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Developmental Biology

Cell cycle-coupled [Ca2+]i oscillations in mouse zygotes and function of

the inositol 1,4,5-trisphosphate receptor-1

Teru Jellerettea,1, Manabu Kurokawaa,1, Bora Leeb,1, Chris Malcuita, Sook-Young Yoon a,

Jeremy Smytha,b, Elke Vermassenc, Humbert De Smedtc, Jan B. Parysc, Rafael A. Fissorea,*

aDepartment of Veterinary and Animal Sciences, University of Massachusetts, Amherst, MA 01003, USAbProgram in Molecular and Cellular Biology, University of Massachusetts, Amherst, MA 01003, USA

cLaboratorium voor Fysiologie, Katholieke Universiteit Leuven, B-3000 Leuven, Belgium

Received for publication 24 October 2003, revised 12 June 2004, accepted 12 June 2004

Available online 31 July 2004

Abstract

Sperm entry in mammalian eggs initiates oscillations in the concentration of free calcium ([Ca2+]i). In mouse eggs, oscillations start at

metaphase II (MII) and conclude as the zygotes progress into interphase and commence pronuclear (PN) formation. The inositol 1,4,5-

trisphosphate receptor (IP3R-1), which underlies the oscillations, undergoes degradation during this transition, suggesting that one or more of

the eggs’ Ca2+-releasing machinery components may be regulated in a cell cycle-dependent manner, thereby coordinating [Ca2+]i responses

with the cell cycle. To ascertain the site(s) of interaction, we initiated oscillations at different stages of the cell cycle in zygotes with different

IP3R-1 mass. In addition to sperm, we used two other agonists: porcine sperm factor (pSF), which stimulates production of IP3, and

adenophostin A, a non-hydrolyzable analogue of IP3. None of the agonists tested induced oscillations at interphase, suggesting that neither

decreased IP3R-1 mass nor lack of production or excessive IP3 degradation can account for the insensitivity to IP3 at this stage. Moreover, the

releasable Ca2+ content of the stores did not change by interphase, but it did decrease by first mitosis. More importantly, experiments revealed

that IP3R-1 sensitivity and possibly IP3 binding were altered at interphase, and our data demonstrate stage-specific IP3R-1 phosphorylation by

M-phase kinases. Accordingly, increasing the activity of M-phase kinases restored the oscillatory-permissive state in zygotes. We therefore

propose that the restriction of oscillations in mouse zygotes to the metaphase stage may be coordinated at the level of IP3R-1 and that this

involves cell cycle stage-specific receptor phosphorylation.

D 2004 Elsevier Inc. All rights reserved.

Keywords: Calcium; Eggs; Fertilization; MAPK; MPF; Sperm; Sperm factor

Introduction

Long lasting intracellular calcium ([Ca2+]i) oscillations

occur in response to fertilization in eggs of all mammalian

species studied to date (Miyazaki et al., 1993; Stricker,

1999). The oscillations initiate, in a stepwise manner,

0012-1606/$ - see front matter D 2004 Elsevier Inc. All rights reserved.

doi:10.1016/j.ydbio.2004.06.020

* Corresponding author. Department of Veterinary and Animal

Sciences, University of Massachusetts Paige Laboratories, Room 411

Amherst, MA 01003. Fax: +1 413 545-6326.

E-mail address: rfissore@vasci.umass.edu (R.A. Fissore).1 These authors contributed equally to the completion of this work.

cellular and molecular events that result in cortical granule

exocytosis, exit from the metaphase II (MII) stage of

meiosis, and progression into interphase; these events are

collectively called egg activation (Schultz and Kopf,

1995). Following progression into interphase and pronu-

clear (PN) formation, zygotes proceed to the first mitosis,

after which recurrent mitotic cycles lead to embryo

development.

While the signaling mechanism whereby the sperm

initiates oscillations in mammalian eggs is not fully

elucidated (Runft et al., 2002), the channel and ligand

responsible for mediating Ca2+ release at fertilization are

274 (2004) 94–109

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109 95

known. Earlier studies have shown that injection of

inositol 1,4,5-trisphosphate (IP3) induces Ca2+ release in

mammalian eggs (Fissore et al., 1992; Miyazaki, 1988).

IP3, along with diacylglycerol, is the product of hydrolysis

of phosphatidylinositol 4,5-bisphosphate by phospholipase

C enzymes (Berridge et al., 1999). IP3 evokes Ca2+ release

by binding to the IP3 receptor (IP3R), a ligand-gated

channel localized mainly in the endoplasmic reticulum

(ER), the Ca2+ reservoir of the cell (Berridge, 2002;

Pozzan et al., 1994; Sousa et al., 1996). Of the three

known IP3R isoforms, mammalian eggs express prepon-

derantly, and abundantly, type 1 (Fissore et al., 1999; He et

al., 1999; Miyazaki et al., 1992; Parrington et al., 1998).

Immunolocalization studies have shown that the receptor is

homogeneously distributed in the ooplasm before matura-

tion, but acquires a more cortical localization at the MII

stage (Mehlmann et al., 1996; Shiraishi et al., 1995), the

stage at which eggs are ovulated and fertilized, and partly

disperses after fertilization (Goud et al., 1999). Fertilization

is also accompanied by a decline in IP3R-1 mass, the loss

of which depends on IP3 production (Brind et al., 2000;

Jellerette et al., 2000). Moreover, inhibition of IP3R-1 by

injection of a function-blocking antibody precludes the

initiation of oscillations and subsequent activation in

fertilized mouse eggs (Miyazaki et al., 1992), thereby

providing direct evidence for the predominant role of

IP3R-1 in the initiation of [Ca2+]i oscillations in mouse

eggs.

IP3-mediated and sperm-induced [Ca2+]i oscillations

occur preferentially in correspondence with the metaphase

stage of the cell cycle. For instance, sperm-initiated

oscillations cease as mouse zygotes progress toward

interphase, approximately at the time of PN formation

(Day et al., 2000; Jones et al., 1995a; Marangos et al.,

2003). Likewise, injection of variable concentrations of IP3,

or bovine sperm factor (bSF), which evokes production of

IP3, induces oscillations if administered at MII but is unable

to do so at interphase (Jones and Whittingham, 1996; Tang

et al., 2000). Progression into interphase and PN formation

are made possible by the simultaneous inactivation of the

meiotic kinases cyclin-dependent kinase 1 (cdk1 or MPF)

and mitogen-activated protein kinase (MAPK), which are

responsible for oocyte maturation and MII arrest (for

reviews, see Abrieu et al., 2001; Ferrell, 1999; Whitaker,

1996). Interestingly, preventing the inactivation of cdk1 and

exit of MII extends the sperm-initiated oscillations in

ascidian and mouse eggs (Levasseur and McDougall,

2000; Nixon et al., 2000). Moreover, the organization of

the ER is also influenced by the activity of cdk1 (FitzHarris

et al., 2003; Stricker and Smythe, 2003; Terasaki et al.,

2001), as the ER reportedly undergoes restructuring

following exit from MII (Kline et al., 1999). Nonetheless,

which of the aforementioned cellular or molecular changes

caused by egg activation underlies the metaphase depend-

ence of oscillations and whether the IP3R-1 itself is the

target of cell cycle regulation have yet to be elucidated.

