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Xenopus oocyte prophase I meiotic arrest is released independently from a decrease in
cAMP levels or PKA activity
Nancy Nader, Raphael Courjaret, Maya Dib, Rashmi P. Kulkarni and Khaled Machaca*
Department of Physiology and Biophysics, Weill Cornell Medicine Qatar, Education City –
Qatar Foundation, Doha, Qatar
Keywords: progesterone, oocyte maturation, cAMP, Xenopus, PKA
* Correspondence:
Khaled Machaca
Weill Cornell Medicine Qatar
Education City - Qatar Foundation– Doha, Qatar
E-mail: khm2002@qatar-med.cornell.edu
Tel: +1-646-797-3222
Fax: +974-4492-8422
Summary Statement: Progesterone releases Xenopus oocyte meiotic arrest without any reduction in
cAMP levels or PKA activity.
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http://dev.biologists.org/lookup/doi/10.1242/dev.136168Access the most recent version at
Development Advance Online Articles. First posted online on 27 April 2016 as 10.1242/dev.136168
ABSTRACT
Vertebrate oocytes arrest at prophase of meiosis I due to high levels of cAMP and PKA
activity. In Xenopus progesterone is believed to release meiotic arrest by inhibiting adenylate
cyclase, lowering cAMP levels, and repressing protein kinase A (PKA). However the exact
timing and extent of cAMP decrease is unclear with conflicting reports in the literature.
Using various in vivo reporters for cAMP and PKA at the single cell level in real time, we fail
to detect any significant changes in cAMP or PKA in response to progesterone. More
interestingly, there was no correlation between the levels of PKA inhibition and the release of
meiotic arrest. Furthermore, we devised condition where meiotic arrest could be released in
the presence of sustained high levels of cAMP. Consistently, lowering endogenous cAMP
levels by over 65% for prolonged time periods failed to induce spontaneous maturation.
These results argue that the release of oocyte meiotic arrest in Xenopus occurs independently
from lowering either cAMP levels or PKA activity, but rather through a parallel cAMP-PKA-
independent pathway.
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INTRODUCTION
Full-grown vertebrate oocytes arrest in a G2-like state at prophase of meiosis I for prolonged
periods of time during which the oocyte grows and stores macromolecular components
needed for future development (Smith, 1989; Voronina and Wessel, 2003). Before ovulation,
oocytes resume meiosis and arrest in metaphase of meiosis II, in a process termed ‘oocyte
maturation’. Oocyte maturation encompasses drastic cellular remodeling, in line with meiosis
progression, which prepares the egg for fertilization. This is characterized by the dissolution
of the nuclear envelope (referred to as germinal vesicle breakdown (GVBD), extrusion of the
first polar body and chromosome condensation (Bement and Capco, 1990; Nader et al., 2013;
Sadler and Maller, 1985; Smith, 1989; Voronina and Wessel, 2003).
Progesterone (P4) is the classical steroid used to mature Xenopus oocytes in vitro. Though a
role for the classical nuclear steroid receptor cannot be ruled out (Bayaa et al., 2000; Tian et
al., 2000), P4-induced maturation is thought to be achieved primarily through a non-classical
plasma membrane P4 receptor (mPR) (Josefsberg Ben-Yehoshua et al., 2007). It is
generally accepted that, upon hormone binding, mPR inhibits adenylate cyclase (AC)
causing a transient dip in cAMP levels, which is believed to act as the trigger to release
oocyte meiotic arrest by lowering protein kinase A (PKA) activity (Cicirelli and Smith, 1985;
Maller et al., 1979).
There is broad consensus that meiotic arrest in vertebrate oocytes is maintained by high levels
of cAMP-PKA, with the most compelling evidence coming from studies of the constitutively
active orphan GPCR, GRP3, in mouse oocytes (Freudzon et al., 2005; Mehlmann et al.,
2004). Knockout of GPR3 leads to premature oocyte maturation in antral follicles and
sterility. In Xenopus oocytes injection of the catalytic subunit of PKA (PKAc) inhibits P4-
induced oocyte maturation (Daar et al., 1993; Eyers et al., 2005; Matten et al., 1994).
Interestingly however even injection of a catalytically dead PKA mutant inhibits maturation
(Eyers et al., 2005; Schmitt and Nebreda, 2002), although it has been argued that this is due
to residual PKA activity (Eyers et al., 2005). Furthermore, phosphodiesterase (PDE)
inhibitors (Andersen et al., 1998; Han et al., 2006), and treatment with cholera toxin that
activates AC, repress maturation (Huchon et al., 1981; Mulner et al., 1979; Schorderet-
Slatkine et al., 1978). Inhibition of AC accelerated P4-induced oocyte maturation (Sadler et
al., 1986). Consistently, injection of PDE, the PKA regulatory subunit (PKAr), or the PKA
pseudo-substrate inhibitor (PKI), all release meiotic arrest (Andersen et al., 1998; Daar et al.,
1993; Eyers et al., 2005). However, others found that inhibition of PKA activity using H-89
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or PKI injection failed to stimulate oocyte maturation (Noh and Han, 1998). Furthermore,
induction of a drop in cAMP levels through stimulation of exogenously expressed Gi-coupled
serotonin receptor did not stimulate oocyte maturation (Noh and Han, 1998).
Whereas some studies detect a 10-60% drop in cAMP levels 15 seconds up to 6 hours after
P4 addition (Bravo et al., 1978; Cicirelli and Smith, 1985; Gelerstein et al., 1988; Maller et
al., 1979; Mulner et al., 1979; Nader et al., 2014; Sadler and Maller, 1981; Schorderet-
Slatkine et al., 1982), others failed to detect any changes in cAMP levels in response to P4
(Noh and Han, 1998; O'Connor and Smith, 1976; Schutter et al., 1975; Thibier et al., 1982).
Furthermore, some studies found that a decrease in cAMP levels was not sufficient to induce
spontaneous maturation, and that an increase in cAMP accelerates maturation (Gelerstein et
al., 1988; Noh and Han, 1998). Consistently, exposing oocytes to P4 for <30 min induced a
decrease in cAMP but did not induce oocyte maturation, which required >30 min incubation
with P4 (Nader et al., 2014). Hence there is strong support for the cAMP-PKA axis
maintaining the prolonged oocyte prophase I meiotic arrest. However, the evidence
supporting a drop of cAMP/PKA activity to release meiotic arrest is poor, with conflicting
reports regarding the timing and extent of the cAMP decrease in response to P4.
