, 2002; 143(7): 2767 - 2774. [Abstract] [Full Text] [PDF]
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C. M. Komar and T. E. Curry Jr Localization and Expression of Messenger RNAs for the Peroxisome Proliferator-Activated Receptors in Ovarian Tissue from Naturally Cycling and Pseudopregnant Rats Biol Reprod, May 1, 2002; 66(5): 1531 - 1539. [Abstract] [Full Text]
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C. M. Komar, O. Braissant, W. Wahli, and T. E. Curry Jr. Expression and Localization of PPARs in the Rat Ovary During Follicular Development and the Periovulatory Period Endocrinology, November 1, 2001; 142(11): 4831 - 4838. [Abstract] [Full Text] [PDF]
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W. J. Murdoch, R. S. Townsend, and A. C. McDonnel Ovulation-Induced DNA Damage in Ovarian Surface Epithelial Cells of Ewes: Prospective Regulatory Mechanisms of Repair/Survival and Apoptosis Biol Reprod, November 1, 2001; 65(5): 1417 - 1424. [Abstract] [Full Text] [PDF]
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F. J. Richard, A. Tsafriri, and M. Conti Role of Phosphodiesterase Type 3A in Rat Oocyte Maturation Biol Reprod, November 1, 2001; 65(5): 1444 - 1451. [Abstract] [Full Text] [PDF]
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C. Knauf, S. Ferreira, M. Hamdane, C. Mailliot, V. Prevot, J.-C. Beauvillain, and D. Croix Variation of Endothelial Nitric Oxide Synthase Synthesis in the Median Eminence during the Rat Estrous Cycle: An Additional Argument for the Implication of Vascular Blood Vessel in the Control of GnRH Release Endocrinology, October 1, 2001; 142(10): 4288 - 4294. [Abstract] [Full Text] [PDF]
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E. Ekerhovd, A. Enskog, K. Caidahl, N. Klintland, L. Nilsson, M. Brannstrom, and A. Norstrom Plasma concentrations of nitrate during the menstrual cycle, ovarian stimulation and ovarian hyperstimulation syndrome Hum. Reprod., July 1, 2001; 16(7): 1334 - 1339. [Abstract] [Full Text] [PDF]
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C. Knauf, V. Prevot, G. B. Stefano, G. Mortreux, J.-C. Beauvillain, and D. Croix Evidence for a Spontaneous Nitric Oxide Release from the Rat Median Eminence: Influence on Gonadotropin-Releasing Hormone Release Endocrinology, June 1, 2001; 142(6): 2343 - 2350. [Abstract] [Full Text] [PDF]
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M.-A. Ledingham, A. J. Thomson, A. Young, L. M. Macara, I. A. Greer, and J. E. Norman Changes in the expression of nitric oxide synthase in the human uterine cervix during pregnancy and parturition Mol. Hum. Reprod., November 1, 2000; 6(11): 1041 - 1048. [Abstract] [Full Text] [PDF]
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H. Matsumi, T. Yano, Y. Osuga, K. Kugu, X. Tang, J. P. Xu, N. Yano, Y. Kurashima, T. Ogura, O. Tsutsumi, et al. Regulation of Nitric Oxide Synthase to Promote Cytostasis in Ovarian Follicular Development Biol Reprod, July 1, 2000; 63(1): 141 - 146. [Abstract] [Full Text]
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K. Mitsube, M. Mikuni, M. Matousek, and M. Brannstrom Effects of a nitric oxide donor and nitric oxide synthase inhibitors on luteinizing hormone-induced ovulation in the ex-vivo perfused rat ovary Hum. Reprod., October 1, 1999; 14(10): 2537 - 2543. [Abstract] [Full Text] [PDF]
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A. Jablonka-Shariff, S. Ravi, A. N. Beltsos, L. L. Murphy, and L. M. Olson Abnormal Estrous Cyclicity after Disruption of Endothelial and Inducible Nitric Oxide Synthase in Mice Biol Reprod, July 1, 1999; 61(1): 171 - 177. [Abstract] [Full Text]
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G. I. Perez, X.-J. Tao, and J. L. Tilly Fragmentation and death (a.k.a. apoptosis) of ovulated oocytes Mol. Hum. Reprod., May 1, 1999; 5(5): 414 - 420. [Abstract] [Full Text] [PDF]
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 A. Jablonka-Shariff, R. Basuray, and L. M. Olson Inhibitors of Nitric Oxide Synthase Influence Oocyte Maturation in Rats Reproductive Sciences, March 1, 1999; 6(2): 95 - 101. [Abstract] [PDF]
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