An additional, although far less clear, association of

oscillations with first mitosis has been suggested (Kono et

al., 1996; Marangos et al., 2003). While a few reports have

shown that sperm-initiated oscillations resume at the time of

first mitosis and that oscillations can be induced at this stage

by injection of bSF (Tang et al., 2000), several other studies

have demonstrated that spontaneous oscillations at this stage

are not observed (Day et al., 2000; Gordo et al., 2002; Tang

et al., 2000; Tombes et al., 1992). Moreover, it is not

altogether clear whether, if present, the oscillations observed

at first mitosis are comparable to those observed at MII.

In the present study, we have examined factors that may

entrain the cell cycle and [Ca2+]i oscillations in mouse eggs.

We have considered and analyzed whether the number of

intact IP3R-1, the amount of releasable Ca2+ in intracellular

stores, IP3 production and degradation, or IP3R-1 modifi-

cations other than degradation may synchronize these

signaling events. Our results show that while several

simultaneously occurring activation-dependent events may

subtly shape the pattern of sperm-initiated oscillations,

modifications in the regulation of IP3R-1 function, for

example, by protein kinases, are likely to play a pivotal role

in imparting the metaphase restriction of [Ca2+]i oscillations

in mouse zygotes.

Materials and methods

Egg or zygote recovery and culture

MII eggs or fertilized zygotes were collected from the

oviducts of 10- to 16-week-old CD-1 and B6D2F1 (C57BL/

6J � DBA/2J) female mice. Females were superstimulated

with an injection of 5 IU PMSG (Sigma, St. Louis, MO; all

chemicals from Sigma unless otherwise specified) and were

induced to ovulate 40–48 h later by injection of 5 IU hCG.

Cumulus masses were recovered 12–14 h post-hCG and

collected in a HEPES-buffered Tyrode-Lactate solution (TL-

Hepes) supplemented with 5% heat-treated fetal calf serum

(FCS; Gibco, Grand Island, NY). Cumulus cells were

removed by a brief incubation with 0.1% bovine testis

hyaluronidase in TL-Hepes. To obtain fertilized zygotes,

female mice were housed overnight with males immediately

after injection of hCG and, after assessing the presence of a

copulation plug, zygotes were collected as above. Eggs

showing no degeneration and extrusion of the first or second

polar body (fertilized zygotes) were transferred into 50-Aldrops of KSOM (Potassium Simplex Optimized Medium;

Specialty Media; Phillipsburg, NJ) under paraffin oil. Eggs

were incubated at 36.58C in a humidified atmosphere

containing 7% CO2.

Microinjection techniques and reagents

Microinjection procedures were carried out as previously

described (Wu et al., 1997). Briefly, eggs were micro-

T. Jellerette et al. / Developmental Biology 274 (2004) 94–10996

injected in 50-Al drops of TL-Hepes supplemented with

2.5% sucrose and 20% FCS using manipulators (Narishige,

Medical Systems Corp., Great Neck, NY) mounted on a

Nikon Diaphot microscope (Nikon, Inc., Garden City, NY).

Eggs were first injected with 0.5 mM Fura-2 dextran (fura-2

D, Molecular Probes, Eugene, OR), a fluorescent dye that

reports changes in [Ca2+]i. When required, eggs were

injected a second time with porcine sperm factor (pSF) or

adenophostin A, a non-hydrolyzable IP3R agonist (Takaha-

shi et al., 1994) (courtesy of Dr. K. Tanzawa, Sankyo Co,

Tokyo, Japan). The reagents were diluted in injection buffer

(IB), which consisted of 75 mM KCl and 20 mM HEPES,

pH 7.0. Reagents were loaded into glass micropipettes by

aspiration and were delivered into the ooplasm by pneu-

matic pressure (PLI-100 picoinjector, Harvard Apparatus,

Cambridge, MA).

Intracytoplasmic sperm injection (ICSI)

ICSI was carried out as previously described (Kimura

and Yanagimachi, 1995; Kurokawa and Fissore, 2003) using

the aforementioned microinjection set up. ICSI was per-

formed in flushing and holding medium (FHM; Specialty

Media) at room temperature (RT) exclusively on B6D2F1

eggs or zygotes. One part sperm suspension was mixed with

one part IB containing 12% polyvinylpyrrolidone (PVP,

M.W. 360 kDa). Sperm, which were collected from the

cauda epididymis from 12- to 24-week-old B6D2F1 males,

were delivered into the ooplasm using a piezo-driven

micropipette unit (Piezodrill; Burleigh Instruments Inc.,

Fisher, NY). In brief, a few piezo-pulses were applied to

puncture the egg plasma membrane following penetration of

the zona pellucida. Mouse sperm heads were separated from

tails by applying a few piezo-pulses to the sperm’s mid-

piece immediately before injection. The separation of heads

and tails in freshly ejaculated boar sperm was accomplished

by brief sonication (XL2020, Heat Systems Inc., Farm-

ingdale, NY).

Preparation of pSF

Cytosolic sperm fractions were prepared from boar

semen as described by Swann (1990) with modifications

(Wu et al., 2001). Semen samples were washed twice with

Dulbecco’s phosphate-buffered saline (DPBS), and the

sperm pellet was resuspended in IB supplemented with

500 AM EGTA, 10 mM glycerophosphate, 1 mM DTT, 200

AM PMSF, 10 Ag/ml pepstatin, and 10 Ag/ml leupeptin, pH

7.0. The sperm suspension, containing approximately 500–

1000 � 106 sperm/ml, was lysed by sonication using a small

probe at a setting of 3 for 10–15 min at 48C. The resulting

suspension was centrifuged twice at 10,000 � g, and the

supernatant collected both times. The supernatant was

centrifuged at 100,000 � g for 45 min at 48C, and the

clear supernatant collected as the cytosolic fraction. Ultra-

filtration membranes (Centricon-50, Amicon, Beverly, MA)

and IB were used to wash and concentrate the supernatants.

The extracts were then precipitated by the addition of a

saturated solution of ammonium sulfate to a final concen-

tration of 50%, followed by centrifugation at 10,000 � g for

15 min at 48C. The precipitates were stored at �808C and

dissolved or aliquoted at the time of use.

Parthenogenetic egg activation

Several parthenogenetic agents were used in these studies

to generate interphase (pronuclear; PN) and first mitosis

(post pronuclear, PPN) zygotes with different amounts of

intact IP3R-1 (Jellerette et al., 2000). The agents used were

ethanol (Et), SrCl2 (Sr), adenophostin A (Ad), and pSF.

Activation by ethanol was accomplished by exposure of 16–

17 h post-hCG MII eggs to 7% ethanol (v/v) for 5 min at

378C. Eggs activated by SrCl2 were exposed for 2 h to 10

mM SrCl2 prepared in Ca2+-free M-16 like medium. Egg

activation by 10 AM adenophostin A or 1 Ag/Al pSF was

accomplished by injection of these products. After com-

pletion of activation treatments, eggs were washed in TL-

Hepes, placed in drops of KSOM, incubated, and monitored

for signs of activation, such as extrusion of the second polar

body, PN formation, and cleavage. Thimerosal was prepared

freshly and used at a final concentration of 200 AM in

KSOM.