Here we use multiple approaches, including cAMP and PKA sensitive channels and FRET
sensors, to follow cAMP and PKA activity in individual oocytes in real time, and could not
detect any change during the induction of oocyte maturation. Unexpectedly, increasing or
buffering cAMP levels in the oocytes did not correlate with the ability of P4 to release
meiotic arrest. Based on these findings we proposed an alternative model where P4 stimulates
oocyte maturation independent of a decrease in cAMP-PKA.
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RESULTS
cAMP decrease in levels does not correlate with the release of meiotic arrest
We and others have previously shown a decrease in cAMP levels after P4 addition using an
ELISA-based assay in a batch of oocytes. However the early cAMP drop was not sufficient to
induce maturation, since exposure to P4 from 15 sec to 10 min was not sufficient to induce
oocyte maturation, rather oocytes required a minimum of 30 min exposure to P4 to induce
significant oocyte maturation (Nader et al., 2014). However, oocytes may require an
integrated sustained level of lower cAMP to inhibit PKA activity and relieve meiotic arrest.
To explore this possibility, we used the ELISA assay to measure cAMP levels in batches of
10 oocytes at time points ranging from seconds (Fig. 1B, left panel), up to 60 minutes (Fig.
1B, right panel) after P4 addition. Briefly, Oocytes were treated transiently with 10-5 M P4 or
ethanol as the carrier control for the indicated time points after which 10 oocytes were
collected for the cAMP-ELISA assay and the rest of the oocytes were washed twice with the
culture media (L15) to remove P4, and oocytes allowed to mature overnight.
When compared to oocytes treated with ethanol, cAMP levels showed a significant drop at 5,
15 sec, 1, 4, 6, 10, 12, 30 and 60 min after P4 (Fig. 1B and 1C). Interestingly, cAMP levels at
1 min and 30 min after P4 where not significantly different from each other (p=0.5071) (Fig.
1C), however oocytes failed to mature after a 1 min exposure to P4, yet achieved full
maturation after 30 min exposure to P4 (Fig. 1D). These results suggest that the cAMP dip
observed within 30 min of P4 treatment is not the trigger for maturation.
Membrane channels do not detect changes in cAMP or PKA in response to P4
An important limitation of the correlation between cAMP levels and maturation is that the
analysis was performed on a batch of 10 oocytes, as typically reported in the literature. This
is problematic because the early stages of oocyte maturation (before GVBD) are not
synchronized at the single oocyte level; rather individual oocytes reach the GVBD stage with
disparate time courses. Therefore, changes in cAMP could be more pronounced at the single
oocyte level but are averaged out in a cell population, which may account for the absence of a
correlation between the cAMP levels and maturation.
We therefore devised approaches to follow cAMP and PKA levels in individual oocytes in
real time after P4 treatment. For that purpose we used two membrane channels as sensors for
cAMP and its effector PKA: the Cyclic Nucleotide Gated channel (CNG), a cAMP-gated
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Ca2+ channel; and the Cystic Fibrosis Transmembrane Regulator (CFTR), a PKA-activated
Cl- channel (Biel and Michalakis, 2009; Hanrahan et al., 1996). Both channels have been
expressed and well characterized in Xenopus oocytes (Bear et al., 1991; Nache et al., 2012;
Webe et al., 2001; Young and Krougliak, 2004). A rise in cAMP gates CNG channels, which
permeate both mono- and divalent cations including Ca2+. Ca2+ in turn stimulates endogenous
Ca2+-Activated Chloride Channels (CaCC) thus amplifying the signal (Fig. 2A and 2B).
Expressing CNG in oocytes was associated with a large CaCC (Fig. 2B), indicating that the
channel is gated at resting cAMP levels. To test the sensitivity of CNG to changes in cAMP
we inhibited AC with 2’,5’-Dideoxyadenosine (DDA), to lower basal cAMP levels (Sadler
and Maller, 1983). DDA treatment significantly (p=0.021) decreased basal CNG-CaCC
current (ICNG-CaCC) (Fig. 2C), indicating that CNG effectively detects a drop in cAMP.
However, when CNG-expressing oocytes were treated with P4, we failed to detect any
significant differences in ICNG-CaCC, up to 30 min after P4 as compared to ethanol (Fig. 2D).
This suggests that basal cAMP levels, at least in the sub-plasma membrane domain detected
by the CNG channel, were unchanged up to 30 min after P4 treatment. However, the CNG
channel clone used is gated by both cGMP and cAMP (Young and Krougliak, 2004), so the
lack of response could be because the effect was masked by cGMP in the oocyte.
To test for changes in PKA activity we used the PKA-gated Cl- channel, CFTR, to follow
PKA activity in real time in the oocyte. In order to test whether CFTR can detect changes in
PKA activity, we injected db-cAMP to activate PKA, this showed a significant and
reproducible, although short lived, induction of the CFTR current (ICFTR) (Supplemental Fig.
1A-B). Inhibition of PKA with PKI under these condition, resulted in a significant decrease
of ICFTR (p=0.015) (Supplemental Fig. 1C and 1D). This shows that CFTR detects changes in
PKA activity in real time in the oocyte.
ICFTR showed significant run-down within a minute of establishing the voltage clamp (Fig.
2E), as previously reported (Button et al., 2001). To minimize ICFTR rundown, we pre-treated
oocytes with 3-isobutyl-1-methylxanthine (IBMX) (Ramu et al., 2007), a phosphodiesterase
inhibitor, to raise cAMP and activate PKA (Sadler and Maller, 1987; Schorderet-Slatkine and
Baulieu, 1982). IBMX pre-treatment significantly (p=0.0002) increased ICFTR, and limited its
rundown to maintain a significant current for up to 25 min (Fig. 2F). Though we cannot rule
out the possibility that IBMX, by inhibiting PDE might blunt changes in cAMP in response to
P4, we failed to detect any significant changes in ICFTR, as compared to the ethanol control,
for up to 30min after P4 treatment (Fig. 2G) in accordance with the CNG data. This argues
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that PKA activity, at least in the sub-plasma membrane compartment sampled by CFTR, does
not change significantly in response to P4. Collectively, membrane bound sensors did not
detect significant changes in cAMP and PKA in response to P4.