Fluorescence recordings and [Ca2+]i determinations

[Ca2+]i measurements were carried out as previously

described (Gordo et al., 2002; Jellerette et al., 2000). Eggs

were injected with fura-2 D or loaded with 1 AM fura-2

acetoxymethylester (AM) supplemented with 0.02% plur-

onic acid (Molecular Probes) for 20 min at RT. Ca2+

values were monitored using a Nikon Diaphot microscope

fitted for fluorescence measurements. [Ca2+]i concentra-

tions, Rmin and Rmax, were calculated according to

Grynkiewicz et al. (1985) and as described (Wu et al.,

1997). Eggs were individually monitored in 50-Al drops ofTL-Hepes supplemented with BSA (1 mg/ml) placed on a

glass coverslip sealed over an opening in the bottom of a

culture dish and covered with mineral oil. Single eggs

were monitored using a modified Phoscan 3.0 software

(Nikon) and fluorescence was averaged for the whole egg

using a photomultiplier. For experiments involving ICSI,

2–10 eggs were measured simultaneously using the

software Image 1/FL (Universal Imaging, Downington,

PA). Images were acquired using an SIT camera (Dage-

MTI, Michigan City, IN) coupled to an amplifier (Video

Scope International Ltd., Sterling, VA). [Ca2+]i values were

not calibrated in this system and are therefore reported as

the ratios of 340/380 nm fluorescence.

[Ca2+]i monitoring for MII eggs started soon after

recovery from the oviducts, 12–14 h post-hCG. [Ca2+]iresponses at interphase were monitored between 6 and 8 h

post-activation and exclusively in the presence of a PN.

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109 97

[Ca2+]i monitoring at first mitosis was conducted 16–18 h

post-activation and immediately after observing the dis-

appearance of the PN envelope. Fluorescence measurements

were taken every 4–16 s and extended for 10–120 min

according to the experiment. Eggs or zygotes were collected

at the same stages of the cell cycle, frozen in liquid N2,

stored at �808C, and used for Western blotting or kinase

assays.

Histone 1 and MBP kinase assays

The activities of MPF and MAP kinase in eggs and in

zygotes were assayed as described (Gordo et al., 2002).

Lysates from five eggs were mixed with a cocktail

consisting ATP, [g-32P]-ATP (Amersham, Arlington

Heights, IL, USA), and histone 1 (H1) as a substrate for

MPF and myelin basic protein (MBP) as a substrate for

MAPK. The reaction was allowed to proceed for 20 min at

RT and terminated by addition of an equal volume of 2�SDS sample buffer. Samples were boiled for 3 min, loaded

onto 15% SDS-polyacrylamide gels, and run at 200 V for 45

min. The gels were then fixed and phosphorylations of H1

and MBP were visualized by autoradiography using

DuPont’s Cronex intensifying screens overnight at �808C.Autoradiographs were scanned and quantified as described

below.

Western blot technique

To assess the levels of intact IP3R-1 protein, lysates

from 15 eggs were thawed and mixed with 15 Al of 2�SDS sample buffer (Laemmli, 1970) as previously

described (He et al., 1999; Jellerette et al., 2000). Samples

were boiled for 3 min to denature proteins, loaded onto a

4% SDS-polyacrylamide gel, and proteins were separated

by electrophoresis followed by transfer onto nitrocellulose

membranes (Micron Separations, Westboro, MA). The

membranes were blocked in PBS containing 0.1% Tween

(PBST) supplemented with 5% non-fat dry milk for 1.5 h

and incubated overnight with one of two rabbit polyclonal

antibodies (Rbt03, Rbt04) raised against the same C-

terminal region of IP3R-1 diluted in PBST (Parys et al.,

1995). The membranes were subsequently washed in

PBST followed by incubation for 1 h with a goat anti-

rabbit secondary antibody coupled to horseradish perox-

idase. Membranes were incubated for 1 min in chemilu-

minescence reagent (NEN Life Science Products, Boston,

MA) and developed according to manufacturer’s instruc-

tions. Pre-labeled molecular weight markers were run in

parallel to estimate the molecular weight of IP3R-1 bands.

The intensities of IP3R-1 bands were assessed using a

Kodak 440 Image Station (Rochester, NY) and plotted

using Sigma Plot (Jandel Scientific Software, San Rafael,

CA). The intensity of the IP3R-1 band from MII eggs was

arbitrarily given the value of 1 and values in other lanes

were expressed relative to this band from MII eggs. When

the anti-phospho-Ser/Thr-Pro, MPM-2 mouse monoclonal

antibody (Upstate, Lake Placid, NY) was used, 60–85 eggs

were loaded per lane. The membranes were probed with a

4 Ag/ml dilution of the antibody; the blotting procedure

was performed as per manufacturer instructions. These

membranes were subsequently stripped by incubation at

508C for 30 min in a stripping solution (62.5 mM Tris, 2%

SDS and 100 mM 2-beta mercaptoethanol) followed by

washing in PBST and reprobed with an anti IP3R-1

antibody.

Statistical analysis

Values from two or more experiments, performed on

different batches of eggs or zygotes, were used for

evaluation of statistical significance. Statistical compari-

sons of the intensity of IP3R-1 bands, kinase assays, and

[Ca2+]i parameters were performed using Student’s t test or

one-way ANOVA and, if differences were observed

between groups, comparisons between treatments were

carried out by applying the Tukey–Kramer test using the

JMP-IN software (SAS Institute, Cary, NC). Significance

was at P b 0.05.

Results

[Ca2+]i oscillations in the first embryonic cell cycle

The molecular mechanism(s) responsible for the termi-

nation of oscillations at interphase in mouse zygotes and

whether the inhibition is mediated upstream or downstream

of IP3R-1 have not been thoroughly investigated. Therefore,

we first ascertained whether the ability to initiate oscillations

by two long-acting agonists and sperm was affected by the

stage of the cell cycle. To accomplish this, we injected pSF,

which stimulates production of IP3 (Jones et al., 1998; Wu

et al., 2001), and adenosphostin A, a non-hydrolyzable IP3R

agonist, into MII eggs and zygotes obtained after in vivo

fertilization. As expected, both agonists initiated oscillations

at the MII stage (Figs. 1A and B, left panels), but

oscillations were precluded at interphase, although a normal

and immediate first rise was observed in all cases (Figs. 1A

and B, center panels). Interestingly, injection of pSF at first

mitosis did not initiate oscillations (Fig. 1A, right panel),

although a first rise was always observed, whereas injection

of adenophostin A evoked low amplitude [Ca2+]i oscilla-

tions (Fig. 1B, right panel). To demonstrate that the

observed responses represented physiological reactions,

we monitored [Ca2+]i responses induced by injection of a

mouse sperm at analogous cell cycle stages. ICSI initiated

normal oscillations in MII eggs (Fig. 1C, left panel), but it

failed to do so at interphase (Fig. 1C, center panel), and

induced low frequency oscillations in half of the first mitosis

zygotes (Fig. 1C, right panel); a delayed first rise was

always elicited. Together, these results show that the

Fig. 1. Initiation of IP3R1-mediated [Ca2+]i oscillations by agonists depends on the stage of the cell cycle in fertilized (Fert) mouse zygotes. MII eggs,

interphase (PN), or first mitosis (PPN) stage zygotes were injected with 1 Ag/Al pSF (A), 10 AM adenophostin A (B; Ad), or with a mouse sperm (C) at the

arrow. High-amplitude [Ca2+]i oscillations were only consistently detected in MII eggs (left panels, top to bottom). Oscillations were nearly completely blocked

in interphase stage zygotes (center panels), whereas at first mitosis adenophostin A initiated oscillations, but with lower amplitude, and the sperm evoked

oscillations in only half of the oocytes and with low frequency (inset shows a representative profile of oscillations). Data were collected from at least three

different replicates. The number of eggs used per experiment (n) is presented as a ratio with the total number of eggs denoted by the number in the denominator,

whereas the number in the numerator represents eggs that showed MII-like oscillations. The graph shown for each treatment is representative of the most

common response observed within treatment and cell cycle stage. The style of data presentation for number of eggs examined per figure and depiction of

representative graphs was maintained throughout the manuscript.

T. Jellerette et al. / Developmental Biology 274 (2004) 94–10998

agonists used in the study faithfully replicate responses by

the sperm, and demonstrate that the inability to initiate

oscillations at interphase, and most likely the cessation of

sperm-induced oscillations, are not dependent on changes in

the concentration of IP3 and, instead, may occur at, or

downstream of, the IP3R-1.

Differential degradation of IP3R-1 but similar inactivation

of meiotic kinases by parthenogenetic procedures

In as much as fertilized zygotes exhibit degradation of

IP3R-1, it is plausible that the number of intact IP3R-1 is

responsible, at least in part, for the decreased oscillatory

ability detected after meiosis exit. While previous studies

have examined a similar question (Day et al., 2000; Jones

et al, 1995a; Tang et al., 2000), a comprehensive

examination of [Ca2+]i responses to long-acting agonists

and to the sperm at different stages of the cell cycle with

concomitant evaluation of IP3R-1 mass has not been

conducted. Therefore, to assess such responses, we first

differentially downregulated the number of intact IP3R-1

(Jellerette et al., 2000). As shown in Figs. 2A and B, left

panels, activation with SrCl2 induced progression into

interphase without degradation of IP3R-1, as has also been

Fig. 2. Egg activation by a variety of parthenogenetic procedures or by fertilization generates zygotes with different numbers of intact IP3R-1, but with similar

activity of M-kinases. Western blots denoting the presence of intact IP3R-1 in whole cell lysates from 15 eggs or zygotes (A) collected at interphase between 6

and 8 h post-activation (PN, left panel) or 16–18 h post-activation at first mitosis (PPN, right panel). These experiments were repeated at least five different

times and quantification of the blots, which was performed as described in Materials and methods, is presented in the form of a bar graph (B; means F SEM).

Please note that the lower band recognized by the antibody in the right panel of A was not related to the IP3R-1, as it was not recognized by another antibody

raised against the same epitope (not shown). Analysis of MPF activity (Histone H1) or MAP kinase (MBP) was carried out in groups of five eggs at the PN

stage, left panel, or at the PPN stage, right panel, and zygotes were collected at the same time using the aforementioned activation procedures. The data are

summarized in bar graphs and presented as means F SEM (C). Bars with different letters in B and C represent values that are significantly different (P b 0.05)

from those obtained with MII eggs. SrCl2 (Sr), adenophostin A (Ad), fertilization (Fert).

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109 99

previously reported to occur following activation with

ethanol (Brind et al., 2000; Jellerette et al., 2000). What is

more, the amount of intact IP3R-1 remained stable

throughout the first cell cycle in these zygotes (Figs. 2A

and B, right panels). Conversely, activation by adenophos-

tin A resulted in degradation of the receptor by the time of

PN formation, with the number of intact receptors

remaining stably low for the rest of the first cell cycle,

as was also the case for fertilized zygotes (Figs. 2A and B,

left and right panels; Parrington et al., 1998). Importantly,

all the treatments had comparable effects on MPF and

MAP kinase activities (see also FitzHarris et al., 2003), for

they induced their inactivation at interphase and allowed

similar activation of MPF when zygotes proceeded to first

mitosis (Fig. 2C). Therefore, a cellular paradigm was

established to investigate whether the number of intact

IP3R-1 affects the ability to initiate oscillations at different

stages of the cell cycle.

IP3R-1 degradation does not underlie the decreased

sensitivity to oscillations after MII exit

With the above system in place, we injected pSF and

adenophostin A at different stages of the cell cycle and

monitored [Ca2+]i responses. Injection of pSF at interphase

failed to induce oscillations regardless of IP3R-1 numbers

(Fig. 3A). Likewise, adenophostin A failed to initiate

oscillations at interphase irrespective of IP3R-1 numbers

(Fig. 3B). Injection of pSF at first mitosis once again did not

initiate oscillations regardless of receptor mass (Fig. 3C),

whereas adenophostin A induced oscillations in mitotic

zygotes generated by either activation procedure, although

Fig. 3. The number of intact IP3R-1 is unlikely to be responsible for the termination of oscillations at interphase stage (PN) or for the partially regained

responsiveness at first mitosis (PPN). SrCl2 (Sr; high IP3R-1 mass) and adenophostin A (Ad; low IP3R-1 mass) zygotes were injected at the arrow with 1 Ag/AlpSF (A and C) or 10 AMAd (B and D) either at the PN stage (A–B) or at PPN (C–D). At interphase, the initiation of oscillations was inhibited for both agonists

regardless of activation treatment. On the contrary, at first mitosis, injection of pSF was unable to initiate oscillations regardless of activation treatment, whereas

Ad initiated oscillations for all activation treatments utilized, although with less consistency and lower amplitude than at MII. These data were collected from at

least three different replicates.

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109100

the oscillations exhibited significantly lower amplitude and

occurred in just over half of the injected zygotes (Fig. 3D).

Moreover, injection of a mouse sperm at analogous stages of

the cell cycle in SrCl2-activated zygotes was unable to

initiate oscillations at either stage of the cell cycle (Figs.