Intracellular FRET sensors do not detect changes in cAMP or PKA in response to P4
Since we failed to detect changes cAMP levels or PKA activity using membrane channels,
we needed to rule out the possibility of P4 affecting cAMP/PKA in sub-cellular
compartments that cannot be sampled by membrane channels. To do this we expressed two
widely used intracellular FRET-based sensors, TEPACVV and AKAR2, to monitor cAMP
levels and PKA activity, respectively (Klarenbeek and Jalink, 2014; Zaccolo et al., 2000).
TEPACVV employs the mTurquoise-YFP FRET pair, and has improved dynamic range as a
function of cAMP concentrations (Fig. 3A) (Klarenbeek et al., 2011), whereas the A kinase
activity reporter (AKAR2) FRET sensor uses Clover-mRuby2, and has a phospho-amino acid
binding domain, a defined docking site for recruiting PKA and a substrate domain, resulting
in increased FRET as a function of PKA activity (Fig. 3D) (Lam et al., 2012; Ni et al., 2006).
Confocal imaging using oocytes expressing TEPACVV and AKAR2 showed that both sensors
are diffusely cytoplasmically distributed (Fig. 3A and 3D). We then calibrated both sensors in
oocytes. To that end, TEPACVV-FRET efficiency and AKAR2-mRuby/Clover ratio change
(ΔFRET%) were measured following injection with db-cAMP (Fig. 3B and 3E). Given that
basal cAMP values in Xenopus oocytes averages ~1.25 pmole/oocyte with a range of 0.7-2.7
pmole/oocyte (Maller et al., 1979; O'Connor and Smith, 1976; Schutter et al., 1975), we
injected 0.2-0.8 pmole of db-cAMP, which represent 10-80% increase in cAMP levels.
Though db-cAMP is a cell permeable analog, we wanted to control the exact amounts of db-
cAMP delivered to the oocyte; hence we opted for injection rather than incubation.
The FRET efficiency of TEPACVV decreased significantly by ~40% (p=0.0094) after injection
of 0.8 pmole db-cAMP (Fig. 3B), and AKAR2-ΔFRET% increased significantly in a dose-
dependent fashion after db-cAMP injection (0.2-0.8 pmole) (Fig. 3E).
However, when oocytes expressing either sensor were treated with P4 for up to 30 min, we
were unable to detect any significant changes in cAMP or PKA levels when compared to
control ethanol-treated oocytes (Fig. 3C and F) though the expression of the FRET sensors
did not affect the GVBD response (data not shown). These results support the data from the
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membrane bound channel sensors, and argue that the release of meiotic arrest induced by P4
occurs independently of changes in cAMP or PKA.
PKA anchoring to AKAP is not essential for releasing meiotic arrest
Although we could not detect a repression of PKA using CFTR or AKAR2, we cannot rule
out the possibility that changes in PKA activity localize to sub-cellular microdomains. To
explore this possibility, we evaluated PKA catalytic subunit (PKAc) subcellular distribution
by immunostaining (Fig. 4A-C). The specificity of the PKAc antibody was confirmed by pre-
incubating the anti-PKA antibody with its peptide antigen (Fig 4A, right panel). PKAc
showed a diffuse cytoplasmic distribution with enrichment below the plasma membrane at
the animal, but not vegetal, pole of the oocyte (Fig. 4A and 4B). Line scan analyses confirm
PKAc enrichment in the sub-plasma membrane domain on the animal but not the vegetal pole
(Fig. 4C). To test whether sub-membrane PKA is specifically inhibited in response to P4, we
prepared membrane and cytosolic fractions by ultracentrifugation before and 30 min after P4
treatment, and tested PKA kinase activity (as described in Figure 5). The efficiency of the
biochemical separation of membrane and cytosolic fractions was confirmed by
immnoblotting for the plasma membrane protein, Na-K-pump (Fig. 4D). We failed to detect
any changes in PKA activity in either the cytosolic or membrane fractions 30 min after P4
treatment (Fig. 4E), or after normalizing for PKAc protein content in the two fractions (Fig.
4F).
To further test whether P4 treatment modulates PKA activity away from the sub-plasma
membrane space, we tested the role of A Kinase-Anchoring proteins (AKAP)s in oocyte
maturation. AKAPs are scaffold proteins that binds PKA regulatory subunit (PKAr), and
anchor it close to its physiological substrates (Colledge and Scott, 1999; Malbon, 2005;
Malbon et al., 2004). PKA interaction with AKAPs was found to be involved in mammalian
oocyte meiotic arrest (Brown et al., 2002; Kovo et al., 2006; Kovo et al., 2002; Newhall et
al., 2006; Nishimura et al., 2013). Injection of an AKAP inhibitory peptide that blocks
AKAP-PKAr interaction in mouse oocytes stimulates oocyte maturation in the presence of
high cAMP (Newhall et al., 2006). We therefore tested the effect blocking AKAP-PKA
interactions on Xenopus oocyte maturation. Oocytes injected with the AKAP inhibitory
peptide matured significantly (p=0.0349) faster as compared to oocytes injected with the
control peptide (Fig. 4G and 4H). However, interfering with AKAP-PKA interaction did not
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affect maximal GVBD levels achieved in response to P4 (Fig. 4H). These results argue that
PKA anchoring modulates the rate of P4-induced oocyte maturation without affecting its
extent or promoting maturation independently of P4.
PKA activity does not change following the release of meiotic arrest
To further monitor total PKA activity at the single oocyte level we used a quantitative
enzymatic PKA assay (Fig. 5A). We first validated the sensitivity of the assay by injecting
individual oocytes with the PKA inhibitor, PKI, or with PKA catalytic subunit (PKAc) (Fig.
5A). Control oocytes show a basal level of PKA activity at 37±4% phosphorylation of the
substrate peptide (Fig. 5A, lower panel). PKI injection lead to a significant (p=0.0274)
decrease in percent phosphorylated peptide to 16±1% (Fig. 5A). Consistently, PKAc injection
increased the percent phosphorylated peptide to 89±3%, (p0.0001) (Fig. 5A). We then
checked the effect of P4 treatment on PKA activity in single oocytes at 15 sec, 1, 2, 12 and 30
min after P4 and could not detect any change (Fig. 5A).