4A–C), while a delayed first rise was always observed. In

light of these results, it could be assumed that the apparent

inability of pSF and sperm to initiate oscillations at first

mitosis may be due to a failure to effectively induce

production of IP3, because adenophostin A, which initiated

[Ca2+]i rises at this stage, circumvents the need to produce

IP3. Nonetheless, this does not appear to be the case,

because although injection of 100 AM IP3, a concentration

capable of inducing oscillations in MII eggs (Fig. 4D),

initiated oscillations in mitotic zygotes, the oscillations

quickly ceased (Fig. 4E). Collectively, these results suggest

that decreased numbers of intact IP3R-1 are not responsible

for the inability of agonists to initiate oscillations at

Fig. 4. The inability of mouse sperm (ICSI) to initiate fertilization-like oscillations at interphase (PN) or at first mitosis (PPN) in non-fertilized zygotes (SrCl2;

Sr) cannot be ascribed to lack of IP3 production. ICSI at MII initiates periodic [Ca2+]i responses (A), whereas at interphase (PN) it evokes a single [Ca2+]i rise

(B). ICSI is also unable to initiate fertilization-like oscillations in the majority of first mitosis (PPN) zygotes (C; inset depicts a representative profile of one case

in which oscillations were detected). Injection of 100 AM IP3, which induces high-frequency, long-lasting [Ca2+]i oscillations at MII (D), is unable to produce

the same response at first mitosis (E). The [Ca2+]i responses were collected from at least three different replicates. The arrow indicates time of injection.

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109 101

interphase or for the decreased responsiveness observed at

first mitosis. Likewise, an impairment of the ability to

produce IP3 is unlikely to be the reason for the limited

oscillatory activity observed at first mitosis, but an accel-

eration of IP3 degradation cannot be excluded.

The size of the ionomycin-releasable Ca2+ pool decreases

during the first cell cycle

We next explored the possibility that the content of the

Ca2+ stores may decrease as eggs exit MII and that this may

explain, at least in part, the decreased oscillatory capacity of

zygotes at interphase and the lower amplitude of the rises

observed at first mitosis. Accordingly, eggs and zygotes

were exposed to ionomycin in Ca2+-free medium and the

amplitude and duration of the induced [Ca2+]i rises

compared. Our results show that regardless of activation

procedure, the magnitude of the ionophore-induced Ca2+

release did not decrease at interphase (Fig. 5A; P N 0.05). In

contrast, although the amplitude of the ionomycin-induced

rise was not greatly affected at first mitosis, the duration of

the rise was markedly decreased (Fig. 5B), suggesting that

the content of the stores changes at this stage regardless of

IP3R-1 mass. Comparable effects of the cell cycle on the

parameters of the first [Ca2+]i rise were observed after

injection of pSF or adenophostin A (data not shown).

Together, these results demonstrate that the Ca2+ content of

the stores cannot explain the decreased oscillatory capacity

at the interphase stage. It could, however, at least in part,

explain the lack of oscillations during first mitosis in

response to sperm or pSF injection, or the low amplitude

of the rises induced by adenophostin A at this stage.

IP3R-1 function changes during progression from MII to

interphase

Because the major impact on oscillatory activity is

observed at interphase and because at this stage the content

of the Ca2+ stores did not decrease, we focused on

investigating the mechanism(s) that might underlie the

inhibition of oscillatory activity at this stage. We chose to

use zygotes generated by exposure to SrCl2 and ethanol, for

these treatments do not promote IP3R-1 downregulation. We

first ascertained if these interphase zygotes were competent

to undergo oscillations in response to thimerosal, a thiol

oxidizing agent. Our results (Figs. 6A and B) confirmed

previous findings showing that thimerosal is able to initiate

oscillations at interphase (Day et al., 2000; Jones et al.,

1995a). Nonetheless, because thimerosal is not a physio-

logical agonist, we proceeded to inject a porcine sperm,

which is able to initiate higher frequency oscillations in

mouse MII eggs compared to mouse sperm (Kurokawa et

al., 2004; and Fig. 6C), presumably due to higher amounts

of sperm factor activity. Notably, porcine sperm evoked

Fig. 6. Interphase (PN) zygotes are competent to undergo oscillations.

Exposure to 200 AM thimerosal (TMS) induced comparable oscillations in

MII eggs (A) or in SrCl2 (Sr) PN stage zygotes (B). Likewise, although the

frequency of oscillations is greatly decreased in comparison to those at MII

(C), injection of a porcine sperm at PN stage induced oscillations (D).

Fig. 5. The ionomycin-releasable Ca2+ pool does not change greatly in

mouse zygotes at interphase (PN), but is decreased by the time of first

mitosis (PPN). Eggs and zygotes were exposed to 5 AM ionomycin in Ca2+-

free medium. The amplitude and duration of the rise elicited by the addition

of the ionophore at PN (A) and PPN (B) were recorded and statistically

compared. Neither the amplitude nor duration was compromised at the PN

stage (P N 0.05), but at PPN the duration was significantly decreased in

relation to MII eggs (P b 0.05). Data were collected from two different

replicates using at least five eggs per activation treatment and per cell cycle

stage. SrCl2 (Sr), adenophostin A (Ad), fertilization (Fert), ethanol (Et).

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109102

oscillations in interphase zygotes as well, albeit of slower

frequency and duration than those initiated at MII by the

same sperm (Fig. 6D). Collectively, these data demonstrate

that interphase zygotes possess the ability to oscillate;

nonetheless, in the presence of physiological concentrations

of agonists, the system is desensitized and unable to mount

oscillations.

The results thus far raised the possibility that the IP3R-1

itself may be one of the determining factors that restricts the

oscillatory ability at interphase. In this vein, previous

studies and our own results (data not shown) have

demonstrated that the sensitivity of IP3R-1 decreases during

progression to interphase (Day et al., 2000; Jones and

Whittingham, 1996). Accordingly, we first examined the

possibility that IP3R-1 may not release Ca2+ as effectively

because of decreased or limited IP3 binding. To address

this, we made use of the finding that degradation of IP3R-1

is naturally induced by ligand binding (Zhu et al., 1999).

Injection of a mouse sperm or pSF into ethanol or SrCl2-

activated zygotes was unable to induce receptor degradation

(Fig. 7B), while they were able to do so when injected at

the MII stage (Fig. 7A) and that, in the case of ethanol-

activated eggs, they managed to induce [Ca2+]i responses at

this stage (data not shown). Besides the obvious interpre-

tation that IP3 cannot bind its receptor, other possible

explanations are that at interphase the degradation machi-

nery or proteasome is not functional, or that despite

adequate binding, IP3 is unable to promote Ca2+ release

or degradation of the receptor. We first addressed the

adequacy of the proteasome by determining whether

injection of 10 AM adenophostin A, which has higher

affinity for the receptor than IP3 (Morris et al., 2002;

Takahashi et al., 1994), is capable of inducing receptor

degradation at this stage. As shown in Fig. 7C, injection of

adenophostin A successfully caused IP3R-1 degradation, as

did 200 AM thimerosal. Therefore, we examined the

remaining alternatives by asking whether IP3R-1 degrada-

tion could be induced by injection of porcine sperm, which

we showed initiate oscillations at this stage. The results

show that under these conditions IP3R-1 degradation at

interphase takes place (Fig. 8A), suggesting effective ligand

binding by the receptor. Nonetheless, the degree of receptor

degradation seemed less at MII, and this could be

interpreted to mean that a less efficient IP3 binding or

production occurs at this stage. To determine whether this

was the case, we ascertained whether the Ca2+-induced

Ca2+ release (CICR) response, which requires a sensitized

IP3R-1 by IP3, could be induced by injection of CaCl2 into

zygotes subjected to interphase ICSI with porcine (Fig. 8B)

or mouse sperm (Fig. 8C). The results reveal that a

sensitized CICR mechanism is prevalent in zygotes injected

with a sperm at interphase, despite the paucity of

oscillations at this stage, demonstrating persistent IP3R

sensitization.