Concomitant with the PKA activity assay, we tested the effect of PKI and PKAc on oocyte
maturation. This was done at sub-threshold (10-8 M) and saturating (10-5 M) P4 concentrations
(Fig. 5B). Although, PKI injection stimulates oocyte maturation, it was significantly less
effective than 10-5M P4 (Fig. 5B). However, PKI significantly (p=0.0044) enhanced the rate
of P4-induced oocyte maturation (Fig. 5C and 5D), as previously reported (Noh and Han,
1998). The poor activation of oocyte maturation by PKI (Fig. 5B and 5C) is surprising
especially given the robust repression of PKA activity in the same batch of oocytes (Fig. 5A).
These data show a lack of correlation between PKA activity and the ability of PKI and P4 to
release meiotic arrest and induce oocyte maturation, hence bringing into question the
accepted view that the release of meiotic arrest is mediated by a drop in PKA activity. It is
worth noting here that previous studies using PKI to induce maturation in Xenopus oocytes,
including a study from our Lab, reported full maturation when compared to P4 treatment
(Maller and Krebs, 1977; Sun and Machaca, 2004). In contrast, microinjection of PKI did not
cause oocyte maturation, but accelerated GVBD in the presence of P4 in another study (Noh
and Han, 1998). The release of meiotic arrest is an asynchronous process in oocytes from the
same female and show significantly variability among different females based on whether
they are Lab bred, or captured in the wild and whether the females were hormonally
stimulated to test for their ability to produce oocytes. All these factors could affect the
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effectiveness of various stimuli to induce oocyte maturation. However, notwithstanding this
inherent variability, our results show a lack of correlation between the effectiveness of PKI
and P4 in inducing oocyte maturation and their ability to inhibit PKA activity in the same
batch of oocytes. PKI induces a robust inhibition of PKA activity (by 42.5%) with poor
maximal maturation rates (33+7.3%). In contrast, P4 treatment did not induce any detectable
change in PKA activity, yet fully induced oocyte maturation in the population (75+7.8%).
Note that even P4 in this batch of oocytes did not fully stimulate maturation, consistent with
the poor maturation levels with PKI.
Finally, PKAc injection abolished P4-induced maturation (Fig. 5B) as previously reported
(Daar et al., 1993; Matten et al., 1994), indicating that high PKA activity is sufficient to block
signals emanating from P4 stimulation that lead to oocyte maturation. Collectively our data
argue that the cAMP-PKA axis maintains oocyte meiotic arrest at steady state, but that it is
not modulated downstream of P4 to release meiotic arrest, suggesting a parallel pathway that
actively releases meiotic arrest despite constant high levels of cAMP/PKA.
Increasing cAMP levels does not affect the levels of oocyte maturation
The above data suggest that the cAMP-PKA axis acts as the ‘brake’ that maintains meiotic
arrest, but that the release of this meiotic arrest does not involve changes in cAMP-PKA. To
further assess this model, we tested the effects of the non-hydrolyzable cAMP analog, db-
cAMP, on the extent and kinetics of oocyte maturation. We used CNG-expressing oocytes to
determine the level of endogenous cAMP in the oocyte and the effectiveness of db-cAMP.
Young and Krougliak have previously characterized in Xenopus oocytes the cAMP dose
dependence of the recombinant CNG channel, x-fA4, we used here (Young and Krougliak,
2004). They showed a steep dose dependence of ICNG on cAMP concentrations with a K1/2 of
1 μM, and a saturating current above 200 μM cAMP (see Fig. 2C in Young and Krougliak,
2004). Consistently injecting oocytes with 0.8 or 400 pmole db-cAMP (which corresponds to
400 μM final cAMP concentration in the oocyte) dose-dependently increased ICNG-CaCC (Fig.
6A). Assuming a linear response of the CaCC to Ca2+ flowing through CNG channels, which
is reasonable given the large density of CaCC in the oocyte and their Ca2+ dependence
(Kuruma and Hartzell, 1998), we could translate ICNG-CaCC into cAMP concentration in the
oocyte using the Young and Krougliak dose response curve. We used the current induced by
400 pmole db-cAMP as the maximal current (Imax). With an I/Imax of 0.3 in control
oocytes, this translates to a cAMP concentration of 0.9 μM, within the range of 0.7-2.7
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μM/oocyte obtained from direct measurements of cAMP by others (Maller et al., 1979;
O'Connor and Smith, 1976; Schutter et al., 1975). This validates the use of I/Imax of ICNG-
CaCC to estimate oocyte cAMP concentration. Furthermore, ICNG-CaCC recordings show that the
injected db-cAMP is stable and results in a sustained increase in cAMP levels for extended
time periods. Injection of 0.8 and 400 pmole db-cAMP dose-dependently and significantly
(p<0.0022) repressed oocyte maturation stimulated with 10-7 M P4 (Fig. 6B). Surprisingly
however, when oocytes were stimulated with the saturating P4 concentration of 10-5 M for
only 1 hour, neither concentration of db-cAMP produced any inhibition of oocyte maturation
(Fig. 6C). Although maximal GVBD levels were not reduced following injection of 400
pmole db-cAMP, the time course of maturation was slowed down (Fig. 6D), requiring 6±0.62
hours to reach 50% GVBD as compared to 3±0.62 hours in control oocytes (p=0.0146) (Fig.
6E). These data indicate that increasing cAMP levels modulate the rate and extent of
maturation. However, increasing the P4-dependent signal overcomes the cAMP-mediated
inhibition and releases meiotic arrest independently from high sustained levels of intracellular
cAMP, of ~400 fold higher levels then resting cAMP concentrations.
Buffering endogenous cAMP doesn’t induce spontaneous maturation
To further test our hypothesis that P4 induces oocyte maturation through an alternative
pathway rather that reducing the levels of cAMP-PKA, we wanted to test whether a reduction
of endogenous cAMP levels is sufficient to induce oocyte maturation independently of P4.
The highest reported P4-dependent reduction in cAMP levels is 60% (Maller et al., 1979).
The FRET sensor TEPACVV binds cAMP and as such, when expressed at high concentrations
in the oocyte (Fig. 7A), would be expected to buffer endogenous cAMP. Expression of
TEPACVV decreased ICNG-CaCC by more than 10-fold (p=0.018), resulting in a 66.7% reduction
of cAMP levels from 0.9 μM to 0.3 μM (Fig. 7B). This reduction in cAMP is in line with the
maximal 60% reduction reported in response to P4 (Maller et al., 1979). Further attesting to
the effectiveness of TEPACVV to buffer cAMP, cells expressing TEPACVV attenuate the
stimulation of ICNG-CaCC following db-cAMP injection by 3-fold (p=0.0413) (Fig. 7B).
Remarkably though, endogenous cAMP buffering by TEPACVV did not release oocyte meiotic
arrest, nor it did enhance maximal GVBD when suboptimal or optimal concentration of P4
were used (Fig. 7C). Additionally, it did not accelerate the rate of oocyte maturation (Fig.
7D). These data support the conclusion that a dip in cAMP is not the trigger to release
Xenopus oocyte meiotic arrest.
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DISCUSSION
The current accepted model of releasing meiotic arrest in frog oocytes postulates a drop in
cAMP leading to inhibition PKA. Our results challenge this model and argue that P4 releases
meiotic arrest through a cAMP-PKA independent pathway. There is no correlation between
cAMP levels and oocyte maturation (Fig. 1). Furthermore, we could not detect any P4-
mediated changes in cAMP or PKA at the single oocyte level using CNG/CFTR channels
(Fig. 2), FRET sensors (Fig. 3) or a PKA kinase assay (Fig. 4 and 5). Despite these results,
we cannot rule out the possibility that the dip in cAMP and PKA is too small, too localized or
too transient to be detected. However several lines of evidence argue against the need for a
drop in cAMP/PKA to release meiotic arrest: 1) Injection of PKI inhibits PKA activity by
42.5% but results in poor maturation levels (Fig. 5). This argues that inhibition of PKA
activity is not sufficient to release meiotic arrest in the absence of a P4 dependent signal. As
discussed in the introduction whether PKA inhibition is sufficient to release meiotic arrest is
controversial with conflicting reports (Andersen et al., 1998; Daar et al., 1993; Eyers et al.,
2005; Noh and Han, 1998). This could be due to the priming state of the donor female as
discussed above. 2) Maintaining high sustained levels of cAMP inhibited oocyte maturation
but this inhibition was reversed by increasing P4 concentration (Fig. 6). A complete block of
maturation could be achieved only with injection of excess PKA catalytic subunit (Fig. 5). 3)
Buffering cAMP by over 65% was insufficient to induce maturation (Fig. 7).
Collectively, these results argue that meiotic arrest is released through a positive signal
downstream of the membrane P4 receptor (mPRβ), while the cAMP-PKA pathway acts a
“brake” (see model in Fig. 8). It is the balance between the ‘positive’ P4-dependent signal
and the ‘negative’ cAMP-PKA inhibitory signal that defines whether the oocyte commits to
maturation. PKA inhibits the maturation signaling pathway through at least two points:
mRNA translation and Cdc25C activation (Duckworth et al., 2002; Matten et al., 1994),
consistent with the idea that it is acting as a safety mechanism to eliminate spontaneous
maturation in the absence of a positive signal. Although challenging the accepted dogma, our
model is supported by earlier studies (Gelerstein et al., 1988; Nader et al., 2014; Noh and
Han, 1998; Schmitt and Nebreda, 2002), and the yin-yang balance between the mPRβ and
cAMP pathways effectively explains the discrepancies in the literature.
cAMP-PKA levels are maintained at high levels for the duration of the meiotic arrest through
the action of a constitutively active Gs-coupled G protein coupled receptor (GPCR) (Rios-
Cardona et al., 2008). GPR185 is the Xenopus homolog of mammalian GPR3, which has
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been shown to maintain meiotic arrest in mouse oocytes (Freudzon et al., 2005; Mehlmann et
al., 2002; Mehlmann et al., 2004; Norris et al., 2009). Surprisingly, knockdown of GPR185 is
not sufficient to induce oocyte maturation (Deng et al., 2008; Rios-Cardona et al., 2008),
arguing for other GPCRs involved in maintaining meiotic arrest. We have previously shown
that blocking exocytosis, while maintaining endocytosis, releases meiotic arrest, that P4
results in the internalization of GPR185, and that a GPR185 mutant that does not internalize
is more effective at maintaining meiotic arrest (El Jouni et al., 2007; Nader et al., 2014).
These data argue that P4 induces internalization of GPR185 and potentially other GPCRs that
are involved in maintaining high cAMP-PKA and as such meiotic arrest. Consistent with the
regulation of GPR185 in Xenopus, a drop in cAMP has been shown to correlate with the
release of meiotic arrest in mouse oocytes, through a cGMP signal from the surrounding
somatic cells (Norris et al., 2009). Furthermore, there is evidence in the mouse oocyte for a
positive signal through an EGF-like pathway downstream of LH stimulation (Ashkenazi et
al., 2005; Park et al., 2004), which may represent the counter part for the P4 signal in the
frog.
The mPRβ (also named PAQR8) has seven-transmembrane and belongs to the progestin and
adiponectin receptor family (PAQR) (Tang et al., 2005). It is still unclear whether mPRβ is a
GPCR (Moussatche and Lyons, 2012; Thomas et al., 2007). For example, mPRβ does not
work through Gi to inhibit AC since pertussis toxin, a specific Gi inhibitor, failed to block
P4-induced maturation (Mulner et al., 1985; Olate et al., 1984; Sadler et al., 1984). This is
consistent with the idea that mPR acts through other pathways to release the meiotic arrest.
mPRs and AdipoQ receptors can functionally couple to the same signal transduction pathway
in yeast, suggesting a common mechanism of action (Kupchak et al., 2007). Adiponectin
receptors signal through multiple pathways, including 1) the adaptor protein APPL1, which
links to the trafficking GTPase Rab5 (Buechler et al., 2010; Mao et al., 2006), or p38MAPK
(Heiker et al., 2010); and 2) possess alkaline ceramidase activity to generate sphingosine 1-
phosphate (S1P) (Moussatche and Lyons, 2012). S1P is known to modulate GPR3 activity,
the homolog of GPR185 in mammals (Uhlenbrock et al., 2002; Zhang et al., 2012), and
ceramide is a potential mediator of P4-induced maturation in Xenopus oocytes (Strum et al.,
1995).