Fig. 8. IP3R-1 degradation and CICR sensitization by sperm injection at

interphase. Injection of a porcine sperm (Por-ICSI) induced IP3R-1

degradation in MII and in SrCl2 (Sr)-generated zygotes (PN, A), as shown

by a representative Western blot; the arrow denotes the presence of the

IP3R-1 band. Samples for Westerns were collected 2 h post-ICSI. These

experiments were repeated twice and quantification is presented as

means F SEM. Bars with a superscript are different from the respective

untreated controls. Injection of a porcine sperm (B) or of a mouse sperm

(C) into Sr-generated zygotes sensitized the Ca2+-induced Ca2+-release

(CICR) response when tested by injection of 2 mM CaCl2 (right panels);

left panels (B and C) depict the negligible increase in [Ca2+]i induced by

the same injection in the absence of sperm injection. Injection of CaCl2occurred approximately 1 h after ICSI.

Fig. 7. Ligand binding to IP3R-1 and IP3R-1 degradation occurs in

interphase zygotes. Bar graphs (means F SEM) summarize quantification

of Western blotting experiments and show that eggs activated by SrCl2 (Sr)

and ethanol (Et) reach interphase (PN) without IP3R-1 degradation, while

injection of a mouse sperm (ICSI) or pSF at MII induced degradation of the

receptor (A). Interestingly, neither ICSI of a mouse sperm nor pSF induced

IP3R-1 degradation when delivered into interphase zygotes (B). Adeno-

phostin A (10 AM; Ad) and thimerosal (200 AM; TMS), however, induced

normal IP3R-1 degradation (C). Experiments were replicated at least three

different times. Bars (means F SEM) with a superscript are statistically

different from the MII group, which were used as the reference treatment

and given the value of 1.

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109 103

Increased cellular phosphorylation promotes oscillations in

mouse zygotes

Because increased activity of M-phase kinases parallels

the presence of oscillations in eggs, we investigated whether

widespread changes in the phosphorylation status of zygotes

could reestablish the oscillatory-permissive state. Accord-

ingly, we exposed interphase SrCl2 zygotes to okadaic acid

(OA), a broad-spectrum phosphatase inhibitor, and injected

pSF after 60 min, which was the time required for the PN

envelope to break down. Injection of pSF into OA-treated

zygotes induced high-frequency oscillations (Fig. 9B),

whereas control interphase zygotes failed to equally respond

to pSF (Fig. 9A). As expected, OA increased the activity of

MAPK and had negligible effects on MPF activity (Fig.

9C), the activation of which seemed dispensable under these

conditions, for pSF-initiated responses also occurred in the

presence of 20 Ag/ml cycloheximide (data not shown),

which prevents cyclin synthesis and activation of MPF

(Moos et al., 1995, 1996). A similar effect of phosphor-

ylation was evident in fertilized zygotes in which exit from

first mitosis was prevented by colcemid (Fig. 9D); under

Fig. 9. Increase in meiotic kinase activity restores the oscillatory capacity of interphase (PN) or first mitosis (PPN) zygotes. Injection of pSF into SrCl2 (Sr)-

interphase generated zygotes was unable to initiate oscillations (A), but it did initiate oscillations in zygotes that had been exposed to 10 AMokadaic acid (OA) for

60 min (B). OA induced rapid and strong activation of MAP kinase (MBP), whereas the activation of MPF (H1) was negligible (C). The generation of persistent

oscillations at first mitosis in response to mouse ICSI was aided by addition of 0.1 Ag/ml colcemid (D). Experiments were repeated at least two separate times.

Fig. 10. IP3R-1 is phosphorylated in mouse eggs. IP3R-1 immunoreactivity

to the MPM-2 antibody is detectable in MII eggs, but it was undetectable in

SrCl2(Sr)-activated interphase (PN) zygotes (upper panel; A); the same blot

was stripped and reprobed for IP3R-1 (lower panel; A). To further confirm

that MPM-2 reactivity derives from the IP3R-1, MII (control), Sr-exposed

(to mimic oscillations), and adenophostin A (Ad)-injected (to induce IP3R-1

degradation) eggs were maintained arrested at MII for 4 h by treatment with

0.1 Ag/ml colcemid (to preserve metaphase arrest and phosphorylation), and

samples collected for Western blots. MPM-2 reactivity (upper panel; right

lane) decreased concomitantly with decreased IP3-R1 reactivity (lower

panel, right lane) (B). As for A, the original blot was stripped and reprobed

with an anti-IP3R-1 antibody. Experiments were repeated at least four

separate times with consistent results.

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109104

these conditions, ICSI initiated oscillations in all zygotes. It

is noteworthy that the effects of OA and colcemid are

unlikely to be caused by redistribution of the IP3R-1,

because both OA (Lu et al., 2002) and colcemid have

negative effects on microtubular organization, and an intact

microtubular network appears required for IP3R-1 redis-

tribution (Mitsuyama and Sawai, 2001; Vermassen et al.,

2003). Other nonspecific effects of these compounds on the

cytoskeleton can also be discounted because SF- and ICSI-

initiated oscillations occur undisturbed in the presence of

colcemid or cytochalasin B, a microfilament inhibitor (Knott

et al., 2003; Kumakiri et al., 2003).

Given the evidence that phosphorylation can modify the

function or conductivity of IP3R (see deSouza et al., 2002;

Krizanova and Ondrias, 2004; Patel et al., 1999; Tang et al.,

2003), we tested whether IP3R-1 was differentially phos-

phorylated during the cell cycle in mouse eggs. Accord-

ingly, we used the phosphorylation-specific MPM-2

antibody that recognizes the phospho serine–threonine-

proline epitope, which is the consensus sequence phos-

phorylated by cdks and MAPK. MPM-2 immunoreactivity

to an approximately 270-kDa band supposed from stripping

and reprobing experiments to be the IP3R-1 was easily

detectable at the MII stage, whereas it was undetectable in

interphase zygotes (Fig. 10A, upper panel) in spite of

equivalent amounts of IP3R-1 in both groups of cells (lower

panel). To further confirm that the MPM-2 reactivity

reflected IP3R-1 phosphorylation, the IP3R-1 mass was

specifically downregulated by adenophostin A while eggs

were kept at MII by exposure to colcemid. MPM-2

immunoreactivity in these eggs was compared to that

observable in eggs with a full complement of IP3R-1, which

were activated by SrCl2 and similarly arrested at MII. As

expected, the decrease in IP3R-1 signal in the adenophostin

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109 105

A condition caused a concomitant decrease in MPM-2

reactivity (Fig. 10B), demonstrating that the MPM-2 signal

was due to IP3R-1. In total, the results show that the IP3R-1

is phosphorylated in a cell-cycle-specific manner and

suggests that such regulation may underlie the metaphase

predisposition of [Ca2+]i oscillations in mouse eggs.