In summary, our results challenge the generally accepted initial signaling pathway
downstream of P4, which assumes a drop in cAMP and repression of PKA activity to release
meiotic arrest. Rather, they argue for the existence of a positive signal downstream of mPRβ
Development • Advance article
to overcome the negative inhibitory signal from cAMP-PKA to release meiotic arrest. As
such in the future it would be of great interest to better define the mPRβ signaling pathway.
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MATERIALS AND METHODS
Molecular Biology. Human CFTR a gift from John Riordan (University of North Carolina)
was amplified using the following primers and subcloned in the Xenopus oocyte expression
vector pSGEM: 5’-CTGCAGGAATTCGATATGCAGAGGTCGCCTCTGGAAAAGGCC3’
(F) and 5’-ATCGATAAGCTTGATCTAAAGCCTTGTATCTTGCACCTCTTCTTC -3’
(R). TEPACVV (mTurq2Del-EPAC(dDEPCD)Q270E-tdcp173Venus(d)EPAC-SH187) a gift
from Kees Jalink (Netherlands Cancer Institute) (Klarenbeek et al., 2011), was subcloned into
the NotI-XbaI sites of pSGEM. The CNG channel chimeric clone X-fA4 was a gift from
Edgar Young (Simon Fraser University, Canada) (Young and Krougliak, 2004). AKAR2
(Lam et al., 2012) was purchased from addgene, and subcloned into the BamHI-EcoRI sites
of pSGEM. All constructs were verified by DNA sequencing and by analytical endonuclease
restriction digestion. mRNAs for all the clones were produced by in vitro transcription after
linearizing the vectors with NheI (CFTR, AKAR2), Not1 (CNG) or Sph1 (TEPACVV) using
the mMessage mMachine T7 kit (Ambion).
Xenopus oocytes. Stage VI Xenopus oocytes were obtained as previously described (Machaca
and Haun, 2002). The donor females were not hormonally stimulated prior to use. The
oocytes were used 24 to 72 hours after harvesting and digestion with collagenase to remove
follicular cells surrounding the oocytes. To study the role of AKAPs, cells were treated with
100 µM of AKAP St-Ht31 inhibitor peptide (Promega, Cat V8211) or its control peptide
(Promega, Cat V8221). Oocytes were injected with RNA and kept at 18°C for 1-2 days after
injection to allow for protein expression.
ELISA cAMP assay. We used the cAMP Complete Enzyme Immunometric Assay kit (Assay
Designs catalog no. 900-163). Ten oocytes were lysed by forcing them through a pipette tip,
in 250 μl of ice-cold 95% ethanol. Extracts were centrifuged at 15,000 g for 15 min at 4°C.
The supernatants were transferred to new tubes and dried under vacuum. The residue was
dissolved, and cAMP was measured according to the kit’s protocol.
CNG and CFTR recording. The ionic currents were recorded using standard two electrode
voltage-clamp recording technique. Recording electrodes were filled with 3 M KCl and
coupled to a Geneclamp 500B controlled with pClamp 10.5 (Axon instruments). The CNG
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currents were measured indirectly by monitoring the activation of endogenous Ca2+-activated
Chloride Channels (CaCC). Those chloride channels work as biological sensors and give a
very precise and amplified indication of the sub-membrane Ca2+ concentration. The currents
were recorded at a 0.1 Hz frequency using a previously described “triple jump” protocol that
allows the measure of Ca2+ influx (Courjaret and Machaca, 2014; Machaca and Hartzell,
1998). The CNG-CaCC current was measured as the difference in the amplitude of the
currents at +40 mV before and after a voltage pulse to -140 mV that increases the driving
force for Ca2+ entry (Fig. 2B). For CNG recordings using DDA and P4 the extracellular Ca2+
concentration was lowered to 0.9 mM to limit Ca2+ influx. The standard extracellular saline
contained (in mM) 96 NaCl, 2.5 KCl, 1.8 CaCl2, 2 MgCl2, 10 HEPES, pH 7.4. CFTR currents
were recorded at a steady state membrane potential of -80 mV. For CNG currents
normalization was done on the last current trace before treatment, for CFTR experiments
normalization was performed using the average current over a 1 min period before treatment.
TEPACVV and AKAR2 -FRET Imaging. Confocal imaging of live cells was performed using
a LSM710 (Zeiss, Germany) fitted with a Plan Apo 40x/1.3 oil immersion objective. Z-stacks
were taken in 0.45 µm sections using a 1 Airy unit pinhole aperture. When using TEPACVV,
FRET efficiency was calculated using the FRET acceptor bleaching technique. Briefly, this
was done by comparing donor fluorescence intensity in the same sample before and after
photobleaching the acceptor. If FRET was initially present, a resultant increase in donor
fluorescence will occur after photobleaching the acceptor, and the FRET efficiency can be
calculated as follow FRET efficiency = (Dpost-Dpre)/Dpost where Dpos is the fluorescence
intensity of the Donor after acceptor photobleaching, and Dpre the fluorescence intensity of
the Donor before acceptor photobleaching. To that end, TEPACVV expressing-oocytes animal
pole was imaged with the pinhole fully open. mTurquoise was excited at 458 nm and
emission detected at 462-520 nm, and YFP was photobleached at 514 nm and detected at
520-620 nm. YFP photobleaching was done by selecting at least 5 regions and FRET
efficiency calculated using ZEN 2008 (Zeiss) software. When using AKAR2 the excitation
was performed at 488 nm and emission detected at 495-560 nm (Clover) and at 588-702 nm
(mRuby). AKAR2 fluorescence was analyzed using ImageJ software (Schneider et al., 2012)
and the mRuby/Clover fluorescence ratio change (ΔFRET%) were calculated.
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PepTag assay. PKA kinase activity was measured in single oocyte using the PepTag® Non-
Radioactive Protein Kinase Assays from Promega according to the kit’s provided protocol.