Discussion

In the present manuscript we have investigated the

factors that restrict [Ca2+]i oscillations to metaphase-like

stages of the first embryonic cell cycle. The main findings of

our study are as follows: (1) the MII stage is susceptible to

the initiation of [Ca2+]i oscillations, whereas the interphase

stage is resistant and the oscillatory-permissive state is only

partially regained during first mitosis; (2) the downregula-

tion of IP3R-1 that accompanies sperm-triggered cell cycle

progression is not responsible for the development of

insensitivity to IP3-initiated oscillations; (3) the size of the

Ca2+ stores, although not the cause of cessation of

oscillations at interphase, may limit the presence and shape

the amplitude of responses at first mitosis; (4) the function

of IP3R-1 is impaired in interphase zygotes despite IP3binding; (5) IP3R-1 is phosphorylated in a cell cycle stage-

specific manner. We therefore conclude that functional

modifications of IP3R-1 by protein kinases associated with

the cell cycle may underlie, at least in part, the cell cycle

specificity of [Ca2+]i oscillations in mouse eggs.

Lack of oscillations at PN stage is caused downstream of

IP3 production

The cessation of fertilization-initiated oscillations at

interphase in mouse eggs was the first noted and most

striking manifestation of entrainment between cell cycle and

oscillations (Jones et al., 1995a); this association was also

noted in other reports (Fujiwara et al., 1993; Jones and

Whittingham, 1996; Mehlmann and Kline, 1994). In

contrast, discrepancies exist among publications in regards

to the mechanism(s) responsible for the termination of

oscillations and as to whether endogenous, sperm-initiated

oscillations resume at first mitosis (Day et al., 2000; Kono et

al., 1996; Tang et al., 2000; Tombes et al., 1992). Our

present results expand on previous observations by demon-

strating that the initiation of IP3R-1-mediated oscillations in

response to a variety of long acting agonists, including the

sperm, is inhibited at interphase/PN stage whereas at first

mitosis, while oscillations reportedly occur, their initiation

may depend on high agonist concentrations (Tang et al.,

2000) or may be facilitated by inadvertent mitotic arrest

induced by the experimental procedure (Day et al., 2000;

Kono et al, 1996). We can therefore conclude that MII and

first mitosis are not analogous in their abilities to support

oscillations. Furthermore, because we made use of adeno-

phostin A, which circumvents IP3 production and is non-

hydrolyzable, we can infer that decreased ambient levels of

IP3 are not responsible for the cessation or inability to

initiate persistent oscillations at the PN stage or at first

mitosis. Nevertheless, we cannot discount the possibility

that under physiological cell cycle transitions (fertilization),

decreased production or increased degradation of IP3 may

affect the persistence of the initiated oscillations (see

below). Lastly, the recently advanced notion that the

sequestration or relocalization to the PN of the sperm’s

putative Ca2+ active factor is responsible for the termination

of oscillations at interphase (Marangos et al., 2003) can be

excluded as playing a role in our study, because adeno-

phostin A is unlikely to undergo such redistribution. What is

more, PN accumulation of PLC~ (Saunders et al., 2002), thepresumed Ca2+ active molecule in the sperm, occurs after

the cessation of oscillations (Yoda et al., 2004). Thus, our

results point to a mechanism downstream of IP3 production

to entrain the cell cycle and [Ca2+]i oscillations.

To uncover the molecular component of the Ca2+-

releasing machinery affected by the cell cycle, we first

investigated whether downregulation of IP3R-1 in zygotes

limited the ability to initiate [Ca2+]i oscillations. Our

findings show that the number of intact IP3R-1 remained

stable after PN formation and that the propensity of

zygotes to initiate oscillations at interphase was not

affected by whether they exhibited degradation of IP3R-

1, indicating that cessation or decreased ability to initiate

oscillations in zygotes is unlikely to be solely the

consequence of decreased numbers of intact IP3R-1. Our

findings agree with the overall conclusions of two previous

studies (Jones and Whittingham, 1996; Tang et al., 2000),

although in neither of those reports was the status of IP3R-

1 mass evaluated. What is more, results in starfish oocytes

showing that the increase in IP3-induced Ca2+ release that

accompanies oocyte maturation is achieved without

changes in IP3R mass (Iwasaki et al., 2002) support the

notion that striking changes in IP3R-1 sensitivity may

occur posttranslationally.

We next ascertained whether the amount of ionomycin-

releasable Ca2+, that is, the Ca2+ content of the stores,

changes during the first cell cycle, because we noted that

the amplitude of adenophostin A-initiated oscillations was

greatly reduced at first mitosis. Our data show that the

amount of ionomycin-releasable Ca2+ did not decrease by

the time of interphase/PN formation, but it did so by the

time of first mitosis, and this could explain, at least in part,

the inability of injected sperm or pSF to initiate MII-like

oscillations at this stage. These results somewhat contradict

those from a recent report showing that the amount of

releasable Ca2+ at first mitosis was decreased in partheno-

genetic, but not fertilized zygotes, although oscillations

were only observed in parthenogenetic zygotes (Tang et

al., 2000). Nevertheless, in another study, in the few cases

that sperm-induced responses were observed at first

mitosis, the detected rises exhibited low amplitude (Day

et al., 2000). Collectively, these results suggest that neither

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109106

the number of intact IP3R-1 nor the Ca2+ content of the

stores can explain the cell-cycle dependence of oscillations

observed here, although it cannot be discounted that

refilling of the stores may play a role in the persistence

of the oscillations.

The function of IP3R-1 is altered in interphase mouse

zygotes

In mouse eggs, increased activities of MPF and MAPK

play an overwhelming role in cell cycle transitions, and

their activation and persistency of activity coincides with

elevated Ca2+ responsiveness and oscillatory susceptibility

(Fujiwara et al., 1993; Jones et al., 1995b; Mehlmann and

Kline, 1994; Nixon et al., 2000). While this association of

M-kinases and Ca2+ release has been noted in eggs of

several other species including ascidians (McDougall and

Levasseur, 1998), sea urchin (Deng and Shen, 2000),

Xenopus (Terasaki et al., 2001; Tokmakov et al., 2001),

starfish (Santella et al., 2003), and nemerteans (Stricker

and Smythe, 2003), the molecular mechanism(s) modified

by the kinases has not been established. One possibility is

that these kinases control the organization of the ER,

favoring the formation of clusters, the localization of

which appears to coincide with aggregation of IP3Rs. In

addition, these clusters form during maturation and achieve

optimal size and cortical localization near the time of

fertilization (Goud et al., 1999; Kume et al., 1997;

Mehlmann et al., 1996; Shiraishi et al., 1995; Terasaki et

al., 2001). It follows that IP3R-1 clustering in oocytes has

been suggested to increase receptor sensitivity, thereby

favoring the progression of Ca2+ waves (Terasaki et al.,

2001), and that in nemertean eggs the cessation of

oscillations coincides with cluster disappearance (Stricker

and Smythe, 2003). Interestingly, in the latter study,

prevention of cluster formation by administration of a

JNK inhibitor did not impede the generation of oscillations

by the sperm. Moreover, in mouse eggs, cortical ER

clusters disintegrate much earlier than the cessation of

oscillations (FitzHarris et al., 2003). Also, in histamine-

stimulated HeLa cells, the presence of bpuffsQ, which are

the elementary IP3R Ca2+ signals upon which waves and

oscillations are built, does not correspond with IP3R

clustering (Thomas et al., 2000). Collectively, we interpret

this information to mean that the presence of ER/IP3R-1

clusters in eggs may be important for the initiation of

oscillations, but once oscillations are established, and the

system is sensitized by the presence of high ambient IP3concentrations, clusters become dispensable.

Alternatively, or concurrently, kinases may directly alter

the function of IP3R-1. Toward this end, we explored the

possibility that IP3 binding is compromised at interphase.

Our results showing that adenophostin A evokes IP3R-1

degradation, in conjunction with the demonstration of an

activated IP3R at this stage, primarily evidenced by the

presence of a sensitized CICR mechanism, suggest that a

functionally deficient IP3R-1, rather than impaired IP3binding, may control the development of the oscillation-

insensitive state at interphase.

The question then arises as to how the function of IP3R-1

is impaired at interphase. It is well documented that

binding of IP3 to its receptor provokes a large conforma-

tional change, which is thought to mediate channel gating

(Mignery and Sudhof, 1990). The same conformational

change is reportedly required for receptor ubiquitination

and degradation (Wojcikiewicz, 2004). Recent evidence

shows that both the N-terminal and C-terminal segments of

the receptor are critical for Ca2+ gating (Uchida et al.,

2003) and that these segments might interact during in

vivo conditions (Boehning and Joseph, 2000), although the

precise sequence of structural changes that brings about

IP3R-1 gating remains unresolved. Nonetheless, it is

possible that under the prevailing conditions at interphase,

the conformational change(s) or interaction of subunits

induced by agonist binding and required for channel gating

or degradation is disrupted. This presumptive mode of

action is analogous to that proposed for the IP3R-1

inhibitor 2-Aminoethoxydiphenyl borate (2-APB), which

abrogates IP3-induced Ca2+ release and receptor degrada-

tion without compromising IP3 binding (Soulsby and

Wojcikiewicz, 2002). Likewise, the observation that

thimerosal induces oscillations regardless of cell cycle

stage (Day et al., 2000; Jones et al., 1995a) further

supports the view of altered IP3R-1 function at interphase.

While the mechanism of action of thimerosal is not

known, it is presumed to do so by oxidizing sulfhydryl

groups in critical cysteine residues (Uchida et al., 2003); it

is plausible that thimerosal introduces wholesale changes

on the receptor that overcome the putative conformational

constrains imposed at interphase. We therefore conclude

that the function or conductivity of IP3R-1 is negatively

impacted by progression into interphase, and that this is

responsible, at least in part, for the cessation of oscil-

lations. We cannot discount the possibility, however, that

decreased IP3 binding may also contribute to the termi-

nation of oscillations.

IP3R-1 phosphorylation may contribute to the cell-cycle

dependence of [Ca2+]i oscillations in zygotes

While the molecular mechanism whereby phosphoryla-

tion modulates IP3R-1 function was not definitively

determined in this study, the evidence that zygote exposure

to OA or colcemid reestablishes oscillatory capacity, along

with the demonstration of cell cycle-specific phosphoryla-

tion of IP3R-1, supports the argument that phosphorylation

of IP3R-1 plays a pivotal part in the entrainment of the cell

cycle and [Ca2+]i oscillations. IP3R-1 contains multiple

phosphorylation consensus sequences for cdks and MAP

kinases. Interestingly, both OA and colcemid stimulate the

activity of these kinases (Moos et al., 1995, 1996; Sablina

et al., 2001), and treatment of somatic cells with these, or

T. Jellerette et al. / Developmental Biology 274 (2004) 94–109 107

similar compounds, produces IP3R-1 phosphorylation

(Joseph and Ryan, 1993; Malathi et al., 2003). Importantly,

in the latter manuscript, two candidate sites, one within the

ligand binding domain and the other within the regulatory

domain of the receptor, were shown to be phosphorylated

by cdc/cyclin B in cells arrested at the mitosis stage,

further supporting the view that phosphorylation may

modulate IP3R-1 function during the cell cycle. Additional

receptor regulation by phosphorylation cannot be dis-

counted, as the IP3R-1 possesses phosphorylation con-

sensus sites for kinases such as PKA, PKC, PKG, and

CaMKII (for reviews, see Krizanova and Ondrias, 2004;

Patel et al., 1999), all of which have been shown to

phosphorylate the receptor in vitro and, in some cases,

with functional consequences (deSouza et al., 2002; Patel

et al., 1999; Tang et al., 2003). Nonetheless, whether these

kinases are active during cell cycle transitions in mouse

zygotes is not known. We acknowledge that OA may

modify IP3R-1 phosphorylation by these kinases as well.

Importantly, additional phosphorylations may occur in

ancillary proteins, such as the group of CaBP (Nadif

Kasri et al., 2003; Yang et al., 2002), or in proteins

involved in Ca2+ homeostasis (Lim et al., 2003), and the

impact of these modifications on oscillations in mouse

zygotes has not been evaluated. Finally, two recent reports,

one using ascidian (Levasseur and McDougall, 2003) and

the other using mouse eggs (Marangos et al., 2003),

examined the interaction of meiotic kinases and [Ca2+]ioscillations. The results in ascidian eggs suggest a

complementary role for cdk1 on IP3R-1 whereas, in the

other manuscript, evidence is presented that suggests that

MPF and MAPK activities may not be required for

persistence of oscillations, because kinase inactivation,

concomitant with inhibition of PN formation, did not

preclude the termination of oscillations in fertilized

zygotes. However, it was not made clear whether the

target proteins modified by these kinases, including the

IP3R-1, were dephosphorylated after the decay in kinase

activity, leaving open the possibility that IP3R may have

achieved a kinase-dependent permissive state before the

decrease in kinase function, and that such a state might not

have been immediately reversed in the presence of used

treatments.

In conclusion, our results show that [Ca2+]i oscillations

and the cell cycle are entrained in mouse eggs and that

IP3R-1 is phosphorylated in a cell cycle-dependent

manner. We therefore propose that although a variety of

factors upstream and downstream of the IP3R-1 may

contribute to regulate the amplitude, periodicity, and

persistence of oscillations, functional modifications of

the IP3R-1 possibly mediated by meiotic kinases play a

pivotal role in confining the oscillations to the metaphase

stage. Future studies should elucidate the kinase(s) that

phosphorylates IP3R-1 during the cell cycle, the site(s) of

phosphorylation, and how these modifications alter IP3R-1

function.

Acknowledgments

The authors acknowledge the technical support of Ms.

Changli He. These studies were supported in part by grants

from the USDA (02-2078), USDA/Hatch, and NIH-RO3 to

R.A.F., a Lalor Foundation fellowship to M.K., and grants

8/99 and 7/04 of the Concerted Actions from KULeuven to

H.D.S. and J.B.P.

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