Briefly, The PepTag® assay uses a highly specific fluorescent PKA peptide substrate that
when phosphorylated by changes the peptide’s net charge allowing easy electrophoretic
separation of the phosphorylated and non-phosphorylated peptide. This allows for
quantitative measurement of PKA catalytic activity in oocyte lysates. Phosphorylated and
non-phosphorylated bands were imaged using Geliance 600 Imaging system and the
intensities (I) of the bands were analyzed and corrected on the blank using Image J software
allowing calculation of % phosphorylation with correction on the negative control in the
absence of lysates or PKA.
Oocyte fractionation. Xenopus oocytes (
100) were lysed in Tris-HCL pH 8 (25 mM, EDTA
0.5 mM, EGTA 0.5 mM, protease inhibitor 1/100) using 5 µl per oocyte, followed by a spin
at 1000 g for 10 min. Supernatants were collected and spun for 1 hour at 150,000 g. The
supernatant was saved as the cytosolic fraction and the pellet (membrane fraction) was
dissolved with 50 µl PKA extraction buffer (freshly added to it 10 mM -mercaptoethanol,
protease inhibitor and PMSF). All spinning steps were done at 4C, and the tubes were kept
on ice during the whole procedures. The equivalent of 2 oocytes was used for the PepTag®
assay and also for the western blot to detect PKAc.
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Western blots. Cells were dounced in MPF lysis buffer (0.08M -glycerophosphate, 20mM
Hepes (pH 7.5), 15mM MgCl2, 20mM EGTA, 1mM Na-Vanadate, 50mM NaF, 1mM DTT,
1mM PMSF and 0.1% protease Inhibitor (Sigma)) and centrifuged twice at 1000 x g for 10
minutes at 4°C to remove yolk granules. The lysates were then incubated with 4% NP40 at
4°C for 2 hours followed by centrifugation at maximum speed for 15 minutes at 4°C and the
supernatant was stored. Supernatants were run on 4-12% SDS-PAGE gels, transferred to
polyvinylidene difluoride) (PVDF) membranes (Millipore), blocked for one hour at room
temperature with 5% Milk in TBS-T buffer (150 mM NaCl and 20 mM Tris; pH 7.6, 0.1%
Tween) and then incubated overnight at 4°C in 3% BSA in TBS-T with one of the following
primary antibodies: anti-PKAc (1/1000, sc-903, Santa Cruz), anti-GFP (1/1000, 2955S, Cell
Signaling), anti-Actin (1/10,000, A1978, Sigma), and anti-Na-K pump (1/1000, 3010S, Cell
Signaling). Blots were washed three times with TBS-T and probed for 1 hour with infrared
fluorescence, IRDye® 800 and 680 secondary antibodies (1/10,000) and the Western Blots
were revealed using the quantitative system LiCor Odyssey Clx Infrared Imaging system.
Oocytes immunostaining. Oocytes were fixed using 4% paraformaldehyde, washed with
phosphate buffered saline containing 30 % of sucrose, and incubated overnight in 50 % OCT
(WVR, clear frozen section compound) with gentle shaking. The oocytes were then
transferred to 100% OCT and frozen into a plastic mold prior to slicing to 7 µm sections on a
cryostat. For immunostaining the oocyte slices were first saturated with 3% BSA and 1 %
goat serum for 1 hour and incubated with anti-PKA antibody (sc-903, Santa Cruz) at a 1/50
dilution for 2 hours. For experiments using the blocking peptide (sc-903p) the primary
antibody was incubated overnight with the blocking peptide at a 5 times concentrations
according to manufacturer’s instructions. The secondary antibody was anti-rabbit IgG
coupled to and Ax488 fluorochrome (A11008, Fisher Scientific) to a 1/500 dilution for 2
hours. Imaging was performed on a Leica SP5 microscope controlled by Leica LAS software
using a 40x/1.3 lens and the pinhole slightly open to an optical slice of 1.3 µm.
Statistics. Values are given as means ± s.e.m. Statistical analysis was performed when
required using student paired and unpaired t-test. P values are indicated as follows: *
(p<0.05), ** (p<0.01), *** (p<0.001), and ns (not significant).
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Ethical approval. Animals were handled according to Weill Cornell Medicine College
IACUC approved procedures (protocol #2011-0035).
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ACKNOWLEDGEMENTS
We thank Dr Lu Sun for helping with the TEPACVV FRET confocal imaging and analysis,
and Drs John J. Riordan, Kees Jalink, Edgar C. Young for sharing the CFTR, TEPACVV and
CNG, clones respectively. We thank the Histology Core of Weill Cornell Medicine Qatar for
contributing to these studies. The Histology Core is supported by the BMRP program funded
by Qatar Foundation. This work was funded by NPRP 7-709-3-195 from the Qatar national
research fund (QNRF). The statements made herein are solely the responsibility of the
authors. Additional support for the authors comes from the Biomedical Research Program
(BMRP) at Weill Cornell Medical College in Qatar, a program funded by Qatar Foundation.
Competing Interests
The authors declare no competing interests
Author Contributions
NN designed and performed experiments, analyzed data and wrote the paper. RC designed
and performed experiments. MD and RPK performed experiments. KM developed the
concepts, analyzed data and wrote the paper.
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Figures
Figure 1. Lack of correlation between cAMP levels and oocyte maturation. A) Current
model by which P4 releases meiotic arrest. B) cAMP levels in oocytes before and seconds
(left panel) or minutes (right panel) after treatment with P4 (n=7) or ethanol (n=4). Data are
normalized to untreated oocytes. C) cAMP levels at 15 sec, 1 and 30 min after P4 treatment,
normalized to untreated oocytes. D) Percentage of oocytes reaching the GVBD stage
following P4 treatment at different time points (n=6). Oocytes were treated transiently with
10-5 M P4 for 15 sec, 1 min, 30 min and overnight (O/N) or with its carrier ethanol O/N.
After the exposure to P4 for 15 sec, 1 min and 30 min the oocytes were immediately washed
twice with the culture media L15. Oocytes were then left overnight and the following day, the
GVBD count was done. P4 (P4), PKA (Protein Kinase A), AC (Adenylate Cyclase), MPF
(Maturation Promoting Factor).
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Figure 2. CNG and CFTR do not detect changes in cAMP or PKA activity after P4. A)
Schematic representation of the regulation of CFTR, CNG and CaCC. B) Oocytes were
injected with CNG-RNA (10 ng/oocyte), and the CNG-CaCC current (ICNG-CaCC) (6 oocytes)
recorded at least 24 hours later as compared to Naïve oocytes (3 oocytes). C) CNG
expressing oocytes were treated with 0.64 mM DDA, and ICNG-CaCC recorded before and after
treatment (4 oocytes). Data are normalized to before treatment. D) CNG expressing oocytes
were treated with P4 (P4) (19 oocytes) or ethanol (EtOH) (18 oocytes) for up to 30 min and
ICNG-CaCC recorded continuously. Data are normalized to untreated oocytes. E) oocytes were
injected with CFTR-RNA (10 ng/oocyte) or water and the CFTR current (ICFTR) recorded at
least 48 hours later (9 oocytes). F) CFTR-expressing oocytes were pretreated with IBMX (1
mM) for 15 minutes and ICFTR recorded in IBMX for 25 min recording (no perfusion). Time
course of current run down (top panel) and current amplitude after 25 min (bottom panel) in
IBMX treated (IBMX, 16 oocytes) or untreated (Control, 7 oocytes) CFTR expressing
oocytes. G) Time course of ICFTR after P4 (6 oocytes) or EtOH (5 oocytes) addition in CFTR
expressing oocytes treated with IBMX.
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Figure 3. cAMP and PKA FRET sensors do not detect changes in response to P4. A and
D) Upper panels: Cartoon of the two FRET sensors TEPACVV (A) and AKAR2 (D) used for
cAMP and PKA respectively. Lower panels: Representative confocal images 48 hrs after
injecting TEPACVV (50ng/oocyte) or AKAR2 (10ng/oocyte) RNAs. B) FRET efficiency from
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TEPACVV expressing oocytes 10 min after db-cAMP (0.8 pmole) injection (3 oocytes). Data
are normalized to the FRET efficiency before db-cAMP injection. C) Time course of
TEPACVV FRET efficiency after P4 or ethanol treatment. E) ΔFRET changes in AKAR2 10
min after injection of increasing amounts of db-cAMP as indicated (3 oocytes). Data are
normalized to ΔFRET before treatment. F) Time course of AKAR2 ΔFRET changes in
response to P4 or ethanol. Data are normalized to before treatment.
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Figure 4. Subcellular PKA distribution and activity. A) Representative confocal images
from oocyte sections immunostained using an anti-PKAαc antibody in the absence (left
panel) or presence of blocking peptide (right panel). B) Bright field (BF) (left panel) and
fluorescence PKAαc immunostaining (right panel) of an oocyte at the equator showing both
the vegetal pole (VP) and the animal pole (AP). C) Zoomed focal planes of an oocyte at the
animal and vegetal poles showing PKA enrichment below the plasma membrane at the
animal pole (left panel). Average intensity of the immunostaining signal along linescans
across the plasma membrane as illustrated on the images in the left panels (right panel). D)
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Na-K-pump Western Blot (WB) after separation of the membrane and cytoplasmic fractions,
before and after P4 treatment. E) PKA kinase activity measured using the PepTag® assay in
whole lysates, membrane and cytoplasmic fractions before and after P4 treatment (n=3). F)
Cytosolic and membrane fractions were examined by western blotting using the PKAαc
antibody (top panel). Relative PKA activity corrected on the amount of PKA protein
quantified from WB (bottom panel). G) Representative GVBD-time course after P4 treatment
in the presence of AKAP inhibitor or control peptide. H) Oocytes were treated with P4 in the
presence of AKAP inhibitor or its control peptide. The time required for 50% of the oocytes
in the population to reach the GVBD (left y axis) and the maximum GVBD (right y axis) are
shown (n=6).
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Figure 5. P4 does not affect PKA kinase activity. A) Oocytes were injected with PKI (60
ng), PKAc (50 ng) or treated with P4 for different time points and PKA activity was
measured using the PepTag® assay. Representative agarose gels, showing the phosphorylated
and the non-phosphorylate peptide are presented on the top panel, and a quantification of
PKA as % phosphorylated peptide in naïve, PKI injected (n=3), PKAc injected (n=3), and
treated with P4 for different time points (n=4) are shown on the lower panel. B) Oocyte
maturation levels following: PKI injection, P4 treatments at the indicated concentrations in
the presence or absence of PKI (60 ng) or PKAc (50 ng) (minimum of n=3). The
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injections/treatments were done early in the morning and GVBD followed throughout the
day. C) Representative GVBD-time course after PKI injection alone and P4 treatment in the
presence or absence of PKI. D) The time required for 50% of the oocytes in the population to
reach the GVBD in oocytes treated with P4 alone or in the presence of PKI (n=3).
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Figure 6. High sustained cAMP slows down but does not block maturation. A) CNG
expressing oocytes were injected with 0.8 (5 oocytes) or 400 pmole (4 oocytes) db-cAMP
and ICNG-CaCC measured. B-C) Maturation levels following overnight P4 treatment at 10-7M
(n=7) (B), or for one hour at 10-5M (n=3) (C) in the presence or absence of 0.8 or 400 pmole
db-cAMP. D) Representative GVBD-time course after P4 (10-5M) in the presence or absence
of 400 pmole db-cAMP. E) Oocyte maturation rates in oocytes treated with P4 alone or in the
presence of 400 pmole db-cAMP (n=4).
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Figure 7. Buffering endogenous cAMP doesn’t induce spontaneous maturation. A)
Oocytes were injected with TEPACVV-RNA (10 ng/oocytes), and 48 hours later, Naïve and
TEPACVV–expressing oocytes were processed and TEPACVV expression was assessed by
western blot using GFP antibody. B) ICNG-CaCC measured in CNG-expressing oocytes in the
presence (4 oocytes) or absence of TEPACVV (5 oocytes), before or after injection of 0.8
pmole db-cAMP. C) Levels of oocyte maturation in the presence or absence of TEPACVV
and/or P4 (10-8 and 10-7M) (minimum of n=5) after overnight incubation. D) Representative
GVBD-time course after P4 treatment (10-7M ) in the presence or absence of TEPACVV. P4
was added in the morning and the time-course GVBD was followed throughout the day.
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Figure 8. Proposed model for the signal transduction cascade leading to the release of
Xenopus oocyte meiotic arrest. See text for details.
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